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Hepatic microsomal bile acid biotransformation : identification of metabolites and cytochrome p450 enzymes… Deo, Anand K. 2009

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HEPATIC MICROSOMAL BILE ACID BIOTRANSFORMATION: IDENTIFICATION OF METABOLITES AND CYTOCHROME P450 ENZYMES INVOLVED by Anand K. Deo B. Pharm., University of Pune, India, 2000 M. Pharm. Sci., University of Mumbai, 2002 M.Sc., The University of British Columbia, 2005 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (PHARMACEUTICAL SCIENCES) THE UNIVERSITY OF BRITISH COLUMBIA Vancouver February 2009 © ANAND K. DEO, 2009 ABSTRACT Bile acids are end-products of cholesterol metabolism and essential for absorption of dietary lipids in the body. Impaired bile flow leads to hepatic bile acid accumulation and liver damage. Hepatic microsomal oxidation offers a potential mechanism for efficient elimination of bile acids. The present study investigated the cytochrome P450 (P450)-mediated hepatic microsomal biotransformation profiles of lithocholic acid, cholic acid and chenodeoxycholic acid using a liquid chromatography-mass spectrometry (LCIMS) based assay. Incubation of lithocholic acid with rat hepatic microsomes resulted in the formation of a major 6J3-hydroxylated metabolite, murideoxycholic acid, followed by isolithocholic acid and 3- ketocholanoic acid. Ursodeoxycholic acid, hyodeoxycholic acid and 6-ketolithocholic acid were identified as minor metabolites. Studies using P450-specific antibodies, chemical inducers, and rat recombinant enzymes showed that formation of murideoxycholic acid and 3-ketocholanoic acid were mediated by CYP3A2 and CYP2C 11. Similar metabolite profiles were obtained by incubation of lithocholic acid with mouse hepatic microsomes generating murideoxycholic acid as the major metabolite. Studies using P450 inducers and chemical inhibitors suggested the involvement of murine CYP3A in murideoxycholic acid and 3-ketocholanoic acid formation, and CYP1A, CYP2B and CYP3A enzymes in ursodeoxycholic acid, hyodeoxycholic acid and 6-ketolithocholic acid formation by mouse liver microsomes. Biotransformation of lithocholic acid by human hepatic microsomes generated 3-ketocholanoic acid as the major metabolite, and hyodeoxycholic acid, ursodeoxycholic acid, 6-ketolithocholic acid and murideoxycholic acid, as minor metabolites. Studies with chemical inhibitors and human recombinant enzymes demonstrated that CYP3A4 catalyzed the formation of all five metabolites. The biotransformation of cholic acid and chenodeoxycholic acid by human hepatic microsomes revealed the formation of a single cholic acid metabolite, 3-dehydrocholic acid. 11 Chenodeoxycholic acid biotransformation generated 7ct-hydroxy-3 -oxo-5 3-cholan-24-oic acid as the major metabolite followed by ‘y-muricholic acid, 7-ketolithocholic acid and cholic acid, respectively. CYP3A4 was found to be the major enzyme involved in the biotransformation of cholie acid and chenodeoxycholic acid in human liver microsomes. A comparison of metabolite profiles demonstrated the dominant role of human CYP3A4 in the oxidation of bile acids at the C-3 position. In contrast, 63-hydroxy1ation catalyzed by multiple P450 (CYP1A, CYP2B, CYP2C and CYP3A) enzymes was the preferred biotransformation pathway in rodent liver microsomes. 111 TABLE OF CONTENTS ABSTRACT ii TABLE OF CONTENTS iv LIST OF TABLES ix LIST OF FIGURES x LIST OF ABBREVIATIONS xiii ACKNOWLEDGEMENTS xiv DEDICATION xv CO-AUTHORSHIP STATEMENT xvi 1. 1 1.1 Bile Acids: Structure And Physicochemical Properties 2 1.2 Bile Acid Biosynthesis: Steps Involved 3 1.2.1 Biosynthesis of primary bile acids 4 1.2.2 Biosynthesis of secondary bile acids 8 1.3 Biliary Bile Acids: Composition And Physiological Relevance 9 1.4 Bile Acids: Enterohepatic Circulation 10 1.5 Bile Acid Toxicity 12 1.6 Cholestasis And Bile Acid Toxicity 13 1.7 Nuclear Receptors In Bile Acid Regulation 18 1.8 Bile Acid Biotransformation By Conjugation 22 1.9 Bile Acid Biotransformation By P450 Enzymes 24 1.10 Rationale 28 1.11 Hypotheses 31 1.1 1.1 Specific research objectives 32 1.12 References 50 iv 2. BIOTRANSFORMATION OF LITHOCHOLIC ACID BY RAT HEPATIC MICROSOMES: METABOLITE ANALYSIS BY LIQUID CHROMATOGRAPHY/MASS SPECTROMETRY 67 2.1 Summary 68 2.2 Introduction 69 2.3 Materials And Methods 71 2.3.1 Chemicals and reagents 71 2.3.2 Animal treatment and preparation of hepatic microsomes 72 2.3.3 Lithocholic acid biotransformation assay 72 2.3.4 Antibody inhibition 74 2.3.5 Analytical methods 74 2.3.6 Data analysis and calculation of enzyme kinetic parameters 75 2.3.7 Statistical analysis 76 2.4 Results 76 2.4.1 Biotransformation of lithocholic acid and metabolite identification 76 2.4.2 Kinetic analysis of hepatic microsomal metabolite formation 78 2.4.3 Effect of P450 inducers on lithocholic acid biotransformation 80 2.4.4 Antibody inhibition studies 80 2.4.5 Biotransformation studies with recombinant P450 enzymes 81 2.5 Discussion 82 2.6 References 99 3. IDENTIFICATION OF HUMAN HEPATIC CYTOCHROME P450 ENZYMES INVOLVED IN THE BIOTRANSFORMATION OF CHOLIC AND CHENODEOXYCHOLIC ACID 104 3.1 Summary 105 3.2 Introduction 106 3.3 Materials And Methods 108 3.3.1 Chemicals and reagents 108 V 3.3.2 Cholic acid and chenodeoxycholic acid biotransformation assays 109 3.3.3 Analytical methods 110 3.3.4 Data analysis and calculation of enzyme kinetic parameters 111 3.4 Results 112 3.4.1 Biotransformation and kinetic analysis of cholic acid metabolites 112 3.4.2 Biotransformation and kinetic analysis of chenodeoxycholic acid metabolites 113 3.4.3 Biotransformation studies with human recombinant P450 enzymes 114 3.5 Discussion 116 3.6 References 130 4. 3-KETOCHOLANOIC ACID IS TifE MAJOR CYP3A4 MEDIATED LITHOCHOLIC ACID METABOLITE IN HUMAN HEPATIC 133 4.1 Summary 134 4.2 Introduction 135 4.3 Materials And Methods 137 4.3.1 Chemicals and Reagents 137 4.3.2 Lithocholic acid biotransformation assay 138 4.3.3 Analytical methods 139 4.3.4 Data analysis and calculation of enzyme kinetic parameters 139 4.3.5 Chemical inhibition studies 140 4.3.6 Statistical analysis 141 4.4 Results 141 4.4.1 Lithocholic acid biotransformation by human liver microsomes 141 4.4.2 Lithocholic acid biotransformation by human recombinant P450 enzymes 142 4.4.3 Chemical inhibition studies 143 4.4.4 Lithocholic acid biotransformation in human hepatic microsomes with varying CYP3A4 levels 143 4.5 Discussion 144 4.6 References 158 vi 5. HEPATIC MICROSOMAL P450 MEDIATED BIOTRANSFORMATION OF LITHOCHOLIC ACID BY MOUSE HIPi4.TIC 1’II1SO1VIES 162 5.1 Summary 163 5.2 Introduction 164 5.3 Materials And Methods 165 5.3.1 Chemicals and reagents 165 5.3.2 Animal treatment and preparation of microsomes 166 5.3.3 Lithocholic acid biotransformation assay 166 5.3.4 Chemical inhibition studies 166 5.3.5 Statistical analysis 167 5.4 Results 167 5.4.1 Hepatic biotransformation of lithocholic acid by mouse hepatic microsomes 167 5.4.2 Effect of P450 inducers in lithocholic acid biotransformation 167 5.4.3 Chemical inhibition studies 168 5.5 Discussion 170 5.8 References 178 6. ENE1J4.IJ DISCIJSSIO1i 181 6.1 Overview 182 6.2 Cholic Acid Biotransformation By Human Hepatic Microsomes 183 6.3 Chenodeoxycholic Acid Biotransformation By Human Hepatic Microsomes 183 6.4 Comparison of Lithocholic Acid Metabolite Formation Patterns By Rat, Human And Mouse Liver Microsomes 184 6.5 Limitations 188 6.6 Overall Significance of Thesis Research 190 6.7 Future Studies 191 6.8 References 200 vii ITIA £OZJ)CIGtsJaJcI’r LIST OF TABLES Table 1.1 Oxidative metabolites of bile acids 33 Table 1.2 Different P450 enzymes and their levels in human and rat hepatic microsomes 34 Table 2.1 Kinetic parameters of lithocholic acid metabolite formation by rat hepatic microsomes 88 Table 2.2 Effect of sex and treatment with P450 inducers on lithocho lie acid metabolite formation by rat hepatic microsomes 89 Table 4.1 Kinetic parameters of lithocholic acid metabolite formation by human hepatic microsomes 149 Table 4.2 Kinetic parameters of lithocholic acid metabolite formation by recombinant CYP3A4 150 Table 4.3 Rate of formation of lithocholic acid metabolites in human liver microsomes with varying CYP3A4 activities 151 Tabl 5 1 Effect of treatment with P450 inducers on lithocholic acid metabolitee formation by mouse hepatic microsomes 174 Table 6.1 Regio-selective oxidation of lithocholic acid by human, rat and mouse hepatic microsomes 195 Table 6.2 Comparison of P450 enzymes involved in lithocholic acid metabolite formation in human and rodent hepatic microsomes 196 ix LIST OF FIGURES Figure 1.1 Cholesterol and bile acid structure 35 Figure 1.2 Chemical structures of unconjugated and conjugated bile acids 36 Figure 1.3 Bile acid levels in human, rat and mouse liver 37 Figure 1.4 Neutral and acidic pathways in bile acid formation 38 Figure 1.5 Bile acid levels in human, rat and mouse bile 39 Figure 1.6 Enterohepatic circulation of bile acids 40 Figure 1.7 Amphipathic nature of bile acids 41 Figure 1.8 Obstruction to the flow of bile acids from the liver to the intestine is termed as cholestasis 42 Figure 1.9 Comparison of bile acid levels in livers of patients with end-stage cholestasis and normal humans 43 Figure 1.10 Bile acid regulation: A complex process involving various receptors, enzymes, and transporters 44 Figure 1.11 Regulation of bile acids by nuclear receptors 45 Figure 1.12 PXR and CAR cross-talk mediated metabolism of lithocholic acid 46 Figure 1.13 Major hepatic P450 enzymes involved in drug metabolism 47 Figure 1.14 Biotransformation of bile acids 48 Figure 2.1 General bile acid structure showing positions available for hydroxylation 90 Figure 2.2 Representative LC/MS chromatogram showing metabolites of lithocholic acid 91 Figure 2.3 Kinetic profiles of hepatic microsomal murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid formation 92 Figure 2.4 Effect of anti-CYP2C IgG and anti-CYP3A IgG on hepatic microsomal murideoxycholic acid (A), isolithocholic acid (B) and 3-ketocholanoic acid (C) formation 93 Figure 2.5 Comparison of murideoxycholic acid (A), isolithocholic acid (B) and 3- ketocholanoic acid (C) formation by a panel of rat recombinant P450 enzymes 94 Figure 2.6 Enzyme kinetic profile of 3-ketocholanoic acid formation by rat recombinant CYP3A2 95 Figure 2.7 Effect of CYP2D6 antiserum on murideoxycholic acid formation by rat hepatic microsomes and recombinant CYP2D1 96 x Figure 2.8 Scheme showing a proposed mechanism for the formation of 3- ketocholanoic acid and isolithocholic acid from lithocholic acid through a geminal diol intermediate 97 Figure 2.9 Scheme showing P450-mediated lithocholic acid biotransformation in rat hepatic microsomes 98 Figure 3.1 Representative LC/MS chromatogram showing metabolites of cholic acid 121 Figure 3.2 Enzyme kinetic profile of 3-dehydrocholic acid formation by human hepatic microsomes 122 Figure 3.3 Representative LC/MS chromatogram showing metabolites of chenodeoxycholic acid 123 Figure 3.4 Enzyme kinetic profiles of 7a-hydroxy-3-oxo 5f3-cholan-24-oic acid (A), y muricholic acid (B), 7-ketolithoeholic acid (C) and cholie acid (D) formation by human hepatic microsomes 124 Figure 3.5 Comparison of 3-dehydrocholic acid formation from cholic acid (A) and 7x- hydroxy-3-oxo-513-eholan-24-oic acid and y-muricholic acid formation from chenodeoxycholic acid (B) by a panel of recombinant human P450 enzymes 125 Figure 3.6 Enzyme kinetic profile of 3-dehydrocholic acid formation from cholic acid by recombinant human CYP3A4 126 Figure 3.7 Enzyme kinetic profiles of 7a-hydroxy-3-oxo 513-cholan-24-oic acid (A) and y-muricholie acid (B) formation from chenodeoxycholic acid by recombinant human CYP3A4 127 Figure 3.8 Scheme showing P450-mediated eholic acid biotransformation by human hepatie mierosomes 128 Figure 3.9 Scheme showing ehenodexycholic acid biotransformation by human hepatic microsomes 129 Fi ure 4 1 Representative LC/MS chromatogram showing metabolites of lithocholicg acid in human hepatie microsomes 152 Figure 4.2 Enzyme kinetic profiles of 3-ketocholanoic acid (A), hyodeoxycholic acid (B), ursodeoxycholic acid (C), murideoxycholic acid (D) and 6- ketolithocholie acid (E) formation by human hepatic microsomes 153 Figure 4.3 Comparison of 3-ketoeholanoic acid, hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholie acid and 6-ketolithocholic acid formation by a panel of human recombinant P450 enzymes 154 Figure 4.4 Enzyme kinetic profiles of 3-ketocholanoic acid (A), hyodeoxycholic acid (B), ursodeoxycholic acid (C), murideoxycholic acid (D) and 6- ketolithocholic acid (E) formation from lithocholic acid 155 Figure 4.5 Chemical inhibition studies using ketoconazole, quercetin, quinidine, and sulfaphenazole on lithocholic acid biotransformation by human hepatic miersomes 156. xi Figure 4.6 Scheme showing lithocholic acid biotransformation by human hepatic microsomes 157 Figure 5.1 Chemical inhibition studies using ketoconazole, quercetin, quinidine, and sulfaphenazole on lithocholic acid biotransformation by mouse hepatic microsomes 175 Figure 5 2 Scheme showing lithocholic acid biotransformation by mouse hepatic • microsomes 177 Fi ure 6 1 Comparison of lithocholic acid metabolite formation patterns obtained fromg • rat, mouse and human hepatic microsomes 197 Figure 6 2 Lithocholic acid biotransformation by rodent and human liver • microsomes 198 Fi ure 6 3 Hepatic microsomal P450-mediated bile acid biotransformation: Route tog alleviate bile acid mediated hepatotoxicity in cholestasis 199 xl’ LIST OFABBREVIATIONS ANOVA analysis of variance ASBT apical sodium-dependent bile salt transporter AUC area under curve BNF beta napthoflavone BSEP bile salt export protein CO corn oil C.V. coefficient of variation CAR constitutive androstane receptor flEX dexamethasone FIC1 familial intrahepatic cholestasis- 1 FXR farnesoid X receptor GC/MS gas chromatography mass spectrometry HSD hydroxy steroid dehydrogenase IgG immunoglobin G IS internal standard ISBT ileal sodium-dependent bile salt transporter LC-MS liquid chromatography mass spectrometry LCIMSIMS liquid chromatography tandem mass spectrometry LXR liver X receptor MC 3- methylcholanthrene MDR multidrug resistance P-glycoprotein MRP multiple resistance associated protein NADPH reduced nicotinamide adenine dinucleotide phosphate NTCP sodium taurocholate cotransporting polypeptide OATP organic anion transporter protein P450 cytochrome P450 PAR peak area ratio PB phenobarbital PBRE phenobarbital response elements PXR pregnane X receptor RXR retinoid X receptor s.c. subcutaneous SEM standard error of mean SKF-525A proadifen SPGP sister of P-glycoprotein SULT sulfotransferase VDR vitamin D receptor XRE xenobiotic response elements xlii ACKNOWLEDGEMENTS I would like to thank my research supervisor Dr. Stelvio Bandiera. Indeed, it was his scientific acumen and mentoring skills that enabled me to take up this work and take it this far. I express my gratitude to him for letting me explore this field of biotransformation and allowing me to choose my own research project. His valuable guidance and keen interest was the key, without which this work could not have been accomplished. I also take this opportunity to thank Dr. Frank Abbott, for providing the LC/MS facility in his lab for the successful completion of this work. My committee members Dr. Wayne Riggs, Dr. Adam Frankel, Dr. David Chen and were always available for comments and suggestions whenever in need, not only during my course work, but also, during various stages of my research. No work can be carried out alone, and it was here that Dr. Eugene Hrycay helped me in getting acclimatized to the lab techniques and adding some basic skill set to my armory. A special mention to Roland Burton and Andras Szietz is inevitable for their help in teaching me the hands on skills in operating the LC/MS. The hardworking fellow graduate students of my lab, Subrata Deb, Claudio Erratico, Sarah Moffat and Si Zhang have been a constant source of motivation in the completion of this study. I express my appreciation and gratitude to the faculty of Pharmaceutical Sciences, UBC for providing me an opportunity to avail top-notch training in terms of research and otherwise, in addition to supporting my living. I acknowledge Merck Research Laboratories for funding my studies and living in the form of the Merck Graduate Traineeship. I would also like to thank Canadian Institute of Health Research for providing support to our lab by funding our research grant titled, “Role of Hepatic Cytochrome P450 Enzymes in Bile Acid Metabolism And Detoxification”. No words can express my sense of gratitude towards my parents and wife, Mugdha for supporting me to pursue my research interests far from home. Last but not least, I remember God for His countless blessings, the power which has given me strength to face all odds in the successful completion of the thesis. To Him I owe everything. xiv AX MXN 0L CO-AUTHORSifiP STATEMENT This thesis incorporates materials that are results of research work carried out under the supervision of Dr. Stelvio M. Bandiera at the University of British Columbia, Faculty of Pharmaceutical Sciences, Vancouver, BC, Canada. As the first author of the two published chapters (chapters two and three) and two unpublished chapters (chapters four and five), I was responsible for identification of the research questions, design and implementation of the experiments, data analysis and preparation of manuscripts presented in this thesis. xvi Chapter 1 INTRODUCTION I 1.1 Bile Acids: Structure And Physicochemical Properties Bile acids (5 13-cholanoic acids) were isolated from bile early in the nineteenth century (Sobotka, 1938), thus initiating the field of bile acid chemistry. Bile acids are hydrophobic steroids consisting of a cyclopentanoperhydrophenanthrene nucleus, composed of saturated rings (A, B, C and D) with a branched isopentanoic side chain attached at C-17 (Fig. 1.1). During bile acid biosynthesis, isomerization of the A5’6 double bond of cholesterol to a double bond occurs. The double bond is then reduced to give a 5 3-cholanoic acid structure in which the A and B rings are in cis configuration (denoted by a 5-oriented hydrogen at the C-5 position). Other chemical modifications that occur during the conversion of cholesterol to bile acids include f3-oxidation of the c-i 7 isooctane aliphatic side chain to a shorter isopentanoic side chain, epimerization of the C-3 hydroxy group, and a- or f3-hydroxylation of one or more carbons on the steroid nucleus. Bile acid differentiation is characterized by the number and position of the hydroxyl groups attached to the steroid nucleus (Fig. 1.1) (Hofmann and Hagey, 2008). Bile acids that are synthesized from cholesterol in the hepatocyte are called primary bile acids. All primary bile acids have a 3cc- and a 7-hydroxyl group. Cholic acid (3cc, 7cc, 12cc- trihydroxy-5f3-cholanoic acid) and chenodeoxycholic acid (3cc, 7a-dihydroxy-53-cholanoic acid) are primary bile acids in most mammalian species (Vlahcevic et al., 1999; Hofmann and Hagey, 2008). Hyocholic acid (3cc, 6cc,7cc-trihydroxy-5f3-cholanoic acid) and muricholic acid are primary bile acids in pigs and mice, respectively. In hyocholic acid, the 1 2cc-hydroxyl group of cholic acid is replaced by a 6ct-hydroxyl group. In cc-muricholic acid (3cc, 613, 7cc-trihydroxy- 5 13-cholanoic acid) and f3-muricholic acid (3cc, 613, 713-trihydroxy-5 f3-cholanoic acid), the 12cc- hydroxyl group of cholic acid is replaced by a 613-hydroxyl group (Hofmann et al., 1992; Hofmann and Hagey, 2008). Bile acids that are formed by bacterial modification of primary bile 2 acids are called secondary bile acids. Secondary bile acids that are found in most species are deoxycholic acid, lithocholic acid and ursodeoxycholic acid. Secondary bile acids retain the 3cL- hydroxyl group but are conspicuous by the absence of the 7-hydroxyl group (e.g. lithocholic acid and deoxycholic acid) or by epimerization of the 7c-hydroxyl group in the primary bile acid to a 73-hydroxyl group (e.g. ursodeoxycholic acid) (Fig. 1.2). Bile acids are mainly found in the body in conjugated form. Bile acids undergo conjugation on a hydroxy group of the steroid nucleus or on the carboxylic acid side chain. The most common bile acid conjugates are amino acid conjugates involving taurine and glycine. Bile acids can also undergo conjugation with sulfate at C-3 and with glucuronidate at C-3 or C- 24. Figure 1.2 shows various bile acid conjugates using lithocholic acid as a prototype. Conjugation alters the solubility of the bile acid by decreasing its pKa value. Unconjugated bile acids have a pKa of 5-6 whereas glycine conjugates have a pKa of 3.9 and taurine conjugates have a pKa value < 1. Conjugation results in complete ionization and increased water solubility but decreased diffusion of the resulting bile acid across plasma membranes (Carey, 1984; Fini and Roda, 1987). Conjugated bile acids typically exist in ionized form in vivo, and hence are called bile salts. 1.2 Bile Acid Biosynthesis: Steps Involved Bile acids are synthesized in the liver, secreted into bile, which is stored in the gall bladder, absorbed from the distal intestine and finally returned to the liver. The total concentration of bile acids in human liver is about 60-61 nmol!g of liver. Chenodeoxycholic acid and cholic acid account for approximately 45% and 30%, respectively, of this amount. Secondary bile acids (deoxycholic acid, lithocholic acid and ursodeoxycholic acid) together 3 account for approximately 25% of the bile acids in liver (Fischer et al., 1996; Setchell et al., 1997). In rat and mouse liver, the primary bile acids are the muricholic acids, cholic acid and chenodeoxycholic acid. The total concentration of bile acids in rat liver is approximately 70 mnol/g of liver, of which the primary bile acids account for 65-67% and the secondary bile acids (lithocholic acid, hyodeoxycholic acid, deoxycholic acid and ursodeoxycholic acid) account for 33-35% of the total bile acids. Similarly in mouse liver, muricholic acids account for 57% of the total bile acid concentration (40 nmollg liver) followed by cholic acid (3 7%) and chenodeoxycholic acid (1%). Secondary bile acids in mouse account for a relatively minor percentage (approximately 5%) (Setchell et a!., 1997; Stedman et al., 2004) (Fig. 1.3). 1.2.1 Biosynthesis of primary bile acids. Biosynthesis of primary bile acids (cholic acid and chenodeoxycholic acid) from cholesterol in liver involves a wide array of enzymes localized in the endoplasmic reticulum, mitochondria, cytosol and peroxisomes (Princen et al., 1979; Vlahcevic et al., 1999; Russell, 2003). Two main pathways are involved in the formation of primary bile acids, the neutral pathway and the acidic pathway (Russell and Setchell, 1992; Russell, 2003; Norlin and Wikvall, 2007) (Fig. 1.4). In the neutral pathway, the bile acid precursors are present as neutral steroids until the final stages of bile acid formation. This pathway starts with the 7cL-hydroxylation of cholesterol, by cholesterol 7ct-hydroxylase (i.e. CYP7A1). 7c*-Hydroxycho1estero1 is then converted to 7a- hydroxy-4-cholesten-3-one by 313-hydroxy-A5-C-27 steroid dehydrogenase (313-HSD) (Wikvall, 1981; Ichimiya et al., 1991; Furster et al., 1996; Schwarz et al., 2000). The 12cc-hydroxyl group is introduced into the steroid nucleus to form 7cc, 1 2cc-dihydroxy-4-cholesten-3-one. This step is catalyzed by hepatic microsomal sterol 1 2c.c-hydroxylase (i.e. CYP8B 1) (Ishida et al., 1992; 4 Eggertsen et al., 1996). The further conversion of 7cc, 1 2ct-dihydroxy-4-cholesten-3-one requires a two step reduction in the cytosol catalyzed by 3-oxo-z4-ster id 5(3-reductase and 3c-HSD, respectively to form 513-cholestane-3a,7a,12a-triol (Kondo et al., 1994; Usui et al., 1994). 513- Reductase and 3o-HSD are also involved in the simultaneous formation of 5(3-cholestane 3cL,7a-diol from 7c-hydroxy-4-cholesten-3-one. Side chain cleavage is initiated by oxidation of the side chain by sterol 27-hydroxylase (CYP27A1), a mitochondrial cytochrome P450 (P450), which introduces a hydroxyl group into the 27-position of 5f3-cholestane-3c,7a,l2a-triol and 5f3-cholestane-3a,7c-diol to form 27-hydroxyl intermediates (Russell and Setchell, 1992; RusselL, 2003). The resulting 27-hydroxylated sterols, 5j3-cholestane-3c,7a,12cL 27-tetrol and 5[3-cholestane-3a,7c,27c-triol, are subsequently oxidized to cholic acid and chenodeoxycholic acid, respectively (Norlin and Wikvall, 2007) (Fig. 1.4). In the acidic pathway, cholesterol is converted to carboxylic acid intermediates, thus retaining a carboxylic acid group throughout the biosynthesis of primary bile acids. This pathway does not involve 7cL-hydroxylation of cholesterol. The CYP27A1 hydroxylation of cholesterol to form 27-hydroxycholesterol marks the first step of the acidic pathway (Wikvall, 1984). Alternatively, 27-hydroxycholesterol is further oxidized by CYP27A1 to form .313- hydroxy-5-cholestenoic acid. 27-Hydroxycholesterol and 3 3-hydroxy-5-cholestenoic acid are further hydroxylated at the 7cc-position by CYP7B 1 to form 7x,27-dihydroxycholesterol and 3 13,7cL-dihydroxy-5-cholestenoic acid, respectively. CYP7B 1 is not capable of 7a-hydroxylation of 27-hydroxycholesterol by itself but requires an additional enzyme, CYP7A1 (Wikvall, 1984). Hydroxy steroid dehydrogenase (313-HSD) oxidizes 7a,27-dihydroxycholesterol to 7c,27- dihydroxy-4-cholesten-3-one, which is further acted upon by CYP27AI to form 7cL-hydroxy-3- oxo-4-cholestenoic acid (Furster et al., 1996; Schwarz et al., 2000). 3f3-HSD is also responsible 5 for formation of 7c-hydroxy-3-oxo-4-cho1estenoic acid from 3 3,7cx-dihydroxy-5-cho1estenoic acid. 7c-Hydroxy-3-oxo-4-cholestenoie acid undergoes subsequent reduction to form chenodeoxycholic acid (Furster et al., 1996; Norlin and WikvaLl, 2007) (Fig. 1.4). Other relatively minor bile acid biosynthetic pathways include the 24-hydroxylase pathway, the 25-hydroxylase pathway, and the 27-hydroxylase pathway. In the 24-hydroxylase pathway, 24-hydroxylation of cholesterol by CYP46A1 in the brain has been proposed (Bjorkhem and Eggertsen, 2001; Bjorkhem and Meaney, 2004; Norlin and Wikvall, 2007). 24- Hydroxylated cholesterol from the brain crosses the blood brain barrier and reaches the liver by hepatic portal circulation. In the liver it is acted upon by CYP7A 1 or by CYP39A 1, another hepatic 7cL-hydroxylase, to form 7o,24-dihydroxycho1esterol (Li-Hawkins et al., 2000; Norlin et a!., 2000; Norlin and Wikvall, 2007). 7x,24-Dihydroxycholesterol is a 3f3-HSD substrate that results in formation of 7a,24-dihydroxy-4-cholestene-3-one (Schwarz et al., 2000). This intermediate is thought to be converted into chenodeoxycholic acid. The enzymes involved in these final steps have not been identified (Russell and Setchell, 1992; Norlin and Wikvall, 2007). The 25-hydroxylase pathway involves simultaneous hydroxylation of 53-cholestane- 3a,7cL, 12o-triol (an intermediate formed in the neutral pathway) by CYP27A1, as well as by CYP3A4ICYP3A5. The 25-hydroxylation of 513-cholestane-3cx,7cz,l2cL-triol was shown to be catalyzed by CYP3A4 in human liver (Shefer et al., 1976; Furster and Wikvall, 1999). CYP3A4-mediated hydroxylation yields 5 3-cholestane-3ci,7c, 1 2a,24,25-pentol which is converted into cholic acid by unidentified soluble enzymes (Shefer et al., 1976; Furster and Wikvall, 1999; Norlin and Wikvall, 2007). In the 27-hydroxylase pathway, 4-cholesten-3-one, a cholestanol precursor, acts as a substrate of CYP27A1 in the formation of 7cL,27-dihydroxy-4-choleten-3-one. It has been 6 recently shown that 7cL,27-dihydroxy-4-cholesten-3-one may be further converted into bile acids (Norlin et al., 2003). Cholestanol is known to be degraded to 5cx-bile acids (allocholic and allochenodeoxycholic acid) by liver enzymes similar to those involved in biosynthesis of 513- bile acids. Allo-bile acids are minor constituents in mammalian bile ranging from 1% or less in human to more that 5% in rabbits and reptiles (Bjorkhem, 1985). Though the relative contribution of various pathways for bile acid biotransformation is not fully assessed, the neutral pathway is considered quantitatively to be the most important pathway for formation of primary bile acids in humans (Bjorkhem and Eggertsen, 2001). Studies in humans reveal that endogenous production of 27-hydroxycholesterol by the acidic pathway amounted to only 9% of total bile acid formation (Duane and Javitt, 1999). It appears that the acidic pathway can compensate for and maintain bile acid formation under conditions in which CYP7A 1 in the neutral pathway is repressed, as in case of liver disease (Axelson and Sjovall, 1990). It is suggested that extrahepatic 27-hydroxylation of cholesterol in the acidic pathway may play an important role in cholesterol elimination in vascular endothelium and macrophages. Some evidence indicates that the acidic pathway may play an important role in human infants (Setchell et al., 1998). However, in rats, the acidic pathway contributes as much as 50% of total bile acid biosynthesis (Vlahcevic et al., 1999). It has been estimated that only 5% of the total bile acids are formed by the 25- hydroxylase pathway in healthy humans (Duane et al., 1988). However, when the 27- hydroxylase pathway for side chain oxidation is blocked as in patients with the inherited lipid storage disease, cerebrotendinous xanthomatosis, the 25-hydroxylase pathway may be the only possibility for bile acid formation. The 27-hydroxylase pathway involving cholestenone accounts for less than 1% of total bile acid formation in humans (Bjorkhem and Eggertsen, 2001). 7 1.2.2 Biosynthesis of secondary bile acids. Secondary bile acids (lithocholic acid and ursodeoxycholic acid) are formed in the colon from primary bile acids by bacterial dehydratases. The 7a-hydroxyl group of primary bile acids is attacked by anaerobic bacteria in the colon. Pure cultures isolated from the human intestinal flora and screened for their ability to remove the 7c-hydroxyl group from cholic acid and chenodeoxycholic acid show the involvement of Strepotococcus faecalis, Eubacterium species, Clostridium bfermentans, Veillonella species and unidentified Gram-positive anerobic bacteria (Hill and Drasar, 1968; Ferrari et al., 1977; Ferrari et al., 1980; Hirano et al., 1981; Pacini et al., 1987; Bortolini et al., 1997). Bacterial dehydratases in cultures of Eubacterium species V.P.I. 12708 increase cholic acid and chenodeoxycholic acid 7a-dehydroxylation by 25- to 46-fold as compared to the ones mentioned earlier (White et al., 1980). By this process, cholic acid, which is a trihydroxylated (C-3, C-7 and C-12) bile acid, is converted to deoxycholic acid with hydroxyl groups at C-3 and C-12. Similarly, 7-dehydroxylation of chenodeoxycholic acid results in formation of the monohydroxy bile acid, lithocholic acid, which contains one hydroxyl group at C-3. Ursodeoxycholic acid is another secondary bile acid that is present in human liver in trace amounts. It is formed in the distal small intestine or the large intestine by bacterial epimerization of the hydroxyl group at the C-7 position of chenodeoxycholic acid (Hofmann, 1999b; Ridlon et al., 2006). Epimerization of the 7a-hydroxyl group of chenodeoxycholic acid is catalyzed by mixed microbial cultures found in the human colon and feces including Clostridium perfringens HS- 10, Clostridium butyricum, Clostridium absonum, Escherichia coli, and Eubacterium lentum (Macdonald and Hutchison, 1982; Huang et al., 1991; Bortolini et al., 1997). 8 1.3 Biliary Bile Acids: Composition And Physiological Relevance Bile acids are the major components of bile, which facilitates digestion and serves as a vehicle for excretion of bilirubin, drugs and toxins. Bile is a complex alkaline solution consisting of inorganic and organic compounds including bile acids, bilirubin, cholesterol, phospholipids, and proteins such as albumin and immunoglobins. Cholic acid (35%) and chenodeoxycholic acid (35%) are the major bile acids in human bile (Reddy and Wynder, 1977; Reddy, 1981; Vlahcevic et al., 1996; Ridlon et al., 2006). Deoxycholic acid, lithocholic acid and ursodeoxycholic acid are minor bile acids (< 10%) (Yanagisawa et al., 1980). Rat bile consists primarily of cholic acid (42%) and muricholic acids (28-32%), followed by chenodeoxycholic acid (9%), ursodeoxycholic acid (7-8%) and other minor bile acids (15%) (Rost et al., 2003). In mice, the major bile acids are cholic acid (—60%) and muricholic acids (—30%) (Wang et al., 2001a) (Fig. 1.5). Bile acids perform several key functions. The first and most important function is the elimination of cholesterol. In the liver, cholesterol is converted to bile acids, which in turn undergo micellar solublization with cholesterol, enabling cholesterol to move from the hepatocyte to the intestinal lumen, ultimately leading to its elimination via the fecal route (Jezequel et a!., 1994; Erlinger, 1997; Hofmann, 1999b). A second important function of bile acids is lipid transport. In the small intestine, bile acids promote absorption of dietary lipids by forming mixed micelles. Such mixed micelle formation accelerates lipid absorption. Unless bile acids are present in micellar form, fat-soluble vitamins (A, D, E, and K1) are not absorbed, and a vitamin deficiency occurs. The third and fourth functions of bile acids are stimulation of bile flow and stimulation of biliary phospholipid secretion. Bile acids are actively transported from the hepatocytes into the biliary canaliculi and, induce bile flow by their osmotic properties. Bile acids promote the transfer of phospholipids from the canalicular membrane into bile. The presence of phospholipids in bile results in a greater fraction of bile acids existing in the form of 9 mixed micelles and a lower monomer concentration of bile acids, thereby preventing bile acids from damaging the bile duct epithelium. A fifth function is the negative feedback regulation of bile acids and cholesterol biosynthesis. The concentration of bile acids in the hepatocyte seems to act as a signal: when hepatocyte bile acid concentration is high, hepatic bile acid synthesis is low and when hepatic bile acid concentration decreases, bile acid synthesis increases up to 15- fold. Bile acids have additional functions in the intestine. Bile acids solubilize polyvalent metals such as iron and calcium in the duodenum, promoting their absorption. Bile acids also stimulate the release of motilin, which coordinates the interdigestive migrating motility complex. Bile acids have bacteriostatic effects and stimulate mucin secretion; these actions are likely to affect intestinal flora of the small intestine. Bile acids, because of their surface activity, could inhibit bacterial adhesion, and bind entero-toxin in the intestinal lumen. Bile acids also affect the absorption of water and electrolytes by the colonic mucosa and affect colonic motility (Hofmann, 1999b). 1.4. Bile Acids: Enterohepatic Circulation After their formation in liver, bile acids are secreted into the bile, which is stored in the gall bladder and released into the intestine. A very small percentage of these bile acids is excreted (--5%). The rest is re-circulated back to the liver. The entire process of bile acid re circulation is mediated by specific bile acid transporters. Unconjugated bile acids are weak acids that are uncharged at physiological pH and thus can traverse cell membranes by passive diffusion. Unconjugated bile acids undergo conjugation with glycine and taurine in the liver before being secreted into bile ducts and entering the gall bladder. Bile acids conjugated with glycine or taurine, i.e. bile acids in salt form, are predominantly anionic, have lower pKa values and require active transport mechanisms for cellular uptake. Amino acid conjugated bile salts 10 are secreted from hepatocytes across the canalicular membrane into bile canaliculi by bile salt export pump (BSEP), an ATP-binding cassette transporter that is also known as sister of P glycoprotein (SPGP) (Wang et al., 2001a; Wang et al., 2001b; Wang et al., 2003). Glucuronide and sulfate bile acid conjugates together with anionic conjugates, are excreted via the canalicular conjugate export pump (MRP2). The phospholipid export pump (MDR3) facilitates excretion of phosphatidyicholine, which forms mixed micelles in bile together with bile salt and cholesterol. Other basolateral forms of the multidrug resistance-associated protein (MRP 1 and MRP3) provide an alternative route for the elimination of bile salts and non-bile salt anionic conjugates from hepatocytes into the systemic circulation. Bile salts are reabsorbed in the terminal ileum via ileal Na-dependent bile salt transporter (ISBT). In the intestine, primary bile salts (cholate and chenodeoxycholate) are converted into toxic secondary bile salts such as lithocholate and deoxycholate. Secondary bile salts are effluxed from the intestine by MRP3. Similar mechanisms exist in proximal renal tubules and cholangiocytes where an additional, truncated isoform (t-ISBT) may be involved in bile salt efflux from cholangiocytes. The bile acid efflux transporter MRP2 is also present in the apical membrane of enterocytes and proximal renal tubules, while MDR1 is also found in intestine and bile ducts (Trauner and Boyer, 2003). Most effluxed bile salts (--95%) are recycled through reabsorption in the ileum and are returned to the liver via the portal vein. Bile salts are effluxed from the ileum and enter the hepatocytes with the help of the sodium taurocholate co-transporting polypeptide (NTCP). The bile salts effluxed from the cholangiocytes enter the hepatocyte with the help of organic anion transport proteins (OATP). This entire process is termed enterohepatic circulation (Fig. 1.6). 