(Cyp8b1) Gene; an Alternative Mechanism for Bile Acid

Home , CYP8B1, Liver X receptor, Oxidoreductase

BHATT, ASMEEN, Ph.D., May, 2006 BIOMEDICAL SCIENCES

CYTOKINE REPRESSION OF THE HUMAN STEROL 12α-HYDROXYLASE

(CYP8B1) GENE; AN ALTERNATIVE MECHANISM FOR BILE ACID

SUPPRESSION OF CYP8B1 (228 pp.)

Director of Dissertation: John Y. L. Chiang, Ph.D.

Bile acids, synthesized from precursor cholesterol molecules, serve as an important mechanism for elimination of excess cholesterol from the body. Bile acid synthesis is tightly regulated since excess bile acids are toxic and can cause liver damage. Bile acids inhibit their own synthesis by inhibiting the bile acid biosynthetic genes. Cholesterol 7α- hydroxylase (CYP7A1) is the rate-limiting enzyme of classic bile acid biosynthesis, which synthesizes two primary bile acids, cholic acid (CA) and chenodeoxycholic acid

(CDCA) in the liver. Sterol 12α-hydroxylase (CYP8B1) catalyzes CA synthesis in the liver and is feedback inhibited by bile acids. In addition to activating farnesoid X receptor

(FXR, nuclear receptor subfamily 1H4, NR1H4), bile acids also induce the release of inflammatory cytokines, like interleukin 1β (IL-1β) from Kupffer cells in the liver. The objective of this study was to investigate the mechanism by which inflammatory cytokines inhibit human CYP8B1 gene transcription. Real time PCR assays revealed that both CDCA and IL-1β markedly reduced CYP8B1, CYP7A1, and nuclear receptor hepatocyte nuclear factor 4α (HNF4α, NR2A1) mRNA expression levels in human primary hepatocytes. However, CDCA induced, but IL-1β reduced the negative nuclear receptor, small heterodimer partner (SHP, NR0B2) mRNA expression. IL-1β inhibited human CYP8B1 reporter activity only in liver cells, and a c-Jun N-terminus kinase (JNK)-specific inhibitor blocked IL-1β inhibition. Activated JNK1 or c-Jun inhibited, whereas their dominant negative forms blocked IL-1β inhibition of CYP8B1 transcription. Mutagenesis analyses mapped an IL-1β response element to a previously identified bile acid response element, which contains an HNF4α binding site.

Furthermore, IL-1β inhibited HNF4α gene transcription, protein expression and binding to the CYP8B1 gene. JNK1 phosphorylated HNF4α and a JNK-specific inhibitor blocked the IL-1β inhibition of HNF4α expression. Expression of c-Jun, a downstream target of the JNK pathway, was induced by both IL-1β and CDCA in primary human hepatocytes. c-Jun inhibited the HNF4α and coactivator peroxisome proliferator-activated receptor γ co-activator-1α (PGC-1α) mediated trans-activation of CYP8B1 reporter activity. Co- immunoprecipitation revealed an interaction between c-Jun and HNF4α, which was confirmed by GST pull down assay. IL-1β increased the ratio of c-Jun or phosphorylated c-Jun bound to HNF4α in HepG2 cells. Chromatin immunoprecipitation (ChIP) assay revealed that c-Jun did not affect HNF4α binding but blocked HNF4α recruitment of

PGC-1α to the CYP8B1 chromatin. This study also showed that HNF4α stimulated the gene expression of SHP by binding to the promoter of the latter. The results suggest that

IL-1β inhibits CYP8B1 gene transcription, in a SHP-independent manner via the JNK pathway that inhibits HNF4α gene expression and its DNA-binding ability. IL-1β also induces c-Jun, which blocks HNF4α recruitment of PGC-1α to CYP8B1 chromatin. This mechanism may play an important role in the adaptive response to inflammatory cytokines and in the protection of the liver during cholestasis.

CYTOKINE REPRESSION OF THE HUMAN STEROL 12α-HYDROXYLASE

(CYP8B1) GENE; AN ALTERNATIVE MECHANISM FOR BILE ACID

SUPPRESSION OF CYP8B1

A dissertation submitted

to Kent State University in partial

fulfillment of the requirements for the

degree of Doctor of Philosophy

Asmeen Bhatt

May, 2006

Dissertation written by

Asmeen Bhatt

M.B.B.S., University of Mumbai, 2000

Ph.D. Kent State University, 2006

Approved by

Chair, Doctoral Dissertation Committee

John Chiang John Y. L. Chiang, Professor

Members, Doctoral Dissertation Committee

Philip Westerman Philip W. Westerman, Professor

James Hardwick James P. Hardwick, Associate Professor

Hans Folkesson Hans G. Folkesson, Associate Professor

Kathleen Doane Kathleen J. Doane, Associate Professor

Accepted by

Robert Dorman Director, School of Biomedical Sciences

John Stalvey Dean, College of Arts and Sciences

TABLE OF CONTENTS

LIST OF FIGURES...... v

LIST OF TABLES...... vii

ACKNOWLEDGEMENTS ...... viii

CHAPTER I: INTRODUCTION...... 1

1. Bile Acids...... 1 a. Structure ...... 2 b. Function ...... 5 c. Synthesis ...... 5 d. Mutations and deficiencies of important enzymes in bile acid biosynthesis...... 9 e. Enterohepatic circulation of bile ...... 13 2. Cholesterol Homeostasis in the liver ...... 19 3. Nuclear Hormone Receptors...... 23 a. Structure ...... 26 b. Response Elements ...... 26 c. Ligands...... 27 4. Important Nuclear Receptors involved in regulation of bile acid biosynthesis...... 28 a. Farnesoid X Receptor (FXR, NR1H4)...... 28 b. Pregnane X Receptor (PXR, NR1I2)...... 29 c. Vitamin D3 Receptor (VDR, NR1I1) ...... 30 d. Hepatocyte nuclear factor 4α (HNF4α, NR2Α1) ...... 31 e. Human α-fetoprotein transcription factor (hFTF, NR5A2) ...... 32 f. Small Heterodimer Partner (SHP, NR0B2)...... 33 g. Peroxisome Proliferator Activated Receptor α (PPARα, NR1C1) ...... 34 h. Liver X Receptor (LXR, NR1H3) ...... 35 5. Important Transcription factors and Coactivators involved in regulation of bile acid biosynthesis...... 36 a. Sterol Regulatory Element Binding Proteins (SREBPs)...... 36 b. c-Jun...... 38 c. Peroxisome Proliferator-Activated Receptor-γ Coactivator 1α (PGC-1α):...... 39 6. Regulation of Bile Acid Biosynthesis...... 40 a. Feedforward Regulation by Cholesterol...... 41 b. Feedback Regulation by Bile Acids...... 41 7. Regulation of CYP8B1 ...... 47 8. Mitogen Activated Protein Kinase (MAPK) Pathways ...... 50 9. Cholestatic liver disease...... 54 10. Proinflammatory cytokine mediated gene regulation in the liver...... 56

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11. Hypothesis, Specific aims, Approaches and Significance...... 60

Chapter II: MATERIALS AND METHODS ...... 64

1. Cell Culture...... 64 Cell line culture:...... 64 Primary cell culture:...... 65 2. Plasmids ...... 65 Reporters:...... 66 Expression Vectors: ...... 66 3. Preparation of Competent Cells...... 67 4. Bacterial Cell Transformation...... 68 5. Preparation of Plasmid DNA ...... 69 Large Scale preparation or Maxiprep: ...... 69 Small Scale DNA preparation or Miniprep: ...... 70 6. Transient Transfection Assay ...... 71 Lipid Based Transfection Method: ...... 72 Luciferase Assay:...... 76 β-Gal activity assay: ...... 76 7. RNA Extraction ...... 77 8. Reverse Transcriptase PCR...... 78 9. Quantitative Real Time PCR (Q-RT PCR)...... 80 SYBR Green Method:...... 80 Taqman Method:...... 84 Data Analysis:...... 86 10. Protein Extraction ...... 86 11. Immunoblot Analysis...... 87 12. Preparation of Nuclear Extracts and In vitro Translated Proteins ...... 88 Nuclear Extracts:...... 89 Invitro Translated Proteins:...... 90 13. Electrophoretic Mobility Shift Assay (EMSA) ...... 91 14. Chromatin immunoprecipitation assay (ChIP) ...... 95 15. GST fusion protein expression...... 100 16. GST Pull Down Assay ...... 101 18. Co-Immunoprecipitation Assay (Co-IP)...... 109 19. Small Interference RNA (siRNA)...... 114

CHAPTER III: RESULTS...... 119

CHAPTER IV: DISCUSSION...... 187

APPENDIX A: ABBREVIATIONS...... 201

REFERENCES...... 208

LIST OF FIGURES

Figure 1. Structure of Cholesterol and Bile Acids...... 4 Figure 2. Bile acid biosynthetic pathways in the liver...... 7 Figure 3. Enterohepatic circulation of bile...... 15 Figure 4. Cholesterol Homeostasis in the liver...... 22 Figure 5. Structure of nuclear hormone receptors...... 25 Figure 6. Bile acid feedback inhibition pathways...... 44 Figure 7. Activation of different MAPK signaling cascades by different extracellular stimuli...... 53 Figure 8. Transient transfection assay...... 75 Figure 9. Quantitative real time PCR (Q-RT PCR)...... 82 Figure 10. Electromobility shift assay (EMSA)...... 93 Figure 11. Chromatin immunoprecipitation (ChIP) assay...... 98 Figure 12. GST fusion protein expression and GST pull down assay...... 104 Figure 13. Invitro kinase assay...... 108 Figure 14. Coimmunoprecipitation (Co-IP) assay...... 113 Figure 15. Small interference RNA (siRNA)...... 117 Figure 16. Effect of IL-1β on the human CYP8B1 gene transcription...... 126 Figure 17. Dose Response and Time Course of IL-1β effect on human CYP8B1 gene transcription...... 129 Figure 18. Effect of MAPK inhibitors and JNK overexpression on the IL-1β inhibition of CYP8B1 reporter activity...... 132 Figure 19. Mapping the region responsive to IL-1β inhibition on the CYP8B1 gene. ... 135 Figure 20. Effect of HNF4α binding site mutation on the IL-1β response of CYP8B1 reporter activity...... 139 Figure 21. Effects of IL-1β on HNF4α stimulation of CYP8B1 reporter activity...... 142 Figure 22. Effects of IL-1β on HNF4α gene transcription and protein expression in HepG2 cells...... 144 Figure 23. Electrophoretic mobility shift assays and chromatin immunoprecipitation assays of the effect of IL-1β on HNF4α binding to the CYP8B1 gene...... 147 Figure 24. Immunoblot analysis of the effects of MAP kinase inhibitors on the IL-1β mediated inhibition of the HNF4α protein expression...... 150 Figure 25. Phosphorylation of HNF4α by JNK1 and the effect of JNK1 overexpression on the HNF4α binding/ recruitment to the CYP8B1 promoter...... 153 Figure 26. Effect of IL-1β and CDCA on the mRNA and protein expression of cJun. . 156 Figure 27. Effect of cJun, dncJun, PGC-1α and HNF4α cotransfection on human CYP8B1 promoter activity...... 159

Figure 28. Effect of IL-1β treatment on the interaction between HNF4α with cJun or phospho cJun in vivo and GST Pull down assay to study the interaction between HNF4α and cJun...... 163 Figure 29. Electromolibility shift assay and chromatin immunoprecipitation assays to study the effect of cJun on the HNF4α binding/ recruitment to the CYP8B1 promoter. 167 Figure 30. Effect of IL-1β and HNF4α on the human SHP gene transcription...... 172 Figure 31. Electromobility shift assay to show the binding of HNF4α to the human SHP gene...... 175 Figure 32. Effect of cJun, dncJun, PGC-1α and HNF4α cotransfection on human CYP7A1 promoter activity...... 178 Figure 33. Chromatin Immunoprecipitation assays to study the effect of IL-1β and CDCA treatment and pcDNA3, cJun or JNK1 overexpression on the HNF4α binding/ recruitment to the CYP7A1 promoter...... 181 Figure 34. Effect of HNF4α small interference RNA (siRNA) transfection with or without IL-1β or CDCA...... 184 Figure 35. Summary of data...... 200

LIST OF TABLES

Table 1. Effect of IL-1β on mRNA expression (dose response) in human primary hepatocytes (HH1088) determined by real time RT-PCR ...... 120 Table 2. Effect of CDCA on mRNA expression (dose response) in human primary hepatocytes (HH1165) as determined by real time RT-PCR...... 121 Table 3. Effect of IL-1β on mRNA expression (time course) in human primary hepatocytes (HH 1209) as determined by real time RT-PCR...... 122 Table 4. Effect of CDCA on mRNA expression (time course) in human primary hepatocytes (HH 1209) as determined by real time RT-PCR...... 124

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ACKNOWLEDGEMENTS

I would like to express my sincere graditute to my advisor, Dr. John Chiang, for his invaluable support, patience and encouragement throughout my graduate studies. Both his scientific insights and editorial skills helped me complete the work for this dissertation.

I also deeply appreciate the valuable time, comments and assistance provided by all my committee members: Dr. James Hardwick, Dr. Hans Folkesson, Dr. Phillip Westerman and Dr. Kathy Doane. Their guidance, scientific suggestions and careful review of my work helped improve this dissertation significantly.

My parents, Mesbah and Suraiya Jahan receive my deepest gratitude and love for always believing in me, standing by me and showing me the importance of a good education. I also want to thank my late grandfather, Dr. Iftekar Jahan, who has been the greatest inspiration in my life.

Last, but not least, I would like to thank my husband, Sandeep, for his understanding and love and all the sacrifices he made, through all these years. His encouragement and the never-ending discussions concerning my research have been the pillars of strength during my graduate work.

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CHAPTER I

INTRODUCTION

Bile acids are physiological detergents that mediate the uptake of dietary lipids and lipid soluble substances from the intestine [1-3]. Bile acids are synthesized from cholesterol in the liver and serve as an important mechanism for elimination of excess cholesterol from the body. Bile acid synthesis is a tightly regulated process since excess bile acids are toxic to the hepatocytes. Bile acids inhibit their own synthesis by inhibiting the genes encoding the bile acid biosynthetic enzymes [1-3]. It has been suggested that the hydrophobicity index of bile acids determines the potency of bile acid inhibition. Sterol

12α-hydroxylase (CYP8B1) is an important enzyme in the bile acid biosynthetic pathway that is known to be inhibited by bile acids [1-3]. CYP8B1 plays a critical role in regulating the ratio of the primary bile acids, cholic acid (CA) and chenodeoxycholic acid

(CDCA), and thus the hydrophobicity of bile. Bile acids induce cytokine release in the liver [4]. The objective of this dissertation is to study how inflammatory cytokines regulate human CYP8B1 gene transcription. This study will lead to a better understanding of the mechanisms by which bile acids inhibit CYP8B1.

1. Bile Acids

Bile acids are the end products of cholesterol catabolism. The amphipathic structure makes bile acids excellent detergents and signaling molecules. The structure and function

1 2

of bile acids as well as their biosynthetic pathways, the important genes involved in bile acid biosynthesis and the enterohepatic circulation of bile are discussed in this section.

a. Structure

Bile acids are classified as primary and secondary bile acids (Fig. 1). The primary bile acids synthesized in the liver are CA (3α, 7α, 12α) and CDCA (3α, 7α) [1-3]. The carboxyl groups in the side chains of these bile acids remain conjugated to taurine or glycine by amide links. The amidation of bile acids lowers the pKa < 3 and improves their solubility in the intestinal lumen. At physiological pH, bile acids are present as sodium salts and known as “bile salts”, which are negatively charged. Upon secretion into the intestine, a fraction of the CA and CDCA are converted to the secondary bile acids, deoxycholic acid (DCA) (3α, 12α) and lithocholic acid (LCA) (3α) respectively, by the 7α-dehydroxylase, present in the bacterial flora in the intestine.

The structure of bile acids is such that they have a hydrophobic surface separated from a hydrophilic surface by a rigid ring. This amphipathic nature of bile acids assists in the absorption and transport of lipids and lipid soluble vitamins. Bile acids act as lipid carriers as they are able to solubilize many lipids by forming micelles - aggregates of lipids such as fatty acids, cholesterol esters and monoglycerides - that remain suspended in water. The phospholipids, cholesterol esters, fatty acids and monoglycerides remain associated with the hydrophobic surface of bile acids, while the hydroxyl groups of the bile acids face the aqueous environment.

Fig.1.

Figure 1. Structure of Cholesterol and Bile Acids.

Cholesterol has saturated tetracyclic ring structures that are denoted as A, B, C and D,

with an aliphatic side chain at C17 of the D ring, methyl groups at C10 and C13, a double

bond in ring B and a hydroxyl group on C3. Bile acids are derived from cholesterol and

have similar structures with variations in side chains and ring modifications. The position

of the hydroxyl groups on the bile acids is indicated within the parentheses. The primary

bile acids are synthesized in the liver and have been hydroxylated at position 7 and/or 12.

The secondary bile acids are synthesized from the primary bile acids in the intestine, by

the 7α-dehydroxylase activity of the intestinal flora.

b. Function

Bile acids are physiological detergents that facilitate the absorption, transport and distribution of lipid soluble vitamins (vitamins A, D, E and K), dietary fats, cholesterol and steroids [1-3]. Recent studies have implicated bile acids as signaling molecules, which activate nuclear receptors and mitogen activated protein kinase (MAPK) signaling pathways and play critical roles in the regulation of lipid homeostasis. Bile acids also play roles in drug metabolism and detoxification of xenobiotics in the liver and intestine. c. Synthesis

The conversion of cholesterol to bile acids involves multiple hydroxylations of the ring structure and the oxidation and shortening of the side chain. Two major bile acid biosynthetic pathways exist in the liver. They are the “Classic or Neutral” pathway and the “Alternative or Acidic” pathway (Fig. 2). The enzymes catalyzing the reactions in each pathway are located in the endoplasmic reticulum, mitochondria, cytosol and peroxisomes [1-3]. i) Classic or Neutral Pathway: This pathway contains intermediates that are neutral steroids. It constitutes the major bile acid synthetic pathway (~ 80% of the total bile acid synthesis) in humans. In this pathway, the modifications of the steroid nucleus of cholesterol, precedes the oxidative cleavage of the three-carbon side chain. In humans, this pathway produces CA and CDCA in approximately equal amounts. The neutral pathway is initiated by the rate-limiting microsomal enzyme, cholesterol 7α-hydroxylase

(CYP7A1), which introduces a hydroxyl group at the C-7 position, forming 7α- hydroxycholesterol [5]. Then, 7α-hydroxycholesterol is converted to

Fig.2.

Figure 2. Bile acid biosynthetic pathways in the liver.

The two major bile acid synthetic pathways, namely the Classic (Neutral) and the

Alternative (Acidic) are shown. The major regulatory enzymes, along with their substrates and products are shown. The enzymes shown in this figure are: cholesterol 7α- hydroxylase (CYP7A1), sterol 12α-hydroxylase (CYP8B1), sterol 27-hydroxylase

(CYP27A1), oxysterol 7α-hydroxylase (CYP7B1) and 3β-hydroxy-C27-steroid

dehydrogenase/isomerase (3β-HSD).

5 7α-hydroxy-4-cholesten-3-one via a microsomal 3β-hydroxy-Δ -C27-steroid

oxidoreductase isomerase (3β-HSD). The 7α-hydroxy-4-cholesten-3-one may be

hydroxylated at the C-12 position by microsomal CYP8B1 to form 7α,

12α-dihydroxy-4cholestan-3-one [1-3], which is the precursor of CA or remain

unhydroxylated at the C-12 position to form CDCA. Other cytoplasmic enzymes then

convert these intermediates to 5β-cholestane-3α, 7α, 12α triol (for synthesis of CA) and

5β-cholestane-3α, 7α diol (for synthesis of CDCA). The steroid side chains of these triols and diols are subsequently converted to a carboxyl group by the mitochondrial sterol 27- hydroxylase (CYP27A1), leading to the synthesis of CA and CDCA respectively [6].

Both CYP7A1 and CYP8B1 are expressed exclusively in the liver. CYP8B1 regulates the ratio of CA and CDCA, which subsequently determines the hydrophobicity of the bile acid pool [7, 8]. ii) Alternative or Acidic Pathway: The existence of the acidic pathway was proposed by the identification of acidic intermediates, which could not be formed by the neutral pathway [9, 10]. This pathway contributes less than 18% of the total bile acid synthesis in humans [11], but the metabolites from the acidic pathway are accumulated in patients with chronic liver diseases, indicating that this pathway may be stimulated during diseases [10]. In primary cultures of rat hepatocytes however, the acidic pathway may contribute as much as 50% of the total bile acids [12]. In this pathway, modification of the side chain of cholesterol precedes biotransformation of steroid nuclear ring. Although

CDCA is the major byproduct of the acidic pathway [13, 14], recent studies have also suggested that the pathway also produces CA [15-17]. The initial reaction is

hydroxylation of cholesterol at C-27 to form 27-hydroxycholesterol, followed by oxidation of the product to 3β-hydroxy-5-cholenoic acid. These reactions are catalyzed by the mitochondrial CYP27A1 [18]. The next step is the addition of a hydroxyl group at the C-7 position of the steroid nucleus by oxysterol 7α-hydroxylase (CYP7B1) [19]. It is believed that the same enzymes that catalyze the modification of the sterol nucleus in the classic pathway are used subsequently in the alternative pathway for primary bile acid synthesis [20]. CYP27A1 is expressed in many tissues in the body.

If bile acids are prevented from returning back to the liver, by creating a bile fistula or by cholestyramine (a bile-acid binding resin) feeding, the neutral pathway is highly stimulated while the acidic pathway is not [21-23].

d. Mutations and deficiencies of important enzymes in bile acid biosynthesis

Since bile acids play an important role in cholesterol homeostasis, any disturbance in bile acid synthesis can lead to hypercholesterolemia and cardiovascular diseases. Inborn errors of bile acid synthesis have been described in infants and children, with clinical presentations such as advanced liver disease, neonatal hepatitis, progressive cholestasis and biliary atresia [24-26]. Defects in bile acid synthesis can lead to decreased bile formation, malabsorption of fats and fat-soluble vitamins, hypercholesterolemia, accumulation of toxic, abnormal steroids in the liver leading to cholestasis and cirrhosis

[27]. Primary defects in bile acid synthesis include defects in the modification of the steroid nucleus, such as CYP7A1 [28], 3β-HSD [29-31] and Δ4-3-oxosteroid-5β-

reductase deficiencies [32, 33] and defects in side-chain oxidation like CYP27A1 gene

mutations [34]. Secondary defects in bile acid synthesis include defects in peroxisomal biogenesis and enzymes in peroxisomal β-oxidation as in Zellweger syndrome, infantile

Refsum disease and neonatal adrenoleukodystrophy [35]. Secondary defects can also arise from a defect in de novo cholesterol synthesis like the defect in 7- dehydrocholesterol Δ7-reductase, as in the Smith-Lemli-Opitz syndrome [36, 37].

i) CYP7A1 mutations: A family of CYP7A1-deficiency has been identified [28]. The

patients have hyperlipidemia, premature coronary and peripheral vascular disease and

premature gallstones. The mutation is a deletion mutation in which two thymidine residues are deleted from codon 1303, leading to a frameshift and a truncated protein of

413 amino acids, which has lost the active site and enzyme function. The homozygous

patients have markedly reduced bile acid synthesis and excretion and show compensatory up regulation of CYP27A1 activity in the alternative pathway. The patients are also

resistant to statins (HMG-CoA Reductase inhibitor).

The Cyp7a1-/- knockout mice on the contrary have a different phenotype [38, 39].

Most mice die by day 18, revealing a poor survival rate. Vitamin supplementation to the

nursing mothers prevents deaths in the early period and bile acid supplementation

prevents deaths in the later period. The mice show phenotypical characteristics such as

oily coats, hyperkeratosis, vision defects and behavioral irregularities, consistent with the

malabsorption of Vitamins A, D and E. The adult Cyp7a1-/- mice showed some abnormal

7α-hydroxylated bile acids, which could be explained by the induction of hepatic Cyp7b1

after weaning. Newborn Cyp7a1-/- mice developed neonatal cholestasis due to the

accumulation of 27-hydroxycholesterol and monohydroxylated bile acids such as 3α- hydroxy-5-cholenoate and 3α-hydroxy-5β-cholenoate [38, 39]. ii) CYP27A1 mutations: Mutations in CYP27A1 in human patients cause a rare autosomal recessive defect called cerebrotendinous xanthomatosis (CTX). The disease manifestations include tendon xanthomatosis, progressive neurological dysfunction, tissue cholesterol accumulation, premature atherosclerosis, osteoporosis and cholesterol gallstones [34, 40, 41]. The synthesis of bile acids primarily CDCA is suppressed and

CYP7A1 is up regulated leading to the accumulation of 7α-hydroxycholesterol and 7α- hydroxy-4-cholesten-3-one; the latter forms cholestanol. The patients develop xanthomas and atherosclerosis despite the normal levels of circulating cholesterol, which may be explained by the reduced elimination of cholesterol from the macrophages by CYP27A1.

CDCA therapy is used in these patients to prevent or reverse the neurological symptoms.

Disruption of the Cyp27a1 gene in mice, however does not accumulate cholestanol or exhibit the neurological defects observed in human CTX patients. The mice have markedly reduced bile acid synthesis and fecal bile acid excretion by 80%

[42]. These mice have enlarged liver and kidneys and have increased triglyceride (TG) levels, fatty acid synthesis, and cholesterol absorption and cholesterol synthesis [43]. The livers of the Cyp27a1-/- mice have increased sterol regulatory element binding protein

(SREBP) expression. CA feeding to the mice reverses hepatomegaly and hypertriglyceridemia. Thus, it is believed that CYP27A1 plays a central role in TG metabolism.

iii) CYP7B1 mutation: An inborn error of bile acid metabolism due to a defect in

CYP7B1 was described in a 10-wk-old boy presenting with severe cholestasis, cirrhosis, and liver synthetic failure [44]. The proband presented with an absence of primary bile acid conjugates, absence of 7α-hydroxylated bile acids, accumulation of 3β-hydroxy-Δ5- cholenoic acids and an increase in 27-hydroxycholesterol levels to 4500-fold higher than normal. The mutation was a cytosine to thymidine transition mutation in exon 5 that converts an arginine codon at position 388 to a stop codon. The mechanism of the liver injury is likely due to the accumulation of hepatotoxic monohydroxylated bile acids, which inhibit bile acid transport across the canalicular membranes and reduces bile flow.

Unlike the human CYP7B1 mutation, Cyp7b1-/- mice are essentially normal [45].

Plasma and tissue levels of 25- and 27-hydroxycholesterol, two oxysterol substrates of this enzyme are markedly elevated in Cyp7b1-/- knockout animals. Parameters of bile acid metabolism as well as plasma cholesterol and triglyceride levels in these mice are normal. The cholesterol contents of major tissues were not altered. The loss of this enzyme in the liver is compensated for by increases in the synthesis of bile acids by other pathways (Cyp7a1 expression is induced). iv) CYP8B1 mutation: A CYP8B1 mutation has not been reported so far in human patients. The Cyp8b1-/- mice display no physical or behavioral abnormalities [46]. The almost complete lack of cholic acid was replaced by substantial amounts of CDCA and muricholic acid. However, the bile acid pool size and fetal bile acid excretion rate increased by 37% and 53% in the knockout mice and there was a significant induction of

Cyp7a1 mRNA levels. Since muricholic acid (hydrophilic bile acid), which is the

predominant bile acid in the mouse bile, is not a Farnesoid X receptor (FXR) (bile acid receptor) ligand, the highly hydrophilic bile in these mice may derepress the Cyp7a1 gene. The intestinal cholesterol absorption decreased by 41% in the Cyp8b1-/- mice, since CA, which is the most efficient bile acid for cholesterol absorption from the intestine was absent. This led to a compensatory two-fold increase in hepatic de novo cholesterol biosynthesis reflected by an increase in the HMG-CoA reductase mRNA levels in the liver.

e. Enterohepatic circulation of bile

Bile acids are synthesized in the liver, excreted as bile and stored in the gallbladder, secreted into the duodenum in response to food intake, and 95% of the bile acids are reabsorbed back in the terminal ileum and returned back to the liver via the portal blood

(only ~0.5g/day is excreted into the feces) [1-3]. This process is termed as the

“enterohepatic circulation” of bile. The bile acids returning to the liver serve as an important physiological function by regulating processes such as cholesterol synthesis, bile acid synthesis, bile acid transport etc. The transporters which mediate the enterohepatic circulation of bile are described below according to their location and function (Fig. 3). Dysfunction of the bile acid transporters can lead to two clinical situations, namely cholestatic liver disease and intestinal bile salt malabsorption [47].

Cholestasis is caused by an “overflow” of bile salts in the liver and systemic circulation due to hepatocellular bile salt transport and excretion defect. The disease presents as elevated serum makers of cholestasis, pruritis, jaundice, intestinal malassimilation, liver

Fig.3.

Figure 3. Enterohepatic circulation of bile.

At the hepatocyte basolateral membrane bile salt uptake takes place via the Na+

taurocholate cotransporting peptide (NTCP) and Na+ independent uptake takes place via

the organic anion transporting peptides (OATPs). At the hepatocyte canalicular

membrane bile salt excretion takes place via the bile salt export pump (BSEP) and the

multidrug resistance related protein 2 (MRP2). Biliary excretion of phospholipids and

cholesterol is mediated by MDR3 and ATP-binding cassette G5/G8 (ABCG5/G8), respectively. In cholangiocytes and enterocytes, the apical Na+ dependent bile salt transporter (ASBT) mediates bile salt uptake across the luminal membrane. Ductular bile is modified by chloride channels such as the cystic fibrosis transmembrane conductance regulator (CFTR) and the chloride/bicarbonate exchanger (anionic exchanger 2, AE2).

Enterocytes also express ABCG5/G8 at their apical membrane for sitosterol excretion.

