Conjugation Reactions In Drug Metabolism: An Integrated Approach

Conjugation reactions in drug metabolism: An integrated approach

Conjugation reactions in drug metabolism: An integrated approach Substrates, co-substrates, enzymes and their interactions in vivo and in vitro

Edited by

Gerard J.Mulder Center for Bio-Pharmaceutical Sciences, University of Leiden, The Netherlands

1990 Taylor & Francis London • New York • Philadelphia

UK Taylor & Francis Ltd, 4 John St, London WC1N 2ET USA Taylor & Francis Inc., 1900 Frost Road, Suite 101, Bristol, PA 19007 This edition published in the Taylor & Francis e-Library, 2005. “To purchase your own copy of this or any of Taylor & Francis or Routledge’s collection of thousands of eBooks please go to www.eBookstore.tandf.co.uk.” Copyright © G.J.Mulder, 1990 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, electrostatic, magnetic tape, mechanical, photocopying, recording or otherwise, without the prior permission of the copyright owner and the publisher. British Library Cataloguing in Publication Data Conjugation reactions in drug metabolism. 1. Drugs. Action. Role of conjugation reactions I.Mulder, Gerard J. 615’.7 ISBN 0-203-48999-3 Master e-book ISBN

ISBN 0-203-79823-6 (Adobe eReader Format) ISBN 0 85066 738.0 (Print Edition) Library of Congress Cataloguing in Publication Data is available

Contents

Preface

v

Contributors

vii

1.

Introduction G.J.Mulder

1

2.

Kinetics of conjugation reactions in eliminating organs K.S.Pang

4

3.

Competition between conjugations for the same substrate G.J.Mulder

39

4.

Glucuronidation G.J.Mulder, M.W.H.Coughtrie and B.Burchell

49

5.

Sulfation G.J.Mulder and W.B.Jakoby

106

6.

Acetylation W.W.Weber, G.N.Levy and D.W.Hein

162

7.

O-Methylation D.R.Thakker and C.R.Creveling

191

8.

N-Methyltransferases S.S.Ansher and W.B.Jakoby

231

9.

S-Methylation J.L.Stevens and J.E.Bakke

249

10.

Amino acid conjugation A.J.Hutt and J.Caldwell

272

11.

Glutathione conjugation B.Ketterer and G.J.Mulder

306

12.

Epoxide hydrolase T.M.Guenthner

365

Preface

‘The conjugations may well be the most important drug biotransformation reactions.’ Thus reads the first sentence in Chapter 1 of this book; and indeed, this may be little realized, if one compares the overwhelming attention paid to cytochrome P-450-mediated biotransformation with what is published on the conjugations. Yet, oxidative metabolism of xenobiotics is usually followed by conjugation of the group created by oxidation. Only then can these compounds be readily eliminated from the organism. In addition, xenobiotics carry many groups which are already acceptor groups for conjugation. This volume summarizes the state of the art for the main conjugations. It should fill the gap between comprehensive monographs of single conjugations on one hand, and the more general reviews of drug metabolism, which often devote only a few pages to the conjugations (often considered to be just detoxication reactions) on the other. The chapters on the various conjugations are the result of a collaborative effort of authors who are familiar with the biochemical aspect of the conjugation and those whose expertise is on the biotransformation in vivo. Therefore, the conjugation is treated as a whole, reflecting the importance of enzymology for the biotransformation of xenobiotics in vivo, in perfusions or in isolated cells. Obviously, the conjugations are involved in the elimination of compounds present in the natural environment, for instance in food. In addition, and this is in fact the most studied aspect, they metabolize xenobiotics made by man, such as medicinal drugs, insecticides, etc. Moreover, many endogenous compounds, such as steroid hormones or prostaglandin derivatives, are substrates for certain forms of the transferases, so that these also play an important role in the homeostasis of the organism. The chapters present an overview, without giving all the details. Also, most chapters contain a brief section on methodology to facilitate the choice of methods for experimental work. The aim of this book is to provide a convenient source of information for both workers in the field of conjugation and those working in related areas. For those

vi

who need all the details, due reference is made to more extensive reviews on specialized subjects. G.J.MULDER February, 1989

Contributors

Sherry S.Ansher

Center for Biologs Evaluation and Research, Food and Drug Administration, Bethesda, MD 20982 (USA) Jerome E.Bakke Metabolism and Radiation Research Laboratory, Agricultural Research Service, USDA, Fargo, ND 58105 (USA) Brian Burchell Department of Biochemical Medicine, Ninewells Hospital and Medical School, Dundee DD1 9SY, Scotland (UK) John Caldwell Department of Pharmacology and Toxicology, St. Mary’s Hospital Medical School, Norfolk Place, London W2 1PG (UK) Michael W.H.Coughtrie Department of Biochemical Medicine, Ninewells Hospital and Medical School, Dundee DD1 9SY, Scotland (UK) Cyrus R.Creveling Laboratory of Bioorganic Chemistry, NIDDK, National Institutes of Health, Bethesda, MD 20892 (USA) Thomas M.Guenthner Department of Pharmacology, University of Illinois College of Medicine, Chicago, IL 60612 (USA) David W.Hein Department of Pharmacology, University of North Dakota School of Medicine, Grand Forks, ND 58201 (USA) Andrew J.Hutt Department of Pharmacy, Brighton Polytechnic, Brighton BN2 4GJ East Sussex (UK) William B.Jakoby Laboratory of Biochemistry and Metabolism, NIDDK, National Institutes of Health, Bethesda, MD 20892 (USA)

viii

Brian Ketterer

Gerald N.Levy

Gerard J.Mulder

K.Sandy Pang James L.Stevens Dhiren R.Thakker

Wendell W.Weber

Biochemistry Department, University College and Middlesex School of Medicine, London W1P 6DB (UK) Department of Pharmacology, University of Michigan Medical School, Ann Arbor, MI 48109 (USA) Division of Toxicology, Center for BioPharmaceutical Sciences, University of Leiden, 2300 RA Leiden (The Netherlands) Faculty of Pharmacy, University of Toronto, Toronto, Ontario M5S 1A1 (Canada) W.Alton Jones Cell Science Center, 10 Old Barn Road, Lake Placid, NY 12946 (USA) Department of Drug Metabolism, Glaxo Research Laboratories, Research Triangle Park, NC 27709 (USA) Department of Pharmacology, University of Michigan Medical School, Ann Arbor, MI 48109 (USA)

Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 1 Introduction Gerard J.Mulder Division of Toxicology, Center for Bio-Pharmaceutical Sciences, University of Leiden, 2300 RA Leiden, The Netherlands.

1.1.

DISCOVERY OF THE CONJUGATIONS

1

1.2.

TRANSFERASES AND CO-SUBSTRATES: BIOCHEMISTRY

2

1.3.

MOLECULAR BIOLOGY OF THE TRANSFERASES

2

The conjugations may well be the most important drug biotransformation reactions. The major reason is that xenobiotics that enter the body are usually lipid soluble and therefore are only slowly excreted in urine, the major excretory pathway in most species. Only after biotransformation in which a hydrophilic moiety has been added, such as the sulfate or glucuronic acid group, is water solubility increased and lipid solubility decreased enough to make urinary excretion possible. Usually, therefore, the major proportion of the administered drug dose is excreted as conjugates in urine and bile. Often conjugation is preceded by Phase I metabolic reactions such as hydrolysis or oxidation. The cytochrome P-450 system plays a major role in the latter. However, for compounds that already have a group available for conjugation when they enter the body, such as paracetamol (acetaminophen), conjugation alone is usually the major fate. 1.1. Discovery of the conjugations Most of the conjugations were discovered in the 19th century, when various compounds were fed to animals or human volunteers and the products in urine were analysed. In spite of the relatively primitive analytical methodology available at that time, glucuronidation, sulfation and formation of

2 CONJUGATION REACTIONS IN DRUG METABOLISM

mercapturates and hippurates were discovered. Of course it was very helpful that the compounds used could be administered in large doses to facilitate analysis, which usually required crystallization and elemental analysis. More systematic research was initiated in the 1930s, in particular byR. T.Williams and his collaborators, when the biotransformation of related series of compounds was investigated. Dose dependence of metabolism, biotransformation in various species, metabolite patterns for sometimes very complex structures and excretory pathways were explored; many of the ‘rules’ were discovered during this time. Nowadays it has become feasible to make certain predictions of the metabolism of a new compound based on its chemical structure, the test species as well as the dose level and the route of administration. 1.2. Transferases and co-substrates: biochemistry Once the metabolites of certain structural elements were known, interest shifted to how they were formed. The biotransformation enzymes thus became a subject of study and purification. First, however, the co-substrates for the conjugations had to be known in order to study the properties of the enzyme in vitro. Thus, the chemical structures of ‘active sulfate’, UDP glucuronate and S-adenosylmethionine were discovered. The enzymes could be purified, although in several cases this turned out to be rather difficult, notably because some enzymes were membrane bound and tended to lose activity when solubilized from the membrane. As the transferases were purified it became clear that invariably each was composed of a family of separate forms with more or less overlapping substrate specificity. It is possible to separate these forms and study their properties in pure form; however, the unavoidable problem of purified enzymes is that the properties of the pure enzyme may be quite different from those of the enzyme in vivo. This certainly applies to an enzyme like UDP glucuronosyltransferase, which has to be solubilized from its normal membrane environment in order to purify it. 1.3. Molecular biology of the transferases As the biochemistry of the transferase systems became known, the main emphasis in this area shifted to molecular biological aspects of these enzymes. Using c-DNA libraries the various enzyme forms can be separately expressed, the control of their transcription can be studied, and their primary structure can be determined. This type of work has, understandably, moved away from more

INTRODUCTION 3

traditional drug metabolism research with a more pharmacological or toxicological perspective. However, it has its impact on our insight in regulation of drug metabolism in vivo. In the present work, an attempt is made to combine the biochemistry of the transferases with the more classical drug metabolism work in vivo or in perfused organ and isolated cell studies.

Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 2 Kinetics of conjugation reactions in eliminating organs K.Sandy Pang Faculty of Pharmacy, University of Toronto, Toronto, Ontario M5S 1A1, Canada

2.1.

INTRODUCTION

5

2.2.

COMPETING CONJUGATION PATHWAYS IN LIVER

8

Kinetic modeling of conjugation pathways

2.3.

9

Influence of kinetic constants

12

Influence of enzymic distributions

14

CONJUGATION OF PHENOLIC SUBSTRATES IN LIVER

15

Substrate concentration and competing pathways

15

Inhibition of sulfation on competing conjugative pathways

16

Estimations of kinetic parameters

18

Uneven distribution of enzymic activities

19

Kinetic modeling of metabolic data

22

Conjugations mediated by the same isoenzyme

23

Sequential conjugation

25

2.4.

CO-SUBSTRATE

27

2.5.

DIFFUSIONAL BARRIERS FOR CONJUGATES

29

2.6.

Liver

29

Kidney

30

FLOW

30

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 5

2.7.

2.8.

EXTRAHEPATIC CONJUGATION

31

Intestinal conjugation relative to liver conjugation

31

Kidney

30

CONCLUDING REMARKS

34

ABBREVIATIONS

34

REFERENCES

35

2.1. Introduction Conjugations are ubiquitous among drug-eliminating organs and require the presence of both the transferases and co-substrates. Generally speaking, the liver is the most important organ involved in the conjugation of xenobiotics, though demonstrable lung (Cassidy and Houston, 1984), intestine (Barr and Riegelman, 1970; Dollery et al., 1971; Bock and Winne, 1975; Pinkus et al., 1977; Shibasaki et al., 1981), kidney (Tremaine et al., 1984) and skin (Pannather et al., 1978) conjugation abilities have also been reported. To adequately address the topic of conjugation kinetics in any eliminating organ, consideration must be given to the factors which influence drug and metabolite clearances. These include the blood perfusion rate to the organ, the transport process for the conjugate and its precursor, the extent of binding to blood (red cell and plasma proteins) components, availability of conjugative enzymes and co-substrate, the enzymic parameters and attendant zonal heterogeneity, the presence of other competing pathways and whether the formed metabolite is subject to further metabolism (Pang and Xu, 1988). The functional heterogeneities of the organ (Gumucio and Miller, 1978), notably, zonation of enzymes, is the most prominent factor. Much is known about the functional metabolic heterogeneity of the liver, the major organ for conjugation. Hepatic drug conjugation is emphasized in this chapter to illustrate the various rate-determining factors: flow, enzyme, cosubstrate availability that control the rate of conjugation within an intact organ. The concepts developed throughout this chapter on the competition between sulfation and glucuronidation for common phenolic substrates within the singlepass perfused liver are extended to other organs, e.g. the intestine and liver (Pang, 1986; Xu et al., 1989a) and can be applied to other competing pathways, e.g. amino acid conjugation versus glucuronidation of arylcarboxylic acids (Chapter 10).

6 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.1. Blood supply of the simple liver acinus and the zonal arrangement of cells. The terminal afferent vessels, hepatic arterioles, and distributing portal veins leave the large portal space (P.S.). Zones 1, 2 and 3 represent areas supplied with blood of first, second, and third quality with regard to oxygen and nutrient contents. Zones 1, 2 and 3 designate the corresponding areas in an acinar unit. Blood from each micro-circulatory unit is collected by at least two terminal hepatic venules (T.h.v.). The perivenous area is formed by the most peripheral portions of Zone 3 of several adjacent acini (reproduced from Rappaport et al., 1954).

For the description of conjugation kinetics in the liver, the structure of the organ and microcirculation must be taken into consideration. The smallest functional unit of the liver is known as the acinus (Rappaport et al., 1954; Rappaport, 1958), which consists of a terminal portal venule and hepatic arteriole, a bile duct, lymph vessels, and nerves. There is a zonal relationship between the cells constituting the acinus and the blood supply. The hepatocytes situated close to the portal space are first supplied with fresh blood (rich in oxygen and nutrients), and these cells are called Zone 1 cells. In contrast, cells located close to the perihepatic venules are poorly supplied by blood in terms of oxygen and are classified as Zone 3 hepatocytes. Cells in between are called Zone 2 cells. These zones are virtually synonymous with the periportal, midzonal and perihepatic venous (pericentral) regions and differ significantly in drug uptake and metabolic activities (Figure 2.1). Upon entry into the liver, a substrate is carried along the direction of flow from the inlet to outlet of the liver. A substrate gains access to the metabolic machinery along the direction of flow into the Disse Space, an interstitium which allows equilibrative exchange with the hepatocytes. Following substrate

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 7

Figure 2.2. The distributed-in-space phenomenon in drug processing. A schematic representation of uneven distribution of drug metabolizing enzymes in the liver, Systems I and II which are either involved in parallel, competing, or sequential metabolic pathways. Drug processing, occurring along the direction of flow of substrate, occurs from left to right, in a distributed-in-space fashion. The enzymic distributions of Systems I and II are described by the median distances, the distance from the inlet of the liver to the median (plane which divides the amount of enzyme into equal halves). As shown, System I is anteriorly localized relative to System II along the direction of flow of substrate.

access into the cells, substrate conversion, biliary excretion or efflux back to the sinusoid occurs. The successive uptake of substrate along the direction of flow may be defined as a distributed-in-space phenomenon (Goresky et al., 1973), inasmuch as transport, recruitment of enzymic activities, excretion, and consumption of co-substrates commensurate with substrate flow (Figure 2.2). A substrate delivered into the liver will first arrive at the anteriorly-located enzyme system (System I) and generate the corresponding metabolites; formation of metabolites by a posteriorly-located enzyme system (System II) occurs when residual substrate arrives downstream for recruitment of such metabolic activities. For these two systems, the relative locations of each system may be described with respect to its median (or centre) of enzymic distribution, the plane which divides total enzymic activity into halves. The median or the median distance serves to interrelate the distance between inlet of the liver and the bulk of the enzyme. System I is an anterior pathway in relation to System II, since its median (or centre) of distribution precedes that for System II (Figure 2.2). The concentration of substrate at any point x is influenced by events at x or preceding x due to removal of substrate and consequently modulating the intrahepatic substrate concentration gradient. At low or intermediate substrate concentrations (relative to the Kms), a greater substrate concentration gradient is found in comparison to higher input substrate concentrations. Model-

8 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 2.1. Known heterogeneities on intercellular distributions of drug metabolizing activities in the liver.

dependent sensitivity is high at these low input concentrations. At higher concentrations, the intrahepatic concentration gradient is much decreased, and all enzymic systems (anterior and posterior) will be recruited by the substrate, rendering the influence of enzymic heterogeneity unimportant. Metabolic zonation of enzyme system(s) within the liver lobule has been well studied within the past few years. Direct and indirect techniques have shown an enriched presence of the cytochrome P-450s, epoxide hydrolase, glutathione Stransferases and UDP glucuronosyltransferases in the perivenous region (Zone 3), and sulfotransferases in the periportal region of the liver (Table 2.1). Elimination of a substrate/metabolite occurs only when it is present along the sinusoid and if it gains access into hepatocytes. Given the marked enzymic heterogeneities noted for Phase I and Phase II reactions, the nature and proportion of conjugates formed as primary or secondary metabolites arising from parallel or sequential pathways is expected to differ. 2.2. Competing conjugation pathways in liver Sulfation and glucuronidation are competing pathways of phenolic substrates. The studies on conjugation of phenols in isolated rat hepatocytes showed that the avid sulfation was eventually taken over by glucuronidation at increasing concentrations of the substrate (see Chapters 4 and 5). This response has been attributed to a detergent-like effect of substrates on the latency of membranebound UDP glucuronosyltransferases. In reality, these differences are readily explained solely on the basis of the differences in Km and Vmax of the

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 9

conjugative systems. By virtue of its high affinity, sulfation is ordinarily the dominant pathway and is prone to saturation upon substrate loading. The latter occurrence creates a substrate-sparing effect and allows for disproportionate increases in glucuronidation rates relative to those seen ordinarily at low substrate concentration, consequently giving a false impression of activation of the glucuronidation system. Within the intact liver, the same phenomenon exists, albeit in a distributedinspace fashion, and drug processing occurs along the direction of flow and according to the zonal enrichments of the drug metabolizing enzyme systems. This topic has been explored quite extensively both theoretically and experimentally (Pang et al., 1983; Morris and Pang, 1987; Pang et al., 1987; Morris et al., 1988a, 1988b; Xu and Pang, 1989a; Xu et al., 1989b). Let us assume for now, that co-substrate within cells and substrate entry into the hepatocytes are not rate-limiting. Kinetic modeling of conjugation pathways An understanding of the kinetics of conjugation has been approached by theoretical modeling of the liver with various enzymic distribution patterns and kinetic constants. The liver can be viewed simplistically as a series of parallel units or sinusoids of length L, each receiving an equal fraction of blood flow and surrounded by single sheets of hepatocytes, one cell thick, on either side. Enzymic activity within any cell located at the same point x, or Vmax, x, cell, may be summed for all hepatocytes at point x, such that a distribution of the overall Vmax, x results for the whole liver (Figure 2.3). The overall metabolic activities (Vmax) is expressed as

Analogously, the distribution pattern of another enzyme system (either for drug removal or metabolite transformation) may be similarly constructed. It is recognized that this assumption describes a slightly lower Vmax than the actual constant due to a lack of consideration of the capillary transit time (Goresky et al., 1973; Pang et al., 1988a). The Km for the metabolic pathway is considered constant for the same system. Steady-state mass balanced rate equations may be written for the change of sinusoidal drug or metabolite concentrations at any point x due to processing of drug and formation of the metabolite(s) upon a single passage of drug through the liver (similar to the single-pass rat liver perfusion). If the metabolite undergoes sequential metabolism, then mass balance equations incorporating such pathways would be included, and the rate of metabolite formation is represented by the sum of the efflux rates (into hepatic vein and in bile) of the

10 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.3. A schematic representation of extreme distribution patterns for sulfation (horizontal lines) and glucuronidation (vertical lines) activities (Models A, B, and C) in liver. The same amounts of enzymes for sulfation and glucuronidation are present for each model (reproduced from Pang et al., 1987).

metabolite and its subsequent metabolites. Variables such as input concentration of drug and direction of flow are used as perturbations to provide simulated data. As an illustration of this simulation procedure, a compound, with a constant unbound fraction in blood (fB) and which undergoes sulfation and glucuronidation, is used (Figure 2.3). Three extreme situations of enzymic distribution patterns are presented: even distribution for both enzymes (Model A); an exclusive anteriorly-located sulfation but a posteriorly-distributed glucuronidation system (Model B); and the reversed distribution of the Model B (Model C). The sulfation system (I) is assigned as the pathway with higher affinity but lower capacity relative to glucuronidation (II). This is true for many phenolic compounds, including harmol (Pang et al., 1981; Pang et al., 1983) salicylamide (Koike et al., 1981; Xu and Pang, 1989a; Xu et al., 1989b), and acetaminophen (Watari et al., 1983). Based on mass balance considerations, the rate of change of drug (dCx) and conjugates and , respectively, for the sulfate and glucuronide conjugates) over a small increment of length, dx, in a single-pass design across the liver preparations are described by following Michaelis-Menten equations

(2.1)

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 11

(2.2) and respectively denote the rates of sulfation and glucuronidation where at point x. At the outlet of the liver , these concentrations represent the output concentrations. Drug disappearance at steady state during a single passage of drug is denoted by the extraction ratio, E

where CIn and COut are the steady-state input and output drug concentrations, respectively. The length-averaged, steady-state, metabolite formation rate, , is estimated by flow, Q, multiplied by the output concentration of conjugate/ metabolite ; biliary excretion of conjugate/ metabolite is treated as zero for the sake of simplicity. Experimentally, the rate of conjugate formation is found by summing the rates of efflux of metabolite in hepatic venous blood and in bile

Upon assignments of Km and Vmax and the enzymic distribution patterns, E and the steady-state rates of sulfation (S) and glucuronidation (G) may be simulated by numerical approximation or by simple FORTRAN programs for a microcomputer. The distribution (Vmax at any point x, Vmax, x), together with the Km for these two systems, dictate the rates of metabolite formation at given inlet substrate concentrations. From Table 2.2, it can be shown that for the assigned constants, the drug is highly extracted (0·996) at low input concentration (0·1 µM) and the values are identical for all three models, indicating that the clearance of the compound is flow-limited. However, the proportion of conjugates formed is highly modeldependent. Model B generates the highest proportion of sulfate conjugate (S) but the smallest proportion of glucuronidate conjugate (G); Model C gives the opposite results of Model B; and Model A predicts intermediate S and G. At 50 µM input substrate concentration (which exceeds but not ), differences in E are observed among three models: Model C predicts the highest value; Model B is the least efficient system; and Model A, again, is intermediate. When input concentration exceeds both and (400 µM), saturation of all metabolic pathways is found: E is decreased to 0·28 for all models and no difference is observed on S and G from different models. Predictions from models of intermediate distributions may be inferred from those of Models B and C serving as extreme distributions and Model A for mean values. A distinct pattern for S and G is found for Models A and B where sulfation activity is distributed identically or is located exclusively anterior to glucuronidation activity, respectively. In these cases, the rate of glucuronidation

12 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 2.2. Simulated substrate removal and conjugation rates for a high-affinity , low-capacity sulfation system and a lowaffinity , high-capacity glucuronidation system for the three enzyme-distributed models (taken from Pang et al., 1987).

a

E is the steady-state hepatic extraction ratio; S is the steady-state rate of sulfation, expressed as a fraction of the input rate; c G is the steady-state rate of glucuronidation, expressed as a fraction of the input rate. b

(a low-affinity system) displays disproportionate increases upon saturation of sulfation at high substrate concentrations (Figure 2.4). The modulation of intrahepatic substrate concentration due to the high-affinity sulfation pathway is the explanation. At high substrate concentration, the concentration gradient is much decreased, whence all enzymic systems (anterior and posterior) are fully recruited by the substrate, and all sulfation and glucuronidation rates are identical regardless of enzyme heterogeneity. Results from the simulation procedure, therefore, can demonstrate how enzymic distributions and the enzymic constants influence metabolite formation relative to the input substrate concentration. Sequential metabolism of one of the generated metabolites may be incorporated in the above equations (eqns. 2.1 or 2.2). Moreover, if the unbound fraction changes during the single passage of drug through the liver, the above equations may be readily modified such that the rate of elimination is based on the unbound moiety (Xu et al., 1989b). Influence of kinetic constants The influence of enzymic parameters of Km and Vmax has also been shown by the same simulation technique. Upon decreasing the Km (Figure 2.4) or increasing the Vmax for glucuronidation (Figure 2.5), the efficiency in glucuronidation is improved and hence the rate of glucuronidation for Models A and B displays even more marked compensatory increases upon saturation of sulfation (Morris and Pang, 1987; Pang et al., 1987). Conversely, when the Km is increased or the Vmax is decreased for glucuronidation, the apparent compensatory effect

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 13

Figure 2.4. The influence of Km on the rates of glucuronidation for Models A to C (Figure 2.3). The parameters for simulation were: the enzymic constants: , nmol min−1 for sulfation and nmol min−1 for glucuronidation. The Km for glucuronidation was (A) 5 µM; (B) 50 µM; (C) 200 µM; and (D) 500 µM (from Morris and Pang, 1987).

observed for glucuronidation rate versus concentration for Models A and B is reduced. Induction effects within an intact organ must be interpreted with caution, inasmuch as heterogeneities in drug metabolizing activities among competitive pathways exist. The effects of induction and inhibition of two competing pathways on conjugate formation can be reasoned. An equal induction of both anterior and posterior pathways leads to an increased formation rate of metabolite from the anterior pathway but may not readily reveal induction of the posterior system. As complete substrate recruitment by upstream hepatocytes occurs, little substrate is spared for recruitment of downstream activity and a reduction in the formation of metabolite from the posterior pathway is observed, although the pathway is induced (Morris and Pang, 1987). However, inductive effects on the posterior pathway become readily detectable at high inlet substrate concentrations when the anterior pathway becomes saturated.

14 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.5. Influence of on the rates of glucuronidation for Models A to C. The parameters for simulation were identical to those used in Figure 2.4, but was (A) 2000 nmol min−1; (B) 1000 nmol min−1; and (C) 100 nmol min−1 (from Morris and Pang, 1987).

Inhibition of drug metabolism pathways is subject to similar considerations. An inhibited posterior pathway will have little influence on an anterior pathway, whereas inhibition of an anterior pathway may bring about increased formation of metabolites for the posterior pathway (Morris and Pang, 1987), which is unaffected by the inhibitor. Again, the reasoning for this draws upon the availability of substrate for enzyme recruitment. Influence of enzymic distributions The influence of enzymic distributions has been illustrated in the above simulation procedure, that Models A, B, and C provide different metabolites (S and G). Model B is the reverse of Model C in terms of enzymic distributions. At low input substrate concentration, the entire system is flow-limited (see condition for 0·1 µM), the extraction ratio remains high and constant, and the intrahepatic concentration gradient is the steepest. With substrate flow from the portal to hepatic vein, little or no downstream conjugate is formed due to the efficient and almost complete removal by the upstream pathway. With a

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 15

reversal in flow direction (cf. Models B and C), the downstream conjugation pathway becomes upstream in terms of the flow of substrate and receives substrate at the entrance to the organ. The metabolic ratios (S/G) will alter dramatically due to the direction of delivery of substrate. At intermediate concentrations (above but below ), saturation of the sulfation pathway occurs, and both E and S/G would change. At high input substrate concentrations which exceed the Kms (the condition of 400 µM), E and S/G would not be affected due to complete recruitment of all enzymic activities within hepatocytes. This type of kinetic modeling has been used to probe the best substrate concentration for discernment of enzymic distributions in normal and retrograde liver perfusions (to be described later). The intermediate concentration is best as both E and S/G ratios are expected to change (cf. Models B and C at the intermediate concentration, 50 µM, Table 2.2). 2.3. Conjugation of phenolic substrates in liver Substrate concentration and competing pathways Several examples are provided herein to illustrate the concept of substrate recruitment of zonal hepatocyte conjugation activities in single-pass rat liver perfusion studies. Three phenolic substrates are presently used as examples in the perfused rat liver preparation: harmol, which forms sulfate and glucuronide conjugates (Pang et al., 1981); gentisamide (GAM) forms two monosulfates (GAM-2S and GAM-5S) and a monoglucuronide conjugate (GAM-5G) (Morris et al., 1988a); its precursor, salicylamide, forms the sulfate (SAM-S) and glucuronide (SAM-G) as well as GAM in once-through rat liver perfusion studies (Figure 2.6; Xu et al., 1989b). At low inlet concentration of harmol, salicylamide and GAM, sulfation (forming harmol sulfate, SAM-S, and GAM-5S and GAM-2S) was the dominant pathway; glucuronidation/hydroxylation rates were comparatively lower (Figure 2.7). At increasing substrate input, saturation of sulfation for all substrates was being approached and was accompanied by disproportionate increases in glucuronidation/hydroxylation rates. During once-through perfusion, E remained constant for harmol (Pang et al., 1981), gentisamide (Morris et al., 1988a) and salicylamide (Xu et al., 1989b) without constancy in proportions of the conjugates (Figure 2.7). For harmol, rates of formation of harmol glucuronide increased more than anticipated at high input rates. Even for GAM rates of GAM-5G formation displayed non-Michaelis-Menten-like

16 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.6. Metabolic pathways of gentisamide and salicylamide in the rat liver.

characteristics as did harmol glucuronide. The same comment applies to salicylamide glucuronidation and hydroxylation. The higher-than-expected increase in steady-state glucuronidation and hydroxylation rates with concentration may be readily explained purely on the basis that sulfation is the higher-affinity pathway relative to glucuronidation and hydroxylation when enzymes are evenly distributed. The glucuronidation and hydroxylation pathways appear to act as backup systems within this concentration range. These examples illustrate that first-order drug disappearance does not mandate first-order metabolite formation. Rather, the apparent first-order behaviour of the drug may elicit different proportions of metabolites formed. Inhibition of sulfation on competing conjugative pathways When the substrate is spared by an inhibition/suppression of the sulfation pathway with a specific inhibitor such as 2, 6-dichloro-4-nitrophenol (DCNP) (Koster et al., 1982), glucuronidation reveals itself as an effective conjugation pathway of harmol and salicylamide. The competing pathways of sulfation and glucuronidation for harmol ((Pang et al., 1983) and salicylamide) (Xu et al., 1989b) with salicylamide hydroxylation, now display formation patterns conforming to what are expected of simple Michaelis-Menten behaviour

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 17

Figure 2.7. Concentration-dependent elimination of harmol, gentisamide and salicylamide in the single-pass perfused rat liver preparation (10 ml min−1). The steadystate rates of metabolite formation for harmol, gentisamide, and salicylamide were concentration-dependent. At increasing substrate concentrations wherein sulfation was apparently becoming saturated for all three substrates, glucuronidation and hydroxylation (for salicylamide only) rates increased more than proportionately with input concentration (from Pang et al., 1981; Morris et al., 1988a; Xu et al., 1989b).

18 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.8. Inhibition of salicylamide sulfation by DCNP (2, 6-dichloro-4-nitrophenol, 40–50 µM) in the single-pass perfused rat liver preparation. The steady-state sulfation rates were markedly suppressed relative to control livers (Figure 2.7). In an absence of sulfation, glucuronidation surfaced as a dominant pathway for salicylamide; the hydroxylation rates of salicylamide (sum of those for unconjugated and conjugated GAM) were only slightly increased over those for controls. The total represents the sum of all metabolic pathways (from Xu et al., 1989b).

(Figure 2.8). The composite observations on the concentration-dependent formation of conjugates, in an absence (Figure 2.7) and presence (Figure 2.8) of the sulfation inhibitor DCNP may be explained purely on the basis of a lower Km for sulfation when all enzymic systems are evenly distributed in the liver. Estimations of kinetic parameters The presence of a lower Km and Vmax for sulfation and a higher Km and Vmax for glucuronidation has been confirmed by parameter estimations of these enzymic constants in perfused liver studies. The enzymic constants for the intact liver have been routinely estimated by fitting the steady-state rate of formation of the conjugate, v, against the logarithmic average concentration, Ĉ of the substrate. The estimated in this fashion will be slightly underestimated due to a lack of consideration of heterogeneity in capillary transit times (Goresky et al., 1973; Pang et al., 1988a) (2.3) where fB is the unbound fraction of substrate in blood, and Ĉ, the logarithmic average concentration, which relates to the steady-state input and output

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 19

concentrations, CIn and COut, respectively, in single-pass liver perfusions (Goresky et al., 1973; Winkler et al., 1973) (2.4) The appropriateness of the method for multi-enzyme systems has been validated by Morris and Pang (1987) and Xu and Pang (1989a). However, some limitations have been found. Parameter estimations will be close to true values when all enzymic systems are evenly distributed. For unevenly distributed enzymic systems, the estimations will be close to their true values when the Kms of all pathways are of similar orders of magnitude; for vastly different Kms among competing pathways, only the high-affinity system will be correctly estimated (Morris and Pang, 1987). For cases where large differences in Km exist for the conjugative pathways, estimations of enzymic constants for the low-affinity, high-capacity pathway by this method are prone to error. Estimations for enzymic parameters of the low-affinity pathways, therefore, may be obtained by acquiring data upon inhibition of the high-affinity sulfation pathway with selective inhibitors such as DCNP. More precise estimations of these kinetic constants for glucuronidation may then be made from metabolic data in the presence of the sulfation inhibitor with eqn. 2.3. Table 2.3 tabulates the kinetic constants obtained for harmol and salicylamide conjugation and hydroxylation in single-pass rat liver preparations; the glucuronidation and hydroxylation (OH) parameters were obtained in the presence of DCNP since and . Because the kinetic parameters for GAM conjugation are similar, parameter estimations obtained from metabolic data without DCNP in the perfused rat liver would serve as true estimates (Morris et al., 1988a). It must be mentioned that for salicylamide, nonlinear drug protein binding is observed with perfusate and the unbound fraction changes during a single passage of salicylamide through the rat liver. For this reason, the unbound logarithmic average concentration Ĉu was used (similar to eqn. 2.4, expressed in terms of unbound input and output concentrations; Xu and Pang, 1989b). Uneven distribution of enzymic activities For an assignment of relative enzymic distribution patterns, further definition is required. The technique of normal (N) and retrograde (R) once-through liver perfusion (Pang and Terrell, 1981), with perfusate entering the liver from the hepatic vein and leaving via the portal vein, has been used as a tool in the examination of zonal, metabolic heterogeneity (Conway et al., 1982, 1984, 1988; Pang et al., 1983; Morris et al., 1988b; Xu and Pang, 1989a). By employing an identical inlet concentration once through the same liver preparation, a substrate entering the liver from the portal vein will first encounter periportal

20 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 2.3. Kinetic constants for harmol, gentisamide, and salicylamide conjugations.

a The input concentrations of GAM were varied (increased or decreased) stepwise at >35 min to the once-through perfused liver to provide different rates of formation of conjugates. b Estimated in the presence of DCNP (40–50 µM).

cells and later perihepatic venous cells. Reversing the flow direction, with substrate entering the liver from the hepatic vein, brings about a reversal in the order in which enzymic activities are encountered by the substrate. Upon altering from N to R perfusion, any inherent heterogeneity in metabolic activity will be readily revealed by an increase or decrease in metabolite formation. The basic tenet of this approach is that an equal number of hepatocytes is accessed by the substrate, now in a reverse fashion to that ordinarily seen in the normal direction (portal vein to hepatic vein). Only recently, the multiple indicator dilution technique (St-Pierre et al., 1989) provided evidence that the intracellular water space of the perfused rat liver preparation had remained unaltered when the direction of flow was changed from N to R perfusion. Additionally, the unaltered metabolic recruitment of ethanol (St-Pierre et al., 1989), a substrate which shares the same space as intracellular water (Goresky et al., 1983), for both N and R perfusions strongly supports the use of the technique for investigations of acinar distributions for conjugative enzymes. It should be pointed out that the ratedetermining step of the metabolic reaction (e.g. co-substrate supply, enzyme activity, or presence of a diffusional barrier) is not identified by the NR technique. The overall reaction rate and enzymic parameters, Km and Vmax, sometimes involving a multiplicity of isoenzymes with overlapping specificities, are assessed. This method differs from other direct techniques such as immunohistochemical and staining techniques (Redick et al., 1982; Chowdhury et al., 1985; Knapp et al., 1988) which reveal the presence of isoenzyme(s) but seldom the associated activities. As discussed earlier, the elucidative properties of these NR studies are best at inlet substrate concentration (CIn) far below Km for both systems, or above Km for sulfation and below Km for glucuronidation (Models B versus C, Figure 2.4, Table 2.2). At CIn far below Km, E is expected to remain constant during both N

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 21

Figure 2.9. Normal and retrograde perfusion of 8 µM gentisamide in a single-pass rat liver preparation. The unchanged 2S/5S ratio during both normal and retrograde perfusions suggests an identical distribution for the GAM 2- and 5-sulfoconjugation systems, whereas a decreased 5S/5G ratio during retrograde flow suggests an anterior sulfation system in relation to the glucuronidation system(reproduced from Morris et al., 1988b).

and R as clearance is flow-limited (elimination within cells is limited by supply of substrate). For intermediate CIn (above Km for sulfation and below Km for glucuronidation) changes in E and the S/G ratio upon alteration of flow direction from N to R is expected when the median distances for the enzyme systems differ. If sulfation activities are more anteriorly located in relation to glucuronidation activities, E would increase and the S/G ratio would decrease (Morris and Pang, 1987). This kind of behaviour was observed in perfusion studies with harmol, GAM (Figure 2.9) and salicylamide (Figure 2.10). For harmol, S/G decreased but E increased with R (Pang et al., 1983). For GAM, GAM-2S/GAM-5S remained constant during NR whereas GAM-5S/GAM-5G decreased during R, suggesting an identical distribution for GAM sulfation activities which are of anterior distribution in comparison to glucuronidation activity. For salicylamide, SAM-S/ SAM-OH and SAM-S/SAM-G were decreased during R, suggesting that sulfation activity is upstream in relation to glucuronidation and hydroxylation

22 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.10. Normal and retrograde perfusion of 130 µM salicylamide in the single-pass rat liver. The decreased SAM-S/SAM-G and SAM-S/SAM-OH ratios suggest an anterior salicylamide sulfation system in relation to salicylamide glucuronidation and hydroxylation systems (from Xu and Pang, 1989a).

activities (Figure 2.10). Similar observations have been found for 7hydroxycoumarin conjugation, with drug disappearance being monitored by micro-fluorescence with light guides inserted directly into the periportal and perihepatic venous regions of the liver (Conway et al., 1982; 1988). A periportal abundance of acetaminophen sulfation activities has also been reported for the rat liver (Pang and Terrell, 1981; Pang et al., 1988a). Kinetic modeling of metabolic data Kinetic modeling of the three substrates, harmol, GAM, and SAM, for their competing pathways may be performed as outlined in the previous section. Various simplified metabolic patterns are used to describe the competing processes. Values of E and steady-state metabolic ratios (S/G or GAM-2S/ GAM-5S, GAM-2S/GAM-5G, SAM-S/SAM-G or SAM-S/SAM-OH, where S, G, and OH represent sulfation, glucuronidation, and hydroxylation rates) are simulated based on plausible combinations of the enzymic patterns and compared to observations. The enzyme-distributed model that fits the data best (Figure 2.11) reveals a periportally concentrated sulfation activity, an evenly distributed glucuronidation activity towards harmol, GAM and salicylamide, and an enriched perihepatic venous salicylamide hydroxylation activity (Dawson et al., 1985; Morris et al., 1988b; Xu and Pang, 1989a). The observed

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 23

Figure 2.11. Enzymic distribution patterns for the metabolism of harmol, gentisamide and salicylamide in the perfused rat liver. The sulfation (decreasing from inlet to outlet), glucuronidation (even), and hydroxylation (increasing from inlet to outlet) systems show the rank order of median distances: sulfation
metabolic data for both GAM and salicylamide are well predicted by the enzymic distributions (Figure 2.12). Conjugations mediated by the same isoenzyme An interesting relationship is seen for the conjugation of GAM. The kinetic data for the two sulfation pathways prompted whether the same isoenzyme of arylsulfotransferase is mediating the sulfation at the 2- and 5-positions of GAM. The behaviour of this kinetic system (Figure 2.13), involving one isoenzyme but two enzyme substrate complexes (ES1 and ES2) for formation of two distinct products (P1 and P2) and the equilibrium concentrations may be expressed as follows

where [E] and [S], [ES1], and [ES2] are the steady-state concentrations of enzyme, substrate, and the two enzyme substrate complexes, respectively, interrelated by the rate constants, k1, k2, k3, k4, k5, and k6, as defined in Figure 2.13. The apparent Kms equal the reciprocal of the sum of the reciprocals of individual Kms divided by the unbound fraction in blood, fB (Potter et al., 1977: Gillette, 1982; Morris et al., 1988b)

24 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.12. Observed salicylamide metabolism rates and predicted rates based on the enzymic parameters (Table 2.3) and enzymic distributions (Figure 2.15) for single-pass rat liver preparations in an absence (A) and presence (B) of DCNP, the sulfation inhibitor. Good match was obtained between observations (points) versus the predictions (line) (reproduced from Xu and Pang, 1989a).

The ratio of the rates of formation of metabolites arising from these two parallel pathways, v1/v2, becomes (2.5) and is a quotient of the ratios of the Vmax/Km for the parallel pathways. A lack of concentration dependence in the ratio of metabolite formation is predicted from eqn. 2.5, as noted by Gillette (1982) and Potter et al. (1977) for the

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 25

Figure 2.13. Formation of multiple metabolites from one isoenzyme, E, and its substrate, S. Two enzyme substrate complexes, ES1 and ES2, are in equilibria with free E and S, with forward and backward rate-constants, k1 and k2 and k4 and k5, respectively. These enzyme substrate complexes give rise to products, P1 and P2, respectively, with corresponding rate constants, k3 and k6 (reproduced from Morris et al., 1988b).

co-processing of substrate by an isoenzyme in microsomes. The additional criterion for multiple product formation arising from a single isoenzyme for the intact liver is an identical distribution of activities for formation of the products. The apparent Kms must be identical but the apparent Vmaxs for these pathways may differ resulting in constant ratios of products’ formation among all CIns and for NR (eqn. 2.5). The coupling between an isoenzyme and substrate, which forms either ES1 or ES2 enzyme-substrate complexes, is kinetically distinct from competition reactions where two or more substrates compete for the same enzyme or enzyme site(s). Sequential conjugation The hydroxylated product of salicylamide, GAM, is found to be exclusively glucuronidated by the perfused rat liver. However, preformed GAM gave rise mostly to GAM-2S and GAM-5S, the monosulfate conjugates at comparable rat liver perfusion studies (Figure 2.14), Model predictions obtained by combining the enzymic models for salicylamide and GAM metabolism, however, failed to explain the discrepant fates of GAM as a preformed or generated metabolite. The predictions persisted to show that GAM sulfations are dominant pathways over GAM glucuronidation after administration of salicylamide (Xu and Pang, 1989a; Xu et al., 1989b). This discrepancy between predictions and observations might be due to a tight coupling or close proximity between hydroxylation and glucuronidation, reactions mediated by membrane-bound enzymes on the endoplasmic reticulum that are not accounted for in modeling. GAM once formed might be glucuronidated rather than being released to the cytosol for

26 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.14. The rate of formation of gentisamide metabolites, expressed as a per cent of total rate of metabolism of gentisamide, after perfusing preformed GAM (0.91 to 482 µM) once-through rat livers at a flow rate of 10 ml min−1 (data excerpted from Morris et al., 1988a and reproduced from Xu et al., 1989b). At low input GAM concentrations, GAM-5S and GAM-2S were formed more than GAM-5G. However, at higher GAM concentrations, the fractional metabolism to form GAM-5G increased whereas those to form GAM-2S and GAM-5S decreased.

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 27

sulfation. However, upon O-deethylation of phenacetin, acetaminophen so generated is predominantly sulfated instead of glucuronidated (Pang and Gillette, 1978; Pang and Terrell, 1981; Pang et al., 1988b), perhaps because of the markedly higher Km for glucuronidation over sulfation, inhibiting formation of acetaminophen glucuronide, or that O-deethylation and glucuronidation are less coupled than hydroxylation and glucuronidation. This aspect remains to be pursued further for an understanding of the basis underlying secondary conjugate formation. 2.4. Co-substrate Very little attention has been spent on the impact of co-substrate concentration and depletion on the kinetics of conjugation. One of the exceptions is a study on the kinetics of glutathione (GSH) conjugation by Chen and Gillette (1988) for the reactive metabolite of acetaminophen. Acetaminophen (A), a hepatotoxin at high doses, is known to form a reactive metabolite (M) which either conjugates with GSH to form acetaminophen GSH conjugate or binds covalently to macromolecules (Gillette, 1982). Chen and Gillette (1988) considered competing metabolic pathways for acetaminophen metabolism (sulfation and glucuronidation), the formation of a reactive metabolite and, subsequently, a GSH conjugate. They related this to GSH loss by means other than formation of acetaminophen GSH conjugate at low doses of acetaminophen, which depleted hepatic GSH but did not result in covalent binding. Essentially, first-order conditions were assumed for all metabolic pathways but a second-order reaction was used to describe formation of the GSH conjugate of acetaminophen. By incorporation of the synthesis and degradation rates of GSH, remarkably good agreement was obtained between the predictions and the observed plasma acetaminophen concentrations and timecourse of intrahepatic GSH depletion described by Jollow et al. (1974). With similar assumptions on rapid equilibration between drug in liver and outflow blood (venous equilibration) and constancy in drug binding (unbound fraction in blood, fB, is constant), the mass balance equations developed by Chen and Gillette (1988) may be modified to reflect events in the single-pass liver perfusion system. The rate of drug removal may be presented as

where Q is the liver blood flow, and CIn and COut are the respective input and output drug concentrations. VA and [A]H are the volume and concentration of acetaminophen in liver, respectively. The superscripts ‘A, Other’ and ‘A’ are the competitive pathways and formation pathway of the reactive metabolite from

28 CONJUGATION REACTIONS IN DRUG METABOLISM

acetaminophen, respectively. Formation clearance of the reactive metabolite, CLA, at steady-state is given by

where is the intrinsic clearance (Vmax/Km) for the pathway mediating formation of the reactive metabolite. For the short-lived reactive metabolite, which does not leave the cell, its rate of change in the liver is (2.6) and are the intrinsic clearances for formation of the where GSH conjugate (bimolecular reaction) and for reactive metabolite loss (firstorder reaction) by pathways other than conjugation with GSH, respectively. VM is the volume of the metabolite, and [M]H and [G]H are the concentratians of the reactive metabolite and free GSH in the liver, respectively. The rate of change of GSH concentration in the liver may be given by the following equation (2.7) is the intrinsic clearance for loss of GSH other than forming where acetaminophen GSH, VG the volume for intrahepatic GSH, is the hepatic synthesis rate of GSH, assumed to be unchanged in the presence of the depletor. The bimolecular reaction describing the formation of acetaminophen GSH conjugate may be time-dependent even under single-pass (steady-state) conditions for the drug when [G]H changes with time (eqn. 2.7) due to the rapid rate of [G]H depletion relative to the rate of synthesis. However, for short periods of time when the rate of change of liver GSH is not changing and the reactive metabolite is solely transformed to acetaminophen GSH conjugate (other pathways do not exist, or ) then eqns. 2.6 and 2.7 may be set as zero and CIn CLA (formation clearance) may be expressed in terms of [G]H Since the formation clearance CLA may be found as the product of the ratio, [acetaminophen GSH conjugate/total acetaminophen metabolites], and the total hepatic clearance of acetaminophen, and may be estimated in studies where CIn is altered in the perfused liver preparation, and is plotted against the associated [G]H. This relationship should hold even when nonenzymic GSH conjugation is a significant component of the reaction. The right side of the equation, representing the rate of GSH loss due to GSH conjugation, equals the rate of formation of the GSH conjugate (left side of equation). The

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 29

relationship also applies to acceptor substrates which form a GSH conjugate directly with GSH without going through formation of a reactive intermediate. 2.5. Diffusional barriers for conjugates Drug conjugates are of comparatively higher polarity than their precursors and are expected to experience membranes as a barrier. The consequence of this barrier effect for relatively lipophilic precursors but polar products is an accumulation of polar metabolites generated within tissues, lending towards an enhancement of elimination of the metabolites primarily through excretion either via the biliary tree or the renal tubules. The same barrier, however, prevents entry of polar, preformed metabolites into tissues and thus prevents their elimination (De Lannoy and Pang, 1987). Liver This type of behaviour is found for styrene oxide GSH conjugates. Upon recirculation of styrene oxide into the perfused rat liver, about 23% dose was recovered as GSH conjugates in bile after one hour (Smith et al., 1983). In contrast, with recirculation of preformed styrene oxide GSH conjugate, only 5 to 6% dose was recovered in bile at the end of two and a half hours (Steele et al., 1981). The same is observed for 4-methylumbelliferone (4MU) sulfate and glucuronide conjugates in rat liver perfusion studies. After the administration of 4-MU, the extents of biliary excretion of the conjugates by the perfused rat liver greatly exceeded those when 4-MU glucuronide or sulfate themselves were administered, and the occurrences are attributed to the hepatocyte membrane posing as a transport barrier (Miyauchi et al., 1988). In reversible conjugation-deconjugation reactions, conjugates effuxed back into the sinusoid are required to re-enter cells downstream along the direction of flow to effect deconjugation. Although the likelihood of this occurring is slim as polar conjugates gain access into hepatocytes only with difficulty, deconjugation, a relatively high Km reaction, has been observed at high 4-MU sulfate concentrations in perfused livers (Anundi et al., 1986) and the deconjugation activity themselves were confirmed in micro-dissected liver tissues (El Mouelhi and Kauffman, 1986). Much deconjugation can also occur at the time of conjugation, as in futile metabolism, before the conjugates are released into the sinusoids.

30 CONJUGATION REACTIONS IN DRUG METABOLISM

Kidney Conjugation is known to occur within cells, and these intracellular species are subject only to transport at the brush border (luminal) membrane. By contrast, conjugates in the circulation are subject to barrier-limited transport (Silverman and Goresky, 1965; Trainor and Silverman, 1982) at both the luminal and the basolateral membrane. As shown for anionic substrates, such as glucuronide and sulfate conjugates of catechol (dihydroxybenzoate) (Rennick and Quebbemann, 1970), the sulfate conjugate of morphine (Watrous et al., 1970) and 5-hydroxyindoleacetic acid, the metabolite of 5hydroxytryptamine (Hakim et al., 1970), which share similar carrier-mediated system(s) as p-aminohippuric acid and probenecid, those generated within cells are not prone to transport inhibition at the basolateral membrane as are preformed species. Furthermore, if a preformed metabolite is prevented from entry into renal cells due to its polar characteristics, a barrier not existing for the intracellular-generated conjugate, different extents of entry or excretion between preformed and generated metabolites are expected (de Lannoy and Pang, 1987; de Lannoy et al., 1989). Bekersky et al. (1980) have also postulated the presence of a diffusional barrier for salicylic acid entry into renal tubular cells in the recirculating perfused rat kidney. Salicylic acid is conjugated with glycine to salicyluric acid in the recirculating rat kidney, rendering both systemic (preformed) and intrarenal (nascently formed) metabolites, which undergo reversible metabolism. Though the presence of saturable protein binding and reversible metabolism complicated data interpretation for these studies, a diffusional barrier for salicylate is strongly indicated since preformed salicylate renal clearance is less than glomerular filtration rate (GFR) but is greater than GFR when formed from salicyluric acid. 2.6. Flow Blood flow, Q, to an organ bears an inverse relationship to the transit time of a noneliminated substrate of volume, V, within the organ: By analogy, a prolonged transit of drug (reduced flow) through the liver promotes contact between drug and enzymes, thereby enhancing drug loss. A faster rate of flow hastens drug transit and reduces drug extraction. These effects of flow on drug extraction and clearance have been thoroughly explored. For a highly cleared substrate, clearance bears a direct relationship with flow, whereas for a poorly cleared compound, flow is devoid of effect on clearance (Wilkinson and Shand, 1975; Pang and Rowland, 1977a, 1977b; Wilkinson, 1987).

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 31

The effect of flow rate on conjugate(s) formation from sequential or parallel pathways, however, is seldom explored (Pang and Mulder, 1989). For sequential metabolism where is consecutively metabolized to one primary and secondary metabolites, as in tracer [14C]-phenacetin studies in the single-pass perfused liver, the effects of flow on the sulfation of [14C]-acetaminophen, a primary metabolite, were similar to those for its preformed counterpart, [3H]acetaminophen, given simultaneously with [14C]-phenacetin to the liver preparations. For reduced flows, a longer sojourn time promoted formation of the primary (indicated by preformed drug species) and secondary metabolites (indicated by the extent of sulfation of generated acetaminophen). Conversely, at increased flows, a shorter sojourn time diminished formation of the conjugates (Pang et al., 1988a). However, flow exerts differential effects on the anterior and posterior pathways in competitive, parallel pathways (Dawson et al., 1985). In the example of harmol, which was delivered at low concentrations (10 µM) and varying flow rates (8, 12, 16 ml min−1/liver) in the single-pass perfused liver, harmol sulfation rates paralleled with flow those for the extraction ratio, that is, a reduced flow brings about higher sulfation rates and higher extraction ratios; this condition leads to reduced recruitment of glucuronidation activities due to lack of substrate. At higher flow rates, the shorter sojourn time renders a reduced sulfation and allows more residual substrate to recruit glucuronidation activities. Thus two opposing effects are operative for the posterior pathway: residual substrate supply and the sojourn time of drug, interacting in opposite directions. For this reason, harmol glucuronidation rates (as a percent of rate in) were seemingly unchanged among all flow changes (Pang and Mulder, 1989). 2.7. Extrahepatic conjugation Many substrates are prone to Phase I and Phase II conjugation reactions in organs other than the liver. These notable extrahepatic organs include the lung (Cassidy and Houston, 1984), intestine (Gugler et al., 1975; Bock and Winne, 1975; Shibasaki et al., 1981), and kidney (Quebbemann and Anders, 1973; Koster and Noordhoek, 1983; Guder and Ross, 1984; Tremaine et al., 1984). The intestine, liver, and lung are arranged in a series anatomically, and concepts on competing anterior and posterior pathways may be extended to describe the kinetics of conjugation from these serially arranged organs (Pang, 1986). Intestinal conjugation relative to liver conjugation Salicylamide metabolism by the rat intestine and liver was recently investigated simultaneously in a vascularly-perfused, single-pass, rat intestine-liver

32 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 2.15. Influence of intestinal metabolism on the steady-state hepatic extraction ratios and total metabolism at 40 µM salicylamide as input concentrations in the singlepass rat intestine-liver preparation. The steady-state overall extraction ratio, intestinal and hepatic extraction ratios are depicted on the left. The total formation rate of metabolites and intestinal and hepatic formation rate of metabolites are expressed as a fraction of the total formation rate of metabolites (right). It was seen that despite the high SAM hepatic extraction ratios, intestinal metabolism modified the contributions of the liver in the overall formation of metabolites (from Xu et al., 1989b).

preparation, in which salicylamide was delivered systemically into the preparation. At low input concentrations of salicylamide, the steady-state hepatic extraction ratio for the intestine was 0·26 to form solely SAM-G whereas that for the liver was 0·99 to form predominantly SAM-S. Since both the intestine and liver contribute to the overall metabolism across these firstpass organs, the intestine, being the anteriorly-placed organ, however, regulates the substrate concentration in the portal vein entering the liver, hence affecting the rate of hepatic elimination. The presence of intestine drug extraction, albeit small, effectively diminished the component of hepatic removal in the overall rate of elimination despite the high hepatic extraction ratio of salicylamide (left panel, Figure 2.15); the hepatic component of the overall elimination was reduced to 74%. This is similar to the example on the competition between two pathways in the liver; the anterior pathway removes substrate and precludes downstream metabolite formation (Pang, 1986). This reduction in the hepatic component in total elimination due to intestinal removal also occurs in vivo. Another important consideration is that distinctly different metabolites have arisen through salicylamide metabolism. If saturability of the metabolic pathways also differ between the intestine and liver, as suggested by the present and previous data on salicylamide hepatic metabolism, the proportions of metabolites formed by both intestine and liver during first pass will change according to the orally administered dose (Pond et al., 1988). Normally, the effects of intestinal and liver metabolism are not

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 33

distinguished, and the overall metabolism is usually construed as hepatic in origin. As exemplified by the data on salicylamide, such an assumption would be grossly in error and undermine the importance of intestinal metabolism. Metabolites of intestinal origin may be misconstrued as hepatic in origin. For other compounds capable of both intestinal and liver conjugation, such as morphine (Iwamoto and Klaassen, 1977), 1-naphthol (Bock and Winne, 1975; Koster et al., 1985), 4-methylumbelliferone (Mulder et al., 1985) and others, it is expected that the intestine will regulate the contribution of the liver in the total first-pass effect for these substrates. Another occurrence worthy of consideration is the participation of the intestine and liver in enterohepatic circulation. Drug and/or metabolite (typically conjugates) excreted in bile enter the intestine for reabsorption. For polar conjugates, reabsorption is facilitated after hydrolysis, effected mostly by bacterial or brush-border enzymes. This enterohepatic cycle presents an effective mechanism of drug storage, either as itself or as metabolites, in the gall bladder/ intestine. Re-entry of the drug prolongs the duration of drug, and in turn, the metabolites in the body. When biliary excretion is significant, enterohepatic circulation can dramatically alter drug kinetics in the body. The kinetics of enterohepatic circulation have been explored to describe the spurious behaviour of concentration versus time profile (Dahlstrom and Paalzow, 1978; Steimer et al., 1982) of phenolic substrates such as morphine (Dahlstrom and Paalzow, 1978) and phenolphthalein (Colburn et al., 1979). These compounds form the respective glucuronide conjugates which undergo avid enterohepatic circulation. When a narrow therapeutic window exists, the extent of biliary excretion/ reabsorption is often found to correlate with toxicity (Duggan and Kwan, 1979). Kidney Renal metabolic abilities are being increasingly studied (Guder and Ross, 1984; Mohandas et al., 1984; Hjelle et al., 1986; Coughtrie et al., 1987; Peters and Jansen, 1988). Glycine conjugations of benzoic acid (Wan and Riegelman, 1972a), para-aminobenzoic acid (Wan and Riegelman, 1972b), salicylic acid (von Lehmann et al., 1973), and sulfation and glucuronidation of 1-naphthol, pnitrophenol (Tremaine et al., 1984; Hjelle et al., 1986), phenol (Quebbemann and Anders, 1973; Coughtrie et al., 1987), and 4methylumbelliferone (Peters and Jansen, 1988) have been reported among animal species, including man. Possible differences in renal handling of drug conjugates, circulating (formed from extrarenal organs in vivo) and nephrogenic (intrarenally generated following uptake and metabolism of precursor), due to the presence of transport barriers has already been discussed. A conjugate in circulation is subjected to all renal removal processes (Bekersky et al., 1980; Hekman et al., 1986; Redegeld

34 CONJUGATION REACTIONS IN DRUG METABOLISM

and Noordhoek, 1986): filtration, metabolism, reabsorption and secretion (basolateral and brush-border membranes). By contrast, nephrogenic conjugates synthesized within renal tubular cells from precursors are not filtered. These factors, transport and filtration, whether present or absent for arterial or nephrogenic conjugates, have led to differential rates of renal handling for the two species, as exemplified by enalaprilat, the esterolysis product of enalapril (de Lannoy et al., 1989). The excretion clearance of the conjugates monitored thus reflects those for the circulating as well as the nephrogenic conjugate. For this reason, the renal clearance so obtained (after precursor administration) can exceed that following administration of the conjugate. 2.8. Concluding remarks Some of the factors influencing conjugation kinetics have been described, mostly emphasizing events in the liver. The conceptual framework of drug conjugation as a distributed-in-space phenomenon in the liver has provided a better understanding of competing pathways. The conjugates formed are influenced by the enzymic constants and enzymic distributions for the pathways and the substrate concentration. Moreover, these concepts can be extended to examine the conjugation kinetics in two serially arranged conjugation organs: the intestine and liver. The kinetics of conjugation in the intestine and kidney, though less explored than the liver, deserve consideration. Due to its anterior placement to the liver, the intestine regulates the substrate entering the liver and the contribution of the latter in the overall first-pass metabolism. Different mechanisms of renal handling of conjugates occur with renal metabolism. Circulating conjugates are prone to filtration, secretion (basolateral and brush border membranes) and to undergo metabolism and reabsorption. Nephrogenic conjugates are not filtered nor subject to membrane (diffusional barrier or carrier-mediated transport) effects at the basolateral membrane. Abbreviations DCNP E GAM GAM-5G GAM-2S GAM-5S

2, 6-dichloro-4-nitrophenol Extraction ratio Gentisamide Gentisamide-5-glucuronide Gentisamide-2-sulfate Gentisamide-5-sulfate

KINETICS OF CONJUGATION REACTIONS IN ELIMINATING ORGANS 35

GSH 4-MU N R SAM SAM-G SAM-OH SAM-S

Glutathione 4-methylumbelliferone Normal direction perfusion Retrograde perfusion Salicylamide Salicylamide-glucuronide Hydroxylated salicylamide or gentisamide Salicylamide-sulfate References

Anundi, I.R., Kauffman, F.C., El-Mouelhi, M. and Thurman, R.G. (1986), Molecular Pharmacology, 29, 599–605. Baron, J., Redding, J.A. and Guengerich, F.P. (1978), Life Science, 23, 2627–32. Baron, J., Redick, R.A. and Guengerich, F.P. (1981), Journal of Biological Chemistry, 256, 15200–3. Barr, W. and Riegelman, S. (1970), Journal of Pharmaceutical Sciences, 59, 154–63. Bekersky, I., Colburn, W.A., Fishman, L. and Kaplan, S.A. (1980), Drug Metabolism and Disposition, 8, 319–24. Bekersky, I., Fishman, L., Kaplan, S.A. and Colburn, W.A. (1980), The Journal of Pharmacology and Experimental Therapeutics, 212, 309–12. Bock, K. and Winne, D. (1975), Biochemical Pharmacology, 24, 859–62. Brezenicka, E.A., Hazelton, G.A., Klaassen, C.D., (1987), Drug Metabolism and Disposition, 15, 133–5. Cassidy, M.K. and Houston, J.B. (1984), Drug Metabolism and Disposition, 12, 619–24. Chen, R. and Gillette, J.R. (1988), Drug Metabolism and Disposition, 16, 373–85. Chowdhury, J.R., Novikoff, P.M., Chowdhury, N.R. and Novikoff, A.B. (1985), Proceedings of the National Academy of Science, USA, 82, 2990–4. Colburn, W.A., Hirom, P.C., Parker, R.J. and Milburn, P. (1979), Drug Metabolism and Disposition, 7, 100–2. Conway, J.G., Kauffman, F.C., Ji, S. and Thurman, R.G. (1982), Molecular Pharmacology, 22, 509–16. Conway, J.G., Kauffman, F.C., Tsukada, T. and Thurman, R.G. (1984), Molecular Pharmacology, 25, 487–93. Conway, J.G., Kauffman, F.C., Tsukuda, T. and Thurman, R.G. (1988), Molecular Pharmacology, 33, 111–9. Coughtrie, M.W.H., Burchell, B. and Bend, J.R. (1987), Biochemical Pharmacology, 36, 245–51. Dahlstrom, B.E. and Paalzow, L.K. (1978), Journal of Pharmacokinetics and Biopharmaceutics, 6, 505–19. Dawson, J.R., Weitering, J.G., Mulder, G.J., Stillwell, R.N. and Pang, K.S. (1985), The Journal of Pharmacology and Experimental Therapeutics, 234, 691–7. de Baun, J.R., Smith, J.Y.R., Miller, E.C. and Miller, J.A. (1971), Science, 167, 184–6. de Lannoy, I.A.M. and Pang, K.S. (1987), Drug Metabolism and Disposition, 15, 51–8.

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de Lannoy, I.A.M., Nespeca R. and Pang, K.S., (1989). Journal of Pharmacology and Experimental Therapeutics, in press. Dollery, C.M., Davis, D.S., Connally, M.B. (1971), Annals of the New York Academy of Science, 79, 108–12. Duggan, D.E. and Kwan, K.C. (1979), Drug Metabolism Review, 9, 21–41. El Mouelhi, M. and Kauffman, F.C. (1986), Hepatology, 6, 450–6. Gillette, J.R. (1982), in Benet, L.Z., Levy, G. and Ferraiolo, B.L. (Eds.), Pharmacokinetics. A Modern View, pp. 235–52, New York: Plenum Press. Goresky, C.A., Bach, G.G. and Nadeau, B.E. (1973), Journal of Clinical Investigation, 52, 991–1009. Goresky, C.A., Gordon, E.R. and Bach, G.G. (1983), American Journal of Physiology, 244 (Gastrointestinal and Liver Physiology, 7), G198–214. Guder, W.G. and Ross, B.D. (1984), Kidney International, 26, 101–11. Gugler, R., Lain, P. and Azarnoff, D.L. (1975), The Journal of Pharmacology and Experimental Therapeutics, 195, 416–22. Gumucio, J.J. and Miller, D.L. (1978), Journal of Laboratory and Clinical Medicine, 91, 350–62. Hakim, R., Watrous, W.M. and Fujimoto, J.M. (1970), The Journal of Pharmacology and Experimental Therapeutics, 175, 749–62. Hekman, P., Russel, F.G.M., van Ginneken, C.A.M. (1986), Drug Metabolism and Disposition, 14, 370–1. Hjelle, J.T., Hazelton, G.A., Klaassen, C.D. and Hjelle, J.J. (1986), The Journal of Pharmacology and Experimental Therapeutics, 238, 150–6. Iwamoto, K. and Klaassen, C.D. (1977), The Journal of Pharmacology and Experimental Therapeutics, 200, 236–44. Jollow, D.J., Thorgeirsson, S.S., Potter, W.Z., Hashimoto and Mitchell, J.R. (1974), Pharmacology, 12, 251–71. Kashiwagi, T., Ji, S., Lemasters, J.J. and Thurman, R.G. (1982), Molecular Pharmacology, 21, 438–43. Kawabata, T.T., Guengerich, F.P. and Baron, J. (1981), Molecular Pharmacology, 20, 709–14. Knapp, S.A., Green, M.D., Tephly, T.R. and Baron, J. (1988), Molecular Pharmacology, 33, 14–21. Koike, M., Sugeno, K. and Hirata, M. (1981), Journal of Pharmaceutical Sciences, 70, 308–11. Koster, A.Sj. and Noordhoek, J. (1983), Biochemical Pharmacology, 32, 895–900. Koster, A.Sj., Frankhuijzen-Sierevogel and Noordhoek, J. (1985), Biochemical Pharmacology, 34, 3527–32. Koster, H., Halsema, I., Pang, K.S., Scholtens, E. and Mulder, G.J. (1982), Biochemical Pharmacology, 31, 3023–38. Meerman, J.H.N. and Mulder, G.J. (1981), Life Science, 28, 2361–65. Miyauchi, S., Sugiyama, S., Iga, T. and Hanano, M. (1988), Journal of Pharmaceutical Sciences, 77, 688–92. Mohandas, J., Marshall, J.J., Duggin, G.G., Horvath, J.S. and Tiller, D.J. (1984), Biochemical Pharmacology, 33, 1801–7. Morris, M.E. and Pang, K.S. (1987), Journal of Pharmacokinetics and Biopharmaceutics, 15, 473–96.

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Morris, M.E., Yuen, V. and Pang, K.S. (1988b), Journal of Pharmacokinetics and Biopharmaceutics, 16, 633–56. Morris, M.E., Yuen, V., Tang, B.K. and Pang, K.S. (1988a), The Journal of Pharmacology and Experimental Therapeutics, 245, 614–52. Mulder, G.J. and Hagedoorn, A.H. (1974), Biochemical Pharmacology, 23, 2101–9. Mulder, G.J., Brouwer, S., Weitering, J.G., Scholtens, E. and Pang, K.S. (1985), Biochemical Pharmacology, 34, 1325–9. Pang, K.S. (1986), Journal of Clinical Pharmacology, 26, 580–2. Pang, K.S. (1989), in Bend, J.R., Hodgson, E., Philpot, R.M. (eds.), Reviews in Biochemical Toxicology, 10, 187–263. Pang, K.S. and Rowland, M. (1977a), Journal of Pharmacokinetics and Biopharmaceutics, 5, 625–53. Pang, K.S. and Rowland, M. (1977b), Journal of Pharmacokinetics and Biopharmaceutics, 5, 655–80. Pang, K.S. and Gillette, J.R. (1978), The Journal of Pharmacology and Experimental Therapeutics, 207, 178–94. Pang, K.S. and Gillette, J.R. (1979), Journal of Pharmacokinetics and Biopharmaceutics, 7, 275–90. Pang, K.S. and Terrell, J.A. (1981), Journal of Pharmacology and Experimental Therapeutics, 216, 339–46. Pang, K.S. and Xu, X. (1988), in Welling, P.G. and Tse, F.L.-S. (eds.), Pharmacokinetics: Regulatory-Industrial-Academic Perspectives, pp. 383–447, New York: Marcel Dekker Inc. Pang, K.S. and Mulder, G.J. (1989), Drug Metabolism and Disposition, in press. Pang, K.S., Xu, X., Morris, M.E., and Yuen, V. (1987), Federation Proceedings, 46, 2439–41. Pang, K.S., Koster, H., Halsema, I.C.M., Scholtens, E. and Mulder, G.J. (1981), Journal of Pharmacology and Experimental Therapeutics, 219, 134–40. Pang, K.S., Cherry, W.F., Accaputo, J., Schwab, A.J. and Goresky, C.A. (1988b), The Journal of Pharmacology and Experimental Therapeutics, 247, 690–700. Pang, K.S., Koster, H., Halsema, I.C.M., Scholtens, E., Mulder, G.J. and Stillwell, R.N. (1983), The Journal of Pharmacology and Experimental Therapeutics, 224, 647–53. Pang, K.S., Lee, W.-F., Cherry, W.F., Yuen, V., Accaputo, J., Fayz, S., Schwab, A.J. and Goresky, C.A. (1988a), Journal of Pharmacokinetics and Biopharmaceutics, 16, 595–632. Pannather, A., Jenner, P., Testa, B. and Etter, J.C. (1978), Drug Metabolism Reviews, 8, 319–43. Peters, W.H.M. and Jansen, P.L.M. (1988), Biochemical Pharmacology, 37, 564–7. Pinkus, L.M., Ketley, J.N., and Jakoby, W.B. (1977), Biochemical Pharmacology, 26, 2359–63. Pond, S., Waschek, J., Mahachai, V., Fielding, R., Effeney, D. and Tozer, T. (1988), The Journal of Pharmacology and Experimental Therapeutics, 246, 291–3. Potter, W.R., Branchflower, R.V., and Trager, W.F. (1977), Biochemical Pharmacology, 26, 549–50. Quebbemann, A.J. and Anders, M.W. (1973), The Journal of Pharmacology and Experimental Therapeutics, 184, 695–708. Rappaport, A.M. (1958), Anatomical Records, 130, 673–86.

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Rappaport, A.M. Borowy, Z.J., Lougheed, W.M. and Lotto, W.N., (1954), Anatomical Records, 119, 11–34. Redegeld, F.A.M. and Noordhoek, J. (1986), Drug Metabolism and Disposition, 14, 622–4. Redick, J.A., Jakoby, W.B. and Baron, J. (1982), Journal of Biological Chemistry, 257, 15200–3. Rennick, B. and Quebbemann, A. (1970), American Journal of Physiology, 218, 1307–12. Shibasaki, J., Konishi, R., Koike, M., Imamura, A. and Sueyasu, M. (1981), Journal of Pharmacobiodynamics, 4, 91–100. Silverman, M. and Goresky, C.A. (1965), Biophysics Journal, 5, 487–509. Smith, B.R., van Anda, J., Fouts, J.R. and Bend, J.R. (1983), The Journal of Pharmacology and Experimental Therapeutics, 227, 491. St-Pierre, M.V., Lee, W.F., Schwab, A.J., Goresky, C.A. and Pang, K.S. (1989), Hepatology, 9, 285–96. Steele, J.W., Yagen, B., Hernandez, O., Cox, R.H., Smith, B.R. and Bend,J. R. (1981), The Journal of Pharmacology and Experimental Therapeutics, 219, 35. Steimer, J.-L., Plusquellec, Y., Guillaume, A. and Boisvieux, J.-F. (1982), Journal of Pharmaceutical Sciences, 71, 297–302. Trainer, C. and Silverman, M. (1982), American Journal of Physiology, 242, (Renal Fluid Electrolyte Physiology, 11), F436–46. Tremaine, L.M., Diamond, G.L. and Quebbemann, A.J. (1984), Biochemical Pharmacology, 33, 419–27. Ullrich, D., Fisher, G., Katz, N. and Bock, K.W. (1984), Chemical and Biological Interactions, 48, 181–90. von Lehmann, B., Wan, S.H., Riegelman, S. and Becker, C. (1973), Journal of Pharmaceutical Sciences, 62, 1483–90. Wan, S.H. and Riegelman, S. (1972a), Journal of Pharmaceutical Sciences, 61, 1278–94. Wan, S.H. and Riegelman, S. (1972b), Journal of Pharmaceutical Sciences, 61, 1288–92. Watari, N., Iwai, M. and Kaneniwa, N. (1983), Journal of Pharmacokinetics and Biopharmaceutics, 11, 245–72. Watrous, W.M., Amy, D.G. and Fujimoto, J.M. (1970), The Journal of Pharmacology and Experimental Therapeutics, 172, 224–9. Wilkinson, G.R. (1987), Pharmacological Reviews, 39, 1–47. Wilkinson, G.R. and Shand, D.G. (1975), Clinical Pharmacology and Therapeutics, 18, 377–90. Winkler, K., Keiding, S. and Tygstrup, N. (1973), in Paumgartner, G. and Preisig, R. (eds.), The Liver: Quantitative Aspects of Structure and Function, pp. 144–55, Basel: Karger. Xu, X. and Pang, K.S. (1989a) Journal of Pharmacokinetics and Biopharmaceutics, in press. Xu, X., Hirayama, H. and Pang, K.S. (1989a), Drug Metabolism and Disposition, in press. Xu, X., Tang, B.K. and Pang, K.S. (1989b), submitted.

Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 3 Competition between conjugations for the same substrate Gerard J.Mulder Division of Toxicology, Center for Bio-Pharmaceutical Sciences, University of Leiden, 2300 RA Leiden, The Netherlands

3.1.

INTRODUCTION

40

3.2.

ACCEPTOR GROUPS FOR WHICH COMPETITION OCCURS FREQUENTLY

41

Phenolic group

41

Carboxylic group

41

Aromatic amino group

42

Hydroxylamines and hydroxamic acids

42

Epoxides

42

3.3.

RACEMIC SUBSTRATES

42

3.4.

ENDOGENOUS FACTORS

43

3.5.

Kinetic parameters of the enzymes

43

Availability of co-substrates

44

Localization of the enzymes and lipid solubility of the substrate

45

Species

46

Sex differences

46

EXOGENOUS INFLUENCES

46

Diet

46

Enzyme induction

47

Route of administration

47

40 CONJUGATION REACTIONS IN DRUG METABOLISM

ABBREVIATIONS

91

REFERENCES

47 3.1. Introduction

When xenobiotics enter the body they can be metabolized by enzymes that (happen to) have sufficient affinity and activity towards these compounds. Both Phase I (oxidative, reductive or hydrolytic), and Phase II (conjugative) reactions may compete for the same substrate. Even for a simple structure like paminosalicylic acid, several pathways compete for biotransformation (Figure 3.1). For example, enzymes may compete for conversion of different groups in the molecule, while at the same time there is also competition for conversion of the same group. The outcome usually is a whole array of metabolites that are excreted mainly in urine or faeces, in a species- and dosedependent pattern. Further metabolism of a primary metabolite (for instance, a glutathione conjugate which is converted in several steps to a mercapturate) may lead to additional metabolites. Several factors determine the outcome of this interplay between enzymes, substrates, co-substrates, the organs in which these processes take place, and the pharmacokinetics of distribution and elimination of the substrates. Obviously the chemical structure of the substrate determines all subsequent events. In this chapter, the main emphasis will be on competition between conjugations for the same acceptor substrate, often for the same acceptor group. The same principles apply, of course, to competitions in which Phase I reactions are involved. In these cases it is primarily a different region of the substrate molecule from that in conjugation that is involved. However, in the case of the aromatic amine group, N-oxidation is a major pathway competing with conjugation, e.g. acetylation or glucuronidation. In addition, Phase I metabolism often creates a new acceptor group for conjugation, so that it further complicates the total pattern of conjugation of a given xenobiotic substrate.

Figure 3.1. Metabolic pathways of p-aminosalicylic acid.

COMPETITION BETWEEN CONJUGATIONS FOR THE SAME SUBSTRATE 41

3.2. Acceptor groups for which competition occurs frequently Phenolic group

Major conjugation reactions at the phenolic hydroxyl group are sulfation, glucuronidation and methylation; occasionally acetylation or phosphate conjugation may occur. In certain species, such as insects, other glycosylation reactions may be important (see Mulder, 1982, for review). Extensive methylation is usually restricted to catechols, such as adrenaline or dopamine. Extensive investigations on the kinetics of these competitive situations in vivo have been performed only for the balance between sulfation and glucuronidation of phenols (see Chapters 4 and 5). The results suggest that, in general, sulfation is the process with the higher affinity but with a lower capacity than glucuronidation. This seems due primarily to the higher affinity of the sulfotransferases than to the UDP-glucuronosyltransferases for the same substrate, but in addition the co-substrate for sulfation becomes more readily depleted than that for glucuronidation. Carboxylic group The major competing reactions are glucuronidation (and, more rarely, other glycosylations with, e.g. xylose or glucose) and activation to the CoA derivative (see Caldwell, 1982a, for a review). The latter can subsequently be transferred to either an amino acid, such as glycine or taurine, or to a less common acceptor, such as glycerol. In arylacetic acid derivatives steric hindrance at the a and β carbon atom plays an important role; amino acid conjugation (i.e. CoA activation) is much more sensitive to such hindrance than glucuronidation (Chapter 10). Amino acid conjugation usually is the reaction with the lower Km and, therefore, is more important at low doses while at high doses the higher capacity glucuronidation takes over. Both conjugations lead to relatively labile conjugates. The acyl-CoA ester reacts with various acceptor groups to form stable conjugates, for instance with amino acids. The xenobiotic may even become ‘hidden’ due to incorporation into triglycerides. Another special feature is that a racemic mixture of a 2arylpropionic acid derivative may be inverted stereo-selectively to yield only the S-enantiomer as occurs with ibuprofen (Chapter 10). The ester glucuronides are relatively reactive as shown by the fact that they spontaneously bind covalently to nucleophiles, e.g. glutathione (Faed, 1984).

42 CONJUGATION REACTIONS IN DRUG METABOLISM

Furthermore, the acyl group may shift from the original C1 position to C2, C3 or C4 positions on the sugar ring. The resulting isomers are not hydrolyzed by βglucuronidase, thus giving the appearance of a new, unknown conjugate, while in fact it is derived from a normal ester glucuronide. At a slightly alkaline urine pH, ester glucuronides may not be stable and large amounts of the aglycone may be found in urine, giving the wrong impression that the aglycone has been excreted unchanged (see Chapter 4). Aromatic amino group This group may be converted by acetylation, methylation, sulfation and glucuronidation, while N-oxidation is also an important pathway especially for aromatic amines (Caldwell, 1982b). Moreover, double conjugates may be found after subsequent acetylation and N-hydroxylation (Chapter 5). Little systemic work has been done on structure-activity relationships or pharmacokinetics for these competing pathways. Besides analytical problems and the complex pattern of metabolites that is often found for aromatic amines, the fact that several metabolites are very labile complicates studies in this area (see Chapters 5 and 6). Hydroxylamines and hydroxamic acids For hydroxylamines the options are conjugation at the hydroxyl group or at the nitrogen atom (sulfation, glucuronidation, acetylation). For hydroxamic acids conjugation is only possible at the N-hydroxy group. Many of the conjugates of these types of compounds are extremely labile and may yield reactive intermediates that can bind covalently to nucleophilic groups (see Chapter 5). Epoxides The two metabolic reactions for epoxides are glutathione conjugation and hydration (hydrolysis) by epoxide hydrolase. Both enzymes have forms that are located in cytosol and the endoplasmic reticulum. While the co-substrate for glutathione conjugation can be depleted, this is obviously not the case for the epoxide hydrolase reaction. The metabolism of epoxides has been reviewed by Hernandez and Bend (1982). 3.3. Racemic substrates Sometimes the substrate studied is a racemic mixture of enantiomers. Increasingly, attention is being paid to the fact that the drug-metabolizing

COMPETITION BETWEEN CONJUGATIONS FOR THE SAME SUBSTRATE 43

enzymes may show pronounced stereo-specificity towards the separate enantiomers of a racemic drug (or other types of stereoisomers). In fact, these have to be considered as separate substrates; the administration of a racemic mixture implies the addition of two potentially competing substrates. For the conjugation reactions some data are available. For example, the glutathione conjugation of α-bromoisovalerylurea (Figure 3.2) shows marked stereoselectivity. The separate enantiomers have a very different affinity and maximal velocity of conversion for the various glutathione transferase isoenzymes (Te Koppele et al., 1988). This results in pronounced differences between the pharmacokinetics of the enantiomers in the rat in vivo. Moreover, in a racemic mixture the conjugation of the enantiomer with the lower affinity and velocity may be inhibited by the other enantiomer. An alternative pathway for αbromoisovalerylurea metabolism, amidase-catalyzed hydrolysis, has its own stereo-selectivity (Te Koppele et al., 1988). 3.4. Endogenous factors Kinetic parameters of the enzymes When two enzymes compete for the same substrate, the rate at which the product of each is formed is determined by the Km and Vmax values of the respective enzymes, if all other factors are optimal. The ratio at which the metabolites will be formed is determined by the concentration of the substrate in relation to the two Km and two Vmax values. At saturation of both enzymes the ratio of product formation will be solely determined by the respective Vmax values. In general, sulfotransferases have a lower Km for their substrates than the UDP-glucuronosyltransferases, while the Vmax for glucuronidation is higher than for sulfation. This implies that at increasing concentration (or dose, in vivo) the relative importance of sulfation decreases as is observed in experiments in vivo and in perfusions or isolated cells. The same applies to amino acid conjugation and glucuronidation of carboxylic acids. Unfortunately, very few Km and Vmax values of two competing transferases for their shared xenobiotic substrates are available. One of the reasons is that too few ‘logical’ series of substrates have been used to allow a quantitative structure-activity relationship (QSAR) analysis. Moreover, in general they have been determined for only one conjugation, often with mixtures of isoenzymes (e.g. a microsomal or cytosolic fraction).

44 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 3.2. Metabolism of a-bromoisovalerylurea by glutathione conjugation and amidase-catalyzed hydrolysis.

Availability of co-substrates Rapid conjugation requires a sufficient supply of co-substrates, which under certain conditions, may become rate-limiting. For instance, the co-substrate may be depleted by a high dose of the compound to be conjugated; this can occur for most of the conjugations (except, of course, epoxide hydrolase). Alternatively, the biosynthesis of the co-substrate may be affected by e.g. disease, or may be deficient for genetic reasons. Thus, in brachymorphic mice, the biosynthesis of 3′phosphoadenosyl-5′-phosphosulfate (PAPS) is deficient because there is a genetic defect in the activating enzymes (Chapter 5). Of special importance is the effect of nutrition, especially for the sulfur-containing co-factors glutathione, PAPS and taurine, and more indirectly, S-adenosylmethionine. The reason is that their synthesis depends on the supply of L-cysteine (eventually generated by transsulfurylation from L-methionine) in the diet. A decrease in the availability of a co-substrate will result in either a decreased rate of conjugation when the concentration of the co-substrate decreases below saturating levels, or a total block of that conjugation when the cell runs out of the co-substrate altogether. The latter will occur when the precursor of the cosubstrate is not available or its synthesis inhibited. As long as a precursor is present, resynthesis may provide the co-substrate (albeit at a low level). When one co-substrate is depleted, the competing reactions may take over. For instance, when inorganic sulfate is depleted and, therefore, sulfation has become impossible, usually glucuronidation compensates by an increased extent

COMPETITION BETWEEN CONJUGATIONS FOR THE SAME SUBSTRATE 45

of that conjugation. The efficiency of such compensation, obviously, is limited by the degree of saturation of a competing pathway. Localization of the enzymes and lipid solubility of the substrate In certain cases the lipid solubility of the substrate has been shown to play a role in the competition between conjugations. Thus, UDP-glucuronosyl transferase activity towards its substrates increases as lipid solubility increases (Chapter 4). As a result a larger percentage of the dose will be converted by glucuronidation, as observed in hepatocyte incubations for a number of nitrophenols that are both glucuronidated and sulfated (Fry and Patterson, 1985). Moreover, binding of substrates to protein or the membrane phase will affect glucuronidation of low water-soluble substrates (see e.g. Whitmer et al., 1984). This may also be favoured by the localization of this transferase in the endoplasmic reticular membrane, possibly with the active site directed towards the lumen of the endoplasmic reticulum (see Chapter 4). Clearly, however, in the competition between sulfation and glucuronidation, lipid solubility is not the main factor, because at low concentration of the substrate, sulfation is the preferred route, even for highly lipid-soluble substrates. Presumably, therefore, kinetic factors are more important than lipid solubility. Moreover, the cell interior is a highly complex system with, probably, rapid cytoplasmic streaming (Wheatley, 1985). As yet it is unknown whether the differential localization of glutathione transferases in the cytosol or in the membrane phase of the endoplasmic reticulum has consequences for the substrate specificity in terms of lipid solubility for the two subsets of the transferase. The various conjugating enzymes are located in different structures or compartments of the cell, such as mitochondria, microbodies, endoplasmic reticulum or the cytosol. The consequences of this localization for competition between two pathways has not been investigated. The distances within the cell between enzyme active sites located, for instance, in the endoplasmatic reticulum or in the cytosol are very small and can rapidly be covered by diffusion. When the distribution of competing enzymes within an organ is nonhomogeneous (for instance in the periportal and pericentral areas in the liver), this will complicate the kinetics of competition (see Chapter 2). Besides accumulation in or binding to membranes, protein binding in the cell may also affect the free concentration of a substrate for conjugation. This has been discussed by Tipping and Ketterer (1981).

46 CONJUGATION REACTIONS IN DRUG METABOLISM

Species Obviously, the outcome of competition will depend on the species chosen for the studies. A number of reasons can account for this. The activities of the transferases involved may be very different relative to each other, and they may show different isoenzyme composition or have different Km values. The cosubstrate supply may vary widely between species. Further, species-dependent pharmacokinetic differences, for instance in distribution in fat tissues or other organs, may affect the substrate concentration available to the conjugating enzymes. Certain conjugations may be almost completely lacking in some species. Sex differences Relatively little is known about sex differences for the conjugations. In certain cases a pronounced sex difference will give rise to a pronounced difference in biological effects. For instance, the male rat has a much higher sulfation activity towards N-hydroxy-2-acetylaminofluorene than the female, which is the reason why only the male rat shows a high level of hepatomas after exposure to this compound (Chapter 5). 3.5. Exogenous influences Diet The diet may affect the outcome of competition between conjugations at various levels; it may be adequate or restricted, its composition (fat, protein, carbohydrate) may be abnormal, specific deficiencies (minerals, vitamins) may occur, or food contaminants may be present. As a result of food preparation procedures, it may contain inducers of drug metabolism (for instance due to charcoal broiling); food preservatives may have effects, and natural components in certain foods may be inhibitors or inducers of drug metabolism. The activities of the transferases and of the enzymes which synthesize the co-substrate may be affected by the diet. The diet may also be deficient in a precursor for a co-substrate. This is especially important for sulfur-containing co-substrates like glutathione, PAPS and taurine. Since glutathione, especially, is an essential detoxifying factor in many cases, this may severely affect the ability of a organism to defend itself against potentially toxic metabolites such as reactive intermediates. If taurine supply becomes limiting, amino acid conjugation may shift to glycine conjugation.

COMPETITION BETWEEN CONJUGATIONS FOR THE SAME SUBSTRATE 47

Enzyme induction Several conjugating enzymes are readily induced, notably the glutathione transferases and UDP-glucuronosyltransferases. To date no inducers of sulfotransferases, methyltransferases or acetyltransferases have been identified. Clearly, induction of glutathione transferases will increase the flux through this pathway as compared to that through epoxide hydrolase. Smoking results in the induction of certain pathways, although little data on conjugation is available. Route of administration When a substrate is administered intravenously, the highest blood concentration can be reached, certainly when the dose is injected as a rapid bolus injection. Any other route of administration of the same dose will yield lower blood concentrations and therefore lower tissue concentrations. When the competition is concentration-dependent, the reaction with the lower Km will be favoured when the dose is given by routes other than i.v. (Table 3.1). Furthermore, administration by certain routes may show first-pass effects. After oral administration first-pass conjugation in the gut mucosa and the liver is possible. For that reason the metabolite pattern of the same compound administered orally or i.v. may be very different. Intraperitoneal administration results in first-pass metabolism in the liver (Lukas et al., 1971). Table 3.1. Effect of route of administration on phenol conjugation in the hen. A dose of 226 µmol kg-1 was administered in all cases. Recovery of the dose in urine was 45–57% for the groups (Capel et al., 1974).