11 1.5 Bile Acid Toxicity The micellar properties of bile acids cause them to be powerful solubilizers of membrane lipids and are responsible for bile acids being cytotoxic when present at high concentrations. The cytotoxic and micellar properties of bile acids are attributed to their structure. A space- filling model of conjugated cholic acid is shown in Fig. 1.7. The conjugated bile acid molecule is relatively planar. It has a hydrophobic face that consists of non-hydroxyl substituents, primarily the steroidal backbone, and a hydrophilic face made up of hydrophilic substituents such as the hydroxyl groups, the amide carbonyl and the ionized acidic group of glycine or taurine (Fig. 1.7). Enterohepatic circulation of bile acids maintains bile acid homeostasis. The efficient efflux of bile acids by BSEP and MRP2, keep bile acid concentrations within the hepatocyte extremely low (< 3 tM). In healthy individuals, intracellular bile acid uptake is followed by rapid elimination. With impaired elimination, bile acids accumulate within the hepatocyte and the bile acid-binding capacity of bile salt transporters is exceeded (Hofmann, 1 999a). The amphiphatic structure enables bile acids to enter other organelles, damaging cell membranes and impairing their functions. Within the hepatocytes, bile acids are known to induce apoptosis and necrosis by damaging the mitochondria (Roberts et al., 1997; Rodrigues et al., 1 998a; Rodrigues et al., 1998b; Hofinann, 2002). In rodents and in humans, increasing concentrations of lithocholic acid, chenodeoxycholic acid, taurochenodeoxycholic acid, glycochenodeoxycholic acid and deoxycholic acid induce mitochondrial permeability transition (a biomarker for mitochondrial damage), release of cytochrome c, increase hydroperoxide generation, impair electron flow on the respiratory chain, and thus stimulate reactive oxygen species leading to cell death (Sokol et al., 1995; Rodrigues et al., 1998b; Rob et al., 2000; Palmeira and Rob, 2004; Sokol et al., 2005). Hepatic cells (HepG2) exposed in vitro to 12 concentrations of chenodeoxycholic acid as low as 50 tM have been shown to undergo significant inhibition of cell proliferation, cytochrome c release, caspase-9 activity, and induction of membrane permeability transition eventually leading to apoptotic cell death (Rob et al., 2004). Within the hepatocyte nuclei, lithocholic acid induces DNA strand breaks, forms DNA adducts, and inhibits DNA repair enzymes (Hamada et al., 1994; Nagengast et al., 1995; Ogawa et al., 1998). The number of hydroxyl groups on the bile acid molecule is inversely related to toxicity so that the fewer the number of hydroxyl groups, the higher the toxicity. The mono hydroxy bile acid, lithocholic acid, is the most hydrophobic cytotoxic bile acid. The dihydroxylated bile acids, chenodeoxycholic acid and deoxycholic acid, bind tightly to tissue, are hydrophobic and hence, highly cytotoxic. The trihydroxy bile acid, cholic acid, has intermediate hydrophobicity and is non-cytotoxic at low concentrations, but cytotoxic at very high concentrations (80-120 nmollg liver) (Scholmerich et al., 1984; Quist et al., 1991; Hofmann, 1999b). 1.6 Cholestasis And Bile Acid Toxicity Cholestasis, defined as impairment of normal excretion of bile into the duodenum, is a manifestation of many diseases. Inborn errors of bile acid biosynthesis, impaired canalicular bile acid secretion, physical obstruction (gall stones) to bile flow and autoimmune destruction of cholangiocytes causes bile acids to accumulate within the liver (See Fig. 1.8). Impaired excretion coupled with bile acid accumulation in the liver promotes liver cell injury due to the inherent cytotoxic properties of bile acids leading to apoptosis and necrosis. Normal bile functions in the intestine such as absorption of fat-soluble vitamins and dietary triglycerides are disrupted. Cholestasis can be acute or chronic, and extrahepatic or intrahepatic. Examples of 13 cholestatic liver disease in adults are primary biliary cirrhosis, primary sclerosing cholangitis, intrahepatic cholestasis of pregnancy and drug-induced cholestatic liver injury, which is a relatively common cause of cholestasis. There is no curative treatment for many chronic cholestatic liver diseases. Treatment is limited to slowing the progression of the disease, controlling symptoms of these diseases, bile acid replacement therapy, or liver transplantation. The most common form of cholestasis is drug-induced cholestasis. Anabolic steroids, non steroidal anti-inflammatory drugs such as nimesulide, antibiotics such as erythromycin, barbiturates, benzodiazepines, neuroleptics such as phenothiazines, angiotensin converting enzyme inhibitors such as fosinopril, antihyperglycemics such as troglitazone, and certain antifungals have been reported to block the bile flow to the gall bladder and ultimately the gut inducing cholestasis (Jezequel et al., 1994). Cyclosporin A, rifampicin, and glibenclamide inhibit BSEP-mediated bile salt transport to the intestine (Stieger et al., 2000). Drug-induced cholestasis may lead to abdominal pain and dark urine (Erlinger, 1997). Cholestasis is also associated with total parenteral nutrition, which is very common for hospitalized patients. Total parenteral nutrition is a practice of feeding a person intravenously, bypassing the usual process of eating and digestion. Complete inactivity of the gastrointestinal tract results in bile stasis in the gallbladder and subsequently in the liver. Cholestasis during pregnancy is also relatively common and is due to the increased sensitivity of bile ducts to estrogen. The estrogen metabolite, estradiol- 1 73-glucuronide inhibits ATP-dependent taurocholate transport of hepatocellular BSEP and MRP2 in rat, thus suggesting the molecular basis of etiology of cholestasis in pregnant women (Stieger et al., 2000). Cholestasis develops during the second and third trimesters of pregnancy and disappears within two to four weeks after the infant’s birth but may reappear again during subsequent pregnancies. The risk of developing pregnancy-induced cholestasis tends to run in families (Hofmann, 1 999b; Hofmann, 2002). 14 Congenital defects involving abnormalities affecting bile acid biosynthesis are less common with fewer than 50 cases identified so far. Inborn errors in bile acid biosynthesis involve deletion or mutations in genes encoding enzymes (CYP7A1, CYP7B1, 313-HSD and CYP27A1) involved in formation of bile acids (See Fig. 1.4). Truncation of the CYP7AI gene results in no enzyme activity leading to high levels of low density lipoprotein cholesterol in the CYP7A1-deficient adult. Individuals with this mutation are found to be hyperlipidemic (Pullinger et al., 2002). Setchell et al. (1998) reported a newborn child with a CYP7BI gene mutation leading to lethal neonatal cholestasis. The patient had high levels of 24-, 25- and 27- hydroxycholesterol and excreted unsaturated monohydroxy bile acids. This patient had a normal CYP7A1 gene. The finding that the CYP7BI gene defect was lethal indicates that CYP7B1 in humans may be important in neonatal development (Setchell et al., 1998). Deficiency in the gene encoding 313-HSD enzyme leads to clinical manisfestations during childhood that includes jaundice, fat malabsorption, fat-soluble vitamin deficiency and progressive cholestasis. These individuals exhibit defective formation of primary bile acids in the liver and accumulation of 3f3-hydroxylated-z5bile acids in blood and urine (Schwarz et al., 2000). Several cases of an inborn defect in 3f3-HSD gene have been reported (Shneider et al., 1994). Oral bile acid substitution therapy is the only cure (Bove et a!., 2000). Defects in bile acid side chain biosynthesis are primarily due to a CYP27A 1 gene mutation. A defect in expression of CYP27A1 presents as cerabrotendinous xanthomatosis, a rare neurologic disease. Since the hydroxylation at C-27 of cholesterol is blocked, various C-25 and C-27 hydroxylated bile alcohols are produced resulting in accumulation of bile alcohols, cholestanol, atherosclerosis, osteoporosis and neurologic impairment (Cali et a!., 1991; Moghadasian et al., 2002; Norlin et al., 2003). Cerabrotendinous xanthomatosis can be successfully treated with oral bile acid therapy (Bove et a!., 2000). 15 A less common but severe and chronic form of cholestasis observed in children involving impaired canalicular secretion of bile acids is progressive familial intrahepatic cholestasis or Byler’s disease (1 in 7000 births). It occurs due to a congenital abnormality in the transport of bile acids due to defects in transporter genes ATP8B1, ABCBI1, and ABCB4. These genes encode the bile acid transporters, BSEP, MDR3 and familial intrahepatic cholestasis (FTC 1). The absence of any one of these transporters leads to cholestatic bile acid retention in the hepatocyte causing necrosis, apoptosis and inflammation. Infants with these symptoms show severe jaundice. The primary symptoms of cholestasis include itching, jaundiced skin or eyes, nausea, vomiting and white stools. Pruritis, a metabolic bone disease, and fatigue are other manifestations of this disorder that contribute to a poor quality of life for patients (Glasova and Beuers, 2002). Fatigue, jaundice, hepatomegaly, splenomagaly and hyperpigmentation are also observed as symptoms of chronic cholestatic liver disease. Chronic fatigue occurs in >80% of patients with cholestatic liver disease. Cholesterol accumulation, fat malabsorption, and elevated bile acid and alkaline phosphatase blood levels are observed in cholestasis. Chronic liver disease frequently progresses to cirrhosis, which can lead to hepatocellular carcinoma. Ultimately, cirrhosis predisposes to liver failure and is the major reason for liver transplantation. In patients with chronic cholestatic liver disease, levels of cytotoxic bile acids increase in liver. In a study carried out by Fischer et al. (1996), hepatic levels of chenodeoxycholic acid (50-53% of bile acids in normal human liver) were increased four-fold in patients with end— stage cholestasis. Similarly, cholic acid (15% in normal human liver) and lithocholic acid (5% in normal human liver) increased 24-fold and 4-fold, respectively (Fig. 1.9) (Fischer et al., 1996). As shown in Fig. 1.9, the concentration of lithocholic acid in livers of cholestatic patients 16 is approximately 6 pM. Lithocholic acid concentrations of 5-10 pM were reported in livers of cholestatic patients and in rat models of biliary cholestasis (Setchell et al., 1997). Ironically, ursodeoxycholic acid (f3-epimer of chenodeoxycholic acid), although a dihydroxylated bile acid, has gained importance as a hepatoprotective bile acid. The United States Food and Drug Administration has approved it for the treatment of cholestasis. Anti cholestatic effects of ursodeoxycholic acid have been reported in intrahepatic cholestasis of pregnancy, cholestasis associated with total parenteral nutrition, liver disease of cystic fibrosis, cholelithiasis, primary biliary cirrhosis, progressive familial intrahepatic cholestasis and chronic graft-versus-host disease (Spagnuolo et al., 1996; Palma et al., 1997; Hofmann, 1999b). Ursodeoxycholic acid accounts for 3-5% of the bile acid pool in humans. It does not bind to tissue, is more hydrophilic as compared to other secondary bile acids, and is devoid of cytotoxic properties (Scholmerich et a!., 1984; Quist et a!., 1991). Displacement therapy using ursodeoxycholic acid decreases the composition of circulating toxic bile acids. Ursodeoxycholic acid therapy results in a remarkable improvement in liver test results in patients with primary biliary cirrhosis. It decreases the proportion of chenodeoxycholic acid and deoxycholic acid, and upregulates canalicular bile salt transport thus decreasing the hepatocyte bile acid concentrations. These results may be due to the orientation of steroid hydroxyl groups on ursodeoxycholic acid. Ursodeoxycholic acid prevents hepatic and mitochondrial glutathione depletion and oxidation resulting from chronic cholestasis and has shown to increase antioxidant defense mechanisms by glutathione. Modulation of mitochondrial membrane perturbation and inhibition of induction of membrane permeability transition caused by cholic acid, chenodeoxycholic acid and lithocholic acid have been the mechanisms proposed for the hepatoprotective actions of ursodexycholic acid (Palmeira and Rob, 2004; Rob et al., 2004). It is proposed that ursodeoxycholic acid may compete for intracellular transport that promotes 17 uptake of accumulated bile acids into organelles thereby reducing apoptosis and necrosis (Beuers et al., 1998). In cholic acid-induced cholestasis in rats, ursodeoxycholic acid prevented impairment of hepatic function by restoring cholic acid disrupted hepatic transporters such as OATP 1 and OATP2 (Rost et al., 2003). The combination of ursodeoxycholic acid and immunosuppresants is believed to be an efficacious therapy for primary biliary cirrhosis (Wolthagen et al., 1994; Wolthagen et al., 1998). 1.7 Nuclear Receptors In Bile Acid Regulation Nuclear receptors are involved in regulating primary bile acid synthesis, transport, and enterohepatic bile acid circulation in the liver and within the intestine. Nuclear receptors play a role in cholesterol and bile acid regulation as sensors to maintain normal homeostasis. Famesoid X receptor (FXR), liver X receptor (LXR), pregnane X receptor (PXR), vitamin D receptor (VDR) and constitutive androstane receptor (CAR) have been identified as bile acid receptors that regulate bile acid metabolism. Xie et al. (2004) propose that specific and overlapping functions of these nuclear receptors in combination with hepatic and intestinal transporters, P450 enzymes and conjugating enzymes increase the complexity of liver metabolism and suggest the existence of functionally redundant defenses against bile acid toxicity (Fig. 1.10). A major role of FXR as a bile acid receptor has been proposed (Makishima et al., 1999). FXR was originally identified as a receptor activated by famesol, an intermediate in cholesterol synthesis. Bile acids appear to be natural ligands for the nuclear receptor FXR. FXR is expressed in liver, intestine, and kidney. Chenodeoxycholic acid is the most potent natural FXR agonist (EC50 7 j.tM) (Tu et al., 2000; Makishima et al., 2002). FXR is also activated by lithocholic acid and its 3-keto metabolite, 3-ketocholanoic acid (Makishima et al., 2002). 18 Synthetic analogues such as 6-ethyl chenodeoxycholic acid and GW4064 have also been shown to activate FXR. In vivo administration of 6-ethyl chenodeoxycholic acid protects rats against cholestasis induced by intraperitoneal administration of estrogen and lithocholic acid (Fiorucci et al., 2005; Rizzo et al., 2005). GW4064, has proven to be hepatoprotective in rat models of intra- and extrahepatic cholestasis (Liu et al., 2003). Upon activation, FXR binds to its heterodimer, retinoid X receptor (RXR), and regulates P450 enzymes involved in cholesterol catabolism and bile acid biosynthesis (Grober et al., 1999). Activated FXR mediates the feedback suppression of bile acids by down-regulating cholesterol 7c-hydroxylase (CYP7A1), the rate-limiting enzyme in bile acid biosynthesis from cholesterol. FXR also participates in the activation of intestinal bile acid binding protein (IBABP), which is involved in the enterohepatic circulation of bile acids (See Fig. 1.11). FXR gene knockout mice have impaired resistance to bile acid-induced hepatotoxicity (Sinal et al., 2000). Thus, FXR constitutes a potential therapeutic target that can be modulated to enhance the removal of cholesterol from the body and alleviate cholestasis. All these studies show that FXR is involved in bile acid regulation and in protection against bile acid toxicity (Xie et al., 2001; Ekins et al., 2002). Lithocholic acid and its 3-keto metabolite (3-ketocholanoic acid) are PXR ligands. Lithocholic acid activates PXR at a concentration 100 iM (Staudinger et al., 2001; Makishima et a!., 2002). Upon entering the cell, lithocholic acid directly activates PXR in the nucleus. The activation triggers the binding of PXR to its heterodimer RXR. The PXRIRXR heterodimer binds to the xenobiotic response elements and increases the transcription of CYP3A genes, which leads to induced expression of CYP3A enzymes and hence, increased metabolism of xenobiotic and endogenous substrates including lithocholic acid (See Fig. 1.12) (Mangelsdorf and Evans, 1995; Ekins et a!., 2002). The PXR knockout mouse model exhibits heightened levels of lithocholic acid toxicity compared with wild-type mice (Xie et al., 2001). PXR protects 19 against liver damage induced by lithocholic acid and thus serves as a physiological sensor of lithocholic acid (Staudinger et al., 2001; Xie et al., 2001). PXR is implicated in regulating the expression of genes involved in the biosynthesis, metabolism and transport of bile acids (Schuetz et al., 2001; Staudinger et al., 2001; Xie et al., 2001; Chiang, 2002; Chiang, 2004; Stedman et al., 2004) and in this sense acts as a physiological bile acid sensor. PXR also represses the CYP7A1 gene involved in the synthesis of bile acids from cholesterol. Furthermore, induction of CYP3A1 1 and OATP2 gene expression is observed in wild-type mice as compared to PXR knockout mice upon lithocholic acid treatment, indicating that PXR functions as a receptor for lithocholic acid and regulates lithocholic acid metabolism in vivo (Staudinger et al., 2001). PXR and VDR are capable of sensing the concentration of secondary bile acids, including lithocholic acid, and inducing their metabolism in the liver and intestine (Staudinger et al., 2001; Xie et al., 2001; Makishima et al., 2002; Matsubara et al., 2008). A key PXR and VDR target is the CYP3A gene which converts lithocholic acid to the non-toxic hyodeoxycholic acid (Makishima, 2005). It has also been shown that VDR is activated at a lithocholic acid concentration (EC50 8 .tM) at least 10 times less than that required to activate PXR. The mechanisms by which nuclear receptors regulate bile acid levels are summarized in Fig. 1.11. CAR is a xenobiotic receptor that normally resides in the cytoplasm of untreated mouse liver and primary rat hepatocytes. Selective androstane metabolites (e.g. 3cc, 5cL-androstanol) act as inverse agonists of CAR activity, which can be reversed by exposure to xenobiotics. Upon exposure to phenobarbital (PB), CAR is translocated from cytoplasm to the nucleus by phosphorylation dependent mechanisms (Kawamoto et al., 1999). There is no evidence that PB binds directly to CAR, but PB-induced activation of CAR leads to its heterodimerization with RXR (Fig. 1.12) (Handsohin and Meyer, 2003; Maglich et al., 2003). The heterodimer binds to 20 phenobarbital response elements (PBRE) found in promoters of PB-inducible CYP2B genes within the nucleus and increases CYP2B expression (Wilson and Kliewer, 2002). Though PB activates CAR in mice and humans, the human isoform of CAR differs from its murine counterpart. For instance, the chemical 1 ,4-bis[2-(3,5-dichloropyridyloxy)]benzene (TCPOBOP), estradiol and estrone are CAR activators in mouse but hardly bind to human CAR and have no effect on CYP2B levels in man (Kawamoto et al., 2000; Moore et al., 2000). Similarly, 6-(4-chlorophenyl-imidazo[2, 1-b] [1 ,3]thiazole 5-carbaldehyde O-(3,4- dichlorobenzyl)oxime, is a human CAR agonist but does not affect mouse CAR (Maglich et al., 2003). Differences in activation of human and mouse CAR are most likely due to the divergent ligand binding domain of CAR orthologs from these species (Moore et al., 2000). The activation of CAR also induces cytosolic sulfotransferase (SULT) thereby increasing lithocholic acid sulfation to reduce hepatotoxicity of Lithocholic acid. CAR regulates SULT expression by binding to the CAR response elements found within the SULT gene promoters (Saini et al., 2004). This function of CAR was also observed to be independent of CYP3A (Saini et al., 2004). A comparison of responses of PXR and CAR to lithocholic acid treatment demonstrates that CAR predominantly mediates induction of CYP3A 11 and the MRP3 transporter. CAR is also a major regulator of the OATP2 gene involved in bile acid transport (Staudinger et al., 2003). Several groups have recently demonstrated the existence of cross-talk between CAR, PXR and their target P450 genes (Fig. 1.12) (Handschin and Meyer, 2003). Such reciprocal regulation is accomplished via recognition of each other’s DNA response elements. The PBRE of CYP2B contains two imperfect DR-4 type nuclear receptor binding sites that show affinity for PXR. PXR was also found to regulate CYP2B in cultured cells and in transgenic mice (Xie et al., 2000). In the same manner, CAR can activate CYP3A through PXR/RXR response elements (Moore et al., 2000; Xie et al., 2000; Xie and Evans, 2001). There also are important 21 species differences in terms of crosstalk between CAR and PXR. For example, TCPOBOP activates mouse CAR and human PXR, but not human CAR or mouse PXR. Similarly, clotrimazole can induce human PXR and inhibit human CAR, but does not affect mouse CAR. The cross-regulation of P450 gene classes provides an explanation for the dual activation property of certain compounds including bile acids. This reciprocal regulation of P450 genes by multiple receptors reveals the existence of a metabolic safety network to protect against the toxic effect of xenobiotics and endobiotics. 1.8 Bile Acid Biotransformation By Conjugation Bile acid biotransformation by conjugation and oxidation play a key role in bile acid clearance and detoxification from the body. Bile acids may undergo conjugation on both the steroid nucleus and the side chain (See Figure 1.2). Conjugation reactions involve covalent attachment of an endogenous polar moiety to bile acids leading to formation of highly water- soluble conjugates that are excreted mainly through the kidneys. Conjugation of bile acids primarily involves amidation with glycine or taurine, sulfation and ester (C-24) or ether (C-3) glucuronidation. (Hofmann and Hagey, 2008). Conjugation with glycine or taurine is catalyzed by bile acid-CoA:amino acid N acyltransferase (BACAT). Conjugation of the carboxylic acid group of bile acids with taurine or glycine involves activation of the bile acid by co-enzyme A, which produces an acyl-CoA thioester that reacts with the amino group of the amino acid to form an amide linkage. Human and rat BACAT have been reported to reside in peroxisomes with variable amounts present in the cytosol (Solaas et al., 2000; He et al., 2003; Solaas et al., 2004). Consequently, it has been proposed that peroxisomal BACAT is required for conjugation of de novo synthesized bile salts, whereas the cytosolic pooi of BACAT is required for reconjugation of deconjugated bile salts returning from the intestine to the liver (Pellicoro et al., 2007). 22 Cytosolic sulfotransferases (SULTs) are also important for bile acid detoxification. SULTs catalyze the transfer of a sulfate group from the co-substrate, 3-phosphoadenosine-5- phosphosulfate (PAPS), to the acceptor substrates to form sulfate or sulfamate conjugates. Lithocholic acid is a preferred substrate for SULT2A9 (hydroxysteroid sulfotransferase) (Chen and Segel, 1985; Radominska et al., 1990; Song et al., 2001). Kitada et al. (2003), identified a role of SULT2A in protection against lithocholic acid induced liver damage (Kitada et al., 2003). This study also reported increased levels of SULT2A in FXR knockout lithocholic acid treated cholestatic mice. Sulfated lithocholie acid shows less cytotoxicity than lithocholic acid when exposed to cells or animals (Leuschner et a!., 1977). Hepatic glucuronidation is perhaps the most studied Phase II biotransformation pathway for a wide array of exogenous and endogenous molecules including bile acids. Because bile acids are cytotoxic, their hepatic glucuronidation is a significant step in their sequential metabolism (Pauli-Magnus et a!., 2005). Uridine diphosphate (UDP)-glucuronosyltransferase (UGT) enzymes catalyze the glucuronidation reaction. The reaction involves the transfer of the glucuronosyl group from uridine 5-diphosphoglucuronic acid (UDPGA) to endogenous and exogenous molecules with hydroxyl, amine, suithydryl or carboxylic acid functional groups. Glucuronide conjugation involves either the 3-hydroxyl group or the 24-carboxyl group of the steroid nucleus of bile acids, resulting in the formation of ether-type or ester-type glucuronides, respectively (Ikegawa et al., 1996; Ikegawa et al., 2000). The resulting glucuronide products are more polar, water soluble, generally less toxic and more easily excreted than the substrate molecules (Mackenzie et al., 2005). In humans, 18 UGT proteins were characterized and categorized into two major families, UGTI and UGT2, according to their amino acid sequence similarity (Mackenzie et al., 2005). The most abundant bile acid glucuronide conjugate reported in human plasma is chenodeoxycholic acid glucuronide, followed by lithocholic acid glucuronide (Back, 1976; 23 Takikawa et al., 1983). The plasma concentrations of chenodeoxycholic acid glucuronide is increased by 50-fold and lithocholic acid glucuronide is increased by 10-fold, respectively, in cholestatic patients (Takikawa et al., 1983). Human UGT1A3 was reported to be the major enzyme catalyzing hepatic formation of lithocholic acid-24-glucuronide and chenodeoxycholic acid-24-glucuronide (Trottier et al., 2006), while UGT2B4 and UGT2B7 play important roles in glucuronide conjugation of hyodeoxycholic acid (Pillot et al., 1993; Gall et al., 1999). 1.9 Bile Acid Biotransformation By P450 Enzymes The P450 enzyme superfamily consists of a large number of enzymes that form the predominant biotransformation pathway in the mammalian body (Fig. 1.13). Table 1.2 compiles the various P450 subfamilies and enzymes, along with their P450 content and their relative contribution to the total P450 enzymes found in human and rat (Shimada et al., 1994; Lewis et al., 1999; Bandiera, 2001; Lewis, 2001; Paine et al., 2006; Hrycay and Bandiera, 2008). Fifty- seven human P450 genes have been identified to date (Stark and Guengerich, 2007). More importantly, P450 enzymes are involved in the biotransformation of lipophilic endogenous compounds such as steroids, arachidonic acid, retinoids and bile acids, which perform important functions in cellular physiology (Guengerich, 2003; Stark and Guengerich, 2007). Studies on biotransformation of bile acids date back to the early 1 950s. Early studies incubated cholic acid, deoxycholic acid and lithocholic acid with rat liver homogenates (Bergstrom and Norman, 1953; Bergstrom et al., 1953a; Bergstrom et al., 1953b). Formation of several unidentified bile acid metabolites and relatively less sensitive techniques such as thin layer chromatography, to identi1’ the structures of closely related bile acid metabolites hampered these earlier investigations. Identification of hepatic microsomal P450 enzymes as 24 potential catalytic agents for oxidation by Klingenberg in 1958 focused the need to investigate hepatic microsomal bile acid metabolism (Klingenberg, 1958). Probably, the first in-depth analysis of lithocholic acid biotransfonnation using rat hepatic microsomes was carried out by Zimniak et al. (1989). Hepatic microsomal metabolism of lithocholic acid by male Sprague-Dawley rats showed murideoxycholic acid as the major metabolite followed by 6-ketolithocholic acid, chenodeoxycholic acid, cz-muricholic acid and 13- muricholic acid, respectively. These metabolites were identified using thin layer chromatography techniques (Zimniak et al., 1989). Though some lithocholic acid metabolites were identified, the P450 enzymes involved in its biotransformation were not studied. The first study to identify the involvement of hepatic P450 enzymes in bile acid biotransformation by hamster liver was performed in the early 1 990s (Chang et al., 1993). Biotransformation of lithocholic acid, to form its 613-hydroxylated metabolite, murideoxycholic acid, was attributed to hamster CYP3A 10 by Chang et al. (1993). Lithocholic acid metabolites were identified using thin layer chromatography techniques. The Vm value for 613- hydroxylation was reported as 164 pmollmin per mg protein with a Km of 25 M (Chang et al., 1993). Hepatic microsomal metabolism of lithocholic acid by male Sprague-Dawley rats also supported the formation of murideoxycholic acid as the major metabolite along with 13- muricholic acid (Dionne et al., 1994). In an attempt to identify human liver enzymes catalyzing the hydroxylation of bile acids, recombinant human P450 enzymes and human liver microsomes were incubated with taurochenodeoxycholic acid and lithocholic acid (Araya and Wikvall, 1999). Metabolite detection was carried out by gas chromatography and mass spectrometry (GC/MS). This study was able to identify only a single 6cL-hydroxylated lithocholic acid and taurochenodeoxycholic acid metabolite. Recombinant CYP3A4 was the only enzyme found to be active in lithocholic 25 acid and taurochenodeoxycholic acid biotransformation (Araya and Wikvall, 1999). A strong correlation was also obtained between 6c-hydroxylation of lithocholic acid, CYP3A4 levels and testosterone 63-hydroxylation. Troleandomycin, an effective inhibitor of CYP3A and testosterone 63-hydroxylation, was shown to inhibit 6-hydroxylation of taurochenodeoxycholic acid almost completely at a concentration of 10 jiM. The apparent Km for 6x-hydroxylation of taurochenodexycholic acid by human liver microsomes was 716 jiM. The 6x-hydroxylated lithocholic acid metabolite, is known as hyodeoxycholic acid. This particular study did not investigate other biotransformation pathways such as formation of the 63-hydroxylated murideoxycholic acid, by previous researchers (Zimniak et al., 1989; Chang et al., 1993). In vitro studies using human liver microsomes carried out by Xie et al. (2001) pointed to hyodeoxycholic acid, followed by murideoxycholic acid and chenodeoxycholic acid, as major metabolites of lithocholic acid. Metabolite detection, isolation and purification was carried out using preparative thin layer chromatography and later subjected to derivatization and subsequent analysis by mass and nuclear magnetic resonance spectrometry (Radominska-Pyrek et al., 1987). Anti-CYP3A antibody totally inhibited lithocholic acid 6a-hydroxylation by human liver microsomes. Recombinant CYP3A4 efficiently hydroxylated lithochollc acid while CYP3A5 was observed to be less active. This study concluded that lithocholic acid hydroxylation by human liver microsomes generated hyodeoxycholic acid as its major metabolite formed primarily by CYP3A enzymes (Xie et al., 2001). Recent studies carried out by Bodin et al. (2005) were focused on bile acid metabolism by recombinant CYP3A4 only. Metabolite analysis was carried out using GC/MS. Metabolism of lithocholic acid by recombinant CYP3A4 identified 3-ketocholanoic acid as a major metabolite followed by hyodeoxycholic acid and two new minor metabolites, namely, 6a- hydroxy-3-oxo-5f3-cholanoic acid and 1f3-hydroxy lithocholic acid (Bodin et al., 2005). 26 However, chenodeoxycholic acid and murideoxycholic acid, the CYP3A4 metabolites identified previously by Xie et al. (2001), were not detected. In vivo studies on lithocholic acid biotransformation have identified 3-ketocholanic acid and isolithocholic acid from human bile (Norman, 1964; Norman and Palmer, 1964; Whitney et al., 1983). The lithocholic acid metabolites identified in different studies and from various species are shown in Table 1.1. Studies on biotransformation of other bile acids such as cholic acid and chenodeoxycholic acid have also been carried out but are limited. Biotransformation of cholic acid was studied in urine of patients with intrahepatic cholestasis using radiolabeled cholic acid. The radioactive labeled and derivatized metabolites detected by GC/MS from the urine of patients include 113,2f3,6cL-hydroxy cholanoic acid, acid; 1 1 2c-tetrahydroxy-cholanoic acid, 3cL,6cL-dihydroxy- I 2cL-acetoxy-7-oxo-5 3- cholanoic acid, 3-hydroxy,12-ketocholanoic acid and deoxycholic acid (See Table 1.1) (Bremmelgaard and Sjovall, 1980). Recombinant CYP3A4 metabolizes cholic acid to 3- dehydrocholic acid as its major product (Bodin et al., 2005). Recently, a new metabolite, 3cL,6,7,12cL-tetrahydroxy-53-cholanoic acid, was identified in the SPGP gene knockout mouse cholestatic model. Tetrahydroxylated bile acid levels were increased in these SPGP knockout mice (Perwaiz et al., 2003). Characterization of the P450 enzymes involved in the formation of these tetrahydroxy cholic acid biotransformations remains to be investigated. The biotransformation of‘4C-labeled chenodeoxycholic acid was studied in patients with intrahepatic cholestasis, by Bremmelgaard and Sjovall (1980). The14C-labeled metabolites were isolated from urine and were identified after subsequent derivatization using gas-liquid chromatography-mass spectrometry. Chenodeoxycholic acid was metabolized to hyocholic acid, x-muricholic acid, taurolithocholic acid, and taurohyodeoxycholic acid as major metabolites (Bremmelgaard and Sjovall, 1980). Recently, in vitro studies performed by Bodin et al. (2005) 27 identified three new metabolites of chenodeoxycholic acid, namely, 3cL,7cL-dihydroxy-3-oxo-5f3- cholanic acid (major), 1 3-hydroxy- and 22-hydroxychenodeoxycholic acid (minor) following incubation with recombinant CYP3A4 (Bodin et al., 2005). The various hydroxylated metabolites of chenodeoxycholic acid obtained from the literature are compiled in Table 1.1. The major metabolites of deoxycholic acid obtained from patients with intrahepatic cholestasis identified using GC/MS were 1 13,3 CL, 1 2ct-trihydroxy-5 13-cholanoic acid, 6ct-hydroxy deoxycholic acid, 3x,6CL, 12c-trihydroxy-513-cholic acid and 1 3-hydroxy-deoxycholic acid (Bremmelgaard and SjovalL, 1980). Recently, 3-dehydrodeoxycholic acid and I 3-hydroxy- deoxycholic acid were identified as major metabolites of deoxycholic acid formed by recombinant CYP3A4 (Bodin et al., 2005). It should be noted that few studies of hepatic microsomal biotransformation of lithocholic acid, cholic acid, chenodeoxycholic acid and deoxycholic acid have been carried out. Conflicting reports of bile acid metabolites and P450-mediated pathways involved in their formation have also emerged. 