Retinoic Acid Related Orphan Nuclear Receptor Α (Rorα) Regulates Diurnal Rhythm and Fasting Induction of Sterol 12Α-Hydroxylase (CYP8B1) in Bile Acid Synthesis

Home , CYP8B1, Farnesoid X receptor, Retinoid X receptor

Retinoic acid related orphan nuclear receptor α (RORα) regulates diurnal rhythm and fasting induction of sterol 12α-hydroxylase (CYP8B1) in bile acid synthesis

A dissertation submitted to the Kent State University towards partial fulfillment of the requirements for the degree of Doctor of Philosophy

By Preeti Pathak

Dissertation written by Preeti Pathak Msc. Pune University, India. 2005 Ph.D. Kent State University 2013

Approved by

_ _Dr. Diane Stroup______, Chair, Doctoral Dissertation Committee

_ _Dr. John YL. Chiang______, Co-Chair, Doctoral Dissertation Committee

_ Dr. Soumitra Basu______, Members, Doctoral Dissertation Committee

_ Dr. Hanbin Mao ______,

_Dr. Yoon Kwang Lee ______,

_ Dr. David Glass______, Graduate Faculty Representative

Accepted by

_Dr. Michael Tubergen______, Chair, Department of Chemistry

_Dr._ James L. Blank ______, Dean, College of Arts and Sciences

TABLE OF CONTENTS

LIST OF FIGURES…………………………………………………………………...vi LIST OF TABLES……………………………………………………………………vii AKNOLEDGEMENTS……………………………………………………………... viii ABBREVIATIONS…………………………………………………………………....ix CHAPTER I: INTRODUCTION 1.1. Bile Acids 1.1.1. Structure and function…………………………………………………..…1 1.1.2. Synthesis …………… …………………………………………………...4 1.1.3. Enterohepatic circulation of bile acids…………………………………… 7 1.2. Bile acid signaling network 1.2.1. FXR/SHP/FGF19………………………………………………………….12 1.2.2. G protein-coupled bile acid receptor 1 (GPBAR1)/ TGR5……………….…12 1.2.3. MAPK/ERK………………...... ……………………………………. .13 1.3. Nuclear Hormone Receptors 1.3.1. Structure and response element……………………….…………………..16 1.3.2. Farnesoid X Receptor (FXR, NR1H4)……………………………………19 1.3.3. Pregnane X Receptor (PXR, NR1I2) …...…………………...…………. 20 1.3.4. Vitamin D3 Receptor (VDR, NR1I1)……..………………………………21 1.3.5. Liver X Receptor (LXR, NR1H3)………………… …...……………….21 1.3.6. Hepatocyte nuclear factor 4α (HNF4α, NR2Α1)…….……………………22 1.3.7. Liver Receptor Homologue 1 (LRH1 NR5A2) …….…………………….22 1.3.8. Small Heterodimer Partner (SHP, NR0B2)………… ………………..….23 1.3.9. Peroxisome Proliferator Activated Receptor α (PPARα, NR1C1)………..24 1.3.10. Retinoic acid related orphan nuclear receptor α (RORα NR1F1)………. 26 1.3.11. Rev-erbα (NR1D1)…… …………………………………….. ……… 27

1.4 Transcription factors and Co-activators 1.4.1. Sterol Regulatory Element Binding Proteins (SREBPs)………………….28 1.4.2. Peroxisome Proliferator-Activated Receptor-γ Co-activator 1α (PGC-1α).29 1.4.3. Cyclic AMP response element binding-binding protein (CBP)…… ….....30

1.5. Transcriptional regulation of CYP8B1 gene expression………………………....30 1.6. Circadian regulation of bile acid synthesis……………………………………….32 1.7. Nutrient regulation………………………………………………………………...33 1.8. Hypothesis and specific aims…………………………………………………….. 38 Chapter II: MATERIALS AND METHODS 2.1. Animals 2.1.1. Circadian rhythm ………………………… …………………………….43 2.1.2. Adenovirus injection……………………… …………………………….43 2.2. Liver mRNA isolation………………………………..…………………………….44

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2.2.1. Reverse transcription………………………….. …………………………44

2.3. Quantitative real time PCR……………………………………………………...45 2.3.1. SYBR Green method………………………………………………….…46 2.3.2. Taq-man Method…………………………………………………….….. 48 2.3.3. Data analysis………………………………………………………….… 48

2.4. Cell Culture………………………………………………………………….……49 2.4.1. Cell storage and freezing ……………………………………….………..50 2.4.2. Primary cell culture……………………………………………….…….. 50

2.5. Plasmids and cloning ……………………………………………..………….……50 2.5.1. Reporters……………………………………………………………… ..50 2.5.2. Site directed mutagenesis…………………………………………….….. 50 2.5.3. Expression Vectors………………………………………………..………51 2.5.4. Preparation of plasmid DNA…………………………………………..….51 2.5.5. Small Scale DNA preparation ……………………………………..….…..51 2.5.6. Ligation and restriction enzyme digestion…………………...... 52 2.5.7. Preparation of Competent Cells…………………………………………...52 2.5.8. Bacterial Cell Transformation……………………………………………..53

2.6. Cloning adenovirus 2.6.1. Principle…………………………………………………………………...54 2.6.2. Cloning pShuttle-CMV vector ……………………………………………55 2.6.3. Electroporation…………………………………………………………….56 2.6.4. Screening positive recombination…………………………………………57 2.6.5. Preparation of primary adenoviral stock………………………………...... 60 2.6.6. Amplification and purification of virus…………………………………...60 2.6.7. Adenovirus titer measurement………………………………………….....61

2.7. Chromatin immuno precipitation assay (ChIP assay)…………………………..63 2.7.1. Isolation of nuclei and crosslinking……………………………………….63 2.7.2. Sonication………………………………………………………………....63 2.7.3. Immunoprecipitation and washing………………………………………..64 2.7.4. ChIP assay using HepG2 cells…………………………………………….64 2.8. Western blot analysis ……………………………………………………………...64 2.9. Measurement of bile acid pool size …………………………………………….....65 2.9.1. Principle………………………………………………………………..….66 2.9.2. Isolation of bile acids from liver, gallbladder and intestine………….…. ..66 2.9.3. Measurement of bile acid concentration…………………………….…….66

2.10. Isolation and measurement of liver triglycerides and cholesterol….………….67

2.11. Electrophoretic Mobility Shift Assay (EMSA)…… …………………………. 68

2.11.1. Annealing the probes……… …………………………………………...68 2.11.2. Radiolabelling probe………………………………….………………… 68 2.11.3. Binding reaction………………………………………………………… 68 2.11.4. Running EMSA gel………………………………………………………69

2.12. Transient Transfection Assay ………………………………………..………….69

2.13. Small Interference RNA transfection…………………………………..………..71

2.14. Immunoprecipitation Assay (IP)………………………………………………...71

2.15. Microsome isolation ……………………………………………….…………….71

2.16. Mammalian two hybrid assay… ………………………………………………..72

2.17. Statistical analysis ………………………………………………………………..73

Chapter III: RESULTS……………………………………………………..…………74

Chapter IV: DISCUSSION………………………………………..…………………122

REFERENCES……………………………………………………………….137

LIST OF FIGURES

Figure 1……………………………………………………………………….……3 Figure 2. …………………………………………………………………….…….5 Figure 3. …………………………………………….……………………….…...10 Figure 4. . ……………………………….…………………………………….…..14 Figure 5. …………………………………………………………………….……17 Figure 6. ……………………………………………………………………..…...36 Figure 7…………………………………………………………………….……. 59 Figure 8. ……………………………………………………………………..……75 Figure 9. ………………………………………………...... 76 Figure 10. ………………………………………………………..……….…..…...79 Figure 11. ……………………………………………..…………………….….…81 Figure 12. ………………………………………………………………..….….…84 Figure 13. ……………………………………………………..……………….….85 Figure 14…………………………………………………………………….…….89 Figure 15. …………………………………………………….…………….….….94 Figure 16. …………… ………………………………………………………..….97 Figure 17. ………………………………………………………………….…..….99 Figure 18. ………………………………………………………………..…….…101 Figure 19……………………………………………………………………….....103 Figure 20. …………………………………………………………………….…..106 Figure 21. ………………………………………………………………………...109 Figure 22. …………………………………………………………..……….……111 Figure 23. ……………………………………………………………………..….114 Figure 24. …… ………………………………………... .……………………….116 Figure 25. ….………………………………………………………………….….120

LIST OF TABLES

Table 1. ………………………………………………………………………………..61 Table 2. …………………………………………………………………………….…104

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ACKNOWLEDGEMENTS

I would like to express my sincere gratitude to 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. All his guidance, scientific suggestions and careful review of my work helped improve this dissertation significantly. I also deeply appreciate the valuable time, comments and assistance provided by Dr. Diane Stroup and all my committee members. I also want to thank my lab mates Dr. Tiangang Li, Dr. Jessica Francl and Shannon Boehme for valuable discussions and suggestions that helped improve my research techniques.

My parents, Satish and Alka Awati receive my deepest gratitude and love for always believing in me, standing by me and showing me the importance of a good education.

Last, but not least, I would like to thank my husband, Chandan, for his understanding and love and all the sacrifices he made, through all these years. His encouragement and support have been the pillars of strength during my graduate work.

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ABBREVIATIONS

3β-HSD: 3β-hydroxy-C27-steroid dehydrogenase/isomerase 7αOHC: 7α-hydroxycholesterol 24OHC: 24-hydroxycholesterol 25OHC: 25-hydroxycholesterol

ABC A1 (G5/G8): ATP-binding cassette protein A1 (G5/G8) AF (1/2): Activation function domain (1/2) AMPK: Adenosine monophosphate activated protein kinase ATP: Adenosine triphosphate cAMP: cyclic Adenosine monophosphate ASBT: Apical sodium-dependent bile salt transporter ATF2: Activating transcription factor 2

BARE (I/II): Bile acid response element (I/II) BSEP: Bile salt export pump

CA: Cholic acid CAR: Constitutive androgen receptor CDCA: Chenodeoxycholic acid ChIP: Chromatin immunoprecipitation assay Co-IP: Co-immunoprecipitation CREB: cAMP response element binding protein CBP: CREB binding protein CYP7A1: Chosterol 7α-hydroxylase CYP7B1: Oxysterol 7α-hydroxylase CYP8B1: Sterol 12α-hydroxylase CYP27A1: Sterol 27-hydroxylase

DBD: DNA binding domain DBP: Albumin D-site binding protein DR: Direct repeat

EMSA: Electromobility shift assay ER: Everted repeat ER: Estrogen receptor ERK: Extracellular signal-regulated kinase

FFA: Free fatty acids

FGF19: Fibroblast growth factor 19 FGFR4: Fibroblast growth factor receptor 4 FTF: α-fetoprotein transcription factor FXR: Farnesoid X receptor

GPCR: G-protein coupled receptor GR: Glucocorticoid receptor

HAT: Histone acetyl transferase HCA: Hyocholic acid HepG2: Human hepatoma cell line HPLC: High-performance liquid chromatography

IRS-1: Insulin receptor substrate-1 JNK: c-Jun N-terminal kinase

LBD: Ligand binding domain LCA: Lithocholic acid LDL: Low-density lipoprotein

LPS: Lipopolysaccharide LRH1: Mouse liver-related homolog1 LXR (α/β): Liver X receptor (α/β)

MAPK: Mitogen activated protein kinase MAPKK: MAP kinase kinase MAPKKK: MAP kinase kinase kinase MDR 1/3: Multidrug-resistance protein 1/3

NRs: Nuclear receptors NTCP: Sodium taurocholate cotransporting polypeptide

OATP: Organic anion transport peptide Ost (α/β): Organic solute transporter (α/β) p38MAPK: p38 mitogen activated protein kinase PCR: Polymerase chain reaction PKA: Protein kinase A PPAR (α, γ): Peroxisome proliferator activated receptor (α, γ) PPRE: Peroxisome proliferator response element 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 SCAP: SREBP cleavage activation protein SHP: Small heterodimer partner siRNA: Small interference RNA SREBP 1a (1c/2): Sterol response element binding protein 1a (1c/2) SRC-1: Steroid receptor coactivator-1 SREs: Sterol response elements

TG: Triglycerides TNFα: Tumor necrosis factor α TR: Thyroid hormone receptor

UDCA: Ursodeoxycholic acid

VDR: Vitamin D receptor

CHAPTER I

INTRODUCTION

1.1. Bile acids

Bile acids are physiological agents that facilitate intestinal absorption of nutrients and fat soluble vitamins. Bile acids also play an important role in the regulation of drug metabolism and help to maintain glucose and lipid homeostasis. Bile acids are synthesized in the liver and stored in the gallbladder. After each meal, gallbladder contraction empties bile acids into the intestinal tract. When passing through the intestinal tract, most bile acids (95%) are reabsorbed in the ileum [1-3]. Excreted bile acids are replenished by de novo synthesis in the liver to maintain a constant bile acid pool.

1.1.1. Structure and function

Bile acids are classified as either primary or secondary bile acids (Fig. 1).

Chenodeoxycholic acid (CDCA) and cholic acid (CA) are two primary bile acids in human bile [4-5]. In humans, the bile acid composition is significantly different from that in the rodents. In rodents, CDCA is further converted into α and β muricholic acids, which are more soluble than CDCA (Fig. 1). Primary bile acids are conjugated to taurine or glycine before secretion [6-7]. In the intestinal lumen, CA and CDCA are converted

into secondary bile acids, deoxycholic acid (DCA) and lithocholic acid (LCA), respectively, by 7α-dehydroxylase activity in the intestinal bacterial flora [8-9].

Amphipathic structure of bile acids facilitates micelle formation with phospholipid and cholesterol. Bile acids act as lipid carriers as they are able to solubilize many lipids by forming mixed micelles containing fatty acids, cholesterol esters and monoglycerides.

Recent study shows that bile acids are endogenous ligands that activate nuclear receptors,

FXR, PXR and VDR, which regulate glucose and lipid homeostasis [10-11].

Interestingly, bile acids activate a G protein coupled receptor, TGR5, to regulate energy homeostasis. Bile acids also play crucial role in the regulation of xenobiotic mechanism and inflammatory signaling pathway [12].

Figure 1. Structure of bile acids

Bile acids are derivatives of cholesterol (upper panel). The cholesterol backbone undergoes hydroxylation and isomerization of steroid ring followed by oxidative cleavage of the steroid side chains and is further conjugated with glycine or taurine.

1.1.2. Bile acid synthesis

In the liver, cholesterol is converted to bile acids by desaturation, hydroxylation and isomerization of steroid ring followed by oxidative cleavage of the steroid side chains. There are two major bile acid synthesis pathways 1) Neutral pathway and 2)

Acidic pathway [1, 6, 13-15] (Fig. 2).

1) Neutral pathway: This pathway is initiated by the rate-limiting enzyme cholesterol 7α-hydroxylase (CYP7A1), which specifically hydroxylates cholesterol at 7α position to form 7α-hydroxycholesterol. For synthesis of CA, 7α-hydroxy-4-cholesten-3- one is hydroxylated at the C-12 position by microsomal steroid 12α-hydroxylase

(CYP8B1) to form 7α, 12α-dihydroxy-4-cholestan-3-one. Oxidation of steroid side chain is catalyzed by a mitochondrial sterol 27-hydroxylase (CYP27A1), leading to the cleavage of three carbon side chain and synthesis of CA and CDCA [16-20].

2) Alternative or acidic pathway: This pathway involves a number of acidic intermediates, which could not be formed by the neutral pathway. This pathway contributes to less than 18% of the total bile acid synthesis in humans [5, 21]. In the acidic pathway, side chain oxidation proceeds steroid nucleus modification. The initial reaction is catalyzed by CYP27A1 to hydroxylate cholesterol to 27-hydroxycholesterol, which is further oxidized to 3β-hydroxy-5-cholenoic acid. The next step is the addition of a hydroxyl group at the C-7 position of the steroid nucleus by oxysterol 7α-hydroxylase

(CYP7B1). CDCA is the major product of the acidic pathway. CYP7B1 and CYP27A1 are expressed in all other tissues such as brain and macrophages to play varied roles [21-

23].

Figure 2. bile acid synthesis pathways

The first step in the neutral bile acid synthesis pathway is conversion of cholesterol into

7α-hydroxycholesterol by the rate limiting enzyme, cholesterol 7α-hydroxylase

(CYP7A1) (left side) followed by various enzymatic steps that lead to synthesis of

CDCA (chenodeoxycholic acid). For synthesis of cholic acids, sterol 12α hydroxylase

(CYP8B1) catalyzes hydroxylation at 12α position and produces cholic acid (CA). In the alternate pathway, initial hydroxylation reaction is catalyzed by CYP27A1 and followed by oxidation to form 3β-hydroxy-5-cholenoic acid. CYP7B1 catalyzes hydroxylation of

3β-hydroxy-5-cholenoic acid to form chenodeoxycholic acid (CDCA).

1.1.3. Enterohepatic circulation of bile acids

Bile acids are synthesized in the liver and immediately conjugated to taurine or glycine to increase their solubility and secretion into bile. After each meal, the gallbladder contracts to empty bile acids into the duodenum. About 95% of the bile acids are reabsorbed in the terminal ileum and returned back to the liver via the portal blood, while the remaining 5% is excreted into the feces. This process is called as “enterohepatic circulation” of bile [1].

The transporters, which mediate the enterohepatic circulation of bile acids, are described below according to their location and function (Fig. 3). Disruption of enterohepatic circulation of bile acids causes cholestatic liver disease and intestinal nutrient malabsorption [24-27].

Taurine and glycine conjugated bile acids are excreted into bile canaliculi from the hepatocytes via the bile salt export pump (BSEP, ABCB11). BSEP is an adenosine tri-phosphate (ATP) binding cassette (ABC) transporter localized in the apical membrane of the hepatocytes [28-32]. A canalicular transporter, multidrug resistant protein 3

(MDR3, ABCC3) effluxes phosphatidylcholine into bile. Multidrug resistant related protein 2 (MRP2) effluxes glycosylated and sulfated bile salts. MRP2 is responsible for the transport of conjugated bilirubin, some drugs toxins and heavy metals [33-34]. These transporters do efflux bile acids, cholesterol, phospholipids and phosphotidylcholine to form mixed micelle for storage in the gallbladder.

Cholangiocytes are epithelial cells in the bile duct. They can uptake bile acids via apical sodium dependent bile acid transporter (ASBT, SLC10A2) [35-36]. ASBT is expressed on the luminal membrane of cholangiocytes and enterocytes [37-38] . The

physiological role of bile salts uptake into the cholangiocytes pertains to the regulatory effect of the bile salts on mucin and bicarbonate secretion [39].

In the intestine, bile acids are reabsorbed by ASBT, also known as ileal bile acid transporter (IBAT). Human ASBT can uptake conjugated and unconjugated bile acids into enterocytes. It has higher affinity for CDCA and CA (unconjugated) than for taurocholate conjugated bile acids. Mutation in human ASBT gene can cause primary bile acid mal-absorption, an intestinal disorder characterized by congenital diarrhea, and reduced plasma cholesterol levels [37, 40]. Another important transporter, which excretes sterol from enterocytes, is ABCG5/ABCG8. Polymorphism of these transporters can lead to the sitosterolemia and gallstone disease [41]. MRP3 is located in the proximal small intestine on the sinusoidal membrane and efflux reabsorbed bile acids into portal blood

[42]. The OSTα and OSTβ dimer is a major bile acid efflux transporter, which transports bile acids from enterocytes to portal blood [43-44].

From portal blood, bile acids are reabsorbed by sodium-bile acid transporter,

NTCP (Na+-taurocholate co-transporting polypeptide) in the liver. NTCP accounts for more than 80% of hepatic uptake of conjugated bile salts across the sinusoidal membrane of hepatocytes. Another transporter located on the apical membrane of hepatocytes and enterocytes is Na+-independent organic anion-transporting polypeptide (OATP). OATP transports bile acids, bilirubin and hormones across the apical membrane in hepatocytes.