MRP3 is a basolateral efflux system with a low affinity for bile salts. The heteromeric organic solute transporter Ostα/Ostβ acts as a basolateral bile acid carrier in the ileum

and other ASBT-expressing tissue.

damage and gallstone formation. Intestinal bile salt malabsorption on the other hand leads to an “underfilling” of the bile salt pool and hence decreased bile salt excretion leading to clinical manifestations such as chologenic diarrhea and contributing to gallstone formation. i) Hepatocellular Bile Salt Excretion: Taurine and glycine conjugated bile salts are excreted into bile from the hepatocytes via the bile salt export pump (BSEP, ABCB11), an adenosine triphosphate (ATP)-binding cassette (ABC) transporter localized in the canalicular or apical domain of the hepatocytes plasma membrane[48, 49]. BSEP has a high affinity to bile salts and belongs to the multidrug resistance (MDR) gene family of

ABC transporters. Since mammals have a canalicular bile salt concentration, which is

1000-fold greater than the portal blood, an active ATP-dependent efflux system is required at the canalicular membrane of the hepatocytes.

Another canalicular transporter is the MDR related protein 2 (MRP2), which transports glucoronidated and sulfated bile salts [50]. MRP2 is localized on the canalicular domain of hepatocytes and the apical domain of enterocytes in the proximal small intestine.

MRP2 also transports conjugated bilirubin, as well as several drugs (chemotherapeutics, antibiotics), glutathione, toxins and heavy metals.

The excretion of the amphipathic bile salts into bile drives phosphatidylcholine and free cholesterol to bile to form mixed micelles in the gallbladder. The oversaturation of cholesterol in the mixed micelles causes cholesterol gallstone formation [51]. The

MDR3 (ABCB4) is a phospholipid export pump on the canalicular membrane of hepatocytes [52, 53]. A heterodimeric ABC transporter, composed of two units coded by

2 oppositely oriented genes called ABCG5 and ABCG8, is involved in the cholesterol excretion into bile [54, 55]. ABCG5/ABCG8 is localized on the canalicular membrane of the hepatocytes and the apical membrane of the enterocytes. ii) Cholangiocyte Bile Salt Transport: Primary hepatic bile is modified as is passes through the biliary tree by organic anion and electrolyte transport proteins present in the biliary epithelial cells. Cholangiocytes can uptake bile acid via Na+-dependent

transporter, known at the apical sodium-dependent bile salt transporter (ASBT,

SLC10A2) [56]. ASBT is expressed on the apical or luminal membrane of

cholangiocytes, the apical membrane of enterocytes and renal proximal tubular cells. The

physiological role of bile salt uptake into the cholangiocytes pertains to the regulatory

effect of the bile salts on the cholangiocyte mucin and bicarbonate secretion. Since

luminal bile acid concentration in the gallbladder is in the millimolar range, the uptake of

bile salts by the cholangiocytes does not affect the biliary bile salt concentration.

Ductular bile is modified by chloride channels such as the cystic fibrosis transmembrane

conductance regulator (CFTR) and the chloride/bicarbonate exchanger (anion exchanger

2/ AE2) [57]. iii) Intestinal Absorption of Bile Salts: Reabsorption of bile salts takes place primarily in the ileum, via the ASBT transporter, sometimes also known at the ileal bile acid transporter (IBAT) [58, 59]. Human ASBT transports conjugated and unconjugated bile salts with higher affinity for CDCA and CA than for taurocholate [60]. A Na+-

independent bile salt uptake takes place via the organic anion transporting peptide 3

(Oatp3, Slc21a7) from the apical surface of jejunal enterocytes in rats [61].

ABCG5/ABCG8 on the apical membrane of the enterocytes, limits intestinal absorption and promotes excretion of plant sterols (sitosterols) [62, 63]. Another important ABC transporter involved in cholesterol transport is ABCA1, which is located on the basolateral membrane of hepatocytes and Caco2 cells (human colonic carcinoma) [64,

65]. ABCA1 mediates cellular cholesterol efflux from peripheral macrophages, which is important for HDL formation and reverse cholesterol transport. Mutations of the ABCA1 gene in man, causes Tangier disease, a familial high-density lipoprotein deficiency [66-

68].

After their uptake into enterocytes, bile salts are shuttled to the basolateral membrane for efflux into the portal blood, via the ileal bile-acid binding protein (IBABP) that is cytoplasmically attached to ASBT [69]. Basolateral extrusion of bile salts is mediated by MRP3 (ABCC3). However, the human MRP3 has a lower affinity for conjugated bile salts than rat Mrp3, suggesting that other transporters may be involved in the basolateral bile salt efflux [70, 71]. Recently, a new candidate basolateral bile acid carrier, the heteromeric organic solute transporter (Ostα/Ostβ) was identified in studies on wild type and Asbt null mice [72]. The mouse Ostα/Ostβ is a heteromeric transporter and acts as a basolateral bile acid carrier and may be responsible for bile acid efflux in the ileum and other ASBT-expressing tissue. The presence of this transporter in humans is still unknown. iv) Hepatocellular Uptake of Bile Salts: Bile salts circulate in plasma tightly bound to albumin and lipoproteins such as HDL [73]. More than 80% of the conjugated bile salts undergo single-pass extraction by the liver from the portal venous blood, mainly via the

Na+-taurocholate cotransporting polypeptide (NTCP. SLC10A1). NTCP has been

localized to the basolateral hepatocytes membrane in rat and human liver [74, 75]. Unlike

ASBT, the substrate specificity of rat Ntcp is not limited to only bile salts but also

includes sulfated sex steroids, thyroid hormones and certain drug conjugates [75, 76].

Na+-independent hepatic uptake of bile acids is mediated by the OATP family of

transporters [77, 78]. Of the different OATP transporters, OATP1A2 (SLC01A2) and

OATP1B1 (SLC01B1) are the two Na+-independent bile salt uptake systems of the

human hepatocytes, where as OATP1B3 (SLC01B3) mediates taurocholate transport to a

lesser extent and OATP2B1 (SLC02B1) does not transport bile salts at all [79].

2. Cholesterol Homeostasis in the liver

Cholesterol is the precursor to bile acids and may also regulate bile acid synthesis. Hence

cholesterol homeostasis is a relevant topic related to my study. Cholesterol is an essential

component of the cell, a precursor to steroid hormones and bile acids and oxidized

cholesterol (oxysterol) serves as important signaling molecules. However, accumulation

of excess cholesterol in arterial blood vessels can cause atherosclerosis [80, 81].

Imbalance of cholesterol metabolism causes hypercholesterolemia in humans, which is a

major risk factor for atherosclerosis and chronic heart disease, ischemic heart attacks,

stroke and cholelithiasis [82-84]. Cholesterol homeostasis is maintained by complex

mechanisms that control both the absorption and elimination of cholesterol.

The liver plays a central role in maintaining cholesterol homeostasis (Fig. 4) [1-

3]. Conversion of free cholesterol to cholesterol ester requires the enzyme acyl-CoA: cholesterol acyltransferase (ACAT), while the reverse reaction requires the enzyme cholesterol ester hydroxylase (CEH). The major cholesterol input mechanisms in the liver include 1) The uptake of serum cholesterol esters via the circulating low-density lipoprotein (LDL) particles through LDL receptor mediated endocytosis [85]; 2) the reverse cholesterol transport from the peripheral tissues to the liver via the uptake of high-density lipoprotein (HDL) by the scavenger receptor subtype B1 (SR-B1) [86]; 3)

The dietary cholesterol absorption via the intestine and its transport to the liver via chylomicron (CM) remnants which are taken up the LDL receptor related protein (LRP or apolipoprotein E receptor) [87, 88] and 4) The de novo synthesis of cholesterol from acetyl-coenzyme A (Acetyl-CoA). The rate-limiting enzyme for this pathway is 3- hydroxy-3-methyl glutyryl-CoA reductase (HMG-CoA reductase) [89].

Cholesterol output from the liver comprises of the assembly of cholesterol esters into very low-density lipoproteins (VLDLs) that are secreted into the circulation. This is termed as normal cholesterol transport. The VLDLs contain triglycerides and cholesterol esters, which are hydrolyzed to free fatty acids (FFA) and glycerol by lipoprotein lipase

(LPL) in the endothelium of blood vessels in the muscle and adipose tissue. The VLDLs decrease in size to form intermediary density lipoproteins (IDLs) that in turn can be hydrolyzed and converted to LDLs. LDL can be taken back up by the liver as mentioned before or it can be oxidized and internalized by macrophages via the scavenger receptor subtype A1 (SR-A1).

Fig.4.

Figure 4. Cholesterol Homeostasis in the liver.

Cholesterol exists in the liver in both the free and esterified form. Conversion of free cholesterol to cholesterol ester requires the enzyme acyl-CoA: cholesterol acyltransferase

(ACAT), while the reverse reaction requires the enzyme cholesterol ester hydroxylase

(CEH). Cholesterol input mechanisms (dark gray arrows): 1) Uptake of low-density lipoprotein (LDL) particles through LDL receptor mediated endocytosis 2) Reverse cholesterol transport of cholesterol extruded from peripheral tissues by the ATP-binding cassette A1 (ABCA1) receptor, and the uptake of high-density lipoprotein (HDL) by the scavenger receptor subtype B1 (SR-B1) 3) Dietary cholesterol absorption via chylomicron (CM) remnants which are taken up via the LDL receptor related protein

(LRP) 4) de novo synthesis of cholesterol acetyl-coenzyme A (Acetyl-CoA) [rate-limiting enzyme: 3-hydroxy-3-methyl glutyryl-CoA reductase (HMG-CoA reductase)]

Cholesterol output mechanisms (light gray arrows): 1) Assembly of cholesterol esters

(CE) into very low-density lipoproteins (VLDLs)→ intermediary density lipoproteins

(IDLs) → LDL. LDL can be internalized by macrophages via the scavenger receptor subtype A1 (SR-A1) 2) Cholesterol is converted into bile acids [rate-limiting enzyme: cholesterol 7α-hydroxylase (CYP7A1)]. Bile acids are secreted into the intestine and reabsorbed into the liver via the transporter Sodium taurocholate cotransporting peptide

(NTCP) 3) Free cholesterol is excreted along with bile acids and phospholipids into the bile and eventually lost in the feces.

Cholesterol is eliminated from the body via three major mechanisms: 1) 50% of the cholesterol is converted into bile acids; 2) 10% is used for the synthesis of steroid hormones; and 3) The remaining 40% is excreted along with bile acids and phospholipids into the bile. Thus, the secretion of unmodified cholesterol and bile acids accounts for

90% of the total cholesterol disposal per day [90]. The unmodified cholesterol is excreted via the bowel in the feces, while 95% of the bile acids are reabsorbed in the terminal ileum and returned to the liver.

3. Nuclear Hormone Receptors

Nuclear hormone receptors are one of the most abundant classes of transcriptional regulators in animals. They regulate diverse functions in the body and serve as ligand activated transcription factors, thus providing a direct link between signaling molecules that control these processes and their transcriptional responses. A number of compounds such as bile acids, oxysterols, fibrates etc regulate bile acid biosynthesis. Several nuclear receptors (NRs) have been identified as mediating the regulation of bile acid biosynthesis by these compounds. CYP8B1 is also known to be regulated by various NRs as will be discussed in a later section. In this section, the general information about the NR structure, response elements and ligands are discussed (Fig. 5).

Fig.5.

Figure 5. Structure of nuclear hormone receptors.

Upper figure shows the domains of a nuclear receptor. From the N –terminal onwards to domains are: Activation function domain 1 (AF1), DNA binding domain (DBD), Hinge region, Ligand binding domain (LBD), and Activation function domain 2 (AF2). The second portion of the figure shows the structure of the DBD which consists of a Zinc finger motif. The third portion shows how nuclear receptors dimerize (either form homodimers or heterodimers) and bind to the hormone response element (HRE) on the target gene DNA. The last portion shows the different orientations of the hexameric half sites in the HRE. They can be oriented as direct repeats (DR), inverted repeats (IR) or everted repeats (ER). Some nuclear receptors bind the DNA as monomers to extended half-sites preceded by an A/T rich sequence.

a. Structure

The NRs have a typical modular structure composed of several domains which are

differentially conserved between the various receptors and have different roles: a variable

ligand-independent activation function-1 (AF-1) region in the N-terminal, followed by a

highly conserved DNA-binding domain (DBD), a variable hinge region and a moderately

conserved ligand binding domain (LBD), and finally a variable ligand-dependent

activation function-2 (AF-2) domain in the C-terminal region [91-93]. The DBD contains

two Cys2/Cys2 Zinc finger motifs where four cysteines within the consensus Cys-X2-

Cys-X1-3-Cys-X2-Cys sequence coordinate with a zinc ion to form a loop structure. The

DBD is responsible for DNA binding and dimerization. The LBD participates in several activities including ligand/ hormone binding, homo- and/or heterodimerization, formation

of the heat-shock protein complex and transcriptional activation and repression by

coregulator interaction. The NRs bind to the consensus hormone response elements

(HREs) on the target genes. The binding of the ligand induces conformational changes in the NRs. The conformational changes that accompany the transition between the liganded and unliganded forms of the NRs affect dramatically their affinity for corepressors and coactivators, which bind to the helix 2 of the NRs [94, 95].

b. Response Elements

A HRE is a specific DNA sequence that a receptor recognizes with markedly increased

affinity and typically contains two consensus hexameric half-sites. The identity of a

response element resides in three features: the sequence of the base pairs in the half-site,

the number of base pairs between the half-sites (nucleotide spacing) and the relative orientation of the two half-sites. Thus each receptor protein dimer that binds the DNA has to recognize the sequence, spacing and orientation of the half-sites within their response element. The half-sites may be oriented as direct repeats (DR), inverted repeats (IR) or everted repeats (ER) [96]. Classic steroids hormone receptors like glucocorticoid receptor

(GR), mineralocorticoid receptor (MR), androgen receptor (AR) and progesterone receptor (PR) bind palindromic AGAACA-N3-TGTTCT sequences, while estrogen receptor (ER) and the non steroid hormone receptors bind to AG(G/T)TCA-like repeats.

Some receptors such as liver X receptor (LXR), farnesoid X receptor (FXR), pregnane X receptor (PXR) etc. bind to their response elements as heterodimers with retinoid X receptor (RXR) [97]. Hepatocyte nuclear factor 4α (HNF4α) binds as a homodimer to a

DR-1 site, where as human α-fetoprotein transcription factor (hFTF) binds to an extended monomeric site of TCAAAGGTCA. Small heterodimer partner (SHP) is a negative nuclear receptor that lacks a DNA-binding domain and thus cannot bind to DNA.

c. Ligands

NRs bind to a variety of ligands. The hormone receptors bind to specific hormones, e.g.

GRs bind to glucocorticoids or thyroid hormone receptors (TR) bind to thyroid hormone.

However a wide variety of other bioactive molecules such as fatty acids, farnesol metabolites, oxysterols, leukotriene B-1, prostaglandin, etc. have been identified as ligands to NRs. Several intermediates of the bile acid synthesis and cholesterol metabolism are endogenous ligands of NRs e.g. oxysterols are ligands of LXR, while bile

acids are ligands of FXR. Many NRs have unknown ligands and remain constitutively active within cells in which they are expressed. These NRs are referred to as “orphan receptors” [91]. HNF4α, chicken ovalbumin upstream promoter transcription factors I and II (COUP-TF I and II), FTF and SHP are examples of orphan receptors.

4. Important Nuclear Receptors involved in regulation of bile acid biosynthesis

Bile acids serve as important signaling molecules to regulate several physiological processes in the body [1-3]. They activate various NRs in the cell, which in turn exert effects on their downstream target genes. Also several NRs are regulated by different ligands to alter the transcription of bile acid biosynthetic genes. The NRs that are activated by bile acids and those that are important regulators of bile acid biosynthesis are described here.

a. Farnesoid X Receptor (FXR, NR1H4)

FXR is an important nuclear receptor involved in bile acid and lipid metabolism regulation. FXR is highly expressed in the liver, intestine, adrenal and kidney [98, 99] .

FXR binds to DNA as a heterodimer with RXR, preferentially to IR-1 motifs [98, 99].

Farnesol, all trans-retinoic acid, juvenile hormone III and 4-[(E)-2-(5,6,7,8-Tetrahydro-

5,5,8,8-tetramethyl-2-naphthalenyl)-1-propenyl]benzoic acid (TTNPB) can activate FXR at high concentrations [100]. Bile acids have been identified as endogenous ligands for

FXR [100-102]. The hydrophobic bile acid, CDCA, is the most potent activator of FXR.

The secondary bile acids, LCA and DCA, are less effective, and hydrophilic bile acids, ursodeoxycholic acid (UDCA) and muricholic acids, are inactive. A FXR agonist,

GW4064 is 200 fold more potent than CDCA in activation of FXR [103].

Bile acid-activated FXR inhibits CYP7A1 transcription without binding to the gene [104], indicating a indirect mechanism involving other factors, namely FTF and

SHP [105, 106]. Bile acid-activated FXR also inhibits CYP8B1 and HNF4α transcription

[107]. FXR also inhibits NTCP [108], while it markedly induces BSEP [109]. Thus, FXR

may play an important role in protecting the liver against excess bile acids. FXR induces

IBABP expression in the intestine [110]. FXR also induces phospholipid transport protein

(PLTP) [111] and apolipoprotein CII (ApoCII) [81, 112] involved in the reverse

cholesterol transport and triglyceride metabolism. The FXR agonist GW4064, stimulates

SHP mRNA expression in rats [105] and overexpression of FXR in the presence of

CDCA stimulates SHP mRNA levels in HepG2 cells [113]. The Fxr-/- mice show

elevated serum bile acids, cholesterol and triglycerides, reduced bile acid pool and fecal

bile acid secretion and lack of bile acid inhibition of Cyp7a1 expression [114].

b. Pregnane X Receptor (PXR, NR1I2)

Mouse PXR or the human ortholog steroid X receptor (SXR) is an important nuclear receptor known to induce the CYP3A family of genes. CYP3A4 is the most abundant cytochrome P450 isozyme expressed in the human liver and intestine and metabolizes

about 60% of the clinically used drugs in the body. PXR is predominantly expressed in

the liver and intestine [115]. PXR heterodimerizes with RXR and binds to different

response elements consisting of DR-3, DR-4, DR-5 or ER-6 in the CYP3A genes. PXR is a promiscuous xenobiotic receptor that is activated by unrelated steroids (e.g. dexamethasone, prenenolone 16α-carbonitrile (PCN)), xenobiotics and drugs (like phenobarbitol, rifampicin) and antibiotics [116, 117]. LCA has recently been identified as a PXR ligand [117, 118]. It was suggested that PXR might function as a bile acid sensor and induce CYP3A4, which is known to convert LCA to a more hydrophilic and less toxic bile acid, hyodeoxycholic acid (HDCA). PXR is also known to induce OATP2

[119] in the sinusoidal membrane of the hepatocytes, which transports LCA into the hepatocytes. PXR has recently been shown to inhibit human CYP7A1 [120, 121] and

CYP8B1 gene expression [121]. Thus, PXR may play a role in hepatotoxicity and cholestasis. PXR also induces MDR1 and MRP2, which transport sulfate-conjugated tauro-CDCA and tauro-LCA to canaliculi [122]. In Pxr-/- mice, PCN does not affect bile acid secretion and the mice develop inflammatory response and liver damage upon LCA treatment [119]. However, the Pxr null mice are responsive to the LCA induction of

Cyp3a [119].

c. Vitamin D3 Receptor (VDR, NR1I1)

VDR is a ligand-activated nuclear receptor that plays a central role in calcium homeostasis and has been implicated in regulating diverse biological functions, including immunity, cellular proliferation and differentiation [123]. VDR is primarily expressed in the bones, kidney, intestine, liver and parathyroid glands [124]. VDR binds to DNA on

DR-3 sites as either a heterodimer with RXR or as a VDR-VDR homodimer. The

physiological ligand of VDR is 1,25-dihydroxyvitamin D [1,25(OH)2D], the active form

of vitamin D3. However, recently LCA has been identified as a VDR ligand [125]. Of the

different LCA derivatives, LCA acetate is the most potent VDR agonists [126]. VDR is an order of magnitude more sensitive to LCA and its metabolites than are other nuclear receptors. Activation of VDR by LCA or vitamin D, induces the in vivo expression of

CYP3A, which helps detoxify LCA in the liver and intestine and serves as a protective mechanism against colorectal cancers in humans [125].

d. Hepatocyte nuclear factor 4α (HNF4α, NR2Α1)

HNF4α is the most abundant nuclear receptor in the liver and is also highly expressed in

the intestine and kidney. It binds DNA via a DR-1 motif as a homodimer [127]. HNF4α is an orphan receptor, with constitutive activity and can transactivate genes without ligand binding [128]. Fatty acyl-CoA thioesters have been shown to activate HNF4α, but

the physiological relevance of these ligands has been questioned [129]. HNF4α regulates

the liver-specific expression of many lipoproteins including ApoA1, ApoB and ApoCIII

[130, 131], and genes in glucose metabolism [127, 132].

HNF4α binds and stimulates rat Cyp7a1 promoter activity [133, 134]. Mutation

of the HNF4α binding site markedly reduced CYP7A1 promoter activity indicating that

HNF4α is crucial for basal level transcription. HNF4α binding sites have also been identified in CYP8B1 [107, 135] and CYP27A1 genes [136]. Disruption of Hnf4α in mice is lethal, as HNF4α is critical for embryonic liver development and differentiation.

Liver specific conditional disruption of Hnf4α gene results in accumulation of lipids in

the liver, decrease in serum cholesterol and triglycerides and accumulation of bile acids

in the serum [137]. These mice show a marked reduction of Cyp7a1, Hnf4α, Apoall,

Apob100, ApocIII, Ntcp, Oatp1 and microsomal triglyceride transport protein (MTP).

This is consistent with the important role HNF4α plays in basal transcription of CYP7A1

and lipoprotein gene. Mutations of HNF4α gene have been linked to maturity onset

diabetes of the young (MODY 1) [138, 139]. HNF4α is also an upstream regulator of the

HNF1α gene, the mutation of which is linked to MODY3 [140, 141].

e. Human α-fetoprotein transcription factor (hFTF, NR5A2)

The Fushi-tarazu-factor 1 (Ftz-F1) family of nuclear receptors encodes FTF which has

many homologues in other species such as rat FTF [142], human CPF [143], hepatitis B

virus enhancer 1 factor [144], human hFTF [145], mouse LRH [146], Xenopus laevis

xFF1 [146] and the zebrafish zFF1 [147]. FTF is expressed in liver, intestine and pancreas and is most related to steroidogenic factor 1 (SF-1) expressed in steroidogenic tissues [148]. FTF binds as a monomer to the core DNA sequence 5'-TCAAGGTCA-3'.

FTF binding sites have been identified in CYP7A1 [143], CYP8B1 [149, 150], SF-1

[151], SHP [152], HNF3β, HNF4α and HNF1 [153]. SF-1 (NR5A2) plays an important role in steroidogenesis, liver growth, endocrine development and differentiation [154,

155].

The FTF binding sites on the CYP7A1 and CYP8B1 genes overlap with HNF4α sites. FTF is a weak transcription factor that when transfected at high concentrations

stimulates CYP7A1 reporter activity by ~2 fold in non-liver cells [143]. Bile acids can induce FTF mRNA expression in rat livers and HepG2 cells [113, 135] and FTF acts as a repressor of human CYP7A1 and rat Cyp8b1 transcription in HepG2 cells [107, 113, 135].

The inhibitory effect of FTF is most likely due to competition for HNF4α binding to overlapping binding sites on the genes. Ftf-/- mice embryos die at embryonic day 6.5-7.5, with features typical of visceral endoderm dysfunction [156]. Adult Ftf+/- mice are hypocholesterolemic, and express liver Ftf at about 40% of the normal level. Cyp7a1 expression is increased in the Ftf+/- mice liver [156].

f. Small Heterodimer Partner (SHP, NR0B2)

SHP is a unique orphan nuclear receptor protein that contains a receptor-interacting domain and a repressor domain, but has no conventional DNA binding domain [157,

158]. SHP acts like a promiscuous inhibitory heterodimer partner of nuclear receptors and inhibits transactivation activity of FTF, RAR, CAR, PXR, HNF4α, estrogen receptor

α and β (ERα and ERβ), peroxisome proliferators activated receptor (PPAR), TR and other receptors [113, 157, 159-163]. The SHP repression of nuclear receptor activity is said to be mediated either because SHP competes with coactivators (like steroid receptor coactivator) for nuclear receptor binding or directly through the C-terminal repressor domain of SHP [159].

The FXR agonist GW4064, repressed Cyp7a1 mRNA but stimulated Shp mRNA expression in rats [105]. Also, overexpression of FXR in the presence of CDCA stimulated SHP mRNA levels in HepG2 cells [113]. SHP can mediate the suppression of

CYP7A1 [113], CYP8B1 [107, 135] and CYP27A1 [136] by bile acids. SHP mutations have been identified in obese Japanese patients with early onset diabetes [164]. Shp-/- mice are normal except for mild defects in bile acid and cholesterol homeostasis [165,

166]. These mice show an increase in bile acid synthesis due to derepression of Cyp7a1 and Cyp8b1. However, these mice respond to bile acid feedback inhibition of bile acid synthesis when fed bile acids suggesting the existence of compensatory repression pathways of bile acid signaling.

g. Peroxisome Proliferator Activated Receptor α (PPARα, NR1C1)

PPARα is a nuclear receptor expressed in the liver, cardiac muscle and adipose tissue

[167, 168]. Fatty acids, eicosinoids and hypolipidemic drugs (like fibrates) are ligands of

PPAR [167, 169-171]. Upon ligand activation PPARα heterodimerizes with RXRα, and

binds to the peroxisomal proliferator response elements (PPREs), which are DR-1 sites,

to regulate gene transcription. Treatment of gallstone and hypercholesterolemia patients

with fibrates, a PPARα agonist reduces the bile acid synthesis and pool size [172, 173].

The PPARα agonist, Wy14,643 suppresses CYP7A1 in HepG2 cells, via a PPRE (DR-1

site), but the PPARα/RXRα heterodimer cannot bind that site [174, 175]. PPAR also

stimulates Cyp8b1 activity and increases CA synthesis in rats, but binds very weakly to

the Cyp8b1 gene [176]. Fatty acid activated PPARα stimulates LXRα expression and

induces the target genes of the latter, namely ABCA1 and ABCG1 in macrophages [177,

178]. This suggests that the PPAR-LXR-ABCA1 cascade is involved in cholesterol efflux

in macrophages. Pparα knockout mice display a defect in the lipid and lipoprotein

metabolisms. In Pparα-/- mice, inhibitory effects of fibrates on bile acid synthesis and

Cyp7a1 and Cyp27a1 expression is abolished [179].

h. Liver X Receptor (LXR, NR1H3)

LXR has two isoforms in the body, LXRα and LXRβ [98, 180-186]. LXRα is expressed

in liver, spleen, adipose tissue, lungs and pituitary, whereas LXRβ is ubiquitously

expressed. Oxysterols have been identified as LXR ligands, of which the naturally

occurring 22 (R)-hydroxycholesterol, 24 (S)-hydroxy cholesterol and 24(S),

25-epoxycholesterol are the most potent LXR ligands [186]. LXRα might function as a

cholesterol sensor and stimulate genes by binding to DR-4 motifs as heterodimer with

RXRα. For example, Lxrα can stimulate rat Cyp7a1 expression to convert excess

cholesterol to bile acids in response to high cholesterol [187]. However, LXRα has much

less effect on hamster and human CYP7A1, which lack DR-4 motifs [188]. Cholesterol

feeding of mice and rats decreases the activity of Cyp8b1, in contrast to the upregulation

of Cyp7a1 [189].

Oxysterol induced LXRα also binds and stimulates the sterol response element

binding protein -1c (SREBP-1c) gene [190], and thus stimulates lipogenesis and leads to

hypertriglyceridemia. LXRα also stimulates ABCA1 [191], which functions as a

cholesterol and phospholipid efflux regulator in HDL synthesis and ABCG5 and ABCG8

half-transporters [192], which regulate biliary sterol efflux by limiting intestinal

absorption and promoting biliary excretion of plant sterols (sitosterols). Other genes

stimulated by LXRα include ABCG1 in macrophages [193], the ApoE gene in

macrophages and adipocytes [194], the human cholesterol ester transfer protein (CETP)

[195] that mediates the exchange of cholesterol esters from HDL to IDL and LDL, the lipoprotein lipase (LPL) gene [196] involved in hydrolysis of triglycerides carried by

VLDL and CM and also LXRα (autoregulation) synthesis [197] in macrophages. Lxrα-/-

mice show normal levels of Cyp7a1 mRNA, but the Cyp7a1 gene is not stimulated by a

high-cholesterol diet, as in the wild-type mice, leading to a massive accumulation of liver

cholesterol [198]. These mice also show a decrease in the Srebp-1 and stearoyl-CoA

desaturase (Scd) mRNA levels [198].

5. Important Transcription factors and Coactivators involved in regulation of bile

acid biosynthesis

a. Sterol Regulatory Element Binding Proteins (SREBPs)

SREBPs are a family of membrane bound transcription factors. They comprise of three

members SREBP-1a, SREBP-1c and SREBP-2. SREBP-1a and 1c are produced by the

same gene by the use of alternative promoters to produce different first exons that are

spliced into a common second exon [199]. A different gene produces SREBP-2. Most

organs including liver and adipose tissues predominantly express SREBP-2 and -1c

[200]. All the three SREBP proteins differ in their amino terminal domain, which affects

their function. SREBP-2 appears to regulate genes involved in the cholesterol

biosynthetic pathways, while SREBP-1a and SREBP-1c (also known as adipocyte

determination and differentiation factor-1 i.e. ADD-1) is specifically involved in the

stimulation of genes involved in the fatty acid biosynthetic pathway, such as acetyl-CoA carboxylase, fatty acid synthase (FAS), stearoyl CoA desaturase-1 (SCD-1), etc [201].