1.10 Rationale Cholestasis, defined as impairment of the normal excretion of bile into the duodenum, leads to accumulation of inherently toxic bile acids in the liver. Chronic cholestatic liver disease frequently progresses to cirrhosis, which can further lead to conditions such as hepatocellular carcinoma. Ultimately, cirrhosis predisposes to liver failure and is the major reason for liver transplantation. Chronic liver disease and cirrhosis are currently the twelfth leading cause of death in the U.S. and Canada, with an age-adjusted death rate of 9.5 per 100,000 population (Vong and Bell, 2004; Aspinall and Adams, 2006). The economic impact of these diseases is 28 high because liver diseases tend to strike adults in their most productive phase of life. Health costs associated with liver disease are considerable in terms of health care required for complications and costs of liver transplantation. There is also an impact on the quality of life of patients in terms of the ability of patients to work and engage in social activities. Considering the socio-economic impact of accumulated bile acids during cholestasis, efficient detoxification pathways for elimination of bile acids need to be investigated. Cholestasis leads to accumulation of chenodeoxycholie acid, cholic acid and lithocholic acid as the major potentially toxic bile acids in human liver (Fig. 1.8 and 1.9). Enzymatic pathways leading to elimination of these bile acids need to be identified. Previously, only a few studies identified the involvement of microsomal P450 enzymes in bile acid biotransformation. Conflicting data and incomplete analysis of bile acid metabolites and the enzymes involved in their formation have been reported. With the exception of recent studies carried out by Bodin et al. (2005), using recombinant CYP3A4 to study bile acid metabolism, no other focused study to identify bile acid metabolites and to investigate P450 enzymes involved in their formation has been carried out to date. The C-24 cholestane steroid backbone structure of bile acids consists of many vulnerable positions for oxidation (Fig. 1.14). Given the wide array of P450 enzymes available in human and rodents (Fig. 1.13 and Table 1.2), and the various positions at which a bile acid could be oxidized, the hepatic microsomal bile acid biotransformation studies carried out so far seem to be restricted by concentrating only on CYP3A-mediated bile acid biotransformation. Knowing that CYP3A enzymes contribute to almost 40% of the total P450 enzymes in humans (Shimada et al., 1994), the involvement of CYP3A-mediated bile acid biotransformation in humans could be easily understood. Bile acids are structurally related to androgens and cholesterol. The role of non-CYP3A enzymes in the biotransformation of testosterone (e.g. CYP2AI, CYP2B, CYP2C9, CYP2CI 1, 29 CYP2CI9) (Ryan and Levin, 1990; Yamazaki and Shimada, 1997) and cholesterol (e.g. CYP7A, CYP8B, CYP27A) have been well documented (Tsuchiya et al., 2005; Norlin and Wikvall, 2007). Thus, a question about the involvement of other major P450 enzymes (e.g. CYP2C and CYP2D) in hepatic bile acid biotransformation remains unanswered and warrants investigation. In rodents such as the rat, CYP2C enzymes contribute to 65% of the total P450 enzymes (Table 1.2). Surprisingly, none of these CYP2C enzymes have been shown to be involved in bile acid biotransformation in rodents. Moreover, cholestatic rodent models to study regulation of bile acids have been used. The results from these studies are often extrapolated to understand the regulation of bile acids in humans. Taking into consideration the variation in percentage of each P450 subfamily in rodents and humans (Table 1.2), an understanding of similarities and differences in hepatic bile acid biotransformation profiles in these species is needed. These various biochemical and physiological considerations of bile acids in cholestatic disorders, highlight these endobiotics as ideal candidates for study in the field of biotransformation. Additionally, the identification of major and minor bile acid metabolites has been limited due to the use of less sensitive techniques such as thin layer chromatography or complex techniques such as GC/MS, which require derivatization and intricate procedures of metabolite extraction. The use of these techniques to isolate and identi1,’ structurally related bile acid acids differing from each other with a minor change in their molecular weight may have resulted in improper identification of these metabolites. Simplified extraction procedures and analytical tools having greater sensitivity combined with the capability of isolating and identifying closely related bile acid metabolites need to be employed. In the present study, the hepatic P450-mediated biotransformation of physiologically important cholestatic bile acids (lithocholic acid, chenodeoxycholic acid and cholic acid) is examined using rat, mouse and human hepatic microsomes. The metabolites formed were 30 identified by a liquid chromatography/mass spectrometry (LC/MS)-based assay. Enzyme kinetic parameters involved in formation of their metabolites were investigated. The P450 enzymes involved in biotransformation were identified using a combination of approaches such as enzyme induction, recombinant enzymes, chemical inhibitors and antibody inhibition studies as applicable. Finally a comparison of the various bile acid biotransformation patterns in humans and rodents was carried out. Knowledge of hepatic bile acid biotransformation generated in this study may eventually help to identify additional pathways as potential targets to develop small molecule therapies that may alleviate cholestatic conditions. 1.11 Hypotheses 1. The major route of hepatic microsomal biotransformation of chenodeoxycholic acid, cholic acid and lithocholic acid by rodent and human liver microsomes is 6f3-hydroxylation of the cholestane ring. 2. Hepatic biotransformation of lithocholic acid, cholic acid and chenodeoxycholic aéid by rodent and human liver microsomes is not exclusively CYP3A- mediated. 31 1.11.1 Specific research objectives 1. Investigate hepatic microsomal bile acid metabolism by: • optimizing and validating a LCIMS-based assay to study hepatic biotransformation of cholic acid, chenodeoxycholic acid, and lithocholic acid in human hepatic microsomes. • identii,’ing the major and minor metabolites of cholic acid, chenodeoxycholic acid and lithocholic acid. • determining enzyme kinetic parameters associated with the formation of major metabolites of cholic acid, chenodeoxycholic acid and lithocholic acid. 2. Characterize the hepatic P450 enzymes involved in bile acid metabolism by: • identifying P450 enzymes involved in the formation of the major and minor metabolites of cholic acid, chenodeoxycholic acid, and lithocholic acid using recombinant P450 enzymes, selective inhibitory antibodies, chemical inhibitors and inducers of P450 enzymes. 3. Study the biotransformation of lithocholic acid in rat and mouse and compare the metabolite profile with lithocholic acid biotransformation in humans. 32 Bile acid Metabolite Trivial name Species/System Research group Chenodeoxycholic acid (5J3-cholanic acid-3a, 7a- diol) 5J3-cholanic acid-3-ol 5-cholanic acid-3c, 6, 7f3-triol 5-cholanic acid-3n, 6a-diol 5-cholanic acid-3a, 613-diol 3oxo 513-cholanic acid 513-cholanic acid-3a, 7c-diol 3ct, 6oxo 513-cholanic acid 5f3-cholanic acid-3oxo-6a-diol 5f3-cholanic acid-I f3, 3(x-diol 513-cholanic acid-3cL, 6cL, 7a-triol 513-cholanic acid-3a, 6J3, 7a-triol 3oxo 513-cholanic acid isolithocholic acid 13-muricholic acid hyodeoxycholic acid hamster rat, rCYP3A4 human Norman and Palmer, 1964 Dionne et al., 1994 Araya and Wikvall, 1999, Xie et al., 2001 Bodin et al., 2005 Dionne et al., 1994; Zimniak et al., 1989 Staudinger et al., 1989, Chang et al., 1993 Xie et al., 2001, Bodin et al, 2005 Sakai et al., 1980 Zimniak et al., 1989 Zimniak et al., 1989, Bodin et al, 2005 Bodin et al, 2005 Bremmelgaard and Sjovall, 1980 Bremmelgaard and Sjovall, 1980 Bodin et al, 2005 Cholic acid 513-cholanic acid-3ct, 12o-diol (513-cholanic acid-3ct, 7a- l2cL- 5f3-cholanic acid-3a, 6a, 73, 12a- tetraol triol) 513-cholanic acid-3a, 6cc, 7a, l2ct- tetraol 5J3-cholanic acid-1, 3a, 7a, 12a- tetraol deoxycholic acid human mice human human Norman and Palmer, 1964 Perwaiz et al., 2003 Bremmelgaard and Sjovall, 1980 Bremmelgaard and Sjovall, 1980 Deoxycholic acid (5J3-cholanic acid-3a, 12a- diol) 5f3-cholanic acid-1f3, 3cz, l2cL-triol 513-cholanic acid-3a, 6cL, 12a-triol 513-cholanic acid-3c, 6a, Ri, 12a- tetraol 5f3-cholanic acid-3oxo, l2cL-ol human, rCYP3A4 human human rCYP3A4 Bremmelgaard and Sjovall, 1980, Bodin et al, 2005 Bremmelgaard and Sjovall, 1980 Bremmelgaard and Sjovall, 1980 Bodin et al, 2005 a cholanoic acid and cholanic acid are used interchaneabIy. * rCYP3A4 — recombinant CYP3A4 Lithocholic acid (513-cholanic acid-3i-ol) human rat human, rCYP3A4* murideoxycholic acid or rat murocholic acid 3-ketocholanic acida chenodeoxycholic acid rat 6-ketolithocholic acid rat rCYP3A4 1 13-hydroxy-lithocholic acid rCYP3A4 hyocholic acid human, pig cL-muricholic acid 3 ketocholanic acid rat rCYP3A4 1 13-hydroxydeoxycholic acid 3-dehydrodeoxycholic acid Table 1.1 Oxidative metabolites of bile acids Specific % Specific %Human P450 Content Total Rat P450 Enzyme Content TotalEnzyme (pmollmg) P450 (pmol/mg) P450 CYP1A (CYP1AI, 1-65 7-18 CYP1A (CYPIAI, 1A2) 5-20 1-3 1A2) CYP2A6 <1-68 2-6 CYP2A (2A1, 2A2) 20-40 3-6 CYP2B6 1-39 <1-7 CYP2B (2B1, 2B2) 5-20 1-3 CYP2C (CYP2C8, 1-100 12-24 CYP2C (CYP2C6, 2C7, 380-650 40-65 2C9, 2C18, 2C19) 2C11, 2C12, 2C13) CYP2D6 1-9 1-3 CYP2D1 28-32 2-4 CYP2E1 7-49 3-9 CYP2E1 60-80 8-10 CYP3A (CYP3A4, 45-147 20-40 CYP3A (CYP3A1, 3A9, 40-100 5-14 3A5, 3A7) 3A18) Not 10 CYP4A 30-3 1 3CYP4A1 I determined (4A1, 4A2, 4A3, 4A8) Table 1.2 Different P450 enzymes and their levels in human and rat hepatic microsomes. Values represent approximate P450 levels reported. Data compiled from (Shimada et al., 1994; Lewis, 1996; Bandiera, 2001; Lewis, 2001; Paine et al., 2006; Hrycay and Bandiera, 2008). 34 Cholesterol Bile acid 21 R4 %%%__%4% 2320 llf_L%1J\ OOH 2 R+T”R3 R2 COOH 24 27 Figure 1.1 Cholesterol and bile acid structure. Bile acids are hydrophobic steroids consisting of a cyclopentanophenanthrene nucleus, composed of saturated rings A, B, C and D and a branched 5-carbon isopentanoic side chain. Conversion of cholesterol to a bile acid involves a possible hydroxylation on the steroid nucleus (e.g. shown at C-7, for chenodeoxycholic acid), modification of the C-17 isooctane side chain to a isopentanoic side chain, epimerization of the C-3 hydroxy group and reduction of the double bond to give 5f3-cholanoic acid structure in which A/B rings exist in cis configuration (Modified from Hofmann and Hagey 2008). 19 18 23 26 17 24 “P I 20 22 :7 OH4 16 15 OH 35 HO 1,0 0 0 Figure 1.2 Chemical structures of unconjugated and conjugated bile acids. The major bile acids in human bile are cholic acid, chenodeoxycholic acid, deoxycholic acid, lithocholic acid and ursodeoxycholic acid. Bile acids undergo conjugation primarily with amino acids (glycine or taurine), uridine diphospho-glucuronosyltransferases and sulfotransferases. Figure shows representative structures of lithocholic acid conjugates. HO Chollc acid Taurolithocholic acid HO acid COOH 0—I r’OH OH HO,1r Lithocholic acid-24-glucuronide HO Deoxycholic acid Lithocholic acid Lithocholic acid-3-glucuronide COOH Ursodeoxycholic acid Sulfolithocholic acid 36 A. Bile acids in human liver Ursodeoxycholic acid Cholic Hyodeoxycholic acidj — - 10% (/7 C. Bile acids in mouse liver Deoxycholic acid 3% Maricholic acids— 57% F Lithocholic acid 5% DeoxychoIic acid 16% Chenodeoxychohc acid 45% 10% Deoxycholic acid 10% / Hyodeoxycholic acid <1% Cheriodeoxycholic acid 4% Cholic acid 31% Chenodeoxycholic acid 1% Cholic acid 3.7% Ursodeoxycholic acid 1% Figure 1.3 Bile acid levels in human, rat and mouse liver. Data compiled from Fischer et al., 1996; Setchell et al., 1997 and Stedman et al., 2004. B. Bile acids in rat liver Lithocholic acid M.iricholic acids 32% Ursodeoxycholic acid 3% 37 cooii HO CHOLEROL HO neutral CYP7A1 HO acid pathway \ CYP7H1 HC(C16< L<COON llcstcrol HO ‘011 HO OH 3HSD 3L7a-dihydtoXy-5-ChOIeszflOiC acid o 0H 3ØJISD’9’L<CHZOH 1.hydroxy-4-choIes(eii-3one o •- OH V 3aJISD 7o,27-dihydroxy4-choJesten3-cne CYP27AI)D< O HO 7a-hydrozy-3-ozo.4cholestcnoic acid7a.12a-dahydroxy- 58-cholcsianc-3a,7a-diol4-cholesten-3-one 5ra_ H9 CYP27AI Ja.HSD CH2O V HcY ‘H ...* -. ‘‘ jCOOH 58-cbolestane-3a,7cz,12a-thol HO OH I T T th I HC” OH CH2O CHENODEOXVCHOLIC ACID dar—LJ ftnal StLP ofsidt H *aãi rkivage HO 5S.choIesane-3cx,7a,12cz.27-teuoi HO OH cHOUC AcID Figure 1.4 Neutral and acidic pathways in bile acid formation. 3 13-HSD, 33-hydroxy-z5C27-ste id dehydrogenase; 3cx-HSD, 3o-hydroxysteroid dehydrogenase; 5 3-reductase, 3-oxo-z4steroid 5 f3-reductase (Taken from Norlin and Wikvall, 2007). 38 A. Bile acids in human bile Lithocholic acid 1% Deoxychollc acid 2% Cholic acid ‘ 35% Other / p,—’ 2% ‘1 Ursodeoxycholic _/ acid 25% B. Bile acids in rat bile Chenodeoxycholic acid 35% cz- and -MurichoIic acids 2832% Deoxycholic C. Bile acids in mouse bile Ursodeoxycholic acid 8% Chenodeoxycholic acid 9% Cholic acid 42-48% cL.j-, and y M uricholic 26% Deoxycholic acid 4% Ursodeoxycholic acid 2% Chenodeoxycholic Figure 1.5 Bile acid levels in human, rat and mouse bile. Data compiled from Reddy and Wynder, 1977; Reddy, 1981; Vlahcevic et al., 1996; Ridlon et al., 2006; Rost et al., 2003; Wang et a!., 2001a. 3% acid £10 66% 39 Hepatobiliary transport systems in liver and extrahepatic tissues in humans. Bile salts (BS) are taken up by hepatocytes via the basolateral NaJtaurocho1ate cotransporter (NTCP) and organic anion transporting proteins (OATPs). Amino acid conjugated BS are excreted via the canalicular bile salt export pump (BSEP). Glucuronidated or sulfated BS together with anionic conjugates (OK) are excreted via the canalicular conjugate export pump (MRP2). The phospholipid export pump (MDR3) facilitates excretion of phosphatidylcholine (PC), which forms mixed micelles in bile together with BS and cholesterol. Cationic drugs are excreted by the multidrug export pump (MDR1). Other basolateral isoforms of the multidrug resistance- associated protein (MRP1 and MRP3) provide an alternative route for the elimination of BS and nonbile salt anionic conjugates from hepatocytes into the systemic circulation. BS are reabsorbed in the terminal ileum via ileal Natdependent bile salt transporter (ISBT) and effluxed by MRP3. Similar mechanisms exist in proximal renal tubules and cholangiocytes where an additional, truncated isoform (t-ISBT) may be involved in BS efflux from cholangiocytes. MRP2 is also present in the apical membrane of enterocytes and proximal renal tubules, while MDR1 is also found in intestine and bile ducts (Taken from Trauner and Boyer, 2003). Cholangiocyt. I4SBT Na -* 8S -oc, MRP3 Figure 1.6 Enterohepatic circulation of bile acids. 40 HYDROPHOBIC SIDE HYDROPHILIC SIDE Figure 1.7 Amphipathic nature of bile acids (Taken from Hofmann, 1999). 3a-OH 7a-OH 12a-OH Amide carbonyl Ionized acidic group of glycine or taurine 41 Transport A Liver Enterohepatic circulation Excretion I’ Intestine Figure 1.8 as cholestasis. Obstruction to the flow of bile acids from the liver to the intestine is termed Cholesterol CYP7AI CYP8BI CV P27A1 Primary bile acids Cholesta ,-- Primary bile acids iF Secondary toxic bile acids - 42 140 120 80 60 40 20 0 Figure 1.9 Comparison of bile acid levels in livers of patients with end-stage cholestasis and normal humans (Modified from Fischer et al., 1996). a, 100 0 E a, ‘I . C-) w • Ursodeoxycholic acid D Lithocholic acid • Chenodeoxycholic acid Deoxycholic acid • Cholic acid End-stage cholestatic human liver Normal human liver 43 Receptors Figure 1.10 Bile acid regulation: A complex process involving various receptors, enzymes, and transporters. CAR, constitutive androstane receptor; CYP, cytochrome P450 enzymes; FXR, farnesoid X receptor; LXR, liver X receptor; MDR, multi-drug resistant protein; PXR, pregnane X receptor; SUET, sulfotransferase enzymes; VDR, vitamin D receptor (Modified from Xie et al., 2004). Bile acid and precursors Ligands (Oxysterols, cholic acid, lithocholic acid) Phase I (e.g. CYPs)Target proteins Physiological Relevance Phase II (e.g. SULT) Transporters (e.g. MDR) Bile acid transport, synthesis, metabolism and elimination 44 Liver Detoxification Intestine Bile acid synthesis is stimulated by liver X receptor (LXR) in rodents. Negative feedback regulation of bile acid synthesis is mediated by farnesoid X receptor (FXR). FXR represses bile acid import in hepatocytes and stimulates their biliary excretion. FXR induces the expression of intestinal bile acid-binding protein. Pregnane X receptor (PXR) and vitamin D receptor (VDR) are involved in detoxification of secondary bile acids (Modified from Makishima, 2005). Excretion Figure. 1.11 Regulation of bile acids by nuclear receptors. 45 OH OH Bile acids such as lithocholic acid (LCA) directly activate pregnane X receptor (PXR) in the nucleus. Constitutive androstane receptor (CAR) is activated upon exposure to phenobarbital (PB). There is no evidence that PB directly binds to CAR. PB triggers translocation of CAR from the cytoplasm to the nucleus with the help of unknown phosphorylating enzymes. Activated PXR and CAR heterodimerize with RXR in the nucleus and bind to their respective response elements, thus increasing transcription of their target genes. The PXRJRXR heterodimer binds to xenobiotic response elements (XRE) which induce CYP3A. Similarly, CAR/RXR heterodimer binds to phenobarbital reponse elements (PBRE) to induce CYP2B. Response elements that are activated by both PXR and CAR allow direct nuclear receptor (NR) cross-talk and reciprocal regulation of P450 genes generating a metabolic safety network to protect against xeno or endobiotic toxicity. Activation of these receptors not only induces P450 enzymes (CYP3A and CYP2B) but also induces the expression of drug transporters such as MRP2 (Modified from Handschin and Meyer, 2003). Fig.1.12 PXR and CAR cross-talk mediated metabolism of lithocholic acid. 46 Substrates Inducers Barbiturates Barbturates Barbiturates Omeprazole Ethanol Rifampicin Rifampin Rifampicn Tobacco Isoniazid Dexamethasone smoke Carbamazepine Figure 1.13 Major hepatic P450 enzymes involved in drug metabolism. Circles reflect the mean size of the pooi of each of the main cytochrome P450 enzymes (P450s) in human liver. The exact pattern will vary among individuals. A few commonly recognized substrates, inhibitors, and inducers of these P450s are indicated (Taken from Guengerich, 2003). TolbutamideMephenytoin PhenytoinOmeprazole Warfarin Caffeine DebrisoquineTheophyline SparteineTacrine Chlorzoxazone Inhibitors Fluconazole Ketoconzaole Furafylline Disultiram Methoxsalen Sulfaphenazole estodene Fluvoxamine Quinidine 47 12a1 p Figure 1.14 Biotransformation of bile acids. Figure shows the conversion of bile acids to their possible hydroxylated metabolites, reactions normally catalyzed by cytochrome P450 (P450) enzymes for steroids. P450 1a/ 21 18 20 23 HO_c1 15 Bile acid NADPH, 02 H 16 cL/f3 3 6a/ 413 Possible bile acid hydroxylation sites 48 1.12 References Araya Z and Wikvall K (1999) 6alpha-hydroxylation of taurochenodeoxycholic acid and lithocholic acid by CYP3A4 in human liver microsomes. Biochim Biophys Acta 1438:47-54. Aspinall AT and Adams DH (2006) Sickness behaviors in chronic cholestasis: an immune- mediated process? Hepatology 43:20-23. 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J Lipid Res 30:907-9 18. 66 Chapter 2 BIOTRANSFORMATION OF LITHOCHOLIC ACID BY RAT HEPATIC MICROSOMES: METABOLITE ANALYSIS BY LCIMS’ ‘A version of this chapter has been published as: Deo AK and Bandiera SM (2008a) Biotransformation of Lithocholic acid by Rat Hepatic Microsomes: Metabolite Analysis by Liquid Chromatography/Mass Spectrometry. Drug Metab Dispos. 36:442-451. Reprinted with permission of American Society for Pharmacology and Experimental Therapeutics. All rights reserved. Copyright © 2008 by The American Society for Pharmacology and Experimental Therapeutics. 67 2.1 Summary Lithocholic acid is a lipid-soluble hepatotoxic bile acid that accumulates in the liver during cholestasis. A potential detoxification pathway for lithocholic acid involves hydroxylation by hepatic cytochrome P450 (P450) enzymes. The purpose of the present study was to identify the hepatic microsomal metabolites of lithocholic acid by liquid chromatography-mass spectrometry and to determine the P450 enzymes involved. Incubation of lithocholic acid with rat hepatic microsomes and NADPH produced murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid as major metabolites, and 6-ketolithocholic acid and ursodeoxycholic acid as minor metabolites. Experiments with hepatic microsomes prepared from rats pretreated with P450 inducers and with inhibitory antibodies indicated that CYP2C and CYP3A enzymes contribute to microsomal murideoxycholic acid formation. Results obtained with a panel of recombinant P450 enzymes and CYP2D6 antiserum showed that CYP2D1 can also catalyze murideoxycholic acid formation. Similar experimental evidence revealed that formation of 3-ketocholanoic acid was mediated primarily by CYP3A enzymes. Isolithocholic acid formation appeared to be catalyzed by a distinct pathway mediated largely by microsomal non-P450 enzymes. On the basis of the results obtained using lithocholic acid and 3-ketocholanoic acid as substrates, a mechanism for the formation of isolithocholic acid involving a geminal diol intermediate is outlined. In conclusion, lithocholic acid was extensively metabolized by multiple P450 enzymes with the predominant biotransformation pathway being hydroxylation at the 613-position. This study provides an insight into possible routes of detoxification of lithocholic acid. 68 2.2 Introduction Lithocholic acid (3cL-hydroxy-53-cholan-24-oic acid) is a hydrophobic bile acid that is formed by the dehydroxylation of chenodeoxycholic acid (3cL,7cL-dihydroxy-513-cholan-24-oic acid). Removal of the 7cL-hydroxyl group of chenodeoxycholic acid is catalyzed by hydratase enzymes associated with anaerobic bacteria that reside in the colon (Hofmann, 1999). Lithocholic acid is absorbed from the colon and transported to the liver, where the acidic moiety is conjugated with taurine or glycine before being excreted as a component of bile (Hofmann, 2002; Hofinann, 2004). Lithocholic acid, together with other biliary bile acids, facilitates the absorption, transport and distribution of lipid-soluble nutrients from the diet and aids in the elimination of cholesterol from the body (Hofmann, 1999). Figure 2.1 shows the chemical structures of lithocholic acid and other bile acids (Hofmann et al., 1992). Lithocholic acid is present in human and rat liver at a concentration of approximately 1 to 5 nmol/g tissue, which represents approximately 4-5% of total bile acids in human and rat liver (Setchell et al., 1997). In comparison, the major bile acids in human liver, which are cholic acid and chenodeoxycholic acid, and the major bile acids in rat liver, which are cholic acid and muricholic acid, are present at concentrations that are 5 to 10 times greater (Setchell et al., 1997). Hepatic lithocholic acid concentrations are elevated in patients with cholestatic liver disease (Jezequel et al., 1994; Fischer et al., 1996; Erlinger, 1997; Hofinann, 2002; Berta et al., 2003; De Gottardi et al., 2004) and in rat models of biliary cholestasis (Setchell et al., 1997; Rost et al., 2003). The accumulation of lithocholic acid and other hydrophobic bile acids in liver has been implicated as a major factor contributing to liver injury in cholestasis because of the inherent cytotoxicity of this hydrophobic bile acid. Hepatic toxicity following chronic and acute administration of exogenous lithocholic acid or its conjugates is well documented in 69 experimental animals (Javitt, 1966; Palmer and Ruban, 1966; Zaki et al., 1967; Miyai et al., 1971; Fischer et al., 1974). The hepatotoxicity associated with lithocholic acid can be attenuated by hepatic biotransformation pathways including hydroxylation reactions catalyzed by the cytochrome P450 (P450) enzymes and Phase II reactions involving conjugation of the 3cL-hydroxyl group with sulfate (Kitada et al., 2003; Hofmann, 2004). The resulting metabolites are more water- soluble and more easily excreted. P450-mediated hydroxylation has been proposed to be an effective detoxification mechanism in rodents and monkeys, whereas sulfate conjugation is considered to be a more important pathway in humans (Hofmann, 2004). There is evidence indicating that P450-mediated hydroxylation of lithocholic acid is also prominent in humans. Lithocholic acid has been reported to be hydroxylated by human hepatic microsomes to hyodeoxycholic acid, murideoxycholic acid and chenodeoxycholic acid (Xie et al., 2001). Formation of hyodeoxycholic acid was shown to be catalyzed primarily by human recombinant CYP3A4 (Araya and Wikvall, 1999; Xie et al., 2001). More recently, a different metabolite, 3- ketocholanoic acid, was identified as the major metabolite of lithocholic acid with human recombinant CYP3A4 (Bodin et al., 2005). In comparison, studies with rat liver microsomes demonstrated that 6f3-hydroxylation of lithocholic acid leading to murideoxycholic acid is the major pathway in the rat (Zimniak et al., 1989; Dionne et al., 1994). However, the P450 enzymes involved in the formation of murideoxycholic acid in the rat were not identified. In the present study, the hepatic metabolism of lithocholic acid was investigated in rat hepatic microsomes using a liquid chromatography/mass spectrometry (LC/MS)-based assay and the various P450 enzymes involved in metabolite formation were identified. Currently, mass spectrometry is one of the most sensitive and specific methods to quantif’ bile acids in tissue and subeellular fractions (Stedman et al., 2004). LC/MS was used to measure formation 70 of the major and minor metabolites of lithocholic acid. Kinetic parameters associated with rates of metabolite formation in hepatic microsomes were also calculated. The hepatic P450 enzymes responsible for metabolite formation were determined by a combination of approaches involving P450 inducer treatments, antibody inhibition experiments, and rat recombinant P450 enzymes. 2.3 Materials And Methods 2.3.1 Chemicals and reagents. Authentic bile acid standards were purchased from Steraloids Inc. (Newport, RI). Bile acid standards were dissolved in methanol as 1 mg/mi stock solutions. Additional dilutions were made in methanol for the biotransformation assay. Deuterated cholic 2,2,4,4-d4 acid, which served as an internal standard, was a generous gift from Dr. Jan Palaty (Children’s & Women’s Health Center, Vancouver, BC, Canada). Sodium phenobarbital (PB) was obtained from BDH Chemicals (Toronto, Ontario, Canada). Dexamethasone (DEX) and 3- methylcholanthrene (MC) were purchased from Sigma-Aldrich Canada Ltd. (Oakville, Ontario, Canada). Baculovirus-insect cell control microsomes containing expressed rat P450- oxidoreductase and baculovirus-insect cell microsomes containing expressed rat P450 enzymes, co-expressed with rat P450-oxidoreductase or with rat P450-oxidoreductase and rat cytochrome b5 (BD SUPERSOMESTM Enzymes) were purchased from BD Biosciences (Oakville, Ontario, Canada). HPLC-grade chemicals and solvents were purchased from Fisher Scientific (Ottawa, Ontario, Canada). Rabbit CYP2D6 polyclonal antiserum (Daiichi Pure Chemicals Co. Ltd., Tokyo) was purchased from BD Biosciences. The reaction of CYP2D6 antiserum with rat CYP2DI was assessed in our laboratory by immunoblot analysis using recombinant rat P450 enzymes. Rabbit anti-rat CYP2C polyspecific immunoglobulin G (IgG) and rabbit anti-rat CYP3A polyspecific IgG were prepared as described previously (Bandiera and Dworschak, 1992; Panesar et al., 71 1996; Wong and Bandiera, 1996). The inhibitory activities of the anti-CYP2C and anti-CYP3A IgG preparations toward rat hepatic microsomal testosterone 2cL- and l6cx-hydoxylation and testosterone 63-hydroxylation, respectively, had been determined previously (Law, 1995; Wong and Bandiera, unpublished results). 2.3.2 Animal treatment and preparation of hepatic microsomes. Male and female Long- Evans, male Wistar and male Sprague Dawley rats (7-8 weeks of age) were purchased from Charles River Canada Inc. (Saint-Constant, Quebec, Canada). Upon arrival, rats were housed in pairs in polycarbonate cages on corncob bedding (Anderson’s Maumee, OH) with free access to water and food (Laboratory Rodent Diet, PMI Feeds Inc., Richmond, IN). Animal quarters were maintained at a constant temperature (23°C) with controlled light (14 h) and dark (10 h) cycles. Rats were cared for in accordance with the principles and guidelines of the Canadian Council on Animal Care. Male or female Long-Evans rats (n 5) were treated with xenobiotics as follows: PB (dissolved in water, 75 mg/kg/day), MC (dissolved in corn oil, 25 mg/kg/day), DEX (dissolved in corn oil, 100 mg/kg/day) or vehicle (0.8 mllkg/day). Compounds were administered by intraperitoneal injection for 4 days and rats were killed by decapitation 24 h after the last treatment. Microsomes were prepared from pooled livers as described previously (Thomas et a!., 1983). Microsomes were also prepared from pooled livers from untreated Wistar rats (n= 16) or Sprague Dawley rats (n=4). Microsomal pellets were suspended in 0.25 M sucrose and aliquots were stored at —75°C, until needed. Protein concentration was measured by the method • of Lowry et al. (Lowry, 1951) using bovine serum albumin as a standard. 72 2.3.3 Lithocholic acid biotransformation assay. Reaction mixtures contained lithocholic acid (0.5—300 jIM), 0.5 mg of hepatic microsomal protein, 50 mM potassium phosphate buffer, pH 7.4, 3 mM magnesium chloride, and 1 mM NADPH in a final volume of 1 ml. After preincubation for 10 mm at room temperature, reactions were initiated with NADPH and allowed to proceed for 30 mm at 37°C. Reactions were terminated with 4 ml of dichloromethane:isopropanol (80:20). A fixed amount (0.4 tg) of internal standard (cholic 2,2,4,4-d4 acid) was then added to each sample. Tubes were vortex-mixed for 1 mm, shaken manually for 1 mm and spun at 2000 g for 5 mm. The top aqueous layer was carefully removed, placed into a clean tube and re-extracted with a second 4 ml aliquot of dichloromethane:isopropanol (80:20). The final aqueous phase was discarded and the organic phase from the second and first extraction were combined and evaporated to dryness with nitrogen. The residue was reconstituted in 0.2 ml of mobile phase (methanol:water:10 mM ammonium acetate, pH 4.6 (67:23:10)) and filtered through a 3 mm, 0.45 .tm syringe polytetrafluoroethylene filter. A 10 j.tl aliquot of each sample was analyzed by LC/MS as described by Stedman et al. (2004) with modifications as outlined below. Control samples devoid of substrate, NADPH or microsomes, and defined mixtures of authentic bile acid standards, were routinely included in each assay. Incubations with rat recombinant P450 enzymes, instead of rat hepatic microsomes, were also carried out. Reaction mixtures contained 30 pmoles of each recombinant P450 enzyme (CYP1A2, CYP2A2, CYP2B1, CYP2C6, CYP2C1 1, CYP2C13, CYP2D1, CYP3A1 or CYP3A2) or, in the case of insect cell control microsomes, an equivalent amount of protein (0.15 mg). To determine if metabolite formation was P450-mediated, preliminary experiments were conducted with carbon monoxide-treated hepatic microsomes or heat-denatured microsomes. Microsome samples were 73 boiled for 5 mm in assay buffer. Carbon monoxide was bubbled into an incubation mixture containing assay buffer and microsomes for 2 mm. Assay conditions were tested using microsomes from untreated male Wistar rats to ensure that substrate and cofactor concentrations were saturating and that product formation was linear with respect to incubation time (ito 60 mm) and protein concentration (0.25 to 2 mg/mi of reaction mixture). 2.3.4 Antibody inhibition. The lithocholic acid biotransformation assay was performed as described above except that microsomes were preincubated with rabbit anti-rat CYP2C IgG, anti-rat CYP3A IgG or control IgG for 15 mm at room temperature prior to addition of substrate. NADPH was added to initiate the reaction. Four concentrations of each IgG (0, 1, 2.5 and 5.0 mg IgG/mg of microsomal protein) were tested. Formation of murideoxycholic acid by rat hepatic microsomes or recombinant CYP2D 1 was measured in the presence of rabbit CYP2D6 antiserum or control serum. Hepatic microsomes or recombinant CYP2D1 were preincubated with various amounts of either rabbit CYP2D6 antiserum or control serum (0, 10, 25, 50 and 100 ui/mi reaction mixture) for 15 mm at room temperature prior to addition of lithocholic acid. Reactions were initiated with NADPH. 2.3.5 Analytical methods. Formation of lithocholic acid metabolites was analyzed by LC/MS. Lithocholic acid and its metabolites were resolved on a XTerra MS C 18 (2.1 mm x 150 mm, 3.5 iim) column (Waters, Milford, MA) at 40°C using a Hewlett-Packard model 1090 II liquid chromatograph (Avondale, PA). The mobile phase consisted of solvent A (methanol:water:10 mM ammonium acetate, pH 4.6 (50:40:10)) and solvent B (methanol:water:10 mM ammonium acetate, pH 4.6 (85:5:10)). A linear gradient was used 74 starting from 100% solvent A to 100% solvent B from 0 to 20 mm, 100% solvent B from 20 to 25 mm, followed by an abrupt return to 100% solvent A at 25 mm, and re-equilibration with 100% solvent A for 10 mm. The flow rate was maintained at 0.15 mI/mm and total run time was 35 mm/sample. The LC was interfaced to a Fisons VG Quattro mass detector (Fisons Instruments, VG-Analytical, Manchester, United Kingdom). The MS was operated in atmospheric pressure electrospray negative ionization mode, with bath gas flow and nebulizer gas flow rates of 250 1/hr and 20 l/hr, respectively, a source temperature of 160°C, and capillary and cone voltages of 3 kV and 40 V, respectively. MASSLYNX® v3.1 software (Micromass, Altrineham, United Kingdom) was used for data acquisition. Metabolites were identified by comparison of their retention times and mass to charge ratios (m/z) with those of authentic standards. Quantitative determination of bile acids was performed by selected negative ion monitoring at m/z of 411, 407, 391, 389, 375 and 373. Under these conditions, the internal standard, cholic-2,2,4,4-d4acid (MW 412.57), typically eluted at 17 mm and was monitored at m/z 411. ct-Muricholic acid (MW 408.57), -muricho1ic acid (MW 408.57), y-muricholic acid (MW 408.57) and cholic acid (MW 408.57) eluted at 12, 13, 15 and 17 mm, respectively, and were monitored at m/z 407. Murideoxycholic acid (MW 392.57), ursodeoxycholic acid (MW 392.57), hyodeoxycholic acid (MW 392.57), chenodeoxycholic acid (MW 392.57) and deoxycholic acid (MW 392.57) eluted at 11, 15, 17, 20 and 20.5 mm, respectively, and were monitored at m/z 391. 6-Ketolithocholic acid (MW 390.56) eluted at 15 mm and was monitored at m/z 389. Isolithocholic acid (MW 376.57) and lithocholic acid (MW 376.57) eluted at 23 and 26 mm, respectively, and were monitored at m/z 375. 3-Ketocholanoic acid (MW 374.56) eluted at 25 mm and was monitored at m/z 373. Metabolites were quantified from calibration plots of the peak area ratio of the authentic standard and internal standard plotted against the concentration of the authentic standard. 75 Method validation studies with respect to limit of detection, extraction efficiency, inter-and intra-day precision and stability of analytes were carried out as described previously (Deo, 2005). 2.3.6 Data analysis and calculation of enzyme kinetic parameters. Data were analyzed using the SigmaPlot® Enzyme Kinetics Module (v.1.1, Systat Software Inc., Richmond, CA). Metabolite formation as a function of substrate concentration was analyzed by nonlinear regression analysis and apparent Km. K’ and V,,, values were generated using the Hill equation (Equation 1) or the substrate inhibition kinetic equation (Equation 2) for the major metabolites, or the Michaelis-Menten equation (Equation 3) for the minor metabolites of lithocholic acid. V x[S] V = K’+[S] (Equation 1) V V =1+ Km /[S]+ [Sj / K1 (Equation 2) V x[S] V = (Equation 3) where v is initial velocity of the reaction, Vm. is the maximal velocity, [Sj is the substrate concentration, K’ is the Hill dissociation constant, n is the Hill coefficient representing cooperativity of the reaction, Km is the Michaelis-Menten constant, and K1 is the dissociation constant of substrate binding to the inhibitory site. 76 2.3.7 Statistical analysis. Comparisons of rates of metabolite formation were made using the unpaired t test with Welch’s correction. A p value of 0.05 was considered statistically significant. 2.4 Results 2.4.1 Biotransformation of lithocholic acid and metabolite identification. A mixture of thirteen bile acid standards (c-muricholic acid, 13-muricholic acid, ‘y-muricholic acid, cholic acid, murideoxycholic acid, ursodeoxycholic acid, hyodeoxycholic acid, chenodeoxycholic acid, deoxycholic acid, 6-ketolithocholic acid, isolithocholic acid, 3-ketocholanoic acid and lithocholic acid) was initially analyzed to ensure that potential metabolites were resolved with the LC conditions selected. Baseline separation of the bile acid standards was achieved, with the exception of a-muricholic acid and f3-muricholic acid, which were distinguished by spiking with authentic standards. Incubation of lithocholic acid with liver microsomes from male Wistar rats yielded seven metabolites that were identified as 13-muricholic acid, murideoxycholic acid, isolithocholic acid, ursodeoxycholic acid, hyodeoxycholic acid, 6-ketolithocholic acid and 3- ketocholanoic acid (Fig. 2.2) by comparison with authentic standards. The major metabolites were murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid. 6-Ketolithocholic acid and ursodeoxycholic acid were minor metabolites, while 13-muricholic acid and hyodeoxycholic acid were produced at levels close to or below the limit of quantification. Four other peaks (M-1, m/z 407; M-2, m/z 407; M-3, ,n/z 391 and M-4, m/z 389) corresponding to trace metabolites were detected but were not identified because the retention times of these metabolites did not match those of the authentic standards. To determine if lithocholic acid 77 biotransformation differed among rat strains, incubations were performed with hepatic microsomes from male Wistar, Long-Evans and Sprague Dawley rats. The same metabolite profile was obtained for all three rat strains (results not shown). Metabolite formation was not observed when lithocholic acid was incubated with carbon monoxide-treated or boiled microsomal preparations, or when NADPH was omitted from the reaction mixture. Although chenodeoxycholic acid was previously reported to be a metabolite of lithocholic acid in the rat (Zimniak., 1989), our results indicate that chenodeoxycholic acid is not a metabolite. A peak at m/z 391 with the same retention time as chenodeoxycholic acid was observed but was determined to be a contaminant of lithocholic acid as it was detected with reaction mixtures that contained substrate and NADPH but not hepatic microsomes and the area of the chenodeoxycholic acid peak did not change following incubation with increasing concentrations of hepatic microsomes. 