OATPs are expressed in many tissues transporting anions, as well as neutral and even cationic compounds. On the other hand, ABCA1 is responsible for efflux cholesterol and phospholipid, which is important in high density lipoprotein (HDL) formation. Mutation

in ABCA1 gene causes high density lipoprotein deficiency (HDL), which is called

Tangier disease [45-46].

Figure 3. Enterohepatic circulation of bile acids.

Bile acids are synthesized in the liver and conjugated bile acids are secreted in bile canaliculi (green oval) by apical membrane bile acid export pump mainly BSEP (Upper panel figure). MRP2 and MDR2/3 transport bile acids and phospholipids, respectively in bile canaliculi. Bile acids are reabsorbed by bile duct epithelial cells, cholangiocytes (not shown in diagram) and recycled to hepatocytes. After a meal, bile acids are excreted from the gallbladder into the intestine (shown by blue arrow on right). In the intestine, enterocytes (lower panel of figure) reabsorbs bile acids via OATP and ASBT

transporters. Bile acids travel across enterocytes and are excreted in the portal vein via

OSTα and OSTβ. Heterodimer ABCG5/G8 does efflux excess of cholesterol and plants steroids to intestinal lumen. Portal blood carries bile acids back to the liver, which are taken up by NTCP and OATP.

1.2. Bile acid signaling network

Recent studies show that bile acids are complex metabolic integrators and signaling molecules.

1.2.1. FXR/SHP/FGF19

Bile acids have been identified as an endogenous ligand for FXR (Fig. 4). The hydrophobic bile acid, CDCA, is the most potent activator of FXR [47]. Bile acid activated FXR inhibits CYP7A1 transcription indirectly via inducing co-repressor small heterodimer partner (SHP) gene expression [48-49]. SHP also suppresses gluconeogenesis via interaction with the glucocorticoid receptor (GR). Recently, it has been observed that bile acids induce the post-prandial hormone FGF19 in humans and

FGF15 in mice via activation of FXR in the intestine [50-51]. FGF19 is excreted into portal circulation. In hepatocytes, FGF19/15 activates fibroblast growth factor receptor 4

(FGFR4) and further leads to activation of the MAPK/ JNK/ERK1/2 pathways [52]. This acts as an endocrine signaling mechanism to inhibit the bile acid synthesis by suppressing

CYP7A1 gene expression.

1.2.2. G protein-coupled bile acid receptor 1 (GPBAR1)/ TGR5

Recently identified bile acid activated G protein coupled receptor called TGR5 is located in liver resident macrophages (Kupffer cells), brown adipose tissue, intestine and gallbladder epithelial. TGR5 is activated by lithocholic acid (LCA), while weakly activated by DCA [53-55]. TGR5 plays a very important role in secretion of glucagon like peptide 1 (GLP1) from entero-endocrine L cells after food intake. GLP1 regulates insulin secretion from pancreatic β cells (as shown in Fig.4). Therefore, direct stimulation

of TGR5 by bile acids or its derivatives is a promising GLP1 based therapeutic approach

[54]. Furthermore, TGR5 in microphages or Kupffer cells stimulates cAMP based signaling pathway, which reduces anti-inflammatory cytokine production [12, 55]. In brown adipose tissue, TGR5 activation leads to the activation of type 2 iodothyronine deiodinase 2 (D2), which converts T4 to T3, an activate form of thyroid hormone [53,

56]. Activation of the thyroid hormone receptor (TR) increases mitochondrial oxidative phosphorylation in muscle and uncoupling in brown adipose tissue (BAT), resulting in enhanced energy expenditure [53].

1.2.3. Signaling pathways

In the colon, concentrations of DCA are very high. DCA is a promoter of colon cancer. DCA activates the MAPK signaling pathway and increases cell survival to promote the cancer cell growth in colon [57-59]. On the other hand, DCA also modulates

P38 kinase activity and increases apoptosis and hepatic cell death [59].

Intestine

Figure 4. Bile acid signaling network

In hepatocytes, bile acids (shown by green squares) activate the nuclear receptor FXR, which activates co-repressor SHP to inhibit CYP7A1 gene expression (lower panel). SHP also interacts with GR and inhibits gluconeogenesis in the liver. In the intestine, bile acids activate FXR, which induces FGF19. FGF19/15 is secreted into the portal circulation and transported to hepatocytes to activate the ERK/JNK pathway in hepatocytes. In the intestine, bile acids also activate the TGR5 receptor signaling

pathway, which activates GLP1 secretion from endocrine L cells. GLP1 stimulates insulin secretion from the pancreas.

(Modified from Current Diabetes Report (2010) 10:70–77).

1.3. Nuclear receptor

Nuclear receptors are ligand activated transcription factors that regulate gene expression in response to hormone or other stimuli [60-62]. Several nuclear receptors regulate bile acid homeostasis therefore it is important to define these nuclear receptors and highlight some of their major functions in bile acid synthesis.

1.3.1. Structure and response element

A typical nuclear receptor has five domains, a variable NH2-terminal (AF1 activation domain), DNA-binding domain DBD (Zinc finger domain), linking hinge domain and ligand binding domain and C-terminal domain. DNA binding domain is the most conserved among all nuclear receptors family protein, which consist of two Zn fingers. The DBD is responsible for DNA binding and dimerization [62-63]. Ligand or hormone binding causes allosteric changes in the nuclear receptor and increases homo or hetero dimerization (Fig. 5). The N-terminal domain of nuclear receptor is variable in length and primary sequence. In many cases, transcriptional activation function domain

(AF-1) is located in N-terminal domain. In contrast, the AF-2 overlaps with the C- terminal ligand binding region and is responsible for the ligand-dependent transactivation. The strong AF-1 domain in PPARα and RORα is often phosphorylated by different kinase enhancing transcriptional activity [64-66]. The DBD of each nuclear receptor recognizes a specific motif on the gene promoter called hormone response element [67-68]. Nuclear receptors and their response elements are summarized in the

Fig. 5.

Figure 5. Nuclear receptor binding sites

Classical hormone nuclear receptors (Type I) binds as a homodimer to a palindrome sequence or inverted repeat of AGGTCA region (second from left in figure) eg: glucocorticoid receptor (GR), estrogen receptor (ER), thyroid hormone receptor (TR).

Type II nuclear receptors, such as FXR and VDR form a heterodimer with retinoic acid X receptor (RXR) and bind to inverted repeats or direct repeats to the target DNA sequence.

Type III nuclear receptors are orphan nuclear receptors, which bind as monomers to the

monomeric sequence with preceding A/T rich region with or without hormone stimuli such as RORα and Rev-erbα.

1.3.2. Farnesoid X Receptor (FXR NR1H4)

FXR is a bile acid activated receptor and plays an important role in the regulation of bile acid synthesis, glucose and lipid metabolism. FXR is highly expressed in the liver, intestine, adrenal gland and kidney (68). FXR forms a heterodimer with RXR and preferentially binds to the IR-1 motifs on the target gene promoter [47, 69]. The hydrophobic bile acid, CDCA, is the most potent activator of FXR [47]. The secondary bile acids, LCA and DCA, are less effective, and hydrophilic bile acids, ursodeoxycholic acid (UDCA) and muricholic acids, are inactive for activation of FXR [70-71]. A FXR agonist, GW4064 is 200 fold more potent activator of FXR than CDCA. The Fxr knockout mice show elevated serum bile acids, cholesterol and triglycerides, and reduced bile acid pool and impaired bile acid inhibition of cyp7a1 gene expression [72]. Enterohepatic circulation of bile acids is severely altered in FXR knock-out mice [73-74]. Bile acid activated FXR inhibits CYP7A1 transcription indirectly, by inducing a co-repressor SHP expression, [48-49] via interaction with LRH1 or HNF4α. FXR protects the liver from bile acid accumulation by inhibiting NTCP gene expression, while inducing bile acid efflux transporter pump BSEP expression [75]. FXR also induces IBABP protein expression in the intestine [76-77]. Another mechanism of FXR dependent regulation of cyp7a1 was discovered recently. In the intestine, FXR induces FGF-19/15, which inhibits

CYP7A1 gene expression but does not alter CYP8B1 gene expression [52, 78]. Human

CYP8B1 promoter has a FXR binding site that activates gene expression after GW4064 and retinoic acid treatment through interaction with the activation domain of G protein pathway supressor-2 (GPS2) [79]. This mechanism is completely absent in rodents.

These results suggest that the FXR/SHP pathway may be the major mechanism for bile acid inhibition of CYP8B1 gene transcription. On the other hand, FXR/FGF19/FGFR4 pathway activates MAPK/ERK pathway and inhibits CYP7A1 gene expression.

1.3.3. Pregnane X Receptor (PXR, NR1I2)

PXR is a promiscuous xenobiotic receptor that is activated by unrelated steroids

(e.g. dexamethasone, 16α-carbonitrile (PCN), xenobiotic drugs (like phenobarbital, rifampicin) and antibiotics. PXR is predominantly expressed in the liver and intestine.

PXR forms heterodimer with RXR and binds to various response elements consisting of

DR-3, DR-4, DR-5 or ER-6 to regulate drug metabolizing enzymes [80-81]. PXR induces

CYP3A4, the most abundant cytochrome P450 isozyme expressed in the human liver and intestine, which metabolizes about 60% of the clinically used drugs in the body. LCA has been identified as a PXR ligand [11, 82]. PXR functions as a bile acid sensor and induce

CYP3A4 and CYP3A11 expression, which converts LCA to a more hydrophilic and less toxic bile acid, hyodeoxycholic acid (HDCA). PXR plays an important role in induction of phase II drug metabolizing enzyme.

1.3.4. Vitamin D3 Receptor (VDR, NR1I1)

VDR is activated by 1α, 25, dihydroxy vitamin D3 and plays a central role in calcium homeostasis. VDR has been implicated in regulating diverse biological functions, including immunity, cellular proliferation and differentiation [83]. VDR is primarily expressed in the bones, kidney, intestine and parathyroid glands. VDR and

RXR heterodimer binds to DR-3 sites [83]. The physiological ligand of VDR is 1α, 25- dihydroxyvitamin D [1α, 25(OH)2 D3], the active form of vitamin D3. However, recently

LCA has been identified as a VDR ligand [10]. LCA acetate is the most potent VDR agonists [10]. VDR is also expressed in the human liver and regulates bile acid synthesis by inhibiting CYP7A1 gene expression [84]. Activation of VDR by LCA or 1α, 25(OH)2

D3, induces the expression of CYP3A4, which detoxifies LCA in the liver and intestine.

This serves as a protective mechanism against colorectal cancers in humans [85].

1.3.5. Liver X Receptor (LXR, NR1H3)

Two isoforms of LXR, LXRα and LXRβ play an important role in cholesterol homeostasis. LXRα is expressed in liver, spleen, adipose tissue, lungs and pituitary, whereas LXRβ is ubiquitously expressed. 22 (R)-hydroxycholesterol, 24 (S)-hydroxy cholesterol and 25-epoxycholesterol are the most potent LXR ligands [86]. LXRα functions as a regulator of cholesterol metabolism and stimulates genes expression by binding to DR-4 motifs as heterodimer with RXRα. In response to high cholesterol diet,

LXRα induces rat Cyp7a1 expression, which converts excess cholesterol in bile acids.

However, LXRα has no stimulating effect on human CYP7A1, which lack DR-4 motifs

[87-89]. In contrast, cholesterol feeding to mice and rats decreases the activity of

Cyp8b1 [90]. Oxysterol activated LXRα also binds and stimulates the sterol response element binding protein-1c (SREBP-1c) gene, and thus stimulates lipogenesis leading to hypertriglyceridemia [91-92]. LXRα also induces ABCA1 gene, which functions as a cholesterol and phospholipid efflux transporter for HDL synthesis, and ABCG5/G8, which regulates biliary sterol efflux [93-94]. Lxrα knock-out 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 cholesterol in the liver. These

mice also show a decrease in the sterol regulatory element-binding protein-1c (SERBP-

1c) [95-96].

1.3.6. 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. HNF4α homodimer binds to the DR-1 motif on the target gene promoter [97]. HNF4α is also involved in human genetic diseases. Mutations in the HNF4α gene cause maturity-onset diabetes of the young 1. Targeted disruption of the HNF4α gene was found to be embryonically lethal, indicating that HNF4α is an essential factor for mammalian development. Liver-specific HNF4α-null mice were produced using the Cre-loxP system [98]. These mice exhibited impaired lipid homeostasis. In addition, these mice have increased serum BA levels and reduced expression of BA transporters, such as NTCP and organic anion transporter polypeptide 1

(OATP1) [99]. Mutation of the HNF4α binding site markedly reduced human CYP7A1 promoter activity indicating that HNF4α is crucial for basal level transcription of

CYP7A1 gene expression [99]. HNF4α binding sites have also been identified in

CYP8B1 genes and stimulate its expression [100].

1.3.7. Liver receptor homolog-1 (LRH-1 NR5A2)

LRH1 is another orphan nuclear receptor highly expressed in liver, intestine and pancreas and is most related to steroidogenic factor 1 (SF-1) expressed in steroidogenic tissues. LRH1, also known as FTF, binds as a monomer to the core DNA sequence, 5'-

TCAAGGTCA-3'. LRH1 binding sites have been identified in the CYP7A1 and CYP8B1 gene promoter, which overlaps with HNF4α binding site in both gene promoters [101-

102]. LRH1 differentially regulates human and mouse CYP7A1 and CYP8B1 gene expression [77, 101]. In liver specific LRH1 gene knockout mice, cyp8b1 mRNA is abolished, while there is no significant effect on basal Cyp7a1 mRNA expression. This study suggests that LRH1 is an important regulator of mouse cyp8b1 gene, which is inhibited by FXR-SHP-LRH1 mechanism [103-104].

1.3.8. Small hetero dimer partner (SHP, NR0B2)

SHP is an atypical nuclear receptor binding protein that contains a nuclear receptor-interacting domain and a repressor domain, but lacks a conventional DNA binding domain. SHP acts as a co-repressor, which inhibits transactivation activity of various nuclear receptors such as LRH1, RAR, HNF4α, estrogen receptor α and β (ERα and ERβ), peroxisome proliferators activated receptor (PPAR), GR and other receptors

[81, 105-107]. SHP competes with co-activators like steroid receptor co-activator (SRC1) for binding to nuclear receptor binding or directly through the C-terminal repressor domain. SHP mediates the suppression of CYP7A1, CYP8B1 and CYP27A1 by interacting with histone methyl transferases and histone deacetylases (HDAC1/HDAC6) [102].

Deletion of the SHP gene in mice increases bile acid pool size. However, bile acid feeding still inhibits bile acid synthesis in SHP knock-out mice. This study suggests that a compensatory pathway for bile acid feedback inhibition exists [108]. SHP activity can be modulated by ligands such as retinoid related molecules, including 4-[3-(1-adamantyl)-4- hydroxyphenyl]-3-chlorocinnamic acid (3Cl-AHPC), which binds to SHP and acts as an agonist to increase SHP activity and repress bile acid synthesis by inhibiting CYP7A1 and

CYP8B1 gene expression [109]. Mice deficient in both Fxr and Shp of both exhibit

cholestasis and liver injury as early as three weeks of age, and this is due to the dysregulation of bile acid synthesis and enterohepatic circulation [110]. Furthermore,

SHP gene mutations have been identified in obese Japanese patients with early onset of diabetes [111].

1.3.9. Peroxisome proliferator activated receptor α (PPARα NR1C1)

. The PPARα form heterodimer with the retinoid X receptor (RXR) and binds to the peroxisome proliferator-response element (PPRE) located in the promoter region of genes. PPARα regulates bile acid metabolism [112]. PPARα has also been shown to play a critical role in the adaptive response to fasting in mice as the induction of several genes involved in lipid catabolism is abolished in the PPARα-null mice [113]. Fibrates are a group of hypolipidemic agents that efficiently lower serum triglyceride levels by affecting the expression of many genes involved in lipid metabolism [114-115]. Fibrates activate PPARα activity and lower plasma cholesterol levels in human. PPARα also induces FGF21 gene expression upon starvation, which regulates ketone body production

[110, 116]. The PPARα agonist, Wy14,643 suppresses CYP7A1 in HepG2 cells via a

PPRE (DR-1 site), but the PPARα/RXRα heterodimer cannot bind to that site. PPARα stimulates Cyp8b1 gene expression and increases CA synthesis in rats, but binds very weakly to the Cyp8b1 gene promoter [66, 112]. Free fatty acid activated PPARα stimulates LXRα expression and induces the target genes of the latter, namely ABCA1 and ABCG1 in macrophages [117-118]. Pparα knockout mice display a defect in the lipid and lipoprotein metabolism. Similarly, these mice also show abolished cholic acid

production and cyp8b1 gene expression. There is a distinct DR1 site on mouse and rat cyp8b1 promoter but, PPRE is absent on human CYP8B1 promoter [66, 112].

1.3.10. Retinoic acid related orphan nuclear receptor (RORα, NR1F1 and RORγ )

Retinoic acid related orphan nuclear receptor alpha (RORα) and Retinoic acid related orphan nuclear receptor gamma (RORγ) are constitutively active nuclear receptors. RORα/γ binds as a monomer to its binding site AGGTCA with A/T rich region flanking at the 5’end. RORα has four splice variants distributed differently among the tissues. All splice variants have common ligand binding and DNA binding domain but, have different N–terminal domain. According to earlier literature, DNA binding specificity varies among the different isoforms of RORα. RORα1 has better DNA binding ability compared to RORα2 and RORα3. RORα1 and ROR4 are expressed in the liver but, ROR4 is the most predominant form [67-68, 119]. All RORα isoforms have an activation domain and activate transcription of target genes mainly by recruitment of co-activators [119]. Various co-activators such as GRIP, PBP, SRC1 and

SRC2 interact with RORα. Some of the interactions are tissue specific and contribute to distinct physiological function of RORα [119-120]. RORα staggerer mice (sg/sg) have an intragenic deletion in the coding region of RORα gene, removing an exon downstream of the DNA binding domain [121]. This results in a frame shift mutation that affects the co- expressed isoforms RORα1 and RORα4. RORα transcript levels are significantly reduced in staggerer (sg/sg) mice. Several circadian gene profiles are altered in sg/sg mice. This suggests a role of RORα in the regulation of circadian rhythm. It has been demonstrated that male and female homozygous staggerer (sg/sg) mice are characterized by hypo-α-

lipoproteinemia and decreased serum cholesterol level (total and high density lipoprotein

(HDL). Decrease in serum cholesterol is due to reduced apolipoprotein A-I (apoA1) expression and apolipoprotein C-III (apoCIII) expression in liver. RORα (sg/sg) mice are susceptible to atherosclerosis on an atherogenic diet [122]. These studies indicate that

RORα is involved in the regulation of lipoprotein homeostasis in mice. When sg/sg mice are fed a high fat diet for 10-weeks, they exhibit a lower weight gain with reduced hepatic triglycerides and notable decrease in white and brown adipose tissues. Candidate- based expression profiling demonstrated that the dyslipidemia in sg/sg mice is associated with decreased hepatic expression of SREBP-1c, and cholesterol efflux transporters,

ABCA1 and ABCG1. This is consistent with the reduced serum lipids. Furthermore, the lean phenotype in sg/sg mice is also characterized by significantly increased expression of PGC-1α, PGC-1β, and lipin1 mRNA in liver and white and brown adipose tissues

[123].

More recently, a role for RORα in regulation of glucose metabolism was characterized. It was found that loss of the co activator SRC-2 in mice led to a phenotype similar to Von Gierke disease, which is associated with severe hypoglycemia and abnormal accumulation of glycogen in the liver [124]. Loss of expression of the enzyme glucose-6-phosphatase (G6Pase) is responsible for 80% of the diagnosed Von Gierke disease cases. Research shows that SRC-2 is required for RORα to regulate G6pase gene expression during fasting [124]. RORα and RORγ double knockout mice show hypoglycemia. This observation indicates that RORα and RORγ have overlapping functions in glucose homeostasis [119, 125].