References Caldwell, J. (1982a), in Jakoby, W.B., Bend, J.R. and Caldwell, J. (Eds.), Metabolic Basis of Detoxication: Metabolism of Functional Groups, pp. 271–91, London: Academic Press. Caldwell, J. (1982b), in Jakoby, W.B., Bend, J.R. and Caldwell, J. (Eds.), Metabolic Basis of Detoxication: Metabolism of Functional Groups, pp. 291–306, London: Academic Press. Capel, I.D., Milburn, P. and Williams, R.T. (1974), Biochemical Society Transaction, 2, 875–7. Faed, E.M. (1984), Drug Metabolism Reviews, 15, 1213–40. Fry, J.R., and Patterson, P. (1985), British Journal of Pharmacology, 84, 133P.

48 CONJUGATION REACTIONS IN DRUG METABOLISM

Hernandez, O. and Bend, J.R. (1982), in Jakoby, W.B., Bend, J.R. and Caldwell, J. (Eds.), Metabolic Basis of Detoxication: Metabolism of Functional Groups, pp. 207–28, London: Academic Press. Lukas, G., Brindle, S.D. and Greengard, P. (1971), Journal of Pharmacology and Experimental Therapeutics, 178, 562–6. Mulder, G.J. (1982), in Jakoby, W.B., Bend, J.R. and Caldwell, J. (Eds.), Metabolic Basis of Detoxication: Metabolism of Functional Groups, pp. 247–69, London: Academic Press. Mulder, G.J. (1986), Chemico-Biological Interactions, 57, 1–15. Te Koppele, J.M., deLannoy, I.A.M., Pang, K.S. and Mulder, G.J. (1987), Journal of Pharmacology and Experimental Therapeutics, 243, 349–55. Te Koppele, J.M., Coles, B., Ketterer, B. and Mulder, G.J. (1988), Biochemical Journal, 252, 137–42. Tipping, E. and Ketterer, B. (1981), Biochemical Journal, 195, 441–52. Wheatley, D.N. (1985), Life Sciences, 36, 299–307. Whitmer, D.I., Ziorys, J.C. and Gollan, J.L. (1984), Journal of Biological Chemistry, 259, 11969–75.

Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 4 Glucuronidation Gerard J.Mulder1, Michael W.H.Coughtrie2 and Brian Burchell2 1

Division of Toxicology, Center for Bio-Pharmaceutical Sciences, University of Leiden, 2300 RA Leiden, The Netherlands 2

Department of Biochemical Medicine, Ninewells Hospital and Medical School, Dundee, DD1 9SY, Scotland, UK

4.1.

INTRODUCTION

50

4.2.

UDPGA SYNTHESIS AND AVAILABILITY

51

4.3.

SUBSTRATES AND STEREOSELECTIVITY

53

4.4.

PROPERTIES OF UDP-GLUCURONOSYLTRANSFERASES

57

The effect of administered xenobiotics: induction of UDPGT

57

Ontogeny

58

Tissue distribution

59

Inherited deficiences of glucuronidation

59

Purified UDPGTs

60

Cloning and expression of UDPGT cDNAs

62

Substrate specificity of UDPGTs and enzyme mechanism

63

Inhibition of UDPGT

64

4.5.

GLUCURONIDATION IN PERFUSED ORGANS AND ISOLATED CELLS

68

Latency of UDPGT: physiological or artefact

69

Liver perfusion

70

Isolated hepatocytes

75

Intestinal glucuronidation in vitro and in vivo

77

Kidney

78

50 CONJUGATION REACTIONS IN DRUG METABOLISM

Other organs 4.6.

78

GLUCURONIDATION IN VIVO

79

Pharmacokinetics and excretion

79

Extrahepatic glucuronidation

82

Effect of internal or external factors

83

4.7.

INHIBITION OF GLUCURONIDATION IN INTACT CELLS AND IN VIVO

85

4.8.

CONJUGATION AND BIOLOGICAL ACTIVITY

86

4.9.

PRACTICAL CONSIDERATIONS

88

ABBREVIATIONS

90

REFERENCES

90 4.1. Introduction

Between 1855 and 1880 it was established that certain compounds, when fed to various animal species, were excreted in urine as glucuronide conjugates; these substances (or their oxidation products) were conjugated with D-glucuronic acid. In 1955 Storey and Dutton reported the structure of the co-factor required for glucuronidation, uridine 5′-diphosphoglucuronic acid (UDPGA; Figure 4.1). Purification of the transferase catalyzing glucuronidation proved difficult because the enzyme was found to be anchored in the membrane of the endoplasmic reticulum. It had to be solubilized before purification, and its activity towards most aglycones turned out to be dependent on the presence of phospholipids. The properties of the purified enzymes, therefore, may show differences from those of the native enzymes in their physological localization. The enzyme activity is latent when it is measured in freshly prepared microsomal fractions; the capacity for glucuronidation is not fully expressed. The enzymes can be activated by various procedures which disrupt the membrane environment of the vesicles in which the enzymes are located after homogenization. There are several forms of the transferase, with different substrate specificity and under different regulatory control. For instance, they can be differentially induced by phenobarbital (PB) and 3-methylcholanthrene (3MC) pretreatment. The co-substrate required for the reaction is synthesized in the cytosol from the precursors glucose-1-phosphate and UTP; the UDP-glucose (UDPG)

GLUCURONIDATION 51

Figure 4.1. Structure of UDPGA.

product is subsequently oxidized by UDPG dehydrogenase to UDPGA. Since UDPGA is also required for the biosynthesis of glycosaminoglycans, it is synthesized in a wide variety of cell types and organs. High activities of xenobiotic glucuronidation occur in the liver, the intestinal mucosa and the kidney; most other organs or tissues possess glucuronidation activity, although it may be very low. UDPGA has to be synthesized in the cell where glucuronidation takes place since it is very unlikely that it will be taken up by intact cells, although this has not been rigorously tested. Glucuronides are excreted in urine and bile. After biliary excretion, the conjugate may be hydrolyzed in the gut by (bacterial) β-glucuronidases. Glucuronides in the blood may also be hydrolyzed after uptake by various tissues such as the liver. In general glucuronides lack the biological effects of the precursor substrate (but see Section 4.8). Very often glucuronidation has been studied with substrates that are also sulfated. For those substrates, the possibility exists that changes in glucuronidation may in fact reflect primary effects on sulfation. For instance, inhibition of sulfation in vivo may result in increased glucuronidation, which could also be misinterpreted as an activation of glucuronidation. Complete, in-depth reviews on glucuronidation have appeared (Dutton, 1980; Burchell, 1981) as well as proceedings of workshops on glucuronidation (Matern et al., 1985; Siest et al., 1988). 4.2. UDPGA synthesis and availability UDPGA is a highly water-soluble, stable compound. It is formed inside the cell in a two-step reaction from glucose-1-phosphate and UTP. The first step is

52 CONJUGATION REACTIONS IN DRUG METABOLISM

catalyzed by UDPG pyrophosphorylase (EC 2.7.7.9) to form UDPG. Subsequently, UDPG is oxidized at C6 to UDPGA, which requires a fourelectron oxidation in two steps. The activity of UDPG dehydrogenase (EC 1.1.1. 22) is controlled by the NAD+/NADH ratio (Siviswami et al., 1972), and this presumably is the reason why ethanol at high concentration (10−20 mM, or ca. 1 µl ml−1) will inhibit glucuronidation in isolated hepatocytes. Ethanol then shifts the NAD+/NADH ratio so that UDPG dehydrogenase is inhibited; UDPGA is decreased, but UDPG remains unchanged (Aw and Jones, 1983). The alcohol dehydrogenase inhibitor, 4-methylpyrazole, prevents this ethanol effect (Bodd et al., 1986).

Adenosine similarly decreases UDPGA in isolated hepatocytes by decreasing the NAD+/NADH ratio (Shipley and Weiner, 1987). Ethanol does not affect glucuronidation in the rat in vivo unless the animals are fasted for 24 h prior to ethanol administration; then UDPGA and glucuronidation in the liver are decreased (Moldeus et al., 1980; Minnigh and Zemaitis, 1982). Chronic ethanol treatment of rats causes a decrease in maximum glucuronidation rates in the perfused rat liver and in vivo, to which a decreased UDPGA level may be contributing (Vendemiale et al., 1984; Reinke et al., 1986; Hong et al., 1987). However, in isolated hepatocytes from rats treated in this manner, glucuronidation was increased (Moldeus et al., 1980). The concentration of UDPGA in liver is usually higher than in any other tissue. Moreover, turnover rates of UDPGA can be very high as studied in the perfused rat liver: while ca. 0.2 µmol UDPGA g- liver is present, glucuronidation rates of 0.1 µmol g-1 liver min-1 could be sustained for more than 60 min (Pang et al., 1981). Similar high fluxes also occur in vivo (Price and Jollow, 1984; Zhivkov and Tosheva, 1986). Under normal conditions the precursors for UDPGA will be readily available from carbohydrate reserves and the UTP pool. The uridine pool can be depleted by a high dose of D-galactosamine. This severely impairs glucuronidation (Moldeus et al., 1979; Singh and Schwarz, 1981; Watkins and Klaassen, 1982a, 1983; Gregus et al., 1983) possibly almost exclusively in the liver (Gregus et al., 1988a). In fasted rats the carbohydrate reserve may be too low, so that glucose added to the liver perfusion or isolated hepatocytes enhances the glucuronidation rate (Schwarz, 1980; Reinke et al., 1981). Compounds that impair the energy state of the hepatocytes, such as potassium cyanide, fructose or 2, 4-dinitrophenol, are inhibitory (Reinke et al., 1981; Dills and Klaassen, 1986; Dills et al., 1987). In isolated hepatocytes, glucuronidation is dependent on sufficient oxygenation; under hypoxic conditions the ATP/ADP ratio and glucuronidation decrease (Aw and Jones, 1982). Fasting will decrease UDPGA content and biosynthesis in the liver as well

GLUCURONIDATION 53

as rates of glucuronidation (Minnigh and Zemaitis, 1982; Price and Jollow, 1988). When the demand for UDPGA becomes too high at high substrate dose, the concentration of UDPGA, for instance in rat liver, will decrease (Hjelle et al., 1985b; Hjelle, 1986; Howell et al., 1986). Doses required were 1 to 4 mmol kg-1 i.p., and liver UDPGA decreased within 15 min while often UDPG was not affected. Hjelle (1986) suggested that the depletion of UDPGA by high doses of paracetamol was ultimately due to inhibition of UDPG dehydrogenase so that conversion of UDPG to UDPGA became rate-limiting in glucuronidation. Also other drugs that are not direct substrates for glucuronidation may deplete UDPGA in the rat liver, such as PB (Douidar and Ahmed, 1987), aflatoxin and a number of anaesthetics like diethyl ether or halothane (Eriksson and Stråth, 1981; Watkins and Klaassen, 1983). The diethyl ether effect was mainly limited to the liver (Watkins and Klaassen, 1983) and was rapidly reversible after a 10 min exposure; after 60 min UDPGA had almost returned to control levels. In isolated hepatocytes from ether-anaesthetized rats, no inhibitory effect was observed (Shipley and Weiner, 1985). However, 30 mM diethyl ether in the incubation medium did inhibit conjugation of paracetamol (Aune et al., 1984). The in vivo effects of diethyl ether exposure may be prolonged, lasting for several hours after administration (To and Wells, 1986). An age-related decrease in hepatic UDPGA has been observed in the rat (Borghoff et al., 1988). 4.3. Substrates and stereoselectivity The mechanism of glucuronidation is a SN2 reaction, the acceptor group of the substrate attacking the C1 of the pyranose ring to which UDP is attached in an α-glycosidic bond (Figure 4.1); the resulting glucuronide has the β-glycosidic configuration. The attacking group should have sufficient nucleophilic character for a high rate of glucuronidation. A wide variety of groups fulfil this requirement, such as phenols, carboxylic acids, thiophenols, alcohols, aromatic amines, hydroxylamines or hydroxamic acids. Even carbon atoms can attack C1 of the glucuronic acid ring if they are sufficiently nucleophilic, such as when carbon atoms are adjacent to two carbonyl groups, or ethynylic carbon atoms (Abolin et al., 1980). The reactivity will depend on the structure, both due to electronic and steric factors. Examples of the various reactions are given in Table 4.1. In some cases more than one group is available for glucuronidation, like in hyodeoxycholate, from which 3 glucuronides may be formed (Zimniak et al., 1988). A second requirement for a high rate of glucuronidation is sufficient lipid solubility of the substrate. These parameters correlate quite well, but bulky

54 CONJUGATION REACTIONS IN DRUG METABOLISM

groups in the ortho-position cause deviations (Mulder and van Doorn, 1975; Illing, 1980; Schaeffer et al., 1981). The thickness or the bulkiness of the substrate molecule is an important factor in determining which UDPGT form will glucuronidate a given substrate; again ortho-substituents may change a substrate’s classification (Okulicz-Kozaryn et al.,1981). Also in isolated hepatocytes a correlation between lipid solubility and glucuronidation rate was apparent (Fry and Paterson, 1985). Only a few series of substrates (mainly phenols) have been tested with microsomal preparations containing a mixture of UDPGT forms as the source of UDPGT. While occasionally Km and Vmax values were determined, in most cases only the glucuronidation rate at one substrate concentration was measured (Mulder and van Doorn, 1975; Mulder and Meerman, 1978; Bansal and Gessner, 1980; Holloway and Zierhut, 1981; Matern et al., 1983; Colin-Neiger et al., 1984; Coughtrie et al., 1986; Radominska-Pyrek, 1986). Thomassin et al. (1987) reported a very detailed quantitative structure-activity analysis of 34 substrates. The effect of side-chain length of bile acids on glucuronidation by human and rat liver fractions was studied by Kirkpatrick et al. (1988). For studies in vivo, in perfusions and in isolated cells, one would prefer substrates that only undergo glucuronidation; ideally specific substrates for every UDPGT isoenzyme should be available. Unfortunately, for most phenols and hydroxamic acids sulfation is the unavoidable companion of glucuronidation. Thus, the use of drugs like paracetamol to characterize glucuronidation in man should be avoided; effects observed might be in fact effects of sulfation, which are reflected in opposite effects on glucuronidation. Alternatively, the relative contributions of both pathways should be assessed. From the group of phenolic substrates, morphine may be used in vivo. In the dog and the Rhesus monkey [14C-]-labelled (racemic) morphine is almost exclusively glucuronidated and kinetics of the process can be followed (Garrett and Jackson, 1979; Rane et al., 1984; Jacqz et al., 1986); it can be glucuronidated both at the 3- and 6-hydroxy positions in an enantiomer and species-dependent manner. Other phenolic model substrates for glucuronidation (also in man) are lorazepam, ciramadol, oxazepam, phenolphthalein and xamoterol (Colburn et al., 1979; Gerkens et al., 1981; Abernethy et al., 1983; Midha et al., 1983; Meacham et al., 1986; Mulder et al., 1987). Ester glucuronides are formed from many carboxylic acids; at this group there is competition with CoA activation and subsequent amino acid conjugation. A number of substrates are almost exclusively glucuronidated in man and animals: iopanoic acid (Cooke and Cooke, 1983), valproic acid (Watkins and Klaassen, 1982a), clofibric acid (Baldwin et al., 1980) and tocainide (Elvin et al., 1980). One of the problems with ester glucuronides is that they are rather βglucuronidase resistant and readily rearrange especially under alkaline

GLUCURONIDATION 55

Table 4.1. Types of glucuronides formed from various chemical groups.

56 CONJUGATION REACTIONS IN DRUG METABOLISM

conditions, a phenomenon known as acyl migration (Hignite et al., 1981; Faed, 1984; Dickinson et al., 1986). With many of these substrates most likely the contribution of only a few forms of UDPGT is measured. Unfortunately, a substrate that is utilized by several UDPGT isoenzymes, 4-nitrophenol, is also an excellent substrate for sulfotransferases. When racemic substrates are used, two different substrates (the two enantiomers) are studied. These may compete but also may be converted by different forms of UDPGT. Stereoselectivity has been investigated with a number of substrates. In rat liver microsomes (−)-morphine was glucuronidated exclusively at the 3-hydroxy position, but (+)-morphine preferentially at the 6-hydroxy position. In PBpretreated rats (−)-morphine-3-glucuronide and (+)-morphine-6-glucuronide formation was significantly increased, but (+)-morphine-3-glucuronide was hardly affected, suggesting that different forms of the enzyme were involved (Rane et al., 1985). Similarly, the E-1-hydroxynortryptiline enantiomers have been studied in vitro with rat and human liver microsomes (Dumont et al., 1987). For fenoterol isomers relatively minor differences in glucuronidation rate were observed (Koster et al., 1986a). The enantiomers of propranolol and 4′-hydroxypropranolol have been extensively investigated, both in man and various animal species (Silber et al., 1982; Walle et al., 1988). After oral administration of propranolol in human volunteers the glucuronide at the alcoholic hydroxyl group was formed equally from both enantiomers. However, the conjugation of the oxidative metabolite, 4′-hydroxy-propranolol, showed pronounced stereo-selectivity. Glucuronidation shows a preference for the (−)-enantiomer, while sulfation favours the (+)-form. Rat and dog liver microsomes show opposite enantioselectivity for propranolol glucuronidation, but no preference for 4′-hydroxypropranolol (Thompson et al., 1981; Wilson and Thompson, 1984). Oxazepam is glucuronidated stereoselectively in vitro and in vivo in several species (Sisenwine et al., 1982). Qualitatively, the ratios of glucuronidation rates of the enantiomers in vivo and in vitro (liver microsomes) were in agreement, but quantitatively there were large differences, both between species and between in vivo/in vitro measurements. Stereo- and regio-selectivity of glucuronidation of vicinal dihydrodiols of polycyclic aromatic hydrocarbons or their phenolic derivatives has been investigated by Armstrong (1987) and Lilienblum et al. (1987); the size and planarity of the aglycone play an important role. 2-Aryl-propionic acid derivatives form ester glucuronides; the size and planarity of the substrates play an important role in determining the rate at which they are glucuronidated. The stereospecificity observed in vivo and in vitro is both structure and species dependent (Mouelhi et al., 1987; review by

GLUCURONIDATION 57

Caldwell et al., 1988). Enantioselectivity of 2-phenylpropionic acid has been characterized (Nakamura and Yamaguchi, 1987; Fournel-Gigleux et al., 1988). Some glucuronides are not or only very slowly hydrolyzed by β-glucuronidase preparations, so that a lack of hydrolysis is not sufficient evidence against a glucuronide structure. 4.4. Properties of UDP-glucuronosyltransferases The evidence is now overwhelming that UDPGTs exist as a multigene family (EC 2.4.1.17), resulting in a range of isoenzymes, each possessing different, but closely related, physical and catalytic properties. The concept of the multiplicity of UDPGTs has directly evolved from studies on the purified enzymes and cDNA clones, and indirectly from differential induction, ontogeny, tissue distribution and various inherited defects of UDPGT activity. The effect of administered xenobiotics: induction of UDPGT The differential induction of UDPGTs by various xenobiotics is well known (see Dutton, 1980), and two major classes of inducing agent have been defined—(a) the polycyclic aromatic hydrocarbon (PAH)-type and (b) the PB-type. PAHtype inducers, such as 3-MC and β-naphthoflavone (β-NF) induce UDPGT activity in rats towards planar phenols such as 1-naphthol, 4-nitrophenol, 4methylumbelliferone (Bock et al., 1973; Lilienblum et al., 1982) and 2aminophenol (Coughtrie et al., 1987a), whereas PB-type inducers stimulate chloramphenicol, morphine and 4-hydroxybiphenyl UDPGT activities (Bock et al., 1973; Lilienblum et al., 1982). There are several other compounds which have been reported to specifically induce UDPGT isoenzymes, such as clofibrate for bilirubin UDPGT (Foliot et al., 1975) and pregnenolone-16α-carbonitrile (PCN) and spironolactone for digitoxigenin monodigitoxoside UDPGT (Schmoldt and Promies, 1982; Watkins et al., 1982). Because of the overlapping substrate specificity of UDPGTs, the glucuronidation of some aglycones may be induced by more than one inducer, e.g. furosemide UDPGT activity is induced by PB 3-MC and PCN (Rachmel and Hazelton, 1986). The induction of UDPGT activities has recently been shown to be due to the increase in UDPGT enzyme protein as determined by immunoblot analysis (Scragg et al., 1985; Koster et al., 1986b; Burchell et al., 1987; Coughtrie et al., 1987a). This increase in enzyme protein results from increases in mRNA levels (Jackson and Burchell, 1986; Mackenzie, 1986a; Iyanagi et al., 1986; Harding et al., 1989). Thus these inducers are likely to be acting to increase the rate of transcription of UDPGT

58 CONJUGATION REACTIONS IN DRUG METABOLISM

genes. The phenomenon of differential induction of UDPGT enzyme activities has been central to the development of the concept of UDPGT heterogeneity. Little is known, however, about the inducibility of human UDPGTs, although there is some evidence that treatment of patients with PB-like drugs (PB, phenytoin) results in induced UDPGT activity. Bock et al. (1984) reported that liver microsomes from patients treated with these compounds exhibited significantly higher UDPGT activity towards 1-naphthol, 4methylumbelliferone and bilirubin. More recently, Bock and Bock-Hennig (1987) showed that PB/ phenytoin treatment resulted in increased glucuronidation of 1-naphthol, paracetamol, benzo-[a]-pyrene-3, 6-quinol and 4methylumbelliferone, whereas the conjugation of morphine and 4-hydroxybiphenyl were unaffected. This is in contrast to rat liver, where morphine and 4-hydroxybiphenyl are the major PBinducible UDPGT enzyme activities (Ullrich and Bock, 1984). Smoking may induce certain UDPGT enzyme activities in man (Bock et al., 1987; Walle et al., 1987), although the number of cigarettes which these subjects were using was high—one to two packs per day. Therefore, there is evidence for differential induction of UDPGT enzyme activities in man also, but there appears to be a different pattern of enzyme activities involved compared with rat. Ontogeny Perhaps the best example of the differential regulation of UDPGT gene expression is the variation in UDPGT activities during development. Lucier and McDaniel (1977) observed that the development of rat UDPGT activities could be divided into two main groups or ‘clusters’ (Greengard, 1971)–the steroidal and non-steroidal groups. The non-steroidal group developed late-foetally and consisted of activities towards planar phenols such as 1-naphthol, 4-nitrophenol and 4-methylumbelliferone, in which adult levels were obtained one day post partum. The other group consisted of activities towards steroids such as estradiol, estrone, testosterone and diethylstilbestrol. Similar findings for rat liver were reported by Wishart (1978) who also demonstrated that the activities in the late-foetal cluster (non-steroid group) could be precociously stimulated by administration of glucocorticoids, suggesting that hormones regulate the expression of these genes in vivo. Since these early investigations it has become apparent that the development of UDPGT activities is extremely complex, and there are now four recognized key periods during which different UDPGT activities develop in rat liver. Firstly, the late-foetal cluster, comprising activities towards planar phenols such as 1-naphthol and 2-aminophenol (Wishart, 1978); second, the neonatal cluster, comprising activities towards testosterone (Lucier and McDaniel, 1977), bilirubin (Wishart, 1978), and morphine (Wishart, 1978; M.Coughtrie et al., in press); thirdly, the postweaning cluster when activity towards androsterone (Matsui and Watanabe,

GLUCURONIDATION 59

1982) and pregnanediol (Fuchs et al., 1977) develop at about 30 days, and the pubertal cluster at 60–65 days where activity towards pregnanediol surges again dramatically (four-fold; Fuchs et al., 1977). It has recently been demonstrated that these variations in enzyme activity during development correlate with changes in the amounts of enzyme protein representing individual UDPGT isoenzymes as determined by immunoblot analysis (Coughtrie et al., 1988). Tissue distribution The liver is quantitatively the most important site of glucuronidation in the body for the majority of compounds. However, the major extrahepatic organs possess UDPGT activity, particularly those directly exposed to the external environment, i.e. intestine (Hartiala, 1973), skin (Coomes et al., 1983) and lungs (Aitio, 1976), although the complement of UDPGT isoenzymes is reduced. Early studies are reported by Dutton (1966). The majority of the work reported concerns the rat, where the kidney lacks some of the major UDPGT activities present in the liver, in particular testosterone (Lucier and McDaniel, 1977) and morphine (Rush et al., 1983). In rat lung, only UDPGT activity towards planar phenols, e.g. 1-naphthol, 4-nitrophenol, appears to be expressed (Aitio, 1974; Coughtrie et al., 1985). These observations have been confirmed at the molecular level by immunoblot analysis (see Burchell and Coughtrie, 1988). The intestinal mucosa obviously plays an important role in the glucuronidation of ingested xenobiotics (Koster et al., 1985b; see Section 4.5), and in the rats the major UDPGT enzyme activity present in this tissue is the planar phenol UDPGT—the major xenobiotic-metabolizing form (Hartiala, 1973; Koster et al., 1986b). (−)-Morphine is also conjugated in the intestine to a limited extent (Koster et al., 1985b), but there is no good evidence for the presence of any steroid-metabolizing UDPGT, nor bilirubin UDPGT. In man, UDPGT activity is present in kidney, where bile acids are readily conjugated (Matern et al., 1984; Parquet et al., 1985; Parquet et al., 1988). The human kidney is not able to metabolize bilirubin (Fevery et al., 1977). The human intestinal mucosa is also capable of glucuronidating bile acids (Matern et al., 1984; Parquet et al., 1985). Again, these variations in extrahepatic glucuronidation have been investigated at the molecular level using a monoclonal antibody directed against a common epitope on human liver UDPGT (Peters et al., 1987). Inherited deficiencies of glucuronidation The development of the concept of UDPGT multiplicity has been greatly aided by several examples of defects in glucuronidation, where certain UDPGT

60 CONJUGATION REACTIONS IN DRUG METABOLISM

activities are affected while others remain normal. This is excellent circumstantial evidence for the existence of multiple forms of these enzymes. The various populations of Wistar rat around the world exhibit a discontinuous variation in UDPGT activity towards androsterone (Matsui et al., 1979; Green et al., 1985; Corser et al., 1987), whereas the activities towards other substrates (e.g. bilirubin, planar phenols and testosterone) were normal (Matsui et al., 1979; Green et al., 1985; Jackson and Burchell, 1986). This deficiency has since been shown to be due to the absence of UDPGT enzyme protein (Green et al., 1985; Matsui and Nagai, 1986; Corser et al., 1987) and mRNA (Jackson and Burchell, 1986; Corser et al., 1987) resulting from a large deletion in the androsterone UDPGT gene (Corser et al., 1987). The Gunn rat exhibits life-long, severe, unconjugated hyperbilirubinaemia as a result of not being able to form bilirubin glucuronides. This is due to the complete absence of bilirubin UDPGT enzyme activity (Schmid and McDonagh, 1978). However, the defect in the Gunn rat is not restricted to bilirubin glucuronidation. UDPGT activity towards many other substrates, in particular planar phenols such as 2-aminophenol, 1-naphthol and 4-nitrophenol, is significantly decreased (to 5–20% of Wistar levels), whereas the glucuronidation of other compounds such as morphine and testosterone is essentially normal (see Mackenzie and Owens, 1983; Boutin et al., 1987; Coughtrie et al., 1987a; Raza et al., 1987). The defect in phenol glucuronidation has recently been shown to be due to the absence of a UDPGT isoenzyme, , from Gunn rat liver microsomes (Scragg et al., 1985; Coughtrie et al., 1987a). The phenol UDPGT gene appears to be intact at the gross level (as judged by limited restriction fragment length polymorphism analysis), but the level of mRNA detected by Northern blot analysis was decreased to about 10–20% of Wistar levels (Harding et al., 1989). This complex deficiency could result from a lesion affecting some factor(s) controlling the transcription of these UDPGT genes and/or mRNA stability which prevents their transcription. Purified UDPGTs Much of the early purification work on UDPGTs has already been reviewed (Dutton, 1980; Burchell, 1981). Solubilization of UDPGTs from the membrane usually results in pronounced increase of their measurable activity (see Section 4.9). It is still unclear whether the physiological activity equals the ‘native’ or fully activated state (see Section 4.5). Recent advances in purification methods have resulted in improved separation of UDPGT isoenzymes, and more homogeneous enzyme preparations. Perhaps the major advance has been the use of column isoelectrofocusing (chromatofocusing), in which protein mixtures are resolved based on their apparent pI on a weak anion exchanger by titrating with a pH gradient generated by a buffer consisting of

GLUCURONIDATION 61

various amphoteric substances of different pKa values. Since UDPGT isoenzymes are difficult to resolve on conventional anion exchange chromatography, chromatofocusing has allowed the separation of previously unresolved UDPGTs. This technique was originally applied by Tephly’s group (Falany and Tephly, 1983) and when combined with affinity chromatography (on UDPhexanolamine Sepharose 4B) and/or ion exchange chromatography has become the method of choice for the purification of UDPGTs. Using this approach, these workers have purified several of the major UDPGT isoenzymes from rat and human liver. From female rat liver, they purified 4-nitrophenol, 3αhydroxysteroid and 17β-hydroxysteroid UDPGTs (Falany and Tephly, 1983). The 17β-hydroxysteroid UDPGT isoenzyme demonstrated marked overlapping substrate specificity with respect to 4-nitrophenol UDPGT, exhibiting activity towards 1-naphthol and 4-nitrophenol as well as testosterone and β-estradiol. The reverse, however, was not observed. This demonstration of the overlapping substrate specificity of UDPGTs illustrates the potential of endogenous UDPGTs for glucuronidating xenobiotics, a situation which has obvious toxicological significance. The two steroid-metabolizing UDPGTs demonstrated marked stereo- and regio-selectivity, as is reflected in the nomenclature of these enzymes. Similarly, Chowdhury et al. (1986) used chromatofocusing to resolve six isoenzymes of UDPGT, although not all the preparations were homogeneous. Recently, two more UDPGTs have been purified from PB-treated rat liver. By using anion exchange chromatography and chromatofocusing, Puig and Tephly (1986) isolated morphine UDPGT . This preparation was active against morphine, but not against 4-hydroxybiphenyl, 4nitrophenol, testosterone, androsterone or bilirubin. These workers managed to separate 4-hydroxybiphenyl UDPGT activity from the morphine activity on chromatofocusing, with the 4-hydroxybiphenyl activity eluting at pH 5.5, compared with pH 7.9 for the morphine enzyme. This dispels the previous assumptions, based on differential induction experiments (see Dutton, 1980, for review), that morphine and 4-hydroxybiphenyl were glucuronidated by the same UDPGT isoenzyme. Other rat UDPGT isoenzymes which have been purified are those conjugating bilirubin (Burchell, 1980) and digitoxigenin monodigitoxoside (von Meyerinck et al., 1985) from liver, and bilirubin and 1-naphthol from kidney (Coughtrie et al., 1987b). Three UDPGTs have also been purified from mouse liver (Mackenzie et al., 1985) with activity towards 3hydroxybenzo[a]pyrene, morphine and 4-nitrophenol (representative substrates), and two isoenzymes, conjugating estrone and 4-nitrophenol, have been purified from rabbit liver (Tukey et al., 1982). UDPGTs catalyzing the glucuronidation of phenols have also been purified from pig liver (Hochman et al., 1981).

62 CONJUGATION REACTIONS IN DRUG METABOLISM

The first report of the purification of human liver UDPGT isoenzymes has recently appeared (Irshaid and Tephly, 1987). Again, using chromato-focusing and affinity chromatography on UDP-hexanolamine Sepharose 4B, these workers separated two UDPGT isoenzymes from human liver. One enzyme glucuronidated 4-nitrophenol, 4methylumbelliferone, 1-naphthylamine and estriol whereas the other was also active against 4-nitrophenol, 4methylumbelliferone and 1-naphthylamine, but was in addition active against 4aminobiphenyl. This pI=6.2 enzyme was not capable of glucuronidating estriol. Thus, purification of these two isoenzymes demonstrates not only the heterogeneity of human liver UDPGT, but also the overlapping substrate specificity characteristic of these isoenzymes. Certain purified UDPGTs have been physically characterized. N-terminal amino acid sequence analysis of 17ß-hydroxysteroid UDPGT from rat liver allowed identification of a rat liver cDNA clone as encoding this isoenzyme (Harding et al., 1987). Comparison of the N-terminal amino acid sequence with the translated nucleotide sequence revealed that this isoenzyme is synthesized as a precursor protein and that a 23 amino acid signal sequence is cleaved resulting in the mature protein. Additionally, Falany et al. (1986) carried out amino acid analysis and peptide mapping studies on three purified rat liver UDPGTs (17ßhydroxysteroid, 3a-hydroxysteroid and 4nitrophenol). These experiments demonstrated high homology between the two steroid UDPGTs, but 4-nitrophenol was significantly different—an observation which has been confirmed from cDNA clone analysis (Iyanagi et al., 1986; Jackson and Burchell, 1986; Harding et al., 1987). Cloning and expression of UDPGT cDNAs Early work on cloning UDPGT cDNAs (Mackenzie et al., 1984; Jackson et al., 1985) indicated (from restriction map analysis) that the various UDPGTs may be grouped into subfamilies, and this has been confirmed at the nucleotide sequence level (Jackson and Burchell, 1986; Iyanagi et al.,1986; Mackenzie, 1987). To date, eight rat and eight human UDPGT cDNAs have been isolated, and some have been identified by e.g. comparison of the derived amino acid sequence with the amino acid sequence of the purified protein (for rat 17βhydroxysteroid UDPGT—Harding et al., 1987) or from the inherited deficiency of androsterone glucuronidation (for rat 3α-hydroxysteroid UDPGT—Jackson and Burchell, 1986). As extensive protein sequence data is not available for the majority of UDPGTs, the preferred method of identification of cDNAs is by expression of the clone in a suitable cell culture system (such as the monkey kidney cell line, COS-7) and determining the substrate specificity of the resulting protein. This has been accomplished for several UDPGT cDNAs from

GLUCURONIDATION 63

different laboratories. Mackenzie (1986a; 1986b; 1987) has expressed three clones in COS-7 cells. The expressed UDPGTr-4 protein catalyzed the glucuronidation of androsterone, etiocholanolone and lithocholic acid. Both the UDPGT proteins resulting from the expression of UDPGTr-2 and UDPGTr-3 had catalytic activity towards testosterone, dihydrotestosterone and β-estradiol. The UDPGTr-2 protein additionally catalyzed the conjugation of chloramphenicol, 4-methylumbelliferone and 4-hydroxybiphenyl. Jackson et al. (1988) expressed a rat kidney UDPGT cDNA with an identical nucleotide sequence to a 3-MC-inducible 4-nitrophenol UDPGT cDNA isolated from rat liver (Iyanagi et al., 1986) in COS-7 cells, and the resulting glycoprotein glucuronidated various planar phenolic aglycones, including 1-naphthol, 4methylumbelliferone and 4-nitrophenol but not morphine, testosterone or androsterone. Recently, two human liver UDPGT cDNAs have been expressed in COS-7 cells: HP1, which catalyzed the glucuronidation of planar phenols such as 1naphthol and 4-methylumbelliferone (Harding et al., 1988), and H25 which, by virtue of its extensive sequence homology to rat liver 17β-hydroxysteroid UDPGT was expected to result in a protein conjugating steroids. However, when COS-7 cells transformed with H25 were assayed for UDPGT activity towards a variety of substrates, only hyodeoxycholic acid was found to be glucuronidated, and the specificity appeared to be limited to the 6α-OH position of bile acids (Fournel-Gigleux et al., 1989). Therefore the expression of UDPGT cDNAs in cell culture provides a useful means of identification of the boundaries of substrate specificity of the individual isoenzymes and a system for study of substrate transport, glucuronidation and export of glucuronides in whole cells in vitro. Substrate specificity of UDPGTs and enzyme mechanism The specificity of purified, and cloned and expressed rat UDPGTs has been assessed towards a limited range of substrates (see Tables 4.2 and 4.3). Expression of cloned rat kidney and human liver phenol UDPGTs has allowed a simple assessment of the size and planarity of phenolic compounds accepted by these enzymes. A restricted substrate specificity towards small and planar phenols was observed. Further, saturation of the aromatic ring attached to the phenolic group almost abolished the acceptibility of the substrate (Harding et al., 1988; Jackson et al., 1988). Electron withdrawing or releasing substituents in the phenolic ring of nitrophenols will also increase the acceptability of the substrates for glucuronidation (Mulder and van Doorn, 1975). A purified ‘phenol’ UDPGT from rat liver has been shown to exhibit considerable kinetic preference for pseudoequatorial hydroxyl groups in both trans- and cis-K region dihydrodiols

64 CONJUGATION REACTIONS IN DRUG METABOLISM

from 3, 4, 5, 6-tetramethyl-9, 10-dihydroxyphenanthrene. Chirality of the active site of the transferase was indicated by a pronounced kinetic discrimination toward the M and P conformational enantiomers of 1, 1′-bi-2-naphthol. These analyses allow tentative predictions to be made about which substrates could be used to identify and assay individual UDPGTs in microsomes and in whole cells and organs. Specific substrates are obvious for the transferases glucuronidating endogenous compounds (see Tables 4.2 and 4.3), although testosterone may be glucuronidated by more than one isoenzyme. There appear to be specific rat liver UDPGTs catalyzing the glucuronidation of digitoxigenin monodigitoxoside and (−)-morphine, but phenolic xenobiotics are glucuronidated by several isoforms and specific substrates may need to be designed (Armstrong et al., 1988; von Meyerink et al., 1985; Puig and Tephly, 1986). The UDPGT reaction mechanisms studied by kinetic analysis before 1983 used hepatic microsomes as the enzyme source and most commonly 4nitrophenol was used as substrate (see Dutton and Burchell, 1977; Dutton, 1980). The results obtained were obviously confused by the problems associated with kinetic measurements of compartmentalized membrane-bound transferases and the existence of multiple UDPGT isoenzymes, several of which could accept phenols as substrates. Using 4-nitrophenol Vessey and Zakim (1972), suggest a rapid-equilibrium random-order, reaction. However, Koster and Noordhoek (1983), measuring the kinetic properties of rat intestinal microsomal 1-naphthol UDP-glucuronosyltransferase, proposed an ordered sequential bireactant mechanism in which 1naphthol binds first and UDPGA second, suggesting that the enzyme mechanisms may not be consistent for different substrates and tissue locations. These minor controversies may be resolved by examination of highly purified UDPGTs, although the purified enzymes need to be carefully reconstituted with phospholipids in attempts to recreate the endogenous membrane environment. One careful study has examined two UDPGTs catalyzing the glucuronidation of steroids (Falany et al., 1986), using bisubstrate kinetic analysis, product inhibition and dead-end competitive inhibition experiments. The cumulative results are consistent with a rapid, random order, sequential kinetic mechanism for glucuronidation of testosterone and androsterone. Inhibition of UDPGT The studies of inhibition of UDPGT are difficult and confused by the microsomal membrane location of UDP-glucuronosyltransferases. Consequently very few compounds have been classically identified as specific UDPGT inhibitors resulting in lower rates of glucuronidation in vivo. Many compounds that have been considered inhibitors of glucuronidation cause membrane

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Table 4.2. Substrate specificity of purified and cloned and expressed Rat UDPGTs.

a

There are apparently at least two UDPGT isoenzymes capable of glucuronidating a range of planar phenolic compounds, and this is also seen in human liver (see Table 4.3). b Abbreviations—dt1=digitoxigenin monodigitoxoside —dt2=digitoxigenin bisdigitoxoside

66 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 4.3. Substrate specificity of purified and cloned and expressed human UDPGTs.

a

Refers to the pH at which the protein elutes from a chromatofocusing column.

disruption or reduce cellular UDPGA concentrations (see Dutton, 1980; and Section 4.7). Studies with microsomal preparations were also complicated by the likely existence of a UDPGA transporter protein facilitating movement of UDPGA to the lumenal side of the membrane (see Burchell and Coughtrie, 1988, for review). For example, N-ethylmaleimide and diazobenzene sulfonate selectively abolish stimulation of glucuronidation by UDP-N-acetylglucosamine (Haeger et al., 1980; Burchell et al., 1983) suggesting interaction and inhibition of a protein component not directly involved in catalysis of glucuronidation. Early studies of the inhibition of microsomal UDPglucuronosyltransferase by nucleotides showed that UDP competitively inhibited UDPGA binding (see Dutton, 1980). Most of the studies were done using higher than physiological concentrations of UDPGA and with undefined or uncontrolled concentrations of divalent cations. In general removal of latency by membrane perturbation increased inhibition of UDPGT activity by UDP and other nucleotides. Hallinan et al. (1979) showed that uridine tri- and diphosphates tend to inhibit UDPGT activity towards 4nitrophenol and estradiol more in sealed microsomes than leaky microsomes, again indicating another membrane component was confusing the interpretation. Studies of the inhibition of UDP-glucuronosyltransferase activity in vitro should be performed using at least optimally detergent-activated microsomes, where the affect of additional components is minimized. The multiplicity of UDPGTs will continue to confuse the interpretation, but some specificity of

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inhibition studies can be achieved using specific aglycone substrates, although additional confirmation of specific inhibition using purified isoenzymes is required. Specific analysis is perhaps more easily achieved by measurement of the endogenous compound glucuronidation such as the inhibition of bilirubin UDPglucuronosyltransferase activity. Novobiocin, which caused unconjugated hyperbilirubinaemia in animals and man was shown to exert non-competitive inhibition of rat microsomal UDP-glucuronosyltransferase in digitonin-activated preparations with either bilirubin or UDPGA (Duvaldestin et al., 1976). Human microsomal bilirubin UDPGT activity was also inhibited by novobiocin, whereas 1-naphthol UDP-glucuronosyltransferase was unaffected (Burchell et al., 1987). Further studies with purified rat liver bilirubin UDPGT suggested that novobiocin competitively inhibited bilirubin binding to the enzyme, but the drug did not appear to be a substrate for the purified enzyme. Competitive substrates Triphenylacetic acid and related compounds have been shown to competitively inhibit rat and human liver microsomal bilirubin UDPGT (Fournel et al., 1986). Among 20 compounds tested 7, 7, 7-triphenylheptanoic acid most strongly inhibited rat liver microsomal bilirubin UDPGT and had a weaker effect on 1naphthol, androsterone and testosterone glucuronidation (Fournel et al. 1986; Fournel-Gigleux et al., 1988b). This arylalkanoic acid exhibited a competitive inhibition towards rat liver microsomal and purified bilirubin UDPGT activities with respective Ki values of 12 µM and 2 µM. The glucuronidation of these compounds was also studied. 7, 7, 7Triphenylheptanoic acid was actively glucuronidated by purified bilirubin UDPGT in contrast to its analogues with decreasing alkyl chain length (Fournel-Gigleux et al., 1988a, 1988b). Studies of purified steroid glucuronidating enzymes have shown that the transferases can be competitively inhibited by steroid analogues which are not substrates for these isoenzymes (Falany et al., 1986). The binding of UDPglucuronic acid to 17ß- and 3a-hydroxysteroid UDPGTs was competitively inhibited by end products UDP, UMP and testosterone or androsterone glucuronides, respectively. Rat hepatic microsomal morphine UDPGT was competitively inhibited by a number of substrate analogues and synthetic narcotics (Sanchez et al., 1978). Codeine was confirmed as an inhibitor of the purified isoenzyme (Puig and Tephly, 1986). Cyproheptadine was determined to be a potent competitive dead-end inhibitor of morphine glucuronidation in rabbit liver (del Villar et al., 1977). Recently 1-naphthylacetic acid was determined to be a potent inhibitor and potential substrate for rat liver morphine UDPGT (M.Coughtrie,

68 CONJUGATION REACTIONS IN DRUG METABOLISM

unpublished work). Oxazepam inhibits morphine UDPGT activity in human foetal liver microsomes (Pacifici and Rane, 1981) and chloramphenicol and 1naphthol were poor competitive inhibitors of a low-affinity morphine UDPGT in adult human liver microsomes (Miners et al., 1988). Non-specific inhibitors Watanabe et al. (1986) reported that organophosphates selectively inhibit phenol glucuronidation but not phenolphthalein or testosterone glucuronidation after administration of these compounds to rats. This apparently non-specific inhibition does not occur in vitro, suggesting that metabolic activation was required to produce inhibitory metabolites. Complex non-specific inhibition of several UDPGT activities has also been observed with furosemide and salicylamide (Boutin et al., 1984). Active-site labelling Design of better substrate inhibitors requires extensive analysis of active sites of purified enzymes, although very little work has been done. Early work using active-site labelling reagents has been reviewed by Dutton (1966, 1980), but none of these studies were done using purified enzymes; therefore the results obtained are only suggestive of the presence of important amino acid side chains and the reactive groups detected may not be present in all of the individual UDPGTs. The use of thiol-specific reagents has indicated that thiol groups may be involved in the active site of UDPGTs (see Dutton, 1966, 1980). Indeed, the presence of 2-mercaptoethanol or dithiothreitol helps to maintain the activity of purified UDPGTs (Burchell, 1981) and was essential to obtain good preparations of bilirubin UDPGT (Burchell et al., 1987). Diethylpyrocarbonate, which selectively binds to histidine residues on proteins, was a potent inhibitor/ inactivator of rat 1-naphthol UDPGT in intact or disrupted microsomes suggesting that a histidine residue may be part of the active site of this transferase (Arion et al., 1984). The single example of the use of the reagent 2, 3-butanedione on purified UDP-glucuronyltransferase (from pig liver) identified the presence of an active site arginine, which is probably involved in the binding of UDP-glucuronic acid (Zakim et al., 1983). 4.5. Glucuronidation in perfused organs and isolated cells Most perfused organs and isolated cell systems that have been tested catalyze glucuronidation of added substrates, although large variations in conjugation rate are found. Even human skin epithelial cells show glucuronidation, albeit at

GLUCURONIDATION 69

low rate, but fibroblasts do not (Rugstad and Dybing, 1975). Human lymphocytes contain UDPGT, and the cells can potentially be used for characterization of glucuronidation in the human population (Li et al., 1982). No special requirements exist for the composition of the incubation medium in these in vitro systems. For studies in vivo, in perfusions and in isolated cells, substrates should be used that only undergo glucuronidation; ideally specific substrates for every UDPGT isoenzyme should be available. Unfortunately, for most substrates sulfation also occurs to some extent, except for the carboxylic acids. Latency of UDPGT: physiological or artefact It is still not clear whether the physiological activity of UDPGT in situ in the intact cell is comparable to that of native or fully activated microsomal fractions; detergents and other agents or treatments that result in a disruption of the microsomal membrane structure can activate UDPGT activity up to ten-fold, dependent on the structure of the substrate and the procedure for preparing the microsomal fraction (see Dutton, 1980, for a review). Also UDP Nacetylglucosamine has a stimulating effect in native microsomes (Burchell et al., 1983; and Section 4.4). An explanation may be that the active site of UDPGT is located at the luminal side of the endoplasmic reticulum and that a UDPGA carrier is involved in intact endoplasmic reticulum (Dutton, 1980; Hauser et al., 1988; Vanstapel and Blanckaert, 1988). Only a few approaches have been made to compare the activity in microsomal fractions with that in vivo, perfused organs or isolated cells (e.g. Hamada and Gessner, 1975; Abou-el-Makarem and Bock, 1976; Andersson et al., 1978; Morrison et al., 1986). However, little has changed since 1980 when Dutton concluded that general agreement seemed to be that the activity in the cell in situ was most likely somewhere between the fully activated state and the native activity. This has been confirmed by later work in which enzymic parameters for glucuronidation of e.g. morphine and analogues were compared with in vivo elimination kinetics of the substrate (intrinsic clearance): in vivo glucuronidation was much faster than anticipated from enzyme kinetic studies (Rane et al., 1984; Mistry and Houston, 1987). An assumption in many in vivo studies is that glucuronidation takes place only in the liver; this blatant disregard of extrahepatic glucuronidation may lead to errors in the interpretation of data (see Section 4.6). Interestingly, lipid peroxidation caused activation of UDPGT in native microsomal preparations (de Groot et al., 1985). If it also does so in the intact cell, this may be the reason why glucuronidation in the liver measured in vivo and in the perfusion was relatively unaffected by CCl4 intoxication (Bock et al., 1977; Desmond et al., 1981): in the intact cell native activity of UDPGT may prevail, and this is activated by CCl4 treatment, which compensates for the loss

70 CONJUGATION REACTIONS IN DRUG METABOLISM

of UDPGT due to toxicity of CCl4. The increased glucuronidation of paracetamol in perfused livers from rats that had been damaged by a high dose of paracetamol may have a similar cause (Poulsen et al., 1985). Liver perfusion Many substrates for UDPGT have been added to the perfused rat liver, either recirculating, or single-pass perfusion; much work has been done with phenols such as 4-nitrophenol (Gessner and Hamada, 1974; Hamada and Gessner, 1975; Reinke et al., 1981; Sonawane et al., 1981; Thurman et al., 1981; Belinsky et al., 1984; Sultatos and Minor, 1985; Morrison et al., 1986), harmol (Pang et al., 1981, 1983; Koster et al., 1982a; Dawson et al., 1985; Angus et al., 1987, 1988), 7-hydroxycoumarin (Conway et al., 1982, 1984, 1985; Reinke et al., 1986; Cha et al., 1987; Hong et al., 1987), morphine, 5-phenyl-5-phydroxyphenylhydantoin (Auansakul and Vore, 1982; Brock and Vore, 1982), xamoterol (Mulder et al., 1987), 4-methylumbelliferone (Miyauchi et al., 1987), lorazepam (Desmond et al., 1981; Branch et al., 1983), paracetamol (Poulsen et al., 1985), etoposide (Hande et al., 1988) and 1-naphthol (Bock and White, 1974; Bock et al., 1977). The subsequent events in glucuronidation as well as potential rate-limiting steps, are shown in Figure 4.2. With most of these substrates sulfation was the main route of metabolism at low concentration, while at higher concentration glucuronidation became the more important pathway. Only with xamoterol, lorazepam, morphine and 5-phenyl-5-phydroxyphenylhydantoin was sulfation negligible. If sulfate is left out of the perfusion medium, sulfation is severely reduced, and only glucuronidation is carried out when the substrate concentration is not too low (Conway et al., 1984). The glucuronides of smaller molecules are excreted predominantly into the perfusion medium (rat), while those of bigger molecules (>200 daltons) are extensively excreted in bile. Thus, 4-nitrophenyl glucuronide has a preference for the perfusion medium (Gessner and Hamada, 1974; Morrison et al., 1986), while the glucuronides of harmol and xamoterol go mainly in bile (Pang et al., 1981; Mulder et al., 1987). 4-Nitrophenyl glucuronide added directly to the perfusion medium was only slowly excreted in bile, presumably because of its slow uptake by the liver (Gessner and Hamada, 1974). Higher molecular weight glucuronides, however, are readily excreted in bile after addition to the perfusion medium. Several groups let the bile flow out into the perfusion medium effluent, which in pharmacokinetic studies is only justified if both pathways are kinetically identical. When the bile duct is ligated or biliary excretion is impaired, the glucuronides will be excreted from the hepatocyte into the perfusion medium (or blood in vivo; see Krijgsheld et al., 1982). Pretreatment with transstilbene

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Figure 4.2. Potentially rate-limiting steps in glucuronidation in the intact cell. (1) blood flow and substrate supply to the cell; (2) substrate concentration in plasma or incubation medium; (3) uptake of the substrate; (4) synthesis of UDP glucose; (5) availability of UDPGA; (6) UDPGT activity; (7) biliary excretion of the glucuronide; (8) elimination of the glucuronide to the blood; (9) competing pathways. (From Mulder, G.J., et al., 1985, with permission).

oxide causes a pronounced decrease in the biliary excretion of e.g. morphine glucuronide, which then appears in the perfusion effluent (Fuhrman-Lane and Fujimoto, 1982). Most substrates for glucuronidation are rather lipid soluble and will be rapidly taken up by cells. Then the UDGPT activity will determine their rate of removal from the perfusion medium as long as sufficient UDPGA is available. However, some substrates may be taken up almost completely during a single pass through the liver. For instance, 7-hydroxycoumarin has an extraction percentage of almost 100% (Conway et al., 1982), while for harmol it is 85% (Pang et al., 1981). In this case it is the blood or perfusion flow through the liver which is rate-limiting, and not the UDPGT enzyme activity. The extraction was 40–50% for 4-nitrophenol (Reinke et al., 1981) and only 15% for xamoterol, which is quite water soluble (Mulder et al., 1987). When glucuronidation becomes saturated (at high substrate concentration, or limiting UDPGA availability, for instance), the extraction will decrease. When the glucuronidation of a substrate is not limited by perfusion flow (an extraction percentage of e.g. 50% or lower), the conjugation rate may be increased when

72 CONJUGATION REACTIONS IN DRUG METABOLISM

UDPGT is induced with PB or 3-MC (Bock and White, 1974; Hamada and Gessner, 1975; Reinke et al., 1981). The localization of glucuronidation within the liver lobule has been studied by various groups, using harmol or 7-hydroxycoumarin as substrate. Harmol was preferentially sulfated at low input concentration in the single-pass perfused rat liver; at increasing substrate concentration sulfation becomes saturated, whereupon glucuronidation increases more than proportionally (Pang et al., 1983). The same was observed for 7-hydroxycoumarin (Conway et al., 1987). The extraction percentage (ca. 85%) remains constant. Studies in which the liver was perfused in a retrograde direction suggested that sulfation is located anterior to glucuronidation (Pang et al., 1983); the extraction percentage was higher during retrograde perfusion than during normal perfusion, while glucuronidation was increased at the expense of sulfation. This is in agreement with the model in which the system with higher affinity is in Zone 1 (sulfation) and that with the lower affinity but higher capacity (glucuronidation) either in Zone 3 or both Zones 1 and 3. Later work by this group (Dawson et al., 1985) in which the flow rate of the perfusion medium was varied both in retrograde and normal perfusions, confirmed these findings. Very similar data have recently been reported for gentisamide (2, 5-dihydroxybenzamide): Kms of ca, 25 µM for sulfation and 70 µM for glucuronidation. Perfusions in the normal and retrograde flow direction confirmed the anterior localization of sulfation to glucuronidation (Morris et al., 1988, 1989). The pharmacokinetics of this differential distribution are discussed by Pang (Chapter 2). Conway et al. (1987) used the micro-light guide technique to study 7hydroxycoumarin conjugation in their single-pass perfusion system with livers obtained from PB-treated rats. The group demonstrated that (at low substrate concentration) conjugation occurs in Zone 1 or 3 more or less selectively during normal and retrograde perfusions, respectively. Sulfation predominated in Zone 1. When 7-hydroxycoumarin was generated in the liver from 7-ethoxycoumarin, it was almost exclusively sulfated, presumably because the substrate concentration in the cells remained very low, so that the system with the higher affinity predominated (Conway et al., 1982). When sulfate was omitted from the perfusion medium so that only glucuronidation occurred, three-to-four-fold higher glucuronidation activities in Zone 3 than Zone 1 were observed. In homogenates from Zone 1 and 3 samples obtained by microdissection, the Km and maximum glucuronidation rates compared quite well with those determined in the perfusions (Conway et al., 1984). Micro-dissection of human liver yielded the same results for the distribution of UDPGT activity (Mouelhi and Kauffman, 1986). However, in untreated or 3-MC-treated rats they observed the same rate of glucuronidation in Zone 1 and Zone 3 (Conway et al., 1988). The UDPGT activity in homogenates was increased by 3-MC, but this did not lead to a

GLUCURONIDATION 73

higher glucuronidation rate in the perfusion. Apparently UDPGT activity was not rate-limiting. UDPGT activity in Zone 1 of untreated rats was much lower than in Zone 3, but after 3-MC treatment this difference disappeared. Knapp et al., (1988) demonstrated that 4-nitrophenol UDPGT was located at highest amounts in Zone 3 by immunohistochemistry. Hydroxysteroid UDPGTs were more homogeneously distributed across the liver. 3-MC induction increased UDPGT in Zone 3 as determined by assay with 1naphthol and immunohistochemistry in micro-dissection studies in rat liver (Ullrich et al., 1984). After PB induction the highest activity of UDPGT towards 1-naphthol shifted to Zone 1. Bengtsson et al. (1987) reported similar findings in hepatocytes isolated selectively from Zone 1 and 3, confirming earlier studies of Tonda and Hirata (1983). James et al. (1981) and Branch et al. (1983) claimed that glucuronidation was located at highest activity in Zone 1 based on liver perfusions and in vivo studies with rats that had been pretreated with allyl alcohol (Zone 1 damage) and CCl4 (Zone 3 damage) because allyl alcohol caused a more pronounced decrease in glucuronidation activity than CCl4. In the light of the previously described observations, it seems more likely that the interpretation of these experiments was incorrect, due to the complex effects of the toxicants in which activation of UDPGT might also play a role (see Section 4.5.). Detailed pharmacokinetic studies of conjugation are possible in the perfused liver. In the single-pass perfusion a stead-state approach is possible if the substrate concentration is kept low so that the co-substrate is not depleted. The kinetics of glucuronidation show aberrancies from Michaelis-Menten kinetics (Pang et al., 1981; Conway et al., 1982): a lag phase was observed (Figure 4.3). This was due to competition by sulfation as demonstrated by the fact that inhibition of sulfation converts this to normal kinetics for glucuronidation (Koster et al., 1982a). Detailed kinetic studies of hepatic transport can be done with the multiple indicator dilution method, as described for 4methylumbelliferone (Miyauchi et al., 1987). When substrates for glucuronidation are generated from precursors they are often sulfated rather than glucuronidated, presumably due to the low substrate concentration which favours the high-affinity sulfation system, e.g. 7ethoxycoumarin (Cha et al., 1987; Hong et al., 1987), paraoxon which is converted to 4-nitrophenol (Sultatos and Minor, 1985), and biphenyl which yields 2- and 4-hydroxybiphenyl (Kahl et al., 1978). The importance of a carbohydrate reserve for efficient UDPGA synthesis has been demonstrated by the fact that in livers from fasted rats glucuronidation was decreased; this could be alleviated by adding glucose to the perfusion medium. In livers from fed rats glucose did not enhance glucuronidation (Reinke et al., 1981; Conway et al., 1985; Angus et al., 1988). Thus, the supply of UDPGA may become rate-limiting only in fasted rat liver. Fasting also makes glucuronidation

74 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 4.3. Concentration dependence of harmol sulfation (S) and glucuronidation (G) in the single-pass perfused rat liver. The steady-state rate of conjugation was measured in the absence (left panel) and the presence of the sulfation inhibitor, 2, 6-dichloro-4nitrophenol (DCNP) (right panel). Note the disappearance of the lag-phase in glucuronidation in the presence of DCNP. (Data taken from Koster et al., 1982a).

much more sensitive to hypoxia (Angus et al., 1988). Several factors that decrease the ATP/ADP ratio (potassium cyanide, fructose, dinitrophenol, hypoxia) inhibited glucuronidation (Reinke et al., 1981; Angus et al., 1987). Glucuronidation in the perfused rat liver could be strongly influenced by adrenergic agonists: adrenaline (100 µM) in the perfusion medium reduced 4nitrophenol glucuronidation, but not its sulfation. This effect could be inhibited by α- but not by β-adrenergic antagonists. An increased Ca2+ influx into the hepatocytes seemed responsible for this effect, which resulted in an activation of microsomal β-glucuronidase activity; a futile cycle of glucuronidation and deglucuronidation seemed to operate. The calcium ionophore A–23187 also had an inhibitory effect on glucuronidation (Belinsky et al., 1984; Sokolove et al., 1984; Dwivedi et al., 1987; Conway et al., 1988). What the relevance of these findings is to the in vivo situation remains to be determined. In the perfused liver, effects of postnatal development (Sonawane et al., 1981), pregnancy (Auansakul and Vore, 1982; Brock and Vore, 1982), chronic ethanol treatment (Reinke et al., 1986; Hong et al., 1987) or diabetes (Morrison et al., 1985) can be evaluated in detail because complicating influences from other parts of the body or of blood flow through the liver are avoided. Moreover, the organ is still intact.

GLUCURONIDATION 75

Table 4.4. Apparent Michaelis-Menten parameters for glucuronidation in isolated hepatocytes.

Isolated hepatocytes Many substrates are glucuronidated in isolated hepatocytes. The UDPGA concentration in the hepatocytes may transiently be decreased after cell isolation but it recovers again during a preincubation (Ullrich and Bock, 1984; Croci and Williams, 1985; Mizuma et al., 1985). In most cases hepatocytes from the rat have been used, but those from mouse have also been used (e.g. Sweeny and Weiner, 1985), guinea pig (Schwenk and Locher, 1985), cow (Suolinna and Winberg, 1985), dog (Bolcsfoldi et al., 1981), fish (Morrison et al., 1985) and man (Grant et al., 1987). Human liver slices form the glucuronide of 4hydroxybiphenyl (Powis et al., 1987a). Omission of sulfate from the incubation medium results in low levels of sulfation so that glucuronidation occurs almost exclusively. High lipid solubility of the substrate favours a high glucuronidation rate (Fry and Paterson, 1985). Studies with hepatocytes allow the determination of apparent kinetic parameters, although deviation from Michaelis-Menten kinetics may preclude this (Koster et al., 1982a; Evelo et al., 1984; Fry, 1987). Table 4.4 lists such values for a number of substrates. The usual preference of sulfation over glucuronidation at low substrate concentration is also observed in hepatocytes with few exceptions (Wiebkin et al., 1978; Suolinna and Mäntylä, 1980; Koster et al., 1981; Evelo et al., 1984; Suolinna and Winberg, 1985; Fry, 1987). Addition of a precursor for a substrate of glucuronidation and sulfation, e.g. 4-methoxy-biphenyl (Fry, 1987), 4nitroanisole (Moldeus et al., 1976; Eacho et al., 1981b) and aniline (Evelo et al., 1984), usually results in the formation of mainly the sulfate conjugate of the product. When 4-nitroanisole oxidation to 4-nitrophenol was enhanced by PB pretreatment, glucuronidation was increased at the expense of sulfation (Moldeus et al., 1976). These results support the concept of high-affinity/low-

76 CONJUGATION REACTIONS IN DRUG METABOLISM

capacity sulfation on one hand, and low-affinity/high-capacity glucuronidation on the other. In most studies freshly prepared hepatocyte suspensions are used. Cryopreservation of isolated hepatocytes is possible, but leads to pronounced loss of glucuronidation capacity (Powis et al., 1987b). Primary cell cultures allow much longer periods of study-days rather than hours. UDPGT activity is reasonably constant in cultured cells for at least four days (Fry and Bridges, 1980; Holme et al., 1983; Croci and Williams, 1985; Suolinna and Pitkäranta, 1986) and it can be increased by induction (Forster et al., 1986; Suolinna and Pitkäranta, 1986). Similar findings were reported for human hepatocyte monolayer cultures: activities increased two- to three- fold during a 72 h culture period (Toribara et al., 1984; Grant and Duthie, 1987). In the majority of established cell cultures tested by Wiebel et al. (1980) UDPGT activity was present. In the human HepG2 cell line induction by PB occurred (Grant et al., 1988). The effect of animal pretreatment such as the effect of low oxygen or metabolic inhibitors, ethanol, D-galactosamine or diethyl ether (Section 4.2) can conveniently be analyzed. Pretreatment of rats with PB or 3-MC enhances the glucuronidation of some substrates, depending on the forms of UDPGT that are induced (Table 4.5). Biological effects can also be studied in isolated hepatocytes. Thus, no aging of glucuronidation of paracetamol in mouse and rat hepatocytes was observed (Sweeny and Weiner, 1985). In streptozotocin-diabetic, fasted rats glucuronidation of 4-nitrophenol was increased in hepatocytes from male, but not from female rats. Treatment with insulin returned the glucuronidation rate to normal (Eacho et al., 1981a, 1981b). Later work by Grant and Duthie (1987) demonstrated a decreased rate of glucuronidation for 1-napthol and phenolphthalein in isolated hepatocytes, presumably due to a decreased UDPGA level. They attributed these controversial results to the use of fasted (Eacho et al., 1981b) or fed rats. They could only find a decrease of native UDPGT activity, not of activated UDPGT (Morrison and Hawksworth, 1984). A similar decrease of 4-nitrophenol glucuronidation was observed in the perfused liver from streptozocin-diabetic rats (Morrison et al., 1985). Glucuronidation of 4-nitrophenol was inhibited by adenine nucleotides like adenosine and dibutyryl cyclic AMP; since millimolar concentrations were required, the physiological relevance of these findings is as yet uncertain (Shipley et al., 1986). Glucuronidation of phenolphthalein was not affected (Banhegyi et al., 1988). Studies on the uptake of glucuronides by hepatocytes suggest that carriers are involved; these have been somewhat characterized in membrane preparations from the hepatocytes (Barnhart and Witt, 1983; Brock and Vore, 1984; Brouwer et al., 1987; Takacs and Vore, 1987).

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Table 4.5. Effect of pretreatment of rats on glucuronidation in isolated hepatocytes.

a

abbreviations: BHA: butylated hydroxyanisole; PCB: polychlorinated biphenyls. more than two-fold; •=significantly increased but less than two-fold; – =no effect.

b +=

Intestinal glucuronidation in vitro and in vivo Many substrates are glucuronidated during absorption after oral administration in most species investigated (see review by Koster, 1985). In the rat glucuronidation predominates over sulfation, while in the dog and man sulfation is the more important reaction (e.g. Rogers et al., 1987). High conjugation rates may also occur in the intestinal area when certain substrates are in the splanchnic blood, as shown in the intact rat for 4 methylumbelliferone (Mulder et al., 1984a), fenoterol (Koster et al., 1985a) and L-Dopa (Landsberg et al., 1975). Glucuronidation in the intestine can be induced, e.g. by butylated hydroxyanisole in mice (Hjelle et al., 1985a): both UDPGT activity towards paracetamol (but not diethylstilboestrol) and UDPGA concentration increased. Distribution of glucuronidation along the rat intestinal tract showed relatively little difference from duodenum to distal ileum. The maximum capacity of 1naphthol glucuronidation in mucosa cells isolated from various parts decreased to about 20% from jejunum to colon (Schwenk and Locher, 1985; Koster et al.,

78 CONJUGATION REACTIONS IN DRUG METABOLISM

1985b). No such difference was observed for morphine as substrate. The reports on distribution of UDPGT in villus and crypt cells are conflicting; whereas Koster et al., (1984a) reported the highest activity in crypt cells, Dubey and Singh (1988) showed that activities of UDPGT were high in upper villus cells and very low in crypt cells. The UDPGA concentration followed the same distribution. Differential inducibility by PB and 3-MC was reported. Glucuronidation can be studied in isolated intestinal segments or the (vascularly) perfused intestine. Different results may be found when a phenolic substrate is added either at the luminal or the serosal side of the gut (Sund and Lauterbach, 1986). Also the efflux of the glucuronide to either side of the intestinal preparation shows rather complex behaviour, depending on substrate, site of addition and type of gut preparation (Josting et al., 1976; Iwamoto and Watanabe, 1983; Koster and Noordhoek, 1983; Wollenberg et al., 1983). Piperine inhibited 3-hydroxybenzopyrene glucuronidation in guinea pig small intestinal mucosa cells, primarily due to a decreased UDPGA concentration, although direct inhibition of glucuronidation may also have played a role (Singh et al., 1986). Glucose stimulated glucuronidation, suggesting that the intestinal cells have no carbohydrate reserve (Koster et al., 1984b). Kidney The kidney contributes to in vivo glucuronidation of various compounds as demonstrated by the SADR technique (Section 4.6). This could be confirmed in perfusions with human and rat kidneys where a number of phenols were glucuronidated and their kinetics of conjugation could be studied as well as the effect of induction (Elbers et al., 1980; Diamond and Quebbemann, 1981; Emslie et al., 1981). Hjelle et al. (1986) studied UDPGT in cortex, outer stripe of the medulla and proximal tubules from rabbit kidney obtained by microdissection. Highest activities of UDPGT and concentrations of UDPGA were found in the proximal tubule area. These results are in agreement with previously observed glucuronidation activities in renal tubular cells or fragments, (e.g. Shirkey et al., 1979). Other organs Glucuronidation can be detected in lung. Human bronchus or bronchoscopy samples metabolize 1-naphthol to the glucuronide and sulfate conjugate (Gibby and Cohen, 1984). Corresponding tumour tissue yields much more of the glucuronide (Gibby et al., 1985). In lung perfusions glucuronidation can be demonstrated (Hogg et al., 1981; Uotila, 1982).

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Figure 4.4. Pharmacokinetic scheme for glucuronidation of a substrate and its pathways of excretion. Note the competition by sulfation. (From Mulder, G.J., 1986, Federation Proceedings, 45, 2229, with permission of the FASEB).

Other tissues where glucuronidation occurs are hamster embryo cells (Nemoto et al., 1978), aortic cells (Yang et al., 1986) and mouse skin strip (Moloney et al., 1982). 4.6. Glucuronidation in vivo Pharmacokinetics and excretion Glucuronidation in vivo has been extensively studied in animals and man. Glucuronides are excreted in both urine and bile (Figure 4.4). After biliary excretion they may be hydrolyzed by β-glucuronidase in the gut, and the aglycone will be taken up again in a species- and substance-dependent manner (Colburn et al., 1979; Garrett and Jackson, 1979; Siegers et al., 1983; Watari et al., 1983; Greenblatt and Engelking, 1988). After instillation of glucuronides into the duodenum, the aglycones can be found in blood after a few hours. In the rat, biliary excretion is a major route of elimination for glucuronides with a high molecular weight, but in man biliary excretion seems less important than in the rat (Klaassen and Watkins, 1984; Verbeeck et al., 1988). When in pharmacokinetic studies only urinary excretion of glucuronides is followed, part of the dose may initially have been excreted in bile as glucuronide and only after reabsorption of the aglycone from the gut has been excreted as sulfate conjugate in urine. Therefore, one cannot use urinary excretion data of glucuronides solely as a reflection of their rate of synthesis. The more so since certain treatments may shift excretion of glucuronides from bile to urine (e.g. Gregus and Klaassen, 1988). Biliary and urinary excretion of glucuronides occur simultaneously (e.g. Gregus et al., 1988a, 1988b) and both excretory pathways are mutually compensatory (Jorritsma et al., 1979; Siegers and Klaassen, 1984). There is

80 CONJUGATION REACTIONS IN DRUG METABOLISM

evidence that for some special compounds the urinary excreted glucuronides reflect extrahepatic glucuronidation (Gregus et al., 1988b). The urinary excretion of glucuronides does not seem to involve active transport but only glomerular filtration (Hekman et al., 1986; see Moller and Sheikh, 1983, for review). Renal failure will result in increased plasma levels of glucuronides and possibly deconjugation to the parent compound (Verbeeck, 1982). The substrate is usually administered orally or intravenously. In the latter case the kinetic analysis is relatively simple since the systemic availability of the substrate is 100%, unless the lung shows high first-pass metabolism (see below). After oral administration only a part of the dose may be absorbed, and first-pass metabolism in the gut, gut mucosa, liver or lung, may further reduce the systemic availability of the dose. The glucuronidation of several drugs has been characterized in vivo. A very detailed study has been undertaken of morphine glucuronidation in the dog. At high dose it had effects on hepatic blood flow and urine flow which complicated the analysis of the data. Extraction by the liver was almost complete so that the clearance was dependent on liver blood flow. Some 14% was excreted in bile as the glucuronide; once the glucuronide was in the blood, it was only excreted in urine and not at all in bile (Garrett and Jackson, 1979). Paracetamol conjugation and conjugate excretion in the rat has been extensively investigated by Watari et al. (1983). They calculated apparent Km and Vmax values for sulfation and glucuronidation by deconvolution; the Km values (plasma concentrations) were 110 µM for sulfation and 920 µM for glucuronidation. Paracetamol plasma disappearance curves showed typical saturation kinetics, due to saturation of sulfation and concomitant depletion of inorganic sulfate. Glucuronidation was not saturated over the dose range used. Paracetamol glucuronide was also excreted in bile when the glucuronide itself was administered intravenously; excretion rates of the glucuronide reflected glucuronidation rates. Hjelle and Klaassen (1984) reported similar findings. Xamoterol is exclusively glucuronidated in the rat, while in the dog sulfation occurs (Mulder et al., 1987; Groen et al., 1988). Kinetics in both species have been reported in relation to sulfation and glucuronidation. Similar pharmacokinetic data are available on isotretinoin glucuronidation in rat (Meloche and Besner, 1986). Man has a limited capacity for glucuronidation of drugs such as salicylic acid and paracetamol (Levy et al., 1972; Slattery and Levy, 1979). Paracetamol conjugation kinetics were analyzed at both therapeutic and toxic doses. More limited pharmacokinetic investigations have been published on many drugs, both phenols and carboxylic acids such as salicylamide (Morad, 1982), propranolol (Midha et al., 1983; Walle et al., 1988), lorazepam, oxazepam (Abernethy et al., 1983, 1985), tocainide (Elvin et al., 1980), oxaprozin (Janssen et al., 1980) and propofol (2, 6-diisopropylphenol; Simons et al.,

GLUCURONIDATION 81

1988). Several of these drugs are racemic and show stereo-selectivity in glucuronidation (see Section 4.3). Often glucuronidation is not the only reaction occurring; because of competing pathways, the pharmacokinetics of elimination therefore need not reflect the glucuronidation rate. First-pass conjugation can occur after oral administration in the gut mucosa and subsequently the liver. Salicylamide was subject to very extensive first-pass conjugation in the rabbit (95%) and the rat (60%). In the rat 1/3 of the firstpass metabolism occurred in the gut, exclusively glucuronidation; the 2/3 in the liver was equally sulfation and glucuronidation (Shibasaki et al., 1981). In the dog gut mucosa phenols were usually sulfated, while in the liver glucuronidation followed (Gugler et al., 1975; Groen et al., 1988). Clofibric acid, the major metabolite of clofibrate, is mainly converted to the ester glucuronide. Biliary excretion was the main initial excretory pathway in the rat. However, due to enterohepatic recirculation, ultimately everything was excreted in urine. PB pretreatment did not increase the glucuronidation rate (Baldwin et al., 1980). Another well-studied substrate is diflunisal, which forms both an ester and an ether glucuronide. Both were saturable in the rat, but ether glucuronidation seemed to have a lower Km and capacity than the formation of the ester glucuronide. The kinetics were influenced by non-linear protein binding of diflunisal. Glucuronidation was the rate-limiting step in the elimination of diflunisal. Since much of the dose was excreted in bile, enterohepatic circulation played a role in its pharmacokinetics (Lin et al., 1985a, 1985b). In man ester glucuronide formation seems the first pathway to become saturated (Loewen et al., 1986, 1988). After pretreatment of rats with several inducers, especially PB, the clearance by ether glucuronidation was enhanced three-fold, and that by ester glucuronidation two-fold. Valproic acid glucuronidation in the rat has been investigated in relation to its induction and inhibition by several agents (Watkins and Klaassen, 1982a; Heinemeyer et al., 1985). PB treatment markedly increased valproate clearance. The glucuronide was mainly excreted in bile; because the ester glucuronide can rearrange by shifting from the 1-position to other positions on the glucuronic acid ring (acyl migration), the analysis of the metabolite excretion can become quite complicated (Dickinson et al., 1986). One striking feature of human drug glucuronidation in vivo is the ability of man and the great apes (e.g. chimpanzees) to form quaternary ammonium glucuronides of tertiary amine drugs, such as cyproheptadine (Fischer et al., 1980), cyclobenzaprine (Hucker et al., 1978) and tripelennamine (Chaudhuri et al., 1976). Metabolism studies in man, great apes, monkeys, and laboratory animals (e.g. Fischer et al., 1980) have demonstrated that man and the great apes are the only species able to form quaternary ammonium glucuronides from these compounds. This is in contrast to the formation of other N-linked glucuronides, such as that of a sulfonamide (sulfodimethoxine) (Adamson et al.,

82 CONJUGATION REACTIONS IN DRUG METABOLISM

1970) where both man and monkeys but not laboratory animals were able to form the N-glucuronide. Human liver microsomes have also been demonstrated to form quaternary ammonium glucuronides from tertiary amines in vitro (Le Bigot et al., 1983). This suggests that there is a UDPGT isoenzyme present in man and the great apes which is responsible for the formation of quaternary ammonium glucuronides from tertiary amines, and that this isoenzyme is different to the one responsible for the formation of other N-glucuronides. This remains to be demonstrated in vitro and would require the purification and/or expression of the individual isoenzyme involved. As with most UDPGTs there are likely to be endogenous substrates for these isoenzymes which remain to be identified. Extrahepatic glucuronidation It is often tacitly assumed that glucuronidation in vivo takes place in the liver. However, in recent years evidence has accumulated that glucuronidation does occur to an appreciable extent extrahepatically. This can be detected by administration of a substrate at various sites, e.g. intraarterially, intravenously, in the hepatic portal vein, and by blood sampling at those sites. Alternatively, the elegant specific activity difference ratio (SADR) technique may be used, in which radio-labelled substrate and unlabelled glucuronide are continuously infused into the general circulation (see Tremaine et al., 1985, for review). This technique can also demonstrate whether hydrolysis of conjugates occurs in vivo. Cassidy and Houston (1984) used different routes of administration of phenol to demonstrate that the lung had an appreciable capacity for conjugation, as did the intestinal mucosa. The liver had a relatively low extraction efficiency for phenol, especially at high concentration. In later work with 1-naphthol in the rat (Mistry and Houston, 1985), the findings with phenol for these three organs were confirmed. The pulmonary extraction ratio was 0.38, to which sulfation and glucuronidation contributed equally. Oral administration increased glucuronidation because in rat intestinal mucosa glucuronidation is the predominant conjugation reaction. The lung did not catalyze the glucuronidation of morphine, naloxone and buprenorphine; their glucuronidation occurred mainly in the liver. A high glucuronidation rate in the gut mucosa was also observed (Mistry and Houston, 1987). More indirect approaches showed that the total body clearance of 4methylumbelliferone, due to glucuronidation, far exceeded liver blood flow. Glucuronidation of this compound in the liver was blood-flow limited because its extraction was almost 100%. 4-Methylumbelliferone was glucuronidated to a high extent extrahepatically, for instance in the intestinal vascular bed (Mulder et al., 1984a, 1985a, 1985b). This was confirmed for fenoterol, a drug which exhibits both a high presystemic and systemic glucuronidation in the intestine

GLUCURONIDATION 83

(Koster et al., 1985a), and may also apply to harmol and 4nitrophenol (Machida et al., 1982; Mulder et al., 1984b). In the dog, Gerkens et al. (1981) and Jacqz et al. (1986) have evaluated the role of the liver in glucuronidation of lorazepam and morphine by hepatic devascularization; they concluded that up to 50% of glucuronidation was extrahepatic. Using the SADR technique, Rush et al. (1983) and Tremaine et al. (1984) concluded that the rat kidney contributed ca. 10% of total body glucuronidation of 4-nitrophenol in the male, and 20% in the female rat. This sex difference was due to differences in UDPGT activities in the kidney for this substrate, which did not exist for the glucuronidation of morphine. A similar high renal contribution to the glucuronidation of 1-naphthol in vivo was observed for male rats. These results show that extensive extrahepatic glucuronidation is very likely with certain compounds, but at present insufficient data are available to assess its overall quantitative importance in most species, including man. It obviously reflects the tissue-specific expression of different UDPGT isoenzymes. Effect of internal or external factors Several treatments that decrease UDPGA availability, as discussed in section 4.2, also decrease glucuronidation in vivo, such as ethionine and fructose (Dills and Klaassen, 1986), D-galactosamine (Watkins and Klaassen, 1982a; Gregus et al., 1983), and diethyl ether (Cooke and Cooke, 1983; Gregus et al., 1983; Dills and Klaassen, 1984; Watkins et al., 1984). Inducers of drug metabolism have differential effects, depending on the substrate and whether UDPGT activity was rate-limiting in elimination. Thus, PB increased chloramphenicol glucuronidation in the rat (Stramentinoli et al., 1974). Butylated hydroxyanisole increased paracetamol glucuronidation in mice sevenfold. UDPGT activity was only increased two-fold, but UDPGA availability sixfold (Hazelton et al., 1986). The effects of a number of inducers on paracetamol conjugation in rats have been tested by Siegers et al., (1980). PB and rifampicin increased glucuronidation while 3-MC or DDT had no effect. Patients taking rifampicin or anticonvulsants also have increased paracetamol glucuronidation. Smoking had little or no effect on paracetamol conjugation (Prescott et al., 1981; Miners et al., 1984; Bock et al., 1987). PB did not have an effect on tocainide glucuronidation (Elvin et al., 1980). Fasting results in decreased UDPGA availability (see Section 4.2) and decreased rate of paracetamol glucuronidation in the rat in vivo (Price and Jollow, 1988). Yet, when a moderate dose of harmol was given to rats even after a 72 h fast, no major effects were seen on glucuronidation; however at high

84 CONJUGATION REACTIONS IN DRUG METABOLISM

substrate load the glucuronidation rate was decreased (Mulder et al., 1982). This seems to agree with data in the perfused rat liver (Reinke et al., 1981). The diet can affect glucuronidation at two levels: the UDPGT activity and UDPGA availability. The observed effects are not easily interpretable. For instance fish oil (but not several other oils) or a low lipid diet increased some UDPGT activities in rat liver considerably (Mounié et al., 1986). At low protein levels in the diet, rat liver UDPGT was increased when compared to a normal or high protein level; food consumption has to be carefully controlled in these studies to prevent misinterpretation (Merrill and Bray, 1982). A low protein diet will contain little cysteine and methionine which may cause a reduced sulfate availability and consequently lower sulfation of a substrate for both conjugations (Jung, 1985). Then glucuronidation may appear increased because more of a phenol will be excreted as glucuronide. A charcoal-broiled beef diet that increased phenacetin oxidative metabolism in man had no effect on paracetamol glucuronidation (Anderson et al., 1983). Glucuronidation rates of lorazepam, oxazepam and paracetamol in man are increased in obesity (Abernethy et al., 1983). Sex differences in glucuronidation do not show a clear pattern (Mulder, 1986). Dependent on assay condition, the substrate and UDPGT isoenzyme, sex differences may be found, but it is difficult to generalize from the often rather conflicting, mainly in vitro data. For phenols male liver tends towards a higher activity than female liver (Niedermeyer and Shapiro, 1988), while for steroids there is an opposite tendency. One of the few clear-cut examples is the sex difference in 4-nitrophenol glucuronidation in the kidney of the rat (Rush et al., 1983). For phenol and 4-nitrophenol, no such differences were found in overall conjugation in the rat in vivo (Meerman et al., 1987). Recently, a sex difference in 1-naphthol glucuronidation in rat liver cubes was described, the male exhibiting more than three-fold higher activity than the females. The activity was under control of androgens and oestrogens (Graham and Skett, 1987). No data on in vivo glucuronidation of 1-naphthol in both sexes are available to indicate the physiological relevance of these findings. A similar sex difference in UDPGT activity in rat liver was reported by Galinsky et al. (1986). In man, sex differences have been noted in the glucuronidation of oxazepam; the clearance was 30% lower in females and little effect due to age was observed between 20 and 85 years (Greenblatt et al., 1980). The glucuronidation of salicylic acid was higher in men than in women; however oral contraceptive steroids increased glucuronidation in women to the male levels, an effect also observed for other substrates (Miners et al., 1986a, and references therein). During pregnancy paracetamol glucuronidation in rats and mice was little affected (Lin and Levy, 1983; Larrey et al., 1986), even though activated UDPGT activity towards paracetamol was decreased by 50% in pregnant mice. In pregnant women glucuronidation of paracetamol after oral administration

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was increased by 75%, while sulfation was unaffected (Miners et al., 1986b). In the perfused liver from pregnant rats little difference in glucuronidation of morphine or 5-phenyl-5-p-hydroxyphenyl hydantoin was observed (Auansakul and Vore, 1982; Brock and Vore, 1982). The glucuronidation of paracetamol has also been determined in pregnant sheep and their foetuses in vivo. Glucuronidation clearance in the foetal lamb was only 18% that of the mother, which agrees with Vmax differences in the liver microsomes (Wang et al., 1985, 1986). The partial clearance of paracetamol to the glucuronide in old (25–26 months) rats was higher than in young adults (5–6 months). Data over the age period 5–90 days for paracetamol in the rat did not show much sex difference in conjugation (Green and Fischer, 1981). In ageing rats glucuronidation efficiency decreased after 6–20 months due to both decreased UDPGA and UDPGT levels (Borghoff et al., 1988). A report by Galinsky and Corcoran (1986) which showed increased glucuronidation with age in the rat is inconclusive since they did not take into account biliary excretion. In order to evaluate the effect of a traumatic injury on drug metabolism, Griffeth et al. (1985) have determined the effect of infrarenal aortic ligation on glucuronidation of chloramphenicol, one to three days after surgery. Chloramphenicol clearance was decreased 30% presumably due to both decreased UDPGA availability and altered pharmacokinetics. Lorazepam glucuronidation was increased in burn-trauma patients (Martyn and Greenblatt, 1988). In neoplastic or preneoplastic tissue, immunochemically detected UDPGT was increased which might explain increased glucuronidation of 1-naphthol in such tissue (Cohen et al., 1983; Fischer et al., 1983). The Gunn rat lacks UDPGT activity towards bilirubin (see Section 4.4) and therefore is jaundiced. Glucuronidation in vivo of other substrates, such as iopanoic acid, may be unaffected (Barnhart et al., 1982). A rat strain has been identified that is deficient in hepatobiliary excretion of organic anions; this has been cross-bred with Gunn rats to result in double mutants (Jansen et al., 1987). 4.7. Inhibition of glucuronidation in intact cells and in vivo Several compounds that inhibit UDPGT in vitro have been discussed in Section 4.4. In intact cell preparations (isolated cells, perfusions, in vivo) glucuronidation may be inhibited by either a decrease of UDPGA availability (Section 4.2) or by inhibition/decrease of UDPGT activity. No highly efficient selective inhibitors of glucuronidation in vivo or in intact cells are available at present. The compounds that have been used are all substrates for glucuronidation themselves. Thus salicylamide has been used; however, it

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inhibits sulfation much more effectively than glucuronidation (Mulder and Scholtens, 1977); in isolated hepatocytes it had relatively little effect on glucuronidation (Andersson et al., 1978; Zaleski et al., 1983). It did not inhibit glucuronidation of tocainide in man (Elvin et al., 1980). Oxazepam is somewhat more effective. It inhibits paracetamol glucuronidation in microsomes and isolated cells and causes a decrease in the clearance of paracetamol in vivo (Dybing, 1976). Other benzodiazepines, like lorazepam and diazepam, can inhibit glucuronidation of ciramadol, but they were relatively ineffective as such in the dog in vivo (Meacham et al., 1986). Clorpromazine and SKF 525A were potent inhibitors of 4-aminophenol glucuronidation in whole rat hepatoma cells in culture but weaker noncompetitive inhibitors of UDP-glucuronyltransferase in cell homogenates (Dybing 1972; Dybing and Rugstad, 1973). Probenicid, which forms an ester-type glucuronide, decreased paracetamol, lorazepam and zomepirac clearance in man (Abernethy et al., 1985; Smith et al., 1985). It may also interfere with renal elimination of glucuronides by active transport (Smith et al., 1985). Cimetidine did not affect glucuronidation of lorazepam and oxazepam in man (Patwardhan et al., 1980). However, ranitidine was reported to strongly reduce paracetamol glucuronidation in the rat in vivo; it also inhibited microsomal UDPGT with a Ki of 40 µM (Rogers et al., 1988). Both ranitidine and cimetidine inhibited paracetamol glucuronidation in isolated hepatocytes (Emery et al., 1985). Interestingly, a commonly used cytochrome P-450 inhibitor, metyrapone, strongly inhibited paracetamol glucuronidation in vivo (Galinsky et al., 1987; Galinsky and Corcoran, 1988). 4.8. Conjugation and biological activity Glucuronidation in general profoundly changes the biological effects of compounds as a result of the addition of a large hydrophilic group so that the metabolite will not easily fit into its receptor binding sites. For endogenous compounds glucuronidation seems to play mainly a catabolic role: steroid hormones, thyroxine and similar endogenous alcohols or phenols are excreted in urine or bile following glucuronidation. Therefore this conjugation mechanism is involved in their elimination. It plays a very important role in the elimination of bilirubin, which is only possible after glucuronidation and some other sugar conjugations (Heirwegh and Brown, 1982). Therefore, jaundice of the unconjugated type results from inhibition of bilirubin glucuronidation as caused by several drugs, e.g. novobiocin (Duvaldestin et al., 1976). Genetic defects in bilirubin UDPGT are known in rat (Gunn rats) and man (Crigler-Najjar, Gilbert syndromes).