2.4.2 Kinetic analysis of hepatic microsomal metabolite formation. Formation of the major and minor metabolites of lithocholic acid was evaluated over a range of substrate concentrations (0.5 to 300 1iM). An incubation time of 30 mm and a microsomal protein concentration of 0.5 mg/ml were found to be optimal and were used in subsequent experiments. A lithocholic acid concentration of 100 or 250 M, depending on the metabolite, was found to be saturating. Hepatic microsomal murideoxycholic acid formation exhibited typical Michaelis Menten kinetics up to a substrate concentration of 100 j.tM. At higher concentrations of lithocholic acid, the rate of murideoxycholic acid formation decreased, possibly as a result of substrate inhibition (Fig. 2.3A). Formation of isolithocholic acid and 3-ketocholanoic acid exhibited sigmoidal kinetic profiles. Decreased isolithocholic acid and 3-ketocholanoic acid formation was observed at substrate concentrations greater than 100 .tM for isolithocholic acid 78 and 250 j.M for 3-ketocholanoic acid (Fig. 2.3B and 2.3C). Hence, saturating substrate concentrations of 100 pM for murideoxycholic acid and isolithocholic acid formation and 250 p.M for 3-ketocholanoic acid formation were selected for further experiments. Nonlinear Eadie Hofstee plots of murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid formation, signif’ing atypical enzyme kinetics, were obtained (Fig. 2.3A, 2.3B and 2.3C). Of the minor metabolites that could be quantified, formation of 6-ketolithocholic acid and ursodeoxycholic acid followed typical Michaelis-Menten kinetics up to a substrate concentration of 100 p.M (results not shown). Apparent Km and Vm values for hepatic microsomal murideoxycholic acid formation were calculated using equation 2. Apparent K’ and Vmax values for hepatic microsomal isolithocholic acid and 3-ketocholanoic acid formation were calculated using equation 1 (Table 2.1). Positive cooperativity (n> 2) was indicated for isolithocholic acid and 3-ketocholanoic acid formation. The apparent V value for murideoxycholic acid formation was approximately 7 and 14 times higher than the values for isolithocholic acid and 3-ketocholanoic acid formation, respectively, demonstrating that murideoxycholic acid was the predominant microsomal metabolite of lithocholic acid in rats. On the other hand, the lower apparent K’value associated with isolithocholic acid formation (29.3 ± 5.9 p.M) suggests that this metabolite would be preferentially formed at low lithocholic acid concentrations. The apparent Vm values calculated for ursodeoxycholic acid and 6-ketolithocholic acid formation (Table 2.1) confirm that formation of these metabolites was quantitatively less important than formation of murideoxycholic acid, isolithocholic acid or 3-ketocholanoic acid. 79 2.4.3 Effect of P450 inducers on lithocholic acid biotransformation. To identify the P450 enzymes involved in lithocholic acid biotransformation, experiments were performed with hepatic microsomes prepared from male and female Long-Evans rats that had been pretreated with MC, PB or DEX. Comparison of rates of formation of the metabolites revealed a distinct sex difference for some of the metabolites (Table 2.2). Rates of murideoxycholic acid and isolithocholic acid formation for control male rats were approximately 2 and 30 times greater, respectively, than for control female rats, whereas the rate of formation of 3-ketocholanoic acid acid was similar for control male and female rats. Treatment with PB or MC decreased formation rates of murideoxycholic acid and isolithocholic acid for male rats. The rate of formation of 3-ketocholanoic acid was slightly, but not significantly, greater for both sexes following DEX treatment, while formation of murideoxycholic acid and isolithocholic acid was not affected. With respect to the minor metabolites, rates of formation of 6-ketolithocholie acid and ursodeoxycholic acid were greater for male than female rats and formation of both metabolites was increased following treatment with DEX, but not PB or MC (data not shown). 2.4.4 Antibody inhibition studies. The role of CYP2C and CYP3A enzymes in lithocholic acid biotransformation was investigated using antibodies against CYP2C and CYP3A enzymes. Rates of formation of murideoxycholic acid and 3-ketocholanoic acid were each inhibited by approximately 50% by anti-CYP2C IgG (at 5 mg IgG/mg protein), whereas isolithocholic acid formation was not affected (Fig. 2.4). Anti-CYP3A IgG (at 5 mg IgG/mg protein) inhibited the rates of formation of murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid by 40 to 60% (Fig. 2.4). Formation of 6-ketolithocholic acid and ursodeoxycholic acid were not affected by anti-CYP2C IgG but were inhibited by approximately 20% and 30%, respectively, following incubation of microsomes with anti-CYP3A IgG (at 5 mg IgG/mg protein, data not shown). 80 2.4.5 Biotransformation studies with recombinant P450 enzymes. The contribution of individual P450 enzymes in lithocholic acid biotransformation was evaluated using a panel of nine rat recombinant P450 enzymes (Fig. 2.5). Initial experiments were conducted to determine P450 concentrations that would ensure linearity of product formation with incubation time. An incubation time of 30 mm and rat recombinant P450 enzyme concentration of 30 pmol P450/ml were found to be optimal. Under the experimental conditions employed, CYP2D1 was the most active P450 enzyme catalyzing murideoxycholic acid formation. In comparison, formation of isolithocholic acid was catalyzed at a relatively low rate by several recombinant P450 enzymes including CYPIA2, CYP2C6, CYP2D1, CYP3A1 and CYP3A2, but not by CYP2A2 or CYP2C 13. The rate of formation of isolithocholic acid by the recombinant P450 enzymes was approximately 4 to 16 times less than the rate obtained with hepatic microsomes, when activity values were expressed per nmol total P450. CYP3A2, followed by CYP3A1 and CYP2C1 1, were the most active P450 enzymes catalyzing 3-ketocholanoic acid formation. The rate of 3- ketocholanoic acid formation by recombinant CYP3A2 was found to be greater, when expressed per nmol total P450, than that obtained with hepatic microsomes (approximately 31 vs 0.6 pmollminlpmol P450, respectively). Formation of 6-ketolithocholic acid and ursodeoxycholic acid by rat recombinant P450 enzymes was also assessed. 6-Ketolithocholic acid formation was catalyzed mainly by CYP3A1 and CYP3A2, while ursodeoxycholic acid formation was catalyzed solely by CYP2A2 (results not shown). There was no evidence of f3-muricholic acid or chenodeoxycholic acid fonnation by the rat recombinant P450 enzymes. Formation of the minor unidentified metabolite M-4 was catalyzed by CYP3A2 and CYP3A2. The other unidentified metabolites, M- 1 to M-3, were not observed using rat recombinant P450 enzymes. To veri1y the involvement of CYP2D1 in murideoxycholic acid formation, antibody inhibition experiments were conducted using CYP2D6 antiserum, which has been reported to 81 cross-react with CYP2D1 (Umehara et al., 1997). Murideoxycholic acid formation by recombinant CYP2D1 was inhibited by 90% at the highest concentration of CYP2D6 antiserum tested (Fig. 2.7). In contrast, CYP2D6 antiserum had no effect on murideoxycholic acid formation by hepatic microsomes. Conversion of lithocholic acid to isolithocholic acid involves epimerization of the hydroxyl group at the 3-position. To determine whether isolithocholic acid formation could occur by a stepwise process, 3-ketocholanoic acid was incubated with hepatic microsomes prepared from control male rats and with a panel of rat recombinant P450 enzymes in the presence of NADPH. Hepatic microsomes catalyzed isolithocholic acid formation from both lithocholic acid and 3-ketocholanoic acid. Formation of isolithocholic acid from 3- ketocholanoic acid was not catalyzed by any of the recombinant P450 enzymes. Formation of 3-ketocholanoic acid by recombinant CYP3A2 was further evaluated at substrate concentrations of 0.5 to 100 iM (Fig. 2.6). Formation of 3-ketocholanoic acid by recombinant CYP3A2 followed typical Michaelis-Menten kinetics, which is in contrast to the atypical sigmoidal kinetic pattern observed with hepatic microsomes. These data suggest that formation of 3- ketocholanoic acid in hepatic microsomes involved more than a single P450 enzyme. A V,, value of 31.5 pmol/minlpmol P450 and a Km value of 18.9 jiM were obtained for 3- ketochólanoic acid formation by recombinant CYP3A2. 2.5 Discussion Lithocholic acid was extensively metabolized by rat hepatic microsomes to three major (murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid) and two minor (6- ketolithocholic acid and ursodeoxycholic acid) products, as determined by LC/MS. The predominant biotransformation pathway involved hydroxylation of lithocholic acid at the 613- 82 position and led to formation of murideoxycholic acid. This result is consistent with previous in vivo and in vitro studies, which described hepatic 6j3-hydroxylation of bile acids as a major pathway in rat (Voigt et al., 1968; Gustafsson, 1978) and identified murideoxycholic acid as a major metabolite of lithocholic acid (Zimniak et al., 1989; Dionne et al., 1994). However, the P450 enzymes catalyzing murideoxycholic acid formation were not identified in previous studies. Our experiments with hepatic microsomes prepared from rats pretreated with P450 inducers and with inhibitory P450 antibodies indicated that CYP2C and CYP3A enzymes contribute to microsomal murideoxycholic acid formation, while results obtained with a panel of rat recombinant P450 enzymes showed that CYP2D1 also catalyzes murideoxycholic acid formation. The lack of effect by CYP2D6 antiserum on hepatic microsomal murideoxycholic acid formation suggests that there is little or no contribution by CYP2D 1 to the activity of hepatic microsomes. This may be explained by the relatively low level of expression of CYP2D 1. Determination of CYP2D 1 levels in hepatic microsomes by immunoblot analysis indicated that CYP2D1 accounted for approximately 2-4% of total P450 in hepatic microsomes prepared from untreated Wistar and Long-Evans rats (results not shown). Collectively, the results imply that several P450 enzymes, which are expressed at higher levels than CYP2D 1, are involved in murideoxycholic acid formation as shown in Fig. 2.5A. Lithocholic acid concentrations of 5 to 10 ,iM have been reported in the liver of cholestatic patients and in rat models of biliary cholestasis (Jezequel et al., 1994; Fischer et al., 1996; Erlinger, 1997; Setchell et al., 1997; Hoflnann, 2002; Berta et al., 2003; Rost et al., 2003; De Gottardi et al., 2004). A lithocholic acid concentration of 100 jtM was found to be saturating for hepatic microsomal murideoxycholic acid formation in the present study. Nevertheless, murideoxycholic acid was the major metabolite obtained in the in vitro biotransformation assay, with a rate of formation of 250-500 pmol/minlmg at a lithocholic acid 83 concentration of 5 jiM, which approximates the physiological hepatic concentration. Thus, our study demonstrates that 63-hydroxylation is the major P450-mediated hepatic pathway of lithocholic acid biotransformation in the rat. In comparison, hydroxylation of lithocholic acid at the 6u-position has been proposed to be the predominant P450-mediated pathway in humans (Araya and Wikvall, 1999; Xie et a!., 2001). Only a trace amount of hyodeoxycholic acid was produced by rat liver microsomes in the present study. Isolithocholic acid, the 313-isomer of lithocholic acid, was the second most abundant microsomal metabolite detected in our study. Isolithocholic acid was reported to be a major metabolite in human feces (Norman and Palmer, 1964; Palmer, 1971) and was identified as an in vivo metabolite of rats fed sulfated lithocholic acid (Palmer, 1971; Zimniak et al., 1989; Dionne et a!., 1994) but was not found to be a metabolite of lithocholic acid produced by rat liver microsomes (Palmer, 1971; Zimniak et a!., 1989; Dionne et al., 1994). In our study, conversion of lithocholic acid to isolithocholic acid was observed with some rat recombinant P450 enzyme preparations but at a relatively low rate. Hepatic microsomal isolithocholic acid formation was not induced by pretreatment with MC, PB or DEX, was not inhibited by anti CYP2C IgG and was slightly inhibited by anti-CYP3A IgG. The results suggest that microsomal enzymes other than P450 enzymes were involved in isolithocholic acid formation. Epimerization of the hydroxyl group on steroid molecules is most often catalyzed by hepatic hydroxysteroid dehydrogenase and steroid oxidoreductase enzymes and is not a common P450- mediated reaction (Penning et al., 1986). A possible mechanism for the formation of isolithocholic acid is shown in Fig. 2.8. We speculate that microsomal isolithocholic acid formation can proceed by two pathways. One pathway involves conversion of lithocholic acid to isolithocholic acid, possibly through a geminal diol intermediate with subsequent loss of a hydroxyl group. The geminal diol is formed by 13—hydroxylation at an existing a—hydroxy 84 position (as suggested by Bodin et a!., 2005). The geminal diol intermediate can spontaneously rearrange to form either isolithocholic acid or 3-ketocholanoic acid. A second pathway involves 313-oxidation of lithocholic acid followed by dehydration to form 3-ketocholanoic acid, with subsequent reduction of 3-ketocholanoic acid to isolithocholic acid. Our experiments conducted using 3-ketocholanoic acid as substrate showed that although isolithocholic acid is a major hepatic microsomal metabolite of 3-ketocholanoic acid, none of the recombinant P450 enzymes contributed significantly to that conversion. Accordingly, we believe that both pathways are mediated largely by non-P450 enzymes. An example of a microsomal enzyme that can catalyze this reaction is 313-hydroxy-A5-C27ster id oxidoreductase enzyme, which was purified from rabbit liver microsomes and was shown to catalyze the reversible oxidation of the 313-hydroxyl group ofC27-steroids (Wikvall, 1981). 3-Ketocholanoic acid was the third most abundant hepatic microsomal metabolite of lithocholic acid in our study. A recent report identified 3-ketocholanoic acid as the major lithocholic acid metabolite (74% of total) formed by human recombinant CYP3A4 (Bodin et a!., 2005), and two earlier studies found 3-ketocholanoic acid as a major in vivo product of rats fed a lithocholic acid-enriched diet (Thomas et al., 1964; Sakai et al., 1980). Formation of 3- ketocholanoic acid from lithocholic acid entails oxidation of a hydroxyl group and can be catalyzed by P450 enzymes by the mechanism described above. In the present study, treatment of male and female rats with DEX led to increased 3-ketocholanoic acid formation suggesting that CYP3A enzymes contribute to formation of this metabolite. Results of the antibody inhibition experiments substantiate the involvement of CYP3A and CYP2C in 3-ketocholanoic acid formation. Of the recombinant enzymes examined, the highest catalytic activity was obtained with CYP3A2. Taken together, the data provide convincing evidence that 3- 85 ketocholanoic acid formation in rat hepatic microsomes was catalyzed primarily by CYP3A. A proposed mechanism for 3-ketocholanoic acid formation was outlined above (see Fig. 2.8). Minor metabolites, namely 6-ketolithocholic acid and ursodeoxycholic acid, as well as metabolites that were detected but could not be quantified, namely hyodeoxycholic acid and 13- muricholic acid, were also reported by Zimniak et al. (1989). Our results suggest that 6- ketolithocholic acid formation was catalyzed mainly by CYP3A enzymes, while ursodeoxycholic acid formation was catalyzed by CYP2A enzymes. Appreciable formation of 13-muricholic acid was apparent only at low substrate concentrations suggesting that 13- muricholic acid was converted to other metabolites. Chenodeoxycholic acid, which was reported to be a metabolite in previous studies (Thomas et al., 1964; Zimniak et al., 1989), was identified herein as a contaminant of lithocholic acid. We found no evidence of chenodeoxycholic acid formation by either rat hepatic microsomes or rat recombinant P450 enzymes. Our study focused primarily on the contribution of P450 enzymes to the biotransformation of lithocholic acid in hepatic microsomes. We are aware that Phase II enzymes such as sulfotransferases and glucuronosyl transferases also play an important role in lithocholic acid biotransformation. Glucuronosyl transferases are microsomal enzymes that may facilitate conversion of lithocholic acid and its metabolites, such as murideoxycholic acid, to their ester glucuronide conjugates, thereby affecting murideoxycholic acid formation. Moreover, nuclear hormone receptors, such as the pregnane X receptor (PXR), constitutive androstane receptor, liver X receptor, and farnesoid X receptor can be activated by bile acids and help regulate bile acid homeostasis (Staudinger et al., 2001; Makishima, 2005). Some of these receptors, such as PXR, are involved in the inducible expression of P450 enzymes and sulfotransferases and glucuronosyl transferases (Tien and Negishi, 2006). Thus, treatment with 86 DEX and other inducers used in the present study could alter formation of metabolites such as murideoxycholic acid through an effect on Phase II enzymes, apart from the effect on P450 enzymes. This may partly explain the decreased murideoxycholic acid formation observed after pretreatment with DEX. The concentration of DEX used in our study (100 mg/kg) is sufficient to activate PXR in rodents (Hartley et al., 2004). In summary, LC/MS proved to be an effective method for resolving and identifying the biotransformation products of lithocholic acid. Lithocholic acid, like many P450 substrates, is metabolized by multiple P450 enzymes, which catalyze overlapping pathways. Our study suggests a major role for hepatic CYP2C and CYP3A enzymes in lithocholic acid biotransformation. In rats and humans, the contribution of individual P450 enzymes to the various metabolic pathways is determined by their level of expression and the tissue concentration of lithocholic acid. CYP2C enzymes are the predominant P450 enzyme subfamily expressed in untreated rat liver and together with CYP3A enzymes, which predominate in human liver, are expected to be the main catalysts of lithocholic acid biotransformation in rats and possibly in humans. Because CYP3A is involved in the formation of most of the major metabolites of lithocholic acid, induction of CYP3A enzymes offers a potential mechanism to lessen the hepatotoxicity associated with high tissue levels of lithocholic acid. On the basis of results obtained, a scheme for P450-mediated formation of major lithocholic acid metabolites in rat hepatic microsomes is proposed in Fig. 2.9. The LC/MS analytical method employed herein provides a sound foundation for future bile acid biotransformation studies using human hepatic microsomes and purified or recombinant P450 enzymes, studies that are currently underway in our laboratory. 87 Apparent V Apparent Apparent n Apparent Metabolite pmol/min/mg Km K’ K1 protein 1tilvI 1uM JilVI Murideoxycholie N/A N/A 194 ± 74.6 acida 5920±1180 56.0±17.8 Isolithocholie 803 ± 91.8 N/A 29.3 ± 5.9 2.3 ± 0.8 N/A acid” 3-Ketocholanoic 413 ± 25.8 N/A 71.6 ± 6.5 2.5 ± 0.6 N/A acidb 6-Ketolithocholic 44.5 ± 2.1 1.6 ± 0.3 N/A N/A N/A acide Ursodeoxycholic 22.4± 1.7 15.3±3.7 N/A N/A N/A acide Table 2.1 Kinetic parameters of lithocholic acid metabolite formation by rat hepatic microsomes. Kinetic parameters were derived from the metabolite formation data presented in Fig. 2.3. Values represent the mean ± SEM of triplicate determinations. a Kinetic parameters for murideoxycholic acid formation were calculated using the substrate inhibition kinetics model (equation 2). b Kinetic parameters for isolithocholic acid and 3-ketocholanoic acid formation were calculated using the sigmoidal kinetics model, Hill equation (equation 1). C Kinetic parameters for 6-ketolithocholic acid and ursodeoxycholic acid formation were calculated using the Michaelis-Menten equation. N/A, not applicable 88 Treatment Rate ofmetabolite formation pmol/min/mg protein Murideoxycholic Isolithocholic 3-Ketocholanoic acid acid acid Male rats Control 2900 ± 302 803 ± 83.0 512 ± 49.7 MC 1010±171* 403±70.0* 486±117 PB 633±150* 231±55.0* 394±106 DEXa 1810 765 1380 Female rats Control 1220 ± 182* 25.3 ± 1.4* 461 ± 19.0 pBa 1250 23.2 585 DEX 781±122 33.2±10.0 1050±169 Table 2.2 Effect of sex and treatment with P450 inducers on lithocholic acid metabolite formation by rat hepatic microsomes. Incubation of hepatic microsomes with lithocholic acid was carried out at saturating substrate concentrations (100 jiM lithocholic acid for murideoxycholic acid and isolithocholic acid and 250 jiM for 3-ketocholanoic acid, respectively) under optimal assay conditions as described in the Materials and Methods section. Hepatic microsomes prepared from vehicle-treated male and untreated female Long-Evans rats were used as controls. Values represent the mean ± SEM of triplicate determinations, except for treatments denoted by a, where values represent the average of duplicate determinations, except for treatments deonoted by a Rates of metabolite formation for hepatic microsomes prepared from male rats pretreated with MC or PB and from control female rats were compared to rates obtained with hepatic microsomes prepared from control male rats. Rates of metabolite formation for hepatic microsomes prepared from female rats pretreated with DEX were compared to rates obtained with hepatic microsomes prepared from control female rats (p < 0.07). p < 0.05, a Values represent the average of duplicate determinations. 89 R421 22 11 2 121 COOH RMTR3 R2 R1 R, R3 R4 Bile acidlmetabolite Chemical name a-OH I-f H H lithocholic acid 3a-hydroxy-5p-cholan-24-oic acid 2. a-OH H a-OH H chenodeoxycholic acid 3a, 7a-dihydroxy-53-choIan-24-oic acid a-OH H a-OH a-OH cholic acid 3a, 7a, 12a-tnhydroxy-513-cholan-24-oic acid 4. a-OH 13-OH a-OH H a-muncholic acid 3o, 613, 7a-trihydroxy- 513-cholan-24-oic acid a-OH 13-OH 13-OH H 13-muricholic acid 3o, 613, 713-trihydroxy-513-cholan-24-oic acid 6. a-OH a-OH H H hyodeoxycholic acid 3a, 6a-dihydroxy-513-cholan-24-oic acid 7. a-OH p-OH H H murideoxycholic acid 3a, 613-dihydroxy-513-cholan-24-oic acid 8. 0 H H H 3-ketocholanoic acid 3-oxo-513-cholan24-oic acid 9. a-OH 0 H H 6-ketolithocholic acid 3a-hydroxy- 6-oxo-513-cholan-24-oic acid 10. u-OH H H H isolithocholic acid 313-hydroxy-513-cholan-24-oic acid 1 1. a-OH H 130H H ursodeoxycholic acid 3a, 713-dihydroxy-513-cholan-24oic acid 12. a-OH H H a-OH deoxycholic acid 313, 1 2a-dihydroxy-513-cholan-24-oic acid Figure 2.1 General bile acid structure showing positions available for hydroxylation. The bile acid nomenclature listed in the figure uses trivial names and abbreviations which are similar to those used by HofiTlann et a!. (1992). Chemical names conform to IUPAC nomenclature. Please note that the words cholanic acid, cholanoic acid and cholan-24-oic acid have been used interchangeably in this thesis and represent the bile acid steroid system. 90 nilz 411 R e a un eoxyc 0 IC —j Ursodeo,choIic acid mlz 391I acid fiM4 /V j / Hyodeoxycholic acide I Chenodeoxycholic 7’ acid ._.1_______::::dt_ m!z 389 mlz 375 Y Lithocholic acid (%) 373 Figure 2.2 Representative LCIMS chromatogram showing metabolites of lithocholic acid. Lithocholic acid metabolites were extracted from a standard reaction mixture after a 30-mm incubation of rat hepatic microsomes (0.5 mg) with 50 M lithocholic acid and 1 mM NADPH. Metabolite identification was performed by co-chromatography and spiking with authentic standards. The internal standard was deuterated cholic-2,2,4,4-d4acid. Peaks identified as 13- muricholic acid and hyodeoxycholic acid were produced at levels close to or below the limit of quantification. Peaks M- 1, M-2, M-3 and M-4 are metabolites that did not correspond to any of the authentic standards. Peaks denoted by * indicate peaks that were present in controls and blanks and are not metabolites. Standard mlz 407 100 Isolithocholic acid, ‘1 Time (mm) 91 4000 c 1000 0 A °600 f: •. •1 0 50 100 150 200 250 300 V’s —0 00 50 100 150 200 250 300 100[Lithocholic acid] (pM) 0 50 100 150 200 250 300 [Uthocholic acid] (Iv1) Figure 2.3 Kinetic profiles of hepatic microsomal murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid formation. Metabolite formation (activity) was plotted as a function of substrate concentration following a 30-mm incubation with rat liver microsomes (0.5 mg). Data points are the mean ± SEM of at least three separate experiments. Lines represent rates modeled by nonlinear regression analysis of the data. The insets depict Eadie-Hofstee plots. Error bars are not shown on the insets to avoid obscuring the data points, which represent mean values. Substrate inhibition was observed at lithocholic acid concentrations greater than 100 iM for murideoxycholic acid formation (A). Sigmoidal kinetics suggestive of homotropic autoactivation were observed for isolithocholie acid (B) and 3-ketocholanoic acid (C) formation. Saturating lithocholic acid concentrations of 100 .tM and 250 pM were determined for isolithocholic acid and 3- ketocholanoic acid formation, respectively. 500 C 0 20 40 60 80 [Lithocholic acid] (pM) f 92 120 A 140 B ioo —•-----___ 120 05 0 .0 !; :: l00 °40 0040 0 2’ 3 20 0 0 0 1 2 3 4 5 6 0 1 2 3 4 5 6 mg IgGImg m,crosomal protein mg IgGImg microsomat protein 120 100 4- u80 60 . 0- .t C ,— 20 0 0 1 2 3 4 5 6 mg IgGImg microsomal protein Figure 2.4 Effect of anti-CYP2C IgG and anti-CYP3A IgG on hepatic microsomal murideoxycholic acid (A), isolithocholic acid (B) and 3-ketocholanoic acid (C) formation. Hepatic microsomes from untreated male rats (0.5 mg) were incubated with various concentrations of rabbit anti-rat CYP2C polyspecific IgG (—0—), anti-CYP3A polyspecific IgG (—V—), and control IgG (—•—) for 15 mm at room temperature before addition of lithocholic acid (100 iiM). Results are expressed as percent of the activity obtained in the presence of 0 mg IgG. The plots shown above contain data from a single experiment. 93 3.0 2.5 2.0 1.5 1.0 eE C. • Figure 2.5 Comparison of murideoxycholic acid (A), isolithocholic acid (B) and 3- ketocholanoic acid (C) formation by a panel of rat recombinant P450 enzymes. Metabolite formation (activity) was measured following a 30-mm incubation of lithocholic acid (100 M) with baculovirus-insect cell microsomes containing expressed rat P450 enzymes (30 pmol). Insect cell microsomes containing expressed P450 enzymes, except for CYP1A2 and CYP2D1, also contained co-expressed P450 oxidoreductase and cytochrome b5. Microsomes containing expressed CYP1A2 and CYP2D1 contained co-expressed P450 oxidoreductase but not cytochrome b5. There was no obvious correlation between rates of metabolite formation and P450P oxidoreductase or cytochrome b5 levels in the recombinant P450 preparations. Plots show the mean values ± SEM of triplicate determinations. A B 0.30 0.25 1 1 Eu I . 0•20J II ii0.15 -I0.10 -I I0.050.00flr r N << mc 0 4< N ‘N N N C.) C.) a. a. 0. 0. N N 0. a. a.)- >- >- >- a. a.>.>. >.C.) 0 C) 0>->- C) C) C)C) C.) C — N N2<< me — — — — N N N C.) 0 N ra. a. a. a. N N a. 0. a. >. >. >. >. a. a.>->- >. C.) 0(3(3 00 C) C 0 .- Ino a. •0 . 0 U 0. C C, N N — ID — C) — CI << m C.) N C C, a. N N a. a. a. >. >- a. a.>- >. >-C.) 0000>->- C) C) C)CC) 94 30 0 E _.. 25 L O 0U) O..20 0E 0- ..3 oE C., 5 0 [Lithocholic acid] (ElM) Figure 2.6 Enzyme kinetic recombinant CYP3A2. profile of 3-ketocholanoic acid formation by rat Metabolite formation (activity) was plotted as a function of substrate concentration following a 30-mm incubation with rat recombinant CYP3A2 (30 pmoles). The inset depicts an Eadie Hofstee plot. Data points are the average of duplicate determinations. V V 0 20 40 60 80 100 95 140 t 120 m E 100 80 Amount of serum added (.dIml of reaction mixture) Figure 2.7 Effect of CYP2D6 antiserum on murideoxycholic acid formation by rat hepatic microsomes and recombinant CYP2D1. Various amounts of rabbit CYP2D6 antiserum (_•_) or rabbit control serum (— V —) were added to reaction mixtures (1 ml final volume) containing hepatic microsomes from untreated male rats (0.5 mg). Similarly, various amounts of CYP2D6 antiserum (—0—) or rabbit control serum (—A—) were added to reaction mixtures containing rat recombinant CYP2D1 (30 pmol). Reaction mixtures were preincubated with antibody for 15 mm at room temperature before addition of lithocholic acid and initiation of the reaction with NADPH. Results are expressed as percent of the activity obtained in the presence of 0 1 of serum. The plot shown above contains data from a single experiment. 0 20 40 60 80 100 120 96 c±9ThCOOH 3-Ketocholanoic acid H2O +[HJ Isolithocholic acid Figure 2.8 Scheme showing a proposed mechanism for the formation of 3- ketocholanoic acid and isolithocholic acid from lithocholic acid through a geminal diol intermediate. Lithocholic acid geminal diol intermediate 97 HO 21 22 23 Ii 7COOH 10 14 15 H03 4 H Lithocholic acid / CYP2CII, CYP3A2 3-Ketocholanoic acid Figure 2.9 hepatic microsomes. Scheme showing P450-mediated lithocholic acid biotransformation in rat Results of the present study suggest that murideoxycholic acid and 3-ketocholanoic acid formation was mediated by CYP2C and CYP3A enzymes, whereas isolithocholic acid formation was mediated largely by non-P450 enzymes. 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Staudinger IL, Goodwin B, Jones SA, Hawkins-Brown D, MacKenzie KI, LaTour A, Liu Y, Klaassen CD, Brown KK, Reinhard J, Willson TM, Koller BH and Kliewer SA (2001) 101 The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc Nat! Acad Sci USA 98:3369-3374. Stedman C, Robertson G, Coulter S and Liddle C (2004) Feed-forward regulation of bile acid detoxification by CYP3A4: studies in humanized transgenic mice. J Biol Chem 279:11336-11343. Thomas PE, Reik LM, Ryan DE and Levin W (1983) Induction of two immunochemically related rat liver cytochrome P-450 isozymes, cytochromes P-450c and P-450d, by structurally diverse xenobiotics. JBiol Chem 258:4590-4598. Thomas PJ, Hsia SL, Matschiner JT, Doisy EA, Jr., Elliott WH, Thayer SA and Doisy EA (1964) Bile Acids. Xix. Metabolism of Lithocholic Acid-24-14c in the Rat. JBiol Chem 239:102-105. Tien ES and Negishi M (2006) Nuclear receptors CAR and PXR in the regulation of hepatic metabolism. Xenobiotica 36:1152-1163. Voigt W, Thomas PJ and Hsia SL (1968) Enzymic studies of bile acid metabolism. I. 6-beta- Hydroxylation of chenodeoxycholic and taurochenodeoxycholic acids by microsomal preparations of rat liver. JBiol Chem 243:3493-3499. Wikvall K (1981) Purification and properties of a 3 beta-hydroxy-delta 5-C27-steroid oxidoreductase from rabbit liver microsomes. JBiol Chem 256:3376-3380. Wong A and Bandiera SM (1996) Inductive effect of Telazol® on hepatic expression of cytochrome P450 2B in rats. Biochem Pharmacol 52:735-742. Xie W, Radominska-Pandya A, Shi Y, Simon CM, Nelson MC, Ong ES, Waxman DJ and Evans RM (2001) An essential role for nuclear receptors SXRIPXR in detoxification of cholestatic bile acids. Proc Nat! Acad Sci USA 98:3375-3380. 102 Zaki FG, Carey JB, Jr., HofThauer FW and Nwokolo C (1967) Biliary reaction and choledocholithiasis induced in the rat by lithocholic acid. JLab Clin Med 69:737-748. Zimniak P, Holsztynska EJ, Lester R, Waxman DJ and Radominska A (1989) Detoxification of lithocholic acid. Elucidation of the pathways of oxidative metabolism in rat liver microsomes. JLiidRes 30:907-9 18. 103 Chapter 3 IDENTIFICATION OF HUMAN HEPATIC CYTOCIIROME P450 ENZYMES INVOLVED IN THE BIOTRANSFORMATIQN OF CHOLIC AND CHENODEOXYCHOLIC ACID2 2A version of this chapter has been published as: Deo AK and Bandiera SM (2008b) Identification of Human Hepatic Cytochrome P450 Enzymes Involved in the Biotransformation of Cholic and Chenodeoxycholic Acid. Drug Metab Dispos.36: 1983-91. Reprinted with permission of American Society for Pharmacology and Experimental Therapeutics. All rights reserved. Copyright © 2008 by The American Society for Pharmacology and Experimental Therapeutics. 104 3.1 Summary Cholic acid (3a,7cz, 1 2a-trihydroxy-5 f3-cholan-24-oic acid) and chenodeoxycholic acid (3ct,7cx-dihydroxy-53-cholan-24-oic acid) are the predominant hepatic and biliary bile acids of most mammalian species including humans. Cholic and chenodeoxycholic acids are synthesized from cholesterol and accumulate in the liver during cholestasis. Biotransformation by hepatic cytochrome P450 (P450) enzymes represents a potentially effective pathway for elimination of these lipid-soluble bile acids. We developed a liquid chromatography/mass spectrometry-based assay to identify and quantify the human hepatic microsomal metabolites of cholic acid and chenodeoxycholic acid, and using a panel of human recombinant P450 enzymes, we determined the P450 enzymes involved. Incubation of cholic acid with human hepatic microsomes and NADPH produced a single metabolite, 3-dehydrocholic (7x, 1 2x-dihydroxy-3-oxo-5 f3-cholan- 24-oic acid). Of the recombinant P450 enzymes tested, only CYP3A4 catalyzed 3- dehydrocholic acid formation. Similar experiments with chenodeoxycholic acid revealed the formation of 7x-hydroxy-3-oxo-5-cho1an-24-oic acid and y-muricholic acid (3c,6ct,7cL- trihydroxy-513-cholan-24-oic) as major metabolites and 7-ketolithocholic acid (3c-hydroxy-7- oxo-513-cholan-24-oic acid and cholic acid as minor metabolites. Among the human recombinant P450 enzymes examined, CYP3A4 exhibited the highest rates of formation for 7cx-hydroxy-3- oxo-513-cholan-24-oic acid and y-muricholic acid from chenodeoxycholic acid. Formation of 7- ketolithocholic acid and cholic acid from chenodeoxycholic acid has not been reported previously and could not be attributed to any of the recombinant P450 enzymes tested. In conclusion, the predominant pathway for the biotransformation of both cholic and chenodeoxycholic acids in human hepatic microsomes was oxidation at the third carbon of the cholestane ring. This study highlights a major role for CYP3A4. and suggests a possible route for the elimination of these two bile acids. 105 3.2 Introduction Cholic acid and chenodeoxycholic acid comprise approximately 70% of hepatic and biliary bile acids in humans (Hofmann, 2002; Ridlon et al., 2006). Cholic acid and chenodeoxycholic acid are biosynthesized from cholesterol in the liver through a tightly regulated multi-enzyme pathway that includes several cytochrome P450 (P450) enzymes such as CYP7A1, CYP7B1, CYP8B1 and CYP27A1 (Norlin and Wikvall, 2007). In the liver of healthy adults, the cholic acid concentration is reported to be 14 to 21 nmollg liver while the chenodeoxycholic acid concentration is 23 to 31 nmol/g liver (Fischer et al. 1996; Setchell et al., 1997). During cholestasis, a pathophysiological condition that results from obstruction of bile flow and is a common manifestation of liver diseases (Hofmann 2002), cholic acid and chenodeoxycholic acid levels are elevated to 120 nmollg liver and 85 nmollg liver, respectively (Fischer et a!., 1996). Hepatic bile acids provide the primary stimulus for canalicular bile flow and facilitate the excretion of excess hepatic cholesterol into the bile (Hofmann, 1999). Bile acids function as highly effective emulsifiers in the small intestine facilitating the solubilization and absorption of dietary lipids and lipid-soluble nutrients and the elimination of phospholipid and cholesterol (Hofmann, 1999; Hofmann, 2002). In addition, bile acids serve as signaling molecules in the liver (Chiang, 2002; Makishima, 2005) and even play a role in normal liver regeneration (Huang et al., 2006). More specifically, chenodeoxycholic acid and its analogues, and to a lesser extent cholic acid, have been identified as farnesoid X receptor (FXR) agonists (Pellicciari et al., 2002; Ellis et al., 2003; Fiorucci et a!., 2005; Rizzo et al., 2005). FXR, a nuclear transcription factor, regulates expression of several genes involved in bile acid biosynthesis and transport (Grober et al., 1999). At high concentrations, chenodeoxycholic acid binds and activates FXR, leading to down-regulation of the biosynthetic enzymes, CYP7A1 and CYP8B1, and up regulation of proteins involved in bile salt trafficking including bile acid export pump 106 (BSEP/ABCB 11) and multidrug resistance related proteins (MDR3/B4 and MRP2/C2) (Grober et a!., 1999). This feedback mechanism helps maintain bile acid homeostasis and provides protection against bile acid toxicity. The role of this autoregulatory mechanism in providing protection against bile acid toxicity was demonstrated with FXR gene knockout mice, which have impaired resistance to bile acid-induced hepatotoxicity (Sinai et a!., 2000). Biotransformation is an additional process, besides bile acid synthesis and transport, for regulating bile acid levels in the liver. Bile acids are subject to multiple metabolic biotransformations in hepatocytes including conjugation with taurine, glycine, glucuronic acid and sulfate, as well as P450-mediated oxidation, while in the colon, bile acids can undergo dehydroxylation, deconjugation and hydroxylation reactions catalyzed by bacterial enzymes. Although much is known about bile acid hydroxylation catalyzed by bacterial systems (Chapter 1, Section 1.2.2), knowledge of bile acid hydroxylation by hepatic P450 enzymes is limited. Hydroxylation increases hydrophilicity and introduces additional functional sites for glucuronide and sulfate conjugation, thereby facilitating excretion of the bile acids. The relatively few in vivo and in vitro studies that identified hydroxylated metabolites of cholic and chenodeoxycholic acids reported different metabolite profiles. Tetrahydroxylated metabolites of cholic acid namely, 3cL, 6a, 7ci, 1 2cc- tetrahydroxy-513-cholan-24-oic acid and 1 , 3a, 7cL, 1 2- tetrahydroxy 5f3-cholan-24-oic acid, were identified in the urine of a patient with biliary cirrhosis (Bremmelgaard and Sjovall, 1980). In contrast, 3-dehydrocholic acid was the sole metabolite found when cholic acid was incubated with recombinant human CYP3A4 (Bodin et al., 2005). Similarly, y-muricholic acid was the only product identified when chenodeoxycholic acid was incubated with human liver microsomes or with recombinant CYP3A4 (Araya and Wikvall 1999). Because y-muricholic acid was found in urine from healthy individuals and patients. with intrahepatic cholestasis (Alme and Sjovall, 1980; Bremmelgaard and Sjovall, 107 1980; Shoda et al., 1990), 6ct-hydroxylation of chenodeoxycholic acid was proposed to be a major hydroxylation pathway in humans (Setchell et al., 1998; Araya and Wikvall, 1999). However, a more recent study reported that y-muricholic acid was a major but not the predominant metabolite formed from chenodeoxycholic acid when incubated with human recombinant CYP3A4 (Bodin et a!., 2005). In the present study, we investigated the biotransformation of cholic acid and chenodeoxycholic acid by human hepatic microsomes using a liquid chromatography/mass spectrometry (LC/MS)-based assay. Metabolites of cholic acid and chenodeoxycholic acid were identified, metabolite formation was quantified, and kinetic parameters associated with metabolite formation were determined. Using a panel of recombinant human P450 enzymes, we also identified the P450 enzymes involved in metabolite formation. 