No work can be carried out alone, and it was here that Dr. Eugene Hrycay ...... Noshiro M, Okuda K and Coon MJ (1992) Purification and characterization of 7.
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 3ketocholanoic 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
1.
xvi
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
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 1.12
Specific research objectives
References
9
32 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
2.4
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
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
3.4
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
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
4.4
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
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
HEPATIC
5.
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
Table 1.2
Different P450 enzymes and their levels in human and rat hepatic microsomes
Table 2.1
Tabl e 5 1 Table 6.1
150
Rate of formation of lithocholic acid metabolites in human liver microsomes with varying CYP3A4 activities Effect of treatment with P450 inducers on lithocholic acid metabolite formation by mouse hepatic microsomes
151 174
Regio-selective oxidation of lithocholic acid by human, rat and mouse hepatic microsomes
Table 6.2
149
Kinetic parameters of lithocholic acid metabolite formation by recombinant CYP3A4
Table 4.3
89
Kinetic parameters of lithocholic acid metabolite formation by human hepatic microsomes
Table 4.2
88
Effect of sex and treatment with P450 inducers on lithocho lie acid metabolite formation by rat hepatic microsomes
Table 4.1
34
Kinetic parameters of lithocholic acid metabolite formation by rat hepatic microsomes
Table 2.2
33
195
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
Comparison of bile acid levels in livers of patients with end-stage cholestasis and normal humans
43
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 acid
Figure 1.9 Figure 1.10
and
CAR
cross-talk
mediated
metabolism
of
lithocholic 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 hydroxylation
Figure 2.2 Figure 2.3 Figure 2.4
Figure 2.5
Figure 2.6 Figure 2.7
acid
structure
showing
positions
available
for 90
Representative LC/MS chromatogram showing metabolites of lithocholic acid
91
Kinetic profiles of hepatic microsomal murideoxycholic acid, isolithocholic acid and 3-ketocholanoic acid formation
92
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
Comparison of murideoxycholic acid (A), isolithocholic acid (B) and 3ketocholanoic acid (C) formation by a panel of rat recombinant P450 enzymes
94
Enzyme kinetic profile of 3-ketocholanoic acid formation by rat recombinant CYP3A2
95
Effect of CYP2D6 antiserum on murideoxycholic acid formation by rat hepatic microsomes and recombinant CYP2D1
96 x
Figure 2.8
Figure 2.9
Scheme showing a proposed mechanism for the formation of 3ketocholanoic acid and isolithocholic acid from lithocholic acid through a geminal diol intermediate
97
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 chenodeoxycholic acid
chromatogram
showing
metabolites
of 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 7xhydroxy-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 Representative LC/MS chromatogram showing metabolites of lithocholic acid in human hepatie microsomes 152
Fi gure 4 1 Figure 4.2
Enzyme kinetic profiles of 3-ketocholanoic acid (A), hyodeoxycholic acid (B), ursodeoxycholic acid (C), murideoxycholic acid (D) and 6ketolithocholie 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 6ketolithocholic 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 Scheme showing lithocholic acid biotransformation by mouse hepatic microsomes Comparison of lithocholic acid metabolite formation patterns obtained from rat, mouse and human hepatic microsomes Lithocholic acid biotransformation by rodent and human liver microsomes Hepatic microsomal P450-mediated bile acid biotransformation: Route to alleviate bile acid mediated hepatotoxicity in cholestasis
Figure 5 2 • Fi g ure 6 1 •
Figure 6 2 • Fi g ure 6 3
175 177 197 198 199
xl’
LIST OFABBREVIATIONS
ANOVA ASBT AUC BNF BSEP CO C.V. CAR flEX FIC1 FXR GC/MS HSD IgG IS ISBT LC-MS LCIMSIMS LXR MC MDR MRP NADPH
NTCP OATP P450 PAR PB PBRE PXR RXR s.c. SEM SKF-525A SPGP SULT VDR XRE
analysis of variance apical sodium-dependent bile salt transporter area under curve beta napthoflavone bile salt export protein corn oil coefficient of variation constitutive androstane receptor dexamethasone familial intrahepatic cholestasis- 1 farnesoid X receptor gas chromatography mass spectrometry hydroxy steroid dehydrogenase immunoglobin G internal standard ileal sodium-dependent bile salt transporter liquid chromatography mass spectrometry liquid chromatography tandem mass spectrometry liver X receptor 3- methylcholanthrene multidrug resistance P-glycoprotein multiple resistance associated protein reduced nicotinamide adenine dinucleotide phosphate sodium taurocholate cotransporting polypeptide organic anion transporter protein cytochrome P450 peak area ratio phenobarbital phenobarbital response elements pregnane X receptor retinoid X receptor subcutaneous standard error of mean proadifen sister of P-glycoprotein sulfotransferase vitamin D receptor 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
M XN 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 A 6 double bond of cholesterol to a ’ 5 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, 12cctrihydroxy-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-trihydroxy5 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 3cLhydroxyl 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 C24. 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
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 (EC 50 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 (EC 50 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 watersoluble 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-5phosphosulfate (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). Fiftyseven 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 13muricholic 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 613hydroxylation 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, 6ahydroxy-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 3dehydrocholic 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 ‘ C-labeled chenodeoxycholic acid was studied in patients with 4 intrahepatic cholestasis, by Bremmelgaard and Sjovall (1980). The 14 C-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-5f3cholanic 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-hydroxydeoxycholic 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
1.