All the SREBP isoforms share a triparate structure, comprising of ~1150 amino acids (AAs) [199]. The amino terminal domain is a helix-loop-helix-leucine-zipper (DNA binding motif) transcription factor comprising of ~480 AAs. This is followed by a transmembrane domain comprising of ~80 AAs, which are arranged as two α–helices separated by a short hydrophilic loop of 31 AAs, which protrudes, into the lumen of the endoplasmic reticulum and the nuclear envelope. Finally, a carboxy terminal regulatory domain consisting of ~590 AAs remains complexed with a membrane protein SCAP

(SREBP cleavage-activator protein) in the endoplasmic reticulum. The proteolytic release of the active amino terminal of SREBPs is dependent upon the oxysterol levels in the body [202, 203]. In sterol-depleted cells SCAP escorts SREBP to the golgi apparatus where Site-1 protease cleaves a peptide bond in the luminal loop, thus separating the two transmembrane helices. The amino terminal domain is then cleaved by Site-2 protease, a zinc metalloenzyme. The active amino terminal domain then enters the nucleus to activate transcription of the target genes. However, in sterol containing cells,

SCAP/SREBP complex remains in the endoplasmic reticulum bound to and inhibited by the three isoforms a protein called insulin-induced gene ( Insig-1, 2a and 2b) [204, 205].

Therefore, the SREBPs remain membrane-bound and inactive [199].

SREBP-1c can be activated by oxysterols and insulin. Oxysterol induced LXRα binds and stimulates the SREBP-1c gene [190]. Insulin stimulates SREBP-1c in the liver

[206, 207], probably by selectively down-regulating the expression of Insig-2a, which is

the predominant Insig transcript in the livers of fed animals [208]. Thus, Insulin induces

SREBP-1c processing, thereby allowing it to stimulate fatty acid synthesis. Co- transfection of Srebp-1a and -1c stimulated rat Cyp8b1 promoter activity, while Srebp-2 did not have any effects [189, 209]. The rat Cyp8b1 promoter contains several sterol response elements (SREs) and E-box motifs, which bind Srebp and stimulate the Cyp8b1 expression [189, 209]. SREBP also suppresses the human CYP7B1 luciferase reporter gene in several cell lines, without binding to the putative SREs [210].

b. c-Jun c-Jun is a member of the activating protein-1 (AP-1) family of transcription factors [211,

212], which also includes the proteins c-Fos [213, 214] and ATF-2 (activating transcription factor 2). These proteins contain a leucine zipper DNA binding motif [215].

Upon activation, these proteins for homodimers or heterodimers to bind DNA and activate transcription [215]. Mitogen activated protein (MAP) kinases are capable of modulating gene expression by phosphorylating transcription factors directly. c-Jun is phosphorylated on serine 63 and 73 by the c-Jun NH2 terminal kinase (JNK) [211,

212]and on the C-terminal inhibitory sites by extracellular response-regulated kinase

(ERK). Phosphorylation at the N-terminal sites results in increased stability of c-Jun and

an increase in its transactivational potential [211, 212]. While phosphorylation of the C-

terminal sites of c-Jun inhibits DNA binding [216, 217].

c-Jun expression has been shown to be induced by bile acids [218]. c-Jun binds an

AP-1 site on the SHP promoter and activates SHP transcription, leading to the inhibition

of the CYP7A1 gene [218]. However, the direct effect of c-Jun on the CYP7A1 gene promoter is yet to be determined. Recent studies have shown that c-Jun can interact with other transcription factors and alter the transcription of a gene without directly binding to the DNA [156]. It has been suggested that c-Jun could form a repressive complex with an unknown factor to inhibit the CYP71 gene transcription [156, 219]. It is likely that c-Jun could affect the transcription of CYP8B1 however no studies related to this have been performed so far. It is not known whether c-Jun can bind the CYP8B1 promoter at a putative AP-1 site either as a homodimer or as a heterodimer with c-Fos.

c. Peroxisome Proliferator-Activated Receptor-γ Coactivator 1α (PGC-1α):

PGC-1α is a coactivator that was first identified as a protein that interacts with nuclear receptor PPARγ. Coactivators are proteins that increase the rate of transcription by interacting with transcription factors however they do not bind the DNA in a sequence-specific manner. Coactivators can unwind and remodel the chromatin in an ATP-dependent fashion by modifying the histone proteins, allowing RNA polymerase II to bind the chromatin to initiate transcription. PGC-1α does not have an intrinsic enzyme activity, but upon binding a transcription factor it induces a conformational change that recruits other coactivator proteins that contain histone acetyltransferase (HAT) activity, such as steroid receptor coactivator-1 (SRC-1) and

CREB (cAMP response element binding protein) binding protein (CBP/p300) [220].

PGC-1α interacts with a host of NRs such as HNF4α, ER, PPARα, RXRα, GR, retinoic

acid receptor (RAR), TR etc [221-226]. PGC-1α regulates various metabolic processes in

the body such as stimulation of mitochondrial oxidative metabolism in brown adipose tissue, fibre-type switching in skeletal muscle and multiple aspects of the fasted response in the liver [227].

PGC-1α expression in the liver is dramatically increased by fasting [222, 228].

PGC-1α stimulates gluconeogenesis by inducing the key gluconeogenic enzyme, phosphoenolpyruvate carboxy kinase (PEPCK) by interacting with the NRs HNF4α and

GR [222]. The interaction between HNF4α and PGC-1α is ligand independent [222] and has also been shown to strongly stimulate CYP7A1 gene transcription [229]. HNF4α is the most important activator of CYP8B1 and hence it is likely that the interaction of PGC-

1α with HNF4α may strongly stimulate the CYP8B1 gene transcription. Recent sudies in our lab have shown that PXR can disrupt the interaction between HNF4α and PGC-1α leading to down-regulation of CYP7A1 gene transcription [120].

6. Regulation of Bile Acid Biosynthesis

Bile acid synthesis is tightly regulated since excess bile acids can be harmful to the liver, leading to cholestatic liver disease. The hydrophobic bile acids, such as CDCA, LCA and

DCA, are highly toxic. Since CYP7A1 is the rate-limiting enzyme in bile acid synthesis, its regulation is the primary mechanism for regulating bile acid synthesis. The regulation of CYP8B1 is discussed in detail in the next section.

a. Feedforward Regulation by Cholesterol.

Cholesterol (or oxysterol) stimulates bile acid synthesis (feedforward stimulation), by activating Lxrα which stimulates mouse Cyp7a1 expression to convert excess cholesterol to bile acids [187]. Lxrα-/- mice, show normal levels of Cyp7a1 mRNA, which are not

stimulated by a high-cholesterol diet, as in the wild-type mice, leading to a massive

accumulation of liver cholesterol [198]. Transfection assays using rat or mouse Cyp7a1

promoter/reporter constructs indicated the presence of a LXR response element that

functionally activates transcription in response to dietary cholesterol and/or oxysterol

agonists. Lxr bound to a DR-4 site, in the bile acid response element I (BARE-I) region

on the rat Cyp7a1 gene and stimulated rat Cyp7a1 transcription [187]. However, LXRα

has much less effect on hamster and human CYP7A1, which lack DR-4 motifs [188].

Thus unlike humans, rats and mice are unique in their ability to convert excess

cholesterol into bile acids, which prevents the development of gallstones and

atherosclerotic diseases.

b. Feedback Regulation by Bile Acids.

Bile acids returning back to the liver via the enterohepatic circulation inhibit bile acid

synthesis. Blockage of the eneterohepatic circulation of bile acids by either biliary

diversion or treatment with bile acid sequestrants, leads to an increase in the rate of bile

acid synthesis and CYP7A1 activity by 3- to 4-fold. On the other hand, an increase in

enterohepatic circulation by intraduodenal infusion of bile acids inhibits the rate of bile

acid synthesis to the normal level. Mechanisms that have been proposed for the bile acid regulation of CYP7A1 and CYP8B1 gene transcription are illustrated in Fig.6. i) SHP dependent mechanism of bile acid suppression:

Bile acid activated Fxr induces the negative nuclear receptor Shp which interacts with

Lrh-1 and prevents its interaction with coactivators leading to the down regulation of the

Cyp7a1 gene transcription [105, 230]. Fxr-/- mice show low levels of Shp expression, significant increase of Cyp7a1 mRNA levels and lack of bile acid inhibition of Cyp7a1 expression [114], supporting the role of FXR and SHP in bile acid negative feedback regulation of bile acid synthesis.

However bile acid feeding to Shp knockout mice reduces Cyp7a1 mRNA levels similar to the wild-type mice, suggesting the existence of mechanisms independent of the

FXR/SHP/FTF pathway in the bile acid feedback suppression of CYP7A1 [165, 166].

Some of the SHP independent mechanisms of bile acid suppression of CYP7A1 are discussed below. ii) SHP independent mechanisms of bile acid suppression:

FXR/FGF-19/FGFR4 pathway:

Fibroblast growth factor (FGF) is a regulator of adult organ homeostasis, embryonic development and wound healing; with FGF receptor 4 (FGFR4) being significantly expressed in mature hepatocytes [231, 232]. It was shown that mice lacking Fgfr4 exhibited depleted gallbladders, and elevated fecal bile acids, bile acid pool size, and expression of Cyp7a1 [233]. It has been shown that agonists of FXR induced expression

Fig.6.

Figure 6. Bile acid feedback inhibition pathways.

In the small heterodimer partner (SHP) dependent pathway, bile acid activated farnesoid

X receptor (FXR) [which heterodimerizes with retinoid X receptor (RXR) to bind the

SHP promoter] induces SHP, which then inhibits α-fetoprotein transcription factor (FTF) and hepatocyte nuclear factor 4α (HNF4α) transactivation of CYP7A1/8B1 genes. In

SHP independent pathways, bile acids activate protein kinase C (PKC) and inflammatory cytokine (IL-1β and TNFα) release from Kupffer cells, and initiate the MAPK/JNK pathway. The JNK pathway can activate c-Jun which can in turn induce SHP. The downstream target of JNK (possibly HNF4α), the direct effects of JNK or c-Jun on

CYP7A1/8B1 are not certain. FXR also induces fibroblast growth factor 19 (FGF19) and its receptor (FGFR4) which leads to activation of JNK. Bile acids can also activate the pregnane X receptor (PXR) which interferes with the recruitment of peroxisome proliferator activated receptor γ coactivator- 1α (PGC-1α) to HNF4α on the bile acid response element (BARE) of the CYP7A1/8B1 chromatin. MEKK1, MAP kinase kinase

1 and MKK4, MAP kinase kinase 4.

of FGF19 at the transcription level in primary human hepatocytes concurrent with

repression of CYP7A1 [234]. In addition, the treatment of cultured hepatocytes with

FGF19 activated the JNK signaling pathway, and repressed CYP7A1 and bile acid synthesis [234]. It was suggested that bile acid activated FXR inducing FGF19 that acts though FGFR4 and activates JNK signaling to mediate feedback repression of bile acid

synthesis [234]. Recently, transgenic mice overexpressing a constitutively active human

FGFR4 (CahR4) in hepatocytes were generated [235]. These mice showed reduced fecal

bile acid excretion, bile acid pool size and Cyp7a1 expression compared to wild-type

mice. Levels of phosphorylated Jnk were lower in Fgfr4–/– livers, whereas were increased

in CahR4 livers. However, cholate still strongly induced phospho-Jnk in Fgfr4–/– livers

[235]. These results suggest that FGFR4 signaling regulates CYP7A1 expression via the

JNK pathway but FGFR4 is not required for activation of JNK signaling by bile acids.

PXR-mediated pathway:

In a study in Shp-/- mice it was shown that activation of the Pxr pathway by PCN can inhibit Cyp7a1 [165]. As mentioned earlier, PXR is a bile acid (LCA) inducible nuclear receptor. Rifampicin, a human PXR agonist, is shown to reduce CYP7A1 and SHP

mRNA expression in primary human hepatocytes and inhibited CYP7A1 reporter activity

[120]. Rifampicin activated PXR was shown to interact with HNF4α and with PGC-1α

[120, 121]. Thus, the activation of PXR by rifampicin promotes PXR interaction with

HNF4α and PGC-1α, and blocks PGC-1α activation of HNF4α and results in inhibition

of CYP7A1 gene transcription.

Cytokine/JNK mediated pathway:

Bile acids induce inflammatory cytokines, such as tumor necrosis factor α (TNFα) and interleukin-1β (IL-1β) from the Kupffer cells (hepatic macrophages), which cross the

sinusoidal membrane and act on the surrounding hepatocytes via their high affinity

receptors on the hepatocytes to suppress the CYP7A1 mRNA expression [4, 236, 237].

Upon feeding a bile acid-containing diet, mice display a rapid induction of TNFα and IL-

1β mRNA expression by Kupffer cells and a concomitant repression of Cyp7a1; while

treatment with Rosiglitazone, a known inhibitor of cytokine activation, blocks both the

induction of hepatic TNFα and IL-1β mRNA expression and suppression of Cyp7a1 by

dietary bile acid feeding [4]. Cytokines have also been shown to inhibit CYP27A1 [238].

Studies have shown that both cytokines [218, 239] and bile acids [218, 240] can activate the JNK pathway. Earlier a study showed that bile acids can activate protein kinase C (PKC), which in turn can activate the JNK pathway [241, 242]. Bile acid or

cytokine activation of JNK has been shown to suppress CYP7A1 gene transcription [218,

237]. However, the events downstream of JNK have not yet been very well established.

One report showed that the BARE-II region, which contains a HNF4α binding site, is

essential in mediating the cytokine suppression of CYP7A1; indicating that nuclear

receptors may be the downstream targets of the cytokine activated signaling pathways

that regulate CYP7A1 gene expression [237]. The role of JNK in regulating HNF4α expression/function to in turn regulate the CYP7A1 gene expression is yet to be determined.

7. Regulation of CYP8B1

CYP8B1 (Sterol 12α-hydroxylase) is an intronless microsomal cytochrome P450 gene expressed exclusively in the liver [243]. Ishida et al. first purified the CYP8B1 enzyme by using anion exchange high-performance liquid chromatography (HPLC) [244]. The purified enzyme required cytochrome b5 and NADPH-cytochrome P450 reductase for full activity [244]. A full-length cDNA clone encoding rabbit CYP8B1 was later isolated

[245]. CYP8B1 has a high homology (43%) with prostacyclin synthase (CYP8A1). So far, the CYP8B1 gene has been cloned in several species like human [243], mouse [243], rat [246], rabbit [245], pig [247] etc. The cDNA of CYP8B1 consists of a 1509-base pair open reading frame encoding 503 amino acid residues which have characteristic conserved domains for heme, steroid and O2 binding. Therefore CYP8B1 was also categorized into the cytochrome P450 family. The cDNA coding region of CYP8B1 shows a high degree of identity among the different species, but the promoter sequences of CYP8B1 shows only a 21% homology among the different species. Mitochondrial

CYP8B1 activity is not detected in adults, however CYP8B1 activity was found both in the microsomal and mitochondrial fractions of the fetal human liver CYP8B1 [248].

CYP8B1 is required for CA synthesis and regulates the hydrophobicity of the bile acid

pool.

CYP8B1 activity exhibits a diurnal rhythm. A marked circadian rhythm

(maximum at 13:00-16:00 and minimum at 1:00) is observed both on the mRNA level

and the activity of CYP8B1 [249]. This rhythm is shifted from that of CYP7A1 showing

a maximum at 22:00 and a minimum at 10:00, and this shift might oscillate the

CA/CDCA ratio, which is increased in the late afternoon and decreased at midnight

[249]. The circadian rhythm of the liver clock gene albumin D-site binding protein (DBP) is said to mediate the stimulation of CYP7A1 [250, 251] and CYP8B1 (Zhang, M PhD dissertation). While DEC2, another circadian clock gene, can inhibit CYP7A1, overwhelming the DBP binding site and also decreases the transcription of CYP8B1

[252]. Glucocorticoid or insulin may also play a role in the diurnal regulation of the bile acid synthesis. Dexamethasone treatment reduces CYP8B1 mRNA but increases

CYP7A1 mRNA levels [246]. Insulin can also suppress the expression of rat Cyp8b1

[246, 253]. Streptozotocin induced diabetic rats show a higher level of Cyp8b1 and the elevated Cyp8b1 can be suppressed by insulin administration [253]. The rhythm of

CYP8B1 is the inverse of the circadian variation of serum insulin level [253].

CYP8B1 mRNA levels and enzyme activity has been shown to be induced by starvation, unlike the CYP7A1 mRNA levels and enzyme activity, which are repressed

[246]. Feeding with clofibrate, a PPARα ligand, could increase Cyp8b1 mRNA levels but suppressed that of Cyp7a1 [246]. A later study revealed that the response to starvation might be mediated through PPARα. Fasting for 24 hours or administration of clofibrate or WY-14,643 (PPARα ligand) did not change the Cyp8b1 activity and mRNA levels in PPARα null mice, whereas a significant increase was seen in wild-type animals

[176]. Co-transfection with PPARα expression plasmids in HepG2 cells induced a 2.5- fold increase in the rat Cyp8b1 promoter activity in a ligand-dependent manner [176].

PPARα was found to directly bind very weakly to a direct repeat (DR-1) motif in the rat

Cyp8b1 promoter [176]. CYP8B1 expression is also regulated by thyroid hormone [254].

Treatment of intact rats with L-thyroxine causes a 60% reduction in the enzyme activity and a 50% reduction in Cyp8b1 mRNA [254]. Thyroidectomy in rats causes a 2-fold increase in the enzyme activity with a 4-fold increase in Cyp8b1 mRNA in the liver

[254].

Similar to the CYP7A1 gene, the CYP8B1 gene is subjected to a negative feedback regulation by bile acids like CA, CDCA and DCA [16]. Cholestyramine feeding or bile fistula can remove bile acids from the intestine and stimulate CYP8B1 [16, 246]. CYP8B1 is differentially regulated by both FTF and HNF4α, which bind to overlapping sites on the promoter. The putative bile acid response element (BARE) on the rat Cyp8b1 promoter has an HNF4α binding site embedded in two overlapping FTF binding sites

[135, 150], while the human CYP8B1 promoter has one HNF4α and one FTF binding site overlapping each other [107] Bile acid/FXR induced SHP interacts with FTF and inhibits rat Cyp8b1 gene transcription [135, 149, 150], but interacts with HNF4α and inhibits the human CYP8B1 gene [107]. In reporter assays, co-transfection with FXR enhances bile acid inhibition of the rat and human CYP7A1 gene [104], but has no effect on human

CYP8B1 gene transcription [107]. Also, bile acid feeding to Shp-/- mice reduces Cyp8b1 mRNA levels as in wild type mice [165, 166]. These results suggest that the FXR/SHP pathway may not be the major mechanism for bile acid inhibition of CYP8B1 gene transcription and other SHP independent pathways may exist. Interestingly however, long term feeding of a bile acid containing diet to the Shp-/- mice, led to reexpression of

Cyp8b1, which is postulated to prevent toxic liver damage by increasing the

hydrophilicity of the bile acid pool [255]. Rifampicin induced PXR (a bile acid receptor) has been shown to inhibit CYP8B1 expression [121].

In contrast to the up-regulation of Cyp7a1 by Lxrα and cholesterol, cholesterol feeding decreases both transcription and activity of Cyp8b1 in rats [16]. Suppression of

CYP8B1 by cholesterol may be significant in maintaining cholesterol homeostasis, since decreased CA production would ensure a decrease in the intestinal absorption of cholesterol, as CA is the most efficient bile acid in absorbing cholesterol. Cholesterol feeding also reduced mRNA levels for Srebp-1 but not for Srebp-2 in rat livers [189]. Co- transfection of SREBP-1a and -1c stimulated Cyp8b1 promoter activity, while SREBP-2 did not have any effects [189, 209]. Several functional SREs and E-box motifs have been identified in the rat Cyp8b1 promoter [189, 209].

8. Mitogen Activated Protein Kinase (MAPK) Pathways

Cells respond to extracellular stimuli through a series of signal transduction events that occur across the cell membrane, through the cytoplasm into the nucleus. The MAPK signaling pathways play a key role in the regulation of cellular processes such as mitogen-induced cell cycle progression through the G1 phase, regulation of embryonic development, cell movement and apoptosis and cell and neuronal differentiation [256,

257]. The evolutionarily conserved MAPK pathways are organized in three protein- serine/threonine kinase modules consisting of a MAP kinase (MAPK), an activator of

MAP kinase (MAP kinase kinase, MAPKK or MEK) and a MAP kinase kinase kinase

(MAPKKK or MEK kinase, MEKK) [258]. A MAPKKK that is activated by extracellular stimuli phosphorylates a MAPKK which in turn phosphorylates and activates a MAPK. The MAPK can then phosphorylate transcription factors and regulate gene expression [258].

The MAPK signaling cascade has been evolutionarily well-conserved from yeast to mammals. Atleast three distinct MAPK signal transduction pathways have been identified in mammals and named after the particular MAPK associated with it. These include the JNK/stress-activated protein kinase (SAPK), ERK and the p38 MAPK [259].

The specific MEKKs and MEKs known to activate each of these MAP kinases are shown in Fig. 7. The MAPK pathways are linked to many G-protein coupled receptors and

Receptor Tyrosine Kinases on the cell surface [260]. Thus, they can be activated by a host of different factors. The ERK pathway is typically stimulated by growth and differentiation factors like epidermal growth factor, platelet-derived growth factor, nerve growth factor, hormones and regulates cell proliferation and differentiation [261]. While the JNK and p38MAPK pathways are response to cellular stress and inflammation and involved in cell differentiation and apoptosis [262]. They are activated by lipopolysaccharides (LPS), cytokines like IL-1 and TNF-α, ionizing and ultraviolet radiation, the translation inhibitors cyclohexamide and anisomycin, tumor promoters, heat shock and hyperosmotic stress [262].

The different MAPK pathways show a significant degree of cross talk between each other. The cells can tightly regulate the activity of the MAPK enzymes by using different combinations of the MEKs, the protein phosphatases that can differentially Fig.7.

Figure 7. Activation of different MAPK signaling cascades by different extracellular stimuli.

Different extracellular stimuli (mitogen) activate different G-protein coupled receptors or

Receptor Tyrosine Kinases on the cell membrane, which then activate one or many of the intracellular MAPK signaling cascades. The ERK, JNK and p38MAPK cascades all contain the same series of three kinases. A MAPKK Kinase or MEK kinase (MEKK) phosphorylates and activates a MAP kinase kinase (MEK), which in turn phosphorylates and activates a MAP kinase (MAPK). The MAPK finally phosphorylates transcription factors and regulates gene expression.

dephosphorylate and inactivathe either the ERK, JNK or p38 MAPK enzymes and by controlling the subcellular localization of each enzyme to elicit the appropriate physiological response [258].

9. Cholestatic liver disease

Cholestasis occurs due to an imbalance in bile salt transport, which occurs as a consequence of malfunctioning hepatic and intestinal bile acid transporters. Cholestasis can result from hereditary defects in bile salt transport, such as in progressive familial intrahepatic cholestasis (PFIC) type 1, 2 and 3; benign recurrent intrahepatic cholestasis

(BRIC) and Dubin-Johnson syndrome [47, 263]. Acquired defects in bile salt transport result from direct inhibition of function or expression of the transporters and include drug induced cholestasis, intrahepatic cholestasis of pregnancy and sepsis associated cholestasis [47, 263]. The changes in transporter expression that result in cholestasis are secondary adaptive changes that result from the impairment of bile flow and retention of biliary constituents within the hepatic parenchyma [264, 265].

Cholestasis is a common complication in patients with sepsis [266-268].

Endotoxin or sepsis induced cholestasis is more common in gram-negative infections and is caused by the effect of the bacterial endotoxin called lipopolysaccharide (LPS), a component of the gram-negative bacterial cell wall. LPS activates macrophages by binding to CD14 receptors [269]. In the liver, the Kupffer cells are the primary targets of

LPS. Kupffer cells activate innate immunity in response to the entrance of bacteria and

bacterial products from the intestinal tract. They produce cytokines in hepatic

inflammatory conditions such as sepsis and alcoholic, viral and autoimmune hepatitis.

Activated Kupffer cells begin the acute-phase response to prevent tissue damage, eliminate the infectious agent and activate repair processes in the cells. Acute phase

response is initiated by production of proinflammatory cytokines like IL-1β, TNFα, and

IL-6 and anti-inflammatory cytokines like IL-10 [269]. These cytokines then bind to receptors on the sinusoidal membrane of the hepatocytes and initiate a complex pathway of intracellular signal transduction reactions, leading to a cascade of events such as upregulation of inducible nitric oxide (NO) synthase, production of reactive oxygen species (ROS), lipid peroxidation, changes in redox potential etc. The hepatocytes respond to these signals by altering gene expression to restore homeostasis [269].

Proinflammatory cytokines cause intrahepatic cholestasis [270], hypercholesterolemia [271, 272], altered lipid metabolism and changes in cytochrome

P450 expressions [273, 274]. The cytokine inhibition of the genes encoding several hepatobiliary transporters [108, 275-279] is the mechanism of cholestatic liver injury.

Among the bile acid transporters, the expression of canalicular organic anion transporter

Mrp2 is decreased profoundly to 11% of controls in the LPS administered rats [269, 278,

280, 281]. MRP is regulated by nuclear receptors, constitutive androgen receptor (CAR,

NR1I3), PXR and RAR [122, 277]. Expression of Rxr is strongly downregulated during

endotoxemia in hamsters and this may lead to the downregulation of Mrp2 [282].

Administration of LPS, TNFα or IL-1β also causes a decrease in the mRNA levels of the hepatocellular bile salt uptake system Ntcp [270]. The downregulation of Ntcp in

endotoxin induced cholestasis can be explained by decreased binding of Hnf1α or

Rar/Rxr heterodimer to the Ntcp gene promoter [277, 283, 284]. Apart from the decrease in RXR expression, IL-1β induced JNK was shown to phosphorylate RXR and the phosphorylated RXR showed decreased heterodimerization capability with RAR [239].

Reduced canalicular secretion of bile salts is caused by a 32% reduction in Bsep mRNA levels [278] and a concomitant decrease in protein [279] in endotoxin-treated rats [269].

TNFα or IL-1β can repress the activity of the ASBT promoter in colonic carcinoma

(Caco-2) and intestinal epithelial cell-6 cells [276]. Inflammation has been shown to be associated with up-regulation, phosphorylation, and nuclear translocation of c-fos, which

then represses ASBT promoter activity via binding to the 3' AP-1 element by a c-fos/c-

jun heterodimer [276]. However, Mrp3 expression is induced after bile duct ligation in wild type mice, possibly to increase sinusoidal bile acid excretion [275].

10. Proinflammatory cytokine mediated gene regulation in the liver

Cytokines play several important roles in the liver such as liver development during early

embryogenesis, hepatic acute phase response, liver regeneration and fibrosis and in liver

cancer and metastasis [285]. Post-infection tissue injury leads to alterations in the

function of organ systems, which is termed as the acute phase response (APR) [286]. The

rates of synthesis of many plasma proteins are changed during acute phase conditions

(acute phase proteins, APPs). Since majority of the plasma proteins are synthesized in the

liver, the hepatocytes are targets of cytokines during APR [287].

The cytokines of APR are classified according to their functions into two major groups- the IL-1 type cytokines and the IL-6 type cytokines. The IL-1 type cytokines include IL-

1α, IL-1β, TNFα and TNFβ. They stimulate positive type-1 APPs such as C-reactive protein, serum amyloid A and haemopexin and inhibit APPs such as albumin. The IL-6 type cytokines include IL-6, IL-11, leukemia inhibitory factor (LIF) and ciliary neurotropic factor (CNTF). They stimulate type-2 APPs such as fibrinogen, α1- antitrypsin, haptoglobin and ceruloplasmin [288].

The major proinflammatory cytokines in the liver, namely TNFα, IL-1β and IL-6, bind to their respective receptors on the basolateral membrane of hepatocytes and trigger complex signal transduction pathways that lead to activation and translocation of transcription factors, which mediate the changes in gene transcription. The intracellular signaling induced by IL-1 type cytokines involves the transcription factors NF-κB and

AP-1 while that induced by IL-6 type cytokines involves the signal transducer and activator of transcription 3 (STAT3) transcription factors [285]. The MAPK pathways can be induced by both the types of cytokines [289].

The Rel/NF-κB proteins are a family of transcription factors consisting of homo and heterodimers characterized by the highly conserved DNA-binding/dimerization domains called the Rel homology region [290]. The members of the family include p50, p52, p65, c-Rel, and RelB. However, the best-characterized members are the p50/p65 heterodimer and the p50/p50 homodimer complexes [291]. NF-κB proteins remain associated with an inhibitory protein called inhibitory κB (IκB) in the cytoplasm [292]. NF-κB activation involves the activation of the IκB kinase (IKK), which rapidly phosphorylates IκB

leading to the ubiquitination and proteasomal degradation of the latter [292]. The nuclear localization site of NF-κB is then exposed, causing the translocation of NF-κB into the nucleus where it binds the DNA of target genes and activates gene transcription [292].

AP-1 transcription factors consist of homodimers and heterodimers of the basic region- leucine zipper (bZIP) proteins that belong to the Jun (c-Jun, v-Jun, JunB, JunD), Fos (c-

Fos, v-Fos, FosB, Fra1, Fra2) and the related activating transcription factor (ATF2,

ATF3/LRF1, B-ATF) subfamilies. The exact subunit composition depends on the extracellular stimulus and the MAPK signaling pathway that is activated (JNK, ERK, etc.). Upon stimulation by the MAPK, regulation of AP-1 activity occurs by activating transcription of these genes as well as through phosphorylation of existing Jun and Fos proteins at specific serine and threonine sites [293-296]. Jun proteins can form dimers

(Jun-Jun) that are stable bind AP-1 DNA consensus sequences, while Fos proteins, on the other hand, cannot form stable homodimers. Instead, they mediate gene expression by forming heterodimers with various Jun proteins. These heterodimers are more stable than the Jun-Jun dimers and possess higher DNA binding activity [297]. Thus, the differential makeup and composition of the different subunits influence and modulate the binding affinity of AP-1 proteins and their subsequent regulation of gene transcription [298].