X-ray crystallographic studies identified that cholesterol and cholesterol sulphate bind to the ligand binding pocket of RORα [64]. It remains unclear if cholesterol, or a derivative of this sterol, is a physiological ligand for RORα. Recently, oxysterols, mainly

7-oxygenated sterols mainly 7α-hydroxycholesterol (7αOHC) has been shown to bind

with RORα and RORγ with high affinity (Ki ∼20 nM). Interestingly, oxysterol binding to

RORα decreases co-activator binding and suppresses transcriptional activity of RORα receptors [126-127]. Therefore, 7αOHC acts as an inverse agonist for both the receptors.

There is no identified physiological ligand that activates transcriptional activity of RORα.

According to a recent microarray analysis, RORα sg/sg mouse liver shows a twofold decrease in cyp8b1 mRNA with no change in cyp7a1 mRNA compared to wild type mice [125]. RORα/γ double knockout mice show a 4 to 5 fold decrease in cyp8b1 mRNA. These data indicate that RORα may regulate cholic acid synthesis. These observations lead me to further investigate the role of RORα in regulation of human

CYP8B1 and mouse cyp8b1 gene transcription.

1.3.11. Rev-erbα (NR1D1)

Rev-erbα is a transcriptional repressor that interacts with nuclear receptor co-repressor 1

(Nco-R1) to suppress target gene expression [128]. Rev-erbα is highly expressed in the liver, skeletal muscle, adipose tissue, and the brain and participates in the organogenesis.

Rev-erbα suppresses bmal1 gene expression and regulates circadian rhythm [128]. It also regulates lipolysis and suppresses APOCIII gene expression. RORα acts as a stimulator, while Rev-erbα acts as a suppressor and regulates various physiological processes [129].

Rev-erbα expression is also regulated at the post-translational level. Recent study

indicates that Rev-erbα is phosphorylated by glycogen synthase kinase (GSK3β), which increases Rev-Erbα protein stability [130].

1.4. Transcription factors and co-activators

Transcription factor are sometimes called a sequence-specific DNA-binding proteins. They bind to the specific DNA sequence and regulate target gene expression.

Co-activators are proteins that increase the rate of transcription by interacting with transcription factors and nuclear receptors; however they do not directly bind to the

DNA in a sequence-specific manner. Co-activators 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 [115, 131]. Some co- activators possess intrinsic histone acetyl (HAT) activity, which acetylates histones and causes chromatin to relax in a limited region allowing increased access to the DNA.

1.4.1. Sterol regulatory element-binding protein (SREBP)

SREBPs are basic helix-loop-helix leucine zipper (bHLHLZ) transcription factors. Three different SREBPs have been described. SREBP-1a and SREBP-1c are produced from a single gene through the use of alternate promoters that produce transcripts with different first exons. On the other hand, SREBP-2 is produced from a separate gene [132-133]. SREBP1a is expressed in hepatic cell lines, while SREBP1c is expressed in human and mouse liver. Newly synthesized SREBPs are inserted into the endoplasmic reticulum (ER) membrane as inactive precursors [134]. SREBP isoform binds to the sterol-sensing protein, SREBP cleavage-activating protein (Scap), which interacts with INSIG 1/2. In cholesterol-poor cells, the SREBP-Scap complex is

transported to the Golgi by dissociation from INSIG 1/2 [91, 135]. In the Golgi, S1P and

S2P proteases are responsible for proteolytic processing of SREBP1c. Insulin stimulates

SREBP-1c in the liver by increasing SREBP1c proteolytic cleavage. [136]. Thus, Insulin induces SREBP-1c processing, thereby allowing it to stimulate fatty acid synthesis.

According to the early literature, both SREBP1a and SREBP1c increase mouse and human CYP8B1 gene transcription, while SREBP-2 suppresses human CYP8B1 gene transcription via interaction with FTF [137]. The rat Cyp8b1 promoter contains several sterol response elements (SREs) and E-box motifs, which bind SREBP and stimulate the

Cyp8b1 gene expression [138].

1.4.2. Peroxisome proliferator-activated receptor γ co-activator 1α (PGC-1α)

Peroxisome proliferator-activated receptor γ co-activator 1α (PGC-1α) plays an important role in activation of nuclear receptors. PGC-1α does not have an intrinsic enzyme activity. Binding of PGC-1α to transcription factors induces a conformational change to recruit other co-activator proteins containing histone acetyl transferase (HAT) activity to stimulate gene transcription. PGC-1α interacts with a host of NRs such as

HNF4α, ER, PPARα, RXRα, GR, retinoic acid receptor (RAR), TR etc. PGC-1α gene expression is induced upon fasting to regulate various metabolic processes in the energy metabolism [139-140]. Acetylation of PGC1α decreases its activity of binding to nuclear receptors. Therefore, PGC1α is activated after deacetylation by Sirtuin 1, (SIRT1) also known as NAD-dependent deacetylase [141].

1.4.3. Cyclic AMP response element binding (CREB)-binding protein (CBP)

CBP/p300 is a transcriptional co-activator, which has histone acetyl transferase activity. CBP is found to interact with phosphorylated cyclic AMP response element binding protein (CREB) and regulates gluconeogenesis [142]. p300 is first found to interact with adenovirus E1A associated protein, which is 86% homologous to CBP

[143]. Another important class of histone acetyl transferase is steroid receptor co- activator family proteins SRC1, SRC2 and SRC3. They play a major role in various metabolic processes. Mice deficient of SRC1 are resistant to the thyroid hormone. This indicates that SRC1 enhances the thyroid hormone receptor (TR) dependent action [144-

145]. Furthermore, absence of SRC2 results in a glycogenopathy resembling Von

Gierke's disease in mice.

1.5. Transcriptional regulation of CYP8B1 gene

As discussed earlier, the CYP8B1 is important for producing cholic acid, determining the hydrophobicity of bile acid and facilitating cholesterol absorption in intestine. In humans, the CYP8B1 gene is located on chromosome 21. Interestingly, the

CYP8B1 gene has no intron. CYP8B1 cDNA is consist of a 1509-base pair open reading frame encoding 503 amino acid residues, which have characteristic conserved domains for heme, steroid and oxygen binding [146-149]. CYP8B1 has a high homology (43%) with prostacyclin synthase (CYP8A1) [146]. CYP8B1 gene knockout mice have an altered bile acid composition with no CA and an increased muricholic acids, a compensatory up-regulation of Cyp7A1 mRNA and increased bile acid pool size. Their serum cholesterol, triglyceride levels and lipoprotein profiles are unchanged compared to

wild type animals [150-151]. Due to the reduced intestinal absorption of cholesterol, the cyp8b1 null mice have increased expression of the genes involved in hepatic de-novo cholesterol synthesis [150]. The role of cholic acid in absorption of cholesterol is also supported by combined feeding of CA and cholesterol for longer periods, which leads to hypercholesterolemia and the formation of cholesterol gallstones in mice [152]. This study indicates that cholic acid is responsible for cholesterol absorption in intestine and cholesterol accumulation in the liver [153-155].

Several studies show that CYP8B1 gene transcription is repressed by bile acids

[100]. The molecular mechanism behind the repression of CYP8B1 gene transcription remains obscure. Bile acids, mainly CDCA, activate FXR, which in turn activates transcription of SHP. SHP interacts with the LRH1/hFTF and down-regulates expression of CYP8B1 gene expression. In intestine, FXR activates FGF-15, which inhibits CYP7A1 gene expression but, does not alter CYP8B1 gene expression [52, 78]. Human CYP8B1 promoter has FXR binding site that induces hCYP8B1 gene expression through interaction with the activation domain of G protein pathway supressor-2 (GPS2). This mechanism is completely absent in rodents [79]. These results indicate that the FXR/SHP pathway may be the major mechanism for bile acid inhibition of CYP8B1 gene transcription.

Cytokine signaling pathways may play a critical role in CYP8B1 gene regulation in cholestasis and liver inflammation [156-157]. CYP8B1 gene transcription is strongly repressed via the MAPK/JNK signaling pathway. Interleukin-1β (IL-1β) inhibits the

HNF4α trans-activation of the hCYP8B1 gene [158].

According to the early studies, LRH1 and HNF4α differentially regulate rat and human CYP8B1 genes expression [100-101]. In rats, SHP interacts with FTF and inhibits rat Cyp8b1 gene expression [159]. In humans, SHP interacts with HNF4α and inhibits the hCYP8B1 gene expression [100]. Studies indicate that LRH1 is an important factor in the inhibition of mouse cyp8b1 gene by FXR-SHP mechanism [103]. According to the early studies, both SREBP’s, SREBP1a and SREBP1c increase mouse and human CYP8B1 gene transcription while SREBP-2 suppresses hCYP8B1 gene transcription via interaction with FTF [160]. Cholesterol and insulin also repress human and mouse cyp8b1 gene transcription. Cholesterol feeding in mice reduces the activity of cyp8b1 significantly and decreases cyp8b1 mRNA. The mechanism behind this observation remains obscure [138,

153]. HNF4α binding site is identified in the mouse Cyp8b1 promoter region that overlaps with PPARα binding site (DR1). PPARα activates cyp8b1 gene transcription in mice and rats but there is no such binding site on the hCYP8B1 gene promoter region

[112].

1.6. Circadian regulation of bile acid synthesis

A circadian rhythm is any biological process that displays an endogenous oscillation of about 24 hours. These rhythms are driven by a circadian clock, and rhythms have been widely observed in plants, animals, fungi and cyanobacteria. These changes are governed by an internal (endogenous) clock. This internal clock consists of an array of genes and the protein products they encode, which regulate various physiological processes throughout the body [129, 161].

Diurnal rhythm is also a cyclic repetition of biological processes that occurs at daily intervals. However, a diurnal rhythm may be called circadian if the rhythm has been shown to persist under constant environmental conditions and thereby can be distinguished from those rhythms that are simply a response to 24-hour environmental changes such dark-light cycles or temperature changes. For practical purposes, however, there is little reason to distinguish between diurnal and circadian rhythms, because almost all diurnal rhythms are found to be circadian regulated [161].

Diurnal rhythm of CYP7A1 is studied in rats housed in a constant dark. This study indicates CYP7A1 gene expression still follows circadian rhythm. Therefore, nutrition status and day and light cycle regulate bile acid synthesis [102, 162].

Circadian clock genes have two limbs; the CLOCK and BMAL1 complex positively regulate gene expression, while the CRY and PER proteins interfere with

CLOCK and BMAL1 complex formation and represses CLOCK/BMAL1 regulated gene expression. RORα regulates the positive limb of clock genes by elevating BMAL1 gene expression. Mutation of the CLOCK gene in mice disrupts cholesterol metabolism and leads to cholesterol accumulation in liver [163]. This observation underlines the importance of diurnal rhythm in cholesterol homeostasis. It is evident that dysregulation of circadian rhythm often cause metabolic syndrome such as obesity [129, 161, 164-165].

Circadian expression of various genes in liver is regulated by light and dark as well as fasting and feeding cycle. Studies indicate that circadian rhythm of cyp7a1 could be dependent on various circadian regulation factors such as DBP, Rev-erbα, DEC1/2 and

E4BP4 [164-166]. There are few reports about circadian rhythm of cyp8b1, which is

regulated by insulin, DBP and indirectly by Rev-erbα via SHP regulation [167-168]. Still, there are many aspects of the diurnal regulation of bile acid synthesis genes and their impact on cholesterol metabolism is unknown.

1.7. Nutrient regulation

Fasting is a complex biological phenomenon in which various nuclear receptors as well as co-activators regulate gluconeogenesis and glycogenolysis. During the initial stages of fasting, body senses lowering of blood glucose level, which in turn activates glucagon secretion from pancreatic α cell. Glucagon is a 29 amino acids polypeptide and circulates as a free form in plasma. It has a short half-life of ῀20 min [169-170]. In liver, glucagon receptor is expressed at a higher level. Glucagon receptor is a G protein coupled receptor

(GPCR) and consists of Gα, Gβ and Gγ subunits [171]. Binding of glucagon to GPCR activates the glucagon receptor and causes dissociation of the Gβ and Gγ subunits from

Gα subunit. Activated Gα subunit activates adenylate cyclase activity to produce cAMP

[172]. Cyclic AMP binds to the regulatory subunit of protein kinase A (PKA) and causes dissociation of the catalytic subunit. Activation of the catalytic subunit takes place by auto-phosphorylation at T197. Activated PKA phosphorylates CREB, which regulates gluconeogenesis by inducing phosphoenol pyruvate carboxykinase (PEPCK) gene expression. In the later stages of fasting, PKA activates lipase to hydrolyze stored triglycerides into fatty acids, increasing free fatty acid concentration in the blood. Free fatty acids can activate PPARα in the liver, which activates fatty acid oxidation for energy production. PPARα induces FGF21, which increases ketone body production in

the liver. In the brain, FGF21 activates torpor state (less energy expenditure and activity)

[110, 116].

There is a significant difference in the fasting metabolism between healthy and diabetic individuals. According to the earlier studies, glucagon metabolism is disrupted in insulin insufficient Diabetes mellitus (type 2) and insulin deficient diabetes (Type 1)

[170, 173-177] in human subjects. In normal individuals, fasting glucagon concentration is 94-100 pg/ml. After glucose or alanine infusion, glucagon levels rapidly decreases to

13 pg/ml. In type 1 diabetes, fasting glucagon concentration increases to 180 pg/ml, but upon glucose or alanine infusion, glucagon level remains very high at 100 pg/ml [176-

178]. In case of type 2 diabetes, fasting glucagon concentration remains in the normal rage 100-110 pg/ml, but arginine or glucose infusion does not decrease the glucagon concentration, which remains at 57 pg/ml. In the post prandial period, endocrine L cells secrete glucagon like peptide GLP1, which increases insulin secretion from pancreatic β cells. Further, Insulin secretion inhibits glucagon production and glucagon secretion from pancreatic cells [170, 179-180]. Studies in human subjects with type 1 diabetes reveal hyperglucagonemia due to complete absence of insulin [181]. In case of diabetes mellitus type 2, the glucagon to insulin ratio is relatively higher than that in normal individuals because insulin resistance is causes secretion of more glucagon from pancreatic α cells [177]. All this studies indicate that glucagon is an important link to study the fasting to feeding transition in normal and diabetic individuals.

Figure 6. Metabolism in fasting and starvation

Fasting is an important biological process that involves all tissues in the body (above diagram). Fasting increases glucagon secretion, which induces gluconeogenesis and glycogenolysis in the liver (left hand side). Although the glucagon receptor is not expressed in mucle, glycogenolysis is stimulated by glucagon action (not shown in the figure). In adipocytes, glucagon also increases the release of free fatty acids. Upon

starvation, free fatty acids can be taken up by the liver to activate PPARα, which induces

FGF21 release. FGF21 is a major fasting induced hormone, which decreases the overall energy of the animal or human being. (Modified from Cell metabolism, (5) 2007, 405-

407)

1.8. Hypothesis and specific aims

Recent studies show that bile acids play a key role in the regulation of glucose, lipid and energy metabolism. Cholic acid determines bile acid composition and cholesterol absorption in intestine. Ablation of the CYP8B1 gene in mice reduces intestinal cholesterol absorption and elevates the de-novo synthesis of cholesterol [150-

151]. Previous reports indicate that elevation of cholic acid in human bile increases formation of gallstone [153]. Abolished cholic acid synthesis decreases atherosclerosis in apolipoprotein E knockout mice [155]. These findings suggest that increase in cholic acid has detrimental effect. In contrast, impaired synthesis of cholic acid is linked to the decreased FXR signaling and liver steatosis in liver specific FoxO1 and LDLR knockout mice [182]. Thus, the exact role of cholic acid in development of hypercholesterolemia and atherosclerosis is still not clear. According to the earlier studies, glucagon metabolism is disrupted in both type 1 and type 2 diabetes [170, 173-177]. Therefore, it is important to study effect of glucagon on bile acid composition and CYP8B1 gene expression. This study may shed light on the underlying mechanism in development of hypercholesterolemia in the diabetic individuals. Overall objective of this work is to understand the molecular mechanism of fasting induced Cyp8b1 gene expression.

RORα is an important nuclear receptor involved in regulation of cholesterol homeostasis, circadian rhythm and bone development [129, 183]. Deletion of RORα and

RORγ in mice significantly decreases cyp8b1 mRNA level in the mouse liver [119]. All these observations suggest that RORα may be a key regulator of cholic acid synthesis

and CYP8B1 gene expression during fasting and circadian rhythm. Three following specific aims are designed to study the hypothesis.

Specific Aim 1: Study the regulation of CYP8B1 expression in fasting.

Bile acid synthesis is regulated by fasting and feeding cycle. Glucagon is fasting induced hormone. Therefore, it is important to study how glucagon signaling affects

CYP8B1 expression and bile acid synthesis during fasting.

Prior studies from our laboratory show that glucagon or cAMP treatment inhibits

CYP7A1 mRNA and decreases bile acid synthesis [184]. This is in contrast to earlier reports that fasting induces CYP7A1 mRNA expression in mice [185]. CYP7A1 and bile acids exhibit a strong circadian rhythm. It has been reported that insulin negatively regulates CYP8B1 gene expression and CYP8B1 may play a role in regulation of circadian rhythm. RORα is activated upon fasting to stimulate glycogen breakdown [34].

Therefore, we wanted to evaluate the possible role of RORα in regulation of CYP8B1 diurnal expression.

Hypothesis: Fasting increases glucagon secretion, and cAMP signaling induces human and mouse cyp8b1 gene expression.

a) Study the effect of glucagon and cAMP on CYP8B1 gene expression in liver cells.

Approach: Primary human hepatocytes and HepG2 cells were treated with glucagon and cAMP analog (8-Br-cAMP) and change in CYP8B1 mRNA levels were monitored by quantitative real time PCR.

b) Study the effect of fasting on cyp8b1 mRNA in mice.

Approach: Wild type (C57Bl/6J) mice were fasted then refed and cyp8b1 mRNA and protein expression were monitored using real time PCR and western blot analysis respectively.

c) Study circadian rhythm of RORα and cyp8b1 in mice.

Approach: Wild type (C57Bl/6J) mice had free access to food or were fasted then refed. A group of mice were sacrificed every 4 h, for 24 h. Liver tissues were collected to monitor Cyp8b1 and RORα mRNA expression using real time PCR.

Specific Aim 2: Study the role of RORα in regulation of CYP8B1 gene expression.

Microarray analysis of intragenic mutation in RORα mice (sg/sg) shows a decrease in

CYP8B1 mRNA expression in mouse liver [119]. This suggests that RORα may be involved in transcriptional regulation of CYP8B1 gene expression. Interestingly, 7α- hydroxycholesterol, which is synthesized by CYP7A1, antagonizes RORα activity.

Therefore, it is important to study the molecular mechanism of how RORα regulates bile acid metabolism.

Hypothesis: human and mouse CYP8B1 promoter may have a RORα binding site.

a) Study the effect of RORα on CYP8B1 gene expression and bile acid synthesis in mice.

Approach: To study the change in bile acid composition and bile acid pool size, adenovirus mediated over-expression of RORα in mouse liver was performed.

b) Study the role of RORα-mediated effect of glucagon or cAMP on CYP8B1 gene expression

Approach: Luciferase assay was performed of human CYP8B1 reporter. To monitor the effect of 8-Br-cAMP on CYP8B1 gene expression in HepG2 cells, RORα deletion was performed using siRNA.

c) Identify a RORα response element on CYP8B1 promoter.

Approach: Luciferase assay was performed using various human CYP8B1 reporter deletion constructs to delineate the region responsible for RORα action. Further, site directed mutagenesis and EMSA assay was performed to confirm the binding of RORα to

RORα response element.

Specific Aim 3: Study the mechanism of RORα regulation of CYP8B1 gene expression.