Hypotheses
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 CYP3Amediated.
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 Lithocholic acid (513-cholanic acid-3i-ol)
Metabolite
Trivial name
5J3-cholanic acid-3-ol 5-cholanic acid-3c, 6, 7f3-triol 5-cholanic acid-3n, 6a-diol
isolithocholic acid 13-muricholic acid hyodeoxycholic acid
5-cholanic acid-3a, 613-diol
murideoxycholic acid or murocholic acid 3-ketocholanic acida
Species/System human rat human, rCYP3A4*
Research group
rat hamster rat, rCYP3A4 human chenodeoxycholic acid rat 6-ketolithocholic acid rat rCYP3A4 1 13-hydroxy-lithocholic acid rCYP3A4
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
Chenodeoxycholic acid 513-cholanic acid-3cL, 6cL, 7a-triol (5J3-cholanic acid-3a, 7a- diol) 513-cholanic acid-3a, 6J3, 7a-triol 3oxo 513-cholanic acid
hyocholic acid
human, pig
Bremmelgaard and Sjovall, 1980
cL-muricholic acid 3 ketocholanic acid
rat rCYP3A4
Bremmelgaard and Sjovall, 1980 Bodin et al, 2005
513-cholanic acid-3ct, 12o-diol Cholic acid (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, 12adiol)
human, rCYP3A4 human human 3-dehydrodeoxycholic acid rCYP3A4
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
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 a
1 13-hydroxydeoxycholic acid
cholanoic acid and cholanic acid are used interchaneabIy.
Table 1.1 Oxidative metabolites of bile acids
Bremmelgaard and Sjovall, 1980, Bodin et al, 2005 Bremmelgaard and Sjovall, 1980 Bremmelgaard and Sjovall, 1980 Bodin et al, 2005 *
rCYP3A4
—
recombinant CYP3A4
Human P450 Enzyme
Specific Content (pmollmg)
% Total P450
Rat P450 Enzyme
Specific Content (pmol/mg)
% Total P450
CYP1A (CYP1AI, 1A2) CYP2A6
1-65
7-18
CYP1A (CYPIAI, 1A2)
5-20
1-3
C.) C)
a.
>. C)
N
2. C.)
>.
I ii
me N a. N a. >.
>.
0(3(3
—
C.)
N
a.
— 0
N
N
r
a. 0. a.>->N
00
a.
>. C)
C C
0
.- In o a. •0
. 0
U 0. C
C,
N
N
-
N
a. >. >0000>->CC)
C)
a. >. C)
C,
>C)
Figure 2.5 Comparison of murideoxycholic acid (A), isolithocholic acid (B) and 3ketocholanoic 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 b . Microsomes 5 containing expressed CYP1A2 and CYP2D1 contained co-expressed P450 oxidoreductase but not cytochrome b . There was no obvious correlation between rates of metabolite formation and 5 P450P oxidoreductase or cytochrome b 5 levels in the recombinant P450 preparations. Plots show the mean values ± SEM of triplicate determinations.
94
30
0
E
L
_..
O U) 0
V
25
V
O..20
E 0 0-
..3
oE
5 C.,
0 0
20
40
60
80
100
[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.
95
140 t
m E
120 100 80
0
20
40
60
80
100
120
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.