STAT3 exists in an inactive state in the cytoplasm. IL-6 binds to the specific cell surface receptor, which activates tyrosine kinases causing phosphorylation of the STAT3 [299,

300]. Janus Kinase (JAK) is a key player in the cytokine signaling pathway, and when activated, binds to the STAT SH2 (Src homology domain-2) domain resulting in phosphorylation of STAT3 [299, 301]. Upon activation, STAT3 can form homodimers or

STAT1 and STAT3 heterodimers [299]. Sometimes, the STAT proteins can also form tetramers [299]. After dimerization, STAT3 translocates to the nucleus, and activates specific target genes [300, 301]. Dephosphorylation of the STAT proteins occurs in the nucleus after approximately 15 minutes and serves as an important signal for its transport back into the cytoplasm in preparation for the next round of signaling [299, 300].

The cytokine induced transcription factors bind to the specific consensus sequences on target genes and stimulate gene transcription. However, these factors can also cause a downregulation in the transcription of several genes. The exact mechanism for the downregulation is not known. It has been speculated that the transcription factors can compete with other stimulatory transcription factors for binding coactivators such as CBP or SRC-1. This may lead to a decrease in the coactivator levels required to stimulate gene transcription. Also, the cytokine induced transcription factors may interact directly with other transcription proteins and prevent them from binding DNA, which can result in the downregulation of the target genes. It is also likely that cytokine induced transcription factors may induce the transcription of inhibitory proteins, such as corepressors resulting in the downregulation of the target genes. CYP7A1 is known to be inhibited by cytokines; however, the mechanism of this suppression is unknown. It would be interesting to determine the likely mechanism by which cytokine induced transcription factors may downregulate CYP7A1 gene transcription.

11. Hypothesis, Specific aims, Approaches and Significance

CYP8B1 is an important enzyme in bile acid biosynthesis, however very little is known about the regulation of the human CYP8B1 gene. During cholestasis there is an excessive accumulation of toxic bile acids in the liver and circulation. Inhibiting bile acid synthesis during cholestasis is a hepatoprotective mechanism of the body. As discussed earlier, inflammatory conditions, that release cytokines, are associated with cholestatic liver diseases [270]. Studies have shown that cytokines can inhibit CYP7A1 gene transcription

[4, 236, 237]. Also, the JNK pathway has been shown to be activated by bile acids and cytokines, leading to the down regulation of CYP7A1 [237]. No studies have been done so far to determine the effect of cytokines on CYP8B1 gene transcription.

Hypothesis: I hypothesize that cytokines may play a critical role in bile acid feedback inhibition of the human CYP8B1 gene, and that the MAPK/JNK pathway might mediate the bile acid inhibition. I further hypothesize that the effect of cytokines on CYP8B1 may occur through the regulation of HNF4α, which is the most important regulator of

CYP8B1 [107].

To test my hypothesis the following specific aims and experimental approaches are designed:

Specific Aim #1. To determine the effect of cytokines on CYP8B1 and study the signal transduction pathway, which mediates the effect. a. To determine the effect of cytokines on CYP8B1 gene transcription in human liver cells.

Approach: Primary human hepatocytes will be used as a model to study the effects of IL-

1β on CYP8B1 mRNA levels by quantitative real time PCR. Also, the effect of IL-1β on the human CYP8B1/luciferase reporter plasmid activity in the human hepatoma cell line i.e. HepG2 cells, will be assayed by transient transfection assay. b. To delineate the signal transduction pathway that mediates the cytokine effect on the

CYP8B1 gene.

Approach: The specific inhibitors of the MAPK pathways will be used in transient transfection assay, to determine which pathway mediates the IL-1β effect on the human

CYP8B1/luciferase reporter. Also, downstream targets of the JNK pathway, JNK and c-

Jun will be ectopically expressed to study their effect on the human CYP8B1/luciferase reporter.

Specific Aim #2. To elucidate the molecular mechanism of cytokine regulation of

CYP8B1 gene transcription. a. To determine the cytokine response element on the CYP8B1 promoter.

Approach: Human CYP8B1/luc reporter deletion constructs and site-directed mutagenesis constructs, will be used in transient transfection assay to identify the region of the CYP8B1 promoter responsive to IL-1β. b. To reveal a possible mechanism by which HNF4α may mediate the cytokine

suppression of CYP8B1.

Approach: IL-1β treated HepG2 cell extracts will be analyzed by immunoblot analysis to

determine the effect on HNF4α protein. The effect of IL-1β on HNF4α DNA binding

activity will be tested with in vitro electromobility shift assay (EMSA) and in vivo, chromatin immunoprecipitation (ChIP) assays. Effect of the specific inhibitors of the

MAPK pathways will also be tested. Also, to prove if a post translational modification of

HNF4α protein is involved in mediating the IL-1β effect on CYP8B1, constitutively active JNK enzyme will be used to test if JNK phosphorylates HNF4α in an in vitro kinase assay. c. To establish the role of c-Jun in mediating the cytokine suppression of CYP8B1.

Approach: The effect of IL-1β on c-Jun mRNA levels will be determined by quantitative real time PCR in human primary hepatocytes. Effect of IL-1β on cJun and phosphorylation of c-Jun levels will also be measured. The possibility that c-Jun and

HNF4α may interact will be tested via coimmunoprecipitation (Co-IP) assay and the effect of IL-1β on this interaction will be determined. GST pull down assay will be done to confirm the interaction of c-Jun and HNF4α. EMSA and ChIP assay will be done to test if c-Jun interferes with the binding of HNF4α to the CYP8B1 promoter.

Specific Aim #3. To determine the effect of cytokines on SHP expression and to observe if the c-Jun mediated suppression mechanism can regulate CYP7A1. a. To determine the effect of cytokines on SHP expression.

Approach: The effect of IL-1β on the human SHP/Luc reporter activity will be measured in HepG2 cells by transient transfection assay. Since the SHP promoter has a putative

DR-1 site (-563TGGACAGTGGGCA-551), the effect of HNF4α on the SHP/luc reporter activity will be tested in the non-liver human embryonic kidney cell line HEK 293. Also,

the HNF4α binding to the DR-1 site on the human SHP promoter will be tested by

EMSA. b. To observe if the c-Jun mediated suppression mechanism can regulate CYP7A1.

Approach: c-Jun and dnc-Jun will be overexpressed along with HNF4α and coactivator

PGC-1α, to determine the combined effect on the CYP7A1/luc reporter activity. ChIP assay will be done to test if c-Jun interferes with the binding of HNF4α to the CYP7A1 promoter. Small interference RNA (siRNA) will be used to knockdown HNF4α in

HepG2 cells and the cells will be treated with IL-1β to determine if CYP7A1 mRNA levels can be further suppressed in the absence of HNF4α.

Significance: This study will help us understand the regulation of CYP8B1 during inflammatory conditions and cholestatic liver diseases. Evidences indicate that CYP8B1 has a significant effect in regulating the hydrophobicity of the bile acid pool which subsequently influences cholesterol homeostasis and bile acid synthesis [302, 303]. Since studies in the Shp-/- mice indicate that Cyp8b1 responds to bile acids [165, 166], my study may shed light on an alternate mechanism for bile acid inhibition of CYP8B1.

Inhibition of CYP8B1 may play an important role in ameliorating disease conditions such as hypercholesterolemia and cholesterol gallstones by decreased intestinal cholesterol uptake due to the lack of CA.

CHAPTER II

MATERIALS AND METHODS

1. Cell Culture

Cell line culture:

The human hepatoma cell line (HepG2, HB8065) and the embryonic kidney cell line

(HEK293, CRL-1573) were obtained from American Type Culture Collection (ATCC,

Manassas, MA). The cells were stored in a –150oC freezer as aliquots. The cells were

thawed quickly in a 37oC water bath and grown in complete growth media consisting of a

1:1 mixture of Dulbecco’s modified Eagle’s medium and F-12 (Life technologies Inc.,

Rockville, MD) supplemented with 100 units /ml penicillin G/streptomycin sulfate

(Celox Corp., Hopkins, MN) and 10% (v/v) heat-inactivated fetal calf serum (Irvine

Scientific, Santa Ana, CA).

Cells were cultured in 37°C in a humidified, 5% CO2 incubator. Medium was

changed 2-3 times a week. When cells reached confluence, 0.5% trypsin in EDTA was

used to pass the cell. Briefly, the media was aspirated from the cells and 4 ml of trypsin

solution was added to cover the bottom of the T 75-cm2 flask. Cells were incubated for

1–2 min or until the cells began to detach from the flask bottom; 4 ml of complete

medium was added to stop the trypsin digestion. To distribute the cells evenly, the

suspension was pipetted up and down at least 20 times to make a single cell suspension,

64 65

and then transferred evenly to 3 to 6 T 75-cm2 flasks. Then 10 – 20 ml of complete

medium was added to the flasks and they were returned to the incubator to continue the

culture.

If the cells needed to be frozen and stored, the cell suspension was centrifuged at

1200 g for 5 min and resuspended in 50% FBS and 10% DMSO to a concentration of 2 X

106cell/ml. One ml was added to the cryo-tube and the temperature was gradually

lowered to -80°C over 4 h and then transferred to -150°C.

Primary cell culture:

Primary human hepatocytes isolated from human donors (Case # HH1088, female, Case

# HH1165, pediatric female and Case # HH1209, 30 yr female) were obtained in 6 well

plates, through the Liver Tissue Procurement and Distribution System (LTPADS) of

NIH (Dr. Steven Strom, University of Pittsburgh, Pittsburgh, PA). Cells were

maintained in HMM modified Williams E medium (Cambrex (Clonetics), East

Rutherford, NJ) supplemented with 10-7 M of insulin and dexamethasone and a mixture

of gentamicin sulfate and amphotericin B (All supplements were obtained from

Cambrex, East Rutherford, NJ). The cells were typically treated on the day of receiving

and used within 24 h after receiving.

2. Plasmids

Reporters:

The wild type human CYP8B1/Luc reporter p8B1–514/+300/Luc, 3’-deletion mutants of the CYP8B1/Luc reporter (p-514/+248/Luc, p-514/+220/Luc, p-514/+200/Luc, p-

514/+137/Luc, p-514/+76/Luc), and the p8B1mutant/Luc reporter that has the HNF4α binding site mutated were previously constructed [107]. Mouse HNF4α/Luc reporter

(pDGT43mHNF4pro744) containing 744 bp 5’-upstream sequence was provided by Dr.

Todd Leff (Wayne State University, Detroit, MI). The human SHP reporter (hSHP/Luc) containing 2277 bp 5’upstream sequence was provided by David Moore (Baylor College of Medicine, Houston, TX). The human CYP7A1/Luc reporter construct ph-298/Luc was constructed previously [304].

Expression Vectors:

The expression vectors were obtained as follows-

JNK (pcDNA3.1neo-HA-46γ, Dr. James Woodgett, Ontario Cancer Institute, Toronto,

Canada), dominant negative JNK (dnJNK, pcDNA3 FlagJNKapf, Dr. Roger Davis,

Univ. of Massachusetts Medical School, Worcester, MA), c-Jun (originally from Dr.

Inder Verma, The Salk Institute, San Diego, CA, subcloned into pcDNA3), dominant negative c-Jun (dnc-Jun, pcDNA3.1HisTAM67, Dr. Nancy Colburn, NCI, Frederick,

MD), dominant negative HNF4α (dnHNF4α, pDGT23.1dnHNF4α-pcDNA3, Dr. Todd

Leff, Wayne State University. Detroit, MI), HNF4α (originally from Dr. William Chin,

Lilly Research Laboratories, Indianapolis, IN, subcloned to pcDNA3), FTF (pCI FTF,

Dr. L. Belanger, Le Centre de Recherche en Cancerologie de l'Universite Laval, Quebec,

Canada) and PGC-1α (pcDNA3- HA-PGC-1α Dr. A. Kralli, The Scripps Research

Institute, La Jolla, CA).

The GST expression vectors were obtained as follows- pGEX-4T-1 encoding the Glutathione S- transferase (GST) protein alone (Amersham

Pharmacia Biotech) and pET23a-GST-HNF4α encoding a fusion protein of GST with full length human HNF4α ( Dr. Todd Leff, Wayne State University, Detroit, MI).

3. Preparation of Competent Cells

The competent DH5α bacteria were prepared by the calcium chloride method. A single colony (2-3 mm in diameter) was picked from a plate freshly grown for 16-20 h at 37oC and transferred into 5 ml of LB broth or SOC medium and incubated over night. 100 μl of the overnight culture was then inoculated into 100 ml medium and incubated for 3 h at 37oC with vigorous shaking (300 cycles/min in a rotary shaker). For efficient transformation, it was essential that the number of viable cells should not exceed 108 cells/ml. To monitor the growth, the O.D.600 of culture was determined every 20-30 min

until it reached 0.3-0.4. The cells were transferred to sterile, disposable, ice-cold 50-ml

polypropylene tubes (Falcon 2070). The cultures were cooled to 0oC by storing the

tubes on ice for 10 min. The cells were recovered by centrifugation at 4000 rpm for 10

min at 4 oC in a Sorvall GS3 rotor (or its equivalent). The media was decanted from the

cell pellets. The tubes were kept in an inverted position for 1 min to allow the last traces

of media to drain away. Each pellet was resuspended in 10 ml of ice-cold 0.1M CaCl2

and stored on ice. The cells were recovered by centrifugation at 4000 rpm for 10 min at

4oC in a Sorvall GS3 rotor (or its equivalent). The fluid was decanted from the cell

pellets. Each pellet from each of the 50 ml original culture was resuspended in 2 ml of

ice-cold 0.1 M CaC12 and incubated for at least 1 h. Glycerol was added quickly to 25%

of the total volume and dispensed into chilled, sterile microfuge tubes. The competent cells were immediately snap-freezed by immersing the tightly capped tubes in liquid nitrogen. The tubes could be stored at -80oC until needed. For most transformations

purposes, 100 μl aliquots of the competent cell suspension were used.

4. Bacterial Cell Transformation

For transformation an aliquot of the competent DH5α cells was removed from -80oC and thawed by leaving it on ice for 30 min. After the cells were thawed, 1μl of the plasmid DNA (10-100ng) was added to the competent DH5α cells and the mixture was kept on ice for 1 h. Following that, the mixture was heat shocked at 42oC for 2 min and

then cooled on ice for 10 min. 800 ml of LB broth or SOC medium was then added to

the mixture and pipetted up and down, and the resulting mixture was incubated at 37oC for 1 hr. After that, the mixture was centrifuged at 4000 rpm for 2 min and 800 μl of the supernatant was removed and discarded. The pelleted cells were resuspended in the remaining medium. These cells were then spread aseptically on a previously prepared

LB agar plate (containing the resistance antibiotic i.e. Ampicillin [50μg/ml LB Agar])

The LB agar plate was incubated upside down in a 37oC incubator for 16 h (overnight).

Colonies of the plasmid containing DH5α cells grew on the LB plate the following morning.

5. Preparation of Plasmid DNA

For all the plasmid DNA preparations, LB broth/medium containing 50μg/ml of ampicillin was used to select for cells containing the plasmids of interest, which have a resistance gene to ampicillin.

Large Scale preparation or Maxiprep:

For large-scale plasmid preparation, Nucleobond AX 500 cartridges (Clontech, Palo Alto,

CA) were used. All the buffers were prepared according to the manufacture’s instruction.

A single bacterial colony from the transformed LB plate was inoculated into 2 ml of LB medium, with shaking at 37oC for 5 h and then pooled into 200 ml LB medium in a 500-

ml flask. The culture was incubated at 37oC, with shaking at 250 rpm overnight. The

next day, the cells were pelleted by centrifugation at 2000 g for 5 min and resuspended

carefully in 12 ml of buffer S1 (50mM Tris-HCl, 10mM EDTA, 100mg/ml RNase A).

The cells were transferred into a 40-ml centrifuge tube and 12 ml of buffer S2 (200mM

NaOH, 1% SDS) was added and the cell suspension was gently and quickly mixed by inverting the tube 6-8 times. This led to the lysis of the cells and the solution became clear. The tube was then allowed to stand at room temperature for 5 min. Buffer S3 (2.8

M KOAc, pH 5.1) was added and the suspension was mixed gently by inverting the tube

6-8 times until a heavy flocculent precipitate was formed. The suspension was incubated

on ice for 30 min and centrifuged at 14,000 g, at 4 oC for 20 minutes. During the time of centrifugation, an AX 500 cartridge was equilibrated with 5 ml of buffer N2 (100 mM

Tris, 15% ethanol, 900 mM KCl, adjusted to pH 6.3 with H3PO4) by passing the buffer

through the cartridge. The cleared lysate (supernatant) of the centrifugation was then

passed through the equilibrated cartridge through a filter paper (to prevent any pelleted

debris to pass through). The cartridge was washed twice, each with 12 ml of buffer N3

(100 mM Tris, 15% ethanol, 1.15 M KCl, adjusted to pH 6.3 with H3PO4). The DNA

was eluted with 12 ml of buffer N5 (100 mM Tris, 15% ethanol, 1M KCl, adjusted to pH

8.5 with H3PO4). The purified DNA was precipitated with 9 ml of isopropanol and

centrifuged at 12, 000 g at 4 oC for 30 min. The DNA pellet was washed with 70%

ethanol by vortexing briefly, repelleted, dried at room temperature for 5-10 min and

redissolved in 500 μl of 1X TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0). The

DNA was quantitated by measuring the O.D.260 , using a GeneQuant pro RNA/ DNA

Calculator spectrophotometer (Biochrom Ltd., Cambridge, England).

Small Scale DNA preparation or Miniprep:

For plasmids which were required in smaller quantities, a small scale DNA preparation

was performed using the Quantum Prep® Plasmid Miniprep Kit (Biorad laboratories,

Hercules, CA). A single bacterial colony from the transformed LB plate was inoculated

into 2 ml of LB medium, with shaking at 37oC for 5 h and then 500 μl of that culture was transferred to 5 ml of fresh LB medium and incubated overnight with shaking at 37oC.

Two ml of the overnight culture of plasmid-containing cells was transferred to a

microcentrifuge tube and pelleted at 14,000 g in 4oC for 30 sec. All subsequent

centrifugation steps were carried out at 14,000 g in 4oC. All the supernatant was removed by aspirating and 200 μl of Cell Resuspension Solution was added to the pellet and vortexed till the cell pellet was completely resuspended. Then 250 μl of Cell Lysis

Solution was added and mixed gently by inverting 10 times. The solution becomes viscous and slightly clear. Then 250 μl of Neutralization Solution was added and mixed gently by inverting 10 times. A visible precipitate forms. The cell debri was then pelleted for 5 min in a microcentrifuge. The cleared lysate (supernatant) is then transferred to a spin column, which is placed in a 2 ml wash tube. Two hundred μl of thoroughly suspended Quantum prep matrix was then mixed to the lysate by pipetting up and down and the mixture was centrifuged for 30 sec. The spin filter is then removed, the filtrate at the bottom of the tube is discarded and the column is replaced back into the tube. 500 μl of wash buffer (which is 50% ethanol) is added to the column and centrifuged again. This wash step is repeated once more, after which 50-100 μl of 1X TE is added to the spin column. The DNA is eluted from the spin column in 1X TE by centrifugation for 1min.

The DNA was quantitated by measuring the O.D.260, using a GeneQuant pro RNA/ DNA

Calculator spectrophotometer (Biochrom Ltd., Cambridge, England).

6. Transient Transfection Assay

Lipid Based Transfection Method:

Cells were cultured in DMEM supplemented with 10% fetal bovine serum in 24-well plates. Confluent cultures of HepG2 cells or subconfluent 293 cells grown in 24-well tissue culture plates were transfected with plasmids. All the plasmid transfections into the cells were performed using a lipid based transfection reagent called Tfx20 (Promega,

Madison, WI). The Tfx™ Reagents are a mixture of a synthetic, cationic lipid molecule

and L-dioleoyl phosphatidylethanolamine (DOPE). Upon hydration with water, these

lipid molecules form multilamellar vesicles that associate with nucleic acids. The

amount of positive charge contributed by the cationic lipid component of the Tfx™

Reagent must exceed the amount of negative charge contributed by the phosphates on

the DNA backbone, resulting in a net positive charge on the multilamellar vesicles

associating with the DNA, which presumably facilitates the interaction of the vesicles

with the negatively charged target cell surface. Charge ratios of 3:1 (i.e. excess positive

charge) were used in cultured cells for all the experiments. The transfections were

performed according to the manufacturer’s instructions. Briefly, the various plasmids are

mixed with 500 μl of serum free medium and the correct amount of Tfx20 reagent is

added to it and this mixture is incubated at room temperature for 15 min. The medium,

in which the cells were being maintained, is removed completely by aspirating and the

plasmid DNA-media-Tfx20 mixture is added drop wise to the cells. The cells are then

returned to the 37oC incubator for 1 h. Following that the cells are overlaid with 1 ml of

serum containing media and returned to the 37oC incubator till lysis.

The luciferase reporter constructs, expression plasmids and pCMV β- galactosidase plasmid (Clontech, Palo Alto, CA) were transfected in each well. The amount of pCMV β-galactosidase plasmid used was 1/10 of the amount of reporter plasmid transfected in each well and served as an internal control for transfection efficiency. The pcDNA3 empty vector was added to normalize the amounts of DNA transfected in each assay. Cells were lysed 40 h after transfection. Cells were treated either with 2 ng/ml or with different doses as in the dose response experiment, of IL-1β in fresh media for 16 h or for the different time periods as in the time course experiment, prior to the cell lysis. In the transfection assays using the inhibitors of the MAPK pathways, the cells were treated for 30 min with the specific inhibitors (Calbiochem, San

Diego, CA) of the MAPK pathways i.e. SP600125 (JNK1 specific inhibitor, 25 μM),

PD98059 (ERK specific inhibitor, 20 μM) or SB203580 (p38MAPK specific inhibitor,

25 μM) prior to treatment with IL-1β. Luciferase activity was assayed after cell lysis using Luciferase Assay System (Promega, Madison, WI) and luminescence was determined using a Lumat LB 9501 luminometer (Berthold Systems Inc., Pittsburgh,

PA). Luciferase activities were normalized for transfection efficiencies by dividing the

relative light units (RLU) by β–galactosidase activity expressed from the cotransfected

pCMV plasmid. Each data point is the average of triplicate assays. Each experiment was

repeated three times. Data are plotted as means ± standard deviation. Statistical analyses of treated vs. untreated controls (vehicle or empty plasmids) were performed using

Student’s t-test. A p<0.05 is considered statistically significant. Fig. 8 is an outline of the method for transient transfection assay.

Fig.8.

Figure 8. Transient transfection assay.

The typical structures of plasmid that are transfected namely, the reporter and expression vector are shown at the top. The lipid based procedure for transfection (as described in

“Materials and Methods”) is depicted in the figure. The last step of transfection is the assay for luciferase and β-galactoside activity.

Luciferase Assay:

Firefly luciferase was assayed by measurement of light production upon addition of luciferin and ATP. This assay is very rapid and sensitive. The medium overlaying the cells was removed completely by aspiration and the cells were washed once with 1x PBS.

To each well, 100 μl of 1X of Cell Culture Lysis buffer (40 mM Tricine, 50 mM NaCl, 2 mM EDTA, 1 mM MgSO4, 5 mM DTT and 1% Triton X-100) was added to lyse the

cells. The attached cells were dislodged by rocking the cell culture plates on a rotating

platform for 20 min in the lysis buffer. For each assay, 20 μl of the clear supernatant lysate was mixed quickly with the luciferin substrate (Promega Corp. Madison, WI) and the light production was measured with a lumat (Wallac Inc. Gaithersburg, MD).

β-Gal activity assay:

An expression plasmid (pCMVβ, Clontech, Palo Alto, CA) containing E.coli β-

galactosidase gene was cotransfected with luciferase reporter gene constructs into the mammalian cells as an internal standard. For each sample, assay buffer was prepared by mixing: 1 μl 100 x Mg Solution; 22 μl 1x ONPG (o-nitrophenyl-β-D-galactopyranoside);

67 μl 0.1 M sodium phosphate (pH 7.5). In a 96 well microplate, 50 μl of cell lysate prepared as described earlier was added to each well. As many as 90 samples were assayed at the same time. Usually six wells were left for negative control, where lysates from cells that were untransfected were added. To each well, 50 μl of the above assay buffer was added with a multichannel pipetter. The plate was incubated at 37oC till a

yellowish color developed. Absorbance at 405 nm was measured for each well with

microplate reader Spectra Max 250 (Molecular Devices, Sunnyvale, CA). The absorbance was measured three times to make sure the O.D.405 of all the wells was in the

linear range of 0.1 to 0.8.

7. RNA Extraction

The primary human hepatocytes in 6-well tissue culture plates were treated with increasing concentrations of IL-1β (0, 2, 5 and 10 ng/ml) or CDCA (0, 5, 10, 25, 50 μM)

for 16 h OR with IL-1β (10 ng/ml or CDCA (50 μM) for 0, 1/2, 1, 2, 6 and 20 h. Total

RNA was extracted from these cells using Tri Reagent TM (Sigma-Aldrich, St.Louis, MO)

according to the manufacturers instructions. The cells were lysed directly on the culture

dish. After aspirating the media off the cells, 1 ml of Tri Reagent was added to each well.

After addition of the reagent, the cell lysate was passed several times through a pipette to

form a homogenous lysate. The lysate was then transferred to 1.5-ml Eppendorf tubes. To

ensure complete dissociation of nucleoprotein complexes, the samples are allowed to

stand for 5 min at room temperature. Following that, 0.2 ml of chloroform was added and the sample was vortexed for 15 sec and allowed to stand at room temperature for 15 min.

The resulting mixture was then centrifuged at 12,000 g for 15 min at 4oC. Centrifugation

separates the mixture into 3 phases: a red organic phase (containing protein), an interphase (containing DNA) and a colorless upper aqueous phase (containing RNA). The aqueous phase was transferred to a fresh 1.5-ml eppendorf tube and 0.5 ml of isopropanol was added to it. The samples were mixed gently by inverting the tubes 10 times. The

samples were then allowed to stand for 10 min at room temperature and then centrifuged at 12,000 g for 10 min at 4oC. The RNA precipitate would now form a pellet at the side

and bottom of the tube. The supernatant was then removed and the RNA pellet was

washed in 1 ml of 75% ethanol (made in DEPC [diethylpyrocarbonate] water) by vortexing. The tubes were then centrifuged at 12,000 g for 5 mins at 4oC. The ethanol

supernatant was then removed and the RNA pellet was air dried for 15 min at room

temperature. The RNA pellet was then dissolved in 50 μl of DEPC water. The RNA was quantitated by measuring the O.D.260, using a GeneQuant pro RNA/DNA Calculator

spectrophotometer (Biochrom Ltd., Cambridge, England).

8. Reverse Transcriptase PCR

DNase treatment of total RNA is an essential step to eliminate any DNA contamination

of the RNA, in order to avoid amplifying DNA templates during reverse transcriptase

PCR. The RNA (10 μg total RNA) was treated with 1 μl of RNase free DNase I

(Concentration: 10 units/μl) (DNA-freeTM, DNAse treatment kit, Ambion, Austin, TX) in

1x DNase reaction buffer, for 1 h at 37oC, after which the DNase was inactivated by

incubation in the DNAse inactivation solution for 2 min at room temperature. The

concentration of the DNase I treated total RNA was determined by UV

spectrophotometry as mentioned above.

For producing the cDNA copies of the target RNA (“first-strand synthesis”), a

DNA primer (in my case an oligo dT primer designed to anneal to the poly-A tail at the

3’ end of mRNA) is annealed to the RNA target and extended by reverse transcriptase.

Reverse transcription of DNase I-treated total RNA (2 μg) was performed using oligo-dT primers (RETROscriptTM first strand synthesis kit for RT-PCR, Ambion, Austin, TX) to

synthesize first strand cDNA in a reaction volume of 20 μl. For each reaction, 2 μg of total RNA (volume made up to 10 μl with nuclease free water) was mixed with 2 μl of oligo-dT primers (50 μM) in a 0.2-ml tube and heated at 72oC for 3 min. All the different

temperature steps were performed using a thermocycler (Gene Amp PCR System 9700,

Applied Biosystems, Foster City, CA). This heating step is necessary in order to eliminate the secondary structure in the target RNA. The tubes are then removed from the

thermocycler and placed on ice for 2 min to cool. Following that a mixture of 2 μl of 10

X RT-PCR Buffer (100 mM Tris-HCL, pH 8.3, 500 mM KCl, 15 mM MgCl2), 4 μl of

dNTP mix (2.5 mM each dNTP), 1 μl of RNase Inhibitor (Placental RNase Inhibitor, 10

units/ μl) and 1 μl of reverse transcriptase enzyme (M-MLV or Maloney-Murine

Leukemia Virus Reverse Transcriptase, 100 units/μl) was pipetted into each reaction. The above mixture was generally made as a cocktail for multiple reactions to minimize pipetting steps. The tubes were spun briefly and then incubated at 42oC for 1 h.

Following that; the tubes were incubated at 92oC for 10 min to inactivate the Reverse

Transcriptase. The tubes were then removed from the thermocycler, diluted 10 fold (with

180 μl of nuclease-free water for a total volume of 200 μl) and stored at –20oC for future

use.

9. Quantitative Real Time PCR (Q-RT PCR)

Quantitative Real Time PCR was performed on the cDNA of the different RNA samples to determine, the expression levels of the target mRNA in each sample. The mRNA of an endogenous reference gene is used as a normalizer from each sample. The Q- RT PCR was performed using two different methods depending upon the availability of different reagents. The dose response experiments were done using the SYBR green method and the time course experiments were done using the Taqman method. Fig. 9 illustrates the principles of both these methods.