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Certain glucuronides are present in the circulating blood and may still be biologically active, such as glucuronides of retinoic acid or 1, 15-dihydroxy vitamin D (Litwiller et al., 1982; Londowski et al., 1985; Zile et al., 1987). The glucuronide as such may be active or it may represent a storage form from which the aglycone can be released by (microsomal) β-glucuronidase. It is possible that further investigation will identify other biologically-active glucuronides. For instance, glucuronidation at the 6-position of morphine or nalorphine enhanced their binding to opioid δ-receptors but decreased that to the µ-receptor; conjugation at the 3-position led to a loss of activity (Oguri et al., 1987). These binding data were confirmed by Pasternak et al. (1987); they also found that morphine-6-glucuronide was a potent µ-agonist more effective than morphine itself when administered by microinjection in the brain. Although it seems that most xenobiotics are detoxified, glucuronidation may also lead to toxic effects in certain cases. A well-investigated example is choleresis by glucuronides. In general glucuronide excretion in bile causes choleresis of ca. 15–20 µl bile µmol−1 glucuronide (Klaassen and Watkins, 1984). The phenolic drug, harmol, however, causes cholestasis at high dose in the rat because harmol glucuronide is excreted to such a high extent in bile that it precipitates and the bile duct becomes obstructed (Krijgsheld et al., 1982). Several steroid D-ring glucuronides are cholestatic in the rat, monkey, and possibly man (Vore and Slikker, 1985), which may be related to occasional cases of jaundice seen with ethinyloestradiol contraceptive use. A-ring glucuronides, or sulfate conjugates at the D-ring are not cholestatic. The cause of the cholestasis is probably an effect on bile formation at the level of carriers or membrane integrity of the hepatocyte biliary membrane (Adinolfi et al., 1984; Durham and Vore, 1986). Another steroid-like compound, lithocholate glucuronide, is cholestatic, as is lithocholate itself (Oelberg et al., 1984). The ester glucuronides may be relatively reactive and could bind covalently to protein (and DNA?). This may lead to certain effects, e.g. allergic reactions; however, thus far the evidence for such effects is rather scanty (Faed, 1984; Wells et al., 1987). As a result of acyl migration these glucuronides become βglucuronidase resistent and can be reabsorbed in intact form into the gut after intrabiliary administration (Dickinson et al., 1985). Finally, it has been proposed that the carcinogenesis of 2-naphthylamine in the bladder is due to the formation of an acid-labile N-hydroxy-N-glucuronide metabolite which may be hydrolyzed in the bladder if the pH is sufficiently acidic to the carcinogenic hydroxylamine. In this case glucuronidation would serve as a detoxication in the liver, but as a shuttle to urine, resulting in local tumours in species with acidic urine such as man and dog (Young and Kadlubar, 1982).

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4.9. Practical considerations Much of the methodology concerning assay of UDPGTs has been reviewed in previous publications (e.g. Dutton, 1980; Burchell, 1981; Dutton et al., 1981; Hallinan, 1983; Bock et al., 1984; and the numerous references contained within these publications). Here, we shall concentrate on some of the more recent methods and their application in the study of the properties and actions of UDPGTs. In freshly prepared microsomal samples, UDPGTs are latent, i.e. their capacity for glucuronidation is not fully expressed in the absence of a membraneperturbing agent (Dutton, 1980). In order to approximate to the maximum value for the UDPGT enzyme activity in a given sample, the activity must be measured in the presence of the optimally-activating concentration of an appropriate membrane-perturbing agent (most commonly used are non-ionic detergents such as Lubrol PX, Brij 58 and Emulgen 911) determined for each aglycone and source of enzyme. If this is not done, then comparisons of UDPGT activity in different samples, e.g. from xenobiotic-treated animals, from different tissues or from different ages, are meaningless. Generally, enzyme source, e.g. microsomes, and detergent are incubated together at 0°C for 30 min prior to assay. Microsomes which have been frozen and thawed once will still exhibit excellent latency if preincubated at 37°C for 30 min to reseal microsomes prior to activation with detergent. Several assays have been described which allow the measurement of UDPGT activity towards a range of aglycones without the need for any methodological variation. The first method published in 1975 by Mulder and van Doorn relies on the formation of pyruvate by the reaction of the by-product of the glucuronidation reaction (UDP) with phosphoenolpyruvate in the presence of pyruvate kinase. The UDPGT activity is then determined from the consumption of NADH (measured spectrophotometrically) in the presence of the formed pyruvate by the action of lactate dehydrogenase. Several criticisms have been levelled at this procedure (Hänninen et al., 1977; Finch et al., 1979), suggesting that UTP recycling would lead to increased UDP concentration and thus to an overestimate of the UDPGT activity; a microsomal NADH oxidase activity would compete with pyruvate kinase for the NADH, leading to an apparently faster reaction; and finally that the assay is not reliable for the measurement of UDPGT activity towards substrates which are turned over very slowly (<1 nmol min−1 mg−1 protein; Antoine et al., 1988). The method has been validated for certain applications by Colin-Neiger et al. (1984) and Antoine et al. (1988), but Triton X-100 is the only detergent which can be used to activate microsomal UDPGT (as it inhibits the NADH oxidase activity), so the method is not

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suitable for e.g. measuring activities of UDPGT towards certain substrates (e.g. bilirubin) and for following enzyme activity during purification of UDPGTs. Three useful methods utilizing radio-labelled UDPGA have been described. Bansal and Gessner (1980) used thin layer chromatography to separate enzymically-formed glucuronides of a range of substrates from other reaction components. Autoradiography localized the glucuronides and quantitation was achieved by liquid scintillation counting of the appropriate area of the chromatogram. This method has recently been used to study the expression of UDPGT activity in tissue culture cells transfected with cloned UDPGT cDNAs (Mackenzie, 1986a, 1986b; Harding et al., 1988; Jackson et al., 1988). A method utilizing HPLC on polar aminocyano bonded phase to resolve labelled glucuronides formed from microsomal incubations performed in the presence of UDP[14C]GA allowed the measurement of UDPGT activity towards more than 20 aglycones (Coughtrie et al., 1986), with all but one of the glucuronides coeluting. The method is sensitive and widely applicable, but the procedure is time-consuming, requiring the scintillation counting of large numbers of samples or the use of an on-line radioactivity monitor. Tephly’s group have developed a method for the measurement of UDPGT activity during purification, involving the separation of labelled UDPGA and glucuronides on a Sep-pak column prior to liquid scintillation counting (Kirkpatrick et al., 1984). Assay procedures for commonly used substrates (such as 2-aminophenol, 1naphthol, 4-nitrophenol, testosterone and other steroids) can be found in reviews by Dutton (1980), Dutton et al., (1981) and Hallinan (1983). Bilirubin UDPGT activity can be measured by the diazo procedure of Heirwegh et al. (1972) or by the more recent and sensitive assay using radioactive bilirubin and alkaline methanolysis as described by Vermier et al., (1984). Morphine glucuronidation is measured by the radiochemical method as originally described by Del Villar et al. (1979) but preferably by the more recent HPLC method of Svensson et al. (1982), which allows the resolution of the vicinal glucuronides produced from the two enantiomers of morphine—the naturally occurring and pharmacologically active (−)-morphine and the inactive synthetic (+)enantiomer. Numerous other methods for assaying specific substrates have been described —particularly using HPLC—e.g. for arylcarboxylic acids (Hamar-Hansen et al., 1986), menthol (Leroy et al., 1986) and retinoic acid (Miller and DeLuca, 1986); most of the HPLC methods can be easily adapted to measure the activity of UDPGT towards other substrates. For investigations on glucuronidation in vivo or in isolated cells and perfused organs, a number of substrates can be used. However, most of them also show sulfation. In studies in perfused organs and isolated cells it is possible to prevent sulfation by leaving out inorganic sulfate from the perfusion or incubation medium. Sulfation in in vivo animal studies can be prevented by pretreatment

90 CONJUGATION REACTIONS IN DRUG METABOLISM

with a selective inhibitor of sulfation, pentachlorophenol (Meerman et al., 1983) or 2, 6-dichloro-4-nitrophenol (Koster et al., 1982b). Convenient substrates are either highly fluorescent or radio-labelled compounds. Harmol and 4-methylumbelliferone have high fluorescence and the conjugates can be easily separated by thin layer chromatography or HPLC. They can be sensitively quantitated by fluorimetry (Mulder and Hagedoorn (1974) for harmol; Mulder et al., (1985a) for 4-methylumbelliferone). Commercially available radio-labelled substrates are for instance, phenol, paracetamol, morphine, 1-naphthol and 4-nitrophenol. Methodology for their determination and the separation of the conjugates can be found in: Howie et al., 1977; Koster et al., 1981; Svensson et al., 1982; Tremaine et al., 1984; and Meerman et al., 1987, respectively. Details about in vivo perfusion and cell isolation methodology can be found in a volume of The Methods in Enzymology series, Vol. 77 (1981). An assay method for UDPGA has been described by Watkins and Klaassen (1982b). Abbreviations 3-MC β-NF PAH PB PCN UDPG UDPGA UDPGT

3-methylcholanthrene β-naphthoflavone polycyclic arylhydrocarbons phenobarbital pregnenolone-16α-carbonitrile UDP glucose UDP glucuronic acid UDP glucuronosyltransferase References

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 5 Sulfation Gerard J.Mulder1 and William B.Jakoby2 1

Division of Toxicology, Center for Bio-Pharmaceutical Sciences, University of Leiden, 2300 RA, Leiden, The Netherlands.

2

Laboratory of Biochemistry and Metabolism, NIDDK, National Institutes of Health, Bethesda, MD 20892, USA. I.

Sulfation: biological aspects Gerard J.Mulder

5.1.

INTRODUCTION

107

5.2.

PAPS SYNTHESIS AND AVAILABILITY OF INORGANIC SULFATE

108

5.3.

SUBSTRATES AND STEREOSELECTIVITY

111

5.4.

SULFATION IN PERFUSED ORGANS AND ISOLATED CELLS

114

Liver perfusion

114

Isolated hepatocytes

116

Intestinal preparations

119

Kidney

119

Lung

120

Other tissues

120

5.5.

SULFATION IN VIVO

120

5.6.

INHIBITION

127

5.7.

SULFATION AND BIOLOGICAL ACTIVITY OF THE SUBSTRATES

129

Sulfolipids

129

Glycosaminoglycans and glycoproteins

130

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Proteins: tyrosine sulfation

131

Peptides

131

Dopamine and other catecholamines

131

Steroid hormones

132

Bile acids

132

Thyroxine and other compounds

133

Role of sulfation in the toxicity of xenobiotics

133

II.

The sulfotransferases William B.Jakoby

5.8.

INTRODUCTION

134

5.9.

SULFOTRANSFERASES IN XENOBIOTIC METABOLISM

136

5.10.

III.

O-Sulfation: alcohol sulfotransferase (EC 2.8.2.2)

137

O-Sulfation: phenol sulfotransferases (EC 2.8.2.1 and 2.8.2.9)

138

N-Sulfation: amine N-sulfotransferase

139

Trans-sulfation: arylsulfate sulfotransferase

141

ASPECTS OF SULFOTRANSFERASE MECHANISM

141

Arylsulfate sulfotransferase

142

Tyrosine-ester sulfotransferase

143

Chemical and catalytic mechanism

144

Methodological aspects REFERENCES

146

ABBREVIATIONS

146

I. Sulfation: biological aspects 5.1. Introduction Sulfation was discovered by Baumann around 1875 when he isolated phenyl sulfate from the urine of a patient who had been treated with phenol. Although

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sulfation was subsequently extensively studied in vivo, its biosynthetic mechanism remained obscure until 1956 when the structure of the co-substrate of sulfation, adenosine 3′-phosphate 5′-phosphosulfate (PAPS or ‘active sulfate’; the systematic name is 3′-phosphoadenylylsulfate) was reported by Lipmann’s group. The enzymes catalyzing the sulfation reaction, the (cytosolic) sulfotransferases, have since been purified from various sources. They are a family of separate enzymes and isoenzymes with differing but often overlapping substrate specificity. Sulfate conjugation is important in the biotransformation of not only xenobiotics but also of many endogenous compounds such as neurotransmitters and steroid hormones. It may play a regulatory role in the biosynthesis of thyroid and certain steroid hormones. Moreover, proteins and peptides are sulfated, resulting in a possible change in their properties. Sulfation occurs in animals, fungi and bacteria. In vivo, PAPS is present in very low concentrations but can be synthesized very rapidly. In the overall process of sulfation the availability of the PAPS precursor, inorganic sulfate, may become rate-limiting. For many xenobiotics, sulfation seems to be a detoxifying reaction. However, a number of compounds are converted into highly labile sulfate conjugates that form reactive intermediates which have been implicated in carcinogenesis and tissue damage. In the first part of this chapter, PAPS generation and overall sulfation in vivo and in vitro will be described. In the second part, the properties of the sulfotransferases will be discussed. The literature on sulfation has been extensively reviewed by Mulder (1981, 1984) and Jakoby et al. (1984). For a detailed treatment of the literature to 1983, the reader is referred to these reviews. 5.2. PAPS synthesis and availability of inorganic sulfate The group-donating co-substrate for sulfation is PAPS (Figure 5.1). It is formed in a two-step reaction from ATP and inorganic sulfate. The effective enzymes, ATP sulfurylase (EC 2.7.7.4) and APS kinase (EC 2.7.1.25), are present in the cytosol of the cell. Sulfate activation has been reviewed by Mulder (1981); recent reports on the activating enzymes are by Seubert et al. (1985), Brion et al. (1987), Geller et al. (1987), Hommes et al. (1987), and Renosto et al. (1987). Inhibition of sulfate activation by chlorate resulted in inhibition of sulfation (see Section 5.6).

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Figure 5.1. Structure of PAPS.

The equilibrium of the first step is very unfavourable in the forward direction, and the equilibrium concentration of PAPS in the cell is rather low. However, the rate of PAPS biosynthesis can be high if it is utilized at a high rate; whereas the PAPS concentration in the liver is 30–70 nmol g−1 liver, its rate of biosynthesis can be as high as 100 nmol min−1 g−1 of liver, as calculated from the steady-state sulfation rate of harmol in the perfused rat liver. That rate could be sustained for at least 60 min (Pang et al., 1981). PAPS is probably synthesized in every animal cell. Its concentration in the liver is the highest of all organs tested (Hazelton et al., 1985; Brzeznicka et al., 1987). In rat kidney, the PAPS concentration was higher in the cortex than in the medulla (Hjelle et al., 1986). PAPS may also be synthesized in intact human blood platelets since these cells could sulfate dopamine, whereas dog platelets could not (Toth et al., 1986; Toth and Elchisek, 1987; Khoo et al., 1988a, 1988b); inorganic sulfate in the incubation medium increased the rate of sulfation. Human platelets contain a very active 3′nucleotidase that breaks down PAPS to APS (Roth et al., 1986; Ramaswamy and Jakoby, 1987c). No sex differences in the concentration of PAPS were observed in rat or dog liver (Brzeznicka et al., 1987). A genetic deficiency in the rate of PAPS biosynthesis, which is most pronounced in the liver, has been identified in brachymorphic mice; the same deficiency found in cartilage is probably responsible for the brachymorphism

110 CONJUGATION REACTIONS IN DRUG METABOLISM

(Sugahara and Schwartz, 1982a, 1982b). It leads to decreased sulfation of compounds such as 4-nitrophenol in vivo (Lyman and Poland, 1983). PAPS, formed in liver cytosol, can be transported into the Golgi system by a carrier (Perez and Hirschberg, 1986). There it is used for sulfation of lipids or proteins. In insects, PAPS is also formed in the cytosol (Isaac et al., 1982) but in Euglena the sulfate-activating system is present in mitochondria (Saidha et al., 1985). Formation of PAPS in rat brain slices was turned off by depolarizing agents (Brion et al., 1987; Gulat-Marnay et al., 1987). The rate of PAPS synthesis in isolated hepatocytes is linearly related to ATP concentration (Aw and Jones, 1985). Surprizingly, however, PAPS concentration in vivo was not readily decreased by inhibitors of mitochondrial energy production such as 2, 4-dinitrophenol or antimycine A (Dills and Klaassen, 1986a). In fact, 2, 4-dinitrophenol caused a pronounced increase of hepatic PAPS, as did two inhibitors of sulfation, pentachlorophenol and 2, 6dichloro-4-nitrophenol (see Sections 5.6 and 5.10). It has not been determined, however, whether these compounds affect PAPS turnover. As expected, substrates for sulfation, e.g. salicylamide, 1-naphthol or paracetamol, decreased hepatic PAPS (Hjelle et al., 1985; Dills and Klaassen, 1986a, 1986b). Ethionine and fructose had the same effect in rat liver in vivo and in isolated hepatocytes (Dills and Klaassen, 1986b); they tended to decrease the excretion of paracetamol sulfate. Chronic feeding of ethanol to rats tended to decrease PAPS biosynthesis in the perfused liver as reflected in a decreased sulfation rate (Reinke et al., 1986). Ethanol (10 mM), incubated with isolated hepatocytes, however, did not affect sulfation (Sundheimer and Brendel, 1984); Efficient synthesis of PAPS requires sufficient inorganic sulfate. Indeed if sulfate is left out of the incubation medium of isolated cells or of the perfusion medium, sulfation is almost completely abolished; only sulfate arising from sulfoxidation of endogenous L-cysteine will be available as long as cysteine is present. When serum sulfate becomes depleted, an infusion of inorganic sulfate rapidly increases the sulfation rate (Galinsky and Levy, 1981; Krijgsheld et al., 1982b). The apparent Km for sulfate in the sulfation of various substrates in isolated hepatocytes is of the order of 0.3–0.5 mM (Koike et al., 1981; Schwarz, 1984; Sundheimer and Brendel, 1984; Sweeny and Reinke, 1988). In isolated guinea pig gut epithelial cells it may be somewhat higher (Schwarz and Schwenk, 1984). In the perfused rat liver, a Km of 0.4–0.8 mM was found (Mulder and Keulemans, 1978). Based on in vivo experiments in the rat in which steady-state harmol sulfation rates were measured, a Km of ca. 0.3 mM could be calculated for serum sulfate (Krijgsheld et al., 1982a). Most likely sulfate is rapidly taken up in cells by carrier-mediated transport (Mulder, 1984). Changes in such transport may be induced by diseases, e.g. fibroblasts from cystic fibrosis patients had a higher Vmax for sulfate uptake (Elgavish and Meezan, 1988).

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L-Cysteine, D-cysteine (Figure 5.2) or L-methionine can be used both in vivo and in vitro as the precursor for inorganic sulfate (Schwarz, 1980; Krijgsheld et al., 1981a; Glazenburg et al., 1983, 1984; Sundheimer and Brendel, 1984; Humphries et al., 1988). Therefore, high glutathione turnover, also requiring Lcysteine, may affect sulfation in vivo (Galinsky, 1986). Surprizingly, 4nitrophenyl sulfate supported sulfation in a sulfate-free incubation medium almost as efficiently as sulfate itself. It provided rates of sulfation in freeze-thawed permeabilized cells that were much higher than those in intact cells in the presence of sulfate (Schwarz, 1984). This may be due to the transfer of the sulfuryl group from 4-nitrophenyl sulfate to a different acceptor substrate (see section 5.10, this chapter). In vivo, inorganic sulfate in blood is immediately available for sulfation in the tissues (Mulder and Scholtens, 1978; Waschek et al., 1986). Serum sulfate varies among species from 0.3 mM in man to 2.4 mM in the goat, and shows diurnal variation (Krijgsheld et al., 1980). It can be decreased by a diet low in sulfurcontaining amino acids (Krijgsheld et al., 1981b; Glazenburg et al., 1983) or by administration of a high dose of a substrate for sulfation such as paracetamol (Krijgsheld et al., 1981b; Lin and Levy, 1981, 1983a). Paracetamol also causes a decrease of sulfate in cerebrospinal fluid in the rat (Morris and Levy, 1984). Fasting does not lead to decreased serum sulfate, presumably because mobilized cysteine from protein catabolism is continuously oxidized. During periods of reduced sulfate availability, conjugation of phenols shifts from sulfation to glucuronidation (Glazenburg et al., 1984; Jung, 1985). Salicylate also decreases serum sulfate due to increased urinary sulfate excretion (de Vries et al., 1985; Morris et al., 1988). Sulfate depletion may occur locally in the intestinal mucosa when a substrate for sulfation is given orally, i.e. if it is conjugated in the gut mucosa. In that case, oral administration of inorganic sulfate may increase first-pass sulfation locally, even if inorganic sulfate in serum had shown no signs of depletion. 5.3. Substrates and stereoselectivity The most common acceptor group for sulfation is the hydroxyl group in phenols (reviewed by Mulder, 1982), alcohols or (N-substituted) hydroxylamines. In addition, sulfation of thiols or amines, resulting in thiosulfates and sulfamates, respectively, has been noted. Almost without exception, there is competition with glucuronidation for the same acceptor group, especially for the phenolic group. O-Methylation may also compete, particularly for catecholamines. Double conjugates can be formed by sulfation subsequent to another conjugation as in disulfates, sulfate-glucuronide conjugates (Musey et al., 1979; Roy et al., 1987a), and sulfate-methyl-conjugates (Qu et al., 1983).

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Figure 5.2. Effect of intravenous injections of L- or D-cysteine on plasma sulfate level in the rat. Rats with a low plasma sulfate (due to a special diet) received 1.5 mmol kg-1 of Lcysteine (O), D-cysteine (?) or sodium chloride (?) at 90 min. (Taken from Glazenburg et al., 1984, with permisison from Pergamon Press.)

Structure activity relationships for series of phenols towards sulfation will be discussed in Section 5.10 of this chapter. In the cell, sulfation of xenobiotics occurs in the cytosol. Sufficient lipid solubility is required to enter the cell but high lipid solubility is not needed for sulfation (Figure 5.3; Fry and Paterson, 1985). Nevertheless, for a series of alcohols, the more lipophilic ones have the lower Kms (Section 5.9). Ideal model substrates for sulfation studies in vivo and for isolated cell preparations or organ perfusions should be exclusively sulfated. However, almost every substrate studied as yet is also glucuronidated or methylated to at least a minor extent. One way to overcome this is to use very low doses of the substrate since sulfation generally has a higher affinity than glucuronidation for the same substrate. For example, a tracer dose of a radio-labelled substrate is often almost exclusively sulfated. Alcohols show a tendency for sulfation rather than for glucuronidation. Additional requirements for a good model substrate of a specific conjugation in vivo have been mentioned in Chapter 10, Section 10.2. The pharmacokinetics of many phenols in many species have been studied: 4nitrophenol, harmol, phenol and paracetamol (see Section 5.5). These are sulfated by various forms of the sulfotransferases, and each substrate reflects the activity of a combination of such forms. Some, such as harmol, have the

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Figure 5.3. Rate of glucuronidation (G) and sulfation (S) of a series of substituted phenols in isolated rat hepatocytes as a function of their lipid solubility. P is the octanol/ 0.1 M sodium phosphate, pH 7.4 partition coefficient. Me=methyl; N=Nitro; Cl=chloro. (Data from Fry and Paterson, 1985.)

advantage of being easily detected by their fluorescence. Other substrates are commercially available in radio-labelled form. When conjugation is studied in relation to kinetics of elimination it is important to administer the substrates intravenously (i.v.) because of high firstpass sulfation that occurs in several species when the drug is administered orally (see Section 5.5.). Therefore, results of studies with substrates such as amethyldopa, salbutamol or salicylamide, for which high first-pass sulfation has been demonstrated in man, should be analyzed carefully. Pronounced species differences are observed in conjugation of phenols. The cat almost exclusively forms sulfates from most of these substrates (Williams, 1974; Mulder, 1981). Sex differences are less common but occasionally may be quite large (Mulder, 1986b). Thus, only female rats sulfated tiaramide at the alcoholic group, whereas N-sulfation of tiaramide occurred only in the female mouse (Noguchi et al., 1982; Iwasaki et al., 1986). At present, no substrates that are exclusively sulfated are available for studies in vivo or in isolated cells and organ perfusion. A further complication is that the sulfotransferases are a collection of (iso)enzymes with different but overlapping

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substrate specificity. Individual substrates will utilize only a limited number of these sulfotransferase forms. Little information is available on stereoselectivity of sulfation. Recently, Binder and Duffel (1988) determined the stereoselectivity of a purified sulfotransferase form from rat liver for the 1-phenylethanol enantiomers; there was a three-fold difference in catalytic efficiency. Christ and Walle (1985) studied the sulfation of 4-hydroxypropranolol enantiomers in a postmicrosomal rat liver supernatant of four species. Some stereoselectivity was observed in the hamster (a ratio of 0.62 for the (−)/(+) enantiomers) but not in the rat, a difference that may be due to the contribution of different isoenzymes in these species. Stereoselectivity of sulfation of 3methoxy-4-hydroxyphenyl ethylene glycol was observed in the dog (Murray et al., 1980). 5.4. Sulfation in perfused organs and isolated cells Liver perfusion In the perfused liver, sulfation of many substrates occurs. Work of a more detailed, mechanistic nature has been done with the phenolic substrates 4nitrophenol, harmol, 4-methylumbelliferone (4-methyl-7-hydroxycoumarin) and 7-hydroxycoumarin. If inorganic sulfate is omitted from the perfusion medium, sulfation will be dependent on the small amount of inorganic sulfate generated in the liver (Mulder and Keulemans, 1978). Under these circumstances little sulfation will occur, an event that is compensated for by increased glucuronidation of the substrate. In the recirculating perfused liver a pre-perfusion period is required to wash out sulfate from the liver, after which the perfusion medium should be replaced by fresh medium. In the single-pass perfused liver, hepatic sulfate will be washed out in the initial period, so that sulfation is no longer possible later on (Conway et al., 1982, 1988). The extraction of substrates for sulfation by the perfused rat liver can be very high. For harmol, 7-hydroxycoumarin and 4-nitrophenol, extractions of the order of 90–100% were measured at low substrate concentrations. Sulfation is higher in the periportal area (Zone 1) of the liver than in the centrilobular (Zone 3) area (see Chapter 2 for extensive discussion). This was first shown in a study on the metabolism of phenacetin to paracetamol and subsequent sulfation of the latter in the liver perfusion during normal or retrograde flow (Pang and Terrell, 1981). Cytochrome P-450-catalyzed oxidative metabolism of phenacetin to paracetamol takes place mainly in Zone 3. When paracetamol is generated during normal perfusion, less is sulfated than when it is

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generated during retrograde perfusion. The reason is that during normal flow it leaves the liver immediately after formation in Zone 3 and sulfation gets little chance. However, during retrograde flow, the formed paracetamol goes to Zone 1, the ‘sulfation area’, before it leaves the liver. Studies with N-hydroxy-2acetylarninofluorene, which becomes toxic once it is activated by sulfation (see Section 5.7), confirm this; necrosis is located in Zone 1 of the liver and can be prevented by selective inhibition of sulfation (Meerman and Mulder, 1981). The relative localization of sulfation and glucuronidation in the liver was extensively investigated with harmol (Pang et al., 1981, 1983; Dawson et al., 1985), a substrate of the M-form of phenol sulfotransferase (Wong and Wong, 1985). A number of approaches confirmed the localization of sulfation anterior to glucuronidation: comparison of harmol conjugation in the normal and retrograde flow direction, dependence on flow rate through the liver, and concentration dependence (see Chapter 2). With gentisamide (2, 5dihydroxybenzamide) similar results were obtained. Several pharmacokinetic models have been used to evaluate the results and approach of those studies (see Chapter 2). Sulfate conjugates, such as harmol sulfate, are often more readily excreted into the effluent perfusate than into bile. This transport is most likely carrier-mediated (de Vries et al., 1985). The kinetics of harmol conjugation show anomalous behaviour in glucuronidation, due to competition, by sulfation (Chapter 4). Such anomaly disappears when sulfation is prevented (Conway et al., 1982; Koster et al., 1982b). Conway et al. (1982) came to the same conclusion on the preferential localization of sulfation in Zone 1. They used the micro-light guide technique by which Zone 1 and 3 can be analyzed separately. At low substrate (7hydroxycoumarin) concentration, sulfation is the predominant reaction, while at higher substrate concentration glucuronidation becomes more important, in agreement with the concept of a low Km and low capacity (Vmax) for sulfation and a relatively high Km and Vmax for glucuronidation for the same substrate (Mulder, 1981). Essentially the same results have been observed with harmol (Pang et al., 1981). When 7-ethoxycoumarin was used to generate 7hydroxycoumarin, it was only sulfated, in both zones, because of the higher affinity of sulfation. El Mouelhi and Kauffman (1986) found somewhat higher activity of sulfotransferase activity towards 7hydroxycoumarin in Zone 1 than in the Zone 3 tissue, which had been separated by microdissection techniques from human liver. Sulfatase activity was equally high in both areas. The claim for a futile cycle was made for sulfation/desulfation and was confirmed by work with 4methylumbelliferone sulfate in the perfused rat liver (Anundi et al., 1986): when the sulfate conjugate was added to the perfusion medium, free 4methylumbelliferone could be detected with the micro-light guides, and 4methylumbelliferone glucuronide was formed in the sulfate-free perfusion. Some 6–13% of the incoming substrate was converted to the glucuronide.

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Paracetamol is mainly sulfated at lower concentrations in the perfused liver; the sulfate conjugate is excreted in the perfusion medium rather than in bile (Grafström et al., 1979; Pang and Terrell, 1981). In livers damaged by a high dose of paracetamol (mainly Zone 3 damage) 12 h prior to perfusion, sulfation of paracetamol was unimpaired, presumably because sulfation takes place primarily in Zone 1 (Poulsen et al., 1985). A rather complex picture arises from perfusions with oestrone and some other steroids. Double conjugates are formed through sulfation of the 3hydroxyl group and subsequent glucuronidation at a hydroxyl group in the D-ring. Desulfation also occurs. The perfused guinea pig liver forms more of the glucuronide conjugates and shows a number of other differences when compared with rat liver (Roy et al., 1987a). Similar studies with polyethylene glycols demonstrated that these, too, are readily sulfated in the perfused rat and guinea pig liver and, therefore, should be avoided as solvents for substrates of sulfation (Roy et al., 1987b). 4-Nitrophenol shows both sulfation and glucuronidation during perfusion. Fasting had relatively little effect on sulfation, while glucose addition increased sulfation in the fasted, PB-treated liver (Reinke et al., 1981). In streptozotocininduced diabetic rats, the sulfation rate was somewhat below normal (Morrison et al., 1985b). Pretreatment with 3-MC seemed to enhance sulfation but had a much more pronounced effect on glucuronidation (Hamada and Gessner, 1975). When 4-nitrophenol is low (4 (µM), the extraction by the perfused liver is 100%, with the sulfate conjugate formed almost exclusively. At higher concentrations extraction is less complete and more glucuronidation occurs. If 4nitrophenol is generated from paraoxon, it is mainly sulfated, presumably because the steady-state concentration of 4-nitrophenol remains very low (Sultatos and Minor, 1985). Similar findings were reported for another precursor, 7-ethoxycoumarin, yielding 7-hydroxycoumarin after oxidative metabolism (Conway et al., 1982; Cha et al., 1987). 4-Hydroxybiphenyl preferentially forms the sulfate conjugate under normal conditions, but after PB treatment this shifts to glucuronidation, presumably due to induction of glucuronidation (Kahl et al., 1978). Sulfation of 4hydroxycoumarin decreased in livers taken from rats under chronic alcohol treatment (Hong et al., 1987). A major factor in this decrease is most likely a decreased rate of generating PAPS (Reinke et al., 1986). Isolated hepatocytes Almost every phenol is sulfated to at least a minor extent in hepatocytes if inorganic sulfate or one of its precursors is present. Inorganic sulfate is immediately available when it is added to the incubation medium (Koike et al., 1981). A short incubation period of the cells with sulfate may ensure sufficient

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PAPS availability (Mizuma et al., 1985). The sulfation rate is not influenced by lipid solubility of the substrate (Fry and Paterson, 1985) if it is high enough to allow a rapid entry of the substrate into the cell. Hepatocytes from all species tested are capable of sulfation. Beside the rat, mouse (Moldéus, 1978), guinea pig (Miller and Jollow, 1987), cow (adult and foetal; Suolinna and Winberg, 1985), dog (Bolcsfoldi et al., 1981) and fish (Morrison et al., 1985a) serve as donor. In cultured rat hepatocytes sulfation survives, although it may show a transient decrease during the first days after start of the culture (Fry and Bridges, 1980; Suolinna and Pitkäranta, 1986). Adult human hepatocytes in primary culture showed a rapid loss of sulfation (Grant et al., 1987) but no data beyond 72 h of culture were reported so that an eventual later increase may have been missed. In hepatocarcinoma cell lines, sulfation of thyroid hormones, for instance, occurs but sulfotransferase activities are low (Sorimachi and Robbins, 1978; Wiebel et al., 1980; Sekura et al., 1981b). Human liver slices also can support sulfation (Powis et al., 1987). The presence of albumin in the incubation medium may affect metabolism of substrates that are strongly bound to albumin. Thus, resorufin is so tightly bound that it becomes essentially unavailable for conjugation within the cell (Burke and Orrenius, 1978). Sulfation, obviously, will be dependent on adequate oxygenation. The oxygen concentration at half the maximum velocity is 2.5 µM for paracetamol sulfation. The sulfation rate changes with the ATP/ADP ratio. Compounds that decrease that ratio, such as ethionine or atractyloside, also decrease sulfation so that sulfation rate correlates with the ATP concentration in the cell (Aw and Jones, 1982, 1985). The Km for PAPS in isolated cells is of the order of 4 µM for the sulfation of paracetamol (Sweeney and Reinke, 1988). Phenols are generally preferentially sulfated at low concentration and undergo increasing glucuronidation at higher concentration (Wiebkin et al., 1978; Suolinna and Mäntylä, 1980; Koster et al., 1981). Several substrates, harmol for example, have been extensively investigated, including the kinetic behaviour of the conjugates. In the cases of paracetamol and harmol it could be established that the half maximal concentration (Km) for the acceptor substrate in the incubation was much lower for sulfation than for glucuronidation (Sundheimer and Brendel, 1983; Mizuma et al., 1985): these values for paracetamol were 30 µM for sulfation and 2100 µM for glucuronidation. A number of kinetic parameters have been collected in Table 5.1. However, a caveat is warrented here; especially at low substrate concentration it is often not the conjugation rates (i.e. initial velocities) that are measured but conversion after only one incubation time (e.g. 10 or 30 min). In the latter case the ‘rate’ is probably not an initial rate since as much as 60–100% of the substrate may have been converted during that incubation period.

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Table 5.1. Some kinetic parameters of sulfation in isolated hepatocytes.

a

from the rat unless indicated otherwise. with the assumption that 106 hepatocytes equals 1 mg protein.

b calculated

When a substrate is generated from a precursor, 4-hydroxybiphenyl from biphenyl as an example, it is primarily sulfated when produced at a low rate. However, after induction by 3-MC or PB, the rate of 4-hydroxybiphenyl generation was much higher so that glucuronidation became more important than sulfation (Wiebkin et al., 1978). Similar findings were reported for 4aminophenol as generated from aniline (Evelo et al., 1984) and for 4hydroxybiphenyl generated from 4-methoxybiphenyl (Fry, 1987). When harmol glucuronide is slowly hydrolyzed by ß-glucuronidase in isolated hepatocytes, it is reconjugated by sulfation (Sundheimer and Brendel, 1983). Sulfation is not induced by the common inducers of microsomal drug metabolizing enzymes (Andersson et al., 1978; Moldéus, 1978; Suolinna and Mäntylä, 1980; Moldeus et al., 1982; Tonda and Hirata, 1983). However, 3-MC does increase 4-nitrophenol and resorufin sulfation by isolated cells (Burke and Orrenius, 1978; Tonda and Hirata, 1983), as it does in perfused rat liver (Hamada and Gessner, 1975). Interference with sulfation by several compounds in isolated hepatocytes has been studied; in most cases, the mechanism is unclear. For instance, dibutyryl cAMP and some other nucleotides decreased sulfation but only at extremely high concentrations (Shipley et al., 1986). Similarly, a high concentration of diethyl ether (30 mM) inhibited sulfation severely, although cell viability was not affected (Aune et al., 1984). Ethanol (10 mM) had no effect on harmol sulfation, whereas a 24 h prior fast caused a slight decrease (Sundheimer and Brendel, 1984). Chronic alcohol treatment of rats had no effect on sulfation of paracetamol (Moldéus et al., 1980). In streptozotocin diabetic rats sulfation was little affected, although insulin treatment of donor rats decreased subsequent sulfation in the hepatocytes significantly (Eacho et al., 1981; Grant and Duthie, 1987). An age-related decrease in sulfation of paracetamol was observed between hepatocytes from 2–4 months and 22 to 26-months-old rats (Sweeny and Weiner, 1985).

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Table 5.2. Sulfation of 1-naphthol in isolated cells from the guinea pig.

Intestinal preparations Sulfation may play an important role in first-pass conjugation of many substrates, particularly in man and dog (see Section 5.5). With intact intestinal preparations from the rat, sulfation of 1-naphthol, 7-hydroxycoumarin, salicylamide or paracetamol could be demonstrated (Iwamoto and Watanabe, 1983; Wollenberg et al., 1983; Wollenberg and Rummel, 1984; Pang et al., 1986; Sund and Lauterbach, 1986, 1987). In general, rat intestinal preparations are more active in glucuronidation than in sulfation. The sulfate conjugate is released at both the luminal and the serosal side of the intact intestinal preparation; carrier transport is involved. Moreover, different results may be found if the substrate is added either in the lumen of the gut or in the blood of vascularly perfused intestine. When 1-naphthol is added at the luminal side, sulfation is higher than when it is added in the perfusing blood (Sund and Lauterbach, 1986). Addition of inorganic sulfate stimulated sulfation; again it differed whether it was added at the luminal or the serosal side (Sund and Lauterbach, 1987). Sulfation in mucosal sheets from normal human jejunum or ileum has been studied in Ussing chambers (Rogers et al., 1987a). Ethinyloestradiol was readily sulfated, but paracetamol much less so. Sulfation also can be studied in isolated intestinal cells (Dawson et al., 1983; Schwarz and Schwenk, 1984; Schwenk and Locher, 1985). Cells from the jejunum sulfated 1-naphthol more actively than those from the ileum (Table 5.2). A high concentration of inorganic sulfate was required for maximum sulfation, so that Schwarz and Schwenk (1984) suggested that oral sulfate intake would increase sulfation of phenols, as has been observed indirectly in man (see Section 5.5). Kidney The perfused rat kidney catalyzes sulfation of various compounds. At increasing substrate concentration of 1-naphthol and 4-dimethylaminophenol in the perfusate, the balance between sulfation and glucuronidation shifts towards more glucuronidation (Elbers et al., 1980; Redegeld et al., 1988). The 1-

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naphthyl sulfate produced is recovered in the perfusion medium rather than in urine, one of the reasons being that it is strongly bound to protein in the perfusate. The isolated perfused human kidney can also sulfate phenol and 4nitrophenol (Diamond et al., 1982), as can slices of human kidney (Powis et al., 1987). Highest activity of sulfation is expected in proximal tubular cells as determined by microdissection studies of rabbit kidney (Hjelle et al., 1986). Isolated kidney cell preparations, containing mainly (proximal) tubular cells, catalyze sulfation (Dawson et al., 1983; Schwenk and Locher, 1985). Pretreatment with 3-MC increased sulfation of paracetamol (Emslie et al., 1981). Lung The isolated perfused rat lung (Hogg et al., 1981) as well as cell preparations catalyze phenol sulfation. Surprizingly, L-cysteine was a better sulfate source than inorganic sulfate (Dawson et al., 1983) in cells. Isolated lung macrophages did not sulfate phenol (Hogg et al., 1981). Human bronchus in short-term explant culture synthesized 1-naphthyl sulfate (Gibby and Cohen, 1984). In tumour tissue from human lung, sulfation of 1-naphthol was almost absent (Mehta et al., 1981). The same was observed in cultured human tumourous colon (Cohen et al., 1983). Other tissues Cultured human aorta smooth muscle or endothelial cells could sulfate 3hydroxy-benzo[a]pyrene; PAPS generation appeared to be rate-limiting (Yang et al., 1986). Human adrenocortical cells in culture contain a dehydroepiandrosterone sulfotransferase that is regulated by AMP and protein kinase C (McAllister and Hornsby, 1988). Mouse and rat skin strips also supported some sulfation (Moloney et al., 1982). 5.5. Sulfation in vivo Sulfation in vivo has been studied mainly with phenolic substrates. In most species, sulfation and glucuronidation of those substrates occur side by side although major species differences exist (Williams, 1974). The extremes are the cat, which shows very high sulfation, and the rabbit or pig, in which it is very low. In general, sulfates are excreted predominantly in urine and relatively less in bile, although this is species dependent; harmol sulfate was excreted in high amounts in bile in the cat and the mouse (Mulder and Bleeker, 1975). Several factors determine the balance between sulfation and glucuronidation (Mulder,

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1984; Mulder et al., 1984a). A major determinant is the dose; at high doses glucuronidation is usually the most important, while at low doses sulfation predominates. Usually the dose used in man is much lower than that in animals; for this reason alone, sulfation would be expected to be more important in man than in other species. The dose dependence can be found in many species and for many phenols (Koster et al., 1981; Hjelle and Klaassen, 1984;, Savides et al., 1984; Sund, 1987). Although sulfate depletion may occur at high dosage, it could be shown that this is not the main cause of the shift of balance. Thus, a shift from sulfation to glucuronidation in the conjugation of phenol in the rat, when the dose increased from 13 to 266 µmol kg−1 (i.v.) was not due to decreased availability of sulfate but could be ascribed to differences in Kms for glucuronidation and sulfation (Weitering et al., 1979). At very high doses of a substrate, of course, sulfate depletion does contribute (see below). With harmol, infused at various rates to steady state, dose dependence has been clearly demonstrated (Koster et al., 1981; Figure 5.4). Extraction by rat liver in vivo is about 60% (Mulder et al., 1984b). Based on the kinetics of harmol elimination from blood it was concluded that the clearance of harmol was much higher than liver blood flow; therefore, appreciable extrahepatic conjugation, in particular sulfation, was present. One such site is the kidney, as shown in vivo in the rat and the chicken (Diamond and Quebbeman, 1981; Tremaine et al., 1984). Hydrolysis of sulfate conjugates also may occur in vivo (Tremaine et al., 1984), although the salicylamide sulfate conjugate was not hydrolyzed in the dog (Waschek et al., 1986). Very high clearances were measured for phenol and 1-naphthol, indicating that conjugation had to occur to a large extent extrahepatically, particularly in the lung (Cassidy and Houston, 1984; Mistry and Houston, 1985). However, these investigators did not report data on sulfation and glucuronid ation separately, so that the role of sulfation cannot be evaluated directly. In the dog, Waschek et al. (1984) and Fielding et al. (1986) found extensive sulfation of salicylamide at low infusion rates; at trace concentrations a clearance equal to cardiac output was found. The lung seemed to contribute greatly. At higher doses, the extrahepatic contribution fell drastically. Blood was sampled at various sites to evaluate the role of a number of organs. Depletion of inorganic sulfate in blood clearly was not the reason for saturation of salicylamide sulfation observed in the dog. An analysis of the data gives confusing results since a steady state is not really reached (Waschek et al., 1988). Blood platelets, an unusual site of conjugation, have a capacity for sulfation (Khoo et al., 1988b). Whether they contribute to overall sulfation capacity is not known. Thus, whether the arterio-venous difference in adrenaline-sulfate measured in the human forearm is due to sulfation by platelets or takes place at other sites in the forearm (Joyce et al., 1982) is unknown. Attempts have been made to correlate sulfotransferase activities in platelets with sulfation activity of

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Figure 5.4. Dependence of sulfation and glucuronidation on substrate infusion rate at steady-state pharmacokinetics in the rat in vivo. Harmol was infused at various rates to a steady state of metabolite excretion. The excretion rate, bile+urine, reflecting synthesis rate, for the glucuronide (vg) and the sulfate (vs) are given. (Adapted from data of Koster et al., 1981.)

the whole individual. The results are somewhat controversial, which is not surprizing: the percentage of an oral dose that is sulfated (a multi-step process!) is compared to sulfotransferase activities in the platelet fraction. A correlation between two different forms of sulfotransferase in the platelets and sulfation of paracetamol was found by Reiter and Weinshilboum (1982), but not by Bonham Carter et al. (1983). The percentage of the dose that is sulfated or glucuronidated differs widely among volunteers, including monozygous twins (Nash et al., 1984). Additional studies were done with α-methyldopa, for which most of the dose is excreted in urine as sulfate conjugates (Campbell et al., 1984). A correlation between platelet sulfotransferase activity and the percentage of α-methyldopa sulfated was only observed when volunteers also took sodium sulfate orally (Campbell et al., 1985). An analysis of the data is not easy; complicating the interpretation is that oral sulfate decreases absorption of α-methyldopa (Campbell et al., 1988). Moreover, different forms of these enzymes behave differently; the activities of the thermostable form in brain and in platelets taken from surgical lobectomy patients correlated, but those of the thermolabile form did not (Young et al., 1985). Familial variation in certain

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platelet sulfotransferase forms suggested genetic differences (van Loon and Weinshilboum, 1984). An important site of sulfation, especially after oral administration, is the intestine (George, 1981). Several drugs (e.g. isoprenaline, steroid hormones, salicylamide or xamoterol) are affected (Shibasaki et al., 1981; Rogers et al., 1987b; Groen et al., 1988). The extent of intestinal sulfation differs greatly among species. Competition for intestinal sulfation between substrates may severely influence the area under the curve (AUC) for the compound in blood: salicylamide prevented first-pass sulfation of isoprenaline so that the AUC in the dog more than doubled (Bennett et al., 1975). In women one gram of paracetamol inhibited first-pass sulfation of ethinyl oestradiol (Rogers et al., 1987b), although paracetamol itself underwent very little sulfation during first pass in the gut (Clements et al., 1984). An interaction between fenoldopam, which in man is converted to the 7- and 8-sulfate conjugates, and paracetamol may be due similarly to interaction at the first-pass intestinal level. High infusion rates of fenoldopam may cause a local sulfate depletion in the gut mucosa (Ziemniak et al., 1987, 1988). Such local sulfate depletion might explain why oral dosing of sulfate decreased the bioavailability of salicylamide in the dog (Waschek et al., 1985). The sulfation rate in vivo is dependent on serum sulfate. Krijgsheld et al. (1982a), using rats that had low serum sulfate due to a special diet, estimated that the Km of serum sulfate for sulfation in vivo was 0.3 mM (Figure 5.5). Steady-state rates of harmol sulfation at various infusions of inorganic sulfate were measured in these experiments. In the dog a decrease of serum sulfate from 0.9 to 0.3 mM also led to decreased sulfation of salicylamide (Waschek et al., 1985) but a much higher than physiological sulfate level did not increase sulfation (Fielding et al., 1986). Serum sulfate is immediately available for sulfation in the organs in vivo since i.v. injected 35S-labelled sulfate mixes very rapidly with the sulfate pool from which sulfation takes place (Mulder and Scholtens, 1978; Waschek et al., 1986). Inorganic sulfate in blood may be depleted by high doses of substrates for sulfation both in experimental animals and in man; even 0·75–1·5 g of paracetamol (0·7–0·14 mmol kg−1) in man leads to a decrease of serum sulfate (Morris and Levy, 1983; Hendrix-Treacy et al., 1986). In rats, virtually complete sulfate depletion results when 2 mmol kg−1 paracetamol are administered (Galinsky and Levy, 1981; Lin and Levy, 1981); sulfate in cerebrospinal fluid is also decreased (Morris et al., 1984). As a consequence, the rate of sulfation of paracetamol decreases in a dose-dependent way, which can be prevented by administration of inorganic sulfate (Lin and Levy, 1986). Both L- and Dcysteine when administered rapidly lead to increased sulfation in rats that are depleted of sulfate (Glazenburg et al., 1984).

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Figure 5.5. Dependence of sulfation in vivo on plasma sulfate concentration. Harmol was infused at various plasma sulfate levels (resulting from infusion of inorganic sulfate to steady state), and the steady state excretion (bile+urine) of harmol sulfate (reflecting rate of synthesis) is given for six rats. (Taken from Krijgsheld et al., 1982a, with permission from Pergamon Press.)

Mice that are deficient in PAPS synthesis, the brachymorphic mice, excrete a lower percentage of paracetamol as sulfate conjugate than normal mice (Lyman and Poland, 1983). A major difference between sulfate and glucuronide conjugates is that sulfates are more tightly bound to protein in blood. Whereas 4methylumbelliferone was 90% bound in rat plasma, 97% was bound for the sulfate conjugate (Mulder et al., 1985b). Paracetamol sulfate was 36% bound but the glucuronide only 4% (Wang et al., 1985). A similar difference was found for chenodeoxycholate (Bartholemew and Billing, 1983). Sulfate conjugates are excreted mostly in urine, although appreciable biliary excretion can occur in certain species (Møller and Sheikh, 1983; Mulder et al., 1985b). Carrier-mediated transport of sulfate conjugates from the hepatocyte to blood and carrier-mediated, active tubular secretion of sulfate conjugates occur (Morris and Levy, 1984; de Vries et al., 1985). The routes of biliary or urinary excretion in the rat are compensatory (Jorritsma et al., 1979). In man, extensive biliary excretion of the sulfate conjugate of 17-α-ethinyloestradiol was found (Maggs et al., 1983). If a sulfate conjugate is given i.v. it may be less readily excreted in bile than when it is synthesized within the liver; administration of

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the morphine sulfate conjugate i.v. in the rat or the cat resulted in very little excretion in bile as compared to the glucuronide (Smith et al., 1973). In general, the glucuronides are excreted in bile to a much higher extent than the sulfates. This implies that in studies on the rate of metabolism of a substrate for both sulfation and glucuronidation, both biliary and urinary excretion should be followed. If only urine is collected, the percentage of sulfation will be overestimated because any glucuronide formed will preferentially be excreted in bile. Upon hydrolysis in the gut the aglycone may be reabsorbed, sulfated, and only then excreted in urine as sulfate conjugate, as occurs for paracetamol (Siegers et al., 1983). Therefore, the primary metabolite, the glucuronide, will be missed, and only the secondary metabolite, the sulfate, will be collected in urine. A number of drugs have been used in man in various studies on sulfation. Investigations on diflunisal revealed that during chronic administration the percentage of the dose that became sulfated increased from 10 to 30%. The dosedependent kinetics of sulfation were determined in volunteers (Loewen et al., 1986, 1988). The sulfate conjugate is not excreted in bile (Verbeeck et al., 1988). High first-pass sulfation of salbutamol by the intestinal mucosa was observed after oral administration. Renal clearance of the parent compound was much higher than that for the sulfate conjugate, presumably because the latter is more extensively protein bound (Morgan et al., 1986). A similar difference in renal clearance, for the same reason, was observed for triamterene and the sulfate conjugate of its hydroxylated metabolite (Hasegawa et al., 1982). The pharmacokinetics of salicylamide in human volunteers (Levy and Matsuzawa, 1967) showed an increased glucuronide/sulfate conjugate ratio at increasing dose. Local sulfate depletion in the gut mucosa leading to decreased first-pass sulfation may have been the cause. Sulfation of α-methyldopa in newborns is high (Cummings and Whitelaw, 1981), and sulfation of paracetamol in neonates is more extensive than in elderly people (Miller et al., 1976). This is in agreement with enzyme activities measured in human foetal and adult tissues (Steiner et al., 1982; Pacifici et al., 1988). Paracetamol kinetics have been extensively studied, both in volunteers and in acutely poisoned patients (Slattery and Levy, 1979; Prescott, 1980). Paracetamol has a short half-life of elimination from the blood (about 2 h), which is prolonged at higher dosage and under conditions of hepatotoxicity. Sulfation is saturated at high doses, so that a greater percentage of paracetamol is converted to the toxic metabolite. A high renal clearance of the sulfate conjugate suggests active secretion into urine. Patients taking anticonvulsants or rifampicin showed no changes in paracetamol sulfation; similarly, smoking, a charcoal-broiled diet (Prescott et al., 1981; Anderson et al., 1983; Miners et al., 1984), gender, or the use of oral contraceptives did not affect paracetamol sulfation (Miners et al., 1983).

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Pharmacokinetics of paracetamol sulfation in the rat have been studied in detail by Watari et al. (1983). They derived a lower overall in vivo plasma Km for sulfation (0·1 mM) than for glucuronidation (0·9 mM), whereas the Vmax for sulfation was two-fold that of glucuronidation. A wide dose range was studied, both with paracetamol and its separate conjugates, to derive a detailed pharmacokinetic model. Little paracetamol sulfate was excreted in bile. The biliary excretion of the sulfate is influenced by anaesthesia (Watkins et al., 1984). In the guinea pig, sulfation of paracetamol is limited mainly by saturation of the sulfotransferases (Miller and Jollow, 1987). Minor age and sex differences were noted in paracetamol sulfation in the rat (Green and Fischer, 1981; Galinsky et al., 1986). Sex differences were observed in organ cytosolic fractions with some substrates but not with others (see Mulder, 1986b, for a review). For sulfation of phenol, 4nitrophenol or harmol, no sex difference was found in the rat, while there was a pronounced difference for N-hydroxy-2-acetylaminofluorene (Meerman et al., 1987). Sex differences in tiaramide sulfation in vivo and in vitro were related to changes in sulfotransferases (Lockley et al., 1982; Noguchi et al., 1982; Iwasaki et al., 1986); age dependence also plays a role (Hobkirk et al., 1983; Kane and Chen, 1987). During pregnancy, sulfation of paracetamol was somewhat decreased in the rat (Lin and Levy, 1983b), although sulfate availability was unaffected. In pregnant mice, however, sulfation was increased due to an increase in sulfotransferase and a decrease in glucuronidation (Larrey et al., 1986). In pregnant women no difference in paracetamol sulfation was observed (Miners et al., 1986). Detailed studies on the metabolism of paracetamol in the maternalplacental-foetal unit in sheep (Wang et al., 1985, 1986) revealed relatively high sulfation in the foetus, while glucuronidation matured late. Little work has been done on interactions between the diet and sulfation. In Section 5.2, studies are mentioned in which a low-protein diet led to reduced PAPS availability and, therefore, reduced sulfation. In a study of rats fasted for 72 h, sulfation of harmol was little affected; presumably, the catabolic state provided enough inorganic sulfate for conjugation (Mulder et al., 1982). In isolated hepatocytes from rats fasted 72 h, sulfation of iodothyronine was also unaffected (Otten et al., 1984). Dietary changes (lipid, carbohydrate and protein) had only minor effects on salicylamide sulfation in human volunteers (Morad, 1982). Sulfation of paracetamol was little affected in streptozotocin diabetic rats (Price and Jollow, 1982; Siegers et al., 1985), although bile salt sulfation increased (Kirkpatrick and Kraft, 1984). In patients with Gilberts syndrome, who have a benign disorder of bilirubin glucuronidation, sulfation of paracetamol was unchanged (Ullrich et al., 1987). A hindlimb ischaemia model in the rat showed that such trauma resulted in reduced sulfation (Griffith et al., 1985).

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The role of sulfation in the conjugation of dopamine and other catecholamines, thyroxine, steroids and bile salts will be discussed in Section 5.7. 5.6. Inhibition Inhibition of sulfation can be achieved by inhibition of the transferases or by prevention of PAPS biosynthesis. The latter can be effected by inhibition of sulfate activation or by decreasing sulfate availability (see Section 5.2). Most information is available from direct inhibition of the sulfotransferases. Obviously, substrate competition is a possibility, but is not effective unless a compound has a very high affinity and very low Vmax. Salicylamide, which has been used frequently in isolated cell experiments, inhibits sulfation of other substrates (Moldéus et al., 1979; Zaleski et al., 1983) but is essentially ineffective in vivo; it is eliminated too rapidly by (mainly) sulfation (Mulder and Scholtens, 1977). Much more effective are 2, 6-dichloro-4-nitrophenol (DCNP) and pentachlorophenol (PCP) (Mulder and Scholtens, 1977). These compounds are dead-end inhibitors of most sulfotransferases, with] values of 0·1–1 µM (Duffel and Jakoby, 1981; Campbell et al, 1987a) and very effectively inhibit the sulfation of many compounds both in vivo and in vitro (Mulder and Scholtens, 1977; Koster et al., 1982a). However, PCP is an effective substrate for amine sulfotransferase, an enzyme which may include in its specificity amines, phenols and alcohols (Ramaswamy and Jakoby, 1987a, 1987b). Many other phenols have been tested as inhibitors (Koster et al., 1979), but none performed better than PCP and DCNP. Their pharmacokinetics and effectivity, as well as characteristics of the inhibitory effect, have been defined (Koster et al., 1982a; Meerman et al., 1983). When they are given intravenously, they immediately arrest sulfation. The effect is reversible when the compounds can be removed, as in the perfused liver (Figure 5.6; Koster et al., 1982a; Mulder, 1986a). PCP and also DCNP have become important tools in the evaluation of the role of sulfation in carcinogenesis by N-hydroxylated aromatic amines or benzylic alcohols such as 1′-hydroxy safrole (Meerman et al., 1980, 1981; Meerman and Mulder, 1981; Boberg et al., 1983, 1987; Kedderis et al., 1984; Graichen and Dent, 1985; Meerman, 1985; Ringer et al., 1985; Fennell et al., 1985; Lai et al., 1985, 1987; Delclos et al., 1986; Furlong et al., 1987; Ringer and Norton, 1987; Wiseman et al., 1987; Yamazoe et al., 1987; Okuda et al., 1988). PCP can be given as sodium salt for a prolonged period in the food or in the drinking water. PCP and DCNP are not inhibitory with every substrate (Mulder, 1984); PCP hardly inhibited sulfation of 7hydroxymethyl-12methylbenz[a]anthracene (Surh et al., 1987)— dehydroepiandrosterone was a more effective inhibitor. Furthermore, these

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Figure 5.6. Inhibition of sulfation by 2, 6-dichloro-4-nitrophenol (DCNP) in the singlepass perfused rat liver. The single-pass perfused rat liver was perfused with a constant concentration of 3H harmol, and the production of harmol sulfate (hs) and harmol glucuronide (hg) was followed (bile+effluent perfusate). From 60 to 120 min 20 µM DCNP was included in the perfusion medium. (From Koster et al., 1982a, with permission from Pergamon Press.)

inhibitors may have other effects. PCP inhibits the acetyl-CoA-dependent acetylation of N-hydroxyarylamines although sulfation is more sensitive (Flammang and Kadlubar, 1986; Shinohara et al., 1986). Both DCNP and PCP led to increased PAPS levels in rat liver (Dills and Klaassen, 1986a), whereas inhibition of PAPS synthesis in brain slices by DNCP was reported (Gulat Marnay et al., 1987). In isolated guinea pig mucosa, DCNP also had effects on the efflux of conjugates from the cells (Sund and Lauterbach, 1987). Sulfotransferase forms respond differently to DCNP: the ‘M’ form is relatively insensitive to DCNP, while the ‘P’ form is very sensitive (Rein et al., 1982). Dopamine sulfation in intact platelets is inhibited by DCNP (Toth and Elchisak, 1987); also the sulfation of the octapeptide cholecystokinin could be inhibited by DCNP in the rat in vivo (Giorgi and Meek, 1985). Sulfation of many phenols in vivo in the perfused liver or in isolated cells can be inhibited almost completely by PCP or DCNP (Koike et al., 1981; Otten et al., 1983, 1984; Fayz et al., 1984; Mulder et al., 1985a; Fry and Patterson, 1987; Miller and Jollow, 1987; Sund and Lauterbach, 1987; de Herder et al., 1988).

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Sulfation of proteins is not inhibited by either DCNP or PCP, but is, in intact cells, by chlorate. Chlorate is an inhibitor of ATP sulfurylase activity (Baeuerle and Huttner, 1986; Hortin et al., 1988; Humphries et al., 1988) and is effective in inhibiting sulfation in the perfused liver (Roy et al., 1988). Several PAPS analogues were quite inhibitory toward the purified human sulfotransferases (Rens-Dominano and Roth, 1987). The rat brain galactosylcerebroside sulfotransferase could be inhibited by a number of triazone aromatic dyes, possibly because the dyes fit the PAPS binding site (Zaruba et al., 1985). More or less potent inhibitors of sulfation seem to be present in tissue extracts, in food and in red wine but remain unidentified (Anderson and Weinshilboum, 1979; Littlewood et al., 1985; Gibb et al., 1987). A number of other compounds have no effect on sulfation in vivo, e.g. D-galactosamine or diethyl maleate (Moldéus et al., 1979; Siegers et al., 1980; Zaleski et al., 1983). Ascorbate inhibits the sulfation of 4-hydroxybiphenyl in isolated hepatocytes but only in extremely high concentrations (Patterson and Fry, 1983); it is an acceptor substrate for alcohol sulfotransferases (EC 2.8.2.2.). Inhibition of steroid sulfotransferases was studied by Horwitz et al. (1986) and Rozhin et al. (1986) with a large series of 2- and 4-substituted estra-1, 3, 5(10) trien-17β-ols. The 4-nitro derivative was most effective with a Ki of 2 µM. Inhibition due to depletion of the co-substrate is possible but is not easily controlled. Rapid depletion will occur with a high dose of a substrate; sulfate levels will return to control levels when the substrate has been eliminated (see Section 5.5). A diet low in cysteine and methionine gave good results in the studies of Krijgsheld et al. (1981b). Adaptive effects to such a diet may occur. Control experiments in which inorganic sulfate is infused in the diet-restricted animals would allow evaluation of such changes in order to distinguish them from those due to sulfate depletion. 5.7. Sulfation and biological activity of the substrates Many low molecular weight, endogenous compounds are sulfated. Their biotransformation usually terminates the biological action of the substrate and leads to excretion of the conjugate in urine. Besides this mainly catabolic role of sulfation, there is an anabolic role, which applies mainly to sulfolipids or to macromolecules such as sulfated glycosaminoglycans, proteoglycans and protein. Sulfolipids Sphingolipids, e.g. cerebrosides, are often sulfated. Bis-sulfated lipids may occur when more sugar residues are present, as in ceramides (Tadano et al., 1982). These are synthesized in many organs including brain, gastric mucosa, kidney and

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salivary glands. In some cases, the sulfotransferases involved have been purified (Benjamins et al., 1982; Tennekoon et al., 1983, 1985; Slomiany et al., 1983; Slomiany et al., 1984; Lingwood et al., 1986). Sulfatides reduce mobility in model membranes due to strong side to side headgroup interactions (Cestaro et al., 1983). Cholesterol 3-sulfate is another sulfolipid that may accumulate as it does in patients with recessive X-linked ichthyosis (Muskiet et al., 1983) or during squamous differentiation of tracheal epithelial cells in vitro (Rearick et al., 1987). Glycosaminoglycans and glycoproteins A high degree of sulfation occurs at the glycosyl residues in glycosaminoglycans or glycoproteins. These macromolecules may be excreted by the cell (cultured in vitro or in vivo), or they may be incorporated into cell membranes. Their sulfation can be demonstrated by the incorporation of 35S-sulfate. The presence of these sulfated macromolecules has been demonstrated in most cell types or tissues, including ovary, kidney, thyroid, endothelial cells, chrondrocytes and hepatoma (Heifetz et al., 1982; Dennis et al., 1984; East and Dean, 1984; Edge and Spiro, 1984; Kato and Gospodarowicz, 1984). Sulfation is also a common feature of several pituitary releasing factors that include (the sugar moiety of) lutropine (LH), follitropine (FSH) and thyrotropine (TSH), and proopiomelanocortine (Hortin et al., 1981; Hoshina et al., 1982; Green et al., 1984, 1985). The participating sulfotransferases appear to be membrane-bound and have been isolated from several sources. They are a collection of enzymes with different substrate specificity, e.g. for the 4- or 6-position of chondroitin (Göhler et al., 1984; Merkle and Heifetz, 1984; Rüter and Kresse, 1984; Munakata et al., 1985; Hortin et al., 1986a, 1986b, 1986c; Sugumaran et al., 1986; Slomiany et al., 1987). Glucosaminoglycan sulfotransferase activity is also found in serum (Inoue et al., 1986). High levels of chondroitin sulfotransferase activity were detected in foetal serum (Sugahara et al., 1985, 1987). The mucus glucoprotein in the small intestine of the newborn rat also had a high degree of sulfation (Shub et al., 1983). Under a number of conditions, the degree of sulfation of these macromolecules may be decreased, e.g. in hepatoma cell lines (Robinson et al., 1984), in patients with Lowe syndrome (Kieras et al., 1984), or when sulfate supply is limiting (Tyree et al., 1986; Esko et al., 1986). Sulfation of certain groups may be essential for biological activities, as in heparin (Lindahl et al., 1983) or in the oligosaccharide side chain of proopiomelanocortine (Bourbonnais and Crine, 1985).

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Proteins: tyrosine sulfation The hydroxyl group of tyrosine in proteins can be sulfated (Huttner, 1987) as has been demonstrated in liver, hepatoma and pituitary. Proteins in basement membranes, as well as collagen and secretory proteins are involved (Hille et al., 1984; Sorkin et al., 1984; Paulsson et al., 1985; Rosa et al., 1985; Liu et al., 1985; Fessler et al., 1986; Hortin et al., 1986a, 1986b; Liu and Baenziger, 1986; Hirose et al., 1988). This post-translational modification occurs in the Golgi system. When glycosylation is inhibited, tyrosine sulfation is increased (Baeuerle and Huttner, 1984, 1987). When the tyrosine-O sulfated proteins are proteolyzed, tyrosine-O-sulfate is released (Liu et al., 1987). Peptides A number of endogenous peptide hormones incorporate 35S-sulfate in vivo: cholecystokinin and the gastrines are sulfated (Brand et al., 1984a, 1984b; Cantor et al., 1986; Vargas and Schwartz, 1987). The sulfate group modifies peptide folding in cholecystokinin (Durieux et al., 1983) and strongly increases biological activities of the hormone (Vinayek et al., 1987). The endorphins were first shown to be sulfated by tyrosine ester sulfotransferase (Sekura and Jakoby, 1981). Dopamine and other catecholamines Catecholamines can be sulfated in man and animals and play a role in the physiological inactivation of these neurotransmitters (Buu, 1985b; Tyce et al., 1986), although the precise contribution of sulfation has not been defined (Roth and Rivett, 1982). Based on studies in human brain tissue, the contribution of sulfation was estimated to be as much as 10% of the metabolism of dopamine and noradrenaline in brain (Rivett et al., 1982; Whittemore and Roth, 1985). Inhibition of mono-amino oxidase leads to an increased contribution of sulfation in rat brain (Buu, 1985a; Buu et al., 1985). Rather different regional distributions of sulfotransferases were reported by Rivett et al. (1984) and Young et al. (1984) in human brain. In human plasma, catecholamines are only present as sulfate conjugates, whereas in rat plasma glucuronides play a major role (Wang et al., 1983). Conversion of dopamine sulfate to adrenaline by dopamine β-hydroxylase (Buu et al., 1981) is unlikely (Demassieux et al., 1987). The role of sulfation of catecholamines in blood pressure regulation has been reviewed (Kuchel et al., 1986). When dopamine or L-DOPA are administered orally in man high levels of dopamine sulfate (both the 3- and the 4-sulfate) are found in human plasma.

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However, after i.v. administration, only L-DOPA led to an increase in sulfate conjugates, whereas dopamine had little effect on dopamine sulfate in plasma (Hashizume et al., 1987). Presumably, orally administered dopamine is sulfated during uptake in the gut mucosa, while dopamine given intravenously does not readily reach (possibly intracellular) sulfation sites. Similar findings were reported in the dog (Oka et al., 1987). Dopamine sulfate has pharmacological effects (Ackerman et al., 1984; Buu et al., 1984). Steroid hormones In man, sulfation is a major conjugation reaction for steriods. Although the sulfates are excreted in urine, there is circumstantial evidence that they may have biological roles as intermediates or storage forms of the parent compound. The area of steroidal sulfates, including the corresponding sulfatases and their biological significance, has recently been reviewed by Hobkirk (1985). Briefly, the oestrogen sulfotransferase plays a role in regulation of the active oestrogen and, consequently, that of the oestrogen receptor. Sulfotransferases in the female reproductive organs have been studied extensively, as have those in the testis. The adrenal cortex and its role in cholesterol or dehydroepiandrosterone sulfate metabolism has also received attention. The latter is quantitatively the most important steroid sulfate in human blood. The level of the steroid sulfates in blood is highly variable, depending on such physiological conditions as the menstrual cycle, and the sulfotransferases are influenced by similar processes. Activities of the steroid enzymes are highest in the liver and most of steroid sulfation may occur in this organ. The review by Hobkirk (1985) and an earlier book on steroid biochemistry (Hobkirk, 1979) provide many details of this extensively studied subject. Although this is discussed in more detail in the section on alcohol sulfotransferases (Section 5.9), it is worth emphasizing that the enzymes active with steroids bearing an aromatic ring, are unique. On the other hand, non-aromatic steroids appear to be the domain of alcohol sulfotransferases, i.e. enzymes active with a wide spectrum of primary and secondary alcohols that include ethanol as substrate (Lyon and Jakoby, 1980). Bile acids Like steroids, bile acids can be sulfated at various positions (Mulder, 1981, 1984). The sulfate conjugates can subsequently be excreted in urine (Goto et al., 1987) or bile. The sulfate conjugates are more slowly excreted in bile than the unconjugated bile acids (Yousef et al., 1987); they seem to utilize separate transport systems (Kuipers et al., 1988). The enzymes involved have been studied and purified to a degree; they are regulated by steroid hormones and show developmental changes (Balistreri et al., 1984; Collins et al., 1987). In

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certain pathological states the excretion of bile acid sulfates in urine or bile is increased (Niessen et al., 1984; Kuipers et al., 1985; Stiehl et al., 1985). In streptozotocin-diabetic rats bile acid sulfotransferases and in vivo sulfation of bile acids were increased (Kirkpatrick and Kraft, 1984). Once sulfate conjugates have been excreted in bile they can be reabsorbed from the gut. Although this is somewhat slower for conjugated as compared with the unconjugated form, both are extensively reabsorbed in the rat. A high calcium concentration in the gut slows down the absorption (Kuipers et al., 1986). The position of the sulfate group critically affects the rate of ileal, i.e. carrier-mediated absorption (Lack et al., 1984). Bacterial sulfatases in the gut may give rise to an enterohepatic recirculation of the bile acids (Robben et al., 1988). The sulfate conjugates are little further metabolized in vivo (Eng and Javitt, 1983; Kuipers et al., 1986). Sulfation reduces the toxicity of the bile acids (Ammon et al., 1985). Uptake of the sulfate conjugate of chenodeoxycholate by isolated hepatocytes is almost as efficient as that for the unconjugated bile acid. Albumin inhibited the uptake of the sulfate conjugate more than that of the unconjugated form, probably because the former is more strongly bound (Bartholomew and Billing, 1983). Some bile acid sulfates, the sulfate conjugate of lithocholate for example, are cholestatic; taurine may prevent this cholestatic effect due to the formation of a taurine-sulfate double conjugate (Dorvil et al., 1983; Mathis et al., 1983). Thyroxine and other compounds Sulfation plays an important role in thyroxine metabolism (Sekura et al., 1981b; Otten et al., 1983, 1984; Mol and Visser, 1985; de Herder et al., 1988). Sulfation of the phenolic group of thyroxine or of the iodothyronines facilitates subsequent deiodination, leading to rapid and irreversible inactivation of thyroid hormones. The interrelationship between glucuronidation, sulfation and deiodination has been studied in isolated hepatocytes and in vivo. The major form of 25-hydroxy vitamin D3 circulating in blood in man is the sulfate conjugate (Axelson, 1985). Role of sulfation in the toxicity of xenobiotics Usually, biological activity of xenobiotics is decreased by sulfation, although any such effects of the sulfate conjugates are generally not investigated. That certain morphine sulfate conjugates are highly active shows that conjugates need not have less pharmacological activity. Morphine 6-sulfate is a 30-fold, more active analgesic than morphine or its 3-sulfate conjugate when injected intraventricularly (Brown et al., 1985). The 6-sulfate also showed increased binding to the δ-receptor (Oguri et al., 1987).

134 CONJUGATION REACTIONS IN DRUG METABOLISM

Sulfation certainly increases the rate of elimination of most lipid-soluble drugs, thereby enhancing the termination of their effects. In a growing number of cases however, sulfation increases toxicity reviewed by Miller et al. (1985) and Mulder et al. (1988). This is due to the formation of very reactive sulfate conjugates from specific groups of compounds that may react with other groups available in their environment or to rearrangement. Examples of the latter are the sulfate of 9-fluorenone oxime (Mangold et al., 1986), which rearranges spontaneously, or the sulfate conjugate of N-hydroxy-2acetylaminofluorene (NOH2AAF). The substrates that become toxic by reason of sulfation are mainly hydroxylamines and hydroxamic acids and benzylic alcohols. Most studied is NOH2AAF which is converted to a reactive nitrenium ion by sulfation. The reactive, electrophilic ions bind to RNA, DNA and protein, or to glutathione and similar nucleophiles, as can be shown with chemically-prepared sulfate conjugate (van den Goorbergh et al., 1985; Smith et al., 1987). For as yet unknown reasons, several carcinogens, when fed for a week, strongly decrease liver sulfotransferase activity (Ringer and Norton, 1987). The role of sulfation of NOH2AAF and a number of other carcinogens, e.g. 1′ hydroxysafrole or 4-aminoazobenzene, could be demonstrated by inhibition of sulfation with PCP, DCNP or inhibitors of alcohol sulfotransferases. Under such inhibitory conditions, the carcinogenic effect, the generation of preneoplastic foci, and the formation of DNA adducts were strongly reduced or totally prevented (Boberg et al., 1983, 1987; Kedderis et al., 1984; Fennell et al., 1985; Lai et al., 1985, 1987; Meerman, 1985; Delclos et al., 1986; Watanabe et al., 1986; Furlong et al., 1987; Surh et al., 1987; Okuda et al., 1988). Sulfate conjugates of certain compounds may serve as direct mutagens in the Ames-salmonella test (Mori et al., 1986; Rogan et al., 1986). Sulfation decreased the mutagenesis of NOH2AAF (Andrews et al., 1978). Sulfation may also be involved in the embryotoxicity of 2AAF (Faustman-Watts et al., 1986). II. The sulfotransferases 5.8. Introduction This portion of the chapter briefly summarizes what we know of the enzymology of sulfate transfer garnered from data in which only highly purified enzymes, preferably homogeneous proteins, were used. The material chosen is focused on the catalytic mechanisms and the substrate range of those enzymes active with xenobiotics. The historical aspects of the discovery of these enzymes has been reviewed in detail elsewhere (Roy, 1971; DeMeio, 1975; Jakoby, 1980). Because

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much of the information stems from work with proteins isolated from common laboratory animals rather than from man, it invites the comment that direct application from other species to man is limited. The argument is meritorious, particularly when applied to specific physical constants such as Km, pI, or size. In fact, the actual value for these factors may differ among organs in a single species and even among the isozymes present in one type of cell; thus, the caveat that a detailed set of properties should not be anticipated to be the same, even for enzymes performing similar functions in the same species. Nevertheless, the general catalytic process is expected to be closely similar, however unrelated the species; certainly the thermodynamics of the overall reaction are immutable. Therefore, the effort made here is to present the essentials of the general case, leaving many of the specific details for sulfotransferases with similar function from different sources to the specialized reviews that are available (Jakoby et al., 1980, 1984; Mulder, 1981). Any discussion of sulfate conjugation must begin with the donor molecule that was characterized in mammalian cells by Robbins and Lipmann (1957) as PAPS (systematically described as 3′-phosphoadenylysulfate). The currently known mammalian sulfotransferases, whatever their substrate specificity, result in the formation of organic sulfates by catalyzing the transfer of the sulfuryl group, −SO3−, from PAPS to an appropriate nucleophilic acceptor, thereby forming adenosine 3′, 5′-bisphosphate (abbreviated as PAP) and the sulfate conjugate. The reaction, shown in eqn. 5.1 for phenol as the acceptor molecule, is written as reversible, accurately reflecting the capability of some, but not all, of the sulfotransferases to catalyze a demonstrable back reaction (Gregory and Lipmann, 1957; Duffel and Jakoby, 1981; Anhalt et al., 1982). These enzymes also appear to catalyze the direct transfer described in eqn. 5.2; the real situation is more complex for mammalian cells (Duffel and Jakoby, 1981), but the equation accurately reflects the reaction catalyzed by certain anaerobes that probably participate in the utilization of sulfate in the intestinal tract (Kim et al., 1986; Kobashi et al., 1987). The mechanism of each of these processes will be presented. (5.1) (5.2) Assays and substrates As a practical matter, working with these enzymes requires assays for a wide range of substrates. Of the many that have been devised, most have been summarized (Roy, 1971; Sekura et al., 1979; Ramaswamy and Jakoby, 1987a) and take many forms that include the use of radioactive receptor molecules (Sekura et al., 1979; Ramaswamy and Jakoby, 1987a), systems particularly

136 CONJUGATION REACTIONS IN DRUG METABOLISM

effective with steroids (Lyon et al., 1981); the general use of a radioactive donor molecule, 35S-PAPS; the separation and identification of products by a number of means including small ion-exchange columns (Borchardt, 1983), thin layer chromatography (Sekura et al., 1979; Ramaswamy and Jakoby, 1987a, 1987b), HPLC (Ramaswamy and Jakoby, 1987b), and adsorption to membrane filters (Lyon et al., 1981); and a colorimetric procedure in which the ion pair of an organic sulfate with methylene blue is formed, extracted and measured spectrophotometrically (Roy, 1956; Ramaswamy and Jakoby, 1987a). Since each method has its special merit, it is useful to consult the specific description although detailed collections of generally applicable procedures are available (Sekura et al., 1979; Ramaswamy and Jakoby, 1987a). Another practical consideration is the lack of purity of many of the readily obtainable substrates. Despite the wealth of phenols, alcohols and amines that are commercially available, it is necessary to emphasize that most acceptor substrates are used at millimolar concentrations, i.e. at concentrations sufficiently high that a 1% contamination will represent micromolar impurity. That is too high and prior purification of potential substrates is critical. Less obvious and more insidious are the impurities present in commercial preparations of PAPS. Inhibition occurs due to contamination by PAP which is effective at micromolar levels for all of the PAPS-linked sulfotransferases. Preparations from one manufacturer had been contaminated by 30% whereras another had only a few per cent PAP (Ramaswamy and Jakoby, 1987c). Working with the more contaminated preparation of PAPS led to an interesting artifact which, as may be imagined, was time consuming to unravel. Two enzymes appeared to be necessary for high activity with amine Nsulfotransferase: the sulfotransferase itself and a second enzyme, (2′)3′, 5′ bisphosphate nucleotidase, whose sole function was to hydrolyze the 3′ phosphate of contaminating PAP, i.e. of adenosine 3′, 5′-bisphosphate, to form the innocuous AMP and inorganic phosphate (Ramaswamy and Jakoby, 1987c). 5.9. Sulfotransferases in xenobiotic metabolism A number of the sulfotransferases are believed to play a role in the metabolism of foreign compounds, and it is this group which is presented here. Not covered are the enzymes that participate in biosynthetic reactions in which macromolecules are substrates, e.g. the formation of heparin (Riesenfeld et al., 1982) or the modification of chondroitin (Nakanishi, 1981). Other sulfotransferases play an important role in sulfating bile acids (Chen and Segel, 1985; Barnes et al., 1986) and the hydroxyl groups on the aromatic ring of certain steroids (Adams and Poulos, 1967); these are viewed as reactions in which metabolic intermediates are made available. Whatever the purpose, we presently know of two functional

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Table 5.3. Rat liver alcohol sulfotransferases: apparent kinetic parametersa.

a

Data from Lyon and Jakoby, 1980; Marcus et al., 1980; and subsequent unpublished work by E.S.Lyon and J.L.Wang.

groups in mammals, hydroxyl and amino, that can undergo sulfuryl group transfer, i.e. sulfation, and our discussion of them will be based largely on whether the product is an O- or an N-sulfate. O-Sulfation: alcohol sulfotransferase (EC 2.8.2.2) There is a division between enzymes catalyzing sulfoconjugation of alcoholic and phenolic hydroxyl groups; the latter is considered separately below. The alcohol sulfotransferases of rat liver, of which at least three have been purified to homogeneous form (Lyon and Jakoby, 1980; Marcus et al., 1980; Lyon et al., 1981), do exactly that. Primary or secondary alcohols, but not tertiary alcohols, whether as low in the series as ethanol or as large as hydrocortisone, are all substrates. As with most enzymes active in the metabolism of xenobiotics, it is the more lipophilic substrates that have the lower Km as is shown in Table 5.3. Among the most effective substrates for the alcohol sulfotransferases, as evaluated by kcat/km, are the hydroxysteroids, and it was under the name 3ßhydroxysteroid sulfotransferase that these enzymes were originally described (Ryan and Carroll, 1976). Subsequently, it was shown that a much larger variety of steroid alcohols were capable of acting as sulfate receptor than simply those bearing a hydroxyl in position 3 (Jakoby et al., 1980) although specific or generic names, glucocorticoid (Singer et al., 1976) or mineralocorticoid sulfotransferase (Singer, 1982) for example, have been used. Finally, straight chain and other secondary alcohols were described as substrates, a list that we now know to include chloramphenicol, retinol (vitamin A), L- and D-

138 CONJUGATION REACTIONS IN DRUG METABOLISM

propranolol, ascorbic acid, ouabain and ephedrine (Jakoby et al., 1980); it will be evident that some of the steroid and aliphatic substrates are secondary alcohols. The alcohol sulfotransferases are not active with phenols or with the aromatic hydroxyl group of estrone; the exception is an enzyme that appears to effectively utilize hydroxysteroids whether or not the hydroxyl group is on an aromatic ring (Adams and McDonald, 1979; 1980). The three purified alcohol sulfotransferases do not use organic hydroxylamines, bile acids, cholesterol or amines as a sulfuryl group acceptor. O-Sulfation: phenol sulfotransferases (EC 2.8.2.1 and 2.8.2. 9) The second group of enzymes active in O-sulfation has been variously listed as phenol or aryl sulfotransferases but must be further subdivided, at least for rat liver from which these species are available in homogeneous form. One category accommodates only low molecular mass phenols (Sekura and Jakoby, 1979), among which are the thyroid hormones and their degradation products (Sekura et al., 1981a), whereas the second, in addition, includes peptides bearing an Nterminal tyrosine residue, as well as organic hydroxylamines, as substrates (Sekura and Jakoby, 1981b). These relationships are illustrated, in part, for three rat liver enzymes in Table 5.4. Aryl sulfotransferase (EC 2.8.2.1.1) is represented by two isoenzymes (I and II) whereas the third enzyme (IV), tyrosine-ester sulfotransferase (EC 2.8.2.9), is responsible for the catalytic activity with tyrosine methylester, N-terminal tyrosine peptides, and organic hydroxylamines. Tyrosine-ester sulfotransferase does not accept tyrosine itself as a substrate, nor will it act on an internal tyrosine residue in a peptide; endorphins are substrates as is the heptapeptide (N-terminal tyrosine), but not the octapeptide of cholecystokinin. The two organic hydroxylamine derivatives that have been tested, N-hydroxy-2-acetylaminofluorene and the model compound, 2cyanoethyl-N-hydroxythioacetamidate, are sulfated by this specific species, tyrosine ester sulfotransferase. The same enzyme has been shown to catalyze sulfation of benzylic alcohols to form electrophilically reactive compounds (Binder and Duffel, 1988); there was an inverse correlation of Km with lipophilicity of the benzylic alcohol. Interestingly, the catalytic efficiency for S(−)-1-phenylethanol was about three-fold that for R-(+)-1-phenylethanol. An enzyme, isolated from rat liver on the basis of its activity with Nhydroxy-2-acetylaminofluorine (Wu and Straub, 1976), is in all probability a tyrosine-ester sulfotransferase. There are now several highly purified preparations of phenol sulfotransferase from a number of species and from tissues as diverse as brain (Pennings et al., 1978; Roth and Rivett, 1982) and platelet (Hart et al., 1979), as well as additional isolates from liver (Borchardt and Schasteen, 1982). As a minimum, substrate studies using the identical spectrum

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Table 5.4. Rat liver aryl sulfotransferases: apparent kinetics constants.a

a

Data from Sekura and Jakoby, 1979, 1981, and adapted from Jakoby et al., 1980, 1984. Determined at pH 6.5 with 0.1 mM PAPS. c Unless otherwise noted under ‘Substrate’, data were obtained at pH 5.5 with 0.1 mM PAPS. d One unit is equivalent to 1 nmol min.-1 e Determined at pH 7.9 with 0.1 mM PAPS. b

of compounds need to be performed in order to evaluate possible relationships among these protein species. Very extensive studies of phenol utilization (Campbell et al., 1987a) are available for two aryl sulfotransferases from human liver although neither has been highly purified (Rein et al., 1982; Campbell et al., 1987b). Human tissue appears to have two types of enzymes that differ in their thermostability, sensitivity to inhibitors and substrate spectrum (Weinshilboum, 1986) although there is a large overlap in specificity (Campbell et al., 1987a). N-Sulfation: amine N-sulfotransferase The third group of sulfotransferases catalyzes the formation of sulfamates from amines and is represented by amine N-sulfotransferase. This activity was originally described three decades ago in extracts from rat and guinea pig liver (Roy, 1956) after the observation that 2-naphthylamine sulfamate was excreted after oral administration of 2-naphthylamine to rats (Boyland et al., 1957). The in vitro formation of the appropriate N-sulfamate has been verified recently (Prosser and Roy, 1982). Subsequently, amine N-sulfotransferase has been purified from guinea pig liver and found to be an enzyme of broad specificity catalyzing sulfuryl group transfer to both primary and secondary amines with diverse carbon skeletons (Table 5.5) (Ramaswamy and Jakoby, 1987b).

140 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 5.5. Guinea pig liver amine N-sulfotransferase.

a

In the presence of enzyme, 2 mM of acceptor amine, 1 mM PAPS, and either 0.1 M potassium bicine at pH 7.2 or 0.2 M glycine at pH 10 for 60 min at 37°C. (Adapted from Ramaswamy and Jakoby, 1987b.)

Since only one example of N-sulfotransferase has been purified, there is no means for assessing the generality of the behaviour of this type of enzyme. Nevertheless, only the unprotonated form of the amine serves as acceptor substrate. This is clearly indicated by the pH-activity profile for the enzyme, detailed elsewhere (Ramaswamy and Jakoby, 1987b) but summarized in Table 5.5, in which arylamines, with their high pKa, are effective substrates only at high pH. Not listed in the table but active as acceptors are aniline, 1naphthylamine, 1-amino-4-nitronaphthylene, 4-tertbutylaniline, Nmethylaniline, tridecylamine, and toluidine. It may be of interest that cyclohexylamine serves as an acceptor to yield its corresponding sulfamate, the sugar substitute known as cyclamate. No evidence of reversibility was found upon incubation of enzyme with 2-naphthylsulfamate and PAP. The following compounds were not found to form sulfamates at the level of detection of the assay method: pyridine and its 4-amino-, 4chloro- or 4-phenyl derivatives; diaminoanthraquinone; ß-phenylethylamine; and saccharine. Despite the high degree of purification attained, the amine N-sulfotransferase displays activity for sulfation of phenols, hydroxysteroids and even estrone as sulfuryl group acceptors. Although additional data favour the conclusion that the isolated N-sulfotransferase is one enzyme of broader than expected activity, encompassing both N- and O-sulfotransferase substrates, these observations serve to emphasize the difficulty in distinguishing between multiple enzymes and multiple activities in one enzyme. Cell-free extracts of guinea pig, rat and rabbit liver were able to form the sulfamate of 2-naphthylamine at rates of 290, 180, and 60 pmol min−1 mg−1 of protein, respectively. Also active were extracts of intestinal mucosa from rat (210 pmol) and rabbit (40 pmol) but not from guinea pig. Kidney from the rat was active (50 pmol) but not from the other two species. Activity in lung was, if

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present at all, below 10 pmol for all three animals, as it was for brain, heart and spleen of the rat. Trans-sulfation: arylsulfate sulfotransferase A novel sulfate conjugating enzyme has been described from an anaerobic bacterium present in the human intestinal tract. The enzyme, arylsulfate sulfotransferase, catalyzes the transfer of the sulfuryl group (i.e. sulfate) from one phenylsulfate to another phenol without mediation by either PAPS or PAP (Kim et al., 1986; Kobashi et al., 1987). As might be expected, not all arylsulfates are donors nor are all phenols acceptors although it is not known whether a substrate’s effectiveness is a function of its affinity for the enzyme or the thermodynamics of the transfer; undoubtedly both have a role. The enzyme does have a broad specificity. Sulfuryl group donors are 4-acetylphenylsulfate, 4methylumbelliferylsulfate, 4-nitrophenylsulfate, picosulfate, estrone sulfate and indoxylsulfate but not PAPS. The following compounds served as acceptors: estradiol, phenol, tyrosine methylester, tyramine, dopamine, epinephrine and 3, 4-hydroxyphenylalanine (Kobashi et al., 1986). Aside from the classical phenols, estrone sulfate, in which the hydroxyl is on the aromatic ring of the steroid, is a donor; although estrone itself was not tried as an acceptor, estradiol was an acceptor and it was the hydroxyl group on the aromatic ring that was esterified (D.-Y.Kim, personal communication). Bile acids and hydroxysteroids do not serve as acceptors. In contrast to tyrosine-ester sulfotransferase, this enzyme is capable of conjugating internal tyrosine residues of peptides, e.g. the octapeptide of cholecystokinin or the angiotensins, in addition to the N-terminal tyrosine residues for which both enzymes are competent. The bacterium, a species of Eubacterium, is a dominant organism in the gut of man and its participation in xenobiotic metabolism must be considered. The activity of the bacterial flora on sulfate metabolism may be appreciable as suggested by the effects of treating rats with antibiotics (Kim and Kobashi, 1986). 5.10. Aspects of sulfotransferase mechanism It may be worthwhile to simply identify and describe the several enzymes that constitute the sulfotransferases, but a major goal of such investigation is an understanding of the mechanisms by which these enzymes act and the ability to predict and manipulate the effectiveness of specific compounds as substrates. Although the availability of highly purified enzymes creates the opportunity for studies of catalytic mechanism, success has been limited. A more extensive

142 CONJUGATION REACTIONS IN DRUG METABOLISM

presentation of the chemical mechanism of sulfation and sulfate ester hydrolysis than is summarized here, is available (Duffel and Jakoby, 1987). There are problems with even preliminary tasks such as obtaining valid kinetic constants for the sulfotransferases. It was with difficulty that tyrosineester sulfotransferase was separated from sulfotransferase III (Duffel and Jakoby, 1981; Sekura and Jakoby, 1981); the latter had very different kinetic constants, sufficient to suggest that two enzyme species were present and, hence, to require separation. Nevertheless, at least two kinetic analyses of phenol sulfotransferases, performed without knowledge of the variety of isoenzymes present in the same tissue, are available from other laboratories. An additional difficulty, already referred to, is that even apparent kinetic constants cannot be obtained easily when high concentrations of substrates are necessary; this brings the possibility of contamination by inhibitors. If Lineweaver Burk plots are curvilinear as they approach the y-axis, is that due to substrate inhibition or to the presence of inhibitors? The information on mechanism—whether kinetic, chemical or enzymic—is limited. For the alcohol sulfotransferases, it will be apparent that certain correlations can be made with regard to substrate lipophilicity. Thus, the Km for each of these enzymes progressively decreases from the simplest substrate, ethanol, through higher alcohols and secondary alcohols to the sterols (Jakoby et al., 1980). This property, that of avidity for the lipophilic substrate may be a general rule for the enzymes involved in the metabolism of xenobiotics (Jakoby, 1980). For the N-sulfotransferase, it is clear that the unprotonated form of either the primary or secondary amine substrate is required for the amine to serve as acceptor (Ramaswamy and Jakoby, 1987b). Arylsulfate sulfotransferase The mechanism of the bacterial sulfotransferase differs in that neither PAP nor PAPS participates. It has been shown that the ratio of absorbance of the purified protein at 260nm to that at 280nm is low (Kim, personal communication) in accord with the absence of such a nucleotide. Evidence has been presented that eqn. 5.2, when carried out in the absence of a phenol as acceptor, gives rise to a sulfate covalently linked to a tyrosine residue of the sulfotransferase. When the enzyme-sulfate was isolated and treated with a phenol, an arylsulfate was not formed. However, the conjugated sulfate was released to an acceptor phenol if an arylsulfate was also present (Kim et al., 1986). Whether this series of reactions represents a direct displacement mechanism is not yet clear.