3.3 Materials And Methods 3.3.1 Chemicals and reagents. Chenodeoxycholic acid, cholic acid, deoxycholic acid, hyodeoxycholic acid, isolithocholic acid, lithocholic acid, cL-muricholic acid, 13-muricholic acid, y-muricholic acid, murideoxycholic acid, ursodeoxycholic acid, 3-dehydrocholic acid, 3- ketocholanoic acid, 6-ketolithocholic acid, and 7-ketolithocholic acid were purchased from Steraloids Inc. (Newport, RI). 1 cL,3 l3,7c, 1 2a-tetrahydroxy-5 (3-cholan-24-oic acid and 3a,6f3,73,12c-tetrahydroxy-5j3-cholan-24-oic acid were generous gifts from Dr. Lee R. Hagey, University of California, San Diego. 7c-hydroxy-3-oxo-5[3-cholan-24-oic acid was custom synthesized by Steraloids Inc. specifically for this study. The identity and purity of 7cc- hydroxy-3-oxo-53-cho1an-24-oic acid were confirmed by Steraloids Inc. Bile acid standards were dissolved in methanol as 1 mg/ml stock solutions and stored at -4°C. Additional dilutions were made in methanol for the biotransformation assay. Cholesterol, 25-hydroxycholesterol, 108 and cholestanol were purchased from Sigma Inc. (Oakville, Ontario, Canada). Stock solutions of cholesterol, 25-hydroxycholestrol, and cholestanol (10 mM) were dissolved in acetone and stored at room temperature. Proadifen (SKF-525AHC1) was kindly provided by Dr. T.K.H. Chang, University of British Columbia, Vancouver, Canada. Pooled human liver microsomes were purchased from Xenotech (Kansas, MO). Baculovirus-insect cell control microsomes containing expressed human P450-oxidoreductase and baculovirus-insect cell microsomes containing expressed human P450 enzymes (BD SUPERSOMESTM Enzymes), co-expressed with human P450-oxidoreductase or with human P450-oxidoreductase and human cytochrome b5, were purchased from BD Biosciences (Oakville, Ontario, Canada). High-performance liquid chromatography-grade chemicals and solvents were purchased from Fisher Scientific (Ottawa, Ontario, Canada). 3.3.2 Cholic acid and chenodeoxycholic acid biotransformation assays. Reaction mixtures contained 50 mMpotassium phosphate buffer, pH 7.4, 3 mM magnesium chloride, 0.5 mg of human hepatic microsomal protein, 1 mM NADPH and varying concentrations (1-800 jtM) of either cholic acid or chenodeoxycholic acid, in a final volume of 1 ml. After preincubation for 10 mm at room temperature, reactions were initiated with NADPH and allowed to proceed for 30 mm at 37°C. Reactions were terminated with 8 ml of dichloromethane/isopropanol (80:20 v/v). A fixed amount (0.4 tg) of murideoxycholic acid, which was the internal standard, was then added to each sample. Sample extraction evaporation, and reconstitution in preparation for analysis by LC/MS were carried out as described previously (Deo and Bandiera, 2008). Reaction mixtures that were devoid of substrate, NADPH or microsomes, as well as reaction mixtures that contained defined concentrations of authentic bile acid standards, were routinely included in each assay. 109 Assay conditions were tested using pooled human microsomes to ensure that substrate and cofactor concentrations were saturating and that product formation was linear with respect to incubation time (1 to 60 mm) and protein concentration (0.25 to 2 mg/mi of reaction mixture). To determine if metabolite formation was P450-mediated, preliminary experiments were conducted with carbon monoxide-treated hepatic microsomes or heat-denatured microsomes or by replacing NADPH with NADH or by adding SKF-525A. Incubations with recombinant human P450 enzymes, instead of human hepatic microsomes, were also carried out. Reaction mixtures contained 30 pmoies of each recombinant P450 enzyme (CYPIA1, CYP1A2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2CI9, CYP2D6, CYP2E1, CYP3A4, CYP3A5 or CYP4A1 1) or, in the case of insect cell control microsomes and reductase control, an equivalent amount of protein (0.15 mg). 3.3.3 Analytical methods. Formation of chenodeoxycholic acid and cholic acid metabolites was analyzed by LC/MS using a procedure described previously for lithocholic acid metabolites (Deo and Bandiera, 2008), with the following modifications. Chenodeoxycholic acid and cholic acid metabolites were detected using a Waters Acquity Ultra Performance Liquid Chromatograph System (UPLC, Waters Corp. Milford, MA) consisting of a Binary Solvent Manager and Sample Manager and connected to a Waters Quattro Premier XE triple quadrupole mass spectrometer (Waters Corp., Milford, MA). The MS was operated in Single Ion Recording mode, using negative electrospray ionization with desolvation gas 600 LIhr and cone gas 52 L/hr respectively, a source temperature of 100°C, and capillary and cone voltages of 3 kV and 20 V, respectively. Waters MassLynx® v4. 1 software (Waters Corp., Milford, IVIA) was used for data acquisition. Metabolites were identified by comparison of their retention times and mass to charge ratios (m/z) with those of authentic standards. A mixture of 17 bile acid 110 standards was prepared. Under these conditions, 1 ct,3 13,7ct, 1 2c-tetrahydroxy-5 13-cholan-4-oic acid (MW 424.57) and 3c,6P,713,12cL -tehydroxy-53-cholan-24-oic acid (MW 424.57) typically eluted at 4 and 5 mm, respectively, and were monitored at m/z 423. cj-Muricholic acid (MW 408.57), f3-muricholic acid (MW 408.57), y-muricholic acid (MW 408.57) and cholic acid (MW 408.57) eluted at 12, 13, 15 and 16 mm, respectively, and were monitored at m/z 407. 3- Dehydrocholic acid (MW 406.58) eluted at 13 mm and was monitored at m/z 405. Murideoxycholic acid (MW 392.57), ursodeoxycholic acid (MW 392.57), hyodeoxycholic acid (MW 392.57), chenodeoxycholic acid (MW 392.57) and deoxycholic acid (MW 392.57) eluted at 11, 15, 17 20, 20.5 mm, respectively, and were monitored at m/z 391. 6-Ketolithoeholic acid (MW 390.6), 7-ketolithocholic acid (MW 390.6) and 7a-hydroxy-3-oxo-5 13-cholan-24-oic acid (MW 390.6) eluted at 15, 16 and 17 mm, respectively, and were monitored at m/z 389. Isolithocholic acid (MW 376.57) and lithocholic acid (MW 376.57) eluted at 23 and 26 mm, respectively, and were monitored at m/z 375. 3-Ketocholanoic acid (MW 374.56) eluted at 25 mill and was monitored at m/z 373. All ion channels were routinely scanned with the help of this standard mixture for identification of metabolites during initial experiments. Metabolites were quantified from calibration plots of the peak area ratio of authentic standard and internal standard plotted against the concentration of the authentic standard. Method validation studies with respect to limit of detection, extraction efficiency, inter-and intra-day precision and stability of analytes were carried out as described in Appendix I. 3.3.4 Data analysis and calculation of enzyme kinetic parameters. Data were analyzed using the SigmaPlot® Enzyme Kinetics Module (v.1.1, Systat Software Inc., Richmond, CA). Metabolite formation as a function of substrate concentration was analyzed by nonlinear 111 regression analysis and apparent Km, K’ and V,,,,, values were generated using the Michaelis Menten (equation 1) or the Hill equation (equation 2). VmaxX[SI K,,,+[S] (equation 1) VmX[SI — K’ + [S] (equation 2) where v is initial velocity of the reaction, Vm is the maximal velocity, [S] is the substrate concentration, K’ is the Hill dissociation constant, n is the Hill coefficient representing cooperativity of the reaction, Km is the Michaelis-Menten constant. Several criteria such as sum of squares, standard deviation of residuals, for each equation, Akaike Information Criterion and visual inspection of the fit were considered in selecting the most appropriate model for each data set (Tracy and Hummel, 2004). 3.4 Results 3.4.1 Biotransformation and kinetic analysis of cholic acid metabolites. Incubation of cholic acid with human liver microsomes yielded a single metabolite identified as 3- dehydrocholic acid, by comparison with authentic standards (Fig. 3.1). A microsomal protein concentration of 0.5 mg/mI and incubation time of 30 mm and were found to be within the linear range with respect to product formation and were used in subsequent experiments. Formation of 3-dehydrocholic acid evaluated over a range of substrate concentrations (1-800 tM) exhibited unsaturable kinetics (Fig. 3.2). Higher cholic acid concentrations were not used routinely because distorted peak profiles due to column overload were observed at concentrations greater than 800 riM. Apparent kinetic parameters were determined using 112 equation 1 (Fig. 3.2). The inset in Fig. 3.2 shows the Eadie-Hofstee plot (velocity versus velocity/substrate concentration). 3.4.2 Biotransformation and kinetic analysis of chenodeoxycholic acid metabolites. Incubation of chenodeoxycholic acid with human liver microsomes yielded two major metabolites identified as 7cL-hydroxy-3-oxo-5f3-cholan-24-oic acid and y-muricholic acid, and two minor metabolites identified as 7-ketolithocholic acid and cholic acid (Fig. 3.3). Identification of metabolites was confirmed by comparing retention times and spiking using authentic standards. An incubation time of 30 mm and a microsomal protein concentration of 0.5 mg/mI were found to be optimal for all four chenodeoxycholic acid metabolites and were used in subsequent experiments. Formation of 7c-hydroxy-3-oxo-53-cholan-24-oic acid and y-muricholic acid was not observed with reaction mixtures that were devoid of substrate, NADPH or microsomes. However, chromatographic peaks with the same m/z values and retention times as 7- ketolithocholic acid and cholic acid were detected when chenodeoxeholic acid was incubated with boiled microsomal preparations or when human liver microsomes or NADPH were omitted from the reaction mixture. We observed that the 7-ketolithocholic acid and cholic acid peaks were approximately two- to three-times larger when substrate was incubated with both NADPH and human liver microsomes, and peak areas increased with increasing microsomal protein concentration or increasing incubation time. Moreover, addition of carbon monoxide or SKF 525A (1 mM) to the reaction mixtures inhibited formation of all four metabolites. Taken together these data indicate that 7-ketolithocholic acid and cholic acid were hepatic microsomal metabolites, as well as contaminants of chenodeoxycholic acid. Hence, quantification of 7- ketolithocholic acid and cholic acid formation necessitated subtraction of peak area ratios for 7- 113 ketolithocholic acid and cholic acid obtained with blank reaction mixtures from peak area ratios measured with complete reaction mixtures. Metabolite formation was evaluated over a range of substrate concentrations (1 to 800 1iM). A chenodeoxycholic acid concentration of 500 jiM was found to be saturating for all four metabolites. Plots of metabolite fonnation versus substrate concentration showed that hepatic microsomal formation of 7c-hydroxy-3-oxo-5 13-cholan-24-oic acid, 7-ketolithocholic acid and cholic acid followed Michaelis-Menten kinetics (Fig. 3.4A, 3.4C and 3.4D, respectively), whereas, formation of y-muricholic acid followed sigmoidal kinetics (Fig. 3.4B). The inset in Fig. 3.4B shows the Eadie-Hofstee plot (velocity versus velocity/substrate concentration), typical of homotropic positive cooperativity and suggestive of substrate autoactivation (Tracy and Hummel, 2O04). Apparent kinetic parameters for 7c-hydroxy-3-oxo-53-cholan-24-oic acid, cholic acid and 7-ketolithocholic acid were determined using equation 1 (Fig. 3.4A, 3.4C and 3.4D). Apparent K’ and Vm values for y-muricholic acid formation were determined using equation 2 (Fig. 3.4B). Rates of hepatic microsomal metabolite formation, expressed as apparent Vm, demonstrate that 7c-hydroxy-3-oxo-5f3-cholan-24-oic acid would be the predominant metabolite of chenodeoxycholic acid in human liver microsomes at high substrate concentrations. 3.4.3 Biotransformation studies with human recombinant P450 enzymes. The contribution of individual P450 enzymes in cholic acid and chenodeoxycholic acid biotransformation was evaluated using a panel of twelve human recombinant P450 enzymes. Initial experiments were conducted to determine P450 concentrations that would ensure linearity of product formation with incubation time. For cholic acid and chenodeoxychloic acid biotransformation, an 114 incubation time of 30 mm and human recombinant P450 enzyme concentration of 30 pmol P450/mi were found to be optimal, at a substrate concentration of 500 jiM. Conversion of cholic acid to 3-dehydrocholic acid was catalyzed solely by CYP3A4 (Fig. 3 .5A). Formation of 3-dehydrocholic acid by recombinant CYP3A4, evaluated over a range of substrate concentrations, gave a sigmoidal kinetic profile exhibiting saturation at 500 .tM (Fig. 3.6). Kinetic parameters for 3-dehydrocholic acid formation by recombinant CYP3A4 were determined using equation 2 (Fig. 3.6). With chenodeoxycholic acid as the substrate, the most active enzyme catalyzing conversion of chenodeoxycholic acid to 7a-hydroxy-3-oxo-5f3-cholan-24-oic acid was CYP3A4 (Fig. 3.5B). CYP3A5 and several other P450 enzymes catalyzed 7cc-hydroxy-3-oxo-53-cholan- 24-oic acid formation at much lower rates. CYP3A4 was the only enzyme that mediated formation of y-muricholic acid (Fig 3 .5B). Formation of 7cL-hydroxy-3-oxo-5 13-cholan-24-oic acid by recombinant CYP3A4, evaluated over a range of substrate concentrations (1-600 tiM), exhibited typical Michaelis-Menten kinetics (Fig. 3.7A). Formation of y-MCA by recombinant CYP3A4 exhibited a sigmoidal kinetic profile as was observed with human liver microsomes (Fig. 3 .7B). Kinetic parameters obtained for formation of 7c-hydroxy-3-oxo-5 13-choian-24-oic acid and y-muricholic acid were determined using equations 1 and 2, respectively. In contrast, conversion of chenodeoxychoiie acid to 7-ketolithocholic acid and cholic acid was not catalyzed by the human recombinant P450 enzymes. Formation of 7- ketolithocholic acid involves oxidation of the 7a-hydroxy group of chenodeoxycholic acid. Steroid ring oxidation at the 7-position occurs in the conversion of cholesterol to 7cc- hydroxycholesterol, which is catalyzed by CYP7A1, and in the conversion of 25- hydroxycholesterol to 7cc-hydroxylated oxysterols, which is catalyzed by CYP7BI (Martin et al., 1993; Schwarz et al., 2001). To determine whether CYP7A or CYP7B enzymes are 115 involved in 7-ketolithocholic acid formation, ehenodeoxycholic acid was incubated with human liver microsomes and NADPH in the presence of cholesterol (a CYP7A1 substrate), 25- hydroxycholesterol (a CYP7B 1 substrate) and cholestanol (a non-specific CYP7A and CYP7B inhibitor) (Martin et al., 1993). Addition of increasing concentrations (100, 200, 250, and 300 jiM) of cholesterol, hydroxycholesterol, or cholestanol to a saturating chenodeoxycholic acid concentration (500 jiM) did not alter formation of 7-ketolithocholic acid or cholic acid (data not shown). 3.5 Discussion Biotransformation of cholic acid by human liver microsomes generated a single metabolite, 3-dehydrocholic acid, as determined by LC/MS. By comparison, chenodeoxycholic acid produced two major metabolites, 7cL-hydroxy-3-oxo-53-cho1an-24-oic acid and ‘y muricholic acid, and two minor metabolites, 7-ketolithocholic acid and cholic acid. Oxidation at the C-3 position was the predominant biotransformation pathway for both cholic acid and chenodeoxycholic acid. Hydroxylation at the 6cL-position was the next most important biotransformation pathway for chenodeoxycholic acid. Both reactions were catalyzed almost exclusively by CYP3A4. Limited information is available regarding cholic acid and chenodeoxycholic acid hydroxylation by human liver microsomes and the information that exists is largely derived from analyses of urinary and fecal metabolites, and more recently from, in vitro incubations performed with recombinant CYP3A4. We initially expected cholic acid to be converted, at least partially, to tetrahydroxylated metabolites because urinary tetrahydroxy-metabolites were identified in a patient with primary biliary cirrhosis who was given labeled cholic acid intravenously (Bremmelgaard and Sjovall, 1980). Consequently, we scanned for but did not 116 detect molecular ions (nv’z 423) corresponding to tetrahydroxylated metabolites of cholic acid. Instead, oxidation of cholic acid by human liver microsomes yielded a single metabolite, 3- dehydrocholic acid, and was catalyzed by CYP3A4. This result is consistent with a previous study that identified 3-dehydrocholic acid as the only product formed when cholic acid was incubated with recombinant CYP3A4 (Bodin et a!., 2005). Oxidized bile acid metabolites containing a keto group at C-3 are found in human feces and are thought to result from bacterial metabolism in the colon (Ridlon et al., 2006). Evidence that hepatic oxidation of cholic acid at the C-3 position occurs in vivo is provided by a report that 3-oxo-bile acids are present, at low levels, in bile from human fetal gall bladder samples (Setchell et al., 1988). The present study is the first to identify 7a-hydroxy-3-oxo-5 3-choIan-24-oic acid as the predominant hepatic microsomal metabolite of chenodeoxycholic acid. This metabolite was detected as a major chromatographic peak by LC/MS but the retention time and molecular ion m/z ratio of the peak did not correspond to those of the commercially available bile acid standards. The m/z ratio suggested that the unidentified peak was an oxo metabolite with oxidation at C-3. On the basis of its chromatographic characteristics and m/z ratio, we predicted the chemical structure of the metabolite and had 7cL-hydroxy-3-oxo-53-cholan-24-oic acid custom-synthesized by Steraloids Inc. (Newport, RI). The retention time and molecular ion m/z ratio of 7a-hydroxy-3-oxo-53-cholan-24-oic acid matched that of the major metabolite peak and its identification was established. As was the case with cholic acid, the predominant biotransformation pathway for chenodeoxycholic acid in human liver microsomes was CYP3A4-eatalyzed oxidation at the C-3 position. The physiological significance of the conversion of chenodeoxycholic acid to 7x-hydroxy-3-oxo-5 13.-cholan-24-oic acid, or of cholic acid to 3-dehydrocholic acid, is unknown. Based on the chromatographic retention times (Fig. 3.1 and Fig. 3.3), it is reasonable to suggest that these metabolites are hydrophilic than their 117 substrates. Trace amounts of 7ct-hydroxy-3-oxo-5f3-cholan-24-oic acid have been reported in human fetal gall bladder bile samples (Setchell et al., 1988) indicating that this metabolite is formed in vivo, as well as in vitro. y-Muricholic acid, a 6a-hydroxylated bile acid, is found at low levels in bile, serum and urine of healthy adult humans (Almé and Sjovall 1980; Shoda et al., 1989; Wietholtz et a!., 1996) but is a major bile acid component in human fetal bile (Setchell et al., 1988). Two in vitro studies (Bodin et a!., 2005) identified y-muricholic acid as a CYP3A4-mediated metabolite of chenodeoxycholic acid. The present study corroborates the role of CYP3A4 in y-muricholic formation. Excretion of y-muricholic acid is increased in patients with hepatobiliary disease and in pregnant women with cholestasis (Summerfield et a!., 1976; (Bremmelgaard and Sjovall, 1980) Nakashima et al., 1990; Shoda et al., 1990) indicating that CYP3A4-catalyzed 6cc- hydroxylation of chenodeoxycholic acid is up-regulated in cholestasis. Our study identified 7-ketolithocholic acid and cholic acid as contaminants of chenodeoxycholic acid and as minor metabolites of chenodeoxycholic acid biotransformation by human hepatic microsomes. These two bile acids were not previously reported to be metabolites of chenodeoxycholic acid. 7-Ketolithocholic acid is considered to be a major intermediate in the conversion of chenodeoxycholic acid to ursodeoxycholic acid in human colon (Fromm et al., 1983) and its formation is known to be catalyzed by bacterial 7- hydroxysteroid dehydrogenases of resident intestinal micorflora (Ridlon et al., 2006). Reduction of 7-ketolithocholic acid to chenodeoxycholic acid and ursodeoxycholic acid by human liver enzymes has been demonstrated (Fromm et a!., 1983; Amuro et al., 1989). We did not observe ursodeoxycholic acid formation in the present study. Formation of 7- ketolithocholic acid by human liver microsomes has not been reported previously. However, hepatic synthesis of 7-ketolithocholic acid has been demonstrated in guinea pig, a species in 118 which 7-ketolithocholic acid constitutes approximately 30-35% of the total biliary bile acid pool (Tint et al., 1990). None of the panel of recombinant P450 enzymes tested catalyzed formation of 7-ketolithocholic acid. In addition, formation of 7-ketolithocholic acid did not appear to be catalyzed by CYP7A and CYP7B enzymes because its formation was not affected by competitive inhibitors of CYP7A1 or CYP7B 1 (i.e. cholesterol, 25-cholesterol or cholestanol). Formation of 7-ketolithocholic acid can possibly be mediated by non-P450 enzymes, but the identity of these enzymes remains unknown. Conversion of chenodeoxycholic acid to cholic acid involves 12c-hydroxylation. This reaction was also not catalyzed by any of the recombinant P450 enzymes tested and was not inhibited by competitive inhibitors of CYP7A1 or CYP7B 1 (i.e. cholesterol, 25-cholesterol or cholestanol). A hepatic microsomal P450 enzyme that was not examined in this study is CYP8B 1. CYP8B 1, also known as sterol 1 2-hydroxylase, catalyses the 1 2x-hydroxylation of 3-oxo-7c-hydroxy-4-cholestene, which is an intermediate step in the multi-enzyme pathway leading to cholic acid formation from cholesterol (Russell and Setchell, 1992), CYP8B 1 has relatively broad substrate specificity in vitro and has been shown to hydroxylate chenodeoxycholic acid at the 1 2cc-position (Andersson et al., 1998). We speculate that CYP8B 1 may be responsible for the formation of cholic acid from chenodeoxycholic acid observed in the present study, but we were unable to assess the involvement of CYP8B 1 because recombinant CYP8B1 or an inhibitory antibody to CYP8B1 were not commercially available. The physiological significance of hepatic P450-catalyzed cholic acid and chenodeoxycholic acid biotransformation in vivo is unknown. Cholic and chenodeoxycholic acid concentrations of approximately 120 M and 85 tM, respectively, have been reported in the liver of patients with chronic cholestasis (Fischer et al., 1996). At these concentrations, the rate of formation of 3-dehydrocholic acid from cholic acid and the rate of formation of 7cL- 119 hydroxy-3-oxo-5f3-cholan-24-oic acid from chenodeoxycholic acid would be approximately 50- 75 and 75-100 pmollmin/mg protein, respectively, as determined by our biotransformation assay. The lower apparent Km value associated with 7-ketolithocholic acid formation by human liver microsomes (27 p.M, Fig. 3 .4C) suggests that this metabolite may be preferentially formed at physiological concentrations of chenodeoxycholic acid in vivo. The present study focused on the contribution of P450 enzymes in the biotransformation of cholic acid and chenodeoxycholic acid by human hepatic microsomes. P450-mediated oxidation probably represents a relatively minor pathway for bile acid elimination compared to conjugation reactions catalyzed by Phase II enzymes such as sulfotransferases, glucuronosyl transferases and amino acid transferases. Based on the results obtained, a scheme for biotransformation of cholic acid and chenodeoxycholic acid in human hepatic microsomes is proposed in Fig. 3.8 and Fig. 3.9, respectively. The results provide compelling evidence that cholic acid and chenodeoxycholic acid can serve as physiological substrates of CYP3A4. CYP3A4 is the most abundant P450 enzyme in human adult liver and small intestine (Guengerich, 1995; Guengerich, 2003) and plays a major role in the oxidation of a large number of structurally diverse xenobiotics and physiological compounds including steroid hormones and bile acids (Hrycay and Bandiera, 2008). CYP3A4 expression is subject to regulation by the pregnane X receptor and constitutive androstane receptor, which are activated by several drugs and other xenobiotics. Induction of CYP3A4 by treatment with drugs such as phenobarbital, rifampicin and phenytoin, or alternately, inhibition of CYP3A4 activity by antifungal agents or macrolide antibiotics can affect cholic and chenodeoxycholic acid biotransformation, in addition to resulting in clinically significant drug interactions. 120 Cholic acid Figure 3.1. Representative LCIMS chromatogram showing metabolites of cholic acid. Cholic acid metabolites were extracted from a standard reaction mixture after a 30-mm incubation of human hepatic microsomes (0.5 mg) with 100 iiM cholic acid and 1 mM NADPH. Metabolite identification was performed by co-chromatography and spiking with authentic standards. In the above figure, the internal standard (murideoxycholic acid, mlz 391), eluted at 12 mm, 3-dehydrocholic acid eluted at 13 mm and cholic acid (m/z 405) eluted at 16 mm, respectively. Peaks denoted by * indicate peaks that were present in controls and blanks and are not metabolites. 100 R e I “ a 0 m!z 407 V e 100 n t e I’ S 100 If (%) 0 m!z 405 .- Internal standard mlz 391 10 Time (mm) 20 25 30 121 400 0 . E U C., Figure 3.2. Enzyme kinetic profile of 3-dehydrocholic acid formation by human hepatic microsomes. Metabolite formation (activity) was plotted as a function of substrate concentration following a 30-mm incubation with human liver microsomes (0.5 mg). Data points are the mean ± SEM of at least three separate experiments. Data were fitted to a one-enzyme Michaelis-Menten model. Lines represent rates modeled by nonlinear regression analysis. Kinetic parameters were calculated using equation 1. The inset depicts the Eadie-Hofstee plot. Error bars are not shown on the insets to avoid obscuring the data points, which represent mean values. Estimated V,. = 756 ± 158 pmol/minlmg protein Estimated Km = 1060±133 liM 300 200 100 0 a 00 a 00 a a 0 200 400 600 800 [Cholic add] (pIV 1000 122 100 Cholic acid / mlz 407 R e a mlz 391 I Internal standard mlz 389 Y 7-Ketolithocholic acid 10 1 Figure 3.3. Representative LCIMS chromatogram showing metabolites of chenodeoxycholic acid. Chenodeoxycholic acid metabolites were extracted from a standard reaction mixture after a 30- mm incubation of human hepatic microsomes (0.5 mg) with 100 jiM chenodeoxycholic acid and 1 mM NADPH. Metabolite identification was performed by co-chromatography and spiking with authentic standards. In the above figure, the internal standard (murideoxycholic acid, mlz 391) eluted at 15.5 mm. y-Muricholic acid (mlz 407) and cholic acid (m/z 407) eluted at 16.5 and 18 mm, respectively. 7-Ketolithocholic acid (mlz 389) and 7c-hydroxy-3-oxo-5f3-cholan- 24-oic acid (m/z 389) eluted at 18 and 18.5 mm, respectively. ‘y-muricholic acid c 0 100 / Chenodeoxycholic1 acid 0 100 0 — 7a- hydroxy-3-oxo- 5 fr cholari-24-oic-acid 20 Time (mm) 2 30 123 AApparent V,,,,,,, = 392 ± 21 prnoilminlmg protein Apparent Km = 244*3IILM B I Apparent Vmar = 94±6 pmoUminlmg protein Apparent K’ = 225±l9iLM I, = 3±0.4— 100 -0 • E p.. a’ a 200 400 800 Chenodeoxycholic acidl (pM) 1000 C 0 200 400 600 800 fChenodeoxycholic acid] (pM) Apparent V,,,,,,, 62 ± 2 pmollmlnlmg protein Apparent K,,, = 27±4ILM D Apparent V,,,,,, = 33 ± 3 pmoilmlnimg protein Apparent K,,, 111±45ILM I . 30 30 10 0 a C 0 a 400 U0 .2 .9 30000 40 c. C 0 200 9 . 100 C’3 — 0 0 70 60 0.4 (6 uE .C0 0 (Chenodeoxycholic acid] (pM) Figure 3.4. Enzyme kinetic profiles of 7a-hydroxy-3-oxo-5f3-cbolan-24-oic acid (A), ‘y muricholic acid (B), 7-ketolithocholic acid (C) and cholic acid (D) formation by human hepatic microsomes. Metabolite formation (activity) was plotted as a function of substrate concentration following a 30-mm incubation with human liver microsomes (0.5 mg). Data points are the mean ± SEM of at least three separate experiments. Lines represent rates modeled by nonlinear regression analysis of the data. Kinetic parameters for 7cx-hydroxy-3-oxo-5 13-cholan-24-oic acid, 7- ketolithocholic acid and cholic acid formation were calculated using equation 1. Kinetic parameters for y-muricholic acid formation were calculated using equation 2. The insets depict Eadie-Hofstee plots. Error bars are not shown on the insets to avoid obscuring the data points, which represent mean values. 30 00 20 .zs . 0 10 200 30 25 20 15 10 S 600 800 0.0 200 0.1 02 0.3 “Is 400 0.4 0.5 600 800 (Chenodeoxycholic acid] (pM) 1000 124 3-Dehydrocholic acid 25 A 20 15 22 I[0 5 0 .5 .5 — N D 00 O O D — — . — N N N N Q N N eon. n.n..o.n.Na. 0.0. Q oc3(.)oOc1 OOU 7x.Hydroxy4.oxo.510.cholan.24-oic acid 30 r-Murlchollc acid 25 B 20 2E 0. 15 .0 10 5 0 . . r.— r—. .5 5 — N D CO Q (0 — (0 — 0 0 0.0.0.0. 0. 0. N 0. 0. 0. 0. o 0)-,->-).>-)-n.)-)-)->-0. C)OUOC)OUQQO Figure 3.5. Comparison of 3-dehydrocholic acid formation from cholic acid (A) and 7o- hydroxy-3-oxo-5f3-cholan-24-oic acid and y-muricholic acid formation from chenodeoxycholic acid (B) by a panel of recombinant human P450 enzymes. Metabolite formation (activity) was measured following a 30-mm incubation of cholic acid or chenodeoxycholic acid (500 jtM) with baculovirus-insect cell microsomes containing expressed human P450 enzymes (30 pmol). Plots show the mean values ± SEM of triplicate determinations. 125 30 0 0. 0— 00 I 0 ‘4, [Cholic acid] (pM) Figure 3.6. Enzyme kinetic profile of 3-dehydrocholic acid formation from cholic acid by recombinant human CYP3A4. Metabolite formation (activity) was plotted as a function of substrate concentration following 30-mm incubation with recombinant CYP3A4 (30 pmol). Data points are the mean ± SEM of triplicate determinations. Lines represent rates modeled by nonlinear regression analysis of the data. Kinetic parameters for 3-dehydrocholic acid formation by recombinant CYP3A4 were calculated using equation 2. The inset depicts the Eadie-Hofstee plot. Error bars are not shown on the inset to avoid obscuring the data points, which represent mean values. Apparent V1f = 26 ± 4 pmollminlpmol P450 Apparent K’ = 161 ± 5 LLM n = 1.725 20 15 10 5 0 . . 30 25 20 15 10 5 0 a 0 100 200 300 400 500 600 700 126 A Apparent = 40±3 pmollmin(pmol P450 ApparentK, = 165±3liiM B 5 4 Apparent Vmx = 5 ± 0.4 pmouninipmol P450 Apparent K’ = 208±20M n = 2±0.3 3 20 25 20 Is 10 5 C 0 Q. . e. o — 2 0.05 0.10 0.55 020 025 030 00 100 200 300 400 500 600 100 200 300 400 500 [Chenodeoxycholic acidj (pM) [Chenodeoxycholic acid] (pM) C 0 40 .2 •0 30 9. 20 a o Xe 9 E 10 r, 0. X0 X 0 r 600 Figure 3.7. Enzyme kinetic profiles of 7a-hydroxy-3-oxo-53-cho1an-24-oic acid (A) and y-muricholic acid (B) formation from chenodeoxycholic acid by recombinant human CYP3A4. Metabolite formation (activity) was plotted as a function of substrate concentration following a 30-mm incubation with recombinant CYP3A4 (30 pmol). Data points are the mean ± SEM of triplicate determinations. Lines represent rates modeled by nonlinear regression analysis of the data. Kinetic parameters for 7cL-hydroxy-3-oxo-5 f3-cholan-24-oic acid formation by recombinant CYP3A4 were calculated using equation 1. Kinetic parameters for y-muricholic acid formation by recombinant CYP3A4 were calculated using equation 2. The insets depict Eadie-Hofstee plots. Error bars are not shown on the insets to avoid obscuring the data points, which represent mean values. 127 HO• P3A4—’ Ox::SoTh OOH Cholic acid 3-Dehydrocholic acid (3a, 7a,, I 2a-tri hydroxy- (7a, I 2a-di hydroxy-3-oxo - 5p-chol an-24-oic acid) 5p-chol an-24-oic acid) Figure 3.8. Scheme showing P450-mediated cholic acid biotransformation by human hepatic microsomes. Results of the present study suggest that CYP3A4 is the only enzyme involved in 3- dehydrocholic acid formation. 128 XD1Th H OH y-Muricholic acid CYP3A4 and 21 CYP3 (3ci;6a,7a-trihydroxy-7cz-hydroxy-3-oxo- 5f3-cholan-24-oic acid)5p-cholan-24-oic acid CYP3A5 12 • 120 ‘7COOH HO4YfOH Chenodeoxycholic acid 513-cholan-24-oic acid) / (3cc,7a-dihydroxy- HO_cTh OH 7-Ketolithocholic acidCholic acid (3x-hydroxy-7-oxo-(3a,7ct,1 2a-trihydroxy- 5p-cholan-24-oic acid)5f3-cholan-24-oic acid) Figure 3.9. Scheme showing chenodexycholic acid biotransformation by human hepatic microsomes. Results of the present study suggest that formation of 7ct-hydroxy-3-oxo 513-cholan-24-oic acid and ‘y-muricholic acid was mediated mainly by CYP3A4. Enzymes involved in the formation of 7-ketolithocholic acid and cholic acid have not been determined. 129 3.6 References Alme B and Sjovall J (1980) Analysis of bile acid glucuronides in urine. Identification of 3 alpha, 6 alpha, 12 alpha-trihydroxy-5 beta-cholanoic acid. J Steroid Biochem 13:907- 916. Bodin K, Lindbom U and Diczfalusy U (2005) Novel pathways of bile acid metabolism involving CYP3A4. Biochim Biophys Acta 1687:84-93. Bremmelgaard A and Sjovall 3 (1980) Hydroxylation of cholic, chenodeoxycholic, and deoxycholic acids in patients with intrahepatic cholestasis. JLipid Res 21:1072-1081. Deo AK and Bandiera SM (2008) Biotransformation of lithocholic acid by rat hepatic microsomes: metabolite analysis by liquid chromatography/mass spectrometry. Drug Metab Dispos 36:442-451. Ellis E, Axelson M, Abrahamsson A, Eggertsen G, Thorne A, Nowak G, Ericzon BG, Bjorkhem I and Einarsson C (2003) Feedback regulation of bile acid synthesis in primary human hepatocytes: evidence that CDCA is the strongest inhibitor. Hepatology 38:930-938. Fiorucci S, Clerici C, Antonelli E, Orlandi S, Goodwin B, Sadeghpour BM, Sabatino G, Russo G, Castellani D, Willson TM, Pruzanski M, Pellicciari R and Morelli A (2005) Protective effects of 6-ethyl chenodeoxycholic acid, a farnesoid X receptor ligand, in estrogen-induced cholestasis. JPharmacol Exp Ther 313:604-612. Fischer S, Beuers U, Spengler U, Zwiebel FM and Koebe HG (1996) Hepatic levels of bile acids in end-stage chronic cholestatic liver disease. Clin Chim Acta 251:173-186. Grober J, Zaghini I, Fujii H, Jones SA, Kliewer SA, Willson TM, Ono T and Besnard P (1999) Identification of a bile acid-responsive element in the human ileal bile acid-binding protein gene. Involvement of the famesoid X receptor/9-cis-retinoic acid receptor heterodimer. JBiol Chem 274:29749-29754. Guengerich FP (1995) Human Cytochrome P450 Enzymes. Plenum Press, New York. 130 Guengerich FP (2003) Cytochromes P450, drugs, and diseases. Mol Interv 3:194-204. Hofinann AF (1999) Bile Acids: The Good, the Bad, and the Ugly. News Physiol Sd 14:24-29. Hofmann AF (2002) Cholestatic liver disease: pathophysiology and therapeutic options. Liver 22 Suppl 2:14-19. Hrycay EG and Bandiera SM (2008) Cytochrome P450 Enzymes, in Preclinal Development Handbook: ADME and Biopharmaceutical Properties (Gad, SC ed), pp 627-696, John Wiley & Sons, Inc., Hoboken, NJ. Martin KO, Budai K and Javitt NB (1993) Cholesterol and 27-hydroxycholesterol 7 alpha hydroxylation: evidence for two different enzymes. JLipid Res 34:5 8 1-5 88. Norlin M and Wikvall K (2007) Enzymes in the conversion of cholesterol into bile acids. Curr MolMed7:199-218. Pellicciari R, Fiorucci S, Camaioni E, Clerici C, Costantino G, Maloney PR, Morelli A, Parks DJ and Wilison TM (2002) 6alpha-ethyl-chenodeoxycholic acid (6-ECDCA), a potent and selective FXR agonist endowed with anticholestatic activity. J Med Chem 45:3569- 3572. Ridlon JM, Kang DJ and Hylemon PB (2006) Bile salt biotransformations by human intestinal bacteria. JLipidRes 47:24 1-259. Rizzo G, Renga B, Mencarelli A, Pellicciari R and Fiorucci S (2005) Role of FXR in regulating bile acid homeostasis and relevance for human diseases. Curr Drug Targets Immune Endocr Metabol Disord 5:289-303. Schwarz M, Russell DW, Dietschy JM and Turley SD (2001) Alternate pathways of bile acid synthesis in the cholesterol 7alpha-hydroxylase knockout mouse are not upregulated by either cholesterol or cholestyramine feeding. JLipid Res 42:1594-1603. 