SYBR Green Method:

In this method, a specific forward and a reverse primer is used to amplify a portion of the cDNA of interest. The method uses a detection system using the SYBR green dye. Direct detection of PCR product is monitored by measuring the increase in fluorescence caused by binding of SYBR green dye to double-stranded (ds) DNA. The SYBR green master mix (Applied Biosystems, Foster City, CA) is a convenient premix of all components to perform real time PCR except primers, template and water. The SYBR green master mix contains SYBR Green I Dye, AmpliTaq Gold DNA Polymerase, dNTPs with dUTP, passive reference and optimized buffer components. The AmpliTaq Gold DNA

Polymerase is used to perform an efficient Hot Start PCR reaction. It is a chemically modified DNA polymerase, which makes the enzyme inactive. Upon thermal activation, the modifier is released, resulting in active enzyme. The high-temperature incubation step

Fig.9.

Figure 9. Quantitative real time PCR (Q-RT PCR).

The principles of the Q-RT PCR are shown in this figure. As described in “Materials and

Methods”, there are two methods use to perform this technique. The Taqman method uses a probe that has a Fluorescent dye (F) attached to its 5’ end and a Quencher (Q) attached to its 3’ end. The SYBR green method uses a fluorescent SYBR green dye. This figure is adopted from www.takaramirusbio.com

required for activation ensures that active enzyme is generated only at temperatures where the DNA is fully denatured.

Primers for quantitative PCR of human CYP8B1 and human HNF4α were designed using the Primer Express software (Applied Biosystems, Foster City, CA) following the manufacturers recommended parameters. The primer sequences are as follows: hCYP8B1F: 5’-GCCGACTCCAGCGTCTCTC-3’ hCYP8B1R: 5’-GCCCgccGTTGCTGAGCT’3’ hHNF4αF: 5’-GGGTGTCCATACGCATCCTT-3’ hHNF4αR: 5’-CATTGTCATCGATCTGCAGCT-3’

Primer sequences for human SHP [305], human CYP7A1 [306] and the human 23-kDa highly basic protein (HBP, used as the internal standard for normalizing RNA) [306] were previously published. All the primers were tested for specificity using a BLAST

® search. A 1 x SYBR Green PCR master mix (Applied Biosystems, Foster City, CA)

was used with 0.3 μM of forward and reverse primers in a total volume of 25 μl. PCR

reactions contained 5 μl (50 ng) cDNA from the 200 μl diluted RT reaction. To confirm

that genomic DNA was absent, an aliquot of each RNA sample that had not been reverse transcribed was amplified using each primer pair. Also, to confirm the absence of any

contamination in the reaction, controls were run in which H2O was substituted for

template for each primer set. All PCR reactions were done in triplicate. PCR

amplification was performed as follows: 50o for 2 min, 45 cycles of 95o for 15 sec and

55o for 1 min using a ABI PRISM 7700 sequence detector (Applied Biosystems, Foster

City, CA). Following PCR the melting curve analysis was done using Dissociation

Curves v1. Ob1 Software (Applied Biosystems, foster City, CA) to confirm the absence of primer dimers and to see if the correct product was amplified.

Taqman Method:

In this method, a specific forward and a reverse primer and a TaqMan probe consisting of an 18-22 bp oligonucleotide probe which is labeled with a reporter fluorophore at the 5'

end and a quencher fluorophore at the 3' end is used. The fluorescence of the reporter

fluorophore is quenched by a quencher. During PCR, the Taqman probe anneals

specifically between the forward and reverse primer. The polymerase then carries out the

extension of the primer and replicates the template to which the TaqMan is bound. The 5'

exonuclease activity of the polymerase cleaves the probe, releasing the reporter molecule

away from the close vicinity of the quencher. As a result, the fluorescence intensity of the

reporter dye increases. This process repeats in every cycle and does not interfere with the

accumulation of PCR product. The Taqman Universal Master Mix (Applied Biosystems,

Foster City, CA) contains AmpliTaq Gold DNA Polymerase, AmpErase UNG, dNTPs

with dUTP, Passive Reference, and optimized buffer components. The AmpErase UNG

(Uracil N-glycosylase) is used to prevent false positive data. When dUTP replaces dTTP

as a dNTP substrate in PCR, the AmpErase UNG can be used, AmpErase UNG treatment

can prevent the reamplification of carryover PCR products in subsequent experiments.

This method uses enzymatic and chemical reactions analogous to the restriction-

modification and excision-repair systems of cells to degrade specifically PCR products from previous PCR amplifications or to degrade mis-primed, non-specific products produced prior to specific amplifications, but does not degrade native nucleic acid templates. The Assays-on-Demand gene expression products (Applied Biosystems,

Foster City, CA) contain two primers for amplifying the sequence of interest and one

Taqman MGB probe (6-FAM dye-labeled) for detecting the sequence of interest.

Assay-on-Demand PCR primers and Taqman MGB probe mix used are: human CYP7A1 (Cat.# Hs00167982 m1) human HNF4α(Cat. #: Hs00230853 m1) human cJun (Cat.#: Hs00277190_s1) human SHP (Cat.#: Hs00222677_m1) human CYP8B1 (Cat.#: Hs00244754_s1) human UBC (Cat.# Hs00824723 m1).

Amplification of ubiquitin C (UBC) was used in the same reactions as an internal reference gene [307]. A 2X Taqman Universal Master Mix (Applied Biosystems, Foster

City, Ca) was used in a total volume of 25 μl. PCR reactions contained 5 μl (50 ng) cDNA from the 200 μl diluted RT reaction and 1.25 μl of Assay-on-Demand primer/ probe mix specific for each gene. To confirm that genomic DNA was absent, an aliquot of each RNA sample that had not been reverse transcribed was amplified using each primer pair. Also, to confirm the absence of any contamination in the reaction, controls were run in which H2O was substituted for template for each primer set. All PCR

reactions were done in triplicate. PCR amplification was performed as follows: 50o for 2

min, 95o for 10 min, 40 cycles of 95o for 15 sec and 60o for 1 min using a ABI 7500 Real

Time PCR System (Applied Biosystems, Foster City, CA).

Data Analysis:

Amplification data was analyzed using the Sequence Detector v1.7 software (Applied

Biosystems, Foster City, CA) to determine the Ct values. Relative mRNA expression levels were calculated using the mathematical formulas (2-∆∆Ct) recommended by Applied

Biosystems (Applied Biosystems, User Bulletin No.2, 1997). Statistical analysis of real

time PCR results were done using mean normalized cycle threshold (∆Ct) values and the

pooled standard deviation of the mean ∆Ct, which were analyzed by one-way ANOVA

followed by Tukey’s Honestly Significant post-hoc test. A p value of <0.05 was

considered as a statistically significant difference.

10. Protein Extraction

Total cellular protein was extracted from confluent cultures of HepG2 cells in 6-well

tissue culture plates treated with increasing concentrations of IL-1β (0, 2, 5 and 10 ng/ml)

for 16 h. In the immunoblot assay using the inhibitors, the cells were treated for 30 min

with the specific MAPK inhibitors SP600125 (25 μM), PD98059 (20 μM) or SB203580

(25 μM) prior to treatment with IL-1β (5 ng/ml) for 16 h. For the time course experiment,

HepG2 cells were treated with IL-1β (5 ng/ml) or CDCA (25 μM) for 0, 1/2, 1, 2, 6 and

20 h. The bottoms of the wells were scraped with a cell scraper and the cells were

collected in 1ml of 1 X PBS by centrifugation at 5000 g for 5 min at 4oC. The supernatant

was discarded and the pellet was then re-suspended with 1 ml of 1 X PBS and centrifuged

again at 5,000 g for 5 min at 4oC. The resulting pellet was then suspended in 500 μl of

lysis buffer (62.5 mM Tris, 2 mM EDTA, 2.3% SDS, 10% glycerol) containing the

protease inhibitors, 1 mM EGTA, 1 mM PMSF, 2 μg/ml each of leupeptin, pepstatin and

aprotinin. The pellet in lysis buffer was sonicated three times using a Branson sonifier

250 with a micro tip (setting 6 for 15 sec at 40% output) to homogenize the protein

sample. Thereafter, the protein samples in lysis buffer were placed in ice for 15 min.

Protein concentration was determined calorimetrically by BCA protein assay kit (Pierce

biotechnology, Rockford, IL) using different dilutions of albumin proteins as standards.

The cellular extracts were stored as aliquots in -80o C.

11. Immunoblot Analysis

The samples for immunoblot were prepared by diluting 5 μg of the total cellular protein

in 1 X SDS Gel loading or protein loading buffer (50 mM Tris-HCl, pH 8.0, 100 mM

DTT (dithiothreitol), 2% SDS, 0.1% bromophenol blue, 10% glycerol and boiling

(between 95oC to 100 oC) for 10 min. The samples were then loaded in the wells of

1.5mm thickness polyacrylamide gels along with molecular weight markers. The SDS-

polyacrylamide gel consists of a 10% resolving gel and a 4% stacking gel.

Electrophoresis was run in a Mini-Protean III gel-running apparatus (BioRad

Laboratories, Hercules, CA) using 1 X Tris-Glycine buffer (25 mM Tris, 250 mM

Glycine pH 8.3, 0.1% SDS) at 100 V for approximately 2 h or till the dye reached the bottom of the resolving gel. The protein was then transferred to a nitrocellulose membrane (Hybond ECL, Amersham Biosciences, Buckinghamshire, England) using a gel transfer electrophoretic apparatus (Biorad, Hercules, CA) in 1 X Transfer buffer (30 mM Glycine, 48 mM Tris, 0.037% SDS, 20% Methanol) at 75 V for 1.5 h. The membrane was then blocked with 5% (w/v) non-fat dry milk in TBS (Tris-buffered saline [100 mM Tris-Cl, pH 7.5 and 0.9% (w/v) NaCl]) for 2 h at room temperature and incubated with the specific primary antibody (Santa Cruz Biotechnology, Santa Cruz,

CA) at a dilution of 1:5000 in TTBS (0.1% Tween 20 in Tris-buffered saline (100 mM

Tris-Cl, pH 7.5 and 0.9% (w/v) NaCl) overnight at 4oC. Membranes were washed five

times in TTBS (5 min each) and incubated with secondary antibody (horseradish peroxidase-conjugated anti-goat IgG) (Santa Cruz Biotechnology, Santa Cruz, CA) at a dilution of 1:3000 in TTBS for 1 h at room temperature. Membranes were washed again for five times in TTBS (10 min each). Immunodetection was done with an enhanced chemiluminescence (ECL) kit (Amersham Biosciences, Buckinghamshire, England).

Membranes were imaged by exposing them to X-ray films. The specific primary antibodies used were against HNF4α, cJun, phospho-cJun and Actin.

12. Preparation of Nuclear Extracts and In vitro Translated Proteins

Nuclear Extracts:

Confluent cultures of HepG2 cells (3 T-175 cm2 flasks per sample) treated with IL-1β (0,

2, 5 and 10 ng/ml) for 16 h were lysed with trypsin and washed twice with 50 volumes of

cold phosphate-buffered saline (PBS) and the cells were pelleted by centrifugation at 170

g for 10 min at 4°C. The washed cells were transferred to a 15-ml graduated centrifuge

tube, resuspended in a small volume (3 ml) of phosphate buffered saline and sedimented

at 300 g for 10 min at 4oC. The packed cell volume (PCV) was measured and the cells

were resuspended in 5 PCV of Hypotonic buffer (10 mM HEPES pH 7.9, 1.5 mM

MgCl2,10 mM KCI, 0.5 mM DTT, 0.5 mM PMSF). The cells were allowed to swell for

10 min on ice, and then pelleted again at 300 g for 10 min at 4°C. The buffer

(supernatant) was poured off and replaced with 2 times the original PCV of Hypotonic

buffer. The cells were broken by six strokes with the tight pestle of a Dounce

homogenizer (Kontes Glass Co) and checked by phase-contrast microscopy for >95%

cell breakage. One-tenth volume Sucrose Restore buffer (9 volumes of 75% RNase free

sucrose and 1 volume of 10 x salts consisting of 500 mM HEPES pH 7.9, 7.5 mM

spermidine, 1.5 mM spermine, 100 mM KCI and 2 mM EDTA, 10 mM DTT) was added

immediately, and the homogenate was mixed with two strokes of the loose pestle of the

homogenizer. The homogenate was immediately spun for 30 sec at 10,000 rpm in a

Sorvall HB-4 rotor (16,000 g) at 4oC with the brake on. The total centrifugation time was

approximately 2.5 min. The supernatant was carefully poured off and the viscous nuclear

pellet resuspended in 3 ml of Nuclear Resuspension buffer/1 x 109 cells and transferred to

a straight-wall polycarbonate centrifuge tube. The final Nuclear Resuspension buffer was

prepared by adding 9 volumes of Nuclear Resuspension solution (20 mM HEPES pH 7.9,

0.15 mM spermine, 0.75 mM spermidine, 2 mM EGTA, 2 mM DTT, 25% glycerol) to 1 volume of saturated (at 4ºC) ammonium sulfate. The extract was then rocked on a

“Lab-Quake” (Tech-Lab Industries, Arlington, TX) for 30 min at 4oC and sedimented by

centrifugation at 2oC for 90 min at 150,000 g (40,000 rpm, Beckman Ti-80 rotor) to pellet

the nuclear debris and chromatin. The supernatant was carefully removed. Solid

ammonium sulfate (0.33 g/ml) was added to the supernatant to precipitate the nuclear

protein from the supernatant. The sample was then rocked on the “LabQuake” for 20 min

after the ammonium sulfate was dissolved. The total time for this step was about 1 h.

The precipitate was collected by centrifugation at 85,000 g for 20 min at 2oC. The pellet

was re-dissolved in up to 1 ml of Nuclear Dialysis buffer (20 mM HEPES pH 7.9, 20%

glycerol, 100 mM KCI, 0.2 mM EDTA, 0.2 mM EGTA, 2 mM DTT)/109 cells, and dia-

lyzed twice overnight, against > 200 volumes of Nuclear Dialysis buffer. The nuclear

extract was quick-frozen in small aliquots and stored in –80oC.

Invitro Translated Proteins:

The T7 TNT Quick Coupled Transcription/Translation System (Promega, Madison, WI) was used for in vitro synthesis of proteins. The TNT Quick Coupled

Transcription/Translation System has RNA polymerase, salts, ribonuclease inhibitor and rabbit reticulocyte lysate. Expression Vector or Circular plasmid DNA (1 μg) containing a T7 promoter is added to an aliquot of the TNT Quick Master Mix (40 μl) and incubated with 1 μl Methionine in a 50µl reaction volume for 90 minutes at 30°C in a thermocycler

(Gene Amp PCR System 9700, Applied Biosystems, Foster City, CA). 5 µl of reaction mixture was in each EMSA reaction or the reaction mixture was quick-frozen in small aliquots and stored in –80oC.

13. Electrophoretic Mobility Shift Assay (EMSA)

EMSA is a useful tool for identifying DNA-protein interactions. This assay is also known

as gel retardation or band shift assay. The assay is performed by incubating radiolabeled

double strand DNA oligonucleotide fragments with either nuclear extracts or purified

DNA binding proteins. The mixture is separated on a nondenaturing polyacrylamide gel.

Oligonucleotide duplexes that are bound by protein migrate slower on the gel than

unbound duplexes and appear as bands that are shifted relative to the bands from the

unbound duplexes. The bands are visualized by autoradiography. Specificity of the DNA-

protein interaction is determined by adding a molar excess of unlabeled specific

oligonucleotide probe (known to bind the specific protein) to the reaction, which

competes out (hence decreases/eliminates) the binding of the radiolabeled probe to the

protein (Competition Reaction). Alternatively, a specific antibody against the protein of

interest is added to the reaction, thereby further retarding the migration of the DNA-

protein-antibody complex causing a supershifted band (Supershift Reaction). The

procedure and principles of EMSA are illustrated in Fig. 10.

Double stranded probes were prepared by annealing equal molar amounts of

complementary oligonucleotides by heating the oligos at 100 °C in 2 × SSC (0.3 M NaCl,

Fig.10.

Figure 10. Electromobility shift assay (EMSA).

The procedure of EMSA is shown in this figure. A 32P labeled oligonucleotide probe is tested for binding the protein of interest. The gel at the bottom shows a typical picture seen on EMSA. If the protein and DNA interact a band is seen. The free probe, which does not bind protein has a higher mobility runs to the bottom of the gel. Addition of antibody against the protein to the reaction retards to mobility of the band forming a supershifted band. A specific cold competitor will decrease the intensity of the band, as it competes with the hot (radiolabeled) probe for binding to the protein. While a no-specific competitor will not change the intensity of the band. However the mobility of the bands does not change in the competition assays.

0.03 M sodium citrate, pH 7.0) for 5 min and then cooling them slowly to room temperature. The concentration for the oligo was 100pMol/μl. The resulting double strand oligos were labeled by filling in a 5’ overhang with the Klenow fragment as follows: 100 pmol ds oligos, 1 mM unlabeled dCTP, dTTP and dGTP, 50 μCi α-32PdCTP

(3000 Ci/mol), 5 μl of 5 X Klenow buffer, 1 μl Klenow enzyme (5 units) and H2O in a

final volume of 50 μl. This mixture was incubated at 37 °C for 30 min and the volume was adjusted to 200 μl with 1 X TE buffer. The labeled oligonucleotides were separated from unincorporated radioisotope label by centrifugation through a 1 ml Sephadex G-25 column at 1000 rpm with a desktop centrifuge for 5 min. Counts of 1 μl of the reaction mixture after the spin-column centrifugation were used to determine the specific activity

(cpm/μl). The same oligonucleotide filled-in with non-labeled dNTPs was used as a cold competitor. The oligonucleotide probe sequences are included with the respective figures in the Results section.

A 5% non-denaturing polyacrylamide gel was pre-run in 1 × TBE buffer at 100 V for

1 h. Meanwhile, the binding reaction was assembled in the binding buffer (12 mM

HEPES pH 7.9, 50 mM KCl, 1 mM EDTA, 1 mM DTT and 15% Glycerol) as follows:

50,000 cpm double strand oligonucleotides probe, 1 μg poly dI-dC (to eliminate non-

specific binding), nuclear extracts 2–4 μg or TNT expressed protein 5 μl, 4μl of 5 X binding buffer and H2O in a final volume of 20 μl. The reaction was incubated on ice for

20 min. For competition EMSA, excess amounts of unlabeled specific and nonspecific competitor oligonucleotides were added before adding the DNA-binding protein. For

supershift assay, a small volume of antibody (~2 μl) (Santa Cruz Biotechnology, Santa

Cruz, CA) was added to the binding reaction and incubated for 15 min before adding the labeled probe. Each binding reaction was loaded onto the prerun gel and electrophoresis was carried out at 200 V for about 1.5 h - 2 h or until the bromophenol blue marker ran to two-thirds of the way down the gel. The gel was dried on to a filter paper (by vacuum drying for 1 h 80°C). The dried gel was placed with an intensifying screen overnight and autoradiographed the following morning using Phosphoimager 445Si (Molecular

Dynamics, Sunnyvale, CA). The images were analyzed using IP Lab Gel software

(Signal Analytics Corp. Vienna, VA).

14. Chromatin immunoprecipitation assay (ChIP)

ChIP assay is a powerful method used to identify regions of the genome associated with specific proteins. The ChIP assay has been used to study both histones and non-histone proteins, such as transcription factors, within the context of the cell. Transcription factors and other DNA binding proteins have a weaker affinity than histones, which generally are tightly associated within the chromatin complex. To avoid dissociation of non-histone proteins from the chromatin binding site, it is necessary to cross-link proteins to DNA.

After cross-linking with formaldehyde, the chromatin are released from the nuclei and sheared into 200-1000bp fragments. The chromatin are then immunoprecipitated with specific antibodies. DNA sequences cross-linked to the protein of interest co-precipitate

as part of the chromatin complex. The DNA/ chromatin/antibody complex is then isolated using Protein A agarose. After reverse cross-linking, the associated DNA is released from the complex and amplified by PCR for analysis. Important steps in ChIP assay are shown in Fig. 11.

HepG2 cells were grown in 100 mm culture dishes to 80% confluence and then treated with 10 ng/ml of IL-1β or 50 μM CDCA for 20 h or left untreated; or alternatively they were grown to 50% confluence and then transfected using Lipofectamine 2000

(Invitrogen Corporation, Carlsbad, CA) with pcDNA3 or JNK1 plasmids or with pcDNA3 or cJun plasmids along with or without a HA tagged PGC-1α plasmid. The cells were used for ChIP assay 48 h after transfection. ChIP assays were performed using a

ChIP Assay kit (Upstate Cell Signaling Solutions, Lake Placid, NY) according to manufacturer’s protocol. Briefly, HepG2 cells were cross-linked in 1% formaldehyde for

10 min and washed twice with ice cold PBS containing protease inhibitors (1 mM PMSF,

1 μg/ml aprotinin and 1 μg/ml pepstanin A) twice. Cells were scraped and incubated in

1% SDS lysis buffer (1% SDS, 10 mM EDTA and 50 mM Tris-HCl, pH 8.1) for 10 min on ice and sonicated using a Branson sonifier 250 with a micro tip (setting 6 for 15 sec at

40% output) to break the DNA into 0.2 to 2 kb fragments. Cell lysates were collected by centrifugation and diluted 10 fold in ChIP dilution buffer (0.01% SDS, 1.1% triton X-

100, 1.2 mM EDTA, 16.7 mM Tris-HCl, pH 8.1 and 167 mM NaCl). 5% of the cell lysate solution in ChIP dilution buffer was kept aside as Input. After preclearing the remaining diluted cell lysate with protein A-agarose, DNA-protein complexes were precipitated by incubating the cell lysates with 10 μg of specific antibody (as indicated in

Fig.11.

Figure 11. Chromatin immunoprecipitation (ChIP) assay.

The basic principle and procedure of a ChIP assay is shown here. The formaldehyde crosslinking of protein to DNA, the shearing (by sonication) of the cells, the addition of antibody against the protein of interest, immunoprecipitation of the DNA-protein complexes by the antibody linked to a protein A/G agarose beads and the reverse crosslinking reaction are some important steps in the process. The DNA is finally analyzed by PCR amplification of a specific fragment of the gene of interest. This figure is adopted from www.activemotif.com (ChIP-ITTM kit).

the respective figures in the results section) or Non- Immune (Normal) antibody (Santa

Cruz Biotechnology, Santa Cruz, CA) overnight followed by incubation with protein A- agarose beads for 3 h. The beads were washed once each with low salt wash buffer (0.1%

SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCL, pH 8.1 and 150 mM NaCl), high salt wash buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCL, pH

8.1 and 500 mM NaCl) and LiCl wash buffer (0. 25 M LiCl, 1% NP40, 1% deoxycholate,

1 mM EDTA, 10 mM Tris-HCl, pH 8.1), and twice with TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0). Samples were eluted twice with freshly prepared ChIP elution buffer (1% SDS and 0.1 M NaHCO3) and the eluants were combined. Reverse

crosslinking of both the eluant and the input fractions was achieved by adjusting the NaCl

concentration to 200 mM and incubating at 650C for 4 h. The eluants were then incubated

at 450C for 1 h in 0.04 μg/μl proteinase K. DNA was extracted using phenol/chloroform

and precipitated using isopropanol. Depending upon whether the human CYP8B1 or the

human CYP7A1 promoter was being studied, the PCR step was performed as follows:

A 251 bp DNA fragment containing the bile acid response element of the human

CYP8B1 promoter was PCR amplified for 40 cycles (primer sequence given below)

using 5 μl of the DNA as template and analyzed on a 1.5% agarose gel. The PCR primers

used were:

L8B1ChIPHNF4F: 5’ AAGCTGGTGAGCAGCTGTGA 3’

L8B1ChIPHNF4R: 5’ CACACTGTTCCCTGGGTGC 3’

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A 391 bp DNA fragment containing BAREI and BAREII (from -432 to – 41) of the human CYP7A1 promoter was PCR amplified for 40 cycles (primer sequence given below) using 5 μl of the DNA as template and analyzed on a 1.5 % agarose gel. The PCR primers used were: forward primer: 5' ATCACCGTCTCTCTGGCAAAGCAC 3’ reverse primer: 5' CCATTAACTTGAGCTTGGTTGACAAAG 3’

15. GST fusion protein expression

The pGEX-4T-1 plasmid encoding the Glutathione S- transferase (GST) protein alone, and the pET23a-GST-HNF4α plasmid encoding a fusion protein of GST with full length human HNF4α were transformed into E. coli BL21 cells (Amersham Biosciences,

Buckinghamshire, England). Cells were grown in large cultures (1 L) to an O.D. of about

0.6 and induced with 1 mM IPTG (Amersham Biosciences, Buckinghamshire, England) for an additional 2 h. Cells were centrifuged and the bacterial pellets were suspended in

Triton X100 lysis buffer (0.05 M Tris-HCl, 5 mM EDTA, 0.05 M NaCl, 0.1% Triton

X100, 10% glycerol) containing the protease inhibitors (1 mM PMSF, 1 μg/ml aprotinin and 1 μg/ml pepstanin A) and sonicated 10 times in pulses of 30 sec each. Cell lysates were clarified by centrifugation at 10,000 g for 10 min at 4°C. Cell lysates (1 ml each) were incubated with immobilized glutathione beads (Amersham Biosciences,

Buckinghamshire, England) at 4° C for 2 h. The beads were washed four times in the same lysis buffer and the proteins bound to the glutathione beads were eluted with an

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elution buffer (50 mM Tris-HCl pH 8.0) containing 10 mM reduced glutathione. The protein samples, mixed with 2 X protein loading buffer were loaded on an SDS-PAGE

(10% resolving and 4% stacking gel). The gel was then stained with Coomassie Blue stain (Biorad Laboratories, Hercules, CA) to confirm the correct protein expressions (Fig.

12A.). Protein concentration was determined by the Coomassie Plus-Better Bradford

Assay Reagent (Pierce Biotechnology, Rockford. IL) using different dilutions of albumin proteins as standards. The proteins were stored as aliquots in -80o C for future use in the

GST pull down assays or the kinase assays.

16. GST Pull Down Assay

The pull-down assay is an in vitro method used to determine physical interaction between

two or more proteins. The minimal requirement for a pull-down assay is the availability

of a purified and tagged protein (the bait), which will be used to capture and ‘pull-down’

a protein-binding partner (the prey). A Gluathione-S-Transferase (GST)-tagged bait

protein is bound to an immobilized glutathione support. In a typical pull-down assay, the

immobilized bait protein is introduced to a protein pool derived from a cell lysates or a

specific purified protein that is suspected to interact with the bait protein. After the

prescribed washing steps, the 'interactors" are selectively eluted. The interacting proteins

are then detected in an SDS-PAGE gel. The principles are steps of a GST pull down

assay are shown in Fig. 12B.

The GST and GST-HNF4α proteins were each incubated with immobilized glutathione

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Fig.12A.

103

Fig.12B.

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Figure 12. GST fusion protein expression and GST pull down assay.

A. GST fusion protein expression. The method used for GST (glutathione-S- transferase) fusion protein expression is discussed in the “Materials and Methods”. This picture shows a Coumassie Blue stained gel showing the staining of the SDS-PAGE in which the purified proteins were run. The molecular weight marker shows the molecular weight of the standard proteins in kilo daltons. Lanes 2 and 3 show the expression of

GST protein alone and lanes 4 and 5 show the expression of GST-HNF4α protein.

B. GST pull down assay. This figure shows the basic principle and procedure for a GST

(glutathione-S-transferase) pull down assay. The procedure is discussed in the “Materials and Methods” section. The GST fusion protein (“bait”) binds to the glutathione sepharose beads and pulls down along with it the 35S labeled protein (“prey”) with which it interacts. If the two proteins interact a band is seen after SDS-PAGE and autoradiography.

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beads (Amersham Biosciences, Buckinghamshire, England) at 4 °C for 2 h. The beads were washed four times in the Triton X100 lysis buffer and the proteins bound to the glutathione beads were divided into equal parts for the number of reactions for each protein. Typically each reaction had approximately 4μg of the protein. The 35S labeled

cJun and PGC-1α were generated using their respective expression vectors in the TNT

rabbit reticulocyte system (Promega, Madison, WI) in the presence of 35S labeled

Methionine. 1μl (20% Input) of the 35S labeled invitro translated proteins were run on an

SDS PAGE (10% resolving and 4% stacking gel) and analyzed for expression. 4μg of the

GST or GST-HNF4α proteins bound to the glutathione beads was added to 5 μl of the 35S labeled cJun or PGC-1α protein (or with 5 μl PGC-1α and 5 μl and 20 μl of cJun in the last two lanes for competition) and incubated with rotation in GST incubation buffer (50 mM KCl, 40 mM HEPES pH 7.5, 5 mM β-mercaptoethanol, 0.1% Tween-20. 1% non fat dry milk) at 4oC for 2h. After incubation the beads were washed three times with the GST

incubation buffer. After the third wash the beads were pelleted at 3,000 rpm and

resuspended in 1X SDS lysis buffer. The tubes were vortexed and then heated at 95oC for

10 min. The beads were then repelleted by centrifugation and the supernatant from each

sample was loaded on an SDS PAGE gel (10% resolving and 4% stacking gel). The gel

was dried on to a filter paper (by vacuum drying for 1 h 80°C). The dried gel was placed with an intensifying screen for 48 h prior to autoradiography using the Typhoon 8610

phosphoimager (Amersham Biosciences, Piscataway, NJ).

17. In vitro kinase Assay

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A kinase is protein enzyme that adds a phosphate onto a molecule, though typically only proteins. The phosphorylated molecule can be another protein, the kinase itself

(autophosphorylation) or any other molecule. The source for the phosphate is a terminal phosphate, which is called the gamma (γ) phosphate from ATP. In an in vitro kinase assay, a purified and active kinase enzyme is used to phosphorylate a protein in a reaction mixture containing radiolabeled-γ ATP (γ-32P ATP). If the protein is phosphorylated by

the kinase, the γ-32P ATP gets incorporated into the protein, and can be visualized by

SDS-PAGE and autoradiography. The principles of an invitro kinase assay are shown in

Fig. 13.