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Figure 5.7 Outline of the kinetic intermediates in the reaction catalyzed by tyrosineester sulfotransferase (E). Included are complexes with 2-chloro-4-nitrophenol (CNP), PAPS, PAP, and 2-chloro-4-nitrophenylsulfate (CNPS).

Tyrosine-ester sulfotransferase The only enzyme of this group for which there is extensive evaluation is the tyrosine-ester sulfotransferase (Duffel and Jakoby, 1981). Kinetically the mechanism of the enzyme-catalyzed reaction of PAPS with 2chloro-4nitrophenol (CNP) to yield PAP and 2-chloro-4-nitrophenylsulfate (CNPS) is consistent with a rapid equilibrium random Bi-Bi mechanism with two dead-end inhibitor complexes. The kinetic analysis has included patterns of inhibition as well as binding studies and is consistent with the scheme shown in Figure 5.7. Parenthetically, a bile salt sulfotransferase was found to have a sequential order Bi-Bi kinetic mechanism with the acceptor substrate as the first to bind (Barnes et al., 1986). The importance of the kinetic data that yielded Figure 5.7 is that the rate constants of each of the participating reactions are known which, in turn, allowed solution of the four Haldane equations in effect for a rapid equilibrium random mechanism. By this calculation, the equilibrium constant for the reaction shown was found to be 0.045±0.009. At pH 7.0, sulfuryl group transfer from PAPS to CNP is an endergonic reaction with a change in free energy of +1. 6 kcal per mole at pH 7 and 25°. Not only is the reaction experimentally reversible, but for 2-chloro-4-nitrophenylsulfate, it is in favour of PAPS formation. For most other arylsulfates, the reaction is far less favourable. Let it be clear that the back reaction is not simply that summarized as eqn. 5.2. Rather, although PAP does not appear in the \ equation, eqn. 5.2 for this enzyme is the sum of eqns. 5.3 and 5.4, i.e. small amounts of PAP are necessary and play a catalytic role (Duffel and Jakoby, 1981). The sulfotransferase has been shown to catalyze one other reaction (eqn. 5.5), the hydrolysis of sulfate esters, although this function is at a very slow rate and of doubtful physiological significance (Duffel and Jakoby, 1981). (5.3) (5.4) (5.5)

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A misconception has arisen about sulfuryl transfer based on the idea of a ‘sulfate potential’ (Gregory and Lipmann, 1957). The suggestion was offered that certain phenols, 1- and 2-naphthols are examples, are not acceptors from phenylsulfate, despite the presence of PAP, because of a low sulfate potential; in contrast, phenol itself is a good substrate. Certainly, the thermodynamics of the system will govern the extent of the reaction with the change in free energy differing for each substrate. In the event, however, the explanation of acceptor activity of the naphthols is more mundane. The enzyme has a low Km for the naphthols (1 to 5 µM), both of which are strong substrate inhibitors in the millimolar range at which they were tested. Phenol, on the other hand, has a Km in the millimolar range at which it was tested as an acceptor and is not inhibitory at these concentrations. Thus, both compounds serve as sulfuryl group acceptors at non-inhibitory concentrations, i.e. at their respective Km (Duffel and Jakoby, 1981). Chemical and catalytic mechanism Sulfation occurs by electrophilic attack on the oxygen of alcohols and phenols by the sulfur of sulfuric acid, sulfuryl halide or of a sulfuric anhydride (Deno and Newman, 1950; Burwell, 1952; Batts, 1966). Only primary and secondary alcohols react, the latter at a much slower pace (Deno and Newman, 1950); tertiary alcohols do not form stable esters. Much of the data that bears on the chemical mechanism of sulfation is the result of investigation of the reverse action, i.e. the hydrolysis of sulfate esters. Because the principle of microscopic reversibility requires both forward and reverse reactions to proceed through the same transition state, study of the hydrolytic reaction was productive. For phenylsulfate and 4–nitrophenylsulfate subjected to acid hydrolysis, the transition state, shown below, has been suggested as being relatively non-polar with many of the characteristics of a sulfate ester. The phenolic oxygen is affected by the

strong electron-withdrawing SO3 group as well as by electron delocalization in the aromatic ring (Benkovic and Benkovic, 1966; Kice and Anderson, 1966). Since para substitution in the phenyl ring affects the rate of hydrolysis less than might be expected, Kice and Anderson (1966) proposed that electronwithdrawal by the SO3 group adjacent to the phenolic oxygen, minimizes resonance interaction with electron-withdrawing para substituents. Their conclusion as to the transition state is consistent with the results of kinetic

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studies of the tyrosine-ester sulfotransferase with which a number of parasubstituted derivatives of phenol were assessed as substrates. The Hammett ρ value for phenol sulfation was −0.25 when σ− values were used (Duffel and Jakoby, 1981), much smaller than expected if the substituent effect was due only to resonance stabilization of the phenol and in accord with a transition state that better resembles a sulfate ester than a phenol. The transition state proposed for acid hydrolysis of phenyl sulfate, a relatively non-polar entity (Kice and Anderson, 1966) could reasonably be stabilized by a hydrophobic environment at the enzyme’s active site. As with most enzymes involved in xenobiotic metabolism, based on the broad range of lipophilic compounds that such enzymes will accept as substrate, a lipophilic environment seems appropriate for the binding site of the acceptor substrate. The process of quantitative structure-activity analysis (Hansch and Leo, 1979) has been carried out for a large number of phenolic substrates of the human liver phenol transferases and compared with values calculated from the apparent kinetic constants obtained previously (Sekura and Jakoby, 1979) for the two rat aryl sulfotransferases (Campbell et al., 1987a). By observation, neither the absolute values nor the rank order of the Km values for substrates of human and rat liver enzymes correlated well; Km differed by three orders of magnitude for the two sets of enzymes. This analytical approach (Hansch and Leo, 1979) is an attempt to correlate Km with a number of parameters among which are steric factors and bulk, hydrophobicity and, for the transferases, the σ− value. An empirical equation was assembled for each sulfotransferase in which constants for each parameter were approximated. Although such approximation led to similar values for the two human enzymes and for the two rat enzymes, it was suggested that there may be substantial differences between the protein groups from the two species in so far as the contribution of specific parameters to the differences in Km (Campbell et al., 1987). Of parenthetical interest is the observation that substituent effects on the rate of phenol sulfation are greatly increased when all ortho and para positions are occupied by electron-withdrawing groups. Both, 2, 6-dichloro 4-nitrophenol and pentachlorophenol (Mulder and Scholtens, 1977) are excellent dead-end inhibitors of rat phenol sulfotransferases with a Ki of 1 µM or less (Duffel and Jakoby, 1981). In contrast, the amine N-sulfotransferase which is also active with phenols and alcohols, actually utilizes pentachlorophenol as a substrate, forming an O-sulfate (Ramaswamy and Jakoby, 1987b). III. Methodological aspects In a recent volume of Methods in Enzymology (vol. 143, 1988), a complete survey is given of methodology in the area of sulfation, such as assay methods for

146 CONJUGATION REACTIONS IN DRUG METABOLISM

inorganic sulfate in plasma, PAPS in tissue extracts, and sulfotransferases as well as their purification. For experiments in isolated cells, perfused organs and in vivo, the same substrates are usually employed as mentioned for glucuronidation. When low concentrations are used, sulfation almost exclusively occurs. At increasing concentration, glucuronidation becomes more important as a competitive pathway. References for methodology can be found in Chapter 4, Section 4.9. Abbreviations 2AAF APS AUC CNP CNPS DCNP 3-MC NOH2AAF PAP PAPS PCP

2-acetylaminofluorene Adenosine 5′-phosphosulfate or adenylylsulfate Area under the curve 2-Chloro-4-nitrophenol 2-Chloro-4-nitrophenyl sulfate 2, 6-Dichloro-4-nitrophenol 3-methylcholanthrene N-hydroxy-2-acetylaminofluorene Adenosine 3′, 5′-bisphosphate Adenosine 3′-phospho 5′-phosphosulfate phosphoadenylylsulfate Pentachlorophenol

or

3′-

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 6 Acetylation Wendell W.Weber1, Gerald N.Levy1 and David W.Hein2 1

Department of Pharmacology, University of Michigan Medical School, Ann Arbor, MI 48109, USA.

2

Department of Pharmacology, University of North Dakota School of Medicine, Grand Forks, ND, USA.

6.1.

INTRODUCTION

163

6.2.

ENZYMOLOGY OF ACETYLATION

166

Tissue distribution

166

Substrates

167

Inhibitors and inducers

168

Developmental variation

169

Species and hereditary variation

169

Physicochemical properties

170

Kinetic properties

171

Multiple acetyltransferase isozymes

173

6.3.

ACETYLATION IN ISOLATED CELLS AND IN VIVO

177

6.4.

THE BIOLOGICAL SIGNIFICANCE OF ACETYLATION

179

6.5.

PRACTICAL CONSIDERATIONS

182

Enzyme purification

182

Enzyme assays and data analysis

182

Measurement of DNA damage induced by aromatic amines

184

ACKNOWLEDGEMENTS

185

NOTES

185

ACETYLATION 163

ABBREVIATIONS

185

REFERENCES

185

6.1. Introduction Toward the end of the last century, physiological chemists discovered that chemicals taken into the body are subjected to definite biotransformations and are usually excreted in the urine as conjugates. During that period, biological acetylation was identified by the German organic chemist, Rudolf Cohn (1893), as one of the major conjugation reactions of humans and animal species. Biological acetylation in humans and animals usually involves the enzymic transfer of the acetyl group from endogenous acetyl coenzyme A (AcCoA) to molecules that contain a primary amine, a hydroxyl or a sulfhydryl group. Acetylation of the primary amine group of arylamines and hydrazines is a major route of biotransformation of these substances. The acetylation of choline and of coenzyme A are singular examples of endogenous substances that involve acetyl transfer to the hydroxyl group and the sulfhydryl group respectively, but the acetylation of those groups in environmental chemicals is much less prevalent (Williams, 1959). Only recently it has been shown that the acetylation of the hydroxyl group does occur for certain carcinogenic Nhydroxylated aromatic amines, but examples of the acetylation of the sulfhydryl group of environmental chemicals still appear to be lacking. The liver is a major site of biological N-acetylation (Klein and Harris, 1938). Studies of pigeon liver in cell-free systems pioneered by Lipmann (1948–1949) pointed to a coenzyme A derivative, ultimately shown to be acetyl coenzyme A (Lynen, 1952–1953), as the essential co-factor of acetylating enzymes. The enzyme catalyzing acetyl coenzyme A-dependent N-acetyl transfer was first characterized in pigeon liver (Lipmann, 1945; Tabor et al., 1953; Johnson, 1954) and was named acetyl coenzyme A: arylamine N-acetyltransferase (NAT) (E.C. 2.3.1.5). Biotransformation through N-acetylation has long been known to alter the disposition and excretion of arylamine and hydrazine drugs and non-drugs, and to affect the responses of individuals to those substances. More recently, Nacetylation has been recognized to play a modulatory role in the metabolic activation of mutagenic and carcinogenic aromatic amines and of various food pyrolysates that are classified as carcinogens. Drugs and other environmental chemicals that do not contain a primary amine but can be transformed to that state can also be N-acetylated, and the N-acetylation of neurotransmitters in the pineal gland and of peptide hormones of the pituitary and hypothalamus has

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stimulated interest in acetylation as a regulatory process of physiological importance. Enzymic acetyl transfer reactions other than N-acetylation contribute to the cytotoxicity, mutagenicity and carcinogenicity of aromatic amines and hydrazines (Figure 6.1). Booth’s observations on rat liver cytosol (1966) that proximate carcinogenic arylhydroxamic acids, such as N-hydroxy-N-acetyl-2aminofluorene, can serve as the acetyl donors for acceptor molecules, such as 2aminofluorene, benzidine, 4-aminobiphenyl and beta-naphthylamine, provided initial evidence for arylhydroxamic acid-dependent N, N-acetyl transfer. King and Phillips (1968) and Bartsch et al. (1972, 1973) observed arylhydroxamic acid-dependent inter- and intramolecular transfers, respectively, of the acetyl group from the nitrogen to the oxygen of N-acetyl-N-hydroxyarylamines (arylhydroxamic acids), N, O-acetyl transfer, as pathways for the formation of carcinogenic N-acetoxyarylamines. Within the past ten years, however, several investigators in laboratories in Japan and the United States have shown the occurrence of acetyl coenzyme A-dependent O-acetyl transfer as another route of formation of carcinogenic N-acetoxyarylamines from mutagenic and carcinogenic N-hydroxyarylamines (reviewed in Kato, 1986; Weber, 1987; Hein, 1988; Kato and Yamazoe, 1988). The N-acetyltransferases of mammalian liver catalyze acetylation according to a ‘ping-pong Bi-Bi’ mechanism. The reaction is customarily written as follows

or as two half reactions

The ping-pong mechanism for acetyl coenzyme A-dependent N-acetyl transfer was characterized for mammalian N-acetyltransferases by studies of the initial velocity and product patterns, of the reversibility of the half reactions and by isolation of the acetyl enzyme intermediate (Weber, 1973). A mechanism similar to that for acetyl coenzyme A-dependent acetyl transfer has also been proposed for arylhydroxamic acid-dependent intramolecular N to O acetyl transfer (King, 1974). However, probing the mechanisms of N, O acetyl transfer is not amenable to analysis by the approach applied to N-acetyl transfer because of the instability of reaction products such as the N-acetoxyarylamines that occur in that reaction (Banks and Hanna, 1979; Hanna et al., 1982; Smith and Hanna, 1988). Investigation of the mechanism of O-acetyl transfer faces the same type of difficulty as that for N, O-acetyl transfer and, as yet, has not been carried out.

Figure 6.1. Acetyl transfer reactions in mammalian tissues. (Reproduced with permission from Hein, 1988.)

ACETYLATION 165

166 CONJUGATION REACTIONS IN DRUG METABOLISM

The identification of persons as ‘rapid’ and ‘slow’ acetylators of amines and hydrazine drugs during the 1950s and of many other chemicals since then added another important dimension to the biology of acetylation. Person-to-person differences in acetylation are attributable to two major acetylator alleles at a single autosomal gene locus which are expressed in liver, gut mucosa and most other tissues. The acetylator gene locus confers large stable differences in acetylating capacity on individuals early in life, probably during infancy, and they also influence the therapeutic, toxicological and pharmacological responses of individuals to drugs and other environmental aromatic amines and hydrazines to a remarkable extent. Genetic epidemiology studies suggest an expanding role for the acetylator trait as a susceptibility determinant to drug-induced toxic states, to cancers induced by aromatic amines and to various spontaneous human disorders (reviewed in Weber, 1987). Our knowledge of the biological basis of the human trait and some of its consequences has been augmented by experimental pharmacogenetic investigations of the hereditary acetylation polymorphisms in rabbits, mice, hamsters and rats and in bacteria during the past ten years. In this chapter we summarize information about the human and animal traits, much of which has been reported within the past five years. More comprehensive accounts of biological acetylation are available (Weber, 1987; Hein, 1988) for those who wish to go beyond the scope of this article. 6.2. Enzymology of acetylation Tissue distribution N-Acetylating enzymes are distributed in many tissues of humans and animals. N-Acetyltransferases of tissues of humans, hamsters, mice, rabbits and rats have been examined in many studies (reviewed in Weber, 1987; Hein, 1988). Extrahepatically, N-acetyltransferases have been identified in intestinal mucosa, colon, lung, thymus, ovary, spleen, uterus, adrenal gland, leukocytes, kidney, bone marrow, salivary gland, pancreas, pineal gland, erythrocytes and brain. Activity is also present in the mucosa of the urinary bladder (Smolen and Weber, 1983; Hein et al., 1987c; Kirlin et al., 1988a, 1989), the human placenta (Juchau et al., 1968; M.Manis and W.Weber, unpublished observations), and skin (Kawakubo et al., 1988) but has not been detected in plasma, skeletal muscle and fat.

ACETYLATION 167

Substrates Liver N-acetyltransferase catalyzes acetyl coenzyme A-dependent N-acetylation of drugs and other chemicals that possess NH2 or NH-NH2 groups attached directly, or via a carbonyl group, to an unsaturated ring; it can also catalyze Nacetylation of the NH2 group attached to ring systems via a short aliphatic carbon side chain. Structure-activity studies of 30 aniline derivatives that differ in hydrophobicity, position and number of substituents, size, and charge show that some ortho-substituted derivatives are not acetylated, probably due to steric hindrance (Andres et al., 1987). The specificity of the enzyme is broad, encompassing many arylamines and hydrazines. Arylamine and hydrazine substrates include acetylhydrazine, 4aminobiphenyl, 2-aminofluorene, aniline (and its ring-substituted analogues), benzidine, methyl bis-2-chloroaniline, methylene dianiline, pphenetidine, paminobenzoic acid, p-aminosalicylic acid, p-aminosulfonic acid, 1naphthylamine, 2-naphthylamine, sulfadiazine, sulfamethazine, sulfanilamide, sulfapyridine, diaminophenylsulfone (Dapsone), isonicotinyl hydrazide (isoniazid), hydralazine, p-nitroaniline, phenelzine, procainamide, and thiazolsulfone (Promizole). Certain biogenic amines such as histamine, tyramine, 5-hydroxytryptamine (serotonin), are substrates. Other substrates include the amine metabolites of acebutolol, caffeine, clonazepam, dipyrone, nitrazepam, sulfasalazine, and various heterocyclic arylamine food pyrolysates such as 2amino-6-methylpyridol[1, 2-a:3′, 2′-d]imidazole (Glu-P-1) (reviewed in Kato, 1986; Hein, 1988). Substrates for acetyl coenzyme A-dependent O-acetylation include hydroxylamine derivatives of aromatic amine carcinogens such as N-hydroxy4aminobiphenyl and N-hydroxy-2-aminofluorene, and the heterocyclic arylamine food pyrolysates such as N-hydroxy-Glu-P-1 (Kato, 1986; Hein, 1988). AcCoA is the endogenous acetyl donor for acetyl coenzyme A-dependent Nacetylation, but arylhydroxamic acids and a number of other acetyl donors can also serve this purpose (Flammang and Kadlubar, 1986; Shinohara et al., 1986a). Cyclohexylamine, glucosamine, phenylalanine, and pyridoxamine are examples of amines that are not acetylated by the enzyme. The specificity of the N, O-acetyltransferase reaction, as discussed elsewhere (Weber and King, 1981), involves two successive steps, i.e. N-deacetylation and O-acetylation. Studies performed mainly with preparations from rat liver or mammary gland show that a wide variety of arylhydroxamic acids are suitable as substrates for both of these reactions (e.g. fluorenyl, biphenyl, benzidine, stilbene, phenanthrene). Microsomal deacetylases are often distinguished from cytosolic N, O-acetyltransferases by the former’s sensitivity to diethyl-pnitrophenyl phosphate (paraoxon; King and Glowinski, 1983). The specificity

168 CONJUGATION REACTIONS IN DRUG METABOLISM

involved in the metabolism of propionylated substrates is qualitatively similar to that of acetylated substrates, but propionylated substrates are usually less rapidly metabolized. Formylated N-arylhydroxyamines can also be activated, but their mechanism of activation is unknown. Inhibitors and inducers N-Ethylmaleimide, iodoacetate and p-chlormercuribenzoate are potent irreversible inhibitors of the N-acetyltransferase (Weber, 1973). Bromoacetanilide, a product analogue, also causes rapid irreversible loss of activity (Andres et al., 1988). Cadmium chloride, arsenite, diisopropylfluorophosphate, and paraoxon have no effect on the activity. Various metal ions including Cu2+, Zn2+, Mn2+ and Ni2+ are inhibitory (Weber, 1973). Pentachlorophenol and 1-nitro-2-naphthol are potent inhibitors of arylamine Nacetyltransferase, N-hydroxyarylamine O-acetyltransferase, and arylhydroxamic N, O-acyltransferase reactions in hamster and rat liver cytosols (Shinohara et al., 1986b). No induction effects have been shown to occur with either barbiturates (pentobarbital) or isoprenaline. Glucocorticoids, such as hydrocortisone, and immunostimulants, such as zymosan and complete Freund’s adjuvant, have been shown to have an inducing effect on N-acetyltransferase activity (summarized in Reeves et al., 1988). Numerous compounds, some of which are structurally similar to substrates or products of the acetylation reaction, are reversible N-acetyltransferase inhibitors. These include various amides of heterocyclic carboxamides, such as salicylamide, 5-bromosalicylamide and 5-methylsalicylamide, and carboxyhydrazides (Johnson and Corte, 1956), folic acid and various 4amino folic acid antagonists (Johnson et al., 1958; Mandelbaum-Shavit and Blondheim, 1981; Hultin and Weber, 1986), and certain beta-carboline derivatives, such as harmine and harmaline (Wright et al., 1979). The inhibition of salicylamide is competitive (Johnson, 1955a) whereas the inhibition of the folic acid antagonists is non-competitive (Johnson et al., 1958). For certain miscellaneous inhibitors, patterns of inhibition may vary between and within species. The best example to illustrate this is dimethylsulfoxide (DMSO) which inhibits the mouse liver enzyme (Mattano and Weber, 1987) but fails to inhibit the rabbit liver enzyme (Mattano and Weber, unpublished observation). Further studies show that its inhibitory effect on slow (A/J) is much greater than that on the rapid (C57BL/6J) liver isozyme (Mattano and Weber, 1987).

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Developmental variation Tissues of foetal and neonatal humans (Fichter and Curtis, 1955; Vest and Salzburg, 1965) and animals are capable of N-acetylation (Uher, 1966; Juchau et al., 1968; Cohen et al., 1973; Sonawane and Lucier, 1975) and patterns of activity accompanying development may differ from one tissue to another. For example, N-acetyltransferase activity of foetal liver and gut of rats and rabbits increases during gestation and continues for two weeks after birth (Sonawane and Lucier, 1975) whereas kidney activity remains relatively constant throughout this period. Lung activity, on the other hand, shows a biphasic pattern distinct from that in other tissues, reaching a peak during gestation that exceeds that of neonatal or adult animals. Immediately after birth, lung activity decreases and then increases again to adult levels by 14 days postnatally. Cohen et al. (1973) have shown that the level of activity per gram liver of one-week-old rabbits is only about 3–5% of adult levels and reaches adult levels in about four weeks. The molecular weight, substrate specificity, inhibitory properties and the mechanism of acetylation of one-week-old and mature rabbits are identical, but the developmental variation is associated with a ‘foetal’ form of N-acetyltransferase that is more heat-labile and has a limiting Km value for isonicotinyl hydrazide (isoniazid) approximately one-third that of the enzyme of mature animals, The transition from the foetal to the adult form occurs during the third week after birth but the mechanism of this change has not been investigated. Species and hereditary variation Species variation in the N-acetylation of drugs and other environmental arylamines and hydrazines is large. The hamster and rabbit possess comparatively high N-acetylating capacities, whereas the dog has little or no appreciable acetylating capacity. Other species, such as the primates (including humans), mouse, and rat, have intermediate capacities. Since the discovery of the human acetylator polymorphism, hereditary variation in N-acetylation has been found in numerous species including the rabbit, Rhesus monkey, deer mouse, house mouse, rat and Syrian hamster. The human trait has been investigated for more than 30 years, but the animal traits have been investigated most intensively only within the past 10 to 15 years. Numerous advances in our knowledge of the characteristics of the hereditary acetylator traits of the rabbit, mouse, and hamster have been achieved during this period. Biochemical studies of animal and human tissues are beginning to provide an improved understanding of the biological basis of the human trait. Epidemiological studies of human populations and toxicological studies of

170 CONJUGATION REACTIONS IN DRUG METABOLISM

animals that express an acetylation polymorphism have led to a better appreciation of the significance of the human trait. Physicochemical properties The liver enzyme of rapid acetylator rabbits is a monomer with a pI of 5·2 and a molecular weight of 32 000–33 000 (Andres et al, 1987; Kabishev and Patrushev, 1987), whereas that of the mouse is slightly smaller (31 500) (Mattano et al., 1989). Studies of the avian liver enzyme show its molecular weight to be 33 000 in the pigeon (Andres et al., 1983a) and 33 000–34 000 in the chicken (Ohsako et al., 1988). Activity of the mammalian enzyme is strongly dependent on ionic strength and on the concentration of specific salts. It is essentially independent of pH over a wide pH range (pH 5·8–8·6). Amino acid analysis of liver N-acetyltransferases of the pigeon (Andres et al., 1983a), of the rapid acetylator rabbit (Andres et al., 1987) and of the rapid acetylator (C57BL/6J) mouse have been determined (Table 6.1; Mattano et al., 1989). Studies of the rabbit liver enzyme indicate that it contains no amino sugars or co-factors and that its N-terminal amino acid is blocked. Partial amino acid sequences have been determined for approximately 55% of the enzyme from tryptic peptide fragments of the rapid rabbit acetylator liver enzyme (Andres et al., 1987). More recently, partial amino acid sequences have also been determined for the chicken liver enzyme (Deguchi et al., 1988), and screening of a ?gt10 cDNA library with an oligonucleotide deduced from those amino acid sequences has enabled the isolation of a cDNA clone encoding the chicken liver N-acetyltransferase (Ohsako et al., 1988). The complete nucleotide sequence determined for the chicken liver cDNA consists of 1302 nucleotides including a 861 nucleotide region coding for 287 amino acids with a molecular weight of 32 914. Comparisons reported show that homologies are present between rabbit and chicken, some as high as 80% or greater within individual tryptic peptides. For the ten tryptic peptides compared, approximately 60% homology exists between corresponding peptides for the liver Nacetyltransferases of the rabbit and chicken (Ohsako et al., 1988). The product analogue, bromoacetanilide, causes irreversible loss of Nacetyltransferase activity proportional to the incorporation of the inhibitor with a 1:1 stoichiometry of inhibitor to enzyme. Amino acid analysis of inhibitortreated enzyme shows that cysteine is the nucleophilic target amino acid of the inhibitor. The reaction with bromoacetanilide is relatively rapid (pseudofirstorder rate constant=1 min−1) providing evidence that the active site cysteine is activated to its thiolate form (Andres et al., 1988).

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Table 6.1. Amino acid compositions of hepatic N-acetyltransferase from rapid acetylator mouse, rabbit, and pigeon, in comparison with a mean composition of 81 proteins.

1

Cysteine and cystine were determined as cysteic acid. was detected at a level of 2·1 residue mol-1 inthe mouse enzyme. 3 The tyrosine value for the mouse enzyme was estimated from the Leu/Tyr ratio determined for the rabbit enzyme. 4 Tryptophan was not determined for the mouse. 5 Data from Andres et al. (1987). 6 Data from Andres et al. (1983a). 7 Data from Holmquist (1978). 2 4-hydroxyproline

Kinetic properties Attempts to determine the biological basis of the acetylator polymorphisms in humans and animals have led to extensive studies of the kinetic properties of the liver acetylating enzymes. Comprehensive studies of the kinetic mechanism of rabbit liver N-acetyltransferases indicates a ‘one-site’ (classical) ping-pong mechanism for biological N-acetylation (Weber, 1973) in contrast to the ‘twosite’ (hybrid) ping-pong mechanism observed for the pigeon liver enzyme (Andres et al., 1983b). Studies of liver N-acetyltransferases of human, monkey and rat give results that are consistent with those of rabbit liver enzyme (Weber et al., 1968). The second half of the overall N-acetylation reaction (see Introduction) is sensitive to the structure, polarity and basicity of the acceptor amine (Andres et al., 1987). For weakly basic acceptor amines, such as p-nitroaniline, the secondhalf reaction is the rate-determining step for acetyl coenzyme A-dependent N-

172 CONJUGATION REACTIONS IN DRUG METABOLISM

acetyl transfer. But most drugs and other environmental aromatic amines are relatively strongly basic substances, and for those the rate of formation of the acetyl-enzyme intermediate (the first half-reaction) is rate-determining. Initial studies of the Michaelis-Menten constants of the human and rabbit enzymes failed to reveal any significant differences between the rapid and slow enzymes for substrates such as isoniazid and sulfamethazine. These observations led to the tentative suggestion that the acetylator polymorphism might be due to quantitative differences in an identical enzyme of the liver in humans and rabbits (Jenne, 1965; Weber et al., 1968). Later, however, we learned that a single enzyme was capable of acetylating the polymorphic substrate, sulfamethazine, and the monomorphic substrate, p-aminobenzoic acid (Patterson et al., 1980; Hein et al., 1982a). We also obtained limited evidence suggesting that the rapid and slow enzymes of rabbit liver exhibited six- to eight-fold differences in the Km for p-aminobenzoic acid (Weber et al., 1978). Further studies yielded convincing evidence that the slow enzyme of rabbit liver has a much lower Km for paminobenzoic acid (Table 6.2) and p-aminosalicylic acid (Hein et al., 1982c), whereas the lack of a significant difference in Km for substrates such as sulfamethazine and procainamide was confirmed (Table 6.2) (Andres and Weber, 1986). A definite difference has not been demonstrated in rapid and slow acetylator mice with p-aminobenzoic acid (Mattano and Weber, 1987), probably for technical reasons, but these mice show a Km difference for 2aminofluorene that is qualitatively similar to that observed for p-aminobenzoic acid in rabbits (Table 6.2). The hamster polymorphism presents a picture that differs from that of the rabbit and mouse polymorphisms. Cytosolic preparations of hamster livers contain two isozymes of acetyltransferase activity, one of which is polymorphically regulated by the acetylator gene locus and one which is not. Slow acetylator cytosols show a much higher Km for p-aminobenzoic acid than rapid acetylator cytosols; however, the slow acetylator cytosols show lower apparent Kms for the arylamine carcinogens 2-aminofluorene and 4aminobiphenyl (Table 6.2). This unusual result can be explained when the two isozymes are separated and studied individually, as discussed later (see Multiple acetyltransferase isozymes). In view of the findings with p-aminobenzoic acid and p-aminosalicylic acid in animals, we reinvestigated the relationship of the acetylator phenotype to the Michaelis-Menten constants in human tissues. Preliminary analysis of results of studies currently in progress indicates the general picture emerging for the rapid and slow enzymes of human liver is highly similar to that observed for the rabbit with p-aminobenzoic acid and sulfamethazine as substrates1. Thus the Kms differ significantly for p-aminobenzoic acid and those for sulfamethazine do not (Table 6.2).

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Table 6.2. Michaelis-Menten constants for liver cytosol N-acetyltransferases of rapid and slow acetylator rabbits, mice, and hamsters compared to rapid and slow acetylator human liver cytosol N-acetyltransferases for arylamines.

1 p-aminobenzoic acid, PABA; 2-aminofluorene, AF; p-aminosalicylic acid, PAS; sulfamethazine, SMZ; 4-aminobiphenyl, ABP.

In summary of the results for the human acetylation polymorphism as well as for the rabbit, mouse and hamster polymorphisms, we can say that each of these traits is determined by kinetic variants of a specific acetylating liver enzyme. Differences in the activity and affinity of the rapid and slow acetylator enzymes for the aromatic amine substrate p-aminobenzoic acid and/or p-aminosalicylic acid are conferred on the enzymes by the acetylator genes of those species. For all species examined so far the data in Table 6.2 suggest that the affinity of the slow acetylator variant significantly exceeds that of the rapid variant for certain aromatic amines, and that the slow variant is less able to catalyze the acetylation of aromatic amines. The data also suggest that the rapid and slow enzymes of each species are isozymes of a nearly identical liver acetylating enzyme expressed in each species, but that the primary structure of the parent enzyme differs from one species to another. Multiple acetyltransferase isozymes Measurement of the N- and O-acetyltransferase activities in hamster tissues provides additional insights into the complex role of acetylator genotype on these enzymes (Table 6.3). N-Acetyltransferase polymorphisms have been identified in hamster liver (Hein et al., 1982b, 1985a), blood lysates (Hein et al., 1986a), intestine, lung, kidney (Hein et al., 1987a),

b nmol

min-1 mg-1 mean values differ significantly (p<0.01) between acetylator genotypes; from Hein et al., 1986b. min-1 mg-1 from Hein et al., 1986b. c nmoles bound/15 min/mg protein/mg DNA; mean values differ significantly (p<0.01) between acetylator genotypes; from Hein et al., 1987a. d nmoles bound/20 min/mg protein/mg DNA; from Hein et al., 1986c.

a nmol

Table 6.3. Relationship of acetylator genotype to arylamine acetyltransferase activities in inbred hamster cytosols.

174 CONJUGATION REACTIONS IN DRUG METABOLISM

ACETYLATION 175

bladder (Hein et al., 1987c), and colon cytosols. In most tissues studied, levels of activity are highest in homozygous rapid acetylators, lowest in homozygous slow acetylators, and intermediate in F1 heterozygous acetylator progeny. Inheritance patterns of N-acetyltransferase activity across F1, F2, and backcross generations are most consistent with simple autosomal Mendelian inheritance of two co-dominant alleles at a single gene locus. Recently, Kato and co-workers (1988) have also identified rapid, intermediate, and slow acetylators among outbred Syrian hamster stocks in Japan and have shown polymorphic expression of arylamine N-acetyltransferase activity in hamster skin (Kawakubo et al., 1988). High levels of polymorphic AcCoA-dependent N-hydroxyarylamine Oacetyltransferase activity are expressed in hamster liver, intestine, kidney, and lung cytosols (Table 6.3). Thus, both 2-aminofluorene N-acetyltransferase and N-hydroxyaminofluorene O-acetyltransferase activities are regulated by acetylator genotype in cytosolic preparations of each tissue (Table 6.3). However, comparable determinations of N-hydroxy-N-acetylaminofluorene N, O-acetyltransferase and N, N-acetyltransferase activities in the same tissue cytosols indicate acetylator genotype-independent levels of activity (Table 6.3). Expression of the polymorphic and monomorphic activities of various arylamine and N-hydroxyarylamine N- and O-acetyltransferases has spurred investigations into the molecular basis for these relationships. Partial purification of inbred hamster liver cytosol has resulted in the identification of two acetyltransferase isozymes that differ in chromatographic properties, genetic control, and kinetic constants (reviewed in Hein, 1988). The two acetyltransferase isozymes of liver cytosol have been distinguished in vitro (Smith and Hanna, 1986) and in vivo (Smith and Hanna, 1988) by mechanism-based inactivators (suicide inhibitors). In addition to the studies of liver cytosol, two acetyltransferase isozymes have been reported in cytosols of hamster intestine (Smith and Hanna, 1986) and bladder (Yerokun et al., 1989) and in peripheral blood lysates (Hein et al., 1986a). Activity of the polymorphic acetyltransferase isozyme is acetylator-phenotype dependent, and that of the monomorphic acetyltransferase isozyme is not (Hein et al., 1985a, 1987a, 1987c). Both isozymes catalyze a diverse set of acetyltransferase reactions, but the relative specificity of the two isozymes differs for these reactions and is also substratedependent for each reaction. Their relative specificity accounts for the genotyperelated expression of the acetylation reactions in vitro and also presumably in vivo. Thus, AcCoA-dependent N-acetyltransferase activity towards paminobenzoic acid, p-aminosalicylic acid, and 2-aminofluorene, and AcCoAdependent O-acetyltransferase activity towards N-hydroxyaminofluorene are acetylator genotype-dependent in hamster liver cytosol because of the relatively high contribution of the polymorphic isozyme. In contrast, AcCoA-dependent N-acetyltransferase activity towards procainamide and isoniazid, AcCoAdependent O-acetyltransferase activity towards N-hydroxy-3, 2'-

176 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 6.2. N-, O-, and N, O-Acetyl transfer probably involves a common intermediate in the mouse.

dimethylaminobiphenyl, and N-hydroxy-N-acetylaminofluorene N, Oacetyltransferase activities are acetylator genotype-independent in hamster liver cytosol because of the relatively high contribution of the monomorphic isozyme. Each of the acetyl transfer reactions catalyzed by the polymorphic isozyme can be shown to be acetylator genotype-dependent when it is separated from the monomorphic one (Hein, 1988). These diverse catalytic activities are illustrated in Figure 6.1. The molecular basis of the hamster acetylator polymorphism thus has much in common with those of the rabbit and mouse. Each species is characterized by a single genetically variant N-acetyltransferase or a family of variant Nacetyltransferases, but the mouse (Hein et al., 1988) and the hamster (Hein et al., 1985b, 1987a, 1987c) also express a monomorphic acetyltransferase isozyme. In the mouse, the homogeneous preparations of the rapid acetylator isozyme catalyzes N-acetylation, O-acetylation and N, O- acetylation of amine carcinogens (Figure 6.2; Mattano et al., 1989), but neither the Nhydroxyarylamine O-acetyltransferase activity nor the N-hydroxyaminofluorene N, O-acetyltransferase activity differs according to the acetylator phenotype (Mattano et al., 1989). In the rabbit, 2-aminofluorene N-acetyltransferase and N-hydroxy-Nacetylaminofluorene N, O-acetyltransferase activities of liver cytosol co-purify to electrophoretic homogeneity (Glowinski et al., 1980), although the Nhydroxyarylamine O-acetyltransferase activity was not tested. The hamster, on the other hand, has two distinct acetyltransferase isozymes, one of which catalyzes N-acetylation, O-acetylation and N, O-acetylation. Observations by Saito et al. (1986b) that these three activities co-purify to electrophoretic

ACETYLATION 177

homogeneity provides support for a common acetyltransferase enzyme in the hamster (Figure 6.1). A genetic acetylation polymorphism has been identified in inbred rat strains, but it has been studied less extensively than the human trait and those of the rabbit, mouse and hamster. A survey of inbred rat strains indicates that paminobenzoic acid N-acetyltransferase activity of liver cytosols varies approximately 25-fold (0·09–2·51 nmol min−1 mg−1 protein) between strains, whereas isoniazid and 4-aminobiphenyl N-acetyltransferase activities vary only approximately three-fold (Weber, 1987). Recent studies have shown that the acetylator trait is expressed in a coordinate manner in the liver and pancreas (Martell and Weber, 1988). While additional studies are necessary to characterize the inheritance and biochemical properties of the rat trait, these results suggest that the rat trait may resemble the hamster trait more closely than that of the rabbit, mouse or human. Further exploration of the function of Nacetyltransferase activity in the pancreas should lead to a better understanding of the relationship between acetylator status and diabetes mellitus. N, O-Acetyltransferase of rat tissues has been extensively investigated. The enzyme is widely distributed in tissues. Highest activities are in liver with significant activity in mammary tissue, gastrointestinal tract, kidney, stomach, spleen and lung (Booth, 1966; Bartsch et al., 1972, 1973; King, 1974; King and Olive, 1975). Multiple forms of the enzyme that differ in substrate specificity have been reported for rat mammary tissue and small intestine (Bartsch et al., 1973; Olive and King, 1975) and liver (Beland et al., 1980; Shirai et al., 1981; Glowinski et al., 1983). Although N, O-acetyltransferase and the genetically variant N-acetyltransferase of liver of rabbits (Glowinski et al., 1980) and mice (Mattano et al., 1989) appear to be associated with a single protein, preliminary evidence for separation of rat liver N- and N, O-acetyltransferases has been reported (Land et al., 1987). 6.3. Acetylation in isolated cells and in vivo The occurrence of acetylation of drugs and other aromatic amines in isolated cells of liver and numerous other tissues is well established. Acetylation of drugs by isolated liver cells has been demonstrated with sulfonamides by several investigators (Govier, 1965; Suolinna, 1980; Olsen and Morland, 1981; Olsen et al., 1981). Govier (1965) originally reported that acetylation of sulfanilamide and p-aminobenzoic acid occurred in the reticuloendothelial cells of rabbit liver. These observations received support from studies of Notter and Roland (1978). Further studies by Suolinna (1980) and by Olsen and his colleagues (1981) of rabbit and rat liver cells show that sulfamethazine, sulfanilamide and p-

178 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 6.4. The acetylation pharmacogenetics of 2-aminofluorene (AF) and 2acetylamino-fluorene (AAF) in rapid (C57BL/6J) and slow (A/J) acetylator mice.

1 Numbers

in parentheses are numbers of animals studied.

aminobenzoic acid are acetylated by hepatocytes while non-parenchymal cells of liver appear to lack this ability. The pharmacokinetics of 2-aminofluorene disappearance and its acetylation to 2-acetylaminofluorene have been studied in primary cultures of hepatocytes isolated from rapid (C57BL/6J) and slow (A/J) acetylator mice (Weber et al., 1984). The disappearance of 2-aminofluorene and the appearance of 2acetylaminofluorene were both time- and concentration-dependent. 2Aminofluorene disappearance was biphasic, following pseudozero-order kinetics at 2-aminofluorene concentrations in the medium greater than 8 µM, and following first-order kinetics below that concentration. Under first-order conditions, 2-aminofluorene disappearance was two to three times faster in rapid acetylator hepatocytes than in slow acetylator hepatocytes (Table 6.4). Under saturating conditions, the rate of 2-acetylaminofluorene formation was also approximately twice as fast in rapid as in slow acetylator hepatocytes (Table 6.4). These rates were determined in the presence of paraoxon, an inhibitor of deacetylation, to avoid the effect of deacetylation by C57BL/6J and A/J mouse hepatocytes (Hultin and Weber, 1987). The influence of N-acetyltransferase activity on 2-aminofluorene elimination has also been investigated in intact rapid and slow acetylator mice (Hultin and Weber, 1984). Elimination rates of 2-aminofluorene from the blood of mice

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administered a single i.p. dose of 30, 50, or 100 mg/kg body weight of 2aminofluorene is dose-dependent in both strains. The average rate of 2aminofluorene elimination at those doses averages about three times faster in rapid acetylator mice than in slow acetylator mice (Table 6.4), and the differences are statistically significant at each dose examined. 6.4. The biological significance of acetylation Conjugation by acetylation and person-to-person differences in the capacity for acetylation exert significant effects on individual responses to drugs and other environmental arylamines and hydrazines (Weber and Hein, 1985; Weber, 1987). Slow acetylators are often more prone than rapid acetylators to drug-induced toxicities—such as isoniazid-induced peripheral nerve damage, to procainamideand hydralazine-induced lupus erythematosus, to phenytoin toxicity if phenytoin and isoniazid are co-administered, to sulfasalazine-induced hematologic disorders and to sulfonamide-induced lesions of the kidney, skin and other tissues. Rapid acetylators, on the other hand, are more likely to show reduced levels of therapeutic drug efficacy that are clinically significant. Such effects are reported for isoniazid-treated tuberculosis patients, for hydralazine-treated hypertensive patients, for procainamide-treated patients with cardiac arrhythmias and for sulfasalazine-treated rheumatoid arthritis patients (reviewed in Weber, 1987). The predisposition of slow acetylators to drug toxicity and of rapid acetylators to drug ineffectiveness is probably explained by the higher serum concentrations of drug attained at any time after drug ingestion in slow compared to rapid acetylators. Human epidemiology studies suggest that certain sporadic disorders may be more prevalent in one acetylator phenotype than the other (Table 6.5; summarized from Weber, 1987). Thus, slow acetylators appear to be more susceptible to bladder cancer (S/R relative mean excess=1·36, , ), but the excess is mainly accounted for by workers occupationally exposed to aromatic amines (S/R relative mean excess=1·70, , ). In contrast, rapid acetylators may be more susceptible to colorectal cancer (R/S relative mean excess=3·03, ). Human colonic mucosa contains high levels of N- and O-acetyltransferase activities (Flammang et al., 1988) that may enhance the genotoxicity of aromatic amine food pyrolysis products (Sugimura and Sato, 1983) in transit through the bowel. Thus in some instances aromatic amines may be involved in the genesis of colorectal cancer, and the rapid acetylator phenotype may predispose individuals to this disorder2.

180 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 6.5. The human N-acetylation polymorphism as a risk factor in various sporadic human disorders1.

1 Summarized 2

from Weber, 1987. S=slow acetylator; R=rapid acetylator.

A strong statistical association suggests that rapid acetylation may be a factor affecting individual susceptibility to Type I diabetes mellitus in Europeans (Weber, 1987). The relationship of human acetylator status to the efficacy and toxicity of arylamine and hydrazine drugs has been firmly established for only a few drugs; claims for such a relationship to other conditions are just emerging, and they must be carefully weighed before being accepted. Association of the acetylator status with drug toxicity, teratogenicity and DNA damage has been investigated in intact rapid and slow acetylator animals, in tissues of these animals and in cells isolated in culture from them. For example, the genetic difference in acetylation is related to differences in neurotoxicity (Hein and Weber, 1984) induced by isoniazid in rapid and slow acetylator (New Zealand White) rabbits, and to differences in procainamideinduced antinuclear antibodies (Tannen and Weber, 1980), glucocorticoidinduced cleft lip (Liu and Erickson, 1986), to phenytoin-induced cleft lip with or without cleft palate (Karolyi et al., 1987) and to aromatic amine carcinogeninduced DNA damage (Weber and Levy, 1988a, 1988b; Levy and Weber, 1988a, 1988b, 1988c) in rapid (C57BL/6J) and slow (A/J) acetylator mice. To improve the genetic analysis of such pharmacogenetic differences, our laboratory (W.W.W.) has constructed congenic rapid (A.B6-Natr) and slow (B6.A-Nats) acetylators from the parental C57BL/6J and A/J inbred mouse strains (Mattano et al., 1988). Studies of those congenic acetylator mouse strains provide much stronger evidence for the acetylator gene as a determinant of susceptibility to aromatic amine carcinogen-induced DNA adduct formation (Levy and Weber, 1988a, 1988b, 1989; Weber and Levy, 1988a, 1988b; Weber et al., 1988) as demonstrated by a high correlation between the extent of DNA adduct formation and metabolic characteristics of the acetylator congenic strains (Table 6.6). There is a six- to seven-fold difference in the extent of 2aminofluorene-induced DNA adduct formation in liver and in the rate of acetyl coenzyme A-dependent N-acetylation of p-aminobenzoic acid and

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Table 6.6. N-acetyltransferase (NAT) activity and DNA adduct formation of liver in rapid and slow acetylator parental and congenic mouse strainsa.

a

B6.A and A.B6 are abbreviations for B6.A-Nats and A.B6-Natr, respectively. The genotypes given refer to homozygous rapid (rr) and homozygous slow (ss) acetylator mice. Numbers in parentheses are numbers of animals studied. Results are expressed as the mean ±standard error. Adapted from Hein et al. (1988) and Weber and Levy (1988b). b PABA=p-aminobenzoic acid; AF=2-aminofluorene.

2- aminofluorene between the congenic strains compared to the 2·5 to 3-fold difference in both of these parameters between parental C57BL/6J and A/J mice (Hein et al., 1988). In addition to the effects of the acetylator gene, it is evident from our findings that other polymorphic genes of the mouse genome are capable of modifying the metabolic and toxic responses observed. The identity of the background genes is yet unknown, although the difference in the genes that control deacetylation in the parental mouse strains are in the appropriate direction and are of such a magnitude (Hultin and Weber, 1987) to suggest that they may be partly responsible for the greater differences in acetylation and DNA-adduct formation observed in the congenic strains compared to the parental strains. Other studies of the congenic acetylator strains suggest that the acetylator gene may influence the susceptibility of mice to glucocorticoid-induced cleft palate (I.J.Karolyi, R.P.Erickson, S.Liu and L.Killewald, unpublished observation). In that situation, the movement of the Nat region from the susceptible A/J strain to the nonsusceptible C57BL/6J strain results in a strain (B6.A-Nats) that is nearly identical in susceptibility to that of the A/ J strain. Hepatocytes isolated from rapid and slow acetylator rabbits have been used to examine the effects of acetylation on toxicity. Thus the DNA damage measured as unscheduled DNA synthesis that is caused by aromatic amine carcinogens, such as 2-aminofluorene and benzidine, is more severe in hepatocytes of the rapid acetylator than the hepatocytes of the slow acetylator (McQueen et al., 1982, 1983). Bacterial strains with deficiencies in their acetylating capacity have also served as experimental models to investigate the influence of acetylation on aromatic amine-induced mutagenesis (McCoy et al., 1983; Saito et al., 1983; Josephy, 1986). Thus, strains of Salmonella typhimurium have been used to determine the extent to which N-, O- and N, O-acetyl transfer may be

182 CONJUGATION REACTIONS IN DRUG METABOLISM

implicated in that process. Recently, Watanabe et al. (1987) have constructed plasmid-containing derivatives of Salmonella typhimurium strain 1538/1, 8-DNP (pYG 121 and 122) with high levels of arylamine N-acetyltransferase. Preliminary studies of responses to benzidine and 2-aminofluorene indicate the resultant strains are highly sensitive to arylamine mutagenicity in the Ames test (Josephy and LeBruin, 1988). 6.5. Practical considerations Enzyme purification Procedures for purification of arylamine N-acetyltransferases of liver are described elsewhere (Weber and King, 1981; Saito et al., 1986b; Andres et al., 1987; Kabishev and Patrushev, 1987; Deguchi et al., 1988; Mattano et al., 1989; Trinidad et al., 1988a). Use of those techniques has achieved near homogeneous preparations of New Zealand White rabbit liver N-acetyltransferase and homogeneous preparations of mouse (C57BL/6J) (Mattano et al., 1989) and chicken (Deguchi et al., 1988) liver N-acetyltransferases. The homogeneous preparation of mouse liver enzyme was achieved by 10 000–18 000-fold purification. A single band of enzyme protein of molecular weight 31 500 was obtained on polyacrylamide gel. The enzyme had N-acetyl, O-acetyl and N, Oacetyl transfer activities in the ratio of 1000–2000/3–4/1, respectively (Mattano et al., 1989). Mature New Zealand White rabbits that can be identified by acetylator phenotype (Weber et al., 1976) are usually available locally. C57BL/6J (rapid acetylator) and A/J (slow acetylator) inbred mice are available from Jackson Laboratories, Bar Harbor, ME (USA). Homozygous rapid acetylator inbred Syrian hamsters (Bio.87.20) are available from the Bio Breeders, Inc., Watertown, MA (USA), whereas homozygous slow (Bio.82.73/H) and obligate heterozygous F1 hybrid hamsters are maintained at University of North Dakota School of Medicine, Grand Forks, ND (USA). Chromatography materials for purification of the rapid and slow acetylator liver isozymes are available from Sigma Chemical Co. and/or the BioRad Co. Mating pairs of selected inbred rat strains are available from the NIH Division of Animal Resources, Bethesda, MD (USA) and from the Department of Pathology, University of Pittsburgh, Pittsburgh, PA (USA). Enzyme assays and data analysis Procedures for measuring acetyltransferase activities are listed in Table 6.7. Acetyl coenzyme A is the best acetyl donor for the N-acetyltransferase reaction. Only a few of many possible acetyl acceptor amine substrates are listed in the Table. The spectrophotometric assay procedure of Andres et al. (1985) is superior to alternative procedures (Weber and Glowinski, 1980; Weber and King, 1981) because it enables measurements of N-acetyltransferase activity at

ACETYLATION 183

Table 6.7. Procedures for measurement of acetyltransferase activity in tissue preparationsa.

a Additional procedures for determining acetyltransferase activities are referenced in Weber (1987). b AT, acetyltransferase. c Different procedures are described for intermolecular and intramolecular N, O-AT activities.

low physiological acetyl coenzyme A concentrations and because it is applicable to a wide range of acceptor amines and biological systems (Andres et al., 1987; Mattano and Weber, 1987; Mattano et al., 1988, 1989). With this procedure, Nacetyltransferase activity is measured by a decrease in Schiff’s base formation with dimethylaminobenzaldehyde, and coenzyme A, the inhibitory product formed during the reaction, is recycled by the AcCoA regenerating system. The procedures listed in Table 6.7 are adapted for use with small quantities of enzyme. In the colorimetric and in certain fluorimetric procedures, the Nacetylated product is not detected and its rate of formation is determined from the rate of disappearance of the acceptor amine substrate. Isotopic methods involve incubation of the enzyme in the presence of labelled acetyl coenzyme A (either 3H- or 14C-acetyl coenzyme A) or the labelled acceptor amine substrate followed by the isolation and measurement of the radio-labelled product. Alternative procedures for measuring N-acetyltransferase activity, such as those with drugs (e.g. isoniazid) or a procedure using dithio(2-nitrobenzoic acid), have been previously summarized (Weber and Glowinski, 1980). Unlabelled acetyl coenzyme A is available from Pharmacia/P-L Bio-chemicals, Inc. and Sigma Chemical Co., and radio-labelled acetyl coenzyme A is available from New England Nuclear Co. Other chemicals for the N-acetyltransferase

184 CONJUGATION REACTIONS IN DRUG METABOLISM

assay recycling procedure (Andres et al., 1985) are available from commercial sources such as Sigma Chemical Co. Sources of specialized chemicals are given in the original publications cited. Statistical analysis of the kinetic data is usually performed according to Wilkinson’s method (1961) with the aid of the HYPER and PING PONG computer programs of Cleland (1963, 1967). Measurement of DNA damage induced by aromatic amines The relation of acetylation of arylamines to DNA damage has been investigated by several methods including unscheduled DNA synthesis, DNA strand breaks and carcinogen-DNA adduct formation. Measurement of DNA-adduct formation has most often been attempted with 3H or 14C labelled carcinogens. Immunological methods employing antibodies directed against a specific carcinogen adducted nucleotide can be used throughout the range of expected adduct formation (Poirier, 1981). A significant limitation on the use of antibodies or radioactive carcinogens is that specific antibodies must be produced or labelled carcinogens must be obtained or synthesized. Additionally, in the case of radio-labelled carcinogens, testing at high dose levels is extremely expensive, or if unlabelled carcinogen is added, specific activity falls, thereby decreasing sensitivity. An alternative approach to measuring adduct formation is 32P-post-labelling with high specific activity [γ-32P]ATP andpolynucleotide kinase (Randerath et al., 1981). With this method, adducts of DNA (or RNA) and a wide variety of compounds, or unfractionated mixtures of compounds, can be measured (Reddy et al., 1984; Levy and Weber, 1988a, 1988b, 1988c). Following the postlabelling of the carcinogen-nucleotide adduct, analysis is carried out by chromatography. In the original procedures (Randerath et al., 1981; Reddy et al., 1984) thin layer chromatography is used. After sequential development of the plates in several solvents, autoradiograms are prepared and areas containing radioactivity are cut out or scraped and counted by scintillation methods. We have recently devised a modification of the 32P-post-labelling technique (Levy and Weber, 1988c, 1989; Weber and Levy, 1988a, 1988b) in which the labelled adducts are analyzed by high performance liquid chromatography (HPLC; see Table 6.6). The latter method resolves adducts by ion pairing on a reversed phase column. The radioactivity of the effluent is measured either by a flowthrough detector or by collecting samples and counting in a liquid scintillation counter. The HPLC method is more rapid than thin layer chromatography and requires considerably less handling of radioactive material.

ACETYLATION 185

Acknowledgements This work was partially supported by USPHS Grants GM 27028, CA 39018, CA 34627, and RR 08248. Notes 1. A preliminary account of this work was presented at the Federation of American Biological Societies, May 1–5, 1988, Las Vegas, NV, USA (Kilbane et al., 1988). 2. Previous data (Bulovskaya et al., 1978; Cartwright, 1984) suggested a relationship between rapid acetylation and breast cancer in women (Evans, 1986). Estimation of the susceptibility of women to breast cancer from more recent reports (Ladero et al., 1987; Philip et al., 1987) combined with early data (Bulovskaya et al., 1978) no longer shows significant association between acetylator status and the overall occurrence of breast cancer (relative mean excess R/S=1·2, X=1·49, p<0·10) (Weber and Levy, unpublished results). However, the data do suggest an increased trend toward rapid acetylation at advanced stages of the disease at presentation (Philip et al., 1987).

Abbreviations NAT DMSO cDNA

N-acetyltransferase dimethylsulfoxide complementary DNA References

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 7 O-Methylation Dhiren R.Thakker1 and Cyrus R.Creveling2 1

Department of Drug Metabolism, GLAXO Research Laboratories, Research Triangle Park, NC 27709, USA 2

Laboratory of Bioorganic Chemistry, NIDDK, NIH, Bethesda, MD 20892, USA

7.1.

INTRODUCTION

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7.2.

THE METHYL DONOR: S-ADENOSYLMETHIONINE

193

7.3.

THE SUBSTRATES

193

7.4.

CATECHOL-O-METHYLTRANSFERASE

195

7.5.

7.6.

Distribution and purification

195

Multiple forms

196

Kinetic properties

197

Active site

198

IN VITRO INHIBITORS OF COMT

201

Analogues of S-adenosylhomocysteine

201

Isosteres of catechol substrates

202

Metal ions

203

THE MULTIPLE ROLES OF COMT IN VIVO

203

The central nervous system

204

Median eminence and pituitary

206

The reproductive system

207

The cardiovascular system

210

Erythrocytes

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The metabolism of catechols in brain

213

L-DOPA therapy of Parkinson’s disease

214

The barrier role of COMT

216

7.7.

O-METHYLATION OF DRUGS IN VIVO

217

7.8.

IN VIVO INHIBITORS

219

7.9.

CONCLUSION

220

ABBREVIATIONS

221

REFERENCES

221 7.1. Introduction

Methyl transfer is one of the most extensively carried out reactions in nature. The wide ranging physiological consequences of the methyl-transfer reaction can be attested by the great diversity of the methyl-acceptor substrates found in the biological systems; these include proteins, nucleic acids, phospholipids, and many small molecules of diverse structures. All of these methyl-transfer reactions utilize one of the two methyl-donor substrates, i.e. Sadenosylmethionine (SAM) or N5-methyltetrahydrofolic acid. When SAM is the methyl-donor, the transfer of the methyl group occurs to a sulfur-, nitrogen-, or oxygen-nucleophile. In the present chapter, we will restrict our discussion to the O-methylation reaction. Among the conjugation reactions, O-methylation is unique in that the products of the reaction are less polar than the substrates. The most commonly encountered O-methylation reaction involves methylation of one of the phenolic groups of a variety of endogenous and xenobiotic catecholic compounds and is catalyzed by the enzyme catechol O-methyltransferase (EC 2. 1.1.6) (COMT) (Axelrod and Tomchick, 1958). Another enzyme, hydroxyindole O-methyltransferase (EC 2.1.1.4), catalyzes O-methylation of the phenolic group of N-acetylserotonin to form melatonin (Axelrod and Weissbach, 1960). A third O-methylation reaction, catalyzed by proteincarboxy O-methyltransferase (EC.2.1.1.24), results in the esterification of aspartic and glutamic acid residues on several proteins. The latter reaction appears to play a significant role in the regulation of the function of highly specialized proteins. Among these O-methylation reactions, the one catalyzed by COMT is the most important metabolic conjugation reaction and will be discussed in detail here.

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7.2. The methyl-donor: S-Adenosylmethionine Since its discovery in 1953 (Cantoni, 1953), SAM has been implicated in numerous transmethylation and transalkylation reactions, several of which are critical for the survival of cells (see Borchardt et al., 1986). The enzyme ATP: Lmethionine S-adenosyltransferase (EC 2.5.1.6) catalyzes the condensation of ATP and L-methionine, which results in the synthesis of SAM and the release of inorganic pyrophosphate as well as phosphate. The enzyme requires divalent cations for its activity and is activated by monovalent cations. The diastereomer of SAM, synthesized by the enzymic condensation, has (S) configuration at the sulfonium centre (de la Haba, et al., 1959; Conforth et al., 1977). This diastereomer is required for all the methyltransfer reactions (de la Haba et al., 1959; Zapia et al., 1969, Borchardt et al., 1976d). Interestingly, any change in the base, sugar or amino acid moiety causes a dramatic increase in the Km values for the methyltransferases (Borchardt et al., 1976d). Although some SAM is produced by the donation of the methyl group of N5methyltetrahydrofolate to S-adenosyl-L-homocysteine, methionine is the major source of the S-methyl group in SAM. This is evident from a rapid increase in the levels of SAM in various tissues after administration of methionine either orally or intraperitoneally (cf. Mulder and Krijgsheld, 1984). It is interesting, however, that the levels of the enzyme methionine adenosyltransferase activity in various organs of rats do not correlate well with the levels of SAM. While the specific activity of the enzyme (pmole product min−1/mg protein) ranged from a low value of 40 in heart and brain of rats to a high value of 7700 in liver (over 180-fold increase), the concentrations of SAM ranged from a low of 25 (nmoles/ g of tissue) in brain to a high of 68 in liver (less than three-fold increase; Eloranta, 1977). Similarly, despite 13-fold higher specific activity of the enzyme in pancreas over that in heart, the levels of SAM in both the organs were similar. Thus, factors other than the level of methionine adenosyltransferase must play an important role in controlling the levels of SAM in different tissues and organs. Furthermore, drugs or xenobiotics, that can be methylated by SAMdependent methyltransferases, can also affect the levels of SAM in various tissues, although such effects are expected to be transient (Fuller et al., 1983). For further details on SAM metabolism and availability, recent reviews are available (Mulder and Krijgsheld, 1984; Borchardt et al., 1986). 7.3. The substrates As shown in Figure 7.1, COMT catalyzes transfer of the methyl group from SAM to either the meta- or the para-hydroxyl group of a substituted catechol

194 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 7.1. The reaction catalyzed by COMT. The methyl group of SAM that is being transferred is shown in bold letters. The terms meta and para indicate meta- and para-Omethylated products when R2 and R3=H. COMT has a broad selectivity for catechol substrates and can accommodate a variety of R1, R2, and R3 substituents.

derivative (R2=R3=H) to form two monomethyl phenols and the demethylated product S-adenosylhomocysteine (SAH). The diverse catecholic substrates of COMT include catecholamines (dopamine, norepinephrine, epinephrine), amino acids (L-dopa), alkaloids (dihydroxyisoquino-lines, apomorphine) and steroids (2- and 4-hydroxyestradiols, 2- and 4-hydroxyestrones). The enzyme accommodates catechol substrates with positively charged, negatively charged or neutral substituents (Creveling et al., 1970; Creveling et al., 1972). Catechols with multiple substituents in the aromatic ring (R1, R2, R3 in Figure 7.1) are also good substrates of COMT as evidenced by low Km values for 2-hydroxy-17βestradiol and 4- hydroxyestrone (15 µM and 20 µM, respectively) compared to the Km value of 300 µM for epinephrine (Ball et al., 1972a). COMT has a stringent, if not absolute, requirement for the catechol functionality. The report by Qu et al., 1983) that dopamine-3-O-sulfate and dopamine-4-O-sulfate can be O-methylated by a rat liver COMT preparation suggests that the enzyme does not require a catechol functionality for its activity; however, the identification of the products was somewhat tentative. Hence, further studies are required to determine if monosulfates of catecholic derivatives can be O-methylated by COMT. O-Methylation of L-ascorbic acid by rat liver COMT to form its 2-Omethylated derivative represents a rare example of a non-catecholic substrate

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being O-methylated by COMT (Blaschko and Hertting, 1971). The 2-Omethylated derivative of L-ascorbic acid could also be detected in urine after administration of 1-[14C]-L-ascorbic acidtorats. The authors have suggested that the enediol form of 3-keto-L-gulonic acid contributes to the structure of Lascorbic acid, and that the enediol functionality, because of its isosterism with a catechol moiety, is methylated by COMT. Arnett et al. (1977) provided another example of bioisosterism when they demonstrated that benzimidazole could be N-methylated by a partially purified bovine liver COMT. The unusual catalytic activity of COMT was rationalized based on the isosterism between the two tautomeric forms of benzimidazole and the two stable conformations of a catechol with an intramolecular hydrogen bond. 7.4. Catechol-O-methyltransferase Distribution and purification COMT is one of the most widely distributed enzymes with its presence extending from yeast (Veser, 1987), through the plant kingdom (Hermann et al., 1987) to various species of invertebrates and vertebrates including humans (Guldberg and Marsden, 1975; Borchardt, 1980; Grossman et al., 1985; Nissinen et al ., 1988a and references therein). Liver is particularly rich in this enzyme, and hence, the liver enzyme from rats and humans has been extensively examined. In all the tissues, the bulk of COMT activity is found in the cytosol or loosely associated with the membranes so that the enzyme activity is recovered in the soluble fractions. However, existence of the membrane-bound COMT in a variety of tissues has also been demonstrated (cf. Borchardt, 1980; Rivett and Roth, 1982; Nissinen, 1984; Grossman et al., 1985 for references). When the membrane-bound COMT from rat liver, brain and heart microsomes was solubilized and partially purified, it appeared to have properties very similar to that of the soluble enzyme (Borchardt et al., 1974a; Borchardt and Cheng, 1978). The enzyme has been partially purified from various sources to different extent with the use of classical protein purification techniques (Nikodijevic et al. 1970; Veser et al., 1979; Jeffrey and Roth, 1985; Veser and Martin, 1986; Veser and May, 1986; Nissinen et al., 1988b; and references in Borchardt, 1980 and Grossman et al., 1985). Various affinity chromatography techniques have also been attempted for rapid purification of COMT (Creveling et al., 1973; Borchardt et al., 1975; Gulliver and Wharton, 1976; Veser and May, 1986). Use of affinity chromatography along with the classical protein purification techniques has allowed the isolation of homogeneous COMT from rat liver (Borchardt et al., 1975; Grossman et al., 1985).

196 CONJUGATION REACTIONS IN DRUG METABOLISM

Multiple forms Because of the wide distribution of COMT among species as well as among tissues in a given species, the question regarding the multiplicity of enzymic forms needs to be addressed. The following observations have been used to suggest the existence of multiple forms of COMT: (1) A variety of purification methods have yielded COMT from a large number of tissues in rats, humans and pigs with apparent molecular weights ranging from 23000 to 29000 daltons (Jeffrey and Roth, 1985; Veser and Martin, 1986; Nissinen et al., 1988b; and references in Borchardt, 1980, and Grossman et al., 1985); the presence of several enzymic forms with much larger molecular weights (45000 to 65000 daltons) also have been reported (Huh and Friedhoff, 1979; Grossman et al., 1985; Veser and Martin, 1986). (2) Isoelectric focusing technique has revealed the existence of COMT molecules with several different pI values in rats (Grossman et al., 1985; Huh and Friedhoff, 1979, and references therein) and in humans (Jeffrey and Roth, 1985). (3) The presence of both soluble (S-COMT) and membrane-bound COMT (MB-COMT) has been demonstrated (cf. references cited in ‘Distribution and purification’ section) (4) Sedimentation during sucrose density gradient ultracentrifugation revealed the presence of two forms of COMT with distinguishable molecular properties such as Stokes radius, sedimentation coefficient, frictional coefficient, and diffusion coefficient (Huh and Friedhoff, 1979). (5) Various forms of COMT from rats, separated on the basis of their molecular weights or their net charge, were found to be immunoreactive to the antibody produced against homogeneous rat liver COMT (molecular weight-23000 daltons) (Grossman et al., 1985; Heydorn et al., 1987). (6) Different ratios of the products formed by meta or para O-methylation of 4substituted catechol substrates by different COMT preparations have been observed (Marzullo and Friedhoff, 1975; Bade et al., 1976). It appears from the above observations that several molecular forms of COMT exist with different molecular weights and different physicochemical properties. However, a functional basis for the existence of such a large number of molecular forms of the enzyme is not at all clear. None of the studies cited above provided evidence for significant differences among the various molecular forms in substrate selectivity, in catalytic mechanism or in kinetic parameters. Although the membrane-bound enzyme appears to have a much lower Km for the catechol substrates when compared with the S-COMT (Assicot and Bohuon, 1971; Tong and D’Iorio, 1977; Rivett and Roth, 1983a, 1983b; Rivett et al., 1983; Jeffrey and Roth, 1984), its higher affinity for catechols is lost upon solubilization (Borchardt and Cheng, 1978; Goldberg and Tipton, 1978). Furthermore, upon purification, the MB-COMT exhibits similar kinetic and biochemical properties as the soluble form (Grossman et al., 1985). A difference in the ratio of meta and para-O-methylated products formed by different preparations of COMT has

O-METHYLATION 197

been used as an evidence for the existence of different enzymic forms of COMT (Marzullo and Friedhoff, 1975; Bade et al., 1976). However, the use of product ratios to demonstrate significant differences in product composition can be misleading. For example, Marzullo and Friedhoff (1975) showed that their two COMT preparations gave a meta/para ratio of 5.6 (COMT-A) and 11·7 (COMTB) when 3, 4-dihydroxybenzoic acid was used as a substrate. They used this as evidence for the existence of two different forms of COMT. However, a change in the meta/para ratio from 5·6 to 11·7 would mean an increase of the meta-Omethylated product from 85% to 92%. It would be inappropriate to propose the existence of a different form of COMT based on a small increase in the percentages of one of the products. The question as to why do we see so many variations in the physicochemical properties of various COMT preparations still awaits a satisfactory answer. Since the enzyme appears to be glycosylated (Creveling, 1988), differences in the glycosylation among different preparations of COMT need to be carefully examined. Of course, a definitive answer can be obtained only by determination of the amino acid sequence(s). Kinetic properties COMT catalyzes transfer of a methyl group from SAM to one of the phenolic groups of a catechol in the presence of Mg+2 ions. Other divalent cations can be substituted for Mg+2 with different degrees of success (Axelrod and Tomchick, 1958). The optimum catalysis takes place in the pH range between 7·3 and 8·2 (cf. Borchardt, 1980). A second pH optimum has been reported in many instances at pH values near 9 (Flohe and Schwabe, 1970; Ball et al., 1972b and references therein). This may represent ionization of a phenolic group of the catechol substrate. Initial velocity studies and product inhibition studies with the soluble and membrane-bound COMT from human brain indicate an ordered mechanism in which SAM binds with the enzyme first followed by the catechol substrate (Jeffrey and Roth, 1985 and references therein). Studies with the soluble enzyme from rat brain also indicate a similar mechanism (Tunnicliff and Ngo, 1983). In contrast, Flohe and Schwabe (1970) and Coward and Slisz (1973) suggested a rapid equilibrium random mechanism for a partially purified rat liver preparation of COMT. Their conclusions were based on initial velocity studies and product inhibition studies, respectively.

198 CONJUGATION REACTIONS IN DRUG METABOLISM

Active site Functional groups Despite many reports on isolation and purification of COMT from a variety of sources, only limited information is available regarding the structure of the enzyme, the nature of the active site, the identity of the functional groups, and the role of the functional groups in the catalytic process. Through the use of functional group reagents (Borchardt and Thakker, 1976) and affinity-labelling reagents (Lutz et al., 1972; Borchardt and Thakker, 1975a, 1975b; Borchardt, 1975; Borchardt et al., 1976a, 1976b, 1976c; Borchardt and Bhatia, 1982; Borchardt and Huber, 1982), the presence of two essential nucleophilic functional groups at the active site has been proposed. At least one of these groups appears to be a sulfhydryl group. The presence of an arginine group at the active site was suggested based on the inactivation of the enzyme by phenyl glyoxal (Tunnicliff and Ngo, 1983). However, the possibility cannot be ruled out that phenyl glyoxal was reacting with a sulfhydryl or an amino group at the active site of COMT. Methyl-transfer reaction The presence of critical nucleophilic groups at the active site and the initial proposal of a ping-pong kinetic mechanism (Borchardt, 1973) raised the possibility that the methyl group was initially transferred from SAM to a nucleophile at the enzyme active site and then to a phenolic group of the catechol substrates. However, elegant studies by Woodard et al. (1980) and Floss and Woodard (1982) established that the methyl transfer from SAM to the catechol substrates was accompanied by inversion of the configuration of the chiral, double-labelled CH2H3H S-methyl group. These results made a strong case for a direct transfer of the methyl group from SAM to a phenolic group of the catechol substrates by a SN2 mechanism. Previously, Higazi et al. (1976) had proposed an SN2 mechanism by demonstrating an inverse α-deuterium secondary isotope effect (VH max/VD max=0.832) during the COMT-catalyzed methyl transfer from SAM to 3, 4–dihydroxyaceto-phenone. Regio-selectivity of COMT The methyl transfer catalyzed by COMT occurs predominantly to the meta phenolic group of catecholamines. This selectivity of COMT is lost when the ethylamine or ethanolamine moiety is replaced by a neutral functionality (Creveling et al., 1970, 1972). Interestingly, the relative amount of the para-Omethylation of norepinephrine increases from < 10% at pH 7 to approximately

O-METHYLATION 199

25% at pH 9·5 (Thakker et al., 1982). This increase in para-O-methylation was attributed to ionization of the para-phenolic group with a pKa of approximately 8·6. This was firmly established when it was demonstrated that in 5fluoronorepinephrine, the increase in para-O-methylation with pH accompanied the ionization of a group with pKa of approximately 7·9 which is also the pKa of the para-phenolic group of this compound. Furthermore, the O-methylation occured predominantly at the para-phenolic group in 5-fluoronorepinephrine (Thakker et al., 1986). Since the presence of a fluoro substituent in the aromatic ring is not expected to affect the steric requirements of the substrate, the binding mode of norepinephrine and its 5-fluoro derivative are expected to be very similar. Hence, a change in the site of predominant O-methylation from the meta-phenolic group in norepinephrine to the para-phenolic group in 5fluoronorepinephrine strongly suggested that the methyl group of SAM is equally accessible to both the phenolic groups of the catechol substrates. Interestingly, the equal distribution of the meta- and para-O-methylated products from 3, 4-dihydroxybenzyl alcohol (an uncharged side chain), remained unchanged over the pH range of 7–9. Active-site model The high selectivity of COMT for meta-O-methylation of catecholamines prompted a proposal that the catecholamine substrates can bind at the active site in two different modes and that one of the two modes is unfavourable because of the interactions of a charged side chain with a hydrophobic pocket in or near the active site (Creveling et al., 1970). According to this model, the methyl group of SAM is accessible to only the meta-phenolic group in the favourable binding mode and to the para-phenolic group in the unfavourable binding mode. Coward and Slisz (1973) also proposed two binding modes with the methyl group accessible to different phenolic groups in the two binding modes. Both these models fail to explain the recent results of the effect of pH on the site of O-methylation (Thakker et al., 1982) as well as the reversal of the regio-selectivity of COMT upon adding a fluoro substituent at the 5-position of norepinephrine (Thakker et al., 1986). Hence, we have proposed a model for the active site of COMT (i) that is consistent with the high selectivity of the enzyme for meta-O-methylation of catechol substrates with a charged functionality in the side chain and the lack of selectivity toward catechols with uncharged side chains like 3, 4-dihydroxybenzyl alcohol; (ii) that takes into account the proposed SN2 mechanism of methyl transfer; (iii) that allows equal access of the methyl group to both the phenolic groups from the same binding mode; and (iv) that provides for a role of a nucleophile in the catalytic process (Thakker et al., 1986, 1988). As in the previous model (Creveling et al. 1970), we also propose the presence of a hydrophobic pocket in or near the active site

200 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 7.2. A model for the catalytic site of COMT. The hydrophobic pocket repels the charged functionality of substrates, such as norepinephrine (A, R=H), and forces them to bind as shown in A. In this binding mode, the meta-hydroxyl group is proposed to be activated by hydrogen bonding with a nucleophile in the vicinity. This explains the predominant meta-O-methylation of substrates such as norepinephrine. According to this model, the 5-fluoronorepinephrine also binds in the same way as norepinephrine (B), but increased acidity of the para-hydroxyl group due to the 5-fluoro substituent results in the predominant para-O-methylation. The 2, 5-difluoronorepinephrine (A, R=F) is predominantly meta-O-methylated because the inductive effect of the 5-fluoro substituent on the para-hydroxyl group is counterbalanced by a similar inductive effect on the meta-hydroxyl group. The catechol substrates with uncharged substituents can bind in two different modes (180° rotation around the axis which bisects the bond between the two oxygen bearing carbons) such that the uncharged substituent is projecting away from or into the hydrophobic pocket (binding mode I and II in C, respectively). Because the meta-and the para-hydroxyl groups are activated in the binding modes I and II, respectively, equal distribution of the meta- and para-O-methylated products are predicted and obtained for substrates like 3, 4-dihydroxybenzyl alcohol.

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of COMT (Figure 7.2). This hydrophobic pocket forces the catechol substrates with charged functionalities (e.g. norepinephrine, R=H in Figure 7.2A) to bind in only one binding mode. In this binding mode, the meta-phenolic group is situated near a nucleophile so that its phenolic proton is hydrogen bonded to the nucleophile (Figure 7.2A and 7.2B). It is proposed that the methyl group of SAM is approximately equidistant from both the phenolic groups, but the metaphenolic group of norepinephrine preferentially mounts a nucleophilic attack on the methyl group because of the δ-charge developed by its interaction with the nucleophile (Figure 7.2A, R=H). The para-phenolic group can compete with the meta-phenolic group only when it is ionized. Hence, as the pH goes up, an increased ionization of this group results in its increased methylation. In 5fluoronorepinephrine (Figure 7.2B) the para-phenolic group is much more acidic because of the ortho-fluoro substituent. Hence, it competes successfully for the methyl group at even lower pH values and is the predominant site of Omethylation. Interestingly, by introducing a second fluoro substituent at the 2position (Figure 7.2A, R=F), the acidities of the two phenolic groups become nearly equivalent, as in the case of norepinephrine, and as expected, the predominant product is the meta-O-methylated product. The catechol substrates with neutral moieties in the side chain bind equally well in both binding modes I and II (Figure 7.2C) in which the meta- and the para-phenolic groups are activated, respectively. Hence, a substrate like 3, 4-dihydroxybenzyl alcohol is converted to a 1:1 mixture of meta- and para-O-methylated products at all pH values (Thakker et al., 1986, 1988). 7.5. In vitro inhibitors of COMT There are three general classes of compounds that inhibit the catalytic activity of COMT. These include (I) analogues of the demethylated product SAH, (II) isosteres of catechol substrates, and (III) divalent and trivalent metal ions. A brief discussion of the structure-activity relationships for each of the classes is given below. Analogues of S-adenosylhomocysteine SAH is a potent inhibitor of COMT and other SAM-dependent methyltransferases with Ki values <50 µM at subsaturating concentrations of SAM (Borchardt et al., 1974b). The inhibition of the methyltransferases by SAH appears to be a feedback control mechanism to regulate the activities of these enzymes. In extensive studies (Coward et al., 1972, 1974; Coward and Sweet, 1972; Coward and Slisz, 1973; Borchardt and Wu, 1974, 1975; Borchardt et al., 1974b, 1976a) in which systematic changes were made in the base, the

202 CONJUGATION REACTIONS IN DRUG METABOLISM

sugar and the homocysteine moieties, the structural features required for effective inhibition of several SAM-dependent methyltransferases were evaluated. These studies revealed that COMT and other methyltransferases show strict specificity for the structural features of SAH. In particular, for effective binding of SAH to COMT, the amino and the carboxy group of Lhomocysteine, the 6-amino group of the adenine moiety, and the 2’hydroxyl group of the ribose moiety are essential. Attempts to design transition state analogues containing both the catechol and the SAH (or SAM) moiety did not result in much success (Anderson et al., 1981; Lever et al., 1984). Isosteres of catechol substrates With one or two exceptions noted earlier, all the methyl acceptor substrates of COMT have a catechol moiety. Hence, several inhibitors have been developed for COMT that contain either a catechol group or a pair of ortho-substituents which are isoelectronic and isosteric with the catechol group. While the former compounds act as alternate substrates, the latter ones are usually dead-end inhibitors. Many inhibitors in this class are O-methylated derivatives of pyrogallol or catechols, and thus resemble products. A few examples of inhibitors from this class are pyrogallol, flavonoids, 8-hydroxyquinolines, pyrones and pyridones, 3-mercaptotyramine, tropolones, etc. These inhibitors and others have been discussed in considerable detail by Guldberg and Marsden (1975) in their excellent review, and hence will not be discussed here. An interesting series of compounds not covered in the above review consists of 3-hydroxy-4methoxy-5-substituted benzaldehydes (5-substituted isovanillins) (Borchardt et al., 1982). Electron withdrawing substituents at the 5-position enhanced the inhibitory activity of these compounds. The 5-nitroisovanillin was particularly potent with Ki=0·7 µM. Recently, 6, 7-dihydroxy-3, 4-dihydroisoquinolines were reported as inhibitors of COMT that could not be methylated by the enzyme despite the presence of catechol functionality (Cheng et al., 1987). The authors argued that the compounds existed as a quinoid tautomer and hence could not be O-methylated, but sufficient evidence was not presented to prove that the catechol tautomer did not exist at all. In contrast to the results by Cheng et al. (1987), Roswadowska et al. (1988) reported that the related (S)- and (R)dideoxynorlaudanosoline-1-carboxylic acids proved to be substrates rather than inhibitors of COMT. 2, 3-Dihydroxypyridine is another catechol inhibitor of COMT which cannot be O-methylated by the enzyme (Raxworthy et al., 1983). In this case, the authors argue that the ring nitrogen alters the susceptibility to O-methylation. In contrast to the compounds described above, the tumour inhibitor, vernolepin, which contains two α-methylene lactone groups, enhances the activity of COMT (from cat erythrocytes) after small molecular weight thiols have been added across the methylene groups (Lantz et al., 1975).