131 Shoda J, Tanaka N, Osuga T, Matsuura K and Miyazaki H (1990) Altered bile acid metabolism in liver disease: concurrent occurrence of C-i and C-6 hydroxylated bile acid metabolites and their preferential excretion into urine. JLipid Res 31:249-259. Sinal CJ, Tohkin M, Miyata M, Ward JM, Lambert G and Gonzalez FJ (2000) Targeted disruption of the nuclear receptor FXRIBAR impairs bile acid and lipid homeostasis. Cell 102:731-744. Tint GS, Xu GR, Batta AK, Shefer S, Niemann W and Salen G (1990) Ursodeoxycholic acid, chenodeoxycholic acid, and 7-ketolithocholic acid are primary bile acids of the guinea pig. JLipidRes 31:1301-1306. Tracy TS and Hummel MA (2004) Modeling kinetic data from in vitro drug metabolism enzyme experiments. Drug Metab Rev 36:231-242. Wietholtz H, Marschall HU, Sjovall J and Matern S (1996) Stimulation of bile acid 6 alpha hydroxylation by rifampin. JHepatol 24:713-7 18. 132 Chapter 4 3-KETOCHOLANOIC ACID IS THE MAJOR CYP3A4 MEDIATED LITHOCHOLIC ACID METABOLITE IN HUMAN HEPATIC MICROSOMES4 4A version of this chapter will be submitted for publication as: Deo AK and Bandiera SM (2009) 3-Ketocholanoic Acid is the Major CYP3A4 Mediated Lithocholic Acid Metabolite in Human Hepatic Microsomes. Drug Metab Dispos. 133 4.1 Summary Biotransformation of endogenous bile acids by hepatic enzymes is a potential pathway for effective elimination of hepatotoxic lithocholic acid. In the present study, unconjugated human hepatic microsomal metabolites of lithocholic acid, and the enzymes involved in their formation, were identified. An optimized method to analyze metabolite formation using liquid chromatography-mass spectrometry (LC/MS) was utilized. Reaction mixtures contained 50 mM potassium phosphate buffer, pH 7.4, 3 mM MgCI2, 0.5 mg of human hepatic microsomal protein, 1 mM NADPH and varying concentrations (1-250 i.iM) of lithocholic acid, in a final volume of 1 ml, incubated for 30 mm. Incubations with a panel of human recombinant P450 enzymes (CYPIA1, CYPIA2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2CI9, CYP2D6, CYP2E1, CYP3A4, CYP3A5 or CYP4A1 1) along with suitable controls were carried out. LCIMS analysis using human liver microsomal incubations revealed the formation of 3- ketocholanoic acid as the major metabolite followed by hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid. Recombinant CYP3A4 was identified as the only enzyme involved in the formation of all five metabolites. Chemical inhibition studies using ketoconazole, quercetin, sulfaphenazole and quinidine showed that ketoconazole inhibited overall CYP3A4 mediated biotransformation of lithocholic acid. Rate of metabolite formation was increased in hepatic microsomal samples containing greater CYP3A4 activity (13700 pmol/minlmg protein) and decreased in samples with lower CYP3A4 activity (650 pmollminlmg protein). The preferred position of lithocholic acid biotransformation by human hepatic microsomal P450 enzymes was the third position on the cholestane ring. 134 4.2 Introduction Lithocholic acid (3c-hydroxy-53-cholan-24-oic acid), together with other biliary bile acids, facilitates the absorption, transport, and distribution of lipid-soluble nutrients from the diet and aids in the elimination of cholesterol from the body (Hofhiann, 1 999b). Lithocholic acid is formed from chenodeoxycholic acid, by the action of bacterial dehyratases in the colon (Hofmann, 1 999b). Lithocholie acid is absorbed from the colon and transported to the liver, where it is conjugated with taurine or glycine before being excreted as a component of bile (Hofmann, 2002; Hofmann, 2004). In spite of its beneficial role, lithocholic acid is cytotoxic (Hoflnann, 1999a). The accumulation of lithocholic acid and other hydrophobic bile acids in liver due to cholestasis has been implicated as a major factor contributing to liver injury. Cholestasis is a common manifestation of many liver disesases such as primary biliary cirrhosis, viral hepatitis and alcoholic liver disease. Hepatic toxicity was also shown in experimental animals following chronic and acute administration of exogenous lithocholic acid or its conjugates (Javitt, 1966; Palmer and Ruban, 1966; Zaki et al., 1967; Miyai et al., 1971; Fischer et al., 1974). The hepatotoxicity associated with bile acids can be attenuated by Phase I biotransformation pathways involving hydroxylation catalyzed by hepatic microsomal P450 enzymes and by phase II biotransformation pathways involving conjugation with sulfate. A previous study showed that incubation of lithocholic acid with human hepatic microsomes yielded hyodeoxycholic acid, murideoxycholic acid and chenodeoxycholic acid as hydroxylated metabolites (Xie et al., 2001). 6a-Hydroxylation of lithocholic acid to form hyodeoxycholic acid by human recombinant CYP3A4 and human liver microsomes was reported to be the major route of lithocholic acid biotransformation (Araya and Wikvall, 1999; Xie et al., 2001). 135 More recently, a different metabolite, 3-ketocholanoic acid, was identified as the major metabolite of lithocholic acid using human recombinant CYP3A4 (Bodin et a!., 2005). Formation of other lithocholic acid metabolites, chenodeoxycholic acid and murideoxycholic acid as suggested by Xie et al. (2001) was not reported by this study. In rat, we demonstrated that hepatic biotransformation of lithocholic acid was extensively catalyzed by multiple P450 enzymes. Lithocholic acid biotransformation resulted in the formation of the 6j3-hydroxylated metabolite, murideoxycholic acid, as the predominant pathway with trace amounts of hyodeoxycholic acid (6cL-hydroxylation) and a complete absence of chenodeoxycholic acid (Deo and Bandiera, 2008a). Results obtained with inhibitory antibodies, recombinant enzymes and P450 enzyme inducers demonstrated that CYP2C, CYP2D and CYP3A enzymes contributed to microsomal lithocholic acid biotransformation. Conflicting data on identification of the major pathway of lithocholic acid biotransformation and its metabolite profile in humans has been reported. Moreover, based on our results obtained using rat hepatic microsomes (Deo and Bandiera, 2008a), a role of multiple P450 enzymes in human hepatic microsomal lithocholic acid biotransformation can be speculated. To resolve these issues, lithocholic acid metabolite formation by human hepatic microsomes was investigated using a liquid chromatography-mass spectrometry (LCJMS) method developed previously (Deo and Bandiera, 2008a; Deo and Bandiera, 2008b). The contribution of P450 enzymes in hepatic microsomal biotransformation of lithocholic acid was assessed using recombinant human enzymes and chemical inhibitors. In addition, kinetic parameters associated with rates of metabolite formation in human hepatic microsomes and recombinant enzymes were also calculated. 136 4.3 Materials and Methods 4.3.1 Chemicals and reagents. Lithocholic acid and unconjugated bile acid standards were purchased from Steraloids Inc. (Newport, RI). la,3f3,7c,12cL-tetrahydroxy-5f3-cholan-24-oic acid and 3cx,6,713,12a-tetrahydroxy-53-cho1an-24-oic acid were generous gifts from Dr. Lee R. Hagey (University of California, San Diego, CA). Bile acid standards were dissolved in methanol as 1 mg/mi stock solutions and stored at -4°C. Additional dilutions were made in methanol for the biotransformation assay. Ketoconazole, SKF-525A, quercetin, quinidine, and sulfaphenazole were generously provided by Dr. T.K.H Chang (University of British Columbia, Vancouver, BC, Canada). Stock solutions of these chemical P450 inhibitors were prepared in methanol. Pooled human liver microsomes were purchased from. Xenotech (Lenexa, KS). Baculovirus-insect cell control microsomes containing expressed human P450-oxidoreductase and baculovirus-insect cell microsomes containing expressed human P450 enzymes (BD SUPERSOMESTM Enzymes), co-expressed with human P450-oxidoreductase or with human P450-oxidoreductase and human cytochrome b5, were purchased from BD Biosciences (Oakvilie, Ontario, Canada). Four human iiver microsome samples from single donors (donor no. 111118, HG95, HH13, HH837) were obtained from BD Biosciences (Woburn, MA). Donor information provided by BD Biosciences is summarized below. HH18 (African American female, 78 years of age) and HH837 (Asian female, 52 years of age) had high CYP3A4- dependent testosterone 613-hydroxylase activity of 12000 pmol/minlmg protein and 13700 pmoilminlmg protein, respectively, HG95 (Hispanic female, 47 years of age) and FIH 13 (Asian male, 55 years of age) had low CYP3A4-catalyzed activities of 650 pmol!minlmg protein and 890 pmollminlmg protein, respectively, as reported by BD GentestTM. High-performance liquid chromatography-grade chemicals and solvents were purchased from Fisher Scientific (Ottawa, Ontario, Canada). 137 4.3.2 Lithocholic acid biotransformation assay. Reaction mixtures contained 50 mM potassium phosphate buffer, pH 7.4, 3 mM magnesium chloride, 0.5 mg of human hepatic microsomal protein, 1 mM NADPH and varying concentrations (1-250 M) of lithocholic acid, in a final volume of 1 ml. After preincubation for 10 mm at room temperature, reactions were initiated with NADPH and allowed to proceed for 30 mm at 37°C. Reactions were terminated with 8 ml of dichloromethane/isopropanol (80:20 v/v). Internal standard, (cholic-2,2,4,4-d4acid, 0.4 pjg), was then added to each sample. Sample extraction, evaporation, and reconstitution in preparation for analysis by LCIMS were carried out as described previously (Deo and Bandiera, 2008a). Reaction mixtures devoid of substrate, NADPH or microsomes, as well as reaction mixtures containing defined concentrations of authentic bile acid standards, were routinely included in each assay. Assay conditions were tested using pooled human liver microsomes to ensure that substrate and cofactor concentrations were saturating and that product formation was linear with respect to incubation time (1 to 60 mm) and protein concentration (0.25 to 2 mg/mI of reaction mixture). To determine if metabolite formation was P450-mediated, preliminary experiments were conducted with carbon monoxide treated hepatic microsomes or heat-denatured microsomes or by replacing NADPH with NADH or by adding SKF-525A, a P450 inhibitor. Incubations with human recombinant P450 enzymes were also carried out. Reaction mixtures contained 30 pmoles of recombinant P450 enzyme (CYP1A1, CYPIA2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2C19, CYP2D6, CYP2E1, CYP3A4, CYP3A5 or CYP4A 11) or, in the case of insect cell control microsomes and reductase control, an equivalent amount of protein (0.15 mg). 138 4.3.3 Analytical methods. LCIMS conditions to resolve and quantif’ the metabolites were used as outlined previously (Deo and Bandiera, 2008a; Deo and Bandiera, 2008b). A mixture consisting of 19 bile acid standards prepared for identification of bile acids. Under these conditions, hx,33,7ct,12c-tetrahydroxy-5J3-cholan-24-oic acid (molecular mass 424.6) and 3c,6I3,713, 1 2c-tetrahydroxy-5 13-cholan-24-oic acid (molecular mass 424.6) typically eluted at 4 and 5 mm, respectively, and were monitored at m/z 423. c-Muricholic acid (molecular mass 408.6), 13-muricholic acid (molecular mass 408.6), y-muricholic acid (molecular mass 408.6) and cholic acid (molecular mass 408.6) eluted at 12, 13, 15 and 16 mm, respectively, and were monitored at m/z 407. 3-Dehydrocholic acid (molecular mass 406.6) eluted at 13 mm and was monitored at m/z 405. Murideoxycholic acid (molecular mass 392.6), ursodeoxycholic acid (molecular mass 392.6), hyodeoxycholic acid (molecular mass 392.6), chenodeoxycholic acid (molecular mass 392.6) and deoxycholic acid (molecular mass 392.6) eluted at 12, 13.4, 15, 19.5, 20 mm, respectively, and were monitored at m/z 391. 6-Ketolithocholic acid (molecular mass 390.6), 7-ketolithocholic acid (molecular mass 390.6) and 7a-hydroxy-3-oxo-5 3-cholan- 24-oic acid (molecular mass 390.6) eluted at 13.4, 14.5 and 16 mm, respectively, and were monitored at m/z 389. Isolithocholic acid (molecular mass 376.6) and lithocholic acid (molecular mass 376.6) eluted at 20.5 and 22.5 mm, respectively, and were monitored at m/z 375. 3-Ketocholanoic acid (molecular mass 374.6) eluted at 21.5 mm and was monitored at m/z 373. The internal standard, cholic-2,2,4,4-d4acid, eluted at 15.4 mm and was monitored at m/z 411. Metabolites were quantified from calibration piots of the peak area ratio of authentic standard and internal standard plotted against the concentration of the authentic standard. 4.3.4 Data analysis and calculation of enzyme kinetic parameters. Data were analyzed using the SigmaPlot® Enzyme Kinetics Module (v.1.1, Systat Software Inc., Richmond, CA). 139 Metabolite formation as a function of substrate concentration was analyzed by nonlinear regression analysis and apparent Km, K’ and Vmax values were generated using the Michaelis Menten equation (Equation 1), or the Hill equation (Equation 2) as described previously (Houston and Kenworthy, 2000). VmX[S] — Km + [S] (equation 1) VmX[S1 — K’ + [S] (equation 2) where v is initial velocity of the reaction, Vmm is the maximal velocity, [Sj is the substrate concentration, K’ is the Hill dissociation constant, n is the Hill coefficient representing cooperativity of the reaction, Km is the Michaelis-Menten constant. Several criteria such as sum of squares (R2), standard deviation of residuals (Sy.X), for each equation, Akaike Information Criterion (AIC) and visual inspection of the fit were used to choose the appropriate fitting model (Tracy and Hummel, 2004). 4.3.5 Chemical inhibition studies. P450 chemical inhibition studies were carried out using human liver microsomes. Chemical inhibitors used were quercetin (2, 20 and 50 .tM, a selective CYP2CS inhibitor), sulfaphenazole (1, 10 and 20 tiM, a selective CYP2C9 inhibitor), quinidine (0.1, 1 and 10 jiM, a selective CYP2D6 inhibitor) and ketoconazole (0.01, 0.1, 1 and 10 jiM, a selective CYP3A4 inhibitor) at final concentrations indicated in parentheses. Reactions were initiated with NADPH after initial pre-incubation of microsomes with lithocholic acid and the P450 inhibitor for 5 mm at 37°C. 140 4.3.6 Statistical analysis. Statistical analysis of data was performed using GrapliPad Prism®. Comparison of chemical inhibition data with the control was carried out using ANOVA. A p value of 0.05 was considered statistically significant. 4.4 Results 4.4.1 Lithocholic acid biotransformation by human liver microsomes. Incubation of lithocholic acid with human liver microsomes yielded five unconjugated metabolites identified as 3-ketocholanoic acid, hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid. Of these, 3-ketocholanoic acid was identified as the major metabolite, followed by hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6- ketolithocholic acid, respectively, as minor metabolites (Fig. 4.1). No metabolite formation was observed in reaction mixtures devoid of substrate, NADPH, or microsomes. Peaks with retention times of 19.5 mm (m/z of 391) and 14.5 mm (m/z 389) corresponded to chenodeoxycholic acid and 7-ketolithocholic acid respectively, which were determined to be contaminants of lithocholic acid as reported previously (Deo and Bandiera, 2008a). Considering that tetrahydroxy metabolites (such as 1 c,3 13,7cc, 1 2a-tetrahydroxy-5 13- cholan-24-oic acid and 3a,6 12x-tetrahydroxy-53-cholan-24-oic acid) of the hydrophobic lithocholic acid could be formed we scanned for, but did not observe any metabolite peaks in the corresponding ion channels with m/z 423. Metabolite formation was evaluated over a substrate concentration range of 1-250 tM. An incubation time of 30 mm and a microsomal protein concentration of 0.5 mg/mI were found to be optimal and were used for all subsequent experiments. A lithocholic acid concentration of 100 jiM was found to be saturating for all metabolites under consideration and this was used for 141 any further experiments. Plots of metabolite formation versus substrate concentration showed that hepatic microsomal formation of 3-ketocholanoic acid, hyodeoxycholic acid, ursodeoxycholic acid, 6-ketolithocholic acid, and murideoxycholic acid followed Michaelis Menten kinetics (Fig. 4.2, A-E). The apparent Vm value (336.2 ± 13.6) for 3-ketocholanoic acid formation was 72 times larger than for 6-ketolithocholic acid formation, 45 times larger than for murideoxycholic acid, 23 times larger than for ursodeoxycholic acid and 7 times larger than for hyodeoxycholic acid formation, respectively. The low apparent Km value (21.9 ± 2.6 iiM) associated with 3- ketocholanoic acid formation and its high Vm value demonstrated that 3-ketocholanoic acid was the predominant microsomal metabolite of lithocholic acid in human liver (Table 4.1). 4.4.2 Lithocholic acid biotransformation by human recombinant P450 enzymes. The contribution of individual P450 enzymes to lithocholic acid biotransformation was evaluated using a panel of twelve human recombinant P450 enzymes as described under Materials and Methods. Initial experiments were conducted to determine P450 concentrations that would ensure linearity of product formation. An incubation time of 20 mm and a recombinant P450 enzyme concentration of 30 pmol P450/ml were found to be optimal at a substrate concentration of 100 M. Conversion of lithocholic acid to 3-ketocholanoic acid, hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid was catalyzed solely by CYP3A4 (Fig. 4.3). Formation of these metabolites by recombinant CYP3A4, evaluated over a range of substrate concentrations (1-250 jtM), exhibited typical Michaelis-Menten type kinetics with the exception of murideoxycholic acid, which followed sigmoidal kinetics (Fig. 4.4 A-E). Kinetic parameters for the formation of these metabolites by recombinant CYP3A4 are listed in Table 142 4.2. 3-Ketocholanoic acid was the major metabolite produced by recombinant CYP3A4 (27.2 pmol/minlpmol P450), followed by hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid. 4.4.3 Chemical inhibition studies. The effect of chemical P450 enzyme inhibitors on lithocholic acid biotransformation was evaluated by incubating lithocholic acid with human hepatic microsomes in the presence of ketoconazole, quercetin, quinidine and sulfaphenazole. The rate of formation of 3-ketocholanoic acid was inhibited by 70% at 10 iM ketoconazole, and by 48% at 50 M quercetin (Fig. 4.5A). The rate of formation of hyodeoxycholic acid was inhibited by 95% using 10 iM ketoconazole and 60% with 50 iM quercetin (Fig. 4.5B). 6- Ketolithocholic acid was not detected at higher concentrations of ketoconazole (10 aiM). The CYP2D6 and CYP2C9 inhibitors, quinidine and sulfaphenazole, respectively, had no effect on rates of formation of any metabolites as compared to the control (no inhibitor). Recombinant CYP3A4 was incubated with lithocholic acid in the presence of quercetin (a CYP2C8 inhibitor) to determine its inhibitory effect on metabolite formation. Formation of all five lithocholic acid metabolites was observed to be decreased in presence of quercetin (20 jiM and 50 jiM final concentration). The rates of formation of 3-ketocholanoic acid and hyodeoxycholic acid were decreased by 60% and 70%, respectively, using 50 jiM quercetin and were decreased by 50% and 40%, respectively, using 20 jiM quereetin (data not shown). 4.4.4 Lithocholic acid biotransformation in human hepatic microsomes with varying CYP3A4 levels. The metabolite profile obtained using hepatic microsomes obtained from individual human hepatic donors HH1 8, HH1 3, HH83 7 and HG95 were determined to relate the rate of metabolite formation to the activity of CYP3A4 in each of the donor samples. Formation 143 of all five lithocholic acid metabolites increased in microsomal samples with relatively high CYP3A4-dependent testosterone 63-hydroxylase activity (HH1 8 and HH837) than in microsomal samples with low CYP3A4-linked activity (HG95 and HH13) (Table 4.3). The correlation between testosterone 613-hydroxylase activities and rates of metabolite formation was greater than 0.95 for each of the five lithocholic acid metabolites suggesting that the same P450 enzyme catalyzed testosterone 63-hydroxylation and lithocholic acid biotransformation. The average 17-fold increase in CYP3A4 activity in microsomal samples with low activity (HG95 and HH 13) as compared to the samples with greater activity (HH 18 and F1H83 7) was reflected with approximately a 6-fold increase in rate of 3-ketocholanoic acid formation, 23-fold increase in hyodeoxycholic acid formation, 6-fold increase in ursodeoxycholic acid formation and 4-fold increase in murideoxycholic acid and 6-ketolithocholic acid formation. 4.5 Discussion Lithocholic acid was extensively biotransformed by human hepatic microsomes to five metabolites. The major metabolite was 3-ketocholanoic acid. The minor metabolites were hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid. The predominant biotransformation pathway was observed to be the oxidation of lithocholic acid on the third carbon leading to the formation of 3-ketocholanoic acid. Formation of 3- ketocholanoic acid could be through a geminal diol intermediate on the third carbon of lithocholic acid which further dehydrates to a 3-ketone as suggested by Bodin et al., 2005. Our results differ from the data published by Xie et al. (2001) and Araya and Wikvall (1999). Both studies reported that lithocholic acid was primarily converted to hyodeoxycholic acid by 6c-hydroxylation. Xie et al. (2001) further reported the formation of murideoxycholic 144 acid (63-hydroxylation) and chenodeoxycholic acid (7cL-hydroxylation) by human hepatic microsomes. Our results indicate that chenodeoxycholic acid is not a lithocholic acid metabolite in human liver microsomes. Thus, we consider that previous reports suggesting chenodeoxycholic acid formation may have overlooked the possibility of having an impure substrate. Previous studies have not reported the formation of other metabolites such as 3- ketocholanoic acid, ursodeoxycholic acid, and 6-ketolithocholic acid from human liver microsomes. The use of a simple LC/MS based assay with single ion recording at various m/z facilitated our studies in this regard, as compared to thin layer chromatography and complex gas chromatography/mass spectrometry procedures used earlier (Araya and Wikvall, 1999; Xie et al., 2001). Our data using recàmbinant human P450 enzymes partially agree with reports showing the involvement of recombinant CYP3A4 in 6ix-hydroxylation of lithocholic acid (Araya and Wikvall, 1999; Xie et al., 2001). However, we also highlight that 6ct-hydroxylation to form hyodeoxycholic acid is a minor pathway of lithocholic acid biotransformation by recombinant CYP3A4. Our studies further add that all five lithocholic acid metabolites including the ones that were not investigated earlier (i.e. 3-ketocholanoic acid, ursodeoxycholic acid and 6- ketolithocholic acid) are formed through recombinant CYP3A4 mediation with 3-ketocholanoic acid being the major metabolite (Fig. 4.3). Chemical inhibition studies were carried out in order to confirm the role of CYP3A4 in lithocholic acid biotransformation. Inhibitor concentrations were chosen in the range reported previously (Liu et al., 2007; Wang et al., 2008). All lithocholic acid metabolites were inhibited by the CYP3A4 inhibitor, ketoconazole. No effect of quinidine (CYP2D6 inhibitor) and sulfaphenazole (CYP2C9 inhibitor) was observed on the formation of any of the metabolites. However, quercetin (CYP2C8 inhibitor) seemed to inhibit lithocholic acid metabolism at higher 145 concentrations (20 iiM and 50 ji.M). To understand its involvement, lithocholic acid was incubated with recombinant CYP3A4. The decrease in formation of metabolites using recombinant CYP3A4 and quercetin as inhibitor is suggestive of non-specific quercetin mediated P450 inhibition. Inhibition by ketoconazole, a widely used drug for treating skin and anti-fungal infections, seems to be of clinical importance. Ketoconazole has been implicated in a number of hepatic dysfunctions. Ketoconazole-induced hepatic cholestatic injury has been reported in humans (Stricker et al., 1986; Bensaude et al., 1988; Findor et al., 1998). In vivo studies carried out in rats treated with ketoconazole (25-50 mg/kg) showed elevated levels of bile acids such as cholic acid, taurocholic acid, chenodeoxycholic acid and taurochenodeoxycholic acid in serum (Azer et al., 1995). Considering the potential of ketoconazole to cause similar effects in human, combined with our results showing the ability of ketoconazole to inhibit CYP3A4-mediated bile acid metabolism, suggest that prescribing ketoconazole in cholestasis may increase propensity towards severe irreversible liver damage. A comparison of metabolite formation patterns of hepatotoxic bile acids (cholic acid, chenodeoxycholic acid and lithocholic acid) reveals that formation of 3-oxo metabolites is the key biotransformation pathway for bile acids in human liver microsomes. 3-Ketocholanoic acid was the major metabolite of lithocholic acid in this study. Earlier we observed the formation of 3-dehydrocholic acid from eholic acid and 7c*-hydroxy-3-oxo-53-cholan-24-oic acid from chenodeoxycholic acid as major biotransformation pathways in human liver microsomes. All these major biotransformation pathways were CYP3A4 mediated. Lithocholic acid concentrations of 5 to 10 iM have been reported in the liver of cholestatie patients and in rat models of biliary cholestasis (Jezequel et al., 1994; Fischer et al., 1996; Erlinger, 1997; Setehell et al., 1997; Hofmann, 2002; Berta et a!., 2003; Rost et al., 2003; De Gottardi et al., 2004). A lithocholic acid concentration of 100 M was found to be 146 saturating for hepatic microsomal 3-ketocholanoic acid formation in the present study. Nevertheless, 3-ketoeholanoic acid was the major metabolite obtained in the in vitro biotransformation assay, with a rate of formation of 40-50 pmol/min!mg at a lithocholic acid concentration of 5 jiM, which approximates the physiological hepatic concentration. Based on the results obtained in this study, a scheme for biotransformation of lithocholic acid in human hepatic microsomes is proposed in Fig. 4.6. The results provide compelling evidence that lithocholic acid can serve as a physiological substrate of CYP3A4 with 3- ketocholanoic acid as its major metabolite. Adult human liver and intestine are the main organs in which CYP3A4 is expressed. CYP3A4 contributes to the metabolism of more than 40% of drugs and endogenous chemicals (Guengerich, 2003). Inter-individual variability in specific content of CYP3A4 in Caucasians has been observed (Shimada et al., 1994). The range of hepatic and intestinal CYP3A4 expression varies as much as 40%. Our results show that the average 17-fold increase in CYP3A4 activity in microsomal samples with low activity (HG95 and HHI3) as compared to the samples with higher activity (HHI8 and 1*1837) was reflected with approximately a 6-fold increase in rate of formation of the major lithocholic acid metabolite, 3-ketocholanoic acid (Table 4.3). Inter-individual variability in CYP3A4 activity affects intrinsic clearance of certain drugs such as midazolam (Lamba et a!., 2002). Similarly, variation in CYP3A4 levels may affect the clearance and elimination of bile acids such as lithocholic acid. Altered levels of CYP3A4 have been observed due to changes in nutrition, diseases, environmental factors and exposure to chemicals such as phenobarbital, rifampicin and phenytoin, or alternately, inhibition of CYP3A4 activity by antifungal agents or macrolide antibiotics. Induction or inhibition of CYP3A4 activity by these factors may change the rates of lithocholic acid metabolite formation. 147 CYP3A4 expression is subject to regulation by the nuclear receptors such as pregnane X receptor (PXR), farnesoid X receptor (FXR) and vitamin D receptor (VDR), which are activated by several drugs and other xenobiotics (Staudinger et al., 2001; Makishima et a!., 2002; Matsubara et al., 2008). Lithocholic acid and its major metabolite, 3-ketocholanoic acid are known to activate PXR, FXR and VDR that in turn regulate bile acid mediated liver toxicity in cholestasis (Staudinger et al., 2001; Makishima et al., 2002). The minor lithocholic acid metabolite, 6-ketolithocholic acid also activates VDR but not FXR (Makishima et al., 2002). Another minor metabolite, ursodeoxycholic acid, is considered to be clinically useful in the treatment of chronic cholestatic disease and biliary cirrhosis (Calmus and Poupon, 1991; Festi et a!., 2007). A potential role for ursodeoxycholic acid in inducing human hepatic CYP3A enzymes and eventually assisting the elimination of hepatotoxic bile acids needs to be investigated. Studies identif’ing the role of hyodeoxycholic acid and murideoxycholic acid either in receptor activation or P450 induction are not available. The data generated in this study highlights the major role of CYP3A4 in human hepatic microsomal biotransformation to all lithocholic acid metabolites. The current work also resolves conflicting issues about the major lithocholic acid biotransformation pathways reported by previous studies (Araya and Wikvall, 1999; Xie et al., 2001; Bodin et al., 2005). The results generated in this study may serve as a useful tool for researchers studying bile acid regulation and assist in identifying mechanisms associated with detoxification of hepatotoxic bile acids. 148 Metabolite Apparent Vmct. Apparent Kmpmol/min/mg protein pM 3-Ketocholanoic acid 336.2 ± 13.6 21.9 ± 2.6 Hyodeoxycholic acid 47.5 ± 4.3 46.3 ± 13.3 Ursodeoxycholic acid 14.6 ± 0.6 32.6 ± 5.2 Murideoxycholic acid 5.5 ± 0.2 18.2 ± 3.2 6-Ketolithocholic acid 4.7 ± 0.2 39.4 ± 4.8 Table 4.1. Kinetic parameters of lithocholic acid metabolite formation by human hepatic microsomes. Kinetic parameters were derived from the metabolite formation data presented in Fig. 4.2. Values represent the mean ± SEM of triplicate determinations. Kinetic parameters of hepatic microsomal metabolite formation were calculated using equation 1 for all five metabolites. 149 Tn i_i. mar m flivie avoil e pmol/min/pmol P450 u&1 pM 3-Ketocholanoic acid 50.0 ± 2.5 31.9 ± 6.2 N/A N/A Hyodeoxycholic acid 4.5 ± 0.4 45.4 ± 13.6 N/A N/A Ursodeoxycholic acid 1.2 ± 0.1 40.1 ± 13.6 N/A N/A Murideoxycholic acid 0.3 ± 0.0 N/A 35.4 ± 5.5 1.7 ± 0.3 6-Ketolithocholic acid 0.4 ± 0.0 29.3 ± 4.4 N/A N/A Table 4.2. Kinetic parameters of lithocholic acid metabolite formation by recombinant CYP3A4. Kinetic parameters were derived from the metabolite formation data presented in Fig. 4.4. Values represent the mean ± SEM of triplicate determinations. Kinetic parameters of all metabolite formation were calculated using equation 1, except for murideoxycholic acid formation, for which equation 2 was used. N/A, not applicable. 150 Human CYP3A4 Rate of formation of metabolites donor activity* (pmol/minlmg protein) (pmol/minlmg protein) Murideoxy- Ursodeoxy- Hyodeoxy- 6-Ketolitho- 3-Keto- cholic acid cholic acid cholic acid cholic acid cholanoic acid HH13 890 2.5 ± 0.1 4.6 ± 0.2 11.3 ± 0.2 0.9 ± 0.0 149.3 ± 9.1 HG95 650 0.7 ± 0.1 8.9 ± 1.34 1.6 ± 0.4 1.7 ± 0.5 54.4 ± 5.6 HH18 12000 11.2±0.5 35.1 ± 1.6 118.0±3.4 10.6±0.4 542.4±9.3 HH837 13700 15.4 ± 0.3 47.3 ± 1.0 160.5 ± 2.2 16.4 ± 0.5 750.1 ± 38.3 r2 value 0.97 0.97 0.98 0.95 0.96 Table 4.3 Rate of formation of lithocholic acid metabolites in human liver microsomes with varying CYP3A4 activities. Details of the microsorne donors are as follows. HH13, Asian male, 55 years of age; H095, Hispanic female, 47 years of age; HH18, African American female, 78 years of age; and HH837, Asian female, 52 years of age. * CYP3A4 activities were determined using testosterone 6-hydroxylase assay by BD Biosciences (Woburn, MA). The r2 value is the correlation coefficient determined for testosterone 613-hydroxylase activities of the four human liver samples and rates of metabolite formation of individual lithocholic acid metabolites. m!z 411lOG —d4-CIio& acid as internal standard 0 . 200 400 6.00 8.00 10.00 12.00 14,00 1600 18.00 20.00 2200 2400 26.00 28.00 30.00 - 3200 34:00 R 100 Hyodeoxycholic acid mlz 391 Ursodeoxycholic acid — Chenodeoxycholic acid a Murideoxycholic acid . * * 0 I 2.00 4.00 6.00 800 10.00 12.00 14.00 16.00 18.00 20.00 22.00 2400 26.00 28.00 3000 32.00 34.00 V mlz 389 e 100 7-Ketolithocholic acid 6-Ketolithocholic acid n I friTr.I,t 2.00 4.00 600 800 10.00 1200 14 00 1600 18.00 2000 22.00 24 DO 26.00 26.00 3000 e m/z375 Lithocholic acid ILt y 2.00 4120 6h0 8120 10.00 1200 1400 1600 1800 20.00 2200 • 2400 26.00 28.00 3000 3200 3400 mlz 373100 — 34cetocholanoic acid O i.0 i’.•p2.00 400 6.00 8.00 10.00 12.00 14.00 1600 18.00 20.00 22.00 24.00 26.00 28.00 30.00 3200 34.00 Time (mm) Figure 4.1. Representative LC/MS chromatogram showing metabolites of lithocholic acid in human hepatic microsomes. Lithocholic acid metabolites were extracted from a standard reaction mixture after a 3 0-mm incubation of human hepatic microsomes (0.5 mg) with 100 iM lithochocholic acid and 1 mM NADPH. Metabolite identification was performed by co-chromatography and spiking with authentic standards. In the above figure, the internal standard (d4-cholic acid, m/z 411) eluted at 15.5 mm. Metabolites, murideoxycholic acid (m/z 391), ursodeoxycholic acid (m/z 391), and hyodeoxycholic acid (m/z 391) eluted at 12.5, 13.5, and 15 mm respectively. 3-Ketocholanoic acid (m/z 373) eluted at 21.5 mm. Chenodeoxycholic acid (m/z 391) and 7-ketolithocholic acid were identified as impurities within the substrate lithocholic acid and were not classified as metabolites. * denotes unidentified peaks that were observed in controls and blanks and were not metabolites. 152 400 A 50 400 300 V 200 100 0 5 10 15 ‘45 20 0 50 100 150 [LthochoIic acid] (pM) 25 200 250 0 20 00. o 10 x C 0 a—. Sc •0’-0. aol oE . .9 >.—<0 OE z 2 50 100 150 (lithocholic acid] (IJM) 200 250 C 0 E .9 300 5-0 2 200 0’- 16 C 0 14 Sc 12 . 10 aol 8 .5 6 >5 — x oE 2 0 C 0 5.5 oE .9 °.o a ‘0 D 5, 4 3 2 0 0.0 50 100 150 200 250 Llithocholic acid] (liM) 6vv V 0.1 02 0.3 0.4 OL 0.6 V’s V V E 0 50 100 150 200 (Lithocholic acid] (pM) 3 2 250 00 5 4 3 2 0 ODO 0.02 0D4 5.06 0118 0.10 0.12 0.14 vls 0 0 50 100 150 200 0 LLithocholic acid] (pM) Figure 4.2 Enzyme kinetic profiles of 3-ketocholanoic acid (A), hyodeoxycholic acid (B), ursodeoxycholic acid (C), murideoxycholic acid (D) and 6-ketolithocholic acid (E) formation by human hepatic microsomes. Metabolite formation (activity) was plotted as a function of substrate concentration following a 30-mm incubation with human liver microsomes (0.5 mg). Data points are the mean ± SEM of triplicate determinations. Lines represent rates modeled by nonlinear regression analysis of the data. The insets depict Eadie-Hofstee plots. Error bars are not shown on the insets to avoid obscuring the data points, which represent mean values. 153 MLrideoxycholic acid Ursodeoxycholic acid Hyodeoxycholic acid . 3-Ketocholanoic acid UItEIflJ 6-Ketolithocholic acid - 0 3 - cq o Co a ‘.c c C) C.) 0 UI c CE c - v- ci c.i ci c.i C.) c.i c. , c’ CE o a a Q.. O G. D Q. C%1 Q. 0.. 0.. 0.. 0 0 >- >- >- >- >- >- Q_ >- >- >- >- 0 C) C) C) C) C.) C) >- C) C) C) C) >(fl C) C)I) 4- 0 C — -V 4) Figure 4.3 Comparison of 3-ketocholanoic acid, hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid formation by a panel of human recombinant P450 enzymes. Metabolite formation (activity) was measured following a 20-mm incubation of lithocholic acid (100 jiM) with baculovirus-insect cell microsomes containing expressed human P450 enzymes (30 pmol). Plots show the mean values ± SEM of triplicate determinations. 154 50 C 0 E 40 . 0 30 5 c a o . 10 a C., 20 •. .• . . 50 40 20 ID C 0 —a. a ii XE 0 0 x . 50 100 150 200 250 ELithocholic acid] (pM) [Lithocholic acid) (pM) C 0 •0 oE 4, 0. •0 0U, D0.36 0.30 0.26 0.20 C 0 o. >.o E 0.10 O. •0 0.15 V. •V . 0.05 0.00 E 0 100 150 (Uthocholic acid) (pM) CLithocholic acid) (pM) V 50 0.30 0.26 200 250 0.4 C 0 (0g. EU, ,D. E 0.20 •5•— 0.15 0.10 0.05 0.3 oQo 0 > 02 0.1 0.0 0.5000.5020.0(14 0.506 0006 0.010 6.612 0014 Ws 60 200100 150 (Lithocholic acid] (pM) Figure 4.4 Enzyme kinetic profiles of 3-ketocholanoic acid (A), hyodeoxycholic acid (B), ursodeoxycholic acid (C), murideoxycholic acid (D) and 6-ketolithocholic acid (E) formation from lithocholic acid. Metabolite formation (activity) was plotted as a function of substrate concentration following a 20-mm incubation with recombinant CYP3A4 (30 pmol). Data points are the mean ± SEM of triplicate determinations. Lines represent rates modeled by nonlinear regression analysis of the data. The insets depict Eadie-Hofstee plots. Error bars are not shown on the insets to avoid obscuring the data points, which represent mean values. 