In the JNK assays, 4 μg of GST, 4 μg of GST-HNF4α or 2.5 μg of ATF2 protein

(Upstate Cell Signaling Solutions, Lake Placid, NY) were incubated at 37o for 30 min

with 20 ng of active JNK1 enzyme (Upstate Cell Signaling Solutions, Lake Placid, NY)

along with 10 μCi of γ-32P ATP in the JNK kinase reaction buffer (500mM Tris-HCl pH

7.5, 1% 2-mercaptoethanol, 20 mM MOPS pH 7.2, 25 mM β-glycerol phosphate, 5 mM

EGTA, 1 mM sodium orthovandate and 1mM DTT) to which 75 mM MgCl2 and 500 μM

ATP were added. An enzyme negative reaction was performed with each protein without

adding active JNK1 enzyme. In the AMPK assays, 4 μg of GST, 4 μg of GST-HNF4α or

100 mM of SAMS peptide (Upstate Cell Signaling Solutions, Lake Placid, NY) were incubated at 37o for 30 min with 1000 mU of active AMPK enzyme (Upstate Cell

Signaling Solutions, Lake Placid, NY) along with 10 μCi of γ-32P ATP in the AMPK

kinase reaction buffer (20 mM HEPES-NaOH pH 7.0, 0.01% Brij-35, 20 mM MOPS pH

7.2, 25 mM β-glycerol phosphate, 5 mM EGTA, 1mM sodium orthovandate and 1 mM

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Fig.13.

108

Figure 13. Invitro kinase assay.

This figure shows the basic principle and procedure for an invitro kinase assay. The procedure is discussed in the “Materials and Methods” section. The active kinase enzyme can add the phosphate group from the 32P -γ-ATP to its particular phosphorylation

recognition sequence (if present) on the GST fusion protein. If the kinase phosphorylates

the GST fusion protein a band is seen after SDS-PAGE and autoradiography.

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DTT either with or without 300 μM AMP) to which 75 mM MgCl2 and 500 μM cold

ATP were added. An enzyme negative reaction was performed with each protein without

adding active AMPK enzyme. After incubation all the samples were mixed with Laemmli

buffer, boiled for 5 min and analyzed by SDS-PAGE (10% resolving and 4% stacking

gel). The gel was dried on to a filter paper (by vacuum drying for 1 h 80°C). The dried

gel was placed with an intensifying screen for 15 min prior to autoradiography using the

Typhoon 8610 phosphoimager.

18. Co-Immunoprecipitation Assay (Co-IP)

Co-IP is an important technique to identify protein-protein interaction in vivo. An

antibody (monoclonal or polyclonal) against a specific target antigen is allowed to form

an immune complex with that target in a sample, such as a cell lysate. The immune

complex is then captured on a solid support to which either Protein A or Protein G has

been immobilized (Protein A or G binds to the antibody, which is bound to its antigen).

The process of capturing this complex from the solution is referred to as precipitation.

The target antigen precipitated by the antibody “co-precipitates” a binding partner/protein

complex from a lysate, i.e., the interacting protein is bound to the target antigen, which

becomes bound by the antibody that becomes captured on the Protein A or G gel support.

Any proteins not “precipitated” by the immobilized Protein A or G support are washed

away. Finally, components of the bound immune complex (both antigen and antibody)

are eluted from the support and analyzed by SDS-PAGE (gel electrophoresis) followed

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by Western blot detection to verify the identity of the antigen and its interacting protein.

The assumption that is usually made when associated proteins are co-precipitated is that these proteins are related to the function of the target antigen at the cellular level. The principle of Co-IP is shown in Fig 14A, while the steps used in this experiment are shown in Fig. 14B.

HepG2 cells were cultured in T 150 cm2 flasks. The cells were treated with IL-

1β (10ng/ml) for 0, 1, 6, and 20 h. The cells were then collected and incubated in modified RIPA buffer (50mM Tris-HCl, 1% NP-40, 0.25%-deoxycholate, 150mM NaCl,

1mM EDTA) containing protease inhibitors (1 mM EGTA, 1 mM PMSF, 2 μg/ml each of leupeptin, pepstatin and aprotinin) for 30 min. Clear cell lysate was collected by centrifugation at 10,000 g at 4oC for 15 min. Clear lysates were precleared with protein A

beads. 5% of the cell lysate solution was kept aside as Input. The remaining cell lysate

was then incubated with 20 μg of Rabbit Anti-HNF4α antibody (Santa Cruz

Biotechnology, Santa Cruz, CA) at 4oC with rotation overnight, followed by an additional incubation for 2 h with protein A beads. The beads were then washed three times with

modified RIPA buffer and were boiled in 1X protein loading buffer for 5 min. Samples

were divided into equal amounts and loaded on SDS PAGE gels for Immunoblot analysis

(details as described above) with Goat Anti-HNF4α, Anti-cJun and Anti-phospho-cJun

antibodies (Santa Cruz Biotechnology, Santa Cruz, CA). Equal amount of the Input from each sample was also run on an SDS PAGE gel and immunoblotted with Anti-Actin antibody (Santa Cruz Biotechnology, Santa Cruz, CA).

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Fig. 14A.

112

Fig.14B.

113

Figure 14. Coimmunoprecipitation (Co-IP) assay.

A. The principle of Co-IP is shown in this figure. To determine the interaction between protein X and protein Y, an antibody against protein X that is bound to a ProteinA/G agarose bead is used to immunoprecipitate both the proteins together. B. This figure shows the procedure used in a Co-IP assay. The immunoprecipitated proteins are finally analyzed by a western blot. This figure is adopted from www.piercenet.com. (Seize

Classic IP kit).

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19. Small Interference RNA (siRNA)

RNA interference (RNAi) is a phenomenon of gene silencing at the mRNA level offering a quick and easy way to determine the function of a gene both in vivo and in vitro [308-

310]. The antisense approach to gene silencing involves injecting an organism with RNA sequence complementary to mRNA transcribed from a target gene. The antisense RNA and sense mRNA hybridizes and blocks the translation and the production of an encoded protein. Hence, the presence of dsRNA duplex leads to a RNA interference effect. In

RNAi exogenous, double-stranded RNAs (dsRNAs) that are complimentary to known mRNAs, are introduced into a cell to specifically destroy that particular mRNA, thereby diminishing or abolishing gene expression. The technique has proven effective in

Drosophila, Caenorhabditis elegans and plants. However, in mammalian cells, long dsRNA was not very successful in inducing RNAi, due to the induction of an interferon response, which resulted in a general inhibition protein synthesis [311]. To make the technique work in cultured mammalian cells small interfering RNAs (siRNAs), which are dsRNAs of some 21-25 nucleotides, are delivered into the cells using a transfection reagent. Artificial siRNAs can be made in the lab by a phage enzyme referred to as

DICER. Duplexes of 19 nucleotides (nt) RNAs with symmetric 2-nt (preferably dT bases) 3' overhangs (siRNA) are introduced into a cell where they associate with specific proteins to form a multi-protein siRNA complex termed RISC (RNA Induced Silencing

Complex). This ribonucleoprotein complex then scans the mRNA cell-content to degrade the corresponding mRNA target in a highly specific manner. Typically, 3-5 double-

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stranded siRNA molecules are designed per gene and used either alone or in combination in order to find a siRNA that has a strong effect. The principle of a siRNA experiment is illustrated in Fig. 15.

To knockdown HNF4α, a SMARTpool Reagent product (Dharmacon Inc, Chicago, IL) was used. The SMARTselection strategies incorporate a multi-component algorithm that identifies siRNAs with a high probability of potent and specific silencing. The

SMARTpool reagents combine 4 different SMART-selection designed siRNAs into a single pool, resulting in a greater probability that the reagent will reduce target mRNA to low levels. The SMARTpool Reagent targeted to knockdown HNF4α (Cat # M-003406-

01-0005) is provided annealed, desalted, pooled and 2’-hydroxylated. The SMARTpool

Reagent (e.g. 5nM) is dissolved in appropriate amount (in 0.25 ml) of 1 X siRNA buffer

(20mM KCl, 6 mM HEPES, pH 7.5, 0.2 mM MgCl2) to obtain a final concentration of 20

μM (20 pmol/μl).

The SMARTpool Reagent was transfected into the cells using a lipid based

transfection reagent called TransIT-siQUEST Reagent (Mirus, Madison, WI) as per the

manufacturer’s instructions. HepG2 cells were seeded in 6 well tissue culture plates, 24 h

before transfection to achieve a confluency of 60-80% at the time of transfection.

Immediately prior to transfection, the original media in the wells was replaced with 1.75

ml of fresh growth media. Then, 15 μl of the TransIT-siQUEST reagent was added to

0.25 ml of serum free media. This mixture was incubated for 20 min at room temperature.

Then, 10 μl of SMARTpool siRNA Oligos in buffer was added to the above mixture of

transfection reagent and media. This amount is sufficient to achieve a final siRNA Oligo

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Fig.15.

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Figure 15. Small interference RNA (siRNA).

The principle of gene knockdown by siRNA is shown in this figure. Duplexes of RNA

(dsRNAs) of some 21-25 nucleotides, are delivered into the cells using a transfection reagent. siRNAs are made within the cell by a phage enzyme referred to as DICER. The antisense strand of the siRNA hybridizes with the corresponding sense strand of a target mRNA. This complex associates with specific proteins to form a multi-protein siRNA complex termed RISC (RNA Induced Silencing Complex). This ribonucleoprotein complex then degrades the corresponding mRNA target in a highly specific manner. This figure is adopted from www.nature.com

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concentration of 100 nM (as recommended by Dharmacon, Inc). This mixture was incubated for another 20 min at room temperature, after which it was added drop wise to the cells. The plates were incubated for 48 h, before the RNA was extracted. In some cases the cells were treated with either 10 ng/ml of IL-1β or 50 μM of CDCA 20 h prior to RNA extraction. DNase treatment, cDNA synthesis and real time PCR using the

Taqman method were performed as described earlier. The following mRNAs were assayed in real time PCR: HNF4α, CYP7A1, PEPCK and SHP.

CHAPTER III

RESULTS

Effect of IL-1β on CYP8B1 mRNA expression in human primary hepatocytes

Quantitative real time PCR was performed to determine the effect of IL-1β on CYP8B1

mRNA expression in human primary hepatocytes. Treatment of increasing amounts of

IL-1β for 20 h markedly reduced CYP8B1 mRNA levels in human primary hepatocytes by about 70 to 90% (Table 1). Similarly, IL-1β reduced CYP7A1 mRNA expression in a

dose-dependent manner. Interestingly, IL-1β also reduced HNF4α and SHP mRNA

levels in human hepatocytes by more than 80% at10 ng/ml. To compare the effect of IL-

1β with bile acids on mRNA expression, I treated human primary hepatocytes with

CDCA, a FXR ligand. As shown in Table 2, CDCA significantly inhibited CYP8B1,

CYP7A1 and HNF4α mRNA levels in a dose-dependent manner by about 40 to 70%. In

contrast, CDCA dose-dependently induced SHP mRNA levels by up to 4 fold at 50 μM.

Quantitative real time PCR assays was also performed to determine the time dependent

effect of IL-1β on CYP8B1 mRNA expression in human primary hepatocytes. A dose of

10ng/ml of IL-1β inhibited CYP8B1 expression by about 70% after 20 h treatment (Table

3). This inhibition was time dependent with approximately a 25% inhibition seen as early

as 1 h post treatment. The same dose of IL-1β also showed a time dependent suppression

119 120

Table 1. Effect of IL-1β on mRNA expression (dose response) in human primary hepatocytes (HH1088) determined by real time RT-PCR

IL-1β (ng/ml) 0 2 5 10

CYP8B1 1 0.3* 0.25* 0.12*

∆Ct±S.D.∆Ct 8.93±0.38 10.67±0.50 10.90±0.57 11.95± 0.03

CYP7A1 1 0.48* 0.26* 0.14*

∆Ct±S.D.∆Ct 9.50±0.24 10.35±0.15 11.21±0.07 12.26±0.23

HNF4α 1 0.39* 0.25* 0.18*

∆Ct±S.D.∆Ct 6.98±0.48 8.31±0.19 8.97±0.13 9.43±0.17

SHP 1 0.19* 0.14* 0.13*

∆Ct±S.D.∆Ct 10.36±0.15 13.79±0.92 14.38±0.43 14.90±0.25

Reference Gene: 23kD Highly Basic Protein (HBP)

Relative expression: Relative expression of gene of interest calculated as 2-∆ ∆Ct

ΔCt = Ct of gene of interest – Ct of reference gene.

2 2 SDΔCt = Square Root [(Stdev of Ct of reference gene) + (Stdev of gene of interest) ].

* Indicates statistically significant difference (p<0.05).

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Table 2. Effect of CDCA on mRNA expression (dose response) in human primary hepatocytes (HH1165) as determined by real time RT-PCR

CDCA(μM) 0 5 10 25 50

CYP8B1 1 0.84* 0.31* 0.32* 0.26*

∆Ct±S.D.∆Ct 4.01±0.23 4.26±0.19 5.68±0.15 5.64± 0.14 5.92±0.18

CYP7A1 1 0.75* 0.69* 0.61* 0.52*

∆Ct±S.D.∆Ct 9.64±0.24 10.06±0.26 10.17±0.14 10.33±0.35 10.58±0.42

HNF4α 1 0.87* 0.43* 0.48* 0.43*

∆Ct±S.D.∆Ct 2.76±0.18 2.95±0.15 3.98±0.13 3.81±0.24 3.96±0.23

SHP 1 1.79* 1.83* 2.61* 3.99*

∆Ct±S.D.∆Ct 2.53±0.20 1.69±0.07 1.66±0.12 1.15±0.21 0.54±0.15

Reference Gene: 23kD Highly Basic Protein (HBP)

Relative expression: Relative expression of gene of interest calculated as 2-∆ ∆Ct

ΔCt = Ct of gene of interest – Ct of reference gene.

2 2 SDΔCt = Square Root [(Stdev of Ct of reference gene) + (Stdev of gene of interest) ].

* Indicates statistically significant difference (p<0.05).

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Table 3. Effect of IL-1β on mRNA expression (time course) in human primary hepatocytes (HH 1209) as determined by real time RT-PCR

IL-1β 0 hr 1/2 hr 1 hr 2 hr 6 hr 20 hr 10ng/ml

CYP7A1 1 0.97 0.84* 0.62* 0.42* 0.27*

∆Ct±S.D.∆Ct 13.00±1.06 13.04±0.40 13.25±0.19 13.67±0.18 14.24±0.27 14.87±0.46

HNF4α 1 1.02 0.74* 0.65* 0.51* 0.47*

∆Ct±S.D.∆Ct 2.78±0.83 2.75±0.20 3.20±0.55 3.40±0.10 3.75±0.13 3.85±0.52

CYP8B1 1 1.01 0.75* 0.59* 0.49* 0.32*

∆Ct±S.D.∆Ct 1.80±0.86 1.79±0.72 2.20±0.73 2.56±0.70 2.83±0.58 3.42±0.49

SHP 1 0.98 0.79* 0.64* 0.53* 0.43*

∆Ct±S.D.∆Ct 5.55±0.75 5.56±0.18 5.87±0.41 6.17±0.25 6.45±0.23 6.76±0.43

Reference Gene: Ubiquitin C (UBC)

Relative expression: Relative expression of gene of interest calculated as 2-∆ ∆Ct

∆Ct=Ct of gene of interest - Ct of reference gene

2 2 S.D.ΔCt = Square Root [(Stdev of Ct of reference gene) + (Stdev of gene of interest) ]

* Indicates statistically significant difference (p<0.05).

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of HNF4α, CYP7A1 and SHP mRNA levels, which were reduced by approximately

50%, 70% and 60% respectively after a 20 h treatment period. To compare the results of

IL-1β treatment I decided to treat primary human hepatocytes with a 50μM dose of

CDCA.As expected, CDCA suppressed the CYP8B1 mRNA expression by approximately expected, CDCA suppressed the CYP8B1 mRNA expression by approximately 75% after 20 h treatment, with a significant level of suppression seen as early as 1 h (Table 4). CDCA treatment also suppressed HNF4α and CYP7A1 mRNA levels by approximately 50% and 75%, 20 h after treatment. Also as expected CDCA treatment showed a huge induction of SHP mRNA levels by as much as 25 fold after 20 h of treatment

These data revealed that both IL-1β and CDCA reduced CYP8B1, CYP7A1 and HNF4α mRNA expression levels, but had different effects on SHP mRNA expression in human primary hepatocytes. CDCA, an FXR agonist induced SHP mRNA expression as expected, since SHP is a negative regulator known to be induced by bile acids to inhibit

CYP7A1 expression [106]. The inhibition of SHP mRNA levels by IL-1β are similar to those seen in mouse livers in which LPS treatment lead to a decrease in FXR and its target genes, SHP and apolipoprotein CII by 70 and 60%, respectively [312]. Previously, it has been reported that CDCA feeding (1%) caused a 40-50% reduction in HNF4α mRNA expression in rat livers [135] and CDCA reduced HNF4α protein expression and gene transcription in HepG2 cells [107].

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Table 4. Effect of CDCA on mRNA expression (time course) in human primary hepatocytes (HH 1209) as determined by real time RT-PCR

CDCA 0 hr 1/2 hr 1 hr 2 hr 6 hr 20 hr 50μM

CYP7A1 1 1.02 0.89* 0.80* 0.45* 0.30*

∆Ct±S.D.∆Ct 12.02±0.29 11.98±0.34 12.18±0.32 12.33±0.96 13.15±0.46 13.72±0.42

HNF4α 1 0.97 1 0.91 0.76* 0.51*

∆Ct±S.D.∆Ct 3.94±0.13 3.97±0.48 3.93±0.23 4.06±0.18 4.33±0.20 4.88±0.50

CYP8B1 1 1 0.88* 0.77* 0.58* 0.25*

∆Ct±S.D.∆Ct 4.66±0.97 4.65±0.28 4.83±0.40 5.02±0.38 5.44±0.90 6.62±0.80

SHP 1 4.15* 6.80* 14.37* 15.56* 24.70*

∆Ct±S.D.∆Ct 3.19±0.94 1.14±0.68 0.42±0.45 -0.65±0.35 -0.76±0.23 -1.43±0.27

Reference Gene: Ubiquitin C (UBC)

Relative expression: Relative expression of gene of interest calculated as 2-∆ ∆Ct

∆Ct=Ct of gene of interest - Ct of reference gene

2 2 S.D.ΔCt = Square Root [(Stdev of Ct of reference gene) + (Stdev of gene of interest) ]

* Indicates statistically significant difference (p<0.05).

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Fig.16.

126

Figure 16. Effect of IL-1β on the human CYP8B1 gene transcription.

Transient transfection assays of the human CYP8B1/Luc reporter, p8B1–514/+300/Luc, were performed in HepG2 cells (A) and HEK 293 cells (B) with or without treatment with IL-1β (2ng/ml) for 16 hrs. Reporter assays were carried out as described under

“Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference between IL-1β treated vs. non-treated control (p<0.05).

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Effect of IL-1β on the human CYP8B1 gene transcription

To determine the effect of IL-1β on CYP8B1 gene transcription, reporter assays were

performed in HepG2 cells transiently transfected with the human CYP8B1/Luciferase

reporter plasmid, p8B1-514/+300/Luc, and treated with IL-1β (2ng/ml) for 16 h. IL-1β

inhibited the CYP8B1 reporter activity by about 80% (Fig. 16A). The same experiment

was repeated in the HEK293 cells. Figure 16B shows that IL-1β did not alter the

CYP8B1 reporter activity in HEK293 cells. These results suggest that liver-specific

factors may be required for the inhibitory effect of IL-1β on CYP8B1 gene transcription.

I then performed a transient transfection assay in HepG2 cells treated with increasing

concentrations of IL-1β for 20 h; human CYP8B1 reporter activity was strongly

suppressed by 75% at concentrations as low as 2ng/ml (Fig.17A). The level of

suppression remained the same in concentrations as high as 50ng/ml. A time course

experiment was then performed to determine the time dependent effect of IL-1β on

CYP8B1 reporter activity in transient transfection assay. Suppression of human CYP8B1

reporter activity was observed as early as 6 hours of incubation with IL-1β (2ng/ml). This

suppression reached its maximum (~75%-80%) at around 20 h (Fig. 17B).

Identification of the IL-1β-activated MAPK signaling pathway involved in inhibition of

the CYP8B1 gene transcription

To determine which of the three MAPK pathways are involved in IL-1β inhibition of

CYP8B1 gene transcription, the specific inhibitor of JNK, ERK or p38MAPK was added

in HepG2 cells transfected with p8B1-514/+300/Luc reporter, prior to treatment with IL-

128

Fig.17.

129

Figure 17. Dose Response and Time Course of IL-1β effect on human CYP8B1 gene transcription.

Transient transfection assays of the human CYP8B1/Luc reporter, p8B1–514/+300/Luc, were performed in HepG2 cells. Cells were then either treated with increasing concentrations of IL-1β for 16 hrs (A) or with IL-1β (2ng/ml) for increasing incubation times (B). Reporter assays were carried out as described under “Materials and Methods”.

The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference between

IL-1β treated vs. non-treated control (p<0.05).

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1β. Pre-incubation with the JNK inhibitor, SP600125 (25μM) completely blocked the inhibitory effect of IL-1β on the CYP8B1 reporter activity (Fig. 18A). However, the ERK inhibitor, PD98059 (20μM), and the p38MAPK inhibitor, SB203580 (25μM) had no such effect on the CYP8B1 reporter activity (Fig. 18A). These results suggest that only the

JNK pathway is involved in mediating the inhibitory effect of IL-1β on CYP8B1 gene transcription. To further study the role of the JNK pathway in the suppression of

CYP8B1 gene transcription by IL-1β, expression plasmids for JNK1 were co-transfected with CYP8B1 reporter in HepG2 cells. As shown in Fig. 18B, ectopically expressed

JNK1 inhibited the CYP8B1 basal promoter activity, whereas a dominant negative form of JNK1 (dnJNK1) did not. When c-Jun, a target of JNK1, was co-transfected, the reporter activity was markedly inhibited, whereas a dominant negative c-Jun (dnc-Jun) markedly stimulated CYP8B1 reporter activity. These results suggest that endogenous c-

Jun in HepG2 cells might inhibit CYP8B1 gene transcription. Thus, any up-regulation of

JNK1 or c-Jun will inhibit the CYP8B1 gene transcription. These results further support the finding that the JNK pathway of the MAPK signaling cascade is responsible for mediating the IL-1β inhibition of CYP8B1 gene transcription.

Mapping the IL-1β responsive sequence on the CYP8B1 promoter and determining the

role HNF4α in mediating the cytokine effect on the CYP8B1 promoter.

To map the DNA sequence on the CYP8B1 gene that is responsive to IL-1β, transient

transfection assay was performed using 3’-deletion mutants of the CYP8B1/Luc reporter.

The inhibitory effect of IL-1β on CYP8B1/Luc reporter activity was diminished when the

131

Fig.18.

132

Figure 18. Effect of MAPK inhibitors and JNK overexpression on the IL-1β inhibition of CYP8B1 reporter activity.

A. Transient transfection assay of CYP8B1/Luc reporter (p8B1–514/+300/Luc) was carried out in HepG2 cells. HepG2 cells were pretreated with JNK1 specific inhibitor

SP600125 (25 μM), ERK specific inhibitor PD98059 (20 μM) or p38MAPK specific inhibitor SB203580 (25 μM) for 30 min. The cells were then treated with IL-1β (2ng/ml) for 16 hrs. B. HepG2 cells were cotransfected with expression plasmids for JNK1, dnJNK1, cJun, dncJun, or pcDNA3 empty vector. The luciferase reporter activity of cells transfected with pcDNA3 (control) is set at 1. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference (p <0.05) between IL-1β treated vs. non- treated control (A) and between pcDNA3 control and cotransfection with the other expression plasmids (B).

133

region from +220 to +200 was deleted (Fig. 19). This IL-1β responsive region contains an overlapping DR-1 and FTF binding site that was previously shown to bind HNF4α and FTF, respectively [107]. To investigate the possible roles of HNF4α and/or FTF on mediating the inhibitory effect of IL-1β on the CYP8B1 gene, I used a mutant reporter plasmid in which the HNF4α binding site was mutated but an FTF binding site was created on the reverse strand of the CYP8B1 gene (p8B1mutant/Luc) and assayed the reporter activity in HepG2 cells. As shown in Fig. 20A, mutation of the HNF4α binding site markedly reduced the basal reporter activity and the mutant plasmid was unable to be transactivated by HNF4α; while there was no change in the effect of FTF on the p8B1mutant/Luc reporter activity. To confirm the binding of HNF4α and FTF to the p8B1mutant sequence EMSA assay was done. The left panel of Fig. 20B shows that

HNF4α is unable to bind the wild type p8B1 mutant sequence unlike its binding to the wild type CYP8B1 oligo (+198 to +227). On the other hand the right panel of Fig. 20B shows that FTF can bind both the wild type CYP8B1 and the p8B1 mutant oligo. These findings suggest that the p8B1 mutant oligo can be used to determine the effect of loss of

HNF4α binding while retaining the FTF binding and effect at the overlapping site. Fig.

20C shows that IL-1β did not inhibit the p8B1 mutant/Luc reporter activity unlike the wild type CYP8B1/Luc reporter. This result suggests that the HNF4α binding site is required for mediating IL-1β inhibition of CYP8B1 gene transcription. On the other hand,

FTF does not play a role in mediating IL-1β effect because this mutant reporter has a FTF binding site and does bind FTF. To further corroborate the role of HNF4α in mediating

134

Fig.19.

135

Figure 19. Mapping the region responsive to IL-1β inhibition on the CYP8B1 gene.

Transient transfection assays with the 3’-deletion mutants of the CYP8B1/Luc reporter in

HepG2 cells treated with IL-1β (2ng/ml) for 16 hrs. Reporter assays were carried out as described under “Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference between IL-1β treated vs. non-treated control (p<0.05).

136

Fig.20.

137

Fig.20.

138

Fig.20.

139

Figure 20. Effect of HNF4α binding site mutation on the IL-1β response of CYP8B1 reporter activity.

A. Transient transfection assay of the p8B1mutant/Luc reporter (containing the mutated

HNF4α site of the CYP8B1 promoter) in HepG2 cells, along with cotransfection of

HNF4α and FTF. Sequences of wild type and mutation introduced into human CYP8B1 promoter/luciferase construct are shown at the bottom. Reporter assays were carried out as described under “Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference (p <0.05) between pcDNA3 control and expression plasmid. B. Electrophoretic mobility shift assays (EMSA) showing the binding of invitro translated HNF4α and FTF proteins to 32P labeled oligonucleotide

probe based on the HNF4α binding site in the human CYP8B1 gene (CYP8B1 oligo) and

the p8B1 mutant. The HNF4α and FTF consensus oligos were used as positive controls and for competition assay. One hundred-fold excess of unlabeled probe was used for competition assay. Sequences of all the oligos are shown at bottom of the figure. C.

Transient transfection assay of the p8B1mutant/Luc reporter in HepG2 cells treated with

IL-1β (2ng/ml) for 16 hrs. Sequences of wild type and mutation introduced into human

CYP8B1 promoter/luciferase construct are shown at the bottom. Reporter assays were carried out as described under “Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3.

The * indicates statistically significant difference between IL-1β treated vs. non-treated control (p<0.05).

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IL-1β effect, a dominant negative form of HNF4α (dnHNF4α) was co-transfected into

HepG2 cells to test its effect on reporter activity. Fig. 21A shows that dnHNF4α strongly inhibited basal reporter activity. This dnHNF4α has a defective DNA binding domain but is able to dimerize with wild type HNF4α [313], thus markedly reducing CYP8B1 gene expression. Addition of IL-1β did not further reduce the reporter activity. These results support our finding that HNF4α is critical for trans-activating the CYP8B1 gene and IL-

1β inhibits HNF4α induction of the human CYP8B1 gene transcription. Furthermore, over-expression of HNF4α stimulated the human CYP8B1 promoter activity as expected, however IL-1β was not able to suppress the CYP8B1 promoter activity stimulated by

HNF4α (Fig. 21B). As a negative control, IL-1β was able to suppress CYP8B1 reporter activity in cells over-expressing FTF. These data further support that HNF4α is involved in IL-1β inhibition of the CYP8B1 gene.

Effect of IL-1β on HNF4α gene expression.

To determine if IL-1β had any direct effect on HNF4α gene transcription, the

HNF4α/Luc reporter was transfected into HepG2 cells, which were subsequently treated

with IL-1β. Fig. 22A shows that IL-1β decreases the HNF4α reporter activity. These

results are consistent with IL-1β inhibition of HNF4α mRNA expression in primary

hepatocytes (Table 1 and 3) and suggested that inhibition of HNF4α gene transcription

by IL-1β may be responsible for the suppression of CYP8B1 by IL-1β. Since IL-1β

treatment decreased HNF4α reporter activity and mRNA levels, it was interesting to see

141

Fig.21.

142

Figure 21. Effects of IL-1β on HNF4α stimulation of CYP8B1 reporter activity.

A. Transient transfection assays of the wild type CYP8B1/Luc reporter (p8B1–

514/+300/Luc) in HepG2 cells cotransfected with expression plasmid dnHNF4α and pcDNA3 (control) and treated with IL-1β (2ng/ml) for 16 hrs. B. Transient transfection assays with p8B1–514/+300/Luc in HepG2 cells cotransfected with expression plasmids for HNF4α, FTF, or pcDNA3 (control) and treated with IL-1β (2ng/ml) for 16 hrs.

Reporter assays were carried out as described under “Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference (p<0.05) between untreated control and IL-1β treated.

143

Fig.22.

144

Figure 22. Effects of IL-1β on HNF4α gene transcription and protein expression in HepG2 cells.

A. Transient transfection assay of the HNF4α/Luc reporter in HepG2 cells treated with

IL-1β (2ng/ml) for 16 hrs. Reporter assays were carried out as described under “Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference (p<0.05) between untreated control and IL-1β treated. B. Immunoblots of

HNF4α protein in HepG2 cells. Total cell proteins were extracted from HepG2 cells

treated with IL-1β in the concentrations indicated. An equal amount of each of the

protein samples was analyzed by immunoblot analysis using polyclonal antibody against

HNF4α (upper panel) or actin (lower panel).