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Whether the carbonyl groups of the lactone moiety and the thioether moieties are involved in the binding of this compound to COMT is uncertain. This is the only example of a compound that activates COMT by direct interaction with the enzyme. Metal ions Because of the requirement by COMT for Mg+2 ions, several metal ions have been tried as inhibitors of this enzyme. Among the alkaline earth metals tried, Ca +2 ions inhibited 60% of the enzymic activity of rat liver COMT at 1 mM concentration in the presence of 1 mM Mg+2 ions (Weinshilboum and Raymond, 1976). At 2 mM concentration, Mg+2 ions did not inhibit COMT. Lanthanum and lanthanides can mimic the effects of calcium and other alkaline earth metals in many biochemical systems. Hence, salts of lanthanum and the lanthanides neodymium and europium were examined as inhibitors of COMT. All three ions, i.e. La+3, Nd+3 and Eu+3, inhibited 50% of rat liver COMT activity at 1–3 µM concentrations (Quiram and Weinshilboum, 1976a). 7.6. The multiple roles of COMT in vivo The functions of COMT in vivo have become more apparent with (i) the expanding knowledge of the specific cellular localization of the soluble form of the enzyme (S-COMT) (for reviews see Inoue et al., 1977; Creveling and Hartman, 1982; Creveling, 1984, 1988); (ii) the distribution and activity of the membrane-bound form of the enzyme (MB-COMT) (Rivett and Roth, 1982; Rivett et al., 1982; Rivett and Roth 1983a, 1983b; Jeffrey and Roth, 1984); and (iii) the remarkable increases in the specificity and sensitivity afforded by high performance liquid chromatography and electrometric detection (Adams, 1979; Adams and Marsden, 1982; Marsden, 1983) for the measurement of COMT activity (Koh et al., 1981; Nissinen and Mannisto, 1984; Schultz et al., 1988), substrate levels and O-methylated and other metabolic products (Wagner et al. 1982; Kaneda et al., 1986; for general reviews of catecholamine metabolism in vivo see Guldberg and Marsden 1975; Saavedra, 1977; Kopin, 1985). Several general aspects about COMT can now be appreciated. First, while there is a wide variation in the level of COMT activity in various tissues, among species and strains, the individual level of activity in most tissues and strains show great similarities. In most cases COMT activity increases rapidly from relatively low levels at birth to a characteristic level early in life and remains essentially constant throughout the adult life (Goldstein et al., 1980; Ladosky et al., 1984, Commissiong, 1983; Rodriguez et al., 1985). Second, in certain tissue sites, such as the epithelial lining of the uterus and in the ductal epithelium of

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breast, the level of S-COMT activity undergoes marked increases in response to pregnancy, lactation, and oestrus (Inoue et al., 1980; Amin et al., 1983; Creveling, 1984). It should be noted that COMT is sexually dimorphic with clear differences in the hypothalamic nuclei related to reproduction, and in the hippocampus in male and female rats. The levels of COMT, characteristic of the male or female, can be altered by exposure to sex hormones at birth; however, once this change is established, the COMT level remains constant throughout life (Ladosky et al., 1984). In peripheral organs of rat, such as the liver, kidney, and heart, S-COMT activity and immunoreactive S-COMT protein increase rapidly at birth, plateau in the young adult and then remain essentially constant throughout life (Goldstein et al., 1980). Finally, COMT is present, and in some cases at remarkably high levels, in certain neoplasms. Several studies have demonstrated the presence of relatively high levels of COMT activity in adenocarcinomas of breast tissue in women (Assicot et al., 1977; Hoffman et al., 1979a, 1979b); in mouse and rat (Amin et al., 1983); and in beta-islet insulinomas of rat and hamster (Feldman et al., 1979). S-COMT activity primarily appears constant and characteristic of the individual at most sites in the adult animal, while in certain cell types COMT activity changes in response to hormonal or physiological cues. These observations strongly suggest that in addition to the now classical function of COMT in the inactivation of circulating catecholamines in the liver, originally described by Axelrod (1966), COMT appears to have a much wider role in the control of the level and distribution of substances bearing the reactive catechol moiety. The central nervous system Developmental aspects COMT activity appears early in development of the brain (for review see: Parvez and Parvez, 1980), reaching a maximum in rat and rabbit as early as foetalday 15 (Parvez et al., 1979), and in the developing spinal cord of the rat (Commissiong, 1983). Enzyme activity declines rapidly in late foetal life and then begins to rise again after birth. In the hypothalamus and spinal cord, the capacity for O-methylation is well developed before the catecholamine levels reach a maximum (Commissiong, 1983). COMT is clearly sexually dimorphic by day 12 after birth in the rat hypothalamus with enzyme activity lower in males than in females, while in the hippocampus the reverse is true (Ladosky et al., 1984). Exposure of the newborn females to androgens or estradiol results, by day 12, in a dose-dependent decrease of COMT in the hypothalamus and an increase in the hippocampus, whereas castration of males at birth results in

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levels of COMT similar to those found in females. It should be noted, however, that alterations of COMT levels, either by surgical or hormonal intervention during the critical period for sexual differentiation, once established, are permanent and are maintained in adulthood (Breuer et al., 1981). Sexual dimorphism in COMT levels appears to be related to gonadotrophin release and is present only in those hypothalamic nuclei involved in gonadotrophin release and not in nuclei unrelated to gonadotrophin control (Ladosky et al., 1984). The distribution of COMT in brain The distribution of COMT in brain is divided between the high-affinity, MBCOMT and the cytosolic S-COMT. The MB-COMT activity varies from 1 to 30% of the total COMT activity in rat and human brain (White and Wu, 1972; Borchardt and Cheng, 1978; Rivett et al., 1983a, 1983b; Jeffrey and Roth, 1984; Kaakkola et al., 1987). The highest levels of MB-COMT are present in the hippocampus, cerebellum and cerebral cortex (Rivett et al., 1983a, 1983b) and striatum (Kaakkola et al., 1987). Studies in striatum following denervation with 6-hydroxydopamine or treatment with kainic acid have clearly indicated that SCOMT is located post-synaptically, probably associated with striatal glial cells, whereas MB-COMT may be present in both neuronal and extraneuronal cells (Rivett et al., 1983a, 1983b; Kaakkola et al., 1987). The availability of a specific polyvalent antiserum to S-COMT (Inoue et al., 1977; Grossman et al., 1985) has permitted immunochemical localization of SCOMT to specific cell types. One of the characteristic sites of S-COMT in brain is its uniform presence in the cytoplasm of the ciliated, cuboidal cells of the ventricular ependyma along the borders of the lateral, third and fourth ventricles of the rat brain (Kaplan et al., 1981a). This apparent barrier to catechols between the cerebrospinal fluid (CSF) and the brain parenchyma is not complete, for at the border of the 3rd ventricle the cells bordering the arcuate nucleus do not contain S-COMT. A similar configuration is present at the other circumventricular organs, and a similar distribution of monoamine oxidase (MAO-B) immunoreactivity was observed in ependymal cells and cells in the matrix of the circumventricular organs (Levitt et al., 1982). The presence of both S-COMT and MAO-B in astrocytes and tanycytes in these areas which lack a blood-brain barrier, can be considered to be in direct contact with cerebrospinal fluid. This morphological configuration may result in differing functional consequences compared to brain areas shielded by the ependymal barrier. In the brain parenchyma proper, S-COMT is found primarily in glial elements (Kaplan et al., 1979). The cell bodies and proximal portions of oligodendrocytes in large fibre tracts exhibited bright S-COMT-specific immunofluorescence, as did the perineuronal satellite oligodendrocytes in

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cortical areas. The cell bodies and extended processes of many fibrous astrocytes were especially immunoreactive (Kaplan et al., 1979). These immunocytochemical results are in agreement with measurements of COMT activity in primary cultures of astrocytes derived from the cerebral cortex of newborn rat (Pelton et al., 1981; Hansson and Sellstrom, 1983). According to Pelton et al. (1981) the cultured astrocytes maintain a K+ ion-dependant membrane potential of −70 mV and exhibit a high-affinity, Na+ ion-dependent uptake system for dopamine with an associated O-methylating system. Other studies (Hansson and Sellstrom, 1983; Hansson, 1984) report only low-affinity, Na+-independent uptakeofcatecholamines. The distribution of COMT activity in primary astroglial cultures from various areas of brain was similar to that found in adult animals with highest activity in hippocampus and lowest in brain stem (Hansson, 1984). The astrocytic endfeet bordering arterioles and larger vessels in the brain contain S-COMT (Kaplan, 1980). In this regard, Spatz et al. (1986) showed that the endothelium of cerebral capillaries, arterioles, and larger vessels contained S-COMT. Thus COMT in conjunction with MAO may provide cerebral capillaries with an enzymic barrier for the passage of catechols. Kaplan et al. (1980, 1981b) reported the immunolocalization of S-COMT in cells of the pia mater and arachnoid. This localization of COMT may provide an ‘arachnoid barrier’ to prevent peripherally derived catecholamines in the dural circulation from entering the subarachnoid CSF and the brain parenchyma (Nielsen and Owman, 1967; Nabeshima and Reese, 1972; Nabeshima et al., 1975). In the cerebellum, in addition to glial elements, the cell bodies of Bergmann cells, adjacent to S-COMT-negative Purkinje cells, contained S-COMT as did the Bergmann fibres ascending through the molecular layer to the pial surface (Kaplan et al., 1979). No evidence of the presence of S-COMT in neurons has been observed with this antisera; on the contrary, attempts to find S-COMT immunoreactivity in neurons have been negative (Kaplan et al., 1979, 1980). It should be emphasized that the possibility of neuronal S-COMT cannot be excluded by immunological methods. Median eminence and pituitary S-COMT has been localized by immunofluorescence techniques in the tanycytes in matrix of the median eminence and infundibular stalk, as well as in the glial-like pituicytes of the neural lobe of the rat pituitary (Kaplan, 1980). The cytoplasm and processes of the tanycytes contain high concentrations of SCOMT as do the pituicytes of the neural lobe. The COMT-containing processes extend to the perivascular space surrounding the portal vessels in the median eminence. No immunoreactive S-COMT could be detected in the intermediate lobe. The epithelial cells (Kagayama et al., 1969) at the border of the

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intermediate and anterior lobes contained high concentrations of S-COMT. In the anterior lobe, S-COMT was found in the network of follicular cells but was absent in the granular secretory cells (Kaplan, 1980). It is clearly established that neural and intermediate lobes contain catecholamines and catecholaminergic fibres primarily of the arcuatehypophyseal system which descend though the median eminence from cell bodies in the hypothalamus (see Dahlstrom and Fuxe, 1966; Bjorklund et al., 1970). A dense dopaminergic innervation is present in the intermediate lobe (Bjorklund et al., 1973). Electrical stimulation of the pituitary stalk in isolated preparations of the rat pituitary results in dopamine release from the neural lobe (Holzbauer et al., 1983, 1984). Further studies have demonstrated a pronounced rate of dopamine synthesis and an even greater rate of MAOmediated intraneuronal conversion of dopamine to dihydroxyphenylacetic acid (DOPAC). More than half of the released DOPAC is then O-methylated to homovanillic acid (HVA), and lesser amounts to 3-methyoxy-4hydroxyphenylethanol (MOPET). Surprizingly, nearly 80% of this dopamine synthesis and metabolism occurs in the intermediate lobe (Racke and Muscholl, 1986, 1989) suggesting that a significant portion of dopamine catabolism must be catalyzed by COMT at the intermediate lobe. The apparent absence of immunologically detectable S-COMT in the intermediate lobe (see above) raises the intriguing possibility that MB-COMT may be unique in being the only form of COMT at this site. Earlier studies by Saavedra et al. (1975) reported low COMT activity in the anterior pituitary but were unable to detect activity in either the intermediate or posterior lobes. An additional aspect of the localization of COMT in the pituitary is the immunological localization of catecholestrogens in the cytoplasm of pituicytes of the posterior lobe and the co-existence of S-COMT and catecholestrogen in tanycytes of the median eminence (Inoue and Yoshizawa, 1986). This distribution suggests that catecholestrogens may provide connecting links between primary oestrogens, catecholamines and the regulation of gonadotropin and prolactin secretion. The co-existence of both S-COMT and catecholestrogens in the ductal cells of the rat parotid gland and in the liver have been demonstrated (Inoue and Yoshizawa, 1985; Inoue et al., 1987). Thus, the immediate presence of COMT may control the local accumulation of catecholestrogen. While the presence of 2- or 4-hydroxyestradiol has been established, their physiological roles are still controversial (Davies et al., 1975; Martucci and Fishman, 1976; Merriam and Lipsett, 1983). The reproductive system Immunological localization of S-COMT in the uterus, placenta, oviduct, mammary gland, and vas deferens and seminal vesicle has led to a greater

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appreciation of the role of COMT in the reproductive process (Inoue et al., 1980; Amin et al., 1983; Inoue and Creveling, 1986). Earlier studies had indicated that COMT activity in rat uterus is higher during oestrus than in diestrus (Giles and Miller, 1967). It increases during pregnancy and persists for a short time after parturition (Wurtman et al., 1964). Others have indicated the presence of COMT activity in both human and rat placenta (Chen et al., 1974; Burba, 1979; Hobel et al, 1981) and in rabbit placenta (Kennedy et al., 1984). COMT activity has been demonstrated in human placenta vera in early and late pregnancy (Casey and MacDonald, 1983). Placental COMT activity was fourto seven-fold higher than that of the myometrium. COMT activity is greater in erythrocytes of pregnant women than in those of men or nonpregnant women (Bates et al., 1978). The appearance and localization of S-COMT was examined immunologically in rat uterus from the initiation of pregnancy to 12 h after delivery of the last foetus (Inoue et al., 1980). On day 5 of pregnancy, S-COMT appeared in endometrial and glandular epithelium. By day 7, S-COMT was present in differentiating decidual cells near the site of the implanting blastocyst. By day 8, the S-COMT-containing cells formed a crescent around the antimesometrial border of the lumen where the blastocyst was in contact with the endometrium. On the mesometrial side, S-COMT-positive polygonal cells were observed near the developing sinusoids. By day 11, S-COMT-positive polygonal cells were present in the walls of the vascular spaces and in stellate cells in the decidua basalis. By day 14 a network of numerous S-COMT-positive stellate cells had formed in the myometrium and remained until term. Following delivery of the last foetus, the S-COMT-containing cells in the myometrium resembled fibroblasts and were associated with the numerous blood vessels. In the rat oviduct, S-COMT was present in the cytoplasm of the epithelial cells lining the lumen. In the infundibulum and ampulla, nonciliated epithelial cells were strongly COMT-positive with lesser amounts in the ciliated cells (Inoue and Creveling, 1986). Epithelial cells in the oviduct are at some distance from the adrenergic innervation of the smooth muscle layer (Paton et al., 1978) suggesting that epithelial S-COMT may function in catechol steroid inactivation, possibly related to fluid production rather than catecholamine inactivation. It is important that S-COMT in the epithelial cells of the oviduct is present in the nonpregnant rat and does not appear to change with the onset of pregnancy as does S-COMT in the uterine epithelium. It has been suggested that one function of epithelial COMT which extends from the fimbria to the junction with the uterus, might be related to the inhibition of implantation of fertilized blastocysts. The porcine blastocyst cannot only synthesize estradiol de novo, but 2-hydroxyestradiol as well (Mondshein et al., 1987). It has recently been shown that 2-hydroxyestradiol is a potent, competitive inhibitor of the Omethylation of 4-hydroxyestradiol (Roy et al., 1988). The 4-hydroxy oestrogens,

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unlike the 2-hydroxy-oestrogens, have potent, long-acting oestrogenic effects (MacLusky et al., 1981). 4-Hydroxyestradiol is capable of inducing implantation in the ovariectomized mouse primed with progesterone (Hoverland et al., 1982). These factors imply a relationship between COMT activity, a 2-hydroxyestradiol-mediated increase in the local concentration and half-life of 4hydroxyestradiol, and the possible interaction of the 4-hydroxy species with the estradiol receptor. An understanding of the role of COMT in events surrounding implantation in uterus as well as of its influence upon factors preventing implantation in the oviduct must await further research. Several factors implicate an important role for COMT in the ovary: adrenergic nerve fibres rapidly infiltrate the theca externa of ovarian follicles (Sporrong et al., 1985; Spicer, 1986); the catecholamines, norepinephrine and epinephrine, are present in follicular fluid, and their concentration increases as ovulation approaches (Bahr and Ben-Jonathan, 1985); the catechol oestrogen, 2hydroxyestradiol, synthesized de novo is present in granulosal cells (Hammond et al., 1986); and epinephrine in combination with catechol steroid acts synergistically to stimulate progesterone production (Spicer and Hammond, 1987). The potency of 2-hydroxyestradiol is similar to that of oestrogen in the simulation of progesterone production in porcine granulosa cells (Spicer and Hammond, 1987). Both catecholamines and catechol steroids are rapidly Omethylated in the ovary and in isolated granulosa cells. Ball et al. (1983) suggested that catecholamine levels may be locally regulated by competition with steroids for COMT in both the ovary and pituitary gland. S-COMT was demonstrated immunologically in macrophages in the corpus luteum of the rat ovary (Inoue and Creveling, 1986) and in macrophages of lymphoid tissue (Inoue and Creveling, 1980). Previously Kirsch et al. (1981) reported that macrophages in the corpora lutea of mice affected luteal production of progesterone in culture. The increase in progesterone production was directly proportional to the number of macrophages present in the luteal cell culture. Specialized contacts between macrophage microvilli and secreting luteal cells were described, suggesting additional functions of macrophages in corpus lutea than heterophagy. Interest in the function of COMT in mammary glands stems from the discovery of elevated levels of COMT activity in human breast tumours, the apparent positive relationship between the COMT activity and the grade of malignancy in primary carcinomas (Assicot et al., 1977) and the demonstration of de novo synthesis of catecholsteroids in breast tumours (Hoffman et al., 1979a, 1979b). Elevated levels of COMT are present in human and rodent breast tumours and cell lines derived from them (Amin et al., 1983; Schneider et al., 1984; Levin et al., 1987). Since more than one third of human breast cancers are oestrogen dependent (Manni and Pearson, 1982) and the major route of oestrogen metabolism appears to proceed through catecholsteroid formation and

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O-methylation (MacLusky et al., 1981) the formation, properties, and enzymatic inactivation of the 2- and 4-hydroxy steroids derived from oestrogen have been the focus of extensive research (Merriam and Lipsett, 1983). While the affinity of 4-hydroxyestradiol is the highest of the catecholsteroids (approximately 1/ 20th that of estradiol), there is evidence that the dissociation rate of 4-hydroxyestradiol from the oestrogen receptor complex is equal to or longer than that of estradiol itself (Merriam et al., 1981). Studies in MCF-7 human breast cancer line suggest that 2hydroxyestrogen acts as an anti-oestrogen, inhibiting oestrogen-stimulated cell growth when protected from rapid metabolism by the presence of an inhibitor of COMT (Schneider et al., 1984). Alternatively, catecholestrogens may possess independent receptor-mediated functions. In this regard, a specific membrane binding site for 2-hydroxyestrone has been characterized in two oestrogen-receptor-positive human mammary carcinoma cell lines which differs in specificity from the classical oestrogen receptor (Vandewalle et al., 1988). However, the question of the function of catecholsteroids remains controversial (Merriam et al., 1981). The site of the elevated levels of COMT was localized immunocytochemically in the cytoplasm of a spontaneous mouse breast adenocarcinoma and in a spontaneous rat mammary ductal hyperplastic tumour (Amin et al., 1983). In breast tissue from normal and lactating rat, S-COMT was present in the cytoplasm of the ductal epithelial cells, with lesser amounts in fibroblasts in connective tissue and endothelial cells lining blood vessels. The level of SCOMT in the lactating rat was significantly greater than in the non-lactating rodent. Immunological examination of several infiltrating ductal human adenocarcinomas of the breast indicated that S-COMT was present throughout the cytoplasm of the malignant cells (Lowe and Creveling, unpublished data). The increased vascularity associated with tumour growth could account for a portion of the increased level of COMT activity in breast tumours. In blood vessels adjacent to the tumours S-COMT was present in the endothelium, in agreement with the vascular localization of S-COMT reported earlier (Lowe and Creveling, 1979). In sections of the rat vas deferens, S-COMT was localized in the cytoplasm of the epithelial cells in the mucosa. No immunoreactive S-COMT was apparent in the lamina propria, muscularis or adventitia (Inoue et al., 1977). In the seminal vesicles S-COMT was present in the cytoplasm of the small cells corresponding to the round basal cell of the epithelium. The cardiovascular system The interrelationship of COMT to the uptake and metabolism of catecholamines in the cardiovascular system has been the subject of intense investigation (Trendelenberg, 1963, 1971). The kinetic analysis of the

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extraneuronal uptake and metabolism of norepinephrine and other catecholamines in the isolated, perfused heart and many other tissues has resulted in the characterization of sites of extraneuronal uptake and Omethylation (Iversen, 1967; Trendelenberg, 1978, 1980). For a long time a satisfactory explanation of the nature of the extraneuronal O-methylation system was frustrated by the apparent requirement of relatively high concentrations (30 to 100 µM) of catecholamines to saturate COMT. The reported measurements of the Km of COMT for catecholamines were in the 100 to 400 µM range. Kaumann (1970) had suggested that the degree of supersensitivity would be explained if the extraneuronal mechanisms of inactivation were saturable and had a high affinity for catecholamines. Many careful studies were performed that clearly indicated that the dissociation constants for the extraneuronal uptake and metabolism via O-methylation required both a low-affinity and a high-affinity form of COMT (Bonisch and Trendelenberg, 1974; Bonisch, 1978; Fiebig and Trendelenberg, 1978; Bryan et al., 1983; Trendelenberg, 1984; Cassis et al., 1986; Magaribuchi et al., 1987). However present evidence points to the participation of a high-affinity MB-COMT in the extraneuronal metabolism of catecholamines (Rivett and Roth, 1982; Reid et al., 1986). In an earlier report, Wrenn et al. (1979) in a study of the beta-adrenergic receptor regulation of a membrane-bound form of COMT activity in myocardium, reported the Km value for norepinephrine to be 3·5 µM. In a recent study in rat heart, Grohmann (1987) measured the steady-state formation of the O-methylated metabolites of dopamine, norepinephrine, isoproterenol and epinephrine and from their apparent rate constants estimated the ‘pseudo-Km’ values for COMT to be 3·5, 3·1, 1·6, and 1·5 µM, respectively. These values suggest that the low-Km, membrane-bound form of COMT is responsible for theO-methylation of catecholamines in extraneuronal tissues exposed to low outside concentrations of catecholamines (Grohmann and Trendelenberg, 1985; Grohmann, 1987). While very little is known about the cellular localization of MB-COMT, studies on the immunocytochemical localization of S-COMT in cardiovascular tissues of rat have been reported (Lowe and Creveling, 1979). S-COMT is present in the cytoplasm of aortic endothelial cells and in myocardial cells. Curiously the smooth muscle cells of the aorta and the coronary vasculature appeared to be nearly devoid of S-COMT. Ultrastructural localization of SCOMT included the cytoplasm, plasma membranes, and basal laminae of the endothelial cells. In myocardial cells the S-COMT positive sites included plasma membranes, external laminae, and the sarcoplasmic cytoplasm (Lowe and Creveling, 1979). The significance of the virtual absence of S-COMT in the adrenergically innervated smooth muscle in arteries and the aorta is not clear, in view of the considerable pharmacological evidence for the presence of an extraneuronal O-methylation system in vascular smooth muscle

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(Trendelenberg, 1980) and the localization of S-COMT in cultured smooth muscle cells from microvessels of the brain (Spatz et al., 1986). However a comparison of the uptake and metabolism of [3H]norepinephrine in dog pulmonary artery clearly indicated that the endothelium was an important component of extraneuronal O-methylation system accounting for as much as 50% of the O-methylation (Rorie and Tyce, 1985). The importance of Omethylation of isoproterenol in the endothelium of the rabbit thoracic aorta has been reported by Head et al. (1986). These workers noted that the large contribution of the endothelial COMT may be related to the O-methylation of blood-borne catecholamines in contrast to other sites of O-methylation which may have a more prominent role in the inactivation of neuronally released catecholamines. A comparison of the relative role of O-methylation for norepinephrine and epinephrine in saphenous vein, mesenteric and renal arteries indicated that the importance of O-methylation varied with the morphology of the vessel and the amine. O-methylation is more important for the inactivation of epinephrine than norepinephrine and is the predominant pathway for both amines in the renal artery (Nunes et al., 1987). Erythrocytes The presence of COMT in erythrocytes (RBC-COMT) and other blood-borne cells has led to the considerations of this COMT activity as a diagnostic device in human disease, genetics, and as a possible reflection of tissue levels of COMT. RBC-COMT is both biochemically and immunologically similar to COMT in other tissues (Axelrod and Cohn, 1971; Quiram and Weinshilboum, 1976a). Initial studies in man indicated a significant bimodal distribution of RBC-COMT activity in siblings (Weinshilboum et al., 1974). A similar bimodal distribution was found in a large (372 subjects), randomly-selected population. In the total population examined 77% were classified as having high RBC-COMT activity and 22% with low RBC-COMT activity. These groups were separated by a clear nadir between the high and low RBC-COMT. The familial factors with regard to the low RBC-COMT led these authors to suggest an autosomal inherited allele for low RBC-COMT. A thermolabile variant of COMT was associated with low RBC-COMT which may be related to inherited structural differences in the enzyme (Scanlon et al., 1979; Baron et al., 1982). In 1982, Floderus et al. reported evidence for a major locus as well as a multifactorial component in the regulation of RCB-COMT activity. Similarly the results of a major study of RBC-COMT activity conducted on five large families composed of 1189 individuals definitively confirmed the bimodal distribution of RBC-COMT activity and strongly suggested that the variation of COMT activity is in part due to the effects of a major gene (Siervogel et al., 1984). A segregation analysis of RBC-COMT activity by Goldin (1985) supported the

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presence of a dominant major locus and also the presence of an additional polygenic component. The location of a single COMT gene on chromosome 22 was obtained from the segregation analysis of COMT in hybrids and subclones of several mouse-human fibroblast cell lines (Brahe et al., 1986). A definitive resolution of the genetic basis for COMT activity(s) must await a determination of the DNA sequence(s) and the structure of the protein(s) (see Grossman et al., 1989). The accessibility of RBC-COMT has resulted in its use as a biological probe in clinical studies. The level of COMT activity in human lung and kidney showed a significant positive correlation to RBC-COMT activity in 29 patients suggesting that measurements of RBC-COMT activity might be of value in predicting tissue levels of COMT and perhaps the capacity to metabolize catechols (Weinshilboum, 1978). A bimodal distribution of RBCCOMT activity was found in normal caucasians and orientals; however, the frequency distribution of high RBC-COMT activity was significantly greater in orientals than in caucasians. This difference may be related to the clinical variation in the tolerance to L-DOPA seen in Parkinsonian patients in these two racial groups (Rivera-Calimlim and Reilly, 1984). Many studies have attempted to relate RBC-COMT activity as a biological marker for affective disorders in man (for a review see Fahndrich et al., 1980). A recent finding indicated that RBC-COMT activity was lower in patients with major depression and in recurrent bipolar depression (Karege et al., 1987). No differences between normal patient RBC-COMT activity were found in thyroid dysfunction (Coulombe et al., 1977), in subjects with alcoholism and under controlled abstinence (Agarwal et al., 1983) and in women taking oral contraceptive steroids (Bates et al., 1979). While many of the early clinical studies appeared to yield contradictory results, stemming primarily from technical deficiencies in the assay of COMT (see Dunner et al., 1971), the applicability of COMT measurements as an estimate of O-methylation capacity via erythrocytes, platelets, and lymphocytes (Bidart et al., 1983; Sladek-Chelgren and Weinshilboum, 1981; de Prada et al., 1984) appears to be a clinically useful device. The metabolism of catechols in brain The central metabolism of catechols in vivo with regard to the role of COMT has been examined by several techniques (for a recent and complete review of catecholamine metabolism in vivo, see Kopin, 1985). Koster et al. (1984a, 1984b) performed a kinetic analysis similar to the procedures applied to the heart by Trendelenburg (1978) following the intraventricular injection of [3H]NE; McCalden et al., (1979) examined the changes in cerebral blood flow during infusion of NE in baboon following osmotic disruption of the blood brain barrier in normal and 6-hydroxydopamine-treated animals; and Buu (1985) followed

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the metabolism of dopamine after the intraperitoneal administration of LDOPA. In these studies an important, extraneuronal role for the metabolism of catechols is ascribed to COMT. The global metabolism of i.c.v. administered [3H]norepinephrine in various regions of rat brain in vivo has been examined by Koster et al. (1984a, 1984b). The effect of inhibition of the COMT-catalyzed metabolism of [3H] norepinephrine was examined under conditions designed to examine extraneuronal O-methylation. Thus, [3H]norepinephrine was administered after (i) the specific intraneuronal vesicular uptake was blocked by reserpine; (ii) neuronal uptake1 was blocked by desmethylimipramine; and (iii) in the presence of carrier amount of norepinephrine. The time course and retention of norepinephrine and its metabolites showed that norepinephrine was rapidly and almost exclusively O-methylated outside of the neuronal compartment. Inhibition of COMT by tropolone primarily affected this initial rapid degradation of norepinephrine but not a slower MAO-dependent metabolism. Koster postulated that a ‘high-affinity methylating system’ may exist in brain. However, this site of extraneuronal uptake followed primarily by O-methylation has no capacity to retain or accumulate unchanged norepinephrine. The high affinity of this system for norepinephrine suggests the operation of MB-COMT rather than the low-affinity S-COMT. The regional distribution of the metabolites and the individual turnover rates suggested the presence of an additional extraneuronal compartment charact erized by a slow, MAOdependent metabolism of NE in contrast to rapid MAO-dependent intraneuronal metabolism. L-DOPA therapy of Parkinson’s disease COMT is an important determinant in the effective use of L-DOPA for the symptomatic therapy of Parkinson’s disease. Ever since the discovery of dopaminergic cell loss, the accompanying decrease in striatal dopamine (Ehringer and Hornykiewicz, 1960), and the introduction of L-DOPA as an effective means for the restoration of central dopamine stores (Cotzias et al., 1967), there has been increasing attention directed towards the COMTcatalyzed formation of 3-O-methyl-DOPA (OMD) which accompanies L-DOPA administration. This interest has been heightened by concern with the decreasing clinical efficacy of chronic L-DOPA therapy and the development of untoward effects (for reviews see Melamed and Hefti, 1984; Nutt and Fellman, 1984; Mouradian and Chase, 1988). The interest in COMT stems from the interference of the delivery of orally administered L-DOPA to the brain by the rapid formation of OMD. OMD is a major metabolite of L-DOPA (Sharpless et al., 1972) and is increased with the combined use of carbidopa, a peripheral decarboxylase inhibitor (Rivera-

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Calimlim et al., 1977) and is readily formed in most tissues. OMD formation after oral administration L-DOPA is of special significance due to the relatively high levels of both S-COMT and MB-COMT in the gut wall (Nissinen et al., 1988a). OMD competes with L-DOPA with greater affinity for the neutral amino acid transport system in both gut and at the blood brain barrier (Wade and Katzman, 1975; Gervas et al., 1983). This, along with the much greater biological half-life of OMD compared to that of L-DOPA, results in ratios of OMD/L-DOPA as high as 14 to 1 (Fahn, 1974; Reches and Fahn, 1984; Marion et al., 1986). A series of animal studies with short-acting inhibitors of COMT, mostly with the competitive substrate 3′, 4′-dihydroxy-2-methylpropiophenone (U-0521), have shown a dose-dependent inhibition of OMD formation and increased levels of both dopamine and DOPAC in striatum following the administration of L-DOPA. Verification of an increased central utilization of LDOPA in these same studies was shown by the potentiation of L-DOPAinduced circling in the rat-nigrostriatal lesion model upon inhibition of COMT (Ungerstedt and Arbuthnott, 1970) and the L-DOPA dependent reversal of reserpine-treated mouse hypoactivity (Reches et al., 1982; Reches and Fahn, 1984; Nuutila et al., 1987; Linden et al., 1988; Nissenin et al., 1988b). Cumming et al. (1987) reported that U-0521 administration decreased the formation of 6-18F-OMD in plasma and increased striatal 6-18F-dopamine and 6-18F-DOPAC in rat following administration of the PET scanning agent, 6-18FDOPA. It should also be noted that tropolone, a noncompetitive inhibitor of COMT, blocks the L-DOPA-induced decrease in SAM and increase in SAH in rat hypothalamus (Fuller et al., 1983). Though not reported, this observation suggests that a similar L-DOPA-induced change in the SAM/SAH ratio may also occur in striatum. Despite the obvious clinical advantage that the suppression of COMT activity might have in the therapy of Parkinson’s disease, none of the known COMT inhibitors, until very recently, have had sufficient specificity, potency, duration of action or lack of toxicity for clinical use. A new class of inhibitors have recently been developed which appear to be selective and potent inhibitors of COMT. One derivative, OR-462 (3-(3, 4-dihydroxy-5nitrobenzylidene)-2, 4-pentanedione) effectively inhibits COMT activity in the gut wall for up to 10 h when given orally to rats. COMT activity in liver and red blood corpuscles (RBC) was only transiently affected and striatal COMT was unaffected (Nissinen et al., 1988b). When given with L-DOPA it produces a long-lasting inhibition of OMD formation both peripherally and in the striatum. With this inhibitor, equivalent levels of striatal dopamine were achieved with one-fourth as much L-DOPA compared to controls given L-DOPA alone. The increased utilization of L-DOPA was verified in the rat nigrostriatal lesion and reserpinized mouse models (Linden et al., 1988). These initial studies and the reported low toxic potential of these compounds in mice and rats (Backstrom et al., 1989) suggest that OR-462 or other compounds of this class may have

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clinical potential in the treatment of Parkinson’s disease, especially in patients exhibiting the untoward effects of chronic L-DOPA therapy. The barrier role of COMT An important role for COMT is its participation as an enzymic barrier for the passage of catechols. The immunological localization of COMT in epithelial cells and similar cell types which separate adjacent fluid compartments had been demonstrated in many tissues (Creveling and Hartman, 1982). This characteristic localization is exemplified in the choroid plexus where the major concentration of COMT is found in the cytoplasm of the epithelial cells (Kaplan et al., 1980). These cells are characterized by continuous, apical tight junctions (Brightman, 1975). The choroid plexus is able to actively take up catechols against a concentration gradient and to rapidly degrade catechols to Omethylated metabolites (Lindvall et al., 1980). This enzymic barrier at the blood-CSF interphase appears to prevent the passage of catechols in both directions, thus preventing the passage of circulating catechols into the CSF and to provide for the excretion of catechols from the CSF via the choroid plexus into the circulation. The presence of relatively high concentrations of S-COMT in epithelial cells is not limited to the choroid plexus but is characteristic of many epithelial cells. The distribution of S-COMT in the ciliary body of the eye is almost identical in appearance to that of the choroid plexus and very probably subserves the same function in these morphologically similar organs (Kaplan et al., 1981a). This proposed barrier analogy is not as precise with regard to the localization of S-COMT in epithelial cells in the vas deferens, oviduct, the ductal cells of breast and salivary gland, the proximal tubules and collecting ducts of the kidney, and the epithelial lining of the uterus. In these sites mucous producing, COMT-negative cells are often interspersed between the S-COMT-containing epithelial cells. However, the activity of S-COMT in isolated epithelial cells of the pregnant rat uterus was equivalent to the level of S-COMT activity in the liver and correlated well with the observed immunocytochemical density of the COMT-specific reaction product (Creveling, 1988). The biochemical measurements of the relatively high SCOMT activity in the chorid plexus also agreed closely with the observed intense S-COMT-specific immunofluorescence in the chorid plexus (Brightman, 1975; Kaplan et al., 1981a). Since the measurement of COMT activity in tissue homogenates necessarily underestimates the local cellular activity, it is safe to assume that the dense immunological reaction seen in epithelial cells is of the same order of activity as that of liver where such limitations are minimal. Another important consideration is cellular localization of MB-COMT. Extensive kinetic studies in isolated perfused organ systems on the relationship between COMT inhibition and the extraneuronal uptake and metabolism of

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catecholamines clearly point to the determinative role of a saturatable, highaffinity form of COMT (Trendelenberg, 1980, 1984; Grohmann, 1987; Nunes et al., 1987). While the relationship between function and morphology is less definitive than in epithelial cells, the localization of a second category of S-COMTcontaining cells suggests that they may provide regional barriers to the diffusion of catechols within the matrix of a tissue. The wide-spread localization of SCOMT-containing glial elements throughout the brain may provide a regional limitation for the diffusion of catechols escaping from the terminal fields of catecholaminergic neurons. A similar function may be ascribed to the presence of S-COMT in Schwann cells and in the satellite cells in sympathetic ganglia (Kaplan, 1980). Perhaps a more definitive example is the dense layer of SCOMT-containing cells that form a continuous ring around the margins of the white and red pulp in the rat spleen (Inoue and Creveling, 1980). 7.7. O-Methylation of drugs in vivo A wide variety of compounds which have the essential catechol configuration are substrates for COMT. This includes catechols ranging in complexity from 1, 2-dihydroxybenzene to large polycyclic catechols of plant origin. COMT plays an important role in the inactivation of drugs which are catechols like levophed, epinephrine, isoprenaline, DOPA, and α-methylDOPA (Guldberg and Marsden, 1975; Kopin, 1985). The decarboxylase inhibitors, carbidopa and benserazide, have both been shown to be substrates for COMT. The metabolism of isoprenaline is almost exclusively through the formation of 3-Omethylisoprenaline (Herting, 1964; Kadar et al., 1978). Other β-agonists, employed as bronchodilators or cardiotonic agents, like isoetharine and rimiterol or dobutamine and butanephrine, are substrates for COMT and have been shown to be metabolized in vivo by O-methylation (Gordonsmith et al., 1982; Raxworthy et al., 1986). An interesting example of the role of COMT in drug inactivation is the comparison of the behavioural effects of the 5, 6- and 6, 7-dihydroxy derivatives of 2-amino-1, 2, 3, 4-tetrahydronaphthalene, two dopamine agonists. In mice, after equal doses, the 5, 6-isomer yields brain concentrations several times those of the 6, 7-isomer, which is reflected in a greater behavioural potency. Inhibition of COMT abolishes this apparent difference in potency (Horn et al., 1981). Later studies demonstrated that the 6, 7isomer is a markedly superior substrate for COMT compared to the 5, 6isomer. Thus, the disparity of the biochemical and behavioural potencies is explained by the rapid O-methylation of the 6, 7-isomer in vivo (Youde et al., 1983). Another interesting example of the inactiviation of a complex molecule by Omethylation is susceptibility of β-lactam antibiotics to metabolism by COMT.

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Ohi et al. (1987) have published a series of studies on the antibacterial activity of 3, 4-dihydroxybenzoyl derivatives of 2phenylacetamidopenicillanic acids, showing that antibacterial activity is inversely related to the affinity of the derivatives for COMT in vitro and in vivo. In addition many drugs and xenobiotics possess mono-phenolic groups in their structure and as such are possible candidates for further hydroxylation by a variety of microsomal hydroxylases resulting in catechol formation (Daly et al., 1965). The simplest example is phenol itself which undergoes aromatic hydroxylation to form catechol which is readily O-methylated by COMT (Daly et al., 1965). In this regard, p-bromophenol is hydroxylated to 4-bromo-catechol and excreted as the non-toxic glucuronide of the O-methylated product (Lau et al., 1984; Monks et al., 1984). A smaller group of drugs bearing aromatic ring structures undergo P-450-catalyzed hydroxylation to yield monophenolic metabolites that then undergo a second hydroxylation to form catechols (Billings, 1985). An example of this pathway occurs with the xenobiotic, biphenyl, which has been shown to undergo sequential aromatic hydroxylation, first to 4-hydroxybiphenyl and then to the COMT substrate, 3, 4-dihydroxybiphenyl in both mouse and rat (Halpaap-Wood et al.). Dihydroxyindoles are formed by microsomal hydroxylases from many hydroxyindoles. One example is the hydroxylation of 5-hydroxyindole to form 5, 6-dihydroxyindole, which then undergoes O-methylation to the 6-methoxy derivative (Daly et al., 1965). Many phenolic phenethylmines can undergo hydroxylation to form catechols, such as synephrine, which is hydroxylated to form p-hydroxyephedrine (Inscoe et al., 1965). An early example of this hydroxylation pathway was the demonstration that norepinephrine can arise in vivo through the β-hydroxylation of tyramine to form norsynephrine, followed by ring-hydroxylation to give rise to norepinephrine (Creveling et al., 1962). A similar series of reactions demonstrating the cytochrome P-450-dependent hydroxylation of amphetamine to form p-hydroxyamphetamine and then αmethyldopamine and to some extent α-methylnorepinephrine was reported by Hoffman et al. (1979a, 1979b). These catechols are rapidly O-methylated both in vitro and in vivo by COMT. More significant perhaps is the formation and Omethylation of catechols from analgesics, such as morphine, nalorphine, levophanol, phenazocine and other potential catecholamine alkaloids (Daly et al., 1965; McKenzie and White, 1973). A recent note suggests that the dopamine receptor agonist, dipropyl-5, 6-dihydroxyaminotetralin, is inactivated by O-methylation (Rollema and Grol, 1983). The putative dopamine autoreceptor agonist, (3-(3hydroxyphenyl)-N-n-propylpiperdine (NilssonandCarlsson, 1982), undergoes aromatic hydroxylation to the 4-hydroxy catechol derivative which is an excellent substrate for COMT, thereby accounting for the rapid inactivation of the drug in vivo (Bhaird et al., 1985).

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The major route for the metabolism of propranolol is through aromatic hydroxylation of the naphthalene ring yielding a series of monohydroxy naphthols (Talaat and Nelson, 1988a, 1988b). The major product of aromatic monohydroxylation in man is 4-hydroxypropranolol with lesser amounts of the 5- and 7-hydroxy derivatives (Walle et al., 1982, 1984). In the rat the major product is 4-hydroxypropranolol with decreasing amounts of the 7-, 5-, and 2hydroxy derivatives (Bond, 1967; Tindell et al, 1972; Walle and Gaffney, 1972). In keeping with the earlier observations of Daly et al. (1965) it is not surprizing that some of the monohydroxyphenols derived from propranolol also give rise to catechols. In 1978, Tindel et al. reported the formation of several catechol-like compounds derived from propranolol in incubations of rat liver preparations in vivo. The presence of two O-methylated catechol-like metabolites of propranolol were shown to be excreted as glucuronides in man (Walle et al., 1978). Later, a minor metabolite of propranolol, 3, 4dihydroxypropranolol (Nelson et al., 1984) was shown to give rise to 3methoxy-4-hydroxypropranolol in man, while in rat both the 3- and 4-methoxy derivative were detected (Gustavson and Nelson, 1988). While not documented, other naphthalene-based drugs, such as nadolol and nafronyl, can be expected to be metabolized to some extent through the formation of catechols. Finally, it should be noted that not only do oestrogens undergo aromatic hydroxylation in vivo to form catechol steroids but several steroid drugs follow the same series of reactions culminating in COMT-catalyzed inactivation. The principal metabolite of the contraceptive hormone, ethynylestradiol, is formed through the hydroxylation to the catechol 2-hydroxyethynylestradiol and rapid Omethylation by COMT (Bolt et al., 1973; Raxworthy and Gulliver, 1982). 7.8. In vivo inhibitors Although many inhibitors of COMT have been identified that are quite effective in vitro, very few of these compounds are selective and effective in vivo inhibitors of this enzyme. Among the three classes of inhibitors discussed previously (Section 7.5), only the isosteres of catechol substrates can be selective inhibitors of COMT because the SAH analogues inhibit other SAM-dependent methyltransferases, and the lanthanide metals are expected to disrupt the catalytic functions of several metal-dependent enzymes. The catechol isosteres suffer from a drawback in that many of them would have a phenolic group and, hence, would be amenable to other metabolic conjugation reactions and rapid elimination. A few in vivo inhibitors of COMT that are currently being used as pharmacological tools include 4-acetamidotropolene, pyrogallol and its derivative RO 4–4602, and 3′, 4′-dihydroxy-2-methylpropiophenone (U-0521 (cf. Guldberg and Marsden, 1975); however, none of these are in clinical use. A

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good in vivo inhibitor of COMT can be very useful in protecting catechol drugs from metabolic inactivation. L-DOPA provides an excellent example of a drug whose effectiveness can be improved by an in vivo inhibitor of COMT. In this regard a new class of catechol isosteres has recently been developed as inhibitors of COMT (Mannisto et al., 1988; Backstrom et al., 1989) consisting of substituted derivatives of 3-nitrocatechol. These derivatives inhibit COMT in vitro in the nanomolar range (Ki=3 to 35 nM) and thus are an order of magnitude more potent than other known inhibitors, such as U-0521 (Backstrom et al., 1989). These derivatives appear to have a specific affinity for the catechol binding site of COMT but do not appear to be substrates. They are characterized by tight binding to the enzyme and produce a long-lasting inhibition of the enzyme in vivo. The properties of this class of inhibitor, modified by a variety of substituents on the unoccupied positions of the aromatic ring of the catechol, can either pass through membrane barriers or are essentially restricted to the gut or periphery (cf. Section 7.7). 7.9. Conclusion In conclusion we would like to emphasize that methyltransferases and the methyl-donor co-substrate, SAM, play a pivotal role in diverse biological systems. In this chapter, we have discussed in considerable detail the role of one such methyltransferase, COMT. It appears that the role of COMT is underestimated. It is now quite apparent that the role of COMT is more extensive than in the inactivation of catechol xenobiotics, circulating catecholamines, and catecholamine neurotransmitters. The function of COMT in the reproductive system, the presence of sexual dimorphism with regard to COMT, the physiological and neoplastic alterations in the activity of COMT, and the extensive localization of COMT clearly point to a significant role in the inactivation of catechol oestrogens as a barrier for the passage of catechols between tissue compartments and the control of other, as yet unrecognized, catechol-mediated functions. A continuing problem is the nature of the multiple forms and isozymes of COMT. We feel that the present evidence for the apparent multiple forms of COMT, including our own, is based on less than definitive evidence. It can be expected that in the near future the DNA and amino acid sequence of COMT will be obtained with the subsequent resolution of the structure of the enzyme or enzymes, and evidence for post-translational modifications of COMT (Grossman et al., 1989). Of growing importance is the need to understand the relationship between abundant low-affinity S-COMT and the lesser amounts of the high-affinity MB-COMT. Are these forms of COMT independent of one another or does S-COMT provide a source for the membranous form? Are the levels and activity of MB-COMT modified by

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physiological or hormonal cues like S-COMT? The study of the role of Omethylation in vivo has long been hampered by the lack of specific, long-lasting, and non-toxic inhibitors of COMT. The recent development by Orion Pharmaceutica of Finland of a series of derivatives of 3-nitrocatechol appears to have provided the research community with a selection of specific, long-lasting, essentially irreversible inhibitors of COMT. Abbreviations CSF COMT DOPA DOPAC HVA MAO-B MB-COMT MOPET NE OMD PET RBC SAH SAM S-COMT

Cerebral spinal fluid Catechol-O-methyltransferase Dihydroxyphenylalanine 3, 4-Dihydroxyphenylacetic acid Homovanillic acid Monoamine oxidase B Membrane-bound COMT 3-Methoxy-4-hydroxyphenylethanol Norepinephrine 3-O-Methyl-Dopa=3 methoxy-y-hydroxy-phenylalanine Positron emission tomography Red blood cells S-Adenosylhomocysteine S-Adenosylmethionine Soluble COMT References

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 8 N-Methyltransferases Sherry S.Ansher1 and William B.Jakoby2 1

Center for Biologs Evaluation and Research, Food and Drug Administration, Bethesda, MD 20982, USA.

2

Laboratory of Biochemistry and Metabolism, NIDDK, National Institutes of Health, Bethesda, MD 20892, USA.

8.1.

INTRODUCTION

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8.2.

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233

8.3.

8.4.

8.5.

The enzymes

233

Substrates

234

Inhibitors

236

Role in detoxication

236

Formation of toxic and carcinogenic products

237

HISTAMINE METHYLTRANSFERASE

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The enzymes

239

Substrates

239

Inhibition

240

PHENYLETHANOLAMINE N-METHYLTRANSFERASE

241

The enzymes

241

Substrates

242

Inhibition

242

Physiological roles

243

METHODS OF DETECTION

244

ABBREVIATIONS

245

232 CONJUGATION REACTIONS IN DRUG METABOLISM

REFERENCES

245

8.1. Introduction N-Methylation may be among the more important reactions by which amino groups are modified, although this functional group is well endowed with several other means for its oxidation and conjugation (Jakoby et al., 1982). Conjugation as a route of metabolism also allows acetylation (Chapter 6); glycosylation, particularly by N-glucuronidation (Chapter 4); N-sulfation to form sulfamates (Chapter 5); and, possibly, carbamylation (Elvin et al., 1980). Recognition of Nmethylation, among the first of the conjugation reactions that was observed, occurred when His (1887) examined the urine of dogs after administration of pyridine and thereby discovered its conversion to the N-methylpyridinium ion. A large group of animals produced the same metabolic product (Abderhalden and Brahm, 1909; Totani and Hoshioi, 1910), and the reaction was subsequently extended to a broader group of substrates (Williams, 1947). First recognized in lung (Brown et al., 1959a, 1959b), enzymes that catalyze methylation at nitrogen have been found in extracts of liver, skin, brain and kidney, among other tissues. N-Methyltransferases appear to participate in two relatively separate functions. Already noted above, the detoxication of pyridine by conversion to a more readily excretable metabolite is a good example of the metabolism of xenobiotics. The reaction with pyridine is one example of catalysis by amine Nmethyltransferase, an enzyme capable of utilizing a large array of primary, secondary and tertiary amines. A second major role is the formation and inactivation of a number of normal amine metabolites that include the neurotransmitters epinephrine and histamine. These reactions are catalyzed by phenylethanolamine N-methyltransferase and histamine N-methyltransferase, respectively. This list of three is not the entire spectrum of N-methyltransferases but only those purified to homogeneity. An enzyme active in the formation of N′methylnicotinamide is known (Swiatak et al., 1973). Still waiting to be purified and characterized is a transferase that catalyzes methylation of theophylline to yield caffeine, a clinically significant reaction since the administration of theophylline to infants for asthma has led to morbidity due to toxicity from endogenously-formed caffeine (Bory et al., 1979). The source of the methyl group that is transferred in each instance is Sadenosyl-L-methionine (AdoMet) (see Chapter 7) and the products are secondary, tertiary or quaternary N-methylamines, in addition to S-adenosyl-Lhomocysteine (AdoHcy). The enzymes considered here are those for which

N-METHYLTRANSFERASES 233

information is available at the level of highly purified proteins. On this basis, the discussion is limited to amine N-methyltransferase, an enzyme of extraordinarily broad spectrum, and to the more specific histamine Nmethyltransferase and phenylethanolamine N-methyltransferase. 8.2. Amine N-methyltransferase Most tissues catalyze the N-methylation of a large number and variety of amines in which the methyl group is donated by AdoMet (Axelrod, 1962a; Mandell and Morgan, 1971; Narasimhachari et al., 1972; Wyatt et al., 1973). The reaction is shown for a primary amine (eqn. 8.1) but both secondary and tertiary amines are substrates. Although the substrate range is broad, if not enormous, as is now expected for enzymes active in the metabolism of xenobiotics (Jakoby, 1982), histamine and phenylethanolamine are conspicuous by their absence. (8.1) Initial reports of N-methylation of amines appropriately attributed the activity to ‘non-specific’ N-methyltransferase (Axelrod, 1962b) so as to distinguish this enzyme from those for which phenylethanolamine and histamine were substrates (Saavedra et al., 1973). When the enzyme was obtained in homogeneous form from rabbit liver as two isoenzymes, A and B, the enzyme was renamed amine N-methyltransferase (Ansher and Jakoby, 1986), the term adopted here. The same enzyme, prior to detailed knowledge of its specificity and its isoenzymes, was labelled as arylamine N-methyltransferase (Lyon and Jakoby, 1982) and, as such, is recorded as EC 2.1.1.49 by the International Union of Biochemistry (1984); the name is to be changed although it is presumed that the number will not. An enzyme purified from rabbit lung, indoleamine N-methyltransferase (Irace et al., 1982) is similar to that from liver; however, although two isoenzymes have been reported (Porta et al., 1979), it is not clear whether the electrophoretically homogeneous preparation contains one or two of the isoenzymes. The term indolethylamine methyltransferase has also been used (Borchardt, 1980). The enzymes It is from the rabbit that highly purified preparations of amine Nmethyltransferase have been obtained. Whether from rabbit liver (Ansher and Jakoby, 1986, 1987) or lung (Irace et al., 1982) the N-methyltransferases have similar molecular mass, 30 000, and a pI of 4·9 and 5·1 for the two isoenzymes. The search for the presence of liver isoenzymes began when differences were noted in the relative activity of partially purified fractions with tryptamine, aniline and desmethylimipramine as substrates. The ratio for the three substrates

234 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 8.1. Apparent kinetic constants for amine N-methyltransferasesa

a b

Data from Ansher and Jakoby (1986), Ansher et al., (1986) and Ziegler et al. (1988). Tris-HCl at 50 mM was used as buffer at the indicated pH.

in the standard assay was approximately 10:4:3 with crude and partially purified preparations. After complete separation of the two isoenzymes by high performance liquid chromatography (HPLC) on a preparative column of DEAEsilica, the ratio was established as 10:2:9 for N-methyltransferase A and 10:7:2 for N-methyltransferase B (Ansher and Jakoby, 1986). The two liver isoenzymes are clearly related since antibody to transferase A produces a precipitin line of identity with transferase B. Substrates The validity of the designation as an amine N-methyltransferase, rather than an arylamine N-methyltransferase or any other more specific label, is evident from the extensive but essentially qualitative data that is now available for the rabbit liver enzymes. Apparent kinetic constants (Table 8.1) have been calculated (Ansher and Jakoby, 1986; Ansher et al., 1986; Ziegler et al., 1988), but work with the enzyme is subject to specific substrate inhibition and to the difficulty of procuring and preparing sufficiently pure amines; the discussion in Chapter 5 on substrate purity applies here as well. With AdoMet as methyl-group donor, the amine N-methyltransferases accommodate primary (eqn. 8.1), secondary and tertiary amines as methyl acceptors whether they be aryl, alkyl, aralkyl or azaheterocyclic (Table 8.2). Tryptamine has been used as the assay substrate; its N-methylation product and 5-methoxytryptamine are also active substrates. Amino acids are not substrates and certain of the amines with central nervous system or related function, e.g. histamine, norepinephrine, dopamine, and octopamine are less than 5% as effective under standard assay conditions as tryptamine at either pH 7·6 or pH

N-METHYLTRANSFERASES 235

Table 8.2. Amine N-methyltransferase substrate spectrum: activity relative to tryptamine (data from Ansher and Jakoby, 1986).

a The relative activity of transferase A at pH 8.5 was 85% of that of transferase B with tryptamine at the same pH. b Activity less than 5% of that with tryptamine.

8·5. It will be noted that pH optima differ for different compounds although not for the two isoenzymes (Ansher and Jakoby, 1986). The methylation of azaheterocycles has been noted as beginning with the observation of conjugation of pyridine by His (1887). On the level of soluble enzymes, Damani and Cundy and their colleagues have used extracts of several tisues to show AdoMet-linked methylation of not only pyridine but also of R(+) nicotine and cotinine (Cundy et al., 1985; Damani et al., 1986a, 1986b). The weakly basic indole is not a substrate, although it had been considered that the indole nitrogen of tryptamine could be methylated (Lyon and Jakoby, 1982); this spurious conclusion was a consequence of an artifact of the fluorescence assay that was attempted to distinguish between methylation of a primary and a secondary amine (Crooks et al., 1986). The insertion of another, more basic nitrogen, into the six-membered ring to form azaindole, provides an effective substrate with methylation at the 7N-position (Crooks et al., 1988). A systematic but qualitative exploration of azaheterocyclic compounds as

236 CONJUGATION REACTIONS IN DRUG METABOLISM

substrates has been conducted (Crooks et al., 1988). Among the more active methyl-group acceptors were dichlorotetrahydro-isoquinoline and phenyltetrahydropyridine. 4-Phenylpyridine, bipyridyls, aminobiphenyl and benzidine are also good substrates for both isoenzymes; substitution at the 2 position of pyridine yields relatively poor substrates, probably due to steric hindrance since the basicity of the two isomers is not that different from other substitutions (Summers, 1984). Inhibitors These enzymes are uniformly competitively inhibited by micromolar concentrations of the common product, AdoHcy. The fully methylated product of tryptamine, N, N-dimethyltryptamine, is also a potent inhibitor with a Ki of less than 10 µM for both isoenzymes. A number of sulfhydryl reagents are inhibitory as well (Ansher and Jakoby, 1986; Lyon and Jakoby, 1982). A naturally occurring inhibitor was discovered quite early in the investigation of methyl transfer to amines and was found to be present in quantities sufficient to void any analysis of methyl-group transfer in crude enzyme preparations prior to dialysis (Axelrod, 1962a; Lyon and Jakoby, 1982). The inhibitor, methinin, has been partially purified from rabbit liver (Lyon et al., 1982, 1987) although it is present in other tissues including brain (Ansher et al., 1986). Methinin is inhibitory for an extensive array of methylation systems which include O, S and N-methylation. Curiously, methinin, over a ten-fold range of substrate concentration is competitive, not with the common substrate AdoMet, but with each of the methyl group acceptors whatever the functional atom (Lyon et al., 1982). Role in Detoxication The detoxication function of those enzymes which seem intended for countering human exposure to xenobiotics appears more tenuous than most for the amine N-methyltransferases. Two means for detoxication can be adduced for most of the conjugating enzymes. The first of these is in neutralizing the pharmacological activity of the unconjugated substrates. Certainly that occurs by methylation although, as with the other conjugating enzymes, the reverse can also be true: apomorphine, as an example, is methylated to form the active morphine. Methylation also accounts for what may be termed a ‘futile’ cycle that is responsible for the very slow rate at which the antidepressant, imipramine, is cleared from the circulation. Imipramine, a methylated tertiary amine, is readily oxidized to desmethylimi pramine which is further metabolized at a slower rate before being excreted. It has been observed, however, that desmethylimipramine is remethylated (Table 8.2) in vivo to imipramine. Thus,

N-METHYLTRANSFERASES 237

the N-methyltransferases seem to have a role in recycling this and probably other drugs. The second, if not primary, role of conjugation reactions in detoxication is generally considered as the conversion of a lipophilic molecule to a more hydrophilic one by reason of the polar group that is thereby affixed to the xenobiotic. Since the methyl group is not polar, its addition to primary and secondary amines produces no expectation of an increased rate of excretion. The exception here is the methylation of a tertiary amine to yield the positively charged quaternary cation that would be removed more readily by the kidney. The formation of quaternary cations by methylation needs emphasis since charged compounds of this sort will not easily cross the blood-brain barrier in either direction (Bodor, 1985). The idea was offered that the dosage of a therapeutic agent intended for the brain could be lowered by taking advantage of a prodrug bearing a tertiary amino group which could be methylated. Pyridine analogues for example, were shown to be methylated by soluble enzyme from human and other mammalian brain (Ansher et al., 1986). This strategy should decrease drug dosage since the efflux of methylated quaternary pyridinium ion would be expected to be slowed in brain. Formation of toxic and carcinogenic products In addition to producing a pharmacologically active compound, methylation may lead to the formation of highly toxic material. Not only the homogeneous liver amine N-methyltransferases, but enzyme preparations from brain were capable of catalyzing the N-methylation of 4-phenyl-1, 2, 3, 6tetrahydropyridine [1] (Ansher et al., 1986) to 1-methyl-4phenyltetrahydropyridine ([2]MPTP) which is in turn converted, probably by a monoamine oxidase (Langston et al., 1984; Markey et al., 1984), to 1-methyl-4phenylpyridinium ion ([3] MPP+). The last compound is implicated in the development of the Parkinsonium syndrome by destruction of dopamine neurons in the substantia nigra (Calne et al., 1985). These N-methyltransferases were also effective in directly converting 4-phenylpyridine [4] to MPP+. Whereas the Km for each (8.2)

of the methylation steps is relatively high and the Vm low (Table 8.2), all are lipophilic compounds with access to brain.

238 CONJUGATION REACTIONS IN DRUG METABOLISM

(8.3) Methylation leading to toxicity is emphasized by pointing to 4, 4′dipyridyl [5] which yields the monomethyl precursor [6] of paraquat [7] (Ansher et al., 1986). The dipyridyl and its monomethyl product are equally toxic in producing pulmonary hemorrhage in acute tests (Groce and Kimbrough, 1982). It is significant that both 4, 4′dipyridyl [5] (Saint-Jalan and Moree-Testa, 1980) and 4-phenylpyridine [4] (Heckman and Best, 1981) are present in tobacco smoke. The activity of the amine N-methyltransferases seems to explain a peculiar situation in which certain aminobiphenyls and their analogues are hepatocarcinogenic, whereas other reasonable candidates for that role are ineffective. For adult rats, it is known that 4-aminoazobenzene and 2aminobiphenyl are not carcinogenic whereas the similar arylamines, benzidine and 4-aminobiphenyl, produce tumours. The importance of N-methyl substituents for hepatocarcinogenicity is emphasized by the early observation that N-methyl-4-aminoazobenzene is a carcinogen (Miller and Miller, 1953). In reviewing the data, Ziegler et al. (1988) accepted the assumption that carcinogenicity of arylamines requires oxidation at nitrogen, and that direct Noxidation in liver would not be a cytochrome P-450-catalyzed reaction (Damani, 1982; Ziegler, 1985) but a two-electron oxidation catalyzed by the flavin-containing monooxygenase (Williams et al., 1984; Ziegler, 1985). Although the flavin enzyme is inactive toward primary amines, it catalyzes the N-oxygenation of secondary and tertiary amines (Kadlubar et al., 1976; Ziegler, 1985). Thus, methylation would be necesary before an arylamine could generate the N-hydroxyarylamine that would be converted to the ultimate carcinogen (Ziegler et al., 1988). The hypothesis requires that the specificity of the two enzymes, N-methyltransferase and monooxygenase, be in accord with the pattern of hepatocarcinogenicity. There is very good agreement. The noncarcinogenic arylamine, 4-aminoazobenzene, is not a substrate for the homogeneous rabbit liver amine N-methyltransferases. However, given synthetic N-methyl-4-aminoazobenzene, which is carcinogenic, the flavin monooxygenase is able to oxidize it. Although benzidine, 4-aminobiphenyl and 2-aminobiphenyl are all methylated by the transferase, only the N-methyl derivatives of benzidine and 4-aminobiphenyl are oxidized by the monooxygenase and only benzidine and 4-aminobiphenyl are carcinogenic. For one group of arylamines, then, the sequential action of methyltransferase and monooxygenase appears necessary to achieve carcinogenicity (Ziegler et al., 1988).

N-METHYLTRANSFERASES 239

8.3. Histamine methyltransferase Methylation of histamine by tissue extracts was initially ascribed to an imidazole N-methyltransferase, the name stemming from the identification of the product, i.e. the methylated derivative at nitrogen in the imidazole ring (Brown et al., 1959b). Since it soon became clear that imidazole itself was inactive as a methylgroup acceptor, the term histamine methyltransferase (EC 2.1.1.8) is presently used. The enzyme catalyzes the transfer of a methyl group from AdoMet (Reaction 8.4) to form 3-N-methylhistamine (tele-methylhistamine; Black and Ganellin, 1974) in a number of tissues. In the kidney, gastric mucosa, skin and erythrocyte, the transferase appears to be a major means of histamine metabolism although attack of histamine by diamine oxidase also occurs (Mehler et al., 1952; Schayer, 1952). In brain, histamine is a putative neurotransmitter and the transferase is the means for inactivation of that signal. A detailed discussion of possible roles for the enzyme in histamine metabolism is available (Verburg and Henry, 1986). (8.4)

The enzymes Histamine N-methyltransferase has been brought to the stage of homogeneity from rat kidney (Bowsher et al., 1983) and guinea pig brain (Matuszewska and Borchardt, 1983). Relatively purified fractions have also been prepared from ox brain (Gitomer and Tipton, 1986), guinea pig skin (Tachibana et al., 1986) and porcine gastric mucosa (Barth et al., 1984), although the distribution of the enzyme is wider. Its ontogeny has been studied (reviewed by Verburg and Henry, 1986). All preparations have a similar Km for histamine (3 to 27 µM) and for AdoMet (10 to 40 µM). Protein size is in the range of 29 000 to 35 000, and the enzyme is monomeric. Antibodies raised against the rat kidney enzyme precipitated the enzymes from rat intestine, rat brain and guinea pig brain (Bowsher et al., 1983). Substrates As noted, the enzyme from all sources is relatively specific, the most specific of the three N-methyltransferases that are discussed in this chapter. Methylgroup acceptor substrates are limited to those compounds in which positions 1, 2 and 3 are unsubstituted and in which a positive charge is close by on the side chain.

240 CONJUGATION REACTIONS IN DRUG METABOLISM

Imidazoles with a negative charge on the side chain are neither substrates nor inhibitors (Barth and Lorenz, 1978; Barth et al., 1980). Substitution of the ethylamine side chain by a propylamine does not change the Km (Dent et al., 1982) but methylating the primary amino group of histamine to Nmethylhistamine reduces the Km by an order of magnitude (Hough et al., 1981). Spinaceamine (4, 5, 6, 7, -tetrahydroimidazo-[5, 4-c]pyridine) and its N-αmethyl derivative are both substrates (Barth et al., 1980). Although the D isomer of AdoMet was inactive as a methyl donor (Zappia et al., 1969), Borchardt and his colleagues have been successful in altering AdoMet and in preparing a number of derivatives, each of which has a much higher Km, although maximum velocity can remain the same as for AdoMet (Borchardt, 1980). Good kinetic evaluations have been difficult to achieve since all except the enzyme from ox brain are severely inhibited by a ten-fold higher than Km concentration of the substrate histamine; all methyltransferases are subject to inhibition at low concentration of one of the products, AdoHcy. The transferase from ox brain, while not homogeneous, is highly purified and stable, in addition to being insensitive to histamine inhibition. Kinetic analysis of that enzyme is consistent with a ternary complex of methyl donor and methyl acceptor with enzyme but without the requirement of a necessary order of binding (Gitomer and Tipton, 1986). Cruder preparations from human skin, a preparation that is subject to histamine inhibition, suggests an ordered steadystate mechanism involving ternary complexes with enzyme (Francis et al., 1980). More difficult to understand is the finding of kinetics consistent with a double-displacement type of kinetic mechanism as reported for a preparation from guinea pig brain (Thithapanda and Cohn, 1978). The last proposal implies formation of an active methyl group-enzyme complex for which a model is difficult to picture. In fact, the methyltransferase-catalyzed reaction, transfer of the methyl group from AdoMet to histamine, has been shown to occur with configurational inversion, a finding that strongly favours direct transfer to histamine without a methyl-enzyme intermediate (Asano et al., 1984). Inhibition In direct contrast to the limited substrate range of the enzyme, histamine methyltransferases are inhibited by many types and a large number of compounds. Included in this array are antimalarial drugs (quinacrine, amodiaquine), H-1 (diphenhydramine, chlorpromazine) and H-2 (metiamide, burimamide, cimetidine) receptor antagonists, thioureas (dimaprit), diuretics (ethacrynic acid, merallutide) and local anaesthetics (dibucaine, procaine). Extensive lists of these inhibitors have been prepared and discussed (Thithapanda and Cohn, 1978; Beaven and Shaff, 1979; Barth et al., 1980; Duch et al., 1984; Verburg and

N-METHYLTRANSFERASES 241

Henry, 1986; Harle and Baldo, 1988; Tachibana et al., 1988). A number of biogenic amines, compounds such as tyramine, tryptamine, dopamine and serotonin, are all competitive with histamine in inhibiting the Nmethyltransferase (Tachibana et al., 1986). Such amines are present as normal components of the inflammatory response, and the inhibitory interaction with histamine may affect that response. 8.4. Phenylethanolamine N-methyltransferase The other relatively specific enzyme of interest is phenylethanolamine Nmethyltransferase, EC 2.1.1.28, (Axelrod, 1962a, 1962b) which has also been labelled as phenethanolamine or norepinephrine (or noradrenaline) methyltransferase. The physiological reaction is methylation of norepinephrine to epinephrine (eqn. 8.5), the terminal step in the biosynthesis of the hormone. This N-methyltransferase is present at its highest concentration in the chromaffin cells of the adrenal medulla, the source from which it has been purified. It is found in specific areas of brain, particularly in the hypothalamus and brain stem (Fuller and Hemrick-Luecke, 1983) but also to some degree in the retina, heart and liver (Kirschner and Goodall, 1957; Axelrod, 1962b; Axelrod and Vesell, 1970). (8.5)

The enzymes The critical role of its natural product, epinephrine, is undoubtedly responsible for the extensive study of this cytosolic enzyme, but only the transferases from the bovine (Connett and Kirshner, 1970) and rabbit (Lee et al., 1978) adrenal gland have been purified to homogeneity. The bovine and human enzymes have been cloned (Batter et al., 1988) which has led to comparison with the amino acid sequence determined by protein chemistry (Weisberg et al., 1988). The amino acid sequence of the human enzyme was highly homologous (88%) to the bovine enzyme. The human gene is on chromosome 17 (Kaneda et al., 1988). Multiple forms of the transferase have been sought or shown for each of several species that range from only one in rat adrenal (Park, 1986), to two in the dog (Pohoracky et al., 1973), to five in the cow (Joh and Goldstein, 1973; Park, 1986) and rabbit (Lee et al., 1978). At least one form is present only in young rabbit adrenal glands whereas another is present only in the adult (Lee et al., 1978).

242 CONJUGATION REACTIONS IN DRUG METABOLISM

The reported Mr has ranged from 35000 to 40000 but the immunoprecipitable protein obtained after translation of bovine mRNA, has an Mr of 31000 (Park et al., 1982; Baetge et al., 1983). The bovine preparation was shown to be glycosylated, and the glycosylated form has been identified as subject to a high degree of stimulation by phosphate ion (Park and Joh, 1984; reviewed by Park, 1986). Antibodies raised against adrenal transferases react well with enzyme from the brain stem of the same species, and there is some cross reactivity among different species (Joh and Goldstein, 1973; Pohoracky and Baliga, 1973; Kondo et al., 1982; Park et al., 1982). Substrates A variety of phenylethanolamines, but not phenylethylamine, serve as methylgroup acceptors with AdoMet as the donor molecule. Thus, the β-hydroxyl group on the alkyl side chain is of major importance although there are exceptions (Rafferty and Grunewald, 1982). Among the naturally occurring substrates are norepinephrine, epinephrine (to form the dimethyl, tertiary amine), octopamine, normetanephrine, metanephrine, and synephrine. As a generalization, primary amines are better substrates than secondary amines. A low level of methyl-group acceptor activity is evident with compounds in which the side chain β-hydroxyl group is substituted with an amino group, or compounds in which a chloro group is in the ortho, meta or para position of the aromatic ring (Fuller et al., 1977). Compounds in which the aromatic ring is replaced by cyclohex-3-enyl, cyclohexyl or cyclooctyl groups are very good substrates (Grunewald et al., 1975). The effect of aromatic substituents on the apparent kinetic constants for the enzyme from bovine adrenal gland are shown in Table 8.3 (Fuller, 1987). It is apparent that the enzyme is particularly efficient with norepinephrine, the physiologically important substrate. Although it is dangerous to correlate affinity with the approximate apparent Km values that were obtained using the rabbit adrenal enzyme, the data suggest that the effectiveness of a substrate for methylation is a function of binding to the enzyme rather than of the catalytic step. Km changes over three orders of magnitude but the maximum velocity is relatively constant. Inhibition Whatever its source, the enzyme is subject to strong substrate inhibition to the extent that high concentrations of catecholamines in the circulation can invalidate the results of enzyme assays (Fuller and Marsh, 1972). One of the products of the reaction, AdoHcy, is a strong inhibitor as it is for all enzymic methylation reactions. The other product, generally a secondary methylated

N-METHYLTRANSFERASES 243

Table 8.3. Effect of aromatic substitution in phenylethanolamine for bovine phenylethanolamine N-methyltransferasea

a

Data from Table 8.1 of Fuller, R.W. (1987).

amine, is also a substrate for the enzyme and, therefore, a competitive inhibitor of the reaction. Sulfhydryl reagents are inhibitory as is methinin (Lyon et al., 1982). Because of its importance in forming epinephrine, phenylethanolamine Nmethyltransferase has been an appropriately tempting target for efforts to modulate the synthesis of epinephrine. A large number of compounds have been tested both in vivo and in vitro in an attempt to assess their therapeutic value, but a detailed discussion of these aspects is beyond the scope of this review. It is of significance in the context of this chapter that many such inhibitors are phenylethylamines, tetrahydroisoquinolines and other amines, i.e. in general, the very substrates of amine N-methyltransferase, the enzyme with the broad substrate spectrum that is discussed at the beginning of this chapter. Physiological roles Epinephrine in the central nervous system is a neurotransmitter participating in the regulation of temperature, reproduction, cardiovascular function and food intake. It also functions as a regulator of blood pressure, of pituitary hormones, and of a2 adrenoreceptors. Enzymes charged with the task of producing such vital regulators of metabolism and physiological responses are themselves strongly regulated. Phenylethanolamine N-methyltransferase is not an exception and is subject to regulation by the product, epinephrine, as well as by a number of hormones and other factors (summarized by Parker, 1986). Marked differences in enzyme activities exist due to gender, during development from foetus to newborn, and in specific organs during development (Padbury, 1983a, 1983b).

244 CONJUGATION REACTIONS IN DRUG METABOLISM

8.5. Methods of detection All of the N-methyltransferases can be assayed by modifications of a procedure (Axelrod, 1962a, 1962b) that utilizes AdoMet radio-labelled in the methyl group. Following incubation at 37° for 15 to 60 min, the reaction is terminated by addition of alkaline sodium or potassium borate. The resulting unionized, methyl-substituted amine is extracted and separated from radio-labelled AdoMet by use of an organic solvent. Borate, rather than KOH, is used to terminate the reaction since lower blanks result, presumably because of its binding to the cis hydroxyl groups of the sugar moiety of AdoMet (Bowsher et al., 1983). None of the methods can be trusted when crude extracts of tissue are used because of the presence of methinin (Lyon et al., 1982). The procedure has been simplified for the amine methyltransferases when primary or secondary amines serve as substrate (Ansher and Jakoby, 1986). The reaction mixture, 200 µl, is incubated in a scintillation vial. After termination with potassium borate, Econofluor, a non-aqueous commercial scintillation fluid, is added directly to the vial which is then shaken to extract the methylated product. Since the scintillation process is one of fluorescence under conditions in which the quantum yield is orders of magnitude greater in organic solvent than in aqueous solution, these vials can be assayed directly, without removal and transfer of the organic phase after extraction. The presence of large amounts of radioactivity due to AdoMet in the aqueous phase provides a slightly higher background. In general background values may be reduced by precipitating radioactive AdoMet before extraction (Fuller and Hunt, 1966) or by extracting after formation of an ion pair (Henry et al., 1975). These methods are not effective for extracting quaternary amines because of the positive charge on the nitrogen. Such incubation mixtures are terminated with boric acid or HCl, to which is added an ion-pairing reagent, e.g. 0.1M sodium dodecylsulfate. The complexed product is extracted with ethylacetate and can be measured for radioactivity, again in the same vial, following addition of Econofluor (Ansher et al., 1986). Specific reaction products are presently identified by HPLC methods. An assay for phenylethanolamine methyltransferase makes use of this approach (Borchardt et al., 1977). The phenylpyridines and bipyridyls can be separated on C18 or cation-exchange columns (Ansher et al., 1987; Crooks et al., 1988). Amino and diazaheterocyclic derivatives have been identified by similar methods (Cundy and Crooks, 1984; Crooks et al., 1988).

N-METHYLTRANSFERASES 245

Abbreviations AdoHcy MPP+ MPTP

S-adenosyl-L-homocysteine 1-methyl-4-phenylpyridinium ion 1-methy1-4-phenyltetrahydropyridine References

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Kondo, Y., Nagatsu, I., Karasawa, N., Kojima, H. and Nagatsu, T. (1982), Biomedical Research, 3, 197–206. Langston, J.W., Irwin, I., Lanston, E. and Forno, L.S. (1984), Neuroscience Letters, 50, 289–94. Lee, H.-S., Schulz, A.R. and Fuller, R.W. (1974), Archives of Biochemistry and Biophysics, 163, 735–41. Lee, H.-S., Schulz, A.R. and Fuller, R.W. (1978), Archives of Biochemistry and Biophysics, 185, 222–7. Lew, J.Y., Matsumoto, Y., Pearson, J., Goldstein, M., Hokfelt, T. and Fuxe, K. (1977), Brain Research, 119, 199–210. Lyon, S.E. and Jakoby, W.B. (1982), Journal of Biological Chemistry, 257, 7531–5. Lyon, E.S., McPhie, P. and Jakoby, W.B. (1982), Biochemical and Biophysical Research Communications, 108, 846–50. Lyon, S.E., Buonocore, L. and Miller, M. (1987), Molecular Cell Biology, 7, 1759–63. Mandell, A.J. and Morgan, M. (1971), Nature, 230, 85–7. Markey, S.P., Johannessen, J., Chiach, C.C. and Burns, R.S. (1984), Nature, 311, 464–67. Matuszewska, B. and Borchardt, R. (1983), Journal of Neurochemistry, 41, 113–8. Mehler, A.H., Tabor, H. and Bauer, H. (1952), Journal of Biological Chemistry, 197, 475–80. Miller, J.A. and Miller, E.C. (1953), Advances in Cancer Research, 1, 339–96. Narasimhachari, N., Plaut, J.M. and Himwich, H.E. (1972), Life Sciences, 11, 221–7. Padbury, J., Hobel, C., Gonzalez, F. and Fisher, D. (1983a), Biology of the Neonate, 43, 205–10. Padbury, J., Lam, R., Hobel, C. and Fisher, D. (1983b), Pediatric Research, 17, 362–7. Park, D.H. (1986), in Boulton, A.A., Baker, G.B. and Yu, P.H. (Eds.), Neuromethods Neurotransmitter Enzymes, p. 117, Cliffon: Humana Press. Park, D.H. and Joh, T.H. (1984), Biochemical Pharmacology, 33, 2911–3. Park, D.H., Baetge, E.E., Kaplan, B.B., Albert, V.R., Reis, D.J. and Joh,T. H. (1982), Journal of Neurochemistry, 38, 410–4. Pohoracky, L.A. and Baliga, B.S. (1973), Archives of Biochemistry and Biophysics, 156, 703–11. Pohoracky, L.A., Baliga, B.S., Wurtman, R.J. and Bartter, F.C. (1973), Endocrinology, 93, 566–74. Porta, R., Espostito, C., Camardella, M., Pietra, G.D. (1979), International Journal of Biochemistry, 10, 919–23. Rafferty, M.F. and Grunewald, G.L. (1982), Molecular Pharmacology, 22, 127–32. Saavedra, J.M., Coyle, J.T. and Axelrod, J. (1973), Journal of Neurochemistry, 20, 743–52. Saint-Jalan, Y. and Moree-Testa, P. (1980), Journal of Chromatography, 198, 188–92. Schayer, R.W. (1952), Journal of Biological Chemistry, 196, 469–75. Summers, L.A. (1984), Advances in Heterocyclic Chemistry, 35, 282–372. Swiatak, K.R., Simon, L.N. and Chas, K.L. (1973), Biochemistry, 12, 4670–4. Tachibana, T., Taniguchi, S., Imamura, S., Fujiwara, M. and Imamura, S. (1986), Experimental Molecular Pathology, 45, 257–69. Tachibana, T., Taniguchi, S., Imamura, S., Fujiwara, M. and Hayashi, H. (1988), Biochemical Pharmacology, 37, 2872–6. Thithapandha, A and Cohn, V.H. (1978), Biochemical Pharmacology, 27, 263–71.

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 9 S-Methylation James L.Stevens1 and Jerome E.Bakke2 1

W.Alton Jones Cell Science Center, 10 Old Barn Road, Lake Placid, NY 12946, USA.

2

Metabolism and Radiation Research Laboratory, Agricultural Research Service, USDA, Fargo, ND 58105, USA.

9.1.

INTRODUCTION

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9.2.

SUBSTRATES

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9.3.

9.4.

9.5.

9.6.

Thiol-containing drugs and xenobiotics

250

Certain substrates for thiol methyltransferases

251

REGULATION OF THIOL METHYLTRANSFERASE ACTIVITY

253

Regulation by co-factor availability

253

Regulation by hormones and enzyme induction

254

THE ENZYMES

254

Non-human thiol methyltransferases

255

Human thiol methyltransferases

256

Pharmacogenetics of thiol methyltransferase activity

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METABOLISM OF CHEMICALS BY THIOL METHYLTRANSFERASES IN VIVO

258

Methylation of thiol-containing xenobiotics

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Methylation of certain thiols

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Tissues involved in thiol methylation in vivo

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Further metabolism of methylthio conjugates in vivo

264

INHIBITION OF THIOL METHYLTRANSFERASES

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9.7.

BIOLOGICAL FUNCTION OF THIOL METHYL TRANSFER

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9.8.

PURIFICATION AND ASSAY PROCEDURES FOR THIOL METHYLTRANSFERASES

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ABBREVIATIONS

267

REFERENCES

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9.1. Introduction Transmethylation from S-adenosylmethionine (SAM) to alcohols, amines and thiols is a well-known route of drug metabolism (See Chapters7 and 8). In the case of thiols, methyl transfer clearly represents a detoxication step (Weisiger and Jakoby, 1980). Besides the multitude of xenobiotic thiols to which humans may be exposed, this enzyme plays a role in the detoxication of endogenous compounds, such as hydrogen sulfide, produced by enteric microorganisms (Weisiger et al., 1980). In this chapter, a review of the thiol methyltransferase enzymes and their roles in the metabolism and detoxication of xenobiotic thiols is provided. It brings together new data on very important but less obvious sources of xenobiotic thiols, of which the metabolism by methyl transfer is very important. Indeed, sulfur can be incorporated into xenobiotics after they enter the body; thus, methylthio metabolites from nonsulfur-containing xenobiotics are often found (Tateishi, 1983; Bakke, 1989). Finally, the current knowledge concerning the identity of the thiol methyltransferases, an area in which there is some disagreement, will be summarized. Others have recently reviewed thiol methyltransferases (Weisiger and Jakoby, 1980; Weinshilboum, 1984). 9.2. Substrates Thiol-containing drugs and xenobiotics Thiol methyltransferases are similar to many of the enzymes of detoxication in that they are promiscuous in their activity with thiols. Other than the presence of the free thiol in the molecule, it appears that the only structural requirement is that the compounds not be too hydrophilic. For example, it appears that glutathione, a very hydrophilic compound, is not a substrate for thiol methyltransferase (Loo and Smith, 1985a), though this was initially reported to be the case (Remy, 1963). Recent reviews provide detailed information on the

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substrate specificity of purified thiol methyltransferase (Weisiger and Jakoby, 1980; Tateishi, 1983; Jakoby et al., 1984); therefore, we will not present an extensive list. However, aromatic thiols are, in general, the best substrates while simple aliphatic thiols, such as mercaptoethanol, are less active. A number of therapeutic compounds are known to undergo thiol methylation. Thiopurine and thiopyrimidine drugs find a number of therapeutic uses and are metabolized by thiol methylation (Remy, 1963; Weinshilboum, 1984; Weinshilboum, 1987). Captopril, an antihypertensive compound, contains a free thiol and is a substrate for thiol methyltransferases (Drummer et al., 1983a, 1983b). Heterocyclic metabolites of cephalosporin antibiotics undergo methylation by human enzymes (Kerremans et al., 1985). The reduction product of antabuse (diethylthiocarbamate) has been isolated as a methylthio compound (Jakubowski and Gessner, 1972; Cobby et al., 1977). Penicillamine is methylated by thiol methyltransferase activity in human red blood cells (Keith et al., 1985). Several model substrates have been used extensively in assays for thiol methyltransferase activity. Thioacetanilide and 2-mercaptoethanol have been used in purification of the rat enzyme (Borchardt and Cheng, 1978; Weisiger and Jakoby, 1979). 6-Mercaptopurine has served as the substrate for purification of human kidney thiol methyltransferase (Woodson and Weinshilboum, 1983). An important consideration in choosing a substrate is the availability of the pure thiol uncontaminated by disulfides, to which thiols are easily oxidized. A second consideration is that the methylthio product may be volatile, as in the case of mercaptoethanol. This can present a safety problem since radioactive products may enter the laboratory environment. The use of thioacetanilide and 6-mercaptopurine circumvents these problems. Certain substrates for thiol methyltransferases Another important class of substrates for thiol methylation are generated from xenobiotics to which sulfur has been added from glutathione. In general, this pathway begins with the addition of glutathione followed by degradation to the cysteine conjugate. The cysteine conjugate is then cleaved via β-elimination by enzymes known as cysteine conjugate β-lyases (Anderson and Schultze, 1965; Tateishi et al., 1978; Stevens, 1984; Tomisawa et al., 1984; Larsen and Stevens, 1986; Stevens et al., 1986b, 1988) to pyruvate, ammonia and the thiol (eqn. 9. 1). The resulting thiol then serves as the methyl acceptor (eqn. 9.2). This pathway has been called the thiomethyl shunt since it diverts cysteine conjugate intermediates, which arise during mercapturic acid biosynthesis, from Nacetylation and excretion to cleavage and thiol methylation (Jakoby et al., 1984). The substrates for methylation produced by the β-lyase pathway, which have been studied most extensively in vivo, are propachlor (Larsen and Bakke,

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1981), benzothiazolyl sulfonamide (Colluci and Buyske, 1965), bromazepam (Tateishi and Shimizu, 1976; Tateishi et al., 1978), paracetamol (Mikov et al., 1988) and halogenated aromatic compounds (Bakke et al., 1981, 1988). (9.1) (9.2) Participation of the thiomethyl shunt pathway is only recognized after the isolation of methylthio metabolic products in excreta. Colluci and Buyske (1965) were the first to describe the introduction of sulfur from glutathione into a xenobiotic and the subsequent metabolism of the glutathione conjugate to the thiol. Schultze and colleagues (Anderson and Schultze, 1965; Saari and Schultze, 1965) first described bacterial and mammalian enzyme activities which cleaved cysteine conjugates to thiols by ß-elimination (eqn. 9.1). Subsequently, Tateishi et al. (1978) partially purified an enzyme from rat liver which catalyzed the ß-elimination of several cysteine conjugates. Homogeneous enzyme from both rat liver (Stevens, 1985) and kidney (Stevens et al., 1989b) have spectra typical of pyridoxal phosphate-dependent enzymes. Other enzymes for which spectral data are not available are inhibited by agents which inactivate pyridoxal phosphate-dependent enzymes (Tateishi et al., 1978; Stevens and Jakoby, 1983; Tomisawa et al., 1984; Larsen and Stevens, 1986). These mammalian enzymes appear to require that the substrate have a strong electron withdrawing group in the thioether moiety. Bacteria resident in the intestine contain ß-lyase activity with a much broader substrate specificity for cysteine conjugates (Saari and Schultze, 1965; Tomisawa et al., 1984; Larsen and Stevens, 1986). Due to this broad specificity, bacterial ß-lyases may be the most important source of substrates for thiol methyltransferases in vivo. Substrates for these enzymes include alkyl and aralkyl conjugates such as S-ethyl-, S-methyl- and S-benzyl-L-cysteine (Larsen and Stevens, 1986). It is germaine to point out that the organization of methylthio product formation via the thiomethyl shunt is complex and involves transport and enterohepatic circulation (Inoue, 1985; Schaeffer and Stevens, 1986; Bakke et al., 1987). A majority of the glutathione conjugates may be formed in the liver from which they exit to the gut via the bile (Chapter 11). Degradation of the glutathione conjugate yields the cysteine conjugate which can be metabolized by the bacterial ß-lyase in the gut or reabsorbed and returned to the liver (Bakke et al., 1987). Cysteine conjugates are acetylated in the liver and exported to the kidney (Inoue, 1985) where they can be further metabolized by deacetylation (Suzuki and Tateishi, 1981) or excreted. Therefore, three organs have the potential to methylate thiol substrates produced by this pathway. Tissue distribution studies show that all three have thiol methyltransferase activity. A

S-METHYLATION 253

recent report suggests that bacteria also contain thiol methyltransferase activity (Drotar and Fall, 1984), but the activity has not been measured in gut microflora. The participation of the various organs in thiol production will be discussed in detail using specific model compounds in Section 9.5. Several recent reviews cover the spectrum of compounds which are substrates for the thiomethyl shunt and discuss in detail the physiology of mercapturic acid biosynthesis (Tateishi, 1983; Jakoby et al., 1984; Inoue, 1985; Bakke et al., 1987; Stevens and Jones, 1989). 9.3. Regulation of thiol methyltransferase activity Regulation by co-factor availability Few data are available on the regulation of thiol methyltransferase activity by the availability of SAM. However, based on correlations with other methyltransfer reactions we may conclude that SAM may be rate-limiting for thiol methyltransferase activity as well. Regulation by co-factor availability can occur via a decrease in SAM content of a tissue or by an increase in the Sadenosylhomocysteine (SAH) concentration, since SAH is a competitive inhibitor of transmethylation (Borchardt and Cheng, 1978; Loo and Smith, 1985b). Loo and Smith (1985b) showed that a reduction in the SAM/SAH ratio in vitro resulted in decreases in both thiopurine and mercaptoethanol methylation. Administration of D, L-homocysteine also decreased the ratio of SAM/SAH in rat liver but only thiopurine methyltransferase activity was reduced. Based on these results, it was suggested that there is compartmentalization of SAM in hepatocytes. Tissue levels of SAM can also be reduced transiently by the administration of methylation substrates (Guldberg and Marsden, 1975). A brief review of SAM biosynthesis appears elsewhere in this volume (Chapter 7), and Mulder and Krijgsheld (1984) have published details of SAM regulation in vivo. For our purposes it is sufficient to point out that the availability of methionine is a critical step in SAM biosynthesis. Transmethylation is inhibited by the administration of methionine analogues that interfere with SAM biosynthesis, such as ethionine or methionine sulfoximine, or nucleoside analogues of SAM, probably through alteration in tissue SAM levels (Shull et al., 1966; Villa-Trevino et al., 1966; Griffith et al., 1979; Kredich, 1980; Tisdale, 1980; Tsukuda et al., 1980; Zimmerman et al., 1980).