RORα interacts with co-activators to stimulate target gene transcription. A recent study indicates that the RORα is phosphorylated by cAMP activated protein kinase A (PKA) at serine 99 position. Therefore, it is important to understand if phosphorylation of RORα by PKA enhances its interaction with co-activators to increase CYP8B1 gene expression.

Hypothesis: Phosphorylation of RORα increases stability of the protein and increases

CYP8B1 gene expression.

a) Study the co-activator interaction with RORα and effect of cAMP on CYP8B1 gene expression

Approach: luciferase assay using co-transfection of different co-activators was performed. Further, RORα and different co-activator interactions were studied by mammalian two hybrid assay. To study RORα recruitment on CYP8B1 promoter, ChIP assay was performed using adenovirus mediated RORα over-expressed mouse liver.

b) Study post-translational modification of RORα upon cAMP treatment.

Approach: RORα protein expression was monitored upon cAMP treatment in HepG2 cells. Phosphorylation of RORα by PKA was studied. Effect of cAMP on RORα protein stability was studied with or without treatment of cycloheximide (CHX) in HepG2 cells.

CHAPTER II

MATERIALS AND METHODS

2.1. Animal

Male wild type C57BL/6J mice were purchased from the Jackson Laboratory (Bar

Harbor, ME). All mice were maintained on a standard chow diet and water ad libitum and housed in a room with a 12 h light (6 am to 6 pm) and 12 h dark (6 pm to 6 am) cycle.

For the fasting and re-feeding study, mice were fasted from 8 a.m. and refed with a standard laboratory chow diet at midnight (12 am). All mice were sacrificed and tissues were frozen on dry ice and stored in -80oC

2.1.1. Circadian Rhythm studies

To study circadian rhythm, mice were housed in a similar condition mentioned above with free access to food and water. A group of 4-5 Mice were sacrificed every 4 h for 24 h at zeitgeber time (ZT), ZT 0 (6 am), ZT 2 (8 am), ZT 6 (12 pm), ZT 10 (4 pm),

ZT 14 (8 pm), ZT 18 (12.00 am) and ZT 22 (4 am). Lights on at ZT 0 and lights off at

ZT 12

2.1.2. Adenovirus injection

Mice (C57BL/6J) were injected with 100 µl adenovirus expressing RORα (Ad-

RORα, 2×109 infectious units / µl) or 100 µl adenovirus null (Ad-null, 1.8×109 infectious

units / µl) as control, via tail vein injection. After injection, these mice were housed in biohazard room with a 12 h light (6 am to 6 pm) and 12 h dark (6 pm to 6 am) cycle for seven days. One day before the experiment, mice were fasted overnight. The following day, mice were sacrificed using isoflurane inhalation and cervical-cord dislocation. All tissues were extracted and frozen immediately in -80o C.

2.2. Liver mRNA isolation and reverse transcription

Livers (0.2 mg) collected from the sacrificed mice were treated with 1 ml Trizol reagent and homogenized. Following that, 0.2 ml of chloroform was added in each tube.

The samples were vortexed for 15 sec and allowed to stand at room temperature. The resulting mixture was then centrifuged at 12,000 rpm for 2 min. Centrifugation separates the mixture into 3 phases: a lower red organic phase (containing protein), inter phase

(containing DNA) and a colorless upper aqueous phase (containing RNA). The aqueous phase was transferred to a 1.5-ml micro centrifuge tube and 0.5 ml of isopropanol was added to it. The samples were mixed gently by inverting the tubes 10 times. Further, this mixture was centrifuged for 15 min at 12000 rpm at 4oC to recover RNA pellet. RNA pellets were washed with 70% ethanol (500 µl) and dissolved in diethyl pyrocarbonate

(DEPC) treated water. In the next step, DNase was added to remove contamination of genomic DNA. Therefore, 0.4 µg of RNA was treated with 1 µl DNase and 5 µl DNAse buffer to a total volume of reaction of 50 µl and incubated at 37oC for 30 min. Further,

DNase enzyme was deactivated using DNase deactivation beads provided by DNase free kit (Ambion Life Technologies, Grand Island, NY). Isolated liver mRNA was dissolved

in DEPC treated water to measure RNA concentration and integrity using spectrophotometer (Thermo Scientific Fisher).

2.2.1. Reverse transcription

Reverse transcription was carried out to generate cDNA from mRNA for further analysis. Reverse transcription reaction (RT) was carried out using reverse transcription kit (Ambion life technologies, Grand Island, NY). First, 0.2 µg RNA was incubated with

1 µl oligo dT, 4 µl dNTP’s and 1 µl RNase inhibitor at 78oC for 5 min. This step allows oligo dT primers to anneal to the poly A tail. Further, 1 µl of reverse transcriptase enzyme was added and incubated at 42oC for 90 min to complete cDNA formation. Total reaction volume was maintained 20 µl. In order to deactivate reverse transcriptase, enzyme mixture was heated at 95oC for 5 min. The reaction mixture was cooled down to room temperature. Twenty µl of the reaction mixture was mixed with 180 µl of nuclease free water.

2.3. Quantitative Real time PCR (Q-RT-PCR)

Quantitative Real Time PCR was performed to determine the relative mRNA expression levels in each RNA sample. The mRNA of an endogenous reference gene was used to normalize from each sample, e.g.: ubiquitin C (UBC) and glyceraldehyde 3 phosphate dehydrogenase (GAPDH). The Q-RT PCR was performed using two different methods depending upon the availability of different reagents.

2.3.1. SYBR Green Method

In this method, a specific forward and a reverse primer were used to amplify a portion of the cDNA of interest. The method uses SYBR Green dye to detect PCR

products. Direct detection of PCR product was 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, Grand Island, NY) 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. Upon thermal activation, the modifier is released resulting in an active enzyme. The high- temperature incubation step required for activation ensures that active enzyme is generated only at temperatures where the DNA is fully denatured. The RORα and β-actin primers were purchased as mentioned in earlier [125]. Primers specific for the mouse cyp8b1 promoter were designed using primer express software. All primers for RT-PCR are listed in the table 1.

Table-1: Primers used for RT-PCR

Mouse cyp8b1 -600 TCCTCCCTGTGCCAGCTAAC (Forward) (F)

(ChIP assay) -416 AGGTTCCTGCCCTTGGACTT (Reverse) (R)

Mouse cyp8b1 -3.5 AGTTTGTTGCCTACAGTCCTATCAGAG (F)

(ChIP assay Control) -3.2 TGCCATCCTAGCGTATTTGCTGCCATTGTGC (R)

Human CYP8B1 +3 CCTTTACATCCTGGACTTTCCAA (F)

(ChIP assay) +255 GCTCTCGACACGCACACTGT (R )

MGB probe-: CTCTGGCACCCAGGG

Human RORα1 TCAGGTAAGGGTGAATGAT (F)

GCAATTCGACGGTTCCATT (R)

Human RORα4 CTCGATCATTGGACTGTTA (F)

CTGCAGTCCAATCGATGATG (R )

Human Β-Actin TCTACAATGAGCTGCGTGTGG (F)

GGAACCGCTCATTGCCAATG (R )

Mouse GAPDH GCCTTCCGTGTTCCTACC (F )

GCCTGCTTCACCACCTTC (R )

2.3.2. Taqman Method:

In this method, a specific forward, reverse primer and a TaqMan probe of 18-22 bp is labeled with a reporter fluorophore at the 5' end and a quencher fluorophore at the 3' end. The fluorescence of the reporter fluorophore is quenched by a quencher. During

PCR, the Taqman probe anneals specifically between the forward and reverse primers.

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. Taqman primer and probe specific for human (h) CYP8B1, mouse (m) cyp8b1, HNF4α, PGC1α, Rev- erbα, hCYP7B1, mcyp7b1 were purchased form Applied Biosystems.

2.3.3. 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-ΔΔCt) recommended by Applied Biosystems (Applied Biosystems, User Bulletin No.2, 1997). ΔCt was calculated by subtracting the Ct value of internal control gene from Ct value of gene of interest. For eg: Calculation of relative expression of RORα in adenovirus infected mouse liver. ΔCt = Ct (of RORα in Ad-RORα infusion)-Ct (of GAPDH in Ad-RORα infusion), ΔΔCt = ΔCt (of RORα in

Ad-RORαinfusing ) – ΔCt (of RORα in Ad-Control infusion). Further, the relative mRNA expression was

-ΔΔCt. calculated using formula 2 In case of animal experiments, each relative mRNA expression was calculated in individual mouse liver with 4/5 mice in each group.

Standard deviation (±SD) was calculated among each group. Results were indicated in standard error (±SE).

In cell culture experiments, relative mRNA expression was calculated with respect to control or vehicle treated cells, which was mentioned in each figure. Each experiment was repeated at least three times and standard deviation was calculated.

Results were indicated in standard error (±SE).

2.4. Cell culture and Treatment

Human hepatoblastoma cell line HepG2 (American Type Culture Collection,

Manassas, VA) was cultured in high glucose (22.5mM) Hyclone media (Thermo scientific, Fischer) with 10% FBS (Atlanta biologic ) and 1% antibiotic in the incubator at 37°C and 5% CO2 incubator. Medium was changed 2-3 times a week. When the cells reached confluence, 0.5 % trypsin in EDTA was added to detach the cells. Briefly, the media was aspirated from the cells and 4 ml of trypsin solution was added to cover the bottom of the T 75 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 and cultured in to 6 well plates or 24 well plates.

To freeze, the cells were detached according to above procedure and centrifuged.

Cell pellet was re-suspended in 50% FBS and 10% DMSO. Before treatment, the

following stock solutions were prepared: glucagon (100 µg/ml) in water, 10 mM 8-Br- cAMP (DMSO), 100 mM PKA inhibitor (H89) and 10 mM Cycloheximide (CHX) dissolved in DMSO. Day before the experiment, cells were cultured in 6 wells plate and treated with 100 ng/ml glucagon, 10 µM 8-Br-cAMP, 10 µM H89, 150 µg/ml CHX and

DMSO as a vehicle as indicated in each experiment.

2.4.1. Human primary hepatocytes culture

Primary human hepatocytes (donors # 1839, 1859 and 1858) were obtained from the Liver Tissue and Cells Distribution System of National Institutes of Health

(University of Pittsburgh, Pittsburgh, PA). Cells were maintained in hepatocyte maintenance medium as described previously [85].

2.5. Plasmids and Cloning

2.5.1. Reporters

Human CYP8B1 luciferase reporter 5’end deletion plasmids pGL3 luc (-

3.5/+300), pGL3 luc (-514/+300), pGL3 luc (-297/+300), pGL3 luc (-164/+300) and 3’ end deletions at pGL3 luc (-514/+248), pGL3 luc (-514/+220) and pGL3 luc (-514/+200) were constructed previously. [186].

In order to clone mouse cyp8b1 promoter, genomic DNA was isolated from wild type C57BL/6J mouse liver using method as follows. First, liver (0.2 µg) was homogenized in TE buffer and incubated at 65o C with proteanse K (2 mg/ml) overnight.

Next morning, DNA was isolated after phenol chloroform extraction followed by ethanol precipitation. Precipitated DNA was reconstituted in 50 µl TE buffer. This genomic DNA was used to amplify 600 bp mouse cyp8b1 (-580/+30) promoter using PCR. DNA was

extracted using gel extraction kit (Qiagen.Inc., Valencia, CA) and was ligated into T- vector (Promega Scientific, Madison, WI) and transformed in bacterial strain DH5α.

Positive colonies were selected using X-gal (Fischer Thermo Scientific.). Positive colonies were indicated white, while negative colonies were purple. Positive colonies were amplified using LB ampicillin media followed by plasmid extraction. This plasmid was digested with restriction enzymes MluI and XhoI (Promega Scientific) extracted from the gel. Similarly, pGL-3 basic vector (promega Scientific) was digested using the same restriction enzymes followed by ligation. Positive clones were screened and plasmid was designated as 600 (-580/+30). Sequence was confirmed by the DNA sequencing (Roswell

Park sequencing facility). Deletion construct, which has 5’ 100bp deletion of mouse

CYP8B1 reporter, was made according to the same protocol.

2.5.2. Site-directed Mutagenesis

Site directed mutagenesis was performed to mutate RORα response element on human CYP8B1 promoter using Site Directed Mutagenesis Kit—Transformer

(Clonetech, laboratories Mountain View, CA) according to the manufacturer’s protocol and cloning method explained above. Sequence was confirmed by the DNA sequencing

(Roswell Park sequencing facility).

2.5.3. Expression Vectors

Human RORα expression plasmid was provided by Dr. Wen Xie (University Of

Pittsburg). Gal4-CBP was a kind gift from Dr. Carolyn Smith (Baylor College of

Medicine Huston). PGC1α, Gal4-PGC1α, Gal4-SRC1, SRC1 plasmids were cloned as reported previously [84].

2.5.4. Preparation of Plasmid DNA

For all the plasmid DNA preparations, LB broth/medium containing 60 μg/ml of ampicillin was used to select ampicillin resistant cells containing the plasmids. To obtain specific plasmid of interest , ampicillin containing LB/ broth medium was used to grow overnight cultures. Bacterial pellet was isolated and plasmid purification was carried out as follows.

2.5.5. Small Scale DNA preparation:

Mini prep plasmid isolation was carried out using a plasmid preparation kit

(Qiagen.Inc., Valencia, CA) according to the manufacturer’s protocol. First step was to re-suspend pelleted bacterial cells in 250 μl buffer P1, which contains RNase. Further, addition of 250 μl Buffer P2 and mixing the content thoroughly was an important step in order to achieve the complete lysis of bacterial cells. Buffer N3 (350 μl) was added and thoroughly mixed to ensure neutralization of basic pH to pH 4.8. This step helped to stabilize the DNA in the solution and separate it from the rest of the cell debris. Cell debris was removed after centrifugation for 10 min at 13,000 rpm (~17,900 xg) in a table- top microcentrifuge. In the next step, Qiagen mini columns were used to achieve DNA extraction from the solution. Those mini columns were placed on a microcentrifuge tubes and content of centrifugation was loaded on the column. Further, these columns were centrifuged and DNA was retained on a column resin bed. In the flowing step, DNA was washed using ethanol based PE buffer and eluted using 50 µl of PE buffer. DNA concentration is measured. All plasmids were purified according to this method.

2.5.6. Ligation

Ligation was performed using enzyme ligase and a buffer provided (Promega scientific Madison, WI) in 10 µl reaction mixture with a ratio of plasmid to insert 1: 3 at

4oC. Restriction enzymes KpnI, HindIII and BamH1 (Promega scientific) were used for digestion of plasmids. In 20 µl reaction mixture plasmids were incubated at 37oC for 2-3 h with restriction enzymes. For adenovirus cloning, PacI and PmeI (New England

Biolabs Ipswich, MA) restriction enzymes were used in 20-50 µl reaction mixture at

37oC.

2.5.7. Preparation of Competent Cells

The competent DH5α cells were prepared by calcium chloride method. A single colony (2-3 mm in diameter) was picked from a plate freshly grown for 16-20h at 37oC and transferred into 5 ml of LB broth and incubated overnight. Further, 100 μl of the overnight culture was then inoculated into 100 ml medium and incubated for 3h at 37o C 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 optical density at 600 nm wavelength of culture was determined every 1h 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

o 4 C. Each pellet was re-suspended in 10 ml of ice-cold 0.1 M CaCl2 and stored on ice.

The cells were recovered by centrifugation at 4000 rpm for 10 min at 4oC. The fluid was decanted from the cell pellets. Glycerol was added quickly to 30% of the total volume and suspended into chilled, sterile microfuge tubes. The competent cells were

immediately snap-freezed by immersing the tightly capped tubes on dry ice. The tubes could be stored at -80oC until needed. For most transformations purposes, 50 μl aliquots of the competent cells suspension were used.

2.5.8. 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, appropriate amount of the plasmid DNA (10-100 ng) was added to the competent DH5α cells and the mixture was kept on ice for 1h. Following that, the mixture was heat shocked at 42oC for

2 min and then cooled on ice for 10 min. Eight hundred ml of Luria Broth (LB) was then added to the mixture and pipette up and down, and the resulting mixture was incubated at

37o C for 1h. After that, the mixture was centrifuged at 4000 rpm for 2 min and 800 μl of the supernatant was removed and discarded. The cell pellets were re-suspended in the

200μl fresh medium.

2.6. Cloning of Adenovirus

2.6.1. Principle

Recombinant adenovirus is an important tool for gene delivery. There are several advantages of adenovirus mediated gene delivery; adenoviruses are capable of infecting a broad range of cell types and infection is not dependent on active host cell division. The most commonly used adenoviral vector is human adenovirus serotype 5. In this adenovirus, replication is made defective by the deletion of the E1 and E3 genes. The E1 gene is essential for the assembly of infectious virus particles. Similarly, E3 gene encodes proteins involved in evading host immunity, which is not essential. These deletions

render the virus incapable of replicating itself, but they also create space for insertion of up to 7.5 kb of foreign DNA. After all these deletions, virus still can infect the host, but is easily removed by the host immune system. Therefore, gene delivery using this adenovirus is not stable transformation.

Two methods have traditionally been used to generate recombinant adenoviruses.

The first involves direct ligation of the gene of interest into the adenoviral genome.

Because the adenovirus genome is large (36 kb) and contains few useful restriction sites, this method is technically very challenging. The second, and more commonly used method, involves cloning the gene of interest into a shuttle vector and transferring the gene into the adenovirus genome by means of homologous recombination. The isolation of recombinant adenovirus by this method involves performing multiple plaque isolations and is extremely laborious and time consuming. This method is slightly modified such that homologous recombination event takes place between co-transformed adenoviral backbone plasmid vector (pAdEasy-1) and a shuttle vector carrying the gene of interest.

All the recombination events take place in E. coli. Later assembly of adenovirus takes place in vivo by an adenovirus packaging cell line AD-293.

2.6.2. Cloning of RORα in pShuttle-CMV vector

All the vectors and competent cells used for adenovirus cloning were purchased from GE, Life Science (Piscataway, NJ). The pShuttle-CMV vector contains a multiple cloning site sandwiched between the CMV promoter and the SV40 polyadenylation signal and is suitable for insertion of a large cDNA (up to 6.6 kb). There are two distinct regions indicated as arms, which are stretches of sequence homology with pAdEasy-1

where the homologous recombination occurs. The R-ITR and L-ITR regions are short inverted terminal repeats (Left and Right), which have a role in replication of the viral

DNA. It has a PacI restriction enzyme site, which is important to test the integrity of homologous recombination.

RORα cDNA was digested with KpnI and BamH1 and DNA fragments were resolved on 10% agarose gel. Exactly 1.5Kb band was isolated from gel and ligated into pGEM-3z vector (Promega Scientific). PCR amplification of RORα was avoided in this method. Further, RORα-pGEM-3z plasmid was subjected to restriction digestion with

Kpn1, HindIII, and 1.5 Kb DNA fragment was extracted from the gel. This fragment was ligated into KpnI and HindIII digested pShuttle-CMV vector overnight. Next day, DH5α bacterial cells were transformed using ligated mixture and plated on 60 µg/ml ampicillin

LB plates. Positive colonies were screened using mini-preparation and restriction digestion method. This plasmid integrity was tested by DNA sequencing.

Approximately 2 μg purified RORα-pshuttle-CMV vector was linearized using

Pme I. This step was important for homologous recombination. It was also ensured that

RORα did not contain PmeI or PacI restriction enzyme sites. Linearized plasmid was resolved on 10% agarose gel. A 9 kb fragment was purified from the gel and eluted in water instead of TE buffer.

2.6.3. Transforming the BJ5183 bacterial cells by electroporation

In this method, the BJ5183 cells (GE Life Science) were co-transformed with the linearized RORα-pshuttle-CMV vector and the pAdEasy-1 vector. A recombination event that takes place in the bacterial cells results in the production of recombinant plasmid

DNA. It is very important that this co-transformation takes place in BJ5183 cells because only the BJ5183 cells have the cellular components necessary to carry out recombination.