145

if it also alters the HNF4α protein levels. HepG2 cells were treated with increasing concentrations of IL-1β and immunoblot analysis with HNF4α antibody was performed to estimate the amount of HNF4α protein in the control and treated cells. The amount of

HNF4α protein in HepG2 cells decreased in a dose-dependent manner by IL-1β (Fig.

22B). Intracellular actin levels were determined as an internal control. Thus, it can be concluded that IL-1β inhibition of HNF4α transcriptional activity leads to a decrease in

HNF4α mRNA and protein levels, and is responsible for the suppression of CYP8B1

gene transcription.

Effect of IL-1β on HNF4α binding to the CYP8B1 gene.

As it was reported previously [107] and as shown in Fig. 20B HNF4α bound to the DR-1 element on the human CYP8B1 promoter. To determine if IL-1β treatment alters the

ability of HNF4α to bind to the DR-1 element, EMSA of an oligonucleotide probe based

on the DR-1 element of the human CYP8B1 gene was performed using nuclear extracts

prepared from HepG2 cells treated with increasing concentrations of IL-1β. As shown in

Fig. 23A, increasing concentrations of IL-1β decreased the HNF4α binding to the probe.

It is likely that IL-1β treatment reduced nuclear HNF4α protein or its DNA binding

activity. This result is consistent with IL-1β inhibition of HNF4α gene transcription

shown in Fig. 22A and supports the hypothesis that IL-1β reduces HNF4α trans-

activation of the CYP8B1 gene. To confirm that IL-1β reduces HNF4α binding to the

CYP8B1 promoter in cells, chromatin immunoprecipitation (ChIP) assay was performed.

146

Fig.23.

147

Figure 23. Electrophoretic mobility shift assays and chromatin immunoprecipitation assays of the effect of IL-1β on HNF4α binding to the CYP8B1 gene.

A. EMSA. Nuclear proteins were extracted from HepG2 cells treated with IL-1β in the

concentrations indicated. Equal amount of proteins was used for EMSA of 32P labeled oligonucleotide probe based on the HNF4α binding site in the human CYP8B1 gene

(sequence shown at bottom of figure). One hundred-fold excess of unlabeled probe was used for competition assay. HNF4α antibody was added for supershift assay of antibody-

DNA-protein complex. B. ChIP assays. HepG2 cells were treated with IL-1β (10ng/ml)

or CDCA (50μM) for 20 hrs. ChIP assays were performed as described under

Experimental Procedures. Anti-HNF4α antibody (upper panel) or non- immune IgG

(middle panel) was used to precipitate the cross-linked chromatins. A 251 bp fragment

containing the HNF4α binding site on the CYP8B1 promoter was PCR amplified and

analyzed on a 1.5% agarose gel.

148

HepG2 cell extracts (control and IL-1β treated) were immunoprecipitated with HNF4α antibody and the amount of the immunoprecipitated DNA fragments containing the

HNF4α binding site on the CYP8B1 promoter was quantified by PCR. As shown in Fig.

23B, IL-1β (10ng/ml) treatment reduces the amount of HNF4α bound to the CYP8B1 chromatin as indicated by reduced levels of anti-HNF4α antibody precipitated chromatins. Also, 50 μM of CDCA reduced the HNF4α bound to the CYP8B1 chromatin.

Non-immune IgG was used as a negative control of ChIP assay. This data is consistent with the real time RT-PCR results (Table 1, 2, 3 and 4). Thus both the EMSA and ChIP assay results suggest that IL-1β reduces the HNF4α binding to the CYP8B1 chromatin.

Effect of the MAPK inhibitors on the IL-1β mediated inhibition of the HNF4α protein expression.

To determine if the JNK pathway had any effect on the HNF4α protein expression levels,

I performed an immunoblot assay. HepG2 cells were preincubated for 30 min with the specific inhibitors of the JNK, ERK or p38MAPK prior to treatment with IL-1β (5ng/ml) for 16 h. Pretreatment with the JNK specific inhibitor SP600125 (25 μM) was able to block the IL-1β-mediated suppression of HNF4α protein levels (Fig. 24). However, treatment with the ERK-specific inhibitor PD98059 (20 μM) and the p38 MAPK-specific inhibitor SB203580 (25μM) did not block IL-1β suppression of the HNF4α protein levels. The results suggest that the JNK pathway mediates the effect of IL-1β on the

CYP8B1 promoter, possibly by the regulation of the HNF4α protein expression levels.

149

Fig.24.

150

Figure 24. Immunoblot analysis of the effects of MAP kinase inhibitors on the IL-1β mediated inhibition of the HNF4α protein expression.

HepG2 cells were pretreated with JNK1 specific inhibitor SP600125 (25 μM), ERK specific inhibitor PD98059 (20 μM) or p38MAPK specific inhibitor SB203580 (25 μM) for 30 min. The cells were then treated with IL-1β (5ng/ml) for 16 hrs and total cell proteins were extracted for immunoblot analysis. An equal amount of each of the protein samples was analyzed by immunoblot analysis using polyclonal antibody against HNF4α

(upper panel) or actin (lower panel).

151

JNK1 phosphorylation of HNF4α and the effect of JNK1 overexpression on the

HNF4α binding/ recruitment to the CYP8B1 promoter.

Apart from the decrease in HNF4α gene expression, I wanted to test the hypothesis that a

post translational modification of HNF4α protein is involved in mediating the IL-1β

effect on CYP8B1. It is known that HNF4α is phosphorylated by protein kinase A (PKA)

[314], ERK [315] and AMP kinase [316, 317], and that phosphorylated HNF4α loses its

DNA binding activity [318]. Since the JNK was shown to be activated in HepG2 cells by

IL-1β [239], I was interested in seeing if JNK1 could directly phosphorylate HNF4α.

Purified GST-tagged HNF4α fusion protein was incubated with active JNK1 in the

presence of γ32P-labeled ATP. JNK1 could phosphorylate the GST: HNF4α fusion

protein, but did not phosphorylate the GST protein (Fig. 25A). Without JNK, HNF4α

was not phosphorylated. As a known positive control target for JNK1 phosphorylation,

JNK1 strongly phosphorylated ATF2. I also performed a positive control for AMPK

phosphorylation of HNF4α. As shown in Fig. 25B, AMP-activated AMPK

phosphorylated HNF4α and the positive control, SAMS peptide

(HMRSAMSGLHLVKRR). Without AMPK, HNF4α and SAMS were not

phosphorylated. These results suggest that JNK1 was able to directly phosphorylate

HNF4α.

To determine if JNK1 overexpression could alter HNF4α binding to the CYP8B1

chromatin in vivo, I overexpressed JNK1 in HepG2 cells and performed a ChIP assay.

152

Fig.25.

153

Figure 25. Phosphorylation of HNF4α by JNK1 and the effect of JNK1 overexpression on the HNF4α binding/ recruitment to the CYP8B1 promoter.

A. and B. In vitro kinase assays of purified GST or GST-HNF4α proteins by active JNK1

(A) or AMPK (B) were performed as described under “Materials and Methods”. ATF2

and SAMS peptide were used as positive controls for JNK1 and AMPK kinase activities,

respectively. Reactions are performed with or without 300 μM AMP in case of AMPK.

The samples were then run on a SDS-PAGE, dried and autoradiographed. C. HepG2 cells

were transfected with pcDNA3 or JNK1 expression plasmids; 48 hrs later ChIP assay

was performed on the cells as described under “Materials and Methods”. The cross-

linked cell chromatins were precipitated with Anti-HNF4α antibody. Non- immune IgG

(middle panel) was used as control to precipitate the cross-linked chromatins. A 251 bp

fragment containing the HNF4α binding site on the CYP8B1 promoter was PCR

amplified and analyzed on a 1.5% agarose gel.

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Fig.25C. shows that cells overexpressing JNK1 show a decreased HNF4α binding to the

CYP8B1 chromatin, in comparison to the pcDNA3 overexpressing cells used as control.

Effect of IL-1β on c-Jun expression.

c-Jun, a downstream target of the JNK pathway has also been shown to be activated by

bile acids [218]. To test the hypothesis that IL-1β may affect c-Jun expression, c-Jun mRNA levels were determined by quantitative real time PCR in human primary hepatocytes treated with IL-1β (10ng/ml) and CDCA (50μM) for increasing lengths of time. Fig. 26A, c-Jun mRNA levels increased upon both IL-1β and CDCA treatment, as early as 2 h after treatment. After 20 h treatment with IL-1β, c-Jun mRNA levels were

increased to 3 fold above the untreated samples, while CDCA treatment increased c-Jun

levels to 5.5 fold above the untreated samples. The mRNA results were confirmed by

immunoblot analysis in HepG2 cells which were treated with IL-1β (5ng/ml) or CDCA

(25μM). As shown in Fig. 26B, IL-1β treatment decreased HNF4α protein levels as

expected (Table 3) and increased c-Jun protein levels in concordance with the mRNA

data of Fig. 26A. Also, since activated JNK can phosphorylate c-Jun, the phosphorylated

c-Jun levels were measured in the samples and a time dependent increase in its levels

were noted. Fig. 26C shows that CDCA treatment of HepG2 cells increases the protein

levels of c-Jun and phosphorylated c-Jun, while decreasing the levels of HNF4α. These

results correlate to the mRNA data in Fig. 26A and Table 4.

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Fig.26.

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Figure 26. Effect of IL-1β and CDCA on the mRNA and protein expression of cJun.

A. Human primary hepatocytes were treated with IL-1β (10ng/ml) or CDCA (50μM) for different time periods as indicated in the figure. Quantitative Real time PCR was performed on the RNA extracted from the cells as described in “Materials and Methods”.

The relative mRNA levels of cJun is calculated with respect to the UBC mRNA expression and expressed as 2–ΔΔCt for each time point is shown in the figure. ΔCt=Ct of

gene of interest-Ct of reference gene. The error bars represent the standard deviation of

2–ΔΔCt from three different experiments. N=3. B. HepG2 cells were treated with IL-1β

(5ng/ml) for the indicated time periods and total cell proteins were extracted for

immunoblot analysis. C. HepG2 cells were treated with CDCA (25μM) for the indicated time periods and total cell proteins were extracted for immunoblot analysis. For both B

and C, an equal amount of each of the protein samples was analyzed by immunoblot

analysis using polyclonal antibody against cJun (first panel), phospho cJun (second

panel), HNF4α (third panel) or actin (fourth panel).

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Effect of c-Jun and dnc-Jun over expression on CYP8B1 gene transcription.

The marked up regulation of c-Jun mRNA and protein levels seen on both IL-1β and

CDCA treatment led me to investigate how this factor played a role in the inhibition of

CYP8B1 gene transcription. In transient transfection assay using the human

CYP8B1/Luciferase reporter (p8B1-514/+300/Luc) in HepG2 cells, over expression of c-

Jun led to 80% suppression of the CYP8B1 reporter activity (Fig.27), while over expression of a dominant negative c-Jun (dnc-Jun) protein showed a greater than 4 fold increase in the CYP8B1 reporter activity. These results are similar to Fig. 18B.

HNF4α over expression alone stimulated the CYP8B1 reporter as expected (more than 3 fold), but along with increasing concentrations of c-Jun showed a significant decrease in

CYP8B1 reporter activity; while along with increasing concentrations of dnc-Jun showed a synergistically high level of stimulation (greater than 20 fold) of the CYP8B1 reporter activity. HNF4α and coactivator PGC-1α overexpressed together, further stimulated the

CYP8B1 reporter activity (more than 6 fold), however this stimulation was overcome by c-Jun overexpression in a dose dependent manner. On the other hand, dnc-Jun was unable to overcome the HNF4α and PGC-1α mediated stimulation of CYP8B1 reporter activity and instead together with these two plasmids showed almost a 30 fold induction in

CYP8B1 reporter activity. This result indicates that c-Jun inhibits the CYP8B1 promoter activity and can prevent the HNF4α and PGC-1α mediated stimulation of the CYP8B1 promoter activity. On the contrary, dnc-Jun causes a huge stimulation of CYP8B1 reporter activity both alone or in conjunction with HNF4α and PGC-1α.

158

Fig.27.

159

Figure 27. Effect of cJun, dncJun, PGC-1α and HNF4α cotransfection on human CYP8B1 promoter activity.

Human CYP8B1 promoter/luciferase construct (p8B1–514/+300/Luc, 0.1 μg) was cotransfected into HepG2 cells with 0.01 μg β-gal expression plasmid. The expression plasmids for cJun, dncJun, PGC-1α and HNF4α were transfected in the concentrations as indicated. Reporter assays were carried out as described under “Materials and Methods”.

The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The *, @, # and % indicate a statistically significant

difference (p<0.05) between the different bars as indicated by the lines.

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Evidence of protein-protein interaction between HNF4α and c-Jun.

Since HNF4α is the only known major factor that stimulates CYP8B1 gene transcription

and since c-Jun cotransfection with HNF4α could negate the stimulatory effect of

HNF4α on the CYP8B1 reporter I decided to test if these two proteins could interact with

each other. I performed a Co-IP assay in HepG2 cells treated with IL-1β (5ng/ml) for

increasing time periods. Cell lysates were immunoprecipitated with anti-HNF4α

antibody, followed by western blots with antibodies against HNF4α, c-Jun and phospho

c-Jun. As expected HNF4α protein levels decreased in a time dependent manner upon IL-

1β (5ng/ml) treatment (Fig.28A). Both c-Jun and phospho c-Jun were found to be

associated with HNF4α both in the absence and presence of IL-1β treatment. However

after IL-1β treatment, I observed that the amount of c-Jun and phospho c-Jun associated

with HNF4α increased time dependently, even though the HNF4α levels decreased. To

quantify my results I divided the intensity of the c-Jun protein bands by that of the

HNF4α protein bands at the corresponding time periods to obtain a relative intensity, and set the relative intensity of time point 0 h at 1. As seen in Fig.28B, the relative intensity

of c-Jun/ HNF4α increased by greater than two – fold after 20 h of IL-1β treatment. This

result is consistent with the result from Fig. 26A and 26B which show that IL-1β

treatment increases the c-Jun mRNA and protein levels significantly. Since IL-1β

activates the JNK pathway, which leads to the increase in phosphorylation of c-Jun by

JNK, the phospho c-Jun levels increase on IL-1β treatment. In Fig.28C, it is shown that

the relative intensity of phospho c-Jun/ HNF4α increased by greater than three – fold

161

Fig.28.

162

Fig.28.

163

Figure 28. Effect of IL-1β treatment on the interaction between HNF4α with cJun or phospho cJun in vivo and GST Pull down assay to study the interaction between HNF4α and cJun.

A. HepG2 cell extracts treated with IL-1β (10ng/ml) for different periods of time were immunoprecipitated with Rabbit Anti-HNF4α antibody as described in “Materials and

Methods”. Immunoblot analysis was performed with Goat Anti-HNF4α, Anti-cJun and

Anti-pcJun antibodies. Equal Amount of the Input (5%) from each sample was also run on an SDS PAGE gel and immunoblotted with Anti-Actin antibody. Rabbit Non-Immune

IgG was used as a negative control. B. The intensity of the cJun bands were divided by the intensity of the HNF4α bands and expressed as relative intensity at each time point.

C. The intensity of the phospho-cJun bands were divided by the intensity of the HNF4α bands and expressed as relative intensity at each time point. D. Bacterially expressed

GST or GST- HNF4α bound glutathione beads were incubated with either 35S labeled

invitro translated cJun, PGC-1α or both in the amounts indicated in the “Materials and

Methods”. The 35S labeled proteins complexed with the GST proteins were run on an

SDS PAGE, dried and autoradiographed. 20% of the total 35S labeled proteins was run as

input on an SDS PAGE.

164

after 20 h of IL-1β treatment. My results here provide evidence that c-Jun and HNF4α interact with each other and that this association is further increased on IL-1β treatment.

To provide an additional proof of the c-Jun interaction with HNF4α, I performed a GST pull down assay. Bacterially expressed GST-HNF4α protein was able to pull down invitro translated c-Jun protein (Fig 28D). Invitro translated PGC-1α, which is a potent co activator known to interact with HNF4α, was used as a positive control. In the last two lanes of the figure, both invitro translated PGC-1α and c-Jun were added, and I noticed that as the amount of c-Jun interacting with HNF4α increases, the amount of PGC-1α associated with HNF4α decreases. This suggests that the c-Jun interaction with HNF4α can reduce the binding of co activator PGC-1α with HNF4α, thus decreasing the

CYP8B1 promoter activity.

Effect of c-Jun on the recruitment of HNF4α to the CYP8B1 chromatin.

After it was determined that c-Jun and HNF4α interact with each other, I wanted to see if

c-Jun could affect the binding of HNF4α to the CYP8B1 promoter sequence. The EMSA

shown in Fig. 29A shows the binding of in vitro translated HNF4α to the CYP8B1 oligo

(+198 to +227) which contains the DR-1 site. c-Jun is known to bind the consensus AP-1

sequence both alone (as a homodimer) or along with c-Fos (as a heterodimer) as shown in

the right hand side of Fig.29A. The CYP8B1 oligo (+198 to +227) oligo was unable to

bind c-Jun or c-Fos or the combination of the two. When c-Jun or c-Fos or the

combination of the two was added along with HNF4α to the reaction mixture, the binding

165

Fig.29.

166

Fig.29.

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Figure 29. Electromolibility shift assay and chromatin immunoprecipitation assays to study the effect of cJun on the HNF4α binding/ recruitment to the CYP8B1 promoter.

A. EMSA showing the binding of invitro translated either HNF4α, cJun or cFos proteins,

alone or in combination to 32P labeled oligonucleotide probe based on the HNF4α

binding site in the human CYP8B1 gene (CYP8B1 oligo) and the AP-1 consensus

sequence (control). HNF4α or cJun antibody was added for supershift assay of antibody-

DNA-protein complex. Sequences of the oligos are shown at bottom of the figure. In B,

C and D, ChIP assays were performed on HepG2 cells as described under “Materials and

Methods”. Non- immune IgG (middle panels) was used as control to precipitate the cross-

linked chromatins. A 251 bp fragment containing the HNF4α binding site on the CYP8B1

promoter was PCR amplified and analyzed on a 1.5% agarose gel. B. Untreated cell

chromatins were precipitated with Anti-cJun antibody. C. Cells were transfected with pcDNA3 or cJun expression plasmids and 48 hrs later the cell chromatins were precipitated with Anti-HNF4α antibody. D. Cells were transfected with HA-PGC-1α expression plasmid and with pcDNA3 or cJun expression plasmids and 48 hrs later the cell chromatins were precipitated with Anti-HA antibody.

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of HNF4α to CYP8B1 oligo remained unaltered. This data proved that even though c-Jun interacted with HNF4α, it did not affect the binding of HNF4α to the CYP8B1 promoter sequence.

Since c-Jun reduced HNF4α trans-activation of CYP8B1 gene transcription, I used a c-

Jun antibody to immunoprecipitate HepG2 cells in a ChIP assay to detect c-Jun in

CYP8B1 chromatin. Fig.29B shows that c-Jun is associated with the CYP8B1 chromatin at the HNF4α binding site, most likely via an interaction with HNF4α, since the EMSA in Fig. 29A showed no evidence of direct binding of c-Jun to the CYP8B1 chromatin.

To determine if c-Jun affect HNF4α binding to the CYP8B1 chromatin in vivo, I over expressed c-Jun in HepG2 cells and performed a ChIP assay. Fig.29C shows that cells overexpressing c-Jun do not display any difference in HNF4α binding to the CYP8B1 chromatin, in comparison to the pcDNA3 over expressing cells used as control. This result indicates that the c-Jun interaction with HNF4α does not affect the DNA binding of HNF4α to the CYP8B1 chromatin and confirms the invitro data in Fig. 29A.

Since HNF4α is known to interact with PGC-1α, I did a ChIP assay using extracts from

HepG2 cells overexpressing HA-tagged PGC-1α, either alone or along with c-Jun overexpression and immunoprecipitated with anti-HA antibody. Fig.29D shows that

PGC-1α was associated with CYP8B1 chromatin, mostly likely by an interaction with

HNF4α, since PGC-1α being a coactivator does not bind DNA directly. Interestingly, cJun overexpression markedly reduced PGC-1α recruitment to the CYP8B1 chromatin.

This data suggests that the interaction between c-Jun and HNF4α may block the PGC-1α

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coactivator recruitment by HNF4α on the CYP8B1 chromatin and result in the inhibition of CYP8B1 gene transcription. This result is consistent with the GST pull down assay data in Fig. 28D.

Effect of IL-1β on SHP expression.

To determine if the IL-1β effect on CYP8B1 is SHP dependent/ independent, it was

important to see the effect of IL-1β on SHP transcription. Transient transfection assay

with the wild type human SHP/Luc reporter was performed in HepG2 cells, followed by

treatment with IL-1β (2ng/ml) for 16 h. I observed that IL-1β attenuated the hSHP/Luc

reporter activity (Fig.30A). This result was consistent with IL-1β inhibition of SHP

mRNA expression in human primary hepatocytes (Table 1 and 3). Since SHP is a

negative nuclear receptor whose expression is inhibited by IL-1β, it would imply that the

IL-1β suppression of CYP8B1 expression is mediated via a SHP independent pathway.

Thus, the IL-1β signaling pathway may serve as an explanation to the bile acid mediated

repression of Cyp8b1 mRNA expression in Shp-/- mice [165, 166]. Analysis of the

human SHP promoter revealed the presence of a DR-1 site (-563TGGACAGTGGGCA-

551). This led me to hypothesize that HNF4α may regulate the transcription of SHP and that the decrease in HNF4α levels caused by IL-1β may in turn reduce the expression of

SHP.

170

Fig.30.

171

Fig.30.

172

Figure 30. Effect of IL-1β and HNF4α on the human SHP gene transcription.

A. Transient transfection assays of the human SHP/Luc reporter, hSHP/Luc, was performed in HepG2 cells with or without treatment with IL-1β (2ng/ml) for 16 hrs. B.

Transient transfection assays of the hSHP/Luc, was performed in Hek293 cells along with cotransfection of increasing amounts of HNF4α expression plasmid as indicated in the figure. C. Transient transfection assays of the hSHP/Luc, was performed in Hek293 cells along with cotransfection of increasing amounts of HNF4α and dnHNF4α expression plasmid as indicated in the figure. All the reporter assays were carried out as described under “Materials and Methods”. The error bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * indicates statistically significant difference between IL-1β treated vs. non-treated control or between pcDNA3 only control and cotransfection with the other expression plasmids

(p<0.05). The # indicates statistically significant difference (p<0.05) between the different bars as indicated by the line.

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Role of HNF4α in regulation of SHP expression.

The hSHP/Luc reporter was transfected in Hek293 cells, along with cotransfection of

increasing amounts of HNF4α expression plasmid. As shown in Fig. 30B, overexpression

of HNF4α increases the hSHP/Luc reporter activity dose dependently, to as much as an

eight-fold increase with 200ng expression. The same experiment was also repeated in

HepG2 cells; even though HNF4α did stimulate the hSHP/Luc activity, the effect was not as pronounced as in the Hek293 cells possibly due to the high levels of HNF4α expression in HepG2 cells which may contribute to an already activated SHP transcription. To confirm the effect of HNF4α on SHP gene transcription; dnHNF4α was transfected along with the HNF4α expression plasmid and the hSHP/Luc reporter in

Hek293 cells. dnHNF4α significantly decreased the stimulatory effect of HNF4α on the hSHP/Luc reporter activity (Fig.30C). Since dnHNF4α has a defective DNA binding domain but is able to dimerize with wild type HNF4α, it would prevent the homodimerization of the wild type HNF4α and decrease the transcription activity of the

SHP promoter.

To determine if HNF4α does bind the putative DR-1 site on the human SHP promoter, an

EMSA was done. In vitro translated HNF4α protein bound the oligo containing the DR-1 site and this binding was confirmed by both antibody supershift with HNF4α antibody and cold competition with HNF4α consensus binding sequence oligo (Fig. 31). Thus my results prove that HNF4α directly binds and transactivates the human SHP gene and in

174

Fig.31.

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Figure 31. Electromobility shift assay to show the binding of HNF4α to the human SHP gene.

EMSA showing the binding of invitro translated HNF4α protein to 32P labeled

oligonucleotide probe based on the HNF4α binding site in the human SHP gene (hSHP

DR-1 oligo), the mutant of that site (M hSHP DR-1 oligo) or the HNF4α consensus oligo

(control). One hundred-fold excess of unlabeled probe was used for competition assay.

HNF4α antibody was added for supershift assay of antibody-DNA-protein complex.

Sequences of the oligos are shown at bottom of the figure.

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the presence of IL-1β the decrease in HNF4α expression could contribute to the decrease in SHP expression.

Effect of c-Jun and dnc-Jun over expression on CYP7A1 gene transcription.

It was shown earlier that IL-1β suppressed the CYP7A1 gene transcription via a JNK mediated pathway, and that this effect was lost upon mutation of the HNF4α binding site on the human CYP7A1 promoter (Jahan A, Li T and Chiang J, unpublished data). The marked up regulation of c-Jun mRNA and protein levels seen on both IL-1β and CDCA treatment and shown to play a role in suppression of CYP8B1, led me to investigate if c-

Jun could also inhibit CYP7A1 gene transcription.

In transient transfection assay using the human CYP7A1/luciferase reporter (ph-298/Luc) in HepG2 cells, over expression of c-Jun led to 80% suppression of the CYP7A1 reporter activity (Fig.32), while over expression of the dnc-Jun protein showed no change in the

CYP7A1 reporter activity. HNF4α over expression alone stimulated the CYP7A1 reporter as expected (more than 2 fold), but along with increasing concentrations of c-Jun showed a significant decrease in CYP7A1 reporter activity; while along with increasing concentrations of dnc-Jun showed no significant difference than HNF4α cotransfection alone. HNF4α and PGC-1α overexpressed together, further stimulated the CYP7A1 reporter activity (more than 4 fold), however this stimulation was overcome by c-Jun overexpression in a dose dependent manner. On the other hand, dnc-Jun was unable to overcome the HNF4α and PGC-1α mediated stimulation of CYP7A1 reporter activity.

This result indicates that the presence of c-Jun down regulates the CYP7A1 promoter

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Fig.32.

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Figure 32. Effect of cJun, dncJun, PGC-1α and HNF4α cotransfection on human CYP7A1 promoter activity.

Human CYP7A1 promoter/luciferase construct (ph-298/luc, 0.1 μg) was co-transfected

into HepG2 cells with 0.01 μg β-gal expression plasmid. The expression plasmids for

cJun, dncJun, PGC-1α and HNF4α were transfected in the concentrations as indicated.

Reporter assays were carried out as described under “Materials and Methods”. The error

bars represent the standard deviation from the mean of triplicate assays of an individual experiment. N=3. The * and # indicate a statistically significant difference (p<0.05)

between the different bars as indicated by the lines.

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activity and can prevent the HNF4α and PGC-1α mediated stimulation of the CYP7A1 promoter activity.

Effect of c-Jun on the recruitment of HNF4α and PGC-1α to the CYP7A1 chromatin.

ChIP assay was used to assay the effect of IL-1β and CDCA on HNF4α recruitment to

CYP7A1 chromatin. An antibody against HNF4α was used to immunoprecipitate HepG2

extracts for PCR amplification of a 391 fragment covering both BARE-I and BARE-II

(HNF4α binding site) of the human CYP7A1 promoter. As seen in Fig.33A, IL-1β

(10ng/ml) [left panel] and CDCA (50μM) [right panel] treatment for 20 h, respectively,

decreased the amount of HNF4α protein associated with the CYP7A1 chromatin. These

results are consistent with the decrease in HNF4α mRNA and protein expression by IL-

1β and CDCA.

Since c-Jun reduced HNF4α trans-activation of CYP7A1 gene transcription, I used a c-

Jun antibody to immunoprecipitate HepG2 cells in a ChIP assay to detect c-Jun in

CYP7A1 chromatin. Fig.33B shows that that c-Jun is associated with the CYP7A1

chromatin at the HNF4α binding site, most likely via an interaction with HNF4α.

To determine if c-Jun affect HNF4α binding to the chromatin, c-Jun was over expressed

in HepG2 cells and a ChIP assay was done. Fig.33C shows that cells overexpressing c-

Jun do not show any difference in HNF4α binding to the CYP7A1 chromatin, in

comparison to the pcDNA3 (empty vector) overexpressing cells used as control. This

result indicates that the c-Jun interaction with HNF4α does not affect the DNA binding

180

Fig.33.

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Figure 33. Chromatin Immunoprecipitation assays to study the effect of IL-1β and CDCA treatment and pcDNA3, cJun or JNK1 overexpression on the HNF4α binding/ recruitment to the CYP7A1 promoter.

ChIP assays were performed on HepG2 cells as described under Materials and Methods.

Non- immune IgG (middle panels) was used as control to precipitate the cross-linked

chromatins. A 391 bp fragment containing the BAREI and BAREII regions of the

CYP7A1 promoter was PCR amplified and analyzed on a 1.5% agarose gel. A. Cells were

treated with IL-1β (10ng/ml) [left panel] or CDCA (50 μM) [right panel] for 20 hrs and

Anti-HNF4α antibody was used to precipitate chromatins. B. Untreated cell chromatins

were precipitated with Anti-cJun antibody. C. Cells were transfected with pcDNA3 or

cJun expression plasmids and 48 hrs later the cell chromatins were precipitated with

Anti-HNF4α antibody. D. Cells were transfected with HA-PGC-1α expression plasmid

and with pcDNA3 or cJun expression plasmids and 48 hrs later the cell chromatins were

precipitated with Anti-HA antibody. E. Cells were transfected with pcDNA3 or JNK1

expression plasmids and 48 hrs later the cell chromatins were precipitated with Anti-

HNF4α antibody.