96
ThCOOH 9 c±
3-Ketocholanoic acid H2O +[HJ
Lithocholic acid geminal diol intermediate Isolithocholic acid
Figure 2.8 Scheme showing a proposed mechanism for the formation of 3ketocholanoic acid and isolithocholic acid from lithocholic acid through a geminal diol intermediate.
97
21
22 23
Ii 14
10
H03
4
COOH 7 15
H
Lithocholic acid
/
CYP2CII, CYP3A2
V CYP3A2 HO
Murideoxycholic acid Isolithocholic acid
m icrosomal non-P450 enzymes
\
3-Ketocholanoic acid Figure 2.9 Scheme showing P450-mediated lithocholic acid biotransformation in rat hepatic microsomes. 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.
98
2.6
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 :4 7-54.
Berta L, Fronticelli Baldelli C, Fazzari A, Radice E, Bargoni A, Frairia R and Gaetini A (2003) Sex steroid receptors, secondary bile acids and colorectal cancer. A possible mechanism of interaction. Panminerva Med 45:261-266. Bodin K, Lindbom U and Diczfalusy U (2005) Novel pathways of bile acid metabolism involving CYP3A4. Biochim Biophys Acta 1687:84-93. De Gottardi A, Touri F, Maurer CA, Perez A, Maurhofer 0, Ventre G, Bentzen CL, Niesor EJ and Dufour JF (2004) The bile acid nuclear receptor FXR and the bile acid binding protein IBABP are differently expressed in colon cancer. Dig Dis Sd 49:982-989. Deo AK (2005) Hepatic Biotransformation of Lithocholic Acid: Role of Cytochrome P450 Enzymes, in M.Sc. Thesis, Faculty of Pharmaceutical Sciences, University of British Columbia, Vancouver, BC, Canada. Dionne S, Tuchweber B, Plaa GL and Yousef IM (1994) Phase I and phase II metabolism of lithocholic acid in hepatic acinar zone 3 necrosis. Evaluation in rats by combined radiochromatography and gas-liquid chromatography-mass spectrometry. Biochem Pharmacol 48:1187-1197. Erlinger S (1997) Drug-induced cholestasis. JHepatol 26 Suppl 1:1-4. Fischer CD, Cooper NS, Rothschild MA and Mosbach EH (1974) Effect of dietary chenodeoxycholic acid and lithocholic acid in the rabbit. Am JDig Dis 19:877-886. Fischer S, Beuers U, Spengler U, Zwiebel FM and Koebe HG (1996) Hepatic levels of bile acids in end-stage chronic cholestatic liver disease. Cliii Chim Acta 251:173-186.
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Gustafsson J (1978) Effect of biliary obstruction on 26-hydroxylation of C27-steroids in bile acid synthesis. JLipid Res 19:237-243. Hofmann AF (1999) The continuing importance of bile acids in liver and intestinal disease. Arch Intern Med 159:2647-265 8. Hofmann AF (2002) Cholestatic liver disease: pathophysiology and therapeutic options. Liver 22 Suppl 2: 14-19. Hofmann AF (2004) Detoxification of lithocholic acid, a toxic bile acid: relevance to drug hepatotoxicity. Drug Metab Rev 36:703-722. Hofmann AF, Sjovall J, Kurz G, Radominska A, Schteingart CD, Tint GS, Vlahcevic ZR and Setchell KD (1992) A proposed nomenclature for bile acids. JLipid Res 33:599-604. Javitt NB (1966) Cholestasis in rats induced by taurolithocholate. Nature 210:1262-1263. Jezequel AM, Benedetti A, Bassotti C and Orlandi F (1994) Drug induced cholestasis. Elsevier Science By, Netherlands. Kitada H, Miyata M, Nakamura T, Tozawa A, Honma W, Shimada M, Nagata K, Sinal CJ, Guo GL, Gonzalez FJ and Yamazoe Y (2003) Protective role of hydroxysteroid sulfotransferase in lithocholic acid-induced liver toxicity. J Biol Chem 278:1783817844. Law E (1995) Effect of Cimetidine On Hepatic Cytochrome P450 1A and P450 2C in Male Rats. in MSc. Thesis, Faculty of Pharmaceutical Sciences, University of British Columbia, Vancouver, BC, Canada. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the folin phenol reagent. JBiol Chem 193:265-275. Makishima M (2005) Nuclear receptors as targets for drug development: regulation of cholesterol and bile acid metabolism by nuclear receptors. JPharmacol Sci 97:177-183.