254 CONJUGATION REACTIONS IN DRUG METABOLISM

Regulation by hormones and enzyme induction Modest increases (1.3-fold) in thiol methyltransferase activity have been observed upon administration of 3-methycholanthrene (Fujita et al., 1973b). The same authors reported that male rats have 1.4-fold higher transmethylation activity than females. Woodson et al. (1981) found that thiopurine methylation in rat kidney was two-fold higher in males compared to females. Both hypophysectomy and orchiectomy reduced circulating levels of testosterone and reduced kidney thiopurine methylation activity. Testosterone administration restored activity to near normal. The pharmacological and toxicological significance of methyltransferase induction is not clear. 9.4. The enzymes Early knowledge of thiol methyltransferases comes from the original observations by Remy (1963) and Bremer and Greenberg (1961a, 1961b). While investigating choline biosynthesis in microsomal fractions from rat, Bremer and Greenberg (1961a) observed that sulfhydryl compounds inhibited choline formation because they served as alternative methyl acceptors for SAM. Subsequently, they described an activity in rat liver microsomes which catalyzed the methylation of a variety of ‘nonphysiological’ thiols, but was inactive with ‘physiological’ substrates such as glutathione, homocysteine and cysteine (Bremer and Greenberg, 1961b). Rat liver and kidney contained equal amounts of the activity. Remy (1963) later described a soluble thiol methyltransferase activity in the rat which was active with thiopurine and thiouracil. Unlike the microsomal activity, inhibition studies with the soluble thiopurine methyltransferase suggested that ‘physiological’ thiols, such as cysteine and glutathione, were substrates for the enzyme. However, methylated products of cysteine and glutathione were not isolated. More recent work by Loo and Smith (1985b) suggests that inhibition is due to a reduction in the assay pH caused by the high concentrations of glutathione. They found no evidence for inhibition of soluble thiol methyltransferase activity when the assay pH was held constant. The soluble thiopurine methyltransferase is reported to be much higher in kidney than liver (Remy, 1963). Fujita and Suzuoki (1973) and Fujita et al. (1973a; 1973b) studied a rat liver enzyme activity, similar to that described by Bremer and Greenberg (1963b), which catalyzed the methylation of tetrahydrofurfuryl mercaptan. The activity was highest in the microsomal fraction of liver followed by kidney and small intestine. There was little or no activity found in the soluble fraction of liver. Therefore, the tissue distribution and subcellular localization were consistent with the thiol methyltransferase described by Bremer and Greenberg (1961a, 1961b). Taken together, these early

S-METHYLATION 255

studies suggested the presence of two enzymes: 1) a soluble thiopurine methyltransferase and 2) a microsomal thiol methyltransferase. More recent work has added substantial knowledge which clarifies the role of thiol methyltranferases in detoxication (Weisiger and Jakoby, 1980). However, there is now disagreement as to the nature and number of thiol methyltransferases. In the following summary of the literature we will draw attention to areas of agreement and disagreement. For clarity we will refer to microsomal and soluble thiolmethyltransferase activity, sometimes called thiopurine methyltransferase (Weinshilboum, 1984). As will become clear, the crux of the disagreement is whether the soluble thiol methyltransferase is in fact the microsomal enzyme which has been released from the microsomes (Weisiger and Jakoby, 1979, 1980). Several facts are key issues in the identity of soluble and microsomal thiol methyltransferase: substrate specificity, inhibitors, and subcellular localization. The fact that the microsomal enzyme can be solubilized by freezing or high concentrations of salt (Weisiger and Jakoby, 1979) and that the substrate specificity of the soluble and microsomal enzymes overlap (Weisiger and Jakoby, 1979; Woodson et al., 1983) precludes drawing a clear distinction at present. In addition, it has been shown that the substrate specificity of the microsomal enzyme can change upon solubilization (Brunner and Tegtmeier, 1983). Obviously, the solution to the problem is the simultaneous purification of both activities to determine if they are in fact the same or different. Unfortunately, at this writing definitive experiments have not been done. Non-human thiol methyltransferases Borchardt and Cheng (1978) solubilized an enzyme from rat liver microsomes which catalyzed transmethylation between SAM and mercaptoethanol. Solubilization from sucrose gradient-purified microsomes was achieved with Triton X-100. The enzyme has a pH optimum between 7·8 and 9·0. However, as will be discussed in Section 9.8, assays are carried out at pH 7·9 due to the instability of SAM at basic pH. Addition of EDTA or metal ions did not alter the enzyme activity unlike that of catechol-O-methyltransferase. However, sulfhydryl active agents, such as N-ethylmaleimide and p-chloromercuribenzoate, inhibited S-methyl transfer under conditions which did not interfere with thiol substrates. A thorough study of inhibition by analogues of SAH and SAM was also done (Borchardt and Cheng, 1978). Weisiger and Jakoby (1979) solubilized and purified rat liver thiol methyltransferase to apparent homogeneity. They observed that freezing liver to −70°C resulted in 30% solubilization of the microsomal enzyme. Therefore, they used a soluble fraction from frozen rat liver for purification of rat liver thiol methyltransferase. As the enzyme has similar properties to that purified by

256 CONJUGATION REACTIONS IN DRUG METABOLISM

Borchardt and Cheng (1978), Weisiger and Jakoby believed it to be the solubilized microsomal enzyme. They concluded that there is one enzyme whose location can be altered by freezing and thawing rat liver. The purified enzyme is a 28000 dalton monomer with an isoelectric point of 6–2 (Weisiger and Jakoby, 1979). A wide variety of xenobiotic thiols serve as substrates, with Km values varying from the submicromolar to the millimolar range. However, Vmax values vary by only eight-fold between 1 and 8 mM. As reported by Bremer and Greenberg (1961b), the enzyme did not use cysteine or glutathione as substrate. Dithiothreitol was reported not to be a substrate for rat liver thiol methyltransferase by Weisiger and Jakoby (1979). However, in a more recent study, Donahue et al. (1988) have shown that the assay used by Weisiger and Jakoby (1979) would not have detected the monomethylated product of dithiothreitol. Dithiothreitol is indeed a substrate for the thiol methyltransferase, albeit a rather poor one . Brunner and Tegtmeier (1983a) studied the solubilization characteristics of pig liver thiol methyltransferase. A number of detergents were effective in solubilizing the activity. A series of alkylsulfides were tested as substrates for the native and solubilized enzymes. When the enzyme was resident in the microsomes, methylsulfide was the best substrate, but the solubilized enzyme had a different specificity with heptanethiol acting as the best substrate. Thus the substrate specificity can change when the microsomal enzyme is solubilized. These authors (Tegtmeier and Brunner, 1983) also studied immobilization of the enzyme by covalently attaching them to agarose beads by co-polymerization with acrylamide. However, the immobilized enzyme was less stable than the soluble enzyme. Human thiol methyltransferases Weinshilboum and co-workers have measured thiol methyltransferase activity in human erythrocytes (Weinshilboum et al., 1978), lymphocytes (van Loon et al., 1987) and kidney (Woodson and Weinshilboum, 1983). An enzyme has been partially purified from the soluble fraction of human kidney (Woodson and Weinshilboum, 1983). The enzyme has an apparent Mr of 36000 daltons and a pH optimum of 6.7 with 6-mercaptopurine as substrate. Aromatic thiols are the best substrates for the enzyme with apparent Km values in the 1–10 µM range (Woodson et al., 1983). Heterocyclic thiols have intermediate activity with apparent Km values in the millimolar range, while alkyl thiols are poor substrates (Woodson and Weinshilboum, 1983). Woodson and Weinshilboum (1983) suggest that the substrate specificity of the soluble thiol methyltransferase differs from the microsomal enzyme; the spectrum of activity is similar to that reported by Weisiger and Jakoby (1979) for the solubilized microsomal enzyme. However, a later report (Woodson et al.,

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1983) demonstrated considerable overlap in substrate specificity. Several other parameters were reported to distinguish soluble from microsomal thiol methyltransferases including subcellular distribution, inhibitor sensitivity and response to inhibitors (Woodson and Weinshilboum, 1983). However, the distinction is murky due to the changes in the substrate specificity of the microsomal enzyme upon solubilization (Tegtmeir and Brunner, 1983). The report that the soluble enzyme can catalyze the methylation of physiological thiols, such as glutathione (Remy, 1963), appears to be incorrect (Loo and Smith, 1985b) and should not be used to distinguish the two activities. Simultaneous purification of both activities and partial sequence analysis of the enzymes has not been done at this time; thus, a final resolution must await such definitive data. Pharmacogenetics of thiol methyltransferase activity Like other enzymes of drug metabolism, methyltransferases show genetic variations. Catechol-O-methyltransferase and thiol methyltransferase activity varies within the population (Weinshilboum et al., 1974; Weinshilboum, 1980, 1984, 1987). Studies by Weinshilboum and associates show that human thiol methyltransferase varies within the general population (Weinshilboum, 1987). Using human red blood cells as a source of enzyme, Weinshilboum and colleagues observed a trimodal distribution with 89% in the high methylator category (13·5±1·86 units ml−1), 11% in the low methylator category (7·2 ±1·08 units ml−1) and 0·3% with undetectable activity (Weinshilboum, 1987; Weinshilboum and Sladek, 1980). There is a good correlation between red blood cell thiol methyltransferase and phenol-O-methyltransferase, though other enzymes do not vary accordingly (Keith et al., 1983). Woodson et al. (1981) showed that the variations in red blood cell thiol methyltransferase correlate well with the activities in visceral tissues which are probably the major site of S-methylation in humans. Lymphocyte thiol methyltransferase activity also correlates with red blood cell activity (van Loon and Weinshilboum, 1982). Inheritance of high, intermediate or low thiol methylation in humans, is autosomal co-dominant (monogenic; Weinshilboum and Sladek, 1980). Segregation of alleles at a single locus, designated TPMTh, for high activity, and TPMTl, for low activity, is responsible for genetic variability in thiol methyltransferase activity. Recently, Otterness and Weinshilboum (1987) reported that a similar monogenic inheritance pattern is apparent in C57BL/ 6J and AKR/J mice. The mouse provides an attractive pharmacogenetic model to investigate the biochemical mechanism of inheritance. Further details can be found in the appropriate references.

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The significance of pharmacogenetic variability in thiol methyltransferase activity lies in the role of methylation as a detoxication mechanism for thiols (Jakoby et al., 1984; Lennard et al., 1987). 6-Thioguanine nucleotides are found as major metabolites of 6-mercaptopurine and azothiopurine (Lennard and Maddocks, 1983). It is believed that thioguanine nucleotides play a role in myelosuppression, a complicating side effect observed in some patients undergoing treatment with thiopurine chemotherapeutic agents (Lennard et al., 1987). A significant correlation was found to exist between myelosuppression and low thiol methyltransferase activity in red blood cells of patients undergoing thiopurine therapy (Lennard et al., 1987). An obvious result of such a hypothesis is that methylated thiopurines are poor .substrates for 6thioguanine nucleotide synthesis. Given that thiols are formed, via microbial and mammalian enzymes and as endogenous metabolites of many chemicals, it is possible that genetic variability could play a role in intoxication from xenobiotic thiols derived from less obvious routes of metabolism (Jakoby et al., 1984; Bakke, 1987; Stevens and Jones, 1989). 9.5. Metabolism of chemicals by thiol methyltransferases in vivo The role of thiol methyltransferase in xenobiotic metabolism and the exogenous sources of substrates are outlined in Figure 9.1. These substrates come from two general sources. One source is thiol-containing xenobiotics which are encountered either deliberately, as in therapeutic drugs, or inadvertently as environmental contaminants. The other source is from thiols produced as a result of the intermediary metabolism of xenobiotics. These metabolicallyproduced thiols can result from hydrolysis of esters, as in metabolism of the Sacetate of spironolactone (Karim et al., 1976), and 0, 0dimethyl phosphorodithioate esters (Gage, 1967; Esser et al., 1968); cleavage of disulfide bonds, as in thiamine tetrahydrofurfuryl disulfide (Suzuoki-Ziro et al., 1967), and tetraethylthiuram metabolism (Gessner and Jakubowski, 1972); and from catabolism of glutathione (GSH) conjugates via the cysteine conjugate β-lyase pathway (Bakke and Gustafsson, 1984). Methylation of thiol-containing xenobiotics The paucity of reports in which in vivo methylations of thiols are discussed probably reflects the number of commercially-used xenobiotics that have free or readily accessible thiol groups; it does not reflect the prevalence of Smethylations in detoxication processes. This is especially true with the discovery of thiol methylation during glutathione conjugate catabolism. The early reports on thiol methylation also show a great variation in the extent of thiol

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Figure 9.1. Possible sources of substrates for thiol methyltransferase (SMT) and the metabolic disposition of thiols and products of SMT-catalyzed reactions.

methylation. For example, subcutaneous doses of [35S]2-thiouracil were excreted as 2-methylthiouracil (6%), sulfate (5%) and unchanged (61%) (Sarctone and Sokal, 1958) but oral doses of [14C]thiophenol were excreted as metabolites in which 40–50% of the thiol groups had been methylated (McBain and Menn, 1969). It may be that differences in the methylation of these two compounds result from the route of dosing; however, this has not been tested. Methylation of certain thiols The GSH-mediated metabolic introduction of thiol groups into xenobiotics (see Figure 9.1) has been of recent interest because of the renal toxicities attributed to some of the thiols that are formed by the β-lyase pathway (Anders et al., 1988; van Bladeren, 1988). It is of interest that methylthio-containing metabolites of nephrotoxic vinyl-halogen compounds have not been detected, probably due to the reactivity of the thiol (Derr and Shultze, 1963; Odum and Green, 1985; Stevens et al., 1986a; Anders et al., 1988). Thiols formed in this pathway are often excreted as the methylated conjugate and/or one of its other oxidation congeners (methylsulfoxides and methylsulfones) as shown in Figure 9.1. This oxidation pathway for methylthio-metabolites appears to be prevalent because at least 35 xenobiotics are known to be excreted as methylthio oxidation congeners (Mulford, 1986). The extent of this metabolism can range from a minor pathway (naphthalene, <5% of the dose; Bakke et al., 1985) to a major pathway (pentachlorothioanisole, 80%; Bakke et al., 1987). Though more data are needed, the current literature indicate that metabolism by prokaryotic

Figure 9.2. Methylthio-turnover (MTT) cycles proposed to explain the near equivalent decreases in 35S specific activities (dpm 35S nmol−1) anddilutions of 13C observed in two faecal metabolites (in boxes) produced during the metabolism of triplelabelled pentachlorothioanisole (13CH335–14C6Cl5). Isotopelabelsare only shown for thedoseandthefaecal metabolites; −SG= glutathione.

260 CONJUGATION REACTIONS IN DRUG METABOLISM

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β-lyases in the gut may be the pathway which produces the most methylthioderived metabolites during mercapturic acid biosynthesis. Possibly an extreme example of the involvement of thiol methyltransferases in the intermediary metabolism of glutathione conjugates is the biodisposition of pentachlorothioanisole (PCTA) in rats (Figure 9.2). PCTA is an environmental pollutant (Quirijns et al., 1979) which is also produced from metabolism of the glutathione conjugates of hexachlorobenzene and pentachloronitrobenzene. After conjugation, the pentachlorothioether of glutathione is degraded to the cysteine conjugate which is acted on by a β-lyase. Methylation yields PCTA. The metabolism of three differently labelled PCTAs in control rats, rats with cannulated bile ducts, and germ-free rats (Bakke et al., 1981, 1987) showed that thiol methyltransferase was more extensively involved in PCTA metabolism than indicated by quantitation of excreted metabolites. The use of ring-labelled PCTA (CH3S-14C6Cl5) invivo established that the bile was the major route for excretion (80%) of the doses and mercapturic acid pathway (MAP) metabolites were the major constituents (Bakke et al., 1981). These biliary metabolites were precursors to the faecal metabolites (CH3S-C6H4SCH3, 44%; CH3S-C6Cl4H, 8%; CH3S-C6Cl5, 5%; and nonextractable, 28%). Quantitation of the faecel metabolites indicated that about half of the dose had added a methylthio group, and because about half the dose was present in the bile as MAP metabolites, the C—S lyase-mediated pathway must have been involved. The administration of methyl-labelled PCTA (14CH3S-C6Cl5) established that about half of the methyl groups had been separated from the ring during metabolism and that these methyl groups were excreted about equally as carbon dioxide (23%) and methanesulfinic acid. Quantitation of the 14C in the individual methyl-labelled faecel metabolites showed about equivalent decreases in the amounts (62–68%) when compared with ring-labelled metabolites. These results indicated turnover of either the methyl group, the methylthio group or both. In either case thiol methyltransferase would be involved. Studies with triple-labelled PCTA (13CH335S-14C6C15) (Bakke et al., 1987) established that methyl turnover was probably not occurring, at least in the metabolism to the extractable metabolites, because the decreases in the specific activities of the 35S were about equivalent to the percent dilution of the 13C for the major faecal metabolites. The methylthio turnover mechanisms shown in Figure 9.2 are proposed to explain these isotope dilutions. This may also be true for the nonextractable fraction where the specific activity also decreased by a similar amount (40%), however, the 13C dilution could not be determined. The methane sulfinic acid in the urine also underwent dilution of the 13C-label. This indicated either methyl turnover, dilution with endogenous methane sulfinic acid or dilution with methane sulfinic acid that resulted from displacement of nonlabelled methylthiol groups that had been introduced into the PCTA during

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Figure 9.3. Metabolism of 2, 6-dichlorobenzonitrile to 2-methylthio-3-hydroxy-6chlorobenzonitrile.

methylthio turnover. These studies showed that by methylthio turnover and/ or multiple conjugations, thiol methyl transfer can be involved more than once in the biodisposition of xenobiotics. Using the triple-labelled compound, it was also possible to detect that metabolism had occurred to some of the parent compound isolated from the faeces. Dilutions of isotopes (Figure 9.2) indicated that about 0·5% of the PCTA had undergone methylthio turnover. This metabolism occurred during enterohepatic circulation because there was no diluton of isotopes in PCTA isolated from faeces from rats with cannulated bile ducts that had been dosed with triple-labelled PCTA. Tissues involved in thiol methylation in vivo In rats the highest concentrations of thiol methyltransferase are in the cecal and colonic mucosae, liver, lungs and kidneys; lower concentrations of enzyme are present in other gastrointestinal mucosae (Weisiger et al., 1980). Liver and kidney contain cysteine conjugate ß-lyase activities which are possible sources of substrates for hepatic and renal thiol methyltransferases; the intestinal contents are juxtaposed to the intestinal mucosa and can supply substrates for mucosal thiol methyltransferases by means of microfloral)8glucuronidase and β-lyases. Demonstration of the participation in vivo of a particular thiol methyltransferase in the metabolism of a xenobiotic has not been reported. Attempts have been made in one of our laboratories (J.B.) to deduce the sites of methylation of thiols formed in the catabolism of GSH conjugates by in situ perfusions. The results of these studies lead to the conclusion that the intestinal mucosa is the major site of thiol methylation; these results are discussed below.

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Figure 9.4. Metabolism of 2-chloro-N-isopropylacetonilide (Propachlor).

Davison (1989) has described methods for in situ infusion of xenobiotic conjugates into the kidneys (perfusion into one kidney, collect urine from both) and hepatic portal veins of rats, pigs and chickens (in all preparations, bile ducts are cannulated to prevent enterohepatic circulation). These preparations work well for the investigation of water-soluble conjugates. The problem is that most of the metabolites must be isolated from biological fluids or from in vitro incubations. This has limited the variety of xenobiotics studied. Nevertheless, it was possible to measure single-pass clearance in the perfused kidney by the difference in excretion of metabolites between the two kidneys. These perfusion methods and other in vivo studies have been applied in an attempt to determine the role of various tissues in the methylation of thiols formed by the β-lyase pathway. The GSH conjugates of PCTA (Figure 9.2), 2, 6dichlorobenzonitrile (DCBN) (Figure 9.3) and propachlor (Figure 9.4) were synthesized or isolated from bile and were used for the perfusions with pigs, chickens and rats. As expected, no methylthio-containing metabolites from propachlor GSH conjugate were excreted after infusion into the renal arteries or portal veins of animals with cannulated bile ducts. Only mercapturic acid was excreted

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because tissue β-lyase does not use the cysteine conjugate of propachlor as a substrate. The GSH conjugates of DCBN and PCTA were excreted as thiols by pig kidneys both on a single pass and after entering systemic circulation. The pig kidney cleared 40% and 70% of the doses, respectively, during a single pass, however, no methylation of the thiols was detected. Chicken kidney also excreted the GSH conjugate of DCBN as the thiol (single pass, 30%, total, 90%) with no methylation of the thiol. The results from these two species indicate that even though thiol methyltransferase is present in chicken and pig kidneys (Larsen, 1985), it did not function with these thiols. Rats excreted only mercapturates in urine (70% of dose) after infusion of DCBN-GSH conjugate into renal arteries. No single-pass excretion was detectable, therefore the doses must have returned to the systemic circulation before excretion. Rat kidney excreted the GSH conjugate of PCTA as the mercapturic acid in urine (25% single pass, 26% total) and bile (62%). Therefore, substrates for thiol methyltransferase were not formed in rat tissues from these GSH conjugates. Infusions of the GSH conjugate of DCBN into the hepatic portal veins of these three species was also negative with respect to the formation of methylthio-containing metabolite. Only in the case of the pig were nonquantifiable traces of the methylthio metabolite detectable in urine by capillary gas chromatography-mass spectrometry. The absence of thiol formations from DCBN and PCTA-GSH conjugates infused into rats is difficult to explain especially because the corresponding thiols are present in good yield (ca. 20%) in bile from rats dosed orally with the parent compounds. Intestinal mucosal thiol methyltransferase is assumed to be involved in the methylation of the DCBN-derived thiol in vivo because control rats excrete the methylthio metabolite in the urine (ca. 15%), whereas rats with cannulated bile ducts excrete none in the urine but excrete the precursors (thiol glucuronide and GSH conjugate) in the bile (Bakke et al., 1989). Because thiol methyltransferase activity has not been detected in the intestinal contents (Weisiger et al. 1980; Larsen, 1985), it was concluded that only thiol methyltransferase in the intestinal mucosa was functioning in DCBN metabolism in rats. Further metabolism of methylthio conjugates in vivo Methylthio-containing metabolites often undergo further metabolism, probably because the metabolites are still very lipophilic. Therefore, in vivo metabolites which arise from methylation are often isolated and include reactions such as oxidation to the methylsulfoxides and methylsulfones. These oxidation congeners of methylthio groups can be displaced by GSH resulting in

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methylthio turnover. The displaced methylthio group can be excreted as methane sulfinic and sulfonic acids, carbon dioxide and sulfate (Bakke et al., 1987). These oxidation congeners can also undergo functionalization and conjugation reactions in other parts of the molecule until they are excreted or excretable in propachlor metabolism (Figure 9.4). Some consequences of thiol methylation are persistent residues in lungs (Bergman et al., 1979), kidneys (Kato et al., 1986), and the environment (Quirijns et al., 1979). These residues are extractable, and some are known to be present in association with specific proteins. Lund et al. (1985) have shown that lung residues of a polychlorinated biphenyl-methylsulfone are protein associated, and the kidney residues of 3, 5- and, 2, 4-dichloro-methylsulfonylbenzene described by (Kato et al., 1986) are also known to be associated with a protein (G.L.Larsen, private communication). The latter methylsulfones were also reported to induce microsomal drug-metabolizing enzymes in rat liver (Kato et al., 1986). 9.6. Inhibition of thiol methyltransferases Inhibitors of thiol methyltransferases have been described. SAdenosylhomocysteine (SAH), the product of methyl transfer from SAM to an acceptor, is an excellent inhibitor of thiol methyltransferases (Borchardt and Cheng, 1978; Loo and Smith, 1985b). Indeed, the slow dissociation of SAH from the enzyme may be responsible for the bizarre kinetic behaviour of the purified protein (Weisiger and Jakoby, 1979). Reagents which react with protein-bound thiols are also effective inhibitors, suggesting a role for a thiol group in enzyme activity (Borchardt and Cheng, 1978; Woodson and Weinshilboum, 1983). A variety of benzoates inhibit thiol methyltransferases with Ki values ranging from the micromolar to millimolar range (Woodson and Weinshilboum, 1983; Ames et al., 1986). Aldehyde analogues of the benzoates are also inhibitors but are not as effective as the benzoates. The catechol Omethyltransferase inhibitor, tropolone, is an inhibitor of both soluble and microsomal-bound thiol methyltransferase in human kidney. In contrast, SKF-525A is effective against red blood cell thiol methyltransferase but was ineffective against the soluble enzyme from human kidney. This difference in inhibitor sensitivity has been suggested to be indicative of separate enzymes (Woodson and Weinshilboum, 1983).

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9.7. Biological function of thiol methyl transfer Thiols are generally considered to be toxic, and the role of thiol methyltransferases as detoxication enzymes has been reviewed (Weisiger and Jakoby, 1980). We have already discussed the toxicological significance of genetic variations in thiol methyltransferase activity (Section 9.4) and the persistence of methylthio metabolites in various tissues (Section 9.5). Recent developments in this detoxication function will be considered here. As was pointed out in the preceding sections, methylthio-containing metabolites are often excreted as their corresponding sulfoxides or sulfones. Thus it may be considered that the coupling of thiol methylation to thioether oxidation facilitates the excretion of these compounds. A likely source of thiol oxidation is the flavin-dependent mixed function oxidase described by Zeigler and colleagues (1984). Whether the oxidation of the thioether can accelerate excretion or prevent further activation is not known. However, its role in recycling of glutathione conjugates by enterohepatic circulation is clear (Bakke et al., 1987). An additional recent development in the study of thiol toxicity is the recognition of the cysteine conjugate β-lyase pathway as a source of toxic thiols. This pathway can play a role in the production of thiols, whose toxicity has been mentioned, and in the formation of reactive electrophiles which are both toxic and mutagenic (Odum and Green, 1985; Anders et al., 1988; van Bladeren, 1988). It seems clear that the free thiol is crucial to the formation of reactive species from products of the β-lyase pathway; thus methylation of these products may constitute detoxication. However, the short half-life of the reactive sulfur species limits methylation (Stevens and Jakoby, unpublished observation). 9.8. Purification and assay procedures for thiol methyltransferases Purification procedures are published for rat (Borchardt and Cheng, 1978; Weisiger and Jakoby, 1979) and human (Woodson and Weinshilboum, 1983) thiol methyltransferases. Weisiger and Jakoby (1979) purified rat liver thiol methyltransferase to homogeneity from the 100 000 x g supernatant from frozen rat livers using 2-thioacetanilide as substrate. The enzyme was highly unstable and lost activity under a variety of conditions. Woodson and Weinshilboum (1983) purified an enzyme from the soluble fraction of human kidney using 6mercaptopurine as substrate. Highly purified human enzyme could be obtained using affinity chromatography on a SAH-ligand column; however, the enzyme was very unstable after this step. Thus, stability appears to be a major problem with purification of the thiol methyltransferases.

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Assay procedures for thiol methyltransferases depend on the availability of [methyl-3H]S-adenosylmethionine (Weisiger and Jakoby, 1980). Procedures have been published for rat (Weisiger and Jakoby, 1979), human (Woodson and Weinshilboum, 1983) and murine enzymes (Otterness et al., 1985). In each case the assay exploits the partitioning of the [3H]-methyl product into organic solvent at basic pH. Product is quantitated by liquid scintillation counting. Most assay procedures are performed near neutrality due to the instability of SAM at alkaline pH. Obviously, this assay procedure is not useful for substrates which are charged at alkaline pH. Dithiothreitol, which is methylated only on one of the thiol groups, will remain charged at alkaline pH and, therefore, extraction is poor. Extraction at acid pH has been used to estimate activity with this substrate. Using this assay modification it has been shown that dithiothreitol is indeed a substrate for rat liver thiol methyltansferase (Donahue et al., personal communcations). We have proposed that cysteine conjugates of xenobiotics may be an important source of thiols in vivo via the β-lyase pathway. In some cases it may be of interest to couple the β-lyase assay to transmethylation from [methyl-3H] SAM. Published procedures for the β-lyase assay are compatible with the methylation conditions (Stevens and Jakoby, 1985). Purification and identification of thiomethylated products from mammalian excretions requires more complex techniques. Several examples can be found in the work of Bakke et al. (1987). An important procedural note is that oxidized thioethers can be produced during the workup, especially in the presence of any peroxides, such as can be found in commercial ether. Therefore, special care must be taken to avoid oxidation during isolation of thioether metabolites. Abbreviations DCBN GSH MAP MTT PCTA SAH SAM SMT TPMT

2, 6-Dichlorobenzonitrile Glutathione Mercapturic acid pathway Methylthio-turnover Pentachloroanisole S-adenosylhomocysteine S-adenosylmethionine S-methyltransferase Thiopurine methyltransferase

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Hiemke, C. and Ghraf, R. (1983), Journal of Neurochemistry, 40, 592–4. Hoffman, D.R., Cornatzer, W.E. and Duerre, J.A. (1979), Canadian Journal of Biochemistry, 57, 56–65. Inoue, M. (1985), in Kinne, R.K.H. (Ed.), Renal Biochemistry: Cells, Membranes, Molecules, pp. 225–69, New York: Elsevier. Jakoby, W.B., Stevens, J., Duffel, M.W. and Weisiger, R.A. (1984), Reviews in Biochemical Toxicology, vol 6, pp. 97–116, Amsterdam: Elsevier. Jakubowski, M. and Gessner, T. (1972), Biochemical Pharmacology, 21, 3073–6. Karim, A., Zagarella J., Hribar, J. and Doolley, M. (1976), Pharmacology and Therapeutics, 19, 158–69. Kato, Y., Kogure, T., Sato, M., Murata, T. and Kimura, R. (1986), Toxicology and Applied Pharmacology, 82, 505–11. Keith, R.A., Abraham, R.T., Pazmino, P. and Weinshilboum, R.M. (1983a), Clinica Chimica Acta, 131, 257–72. Keith, R.A., van Loon, J., Wussow, L.F. and Weinshilboum, R.M. (1983b), Clinical Pharmacology and Therapeutics, 34, (4), 521–8. Keith, R.A., Otterness, D.M., Kerremans, A.L. and Weinshilboum, R.M. (1985), Drug Metabolism and Disposition, 83, 669–76. Kerremans, A.L., Lipsky, J.J., van Loon, J., Gallego, M.O. and Weinshilboum, R.M. (1985), The Journal of Pharmacology and Experimental Therapeutics, 235, 382–8. Kredich, N.M. (1980), Advances in Enzyme Regulation, 18, 181–91. Larsen, G.L. (1985), Xenobiotica, 15, 199–209. Larsen, G.L. and Bakke, J.E. (1981), Xenobiotica, 11, 473–80. Larsen, G.L. and Stevens, J.L. (1986), Molecular Pharmacology, 29, 97–103. Lennard, L. and Maddocks, J.L. (1983), Journal of Pharmacy and Pharmacology, 35, 15–8. Lennard, L., van Loon, J.A., Lilleyman, J S. and Weinshilboum, R.M. (1987), Clinical Pharmacology and Therapeutics, 41, 18–25. Lindsay, R.H., Hulsey, B.S. and Aboul-Enein, H.Y. (1975), Biochemical Pharmacology, 24, 463–8. Loo, G. and Smith, J.T. (1985a), Biochemical and Biophysical Research Communications, 128, 1201–7. Loo, G. and Smith, J.T. (1985b), Biochemical and Biophysical Research Communications, 128, 965–71. Lund, J., Brandt, I., Poellinger, L., Bergman, A., Klaason-Wehler, E. and Gustafsson J.-A. (1985), Molecular Pharmacology, 27, 314–23. Mikov, M., Caldwell, J., Dolphin, C.T. and Smith, R.L. (1988), Biochemical Pharmacology, 37, 1445–9. McBain, J.B. and Menn, J.J. (1969), Biochemical Pharmacology, 18, 2282–5. Mulder, G.J. and Krijgsheld, K.R. (1984), in Roe, D.A. and Campbell, T.C. (Eds.), Drugs and Nutrients, pp. 119–78, New York: Marcel Dekker, Inc. Mulford, D.J. (1986), in Fate of Methylthio-Containing Xenobiotics in Rats, PhD Dissertation, North Dakota State University, Fargo, ND. Odum, J. and Green, T. (1985), Chemico Biological Interactions, 54, 15–31. Otterness, D.M. and Weinshilboum, R.M. (1986), The Journal of Pharmacology and Experimental Therapeutics, 240, 817–24. Otterness, D.M. and Weinshilboum, R.M. (1987), Journal of Pharmacology and Experimental Therapeutics, 240, 817–24.

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Otterness, D.M., Keith, R.A. and Weinshilboum, R.M. (1985), Biochemical Pharmacology, 34, 3823–30. Otterness, D.M., Keith, R A., Kerremans, A.L. and Weinshilboum, R.M. (1986), Drug Metabolism and Disposition, 14, 680–8. Quirijns, J.K., van der Paauw, C.G., Ten Noever de Brauw, M.C. and de Vos, R.H. (1979), Science of the Total Environment, 13, 225–33. Remy, C.N. (1963), Journal of Biological Chemistry, 238, 1078–84. Saari, J.C. and Schultze, M.O. (1965), Archives of Biochemistry and Biophysics, 111, 593–602. Sarctone, E.J. and Sokal, J.E. (1958), Journal of Biological Chemistry, 231, 605–12. Schaeffer, V.H. and Stevens, J.L. (1987), Molecular Pharmacology, 32, 293–8. Shull, K.H., McConomy, J., Vogt, M., Costillo, A. and Farber, E. (1966), Journal of Biological Chemistry, 241, 5060–70. Stevens, J.L. (1985), Journal of Biological Chemistry, 260, 7945–50. Stevens, J.L. and Jakoby, W.B. (1983), Molecular Pharmacology, 23, 761–5. Stevens, J.L. and Jakoby, W.B. (1985), Methods in Enzymology, 113, 510–5. Stevens, J.L. and Jones, D.P. (1989), in Dolphin, D., Poulsen, R. and Avromovic, O. (Eds.), Glutathione: Chemical, Biochemical and Medical Aspects, Vol III B, pp. 45–85, New York: John Wiley and Sons. Stevens, J.L., Hayden, P.H. and Taylor, G. (1986a), Journal of Biological Chemistry, 261, 3325–32. Stevens, J.L., Robbins, J.D. and Byrd, R.A. (1986b), Journal of Biological Chemistry, 261, 15529–37. Stevens, J.L., Ayoubi, N. and Robbins, J.D. (1988), Journal of Biological Chemistry, 263, 3395–401. Suzuki, S. and Tateishi, M. (1981), Drug Metabolism and Disposition, 9, 573–7. Suzuoki-Ziro, K., Murakami, S., Kikuchi, K. and Numata, M. (1967), Journal of Pharmacology and Experimental Therapeutics, 158, 353–64. Tateishi, M. (1983), Drug Metabolism Reviews, 14, 1207–34. Tateishi, M. and Shimizu, H. (1976), Xenobiotica, 6, 431–9. Tateishi, M., Suzuki, S. and Shimizu, H. (1978a), Journal of Biological Chemistry, 253, 8854–9. Tateishi, M., Suzuki, S. and Shimizu, H. (1978b), Biochemical Pharmacology, 27, 809–10. Tegtmeier, F. and Brunner, G. (1983), Enzyme, 30, 185–95. Tisdale, M.J. (1980), Biochemical Pharmacology, 29, 501–8. Tomisawa, H., Suzuki, S., Ichihara, S., Fukazawa, H. and Tateishi, M. (1984), Journal of Biological Chemistry, 259, 2588–93. Tsukuda, K., Yamano, H., Abe, T., and Okada, G. (1980), Biochemical and Biophysical Research Communications, 95, 1160–7. van Bladeren, P.J. (1988), Trends in Pharmacological Sciences, 9, 295–9. van Loon, J.A. and Weinshilboum, R.M. (1982), Biochemical Genetics, 20, 637–58. van Loon, J.A., Weinshilboum, R.M. (1989), Journal of Pharmacology and Experimental Therapeutics, 242, 21–6. Villa-Trevino, S., Shull, K.H. and Farber, E. (1966), Journal of Biological Chemistry, 241, 4670–6. Walker, R.C., Woodson, L.C., and Weinshilboum, R.M. (1981), Biochemical Pharmacology, 30, 115–21. Weinshilboum, R.M. (1980), Trends in Pharmacological Sciences, 378–80.

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 10 Amino acid conjugation Andrew J.Hutt1 and John Caldwell2 1

Department of Pharmacy, Brighton Polytechnic, Brighton, East Sussex BN2 4GJ, UK.

2

Department of Pharmacology and Toxicology, St Mary′s Hospital Medical School, Norfolk Place, London W2 1PG, UK.

10.1.

INTRODUCTION

273

Historical aspects

274

10.2.

AMINO ACIDS UTILIZED FOR CONJUGATION

276

10.3.

STRUCTURE-METABOLISM RELATIONSHIPS

280

10.4.

10.5.

Aliphatic carboxylic acids

281

Aromatic acids

282

Arylacetic acids

284

Aryloxyacetic acid derivatives

286

BIOCHEMICAL MECHANISMS OF AMINO ACID CONJUGATION

287

Acyl-CoA synthetases

287

Acyl-CoA: amino acid N-acyltransferase

288

Tissue location

290

Subcellular distribution

292

CONJUGATION IN VIVO

293

Effect of dose size

293

Induction

295

Age

295

AMINO ACID CONJUGATION 273

10.6.

AMINO ACID CONJUGATION AND BIOLOGICAL ACTIVITY

296

10.7.

FURTHER METABOLISM OF AMINO ACID CONJUGATES

297

10.8.

METHODOLOGICAL NOTES

299

Isolation and characterization of metabolites

299

Examination of enzyme systems

299

ABBREVIATIONS

299

REFERENCES

300 10.1. Introduction

Conjugation with amino acids is an important route in the biotransformation of a variety of xenobiotic carboxylic acids in a number of animal species. These reactions involve the formation of an amide or peptide bond between the carboxyl group of the xenobiotic and the amino group of an endogenous amino acid. Unlike the majority of the conjugation reactions, described elsewhere in this volume, in which the xenobiotic reacts with a ‘high-energy’ endogenous molecule, generally a nucleotide, the xenobiotic acid is activated in the carboxyl moiety to a ‘high-energy’ intermediate prior to the transfer of the acyl group to the endogenous molecule. The major metabolic transformations of xenobiotic carboxylic acids, namely conjugation either with an amino acid or with glucuronic acid, have been known since the last century (Williams, 1959; Conti and Bickel, 1977). However it is only relatively recently that several alternative metabolic fates of this functional group have come to light. It is therefore important to briefly review the metabolic fate of the carboxylic acids as (a) the structure of the acid and particularly the nature of substituents adjacent to the carboxyl group influence the ultimate fate of the compound and (b) the reaction sequence involved in the amino acid conjugation reaction results in the generation of a highly reactive coenzyme A thioester intermediate which may result in the transfer of the acyl moiety to functionalities other than the amine group of an amino acid. The amino acid conjugations are generally accepted to be two-stage processes as shown below (Killenberg and Webster, 1980; Caldwell, 1982) (10.1) (10.2)

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(10.3) The initial step is the activation of the carboxyl group to a reactive Coenzyme A thioester (10.1 and 10.2 above) followed by acyl transfer to an amino acid residue (10.3). Specificity may thus be exerted at the activation and/or acyltransfer steps. The formation of the acyl Coenzyme A thioester is of critical importance in the biochemistry of xenobiotic carboxylic acids and in intermediary metabolism. The active acyl group may become involved with the intermediates of lipid biosynthesis (Caldwell, 1984) and undergo transfer to oxygen with formation of esters to yield hybrid triacylglycerols (Hutson, 1982; Fears, 1985), acylcarnitine derivatives (Vickers et al., 1985; Quistad et al., 1986) or acylated steroids, e.g. with cholesterol (Fears et al., 1982) or bile acids (Quistad et al., 1982); transfer to carbon resulting in chain elongation, i.e. addition of single or multiple two carbon units (Caldwell and Marsh, 1983) possibly followed by incorporation into hybrid triglycerides (Hutson, 1982; Caldwell, 1985) as well as transfer to nitrogen to yield amino acid conjugates. In the case of the 2-arylpropionic acid non-steroidal anti-inflammatory drugs, the acyl-CoA thioesters undergo inversion of configuration of the propionic acid moiety, which in vivo results in the formation of the pharmacologically active S-isomers from the weakly active or inactive R-enantiomers (Hutt and Caldwell, 1983; Caldwell et al. (1988). The pharmacological and toxicological significance of these transformations, many of which are quantitatively minor, have been discussed elsewhere (Fears, 1985; Hutson et al., 1985). However it is of significance that the formation of the acyl-CoA thioesters appears to be the key step for many of these transformations. The structures of the various xenobiotic acids referred to in the text are to be seen in the panels of Figure 10.1. Historical aspects There can be little doubt that, amongst the conjugation reactions, and indeed the reactions of drug metabolism in general, the conjugation of benzoic acid with glycine to yield hippuric acid or benzoylglycine, was the Continued Figure 10.1. Continued first to be discovered (Smith and Williams, 1970; Conti and Bickel, 1977; Caldwell, 1986). Roulle, in 1773, reported what he believed to be benzoic acid in the urine of cows and camels. Proust, in 1801, showed the acid to be similar to, but not identical with, benzoic acid, and Wohler, in 1824, fed a dog with benzoic acid and isolated what he believed to be the unchanged acid from urine. It was not until 1829 when Liebig (1829) isolated a compound from horse urine, which he called hippuric acid (Gk: acid from horse urine), that the compound was shown to contain a benzoyl radical and nitrogen. Whilst this observation was of considerable importance in the

AMINO ACID CONJUGATION 275

Figure 10.1. Structures of the various xenobiotic acids referred to in the text.

history of drug metabolism, the first demonstration of hippuric acid formation from exogenously administered benzoic acid to the horse was reported by Marsh et al. (1981). Ure, in 1841, using benzoic acid as a treatment for gout, isolated hippuric acid from the urine of volunteers dosed with the acid. The first clear proof Figure 10.1. Continued that hippuric acid is a product of benzoic acid metabolism in man is usually attributed to Wohler’s pupil Keller, who in 1842 took benzoic acid and isolated hippuric acid from his urine. The structure of hippuric acid was determined by Dessaignes (1845), who showed that the compound yielded both benzoic acid and glycine on treatment with inorganic acids (Conti and Bickel, 1977). The importance of xenobiotic metabolism, particularly that of the carboxylic acids, to early developments in biochemistry cannot be overstated (Hopkins, 1913). For example, Knoop and Dakin applied the higher homologues of both benzoic and phenylacetic acids for the examination of ß-oxidation of fatty acids (Dakin, 1922). More recently glycine was shown to be a constituent of hippuric acid before it was shown to be present in the bile acid, glycocholic acid (Young, 1977).

276 CONJUGATION REACTIONS IN DRUG METABOLISM

Since these early observations of glycine as a conjugating agent, numerous other examples of the utilization of other amino acids in conjugation reactions have been reported and these are summarized in Table 10.1. 10.2. Amino acids utilized for conjugation The nature of the amino acid utilized for conjugation is highly dependent on both the animal species and the structure of the xenobiotic carboxylic acid under investigation. The most frequently observed amino acid conjugates are those with glycine, which is utilized by the majority of animal species and for the conjugation of a wide variety of carboxylic acids, including aliphatic, aromatic, heteroaromatic and phenylacetic acid derivatives. Exceptions to this generality appear to be restricted either to particular animal species or types of acid. Whilst conjugates of several other amino acids have been reported in in vivo studies (see Tables 10.1–10.3) much of our current knowledge concerning the biochemical aspects of the amino acid conjugations has been determined by an examination of systems using glycine as the acyl acceptor, and relatively little is known of the biochemistry of the other amino acid conjugations.

AMINO ACID CONJUGATION 277

278 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 10.1. Historical aspects of amino acid conjugation.

The first report of an alternative to glycine as an acyl acceptor was that of Jaffe (1877) who found benzoic acid to be conjugated with ornithine in the hen. Ornithine conjugation, unlike the other amino acid conjugations, involves the combination of two xenobiotic acyl residues to the amino acid, i.e. both the amino groups of ornithine undergo acylation; in the case of benzoic acid this yields N2, N5-dibenzoylornithine or ornithuric acid. Conjugation with ornithine has been found to occur in other avian species and in some reptiles (Smith, 1958) and appears to be associated with uricotelic species (Killenberg and Webster, 1980), species which excrete uric acid as a major breakdown product of amino acid metabolism. This process is characteristic of terrestrial species which develop within a shell where they store their excretory products in this insoluble form. Ornithine conjugation is not a general reaction of all avian species as was once thought. Baldwin et al. (1960) examined the fate of benzoic acid in a variety of birds and found that Galliformes (e.g. domestic fowl) and Anseriformes (ducks, geese) yield ornithine conjugates; Columbiformes (e.g. pigeons, doves) excreted hippuric acid, whilst Passeriformes and Psittaciformes (parrots) produced neither amino acid conjugate. Recently an N-acetylornithine conjugate of 3-phenoxybenzoic acid was reported in the chicken (Huckle et al., 1982). Another alternative to glycine conjugation is that with glutamine, first reported for phenylacetic acid in man by Thierfelder and Sherwin (1914). Glutamine conjugation appears to be restricted to arylacetic acids, e.g. phenylacetic acid and related compounds, in mammals. An exception to this is diphenylmethoxyacetic acid, a metabolite of diphenhydramine, which yields a

AMINO ACID CONJUGATION 279

Table 10.2. Amino acid conjugations of restricted occurrence.

glutamine conjugate in the Rhesus monkey (Drach and Howell, 1968; Drach et al., 1970). Glutamine conjugation of phenylacetic acid, and the related compounds 4chlorophenylacetic and indol-3-ylacetic acids, is a reaction which was until recently thought to be restricted to anthropoid apes, Old and New World monkeys and man (James et al., 1972a, 1972b; Bridges et al., 1974). Recently, examples of non-primates forming glutamine conjugates have been reported. Thus phenylacetic and 4-chlorophenylacetic acids yield small quantities of the corresponding glutamine conjugates in the ferret (Hirom et al., 1977; Idle et al., 1978), and 2-naphthylacetic acid undergoes extensive glutamine conjugation in the rat, ferret and rabbit (Emudianughe et al., 1977, 1978). The formation of glutamine conjugates of benzoic acid derivatives in house flies and arachnids has

280 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 10.3. Xenobiotic acids forming dipeptide conjugates.

* Sequence unknown.

also been reported (Smith, 1962; Hitchcock and Smith, 1964; Esaac and Casida, 1968). Whilst not strictly an amino acid, taurine does form peptide bonds with xenobiotic acids. Taurine conjugation was thought to be restricted to arylacetic acids, particularly in carnivorous species, but is now known to be well developed in some aquatic species (James, 1986). In addition some aromatic acids have been shown to undergo this transformation, e.g. 3phenoxybenzoic acid (Hutson and Casida, 1978; Huckle et al., 1981c). The four amino acids cited above represent the most commonly encountered metabolic conjugation options, but there are in addition several examples of other amino acids being involved in conjugation. These appear to be restricted both in terms of their species occurrence and the carboxylic acid utilized (Table 10.2) (Caldwell et al., 1980; Quistad, 1986). In addition certain dipeptide conjugates have also been reported (Table 10.3). The formation of polyglutamyl conjugates of the folic acid antagonist methotrexate has been reported in rats and mice (Baugh et al., 1973; Shin et al., 1974; Israili et al., 1977) and man (Jacobs et al., 1975). The polyglutamates involve the addition of one or two glutamic acid residues to the methotrexate moiety. The importance of this route of metabolism is unknown, but it has been reported that the diglutamate of 4-amino-4deoxypteroic acid is as effective as methotrexate in the inhibition of dihydrofolic reductase (Montgomery et al., 1979). 10.3. Structure-metabolism relationships The metabolic fate of a xenobiotic carboxylic acid is dependent on the size and nature of substituents around/adjacent to the carboxyl group. In addition, the nature of other functionalities within the molecule and their possible metabolic

AMINO ACID CONJUGATION 281

transformations also influences the ultimate fate of the acid. There have been few systematic attempts to examine the structure-metabolism relationships of amino acid conjugation; the majority of the literature stems from an examination of the fate of a particular molecule in a range of animal species. This approach is often handicapped by species variability in the amino acid utilized for the conjugation of a particular carboxylic acid. At a biochemical level the situation is complex since, due to the dual nature of the reaction process, i.e. formation of the activated thioester followed by the acyl-transfer step, specificity or selectivity may be exerted at two stages. For example, whilst substituted phenylacetic acid derivatives are known to undergo amino acid conjugation in a variety of species, substitution at the carbon a to the carboxyl group, e.g. in the 2-arylpropionic acids, was thought at one time to prevent formation of the thioester intermediates, hence explaining their lack of amino acid conjugation in the majority of species (Caldwell, 1978). The only exceptions to this general observation were carnivorous species, which yield taurine conjugates of the 2-arylpropionic acids, e.g. hydratropic (2phenylpropionic) acid and suprofen undergo taurine conjugation in the ferret and dog, respectively (Idle et al., 1978; Sakai et al., 1984). More recent studies indicate that acyl Coenzyme A thioesters are the critical intermediates for the chiral inversion of the 2arylpropionic acids and that the formation of the thioesters is stereo-specific for the pharmacologically inactive R-enantiomers (see Hutt and Caldwell, 1983; Caldwell et al., 1988). Such studies would imply that the acyl transfer to an amino acid is the determining step to explain the lack of amino acid conjugation of these compounds. Furthermore, examples of 2arylpropionic acids undergoing amino acid conjugation in non-carnivores have been reported, e.g. the pyrrole oxidation product of pirprofen has been found to yield a taurine conjugate in the rat and mouse (Egger et al., 1982). The biochemistry of the structure-metabolism relationships of amino acid conjugation are also poorly understood, probably due to the type of methodology used. Ideally the two stages in the reaction sequence should be investigated individually to determine the structural specificity for both activation and acyl transfer. Although such studies have been undertaken in a few cases (see later), in most examples in the literature only the total reaction has been investigated so that the specificity of each step remains essentially unknown. Aliphatic carboxylic acids There are relatively few examples of aliphatic xenobiotic carboxylic acids, or 3aryl substituted acids containing three carbon atoms in a straight chain, undergoing conjugation, presumably due to their facile β-oxidation. Several 3-arylpropionic acid derivatives have been reported to yield amino acid conjugates, e.g. the carboxylic acid metabolites of the antihistamines

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brompheniramine and chlorpheniramine have been reported to yield amino acid conjugates in the urine of both man and dog (Bruce et al., 1968). Similarly glycine conjugates of the unsaturated acid, cinnamic acid, and the related compounds, β-methylcinnamic and 3, 4-dimethyoxycinnamic acids, have been reported (Williams, 1963; Solheim and Scheline, 1976; Hoskins, 1984). Amino acid conjugation of a variety of short-branched chain endogenous aliphatic acids is known to be an important metabolic transformation in several metabolic diseases which result in acidaemia, involving in some instances less commonly utilized amino acids, e.g. sarcosine (Lehnert, 1983). Several studies using purified enzyme systems for the formation of acyl-CoA thioesters and Nacyl transferase activity have indicated that short-chain aliphatic acids may form amino acid conjugates (Mahler et al., 1953; Schachter and Taggart, 1954; Nandi et al., 1979). It is therefore surprizing that relatively few examples of aliphatic xenobiotic acid amino acid conjugates are known. A glycine conjugate of isopropoxyacetic acid has been found in the urine of both the rat and dog following administration of isopropyl oxitol (Hutson and Pickering, 1971), and the glycine conjugate of cyclopropylcarboxylic acid, itself a metabolite of the miticide hexadecylcyclopane carboxylate, has been reported in the urine of rats, cows and dogs (Quistad et al., 1978a, 1978b, 1978c). The glutamic acid conjugate of trans-3-(2, 2-dichlorovinyl)-2, 2dimethylcyclopropane carboxylic acid has been found in the cow, and both geometrical isomers have been reported to yield glycine, serine and glutamic acid conjugates in insects (Unai and Casida, 1977). Of particular interest is the recent observation of four taurine conjugates of the PGE2 analogue trimoprostil in rat bile (Kolis et al., 1986; Figure 10.2). The formation of glycine conjugates of several cyclohexanoic acid derivatives, namely hexahydrohippuric acid and 3, 4, 5, 6-tetrahydrohippuric acid, in addition to hippuric acid, have been reported in perfused rat liver preparations and whole animals following administration of cyclohexanoic acid and shikimic acid (Brewster et al., 1977a, 1977b; 1978). Hexahydrohippurate has also been reported to occur in the urine of cattle, horses and elephants (Balba and Evans, 1977). Aromatic acids Benzoic and heterocyclic aromatic acids are mainly conjugated with glycine in mammalian species. Other amino acid conjugates do occur but their occurrence is restricted in terms of species. Between 1944 and 1955 Bray and co-workers examined the fate of a variety of substituted benzoic acids in the rabbit, and these data (summarized by Williams, 1959) remain the only consistent information on the conjugation of substituted aromatic acids with both glycine and glucuronic acid. Hansch et al. (1968),

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Figure 10.2. Structure of trimoprostil and the taurine conjugates found in rat bile.

using Bray’s data for the fate of para-substituted aromatic acids in the rabbit, found a parabolic relationship between the log percentage dose conjugated with glycine and the lipid solubility of the compound as determined by its log P value (Figure 10.3). The most obvious physicochemical parameter potentially useful in determining conjugation is probably the pKa value of the carboxylic acid. Examination of Bray’s data for the three monochlorobenzoic acid derivatives would indicate that the extent of glycine conjugation also increases with pKa since a change of ca. 1 pKa unit results in a 10- to 12-fold increase in glycine conjugation (Table 10.4). However examination of the corresponding monomethyl analogues, compounds with a narrower range of pKa values, would indicate that the position of substitution is of greater importance (Table 10.4). Closer examination of the data indicates that for ortho-substituted compounds the steric bulk of the substituent, as determined by Taft’s steric parameter Es (Idle, 1976; Caldwell et al., 1980) is the determining factor. The effect of steric bulk on amino acid conjugation is illustrated in Figure 10.4. Originally, Caldwell et al. (1980) interpreted this finding in terms of steric hindrance by the ortho-substituent of the attack of the thiol group of Coenzyme A upon the carbonyl carbon of the carboxyl group. However, it is now clear that many xenobiotic acids which do not give rise to amino acid conjugates are

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Figure 10.3. Relationship between hippuric acid formation in vivo in the rabbit and log partition coefficient for a series of para-substituted benzoic acid derivatives. Where MR=% dose undergoing glycine conjugation. The line of best fit is given by . (Drawn from data presented in Hansch et al., 1968).

converted to CoA thioesters transiently, most notably the 2-arylpropionic acids where the CoA thioesters are the obligatory intermediates in their chiral inversion (see Caldwell, 1984; Caldwell et al., 1988, and references cited above). It thus seems more likely that the marked structural requirement for amino acid conjugation is exerted at the final step of acyl transfer rather than the activation of the xenobiotic acid. Arylacetic acids The effect of chemical structure on the amino acid conjugation of a series of arylacetic acids is presented in Table 10.5. The conjugation of these xenobiotic acids with glycine is a common route for the metabolism of these compounds in the rat, but substitution at the a-carbon atom, i.e. with 2-phenylpropionic and diphenylacetic acids, has a dramatic effect on their fate. These two acids undergo conjugation with glucuronic acid rather than glycine conjugation (Dixon et al., 1977b, 1977c).

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Table 10.4. Effect of pKa and substitution on the conjugation of aromatic carboxylic acids.

Figure 10.4. Relationship between hippuric acid formation in vivo in the rabbit and Taft’s steric parameter (Es) for a series of ortho-substituted benzoic acids.

The conjugation patterns of the regioisomers 1- and 2-naphthylacetic acids are of interest. The major metabolic pathway of the 1-isomer in man, rat and rabbit is the formation of the glucuronide; the cat yields both glycine and taurine conjugates and the glycine conjugation of the compound is relatively low compared to the substituted phenylacetic acids (Dixon et al., 1977a; Table 10.5). The 2-isomer, however, undergoes three amino acid conjugations simultaneously with glycine, taurine and glutamine, as well as glucuronic acid conjugation in the rat, rabbit and ferret (Emudianughe et al., 1978). The total amino acid conjugation is generally much greater than that of the 1-isomer. Also of interest is the reduced recovery of the 2-isomer compared to its 1-isomer in 0–24 h urine samples. This difference in amino acid conjugation may be due

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Table 10.5. Amino acid conjugation of arylacetic acidsa

a adapted from Caldwell et al. (1980), Dixon et al. (1977a, 1977b, 1977c) and Emudianughe et al. (1978). b also undergoes conjugation with taurine (10%) and glutamine (6%) in the rat (see text). c undergoes metabolic chiral inversion in the rat (Fournel and Caldwell, 1986; see text).

to steric hindrance in the case of 1-naphthylacetic acid for the formation of either the Coenzyme A thioester or N-acyltransferase. It would appear in the case of the 2-isomer that the acid undergoes activation readily but that selectivity is exerted in the transferase step. These data suggest that 2naphthylacetic acid may be a useful probe compound for investigating the versatility of these mechanisms. Marsh et al. (1981) have examined the metabolism of 2-naphthylacetic acid in the horse, and whilst the elimination of the acid was low, both glycine and taurine conjugates were found in the urine. 2-Phenylpropionic acid metabolism is also of interest as this compound undergoes chiral inversion in the rat (Fournel and Caldwell, 1986). It appears that the initial step in this transformation involves the formation of a Coenzyme A thioester (see Hutt and Caldwell, 1983), and as an amino acid conjugate is not observed, in this case selectivity seems to occur at the level of the transferase. The effect of structure on glutamine conjugation is also illustrated in Table 10.5. As with glycine conjugation, this reaction appears to be particularly sensitive to substitution adjacent to the carboxyl group in these primate species. Aryloxyacetic acid derivatives Aryloxyacetic acid derivatives, such as the herbicides 2, 4dichlorophenoxyacetic (2, 4-D) and 2, 4, 5-trichlorophenoxyacetic (2, 4, 5-T) acids and the hypolipidaemic agent clofibric acid, are known to undergo taurine conjugation to a minor extent in a variety of species, particularly in carnivorous and marine species (James and Bend, 1976; James, 1982; Emundianughe et al.,

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1983). Recently the related compound 3, 4-dichlorobenzyloxyacetic acid, a potential agent for the treatment of sickle cell anaemia, has been shown to be extensively metabolized (60% of dose) to a taurine conjugate in rats (Peffer et al., 1987), and this compound may well prove to be a useful probe for taurine conjugation in other species. 2, 4-D and 2, 4, 5-T yield relatively low quantities of amino acid conjugates following administration to rodents (Grunow and Bohme, 1974). An explanation for this observation, in terms of the affinity of the corresponding Coenzyme A thioesters, for the glycine N-acyltransferase enzyme, has been proposed by Kelley and Vessey (1986; see later). 10.4. Biochemical mechanisms of amino acid conjugation As mentioned above, the formation of an amino acid conjugate is a two-stage process involving activation of the carboxylic acid followed by acyl transfer to an amino acid. A problem associated with an examination of the available data on the biochemistry of these conjugations is that most workers report data for the overall reaction, i.e. the two-step process, whilst others examine the individual steps, i.e. the formation of the Coenzyme A thioesters and the acyl transfer. These two steps will be examined in turn. Acyl-CoA synthetases The initial reaction, formation of an acyl Coenzyme A thioester, is mediated by an acyl-CoA synthetase or ATP-dependent acid: CoA ligase. These ligases may be divided into three ATP-dependent systems, i.e. short chain or acetyl-CoA synthetase or acetate: CoA ligase (AMP) (EC 6.2.1.1); medium chain or butyryl-CoA synthetase or medium-chain fatty acid: CoA ligase (AMP) (EC 6.2. 1.2); long-chain fatty acyl-CoA synthetase or acyl-CoA synthetase, or longchain fatty acid: CoA ligase (EC 6.2.1.3), and one GTP-dependent, mediumlong chain fatty acid: CoA ligase (GDP) (EC 6.2.1.10) system. Of these the enzyme principally associated with the activation of benzoic and phenylacetic acids appears to be the medium-chain fatty acid: CoA ligase (AMP) (EC 6.2.1. 2), but the involvement of the other synthetases in the metabolism of xenobiotic carboxylic acids should not be discounted. The medium-chain fatty acid: CoA ligase was initially purified by Mahler and coworkers (1953) from beef liver mitochondria and showed broad substrate specificity for straight chain aliphatic acids, from C4 to C12 with optimal activity at C7. Aromatic carboxylic acids, e.g. benzoic and phenylacetic acids, and several branched chain aliphatic acids were also found to be activated. However, neither salicylic or p-amino-salicylic acids were substrates (Schachter

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and Taggart, 1954). Killenberg et al. (1971), using the same tissue preparation, were able to resolve two medium-chain ligases, one of which was unable to activate salicylate and p-amino salicylate (similar to Mahler’s enzyme) whilst the other could. These same workers also noted differences in the stability of the enzymes, the salicylate-activating enzyme being much less stable than Mahler’s enzyme (Killenberg et al., 1971). More recently three soluble synthetases have been isolated from guinea pig liver mitochondria (Groot 1976). These enzymes comprized a medium-chain acyl-CoA synthetase, a ‘salicylate’ enzyme and a propionyl-CoA synthetase. Due to a lack of homogeneous preparations of these enzymes, relatively little is known concerning their properties, e.g. the molecular weight of Mahler’s enzyme has been estimated to be between 30 000 and 60 000 (Mahler et al., 1953). Acyl-CoA: amino acid N-acyltransferase The transfer of the acyl group from a Coenzyme A thioester is catalyzed by an Nacyltransferase. The first example of an enzyme of this type to be partially purified was glycine N-acylase or glycine N-acyltransferase (EC 2.3.1.13) from beef liver mitochondria (Schachter and Taggart, 1954). This enzyme was found to show absolute specificity with respect to the amino acid but to catalyze the acyl transfer of a variety of both aliphatic (C2-C10) and aromatic acyl groups. At about the same time, Moldave and Meister (1957) partially purified a glutamine N-phenylacetyltransferase (EC 2.3.1.14) from the soluble fractions of human liver and kidney tissue obtained at autopsy. The enzyme system was found to catalyze both the phenacylation of glutamine and the benzoylation of glycine, although this latter reaction was carried out at a much slower rate. Moldave and Meister (1957) also found that coenzyme A and ATP were required for the reaction to take place and proposed a mechanism for the overall reaction to be similar to that of glycine conjugation

The purified kidney enzyme had a higher specific activity than that obtained from the liver. Glycine N-acyltransferases have been purified from human (Tishler and Goldman, 1970) and beef (Forman et al., 1971) liver mitochondrial preparations. Both preparations were found to transfer salicyl and benzoyl groups from their corresponding coenzyme A thioesters to yield salicyluric and hippuric acids, respectively.

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More recently Webster et al. (1976) isolated and purified two acyl-CoA: amino acid N-acyltransferases of similar molecular weight (ca. 24 000) from Rhesus monkey and human liver mitochondrial fractions. The enzymes were shown to exhibit acyl-acceptor specificity towards either glycine or glutamine, the ‘glycine’ enzyme showing acyl-donor specificity for benzoyl and salicyl Coenzyme-A, whilst the ‘glutamine’ enzyme utilized either phenylacetyl or indolylacetyl Coenzyme A (Webster et al., 1976). The presence of only one glutamine transferase enzyme was indicated by the virtually constant ratio of phenylacetyl and indolylacetyl transferase activities during the purification of the Rhesus monkey enzyme. These results contrast markedly with those of Moldave and Meister (1957) who found phenylacetylglutamine transferase activity in cytosolic fractions. Their enzyme would not catalyze the formation of indolylacetylglutamine but did form hippurate. These differences are presumably due to developments in methodology, as the results of Webster et al. (1976) are in agreement with the in vivo observations of James et al. (1972a) and Bridges et al. (1974). Webster et al. (1976) also observed that the amino acid N-acyltransferase activity of both enzymes was inhibited by the acyl donors for the other enzyme, i.e. the formation of hippuric acid from benzoyl-CoA and glycine by the glycine N-acyltransferase was inhibited by the presence of both phenylacetyl- and indolylacetyl-CoA, and the formation of phenylacetylglutamine was similarly inhibited by benzoyl-CoA. These studies have been extended to an examination of enzyme systems of beef liver mitochondria. Similar to the above findings, Nandi et al. (1979) isolated two acyl-CoA: amino acid N-acyltransferases showing benzoyl and phenylacetyl-CoA transferase activity. However, glycine was the preferred acyl acceptor for both enzymes, while L-asparagine and glutamine served as very weak acyl acceptors. The molecular weights of the two enzymes were similar, ca. 33 000, results in broad agreement with those of Lau et al. (1977) of 36 000 and Forman et al. (1971) of 32 000, and the acyl donor substrates for one enzyme were inhibitors of the other (Nandi et al., 1979). The differences in the acyl acceptor for the phenylacetyl transferase enzyme isolated from beef liver mitochondria and both Rhesus monkey and human tissue presumably account for the observed differences in the conjugates observed in vivo and may also be of significance from an evolutionary viewpoint. Kelley and Vessey (1986) have recently examined the specificity of the two beef liver mitochondrial transferases with respect to the coenzyme A thioesters of 2, 4-dichlorophenoxyacetic (2, 4-D), 2, 4, 5-trichlorophenoxyacetic (2, 4, 5T) and phenoxyacetic acids using benzoyl- and phenylacetyl-CoA as standards for transferase activity. It was found that whilst phenoxyacetyl-CoA was a substrate for the phenylacetyltransferase, the two halogenated compounds differed in their specificity, the 2, 4-D-CoA being a substrate for both enzymes

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and 2, 4, 5-T-CoA a substrate for the benzoyltransferase. However, the reaction rates were much slower for the herbicide-CoA thioesters. Both enzymes had a high affinity for the coenzyme A thioesters of the herbicides, and the slow reaction rates were found to be due to increased KM values for glycine compared to the normal substrates (Kelley and Vessey, 1986). These results may explain the low yields of amino acid conjugates of both 2, 4-D and 2, 4, 5-T observed in vivo in rats and mice (Grunow and Bohme, 1974). Tissue location Although the foregoing discussion has dealt with hepatic enzymes exclusively, as early as 1870 it was known that the kidney was a major site of synthesis of amino acid conjugates. Schmeideberg and his associates, using elegant surgical preparations well in advance of their time, showed that hippuric acid synthesis in the dog was effected by the kidney rather than the liver, and these findings were confirmed and extended by Quick (1931). More recent studies have confirmed the importance of these two sites, in animals and man, as evidenced by a comparison of the activity of crude tissue homogenates, by isolation of enzyme systems and by in vivo pharmacokinetic studies. Activity has also been reported for other tissues, but this is very low. Strahl and Barr (1971) have shown that rat intestine slices and everted sections of gut are able to synthesize hippuric acid, and Irjala (1972) reported p-aminohippurate synthesis in tissue homogenates of rat and guinea pig duodenum. Rabbit small intestine and lung preparations show extremely low and no glycine Nacyltransferase activity, respectively (James and Bend, 1978a). However both preparations hydrolyzed phenylacetyl-CoA to phenylacetic acid. Similarly human brain, lung, intestine and heart preparations obtained at post-mortem are unable to synthesize hippuric acid from benzoic acid (Caldwell et al., 1976). The relative contribution of the kidney and liver to the formation of amino acid conjugates varies with both species and substrate. Irjala (1972) investigated the formation of the glycine conjugates of p-aminobenzoic, benzoic and salicylic acids using tissue slices, homogenates and mitochondria from the kidney and liver of the rat, guinea pig, cat and dog. Considerable species variation occurred in terms of the extent of conjugation. In addition the relative contribution of each organ to the formation of the hippurates varied considerably. Thus, the conjugation of p-aminobenzoic acid was greater in the kidney tissue slices than the corresponding liver preparations in all four species. Benzoic acid conjugation using kidney slices was greater than in liver of dog and cat but similar in the rat and guinea pig, whilst with salicylic acid the activity of the kidney was greater than the liver in rat, guinea pig and dog. Using homogenates, p-aminobenzoic and benzoic acid conjugation was greater in liver than kidney in rat and guinea pig, but this was reversed in the

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cat and dog. With salicylic acid, only trace activity was found in rat liver homogenates; in the guinea pig, hepatic activity was greater than renal, whilst in the dog, kidney tissue was more active than liver. Using rat liver and kidney mitochondria, the activity of liver preparations was greater than that of the kidney for the conjugation of p-aminobenzoic acid. James and Bend (1978a) have shown that hepatic phenylacetyl-CoA: glycine N-acyltransferase specific activity was lower than that of kidney in both rat and rabbit tissue preparations. Hippuric acid formation has also been shown to occur predominantly in renal tubule cell preparations rather than hepatocytes in both dog and ferret, whilst in the rat and hamster, activity is found in both tissues (Kao et al., 1978). Using human cadaver tissues Caldwell et al. (1976) demonstrated that hippuric acidforming activity was similar in kidney cortex and liver and lower in renal medulla. The activity of both liver and kidney preparations from five species (rat, mouse, hamster, gerbil and ferret) have been examined using 3-phenoxybenzoic acid as a substrate (Huckle et al., 1981b). These workers examined the overall reaction, i.e. the conjugation of 3-phenoxybenzoic acid to yield its glycine conjugate, and the formation of the coenzyme A thioester and the glycine Nacyltransferase activity individually. The formation of the glycine conjugate, as determined by an examination of the overall reaction, indicated that in the ferret and mouse, kidney activity was greater than that in the liver; the activities of both tissues were similar in the gerbil and hamster, and in the rat the liver had greater activity than the kidney. The activation of the acid by formation of the coenzyme A thioester showed a similar species/tissue variation. However, assay of glycine N-acyltransferase activity indicated that in the ferret, gerbil and rat activity in kidney was greater than in the liver, whilst in the mouse and hamster the situation was reversed (Huckle et al., 1981b). It is of interest to note that the acyltransferase was found to be between 10 to 300 times more active than acyl-CoA formation (Huckle et al., 1981c). These results are in broad agreement with those of Forman et al. (1971) who found 1000-fold greater activity of the acyltransferase relative to acyl-CoA formation for the synthesis of salicyluric acid. It seems clear that in this case, the ratelimiting step in the overall conjugation process is the formation of the CoA thioester. Comparison of in vitro data for 3-phenoxybenzoic acid glycine conjugation with the observed in vivo metabolism of the compound indicates that the species with the highest in vitro activity, the gerbil and ferret, excrete the glycine conjugate as the major urinary metabolite (Huckle et al., 1981c), whilst the species (hamster) with the lowest in vitro activity yields the least quantity of the glycine conjugate in vivo. However, rat and mouse both show conjugation activity in vitro but only excrete very small amounts of the glycine conjugate in vivo (Huckle et al., 1981c). The authors accounted for the in vitro/in vivo discrepancy

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in terms of differences in the tissue distribution of metabolic activity, since the major route of metabolism in the rat involves hepatic oxidation. The high renal acyltransferase activity may thus have very little significance for the in vivo disposition of the compound. Riegelman and co-workers have developed a pharmacokinetic method to determine the extent of renal compared to hepatic metabolism in amino acid conjugation in vivo (Wan and Riegelman, 1972a, 1972b; Wan et al., 1972; von Lehmann et al., 1973). Their method is based on the determination of both the apparent clearance rate and the true renal clearance of both the acid and its corresponding glycine conjugate following the intravenous infusion of both compounds. Using this technique the conjugation of benzoic and paminobenzoic acids has been determined in the rabbit and the conjugation of salicylic acid in both the Rhesus monkey and man. In the rabbit, the renal contribution to the total glycine conjugation of both benzoic and paminobenzoic acids was greater than that of the liver, the extent varying between compounds. The conjugation of salicylic acid was found to be totally renal in the Rhesus monkey, while the kidneys account for ca. 68% of its total glycine conjugation in man. The approach taken in these studies has been the subject of some criticism in the literature (Kamath and Levy, 1974). Lowenthal et al. (1974) have investigated the elimination kinetics of salicylate in anephric patients and were unable to show any statistically significant differences in the pharmacokinetic parameters between the patients and normal subjects. In particular, the plasma salicyluric acid concentrations were found to be greater in the anephrics than the normal subjects, presumably due to the absence of renal excretion. However the results obtained do not support the view of von Lehmann et al. (1973) that the kidney makes a significant contribution to the glycine conjugation of salicylic acid in man. Subcellular distribution As indicated above the early studies of Moldave and Meister (1957) indicated that amino acid acylation activity was located in the soluble fractions of liver and kidney preparations. There is now, however, considerable evidence to indicate that the mitochondria are the major subcellular location for the formation of amino acid conjugates, both the activating and the acyl transferase activity having been purified from hepatic and renal mitochondrial preparations (Killenberg et al., 1971; Groot, 1976; Webster et al., 1976; Lau et al., 1977; James and Bend, 1978a; Nandi et al., 1979). The enzyme systems responsible for the synthesis of hippuric acid have been located in the mitochondrial matrix. Gatley and Sherratt (1976) examined the synthesis of hippuric acid using intact and sonicated rat liver mitochondrial

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preparations and found a greater activity in the latter. In addition mitochondria treated with digitonin, to remove the outer membrane and the intermembrane space contents, continued to produce hippurate. Similarly, other techniques which disrupt the inner mitochondrial membrane (sonication, freeze-thawing, lysis with Triton X-100), also resulted in increased benzoyl-CoA synthetase and glycine N-acyltransferase activity (Gatley and Sherratt, 1977). James and Bend (1978a) investigated the glycine N-acyltransferase activity of liver and kidney preparations of both the rat and rabbit. These workers developed a radiochemical technique using 14C-labelled phenylacetyl-CoA and measured the formation of 14C-phenylacetylglycine. Prior to this report the disappearance of the substrate, the acyl coenzyme A thioester, had been determined spectrophotometrically by measuring absorbance changes due to cleavage of the thioester bond. This technique is obviously not ideal as hydrolytic activity is often extensive in crude tissue preparations. An alternative method involves the use of radio-labelled carboxylic acids and the determination of the conjugate formed. However, in this case data for the total reaction only is generated and is of limited value for an examination of the amino acid N-acyltransferase activity. Using labelled phenylacetyl-CoA, James and Bend (1978a) found that glycine N-acyltransferase activity was located in the mitochondrial matrix of tissue derived from the liver and kidney of both species. The activities of both the Nacyltransferase and of glutamate dehydrogenase, a marker enzyme for the mitochondrial matrix, paralleled one another and were highest in Tritonsolubilized preparations. Similarly, Huckle et al. (1981b), using 3phenoxybenzoic acid and 3-phenoxybenzoyl-CoA, have shown increased activity using liver and kidney preparations from a variety of species both for the overall reaction, i.e. synthesis of the glycine conjugate, and for the formation of the coenzyme A thioester, using tissue treated with Triton X-100. Very little is known of the enzymology of taurine conjugation. Taurine transferase activity towards phenylacetyl-CoA also appears to be located in the mitochondrial matrix of rat liver and kidney preparations, the activity of the renal enzyme being greater than that of the liver (James, 1978). 10.5. Conjugation in vivo Effect of dose size The two major metabolic options for xenobiotic carboxylic acids involve conjugation with an amino acid or with glucuronic acid. In vivo, the dose of the xenobiotic has repeatedly been shown to be a determinant of the major pathway.

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At low doses, simple xenobiotic acids tend to undergo amino acid conjugation almost quantitatively, but as the dose increases glucuronic acid conjugation becomes more important. Thus the amino acid conjugation mechanism appears to be a high-affinity, low-capacity system, whereas glucuronic acid conjugation tends to be a high-capacity system with a broader substrate selectivity. The in vivo amino acid conjugation of both benzoic and phenylacetic acids is dose-limited. At high doses of benzoate the formation of hippurate is limited with a compensating increase in the excretion of benzoylglucuronide unless glycine is co-administered. In contrast, the formation of salicyluric acid appears to be dose-limited irrespective of the dose of either salicylic acid or glycine administered. It would appear, therefore, that glycine conjugation of benzoic acid is restricted by the availability of the amino acid, whilst conjugation of salicylic acid is limited by the activation of the acid to the thioester. Supporting data for the limited conjugation of benzoic acid being due to the restricted availability of glycine, arises from the higher apparent KM values for Nacyltransferase activity for the amino acid acceptor substrates than for the acylCoA donor substrates. It is also of interest to note that mitochondria also contain a benzoyl-CoA hydrolase with a high KM which is thought to protect against free coenzyme A depletion if insufficient amino acid acceptor is available for conjugation (Gatley and Sharratt, 1977). The effect of dose size on the in vivo conjugation of a xenobiotic is exemplified by 1-naphthylacetic acid in the rat (Dixon et al., 1977a). At low doses the acid undergoes conjugation with both glycine and glucuronic acid, the majority of the material being excreted in urine as the glycine conjugate, whilst the majority of material excreted via the bile is the glucuronide. As the dose of acid is increased, the percentage of compound eliminated in the urine decreases, as does the proportion undergoing glycine conjugation, with a corresponding increase in glucuronic acid conjugation. The biliary excretion of the compound increases and the major metabolite is the glucuronide at all doses (Dixon et al., 1977a). Thus the saturation of the amino acid conjugation pathway results in a change in the pattern of metabolism and in the route of elimination. Dose may also influence the nature of the amino acid conjugate formed. For example phenylacetic acid yields both glycine and taurine conjugates in the ferret (Hirom et al., 1977). At low doses the glycine: taurine ratio is ca. 9:1 whereas at higher doses the ratio becomes 1:1, the total proportion of the material eliminated as these two conjugates remaining essentially constant. In addition small quantities of phenylacetylglutamine have also been detected (Hirom et al., 1977). The pharmacokinetic implications of capacity-limited amino acid conjugation in man are readily apparent in the metabolism of salicylate. The glycine conjugation of salicylic acid to salicyluric acid is readily saturable at therapeutic dose levels (Levy, 1965a, 1965b). At higher doses the formation of the phenolic

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glucuronide is also saturable (Levy et al., 1972). The limited capacity of these two pathways results in dose-dependent non-linear pharmacokinetics in man. It is only relatively recently that our understanding of the pharmacokinetics of salicylate has been able to explain the observed urinary disposition of these compounds and make a contribution to rational dosage regimes (Levy and Giacomini, 1978). Induction There are several reports of attempts to induce amino acid conjugation. Irjala (1972) found no significant differences in the formation of p-aminohippurate by rat liver or kidney homogenates from animals pretreated for six days with either phenobarbital or p-aminobenzoic acid. These results are in agreement with those of Brandt (1964) who fed newborn rats for 30 days on a diet containing 1% paminobenzoic acid. James and Bend (1978b) reported that pretreatment with phenobarbital increased phenylacetyl-CoA: glycine N-acyltransferase activity of rat kidney but not liver preparations. Pretreatment of animals with salicylic acid, however, yields somewhat different results. Irjala (1972) found that pretreatment of rats with salicylic acid significantly increased the formation of p-aminohippurate in both liver and kidney preparations, whilst salicyluric acid formation was increased in the kidney. That such induction may be species dependent was demonstrated by James and Bend (1978b) who found no increase in glycine N-acyltransferase activity in either liver or kidney preparations following the oral administration of salicylic acid to young rabbits. There occurred a ca. 1.5-fold increase in activity in both renal and hepatic preparations from adult rats. Pretreatment with benzoic acid had little or no effect on glycine N-acyltransferase activity in either renal or hepatic tissue (James and Bend, 1978b). There is also evidence that salicylate may include its own metabolism in man. Furst et al. (1977) demonstrated that pretreatment with aspirin for three days resulted in an increase in salicyluric acid formation rate in 26 volunteers, and Day et al. (1983) have shown an increased rate of salicyluric acid synthesis, primarily in the first two weeks of treatment, in rheumatic patients. It has been postulated that autoinduction may help explain why some patients fail to maintain adequate serum salicylate concentrations during therapy (Day et al., 1983) . Age The ontogeny of the amino acid conjugation mechanism also seems to be species dependent. Glycine N-acyltransferase activity in liver and kidney preparations from foetal and newborn rabbits is low in comparison with adult

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levels, whereas in the rat adult activity is present at birth (James and Bend, 1978b). The ontogenesis of enzyme activity in the rabbit was found to be tissue dependent, hepatic activity reaching adult levels within four weeks of birth whilst renal activity takes four months (James and Bend, 1978b). Similarly paminohippurate synthesis is low or absent in foetal or neonatal liver tissue of rats, mice and rabbits (Brandt, 1964; Gorodisher et al., 1971; Irjala, 1972). Glycine conjugation of benzoic, p-aminobenzoic and salicylic acids has been demonstrated in human foetal liver, kidney and intestine tissue within the tenth week of gestation with activity comparable to that of adult tissue (Irjala, 1972). Similarly Caldwell et al. (1976) have shown adult levels of hippuric acid synthesis in human cadaver liver and kidney tissue from an infant born at 31 weeks gestation. In old age there is an apparent decrease in glycine conjugation, the quantity of hippuric acid excreted following the administration of benzoic acid being reduced in geriatrics compared to young adults (Stern et al., 1946; Binet et al., 1950), possibly due to the reduced availability of glycine with age since administration of the amino acid restores conjugation activity. 10.6. Amino acid conjugation and biological activity The function of the conjugation reactions is generally thought of as a detoxication process, the increased aqueous solubility of the conjugate compared to that of the parent xenobiotic facilitating its more rapid elimination. This view is obviously outdated with the realization of the enhanced reactivity and toxicity of various conjugates arising from sulfation, glucuronidation, acetylation and glutathione conjugation. In contrast, very little is known of the toxicity of amino acid conjugates. This is important for two reasons: (1) numerous drugs and other foreign chemicals undergo this metabolic transformation and (2) man may be exposed to low levels of these conjugates in foods. Plants, for example, conjugate xenobiotic acids with a variety of amino acids to yield conjugates which are not normally found in animals, e.g. 2, 4-dichlorophenoxyacetic acid yields valine, leucine, phenylalanine and aspartic acid conjugates (Climie and Hutson, 1979). Table 10.6 presents data for the acute toxicity of several acids and their conjugates in both rats and mice. Whilst the data are somewhat limited (there being little published information), they are of interest. If the data are compared on a purely mass basis, then the acute toxicity of the conjugates would appear to be either lower than, or equal to, that of the parent acid. If, however, the data are compared on a molar basis, then the differences in toxicity are much less marked and, in the case of 2, 4-dichlorophenoxyacetic acid, the conjugates appear to be slightly more toxic than the parent acid.

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Caldwell et al. (1980) have proposed that the formation of amino acid conjugates is associated with the inactivation of reactive acyl-CoA thioesters so that an examination of acute toxicity data of the type presented may not provide insight into the functional significance of these reactions. Hippuric acid has recently been shown to modulate the activity of γ-glutamyl transpeptidase in vivo and to be formed endogenously from phenylalanine (Thompson and Meister, 1980). Hence as γ-glutamyl transpeptidase has a detoxication function, hippuric acid and structurally related compounds may have considerable biochemical significance. 10.7. Further metabolism of amino acid conjugates The further metabolism of amino acid conjugates is also of interest, particularly with reference to the products of plant metabolism. The metabolism of both the valine and aspartic acid conjugates of 2,4dichlorophenoxyacetic acid (2, 4-D) have been investigated in the rat (Buly and Mumma, 1984). It was found that the compound-related material was extensively excreted in the bile in both cases and the major products of excretion were the parent conjugate and 2, 4-D. In the case of the valine derivative, six other minor products were also detected but not characterized. The most extensively investigated conjugate in terms of its metabolism is probably salicyluric acid, the major metabolite of salicylate in man (Hutt et al., 1986). Shibasaki et al. (1985) have noted the hydrolysis of salicyluric acid to salicylic acid by rabbit intestinal microorganisms, but this does not appear to occur in man, the urinary recovery of both intravenous and oral doses being almost quantitative (Levy et al., 1969; Bochner et al., 1981). Bekersky et al. (1980a, 1980b) have shown, using a perfused rat kidney preparation, that hydrolysis of salicyluric acid to salicylic acid and interconversion of the two forms occurs. Interestingly, the urinary excretion of salicylic acid formed by hydrolysis is more rapid than that of the administered compound (Bekersky et al., 1980b). Similarly, the hydrolysis of hippuric and paminohippuric acids has been reported to occur in the kidney of the dog and rat, respectively (Quick, 1932; Malyusz et al., 1972). Salicyluric acid has also been reported to undergo conjugation with glucuronic acid, at the free phenolic hydroxyl group, to yield salicyluric acid phenolic glucuronide. This compound has been identified as an endogenous constituent of the body fluids of a uremic patient (Zimmerman et al., 1981) and as a urinary metabolite of salicylate following the administration of aspirin to man (Hutt et al., 1982, 1986).

Data from (1)Williams (1959) and (2)James et al. (1972b), LD50 in mice, (3)Mumma (1983) unpublished, cited in Buly and Mumma (1984), LD50, in rats.

Table 10.6. Acute oral toxicity of amino acid conjugates.

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10.8. Methodological notes Isolation and characterization of metabolites Following the administration of xenobiotic carboxylic acid to either an animal or man, the compound may undergo a number of transformations giving rise to a variety of conjugates. Thus biological fluids may contain one or more amino acid conjugates, together with a glucuronic acid conjugate, the free acid and products which may arise from Phase 1 metabolic transformations. The majority of conjugates are highly polar, water-soluble compounds of low volatility, properties which frequently render their isolation and characterization difficult. We have previously examined the methodology available for the characterization of such compounds, detailing the chromatographic and spectroscopic techniques which may be applicable and would refer the interested reader to this review and the references therein (Caldwell and Hutt, 1986). Examination of enzyme systems Detailed accounts for the purification of the benzoyl- and phenylacetyl-CoA: amino acid N-acyltransferases have been presented by Webster in the Methods in Enzymology series (Webster, 1981), in addition to several papers in the same area (Webster et al., 1976; Nandi et al., 1979). As pointed out previously the amino acid conjugation reaction is frequently examined as an entire process rather than as two individual steps. An examination of the N-acyltransferase obviously requires the synthesis and purification of the required CoA thioesters. Methods for the synthesis and purification of the thioesters of benzoic, o-, m- and p-hydroxybenzoic and phenylacetic acids have been published (Webster and Killenberg, 1981) and, in addition, the physical properties, stability and characterization of these compounds have been reported (Mieyal et al., 1974; Webster et al., 1974; Webster and Killenberg, 1981). Abbreviations CoA 2, 4-D 2, 4, 5-T PGE2

Coenzyme A 2, 4-dichlorophenoxyacetic acid 2, 4, 5-trichlorophenoxyacetic acid Prostaglandin E2

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Quistad, G.B., Staiger, L.E. and Schooley, D.A. (1982), Nature, 296, 462–4. Quistad, G.B., Staiger, L.E. and Schooley, D.A. (1986), Drug Metabolism and Disposition, 14, 521–5. Reif, V.D. and Sinsheimer, J.E. (1975), Drug Metabolism and Disposition, 3, 15–25. Rothstein, M. and Greenberg, D.M. (1957), Archives of Biochemistry and Biophysics, 68, 206–14. Sakai, Y., Mori, Y., Toyoshi, K., Horie, M. and Baba, S. (1984), Drug Metabolism and Disposition, 12, 795–7. Sallustio, B.C., Meffin, P.J. and Knights, K.M. (1988), Biochemical Pharmacology, 37, 1919–23. Schachter, D. and Taggart, J.V. (1954), Journal of Biological Chemistry, 208, 263–75. Shibasaki, J., Inoue, Y., Kadosaki, K., Sasaki, H. and Nakamura, J. (1985), Journal of Pharmacobio-Dynamics, 8, 989–95. Shin, Y.S., Buehring, K.U. and Stokstad, E.L.R. (1974), Journal of Biological Chemistry, 249, 5772–7. Smith, J.N. (1958), Biochemical Journal, 69, 509–16. Smith, J.N. (1962), Nature, 195, 399–400. Smith, R.L. and Williams, R.T. (1970), in Fishman, W.H. (Ed.), Metabolic Conjugation and Metabolic Hydrolysis, Vol. 1, pp. 1–19, London: Academic Press. Solheim, E. and Scheline, R.R. (1976), Xenobiotica, 6, 137–50. Stern, K., Tyhurst, J.S. and Askonas, B.A. (1946), American Journal of the Medical Sciences, 212, 302–5. Strahl, N.R. and Barr, W.H. (1971), Journal of Pharmaceutical Sciences, 60, 278–81. Thierfelder, H. and Sherwin, C.P. (1914), Berichte des Deutschen Chemischen Gesellschaft, 47, 2630–4. Thompson, G.A. and Meister, A. (1980), Journal of Biological Chemistry, 255, 2109–13. Tishler, S.L. and Goldman, P. (1970), Biochemical Pharmacology, 19, 143–50. Unai, T. and Casida, J.E. (1977), in Elliot, M. (Ed.), Synthetic Pyrethroids, ACS Symposium Series No. 42, pp. 194–200. Vickers, S., Duncan, C.A. H., White, S.D., Ramjit, H.G., Smith, J.L., Walker, R.W., Flynn, H. and Arison, B.H. (1985), Xenobiotica, 15, 453–8. von Lehmann, B., Wan, S.H., Riegelman, S. and Becker, C. (1973), Journal of Pharmaceutical Sciences, 62, 1483–6. Wallcave, L., Bronczyk, S. and Gingell, R. (1974), Journal of Agricultural and Food Chemistry, 22, 904–8. Wan, S.H. and Riegelman, S. (1972a), Journal of Pharmaceutical Sciences, 61, 1278–84. Wan, S.H. and Riegelman, S. (1972b), Journal of Pharmaceutical Sciences, 61, 1284–7. Wan, S.H., von Lehmann, B. and Riegelman, S. (1972), Journal of Pharmaceutical Sciences, 61, 1288–92. Webster, L.T. (1981), in Jakoby, W.B. (Ed.), Detoxication and Drug Metabolism, Conjugation and Related Systems, Methods in Enzymology, Vol. 77, pp. 301–8, New York: Academic Press. Webster, L.T. and Killenberg, P.G. (1981), in Jakoby, W.B. (Ed.), Detoxication and Drug Metabolism, Conjugation Related Systems, Methods in Enzymology, Vol. 77, pp. 430–6, New York: Academic Press. Webster, L.T., Mieyal, J.J. and Siddiqui, U.A. (1974), Journal of Biological Chemistry, 249, 2641–5.

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Webster, L.T., Siddiqui, U.A., Lucas, S.V., Strong, J.M. and Mieyal, J.J. (1976), Journal of Biological Chemistry, 251, 3352–8. Williams, R.T. (1959), Detoxication Mechanisms, pp. 348–367, London: Chapman and Hall. Williams, R.T. (1963), in Bernfeld, P. (Ed.), Biogenesis of Natural Compounds, pp. 427–74, Oxford: Pergamon Press. Williams, K., Day, R., Knihinicki, R. and Duffield, A. (1986), Biochemical Pharmacology, 35, 3403–5. Young, L. (1977), in Parke, D.V. and Smith, R.L. (Eds.), Drug Metabolism from Microbe to Man, pp. 1–11, London: Taylor and Francis. Zimmerman, L., Jornvall, H., Bergstrom, J., Furst, P. and Sjovall, J. (1981), FEBS Letters, 129, 237–40.

Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 11 Glutathione conjugation Brian Ketterer1 and Gerard J.Mulder2 1

Department of Biochemistry, University College and Middlesex School of Medicine, London W1P 6DB, UK.

2

Division of Toxicology, Center for Bio-Pharmaceutical Sciences, University of Leiden, 2300 RA Leiden, The Netherlands.

11.1.

INTRODUCTION

308

11.2.

GLUTATHIONE SYNTHESIS AND AVAILABILITY

309

Biosynthesis, degradation and distribution

309

Maintenance of cellular GSH

311

Depletion of cellular GSH

312

Interorgan transport of GSH

313

Biliary excretion

315

11.3.

GLUTATHIONE CONJUGATION

317

11.4.

NOMENCLATURE OF THE GSH TRANSFERASES: A SUPERGENE FAMILY

318

11.5.

GSH Transferases in the rat

318

GSH Transferases in man

319

GSH Transferases in the mouse

321

CONJUGATION REACTIONS CATALYZED BY GSH TRANSFERASES

322

Nucleophilic displacement from saturated carbon

322

Nucleophilic displacement from aromatic carbon

324

Reactions of GSH by Michael addition

325

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11.6.

11.7.

Reactions of GSH with xenobiotics containing strained oxirane rings

325

Other GSH transferase-catalyzed reactions

330

ENZYMOLOGY OF GLUTATHIONE TRANSFERASES

331

Catalytic constants, catalytic specificity and rate enhancement

331

The kinetic mechanism and the active site

332

Inhibition

333

DISTRIBUTION OF GSH TRANSFERASES IN TISSUES AND CELLS

334

Tissue distribution

334

GSH Transferases during ageing

336

Induction

336

GSH Transferase gene expression during carcinogenesis

338

11.8.