Two aliquots of 40 μl BJ5183 competent cells were taken from –80°C storage and thawed on ice. In one tube, 1 μl (1 μg) of linearized shuttle vector and 1 μl of pAdEasy-1 supercoiled vector (100 ng/μl) were added, while in other tube 1 μl of linearized shuttle vector was added for control. This mixture was mixed together on ice. Electroporator

(Bio-Rad, Hercules, CA.) was set on 200 Ω, 2.5 kV and 25 μF current. This high voltage and current allowed entering the DNA more efficiently than heat shock method in bacterial cells. Right after the electroporation, cells were removed immediately and super optimal broth (SOB) media was added and cells were kept at 37oC shaking for 1-2h.

These cells were cultured on 60 µg/ml kanamycin agar plate overnight.

2.6.4. Screening Colonies for positive Recombination

Next day, transformation plates were examined. The plates containing the transformation of linearized shuttle vector alone had uniform-sized colonies arising from uncut/re-circularized vector, while a shuttle vector and pAdEasy-1 vector transformed plate had small circular colonies. These colonies may signify recombination event was occurred successfully. All colonies were grown in kanamycin LB agar media overnight for isolation of DNA the next morning.

The purified DNA was subjected to the PacI digestion. Restriction digestion of recombinant adenovirus DNA with PacI should yield a large fragment of ~30 kb, and a smaller fragment of 4.5 kb ( recombination took place at the origins of replication). Uncut recombinants will give a large smear at the top of the gel very close to the wells (often

have a smaller band that runs just below 23 kb). The positive clone was identified and the same culture was grown in 250 ml. The DNA was isolated from 250 ml culture to follow the further step.

Figure 7. Confirmation of RORα insertion in adenovirus genome

DNA fragments were resolved on 0.8% agarose gel after viral DNA extraction (5 µg) from bacterial colonies and digested with or without restriction enzyme PacI (2 µl) digestion and buffer 1(5 µl) provided with the enzyme in 50 µl reaction volume for 3h.

DNA ladder (M) was on the right hand side and designated as M. PacI digest at 4.5Kb indicated positive recombination and insertion of RORα open reading sequence in the viral genome.

2.6.5. Preparation of primary adenoviral stock

Viral recombinant DNA obtained from the above protocol was digested with PacI and DNA fragments were resolved on 10% agarose gel. Linearized DNA was gel purified and kept aside for transfection in HEK-293A cells. HEK-293A cells were modified with

E3 protein important for the adenovirus packing. These cells were maintained in high glucose HYCLON media (Thermo scientific, Fisher) with 10% FBS (Atlanta Biolabs,

CA.) and 1% penicillin streptomycin (Fisher, Thermo scientific), cultured in 100 mm plate (BD, Falcon, VA.). Plasmid was transfected using using X-treme GENE HP DNA transfection reagent (Roshe Applied Science, Indianapolis, IN.) according to the manufacturer’s protocol. Three µg of linearized DNA was mixed with 9 µl of the above mentioned transfection reagent in 100 µl of optimum media (does not have glucose, FBS/ pen-strep) and incubated at room temperature for 30 min. This DNA mixture was added to the 5ml of normal media for 24 h. It takes 10 days to complete packaging of the virus.

Therefore, it is important to check cells every day for cell viability. After almost 10 days cells were collected for virus purification. Cell pellet was frozen in the mixture of dry ice and ethanol where temperature was close to -78oC in the special bath and thawed at 37oC.

This procedure was repeated twice. This freezing and thawing allows rupturing of the cells, which releases the virus. This media was a primary viral stock and stored frozen at -

80oC.

2.6.6. Amplification and purification of adenovirus

In this step, HEK293A cells were cultured in tissue culture plates and grown to

60-70% confluent. The virus extracted from the above method was added to the cells and incubated for 5-7 days. This step was important to amplify the virus. After 5-7 days, 70-

90% cells were floating in the media. This was an end point of the amplification.

Virus was purified using cesium chloride (CsCl) density gradient. CsCl density gradient was performed after dissolving 1.25 mg/ml and 1.35 mg/ml CsCl in 10 mM Tris

–HCl (PH 8.1). In 15 ml ultra-centrifuge tube, first 3.5 ml of 1.25 mg/ml CsCl solution was carefully overlaid with 2 ml of 1.35 mg/ml CsCl solution using glass pipette. The interface between two layers was very clearly visible. On the top of this CsCl layers, 5 ml of cell extract was added, which was prepared by the method mentioned above. The tubes were shifted in the swinging bucket rotor. This rotor was replaced carefully into ultra- centrifuge machine and run at 26,000 rpm for 2 h at 4oC. After centrifugation, the tube was removed carefully and the white turbid layer was visible between two CsCl density gradients. This turbid layer was taken out using syringe, which was approximately 1 ml.

In the next step, CsCl was removed from the virus using desalting column (GE Life

Science, PA.).

2.6.7. Adenovirus titer measurement

Titration of adenoviral stock was important for maintaining consistency and correct level of expression. The titer of the adenovirus was performed using Adeno-X rapid titer kit (Clonetech Laboratories.inc. Foster city, CA.) This method was based on primary anti hexon antibody, which detects the hexon protein of virus, and secondary

antibody with horseradish peroxidase conjugated. This secondary antibody developed black color spots upon developing.

In the first step, HEK293A cells were cultured in collagen treated 12 well tissue culture plates. Further, the virus obtained from above method was diluted until final concentration of virus is 10-2, 10-3, 10-4, 10-5 and 10-6 ml using successive dilution method. Hundred µl of diluted virus was added to each well, which contains 1ml of growth media. 10-6 and 10-7 dilution were performed in duplicates. After 48h, cells were treated with chilled methanol and incubated for 10 min in -20oC. This step was important to fix the cells. Further, the cells were washed with PBS containing 1% BSA three times.

Cells were incubated in PBS containing 1% BSA with anti-Hexon antibody for 1h at

37oC. Further, this primary antibody was removed and cells were washed using PBS with

1% BSA for 3 times. In the next step, secondary anti mouse antibody horseradish peroxidase (HRP) conjugated was added in 1:500 dilution and incubated at 37oC for 1h.

Antibody was removed and cells were washed with PBS containing 1% BSA for 3 times.

To develop antibody reaction, 1X working 3-3’-diaminobenzidine (DAB) solution was prepared after dilution with 1X peroxidase buffer. This 1X DAB buffer was added in each well till brown coloration was observed. DAB buffer was removed and the cells were observed. Each well was observed using 20X lenses under microscope. Twenty fields were counted and mean number of black brown spots was calculated in 10-6 diluted well. Virus titer was calculated in infectious units/ml (ifu/ml) using the equation mentioned in the manufacturer’s protocol. This virus is very pure and further used for in- vitro experiments to test the RORα expression level.

2.7. Chromatin immunoprecipitation assay (ChIP assay)

Chromatin immune-precipitation assay was performed to test the binding of transcription factors to gene promoter.

2.7.1. Isolation of nuclei and cross linking

First, mouse liver 0.2 mg (n=4) was weighed and homogenized into nuclei isolation buffer (15 mM NaCl, 5 mM MgCl2, 0.1 mM EGTA, 15 mM Tris –Hcl (pH 7.5),

0.1 mM PMSF, 3.6 ng/ml aprotinin) using glass homogenizer. The nuclei were isolated from HepG2 cells. In the next step, 37% formaldehyde (270 µl) was used for cross linking at vigorous shaking for 15 min. and 1.5 M glycine (1 ml) was added to stop cross- liking. The mixture was centrifuged at 4000 rpm for 10 min. The supernatant was discarded and same steps were followed twice. In the next step, 10 ml of nuclei isolation buffer was added and centrifuged 8000 rpm for 30 min. The nuclei were re-suspended in

5 ml in PBS. These nuclei were stained using bromo-thymol blue reagent and observed under the microscope at 20 X resolution. Further, 1 ml of PBS suspension was used for the next step.

2.7.2. Sonication

One ml mixture from the above step was centrifuged and supernatant was discarded. Then, 500 µl RIPA buffer was added in the pellet. This mixture was further sonicated at 80 duty cycle, 1 sec. pulse each time 7 pulse/sec and repeated 8 times on ice.

Mixture was centrifuged at 13000 rpm and supernatant was diluted (1:200) using ChIP dilution buffer provided by Upstate biological inc. This diluted solution was pre-cleaned

using 80 µl salmon sperm coated agarose beads and divided into five 1.5 ml eppendorf tubes.

2.7.3. Washing and recovery of DNA

ChIP assay was performed with RORα (BioLegend. San Diego, CA.), CBP,

HNF4α (SantaCruz, Biotechnology, CA.) or acetylated histone (Cell Signaling

Technology Boston, MA), and non-immune control IgG (Millipore Billerica, MA.,) was used as control. The samples were incubated with antibodies overnight, shaking at 4oC.

Salmon sperm coated agarose beads were added to pull down antibody for 1h. Beads were washed and supernatant was discarded. Further, beads were washed using 1 ml of low salt buffer, high salt buffer, lithium chloride and twice with 1 ml TE buffer. After all the washings, DNA was eluted using freshly prepared elution buffer (0.1 M NaHCO3, 1%

SDS) at room temperature. After elution, 5 M NaCl (20 µl) was added to the mixture and heated at 65o C for 4 h. This step is important for uncoupling of protein DNA complex. In the next step, denaturation of protein was carried out using proteanase K (2 mg/ml) digestion at 45o C for 2h and DNA was recovered using 500 µl of Phenol-chloroform extraction and ethanol precipitation and dissolved in nuclease free water for RTPCR.

2.8. Western blot analysis

Approximately 200 mg of mouse livers were subjected to lysis using 500 µl RIPA buffer (Cell Signaling Technology Boston, MA). Livers were homogenized in RIPA buffer with protease inhibitor and phosphatase inhibitor sodium ortho vanadate (2 µM

Na3VO4) and incubated on ice for 1h to ensure complete lysis. Further, samples were sonicated on 50 duty cycles with 1 second pulse. This step is important to ensure

complete nuclear protein isolation. In the next step, the samples were centrifuged at

13,000 rpm for 10 min at 4oC. The pellet was discarded and supernatant was used for protein concentration measurement. Protein concentration was determined using BSA protein concentration kit (Bio Rad. inc).

Exactly 40 or 50 µg of the protein was resolved on the polyacrylamide gel (10%).

Electrophoresis was performed at 200 volt in SDS running buffer. Gel was transferred on nitrocellulose membrane on semidry transfer instrument (BioRad Laboratories, Inc.).

Further, membrane (nitrocellulose) was blocked in 5% non-fat dry milk overnight or for

2h. Antibodies were incubated in 5% milk with or with or without RORα (BioLegend.

Inc.), PKA (Millipore, Billerica, MA.), Total PKA (Cell Signaling Technology Boston,

MA) and Histone (Millipore, Billerica, MA) antibodies as mentioned in each experiment.

2.8. Measurement of bile acid pool size

2.8.1. Principle

Bile acid pool size is the total amount of bile acids in liver gallbladder and intestine. In this method, total bile acids were extracted and measured from tissues using bile acid assay kit (Diazyme Laboratories, CA.) In the presence of thio-NAD, 3α- hydroxyl steroid dehydrogenase (3α-HSD) converts bile acids to 3-ketosteroids and thio-

NADH. This reaction is reversible in presence of thio-NADH. The enzyme 3-αHSD can convert 3-Keto steroids in to bile acids. In presence of thio-NAD and thio-NADH, the enzyme cycling occurs efficiently and rate of formation of Thio-NADH is determined by measuring specific change of absorbance at 405nm.

2.8.2. Isolation of bile acids from liver, intestine and gall bladder

Liver isolated from AD-null (n=4) and Ad-RORα (n=4) infected mice were weighed (100 mg) and homogenized in 95% ethanol and extract was incubated at 60oC for 3h. Next, the samples were centrifuged 8000 rpm for 5 min. The supernatant have all the bile acids, which could be transferred in a fresh tube and stored at -80oC. Five µl of the sample was used directly for bile acid concentration measurement.

Next step was isolation of bile acids from intestine, which has the most amounts of bile acids. Whole intestine was carefully transferred in 15 ml tube. Three ml of 95% ethanol was added and homogenized immediately. Further, samples were incubated at

60o C in water bath for 3h. In the next step, samples were centrifuged at 8000 rpm for 10 min. Supernatant was collected in fresh tubes. Further, pellet was re-suspended in 80% ethanol and incubated at 60oC for 3h followed by centrifugation at 8000 rpm. All the supernatants were combined together. Further, Pellet was re-suspended in chloroform and methanol (2:1) and incubated at room temperature followed by centrifugation at 8000 rpm. Precipitated debris was discarded and all supernatants were combined. This sample was diluted 200 times in ethanol and 5 µl of sample was used for bile acid detection.

Similarly, gallbladder bile acids were isolated using 75 % ethanol followed by sonication in water bath. The sample was diluted 500 times in ethanol and only 5 µl sample were used for bile acid determination.

2.8.3. Measurement of bile acid concentration

First different concentrations of standard bile acids were prepared and 5µl of each standard and sample from above method was aliquoted in 96 wells plate. To each well,

270 µl of R1 buffer (Thio-NAD) provided with the kit was added and incubated at 37oC

for 5min. The absorbance of the plate was measured using Synergy-4 (BioTek,

Instruments). Absorbance was measured right after 2 min and procedure was repeated twice. Buffer R2 (90 µl) with 3α-HSD was added and absorbance was measured right away. Further, absorbance was measured every min for 5 min. Absorbance after addition of R1 was subtracted from absorbance after R2 and standard curve was plotted absorbance (R2-R1) vs. concentration. The straight line was obtained and concentration of bile acids from the sample (unknown) was calculated with respect to standard graph.

2.8.4. Isolation and measurement of liver triglycerides and cholesterol

Liver tissues (0.2 mg) were homogenized, and lipids were extracted after incubation of a mixture of chloroform and methanol (2:1) overnight. Further, this mixture was dried, and dissolved in 5 % Triton X-100 in isopropanol. Cholesterol and TG were determined with cholesterol assay kits (BioVision, San Diego, CA) and a TG-SL assay kit (Genzyme Diagnostic, Framingham, MA) according to the manufacturer’s protocol.

2.9. Electrophoretic mobility shift assay (EMSA assay)

An electrophoretic mobility shift assay (EMSA) is also referred as a gel shift assay. It determines protein–DNA interaction. Different molecules move through the gel is determined by their size and charge, and to a lesser extent, their shape. This technique detects protein-DNA binding, which is observed as upward shift on the gel.

2.9.1. Annealing the probes

The double stranded oligos were designed with 5’and 3’ end overhangs (two G at

5’ and 3’end, respectively) spanning the region between hCYP8B1 and human APOCIII promoter. These oligos were diluted in TE buffer to (1000 pM/µl). In the next step, 1 µl

of each oligo was incubated in 10 µl of 20 X SSC solution and 88 µl of water. Further this mixture was heated at 95-100oC and cooled slowly to the room temperature. This step allows the complementary oligo to anneal and form double stranded probes.

2.9.2. Radiolabelling probes

P32 dCTP was purchased and used for radiolabelling of probes. Above ds-oligos

(9 µl) were mixed with 10X DNA polymerase buffer (2µl), 1 µl Klenow polymerase, 2 µl

P32 dCTP and 6 µl of water. This reaction mixture was incubated at 37oC for 30 min. and diluted with 180 µl of TE. Further, probes were purified using columns (G-50 microcolumns) purchased from GE Health Care and activity was measured using 1µl of the above mixture. The probes were diluted using TE buffer to dilute the activity to

50,000 cpm/µl. The probes were stored at 4oC.

2.9.3. Binding reaction

In-vitro transcription and translation kit (Promega, Scientific.) was used to synthesize protein for EMSA assay. The kit contains aliquots of rabbit reticulocyte lysate

(TNT) transcription and translation system. Forty µl of TNT lysate was incubated with 2

µg of cDNA of RORα for 5h. The protein translation was tested using western blot analysis. The protein aliquot (10 µl) made using this method was incubated in 4X binding buffer and 1 µl dIdC polymer was added to avoid nonspecific binding. Lastly, P32 labeled

CYP8B1 and APOCIII probes (1 µl) were added in total reaction volume 20 µl and incubated at room temperature for 1h or 5h.

2.9.4. Running EMSA gel

Non-denaturing polyacrylamide (10%) gel was used for electrophoresis. All 20 µl of the above mixture was resolved on the gel for about 2h. This gel dried using pump for

30 min. Gel was completely covered with plastic wrap to avoid radioactive contamination. Further, the gel was exposed to phosphore imaging cassette overnight and developed on phosphore imager (Typhoon, imager system).

2.10. Transient transfection assay

Cells were cultured in Hyclone media supplemented with 10% fetal bovine serum in 24-well plates. Confluent cultures of HepG2 cells or sub-confluent cells were grown in

24-wells tissue culture plates. Cells were transfected with different reporter plasmids. All the plasmid transfections were performed using a lipid based transfection reagent called

Tfx20 / Tfx™ (Promega Scientific Madison, WI). The Tfx™ Reagents is a mixture of synthetic, cationic lipid molecules and L-dioleoyl phosphatidylethanolamine (DOPE).

Upon hydration with water, these lipid molecules form multilamellar vesicles associate with nucleic acids. The amount of positive charges contributed by the cationic lipid component of the Tfx™ Reagent must exceed the amount of negative charges contributed by the phosphates on the DNA backbone, resulting in net positive charges 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 were used in cultured cells for all the experiments. The various plasmids were mixed with 500 μl of serum free medium and the correct amount of Tfx™ reagent was added to it and this mixture was incubated at room temperature for 15 min. The medium was removed completely by aspirating and the plasmid DNA-media-Tfx™ mixture was added

drop wise to the cells. The cells were then returned to the 37o C incubator for 1h.

Following that the cells were overlaid with 1 ml of serum containing media and returned to the 37o C incubator. Total DNA concentration was kept constant in each experiment using addition of pcDNA3 empty plasmid (Promega Scientific Madison, WI). An expression plasmid pCMVβ (Clontech, Palo Alto, CA) containing E.coli β-galactosidase gene was co-transfected to measure transfection efficiency.

Various luciferase reporter constructs (mentioned in the each figure) and expression plasmids, pCMV β-galactosidase plasmid (Clontech. Laboratories, Palo Alto,

CA) with 1/10 ratio along with or without pcDNA3 empty vector (negative control) were transfected in each well (according to above protocol). After 24h incubation in growth media, cells were supplanted with serum free media for 10h and treated with above mentioned chemicals in each well for another 12h. After 48h, cells were washed with

PBS and lysed using 100 µl of lysis buffer (Promega Scientific Madison, WI). After the lysis was performed, 20 µl of lysate were added in 96 well plate and activity were measured after addition of 100 µl of LARII reagent (Promega Scientific). In 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 micro plate, 20 μl of cell lysate was prepared as described earlier was added to each well. In each well, 40 μl of the above assay buffer was added and plate was incubated at

37oC till a yellowish color developed. Absorbance at 405 nm was measured for each well.

Luciferase activity was normalized by β-gal activity. Relative luciferase activity was expressed relative luciferase units (RLU/β-gal units) and graphs were plotted with

relative luciferase activity on X axis and a variable Y axis. Each transfection was performed in triplicates. Each experiment was repeated atleast three times. Results were expressed as mean ± S.E.

2.11. siRNA knock-down assay

siRNA specific for RORα and scrambled nonspecific siRNA were purchased

(Fisher Scientific) and dissolved in RNase free water to get 50 µM stock solution.

Further, RNA concentration was measured. Exactly 180 ng and 300 ng concentration of siRNA were transfected using HiPerFect siRNA transfection reagent (Qiagen.Inc.,

Valencia, CA). HepG2 cells were maintained in Hyclone media without serum or penicillin/streptomycin in 6 wells plate. Further, 20 µl of transfection reagent was mixed with siRNA and above mentioned media in a micro centrifuge tube for 30 min. This allows the formation of siRNA and lipid complex. Further, this mixture was added to the

6 wells plate. After 1h transfection reagent incubation, cells were supplemented with growth media for 4h or 8h. Further, RNA and protein were isolated to check the siRNA effect using above mentioned protocol.