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Miyai K, Price VM and Fisher MM (1971) Bile acid metabolism in mammals. Ultrastructural studies on the intrahepatic cholestasis induced by lithocholic and chenodeoxycholic acids in the rat. Lab Invest 24:292-302. Norman A and Palmer RH (1964) Metabolites of Lithocholic Acid-24-C-14 in Human Bile and Feces. JLab Clin Med 63:986-1001. Palmer RH (1971) Bile acid sulfates. II. Formation, metabolism, and excretion of lithocholic acid sulfates in the rat. JLipidRes 12:680-687. Panesar SK, Bandiera SM and Abbott FS (1996) Comparative effects of carbamazepine and carbamazepine- 10,11 -epoxide on hepatic cytochromes P450 in the rat. Drug Metab Dispos 24:619-627. Penning TM, Smithgall TE, Askonas U and Sharp RB (1986) Rat liver 3 aipha-hydroxysteroid dehydrogenase. Steroids 47:221-247. Rost D, Herrmann T, Sauer P, Schmidts HL, Stieger B, Meier PJ, Stremmel W and Stiehi A (2003) Regulation of rat organic anion transporters in bile salt-induced cholestatic hepatitis: effect of ursodeoxycholate. Hepatology 38:187-195. Sakai K, Makino T, Kawai Y and Mutai M (1980) Intestinal microflora and bile acids. Effect of bile acids on the distribution of microflora and bile acid in the digestive tract of the rat. Microbiol Immunol 24:187-196. Setchell KD, Rodrigues CM, Clerici C, Solinas A, Morelli A, Gartung C and Boyer J (1997) Bile acid concentrations in human and rat liver tissue and in hepatocyte nuclei. Gastroenterology 112:226-235. 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)
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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-betaHydroxylation 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.
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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.
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Chapter 3
IDENTIFICATION HUMAN OF HEPATIC CYTOCIIROME P450 ENZYMES INVOLVED IN THE OF BIOTRANSFORMATIQN CHOLIC AND CHENODEOXYCHOLIC ACID 2
A version of this chapter has been published as: 2 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-cholan24-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,7cLtrihydroxy-513-cholan-24-oic) as major metabolites and 7-ketolithocholic acid (3c-hydroxy-7oxo-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-3oxo-513-cholan-24-oic acid and y-muricholic acid from chenodeoxycholic acid. Formation of 7ketolithocholic 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-5 13-cholan-24-oic acid and 1 , 3a, 7cL, 1 2tetrahydroxy 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, 3ketocholanoic 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 , were purchased from BD Biosciences (Oakville, Ontario, Canada). High-performance liquid 5 b 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
(equation 1)
K,,,+[S]
—
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 3dehydrocholic 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 7ketolithocholic 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 7ketolithocholic 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 iM). A chenodeoxycholic acid concentration of 500 jiM was found to be saturating for all four 1 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-cholan24-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 7cchydroxycholesterol, which is catalyzed by CYP7A1, and in the conversion of 25hydroxycholesterol 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), 25hydroxycholesterol (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, 3dehydrocholic 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 6cchydroxylation 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 7hydroxysteroid 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 7cL119
hydroxy-3-oxo-5f3-cholan-24-oic acid from chenodeoxycholic acid would be approximately 5075 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
m!z 407
100
R
e I
“
a 0 V
e
m!z 405
100
n t
e I’ S
0 100
.-
If
Internal standard
mlz 391
(%)
10
20
25
30
Time (mm)
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.
121
400
Estimated V,. = 756 ± 158 pmol/minlmg protein Estimated Km
=
1060±133 liM
0
300
.
E 200 a
U 00
a 00
100 a
C.,
a
0
0
200
400
600
800
1000
[Cholic add] (pIV
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.
122
100
/
R e
Cholic acid
mlz 407
‘y-muricholic acid
c
a 0 100
1
Internal standard
I
Chenodeoxycholic / acid
mlz 391
0 100 —
7-Ketolithocholic acid
Y
7a- hydroxy-3-oxo5 fr cholari-24-oic-acid
mlz 389
0 10
1
20
2
30
Time (mm)
Figure 3.3. Representative chenodeoxycholic acid.