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of HNF4α to the CYP7A1 chromatin. Since HNF4α is known to interact with PGC-1α and recruit the latter to the CYP7A1 chromatin [120], I did a ChIP assay using extracts from HepG2 cells overexpressing HA-tagged PGC-1α, either alone or along with c-Jun overexpression and immunoprecipitated the cells with a anti-HA antibody. Fig.33D shows that PGC-1α was associated with CYP7A1 chromatin, mostly likely by an interaction with HNF4α, since PGC-1α does not bind to DNA and that c-Jun overexpression markedly reduced PGC-1α recruitment to the chromatin. These data suggest that the interaction between c-Jun and HNF4α may block the PGC-1α coactivator recruitment by HNF4α on the CYP7A1 chromatin resulting in the inhibition of CYP7A1 gene transcription. Thus, I proved that the c-Jun mediated mechanism of suppression of CYP8B1, was also applicable to the CYP7A1 gene.

To determine if JNK1 overexpression could alter HNF4α binding to the CYP7A1 chromatin in vivo, I overexpressed JNK1 in HepG2 cells and performed a ChIP assay.

Fig.33 E. shows that cells overexpressing JNK1 show a decreased HNF4α binding to the

CYP7A1 chromatin, in comparison to the pcDNA3 overexpressing cells used as control.

Effect of siRNA mediated downregulation of HNF4α on CYP7A1 transcription.

HNF4α levels were knocked down in HepG2 cells using an oligobased siRNA method,

to determine the effect on CYP7A1 gene transcription. HNF4α mRNA levels were

decreased by 70% in siHNF4α transfected HepG2 cells (Fig. 34A). The same cells

showed a 60% repression of CYP7A1 mRNA levels (Fig. 34A); once again underscoring

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Fig.34.

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Figure 34. Effect of HNF4α small interference RNA (siRNA) transfection with or without IL-1β or CDCA.

Double-stranded HNF4α siRNA oligo (100nM) transfection of HepG2 cells was

described in “Materials and Methods”. The RNA was collected, 48 hrs post transfection

and quantitative real time PCR for HNF4α, CYP7A1, PEPCK and SHP mRNA

expression was done (A). Results indicate that HNF4α, CYP7A1 and PEPCK mRNA

expression were repressed while SHP mRNA expression remained unchanged in HNF4α

siRNA oligo (100nM) transfected HepG2 cells. In Fig 34 B. and C., the cells were

treated with IL-1β (10ng/ml) or CDCA (50μM) for 20 hrs before harvesting of the cells.

RNA was collected and quantitative real time PCR for HNF4α (B) and CYP7A1 (C) mRNA expression was done. Results indicate that IL-1β (10ng/ml) and CDCA (50μM) treatment of HNF4α siRNA oligo (100nM) transfected cells suppressed the already reduced HNF4α and CYP7A1 mRNA levels even further. Relative mRNA levels were calculated with respect to UBC mRNA expression and expressed as 2–ΔΔCt. (ΔCt=Ct of

gene of interest-Ct of reference gene; Error bars indicate the SDΔCt calculated as SDΔCt =

Square Root [(Stdev of Ct of reference gene)2 + (Stdev of gene of interest)2]; * p<0.05

compared to untreated cells; † p<0.05 compared to untreated HNF4α siRNA oligo

(100nM) transfected cells)

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the vital role played by HNF4α in transactivating and maintaining the basal transcriptional activity of CYP7A1.

Phosphoenolpyruvate carboxykinase (PEPCK) is the rate-limiting enzyme of the gluconeogenesis. PEPCK is a highly inducible protein, which is usually expressed at very low levels in hepatocytes. In the fasting condition, PEPCK gene transcription is stimulated several fold to increase glucose synthesis by the liver. Glucagon, epinephrine, insulin, glucocorticoids, and metabolic acidosis regulate PEPCK gene transcription [319].

HNF4α is known to bind and strongly stimulate PEPCK gene transcription [320]. Hence,

PEPCK mRNA levels were measured. PEPCK mRNA levels were decreased by almost

80% in siHNF4α transfected HepG2 cells, indicating that HNF4α knockdown decreased

PEPCK gene transcription (Fig. 34A). Interestingly, mRNA levels of SHP remained unchanged in siHNF4α transfected HepG2 cells (Fig. 34A). This indicated that the SHP gene was not strongly regulated by HNF4α in HepG2 cells, similar to my earlier results in transfection assays. The SHP mRNA levels and the unchanged Ct values of the internal

reference gene UBC, indicated that the siHNF4α mediated response was specific to the

siRNA used.

Upon IL-1β (10ng/ml) and CDCA (50μM) treatment of siHNF4α transfected cells the

already reduced HNF4α and CYP7A1 mRNA levels were suppressed even further (Fig.

34B and 34C). Since the HNF4α levels were not completely knocked down by the

siRNA oligo (Fig. 34B and 34C), the remaining HNF4α still being expressed, was

regulated by IL-1β and CDCA. This could partly account for the suppression of CYP7A1

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by these compounds in siHNF4α transfected cells. However another possibility is the presence of factors other than HNF4α that may be responsible for the suppression of

CYP7A1 by both IL-1β and CDCA. One such factor is c-Jun, whose levels increase on treatment with these compounds and may mediate suppression of CYP7A1.

CHAPTER IV

DISCUSSION

An important component of bile acid homeostasis is the feedback inhibition of bile acid biosynthesis by bile acids that return to the liver via the portal blood. The enterohepatic circulation of bile is an important physiological process that regulates bile acid synthesis and maintains cholesterol homeostasis. Disruption of the enterohepatic circulation of bile acids can lead to gallstone disease, chronic liver diseases such as cirrhosis and fatty liver; as well as contribute to cardiovascular diseases such as atherosclerosis, coronary artery disease and stroke.

The objective of my study was to understand the effect of cytokines on CYP8B1 gene transcription. Previously, cytokines have been shown to suppress CYP7A1 (neutral pathway) [4, 236, 237] and CYP27A1 (acidic pathway) [238]. It was important to study the effect of cytokines on CYP8B1, which is the rate-limiting enzyme for CA synthesis.

This study established primary human hepatocytes as an excellent model to study bile acid biosynthesis. Primary human hepatocytes have the closest known morphology to the human liver since they are derived from the hepatic parenchyma. IL-1β inhibited

CYP8B1 mRNA expression in both a dose and time dependent manner. IL-1β inhibited

CYP8B1 gene reporter activity in HepG2 cells, but not in the non-liver Hek293 cells, suggesting the involvement of liver specific transcription factors in mediating the effect.

187 188

Since CYP8B1 is exclusively expressed in the liver tissue, its regulation in non-liver tissue is not warranted.

The inverse regulation of SHP levels by cytokines and bile acids revealed an important mechanistic difference in the action of cytokines and bile acids in regulating

CYP8B1. CDCA induces SHP which can interact with a trans-activating nuclear receptor such as HNF4α and inhibit CYP8B1 gene transcription. IL-1β on the other hand inhibits

SHP and hence the inhibitory effect of IL-1β on the CYP8B1 gene can occur independent of SHP, since inhibition of a negative factor (SHP) would induce CYP8B1 gene transcription. Since bile acids are known to induce cytokine release in the liver [4] , the inhibition of Cyp8b1 by cytokines may explain why bile acid feeding to Shp knockout mice reduces Cyp8b1 mRNA levels similar to the wild-type mice [165, 166] .

Using the specific inhibitors of the MAPK pathway, I was able to delineate the

JNK pathway as the cytokine signaling pathway that inhibits CYP8B1 gene transcription.

The JNK pathway is known to be a stress activated pathway in mammalian cells. Studies have shown that both cytokines [218, 239] and bile acids [218, 240] can activate the JNK pathway. Bile acid induced FGF19/FGFR4 [234, 235] and PKC pathways [241, 242], can both stimulate the JNK pathway. Bile acid/cytokine activated JNK has been shown to suppress CYP7A1 gene transcription, while this suppression is blocked by the overexpression of a dominant negative form of JNK [218, 237].

In this study I demonstrated that JNK1 and c-Jun markedly decreased CYP8B1 gene transcription and that a dominant negative JNK1 and c-Jun blocked the inhibition.

An interesting observation of this experiment was that dnc-Jun overexpression stimulated

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the CYP8B1 gene transcription very strongly (>4 fold). This suggests that the c-Jun present within HepG2 cells may have an inhibitory effect on the basal promoter activity of CYP8B1 and that the dnc-Jun could counteract this effect causing a marked stimulation of CYP8B1 gene transcription.

Previous studies show that HNF4α is a strong activator of the human CYP8B1 gene and that the HNF4α binding site mediates bile acid inhibition [107]. My mutagenesis studies on the human CYP8B1 promoter indicated that among the nuclear receptors known to bind the promoter so far (i.e. HNF4α and FTF), only mutation of the

HNF4α binding site can completely abolish the IL-1β suppression of the CYP8B1 gene transcription. This proves that the bile acid response element is also a cytokine response element on the CYP8B1 gene promoter. The finding that dnHNF4α inhibits the CYP8B1 gene and abrogates the inhibitory effect of IL-1β on CYP8B1 gene transcription supports the theory that HNF4α is involved in IL-1β inhibition of the human CYP8B1 gene.

Studies in our lab have also shown that IL-1β suppressed the CYP7A1 gene transcription via a JNK mediated pathway, and that this effect was lost upon mutation of the HNF4α binding site on the human CYP7A1 promoter (Jahan A, Li T and Chiang J, unpublished data). This is consistent with several recent reports that have also emphasized on the importance of HNF4α as being the central regulator of human

CYP7A1 [120, 237]. The siRNA knockdown of HNF4α in HepG2 cells also showed the importance of HNF4α in transactivation of CYP7A1 gene.

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The results from the ChIP assays show that both IL-1β and CDCA decrease the recruitment of HNF4α to the CYP8B1 and CYP7A1 chromatin. The observation that IL-

1β inhibits the HNF4α reporter activity, mRNA and protein levels provides an explanation for the decreased HNF4α binding to the CYP8B1 and CYP7A1 chromatin.

The fact that a JNK-specific inhibitor blocked the IL-1β inhibition of HNF4α expression further supports the model that HNF4α is a downstream target of the cytokine activated

JNK pathway and that cytokines inhibit the HNF4α trans-activation of the CYP8B1 gene.

This experiment was crucial in linking two important aspects of my study, the signal transduction pathway and the downstream nuclear receptor that mediate the inhibiton of

CYP8B1 gene transcription by IL-1β. The exact mechanism by which the JNK pathway regulates the expression of HNF4α is yet to be determined.

Phosphorylation of nuclear receptors has been suggested as an additional mechanism by which cytokines can decrease gene transcription. My study proves that cytokine and bile acids can regulate HNF4α at the transcriptional, translational and posttranslational levels. It is known that HNF4α is phosphorylated by protein kinase A

[314], ERK [315] and AMP kinase [316, 317], and that phosphorylated HNF4α loses its

DNA binding activity [318]. The invitro kinase assay showed that constitutively active

JNK1 can directly phosphorylate HNF4α and the ChIP assays showed that overeexpression of JNK1 in HepG2 cells decreased the HNF4α binding to the CYP8B1 and CYP7A1 chromatin. It is more than likely that overexpressing JNK1 in cells would increase the phosphorylation of HNF4α, which could result in the decreased binding of

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HNF4α to the DNA. This would result in a reduction of the HNF4α transactivation of the

CYP8B1 and CYP7A1 gene transcription. Since phosphorylation is a quick physiological process, the HNF4α phosphorylation by JNK may account for the rapid downregulation of CYP8B1 and CYP7A1 by cytokines and bile acids. However, the more long term effects may be due to the FXR/SHP pathway that requires changes in protein synthesis, which is a much slower process. Thus, under normal physiological conditions, the

FXR/SHP pathway may be the major mechanism to control bile acid synthesis, while during conditions of cellular stress associated with acute inflammation, the phosphorylation of HNF4α may play a more significant role.

Another interesting finding of this study was the strong induction of c-Jun on IL-

1β treatment. c-Jun has previously been shown to be activated by bile acids [218]. In the same study it was shown that c-Jun activates the SHP gene transcription, which could inhibit the CYP7A1 gene [218]. Studies have suggested that c-Jun can interact with other transcription factors and alter the transcription of a gene without directly binding the

DNA [156]. A study showed earlier that the α-fetoprotein (AFP) gene, which is a negative acute phase reactant [321], is suppressed by c-Jun via the FTF-binding site on its promoter [321-323]and that c-Jun interacts strongly with FTF in GST pull down assays

[156]. Based on this knowledge it was suggested that c-Jun forms a repressive complex with FTF to inhibit the CYP7A1 gene [156].

I showed that c-Jun coexpression with HNF4α, abolished the capacity of the latter to activate both the CYP8B1 and the CYP7A1 promoter. A novel finding of this study is that c-Jun interacts strongly with HNF4α as demonstrated by in vivo (CO-IP assay) and

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in vitro (GST pull down assay). This interaction existed independent of the phosphorylation state of c-Jun. However, IL-1β enhanced the HNF4α interaction with both c-Jun and phosphorylated c-Jun despite the decrease in the HNF4α protein levels.

Since HepG2 cells expresses high basal levels of HNF4α, the c-Jun interaction with

HNF4α may also account for the repression of the basal reporter activities of the CYP8B1 and CYP7A1 genes (Fig. 18B, 27, 32) upon overexpression of c-Jun alone in HepG2 cells.

My study also revealed that the c-Jun interaction with HNF4α may provide a possible mechanism for suppression of CYP7A1 and CYP8B1 by bile acids and cytokines.

Since the HNF4α interaction with PGC-1α is critical for the induction of the human

CYP8B1 and CYP7A1 gene, c-Jun may compete with PGC-1α for binding to HNF4α and result in inhibition of the CYP8B1/CYP7A1. Thus, the lack of coactivator recruitment onto the chromatin provides an additional mechanism via which cytokines and bile acids can suppress bile acid biosynthetic gene transcription.

In this study I found that IL-1β markedly reduced SHP mRNA levels in primary human hepatocytes. This data is consistent with a previous study which showed that LPS treatment suppresses the expression of Shp mRNA in mouse livers and that TNF and IL-1 treated Hep3B cells (human hepatoma cells) expressed lower levels of SHP mRNA

[312]. The same study also showed that cytokine treatment lead to the suppression of

FXR and hence its target genes such as SHP, Apo CII etc were also suppressed [312].

SHP gene transcription is under a complex regulation by many transcription factors, including FXR, AP-1, LXRα, ERα, SREBP-1c and FTF [3, 305, 324]. Species

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differences in LXRα and SREBP-1c regulation of CYP7A1, CYP8B1 and SHP genes are well documented [3, 305, 324]. My data shows that HNF4α is yet another regulator of

SHP gene transcription and it is likely that IL-1β inhibition of the HNF4α gene accounts

for the inhibition of the SHP gene by IL-1β in the human liver.

SHP mutations have been identified in obese Japanese patients with early onset

diabetes [164]. Mutations of HNF4α gene have been linked to MODY 1 [138, 139]. It is

likely that since both HNF4α and SHP are expressed in pancreatic tissue, the regulation of SHP by HNF4α may play an important role in the development of the pathology of

MODY. It is possible that IL-1β inhibition of the SHP gene may also be an adaptive

response to inflammation and cholestasis. Inhibition of this negative nuclear receptor may

promote nuclear receptor activation of gene transcription during hepatocyte regeneration

and differentiation.

It was previously reported in our lab that SHP could interact with HNF4α and

inhibit human CYP8B1 gene transcription [107]. However, the SHP interaction with

HNF4α is much weaker than with FTF. Furthermore, bile duct ligation alone reduced

Cyp8b1, induced Cyp7a1, but did not have any effect on Shp mRNA expression levels.

In α-naphthylisothiocyanate induced intrahepatic cholestatic rats, Cyp8b1 mRNA

expression is reduced by 80%, but Cyp7a1 and Shp mRNA expression is not altered

[325]. Also Cyp8b1 mRNA expression is not altered in a recent study of bile acid metabolism in Fxr null mice [326]. These results support the conclusion that the

FXR/SHP pathway may not play a major role in bile acid inhibition of CYP8B1 gene

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transcription. It is clear that redundant mechanisms are involved in bile acid inhibition of both Cyp7a1 and Cyp8b1 and explain the finding that Cyp7a1 and Cyp8b1 mRNA expression remain inhibited by bile acid feeding in Shp null mice [165, 166].

It is intriguing that Cyp8b1, but not Cyp7a1, is re-expressed in Shp null mice fed with a diet containing CA or CA plus cholesterol for 12 weeks [255]. It was seen that the long term feeding on these diets was associated with decreased hepatic accumulation of cholesterol and triglycerides in the Shp null mice, making these mice relatively resistant to the hepatotoxicity associated with short term feeding on the same diets [165]. It was suggested that the re-expression of Cyp8b1 could contribute to the decreased toxicity of the chronic bile acid treatment by increasing the hydrophilicity of the bile acid pool. All these results suggest that Cyp7a1 and Cyp8b1 genes are regulated by somewhat different mechanisms by bile acids.

Obesity and Type II Diabetes are two clinical diseases associated with chronic inflammation. These conditions are characterized by abnormally high production of

TNFα which mediates insulin resistance in them [327]. TNFα is produced by adipocytes and its expression is increased in adipocytes isolated from several genetic models of rodent obesity and from obese humans [328]. TNFα can inhibit the insulin-stimulated tyrosine phosphorylation of both the insulin receptor (IR) and insulin receptor substrate

(IRS)-1 and can downregulate the insulin-sensitive glucose transporter, GLUT4, in adipocytes [327-329]. TNFα has also been shown to activate the JNK pathway in the β- cells of the pancreatic islets causing apoptosis leading to a deficiency in insulin levels

[330]. This effect has been shown to be inhibited significantly by the JNK specific

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inhibitor, SP600125 [330]. As mentioned earlier HNF4α can induce the transcription of

HNF1α which in turn can activate the insulin gene [331, 332]. It is likely that the cytokine suppression of HNF4α, as seen in my study could in the pancreas lead to suppression of the insulin gene transcription leading to impaired glucose and fatty acid metabolism. This could be an additional mechanism by which cytokines could mediate the metabolic changes in diabetes. Also, an inhibitor of the JNK pathway could play a beneficial role by counteracting the actions of the cytokines and ameliorating the homeostatic imbalance of the cells during diabetes.

Studies have shown that insulin can suppress the rodent CYP8B1 gene [246, 253] however the effect of insulin on the human CYP8B1 gene is unknown. According to our data, cytokines released in an inflammatory condition could suppress the CYP8B1 levels and possibly also the insulin levels (indirectly via HNF4α), implying that the inverse relationship between the expression of CYP8B1 and insulin that is present in rodents may not exist in humans. This discrepancy could be attributed to a species difference in gene regulation.

It is known that changes in lipid and lipoprotein metabolism, such as increase serum triglygerides and cholesterol, increase in hepatic lipogenesis and enhanced VLDL production [272], are seen during infection and inflammation. The hypercholesterolemia during inflammation is known to be caused by increase in the mRNA levels, protein mass and activity of HMG-CoA reductase, the rate-limiting enzyme in cholesterol synthesis in the liver [333]. The decrease in CYP7A1 gene transcription in response to cytokines, would lead to the reduction in bile acid synthesis, but may contribute to the

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hypercholesterolemia observed in cholestatic liver disease patients. On the contrary, the inhibition of CYP8B1 gene transcription, during inflammation, would decrease CA levels, which in turn would lead to decrease in the intestinal absorption of cholesterol and help maintain cholesterol homeostasis. It has been shown that bile acid activated FXR can inhibit SREBP-1c gene transcription, leading to decrease in triglyceride synthesis and secretion, and activate Apo CII and PLTP gene transcription leading to increased triglyceride clearance [112, 334]. Thus, bile acids can lower the plasma triglyceride levels explaining why gallstone patients treated with cholestyramine show increased levels of triglycerides in their serum. Under inflammatory conditions, the suppression of

CYP7A1, would decrease the bile acid pool size and could contribute to the increase in serum triglycerides seen during an infection.

During cholestasis, the excess bile acids can suppress CYP7A1 and decrease the bile acid pool size which can help ameliorate the condition. However, the suppression of

CYP8B1 by bile acids would lead to the accumulation of the more hydrophobic and toxic bile acids such as CDCA and LCA, which could further damage the hepatocytes. The suppression of bile acid synthesis during cholestasis, leads to accumulation of excess cholesterol and predisposes to cholesterol gallstone formation. Under such conditions the inhibition of CYP8B1 gene transcription would again lead to decrease in the intestinal absorption of cholesterol and reduce the possibility of gallstone formation.

A mutation in the CYP8B1 gene of human patients has not yet been identified.

Due to the differences in bile acid and cholesterol synthesis and regulation seen in mice and humans, one could predict that a CYP8B1 mutation in humans may produce a

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different phenotype compared to the Cyp8b1 null mice. It is likely that a CYP8B1 mutation in a human patient could lead to decreased intestinal cholesterol absorption due to lack of CA, which as mentioned before is a more efficient bile acid for cholesterol absorption. Also since humans do not produce any muricholic acid, lack of CYP8B1 activity will lead to overproduction of hydrophobic bile acids such as CDCA, LCA etc. contributing to hepatotoxicity and liver damage. It cannot be determined if CYP8B1 can be used as a target for drug/genetic therapy in any disease conditions, unless the phenotype for a human CYP8B1 gene mutation is known.

Based on my data in this study, future studies can be focussed in several different directions. Since the JNK pathway has been shown to be a major regulator of bile acid biosynthesis, other activators of the pathway such as UV radiation, translation inhibitors, tumor promoters, heat shock and hyperosmotic stress can be tested for their possible regulatory effects on bile acid homeostasis. HNF4α is a key regulator of various genes involved in processes such as gluconeogenesis, lipoprotein metabolism, embryonic development etc. The cytokine inhibition of HNF4α by multiple mechanisms could possibly lead to the regulation of other HNF4α regulated genes by cytokines and play a role in disease pathogenesis. The interaction of c-Jun with HNF4α and its inhibition of coactivator recruitment for gene transcription, unfolds a new mechanism by which this transcription factor can regulate genes. It is likely that c-Jun can interact with several other nuclear receptors and affect gene transcription in a similar manner.

In summary, my study suggests that cytokines inhibit CYP8B1 gene transcription via activation of the JNK pathway, which reduces HNF4α gene

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transcription and phosphorylates HNF4α, resulting in the inhibition of the human

CYP8B1 gene (Fig. 35). The mechanism of this inhibition is SHP independent. I provide the first evidence that c-Jun expression is induced by cytokines and that c-Jun can interact with HNF4α, and decrease the recruitment of PGC-1α by the latter, thereby suppressing

CYP8B1 and CYP7A1 gene transcription. This is a novel mechanism in studying the suppression of bile acid biosynthetic genes by both cytokines and bile acids. Inhibition of bile acid synthesis is a crucial adaptive response to inflammation and a protection of the liver from accumulating toxic bile acids and developing intrahepatic cholestasis.

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Fig.35.

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Figure 35. Summary of data.

Bile acids induce cytokine (IL-1β, TNFα) release from the Kupffer cells in the hepatic

sinusoids. These cytokines act on the surrounding hepatocytes. My data suggests that the

bile acid suppression of CYP8B1 is mediated by the c-Jun NH2 terminal kinase (JNK)

and not by the extracellular signal-regulated kinase (ERK) or the p38 mitogen activated

protein kinase (p38MAPK). The JNK pathway can decrease the HNF4α protein levels

and phosphorylate HNF4α; both of these events cause the downregulation of the HNF4α

trans-activation of CYP8B1/7A1. Also both cytokines and bile acids can induce c-Jun.

Activated JNK pathway can also phosphorylate the c-Jun protein. c-Jun, in its

phosphorylated or unphosphorylated form interacts with HNF4α, and interferes with the

recruitment of the coactivator PGC-1α to the chromatin. Cytokines also inhibit the

expression of SHP.

APPENDIX A

ABBREVIATIONS

3β-HSD: 3β-hydroxy-C27-steroid dehydrogenase/isomerase

AAs: Amino acids

ABC (A1/G5/G8): ATP-binding cassette protein (A1/G5/G8)

ACAT: acyl-CoA-cholesterol acyltransferase

Acetyl-CoA: acetyl-coenzyme A

ADD-1: Adipocyte determination and differentiation factor-1

AE2: Anionic exchanger 2

AF (1/2): Activation function domain (1/2)

AFP: α-fetoprotein

AMPK: Adenosine monophosphate activated protein kinase

AP-1: Activation protein-1

Apo (A1, B, B100, CII, CIII): Apolipoprotein (A1, B, B100, CII, CIII)

AR: Androgen receptor

ASBT: Apical sodium-dependent bile salt transporter

ATP: Adenosine triphosphate

ATF2: Activating transcription factor 2

BARE (I/II): Bile acid response element (I/II)

201 202

BRIC: Benign recurrent intrahepatic cholestasis

BSEP: Bile salt export pump

CA: Cholic acid

CAR: Constitutive androgen receptor

Caco-2: Colonic carcinoma cell line

CahR4: Constitutively active human FGFR4

CDCA: Chenodeoxycholic acid

CEH: Cholesterol ester hydroxylase

CETP: Cholesterol ester transfer protein

CFTR: Cystic fibrosis transmembrane conductance regulator

ChIP: Chromatin immunoprecipitation assay

CM: Chylomicron

Co-IP: Coimmunoprecipitation

COUP- TF (I/II): Chicken ovalbumin upstream promoter transcription factors (I/II)

CPF: CYP7A1 promoter factor

CREB: cAMP response element binding protein

CBP: CREB binding protein

CTX: Cerebrotendinous xanthomatosis

CYP3A (4): Cytochrome P450 3A family (4)

CYP7A1: Chosterol 7α-hydroxylase

CYP7B1: Oxysterol 7α-hydroxylase

CYP8B1: Sterol 12α-hydroxylase

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CYP27A1: Sterol 27-hydroxylase

DBD: DNA binding domain

DBP: Albumin D-site binding protein dnc-Jun: Dominant negative c-Jun dnHNF4α: Dominant negative HNF4α dnJNK1: Dominant negative JNK1

DR: Direct repeat

EMSA: Electromobility shift assay

ER: Everted repeat

ER: Estrogen receptor

ERK: Extracellular signal-regulated kinase

FAS: Fatty acid synthase

FFA: Free fatty acids

FGF19: Fibroblast growth factor 19

FGFR4: Fibroblast growth factor receptor 4

FTF: α-fetoprotein transcription factor

FXR: Farnesoid X receptor

GLUT-4: Glucose transporter-4

GPCR: G-protein coupled receptor

GR: Glucocorticoid receptor

GST: Glutahione S-transferase

HAT: Histone acetyl transferase

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HDCA: Hyodeoxycholic acid

HDL: High-density lipoprotein

Hek293: Human embryonic kidney cell line

Hep3B: Human hepatoma cell line

HepG2: Human hepatoma cell line

HMG-CoA reductase: Hydroxy-3-methyl glutyryl-CoA reductase

HNF (1/3β/4α): Hepatocyte nuclear factor-(1/3β/4α)

HPLC: High-performance liquid chromatography

HREs: Hormone response elements

IBABP: Ileal bile acid binding protein

IBAT: Ileal bile acid transporter

IDL: Intermediary density lipoprotein

IL (1β/6/10): Interleukin-(1β/6/10)

Insig (1, 2a, 2b): Insulin-induced gene (1, 2a, 2b)

IR: Inverted repeat

IR: Insulin receptor

IRS-1: Insulin receptor substrate-1

JNK: c-Jun N-terminal kinase

LBD: Ligand binding domain

LCA: Lithocholic acid

LDL: Low-density lipoprotein

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LPS: Lipopolysaccharide

LRH: Mouse liver-related homolog

LRP: LDL receptor related protein

LXR (α/β): Liver X receptor (α/β)

MAPK: Mitogen activated protein kinase

MAPKK: MAP kinase kinase

MAPKKK: MAP kinase kinase kinase

M-BAR: Membrane-bile acid receptor

MDR (1/3): Multidrug-resistance protein (1/3)

MEKK1: MAP kinase kinase 1

MKK4: MAP kinase kinase 4

MODY (1/3): Maturity onset diabetes of the young (1/3)

MR: Mineralocorticoid receptor

MRP (2/3): MDR related protein (2/3)

MTP: Microsomal triglyceride transport protein

NO: Nitric oxide

NRs: Nuclear receptors

NTCP: Sodium taurocholate cotransporting polypeptide

OATP (1A2/1B1/1B3/2B1/3): Organic anion transport peptide (1A2/1B1/1B3/2B1/3)

Ost (α/β): Organic solute transporter (α/β) p38MAPK: p38 mitogen activated protein kinase

PCN: Pregnenolone 16α-carbonitrile

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PCR: Polymerase chain reaction

PFIC (type1/ 2): Progressive familial intrahepatic cholestasis (type1/ 2)

PKA: Protein kinase A

PLTP: Phospholipid transfer protein

PPAR (α, γ): Peroxisome proliferator activated receptor (α, γ)

PPRE: Peroxisome proliferator response element

PR: Progesterone receptor

PXR: Pregnane X receptor

Q-RT-PCR: Quantitative reverse transcriptase PCR

RAR: Retinoic acid receptor

ROS: Reactive oxygen species

RT-PCR: Reverse transcriptase PCR

RXR: Retinoid X receptor

SAPK: Stress-activated protein kinase

SCAP: SREBP cleavage activation protein

SCD-1: Stearoyl-CoA desaturase-1

SF-1: Steroidogenic factor 1

SHP: Small heterodimer partner siRNA: Small interference RNA

SR (B1/A1): scavenger receptor (subtype B1/A1)

SRC-1: Steroid receptor coactivator-1

SREs: Sterol response elements

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SREBP (1a/1c/2): Sterol response element binding protein (1a/1c/2)

SXR: Steroid X receptor

TG: Triglycerides

TNFα: Tumor necrosis factor α

TR: Thyroid hormone receptor

UDCA: Ursodeoxycholic acid

VDR: Vitamin D receptor

VLDL: Very low-density lipoprotein

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