MOLECULAR BIOLOGY OF CYTOSOLIC GSH TRANSFERASES

339

11.9.

MEMBRANE-BOUND GSH TRANSFERASES

340

11.10.

THE METABOLISM OF GSH CONJUGATES

341

11.11.

CONJUGATION IN ISOLATED CELLS AND PERFUSED ORGANS

341

GSH transferase content of isolated cells

341

Substrate utilization by isolated hepatocytes and the perfused liver

342

Other organs

344

11.12.

GSH CONJUGATION IN VIVO

345

11.13.

INHIBITION OF CONJUGATION IN VIVO

346

11.14.

CONJUGATION AND BIOLOGICAL ACTIVITY.

347

11.15.

Leukotriene biosynthesis

347

Conjugation leading to toxicity.

348

USEFUL TECHNIQUES

349

308 CONJUGATION REACTIONS IN DRUG METABOLISM

11.16.

CONCLUSIONS

350

ACKNOWLEDGEMENTS

350

ABBREVIATIONS

351

REFERENCES

351 11.1. Introduction

Glutathione conjugation is the formation of a thioether link between glutathione and a compound with an electrophilic centre. This process is usually associated with detoxication and excretion but is also involved in the biosynthesis of endogenous compounds, for instance the chemical mediator, leukotriene C4. Work in this field began in 1879 when a urinary excretory product of bromobenzene was isolated and shown to be (N-acetyl-cystein-S-yl) 4bromobenzene (Baumann and Preusse, 1879; Jaffé, 1879). Although since then mercapturic acids of other foreign compounds have also been isolated and characterized, their origin as metabolites of glutathione (GSH) conjugates was not established until 1959 when a correlation was observed between GSH depletion and mercapturate excretion, and enzymes for the conversion of GSH conjugates to mercapturates were reported (Barnes et al., 1959; Bray et al., 1959). Two years later enzymic catalysis of GSH conjugation by liver extracts was described by Booth et al. (1961). This activity was eventually shown to be due to a number of enzymes with overlapping substrate specificities (Habig et al., 1974). GSH was not fully characterized until 1934 when analysis indicated that it was γ-glutamyl cysteinyl glycine (Hopkins, 1929), a structure which was proved by chemical synthesis (Harington and Mead, 1935). The compounds with which GSH reacts to give conjugates may be electrophiles per se (e.g. certain halogenated hydrocarbons and alkylating anticancer drugs), but they usually arise during metabolism, mixed-function oxygenases often being an essential step. However certain endogenous electrophiles, such as cholesterol epoxide and hydroxyalkenals, arise during the course of lipid peroxidation, while leukotriene A4, a precursor in the biosynthesis of the highly potent chemical mediators leukotrienes C4, D4 and E4, is formed by a specific enzyme, namely 5lipoxygenase. Once synthesized GSH conjugates are usually either excreted via the bile or converted to mercapturates and excreted in the urine. Occasionally mercapturates are further metabolized to toxic methyl thio compounds. A

GLUTATHIONE CONJUGATION 309

recent monograph on many aspects of GSH conjugation is that edited by Sies and Ketterer (1988). 11.2. Glutathione synthesis and availability Biosynthesis, degradation and distribution GSH is synthesized from the amino acids L-glutamic acid, L-cysteine and glycine in a two-step reaction catalyzed successively by γ-glutamyl cysteine synthetase (EC 6.3.2.2) and glutathione synthetase (EC 6.3.2.3), each step consuming a mole of ATP (reactions 11.1 and 11.2). (11.1) (11.2) Its degradation is initiated by γ-glutamyl transpeptidase (Meister et al., 1981). This enzyme can catalyze two types of reaction, namely hydrolysis and transpeptidation (see reactions 11.3 and 11.4). (11.3) (11.4) Breakdown is completed by the action of dipeptidases (reaction 11.5), either membrane-bound (Kozak and Tate, 1982) or cytosolic (Das and Radhakrishnan, 1973). (11.5) Reaction (11.1) is inhibited both by GSH and a selective inhibitor, buthionine sulfoximine (L-BSO; S-n-butyl-L-homocysteine sulfoximine; Griffith and Meister, 1979; Griffith, 1982). BSO is phosphorylated by γ-glutamylcysteine synthetase to give a product which apparently irreversibly inhibits the enzyme (Griffith, 1982). There are several inhibitors of reactions 11.3 and 11.4 including L-serineborate (Tate and Meister, 1978), a transition state analogue, and AT 125 or acivicin (L-(αS, 5S)-α-amino-5(3-chloro-4, 5-dihydro-isox azoyl acetic acid) (Meister et al., 1981). GSH is consumed by conjugation reactions and the reduction of H2O2 and organic peroxides. These reactions depend on the nucleophilic thiol of its cysteinyl residue. Free cysteine cannot replace GSH because of its ease of oxidation in the presence of transition element ions with the formation of free radicals. The much lower susceptibility of GSH to autoxidation and its consequent lack of toxicity is apparently due to intramolecular interactions of the functional groups of the glutamyl residue with the cysteinyl thiol (Misra, 1974; Stark et al., 1987).

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GSH is involved in various metabolic pathways. Thus it is essential in the protection of cells against damage, especially oxidative damage induced by various agents and diseases (Reed and Fariss, 1984). For instance, depletion of GSH in glandular gastric mucosa in the rat by diethyl maleate (DEM) results in profuse ulceration (Boyd et al., 1981). Several genetic disorders in the metabolism of GSH, leading to a lower availability of GSH in man have been described (Meister, 1983). Such patients are at an increased risk to the toxicity of certain drugs, such as paracetamol (Spielberg and Gordon, 1981). Because of its important role in homeostasis, GSH is present in most cells, its intracellular concentration varying with the cell type. It is rich in the liver having a value of approximately 5 µmol GSH per gram of tissue in both rat and man (Shi et al., 1982). This high level is presumably required for the specialized role of the liver in the detoxication and excretion of xenobiotics. Human erythrocytes contain 3 µmol GSH per ml of packed cells which, although available for the conjugation of drugs, probably is present primarily to prevent oxidative damage consequent upon the oxygen-carrying function of these cells (Griffith, 1981); GSH deficiency shortens their life span (Tucker et al., 1981). In many other cells and tissues the levels of GSH are 1–2 µmol/gram tissue. Within a tissue, the GSH may not be homogeneously distributed. For example, in brain, GSH is high in astroglial and ependymal cells but absent from the neuronal stroma (Slivka et al., 1987). In the kidney it is high in the cortex and low in the inner medulla (Mohandas et al., 1984). Within the liver hepatocyte population there appears to be a higher GSH synthesis in Zone 1 (periportal) than in Zone 3 (perivenous) cells (Kera et al., 1988). GSH has been shown to be present in the cytosol, mitochondria and nucleus. Administration of diethyl maleate (DEM), leads to a depletion of GSH which never exceeds 90–95% of liver GSH because it has no effect on the small GSH pool in the mitochondria (Wahlländer et al., 1979; Meredith and Reed, 1982; Ecobichon, 1984; Gaetjens et al., 1984; Schnellman et al., 1988). Ethacrynic acid (Meredith and Reed, 1983) and phorone (2, 6-dimethyl-2, 5-heptadien-4one; Romero and Sies 1984), which are able to penetrate the mitochondria, also deplete this GSH pool. The nucleus, when isolated by techniques designed to minimize the loss of small molecular components, appears to have a GSH content similar to that of the cytoplasm (Tirmenstein and Reed, 1988). In vitro studies predict that as much as 20–30% of rat liver GSH is bound to high-affinity sites on cytosolic proteins (Sugiyama and Kaplowitz, 1984; Higashi et al., 1985). A similar level of binding to macromolecules may also occur in other organelles, such as the nucleus (Tirmenstein and Reed, 1988). GSH and oxidized GSH (GSSG) levels in the liver and other organs undergo diurnal variation (Jaeschke and Wendel, 1985; White et al., 1987). GSH content is also age dependent: in human lymphocytes and erythrocytes, GSH

GLUTATHIONE CONJUGATION 311

levels are highest in the age group 25–40 years, while in mice GSH levels fall away at the age of two years (Hazelton and Lang, 1980; Stohs et al., 1980). Maintenance of cellular GSH Studies of the effect of BSO on a number of cell types in culture show that they have the capacity to synthesize GSH; however the major site of the de novo synthesis of GSH is in the liver. Even in human foetal liver the capacity to synthesize GSH is already high (Rollins et al., 1981). The upper limit of cellular GSH concentration is controlled by feedback inhibition of γ-glutamylcysteine synthetase by GSH (Meister, 1984). Administration of γ-GluCys will overcome this inhibition, since it by-passes the regulatory step. Several inducers of drug metabolism increase hepatic GSH: by 30% for phenobarbital (Kaplowitz et al., 1980) and 140% for butylated hydroxyanisole (Jaeschke and Wendel, 1985b, 1986). Similarly, some metal compounds, such as CoCl2, ZnCl2 or As2O3, may increase GSH in liver (or gut mucosa) (Sasame and Boyd, 1978; Pisciotto and Graziano, 1980; Wong and Klaassen, 1981). However, these compounds have many other effects besides that on GSH. SH-containing agents do not affect the ‘total glutathione’ (GSH+GSSG) concentrations in various organs unless they are precursors of GSH (Jaeschke and Wendel, 1985b). Essential for GSH synthesis is a sufficient supply of L-cysteine. L-Methionine can be converted to L-cysteine through the cystathionine pathway in certain tissues, most notably the liver, and thereby meet demands for cysteine (see e.g. Tateishi et al., 1981). Thus, in isolated hepatocytes, both L-methionine and Lcysteine are precursors for GSH. Propargylglycine, an inhibitor of the cystathionine pathway, inhibits the incorporation of methionine into GSH (Beatty and Reed, 1980, 1981). In cells such as isolated rat kidney cells (Moldeus et al., 1981), a murine lymphoma cell line (Brodie et al., 1981), Clara cells, Type 2 pneumocytes and macrophages isolated from rabbit lung (Horton et al., 1987), the cystathionine pathway is absent and methionine is not a precursor of GSH. In rats on a diet in which protein was completely replaced by pure amino acids, variation of the cysteine or methionine content showed that a minimum amount of L-cysteine+L-methionine was required for normal hepatic GSH levels (Glazenburg et al., 1983). This confirmed earlier data by Tateishi et al. (1977) who used less-defined diets low in cysteine-containing protein. They concluded that part of the GSH in rat liver (30–50%) is metabolically labile and released into the blood to serve as a precursor of cysteine in extrahepatic organs where it is released on demand by γ-glutamyltranspeptidase. L-2-Oxo-thiazolidine-4-carboxylate, a cysteine precursor that is more stable than L-cysteine itself, can be used to increase GSH levels in the liver (Williamson et al., 1982; Glazenburg et al., 1984). So too, can ethyl or methyl

312 CONJUGATION REACTIONS IN DRUG METABOLISM

esters of GSH: whereas GSH cannot enter hepatocytes or the liver unchanged, its esters are readily taken up and release GSH as a result of hydrolysis by intracellular esterases (Puri and Meister, 1983; Wellner et al., 1984; Anderson et al., 1985). The isolated perfused rat liver and lung do not take up unchanged GSH (Hahn et al., 1978; Berggren et al., 1984; Joshi et al., 1986). However, GSH entrapped in liposomes is taken up by the spleen (Wendel and Jaeschke, 1982) and also leads to increased GSH in the liver, but the biosynthetic pathway is still required, since BSO prevents this increase. Certain tissues can take up unchanged GSH. Examples are everted intestinal sacs (and intestinal brush border vesicles) and the human buccal mucosa (Linder et al., 1984; Hunjan and Evered, 1985). The epithelial transport of GSH is most likely carrier-mediated, sodium-dependent and can be inhibited by various peptides (Hagen and Jones, 1987). Although a high oral dose of GSH was reported not to affect the plasma concentration of GSH in fasted rats (Yoshimura et al., 1982), recent work strongly suggested that unchanged GSH is absorbed after oral dosage in the rat in vivo (Hagen and Jones, 1987). Equivalent doses of L-cysteine or cystine did not lead to increased plasma GSH suggesting that GSH itself was absorbed unchanged. An experiment with BSO to inhibit resynthesis of GSH would confirm or refute this, but this has yet to be done. Depletion of cellular GSH A diet lacking protein, and hence cysteine and methionine, results in a GSH deficiency, such as is seen in human protein malnutrition (Shi et al., 1982). This becomes very apparent if there happens to be a high demand of GSH due to, e.g. administration of a high dose of a substrate for GSH conjugation (Glazenburg et al., 1983; Jung, 1985). In the rat fasting results in a 50% decrease in hepatic GSH after one day. However, after a fast for more than 48 h, GSH returns to almost normal concentrations, presumably due to cysteine mobilization from extrahepatic sources. Curiously in the rat GSH depletion due to a protein-restricted diet is sex dependent. For example, the GSH concentrations in the liver of male and female rats at 7 and 12 weeks of age on a normal diet were very similar (Igarashi et al., 1983). However, a low protein diet (containing 5% casein instead of 20%) decreased hepatic GSH approximately 70% and 40% in males and females, respectively (Mainigi and Campbell, 1981). BSO causes GSH depletion within a few hours. When it is given for several weeks in the drinking water, it causes a severe GSH depletion in the liver and several other organs (Sun et al., 1985). Upon termination of treatment, GSH returns to control levels with no signs of major liver or lung toxicity or acute effects on drug-metabolizing enzymes. The kidney responds more rapidly to BSO

GLUTATHIONE CONJUGATION 313

than the liver (Griffith and Meister, 1979; Drew and Miners, 1984; Sun et al., 1985). Compounds which are rapidly conjugated by GSH and which can be administered in high dose acutely deplete GSH in vivo and in isolated cells. DEM and phorone are two such compounds which are commonly used (Boyland and Chasseaud, 1970; Richardson and Murphy, 1975; Romero et al., 1984; Costa and Murphy, 1986; Younes et al., 1986; Dogterom et al., 1988). The depletion is rapidly reversible if a sufficient source of L-cysteine is available for resynthesis. Such treatments may have temporary side effects on organ functions that are directly or indirectly dependent on GSH. Protein synthesis and various enzymes are inhibited, while certain other activities are induced (Costa and Murphy, 1986; Dicker and Cederbaum, 1986; Oguro et al., 1987; Yoshida et al., 1987, 1988). Cytochrome P-450 activity may be affected (Anders, 1978). Certain drugs, for example paracetamol, which are effective in depletion are also hepatotoxic at the same dose required for depletion (Davis et al., 1974). Paracetamol may deplete GSH in man (Slattery et al., 1987). Ethanol decreases GSH in isolated hepatocytes, presumably due to its metabolism to acetaldehyde since this effect can be prevented by the alcohol dehydrogenase inhibitor pyrazole (Viña et al., 1980). Presumably acetaldehyde reacts with GSH to give a thiohemiacetal. Acute alcohol administration in vivo leads to a 35% decrease in hepatic GSH due to an increased GSH efflux from the liver. Inhibition of GSH synthesis has also been observed (Lauterburg et al., 1984; Speisky et al., 1985). Chronic alcohol treatment did not change the rate of elimination of GSH (Callans et al., 1987). Physical exercise will deplete GSH in the liver but not in muscles (Pyke et al., 1986). A high dose of L-cysteine given orally or intraperitoneally may also decrease liver GSH in the rat in vivo, but N-acetylcysteine (orally) does not have such an effect (Estrela et al., 1983). The effects of exercise and cysteine on GSH depletion may be related since both give rise to oxygen radicals, the former during respiration and the latter by autoxidation in the presence of transition metal ions. Salicylate decreases hepatic GSH by an unknown mechanism (Kaplowitz et al., 1980). Interorgan transport of GSH GSH in blood is present in the reduced form at approximately 20 µM (Anderson and Meister, 1980). When GSH was injected i.v. in the rat it was rapidly cleared from plasma with marked dose-dependent pharmacokinetics; the terminal elimination phase showed a t1/2 of 52 min at 50 µmol kg-1, and of 11 min at 300 µmol kg -1 (Ammon et al., 1986). Before this slow phase, there was a rapid elimination phase with a t1/2 of 2–3 min, both in the rat and in man (Wendel and Cikryt, 1980). The clearance of GSH in the rat was 30–40 ml min-1 kg-1,

314 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 11.1. Relationship between GSH in arterial blood and hepatic GSH in the rat. Variously treated groups of rats were used: □, controls; fasted; ∆, DEM-treated; ∇, Paracetamoltreated (taken from Adams et al., 1983 with permission).

with a volume of distribution of 150 ml kg-1 (Lauterburg et al., 1984; Callans et al., 1987). In selenium-deficient rats the plasma GSH was increased but the rapid t1/2 of elimination was not changed; the higher plasma GSH levels appeared to be due to an increased efflux from the liver (Hill and Burk, 1982, 1985). GSH in blood seems mainly derived from the liver (Lauterburg et al., 1984), due to a constant GSH efflux from the liver into blood (or into the perfusion medium in the perfused liver; Sies et al., 1978). GSSG, at levels 10% those of GSH, is also released from the liver into blood or the perfusion medium. However, when various substrates of cytochrome P-450 were added to the perfusion, the efflux of GSSG was increased while GSH efflux remained unchanged. Plasma GSH seems to reflect the hepatic GSH content while no such correlation exists for GSSG (Adams et al., 1983). For instance, fasting or pretreatment with DEM or paracetamol decreased liver and plasma GSH in parallel (Figure 11.1). The efflux of GSH from the liver most likely is a carriermediated process which operates close to saturation (80% of maximum) at normal liver GSH concentration (Ookhtens et al., 1985). Therefore, blood (or plasma) GSH may correspond with hepatic GSH as long as the liver GSH is decreased, but the relation would no longer hold if hepatic GSH were increased. The loss of GSH by efflux and its utilization in GSH conjugation necessitates continuous resynthesis. In the rat a high turnover rate of GSH was detected in kidney, liver and pancreas (Griffith and Meister, 1979).

GLUTATHIONE CONJUGATION 315

Lauterburg et al., (1980) have used paracetamol as a tool to follow GSH turnover in the liver in the rat. Paracetamol first has to be metabolized to an electrophile before it can be conjugated with GSH. By monitoring the specific radioactivity of this GSH conjugate in bile after addition of the three radiolabelled amino acid precursors, they determined a t1/2 of the specific radioactivity for the GSH conjugate of 2–4 h in the rat. The turnover rate decreased with age. Also in the perfused rat liver a decreased GSH efflux was observed with ageing (Ookhtens et al., 1987). DEM administration stimulated turnover due to the depletion of GSH. At high doses of paracetamol that are hepatotoxic, GSH turnover was also affected. The fractional rate of turnover in fasted rats was much higher than in fed rats. Interestingly, Kondo et al. (1984) reported that in human erythrocytes the GSH conjugate of CDNB increased turnover of GSH by activation of -GluCys synthetase activity. Feeding after fasting gave an overshoot of GSH in the liver. GSH in kidney, erythrocytes or muscle was much less if at all affected by fasting (Brooks and Pong, 1981; Cho et al., 1981; Lauterburg and Mitchell, 1981; Lauterburg et al., 1984). After fasting for 48 h the sinusoidal efflux of GSH from the liver had increased by 100%, while biliary efflux had not changed. The increased GSH turnover after such a fast was due to this increased efflux and not to increased break down. Fasting increased extrahepatic GSH catabolism (Lauterburg et al., 1984). GSH is degraded by the kidney at both the luminal surface of the renal brush border membrane (Hahn et al., 1978) and the basolateral side of the proximal tubule, as demonstrated in very elegant double-label experiments in the in vivo single-pass perfused rat kidney by Abbott et al. (1984). The γ-glutamyl transpeptidase inhibitor, acivicin, prevents the breakdown of GSH at the basolateral side in particular, although it also works at the luminal side. Little GSH is taken up by the kidney in the unchanged form—at most 10% (Curthoys, 1986). Approximately 90% of plasma GSH is extracted in a single pass through the kidney. Reapsorption of GSH from the tubular lumen is almost complete, in the form of its break-down products. Treatment with acivicin resulted in a 3000fold increase in urinary excretion of GSH (Curthoys, 1986). Isolated rat kidney tubular cells readily metabolize GSH by the γ-glutamyl transpeptidase pathway (Jones et al., 1979). In spite of this rapid metabolism of GSH by the kidney, 30–50% of GSH in plasma may ultimately be utilized for extrarenal metabolism (rat), providing other tissues with cysteine (Griffith and Meister, 1979; Hill and Burk, 1985). Biliary excretion The liver can also release GSH and GSSG into bile, although quantitatively it is much less; ten-fold more is released at the sinusoidal side than at the biliary side in the hemoglobin-free isolated perfused rat liver (Akerboom et al., 1982a).

316 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 11.2. Relationship between biliary output of GSSG and hepatic GSSG content in the perfused rat liver. Variously treated rats were used: ■, controls; ▲, pargyline-treated; ○, benzylamine-treated; ∆, pargyline+benzylamine-treated; ●, nitrofurantoine-treated; □, t-butylhydroperoxide-treated (taken from Akerboom et al., 1982a, with permission).

There is a correlation between hepatic GSH and GSSG and their biliary output (Figure 11.2). Thus, treatment of rats with DEM or BSP depleted hepatic GSH and decreased biliary efflux of GSH to the same extent. Phenobarbital seems to increase only biliary output since sinusoidal efflux was unaffected (Sies et al., 1978; Kaplowitz et al., 1983). The effect of several hepatotoxins, such as paracetamol, chloroform, and dimethylnitrosamine, and also oxidative stress in the liver, on biliary excretion of GSH and GSSG, has been reported (Akerboom et al., 1982a; Dubin et al., 1983; Lauterburg et al,. 1984). Anoxia leads to a decreased GSSG efflux, while GSH is unaffected (Cummings et al., 1988). The carrier-mediated transport involved in the transport of GSH and GSH conjugates into bile and blood has been investigated in isolated canalicular and sinusoidal membrane vesicles from rat liver (Inoue et al., 1983, 1984; Akerboom et al., 1984). It is very likely that GSH, GSSG and GSH conjugates all share the same carrier system. The GSH in bile is very rapidly oxidized to GSSG and should be determined immediately upon collection (Eberle et al., 1981). It is also metabolically labile in the bile duct: γ-glutamyltranspeptidase activity in the ductular epithelium readily metabolizes GSH in bile to various products, and leads to underestimates of biliary GSH output if only GSH and GSSG are measured (Abbott and Meister, 1986; Ballatori et al., 1986, 1988). Thus treatment with acivicin leads to much higher levels of GSH and less of its break-down products in bile. In young rats (two to ten weeks) the increase in γ-glutamyl transpeptidase activity

GLUTATHIONE CONJUGATION 317

was parallel to the increase in GSH-hydrolysis products in bile (Gregus et al., 1987). The output of GSH and GSSG in isolated hepatocytes comprizes both sinusoidal and biliary efflux. The GSH efflux can be inhibited by methionine and analogues like ethionine. The mechanism seems to be inhibition through an interaction on the outside of the cell (Aw et al., 1984, 1986a, 1986b). A similar inhibition by L-cysteine was observed in the perfused rat liver (Glazenburg et al., 1984). Various compounds, such as ethylmorphine, MPTP and menadione, induce the release of GSH and GSSG from isolated cells (Eklöw et al., 1981; di Monte et al., 1987). The liver is not unique in its GSH efflux. The isolated perfused rat heart also released GSH and GSSG in the perfusate and challenge with tbutylhydroperoxide led to an increased GSSG efflux (Ishikawa and Sies, 1984). 11.3. Glutathione conjugation Although GSH react with electrophilic carbon, oxygen, nitrogen and sulfur atoms, the term GSH conjugate usually refers to the product of the attack of GSH on an electrophilic carbon. Common reactions are nucleophilic displacement from saturated and aromatic carbons, Michael addition and nucleophilic attack on strained oxirane rings (Douglas, 1988; Ketterer et al., 1988). Each of these reactions is illustrated below. These reactions have spontaneous rates which vary considerably depending on the reactivity of the electrophile. For example, the second-order rate constant for the reaction of the carcinogen metabolite 1-nitropyrene-4, 5oxide (NP-4, 5-oxide) with GSH is 1.7×10−1 M−1 s−1 (Djuric et al., 1987), while that for the highly polarizable β unsaturated ketone N-acetyl-p-benzoquinone imine (NAPQI), which is a metabolite of paracetamol (acetaminophen) is approximately 3×104 M−1 s−1 (Coles et al., 1988), four orders of magnitude more reactive. Whether the spontaneous rate is high or low, such reactions are frequently, but not always, catalyzed by the GSH S-transferases (E.C. 2.5.1.18). Since electrophiles are usually toxic, the catalysis of their reaction with GSH in vivo is advantageous to the cell for three reasons. Firstly, the spontaneous rate with GSH may be too slow to compete with the reaction of electrophiles with critical cellular nucleophiles. Secondly, since the Km for GSH of GSH transferases is of the order of 0.1 mM (Jakoby et al., 1976), enzyme-catalyzed reactions, unlike the spontaneous reaction are not influenced by normal variations in intracellular GSH concentration. Enzymic rates are expected to diminish significantly only when the GSH concentrations approach the Km, a situation which arises in severe and highly pathological GSH depletion.

318 CONJUGATION REACTIONS IN DRUG METABOLISM

Thirdly, enzymic reactions may impose an advantageous regio- and/or stereoselectivity on the reaction. There is a range of nucleophilic species in vivo which differ in their reactivity with electrophiles. At one end are GSH and protein thiols, which are similar to GSH in their susceptibility to electrophiles, while at the other end are the nucleophilic sites in DNA, which are susceptible to those electrophiles which tend to react poorly with GSH (Coles, 1984). For example NAPQI which reacts well with GSH also reacts well with protein thiols and is cytotoxic, while NP-4, 5-oxide, which reacts poorly with GSH, reacts significantly with nucleophilic sites on DNA and is genotoxic (Djuric et al., 1986). The success of GSH in detoxication depends on its ability, through either spontaneous or enzymic reaction, to compete with critical cellular nucleophiles for reaction with potentially toxic electrophiles. 11.4. Nomenclature of the GSH transferases: a supergene family The GSH transferases have been found in both the plant and animal kingdoms (Ketterer et al., 1988). Membrane-bound forms have been detected (Kraus 1975; Morgenstern and DePierre, 1988), but soluble forms have been studied most. In those species so far investigated, there is a multiplicity of the soluble forms, and amino acid sequences deduced from cDNA clones suggest that all soluble GSH transferases are dimers of subunits of molecular weight from 20 000 to 25 000 (Taylor et al., 1987). Within a dimer, each subunit functions independently of the other (Mannervik and Jensson, 1982). Of all species, the most thoroughly studied is the rat followed by man and the mouse; beyond these, our knowledge is limited. GSH transferases in the rat At least 11 subunits have been characterized in the rat, although more are known to exist. Two nomenclatures and some of the distinguishing characteristics of these subunits are shown in Table 11.1. The nomenclature used in this chapter gives each subunit a number based on the chronological order of its characterization (Jakoby et al., 1984). The other nomenclature is an extension of one based on the mobility of subunits on sodium dodecyl sulfate polyacrylamide gel electrophoresis. The GSH transferases are now known to be a supergene family and can be further classified according to the multigene family to which they belong. Thus, complete primary structures are known for subunits 1, 2, 3, 4 and 7 (Lai et al., 1984; Pickett et al., 1984; Taylor et al., 1984; Ding et al., 1985; Suguoka et al., 1985; Telakowski-Hopkins et al., 1985; Pemble et al., 1986; Rothkopf et al.,

GLUTATHIONE CONJUGATION 319

Table 11.1. GSH transferase subunits in the rat.

1

Some of these values are unpublished results from E.Lalor, B.Coles, D.J.Meyer, K.H.Tan and B.Ketterer. 2 Isoelectric points refer to dimers of the subunit in question. 3 See Ostlund Farrants et al., 1987. 4 NN—not named.

1986) and fall into three groups which have been named alpha (subunits 1 and 2), mu (subunits 3 and 4) and pi (subunit 7), respectively (Mannervik et al., 1985a, 1985b). Since the identity in sequence between subunits 1 and 2 is 69% and between subunits 3 and 4 is 77% and sequence identity between the groups is only approximately 30%, each group is a separate gene family. On the basis of data including complete or partial amino acid sequences, enzymic properties and immunological cross-reactivity, the relationship between all 11 subunits is believed to be as follows: alpha family, subunits 1, 2, 8 and 10; mu family, subunits 3, 4, 6, 9 and 11; and pi family, subunit 7 only. Subunit 5 has yet to be assigned to a gene family (Meyer et al., 1984; Ketterer et al., 1985, 1986; Scott and Kirsch, 1987; B.Coles, D.J.Meyer, E.Lalor and B.Ketterer, unpublished information). Within a multigene family, subunits may form heterodimers; so far, GSH transferases 1–2, 3–4, 3–6, 3–9, 4–6, 4–9 and 6–9 have been identified (Ketterer et al., 1986). GSH transferases in man Studies on GSH transferases of the rat and other laboratory animals have been undertaken in the expectation that they might provide suitable models for the human. As more is known about human GSH transferases, it becomes apparent that much of the information obtained with the rat is indeed relevant to man. The same multigene families are seen, and there is considerable identity in primary structure across the two species.

320 CONJUGATION REACTIONS IN DRUG METABOLISM

The first separation of human liver GSH transferases involved a tissue sample from a single individual. Cationic forms were separated (Kamisaka et al., 1975) which are now known to be related to the alpha family of rat GSH transferases. In later studies of human liver from various sources, near neutral and anionic forms were found and shown to be homologous with the mu and pi families in the rat, respectively (Mannervik et al., 1985a; Hussey et al., 1986; Soma et al., 1986; Tu and Qian, 1986; Kano et al., 1987; Rhoads et al., 1987; Cowell et al., 1988). It is possible to separate up to 15 forms of the alpha class using chromatofocusing fast protein liquid chromatography (FPLC) (Vander Jagt et al., 1985; Ostlund Farrants et al., 1987). Isoenzymes at each extreme of the range of isoelectric points have been shown to be homodimers, while those in between are heterodimers. The subunits have been named either B1 and B2 (Stockman et al., 1987), Y1 and Y4 (Soma et al., 1986), or ax and ay (Ostlund Farrants et al., 1987); B1, Y1 and ax are the more basic of the two subunits in each nomenclature [an additional nomenclature refers to alpha-class enzymes as GST2 (Board, 1981) but does not as yet distinguish between subunits]. So far only two full-length cDNA clones for human alpha-family subunits, referred to as Ha1 and Ha2, have been described (Tu and Qian, 1986; Rhoads et al., 1987). On present evidence two homodimers and one heterodimer are the only forms so far predicted, and there is as yet no explanation for the multiplicity of forms separated by chromatofocusing FPLC. Comparisons of deduced amino acid sequences show 75% identity between human and rat alpha-class enzymes. Southern blots suggest that the human mu-class isoenzymes may be as numerous as those in the rat (J.T.Taylor, S.Pemble, R.Sherrington, J.Oliver and B.Ketterer, unpublished information) but so far only two have been isolated, one from the liver referred to as GSH transferase µ or GST1 and the other from muscle referred to as GST4. Studies of human liver show that, while the alphafamily GSH transferases are abundant, mu-class enzymes are either of low abundance or not present at all. From genetic studies of 179 liver samples from Chinese, Indian and Caucasian subjects, it was concluded that the GSH transferase µ or the GST 1 locus has two expressing genes and a null allele (Board, 1981). This conclusion has been supported by other population studies (Laisney et al., 1984; Strange et al., 1984; Harada et al., 1987). cDNA sequences for members of the human mu gene family have yet to be reported; however, comparison between amino acid sequences and a partial genomic sequence available for human GSH transferases of the mu family, once more indicate considerable” homology between the species (C.Southan, B.Coles, J.Hayes, D.J. Meyer, J.T.Taylor, S.E.Pemble, J.Oliver and B.Ketterer, unpublished information).

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Table 11.2. Nomenclature and physicochemical properties of human glutathione transferases.

a

Kamisaka et al., 1975. del Boccio et al., 1987. c P.Johnson, J.D.Hayes, D.J.Meyer, unpublished information. d Ostlund Farrants et al., 1987. e Not determined. b

The only pi-class enzyme so far found in humans is GSH transferase p which shows very close homology with rat subunit 7 cDNA (Kano et al., 1987; Cowell et al., 1988). For the properties and nomenclature of human GSH transferases, see Table 11.2. GSH transferases in the mouse The alpha, mu and pi multigene families have also been recognized in the mouse (Mannervik et al., 1985a; Daniel et al., 1987; Pearson et al., 1988); one alpha, two mu and three pi forms have so far been isolated (Hayes et al., 1987). cDNA clones of two mu-class enzymes have been isolated (Pearson et al., 1983), and cDNA clones encoding two alpha forms have been described (Daniel et al., 1987; Pearson et al., 1988); this should prompt the search for at least one other alpha-class enzyme in tissue extracts. No cDNAs for the pi family are available to date and, therefore, it is not known whether the three pi forms which have been isolated are artifacts, post-translational modifications or products of more than one gene. In the mouse, unlike rat and man, pi is present in the liver (Hatayama et al., 1986; Hayes et al., 1987) but apparently absent in many extrahepatic tissues (Hayes et al., 1987). Hepatic pi is much higher in males than females. For the properties and nomenclature of mouse GSH transferases, see Table 11.3.

322 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 11.3. Nomenclature and physicochemical properties of soluble GSH transferases in the mousea.

a

Mannervik and Danielson, 1988. and d refer to the mouse strains CD-1, DBA-2J and NMR1, respectively. Strain differences have yet to be detected. b, c

11.5. Conjugation reactions catalyzed by GSH transferases It is sometimes assumed that all GSH conjugations are enzymically catalyzed, but this is not always the case. For example, the aromatic amine carcinogen, Nmethyl-4-aminoazobenzene (MAB), yields two electrophilic metabolites, namely N-sulfonyloxy-MAB and 4-aminoazobenzene methimine, neither of which appear to be enzymically conjugated even though GSH conjugates are formed and excreted in vivo (Ketterer et al., 1979, 1982; Kadlubar et al., 1980). Nevertheless, the enzymic catalysis of the conjugation of a growing number of compounds has been demonstrated to occur in vitro. Nucleophilic displacement from saturated carbon An example of a substrate of this type is p-nitrobenzyl chloride, where chloride is displaced by GSH to give 1-(glutathion-S-yl)methylene-4- nitrobenzene. This reaction results in an optical density change which is used as the basis for a convenient spectrophotometric assay of GSH transferases, especially for subunit 3 which utilizes it particularly well. A number of substrates of pharmacological or toxicological importance in this class have also been studied. All but one of them are conjugated by displacement of halide. One is Melphalan which is a bifunctional N-mustard used in cancer chemotherapy. This gives two conjugates, a mono- and a diconjugate, resulting from the displacement from the mustard moiety of one and both of the chlorides, respectively (see Figure 11.3; Dulik et al., 1986). Another substrate from which chloride is displaced by glutathione is hexachlorobutadiene, an industrial chemical which gives 1-(glutathion-S-yl)1, 2, 3, 4, 4-pentachlorobutadiene (Wolf et al., 1984; Dekant et al., 1988). Bromide is displaced from the narcotic α-bromoisovaleryl urea (BIU), to give N(aminocarbonyl-2-glutathion-S-yl)-3-methylbutanamide (see Figure 11.4).

GLUTATHIONE CONJUGATION 323

Figure 11.3. GSH transferase catalyzed conjugation of Melphalan (Dulik and Fenselau, 1987).

Bromide is also displaced from ethylene dibromide (EDB), an industrial chemical which gives either a mono- or di-conjugate according to whether one or both bromides are displaced. The mono-conjugate is a sulfur mustard and therefore, in this case, GSH conjugation leads to toxification reaction rather than a detoxication (Peterson and Guengerich, 1988). Displacement of sulfate occurs in the case of 7-sulfonyloxymethyl, 12-hydroxymethyl benz[a]anthracene (Watabe et al., 1987) the carcinogenic metabolite of dimethyl benz[a] anthracene is detoxified to give 7-glutathionS-methyl, 12-hydroxymethyl benz [a]anthracene. Microsomes have been shown to catalyze the conjugation of Melphalan; microsomes and cytosol catalyze the conjugation of hexachlorobutadiene (Wolf et al., 1984; Dulik et al., 1986) and cytosol catalyzes the conjugation of 7sulfonyloxymethyl, 12-hydroxymethyl benz[a]-anthracene. The only substrates in the above list for which purified enzymes have been used are BIU and EDB. EDB is utilized to some extent by enzymes containing subunits 1, 2, 3, 4 and 7 from the rat and α, µ and π from man but is best utilized by rat subunit 2 and human α (J.Cmarik, D.J.Meyer, B.Ketterer, and F.Guengerich, unpublished information). BIU is a substrate for the major isoenzymes of the rat liver (those containing subunits 1, 2, 3 and 4; see Table 11.4). These GSH transferases are stereoselective with respect to BIU which is enantiomeric about the 2-carbon atom. Selectivity depends on the gene family; thus, the two alpha-family enzymes prefer the R-enantiomer and subunit 3 from the mu-family enzymes prefers the S-enantiomer (Te Koppele et al., 1988b).

324 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 11.4. GSH transferase catalyzed conjugation of α-bromoisovaleryl urea (Te Koppele et al., 1988).

The presence of an enantiomeric carbon does not always result in enantioselective conjugation. For example, enantio-selectivity has not been demonstrated for 2-iodo- and 2-bromo-octane (Ridgewell and Abdel-Monem, 1987), although it is appreciable with 1-chloro-1-phenylethane (Mangold and Abdel-Monem, 1980, 1983). Table 11.4 shows the specific activities available for the catalysis of EDB and BIU by rat GSH transferases. Nucleophilic displacement from aromatic carbon Two model substrates which illustrate nucleophilic displacement from aromatic carbons are 1-chloro-2, 4-dinitrobenzene (CDNB) which gives 1(glutathione-Syl)-2, 4-dinitrobenzene (DNB-SG) and 1, 2-dichloro-4-nitro-benzene (DCNB) which also gives a 1-(glutathion-S-yl) adduct. CDNB is used as a general substrate for the estimation of GSH transferase activity in unfractionated tissue extracts. It is a good substrate for all rat subunits except 5 and 9. Since subunits 5 and 9 are usually present in small amounts, the inability of CDNB to detect them is not of great importance. DCNB on the other hand is utilized principally by rat subunit 3 and is more specific than p-nitrobenzyl chloride. The toxicologically significant xenobiotics which fall into this category are the carcinogen, 4-nitropyridine-N-oxide, which gives 4-(glutathion-S-yl) pyridineN-oxide (Chasseaud, 1979) and the anticancer drug Melphalan, which, in addition to the two conjugates resulting from attack on saturated carbons described above, gives p-(glutathion-S-yl) phenylalanine resulting from attack on carbon 4 of the benzene ring of the phenylalanine moiety (Figure 11.3; Dulik and Fenselau, 1987). See Table 11.5 for specific activities available for the catalysis of CDNB and DCNB by rat GSH transferases.

GLUTATHIONE CONJUGATION 325

Reactions of GSH by Michael addition A model substrate which illustrates Michael addition is trans-4-phenyl-3buten-2one. This is specific for both rat subunit 4 and also human µ. Examples of xenobiotics which react with GSH by Michael addition are the following: Nacetyl-p-benzoquinone imine (NAPQI) the toxic metabolite of the analgesic paracetamol (acetaminophen) which gives 3-(glutathion-S-yl)paracetamol (Hinson et al., 1982; Figure 11.5; diethyl maleate (DEM), a compound used to deplete GSH (Section 11.2), which gives 1-(glutathion-S-yl)butandioic acid diethyl ester (Boyland and Chasseaud, 1967); and ethacrynic acid—a diuretic drug in man and used experimentally to deplete both cytosolic and mitochondrial GSH giving 2, 3-dichloro-4-(2-glutathion-S-yl-methylbutyryl) phenoxy-acetic acid (Boyland and Chasseaud, 1967). Of these substrates only ethacrynic acid and NAPQI have been tested with purified enzymes. Ethacrynic acid is utilized to some degree by a number of isoenzymes but is usually regarded as a marker for the pi class and is a very good substrate for rat subunit 8 of the alpha class (Kispert et al., 1989). The spontaneous reaction of NAPQI with GSH gives the reduction product, paracetamol, in addition to the GSH conjugate. Both reactions are catalyzed by the alpha-family enzymes, but the pi family is selective for GSH conjugation (Coles et al., 1988). 4-Hydroxynon-2-enal, a major cytotoxic and genotoxic product of the decomposition of endogenous peroxidized lipids, also belongs to this group, the product being 2-(glutathion-S-yl)-4-hydroxynon-2-anal (Boyland and Chaussead, 1967; Esterbauer et al., 1975; see Figure 11.6). In general hydroxyalkenals are remarkably good substrates for rat subunit 8 where rates increase in proportion to alkyl chain length and are the highest rates so far recorded for a GSH transferase (Danielson et al., 1987). See Tables 11.4 and 11.5 for some of the specific activities in catalysis of some of these substrates by rat GSH transferases. Reactions of GSH with xenobiotics containing strained oxirane rings A model substrate with an oxirane ring is 1, 2-epoxy-3-(p-nitro-phenoxy) propane, which is used as a marker substrate for rat subunit 5. The attack of GSH on an oxirane ring results in a glutathion-S-yl adduct on one carbon and the formation of a hydroxy group on the other and thus the possibility of two regio-isomeric products. Oxirane rings in xenobiotics usually result from metabolism of alkenes and arenes by mixed-function oxygenases (most commonly by the cytochrome P-450 system). Thus, styrene, which is used in the manufacture of polystyrene plastics, is an alkene which can be oxidized to (+)(R) and (−)(S) styrene-7, 8-oxide. The non-enzymic reaction with GSH

b

Mannervik et al., 1985b. Coles et al., 1988. c Coles et al., 1985. d Nemoto et al., 1975. e Robertson et al., 1986. f Djuric et al., 1987. g Te Koppele et al., 1988b. h J.Cmarik, D.J.Meyer, B.Ketterer and F.Guengerich, unpublished information. i Tsuchida et al., 1987. j nd=not determined.

a

Table 11.4. Rat GSH transferases. Activities (µmol min-1mg-1) towards substrates of biological importance.

326 CONJUGATION REACTIONS IN DRUG METABOLISM

b

Jensson et al., 1986; Ketterer et al., 1988. Mannervik et al., 1985b. c nd=not determined.

a

Table 11.5. Rat GSH Transferases. Activities (µmol min-1 mg-1) towardsmodel substrates some of which are used to specify subunits.

GLUTATHIONE CONJUGATION 327

328 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 11.5. GSH transferase catalyzed conjugation and reduction of the paracetamol (acetaminophen) metabolite N-acetyl-benzoquinone imine (Coles et al., 1988).

Figure 11.6. GSH transferase catalyzed conjugation of 4-hydroxy-non-2-enal (Alin et al., 1985).

gives all four possible diastereoisomers but is regio-selective, favouring reaction at the more electrophilic benzylic carbon. GSH transferases in liver, testis, intestine, lung and heart (particularly heart) catalyze a similar attack on the benzylic carbon of both enantiomers, but enzymic activity in the kidney and spleen favours reaction at the less electrophilic non-benzylic carbon (Dostal et al., 1986). Another example of an epoxide derived from a xenobiotic is exo-aflatoxin B1 (AFB 1)-8, 9-oxide. Non-enzymic GSH conjugates have not been reported, but enzymic GSH conjugation is directed by electrophilicity, giving 8-(glutathion-Syl)-9-hydroxy-8, 9-dihydro-AFB1. This reaction is catalyzed by rat and human GSH transferases of the alpha class only (Neal and Green, 1983; Coles et al., 1985). Benzo[a]pyrene (BP) is metabolized to a number of epoxides, one being the carcinogenic electrophile (+)anti-7, 8-diol-9, 10-oxide (7R, 8S, 9S, 10R). This compound has an electrophilic benzylic carbon, namely carbon 10, which is

GLUTATHIONE CONJUGATION 329

Figure 11.7. GSH transferase catalyzed conjugation of (+)antibenzo[a]pyrene-7, 8-diol-9, 10oxide (Roberton and Jernstrom, 1986).

selectively conjugated by GSH transferases (see Figure 11.7). Additional enantio-selectivity is also possible. When rat GSH transferases 1–1, 4–4 and 7–7 and a, µ and π enzymes from the human are tested with a racemic mixture, they are all selective for the (+) enantiomer (i.e. attack the benzylic carbon in the Rconfiguration in the order 7 and π>4 and µ >1 and α (Robertson et al., 1986). The most abundant epoxide metabolite of BP is BP–4, 5–oxide (BPO). Both carbons are benzylic and no selectivity is shown in the spontaneous reaction with GSH. In enzymic reactions, both regio- and enantio-selectivity are observed. GSH transferases in the liver, testis, intestine and lung attack the Rconfigured carbons and prefer the (4R, 5S)-BPO enantiomer. Enzymic activity in the heart also attacks R-configured carbon but shows strong preference for the (4S, 5R)-BPO enantiomer. Enzymic activity in the kidney and spleen is quite different in being selective for the S-configured carbon, although, like liver, testis, intestine and lung it prefers the (4R, 5S)-BPO enantiomer. Results concerning the stereoselectivity of purified rat GSH transferases are limited to those from GSH transferase 3–4 which is greater than 99% stereoselective for the R-configured carbon. More information is available from purified human enzymes. Thus, GSH transferase µ (the same gene family as rat subunits 3 and 4) and less so GSH transferases of the alpha family (the same gene family as rat subunits 1 and 2), preferentially catalyze attack on the R-configured carbon of (4R, 5S)-BPO. The opposite stereoselectivity (i.e. a preference for S-configured carbons) is shown by GSH transferase π (the rat equivalent being subunit 7; Hernandez et al., 1980; Cobb et al., 1983; Boehlert and Armstrong, 1984; Dostal et al., 1986). These results are in accord with the findings with rat tissue samples. The difference in behaviour of kidney and spleen compared with other tissues can be explained by their high content of a GSH transferase subunit 7 with analogous stereoselectivity to its human homologue GSH transferase π. 1-Nitropyrene (NP) gives both a 4, 5- and 8, 9-oxide. Presumably each of these has two enantiomers; however, neither they nor the four enantiomeric GSH conjugates which would result have been analyzed. The GSH transferase 3–3 catalyzed reaction of NP-4, 5-oxide with GSH gives a 1:1 mixture of the 4and 5-glutathionyl conjugates, respectively, but that of NP-9, 10- oxide gave a 2: 1 mixture of the respective 9- and 10- conjugates. This suggests that steric interference by the 1-nitro group hinders attack at the 10-carbon (Djuric et al.,

330 CONJUGATION REACTIONS IN DRUG METABOLISM

Table 11.6. Human GSH transferases. Activities (µmol min-1mg-1) towardssubstrates used to specify subunits.

a

Ketterer et al., 1988. Mannervik and Danielson, 1988. c Nemoto et al., 1975. d Robertson et al., 1986. c J.Cmarik, D.J.Meyer, B.Ketterer and F.Guengerich, unpublished information. f nd=not determined. b

1987). Note the differing Kms in Table 11.8. Both of these nitropyrene epoxides are best utilized by rat subunits 3 and 4. For data of the specific activities of GSH transferases for some of these substrates see Table 11.4. Other GSH transferase-catalyzed reactions Two reactions which do not result in conjugate formation should be mentioned because they are important marker substrates for GSH transferases. One is the reduction of organic peroxides, such as cumene hydroperoxide which is a marker for rat subunits 1 and 2 and alpha-family enzymes in man and mouse, and the other is the isomerization of androstene-3, 17-dione which is a marker for rat subunit 1 and may also be specific to certain members of the alpha family in other species (Ketterer et al., 1988). Within the same family, rat, human and mouse isoenzymes are similar but not identical in substrate specificity (Ketterer et al., 1988). Subunits ax and ay from humans and subunit 4 from mouse are associated with Se independent GSH peroxidase activity. Human GSH transferase µ, like rat subunit 4, has activity towards trans-4-phenyl-3-butene-2-one. Mouse subunit 1 like rat subunit 3 has activity towards DCNB. Human GSH transferase p and mouse subunit 3, like rat subunit 7, are good enzymes for ethacrynic acid and reduce linoleate

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Table 11.7. Mouse liver glutathione transferasesa. Activities (µmoles min-1 mg-1) towards a range of substrates.

a Warholm

et al., 1986. Mannervik and Danielson, 1988. c nd=not determined. b

hydroperoxide better than they do cumene hydroperoxide (see Tables 11.6 and 11.7). 11.6. Enzymology of glutathione transferases Catalytic constants, catalytic specificity and rate enhancement The ratio of kcat to Km describes the specificity of an enzyme for its substrate. This ratio, which has the dimensions of a second-order rate constant, cannot be greater than any second-order rate constant on the forward reaction pathway and, therefore, sets a lower limit on the rate constant for the association of an enzyme with its substrate. In very efficient enzymes, kcat/Km can approach the value for a diffusion-controlled rate of reaction between enzyme and substrate, e.g. for crotonase and fumarase the ratios are 2.8×108 and 1.6×108 M-1 s-1, respectively (Fersht, 1985). Catalytic specificities for several electrophiles of biological importance are presented in Table 11.8. It can be seen that these range from the very efficiently catalyzed substrate NAPQI, which has a value of 5×107 M-1s-1, to the very poorly catalyzed substrate BIU, which has a value of 167 M-1s-1 (Te Koppele et al., 1988b). The majority of values are in the region of 105 M-1s-1. The ratio of kcat/Km to k2, the latter being the second-order rate constant for the spontaneous reaction with GSH, has been mentioned as a possible indicator of rate enhancement brought about by an enzyme (Douglas, 1988). In the case of NAPQI and GSH transferase 2–2, this value is 1.7×103, but in the case of 1-

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Table 11.8. Some kinetic constantsa.

a

Ketterer et al., 1988.

nitropyrene-4, 5-oxide and GSH transferase 4–4 it is 6.5× 105 which is more than two orders of magnitude greater. Thus, although the catalytic specificity of the GSH transferase 2–2 catalyzed reaction of NAPQI with GSH is very high, with respect to rate enhancement the advantage is offset by the very high rate for the spontaneous reaction. The kinetic mechanism and the active site Most studies have involved the catalysis by GSH transferase 3–3 of CDNB conjugation. In steady-state studies marked deviations from Michaelis-Menten kinetics were encountered which led to a scheme in which the two substrates may be bound in random order (Jakobsen et al., 1977). Each subunit has an independent active site (Mannervik and Jensson, 1982) which is composed of a glutathione binding site (G-site) and a hydrophobic electrophile binding site (H-site). The specificity of the G-site depends on the subunit. A number of GSH analogues, modified in the ?-glutamyl moiety, have been tested as substrates (Adang et al., 1988; A.E.P. Adang, D.J.Meyer, J.Brussee, B.Ketterer, A.van der Gen and G.J. Mulder, unpublished information). It has been found that subunit 8 is the most stringent giving very little activity with anything but GSH; subunit 7 has in addition low activity with a-L- and a-D-glutamyl analogues; subunits 1 and 2 have appreciable activity with both the a-L-glutamyl and the glutaryl analogues; while subunits 3 and 4 utilize a much wider range of analogues including not only the glutaryl, aL- and a-L-glutamyl but also the a-D-and ß-D- aspartyl analogues. Subunit 3 will also use the ?-D-glutamyl analogue. With respect to the H-site it has already been demonstrated that while some electrophilic substrates are utilized by a number of subunits (in the case of CDNB), many others are utilized very selectively (see Table 11.4). Some idea of the enzyme mechanism is beginning to emerge. In spontaneous reactions it is clear that GS− rather than GSH is the reactive species (Coles et al., 1988; Douglas, 1988) and in the enzymic reaction a very important component is reduction of the pKa of the GSH thiol. Spectroscopic studies show

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that the pKa of the thiol of GSH when bound as a substrate to glutathione transferase 4–4 is approximately pH 6–8, that is approximately 2 pH units below that of GSH in aqueous solution. Thus, at the active site, the ratio of GS− to GSH is increased 100-fold the ratio found in free solution (Graminski et al., 1987). Such activation might involve an appropriately placed cationic amino acid residue. This proposition gains some support by work on a human mu-class enzyme in which it was shown that modification of one histidyl residue by diethyl pyrocarbonate, resulting in loss of activity, occurred in the absence but not the presence of GSH (Awasthi et al., 1987). Modification of arginine residues has also been shown to cause loss of activity. Since inactivation does not occur in the presence of ophthalmic acid, a GSH analogue in which cysteine is replaced by α-amino butyric acid, it is assumed that the arginine is associated with the G-site and presumed to function for GSH anion recognition rather than thiol activation. Activation of the electrophilic substrate may also occur. For example, stereoselectivity of catalysis might be due to differences in accessibility of the electrophilic centre of a substrate to a group in the enzyme which promotes its polarization. Inhibition GSH transferase activity is highly sensitive to inhibition by numerous compounds, both exogenous and endogenous. Endogenous metabolites, which are strong inhibitors of GSH transferase activity and may modulate enzyme activity in vivo, include bilirubin (Simons and Vander Jagt, 1980; Vander Jagt et al., 1985), haematin (Yalcin et al., 1983; Vander Jagt et al., 1985), bile acids (Hayes and Chalmers, 1983; Hayes and Mantle, 1986), unsaturated fatty acids (Meyer and Ketterer, 1987) and leukotriene C4 (Sun et al., 1985). Exogenous inhibitors are numerous and include trialkyl tin salts (Henry and Byington, 1976; Tipping et al., 1979), tetrabromosulfthalein (Clark et al., 1967) and Cibacron Blue F3GA (Tahir et al., 1985), plant phenols, such as ellagic acid and quercetin (Das et al., 1984), and the herbicides di- and trichlorophenoxyacetic acid. Some inhibitors are highly specific for particular GSH transferase isoenzymes and are used to discriminate between them (Tipping et al., 1979; Yalcin et al., 1983; Tahir et al., 1985). Since the subunits of GSH transferase dimers are kinetically independent (Danielson and Mannervik, 1985), these inhibitors may be used to distinguish between homodimeric and heterodimeric forms, provided inhibition is of the pure competitive or pure-noncompetitive type (Tahir and Mannervik, 1986). Recent studies on the inhibition by fatty acids, bile acids or haematin of CDNB conjugation by rat GSH transferase 1–1, 2–2, 3–3, 4–4 and 7–7 yielded

334 CONJUGATION REACTIONS IN DRUG METABOLISM

non-competitive inhibition in all cases (D.J.Meyer, unpublished information). Pure competitive inhibition was only obtained with linoleic acid during the GSH transferase 1–1 catalyzed reduction of linoleic acid hydroperoxide. Large variations in inhibition kinetics have been observed with a relatively small change in the structure of the substrate. For example linoleic acid is a pure inhibitor of the activity of GSH transferase 3–3 towards CDNB with an I50 of approximately 400 µM; however, with DCNB as substrate, it is only a partial inhibitor but with a much lower I50 of 10 µM (Meyer and Ketterer, 1987). Inhibition data suggest that conformational changes involving the active site occur as a result of substrate/inhibitor binding. In general, GSH conjugates are inhibitors of GSH transferases. In a homologous series, the more hydrophobic the S-substituent the greater the inhibition. For example, with S-n-alkyl substituents, the inhibitory effect increases with chain length (Askelöf et al., 1975). Similar effects are seen with the polyaromatic hydrocarbon GSH conjugates, and with these inhibitors stereoselectivity of inhibition has also been observed (Chen et al., 1986). 11.7. Distribution of GSH transferases in tissues and cells Tissue distribution GSH transferase activity has been found in every rat tissue so far examined and ranges from high values in the liver and testis to very low values in the nonlactating mammary gland. Striking differences in isoenzyme distribution occur from one tissue to another. HPLC analyses by the method of Ostlund Farrants et al. (1987) of the liver, kidney, lung, interstitial cells and spermatogenic tubules of the testis are shown in Figure 11.8. In the liver, the organ most active in GSH conjugation, subunits 1, 2, 3, 4, 6, 8 and 9 are clearly distinguished (both subunits 1 and 8 resolve into two forms). Noteworthy for its virtual absence in the normal liver is subunit 7 which has high activity towards BPDE and NAPQI and is found in many extrahepatic tissues. Particular subunits may predominate in certain tissues: subunit 8 in the erythrocyte (E.Lalor, B.Coles, D.J.Meyer, P.Alin, B. Mannervik and B.Ketterer, unpublished information), subunit 7 in the small intestine (Tahir et al., 1988) and subunit 2 in both the adrenal and lactating mammary gland. In some tissues subunits encountered elsewhere in small amounts may be particularly abundant. Thus, the testis is rich in subunits 6, 9 and 11 and the brain in subunit 6 (Figure 11.8). Immunohistochemistry has revealed interesting differences within tissues; it has been shown that in the normal liver, subunit 7 is present in all bile duct cells and in occasional hepatocytes (Tatematsu et al., 1985). Although subunits

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Figure 11.8. GSH transferase subunit composition of the soluble fraction of a) liver, b) kidney, c) lung, di) spermatogenic tubules; dii) interstitial cells of testis in the rat (Ketterer et al., 1988).

1, 2, 3 and 4 are present in all hepatocytes, they are more abundant in the area around the central vein than in the periportal region (Redick et al., 1982; Tatematsu et al., 1985). In the brain, GSH transferases have been detected in the astroglial and ependymal cells but not the neuronal stroma (Abramovitz et al., 1988). Human tissues in most cases have more GSH transferase activity than equivalent tissues in the rat (Baars et al., 1981). In human studies which, unlike

336 CONJUGATION REACTIONS IN DRUG METABOLISM

studies on experimental animals, involve outbred populations, interindividual differences are striking. Some results for the liver are shown in Figure 11.9 (P.Johnson, D.J.Meyer, F.Guengerich and B.Ketterer, unpublished information). Data concerning individual variation in other tissues are rarely available. However, in general the human kidney, like the rat kidney, has multiple forms of the alpha class, small amounts of an enzyme of the mu class and substantial representation of the pi class (Singh et al., 1987; Tateoka et al., 1987). In many other tissues, GSH transferase π is usually abundant and sometimes predominant. Examples are the erythrocyte, platelet (Loscalzo and Freedman, 1986), thyroid (del Boccio et al., 1987), placenta, lung, heart, spleen, kidney, pancreas and liver (Tateoka et al., 1987). Three other enzymes which have been isolated from extrahepatic tissues are GST4 from muscle, another member of the mu class (Laisney et al., 1984; Board et al., 1988), GST5 from brain (Suzuki et al., 1987) which has yet to be classified, and the alpha-class enzyme from skin, GSH transferase 9·9 (del Boccio et al., 1987). The substrate trans-stilbene oxide, which is a sensitive means for detecting GSH transferase µ, has been used to gather information about its distribution in a human population. Its presence or absence might govern the susceptibility of an individual to those drugs and toxins which are substrates. For example, assays of trans-stilbene oxide—GSH transferase in mononuclear leukocytes from smokers has shown that this activity is less common in those subjects with lung cancer than those without (Seidegard et al., 1985, 1987). GSH transferases during ageing The effect of age on the expression of GSH transferase activity has been investigated in the female mouse. A study of liver, lung and intestine between the ages of 2 weeks and 18 months showed that in the nine months following birth, the activity in the liver increased from 40 nmol/mg protein min−1 to 160 nmol/mg protein min−1 and then fell away sharply to post-natal levels by 15 months. Similar changes were seen in lung and intestine (Fujita et al., 1985). Reduced GSH transferase levels in the latter phase of life may occur not only in mouse but in other species including man and may be associated with the increased susceptibility to disease and drugs which occur with advanced age (Stohs et al., 1982). Induction A wide range of chemicals, both naturally occurring and synthetic, induce GSH transferases in the rat liver, including barbiturates, polyaromatic hydrocarbons (Igarashi et al., 1987), certain antioxidants, such as butylated hydroxytoluene (BHT), butylated hydroxyanisole (BHA) and ethoxyquin (Benson et al., 1978,

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Figure 11.9. Individual variations in GSH transferase subunit composition of human liver (Ketterer et al., 1988).

338 CONJUGATION REACTIONS IN DRUG METABOLISM

1979; Sato et al., 1984), the substrate trans-stilbene oxide (di Simplicio et al., 1983) and natural products, such as dithiolthiones which occur in Brassicas (Pearson et al., 1983). These inducers tend to be selective for subunits 1 and 3. The above inducers are anticarcinogenic in those cases where the genotoxic electrophile is a substrate for GSH transferase subunits 1 and 3. An example is AFB1-8, 9-oxide which is a substrate for subunit 1 (see Table 11.4). The powerful inducer ethoxyquin results in a four- to five-fold increase in the biliary excretion of AFB1-GSH conjugate and a 90% reduction in DNA adducts and greater than 95% reduction in the percentage of liver occupied by preneoplastic foci (Kensler et al., 1986). The mouse gives larger responses than the rat and has been used to detect inducing activity in a much wider range of compounds. For example, phenobarbital, BHA and BHT can increase GSH transferase levels in the liver up to ten-fold (Benson et al., 1978, 1979). A number of other compounds induce two- to five-fold, e.g. the coffee bean diterpene esters, kahweol—and cafestrol—palmitate (Lam et al., 1982), angelica lactone, coumarin, disulfiram, indole-3-carbinol and indole-3-acetonitrile (Sparnins et al., 1982a, 1982b), while still others, like streptozotocin, diethylnitrosamine (Agius and Gidari, 1985) and chronic ethanol induce up to two-fold (David and Nerland, 1983). Induction by crude extracts of Brassicas and tea leaves has also been observed (Lam et al., 1982; Sparnins et al., 1982a). Many of these inducers are active not only in the liver but also in the oesophagus, fore-stomach and intestine (Lam et al., 1982; Sparnins et al., 1982a; Parchment and Benson, 1984). Induction of GSH transferase activity in the fore-stomach has been shown to reduce its carcinogenic response (Sparnins and Wattenberg, 1981). GSH transferase gene expression during carcinogenesis Preneoplastic foci in rat liver resulting from treatment with hepatocarcinogens, such as MAB or AFB1, have enhanced levels of subunits 1 and 3 and express de novo subunit 7 (Kitahara et al., 1984; Sato et al., 1984; Power et al., 1987); many develop into nodules. Most of these foci and nodules redifferentiate into apparently normal liver tissue, but some persist and become neoplasms (Tatematsu et al., 1983; Meyer et al., 1985). Thus, livers containing preneoplastic foci, hyperplastic nodules or early tumours have an enhanced capacity to conjugate potentially toxic substrates for subunits 1 and 3 and an expanded substrate range due to the acquisition of subunit 7. These changes are both advantageous to the altered cells and also provide tumour markers; for example, subunit 7 is one of the earliest and most reliable markers for hepatocarcinogenesis in the rat. Human hepatocellular carcinomas, unlike those of the rat, do not express GSH transferase p. In a human liver it is diagnostic of

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either a cholangiocarcinoma or a metastasis, the latter most likely originating in the colon (Kodate et al., 1986; Soma et al., 1986; Hayes et al., 1987). However, human tumours arising in many other tissues show enhanced expression of GSH transferase p. This has been demonstrated immunohistochemically in cancer of the colon, stomach, pancreas and uterine cervix (Kodate et al., 1986; Sato et al., 1987) and by immunoblotting extracts from adenocarcinoma of the breast and lung, nodular small cell lymphoma and mesothelioma (Shea and Henner, 1987) and metastatic melanoma (Mannervik et al., 1987). It has also been demonstrated by isoelectric focusing in renal cortex tumours (di Ilio et al., 1987). Expression of high levels of GSH transferase p is often associated with malignancy. Many tumour cell lines also have elevated GSH transferase p including both small cell (Awasthi et al., 1988) and non-small cell lung carcinoma, ovarian carcinoma and the EJ6 bladder carcinoma cell lines (Wolf et al., 1987). In human tumour cell lines which express very low levels of GSH transferases, for example, the mammary carcinoma cell line MCF7, induction of resistance to adriamycin resulted in a 45-fold increase in the levels of GSH transferase p (Batist et al., 1986), although adriamycin is not itself a substrate for GSH transferase p. 11.8. Molecular biology of cytosolic GSH transferases The amino acid sequences of rat subunits 1, 2, 3, 4 and 7, two human alpha forms and GSH transferase p have all been deduced from cDNA clones and, when compared, demonstrate the existence of three multigene families (see above). In the alpha and mu families the multiplicity of expressing genes and the possible presence of pseudogenes leads to complex Southern blots. Rat subunit 7 and human GSH transferase p of the pi family each have only one expressing gene; whereas GSH transferase p gives the expected simple Southern blot, that for subunit 7 is complex indicating the presence of pseudogenes (Okuda et al., 1987; Cowell et al., 1988). Genomic clones are available for rat subunits 1 (Telakowski-Hopkins et al., 1985; King et al., 1989) and 7 (Okuda et al., 1987; Sakai et al., 1988), human p (Cowell et al., 1988), and a mouse alpha form (Daniel et al., 1987). Partial genomic sequences are also known for enzymes of both the rat and human mu families. The mu genes not only show the expected exon conservation, both within and across species, but are remarkable for inter- and intra-specific conservation of some intron structure (J.B.Taylor, S.E.Pemble, J.Oliver, R.Sherrington and B.Ketterer, unpublished information). The upstream regulatory regions of these genes are clearly of interest with respect to tissue specific expression and enzyme induction. Those for rat subunits 1 and 7 and human IT are in the process of being mapped.

340 CONJUGATION REACTIONS IN DRUG METABOLISM

Most is known about the subunit 7 gene which has multiple regulatory elements including a GC box and a phorbol ester responsive element (TRE box) at 47 and 61 bases upstream, a silencer element 400 bases upstream and an enhancing element between 2.2 and 2.5 kilobases upstream which contains another TRE box together with simian virus 40 and polyoma virus enhancer core-like elements. The latter TRE box appears to be more responsive to TPA than the former (Okuda et al., 1987; Sakai et al., 1988). Since TRE boxes have been associated with responsiveness to the ras oncogene (Imler et al., 1988), these particular elements may be involved in the de novo expression of subunit 7 which occurs in hepatocarcinogenesis. With respect to the general enhancer and the silencer region, the former may be involved in the widespread expression of subunit 7 and the latter may be important in tissues, such as the liver, which do not normally express subunit 7. The human π gene also contains a phorbol ester responsive element in its upstream regulatory sequence which may be associated with its enhanced expression in malignant cells (Cowell et al., 1988). Expression of the gene for subunit 1 is inducible by a number of compounds including phenobarbital and 3-methylcholanthrene. Two cis-acting regulatory elements have been identified; one is necessary for basal level expression and the other for responsiveness to β-naphthoflavone, a compound which, like 3methylcholanthrene, binds to the dioxin receptor. It has been shown in transfection experiments that a functional dioxin receptor is required for responsiveness to β-naphthoflavone (Telakowski-Hopkins et al., 1985; King et al., 1989), but it is not yet clear if the receptor ligand complex activates the subunit 1 gene by a direct or an indirect mechanism. 11.9. Membrane-bound GSH transferases Membrane-bound forms have been much less studied than soluble forms. They have been observed in hepatic microsomes and mitochondria and in membrane fractions from other cells (Morgenstern and DePierre, 1988). When isolated from rat liver, microsomes contain both strongly adsorbed soluble GSH transferases and at least one intrinsic enzyme (Ketterer et al., 1988). The major intrinsic enzyme is referred to as microsomal GSH transferase (Morgenstern and DePierre, 1988). It has a molecular weight of 17 200 and a primary structure which has no apparent homology with the amino acid sequences of known soluble GSH transferase subunits. Rat and human microsomal enzymes show 95% conservation in amino acid sequence. Southern blots indicate a single gene rather than a multigene family. Northern blots show that the enzyme is most abundant in the liver, abundant in the kidney and testis and present in smaller amounts in lung, spleen, seminal vesicle and brain (DeJong et al., 1988).

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Microsomal GSH tranferase is activated by compounds which react with thiols, including N-ethylmaleimide, disulfides, and perhaps also quinones and products of carbon tetrachloride metabolism. It has appreciable activity towards CDNB and some other model substrates and low activity towards two carcinogenic electrophiles, namely 1-nitropyridine-N-oxide and benzo[a] pyrene-4, 5-oxide (10 and 2.4 nmol min−1/mg microsomal protein, respectively) and also towards hexachlorobutadiene (Wolf et al., 1984; Morgenstern and DePierre, 1988). As yet there is/no clear view as to its function relative to that of the soluble GSH transferases. Other membrane enzymes have been isolated from leukocyte tumour cell lines which catalyze the conjugation of leukotriene A4 and are presumed to represent specific leukotriene C4 synthases (Bach et al., 1984; Söderström et al., 1988). 11.10. The metabolism of GSH conjugates Before excretion, GSH conjugates may be broken down to cysteinyl conjugates and the glutamic acid and glycine released retrieved by the body. The first step involves -glutamyl transpeptidase and produces a cysteinyl glycine conjugate. In the case of GSH conjugates excreted in the bile, this step may begin at the bile duct epithelium and continue in the gut. Those GSH conjugates entering the blood are filtered by the kidney and encounter γ-glutamyl transpeptidase at the surface of the proximal tubules (Meister, 1983). The second step is hydrolysis of the cysteinyl link by cysteinyl glycine dipeptidase (EC 3.4.13.6) or aminopeptidase M (EC 3.4.11.2). This too can commence in the bile duct, continue in the intestine and also take place in the kidney tubules. The result is a cysteine S-conjugate. Such conjugates may then either undergo enterohepatic circulation or be taken up by the kidney tubules. In both cases the action of N-acetyl transferases give an N-acetyl derivative, i.e. a mercapturic acid. The mercapturates are excreted in the urine, and as mentioned in the Introduction, some 100 years ago provided the first evidence for the pathway of detoxication initiated by GSH conjugation (Meister, 1983). 11.11. Conjugation in isolated cells and perfused organs GSH transferase content of isolated cells Isolated perfused organs or freshly prepared isolated cells may still closely resemble the organ in vivo. Cells in culture, however, may show pronounced changes in isoenzyme content. This is unfortunate since cells in culture otherwise would be

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a valuable alternative to the whole animal as a test system for the toxicity of drugs, etc. Thus, hepatocytes in primary culture undergo a remarkable change in gene expression within 48 h. GSH transferase subunits 1 and 2 fall to very low levels, subunit 3 is increased, subunit 4 remains unchanged and subunit 7 is expressed de novo (Vanderberghe et al., 1988). If cultured in the presence of dimethyl sulfoxide, hepatocytes retain a more normal differentiation. That is, subunits 1 and 2 remain at normal levels and subunit 7 is not expressed (Y.Vanderberghe, A.Guillouzo, C.Guigen-Guillouzo, J.B.Taylor, S.Pemble and B.Ketterer, unpublished information). Foetal hepatocytes, which are more easily cultured than adult hepatocytes, have also been used for toxicity testing, but these are even less appropriate since during the course of its development the foetal liver expresses both subunits 7 and 10, neither of which are found in the adult hepatocyte (K.Gilmore, D.J.Meyer, B.Ketterer and G.Yeoh, unpublished information). Substrate utilization by isolated hepatocytes and the perfused liver Several substrates have been used in these studies. Bromsulfthalein (BSP) is the substrate that has been investigated extensively, both in animal studies and in man. It is excreted exclusively in bile as the GSH conjugate and in unchanged form. Uptake by hepatocytes is carrier-mediated with a high affinity Km of ca. 5 µM (Schwenk et al., 1976; Orzes et al., 1985). Whether binding of BSP to albumin in blood plays a major role in the uptake process is controversial (Berk et al., 1987). In isolated rat hepatocytes rapid conjugation of BSP takes place (Schwarz, 1982). This can be completely prevented by addition of styrene oxide which depletes GSH and may inhibit BSP conjugation also by substrate competition at the level of the GSH transferases. In the perfused rat liver a major sex difference in biliary excretion of the GSH conjugate was observed, clearance in females was 40% higher than in males (Sorrentino et al., 1987). BSP has been shown to be taken up by Zone 1 (periportal) cells preferentially if it is added at a low concentration to the perfusate in the absence of albumin. The extraction of 0.01 mM was virtually 100% (Chen et al., 1984). In the retrograde perfusion BSP has been shown to be taken up in Zone 3 (perivenous cells). In both cases ca. 78% of the BSP excreted in bile was in the conjugated form, and there was no difference in the conjugation rate in Zone 1 and 3. Moreover, GSH availability was similar in both areas (Schön et al., 1988). This seems to disagree with micro-dissection and immunohistochemical studies described earlier in which the Zone 3 area was more densely stained for antibodies against GSH transferase subunits 1, 3 and 5 (Redick et al., 1982). Enzyme assays by Bengtsson et al. (1987) of hepatocyte fractions from the periportal (Zone 1) and perivenous (Zone 3) area isolated by the digitonin

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collagenase liver perfusion, confirmed the preferential Zone 3 location of GSH transferase (Bengtsson et al., 1987). However, Harris and Thurman (1986) confirmed with another substrate, DCNB, that GSH conjugation was equally effective in Zone 1 and 3. With CDNB they found much higher activities in Zone 1 than in Zone 3, where, in fact, little conjugation took place. When the flow direction was reversed, GSH conjugation of both substrates (measured in livers from phenobarbital-treated rats) took place preferentially in Zone 3 until GSH was depleted in that zone and conjugation shifted to Zone 1. At low substrate concentration substrate availability may be rate-limiting; for that reason no difference between Zone 1 and 3 is found. It seems, therefore, that substrate availability, zonal distribution of GSH transferases and GSH and flow direction all play a role in the final contribution of each zone in overall GSH conjugation. PB treatment tends to decrease differences in GSH transferase activity between Zone 1 and Zone 3 (Bengtsson et al., 1987). CDNB conjugation in the perfused rat liver leads to preferential biliary excretion of the conjugate, DNB-SG (Wahlländer and Sies, 1979). At high substrate influx GSH depletion occurs, leading to a decrease in conjugation rate; a residual conjugation rate of some 30 nmol min−1/g liver presumably reflects the rate of biosynthesis of GSH, as was confirmed by Harris and Thurman (1986). Since the perfusion medium in these studies did not contain sulfur-containing amino acids, the rate may be (far) below maximum. The GSH transferase activity towards CDNB is very high so that it does not become rate-limiting. If the DNB-SG conjugate is added to the perfusion medium, it is hardly excreted in bile; only the GSH conjugate synthesized in the liver is rapidly excreted, causing a choleresis of approximately 20 µl bile per µmol of conjugate. At low concentration CDNB uptake by the perfused liver is essentially complete (Lindwall and Boyer, 1987). Analysis of the excretion kinetics of the DNB-SG conjugate from isolated hepatocytes after its biosynthesis from CDNB showed that carrier-mediated transport was involved and that GSH conjugates could inhibit each other’s excretion, presumably by competition (Lindwall and Boyer, 1987). Similar competition for biliary excretion between the conjugates of DEM and CDNB has also been observed in the rat in vivo (Akerboom et al., 1982a; 1982b); GSSG utilizes the same carrier. The GSH conjugation of CDNB is very constant in primary monolayer culture of human hepatocytes for at least 72 h. Also GSH levels remain constant (Grant et al., 1987). GSH conjugation in hepatocytes from streptozotocin-induced diabetic rats was decreased due to both a lower GSH concentration and Vmax of GSH transferase in the hepatocytes (Grant and Duthie, 1987). GSH conjugation has also been studied in dog hepatocytes (Bolcsfoldi et al., 1981). The racemic substrate α-bromoisovalerylurea (BIU) has been extensively characterized in the rat in vivo, the isolated liver perfusion and isolated

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hepatocytes (Te Koppele et al., 1986a, 1986b, 1987, 1988a, 1988b; Polhuijs et al., 1989a; 1989b). The separate stereoisomers are conjugated at very different rates, and the GSH conjugates are excreted in bile in the perfused liver at correspondingly different rates. This was confirmed in isolated hepatocytes where a competing amidase-catalyzed hydrolysis yielded the α-bromoisovaleric acid (BI) product. The stereoisomers of BI are also conjugated with GSH in the rat in vivo or in isolated hepatocytes in a stereoselective fashion (Te Koppele et al., 1988a; Polhuijs et al., 1989a, 1989b). With another racemic substrate, aziridine carboxylic acid, a pronounced stereoselectivity in GSH conjugation by rat hepatocytes was observed; this was due to selectivity of uptake of the separate enantiomers by the cells, because a cytosolic fraction conjugated the enantiomers at the same rate (Hata et al., 1988). Styrene oxide is converted to the GSH conjugate and the epoxide hydrolysis product styrene glycol (Steele et al., 1981; Smith et al., 1983). At increasing dose of styrene oxide in the perfused liver, hepatic GSH decreased; however, GSH conjugation remained more important than epoxide hydrolase conjugation (van Anda et al., 1979). Only little of the styrene oxide-GSH conjugate was excreted in bile when the conjugate was added in the perfusion medium (Steele et al., 1981). After DEM pretreatment GSH conjugation is severely reduced (Smith et al., 1983). Busulfan [1, 4-bis(methane sulfonoxy)butane] is converted mainly to the GSH conjugate in the perfused rat liver. In this case the GSH conjugate excreted in bile is a sulfonium ion which decomposes to tetrahydrothiophene (Hassan and Ehrsson, 1987). From various other compounds, such as aromatic epoxides (Smith and Bend, 1979; Dock et al., 1987) and paracetamol (Moldeus et al., 1978), GSH conjugates are synthesized in the perfused liver or hepatocytes, but their formation has not been further characterized or is indirect. The reactive aromatic epoxides, although they are reactive, do leave the hepatocyte as demonstrated by their trapping outside isolated hepatocytes with [3H] labelled GSH (Richieri and Buckpitt, 1987). Other organs Little data are available about GSH conjugation in other perfused organs or cells from those organs. In the perfused rat kidney, paracetamol is converted to the GSH conjugate (Emslie et al., 1981). GSH conjugates of paracetamol and styrene oxide added to the perfusate were converted to the cysteine conjugate or the mercapturate and were excreted in urine (Steele et al., 1981; Newton et al., 1986). The differential distribution of GSH and GSH transferase activity in rabbit kidney has been investigated with special attention for its role in analgesic nephropathy (Mohandas et al., 1984).

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In the perfused rat lung or isolated lung cells, CDNB and epoxides are converted to GSH conjugates; GSH in the perfused medium or incubation medium stimulated conjugation, suggesting that the lung can utilize circulating GSH (Mehendale et al., 1981; Dawson et al., 1984). The GSH conjugate of styrene oxide was not metabolized by the perfused lung when it was added to the perfusion medium (Steele et al., 1981). 11.12. GSH conjugation in vivo Recenty Mulder and Te Koppele (1988) summarized the requirements for ideal model substrates to characterize the properties of glutathione conjugation in vivo. Briefly, these are 1) GSH conjugation should be the only metabolism; 2) GSH conjugation should be rate-limiting in the elimination of the substrate; 3) the substrate should be recovered quantitatively and the pathway(s) of primary excretion should be studied (in many cases bile rather than urine); 4) protein binding of the substrate should not be so high that it interferes with uptake by the cell; 5) it should be possible to administer the substrate intravenously; and 6) the substrate should not have effects that interfere with the physiology or biochemistry of the cell or animal, such as effects on cardiac output, liver blood flow or GSH biosynthesis. A chosen substrate will most likely be converted by only a limited number of GSH transferase subunits. Therefore, the pharmacokinetics of a substrate reflect only the properties of those subunits. Although many xenobiotics have been shown to be converted to GSH conjugates in vivo, only few substrates have been studied in detail: BSP, BIU, DEM, ethacrynic acid and allylisothiocy anate. BSP is mainly excreted in bile as the GSH conjugate; the percentage conjugated varies between species from 35–90% (Whelan, 1980; Foliot et al., 1984; Jimenez et al., 1988). When rats were pretreated with drugs that depleted GSH, a much smaller percentage was conjugated. DEM also inhibits biliary excretion of the infused BSP-GS conjugate, presumably because both conjugates compete for biliary excretion (Whelan, 1980). Clofibrate-treated rats had a lower GSH transferase activity and, therefore, a reduced rate of GSH conjugation of BSP in vivo (Foliot et al., 1984). Phenobarbital (PB) treatment increases GSH transferase activity and the rate of biliary excretion of BSP as the BSP-GS conjugate; it leads, however, to an increased biliary excretion of dibromosulfthalein, a non-metabolized analogue of BSP (Klaassen, 1970). The complexity of the effect of inducers like PB is illustrated by the finding that PB treatment specifically increases the uptake of the BSP-GS conjugate as detected in the liver perfusion (Sorrentino et al., 1987). Butylated hydroxyanisole and trans-stilbene oxide similarly induced GSH transferase activity, and the conjugation of BSP in vivo was enhanced (Gregus and Klaassen, 1982). Vitamin

346 CONJUGATION REACTIONS IN DRUG METABOLISM

A deficiency in the rat led to increased GSH transferase activity in the liver and an increased conjugation in vivo (Siddik et al., 1980a, 1980b). The kinetics of elimination and GSH conjugation of BIU have been studied in the rat (Kiwada et al., 1983; Te Koppele et al., 1986a, 1986b, 1988a). The racemic drug was excreted in bile as the diastereomeric GSH conjugates and in urine as the diastereomeric mercapturates. There was pronounced stereoselectivity in the conjugation of the separate stereoisomers of BIU which was reflected in their t1/2 of elimination from the blood and the t1/2 of the biliary excretion of the corresponding GSH conjugates (Te Koppele et al., 1986b; Polhuijs et al., 1989a). When the bile duct was ligated, the total dose was secreted into urine. The hydrolysis product of BIU, α-bromoisovaleric acid (BI) was also stereoselectively conjugated with GSH in vivo (Polhuijs et al., 1989b). Interestingly, the stereoselectivity towards BIU and BI was exactly the opposite, which could be explained by the stereoselectivity of the various GSH transferase isoenzymes involved in conjugation (Te Koppele et al., 1988b). Increase of GSH transferase activity by PB led to an increased conjugation rate of only the slowly conjugated enantiomers in vivo (unpublished data). Ethacrynic acid is mainly conjugated with GSH. The conjugate is subsequently excreted in bile (Klaassen and Fitzgerald, 1974). DEM decreases the clearance of ethacrynic acid and also reduces the biliary excretion of the GSH conjugate. The administration of DEM causes a choleresis because its GSH conjugate is excreted in bile (Barnhart and Combes, 1978). PB induced the GSH transferase form conjugating ethacrynic acid and enhanced its clearance in the rat (Wallin et al., 1978). Allyl isothiocyanate and comparable isothiocyanates form GSH conjugates in a reversible way; rapid dissociation may occur (Bruggeman et al., 1986). The disposition of allyl isothiocyanate in rat and mouse showed that most was excreted ultimately as mercapturate in urine (Ioannou et al., 1984). Biliary excretion occurred to an appreciable extent (Borghoff and Birnbaum, 1986). The stereoselective metabolism of R- and S- styrene oxide in the rat in vivo was studied by Watabe et al. (1982); very different behaviour was observed for the two stereoisomers. Elmhirst et al. (1985) reported on the conjugation of benzo[a]pyrene 4, 5-oxide in the rat and found a GSH conjugate in bile which underwent enterohepatic recirculation and further metabolism. 11.13. Inhibition of conjugation in vivo Inhibition of GSH conjugation in vivo can be attained by either a decrease of GSH concentration or by inhibition of GSH transferase activity. The decreased GSH availability can be induced by DEM or phorone, as outlined in Section 11.2; however, both are also substrates of GSH transferase so that they

GLUTATHIONE CONJUGATION 347

Figure 11.10. Conjugation of leukotriene A4 to give leukotriene C4 (Hammarström et al., 1979).

will act at two levels simultaneously. Treatment with BSO is preferable since it is not a substrate. Selective inhibitors of GSH transferase that work in the intact cell have not yet been found, other than substrates that inhibit by competition for conjugation. Thus, ethacrynic acid inhibits the GSH conjugation of busulfan in the perfused rat liver (Hassan and Ehrsson, 1987). 11.14. Conjugation and biological activity Leukotriene biosynthesis The peptido-leukotrienes are highly active chemical mediators derived from arachidonic acid by a biosynthetic pathway involving GSH conjugation. The first step involves the action of 5-lipoxygenase, giving (S)-5hydroperoxy-6trans-8, 11, 14, eicosatetraenoic acid (5-HEPTE) which is then converted to 5, 6-trans-oxido-7, 9-trans-11, 14-cis-eicosa-tetraenoic acid (LTA4) by LTA4 synthase, both activities residing in the same enzyme molecule (Hammarström, 1983; Samuelsson, 1983; Samuelsson et al., 1987). LTA4 can be hydrolyzed to the chemical mediator LTB4 or conjugated with GSH to give LTC4 (Hammarström, 1983; Samuelsson, 1983; Radmark et al., 1984; Samuelsson et al., 1987; see Figure 11.10) and is further metabolized to LTD4 and LTE4 (Hammarström et al., 1985; Bernström and Hammarström, 1986). The GSH conjugation reaction is enzymically catalyzed. The soluble GSH transferases described above, particularly those containing GSH transferase subunits 4 and 6, catalyze this reaction but utilize the methyl ester of LTA4 better; in addition they are not specific to the 5, 6-oxide, but will also conjugate the 11,12- and 14, 15-epoxide isomers which are not products of 5-lipoxygenase/ leukotriene A4 synthetase (Mannervik et al., 1984; Tsuchida et al., 1987). More active and more specific leukotriene synthases are membrane bound, distinct from microsomal GSH transferase, and abundant in lung and spleen in rat and hamster (Izumi et al., 1988) and in various blood cells and leukaemia cell lines (Bach et al., 1984; Abe et al., 1985; Yoshimoto et al., 1985; PaceAsciak et al., 1986; Söderström et al., 1988). These membrane-bound enzymes appear to be very labile and have yet to be obtained pure. They utilize the 5, 6-

348 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 11.11. GSH transferase catalyzed reaction of GSH with ethylene dibromide to give a genotoxic mono-conjugate and a non-toxic di-conjugate.

oxide only and prefer the free acid LTA4 to its methyl ester (Izumi et al., 1988; Söderström et al., 1988). LTC4 is metabolized by the mercapturic acid pathway. The action of γ-glutamyl transpeptidase gives the cysteinyl glycine derivative, LTD4, a specific LTD4 dipeptidase gives the cysteinyl derivative, LTE4, which can then be acetylated to N-acetyl LTE4. LTE4, N-acetyl-LTE4 and further metabolites are excreted in bile and urine (Huber and Keppler, 1988). LTC4 and its derivatives have a wide range of biological activities including effects on bronchial and vascular smooth muscle, an important role in inflammation, effects on mucous secretion, a neuroendocrine role in luteinizing hormone secretion and stimulation of growth of astroglial cells in culture (Huber and Keppler, 1988). Conjugation leading to toxicity Although the great majority of the reactions of electrophiles with GSH are detoxication processes, there are several interesting exceptions. For example, it has been mentioned that ethylene dibromide reacts enzymically with GSH to form 1-bromo-2-(glutathione-S-yl) ethane which is a sulfur mustard forming an episulfonium ion which reacts with the N-7 position of guanine in DNA (van Bladeren et al., 1980; Ozawa and Guengerich, 1983; Schasteen and Reed, 1983; Inskeep and Guengerich, 1984), a reaction thought to be responsible for the genotoxicity of ethylene dibromide (see Figure 11.11). Another example is that of the GSH conjugate of hexachlorobutadiene, S- (1, 2, 3, 4, 4-pentachlorobutadienyl)-glutathione, which is not mutagenic itself but becomes mutagenic

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after metabolism by the mercapturic acid pathway to cysteinyl derivatives followed by the action of β-lyase to give a mutagenic vinylic mercaptan (Wolf et al., 1984; Dekant et al., 1988). 11.15. Useful techniques GSH and GSSG either individually in or when both present in a mixture can be measured by the methods of Tietze (1969) and Griffith (1980). Stable GSH conjugates can be separated by thin layer chromatography or HPLC and recognized by polarity, reactivity with ninhydrin and convertibility to a less polar ninhydrin-reacting component upon incubation with γ-glutamyl transpeptidase. Once purified, mass spectrometry and NMR can be used to confirm that the compound is a GSH conjugate and define its structure (Ketterer et al., 1979, 1982; Coles et al., 1988). GSH transferase purification is greatly assisted by affinity chromatography. Both S-hexyl GSH-Sepharose (Mannervik and Guthenberg, 1981) and GSH-Sepharose (Simons and Vander Jagt, 1980) have proved useful. All rat GSH transferases apart from GSH transferase 5–5 and a proportion of GSH transferase 1–1 are retained. Further purification of the eluted GSH transferase fraction can be achieved by using chromatofocusing followed by chromatography on hydroxyapatite. The latter, in addition to carrying out useful separations, removes ampholytes present in fractions from the chromatofocusing column. Where acidic transferases are concerned, chromatofocusing at low pH where the enzyme may be unstable is inadvizable and an ion exchange chromatography is preferred. Some other useful observations are as follows: GSH transferases 1–1 and 8–8 elute early from GSHSepharose and GSH transferases 3–3 and 4–4 late; GSH transferase 7–7 elutes early from hydroxyapatite and GSH transferase 3–3 late (Beale et al., 1982, 1983; Meyer et al., 1984, 1985; D.J. Meyer and B.Ketterer, personal communication). The qualitative and quantitative subunit analysis of mixtures of GSH transferases derived from tissue extracts or fractions in purification procedures can be obtained using reverse phase HPLC (Ostlund Farrants et al., 1987). Assay techniques utilizing spectrophotometric changes are conjugate formation in the case of the general substrate CDNB and the selective substrates DCNB, trans-4-phenyl-buten-2-one and 4-hydroxy-non-2-enal and isomerization of androstene-3, 17-dione. In the case of reduction of cumene and linoleate hydroperoxides the reaction is coupled with NADPH-dependent GSH reductase (see Beale et al. 1982, 1983; Meyer et al., 1985). High levels of sequence homology in exon structure between species enables cDNAs derived from one species to be used as successful probes in another (Cowell et al., 1988).