LCIMS
chromatogram
showing
metabolites
of
Chenodeoxycholic acid metabolites were extracted from a standard reaction mixture after a 30mm 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-cholan24-oic acid (m/z 389) eluted at 18 and 18.5 mm, respectively.
123
C
0
A
a 400
B
Apparent V,,,,,,, = Apparent Km
392 ± 21 prnoilminlmg
=
protein
244*3IILM
U
0
.9 .2 00
—
300
I
40
c.
Apparent Vmar =
94±6 pmoUminlmg protein
Apparent K’
=
225±l9iLM
=
3±0.4
100
I,
-0
C
0
200
9
C’3
.
a’
100
• E
a
—
p..
0 0 200
400
800
0
1000
200
C 70
600
800
D
Apparent V,,,,,,, Apparent
400
fChenodeoxycholic acid] (pM)
Chenodeoxycholic acidl (pM)
K,,,
Apparent V,,,,,, = 33 ± 3 pmoilmlnimg protein
62 ± 2 pmollmlnlmg protein =
Apparent
27±4ILM
K,,,
111±45ILM
60 30
I
0.4 (6
00
.
uE
.zs a
.C0
20
30
.
25 20
0
30
15
30
10
10
10 S
0
0.0
0.1
02
0.3
0.4
0.5
“Is
0 200
600
(Chenodeoxycholic acid] (pM)
800
200
400
600
(Chenodeoxycholic acid]
800
1000
(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, 7ketolithocholic 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.
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.n..o.n.Na.
N
—
eon. Q
oc3(.)oOc1
—
0.0. OOU
7x.Hydroxy4.oxo.510.cholan.24-oic acid r-Murlchollc acid
30 B
25 20 2E 0. 15 .0
10 5 0
r.—
r—.
.
.
.5
5
0
0 0.0.0.0. 0. 0. N 0. 0. 0. 0. 0)-,->-).>-)-n.)-)-)->-0.
o
—
N
D
CO
Q
(0
—
(0
—
C)OUOC)OUQQO
Figure 3.5. Comparison of 3-dehydrocholic acid formation from cholic acid (A) and 7ohydroxy-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
Apparent V f 1 Apparent K’
0
25
n
=
26 ± 4 pmollminlpmol P450
=
161 ± 5 LLM
=
1.7
20 30 0.
25
15
0— 00 I
0 ‘4,
.
20 15
10
. a
10 5 0
5 0 0
100
200
300
400
500
600
700
[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.
126
C 0
A
B
40
Apparent
=
40±3 pmollmin(pmol P450
.2
ApparentK,
=
165±3liiM
5
•0
C 0
30
Apparent Vmx
=
5 ± 0.4 pmouninipmol P450
Apparent K’
=
208±20M
=
2±0.3
n
4
9.
Q. 3 20
20
a
25
o
Is
20
Xe
9 E
r, 0.
.
10
o
e. — —
2
10 5
X
0 0.05
X r
0.10
0.55
020
025
030
0 0
100
200
300
400
500
[Chenodeoxycholic acidj (pM)
600
0
100
200
300
400
500
600
[Chenodeoxycholic acid] (pM)
Enzyme kinetic profiles of 7a-hydroxy-3-oxo-53-cho1an-24-oic acid (A) and Figure 3.7. 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
OOH P3A4—’
::SoTh 4 Ox
HO• Cholic acid (3a, 7a,, I 2a-tri hydroxy5p-chol an-24-oic acid)
3-Dehydrocholic acid (7a, I 2a-di hydroxy-3-oxo 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 3dehydrocholic acid formation.