155 A C 0 00 U .. . C C C . . — 0 Q . C C . 0 0 0 ‘ u 0 o 0 0 0 .0 .0 .0o 0 0 0 0 0 00 0 00 az • 22 222 - 100 80 * 40 20 . .8 .8.8 0 C I .8.8.8 — 0 0 0 0 = 0 C C . 0 0 0 .0 0 C C C C C C C C C . 0 o 0 0 Qo 000 000 0.0.0. z W 222 222 . . . . 0 0 0 e* Figure 4.5 Chemical inhibition studies using ketoconazole, quercetin, quinidine, and sulfaphenazole on lithocholic acid biotransformation by human hepatic micrsomes. Formation of the major metabolites 3-ketocholanoic acid (A) and hyodeoxycholic acid (B) was decreased by ketoconazole significantly up to 60% and 90% by ketoconazole. Quinidine and sulfaphenazole did not seem to have any effect on formation of lithocholic acid metabolites as compared to the control (* p 0.05). Similar effect was observed with the minor metabolites ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid. The minor metabolites murideoxycholic acid and 6-ketolithocholic acid were not detected at higher concentrations of ketoconazole (10 .tM) (data not shown). ‘4 ,, - B 0 156 0•1 2 101 HOI Lithocholic acid (3c—iWdroxy-5p-choIan-24-oic acid) Figure 4.6 microsomes. Murideoxycholic acid (3c6p-dihydroxy- 5D-cholan-24-oic acid)’ Scheme showing lithocholic acid biotransformation by human hepatic The major metabolite was identified as 3-ketocholanoic acid formed by oxidation of the third carbon of the cholestane ring. The minor metabolites were identified as hyodeoxycholic acid, ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid, respectively. Results of the present study show that formation of all five hepatic microsomal metabolites of lithocholic acid was mediated mainly by CYP3A4 in humans. 21 22 12 11 3-Ketocholanoic acid (3-oxo- 5p-cholan-24-oic acid) p CYP3A4 ‘p’ Hyodeoxycholic acid Cyp3A4 (36a-dihydroxy-5-choian-24-oic acid) —CYP3A4--’ HO Ursodeoxycholic acid (3a,7p-dihydroxy- 5p-cholari-24-oic acid) N\CYP3A4 HO 6-Ketocholanoic acid (3ct—hydroxy-6-oxo- 5p-choian-24-oic acid) 157 4.6 References Araya Z and Wikvall K (1999) 6alpha-hydroxylation of taurochenodeoxycholic acid and lithocholic acid by CYP3A4 in human liver microsomes. Biochim Biophys Acta 1438:47-54. Azer SA, Kukongviriyapan V and Stacey NH (1995) Mechanism of ketoconazole-induced elevation of individual serum bile acids in the rat: relationship to the effect of ketoconazole on bile acid uptake by isolated hepatocytes. J Pharmacol Exp Ther 272:1231-1237. 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JPharmacol Exp Ther 270:414-423. Staudinger JL, Goodwin B, Jones SA, Hawkins-Brown D, MacKenzie K!, LaTour A, Liu Y, Klaassen CD, Brown KK, Reinhard J, Willson TM, Koller BH and Kliewer SA (2001) The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc NatlAcad Sci USA 98:3369-3374. Stricker BH, Blok AP, Bronkhorst FB, Van Parys GE and Desmet VJ (1986) Ketoconazole associated hepatic injury. A clinicopathological study of 55 cases. JHepatol 3:399-406. Tracy TS and Hummel MA (2004) ModeLing kinetic data from in vitro drug metabolism enzyme experiments. Drug Metab Rev 36:231-242. Wang L, Christopher U, Cui D, Li W, Iyer R, Humphreys WG and Zhang D (2008) Identification of the human enzymes involved in the oxidative metabolism of dasatinib: an effective approach for determining metabolite formation kinetics. Drug Metab Dispos 36:1828-1839. Xie W, Radominska-Pandya A, Shi Y, Simon CM, Nelson MC, Ong ES, Waxman DJ and Evans RM (2001) An essential role for nuclear receptors SXRIPXR in detoxification of cholestatic bile acids. Proc NatlAcadSci USA 98:33 75-3380. Zaki FG, Carey JB, Jr., Hoffbauer FW and Nwokolo C (1967) Biliary reaction and choledocholithiasis induced in the rat by lithocholic acid. JLab Clin Med 69:737-748. 161 Chapter 5 CYTOCHROME P450 MEDIATED BIOTRANSFORMATION OF LITHOCHOLIC ACID BY MOUSE HEPATIC MICROSOMES4 4This chapter forms part of a manuscript in preparation. Deo AK and Bandiera SM. Cytochrome P450 Mediated Biotransformation of Lithocholic Acid By Mouse Hepatic Microsomes: A Comparison With Human And Rat. 1 2 5.1 Summary In the present study, we investigated the cytochrome P450 (P450) enzymes involved in lithocholic acid biotransformation in the mouse. Lithocholic acid was incubated with mouse liver (C57BL/6) microsomes and metabolite analysis was carried out using liquid chromatography / mass spectrometry (LC/MS)-based assay. The major metabolite of lithocholic acid was identified as murideoxycholic acid. Minor lithocholic acid metabolites were identified as ursodeoxycholic acid, 3-ketocholanoic acid, isolithocholic acid, hyodeoxycholic acid and 6-ketolithocholic acid. To identif’ the P450 enzymes involved, hepatic microsomes were prepared from mice treated with prototypical P450 inducers including phenobarbital (PB, 102 mg/kg bw/day), dexamethasone (DEX, 50 mg/kg bw/day) and beta-napthoflavone (BNF, 100 mg/kg bw/day) and corn oil (CO. 10 mi/kg/day) for three days. Murideoxycholic acid formation decreased significantly with DEX-treatment. Formation of 3-ketocholanoic acid, ursodeoxycholic acid, hyodeoxycholic acid and 6-ketolithocholic acid increased with PB or DEX treatment. Formation of 3-ketocholanoic acid increased significantly with BNF treatment as well. Chemical inhibition studies were carried out by incubating hepatic microsomes with a combination of mouse and human P450 inhibitors (ketoconazole, quercetin, sulfaphenazole and quinidine). Murideoxycholic acid, 3- ketocholanoic acid and isolithocholic acid formation decreased with ketoconazole inhibition only. Decreased formation of ursodeoxycholic acid, hyodeoxycholic acid and 6-ketolithocholic acid was observed with the inhibitors, ketoconazole and quercetin. Results of chemical induction and inhibitor studies suggest the involvement of CYP1A, CYP2B and CYP3A enzymes in lithocholic acid oxidative biotransformation in the mouse. 163 5.2 Introduction Lithocholic acid (3a-hydroxy-53-cholan-24-oic acid) is the most toxic hydrophobic bile acid (Hoflnann, 2004). It is formed by removal of the 7a-hydroxyl group of chenodeoxycholic acid (primary bile acid, 3c,7c-dihydroxy-5 f3-cholan-24-oic acid) by the action of bacterial dehydrogenases. In the hepatocytes, lithocholic acid can undergo conjugation with amino acids (glycine or taurine) at the C-24 position and with sulfate at the C-3 position. The resulting sulfolithocholyiglycine and sulfolithocholyltaurine conjugates are excreted into bile and are not efficiently re-absorbed from the small intestine. As a result, lithocholic acid is eliminated from the body and does not represent more than 5% of biliary bile acids. Efficient elimination of lithocholic acid is beneficial because of its high hepatotoxicity (Fischer et al., 1996; Hoflnann, 2002; Hofinann, 2004). Inborn errors of bile acid biosynthesis, physical obstruction to bile flow and autoimmune destruction of cholangiocytes lead to impaired enterohepatic bile acid circulation (cholestasis) causing accumulation of bile acids in the liver. Hence, hepatic bile acid elimination pathways play an important role in maintaining bile acid homeostasis. Cytochrome P450 enzymes biotransform bile acids to their respective hydroxylated water soluble metabolites, thus assisting their efficient elimination. Our studies using human hepatic microsomes revealed the dominant role by a single P450 enzyme, CYP3A4, in the biotransformation of lithocholic acid to multiple metabolites, mainly, 3-ketocholanoic acid (Deo and Bandiera, 2009). Similarly, CYP3A4 was identified as the only human P450 enzyme involved in hepatic microsomal biotransformation of cholic acid (3a,7ct,12c-trihydroxy-53-cholan-24-oic acid) and chenodeoxycholic acid (Deo and Bandiera, 2008b). 164 Experiments involving P450 inducer treatments, recombinant P450 enzymes and antibody inhibition studies showed that multiple P450 enzymes catalyzed the formation of murideoxycholic acid (CYP2C1 1, CYP3A2 and CYP2D1), 3-ketocholanoic acid (CYP2C1 1 and CYP3A2) and isolithocholic acid (non microsomal enzymes) as hepatic microsomal lithocholic acid metabolites in the rat (Deo and Bandiera, 2008a). Comprehensive data sets for individual P450 levels in mice are not well documented. Different P450 levels have been obtained depending on the mouse strain used (Ford et al., 1979; Noshiro et al., 1986; Negishi et a!., 1991; Park et al., 1999). Inspite of this, the practical ease and cost effectiveness in handling and housing mice has enabled the frequent and preferred use of this rodent (over rat) for research purposes. Humanized mouse has been identified as a clinically relevant model to study P450-dependent metabolism (Muruganandan and Sinai, 2008). Nuclear receptor (PXR, CAR) knock out mouse choiestatic models have also been widely used to investigate bile acid regulation (Staudinger et al., 2001; Xie et al., 2001; Kitada et al., 2003). However, hepatic microsomal P450-mediated bile acid biotransformation data in mice has not been reported. The present study investigated the hepatic biotransformation of lithocholic acid in the mouse and identified the P450 enzymes involved in formation of its metabolites using P450 inducers and chemical inhibitors. Metabolites were identified by previously published assay and analytical methods using liquid chromatography mass spectrometry (LC/MS) (Deo and Bandiera, 2008a; Deo and Bandiera, 2008b; Deo and Bandiera, 2009). 5.3 Materials and Methods 5.3.1 Chemical and reagents. Lithocholic acid and all unconjugated bile acid standards (See Material and Methods, Chapter 2 and 4) were purchased from Steraloids Inc. (Newport, RI). 165 Ketoconazole, SKF-525A, quercetin, quinidine, and sulfaphenazole were generously provided by Dr. T.K.H Chang (University of British Columbia, Vancouver, BC, Canada). 5.3.2 Animal treatment and preparation of microsomes. Female 8-week old C57BL/6 mice (n104, average weightl8.2±l.l g) were obtained from Harlan (Indianapolis, [N). Mice received intraperitoneal injections of sodium phenobarbital (PB) in saline (n25, 102 mg/kg/day), dexamethasone (DEX) in corn oil (CO) (n=25, 50 mg/kg/day), 13-napthoflavone (BNF) in CO (n=27, 27 mg/kg/day) or CO alone (n=27, 10 mi/kg/day) for 3 consecutive days. Microsomes were prepared as described previously (Kania-Korwel et al., 2008). Protein concentrations were measured by the method of Lowry et al. (1951) using bovine serum albumin as a standard. 5.3.3 Lithocholic acid biotransformation assay. Lithocholic acid metabolites were analyzed using the LC/MS based assay described previously (Deo and Bandiera, 2008a; Deo and Bandiera, 2008b; Deo and Bandiera, 2009). An incubation time of 30 mm and a microsomal protein concentration of 0.5 mg/ml were found to be optimal for all six lithocholic acid metabolites and were used in subsequent experiments. Reaction mixtures contained 50 mM potassium phosphate buffer, pH 7.4, 3 mM magnesium chloride, 0.5 mg of mouse (female C57BL/6) hepatic microsomal protein, 1 mM NADPH and saturating concentrations (100 jiM for all metabolites, except 3-ketocholanoic acid (250 jiM)) of lithocholic acid, in a final volume of 1 ml. After preincubation for 10 mm at room temperature, reactions were initiated with NADPH and allowed to proceed for 30 mm at 37°C. Reactions were terminated with 8 ml of dichloromethane/isopropanol (80:20 v/v). Internal standard, (cholic-2,2,4,4-d4acid, 0.4 jig), was then added to each sample. Sample extraction, evaporation, and reconstitution in preparation 166 for analysis by LC/MS were carried out as described previously (Deo and Bandiera, 2008a). Reaction mixtures devoid of substrate, NADPH or microsomes, as well as reaction mixtures containing defined concentrations of authentic bile acid standards, were routinely included in each assay. 5.3.4 Chemical inhibition studies. Chemical inhibition studies were carried out using quercetin (2, 20 and 50 M, final concentrations), a mouse CYP1A1 (1C50 0.01 jiM), CYP1A2 (IC50 0.01 1iM) and CYP1B1 (1C50 0.02 iM) inhibitor, along with ketoconazole (0.01, 0.1, 1 and 10 jiM, final concentrations), a mouse CYP3A 11 inhibitor (IC50 0.02 jiM) (McLaughlin et a!., 2008). Additionally, human CYP2C9 inhibitor, sulfaphenazole (1, 10 and 20 jiM, final concentrations) and CYP2D6 inhibitor, quinidine (0.1, 1 and 10 jiM, final concentrations) were also used. Reactions were initiated with NADPH after initial pre-incubation of microsomes with lithocholic acid and the P450 inhibitor for 5 mm at 37°C. 5.3.5 Statistical analysis. Statistical analysis of data was carried out using GraphPad Prism® 4. Comparisons of chemical induction and inhibition data with control samples were carried out using ANOVA. A p value of 0.05 was considered statistically significant. 5.4 Results 5.4.1 Hepatic biotransformation of lithocholic acid by mouse hepatic microsomes. Incubation of lithocholic acid with liver microsomes from female C57BL16 mice revealed murideoxycholic acid as the major metabolite followed by ursodeoxycholic acid, 3- ketocholanoic acid, isolithocholic acid, hyodeoxycholic acid and 6-ketolithocholic acid as minor 167 metabolites (Table 5.1). No metabolite formation was observed in reaction mixtures devoid of substrate, NADPH or microsomes. Metabolite peaks, retention times and mass to charge ratios matched with the ones observed previously and were confirmed using authentic bile acid standards (Deo and Bandiera, 2008a). A peak of j3-muricholic acid (3-MCA) was found in all samples including control samples in absence of the substrate, lithocholic acid. Thus, the possibility of f3-muricholic acid being a metabolite was ruled out. Chenodeoxycholic acid metabolite formation was not observed. Minor metabolite peaks were identified as M-3 and M-4 similar to those observed previously in rat (Chapter 2, Figure 2.2) (Deo and Bandiera, 2008a). 5.4.2 Effect of P450 inducers in lithocholic acid biotransformation. To identifS’ the P450 enzymes involved, hepatic microsomes were prepared from female C57BL/6 mice treated with inducers including PB (CYP2B inducer), DEX (CYP3A inducer), BNF (CYP1A inducer), and CO as described under Materials and Methods. Reaction rates were calculated and compared with CO treated controls (Table 5.1). Murideoxycholic acid formation rate decreased significantly with DEX treatment. In case of PB treatment, the rate of formation of murideoxycholic acid was lower than the control but the decrease in activity was not significant. BNF treatment did not change the rate of murideoxycholic acid formation. Reaction rate for 3-ketocholanoic acid increased with PB, DEX and BNF treatment significantly. Isolithocholic acid formation rates were not altered with any of the inducer treatments except with a slight increase with BNF treatment as compared to the control. Ursodeoxycholic acid reaction rates almost doubled with PB and DEX treatment, but did not differ significantly as compared to the CO control. Hyodeoxycholic acid formation showed significant increase and was almost 3 and 5.5 times greater with PB and DEX treatment, respectively. The formation of 6-ketolithocholic acid showed significant increase as well and was 4 and 11 times greater upon treatment with PB and 168 DEX, respectively (Table 5.1). No increase in 13-MCA formation was observed by inducer treatments. Effects on the rate of formation of the minor unidentified metabolites M-3 and M-4 were not studied. 5.4.3 Chemical inhibition studies. The effect of chemical inhibitors was evaluated on the formation of oxidative metabolites of lithocholic acid. Metabolite formation was studied using increasing concentrations of ketoconazole (0.01 jiM, 0.1 jiM, 1 jiM and 10 jiM), quercetin (2 jiM, 20 jiM and 50 jiM), quinidine (0.1, 1 and 10 jiM) and sulfaphenazole (1, 10 and 20 jiM). Murideoxycholic acid formation decreased significantly only at higher concentrations of ketoconazole (up to 30% at 10 j.tM) (Fig. 5. 1A). In case of 3-ketocholanoic acid, a significant decrease in formation was observed at 1 jiM and 10 jiM ketoconazole, respectively (Fig. 5.1 B). Isolithocholic acid formation significantly decreased at 1 jiM and 10 jiM ketoconazole (Fig. 5. 1C). Murideoxycholic acid, isolithcholic acid and 3-ketocholanoic acid formation did not show any change with increasing quercetin concentrations (data not shown). Ursodeoxycholic acid formation decreased significantly with increasing inhibitor concentrations of 0.1 jiM, 1 jiM and 10 jiM ketoconazole and with 20 jiM and 50 jiM of quercetin (Fig. 5.1D). Hyodeoxycholic acid formation decreased significantly at 0.01 jiM and 0.1 jiM ketoconazole, and was not detected at higher concentrations of ketoconazole (1 jiM and 10 jiM). Significant decrease in hyodeoxycholic acid formation was also observed with increasing quercetin concentrations of 20 jiM and 50 jiM, respectively (Fig. 5.IE). Similarly, 6-ketolithocholic acid formation significantly decreased with 1 jiM and 10 jiM ketoconazole and 20 jiM and 50 jiM with quercetin (Fig. 5.1 F). No effect of quinidine and sulfaphenazole inhibition was observed in formation of any lithocholic acid metabolites by mouse microsomes (data not shown). 169 5.5 Discussion The focus of this discussion is P450-mediated hepatic microsomal biotransformation of lithocholic acid in mice. A detailed comparison of hepatic microsomal lithocholic acid biotransformation patterns in mouse, rat and human is discussed in Chapter 6. The major metabolite of lithocholic acid biotransformation in C57BL/6 mice was murideoxycholic acid. The minor metabolites were ursodeoxycholic acid, 3-ketocholanoic acid, isolithocholic acid, hyodeoxycholic acid and 6-ketolithocholic acid. Quantitative determinations and identification of metabolites were carried out by LCIMS. The predominant pathway of biotransformation was hydroxylation at the 613-position of lithocholic acid leading to the formation of murideoxycholic acid. This is probably the first report of hepatic lithocholic acid biotransformation carried out in vitro in mouse, and the metabolite profile obtained is consistent with previous in vitro studies carried out in rat (Zimniak et al., 1989; Deo and Bandiera, 2008a). Chemical inducers such as PB (CYP2B inducer), DEX (CYP3A inducer) and BNF (CYP1A inducer) were used to identify mouse P450 enzymes involved in lithocholic acid biotransformation. Specific mouse P450 inhibitors have not been well documented. A previous study identifies ketoconazole (1C50 0.02 riM) as a specific mouse CYP3A1 1 inhibitor and quercetin (1C50 values 0.01-0.02 jiM), as a mouse CYP 1A1, CYP 1 A2 and CYP 1 B 1 inhibitor (McLaughlin et al., 2008). Human P450 specific inhibitors, quinidine (human CYP2D6 inhibitor) and sulfaphenazole (human CYP2C9 inhibitor) were used in the assay with the expectation that they would inhibit the same respective P450 subfamily in mouse microsomes. A specific chemical inhibitor to inhibit mouse CYP2B is not available and hence could not be used. Murideoxycholic acid formation did not increase with PB, DEX and BNF treatment, indicating that mouse CYP2B, CYP3A and CYP1A enzymes are not involved in its formation 170 (Table 5.1). Chemical inhibition experiments showed that murideoxycholic acid formation was decreased at higher concentrations of ketoconazole (30% at 10 pM) (Fig. 5. 1A). Chemical inhibition and induction experiments suggest a partial involvement of CYP3A enzymes in murideoxycholic acid formation. Similar results showing no increase in murideoxycholic acid formation with PB and DEX-treatment along with a partial inhibition by anti-CYP3A antibodies were observed in rat (Deo and Bandiera, 2008a). The data on murideoxycholic acid formation in mouse is similar to the hepatic microsomal data obtained in rat (Deo and Bandiera, 2008a). A partial involvement of mouse CYP3A along with other mouse P450 enzymes (CYP2D, CYP2E and CYP4A) in murideoxycholic acid formation cannot be ruled out. Formation of 3-ketocholanoic acid increased significantly with PB, DEX and BNF treatment suggesting CYP2B, CYP3A and CYP1A mediated 3-ketocholanoic acid formation, respectively (Table 5.1). The CYP3A inhibitor, ketoconazole gradually decreased the formation of 3-ketocholanoic acid, supporting the involvement of CYP3A enzymes in its formation (Fig. 5. 1B). Increasing quercetin (CYP1A inhibitor) concentrations did not inhibit 3-ketocholanoic acid formation and thus BNF-treated CYP1A induction data could not be supported. Though induction studies also suggest the role of CYP2B mediated 3-ketocholanoic acid formation, CYP2B chemical inhibition studies are required to support the data. The induction and inhibition studies confirm the role of mouse CYP3A enzymes in 3-ketocholanoic acid formation. Isolithocholic acid formation did not increase significantly with PB, DEX and BNF treatment (Table 5.1). Chemical inhibition studies showed up to 30% decrease at 10 jiM ketoconazole (Fig 5.1 C). This effect may be due to non-specific inhibition of P450 enzymes by ketoconazole at higher concentrations. Chemical inhibition and induction studies suggest that none of the hepatic microsomal P450 enzymes may be involved in isolithocholic acid formation 171 in mouse. It should be noted that formation of isolithocholic acid from lithocholic acid by rat hepatic microsomes was not attributed to any P450 enzymes (Deo and Bandiera, 2008a). The mechanism for the formation of isolithocholic acid has been discussed previously (Deo and Bandiera, 2008a). A role for human liver cytosolic enzymes has been suggested in the formation of isolithocholic acid by Amuro et al. (1985) (Amuro et a!., 1985). Similar involvement of cytosolic mouse hepatic enzymes in isolithocholic acid formation in mouse cannot be ruled out. The formation of ursodeoxycholic acid, hyodeoxycholic and 6-ketolithocholic acid were increased with PB and DEX treatment (Table 5.1). Though the increase was not significant in case of ursodeoxycholic acid, the inducer studies may suggest mouse CYP2B and CYP3A involvement in formation of all three metabolites. Chemical inhibition studies show that formation of ursodeoxycholic acid, hyodeoxycholic acid and 6-ketolithocholic acid decreased with increasing ketoconazole and quercetin concentrations (Fig. 5. iD-F). Overall, the chemical induction and inhibition studies support the role of CYP3A-, CYP2B- and CYP lA-mediated ursodeoxycholic acid, hyodeoxycholic and 6-ketolithocholic acid formation. Specific mouse CYP2B chemical inhibition studies may further be required to confirm the role of CYP2B enzymes in the formation of these metabolites. Chemical inhibitors (especially of CYP2B and CYP2C) for mouse P450 enzymes are not well documented. Also, mouse recombinant P450 enzymes are not yet commercially available. Thus, conclusive data regarding identification of specific mouse P450 enzymes of these subfamilies in lithocholic acid biotransformation could not be obtained. The results discussed here serve as pointers in the process of identification of mouse P450 enzymes involved in lithocholic acid biotransformation. However, a role of more than one P450 enzyme in mouse lithocholic acid biotransformation can be definitely confirmed from our data. Based on the results obtained in this study, a scheme for biotransformation of lithocholic acid in mouse hepatic microsomes is proposed in Fig. 5.2. A role for the involvement of 172 multiple P450 enzymes (CYP3A, CYP2B and CYP1A) in mouse as compared to the predominant involvement of a single enzyme, CYP3A4, in lithocholic acid biotransformation in humans was supported by these findings. Hepatic lithocholic acid biotransformation data generated in this study may be useful for investigating various biochemical bile acid detoxification pathways in transgenic mouse models of cholestatic conditions observed in humans. 173 Rate of metabolite formation in mice treated with inducers pmol/min/mg protein Lithocholic acid metabolite Control PB DEX BNF Murideoxycholicacid 1670±53 1197±54 528±29* 1609± 104 Ursodeoxycholic acid 138 ± 50 261 ± 56 236 ± 80 130±31 3-Ketocholanoic acid 118 ± 10 552 ± 22* 772 ± 35* 269 ± 12* Isolithocholic acid 95 ± 15 95 ± 10 88 ± 10 129 ± 9 Hyodeoxycholicacid 46±1 154±4* 260±18* 37±2 6-Ketolithocholic acid 8 ± 1 35 ± 4* 90 ± 7* 9 ± 1 Table 5.1 Effect of treatment with P450 inducers on lithocholic acid metabolite formation by mouse hepatic microsomes. Incubation of mouse hepatic microsomes with lithocholic acid was carried out at saturating substrate concentrations (100 p.M lithocholic acid for all metabolites except 3-ketocholanoic acid (250 p.M) under optimal assay conditions as described in the Materials and Methods section. Hepatic microsomes prepared from vehicle-treated female C57BL/6 mice were used as controls. * p 0.05. 174 C 0 : — 120 100 80 (ci- 5Q 40 0 20 0 Figure 5.1 Chemical inhibition studies using ketoconazole, quercetin, quinidine, and sulfaphenazole on lithocholic acid biotransformation by mouse hepatic microsomes. Formation of the metabolites murideoxycholic acid (A), 3-ketocholanoic acid (B), isolithocholic acid (C), ursodeoxycholic acid (D), hyodeoxycholic acid (E) and 6-ketolithocholic acid (F). Quercetin did not show any inhibitory effect on murideoxycholic acid, 3-ketocholanoic acid and isolithocholic acid formation. Chemical inhibitors, quinidine and sulfaphenazole also did not show any effect on formation of lithocholic acid metabolites as compared to the control (data not shown). * p 0.05, n.d.- not detected. 175 120 C 0 100 2 : 80 2 60 CO o- 40 =0 20 I 0 .0 C 0 z 0 • • • 0 0 N N N C C C C o 0 0 0 9 9 0 0 0 0 0 0 t — C I- 0 .0 C 0 z • • 0 0 • 0 N N N N 0 0 C C C C 0 0 0 0U Q U U 0 0 0 0 ‘‘I’ — H yo de ox yc ho lic a c id fo rm at io n (% o fc o n tr ol ac tiv ity ) 7.3 . a 00 0 7.3 0 0 0 0 0 0 0 m C C) 0 CD 9’ U rs od eo xy ch ol ic a c id fo m ia tio n (% o fc o n tr oi ac tiv ity ) N o in hi bi to r 10 pM K et oc on az ol e I pM K et oc on az oi e 0.1 pM K et oc on az oi e 0. 01 pM K et oc on az oi e 50 pM Qu erc eti n 20 pM Qu erc eti n 2 pM Qu erc eti n * N o in hi bi to r 10 pM K et oc on az oi e 1 pM K et oc on az oi e 0. 1 pM K et oc on az oi e 0. 01 jAV iK et oc on az oi e 50 1iV iQ uer cet in 20 pM Qu er ce tin 2 p M Qu erc eti n 54 (et oii tho ch oli c a ci d fo rm at io n (% o fc o n tr ol ac tiv ity ) o 0 0 0 o No in hI bi to r _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ 10 pM K et oc on az oi e I pM K et oc on az oi e 0.1 pM K et oc on az oi e 0.0 1 pM K et oc on az oi e 5O pM Qu erc eti n F* 20 pM Qu erc eti n I— * 2 jdV i Q ue rce tin NIA IiIN iF— * ON Murideoxycholic acid (3a,6p-dihydroxy- 5p-cholan-24-oic acid) 4 CYP3A and other unidentified P450s I Lithocholic acid (3a—hydroxy-5-choIan-24-oic acid) CYP1A, CYP2B I and CYP1A, CYP2B and CYP3A 9 (3cL,6a-dihydroxy- 5p-cholan-24-oic acid) Figure 5.2 microsomes. Scheme showing lithocholic acid biotransformation by mouse hepatic Results of the present study suggest that formation of hepatic microsomal metabolites of lithocholic acid was mediated by several P450 enzymes (CYP1A, CYP2B and CYP3A) in mouse. The major metabolite formed in mouse was murideoxycholic acid. 3-Ketocholanoic acid (3-oxo- 513-cholan-24-oic acid) 12 CYP3A CYP2B? / CYPIA, CYP2B and CYP3A Isolithocholic acid (313—hydroxy-5f3- cholan-24-oic acid) OH Ursodeoxycholic acid (3a,7-dihydroxy- 5-choIan-24-oic acid) Hyodeoxycholic acid 6-Ketolithocholic acid (3a—hydroxy-6-oxo- 5-choIan-24-oic acid) 177 5.8 References Amuro Y, Yamade W, Maebo A, Hada T and Higashino K (1985) Reduction of 3-keto-5 beta cholanoic acid to lithocholic and isolithocholic acids by human liver cytosol in vitro. Biochim Biophys Acta 837:20-26. Deo AK and Bandiera SM (2008a) Biotransformation of lithocholic acid by rat hepatic microsomes: metabolite analysis by liquid chromatography/mass spectrometry. Drug Metab Dispos 36:442-45 1. Deo AK and Bandiera SM (2008b) Identification of Human Hepatic Cytochrome P450 Enzymes Involved in the Biotransformation of Cholic and Chenodeoxycholic Acid. Drug Metab Dispos 36:1983-1991. Deo AK and Bandiera SM (2009) 3-Ketocholanoic acid is the major CYP3A4 mediated lithocholic acid metabolite in human hepatic microsomes. Drug Metab Dispos, submitted. Fischer S, Beuers U, Spengler U, Zwiebel FM and Koebe HG (1996) Hepatic levels of bile acids in end-stage chronic cholestatic liver disease. Clin Chim Acta 251:173-186. Ford HC, Lee E and Engel LL (1979) Circannual variation and genetic regulation of hepatic testosterone hydroxylase activities in inbred strains of mice. Endocrinology 104:857- 861. Goodwin B and Kliewer SA (2002) Nuclear receptors. I. Nuclear receptors and bile acid homeostasis. Am JPhysiol Gastrointest Liver Physiol 282:G926-93 1. Holfriann AF (2002) Cholestatic liver disease: pathophysiology and therapeutic options. Liver 22 Suppi 2:14-19. Hofmann AF (2004) Detoxification of lithocholic acid, a toxic bile acid: relevance to drug hepatotoxicity. Drug Metab Rev 36:703-722. 178 Kania-Korwel 1, Hrycay EG, Bandiera SM and Lehmler HJ (2008) 2,2’,3,3’,6,6’- Hexachiorobiphenyl (PCB 136) Atropisomers Interact Enantioselectively with Hepatic Microsomal Cytochrome P450 Enzymes. Chem Res Toxicol. Kitada H, Miyata M, Nakamura T, Tozawa A, Honma W, Shimada M, Nagata K, Sinai CJ, Guo GL, Gonzalez FJ and Yamazoe Y (2003) Protective role of hydroxysteroid sulfotransferase in lithocholic acid-induced liver toxicity. J Biol Chem 278:17838- 17844. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the folin phenol reagent. JBiol Chem 193:265-275. Makishima M, Lu IT, Xie W, Whitfield GK, Domoto H, Evans RM, Haussier MR and Mangelsdorf DJ (2002) Vitamin D receptor as an intestinal bile acid sensor. Science 296: 13 13-13 16. McLaughlin LA, Dickmann U, Wolf CR and Henderson CJ (2008) Functional expression and comparative characterization of nine murine cytochromes P450 by fluorescent inhibition screening. Drug Metab Dispos 36:1322-1331. Muruganandan S and Sinai CJ (2008) Mice as clinically relevant models for the study of cytochrome P450-dependent metabolism. Clin Pharmacol Ther 83:818-828. Negishi M, Burkhart B and Aida K (1991) Expression of genes within mouse hA and lID subfamilies: simultaneous measurement of homologous P450 mRNAs. Methods Enzymol 206:267-273. Noshiro M, Serabjit-Singh CJ, Bend JR and Negishi M (1986) Female-predominant expression of testosterone 16 aipha-hydroxylase (“I”-P-450( 1 6)alpha) and its repression in strain 129/J. Arch Biochem Biophys 244:857-864. 179 Park SH, Liu X, Hennighausen L, Davey HW and Waxman DJ (1999) Distinctive roles of STAT5a and STAT5b in sexual dimorphism of hepatic P450 gene expression. Impact of STAT5a gene disruption. JBiol Chem 274:7421-7430. Rizzo G, Renga B, Mencarelli A, Pellicciari R and Fiorucci S (2005) Role of FXR in regulating bile acid homeostasis and relevance for human diseases. Curr Drug Targets Immune Endocr Metabol Disord 5:289-303. Staudinger JL, Goodwin B, Jones SA, Hawkins-Brown D, MacKenzie KI, LaTour A, Liu Y, Klaassen CD, Brown KK, Reinhard J, Wilison TM, Koller BH and Kliewer SA (2001) The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc NatlAcad Sd USA 98:3369-3374. Tien ES and Negishi M (2006) Nuclear receptors CAR and PXR in the regulation of hepatic metabolism. Xenobiotica 36:1152-1163. Wang R, Salem M, Yousef IM, Tuchweber B, Lam P, Childs SJ, Helgason CD, Ackerley C, Phillips MJ and Ling V (2001) Targeted inactivation of sister of P-glycoprotein gene (spgp) in mice results in nonprogressive but persistent intrahepatic cholestasis. Proc Nat! AcadSci USA 98:2011-2016. Xie W, Radominska-Pandya A, Shi Y, Simon CM, Nelson MC, Ong ES, Waxman DJ and Evans kM (2001) An essential role for nuclear receptors SXRIPXR in detoxification of cholestatic bile acids. Proc NatlAcadSci USA 98:3375-3380. Zimniak P. Holsztynska EJ, Lester R, Waxman DJ and Radominska A (1989) Detoxification of lithocholic acid. Elucidation of the pathways of oxidative metabolism in rat liver microsomes. JLipidRes 30:907-9 18. 180 Chapter 6 GENERAL DISCUSSION 181 6.1 Overview The current research was based on the premise that hydrophobic bile acids are vulnerable to oxidation at various positions on the cholestane ring to form water-soluble metabolites thereby, facilitating their clearance from the body. Hepatic microsomal P450 enzymes are involved in the biotransformation of lipophilic endogenous compounds such as steroids, eicosanoids, retinoids and bile acids. The goal of the research work was to develop a better understanding of hepatic microsomal bile acid biotransformation including characterization of bile acid metabolite profiles and identification of the P450 enzymes involved in metabolite formation in mouse, rat and human. Studies were carried out to test two hypotheses namely, that the major route of rodent and human hepatic bile acid biotransformation is through 63-hydroxylation of the cholestane ring and that hepatic bile acid biotransformation in rodents and humans would not be mediated exclusively by CYP3A enzymes. A detailed discussion has already been included in each of the four previous chapters. The current chapter is a general discussion of the data obtained with relation to the hypotheses stated. This includes a summary of cholic acid and chenodeoxycholic acid biotransformation data obtained using human hepatic microsomes and enzymes. Additionally, a comparison of lithocholic acid biotransformation data obtained using rodent (mouse and rat) and human hepatic microsomes is discussed. The chapter concludes by highlighting the limitations and significance of the work and proposes future studies based on the data generated and current available literature. 182 6.2 Summary of Cholic Acid Biotransformation By Human Hepatic Microsomes Human hepatic microsomal cholic acid biotransformation generated a single metabolite, 3-dehydrocho lie acid. Formation of this metabolite was through an oxidation reaction on the C- 3 carbon to form the 3-oxo-metabolite. Studies carried out using a panel of recombinant human P450 enzymes suggested that CYP3A4 was the only enzyme involved in the formation of 3- dehydrocholic acid (Chapter 3, Fig. 3.5A and Fig. 3.8). Though previous studies of cholic acid biotransformation had identified 3-dehydrocholic acid as a cholic acid metabolite, the only P450 enzyme investigated was recombinant CYP3A4 (Bodin et al., 2005). The present study is the first to investigate the contribution of a panel of human P450 enzymes and to clearly show that CYP3A4 is the only enzyme involved in cholic acid biotransformation. 6.3 Summary of Chenodeoxycholic Acid Biotransformation By Human Hepatic Microsomes Biotransformation of chenodeoxycholic acid by human hepatic microsomes generated four metabolites. Of these, 7ct-hydroxy-3-oxo-53-cholan-24-oic acid, which was the major metabolite of chenodeoxycholic acid, was identified as a new metabolite. The other metabolites were y-muricholic acid, 7-ketolithocholic acid and cholic acid (Chapter 3, Fig. 3.3). Studies carried out using human recombinant P450 enzymes indicated that CYP3A4 was the only enzyme involved in the formation of 7c-hydroxy-3-oxo-5 13-cholan-24-oic acid and y-muricholic acid. The formation of cholic acid and 7-ketolithocholie acid was not attributed to any of the human P450 enzymes investigated (CYP1A1, CYP1A2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2C19, CYP2D6, CYP2E1, CYP3A4, CYP3A5 and CYP4AI 1) (Chapter 3, Fig. 3.SB). A role for CYP7A and CYP7B in 7-ketolithocholic acid formation and the involvement of CYP8B 1 (sterol 1 2c*-hydroxylase) in cholic acid formation were speculated. Analysis of 183 enzyme kinetic parameters associated with formation of chenodeoxycholic acid metabolites and screening of a panel of human P450 enzymes for potential involvement in the formation of chenodeoxycholic acid were important aspects of this study. 6.4 Comparison of Lithocholic Acid Metabolite— Formation Patterns By Rat, Human And Mouse Liver Microsomes Hepatic microsomal lithocholic acid biotransformation was studied using rat (Chapter 2), human (Chapter 4) and mouse (Chapter 5) liver microsomes. A detailed comparison of (a) lithocholic acid metabolite patterns, (b) P450 enzymes involved and (c) kinetic parameters of metabolite formation in these species is discussed. The comparative data are presented in Tables 6.1 and 6.2 and in Figures 6.1 and 6.2. (a) Biotransformation by mouse hepatic microsomes revealed the formation of murideoxycholic acid as the major 63-hydroxylated metabolite of lithocholic acid. Ursodeoxycholic acid, 3-ketocholanoic acid, isolithocholic acid, hyodeoxycholic acid and 6- ketolithocholic acid were minor metabolites of lithocholic acid in mouse (See Fig. 6.1). Similarly, in rat hepatic microsomes, the major metabolite was murideoxycholic acid. Isolithocholic acid and 3-ketocholanoic acid were the next most abundant metabolites formed. 6-Ketolithocholic acid, hyodeoxycholic acid and ursodeoxycholic acid were relatively minor lithocholic acid metabolites (Deo and Bandiera, 2008a). In the case of human hepatic microsomes, the major metabolite was the 3-oxo metabolite, 3-ketocholanoic acid. The 6a- hydroxy metabolite, hyodeoxycholic acid, was the second most important metabolite formed (in quantitative terms) and ursodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid, were minor metabolites (Fig. 6.1). 