2.12. Immunoprecipitation

First, HepG2 cells were grown in 100 mm tissue culture dish using the method mentioned above. Further, the cells were treated with 8-Br-cAMP, H89 (PKA inhibitor), wortmannin (PI3K inhibitor) for 6hr. Cells were washed and lysed using lysis buffer (20 mM, Tris HCl pH 7.5, 100 mM NaCl, 0.5% NP-40, 0.5 mM EDTA, 0.5 mM PMSF, 0.5 mM protease inhibitor and 0.1% Na3VO4 (phosphatase inhibitor). In the next step, cells were sonicated briefly. The sonication step allows lysis of nuclei. Further, cells were

centrifuged 13,000 rpm for 10min. Supernatant were transferred in a fresh tube. The protein concentrations of each sample were measured. Exactly 500µg of protein were transferred in a fresh tube and RORα antibody (10 µg) was added and incubated with end to end rotation overnight. Next day, 50µl protein agarose beads were added with end to end rotation for 1h. Beads were isolated using centrifugation at 3000 rpm. Beads were washed three times using lysis buffer and LiCl (10 mM, 20 mM Tris HCl pH 7.5) buffer.

Separated agarose beads were resolved using 10% acryl amide gel. Western blot were performed using p-Serine specific anibody as mentioned above.

2.13. Microsomes isolation

CYP8B1 protein is located in the microsome. To assay CYP8B1 protein expression, microsomes were isolated from the liver.

Adenovirus infected (Ad-RORα and Ad-null) mice liver (100mg) n=4 were used for microsome isolation. Around 400 mg of all livers were homogenized using 4 ml ice cold homogenization buffer (0.05M Tris Hcl pH7.2 and 0.15M KCl) in glass homogenizer. Further, cells were centrifuged at 10,000 rpm for 10 min at 4oC. Pellet was re-suspended in fresh microsome isolation buffer. In the next step, the suspension was centrifuged 100,000 xg in the ultra-centrifuge. Further, pellet was re-suspended in RIPA buffer followed by lysis. Microsome proteins were resolved on 10% SDS gel. Western blot was performed using CYP8B1 antibody (Santa cruz, CA).

2.14. Mammalian two hybrid assay

Mammalian two hybrid assays is used to study protein-protein interactions in the cells. DNA binding domain and transcription activation domain were cloned in pBIND

vector (Promega Scientific, Madison, WI ) contains GAL4 DNA binding domain while, pACT vector (Promega Scientific, Madison, WI ) contains the herpes simplex virus VP16 activation domain.. Mammalian two hydrid assay was used to detect interaction between

RORα and different co-activators.

VP16-RORα plasmid was cloned in pACT plasmid. Human RORα was cloned in KpnI and BamHI restriction enzyme sites using above mentioned protocol. Similarly GAL4-

RORα fusion plasmid was cloned in KpnI and BamHI restriction enzyme sites for mammalian two hybrid assay. Transient transfection of pG5luc, Beta-Gal vector with or without RORα-VP16, Gal4-PGC1α, Gal4-SRC1, Gal4-CBP were performed using the method mentioned above. After 48h of transfection, cells were lysed and luciferase assay was performed. Data were normalized using beta-gal activity assay using method mentioned previously.

2.15. Statistical analysis

Results were expressed as means plus standard errors (±SE) or standard deviation

(±SD) as indicated. Each experiment was repeated at least three times and statistical analysis was performed with the Student’s t test. P < 0.05 was considered statistically significant.

CHAPTER III

RESULTS

3.1 Glucagon and cAMP increases CYP8B1 mRNA expression.

Bile acid synthesis is regulated by fasting and feeding cycle. Glucagon is an important hormone in the regulation of fasting process. Therefore, to study the effect of glucagon on bile acid metabolism and cyp8b1 mRNA expression, human primary hepatocytes were treated with glucagon (100 ng/ml) for 2h, 8h and 18h. Isolation of mRNA followed by real-time PCR was performed. Fig.8 shows that glucagon treatment increased CYP8B1 mRNA expression by 2-3 fold compared to untreated cells. On the other hand, CYP7A1 mRNA was suppressed upon glucagon treatment, which was consistent with a previous report. [187]. There was no significant effect of glucagon on

CYP27A1 and CYP7B1 mRNA levels. Results also suggest that glucagon treatment specifically increased CYP8B1 gene expression.

Glucagon increases cAMP production therefore HepG2 cells were treated with cAMP analog, 8-Br-cAMP (10 µM). Since, 8-Br-cAMP is more stable than glucagon or cAMP and can efficiently penetrate into cells; 8-Br-cAMP was used in the cell culture experiments. Isolation of mRNA and real time PCR was carried out. Fig. 9 shows that 8-

Br-cAMP treatment increased CYP8B1 mRNA, while CYP7A1 mRNA was suppressed.

Figure 8. Glucagon increases cyp8b1 mRNA, but suppresses cyp7a1 mRNA in human primary hepatocytes. Human primary hepatocytes (batch numbers 1839, 1958 and 1959) were treated with glucagon (100 nM) at different time points as indicated. Relative mRNA expression of CYP8B1, CYP7A1, CYP7B1, CYP27A1 were measured by quantitative real-time PCR. Relative mRNA expression was calculated with respect to vehicle treated cells. Results were expressed as mean ± S.E.; n=3; an “*” indicates statistical significance vs. vehicle treated cells, p< 0.05

Figure 9. 8-Br-cAMP increases cyp8b1 mRNA, but suppresses cyp7a1 mRNA in

HepG2 cells. HepG2 cells were treated with 8-Br-cAMP (10 µM) at different time points as indicated. Relative mRNA expression of CYP8B1, CYP7A1, CYP7B1 and CYP27A1 were measured by real-time PCR. Relative mRNA expression was calculated with respect to vehicle (DMSO) treated cells. Results were expressed as mean ± S.E.; n=3; an “*” “#” indicates statistical significance vs control (untreated cells), p< 0.05.

3.2. Fasting induces cyp8b1 mRNA and protein expression.

To study the effect of fasting and refeeding on genes involved in bile acid synthesis in vivo, wild type mice (C57Bl/6J) (12 -16 weeks old) were fasted for 16h from

8 am to 12 pm, and fed for 6h. Liver mRNA was isolated and real time PCR was performed. Fig. 10A shows that cyp8b1 mRNA was elevated upon fasting. To understand the change in cyp8b1 protein level, microsomes were isolated from liver tissues of fasted and fed mice. Western blot was performed using cyp8b1 specific antibody. Fig. 10B showed an increased in cyp8b1 protein level in the liver microsome extract of fasted mice compared to the fed mice. These results indicate that fasting induces cyp8b1 mRNA and protein expression in the mouse liver.

Figure 10. Fasting induced Cyp8b1mRNA and protein expression

Male C57BL6J mice (12 weeks old) were fasted for 16 h from 8 am to 12 am and were fed with normal chow diet for 6h. (A) Liver mRNA expression was measured by real- time PCR. Results were expressed as mean ± S.E.; n=4; an “*” indicates statistical significance vs. relative mRNA expression of fed mice. (B) Western blot analysis of

CYP8B1 protein in microsome extract pooled from fasted (n=4) and fed (n=4) mouse liver (upper panel). (C) Quatification of CYP8B1 protein expression in fasted and fed mouse liver.

3.3. Fasting induces RORα protein expression, while mRNA expression remains unchanged.

We wanted to investigate if fasting and re-feeding changes RORα mRNA and protein expression. Mice were fasted for 16h and fed for 6h, and liver mRNA and protein were isolated for analysis. Fig. 11A shows that there was no change in RORα mRNA expression in fasted compared to fed mouse liver. However, RORα protein expression was significantly higher in fasted mouse liver (Fig. 11B). Further, to understand the signaling pathway activated during fasting and feeding, western blot was performed using p-PKA, and p-AKT antibodies. Fig. 11B shows that phosphorylation of PKA was increased during fasting, while phosphorylation of AKT, an insulin activated protein kinase B, was abolished upon feeding. Rev-erbα protein expression was elevated after feeding, which was consistent with previous observation that Rev-erbα was stabilized after phosphorylation by glycogen synthase kinase 3β (GSK3β), an insulin activated kinase. These data suggested that fasting and glucagon signaling induced CYP8B1 mRNA and protein expression. However, fasting induced RORα protein without increasing RORα mRNA expression.

A RORα

Figure 11. RORα protein is induced upon fasting.

Male C57BL6J mice (12 weeks old) were fasted for 16h from 8 a.m. (ZT 2) to 12 a.m.

(ZT 18) and then re-fed a regular chow diet for 6h. Liver mRNA was isolated and (A)

Quantitative real-time PCR was performed. Relative RORα mRNA expression was calculated. Results were expressed as mean ± S.E.; n=4 (B) Western blot analysis of

RORα, Rev-erbα, PKA, p-PKA, AKT, and p-AKT in total liver extracts of fasted (16 h) and fed (6 h) mice. Each lane was loaded with liver extracts from different mouse in each group. Histone 3 was used as a loading control.

3.4. Fasting alters CYP8B1 circadian rhythm and RORα protein expression.

RORα is a circadian gene and regulated by dark and light cycle. Although bile

CYP8B1 expression also shows strong circadian rhythm, factors responsible for it are still obscure. To understand if cyp8b1 mRNA is regulated during the 24 h of the day, free fed cyp8b1 and RORα mRNA expression was monitored every 4h for 24h. Relative mRNA expression is calculated with respect to ZT 0 (6.00 am) liver samples. Fig. 12A shows that cyp8b1 mRNA peaked at the beginning of the light cycle ZT 2 (8.00 am), whereas RORα mRNA peaked at ZT 18 (12.00 pm) in the dark cycle (Fig. 12B). Further, to understand if fasting changes circadian rhythm of cyp8b1 mRNA expression, mice were fasted for 12h from ZT 2 (8.00 am) to ZT 14 (8.00 pm) and re-fed at ZT 14 (8.00 pm). Mice were sacrificed every 4h. Liver mRNA was isolated and RORα and CYP8B1 mRNA expression was monitored. Fig. 12A shows that circadian expression of cyp8b1 mRNA was disrupted after fasting. There was a continuous increase in cyp8b1 mRNA due to fasting and peaked at ZT 14 (8.00 pm) while, it peaked at ZT 2 (8.00 am) in free fed mice. This data indicates that fasting induces, while feeding suppresses cyp8b1 mRNA expression. There was no significant change in RORα mRNA level after fasting and feeding compared to free fed animals. Thus, we want to assay circadian expression of

RORα protein. Fig. 13A shows that RORα protein level peaked at ZT 0 in free fed mice, while fasting significantly increased RORα protein expression, which was suppressed right after feeding. This experiment indicates that RORα protein expression but, not mRNA expression was altered by the fasting and expression of RORα protein level is correlated to the CYP8B1 mRNA level.

Figure 12. Diurnal rhythm of RORα and CYP8B1 in mouse liver.

(A) Liver mRNA expression of cyp8b1 was measured in mice sacrificed (n=4/5) at

different time points of the day (shown in the figure). Relative mRNA expression was

calculated with respect to ZT 0 (6.00 am) scarified mice. Results were expressed as mean

± S.E.; n=4/5; an “*” indicates statistical significance vs. free-fed mice sacrificed at

respective time points p< 0.05. (B) Liver mRNA expression of RORα was measured in

mice sacrificed (n=4/5) at different time points of the day as indicated in the figure.

Results were indicated as per the explanation under panel (A).

Figure 13. Diurnal rhythm of RORα protein expression

(A) Western blot analysis of RORα protein using total liver lysates isolated from mice sacrificed at different time points as indicated. Samples (three each time point) were loaded onto two separate gels run at the same time. (B) Intensity (A.U.) of RORα protein band (averaging of three samples) was normalized to histone as a loading control at each time point. Results were expressed as mean ± S.E.; n=3

3.5. Adenovirus mediated over-expression of RORα increases cyp8b1mRNA level

To study the effect of RORα on CYP8B1 expression and bile acid composition, adenovirus expressing RORα and adenovirus virus (null) were injected via tail vein.

Seven days post injection liver mRNA was isolated and real time PCR was performed.

Fig. 14A shows that Ad-RORα infusion increased RORα mRNA and protein expression

(Fig. 14B). Further, over-expression of RORα increased CYP8B1 mRNA expression.

Similarly, RORα target gene CYP7B1 mRNA was also induced upon RORα over- expression. Overall, data indicates that RORα is a strong activator of mouse CYP8B1 gene expression and could regulate cholic acid synthesis. To measure the change in

CYP8B1 protein level, liver microsomes were isolated from mice infused with Ad-RORα and Ad-null. Fig.14B clearly indicates that over-expression of RORα increased CYP8B1 protein expression in liver microsomes. Further, measurement of total bile acid pool size was performed by adding total bile acids in the liver, intestine and gallbladder in the Ad-

RORα and Ad-null infected mice. Fig 14C shows that total bile acid pool size did not change significantly in RORα over-expressed mice compared to the control mice.

Interestingly, RORα over-expression increases bile acid content in liver and intestine but, decreased in the gallbladder.

Since CYP8B1 regulates cholesterol metabolism, serum cholesterol and triglyceride levels were measured in the mice over-expressing RORα and control. Fig.14

D shows that RORα over-expression significantly elevated serum cholesterol level and

hepatic cholesterol level tends to be higher. On the other hand, there was no change in serum and hepatic triglyceride levels compared to Ad-null infused mice.

Figure 14. RORα induced cyp8b1 mRNA in mouse liver.

Mice were injected (intravenously) with adenovirus (29 ifu/mouse) expressing either empty adenovirus (Ad-null) as control or expressing RORα (Ad-RORα). (A) Liver mRNA expression of cyp8b1, cyp7b1, and RORα measured in overnight-fasted mice.

Results were expressed as mean ± S.E.; n=6; an “*” indicates statistical significance vs.

Ad-null infused mice, p< 0.05. (B) Western blot analysis of RORα protein using mouse total liver extract pooled together from four mice expressing adenovirus RORα (Ad-

RORα) and adenovirus null (Ad-null). Western blot analysis of cyp8b1 using microsomes extracted from control and RORα adenovirus expressing mice. (C) Total bile acids

extracted from liver, intestine, gallbladder and measured pool size in adenovirus expressing RORα (Ad-RORα) and adenovirus null injected mice (Ad-null). Results were expressed as mean ± S.E.; n=6; an “*” indicates statistical significance vs. bile acids measured in Ad-null infused mice, p< 0.05. (D) Total serum cholesterol, triglycerides

(TG), Liver cholesterol and triglycerides in Ad-RORα and Ad-null infused mice were measured. Results were expressed as mean ± S.E.; n=6; an “*” indicates statistical significance vs. serum cholesterol in Ad-null infused mice, p< 0.05.

3.6. Over-expression of RORα changes bile acid composition.

We analyzed the bile acid compositions in the gallbladder. In mice, about 90% of the bile acids are taurine (T)-conjugated. Table 2 shows that TCA and TDCA contents were increased from 38% and 2.9% in Ad-null to 54% and 6.2% in Ad-RORα mice, respectively. TCDCA was reduced from 4.7% in Ad-null to 2.8% in Ad-RORα mice. In mouse liver, CDCA (3α, 7α) is converted to TαMCA (3α, 6β and 7α), TβMCA (3α, 6β and 7β), tauro-hyocholic acid (THCA, 3α, 6α, and 7α) or tauro-ursodeoxycholic acid

(TUDCA, 3α, 7β) by 6-hydroxylation catalyzed by Cyp3a11 and/or epimerization.

TαMCA and TβMCA were decreased from 18.8% and 13.1% in Ad-null mice to 10.1% and 5.3% in Ad-RORα mice, respectively. It was interesting that THCA was markedly increased from 0.8% in Ad-null to 9.4% in Ad-RORα mice. The ratio of total 12α- hydroxylated bile acids (TCA, TDCA, CA, and DCA) to non-12α-hydroxylated bile acids

(the rest of bile acids) was doubled in Ad-RORα mice from Ad-null mice, as the result of increased CYP8B1 activity in Ad-RORα mice.

Table 2 Gallbaldder bile acid analysis

Bile acids (%) Ad-null Ad-RORα

12α-OH TCA 38.2 ± 1.36 53.9 ± 2.28 TDCA 2.9 ± 0.032 6.22 ± 0.34 CA 0.52 ± 0.184 0.71 ± 0.032 DCA 0.13 ± 0.033 0.27 ± 0.041 Total 41.10 ± 1.609 60.12 ± 2.693 Non-12α-OH Ad-null Ad-RORα TCDCA 4.65 ± 0.88 2.88 ± 0.137 TαMCA 18.8 ±1.89 10.13 ±2.01 TβMCA 13.1 ± 2.88 5.34 ± 2.36 TUDCA 3.4 ±0.641 2.5 ± 1.197 THCA 0.8 ± 0.274 9.36 ± 1.36 TMDCA 2.0 ± 0.063 0.4 ± 0.142 TLCA 1 ± 0.05 0.09 ± 0.253 ωMCA 5.4 ± 0.72 1.41 ± 0.33 αMCA 1.0 ± 0.0272 1.9 ± 0.3148 CDCA 0.02 ± 0.097 0.18 ± 0.105 LCA 0.8 ± 0.073 0.24 ± 0.025 Total 50.97 ± 7.595 34.416 ± 8.233

Ratio 12α OH/non 12 αOH 0.81 ± 0.21 1.76 ± 0.32 *

Gallbaldder bile acid analysis (%) of Ad-null (n=2) and Ad-RORα (n=7) injected mice

(seven days). % bile acids were calculated with respect to total bile acids measured

22284.65 (pM) in Ad-null and Ad-RORα infused mice. Abbreviations -: CA (Cholic acid), DCA (Deoxycholic acid), TCA(Tauro-cholic acid), TDCA(tauro deoxycholic acids), TCDCA (tauro-chenodeoxycholic acid), TαMCA (Tauro-α muricholic acids),

TβMCA (tauro-β-muricholic acids), TUDCA (Tauro-urodeoxycholic acids), THCA

(tauro-hyocholic acid), TMDCA (tauro-murideoxycholic acid ), TLCA (tauro-lithocholic acid). Data is represented as ± SE. An “ * “indicates statistical significance vs ratio between 12α OH/non 12 αOH Ad-null injected mice.

3.7. RORα specifically regulates cAMP mediated stimulation of CYP8B1 reporter activity

Above results indicate that RORα may directly regulate CYP8B1 gene transcription. Reporter assay was used to study the effect of RORα on cyp8b1 reporter activity. Human CYP8B1 reporter plasmid was co-transfected with transcription factors expressing plasmid in HepG2 cells. Fig. 15 shows that human CYP8B1 reporter activity was stimulated by co-transfection of RORα and RORγ. Further 8-Br-cAMP treatment increased reporter activity upon RORα transfection. On the other hand, transfection of

FoxO1, which is an insulin regulated transcription factor, decreased the CYP8B1 reporter activity. CREB (cyclic AMP response element binding protein) had no effect on

CYP8B1 reporter activity. In contrast, PPARα suppressed human CYP8B1 reporter activity. Overall, this experiment suggests that RORα or RORγ may induce CYP8B1 gene transcription.

Figure 15. RORα stimulates cyp8b1 reporter activity

HepG2 cells were co-transfected with 0.2 µg of hCYP8B1 (-514/+300), and RORα,

RORγ, FoxO1, CREB, PPARα (0.1 µg) expression plasmids and treated with 8-Br-cAMP

(10 µM) or vehicle (DMSO). Each transfection was performed in triplicates. Raw firefly luciferase activity (RLU) was normalized by β-gal activity. Results were expressed as mean ± S.D.; n=3; an “*”and “**” indicates statistical significance vs. vehicle treated

(DMSO) luciferase activity without RORα transfection and with RORα transfection respectively. p< 0.05.