128
XD T 1 h H
OH
7cz-hydroxy-3-oxo5p-cholan-24-oic acid
CYP3A4 and CYP3A5
CYP3
21 12
•
120
y-Muricholic acid (3ci;6a,7a-trihydroxy5f3-cholan-24-oic acid)
‘1 C 7 OOH
Y 4 HO O H f
Chenodeoxycholic acid
/
(3cc,7a-dihydroxy-
513-cholan-24-oic acid)
HO_cTh OH Cholic acid (3a,7ct,1 2a-trihydroxy5f3-cholan-24-oic acid)
7-Ketolithocholic acid (3x-hydroxy-7-oxo5p-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 13:907916. 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. J Lipid 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 45:35693572. 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 MICROSOMES 4
A version of this chapter will be submitted for publication as: 4 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 MgCI , 0.5 mg of human hepatic microsomal 2 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 3ketocholanoic 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 b , were purchased from BD Biosciences 5 (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 CYP3A4dependent 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 ,4-d acid, 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-cholan24-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-d 4 acid, 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]
VmX[S1 K’ + [S]
(equation 1)
(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 2 (R ) , 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 6ketolithocholic 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 13cholan-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 3ketocholanoic 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). 6Ketolithocholic 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 3ketocholanoic 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 3ketocholanoic 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 6ketolithocholic 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 3ketocholanoic 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
3-Ketocholanoic acid
Apparent Vmct. pmol/min/mg protein
Apparent Km pM
336.2 ± 13.6
21.9 ± 2.6
Hyodeoxycholic acid
47.5
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
±
±
4.3
46.3
±
13.3
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. ivie avoil
e
mar
m
fl
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
N/A
N/A
Murideoxycholic acid
0.3
35.4 ± 5.5
1.7 ± 0.3
6-Ketolithocholic acid
0.4 ± 0.0
N/A
N/A
±
±
0.0
±
13.6
N/A 29.3
±
4.4
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 donor
CYP3A4 activity* (pmol/minlmg protein)
Rate of formation of metabolites (pmol/minlmg protein)
Murideoxycholic acid
Ursodeoxycholic acid
Hyodeoxycholic acid
6-Ketolithocholic acid
3-Ketocholanoic acid
HH13
890
2.5
0.1
4.6 ± 0.2
11.3
0.2
0.9 ± 0.0
149.3
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
2 value r
±
0.97
0.97
±
±
0.98
0.95
±
±
9.1
38.3
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 r 2 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.
lOG
0
.
200
400
6.00
R
8.00
10.00
12.00
14,00
1600
18.00
20.00
2400
2200
26.00
28.00
30.00
Hyodeoxycholic acid
100
a
Murideoxycholic acid
—
3200
34:00
*
.
Chenodeoxycholic acid
*
0 2.00
4.00
6.00
800
10.00
12.00
14.00
16.00
18.00
20.00
2400
22.00
26.00
28.00
3000
V
e
-
mlz 391
Ursodeoxycholic acid
I
m!z 411
-CIio& acid as internal standard 4 d
—
32.00
34.00
mlz 389
100
7-Ketolithocholic acid 6-Ketolithocholic acid
n
t e
2.00
4.00
600
800
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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 (d -cholic acid, m/z 411) eluted at 4 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
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100
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(lithocholic acid] (IJM)
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Llithocholic acid] (liM)
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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 data points, the which represent mean values.
153
MLrideoxycholic acid Ursodeoxycholic acid Hyodeoxycholic acid
.
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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
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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
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222 .
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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).
156
0 3-Ketocholanoic acid (3-oxo- 5p-cholan-24-oic acid)
p
CYP3A4 ‘p’
21
Cyp3A4
22
Hyodeoxycholic acid (36a-dihydroxy-5-choian-24-oic acid)
12 11 •1 2
101
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Lithocholic acid
HO
(3c—iWdroxy-5p-choIan-24-oic acid)
\
CYP3A4
Ursodeoxycholic acid (3a,7p-dihydroxy- 5p-cholari-24-oic acid)
N
HO
Murideoxycholic acid (3c6p-dihydroxy- 5D-cholan-24-oic acid)’
6-Ketocholanoic acid (3ct—hydroxy-6-oxo- 5p-choian-24-oic acid)
Figure 4.6 Scheme showing lithocholic acid biotransformation by human hepatic microsomes. 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.
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. Bensaude Ri, Furet Y, Autret E, Brottes H, Billard JL and Breteau M (1988) [Cholestatic hepatitis caused by ketoconazole]. Ann Gastroenterol Hepatol (Paris) 24:55-57. Berta L, Fronticelli Baldelli C, Fazzari A, Radice E, Bargoni A, Frairia R and Gaetini A (2003) Sex steroid receptors, secondary bile acids and colorectal cancer. A possible mechanism of interaction. Panminerva Med 45:26 1-266. Bodin K, Lindbom U and Diczfalusy U (2005) Novel pathways of bile acid metabolism involving CYP3A4. Biochim Biophys Acta 1687:84-93. Calmus Y and Poupon R (1991) Ursodeoxycholic acid (UDCA) in the treatment of chronic cholestatic diseases. Biochimie 73:1335-1338. De Gottardi A, Touri F, Maurer CA, Perez A, Maurhofer 0, Ventre G, Bentzen CL, Niesor EJ and Dufour JF (2004) The bile acid nuclear receptor FXR and the bile acid binding protein IBABP are differently expressed in colon cancer. Dig Dis Sd 49:982-989. 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.