184 Lithocholic acid was preferentially metabolized at position C-6 by mouse liver microsomes to form murideoxycholic acid, 6-ketolithocholic acid and hyodeoxycholic acid. Similarly, oxidation of lithocholic acid at position C-6 to form murideoxycholic acid, 6- ketolithocholic acid and hyodeoxycholic acid was preferred over oxidation at C-3 (to form 3- ketocholanoic acid and isolithocholic acid) in rat liver microsomes. Oxidation of lithocholic acid at C-7 to form ursodeoxycholic acid was the least preferred site of oxidation in mouse and rat. Thus the preferred sites for lithocholic acid oxidation by rodent liver microsomes was C-6> C-3 > C-7, in that order (See Table 6.1). In the case of lithocholic acid biotransformation by human liver microsomes, oxidation at the C-3 position to form 3-ketocholanoic acid was the preferred pathway. Oxidation at C-6 position to form hyodeoxycholic acid, murideoxycholic acid and 6-ketolithocholic acid, and oxidation at the C-7 position to form ursodeoxycholic acid were less preferred pathways in human hepatic microsomes. 3 13-Epimerization of lithocholic acid at the C-3 position to form isolithocholic acid was not observed in human hepatic microsomes. Thus, the preferred sites of lithocholic acid oxidation by human liver microsomes was C-3 > C-6> C-7, in that order. The C-7 position was the least preferred position for oxidation in all three species (Table 6.1). A similar regio-selective preference for oxidation of cholic acid and chenodeoxycholic acid by human liver microsomes was observed in our studies (Chapter 3). The only metabolite formed from cholic acid was the 3-oxo metabolite, 3-dehydrocholic acid. In the case of chenodeoxycholic acid, the major metabolite was 7a-hydroxy-3-oxo-5f3-cholanoic acid, which was oxidized at the C-3 position (Deo and Bandiera, 2008b). Thus, oxidation at C-3 was the preferred pathway for cholic acid, chenodeoxycholic acid and lithocholic acid metabolite formation in humans (Table 6.1). 185 (b) A comparison of various P450 enzymes involved in lithocholic acid biotransformation in rat, mouse and humans is summarized in Table 6.2. Though most of the metabolites of lithocholic acid formed by rodent and human hepatic microsomes are similar, the relative prominence of the major metabolites (3-ketocholanoic acid in humans and murideoxycholic acid in rat/mouse), differ. The differences in the P450-mediated oxidation of lithocholic acid could be attributed to the relative levels of individual P450 enzymes expressed in rodent and human liver. In rat liver, the CYP2C subfamily is the predominant subfamily (up to 65%) followed by the CYP3A subfamily (up to 14%). The other rat P450 enzymes (e.g. CYP2A, CYP2E and CYP4A) form only 10-15% of the total P450 enzyme pool (Chapter 1, Table 1.2). Thus, results obtained using chemical inducers, antibody inhibition studies and recombinant enzymes (Chapter 2) support a predominant role for CYP2C11 followed by CYP3A2 in murideoxycholic acid formation in rat. Specific data about mouse hepatic microsomal enzymes are not available. The involvement of CYP1A, CYP2B and CYP3A enzymes in lithocholic acid biotransformation in mouse can be due to a similar variation in mouse P450 levels. In humans, the CYP3A subfamily is quantitatively the most important subfamily of enzymes (up to 40%) followed by the CYP2C subfamily (up to 25%) (Chapter 1, Table 1.2). CYP3A4 accounts for 30-40% of the total P450 enzymes in human liver (Shimada et al., 1994) and hence its contribution as the major P450 enzyme in human hepatic microsomal lithocholic acid biotransformation to form 3- ketocholanoic acid can be explained. Formation of chenodeoxycholic acid as a lithocholic acid metabolite in rat or human hepatic microsomes was suggested by previous studies. Our studies using rat, mouse and human hepatic microsomes with the substrate lithocholic acid completely rule out this possibility, as we did not observe chenodeoxycholic acid as a metabolite in any of the incubations. Previous studies relied on less sensitive techniques such as thin layer chromatography and may have used 186 substrate that was contaminated with chenodeoxycholic acid (Zimniak et al., 1989; Xie et al., 2001) (c) A comparison of the kinetic parameters of lithocholic acid metabolite formation in rat, mouse and human reveal that the rate of formation of these metabolites was higher in rat hepatic microsomes. For instance, the rate of formation of murideoxycholic acid was 2900 pmol/minlmg protein in rat liver microsomes and was 1670 pmollminlmg protein in mouse liver microsomes. The rate of murideoxycholic acid formation in human liver microsomes was 5.5 pmollminl mg protein. Also, the rate of formation for 3-ketocholanoic acid, the major metabolite in human, was 512 pmol/minlmg protein in rat liver microsomes followed by 336 pmol/min!mg protein in human liver micromes and 118 pmol/min!mg protein in mouse liver microsomes. The lowest Km values were obtained for 6-ketolithocholic acid formation, in rat and human hepatic microsomes (rat 1.6 jiM and human 29.1 jiM) (Chapter 2, 4 and 5; Tables 2.1, 4.1 and 5.1). These rates of metabolite formation suggest a greater metabolic rate for bile acid biotransformation in rat than in human or mouse. Another comparison of bile acid metabolites formed by human hepatic microsomes shows that the number of metabolites formed was greater when the substrate was a hydrophobic bile acid. The more hydrophobic the substrate, the larger the number of metabolites formed. For instance, lithocholic acid (log P 6.6) is the most hydrophobic bile acid and generated six metabolites. This was followed by the more hydrophilic bile acid, chenodeoxycholic acid (log P 4.9), that generated four metabolites and finally, cholic acid (log P 4.1), which produced only one metabolite. In summary, a comparative analysis of lithocholic acid biotransformation was possible using human and rodent hepatic microsomes. Based on the results obtained, a scheme for the 187 biotransformation of lithocholic acid in human and rodent hepatic microsomes is proposed in Fig. 6.2. The results of this research project partially support the hypotheses stated in Chapter 1 (Section 1.11). Hepatic microsomal lithocholic acid biotransformation in rodents is mediated primarily through 63-hydroxylation, but is not exclusively mediated by CYP3A enzymes. CYP1A, CYP2B, CYP2C enzymes play a significant role in the biotransformation of lithocholic acid in rodents. Human hepatie microsomal biotransformation of lithocholic acid, cholic acid and chenodeoxycholic acid is mediated exclusively by CYP3A4. However, the major route of human hepatic bile acid biotransformation is not 613-hydroxylation, but oxidation at C-3 position on the steroid nucleus. 6.5 Limitations Though the major research objectives of this research project were achieved, the current study had some limitations as stated below. (a) The LC/MS method utilized in the current research work was robust enough to extract, resolve and identify unconjugated bile acid metabolites. Identification of the bile acid metabolites in our study was mainly based on spiking and co-chromatographic retention times of the authentic standard. However, structural identification of unknown bile acid metabolite peaks posed a problem. A method generally utilized for structure identification is collision induced fragmentation. Electrospray and atmospheric pressure ionization techniques combined with collision-induced dissociation do not facilitate fragmentation of bile acid metabolites because of their rigid steroid structures. To overcome this limitation, fractions of the mobile 188 phase eluting from the column can be collected at the respective retention time of the unknown metabolites and further subjected to structure analysis by nuclear magnetic resonance spectrometry. (b) Isolithocholic acid was a non-P450 mediated metabolite of lithocholic acid in rodents. The identity of the enzyme involved in its biotransformation was not revealed by our studies. A role for cytosolic enzymes has been suggested in the formation of isolithocholic acid and needs to be investigated further. (c) Enzymes involved in the formation of 7-ketolithocholic acid and cholic acid from chenodeoxycholic acid were not identified. A role for CYP7A or CYP7B in 7-ketolithocholic acid and CYP8B in cholic acid fonnation was speculated. Commercial availability of recombinant enzymes and inhibitory antibodies of CYP7A, CYP7B and CYP8B enzymes would assist further investigation. (d) Biotransformation studies carried out in mouse identified various P450 subfamilies involved in hepatic microsomal lithocholic acid metabolism. However, the identity of the specific P450 enzymes in formation of lithocholic acid metabolites was not established. The availability of specific mouse P450 inhibitors and inhibitory antibodies may help address this problem. 189 6.6 Overall Significance of Thesis Research Bile acids are implicated in liver diseases such as hepatic cholestasis and liver necrosis. Progress in controlling and treating cholestatic liver disease depends largely on advances in understanding these diseases through basic biomedical research performed in animals and comparing the data with the data obtained in humans. Previous studies on bile acid biotransformation were limited and generated conflicting results due to the complexity of bile acid hydroxylation profile, less sensitive techniques and isolation and purification procedures for bile acid metabolite identification. Moreover, the identity of P450 enzymes in bile acid biotransformation in rodents and humans was limited to CYP3A enzymes only. The increasing use of animal models to study bile acid regulation (Staudinger et al., 2001; Wang et a!., 2001; Goodwin and Kliewer, 2002; Makishima et al., 2002; Rizzo et al., 2005; Tien and Negishi, 2006) stressed the need to investigate and compare bile acid biotransformation patterns in human and rodent hepatic microsomes. Studies to address these issues were designed, implemented and accomplished. A simple LC/MS based assay for identification of bile acid metabolites from hepatic microsomes in rodents and human was developed. Using this method we showed that hepatie microsomal bile acid (especially, lithocholic acid) metabolites in human, rat and mouse are similar. However, the contribution of the specific P450 enzymes, and the major metabolites, involved in bile acid biotransformation within these species differ. Similar metabolite formation profiles may be obtained for cholic acid and chenodeoxycholic acid demonstrating the involvement of multiple P450 enzymes in rodents. These studies were the first to show a dominant involvement of human hepatic microsomal CYP3A4 in biotransformation of cholic acid, chenodeoxycholic acid and lithocholic acid to all of their respective metabolites. The studies carried out in rodents were also the first to identify the role of CYP1A, CYP2B, CYP2C, CYP2D and CYP3A mediated pathways for bile 190 acid biotransformation. Data suggest that induction of CYP3A enzymes in humans and CYP1A, CYP2B and CYP3A enzymes in rodents may offer a potential mechanism to lessen the hepatotoxicity associated with high tissue levels of bile acids (See Fig. 6.3). The hepatic bile acid biotransformation data obtained from rodents and humans may provide additional insights into understanding the pathways involved in bile acid elimination. Thus, we hope that this research will be taken into consideration while identifying potential targets of cholestatic liver disease and aid in the development of drug therapy that stimulates bile acid detoxification pathways. 6.7 Future Studies (a) Formation of oxidative metabolites and reactive oxygen species. Hepatotoxic bile acids impair mitochondrial function, leading to inhibition of oxidative phosphorylation and enhanced formation of toxic oxygen species by the mitochondrial respiratory chain. This results in ATP depletion, increased Ca2 concentration with stimulation of hydrolases, and oxidative stress (Sokol et al., 1995; Rodrigues et al., 1998; Rob et al., 2000; Sokol et al., 2005). Oxidative stress could lead to hydrolysis of lipid membranes and structural proteins causing cell death by necrosis (Roberts et al., 1997; Rodrigues et al., 1998). Thus, a mechanism by which bile acids can cause liver damage is by stimulating production of reactive oxygen species in the tissue. Reactive oxygen species can be generated during P450-mediated formation of oxo- and hydroxy-bile acid metabolites. The role of these metabolites in the generation of reactive oxygen species needs to be investigated. Formation of keto or oxo metabolites of drug substrates such as valproic acid has been linked to reactive oxygen species. Our studies show the formation of oxo metabolites of bile acids (e.g. 3-ketocholanoic acid). 191 Therefore, it is possible that P450-mediated bile acid metabolism may play a role in eliciting toxicity, as opposed to serving a beneficial role in detoxification. (b) Investigation of mechanisms of formation of 3-keto bile acid metabolites and isolithocholic acid. The formation of 3-ketocholanoic acid from lithocholic acid, 3-dehydrocholic acid from cholic acid and 3-oxo-7c-hydroxy-5f3-cholan-24-oic acid from chenodeoxycholic acid may proceed through the formation of geminal diol intermediates. Studies to investigate the formation of a P450—mediated geminal diol intermediate need to be carried out. This can be achieved by incorporation of 180 into the reaction system. Ideally, if a P450-mediated geminal diol intermediate would be formed the in the reaction, 50% incorporation of ‘O into the product would be expected. A mechanism suggesting the incorporation of 180 to form a geminal diol intermediate was reported in the formation of androstenedione from testosterone (Wood et al., 1988). The formation of isolithocholic acid from 3-ketocholanoic acid in rodents may be mediated by hydroxysteroid dehydrogenase/epimerase enzyme. Another mechanism involving dehydration of lithocholic acid to a alkene intermediate followed by direct 3-epimerization to form isolithocholic acid has been suggested but needs to be investigated further (Bortolini et al., 1997). (c) Bile acid structural analogues for treatment of cholestasis. Cytotoxicity of hydrophobic bile acids to hepatocytes has been usually attributed to membrane-disruptive effects from their detergent properties. Ironically, certain hydrophilic bile acids such as ursodeoxycholic acid and chenodeoxycholic acid analogues (6-ethyl chenodeoxycholic acid and GW4064) protect against cholestasis and the hepatotoxicity induced 192 by hydrophobic bile acids (Beuers et al., 1998; Makishima et al., 2002) (Liu et al., 2003). One possibility is that hydroxylated bile acids such as ursodeoxycholic acid increase expression of bile acid transporters which efflux the toxic bile acid out of the hepatocyte. Ursodeoxycholic acid was the most efficacious bile acid tested in terms of inducing CYP3A4 in human hepatocytes (Schuetz et al., 2001). Thus, the reversal of cholestasis in humans by ursodeoxycholic acid may include induction of CYP3A enzymes and possibly transporter genes that result in enhanced metabolism and efflux of hepatotoxic bile acids. A similar investigation on reversal of bile acid toxicity upon administration of chenodeoxycholic acid analogues needs to be carried out. (d) Identification of conjugating enzymes in bile acid biotransformation. Conjugation via glucuronidation and sulfation of bile acids such as lithocholic acid and chenodeoxycholic acid has been already reported. Conjugation of bile acids to form hydrophilic metabolites is a major pathway of bile acid biotransformation and hence, biotransformation studies of other secondary bile acids such as ursodeoxycholic acid and deoxycholic acid by conjugating enzymes need to be carried out. (e) Bile acid metabolites as nuclear receptor ligands. It should be noted that the major metabolite of lithocholic acid, 3-ketolithocholic acid, is a bioactive metabolite and acts as a ligand for the nuclear receptor PXR. Another minor lithocholic acid metabolite, 6-ketolithocholic acid, activates the VDR. Similar studies using hepatic microsomal metabolites of chenodeoxycholic acid and cholic acid should be conducted. It is likely that the major or minor metabolites of these bile acids could bind to receptors such as PXR, FXR and LXR, and in turn regulate P450 enzymes. 193 (f) The current LC/MS method is applicable to quantify unconjugated and conjugated bile acids in liver, bile and serum. Identification of individual bile acid levels in these biomatrices can be useful for determining biliary and plasma levels of bile acids in liver disorders. In addition, modifications in the method can facilitate fragmentation of conjugated bile acids and subsequent detection of metabolites could be achieved by LC/MSIMS. 194 Metabolite Oxidation Human Mouse Rat position Murideoxycholic acid C-6 1% 80% 62% 6-Ketolithocholic acid C-6 1% < 1% 1% Hyodeoxycholic acid C-6 12% 2% < 1% 3-Ketocholanoic acid C-3 82% 6% 15% Isolithocholic acid C-3 N/A 5% 22% Ursodeoxycholic acid C-7 4% 7% < 1% Preference C-3 > C-6 > C-7 C-6 > C-3 > C-7 C-6 > C-3 > C-7 Table 6.1 Regio-selective oxidation of lithocholic acid by human, rat and mouse hepatic microsomes. The values in the table are approximate percentages of lithocholic acid metabolite formation based on individual rates of formation of each metabolite obtained using hepatic microsomal data from Chapters 2, 4 and 5. The preferred position of oxidation in human liver microsomes is C-3 > C-6> C-7. The preferred position of oxidation in rodent liver microsomes is C-6 > C-3> C-7. Data for rat and human modified from (Deo and Bandiera, 2008a; Deo and Bandiera, 2009). 195 Metabolite Human Rat Mouse Murideoxycholic acid CYP3A4 CYP2C1 1, CYP3A2, CYP3A + CYP2D? CYP2D1 6-Ketolithocholic acid CYP3A4 CYP3A2 CYP2B, CYP3A, CYP 1 A Hyodeoxycholic acid CYP3A4 CYP3A2 CYP2B, CYP3A, CYPIA 3-Ketocholanoic acid CYP3A4 CYP2C1 1, CYP3A2 CYP2B, CYP3A Ursodeoxycholic acid CYP3A4 CYP3A2 CYP2B, CYP3A, CYP1A Isolithocholic acid Not formed Microsomal non-P450 Microsomal non-P450 enzymes enzymes Table 6.2 Comparison of P450 enzymes involved in lithocholic acid metabolite formation by human and rodent hepatic microsomes. Data compiled from Deo and Bandiera, 2008a; Deo and Bandiera, 2009 and Chapter 5. 196 Human Mouse HUCA 2% 6KLCA <1% \\ MDCA 80% Carbon position Metabolite •MDCA C6 u6KLCA •HDCA LJ3KCA EIILCA C7 oUDCA Figure 6.1 Comparison of lithocholic acid metabolite formation patterns obtained from rat, mouse and human hepatic microsomes. Murideoxycholic acid (MDCA) is the major metabolite of lithocholic acid in rat and mouse hepatic microsomes but in human hepatic microsomes, MDCA is formed as a very minor metabolite. 3-Ketocholanoic acid (3KCA) is the major metabolite formed in human hepatic microsomes, but a relatively minor metabolite in rodents. Ursodeoxycholic acid (IJDCA), 6- ketolithocholic acid (6KLCA) and hyodeoxycholic acid (HDCA) are minor metabolites in rat, mouse and human hepatic microsomes. Isolithocholic acid (ILCA) is not formed by human hepatic microsomes. Data for rat and human modified from (Deo and Bandiera, 2008a; Deo and Bandiera, 2009). Relative metabolite formation is expressed in terms of approximate percentages and was calculated from individual rates of formation of each metabolite obtained using hepatic microsomal data from Chapters 2, 4 and 5. UOCA 4% 6KLCA 1% MOCA 1% ILCA UDCA 3KCA / 7% Rat LCA 22% 3KCA 15% UDCA [<1% HDCA/\ <1% 6KLCA 1% MDCA 62% 197 ThCOOH 3-Ketocholanoic acidMurideoxychohc acid (3-oxo- 53-choIan-24-oic acid)(3a,63-dihydroxy- 53-cholan-24-oic acid) major pathway for rat! major pathway for mouse liver microsomes human liver microsomes 21 22 23 .%.ICOOH ICOOH only in rat! HO3 4H6 14 15 Isolithocholic acid Lithocholic acid (33—hydroxy-53- (3x—hydroxy-53-cholan-24-oic acid) cholan-24-oic acid) i minor pathways for rat, mouse and human liver microsomes HOXTh HOXtTh HOqTh Ursodeoxycholic acid 6-Ketolithocholic acid Hyodeoxycholic acid (3a,713-dihydroxy- (3x—hydroxy-6-oxo- (3ct,6ct-dihydroxy- 513-cholan-24-olc acid) 5p-cholan-24-oic acid) 513-cholan-24-oic acid) Figure 6.2 Lithocholic acid biotransformation by rodent and human liver microsomes. Murideoxycholic acid formation is the major lithocholic acid biotransformation pathway by rodent (rat and mouse) liver microsomes. Formation of 3-ketocholanoic acid is the major lithocholic acid biotransformation pathway by human hepatic microsomes. Ursodeoxycholic acid, 6-ketolithocholic acid and hyodeoxycholie acid are minor metabolites formed by rat, mouse and human hepatic microsomes. Isolithocholic acid is the only metabolite that is not formed by human hepatic microsomes. 198 Liver circulation Intestine Figure 6.3 Hepatic microsomal P450-mediated bile acid biotransformation: Route to alleviate bile acid mediated hepatotoxicity in cholestasis. Various P450-mediated biotransformation pathways of hepatotoxic cholestatic bile acids (lithocholic acid, chenodeoxycholic acid and cholic acid) in rodent and human hepatic microsomes were identified in this study. Induction of P450-mediated biotransformation pathways may eventually help to alleviate hepatotoxic effects of bile acids observed in cholestasis. 199 6.8 References Beuers U, Boyer JL and Paumgartner G (1998) Ursodeoxycholic acid in cholestasis: potential mechanisms of action and therapeutic applications. Hepatology 28:1449-1453. Bodin K, Lindbom U and Diczfalusy U (2005) Novel pathways of bile acid metabolism involving CYP3A4. Biochim Biophys Acta 1687:84-93. Bortolini 0, Medici A and Poli S (1997) Biotransformations on steroid nucleus of bile acids. Steroids 62:564-577. Deo AK and Bandiera SM (2008a) Biotransformation of lithocholic acid by rat hepatic microsomes: metabolite analysis by liquid chromatography/mass spectrometry. Drug Metab Dispos 36:442-451. Deo AK and Bandiera SM (2008b) Identification of Human Hepatic Cytochrome P450 Enzymes Involved in the Biotransformation of Cholic and Chenodeoxycholic Acid. Drug Metab Dispos 36:1983-1991. Deo AK and Bandiera SM (2009) 3-Ketocholanoic acid is the major CYP3A4 mediated lithocholic acid metabolite in human hepatic microsomes. Drug Metab Dispos, submitted. Goodwin B and Kliewer SA (2002) Nuclear receptors. I. Nuclear receptors and bile acid homeostasis. Am JPhysiol Gastrointest Liver Physiol 282:G926-93 1. Liu Y, Binz J, Numerick MJ, Dennis S, Luo G, Desai B, MacKenzie KI, Mansfield TA, Kliewer SA, Goodwin B and Jones SA (2003) Hepatoprotection by the farnesoid X receptor agonist GW4064 in rat models of intra- and extrahepatic cholestasis. J Clin Invest 112:1678-1687. 200 Makishima M, Lu TI’, Xie W, Whitfield GK, Domoto H, Evans RM, Haussler MR and Mangelsdorf DJ (2002) Vitamin D receptor as an intestinal bile acid sensor. Science 296: 13 13-13 16. Rizzo G, Renga B, Mencarelli A, Pellicciari R and Fiorucci S (2005) Role of FXR in regulating bile acid homeostasis and relevance for human diseases. Curr Drug Targets Immune Endocr Metabol Disord 5:289-303. Roberts LR, Kurosawa H, Bronk SF, Fesmier PJ, Ageilon LB, Leung WY, Mao F and Gores GJ (1997) Cathepsin B contributes to bile salt-induced apoptosis of rat hepatocytes. Gastroenterology 113:1714-1726. Rodrigues CM, Fan G, Ma X, Kren BT and Steer CJ (1998) A novel role for ursodeoxycholic acid in inhibiting apoptosis by modulating mitochondrial membrane perturbation. J Clin Invest 101:2790-2799. Rob AP, Oliveira PJ, Moreno AJ and Paimeira CM (2000) Bile acids affect liver mitochondrial bioenergetics: possible relevance for cholestasis therapy. Toxicol Sci 57:177-185. Schuetz EG, Strom S, Yasuda K, Lecureur V. Assem M, Brimer C, Lamba J, Kim RB, Ramachandran V, Komoroski BJ, Venkataramanan R, Cai H, Sinai CJ, Gonzalez FJ and Schuetz JD (2001) Disrupted bile acid homeostasis reveals an unexpected interaction among nuclear hormone receptors, transporters, and cytochrome P450. JBiol Chem 276:39411-39418. Shimada T, Yamazaki H, Mimura M, Inui Y and Guengerich FP (1994) Interindividual variations in human liver cytochrome P-450 enzymes involved in the oxidation of drugs, carcinogens and toxic chemicals: studies with liver microsomes of 30 Japanese and 30 Caucasians. JPharmacol Exp Ther 270:414-423. Sokol RJ, Dahl R, Devereaux MW, Yerushalmi B, Kobak GE and Gumpricht E (2005) Human hepatic mitochondria generate reactive oxygen species and undergo the permeability 201 transition in response to hydrophobic bile acids. JPediatr Gastroenterol Nutr 41:235- 243. Sokol RJ, Winkihofer-Roob BM, Devereaux MW and McKim 3M, Jr. (1995) Generation of hydroperoxides in isolated rat hepatocytes and hepatic mitochondria exposed to hydrophobic bile acids. Gastroenterology 109:1249-1256. Staudinger JL, Goodwin B, Jones SA, Hawkins-Brown D, MacKenzie K!, LaTour A, Liu Y, Klaassen CD, Brown KK, Reinhard 3, Wilison TM, Koller BH and Kliewer SA (2001) The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc NatlAcad Sci USA 98:3369-3374. Tien ES and Negishi M (2006) Nuclear receptors CAR and PXR in the regulation of hepatic metabolism. Xenobiotica 36:1152-1163. Wang R, Salem M, Yousef TM, Tuchweber B, Lam P, Childs SJ, Helgason CD, Ackerley C, Phillips MJ and Ling V (2001) Targeted inactivation of sister of P-glycoprotein gene (spgp) in mice results in nonprogressive but persistent intrahepatic cholestasis. Proc Nat! AcadSci USA 98:2011-2016. Wood AW, Swinney DC, Thomas PE, Ryan DE, Hall PF, Levin W and Garland WA (1988) Mechanism of androstenedione formation from testosterone and epitestosterone catalyzed by purified cytochrome P-450b. JBiol Chem 263:17322-17332. Xie W, Radominska-Pandya A, Shi Y, Simon CM, Nelson MC, Ong ES, Waxman DJ and Evans RM (2001) An essential role for nuclear receptors SXR!PXR in detoxification of cholestatic bile acids. Proc NatlAcadSci USA 98:3375-33 80. Zimniak P, Holsztynska EJ, Lester R, Waxman DJ and Radominska A (1989) Detoxification of lithocholic acid. Elucidation of the pathways of oxidative metabolism in rat liver microsomes. JLipidRes 30:907-918. 202 7. APPENDIX I 7.1 Cholic acid and chenodeoxycholic acid assay validation To validate the assay for cholic acid and chenodeoxycholic acid metabolites, metabolite stability, limit of detection, inter- and intra-assay variability, and extraction efficiency were determined. Hepatic microsomal samples prepared from human liver microsomes were used for assay validation. 7.1.1 Methods a) Standard (calibration) curves Various concentrations of calibration standards were included in each experiment in duplicate and treated in the same manner as test samples. Calibration standard mixtures were prepared in the range of 500-5,000 pmollml of reaction mixture for quantitation of cholic acid, 7-ketolithocholic acid and 7c-hydroxy-3-oxo-53-cholan-24-oic acid were prepared. Calibration standard curves using concentrations of 2,000-20,000 pmollml of reaction mixture were prepared for quantitation of y-muricholic acid. Calibration standard curves using concentrations of 500-10,000 pmol/ml of reaction mixture were prepared for quantitation of 3-dehydrocholic acid. Murideoxycholic acid was used as the internal standard (I.S.). The calibration curves were determined by plotting peak area ratios (PARs; AUC of metabolite / AUC of I.S.) obtained using LC/MS versus concentrations of individual analytes. b) Stability of analytes Bench-top stability of metabolites standards was assessed using human hepatic microsomes with metabolite standards mentioned above at concentrations of 500, 2,000 and 203 5,000 pmollml of reaction mixture in the absence ofNADPH. These spiked microsomal samples were left on the bench-top overnight (24 h at room temperature at approximately 20°C) and analyzed the next day. The relative stability of each analyte was determined by comparing the PARS of all metabolites with the PARs of freshly prepared reference standards of the same concentrations, in the absence of hepatic microsomes. The freshly prepared reference standards were considered to be 100% stable. The stability was measured as follows: Stability of Sample = (PAR sample/PAR reference standard) x 100 The analytes were considered ‘stable’ if the measured PARs were within ± 10% of the reference PARs. c) Limit of detection To determine the limit of detection, standard mixtures of cholic acid, 7-ketolithocholic acid, 7cL-hydroxy-3-oxo-53-cholan-24-oic acid, y-muricholic acid and 3-dehydrocholic acid (n = 6) were serial diluted to concentrations of 50, 5, 2.5, 2 and I pmoLlml of reaction mixture. Samples were injected on the LC/MS. The PARs of the metabolite standards at individual concentrations were obtained. To determine the inherent noise in the assay, peaks occurring at the same retention times as authentic standards were quantified using samples devoid of metabolite standards (blank). Limit of detection, the smallest concentration of the analyte that can be distinguished from the noise level, was determined at the concentration of analyte that gave a signal equal to the blank plus three times the standard deviation of the blank was identified from the calculated results (Armbruster et a!., 1994). Limit of detection = mean blank value + 3 (standard deviation of blank). 204 d) Variability of the assay The reproducibility of the assay was determined by evaluating inter- and intra-assay variability (coefficient of variation, C.V.) for the following metabolites, cholic acid, 7- ketolithocholic acid, 7c-hydroxy-3-oxo-5 13-cholan-24-oic acid, ‘y-muricholic acid and 3- dehydrocholic acid. Six sets of calibration curves were analyzed on 6 separate days (inter-assay) and on the same day (intra-assay). The reproducibility of the assay (C.V.) was determined from the variance observed for the mean of the replicates and a variability of < 20% for inter-assay and < 10% for intra-assay, was considered to be acceptable. e) Extraction recovery The percent recovery of metabolite standards from the hepatic microsomal reaction mixture was determined. Metabolite standards of cholic acid and chenodeoxycholic acid were chosen in the concentration range from 500-20,000 pmol/ml of reaction mixture as applicable. Two sets of samples, the non-extracted samples and the extracted samples, were prepared in duplicate. In the extracted samples, analytes were extracted by a liquid-liquid single-extraction procedure using dichloromethane: isopropanol (80:20), as mentioned in Chapter 2 and 3. The choice of the extraction solvent mixture was taken from Stedman et al. (2004). The non- extracted samples containing the internal standard were prepared in mobile phase and injected directly onto the LC. The extraction recovery was determined as follows % Extraction recovery = (PARs of extracted samples from hepatic microsomal mixture / PARs of non-extracted samples) x 100 Recovery of extracted analytes within 15% of pure non-extracted standards were considered acceptable (Causon, 1997). 205 7.1.2 Results a) Standard (calibration) curves The calibration curves of authentic chenodeoxycho lie acid and cholic acid metabolite standards are shown in Fig. 7.1 and 7.2, respectively. The response obtained by LC/MS detection for 7cL- hydroxy-3-oxo-5f3-eholan-24-oic acid, y-murieholic acid, 7-ketolithocholic acid, cholic acid and 3-dehydrocholic acid was linear with respect to the respective concentration ranges chosen. The mean slope values for the metabolite standards are summarized in Table 7.1. y = 0.0007x 4.0 R’ = 0.9964 U 10.0g y 0.0006x9 < 3.0 8.0 2.5 0.9792777 _ . 6.02 2.0 01.5 . 4Q1.0 0.5 . 2.0 0.0 0 2000 4000 6000 0.0 0 5000 10000 15000 20000 250007cz-hydroxy-3-oxo-53 -cholan-24-oic acid (pmol/mI reaction mix.) y-Muricholic acid (pmollml reaction mix.) C) 4.5 4.0 (j4Q 35 y 0.000Bx: 3.0 2.5 3.5 y 00007x < 3.0 2.0 0.997 2.5 0.9918 1.5 2.0 1.0 0.5 0.0 0.5C, 0 2000 4000 6000 o.o 7-Ketolithocholic acid (pmollml reaction 0 2000 4000 6000 mix.) Cholic acid (pmollml reaction mix.) Fig. 7.1 Representative calibration curves for chenodeoxycholic acid metabolites The figure shows the calibration curves for 7a-hydroxy-3-oxo-5f3-cholan-24-oic acid, y muricholic acid, 7-ketolithocholic acid and cholic acid. The response obtained by LC/MS detection was linear over the range of concentration (500-5000 pmol/ml reaction mixture for 206 7c-hydroxy-3-oxo-5 f3-cholan-24-oic acid, cholic acid and 7-ketolithocholic acid; 2000-20,000 pmol/ml of reaction mixture for y-murichotic acid) chosen. Fig. 7.2 Representative calibration curve for cholic acid metabolite The figure shows the calibration curve for 3-dehydrocholic acid. The response by LC/MS detection was linear over the range of concentration (500-10,000 pmol/ml of reaction mixture) chosen for the metabolite. Metabolite 7ce-Hydroxy- y-Muricholic 7-Ketolitho- Cholic 3-Dehydro Standard 3-oxo-5 13- acid cholic acid acid cholic cholan 24-oic acid acid Mean Slope 0.0007 0.00065 0.0007 0.00078 0.00032 n6 concentration 500-5000 500-20,000 500-5000 500-5000 500-10,000 range in samples (pmol/ml of raction mixture) Table 7.1 Mean slopes of cholic acid and chenodeoxycholic acid metabolite standards 3 2.5 4 U(V U 1.5 U 0 . 1 >‘ G) 9 0.5 C.) < 0 y = 0.0003x R2 = 0.9838 0 2000 4000 6000 8000 10000 12000 3-Dehydrocholic acid (pmollml reaction mix.) 207 b) Stability of analytes Bench-top stability of metabolite standards was assessed in the concentration range of 500-5,000 pmollml of reaction mixture. The stability values calculated are shown in Table 7.2 below. Considering the stability of the reference standards at 100%, it can be seen that the analytes were stable in the presence of human liver microsomes after 24 h at room temperature (approximately 20°C) and were within ± 10% of the PARs of the reference samples. Concentration Stability of sample expressed as (PAR Sample I PAR Reference standard) x in pmol/ml of 100 reaction (%) mixture 7cz-Hydroxy-3- ‘y-Muricholic 7-Ketolitho- Cholic 3-Dehydro oxo-513-cholan acid cholic acid acid cholic 24-oic acid acid 500 94.1 95.9 92.2 105 97 2000 97.8 99.1 91.7 103 96 5000 97.9 92.4 96.1 101 101 Table 7.2 Stability of cholic acid and chenodeoxycholic acid metabolites in presence of human liver microsomes. c) Limits of detection Limit of detection of 7a-hydroxy-3-oxo-53-cho1an-24-oic acid, y-muricholic acid and cholic acid was found to be 2.5 pmollml of reaction mixture and for 3-dehydrocholic acid and 7- ketolithocholic acid limit of detection was found to be 5 pmollml of reaction mixture. 208 d) Variability of the assay The reproducibility of the assay was determined by evaluating inter- and intra-assay variability. The reproducibility (in terms of coefficient of variation, C.V.) of the assay for all cholic acid and chendeoxycholic acid metabolites was measured as described previously. The intra-assay and inter-assay variability values for the major metabolites are listed in Tables 7.3 and 1.4, respectively. The inter-assay variability for the metabolites in the concentration range of the standard curve was less than 15% (Table 7.3). Similarly, the intra-assay variability for the metabolites in the concentration range of the standard curve was less than 10% (See Table 7.4). Metabolite standard Concentration of metabolite standard in pmoLfml of _____ reaction mixture 500 2,000 5,000 10,000 20,000 C.V.(%) 7a-Hydroxy-3-oxo-53-cho1an-24- 10.6 9.5 11.5 N/A N/A oic acid y-Muricholic acid 5.2 11.6 N/A N/A 11.2 7-Ketolithocholic acid 5.6 8.7 6.7 N/A N/A Cholic acid 8.5 11.1 10.9 N/A N/A 3-Dehydrocholic acid 9.1 10.0 N/A 11.2 N/A Table 7.3 Inter-assay variability (n = 6) The inter-assay variability of cholic acid and chenodeoxycholic acid metaboltes was assessed at the concentrations mentioned above. The C.V. (%) was observed to be less than 15%. N/A-not applicable. 209 Metabolite standard Concentration of metabolite standard in pmo1/mi of reaction mixture 500 2,000 5,000 10,000 20,000 C.V.(%) 7cc-Hydroxy-3-oxo-53-cholan-24- 6.5 7.8 6.5 N/A N/A oic acid ‘y-Muricholic acid 4.9 2.4 N/A N/A 6.5 7-Ketolithocholic acid 8.5 4.5 4.4 N/A N/A Cholic acid 6.3 4.3 5.6 N/A N/A 3-Dehydrocholic acid 3.2 2.3 N/A 4.5 N/A Table 7.4 Intra-assay variability (n = 6) The inter-assay variability of cholic acid and chenodeoxycholic acid metaboltes was assessed at the concentrations mentioned above. The C.V. (%) was observed to be less than 10%. N/A-not applicable. e) Extraction recovery The extraction efficiency of all metabolites of cholic acid and chenodeooxycholic acid was observed between 90-100%. The extraction efficiency of the internal standard, murideoxycholie acid was calculated independently and was found to be 100%. 210 7.2 References Armbruster DA, Tiliman MD and Hubbs LM (1994) Limit of detection (LQD)/limit of quantitation (LOQ): comparison of the empirical and the statistical methods exemplified with GC-MS assays of abused drugs. Gun Chem 40:1233-1238. Causon R (1997) Validation of chromatographic methods in biomedical analysis. Viewpoint and discussion. J Chromatogr B Biomed Sci Appi 689:175-180. 211

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