3.8. RORα is required for cAMP mediated induction of CYP8B1mRNA expression.

To further confirm that RORα is involved in the stimulation of CYP8B1 gene expression upon 8-Br-cAMP treatment, RORα specific siRNA were used to knock down

RORα expression in HepG2 cells and assayed CYP8B1 mRNA expression levels. Fig. 16 shows that RORα-siRNA efficiently knocked down RORα mRNA (Fig. 16A) and protein

(Fig. 16B) levels in HepG2 cells. RORα-siRNA also efficiently knocked down CYP8B1 and CYP7B1 mRNA levels by ~70% compared to cells transfected with scrambled siRNA. Further, knock down of RORα also prevented 8-Br-cAMP mediated induction of

CYP8B1 mRNA. These data suggest that cAMP induction of CYP8B1 is RORα- dependent.

Figure 16. RORα siRNA knockdown abolished cAMP mediated induction of CYP8B1 mRNA in HepG2 cells (A) Relative mRNA expression of CYP8B1, CYP7B1 and RORα in HepG2 cells transfected with siRORα (180 ng and 300 ng) and non-target scrambled siRNA (siControl 300 ng) for 8h. Results were expressed as mean ± S.D.; n=3; an

“*”indicates statistical significance vs. siControl transfected cells. (B) Western blot analysis of RORα protein (10% acryl amide gel) using total HepG2 cell lysate transfected with RORα siRNA (siRORα 180 ng and 300 ng) and non-target scrambled siRNA (300 ng) (siControl) after 10h. (C) Relative mRNA expression of CYP8B1 in HepG2 cells transfected with siRORα (180 ng and 300 ng) and non-target scrambled siControl

(300ng) 4h followed by treatment of 8-Br-cAMP (10 µM) for 8h. Results are expressed as mean ± S.D.; n=3; an “*”indicates statistical significance vs siRORα transfected cells, p< 0.05.

3.9. RORα stimulates mouse and human CYP8B1 reporter activity.

To elucidate the role of RORα in regulation of CYP8B1 gene transcription, human

CYP8B1 (-514/+300) and mouse cyp8b1 (-580/+30) reporters were co-transfected with increasing concentrations of RORα expression plasmid in HepG2 cells. Fig. 17A and 17B show that RORα stimulated both human and mouse CYP8B1 reporter activities by up to five-fold in a dose dependent manner. A recent study indicates that RORα binds to 7α- hydroxycholesterol (7αOHC) and acts as an inverse agonist that alters RORα and co- activator interaction [126]. To study the effect of 7αOHC, cells were co-transfected with

RORα, CYP8B1 reporter plasmid followed by treatment of 7αOHC. Reporter assay results indicate that 7αOHC decreased CYP8B1 reporter activity by 20%. Therefore,

7αOHC treatment antagonized RORα mediated induction of CYP8B1 reporter activity.

Figure 17. Treatment of 7αOHC suppresses CYP8B1 reporter activity

(A) HepG2 cells were co-transfected with 0.2 µg of hCYP8B1 (-514/+300), (B) mouse cyp8b1 (-580/+30) and various concentrations of RORα plasmid (0.05 to 0.2 µg) as designated with or without treatment of 7α-hydroxycholesterol (7αOHC, 20 µM) or vehicle (DMSO) treatment. Each transfection was performed in triplicates. Raw firefly luciferase activity (RLU) was normalized by β-gal activity. Results were expressed as mean ± S.D.; n=3; an “*” indicates statistical significance vs. vehicle (DMSO) treated luciferase activity, p< 0.05

100

3.10. Treatment of 7α-hydroxycholesterol decreases CYP8B1 mRNA expression.

Further, to study the effect of oxysterols on CYP8B1 mRNA expression, HepG2 cells were treated with increasing concentrations of 25-hydroxycholesterol (25OHC) and

7αOHC. Fig. 18 shows that relative mRNA expression of CYP8B1 was suppressed significantly after 7αOHC treatment, while CYP8B1 mRNA levels remain unaltered upon 25OHC treatment. Overall, this data indicates that 7αOHC suppressed CYP8B1 mRNA level via suppressing RORα transcription activation.

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101

Figure 18. 7α-hydroxycholesterol decreases CYP8B1 mRNA expression.

CYP8B1 mRNA expression in HepG2 cells treated with different concentrations of 25- hydroxycholesterol (25OHC) and 7αOHC (as indicated) for 10h. Relative mRNA expression was calculated with respect to vehicle (DMSO 15 μM) treated cells. Results were expressed as mean ± S.E.; n=3; an “*” indicates statistical significance vs vehicle

(DMSO) treated cells p< 0.05.

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3.11. Identification of RORα response element on human CYP8B1 promoter

To locate RORα response element on human CYP8B1 promoter, reporter assay using various deletion constructs of human CYP8B1 reporter was performed [186].

Fig.19A shows that 3’ deletion of CYP8B1 reporter abolished the stimulatory effect of

RORα. This shows that the RORα response element could be located in the 3’ end (+8 to

+248 nucleotide sequence) of the promoter. According to a previous report, Rev-erbα and

RORα shared an overlapping binding site [186]. To confirm RORα response element, site directed mutagenesis was performed on previously reported Rev-erbα binding site located on human CYP8B1 promoter as shown in Fig. 19B (upper panel). To study the effect of RORα response element mutation, reporter assay was performed with or without co-transfection of RORα plasmid. Fig. 19B clearly shows that stimulatory effect of

RORα was abolished upon mutagenesis. It is well known that HNF4α stimulates human

CYP8B1 reporter activity [100]. Therefore as a control experiment, HNF4α was co- transfected with wild type and mutant human CYP8B1 reporter. Mutation of RORE did not affect the stimulatory effect of HNF4α. Similarly, co-transfection of RORα and

HNF4α together showed no synergistic effect of wild type CYP8B1 reporter plasmid.

Therefore, these data suggests that RORα regulates human CYP8B1 gene expression independent of HNF4α.

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Figure 19. RORα response element is located on human CYP8B1 promoter

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(A) HepG2 cells were co-transfected with 0.2 µg of various deletion constructs of human

CYP8B1 reporter (as indicated) and RORα (0.1 µg). Each transfection was performed in triplicates. Raw firefly luciferase activity (RLU) was normalized by β-gal activity.

Results were expressed as mean ± S.D.; n=3; an “*” indicates statistical significance vs. control (without RORα transfection), p< 0.05.(B) Reporter assay using wild type and

RORE mutant human CYP8B1 reporter (upper panel ) co-transfected with RORα (0.1µg) and HNF4α (0.1µg). An “*” and “**” indicates statistical significance vs. control

(without RORα transfection) and wild type human CYP8B1 reporter activity respectively as indicated, p< 0.05.

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3.12. RORα binds on human CYP8B1 promoter

To confirm that RORα binds to human CYP8B1 promoter, electrophoretic mobility shift assay (EMSA) was performed using P32 radiolabelled probes. Monomeric

RORα has promiscuous binding activity to AGGTCA. A canonical RORα binding site from the human APOCIII gene promoter was used as a positive control for EMSA (Fig.

20). Mutant RORα binding sites M1 (A/T region mutation) and M2 (half site mutation) were designed to test binding specificity. Fig. 20 shows that RORα binds as a monomer to the wild type CYP8B1 and APOCIII probe (lane 2 and 3), while binding is abolished in the presence of excess unlabeled APOCIII probe (lane 5). Mutation in A/T rich region

(M1) and RORα half site AGGTCA (M2) abolished RORα binding on CYP8B1 probes

(lane 3 and 4). Further RORα protein binding was confirmed by the antibody super shift assay (lane 6). Results suggest that RORα half site and A/T rich region both were required for RORα binding.

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-267 -235 APOCIII Probe gggatccgatataaaacaggtcagaaccccta +9 +39 CYP8b1WT probe ggagagctgtctgaaaaggtcagagcaaagca CYP8b1-Mutant -1(M1) +9ggagagctgtctggggaggtcagagcaaagca+39 CYP8b1-mutant -2 (M2) +9 ggagagctgtctgaaaaccccagagcaaagca+39

Figure 20. RORα binds to RORα response element on human CYP8B1 promoter.

Electrophoretic mobility shift assay (EMSA) of RORα binding to RORα response element on CYP8B1 promoter. 32P-labeled probes (specific activity=80,000 cpm) were mixed with in vitro transcription and translation lysates programmed with a human

RORα expression plasmid in binding buffer (80 mM HEPES, 0.8 mM EDTA, 40% glycerol, 16 mM MgCl2, with 10 mM DTT and 1 µl dI-dC added just before the reaction). A/T rich region mutant (M1) and RORE half-site mutant (M2) were tested for

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RORα binding. For cold completion, non-radiolabeled APOCIII probe (100x) was added.

For antibody super shift assay, a RORα antibody (3 μl) was added in binding buffer without DTT (lane 6) in EMSA assay mixture. Samples were loaded on a non-denaturing gel (10% polyacrylamide) for electrophoresis.

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3.13. Identification of RORα response element on mouse cyp8b1 promoter.

Further, to elucidate if RORα binding site is located on mouse cyp8b1 promoter, mouse cyp8b1 promoter and two 5’ deletion constructs were cloned in pGL3 basic luciferase reporter. These reporter plasmids were transfected along with RORα expression plasmid in HepG2 cells. Fig. 21 shows that RORα stimulated mouse cyp8b1 reporter (-580/+30) activity, while deletion construct of mouse cyp8b1 reporter (-

480/+30) abolished the stimulatory effect of RORα. Therefore, the RORα binding site may be located between nt -580 and -480 on mouse cyp8b1 promoter.

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Human …..+9ggagagctgtctgaaa aggtcagagcaaagca+39 Mouse --546aaatcttaaacaaaata aggtgaagattttgc -512

Figure 21. RORα response element is located upstream of the mouse CYP8B1 promoter.

HepG2 cells were co-transfected with wild type or deletion mutant of RORα binding site

(explained in upper panel) and wild type mouse CYP8B1 promoter. (A) Reporter assay of mouse cyp8b1 reporter with or without RORα plasmid transfection. Raw firefly luciferase data (RLU) was normalized by Beta-gal colorimetric readings. Each transfection was performed in triplicates. Raw firefly luciferase (RLU) activity was normalized by β-gal activity. Results were expressed as mean ± S.D.; n=3; an “*” indicates statistical significance vs. control (without RORα transfection), p< 0.05.

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3.14 RORα interacts with CBP.

Co-activators are important class of proteins that increase the rate of transcription by interacting with various nuclear receptors. However they do not directly bind to the

DNA in a sequence-specific manner. RORα interacts with various co-activators and binds to the target gene promoter to increase gene transcription [188]. To study RORα interaction with co-activators, mammalian two hybrid assay was performed. Mammalian two hybrid assay was an important technique to study protein –protein interaction. First,

VP16-RORα was generated and co-transfected with Gal4-PGC1α, Gal4-SRC2, Gal4-

SRC1 and Gal4-CBP. Fig. 22 shows that reporter activity was highest after co- transfection of VP16-RORα with Gal4-CBP. Results indicate that RORα could specifically interact with co-activator CBP. Interestingly, interaction of RORα and CBP increased significantly upon 8-Br-cAMP treatment. On the other hand, 7αOHC an inverse agonist of RORα, decreased the interaction of RORα and CBP. Cyclic AMP may elevate

RORα and CBP interaction, while 7αOHC decreases RORα and CBP association.

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Figure 22. RORα specifically interact with co-activator CBP

Mammalian two hydrid assay was performed using HepG2 cells transfected with 0.2 µg

PSG5 plasmid (5copies of Gal4 DNA binding sites), 0.1 µg of VP16-RORα and 0.1 µg of with or without Gal4-PGC1α, Gal4-SRC1, Gal4-CBP and Gal4-SRC2 constructs as shown in figure. Each transfection was performed in triplicates. Raw firefly luciferase activity (RLU) was normalized by β-gal activity. Results were expressed as mean ± S.E.; n=3; an “*”, “ #” and “**” indicates statistical significance vs. vehicle (DMSO) treated cells and 7α OHC treated cells respectively p< 0.05.

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3.15. RORα recruits CBP on CYP8B1 promoter.

To confirm RORα binding on the CYP8B1 promoter in vivo, chromatin precipitation assay approach (ChIP) was used. ChIP assay is a useful technique to detect structure of the chromatin and chromatin binding proteins. ChIP assay was performed using mouse liver over-expressing RORα (Ad-RORα infused) and control adenovirus

(Ad-null infused). Liver nuclear extracts were isolated for ChIP assay and a specific antibody against RORα or CBP were used to detect RORα or CBP chromatin recruitment. Primers were designed to amplify a fragment from nt -600 to -416 region containing the RORα response element (RORE). Fig. 23 (upper panel) shows that RORα binding to CYP8B1 promoter was significantly increased in liver of Ad-RORα infused mice compared to assay using Ad-null mice. CBP is a co-activator that interacts with

RORα, thus RORα recruits CBP to CYP8B1 promoter. Since, CBP has histone acetylase activity, the histone acetylation status on CYP8B1 gene promoter was increased significantly (Fig. 23). A region 5’ upstream was assayed as a negative control for ChIP assay (Fig. 23A, lower panel).

To study RORα and CBP recruitment on human CYP8B1 promoter, HepG2 cells were treated with 8-Br-cAMP and ChIP assay was performed. Fig. 23B shows that RORα and CBP occupancy increased on human cyp8b1 promoter upon 8-Br-cAMP treatment.

On the other hand, HNF4α chromatin occupancy was decreased which was consistent with the previous studies [100].

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Figure 23. RORα recruits co-activator CBP on cyp8b1 promoter.

(A) ChIP assay of Ad-RORα and control adenovirus (Ad-null) infused mouse liver using antibodies specific for RORα, HNF4α, CBP and control IgG. The ChIP was performed using two regions of mouse cyp8b1 promoter as explained in the upper panel of the figure using real time PCR. Data was normalized using 10% input and relative abundance was calculated with respect to Ad-null infused mouse liver ChIP DNA. Results were expressed as mean ± S.E.; n=5; an “*” indicates statistical significance vs. Ad-null infected mice, p< 0.05. (B) HepG2 cells were treated with 8-Br-cAMP (15 µM) for 18h and ChIP assay was performed. DNA was amplified using real time PCR and data was normalized to 10% input and relative abundance was calculated using DNA obtained from precipitation of control IgG. Results were expressed as mean ± S.E.; n=3; an “*” indicates statistical significance vs. vehicle treated cells, p< 0.05.

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3.16. RORα is phosphorylated by PKA

Phosphorylation is a key process, which regulates the function of various nuclear receptors. RORα regulates CYP8B1 gene expression upon 8-Br-cAMP treatment. We want to investigate if RORα is phosphorylated by PKA. Therefore, HepG2 cells were transfected with catalytic domain of PKA. Immunoprecipitation (IP) followed by immunoblotting (IB) was performed to detect phosphorylation of RORα using P-Serine specific antibody. Fig 24A shows that constitutively active PKA plasmid transfection induced phosphorylation of RORα. Further, HepG2 cells were treated with 8-Br-cAMP and PKA inhibitor (H89) and IP assay was performed to detect phosphorylation of

RORα. Fig. 24B shows that RORα phosphorylation increased after 8-Br-cAMP treatment. As expected the phosphorylation of RORα was decreased after PKA inhibitor

H89 treatment, while wortmannin, a PI3 kinase inhibitor had no effect on RORα phosphorylation. This experiment confirms the reported finding that RORα is phosphorylated by PKA likely at Serine 99 position.

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Figure 24. RORα is phosphorylated by PKA

HepG2 cells were transiently transfected with catalytically active PKA plasmid (0.1 µg) for 48h. (A) IP was performed using RORα specific antibody and phosphorylation of

RORα was detected using phosphorylated serine (p-serine) specific antibody. (B) HepG2

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cells were treated with 8-Br-cAMP (20 µM), H89 (5 µM) and wortmannin (5 µM). IP assay using RORα specific antibody and phosphorylated RORα was detected using p- serine antibody.

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3.17 Phosphorylation of RORα increases stability of RORα protein

To study RORα protein half-life, HepG2 cells were treated with protein synthesis inhibitor, cycloheximide (CHX) with or without 8-Br-cAMP at different time points (Fig.

25A). Treatment of CHX decreased RORα protein level rapidly, but treatment of CHX and 8-Br-cAMP together maintained the protein level. This experiment suggests that cAMP may increase the stability of RORα protein. Further, experiment was carried out with treatment of 8-Br-cAMP for 6h followed by treatment of CHX. RORα protein was monitored after every 2h. Fig. 25B shows that RORα protein level was elevated after 6h of 8-Br-cAMP treatment, but decreased slowly within 16h after CHX treatment. This experiment also indicated that 8-Br-cAMP treatment activated PKA, which phosphorylates RORα and increased stability of RORα protein.

Half-life of RORα Protein was calculated using first order reaction plot ln

(RORα/histone) intensity vs. time as shown in Fig. 25C. Slope of the graph was calculated, which was a first order rate constant (K). Half-life period (T1/2) of RORα protein was calculated using formula ln2/ (-K). The calculated T1/2 of RORα protein is

4.71h. Treatment of 8-Br-cAMP increased half-life of the protein to 7.58h. This experiment indicates that phosphorylation increased the half-life of the protein.

Proteins are degraded by proteosome mediated pathway. To study the effect of 8-

Br-cAMP on proteosome mediated degradation of RORα, antibody against 20S proteosome was used to co-IP RORα. RORα was detected using immune-blot analysis.

Fig. 24D shows that 8-Br-cAMP treatment significantly reduced RORα association with

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proteosome, which suggest that phosphorylation of RORα may reduced proteosome degradation.

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Figure 25. Phosphorylation of RORα increases stability of the protein

HepG2 cells were treated with cycloheximide (CHX 20 µM) only or together with 8-Br- cAMP (15 µM) at various time points as indicated in each figure. (A) and (B) HepG2 cells were pretreated with 8-Br-cAMP (15 µM) for 6h followed by new media supplanted with cycloheximide (CHX 20 µM).Total cell extract were used for western blot analysis, which was performed using RORα and histone-3 antibody (C) First order reaction plot ln

(RORα/ histone intensity) vs. time (h). First order rate constant was calculated (Slope).

Half-life period was calculated T1/2 = ln2/(-K). (D) Co-IP assay using 20S proteosome antibody and IB with RORα antibody with or without 8-Br-cAMP treatment.

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CHAPTER IV

DISCUSSION

Bile acid synthesis is regulated by fasting and feeding cycle and exhibits a distinct circadian rhythm. After food intake, bile acids play a more prominent role, since bile acids absorb fat soluble vitamins from the intestine and regulate cholesterol homeostasis

[184]. There are contradictory findings whether bile acid synthesis is inhibited during fasting. Earlier reports showed that fasting induces bile acid synthesis and CYP7A1 gene expression in mice [184-185]. However, recently it was reported that fasting suppresses bile acid synthesis by inhibiting CYP7A1 gene expression [184]. The observed inhibition of CYP7A1 gene expression in the fasted animals could be the effect of glucagon.

Similarly, there is limited study on how glucagon regulates other bile acid synthesis genes. Therefore, the main objective of this work is to understand how glucagon changes the overall CYP8B1 gene expression and bile acid composition.

RORα is a fasting induced clock gene plays a critical role in regulation of circadian rhythm. RORα is a constitutively active nuclear receptor, which binds to bile acid synthesis intermediate 7α-hydroxycholesterol. Interestingly, 7α-hydroxycholesterol binding suppresses RORα transcription activity. This observation suggests that RORα may regulate bile acid composition and fasting induced CYP8B1 gene expression.

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Therefore, another objective of this work is to study if RORα regulates CYP8B1 circadian rhythm.