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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.
Erlinger S (1997) Drug-induced cholestasis. JHepatol 26 Suppl 1:1-4. Festi D, Montagnani M, Azzaroli F, Lodato F, Mazzella G, Roda A, Di Biase AR, Roda E, Simoni P and Colecchia A (2007) Clinical efficacy and effectiveness of ursodeoxycholic acid in cholestatic liver diseases. Curr Clin Pharmacol2:155-177. Findor JA, Sorda JA, Igartua EB and Avagnina A (1998) Ketoconazole-induced liver damage. Medicina (B Aires) 58:277-281. Fischer CD, Cooper NS, Rothschild MA and Mosbach EH (1974) Effect of dietary chenodeoxycholic acid and lithocholic acid in the rabbit. Am JDig Dis 19:877-886. 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. Guengerich FP (2003) Cytochromes P450, drugs, and diseases. Mo! Interv 3:194-204. Hoflnann AF (1999a) Bile Acids: The Good, the Bad, and the Ugly. News Physiol Sci 14:24-29. Hofmann AF (1 999b) The continuing importance of bile acids in liver and intestinal disease. Arch Intern Med 159:2647-2658. Hofmann AF (2002) Cholestatic liver disease: pathophysiology and therapeutic, options. Liver 22 Suppl 2:14-19. Hoflriann AF (2004) Detoxification of lithocholic acid, a toxic bile acid: relevance to drug hepatotoxicity. Drug Metab Rev 36:703-722. Houston JB and Kenworthy KE (2000) In vitro-in vivo scaling of CYP kinetic data not consistent with the classical Michaelis-Menten model. Drug Metab Dispos 28:246-254.
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Jezequel AM, Benedetti A, Bassotti C and Orlandi F (1994) Drug induced cholestasis. Elsevier Science By, Netherlands. Lamba JK, Lin YS, Schuetz EG and Thummel KE (2002) Genetic contribution to variable human CYP3 A-mediated metabolism. Adv Drug Deliv Rev 54:1271-1294. Liu YT, Hao HP, Liu CX, Wang GJ and Xie HG (2007) Drugs as CYP3A probes, inducers, and inhibitors. Drug Metab Rev 39:699-72 1. Makishima M, Lu TT, 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: 1313-1316. Matsubara T, Yoshinari K, Aoyama K, Sugawara M, Sekiya Y, Nagata K and Yamazoe Y (2008) Role of vitamin D receptor in the lithocholic acid-mediated CYP3A induction in vitro and in vivo. Drug Metab Dispos 36:2058-2063. Miyai K, Price VM and Fisher MM (1971) Bile acid metabolism in mammals. Ultrastructural studies on the intrahepatic cholestasis induced by lithocholic and chenodeoxycholic acids in the rat. Lab Invest 24:292-3 02. Palmer RH and Ruban Z (1966) Production of bile duct hyperplasia and gallstones by lithocholic acid. JC1In Invest 45:1255-1267. Rost D, Herrmann T, Sauer P, Schmidts HL, Stieger B, Meier PJ, Stremmel W and Stiehl A (2003) Regulation of rat organic anion transporters in bile salt-induced cholestatic hepatitis: effect of ursodeoxycholate. Hepatology 38:187-195. Setchell KD, Rodrigues CM, Clerici C, Solinas A, Morelli A, Gartung C and Boyer J (1997) Bile acid concentrations in human and rat liver tissue and in hepatocyte nuclei. Gastroenterology 112:226-235. 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, 160
carcinogens and toxic chemicals: studies with liver microsomes of 30 Japanese and 30 Caucasians. 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 MICROSOMES 4
This chapter forms part of a manuscript in preparation. 4 Deo AK and Bandiera SM. Cytochrome P450 Mediated Biotransformation of Lithocholic Acid By Mouse Hepatic Microsomes: A Comparison With Human And Rat.