4.1. Circadian rhythm of CYP8B1.

Circadian rhythm is known to regulate metabolism [130,142]. Disruption of circadian rhythm often leads to metabolic disorders such as obesity and type 2 diabetes

[161, 189-190]. Deletion of CLOCK gene in mice disrupts cholesterol metabolism and increases cholesterol accumulation in the liver [191]. This study underlines the importance of circadian rhythm to maintain cholesterol homeostasis. It is well established that RORα regulates circadian clock by inducing BMAL1 gene expression [125, 129].

Although RORα regulates several metabolic process, expression of RORα is significantly altered during the day and night cycle [125].

Present work sheds light on the underlying molecular mechanism of diurnal rhythm of CYP8B1 gene expression. CYP8B1 expression exhibits a diurnal rhythm that is opposite to that of CYP7A1. In normal mouse physiology (free-fed), CYP8B1 mRNA shows a diurnal rhythm that peaks at early light cycle (ZT 2) and has a nadir at early dark cycle (ZT 14). According to previous reports, CYP7A1 mRNA shows circadian rhythm that peaks at middle of the dark cycle (ZT 18) and shows a nadir at late light cycle [192].

There is a large difference in the circadian rhythms of these two genes. Interestingly,

RORα mRNA peaked at late dark cycle (ZT 22), while protein is elevated at early light cycle (ZT 0). There is a correlation between RORα protein and CYP8B1 mRNA expression. This observation indicates that RORα may regulate the circadian rhythm of

CYP8B1 mRNA in mouse liver.

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Prior studies indicate that fasting elevates serum concentration of cholic acid in human subjects [193-195]. Elevation of cholic acid in the serum is a marker of enhanced cholic acid synthesis and induction of CYP8B1 gene expression. These human studies are consistent with our fasted mouse model, which shows increase in CYP8B1 gene expression and cholic acid synthesis.

The diurnal rhythm of CYP8B1 mRNA and protein and RORα protein is completely disrupted after 12h of fasting. Feeding inhibits CYP8B1 mRNA expression and RORα protein expression. Feeding had no effect on diurnal rhythm of RORα mRNA expression. Interestingly, feeding strongly reduced RORα protein expression level, this is correlated to reduction of CYP8B1 expression. Thus, regulation of RORα protein stability plays a key role in diurnal rhythm and fasting-re-feeding response of CYP8B1.

Glucagon is a major hormone secreted from pancreatic α cells during fasting. The present work shows that glucagon suppresses cyp7a1 mRNA in a time dependent manner, which is consistent with previous findings [187]. CYP7A1 is a rate limiting enzyme in bile acid synthesis. Thus bile acid synthesis is reduced during fasting. On the other hand, glucagon increases CYP8B1 mRNA expression, an important branch point enzyme, which produces cholic acid. Therefore, fasting facilitates change in bile acid composition. Bile acid composition is very important to facilitate intestinal absorbtion of cholesterol in post parendial state. Fasting induced higher level of cholic acid , which has low critical micelle concentration and facilitate bile storage in as mixed micelle in gallbladder.

In contrast to RORα, Rev-erbα is a negative clock gene that recruits a nuclear receptor.co-repressor 1 (NCoR1) and histone deacetylase 3 to inhibit target gene

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expression by binding to the same response element as RORα. Thus, Rev-erbα and RORα oscillate to regulate the circadian rhythm of CYP8B1, which is opposite to that of

CYP7A1 mRNA expression.

Regular fasting and feeding cycle is important to maintain cholesterol homeostasis in the healthy individuals [161]. Recent studies show that intermittent or habitual fasting often increases the tendency of eating high cholesterol or high fat food in the diet, which increases the risk of hypercholesterolemia and atherosclerosis in human subjects [190, 196]. This study suggests that prolonged fasting can enrich CA in the bile, which may increase dietary cholesterol absorption in the successive meal. Therefore, prolonged and intermittent fasting followed by cholesterol rich food consumption may increase the risk of hypercholesterolemia in healthy human subjects.

4.2. RORα regulation of CYP8B1 gene expression.

Fasting is a complex biological process, which is regulated by various transcription factor such as cyclic AMP response element binding protein (CREB),

Forkhead box protein O1 (FoxO1) and PPARα. FoxO1 indirectly regulates insulin sensitivity in the liver. Liver specific FoxO1 ablation shows abnormal lipid profiles in mice. Insulin signaling phosphorylates FoxO1 causing its nuclear exclusion and degradation to inhibit gluconeogenesis and glycogenolysis [197-198]. According to a recent report, loss of hepatic FoxO1 and Ldlr results in altered bile acid composition due to ablation of CYP8B1 gene expression [182]. This study suggests that FoxO1 induces

CYP8B1 gene expression in mice. On the other hand, FoxO1 inhibits CYP7A1 gene expression and bile acid synthesis [199]. Insulin treatment increases phosphorylation of

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FoxO1 and induces CYP7A1 gene expression. FoxO1 inhibits CYP7A1 gene expression via interaction with HNF4α to decrease interaction of HNF4α and PGC1α on CYP7A1 promoter [198]. Present study indicates that FoxO1 suppresses human CYP8B1 reporter activity. Therefore, FoxO1 may inhibit human CYP8B1 gene expression in contrast to expected inhibition of CYP8B1 mRNA expression in FoxO1/ LDLR knockout mice. The detailed role of FoxO1 in the regulation of CYP8B1 gene expression remains unclear.

PPARα was identified as an important nuclear receptor, which stimulates mouse and rat cyp8b1 mRNA expression. Loss of PPARα in mice reduces cyp8b1 mRNA upon fasting [66, 112]. It is very well known that PPARα and RXR binds to the DR1 site of the target gene promoter [7]. Our study indicates that PPARα inhibits human CYP8B1 reporter activity, consistent with previous reports [66]. Therefore, PPARα may inhibit human CYP8B1 gene expression, while it induces mouse CYP8B1 gene expression.

Although the mouse CYP8B1 promoter has a distinct PPARα binding site, which may be absent on human CYP8B1 promoter. This observation again reveals the differential actions of PPARα between mouse and human CYP8B1 gene regulation.

Ablation of RORα and RORγ in mice shows abolished cyp8b1 mRNA expression

[119, 125]. Present study indicates that RORα and RORγ both stimulate CYP8B1 reporter activity. However, RORα further stimulates human CYP8B1 reporter activity upon cAMP treatment. These results indicate that RORα plays a major role in stimulation of CYP8B1 gene expression after glucagon treatment. RORα is more abundantly expressed in the liver than RORγ. Therefore, the overall goal of this work is to study role of RORα in regulation of CYP8B1 gene expression.

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Results show that RORα stimulates human and mouse CYP8B1 reporter activity.

Treatment of 7αOHC, an inverse agonist of RORα, decreases the human CYP8B1 reporter activity and CYP8B1 mRNA level. 7αOHC is produced in the first step of bile acid synthesis by CYP7A1 from cholesterol. Interestingly, 7αOHC may regulate cholic acid synthesis by decreasing RORα activation and CYP8B1 mRNA level. Therefore,

CYP7A1 activity can regulate cholic acid synthesis. In bile acid synthesis, the CYP7A1 mRNA expression is elevated at early dark cycle, while CYP8B1 mRNA expression peaks at early light cycle. This observation also supports our finding that 7αOHC produced in the first step of bile acid synthesis may inhibit RORα activity, CYP8B1 gene expression and bile acid composition.

RORα binding site is conserved in human and mouse CYP8B1 promoter. Site directed mutagenesis and EMSA assay confirms that the human CYP8B1 promoter has a distinct RORα response element. Half site AGGTCA and 5’ A/T rich region are required for RORα binding. This response element is recognized by other monomeric receptors such as LRH1, Nur77 and Rev-erbα. EMSA assay shows Rev-erbα binds to the RORα response element on the human CYP8B1 promoter (data not shown). Several lines of evidence indicate that Rev-erbα and RORα share the same binding site on the target gene promoters [128-129]. RORα is a constitutively active receptor, while Rev-erbα acts as a repressor. Therefore, RORα and Rev-erbα both may regulate CYP8B1 gene expression in fasting and feeding, respectively. Fasting stimulates glucagon/ cAMP / PKA, which stabilizes RORα protein by phosphorylation. On the other hand, feeding stimulates

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GSK3β phosphorylation, which in turn induces Rev-erbα phosphorylation to increase

Rev-erbα protein stability [130, 187].

Deletion of RORα in HepG2 cells suppresses cyp8b1 mRNA expression and induction of cyp8b1 mRNA due to 8-Br-cAMP treatment. This data strongly indicates that the RORα is an important factor, which regulates CYP8B1 gene expression after glucagon/cAMP treatment. HNF4α stimulates CYP8B1 and CYP7A1 gene expression.

Results indicate that there is no synergistic effect of HNF4α and RORα on human

CYP8B1 reporter activity. Therefore, RORα activates CYP8B1 gene expression independent of HNF4α.

4.3. Regulation of bile acid composition

To study the regulation of bile acid composition in mice, adenovirus expressing

RORα is generated. Over-expression of RORα increases CYP7B1 mRNA level, which is involved in alternate bile acid synthesis pathway. Alternate bile acid synthesis pathway, mainly produces chenodeoxycholic acid (CDCA) [1, 200]. Activation of alternate bile acid synthesis pathway produces many acidic intermediates, which increases liver inflammation [200]. Studies indicate that activation of pro-inflammatory cytokines induces VLDL secretion and may increase risk of hypertriglyceridemia [201-202].

Over-expression of RORα increases CYP8B1 gene expression, while bile acid pool size remains unchanged. This is expected since CYP7A1 mRNA remains unchanged upon RORα over-expression. Further, RORα over-expression significantly increases serum cholesterol level and tends to increase liver cholesterol level. RORα is known to induce APOCIII gene expression in the mouse liver [121]. According to previous reports,

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APOCIII is an inhibitor of lipoprotein lipase, which may cause an increase in the serum triglyceride levels [203-204]. However, the present study shows that serum triglyceride

(TG) level remain unaffected after RORα over-expression.

Lithogenic diet containing cholic acid is known to induce atherosclerosis or hypercholesterolemia. Cholic acid feeding increases bile acid pool size and alters bile acid composition by increasing unconjugated bile acids in the bile acid pool. Similarly, cholic acid feeding alters serum TG levels [75, 153, 205]. Here, we observed that RORα increases cholic acid synthesis and alters bile acid composition. There is an increase in tauro cholic acid (TCA) and taurodeoxycholic acid (TDCA), while muricholate, and taurochenodeoxy cholic (TCDCA) acids are decreased. TCDCA is the physiological ligand of FXR. Although taurocholic acid is a weak ligand of FXR, decrease in TCDCA may alter the FXR signaling upon RORα overexpression. This RORα-mediated CYP8B1 over-expressing mouse model could be useful for studying the pathogenesis of nonalcoholic fatty liver disease (NAFLD), atherosclerosis and diabetes. This mouse model is opposite of the CYP7A1 transgenic mouse model, which is resistant to western high fat diet-induced insulin resistance and obesity. In CYP7A1 transgenic mice, bile acid pool size increased two-fold and CYP8B1 was inhibited and resulted in CDCA as the predominant bile acid with very little CA in bile acid pool [206]. A previous study shows that FXR plays an important role in triglyceride metabolism. CYP8B1 over expression my reduce FXR signaling by decreasing FXR ligand CDCA/TCDCA. Reduced FXR signaling may decrease TG clearance in fatty liver disease [48].

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Thus, RORα induction may have higher liver inflammation and higher risk of

NAFLD. This study identifies a novel regulatory function of RORα in regulation of cholic acid synthesis and pathways of NAFLD.

Bile acids, mainly TDCA and DCA increase inflammation in the liver and colon

[28, 57]. Increase in DCA induces apoptosis in liver cells and increases cytokine production in macrophages [207]. RORα over expression elevates TDCA/DCA in the bile acid pool, which may increase inflammation in liver and colon. Therefore, antagonizing RORα could be a novel therapy to reduce inflammation in the liver.

In this work, it was observed that RORα over-expression increases hyocholic acids. Hyocholic acids are the hydroxylated bile acids, which are synthesized from

CDCA by CYP3A11 enzyme in the liver [208]. Induction of CYP3A11 gene expression by PXR protects against bile acid induced hepatic toxicity [11]. Although LCA or derivatives of LCA are potent ligands of PXR, DCA or TDCA may also activate PXR

[209]. Previous findings show that PXR activation inhibits CYP7A1 gene expression.

Therefore, prolonged increase in DCA/TDCA may inhibit CYP7A1 gene expression by activation of PXR. This study also suggests a cholic acid signaling network, which may inhibit FXR activation that may activate PXR signaling.

Furthermore, dietary cholesterol absorption is one of the important functions of cholic acid. An elevated level of cholic acid is responsible in formation of gallstone in human subjects. ABCG5 and ABCG8 are cholesterol transporters, which efflux cholesterol into bile [41]. FXR induces ABCG5/G8 gene expression and regulates cholesterol secretion in the bile [210]. Over-expression of ABCG5/G8 decreases intestine

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cholesterol absorption, while elevates cholesterol secretion in the bile [211]. This may reduce cholesterol secretion in the bile, while may increase dietary cholesterol absorption in the intestine. Therefore, increase in cholic acid may elevate risk of dyslipidemia.

Overall, this study indicates that a balance in bile acid composition is very important for cholesterol homeostasis.

RORα acts as a constitutively active nuclear receptor, which interacts with different co-activators [60, 122, 212]. The crystal structure of RORα shows that the cholesterol or cholesterol sulfate was bound to RORα ligand binding pocket. However, the role of cholesterol or cholesterol sulfate in activation of RORα remains unclear.

Therefore, RORα may act as a constitutively active nuclear receptor without endogenous ligand, while ligand binding may reduce RORα activation [213]. Oxysterols, mainly

7αOHC, strongly reduce RORα activation. On the other hand, oxysterols such as 24- hydroxy cholesterol (24OHC) and 25OHC weakly reduce RORα activation [127].

Twentyfour-hydroxy cholesterol (24OHC) activates nuclear receptor LXR, which regulates lipogenesis by inducing SREBP1c gene expression [123]. Therefore, oxysterols have opposite effect on LXR and RORα. According to previous report, RORα activates

SREBP1c gene expression in the liver [123]. However, SREBP1c mRNA expression remains unaltered upon RORα over expression (data not shown).

4.4. RORα phosphorylation by PKA

Recent report reveals that a major liver isoform of RORα (RORα4) is regulated by phosphorylation [65]. Although there are five PKA consensus phosphorylation sites

RXXS or RXS in RORα protein, serine 99 located on the hinge domain region of the

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protein is the most important one [65]. The physiological role of phosphorylation is not clear. Our work is consistent with the observation that PKA indeed phosphorylates RORα protein. Interestingly, treatment of 8-Br-cAMP activates PKA and phosphorylates RORα and increases RORα protein stability which increases the half-life of RORα protein. Most proteins undergo ubiquitination followed by proteosome mediated degradation. Present work reveals that phosphorylation of RORα decreases RORα association with proteosome to increase its half-life.

RORα interacts with different co-activators. Therefore, mammalian two hybrid assay was performed to identify a co-activator for RORα. Results show that RORα and

CBP interaction is strongest among all the co-activators. Interestingly, RORα and CBP interaction is reduced upon 7αOHC treatment. ChIP assay indicates significant increases in RORα and CBP occupancy on human and mouse cyp8b1 chromatin. RORα co- localizes with CBP, which supports initial finding that RORα specifically binds to the co- activator CBP. CBP is a histone acetyl transferase protein and significantly increases acetylation of histone, which in turn activates CYP8B1 gene expression.

Overall, this study investigates a novel mechanism by which glucagon/cAMP/PKA phosphorylates RORα to increase its stability and inhibits its association with the proteosome. Previously, serine 99 position was identified as a PKA phosphorylation site on RORα [65]. We speculate that this serine 99 position could be the site of ubiquitination [130]. Phosphorylation at this serine site could inhibit ubiquitination, which could also contribute to the stabilization of RORα protein. Further, we also observed that phosphorylation of RORα may increase co-activator CBP

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association. Although phosphorylation site of PKA is located in the hinge domain of

RORα, phosphorylation may lead to the higher interaction with co-activator CBP.

According to several reports PKA activates other signaling pathway such as

MAPK/ERK pathway in different tissues and cell lines. Interestingly, ERK pathway is activated in the mouse liver after feeding, which was reported previously [184].

Overall, this study investigates a novel mechanism in which phosphorylation of

RORα by PKA increases stability of the protein and enhances its interaction with CBP

(co-activator). Increase in RORα and CBP interaction stimulates histone acetylation and activates CYP8B1 gene transcription in response to fasting.

4.5 Physiological Significance

Glucagon, the product of pancreatic α-cells, is a major counteracting hormone to insulin in regulating glucose homeostasis during fasting or hypoglycemic conditions.

Glucagon promotes hepatic gluconeogenesis and glycogenolysis, while inhibiting glycogen synthesis and glycolysis [170, 175]. In addition, glucagon has been shown to play a major role in the development of hyperglycemia in both type 1 and 2 diabetes mellitus [174]. Normally, glucagon level decreases after meal, but according to recent studies, glucagon level is high in human subjects with type 1 and type 2 diabetes even after mealtime, causing hyperglycemia [170].

Type 2 diabetes have higher risk of gallstone disease. Higher level of cholesterol retention in the gallbladder causes gallstone formation. cholic acid has lower critical micelle concentration increases cholesterol crystal formation in the gall bladder. Studies

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indicate that higher level of cholic acid in bile increases risk of gallstone formation [153].

Therefore, antagonizing RORα may be a novel approach to decrease gallstone formation.

Hydrophobic bile acids are toxic and can cause inflammation and injury in the liver and intestine. The balance of bile acid pool size with proper bile acid composition may be important in activating FXR and TGR5 signaling for in anti-inflammation in the digestive system. On the other hand, enlarging the bile acid pool with CA may cause dyslipidemia and contribute to cholesterol gallstone disease and nonalcoholic fatty liver disease, which have high prevalence in type II diabetic patients.

A prolonged increase in cholic acid causes hypercholesterolemia, and increases the risk of atherosclerosis [150-151, 155]. This work suggests that the increase in cholic acid in diabetes may be the result of glucagon stimulated RORα induction of CYP8B1 gene expression. This work also suggests that RORα could be a novel drug target to treat hypercholesterolemia.

This study identifies a novel cholic acid signaling network and may suggest the detrimental effect of cholic acid in development of non-alcoholic fatty liver disease

(NAFLD).

Disruption in circadian rhythm often leads to metabolic disorders such as obesity and type 2 diabetes [161]. Present work shows that fasting and feeding altered circadian rhythm of mouse CYP8B1 expression. Although previous reports indicate that insulin is a major regulator of CYP8B1 circadian expression [167], present study reveals that glucagon is a key regulator of CYP8B1 circadian rhythm. Another novel finding of this work is how a change in fasting and feeding cycle may alter bile acid composition, which

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may disrupt dietary cholesterol homeostasis. Further, a prolonged increase in cholic acid is detrimental, which could happen during prolonged fasting [190, 196].

Present study contributes to our understanding of the molecular mechanism by which bile acid synthesis and composition regulates hepatic metabolic homeostasis.

Enrichment of cholic acid in bile acid pool may alter the bile acid synthesis by inhibiting

CYP7A1 gene expression and disrupt the cholesterol homeostasis. Overall, our study indicates that antagonizing RORα may improve NAFLD disease condition. Therefore,

RORα could be a novel drug target for gallstone disease, NAFLD related inflammation.

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4.6. Future Direction

Present study indicates that bile acid composition is very important for dietary cholesterol absorption. Increase in cholic acid in the bile acid pool may alter FXR and

PXR signaling pathways, which may regulate hepatic cholesterol and serum cholesterol level. Therefore, it is important to study the transgenic CYP8B1 mouse model.

Previously, studies from our lab showed that CYP7A1 transgenic mice are resistant to western high fat diet induced insulin resistance and obesity [206]. This transgenic

CYP8B1 mouse model could be opposite to our CYP7A1 model. Therefore, it is important to identify the role of cholic acid in hypercholesterolemia and atherosclerosis development.

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