12
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, 3ketocholanoic 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-d 4 acid, 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 (1C 50 0.01 jiM), CYP1A2 iM) and CYP1B1 (1C 50 0.01 1 (IC 50 0.02 iM) inhibitor, along with ketoconazole (0.01, 0.1, 1 and 10 jiM, final concentrations), a mouse CYP3A 11 inhibitor (IC 50 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, 3ketocholanoic 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 (1C 50 0.02 riM) as a specific mouse CYP3A1 1 inhibitor and quercetin (1C 50 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
56
236 ± 80
130±31
3-Ketocholanoic acid
118 ± 10
±
35*
269 ± 12*
±
10
Isolithocholic acid Hyodeoxycholicacid 6-Ketolithocholic acid
95
±
±
50
15
46±1 8
±
1
261
±
552 ± 22* 95
±
10
154±4*
35
±
4*
772 88
260±18* 90
±
7*
129
±
9
37±2
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
120 C 0
C 0
100
2:
: —
80
2 60 CO
o=0
40 20 0
•
I
0 .0
C 0
z
•
0
N
N
C
C 0 9 0
C 0 0 0
N
o 9 0
t
I-
• 0 C 0 0 0
0
0
.0
0
C 0
•
•
•
0
N
N
C 0 U
C 0 Q
0
0
0
N
N
C 0 U
C 0
0
0
U
0
z
‘‘I’ —
C
—
120 100 80 (ci-
0
5Q 40 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
ON
2 jdVi Quercetin
20pM Quercetin
5OpM Quercetin
0.01 pM Ketoconazoie
0.1 pM Ketoconazoie
I pM Ketoconazoie
10pM Ketoconazoie
No inhIbitor
F
* *
o
I— NIAIiINiF—
0
2 pM Quercetin
2 pM Quercetin
0
20 pM Quercetin
20 pMQuercetin
0
50 pM Quercetin
50 1 iVi Quercetin
o
0.01 pM Ketoconazoie
0.01 jAVi Ketoconazoie
54(etoiithocholic acid formation (% of control activity)
0.1 pM Ketoconazoie
0.1 pM Ketoconazoie
9’
0
I pM Ketoconazoie
m
7.3 0
1 pM Ketoconazoie
0 0
10 pM Ketoconazole
00 0
10 pM Ketoconazoie
0
a
CD
0
.
No inhibitor
7.3 0
No inhibitor
0
Hyodeoxycholic acid formation (% of control activity)
C)
C
* *
Ursodeoxycholic acid fomiation (% of controi activity)
Murideoxycholic acid (3a,6p-dihydroxy5p-cholan-24-oic acid)
4
CYP3A CYP2B?
CYP3A and other unidentified P450s
I
3-Ketocholanoic acid (3-oxo- 513-cholan-24-oic acid)
/
12
CYPIA, CYP2B and CYP3A Isolithocholic acid (313—hydroxy-5f3cholan-24-oic acid)
OH Ursodeoxycholic acid (3a,7-dihydroxy5-choIan-24-oic acid)
Lithocholic acid (3a—hydroxy-5-choIan-24-oic acid)
I
CYP1A, CYP2B and
CYP1A, CYP2B and CYP3A
9 Hyodeoxycholic acid (3cL,6a-dihydroxy5p-cholan-24-oic acid) 6-Ketolithocholic acid (3a—hydroxy-6-oxo5-choIan-24-oic acid)
Scheme showing lithocholic acid biotransformation by mouse hepatic Figure 5.2 microsomes. 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.
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 104:857861. 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 278:1783817844. 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 C3 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 3dehydrocholic 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 6ketolithocholic 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 6ahydroxy 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, 6ketolithocholic acid and hyodeoxycholic acid was preferred over oxidation at C-3 (to form 3ketocholanoic 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 3ketocholanoic 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 Ca 2 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.
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(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.
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Metabolite
Oxidation position
Murideoxycholic acid
Human
Mouse
C-6
1%
80%
62%
6-Ketolithocholic acid
C-6
1%
C-7
C-6 > C-3
Rat
C-7
C-6
>
1%
1%
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
Murideoxycholic acid
Human
Rat
CYP3A4
CYP2C1 1, CYP3A2,
Mouse
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
MOCA
ILCA
1% UOCA
6KLCA 1%
4%
/
3KCA
UDCA 7%
HUCA 2% 6KLCA
\