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11.16. Conclusions GSH conjugation is only one of the functions of GSH. The detoxication of H2O2 and the provision of a non-toxic reservoir and transport form for cysteine required for protein synthesis are two others. GSH transferases not only catalyze GSH conjugation but also reduction of electrophilic oxygen, nitrogen and sulfur and certain isomerizations (Ketterer et al., 1988) and may also be important in intracellular transport of endogenous and xenobiotic lipophiles (Tipping and Ketterer, 1981). However, GSH conjugation is one of the most important functions they perform. The enzymes as proteins are relatively stable and have proved easier to study than the reactions of importance which they catalyze. Toxicologicallyimportant substrates are often reactive and labile and therefore difficult to synthesize chemically or isolate and identify when produced biosynthetically. Whereas the techniques used in studies of the enzyme protein are standard, those used to identify unstable intermediates may require very sophisticated and expensive technology. Although the rate of advance in this area is very gratifying, there are still a number of apparent anomalies which require explanation. For example, in the rat there is a remarkably specific tissue distribution of GSH transferase subunits suggesting the need for a particular constellation of isoenzymes for the proper function of each tissue. However when a comparison is made between species of the distribution of GSH transferase gene families in the liver (and perhaps other tissues), such big differences are found that doubts are raised about the significance attached to tissue-specific distribution. Thus alpha and mu families are found in the rat liver, the alpha family predominates in the human liver, the mu family in the hamster liver; but in mouse liver alpha, mu and pi families are all well represented (the pi enzyme, particularly so in the male). Evidence so far available suggests that between species homologous enzymes have similar substrate specificities. It may be that variations in gene expression are possible providing sufficient activity towards endogenous toxic substrates which are ubiquitous, such as those derived from lipids (lipid peroxides and hydroxyalkenals), is always available. Acknowledgements Brian Ketterer wishes to thank the Cancer Research Campaign, of which he is a Life Fellow, for their generous support and Miss Lucia Christodoulides for secretarial assistance.

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Abbreviations AFB BHA BHT BI BIU BP BPDE BPO BSO BSP CDNB DCNB DEM DNB EDB FPLC GSH GSSG HPLC LT MAB NAPQI PB -SH

Aflatoxin B1 Butylated hydroxyanisole Butylated hydroxytoluene α-Bromoisovaleric acid α-Bromoisovalerylurea Benzo[a]pyrene Benzo[a]pyrene-7, 8-dihydrodiol-9, 10-epoxide Benzo[a]pyrene-4, 5-oxide Buthionine sulfoximine Bromosulfthalein 1-Chloro-2, 4-dinitrobenzene 1, 2-Dichloro-4-nitrobenzene Diethylmaleate -S Glutathione conjugate of CDNB Ethylene dibromide Fast protein liquid chromatography Glutathione Oxdized glutathione High performance liquid chromatography Leukotriene NMethyl 4-aminoazobenzene NP 1-Nitropyrene N-Acetyl-p-benzoquinone imine Phenobarbital Thiol group References

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Conjugation reactions in drug metabolism Edited by G.J.Mulder © 1990 Taylor & Francis Ltd

CHAPTER 12 Epoxide hydrolases Thomas M.Guenthner Department of Pharmacology, University of Illinios College of Medicine, Chicago, IL 60612, USA.

12.1.

INTRODUCTION

366

12.2.

MICROSOMAL EPOXIDE HYDROLASE

367

12.3.

Substrates

368

Inhibitors and inducers

369

Enzyme purification and mechanism

370

Toxicological implications

373

CYTOSOLIC EPOXIDE HYDROLASE

375

Substrates

375

Inhibitors

377

Enzyme purification and mechanism

378

Induction

381

Biological role

382

12.4.

EXPERIMENTS IN ISOLATED CELLS, PERFUSIONS AND IN VIVO

383

12.5.

ENDOGENOUS SUBSTRATES

385

Cholesterol 5, 6-oxide hydrolase

385

Leukotriene A4 hydrolase

391

12.6.

PRACTICAL CONSIDERATIONS

396

ABBREVIATIONS

397

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REFERENCES

397 12.1. Introduction

The epoxide hydrolases are a group of catalytically-related enzymes, each with its own unique structural and biochemical characteristics. Because they are related but distinct enzymes, the problem of nomenclature is somewhat difficult. Early descriptions of enzymic epoxide hydrolysis named the enzyme involved ‘epoxide hydrolase’ (Maynert et al., 1970) or ‘epoxide hydrase’ (Jerina et al., 1968a). Other contemporary descriptions also used the term ‘epoxide hydratase’. In 1978, the Nomenclature Committee of the International Union of Biochemistry recommended that the enzyme be called ‘epoxide hydrolase’ (EC 3. 3.2.3). At that time it was not widely recognized that more than one epoxide hydrolase existed, and it was assumed that the enzymic hydrolysis of all epoxides was catalyzed by a single enzyme located in the endoplasmic reticulum, or perhaps by the same enzyme adventitiously present to a small degree in the cytosolic fraction. With the subsequent demonstration of a unique epoxide hydrolase located primarily in the cytosol and peroxisomes, which had distinctly different biochemical properties from those of the microsomal enzyme, the terms ‘microsomal epoxide hydrolase’ or ‘mEH’, and ‘cytosolic epoxide hydrolase’ or ‘cEH’, were informally adopted to differentiate the two distinct hydrolases. Further complications arose with the demonstration that mEH was found in cytosol and nuclei, and that cEH could be found in microsomes (see below). Furthermore, a second unique microsomal epoxide hydrolase, which hydrolyzes cholesterol 5, 6-oxides, and a second unique cytosolic epoxide hydrolase, which hydrolyzes leukotriene A4 have been discovered. The problem of nomenclature for these enzymes has yet to be resolved; suggestions include that they be differentiated by subcellular location, by substrate specificity, or simply by unique numbers. Due to the lack of completely rigorous subcellular localization or substrate specificity, the use of a system of unique numbers or letters for each epoxide hydrolase, as is now the accepted system for cytochrome P-450 nomenclature, appears to be the most systematic. However in the case of this review, the following definitions will be subsequently used: ‘Microsomal epoxide hydrolase’ or ‘mEH’ will be used to describe the form of epoxide hydrolase that is found predominantly in the endoplasmic reticulum and that catalyzes the hydrolysis of a broad spectrum of xenobiotic epoxides, specifically including arene oxides. This enzyme has also been referred to as ‘mEHb’ (Oesch et al., 1984), ‘EH1’ (Guenthner and Oesch, 1983) and ‘mCSO’ (Finley and Hammock, 1988). ‘Cytosolic epoxide hydrolase’ or ‘cEH’ will be used to describe that form of epoxide hydrolase that is primarily found in cytosol and

EPOXIDE HYDROLASES 367

peroxisomes and that specifically catalyzes the hydrolysis of trans-substituted styrene oxides. This enzyme has also been referred to as ‘EH2’ (Guenthner and Oesch, 1983) and ‘cTSO’ (Finley and Hammock, 1988). ‘Cholesterol 5, 6-oxide hydrolase’, or ‘ChEH’ will be used to describe that epoxide hydrolase that specifically catalyzes the hydrolysis of cholesterol 5, 6α-and 5, 6β-oxides. This enzyme has also been referred to as ‘mEHch’ (Oesch et al., 1984) and ‘mCE’ (Finley and Hammock, 1988). Finally, ‘LTA4 hydrolase’ or ‘LTA4H’, will be used to define that epoxide hydrolase that stereo-selectively converts leukotriene A4 to leukotriene B4. 12.2. Microsomal epoxide hydrolase The existence of epoxides as intermediates in the metabolic formation of dihydrodiols was postulated in the 1950s (Boyland, 1950, Boyland and Wolf, 1950). In the late 1960s it was demonstrated that epoxides are formed from the cytochrome P-450-dependent oxygenation of aromatic and olefinic hydrocarbons (Jerina et al., 1968a; Jerina et al., 1970a; Leibman and Ortiz, 1970). That an enzymic process is involved in the conversion of these epoxides to dihydrodiols was systematically demonstrated at about the same time for both arene and alkene oxides (Jerina et al., 1968b; Maynert et al., 1970; Oesch et al., 1971a; Watabe et al., 1971). The enzyme that catalyzes this reaction was characterized and partially purified (Oesch and Daly, 1971) and eventually isolated as a single, homogeneous protein species (Bentley and Oesch, 1975; Lu et al., 1975). Mainly because of the participation of this enzyme in metabolic pathways that are crucial to the activation and deactivation of polycyclic aromatic hydrocarbons to and from mutagens and carcinogens, the microsomal xenobiotic epoxide hydrolase (mEH) has attracted great scrutiny from chemists, pharmacologists and toxicologists. It has been the subject of several extensive reviews over the last 15 years (Oesch, 1973; Lu and Miwa, 1980; Guenthner and Oesch, 1981b; Seidegård and DePierre, 1983; Wixtrom and Hammock, 1985; Armstrong, 1987; Yang 1988); the reader is referred to these reviews for more indepth discussions of the physical, chemical, and toxicological properties of mEH than will be offered here. mEH catalyzes the conversion of a fairly wide-ranging group of arene and alkene oxides to vicinal dihydrodiols by hydrolytic cleavage of the oxirane ring. The toxicological significance of this conversion lies in the fact that it represents the conversion of usually highly reactive, electrophilic oxirane species to less reactive, non-electrophilic vicinal dihydrodiols (Jerina and Daly, 1974; Sims and Grover, 1974; Lu and Miwa, 1980; Guenthner and Oesch, 1981b; Wixtrom and Hammock, 1985). The enzyme is widely distributed throughout the animal kingdom (Walker et al., 1978; Balk et al., 1980; Knight

368 CONJUGATION REACTIONS IN DRUG METABOLISM

and Walker, 1982; Glatt and Oesch, 1987). Generally speaking, levels are low in more primitive vertebrates and higher in rodents and larger mammals. mEH activity is also found in one-cell organisms (Kolattukudy and Brown, 1975), insects (Jansen et al., 1986), and higher plants (Croteau and Kolattukudy, 1975). The organ distribution of mEH is broad. Activity was sought in 25 organs from the rat and was measurable in all (Oesch et al., 1977). In rat, the highest activity is seen in liver, followed by testis, kidney, ovary and lung (Oesch et al., 1977; Seidegård and DePierre, 1983). Subfractionation of the hepatocyte reveals that mEH is predominantly, but not exclusively, located in the endoplasmic reticulum, represented by the microsomal fraction in vitro. Significant mEH activity is also found in the nuclear membrane, and lesser amounts are seen in plasma membrane and Golgi fractions (Stasiecki et al., 1980). In some organs and species, particularly human liver (Wang et al., 1982; Schladt et al., 1988b) and lung (Guenthner and Karnezis, 1986a, 1986b), significant amounts of mEH are found in the cytosolic fraction. While mEH is no longer thought of as exclusively membrane bound, it nevertheless can be considered primarily and predominantly a microsomal enzyme. Surveys of mEH activities in a large number of inbred and outbred strains of rats and mice revealed minor quantitiative differences, but no obvious quantitative genetic polymorphism was seen (Oesch et al., 1983; Glatt and Oesch, 1987). A qualitative polymorphism, reflected as an altered pH optimum, has been observed in some inbred mouse strains (Lyman et al., 1980). This difference probably represents enzyme microheterogeneity but has not been shown to be of toxicological significance. Substrates The best substrates for the enzyme are oxiranes with one or two hydrophobic substituents. The monosubstituted oxirane, styrene oxide (Oesch et al., 1971a), and the disubstituted oxirane, cis-stilbene oxide (Mullin and Hammock, 1980), are model substrates used widely in the measurement of mEH activity. While cisdisubstituted oxiranes are generally good substrates for mEH, trans-disubstituted, as well as tri- and tetrasubstituted oxiranes, are poorly hydrolyzed (Oesch et al., 1971b). Arene oxides, including those formed from polycyclic aromatic hydrocarbons, are cis-disubstituted oxiranes and are particularly good substrates for mEH (Bentley et al., 1976; Jerina et al., 1977; Lu et al., 1977). Benzo[a] pyrene 4, 5-oxide is used in both radiometric (Schmassmann et al., 1976) and fluorometric (Dansette et al., 1979) assays of mEH activity and is particularly useful in specifically measuring mEH activity in the presence of cEH (Guenthner and Karnezis, 1986a). While both K-region and non-K-region arene oxides are good substrates for mEH (Lu et al., 1977), bay-region dihydrodiol epoxides appear to be very poor substrates (Wood et al., 1976; Bentley et al., 1977; Guenthner and Oesch, 1981a). Epoxides formed from C16-unsaturated steroids

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are good substrates for mEH. These include the 16, 17 a- and ß-oxides formed from estratetraenol and androstadienone (Breuer and Knuppen, 1961; Bindel et al., 1979). On the other hand, epoxides formed at the 5, 6 double bond of cholesterol are not substrates for mEH (Levin et al., 1983; Oesch et al., 1984). They are, however, rapidly hydrolyzed by another microsomal epoxide hydrolase specific for cholesterol oxide which is discussed below. Halogenated hydrocarbon epoxides are also substrates for mEH. Examples include epoxides formed from halogenated cyclodiene insecticides (dieldrin analogues; Brooks et al., 1970) and the epoxide formed by monooxygenation of vinyl chloride (Guengerich et al., 1979a), Inhibitors and inducers The best inhibitors of mEH activity are generally epoxides that are poorly hydrolyzed by the enzyme (Oesch et al., 1971b), such as trichloropropene oxide and cyclohexene oxide. These compounds do not appear to be strictly competitive inhibitors, however (Oesch et al., 1971b), which may reflect allosteric interaction of the epoxide inhibitors with nucleophilic sites on the enzyme. Metal ions, such as Hg+2, Zn+2, and Cd+2 inhibit mEH (Parkki et al., 1980). Compounds that bind sulfhydryl groups are weak inhibitors (Oesch and Daly, 1971), while 2-bromo-4′-acetophenone, a compound that binds strongly to imidazole nitrogen (DuBois et al., 1978), is a potent inhibitor. A series of cyclopropyl oxiranes were synthesized as proposed mechanism-based irreversible inhibitors of both mEH and cEH activity (Prestwich et al., 1985b). While mEH activity was inhibited by these compounds, it was in a reversible, competitive manner rather than by suicide inhibition. The activity of mEH can be selectively stimulated, either by enzyme induction in vivo, or by direct stimulation of enzyme activity in vitro. Levels of mEH in microsomes are induced by administration of a wide variety of chemical compounds, including phenobarbital, Arochlor 1254, 2(3)-t-butyl-4hydroxyanisole (BHA), 3, 5-di-t-butyl-4-hydroxytoluene (BHT), 2acetylaminofluorene (for review, see Seidegård and DePierre, 1983, and Meijer and DePierre, 1988); heterocyclic compounds including benzofuran, coumaran, trimethylene oxide, trimethylene sulfide, and indole 3-carbinol (Heine et al., 1986); mercuric ion (Kroll et al., 1988); nitrosamines (Craft et al., 1988); and stilbene congeners including benzil, cis-stilbene oxide, and trans-stilbene oxide (Seidegård et al., 1981). Many of these compounds also produce a generalized proliferation of endoplasmic reticulum and an increase in the levels of mutiple microsomal enzymes. Where specific mechanisms of induction of mEH were investigated, it was determined that exposure to the inducing agent results in an increase in mEH-specific mRNA, followed by increased synthesis of enzyme protein (Craft et al., 1988). The 3methylcholanthrene-type inducers, which

370 CONJUGATION REACTIONS IN DRUG METABOLISM

specifically induce cytochrome P-450I isozymes, induce mEH poorly or not at all in the liver (Oesch et al., 1970; Oesch, 1976). However, β-naphthoflavone is reported to induce mEH activity in mouse mammary gland (Silva et al., 1988). Compounds classified as ‘peroxisome proliferators’, including the antihyperlipidemic drugs clofibrate (Loury et al., 1985) and tiadenol (Schladt et al., 1987), as well as halogenated phenoxyacetic acid herbicides (Lundgren et al., 1987a), induce increases in mEH activity ranging from slight to highly significant. These compounds also produce increases in total microsomal protein, so the increase in mEH activity is probably not strictly related to peroxisome proliferation. Stimulation of mEH activity in vivo is also seen, particularly by heterocyclic compounds. Compounds that enhance or stimulate mEH activity include flavones (Ganu and Alworth, 1978), substituted imidazoles (James and Sloan, 1985), harmane, norharmane (Vaz et al., 1981), metyrapone, isoquinoline and clotrimazole (Seidegård et al., 1986). Non-heterocyclic compounds, such as chalcone oxide, 9-fluorenone (Ganu and Alworth, 1978) and several substituted benzil derivatives (Seidegård and DePierre, 1983), also stimulate mEH activity in vitro. A unified hypothesis for the mechanism of stimulation is difficult to propose because the degree of stimulation varies greatly depending on the substrates investigated. For example, clotrimazole, which increases the rate of hydration of some substrates, actually inhibits the hydration of others (Seidegård et al., 1986). Enzyme purification and mechanism The enzyme has been purified from a number of tissue sources, and its biophysical and structural properties are well characterized (Bentley and Oesch, 1975; Lu et al., 1975; Guengerich et al., 1979b; Lu et al., 1979). It is a monomer with a molecular weight estimated at approximately 50 kDa by SDSpolyacrylamide gel electrophoresis. In the absence of detergent, the isolated enzyme forms large oligomeric aggregates (Guengerich et al., 1979b). The enzyme has no prosthetic groups and is not a metalloenzyme or a glycoprotein. The complete amino acid sequence of mEH has been directly determined (Heinemann and Ozols, 1984) or deduced (Porter et al., 1986; Jackson et al., 1987; Skoda et al., 1988), and as would be expected from its membrane-bound nature, it contains a high proportion of hydrophobic amino acids. A hydrophobic sequence at the N-terminus (Heinemann and Ozols, 1984) may act as a signal sequence for membrane insertion. In addition, three (Heinemann and Ozols, 1984) or six (Porter et al., 1986) membrane-spanning domains have been identified. Both the directly determined sequence of rabbit liver mEH and the deduced sequences of rat and human liver mEH show a single polypeptide chain containing 455 amino acids, with a calculated molecular weight of 52 581 (Porter et al., 1986), 52 691 (Heinemann and Ozols, 1984), or 52 956 (Skoda et

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al., 1988). Rat liver mEH is 81% homologous to the rabbit liver enzyme (Heinemann and Ozols, 1984) and 82–84% homologous to human liver mEH (Jackson et al., 1987; Skoda et al., 1988). This degree of sequence conservation lies between those observed for cytochrome b5 (89%) and cytochrome P-450b (77%; Porter et al., 1986). Complete DNA coding sequences have been determined for rat (Porter et al., 1986) and human (Jackson et al., 1987; Skoda et al., 1988) liver mEH, and the complete structural gene for rat liver mEH has been isolated and partially characterized (Falany et al., 1987). Only a single gene coding for mEH is identified in the rat (Falany et al., 1987) or human (Skoda et al., 1988) genome, and gene mapping experiments have shown the mEH structural gene to be located on Chromosome 1 in both the mouse (Lyman et al., 1980; Simmons et al., 1985) and human (Skoda et al., 1988). A cDNA coding for human mEH has been transfected into yeast (Jackson and Burchell, 1988), and the transfected yeast produce a functional mEH with apparent molecular weight of 50 kDa that is recognized by antiserum to human mEH. The existence of multiple mEH-containing fractions eluting from ionexchange columns during enzyme purification procedures has produced speculation that multiple forms of mEH exist in a single tissue. In several cases, these different fractions had different kinetic and catalytic properties (Guengerich et al., 1979b; Bulleid et al., 1986; Moody and Hammock, 1987). Different immunochemical properties have been claimed for these different mEH fractions (Guengerich et al., 1979c). The most conclusive evidence suggests, however, that these multiple mEHs are in fact the same polypeptide chain associated with different amounts of phospholipid and are an artefact of membrane solubilization (Bulleid et al., 1986). The best evidence against the existence of multiple polypeptide chains is provided by the survey of a complete rat liver genomic library which demonstrated only a single gene coding for mEH (Falany et al., 1987). Significant post-translational modification of mEH cannot be rationalized, as the multiple forms have very similar molecular weights and are not known to contain a significant carbohydrate component. The chemical reaction catalyzed by mEH is hydrolysis of the oxirane ring by the trans-addition of water, with inversion of configuration at the attacked carbon atom (see Armstrong, 1987, for thorough review and discussion). The enzyme-catalyzed reaction is highly regio- and stereo-specific; this catalytic specificity has provided insight into the molecular mechanism of enzymic hydrolysis. Hydrolysis by mEH always proceeds with inversion of configuration. For rigid systems, such as arene oxides, this means that trans-dihydrodiols are the sole product (Jerina et al., 1970a; Armstrong, 1987). Enzymic hydrolysis of nonrigid cis-disubstituted oxiranes always yields threo-dihydrodiols (Watabe et al., 1971; Watabe and Akamatsu, 1972; Hanzlik et al., 1976), while hydrolysis of trans-disubstituted oxiranes yields erythro-dihydrodiols (Watabe and Akamatsu, 1972). This is primary evidence for general-base-catalyzed nucleophilic (SN2)

372 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 12.1. Participation of microsomal epoxide hydrolase (mEH) in the formation of a bay-region dihydrodiol epoxide from benzo[a]-pyrene. (A) Benzo[a]pyrene. (B) (7R, 8S)Benzo[a]pyrene-7, 8-oxide. The arrow indicates the preferred site of water addition. (C) 7R, 8R-Benzo[a]pyrene-7, 8-dihydrodiol. (D) 7R, 8R, 9S, 10R-Benzo[a]pyrene-7, 8dihydrodiol-9, 10-oxide.

addition of water to the oxirane ring, a mechanism that is consonant with all experimental observations. An acid-catalyzed mechanism, which would involve cleavage of the oxirane ring and carbocation formation prior to the addition of water, would be expected to result in a mixture of dihydrodiol enantiomers. In addition, the regio-selectivity of mEH predicts an SN2-type hydrolysis. Addition of water always occurs at the least hindered carbon atom, for both olefinic (Watabe et al., 1971, 1983; Hanzlik et al., 1976, 1978) and aromatic (Jerina et al., 1970a; Armstrong 1987) epoxides. Other investigations have used studies of hydrolysis in the presence of nucleophiles other than water (Hanzlik et al., 1976, 1978) and enzymic hydrolysis of mechanism-based inhibitors (Prestwich et al., 1985b), or substrates with a nucleophilic trap to preclude carbocation formation (Bellucci et al., 1981) as a crucial step in mEH-catalyzed hydrolysis. Studies using 2-bromo-4′-acetophenone to block histidine nitrogen implicate the necessary presence of histidine at the active site (DuBois et al., 1978). All experimental evidence suggests that unionized histidine at the active site partially deprotonates the attacking water molecule, enhancing its ability to act as an attacking nucleophile at the backside of the least hindered oxirane carbon atom (Dubois et al., 1978; Armstrong, 1987). The presence at the active site of an acidic amino acid which assists the reaction by partially protonating the oxirane oxygen and destabilizing the oxirane ring has been suggested (Hanzlik et al., 1976; DuBois et al., 1978) but not proven (Armstrong, 1987). mEH shows a high degree of both regio-selectivity and stereo-selectivity. Styrene oxide (Watabe et al., 1983) and naphthalene oxide (Jerina et al., 1970a) are always hydrated at the allylic oxirane carbon atom rather than at the relatively more hindered benzylic carbon atom. In the case of arene oxides formed from more complex polycyclic aromatic hydrocarbons, the same regioselectivity is evident. For simple non-K-region epoxides, the allylic carbon atom is always the preferred site of addition (see, for example, selective addition at C8 of benzo[a]-pyrene 7, 8-oxide, Figure 12.1; Armstrong, 1987). In the case of Kregion epoxides, where the two oxirane carbons are more-or-less sterically equivalent, hydration can occur at either position (Yang, 1988). Although regio-

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Figure 12.2. Stereo-selectivity of microsomal epoxide hydrolase (mEH) towards K-region epoxide substrates. (A) Enantiomeric benzo[a]pyrene 4, 5-oxides. The curved arrows indicate the preferred site of water addition, with the percent of corresponding dihydrodiol enantiomer obtained as indicated. (B) Enantiomeric benzo[c]phenanthrene 5, 6-oxides. The curved arrows indicate the preferred site of water addition, with the percent of corresponding dihydrodiol enantiomer obtained as indicated.

chemical preference is not a significant factor in the hydration of these epoxides, the enzyme nevertheless still exhibits stereo-specificity (Figure 12.2). It was originally suggested that in the case of many K-region epoxides the preferred site of hydration was at the carbon with an S-configuration, regardless of its position in the molecule (Figure 12.2A; Armstrong 1987). It was later pointed out, however, that this rule of thumb applies only for K-region epoxides derived from planar polycyclic hydrocarbons, such as benzo[a]pyrene 4, 5-oxide. In the case of some non-planar K-region oxides, such as benzo[c]phenanthrene 5, 6-oxide (Figure 12.2B), the R-centre is the preferred site of hydration. For other non-planar K-region oxides, hydration occurs at either centre to almost equal degrees (Yang, 1988). Thus while the enzyme exhibits a definite substrate enantio-selectivity, a simple, general rule of thumb to describe it cannot be formulated. Toxicological implications The toxicological significance of mEH is indicated by the large number of epoxides, which humans are directly exposed to or which are produced from environmental chemicals, that are metabolized to some degree by mEH. The list of known or potential epoxide substrates for mEH is extensive and includes those derived from chemicals encountered occupationally, such as styrene (Vainio et al., 1976), butadiene (Malvoisin and Roberfroid, 1982), vinyl chloride (Guengerich et al., 1979a), and epichlorohydrin (Rossi et al., 1983); epoxides derived from herbicides (Magdalou and Hammock, 1987) or insecticides (Brooks et al., 1970); and epoxides derived from therapeutic drugs, such as

374 CONJUGATION REACTIONS IN DRUG METABOLISM

phenytoin (Spielberg, 1984) and carbamazepine (Bellucci et al., 1987). An experimental system that both provides for modulation of the activities of mEH and other related enzymes and provides a measurable, unequivocal, toxic endpoint in vivo has been elusive. Because of the complex inter-relationships among toxic intermediate production, detoxication, and activity at target sites, perturbation of the activity of one enzyme, usually by a chemical agent, often has multiple effects in vivo. Furthermore, most of the toxic epoxides can be detoxified by multiple routes, specifically by glutathione conjugation (Chapter 11). In the face of these factors that complicate in vivo experimentation, several more simplified approaches have been derived for estimating the contribution of mEH versus that of other cytoprotective enzymes to the overall detoxication process. These include isolation of mEH-containing cell fractions from animals that have low glutathione levels, (Guenthner et al., 1980), and the use of more-or-less specific inhibitors of mEH activity in in vitro incubations (Oesch and Guenthner, 1983). These conditions can be applied in vitro, and relevant toxic endpoints can be measured. For most toxic epoxides genotoxicity is important and therefore mutagenicity to bacteria is often measured, as is the formation of covalent adducts with DNA. The role of mEH in altering the toxicity of several specific environmentally-derived epoxides has been studied, and the relative contributions of epoxide hydration and glutathione conjugation to detoxication have been estimated. For example, the genotoxicity of epichlorohydrin (chloromethyloxirane) in vitro is reduced under conditions of high mEH activity but not by added glutathione, implying that mEH, but not glutathione conjugation, plays the predominant role in detoxication in vivo (Rossi et al., 1983). By contrast, trichloropropene oxide, a compound very similar in structure to epichlorohydrin, is primarily metabolized by glutathione conjugation and only to a very small degree by mEH, suggesting that for this compound mEH does not play a significant role in detoxication but rather glutathione conjugation is the primary mechanism (Sinsheimer et al., 1987). mEH plays only a minor role in the detoxication of aflatoxin B1 5, 6-oxide, the genotoxic species derived from the hepatocarcinogen aflatoxin B1; in this case, glutathione conjugation is the major protective process in vivo (Monroe and Eaton, 1987, Jhee et al., 1988). In the case of K-region epoxides of polycyclic aromatic hydrocarbons (examples shown in Figure 12.2), addition of purified mEH to the test system reduces either mutagenicity (Glatt et al., 1983) or DNA binding (Guenthner and Oesch, 1981a; Oesch and Guenthner, 1983). Conversely, inhibition of mEH activity produces an increase in the DNA binding of K-region epoxides (Guenthner and Oesch, 1981a, Oesch and Guenthner, 1983), suggesting that mEH plays a major detoxifying role in vivo. Glutathione conjugation also plays a major role in the detoxication of K-region oxides, expressed as a decrease in mutagenicity (Glatt et al., 1983) or as a decrease in covalent binding of the K-region epoxides to

EPOXIDE HYDROLASES 375

DNA (Guenthner et al., 1980, Hesse et al., 1980). This contrasts with the detoxication of another class of epoxides formed from polycyclic aromatic hydrocarbons, the bay-region dihydrodiol epoxides. In this case, in vitro experiments similar to those described for the K-region oxides demonstrate that mEH plays a role in the formation, but almost no role in the detoxication of these bay-region dihydrodiol epoxides (Figure 12.1; Guenthner and Oesch, 1981a; Oesch and Guenthner, 1983; Oesch, 1984). On the other hand, the presence of glutathione and/or additional purified glutathione S-transferases in the test system diminishes both mutagenicity (Glatt et al., 1983) and DNA binding of these intermediates (Guenthner et al., 1980; Hesse et al., 1980), suggesting that glutathione conjugation is a major detoxication mechanism. 12.3. Cytosolic epoxide hydrolase Early studies of mEH in microsomes also noted that a relatively small amount of hydrolytic activity could be demonstrated in the cytosolic cell fraction (Maynert et al., 1970; Oesch et al., 1971a). At the time, the significance of this cytosolic enzyme activity was disregarded, perhaps as an artefact of cell fractionation. Definitive evidence of the existence of an epoxide hydrolase located primarily in the cytosol rather than in the endoplasmic reticulum was first obtained in the course of studies on the mammalian metabolism of synthetic insect juvenile hormone mimetics which contain an epoxide moiety (Gill et al., 1974). These alkyl epoxides are rapidly hydrolyzed by liver cytosol at neutral pH but are quite resistant to hydrolysis by microsomal enzymes. As the nature of cytosol-catalyzed epoxide hydrolysis was further studied, it became apparent that it was attributable to a unique cytosolic epoxide hydrolase (cEH) with catalytic and structural properties quite different from those of the, by now, well-characterized mEH. Substrates Like mEH, cEH metabolizes synthetic and endogenous epoxides with a fairly broad spectrum of chemical structures. The two epoxide hydrolases are in some ways complementary in that many epoxides that are poor substrates for mEH are good substrates for cEH, and vice versa. mEH is known to metabolize monosubstituted oxiranes well, and tri- or tetrasubstituted oxiranes poorly. cisDisubstituted oxiranes are metabolized well by mEH, while trans-disubstituted oxiranes are poorly metabolized (Oesch et al., 1971b). By comparison, cEH also metabolizes many mono- and disubstituted oxiranes well (Mumby and Hammock, 1979; Gill et al., 1983a; Hammock and Hasagawa 1983). The substituted geranyl epoxide insect hormone mimetic shown in Figure 12.3C, is a

376 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 12.3. Substrates and inhibitors for cytosolic epoxide hydrolase (cEH). (A) transStilbene oxide. (B) trans-β-ethylstyrene oxide. (C) 1-(4′-ethylphenoxy)-3, 7-dimethyl-6, 7-epoxy-trans-2-octene (ethyl epoxide). (D) cis-epoxymethylstearate. (E) 5(S)-cis-5, 6oxido-8, 11, 14-cis-eicosatrienoic acid (5, 6-epoxyeicosatrienoic acid, 5, 6-EET). (F) Trichloropropene oxide. (G) Chalcone oxide. (H) 4′-Phenylchalcone oxide. (I) 2Amino-4-(4′-hydroxyphenyl)-5-phenylthiazole. (J) Nordihydroguaiaretic acid.

trisubstituted oxirane that is metabolized approximately ten times faster by cEH than by mEH (Gill et al., 1974). Investigation of a series of β-substituted styrene oxides, comprising both cis- and trans-disubstituted oxiranes, showed that for all ‘geometric isomer pairs, the trans-compound was metabolized more rapidly by cEH (Gill et al., 1983a; Hammock and Hasagawa, 1983). Rate of metabolism also increases with hydrophobicity of the β-substituent; trans-β-propylstyrene oxide is metabolized most rapidly, followed by trans-β-ethyl-(Figure 12.3B), trans-β-phenyl-(Figure 12.3A), trans-β-methyl- and cis-β-phenyl- and cis-βmethylstyrene oxides. However, many cis-disubstituted oxiranes are readily metabolized by cEH. In the case of epoxy geranyl ether compounds analogous to compound 3C, cis-geometric isomers at the oxirane ring are favoured over trans (Gill et al., 1974). Epoxides of long chain fatty acids are rapidly hydrolyzed by cEH; those with either cis or trans configurations at the oxirane ring are good substrates. Epoxyeicosatetraenoic acids (EET) analogous to Figure 12.3E are cisdisubstituted oxiranes that are generally excellent substrates for cEH (Chacos et al., 1983, see below). Cis- and trans-isomers of epoxymethylstearate (Figure 12.3D) are metabolized by cEH with equal facility (Gill and Hammock, 1979). Substrate hydrophobicity seems to be important. A series of epoxycycloalkanes, ranging in size from cyclopentene oxide to cyclododecene oxide were tested as cEH substrates (Magdalou and Hammock, 1988). No

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activity was seen for compounds smaller than cyclooctene oxide and maximal activity was seen for cyclodecene oxide. In contrast, the smaller compounds (C7 and C8) were the best mEH substrates. The selectivity of cEH for trans-over cissubstituted styrene oxides and the complementary selectivity of mEH for the cisisomers has led to the use of a number of these compounds as selective substrates for the measurement of cEH activity. trans-β-Ethylstyrene oxide (Figure 12.3B) has been used due to its rapid rate of metabolism by cEH, but trans-stilbene oxide (Figure 12.3A) is preferred due to its low volatility and resistance to spontaneous hydrolysis in aqueous solution. Styrene oxide, often used to assay mEH activity, is not particularly useful as a means of selectively measuring mEH activity in the presence of cEH activity because it is well hydrolyzed by both enzymes (Gill et al., 1983a; Hammock and Hasagawa, 1983). Metabolism of arene oxides by cEH is practically nonexistent (Hammock and Hasagawa, 1983). Therefore substrates like benzo[a]pyrene 4, 5-oxide are very useful in selectively determining mEH activity in the presence of cEH, while trans-stilbene oxide is very selective for cEH in the presence of mEH. Inhibitors In accord with the many differences in substrate specificity betwen mEH and cEH, differences in sensitivity to inhibitors also exist. Cyclohexene oxide and trichloropropene oxide (Figure 12.3F), two commonly employed inhibitors of mEH, are very weak inhibitors of cEH (Mullin and Hammock, 1982; Magdalou and Hammock, 1988). In two related studies, 41 compounds (Hammock and Hasagawa, 1983) and 150 compounds (Mullin and Hammock, 1982) were screened as inhibitors of cEH. As is the case for mEH, poorly metabolized epoxides proved to be the best inhibitors. Chalcone oxide (Figure 12.3G) inhibits cEH with an I50 of 8 µM but is not hydrolyzed by the enzyme (Mullin and Hammock, 1982). Substituted analogues of this compound proved to be the most potent cEH inhibitors; 4′-phenylchalcone oxide (Figure 12.3H) inhibits with an I50 of 700 nM (Mullin and Hammock, 1982). Chalcone oxides only slightly inhibit mEH at the highest achievable concentrations (Mullin and Hammock, 1982; Guenthner, 1986a), and in fact in some cases activate mEH activity in vitro (Seidegård et al., 1986). Substituted (non-epoxide) chalcones also inhibit cEH activity but at higher concentrations than the corresponding epoxides (Mullin and Hammock, 1982; Hammock et al., 1983). Other nonepoxide inhibitors of cEH have been identified. α-Phenylketones, including the chalcone oxides, as well as various benzo- and acetophenones and flavones (specifically naphthoflavones; Meijer and DePierre, 1985b) generally inhibit cEH activity (Mullin and Hammock, 1982; Hammock and Hasagawa, 1983). A series of 15 α-β-epoxyketones, again including chalcone oxides, were tested and all proved to be effective inhibitors (Prestwich et al., 1985a). Some sulfhydryl

378 CONJUGATION REACTIONS IN DRUG METABOLISM

reagents, including organomercurials (Meijer and DePierre, 1985b; Prestwich et al., 1985a) and inorganic cations (Gill and Hammock, 1980), are good cEH inhibitors. Cyclopropyl oxiranes are a specifically designed and synthesized family of suicide inhibitors of cEH (Prestwich et al., 1985b). Their covalent binding to, and deactivation of, the active site of cEH is mediated by formation of a transient reactive intermediate during enzyme-catalyzed oxirane ring opening. 7-Methoxycitronellyl thiol is a non-epoxide, weak inhibitor of cEH that has been used as an affinity-column ligand to effect enzyme purification (Prestwich and Hammock, 1985; Hammock et al., 1986). A number of commonly used laboratory solvents inhibit cEH activity at concentrations of 1– 5%; ethyl acetate, tetrahydrofuran and acetonitrile are the most effective inhibitors (Mullin and Hammock, 1982; Meijer and DePierre, 1985b). A final interesting group of non-epoxide compounds that inhibit cEH activity in vitro are compounds first recognized as inhibitors of catalase activity. These compounds, including the hydroxylated metabolite (Figure 12.5I) of 2-amino-4, 5-diphenylthiazole (Figure 12.3I) and nordihyd-roguaiaretic acid (Figure 12.3J), inhibit cEH activity with approximately the same potency with which they inhibit catalase (Guenthner et al., 1988a, 1988b). Enzyme purification and mechanism Enzyme inhibitor studies have been helpful in characterizing the mechanism of cEH-catalyzed hydrolysis and in comparing it to the mechanism of mEH. As is the case for mEH, hydrolysis occurs by the backside attack by water at the least hindered oxirane carbon (Hammock et al., 1980), with resulting trans addition and inversion of configuration at that carbon (Mumby and Hammock, 1979). Enzymic hydration of cis-disubstituted epoxides yields threo diols, while hydration of trans-disubstituted epoxides yields erythro diols (Mumby and Hammock, 1979). In the case of mEH, an active site histidine group is thought to participate in increasing the nucleophilicity of the attacking water species (DuBois et al., 1978). In the case of cEH, however, compounds that inactivate amine groups, such as picrylsulfonic acid, or 2′-bromo-6-nitroacetophenone, are not particularly potent inhibitors of hydrolysis (Mullin and Hammock, 1982), indicating the absence of a crucial amino group at the active site. Sulfhydrylbinding reagents are potent inhibitors of cEH, however, indicating that a cysteine thiol may play a role in activating the hydrating species (Mullin and Hammock, 1982). Evidence obtained with studies using α-β-epoxyketone inhibitors (Prestwich et al., 1985a) provides support for existence of an acidic moiety at the active site of cEH that protonates and activates the oxirane oxygen, facilitating C-O bond cleavage. Thus while general similarities in the mechanism of hydration by cEH and mEH exist, the two are not identical.

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cEH has been purified from a number of different sources, including mouse (Gill, 1983; Meijer and DePierre, 1985a, 1985b; Prestwich and Hammock, 1985; Hammock et al., 1986), rabbit (Waechter et al., 1982), rat (Schladt et al., 1988a), and man (Wang et al., 1982; Silva and Hammock, 1987; Guenthner et al., 1988a), and its physicochemical properties have been well characterized (for review see Meijer and DePierre, 1988). It has a native molecular weight of approximately 120 kDa and is a dimer comprising two apparently identical 60 kDa subunits. It is an acidic protein with a pI in the 5.0–6.1 range. Its amino acid composition has been characterized and varies among species (Waechter et al., 1982; Meijer and DePierre, 1985a; Hammock et al., 1986; Guenthner et al., 1988a), but no N-terminal sequence has been determined as the N-terminus is blocked (Meijer and DePierre, 1988). Its structural distinction from mEH is apparent upon comparison of amino acid compositions of the two enzymes. Accordingly, antibodies prepared to mEH do not cross- react with cEH and can be used to distinguish the two enzymes (Guenthner et al., 1981; Guenthner and Oesch, 1983). By the multiple criteria of substrate and inhibitor selectivity (see above), structural and immunochemical properties, and evidence of genetic control arrived at through induction studies, a complete distinction between mEH and cEH can be drawn. The distribution of cEH in different organs and tissues and the degree of difference in cEH levels among different species have been extensively studied and have been thoroughly discussed in a recent review article (Meijer and DePierre, 1988). A few salient points will be mentioned briefly here. Most surveys of cEH activities in liver cytosol derived from different laboratory animal species have concluded that cEH activity is highest in mouse, intermediate in rabbit or guinea pig, and very low in rat (Gill and Hammock, 1980; Ota and Hammock, 1980; Gill et al., 1983a; Meijer et al., 1987; Meijer and DePierre, 1988). The fact that cEH activity is very low in rat liver, the source of tissue most commonly employed in drug metabolism studies, probably accounts for the initial conclusions that epoxide hydrolysis was primarily catalyzed by microsomal enzymes and that cytosolic activity was negligible. Human cEH activity levels lie between those of mouse and rat and are comparable to those of guinea pig (Gill et al., 1983b; Mertes et al., 1985; Guenthner and Karnezis, 1986a). It should be noted that some microstructural differences in cEH proteins isolated from different species exist. Antiserum to purified mouse liver cEH cross-reacts with hepatic cytosol from mouse, rat, and hamster but not from guinea pig, rabbit, monkey, or man (Meijer et al., 1987). Similarly, antiserum to purified human cEH cross-reacts weakly with a purified cEH preparation derived from guinea pig liver (Guenthner et al., 1988a), and antiserum to rat liver cEH cross-reacts with a cytosolic protein from mouse, guinea pig, hamster and rabbit but not from monkey (Schladt et al., 1988a). The amino acid compositions of purified cEH derived from mouse, rabbit and man differ markedly (Waechter et

380 CONJUGATION REACTIONS IN DRUG METABOLISM

al., 1982; Meijer and DePierre, 1985a; Hammock et al., 1986; Guenthner et al., 1988a). In mouse, the animal with the highest reported levels of hepatic cEH, the rank order of organ distribution has been reported as liver>kidney>lung>testis>spleen (Gill and Hammock, 1980; Loury et al., 1985). In rat, an animal with very low hepatic cEH levels, a different order is seen, namely, heart=kidney>liver> brain>lung>testis>spleen (Gill et al., 1983b). In humans, specific cEH activity is approximately the same in lung and liver (Guenthner and Karnezis, 1986a). cEH activity has also been measured in human mononuclear leukocytes (Seidegård et al., 1984) and in human foetal organs (Pacifici et al., 1988). In homogenized cell and tissue preparations subjected to differential centrifugation, the preponderance of cEH is found in the cytosolic (100000×g supernatant) fraction (Gill et al., 1974; Gill and Hammock, 1979; Ota and Hammock, 1980). Smaller amounts of cEH-like activity are also found in microsomal (Guenthner and Oesch, 1983) and nuclear membrane (Guenthner, 1986b) fractions. Some substrates, like styrene oxide (El-Tantawy and Hammock, 1980; Gill et al., 1983a), or the cis-epoxide of stearic acid (Gill and Hammock, 1979; Watabe et al., 1983), are hydrolyzed by both cytosolic and microsomal enzymes in varying ratios. Such observations are probably attributable to lack of complete substrate selectivity by mEH and cEH. However, metabolic and immunochemical studies have shown that integral membrane proteins exist with catalytic and structural properties like those of cEH and unlike those of mEH, leading to the conclusion that membrane-bound forms of cEH exist (Guenthner and Oesch, 1983). This particularly appears to be the case in human tissues where the distinction between microsomal and cytosolic forms of both cEH and mEH almost disappears. In both human lung and liver, substantial amounts of mEH-like activity are found in the cytosolic fraction and substantial amounts of cEH-like activity are found in microsomes (Wang et al., 1982; Gill et al., 1983b; Guenthner and Karnezis, 1986a, 1986b; Schladt et al., 1988b). Obviously the designation of such activities as microsomal epoxide hydrolase or cytosolic epoxide hydrolase ceases to be semantically precise in such a case. Early studies also observed high amounts of mouse hepatic cEH activity sedimenting with heavy and light mitochondrial fractions. (Gill and Hammock, 1980, 1981a, 1981b). Although it was originally concluded that this activity was mitochondrial in origin, later studies demonstrated that this activity could be attributable to enzymes in peroxisomes that co-sedimented with the mitochondrial fractions (Hammock and Ota, 1983; Waechter et al., 1983; Loury et al., 1985; Kaur and Gill, 1986). In fact, the high specific activity of cEH in peroxisomes, coupled with the observation that cEH levels are induced by compounds that cause peroxisomal proliferation, has led to the tentative conclusion that cEH in situ may be a peroxisomal enzyme which artefactually appears in the cytosol due to lysis of peroxisomes during cell

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homogenization (Hammock and Ota, 1983; Waechter et al., 1983; Loury et al., 1985; Meijer and DePierre, 1988). Induction Induction studies have strengthened the argument that cEH is peroxisomal in origin. Several dozen compounds of diverse chemical structure, many of which are known to be good inducers of mEH or other microsomal xenobioticmetabolizing enzymes, have been assayed as possible inducers of cEH activity in vitro (see Meijer and DePierre, 1988). All compounds that induce cEH also increase both the number and size of hepatic peroxisomes. These peroxisomeproliferating agents that also enhance the levels of cEH recovered in cytosol include such hypolipidemic agents as clofibrate (Hammock and Ota, 1983; Loury et al., 1985; Schladt et al., 1986; Meijer and DePierre, 1987), nafenopin (Waechter et al., 1984), 1-benzylimidazole, tiadenol, and fenofibrate (Schladt et al., 1987). Probucol, an antihypercholesterolemic drug that does not cause peroxisomal proliferation, also does not induce cEH levels (Hammock and Ota, 1983; Schladt et al., 1987). The inductive effects of these agents appear, therefore, to be linked to their ability to enhance peroxisome synthesis rather than to be secondary to their hypolipidemic effects. A number of environmental compounds also coordinately induce peroxisome levels and cEH activity, including the plasticizer di-(2-ethylhexyl)-phthalate (Hammock and Ota, 1983) and structurally related compounds (Hammock and Ota, 1983; Meijer and DePierre, 1987, 1988; Lundgren et al., 1987b), and several halogenated herbicides (Lundgren et al., 1987a; Moody and Hammock, 1987). In the case of one of these latter compounds, 2, 4-dichlorophenoxyacetic acid, immunochemical quantitation of enzyme protein levels showed that the enhancement of enzyme activity was due to a true ‘induction’, that is, an increase in the total amount of cEH protein in the cell (Lundgren et al., 1987a). The cEH proteins isolated from untreated animals or from animals whose cEH levels are enhanced by clofibrate treatment are identical in all respects (Loury et al., 1985; Meijer and DePierre 1985a; Hammock et al., 1986). Compounds that neither induce cEH nor cause proliferation of peroxisomes include phenobarbital, 3-methylcholanthrene, 2, 3, 7, 8-tetrachloro-p-dibenzodioxin, Arochlor 1254, and 2-acetylaminofluorene (Hammock and Ota, 1983; Schladt et al., 1986; Meijer and DePierre, 1987). In no case did a compound induce cEH levels without concomitantly inducing peroxisome proliferation. This evidence indicates that induction of cEH levels in vivo is due to the increased synthesis of peroxisomes and peroxisomal enzymes. cEH levels in animals treated with peroxisome proliferators are not increased to any greater degree, and in some cases to a lesser degree, than levels of other peroxisomal enzymes (Moody and Hammock, 1987; Lundgren et al., 1987a; Schladt et al., 1987). Thus, the evidence

382 CONJUGATION REACTIONS IN DRUG METABOLISM

would indicate that these cEH-inducing compounds probably do not activate a cEH-specific receptor or a single genetic locus specifically controlling cEH levels, but rather act in a general fashion to increase synthesis of a large number of peroxisomal proteins which includes, but is not limited to, cEH. Biological role The putative peroxisomal origin of cEH may provide some insight into its overall functional role in the cell. It has been proposed (Chacos et al., 1983; Hammock and Hasagawa, 1983) that lipid epoxides, specifically epoxides formed from unsaturated fatty acids, may be the natural endogenous substrates of cEH. For example, both cis- and trans-epoxymethylstearate are metabolized by cEH (Gill and Hammock, 1979). The four epoxide derivatives of arachidonic acid (5, 6-, 8, 9-, 11, 12-, or 14, 15-epoxyeicosatetraenoic acid, EET; Figure 12.3E) are all substrates for cEH (Chacos et al., 1983). The Km of 14, 15-EET for cEH is 8 µM, which is only slightly higher than that of the model substrate transstilbene oxide (Chacos et al., 1983). Specific activities of purified cEH using the four EETs as substrates range from 69 nM min−1 mg−1 for 5, 6-EET to 1260 nM min−1 mg−1 for 14, 15EET (Chacos et al., 1983). By comparison, the specific activity of the pure enzyme for trans-stilbene oxide hydrolysis is approximately 900 nM min−1 mg−1 (Hammock et al., 1986). Two observations suggest that the ability of the cell to metabolize fatty acid epoxides may be toxicologically or physiologically significant. The first observation is that these compounds may be formed under conditions conducive to lipid peroxidation (Sevanian et al., 1979). Peroxisomal oxidation of lipids produces hydrogen peroxide as a byproduct; therefore under conditions of high peroxisomal activity, increased rates of lipid peroxidation and fatty acid epoxide formation might be expected. Fatty acid epoxides (EETs in particular) have multiple diverse hormonal activities; EETs have been reported to be vasodilators (Proctor et al., 1987), to inhibit ATPase (Schwartzman et al., 1985), to promote calcium release from the endoplasmic reticulum (Kutsky et al., 1983), and to serve as releasing factors for a number of hypothalamic (Falck et al., 1983), pituitary (Cashman et al., 1987), and pancreatic (Capdevila et al., 1983) peptide hormones. cEH might be thought of, therefore, as a protective enzyme which is present in peroxisomes to protect the cell from the possible deleterious effects of fatty acid epoxide formation arising from an overproduction of hydrogen peroxide. In this regard, cEH can be compared to another peroxisomal enzyme, catalase, which also is protective against high levels of hydrogen peroxide. A related, as yet unanswered, question, is whether fatty acid epoxides might also be toxicologically significant as endogenously generated electrophiles. One study has shown that several fatty acid epoxides are not mutagenic in the Ames test

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(Sevanian and Peterson, 1986), but extensive study of the possible genotoxicity of this family of compounds has not yet been carried out. Two final functions of cEH in the cell bear mentioning. The first relates to its ability to convert leukotriene A4 (Figure 12.5A) to a vicinal 5(S), 6(R)-diol (Figure 12.5D; Haeggström et al., 1986). This enzymic pathway provides a third alternative metabolic fate for LTA4, in addition to its conversion to LTB4 or LTC4. Two different studies using rat liver or rat kidney fractions demonstrated that the 5(S), 6(R)-diol formed by cEH is, in fact, the major metabolite formed when LTA4 is incubated with cytosol (Medina et al., 1987; Haeggström et al., 1988b). cEH appears, therefore, to play a significant role in the control of the leukotriene metabolic pathway by providing an alternative route for LTA4 catabolism. The 14, 15-epoxy analogue of LTA4 (14, 15-LTA4) is also readily hydrolyzed by cEH to the vicinal diol (Wetterholm et al., 1988). An additional role for cEH, also related to eicosanoid metabolism, has been suggested by studies showing that lipoxin A, a physiologically active eicosanoid triol shown in Figure 12.5K, is formed by enzymic hydrolysis of a 5, 6-epoxide precursor. The regio- and stereo-selectivity of this enzymic hydrolysis, as well as its catalysis by a semi-purified cEH preparation, indicate that cEH is the enzyme primarily responsible for lipoxin A formation (Puustinen et al., 1986). Thus, while cEH plays no direct role in the hydrolytic conversion of LTA4 to LTB4, its importance in multiple pathways of eicosanoid metabolism is becoming increasingly apparent. 12.4. Experiments in isolated cells, perfusions and in vivo The vast majority of studies characterizing the toxicological and physiological role (s) of epoxide hydrolases have utilized broken cell preparations, isolated subcellular fractions, and partially or totally purified enzyme preparations. Studies using whole animals, isolated perfused organs, or whole cells in which epoxide hydrolase activities are manipulated and physiological or toxicological endpoints are measured appear infrequently in the literature. This may be due in part to the absence of methods for cleanly and selectively altering epoxide hydrolase levels in vivo. For example, attempts have been made to assess the protective role of mEH against dermatocarcinogenic aromatic hydrocarbon epoxides by co-administering trichloropropene oxide (thereby inhibiting mEH) along with the carcinogen 3-methylcholanthrene (Bürki et al., 1974). Although co-administration of trichloropropene oxide enhances carcinogenesis, it is difficult to ascribe this effect merely to its inhibition of the cytoprotective effects of mEH, as trichloropropene oxide itself has genotoxic properties (Sinsheimer et al., 1987). Likewise, induction of mEH levels in vivo by compounds such as trans-stilbene oxide does not permit clean, selective

384 CONJUGATION REACTIONS IN DRUG METABOLISM

manipulation of mEH activity. No truly selective inducer of mEH activity exists; carcinogen-activating enzymes (e.g. specific cytochrome P-450 isozymes) as well as deactivating enzymes are simultaneously induced (Bücker et al., 1979). Data gathered from surveys of mEH activity in different strains and species have led to inferences regarding its overall protective role in the whole animal. Mice are highly susceptible to spontaneous tumour formation, and in general have the lowest mEH levels of any species (Walker et al., 1978; Glatt and Oesch, 1987). Individual mouse strains with the lowest mEH levels (Walker et al., 1978) are the most susceptible to benzo[a]pyrene hepatocarcinogenesis (Glatt and Oesch, 1987). The validity of the hypothesis that susceptibility to carcinogenesis is primarily, or even partially, determined by mEH levels is cast in doubt by studies showing that highly hepatocarcinogen-susceptible male C3H×A/J mice and highly hepatocarcinogen-resistant female C3H×A/J mice have indistinguishable mEH (or glutathione transferase or cytochrome P-450) levels (Glatt and Oesch, 1987). Studies with freshly isolated hepatocytes have been used to investigate the role of mEH in determining the level of covalent binding of benzo[a]pyrenederived epoxides to DNA. The mEH inhibitor trichloropropene oxide (Figure 12.3F) increases the total amount of benzo[a]pyrene-derived DNA adducts in whole cells (Burke et al., 1977) and specifically enhances the binding of K-region oxides (Jernström et al., 1978). However, these results must be interpreted with the realization that trichloropropene oxide may also inhibit specific glutathione S-transferases and therefore may have multiple effects in the cell. Similar conclusions that mEH prevents the genotoxicity of K-region oxides in vivo are drawn from host-mediated mutagenesis studies. In these experiments, tester bacteria are injected into a mouse host with a mutagen which is activated and/or inactivated by host enzymes. The bacteria are then recovered and allowed to form colonies in vitro. In such studies, mice were able to totally deactivate benzo[a]pyrene 4, 5-oxide, preventing its ability to cause mutation of bacterial DNA (Glatt et al., 1985). It may be assumed that mEH plays a major role in this deactivation, but it should be recognized that benzo[a]pyrene 4, 5oxide is also rapidly deactivated by specific glutathione S-transferases (Bend and Smith, 1979). The best assessments of the relative contributions of individual detoxication pathways (including mEH) in the deactivation of cytotoxic epoxides are offered by isolated organ perfusion studies. These studies demonstrate that for both liver and lung, benzo[a]pyrene 4, 5-oxide is converted to the diol and to the glutathione conjugate at approximately equal rates, suggesting that both detoxifying systems contribute significantly to detoxication of this epoxide, with the role of mEH especially important under conditions of glutathione depletion (Smith and Bend, 1979; Smith et al., 1980). The detoxication of styrene oxide in the isolated perfused liver has also been studied. In normal livers, the ratio of thioether to dihydrodiol metabolites ranges from 1/ 1 to 2/1 (VanAnda et al., 1979). However, in livers that are depleted of cellular

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Figure 12.4. Cholesterol 5, 6-oxides and chemically-related compounds. (A) Cholesterol 5, 6α-oxide. (B) Cholesterol 5, 6β-oxide. (C) Cholestane 3β-5α-6β-triol. (D) 5, 6α-imino-5α-cholestane-3β-ol. (E) 7-Dehydrocholesterol 5, 6β-oxide.

glutathione, mEH activity alone is sufficient to prevent the cytotoxic effects of styrene oxide (Smith et al., 1983). These results again suggest that both pathways are important and that mEH may be particularly important when cellular glutathione levels are low. 12.5. Endogenous substrates Cholesterol 5, 6-oxide hydrolase Cholesterol 5, 6-oxide hydrolase (ChEH) is a microsomal epoxide hydrolase that catalyzes the hydration of cholesterol 5, 6α-oxide (ChαO; Figure 12.4A) and cholesterol 5, 6β-oxide (ChβO; Figure 12.4B), as well as several other steroid 5, 6-oxides by trans-addition of water. In the case of both cholesterol epoxides, a single identical stereoisomer results from enzymic hydrolysis, namely, cholestane 3β, 5α, 6β-triol (Figure 12.4C) (3, 5, 6-triol; Aringer and Eneroth, 1974; Watabe et al., 1981; Nashed et al., 1985). This epoxide hydrolase specifically hydrolyzes steroid 5, 6-oxide substrates and is demonstrably distinct, both structurally and catalytically, from mEH. Cholesterol 5, 6-oxides, and the enzyme that catalyzes their hydrolysis, are of interest because of their ubiquitous presence in trace amounts as products of cellular lipid peroxidation and because they have been demonstrated, albeit somewhat controversially, to be cytotoxic and mutagenic. Gray et al., (1971) isolated and identified ChαO from human serum extracts, while Black and co-

386 CONJUGATION REACTIONS IN DRUG METABOLISM

workers demonstrated the presence of this compound in skin after irradiation by ultraviolet light (Black and Douglas, 1972; Chan and Black, 1974) as well as in rodent liver (Black and Laseter, 1976). Cholesterol 5, 6-oxides are formed from cholesterol in in vitro systems containing microsomes, oxygen, ferrous iron, and NADPH or ATP by lipid peroxidation (Johansson, 1971; Mitton et al., 1971; Mitropoulos and Balasubramaniam, 1972; Aringer and Eneroth, 1973, 1974), and are not formed by cytochrome P-450-dependent oxygenation of cholesterol (Mitropoulos and Balasubramaniam, 1972; Watabe et al., 1980b). The enzyme The enzymic hydrolysis of ChαO and ChβO to form the 3, 5, 6-triol was demonstrated almost simultaneously with the first demonstration of the presence of cholesterol epoxides in biological samples. Feeding ChαO to rats resulted in enhanced 3, 5, 6-triol levels in intestinal lipids (Fioriti et al., 1970). 3, 5, 6-Triol formation from ChαO in vitro requires a microsomal enzyme (Mitton et al., 1971; Martin and Nicholas, 1973; Aringer and Eneroth, 1974). This enzyme was shown to be a classic epoxide hydrolase (Watabe et al., 1980a, 1980b; Nashed et al., 1985) located primarily in the endoplasmic reticulum with minor amounts observed in mitochondrial cell fractions, but not in cytosol (Aringer and Eneroth, 1974; Åström et al., 1986; Sevanian and McLeod, 1986). It has been found in all mammalian species investigated, including rat (Aringer and Eneroth, 1974), mouse (Black and Lenger, 1979), cow (Watabe and Sawahata, 1979), rabbit, hamster and man (Nashed et al., 1985). The highest hepatic activity is found in rat and human, the lowest in rabbit. A survey of its organ distribution in rat (Åström et al., 1986) showed the highest specific activity in liver microsomes. Accordingly, the preponderance of total activity is found in liver, followed in order of much smaller amounts by intestine>testis=kidney>brain>lung>spleen. The enzyme has also been found in skin (Chan and Black, 1974, 1976) and in human intestinal flora (Hwang and Kelsey, 1978). ChEH can be distinguished from mEH on both a catalytic basis, using diagnostic substrates and inhibitors, and on a structural basis, using mEHspecific antibodies. Purified mEH or cEH preparations have no activity toward ChαO or ChβO, demonstrating that all such activity in the microsomes is attributable to ChEH (Levin et al., 1983; Oesch et al., 1984; Kaur and Gill, 1986). Several compounds that are known to either inhibit or be good substrates for mEH, including octene 1, 2-oxide, benz[a]anthracene 5, 6-oxide and styrene oxide, have no effect on ChEH activity (Nashed et al., 1985; Sevanian and McLeod, 1986). Trichloropropene oxide (Figure 12.3F) does not inhibit but rather enhances ChEH activity to varying degrees (Watabe et al., 1980b; Oesch et al., 1984). A series of heterocyclic compounds that enhance mEH activity in

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vitro, including ellipticine, harmane, and norharmane, inhibit ChEH activity in a non-competitive manner (Palakodety et al., 1987). Similarly, compounds that specifically inhibit ChEH activity, such as 5, 6α-imino-5α-cholestane-3β-ol (iminocholestanol; Figure 12.4D) or 7-ketocholestanol exhibit no effect on mEH activity (Watabe et al., 1980b, 1983; Sevanian and McLeod, 1986). A mechanism-based suicide inactivator of ChEH, 7-dehydrocholesterol 5, 6βoxide (Figure 12.4E) does not bind to, nor deactivate, a purified mEH preparation (Nashed et al., 1986). Some inducers of mEH and/or cytochrome P-450, including trans-stilbene oxide and Arochlor 1254, slightly decrease ChEH activity, while other compounds, including phenobarbital, 2acetylaminofluorene, and ChαO itself, have no effect (Levin et al., 1983). None of the compounds that induce mEH activity co-induce ChEH activity (Levin et al., 1983). The peroxisome-proliferating agents, clofibrate and ciprofibrate, compounds that induce cytosolic epoxide hydrolase (cEH) activity (see above), produce a 75% increase in ChEH levels in rabbit and mouse liver microsomes, whereas no concomitant increase in mEH levels is seen. Induction of ChEH by these compounds is observed in rat kidney, out not in rat liver (Finley and Hammock, 1988). Antibodies to mEH remove from detergent-solubilized microsomes all mEH activity toward styrene oxide or benz[a]anthracene 5, 6oxide, without affecting ChEH activity (Levin et al., 1983; Oesch et al., 1984). Finally, mEH and ChEH activities elute in different fractions during chromatography (Watabe et al., 1986). It is therefore readily apparent that ChEH is distinct from cEH and from mEH. Substrate specificity ChEH appears to have a narrow substrate specificity; the only substrates known at this time are steroid 5, 6-oxides. This restricted structural requirement for interaction with the enzyme’s catalytic site is further evidenced by the fact that only analogues of cholesterol 5, 6-oxide(s) have, as yet, proven to be competitive inhibitors of ChEH activity. Known substrates for ChEH include ChαO, ChβO, and the 5, 6α- and 5, 6β-oxides of 5-cholestene, 20-methyl-5pregnen-3β-ol, and pregnenolone (Watabe et al., 1981, 1983). In the cases of all four steroids, both the α-Oxide and the β-oxide are converted to the same product, the corresponding 3β, 5α, 6β-triol. Although it has been inferred that cisepoxymethylstearate is a substrate for ChEH (Sevanian et al., 1980), later evidence has shown this not to be the case (Watabe et al., 1983; Nashed et al., 1985). Kinetics of the enzymic hydration of ChαO and ChβO have been studied in several laboratories, with somewhat varying results. The enzyme exhibits optimal activity in the pH range 7.0–7.4 (Aringer and Eneroth, 1974; Black and Lenger, 1979; Sevanian et al., 1980). Estimates of the Michaelis constant for ChαO hydrolysis range from 1–375 µM (Black and Lenger, 1979; Sevanian et

388 CONJUGATION REACTIONS IN DRUG METABOLISM

al., 1980; Nashed et al., 1985; Sevanian and McLeod, 1986). The discrepancy in these results may be due to the limited water solubility of this lipophilic substrate. The solubility limit for ChαO in one aqueous system has been estimated at 62 µM (Sevanian and Peterson, 1984). Accordingly, the manner and solvent in which the epoxide is added have a major effect on kinetic estimates (Levin et al., 1983; Sevanian and McLeod, 1986). The amount of microsomal lipid also greatly affects hydrolysis kinetics; the Km for ChαO is a linear function of mierosomal protein (and by extension, microsomal lipid; Nashed et al., 1985). Similarly, inhibition of enzyme activity by 5, 6αiminocholestanol is dependent not only on the concentrations of substrate and inhibitor, but on that of microsomal lipid as well (Nashed et al., 1985). Kinetic measurements are further complicated by the fact that the 3, 5, 6-triol product inhibits the enzyme (Nashed et al., 1985; Sevanian and McLeod, 1986). Various estimates have been made for the relative rates of hydrolysis of ChαO and ChβO, and the Vmax values for both epoxides seem to be approximately equal −0.48 nM mg−1 min−1 for ChαO and 0.37 nM mg−1 min−1 for ChβO (Sevanian and McLeod, 1986), or 0.45 nM mg−1 min−1 for both epoxides (Nashed et al., 1985). Inhibitors Compounds that are good inhibitors of mEH generally have no effect on ChEH (Levin et al., 1983; Sevanian and McLeod, 1986). Trichloropropene oxide, one of the most potent mEH inhibitors, actually enhances ChEH activity (Watabe et al., 1980b; Oesch et al., 1984). Nonionic detergents partially or completely irreversibly deactivate ChEH (Watabe et al., 1986). Because membrane solubilization is required for enzyme purification, this deactivation has complicated efforts (as yet unsuccessful) to purify the enzyme. Sodium cholate only partially deactivates the enzyme (Oesch et al., 1984; Watabe et al., 1986). Analogues to the cholesterol epoxides are the most selective and potent inhibitors of ChEH. 6-Ketocholestanol and 7ketocholestanol inhibit hydrolysis of the epoxides competitively, with Ki alues of 2–5 µM (Sevanian and McLeod, 1986). The 3, 5, 6-triol also inhibits the reaction, with a Ki of 6–11 µM (Sevanian and McLeod, 1986). Other studies show that all four triol stereoisomers at the 5, 6 position inhibit ChαO hydrolysis with Kis ranging from 30–110 µM (Nashed et al., 1985). Non- or very weakly inhibitory analogues include cholesterol and oxidosqualene (Nashed et al., 1985; Sevanian and McLeod, 1986). 5, 6α-and 5, 6β-iminocholestan-3β-ol, two aziridine-containing analogues of the cholesterol 5, 6-oxides, provided the first examples of selective ChEH inhibitors (Watabe et al., 1980b). The 5, 6α-imine (Figure 12.4D) is a more potent inhibitor of both stereoisomeric epoxides than is the 5, 6β-imine (Watabe et al., 1980b). 5, 6α-iminocholestanol also competitively inhibits the

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hydrolysis of the alpha-and beta-stereoisomeric epoxides of 5-cholestene, pregnenolone, and 20-methylpregnenolone (Watabe et al., 1983). As mentioned above, this compound does not inhibit microsomal hydrolysis of a variety of substrates for mEH (Watabe et al., 1983). The Ki of 5, 6αiminocholestanol for ChEH is 85 nM, 120 times smaller than the Km of either epoxide for the enzyme (Nashed et al., 1985). A synthetic suicide inhibitor of ChEH, 7-dehydrocholesterol 5, 6β-oxide (Figure 12.4E; Michaud et al., 1985; Nashed et al., 1986), has been employed not only as an effective inhibitor of the enzyme, but as a tool for understanding its mechanism as well (see below). This compound inactivates the enzyme by forming a reactive allylic epoxide intermediate (Michaud et al., 1985) that covalently binds to the active site of the enzyme (Nashed et al., 1986). The I50 concentration for this inhibition is 48 nM (Michaud et al., 1985). Enzyme mechanism The mechanism of cholesterol 5, 6-oxide hydrolysis by ChEH has been deduced primarily by examination of the stereochemistry of the triol product(s) and through the use of the mechanism-based 7-dehydrocholesterol 5, 6β-oxide inhibitor. The enzyme-catalyzed hydrolysis exhibits a unique stereo-selectivity in that ChαO undergoes backside attack and hydroxylation at C6, and ChβO undergoes backside attack and hydroxylation at C5, such that in both cases the resulting triol has the identical 3β, 5α, 6β-configuration (Figure 12.4C; Watabe et al., 1980a, 1981; Nashed et al., 1985). The stereo-selectivity of the enzymecatalyzed reaction reflects a similar selectivity of product formation by both acid- and base-catalyzed hydrolysis of either epoxide in aqueous solution; in both cases, the 3β, 5α, 6β-triol is the only solvolysis product (Nashed et al., 1985). The enzyme appears to be merely facilitating the solvolytic process and does not appear to alter its stereo-selectivity. This product configuration is also the only one observed in the case of ChEH-catalyzed hydrolysis of the 5, 6α- and 5, 6βoxides of 5-cholestene, 20-methylpregnenol, and pregnenolone (Watabe et al., 1981). It has been suggested that the enzyme assists protonation of the oxirane oxygen, which polarizes the C-O bond and activates both oxirane carbons (Watabe et al., 1980a; Nashed et al., 1985). In the case of ChαO, backside hydroxylation at C6 is favoured, probably due to steric hindrance of hydroxylation at C5 exerted by the nearby C19 methyl group. In the case of ChβO, backside attack at C5 is favoured, due to the unfavourability of the alternative which would result in a cis configuration at the C5-C10 ring junction. Thus in both cases, formation of the resulting 5, 6-diol in the 5α, 6β configuration is strongly favoured. Studies wherein all four possible 5, 6-diols were synthesized and used to determine product selectivity showed that, with microsomes isolated from five different species, 93%–100% of the total product

390 CONJUGATION REACTIONS IN DRUG METABOLISM

was in the 5α, 6β configuration (Nashed et al., 1985). These conclusions are supported by studies showing similar patterns of addition by other nucleophiles (Blackburn et al., 1979; Watabe et al., 1979; Michaud et al., 1985). Protonation of the oxirane oxygen as a key step in catalysis is suggested by inhibitor studies. The observed Ki for 5, 6α-iminocholestanol inhibition of ChEH is lower than the Km for either epoxide by a factor of 120 (Nashed et al., 1985). The high affinity of this protonated species for the active site suggests that it is a transition state analogue, which in turn implies that the transition intermediate in epoxide hydrolysis contains a protonated oxygen species. This general mechanism is also suggested by the ability of the 7-dehydro analogue of ChαO (Figure 12.4E) to covalently modify and inactivate the enzyme. The catalysis-dependent covalent binding of this compound to the enzyme active site is mediated by the formation of an allylic carbocation which can only result from protonation of the oxirane oxygen and the resulting cleavage of the C-O bond (Michaud et al., 1985; Nashed et al., 1986). It does not appear, however, that cleavage of the C-O bond prior to nucleophilic attack occurs in the case of ChαO and ChβO hydrolysis; otherwise, formation of a tertiary carbocation at the C5 position would always be favoured and hydration would only (or primarily) occur at C5. The proposed mechanism suggests a somewhat unorthodox active site that either binds the steroid ring system in a single orientation but can activate the oxirane oxygen regardless of the side of the ring on which it occurs, or activates the oxirane oxygen on only one side of the steroid ring but has a non-restrictive substrate binding site that allows the ring system to bind in two different orientations. Biological role ChEH has been proposed as a detoxifying enzyme which serves to protect the cell against the deleterious effects of the 5, 6-oxides of cholesterol (for review see Black, 1980, and Sevanian and Peterson, 1986). This hypothesis assumes that cholesterol 5, 6-oxide poses a cytotoxic and/or genotoxic hazard to the cell at its physiological concentration and that hydrolysis of the epoxide mitigates this hazard; such an assumption is defensible, but not undebatable. The possible involvement of cholesterol epoxides as etiological agents in both intestinal and skin cancer has been raised (Black, 1980; Petrakis et al., 1981), but data from animal experiments do not allow a definitive conclusion. Subcutaneous injection of large amounts of ChaO led to tumours at the injection site in 5–20 % of animals so treated (Bischoff, 1969). Other studies where ChaO was fed or installed intrarectally report no increase in tumour incidence (Seelkopf and Salfelder, 1962; Black, 1980). The cytotoxicity of ChaO in cell culture has been clearly demonstrated. However, at equivalent doses, the cytotoxicity of the 3, 5, 6-triol is greater than that of the parent epoxide (Kelsey and Pienta, 1981;

EPOXIDE HYDROLASES 391

Sevanian and Peterson, 1984, 1986). ChaO is genotoxic in mammalian cell transformation assays; it produces transformation of hamster embryo cells at frequencies equivalent to those produced by similar doses of 3methylcholanthrene (Kelsey and Pienta, 1979, 1981). It should also be noted, however, that the 3, 5, 6-triol produces similar, or slightly higher, transformation frequencies (Kelsey and Pienta, 1981). ChaO has been reported to form covalent adducts with DNA (Blackburn et al., 1979). However, this report supplies little supporting data, and the observation of covalent binding to DNA could not be repeated in two other laboratories (T.Guenthner, unpublished data; W. Lutz, personal communication). The mechanism of cholesterol oxide genotoxicity has been somewhat clarified by recent studies (Peterson et al., 1988) showing that 6-thioguanine resistance, but not ouabain resistance, results from exposure of V79 cells to either cholesterol oxide. The authors suggest that cholesterol oxides act epigenetically or by repression of the HGPRT gene rather than as direct mutagens. This study also provides evidence that cholesterol oxides interact chemically with DNA but that this interaction does not involve covalent adduct formation. Leukotriene A4 hydrolase The enzyme Leukotriene A4 (LTA4) (5(S)-trans-5, 6-oxido-7, 9-cis-11, 14-transeicosatetraenoic acid; Figure 12.5A) is a semi-stable epoxide derived from arachidonic acid via lipoxygenase-catalyzed oxygenation. It is a key intermediate in the biosynthesis of all leukotrienes, which play important roles in a number of immunopathological responses (for review, see Samuelsson, 1983, and Goetzl et al., 1984). LTA4 can undergo nonenzymic hydrolysis to form epimeric 5, 6diols (Figure 12.5D) and 5, 12-diols (Figure 12.5E; Borgeat and Samuelsson, 1979a, 1979b; Maycock et al., 1982; Fitzpatrick et al., 1983) that are not presently known to be physiologically active. It can also undergo glutathione conjugation at C6 which leads to leukotriene C4 (Figure 12.5C) formation (Murphy et al., 1979; Rådmark et al., 1980a). Finally, of interest to this discussion, it can undergo stereo-selective enzymic hydrolysis, catalyzed by a specific epoxide hydrolase, to form 5(S), 12(R)-dihydroxy-6, 14-cis-8, 10-transeicosatetraenoic acid, leukotriene B4 (LTB4; Figure 12.5B). LTA4H is a true epoxide hydrolase in that it catalyzes hydrolytic cleavage of the LTA4 oxirane ring. However, it differs from other known epoxide hydrolases in that the addition of water does not result in a vicinal (α, β) diol. Rather, the product is the 5, 12-diol shown in Figure 12.5B. The addition of water at a position distant from the oxirane ring is attributable to the formation of a

392 CONJUGATION REACTIONS IN DRUG METABOLISM

Figure 12.5. Leukotriene A4 and chemically-related compounds. (A) Leukotriene A4. (B) Leukotriene B4. (C) Leukotriene C4. (D) 5(S), −6(R or S)-dihydroxy-7, 9-trans-11, 4cis-eicosatetraenoic acid. The wavy line indicates 2 possible epimers at C6. (E) 5(S), 12 (R or S)-dihydroxy-6, 8, 10-trans-14-cis-eicosatetraenoic acid. The wavy line indicates two possible epimers at C12. (F) Leukotriene A3. (G) Leukotriene A5. (H) Inhibitor I. (I) Inhibitor II. (J) Inhibitor III. (K) Lipoxin A4.

delocalized carbocation intermediate in the conjugate triene structure which results in activation of the C12 carbon (see below). LTA4H has been identified and characterized in a number of tissues and species. It is widely distributed. Initially the enzyme was identified in human leukocytes (Rådmark et al., 1980b) and rat basophilic leukaemia cells (Maycock et al., 1982; Jakschik and Kuo, 1983), as well as in cell-free plasma obtained from many species (Fitzpatrick et al., 1983). Subsequent investigations showed the enzyme to be present in rodent (Haeggström et al., 1985; Pace-Asciak et al., 1985; Medina et al., 1987) or human (Haeggström et al., 1985; Ohishi et al., 1987) liver, lung, and kidney, and human erythrocytes (McGee and Fitzpatrick, 1985) and nervous tissue (Shimizu et al., 1987). Investigation of the distribution of enzyme activity in cytosolic fractions derived from 13 guinea pig tissues showed specific activities (enzyme activity per milligram of cytosolic protein) in the following rank order: small intestine>adrenal>kidney>stomach>leukocytes> heart>brain>spinalcord>liver (Izumi et al., 1986). No activity was found in erythrocytes or plasma. The enzyme is exclusively cytosolic. LTA4H activities have been measured in the cytosolic and membrane fractions of various tissues, and no significant activity has been seen in any membrane fraction (Jakschik

EPOXIDE HYDROLASES 393

and Kuo, 1983; Rådmark et al., 1984; Haeggström et al., 1985; Pace-Asciak et al., 1985; Izumi et al., 1986; Medina et al., 1987). The enzyme has been purified from human leukocytes (Rådmark et al., 1984), human erythrocytes (McGee and Fitzpatrick, 1985), rat neutrophils (Evans et al., 1985a), human lung (Ohishi et al., 1987), and guinea pig liver (Haeggström et al., 1988a). LTA4H can be distinguished from other known epoxide hydrolases on the basis of its subcellular location, substrate specificity (it does not hydrolyze either styrene oxide or trans-stilbene oxide; Evans et al., 1985a; McGee and Fitzpatrick, 1985), mechanism and product stereo-selectivity (see below) or sensitivity to in vitro inhibition by various synthetic compounds (McGee and Fitzpatrick, 1985, see below). Evidence that it is not structurally related to other epoxide hydrolases has been obtained by isolating the pure enzyme and determining its biophysical properties (Rådmark et al., 1984; Evans et al., 1985a; McGee and Fitzpatrick, 1985; Ohishi et al., 1987; Haeggström et al., 1988a). Although various purified LTA4H preparations have slightly different properties, those obtained from human or rat leukocytes and human lung are quite similar. The enzyme is a monomer, with a molecular weight of 68–71 kDa, as determined by SDS-polyacrylamide gel electrophoresis (Rådmark et al., 1984; Evans et al., 1985a; Ohishi et al., 1987). Gel filtration provides an apparent molecular weight value of 49–51 kDa; the discrepancy between the two values indicates that the protein is non-globular. The enzyme has an isoelectric point of 5.1–5.7 and its amino acid composition and N-terminal sequence have been determined (Rådmark et al., 1984; Ohishi et al., 1987). The pH optimum of purified LTA4H has been estimated as 7–8 (Evans et al., 1985a) or 8–9 (Rådmark et al., 1984). The enzyme obtained from human erythrocytes (McGee and Fitzpatrick, 1985) is similar to the other preparations in respect to isoelectric point, pH optimum and kinetic properties but was determined to have a different molecular weight (54 kDa) by SDS-polyacrylamide gel electrophoresis. The enzyme is very stable upon purification, with no loss of activity seen after incubation of the purified preparation at 4° for one week (Rådmark et al., 1984), or at 37° for 72 h (McGee and Fitzpatrick, 1985). LTA4H, therefore, differs from microsomal epoxide hydrolase(s) by virtue of its molecular weight, amino acid composition, acidic isoelectric point and location in the cytosol and differs from cEH in respect to its basic pH optimum, monomeric composition, molecular weight and amino acid composition. Neither purified microsomal epoxide hydrolase nor purified cytosolic epoxide hydrolase is capable of converting LTA4 to LTB4 (Haeggström et al., 1986). DNA elements that code for LTA4H have been isolated from human placenta (Funk et al., 1987) and spleen (Minami et al., 1987) genomic libraries and their nucleotide sequences have been determined. Both studies agree that the gene product contains 610 amino acids, with a deduced molecular weight of 69 140 (Funk et al., 1987), or 69 153 (Minami et al., 1987). No significant homology between this sequence and the known

394 CONJUGATION REACTIONS IN DRUG METABOLISM

sequence of human mEH could be detected. This human genetic element has been transfected into Escherichia coli, and the transfected bacteria produce an active enzyme protein with a molecular weight of 70 kDa, with similar catalytic properties to those of the isolated human enzyme (Minami et al., 1988). Kinetics of LTA4 hydrolysis have been determined using homogeneous enzyme preparations. Estimates of the pH optimum of the purified enzyme (Rådmark et al., 1984; Evans et al., 1985a; McGee and Fitzpatrick, 1985) generally correspond to the pH optimum of 7–8.5 observed with unfractionated liver cytosol (Haeggström et al., 1985). The Km of the enzyme for LTA4 has been estimated as 10–40 µM (Rådmark et al., 1984; Evans et al., 1985a; McGee and Fitzpatrick, 1985; Ohishi et al., 1987). Estimation of Vmax values for the purified enzyme range from 0.3–3 µmoles product mg−1 min−1. Difficulties in measuring initial enzyme velocity occur because of the short period (much less than 1 min) during which a constant initial rate of substrate conversion occurs. The enzyme undergoes rapid suicide inactivation during LTA4 hydrolysis, decreasing the total rate of substrate conversion and complicating estimation of enzyme velocity (Jakschik and Kuo, 1983; McGee and Fitzpatrick, 1985; Ohishi et al., 1987). A covalent adduct is formed between the substrate and enzyme (Evans et al., 1985b; Nathaniel et al., 1985); it is surmized that the electrophilic delocalized carbocation mentioned above is the reactive intermediate involved in covalent binding. Analogues of LTA4 that inhibit its hydrolysis [e.g. LTA3 (Figure 12.5F) and LTA5 (Figure 12.5G)] also inhibit its covalent binding to the enzyme (Evans et al., 1985b; Nathaniel et al., 1985) apparently by competitively forming covalent adducts themselves. A series of LTA4 analogues were synthesized, some of which were structurally capable of forming a delocalized carbocation intermediate, some of which were not. Only those compounds that could form such an intermediate inactivated the enzyme (Ohishi et al., 1987). The fact that this enzyme is deactivated by its endogenous substrate may insure that the production of LTB4 is tightly regulated and easily limited, but the mechanism of this limitation appears to be rather an inefficient one for the cell. It is not known whether the covalently-bound enzyme can be reactivated by the cell, or whether de novo enzyme synthesis is required to regenerate enzyme activity. A similar example is observed in the case of a nuclear enzyme that repairs alkylated DNA bases by a trans-alkylation process that irreversibly inactivates the enzyme (Harris et al., 1983). Inhibitors and substrates The apparent mechanism of suicide inactivation of the enzyme is highly suggestive that formation of a free carbocation, which can incidentally bind covalently to the enzyme, is an important step in LTA4 hydrolysis. The theoretical stability of this intermediate due to delocalization of its charge in the

EPOXIDE HYDROLASES 395

conjugated triene structure supports its postulation. However, the postulation of enzyme-assisted formation of such an intermediate by protonation of the oxirane oxygen and cleavage of the C-O bond followed by hydration of the resulting ionic species is not sufficient to explain the stereo-selectivity of LTB4 formation. Hydrolysis of LTA4 under acid conditions, presumably occurring by the above mechanism, results in the formation of four diol products, two 5, 6-epimers (Figure 12.5D) and two 5, 12-epimers (Figure 12.5E; Borgeat and Samuelsson, 1979a, 1979b; Maycock et al., 1982; Fitzpatrick et al., 1983). That the enzyme produces a single product of defined stereochemical configuration suggests that attack of water at the C12 position probably occurs stereo-selectively and simultaneously with intermediate formation. cEH also hydrolyzes LTA4 but with a different regio- and stereo-selectivity, resulting in the formation of a 5, 6-diol, the 5(S), 6(R)-dihydroxy-7, 9-trans-11, 14, -cis-eicosatetraenoic acid (Haeggström et al., 1986, 1988b). This other hydrolase is known to proceed by a base-catalyzed SN2 mechanism (Mullin and Hammock, 1982), and consonant with this mechanistic difference between the two enzymes, no suicide inactivation of cEH by LTA4 or by an electrophilic intermediate derived therefrom occurs (Haeggström et al., 1986). Unlike the rather non-substrate-specific hydrolases, mEH and cEH, the catalytic activity of LTA4H is narrowly confined to LTA4 and analogues that closely resemble it. Leukotriene A5 is converted to the corresponding 5(S), 12 (R)-diol, but LTA3 is not (Evans et al., 1985b; Ohishi et al., 1987). The methyl ester of LTA4 is not metabolized (Maycock et al., 1982; Fitzpatrick et al., 1983; Haeggström et al., 1985; Ohishi et al., 1987), nor is the analogous leukotriene 14, 15-LTA4, which has an epoxide function at the 14, 15-rather than the 5, 6position (Rådmark et al., 1984; Evans et al., 1985a; Ohishi et al., 1987). 5, 6EET, a non-conjugated arachidonic acid epoxide (Figure 12.3E) is not hydrolyzed by LTA4H, while the corresponding 14, 15-EET and 11, 12-EET are, albeit extremely slowly (McGee and Fitzpatrick, 1985). Other known epoxide hydrolase substrates, such as styrene oxide and trans-stilbene oxide (Figure 12.3A) are not hydrolyzed by LTA4H (Evans et al., 1985a; McGee and Fitzpatrick, 1985; Ohishi et al., 1987), A series of analogous 5, 6-oxides of fatty acids, which either have a different stereochemistry in the triene system or in which the conjugated system is 2, 3-rather than 1, 2-to the oxirane ring, are not metabolized by the enzyme (Ohishi et al., 1987). The enzyme appears to require as a substrate an unesterified fatty acid with an epoxide at the 5, 6-position which is alpha to a 7, 9-trans-11,14, -cis-conjugated double bond system. Accordingly, the most effective inhibitors of the enzyme are analogues of LTA4. 5, 6-EET (Figure 12.3E), 11, 12-EET, and 14, 15-EET are poor inhibitors (McGee and Fitzpatrick, 1985). The cEH inhibitors trans-stilbene, chalcone oxide, and trans-1, 2-dibenzoylethylene do not inhibit LTA4H (McGee and Fitzpatrick, 1985). Compounds that bind sulfhydryl groups inhibit the enzyme

396 CONJUGATION REACTIONS IN DRUG METABOLISM

(Ohishi et al., 1987), as do high concentrations of trans-stilbene oxide (McGee and Fitzpatrick, 1985) and arachidonic acid (Haeggström et al., 1985). More potent inhibitors are leukotriene A3 and leukotriene A5, which, as mentioned above, compete with LTA4 at the catalytic site and deactivate the enzyme (Evans et al., 1985b; Nathaniel et al., 1985; Evans et al., 1986). The other LTA4 analogues mentioned above (Ohishi et al., 1987) which are not substrates for the enzyme nevertheless inhibit the enzyme by deactivating it. It appears that the major structural criterion for enzyme deactivation is an epoxide alpha to a conjugated diene system, regardless of stereochemistry, again indicating the probable importance of delocalized carbocation formation. A series of LTA4related compounds, shown in Figures 12.5H-12.5J, were synthesized as possible inhibitors of LTA4H (Evans et al., 1986). The 7, 9-trans- diene (12.5H) gave the most complete inhibition followed by the substituted styrene analogue (12. 5I) and the analogue containing the neighbouring triple bond (12.5J). The possibility of enhancing LTA4H activity in vivo has also been considered. The exposure of human neutrophils to phorbol myristate acetate results in the stimulation of LTA4H activity in the presence of exogenous arachidonic acid. This stimulation corresponds with phosphorylation of a 68 kDa protein in the cell, suggesting that LTA4H may be regulated by protein kinase C-dependent phosphorylation (McColl et al., 1987). 12.6. Practical considerations Methods for the determination of epoxide hydrolase activities are found in many of the publications referenced above. Measurement of mEH activity is most easily performed by generating a radio-labelled diol product from a labelled epoxide substrate, separating the two by solvent extraction and quantitating product formation by scintillation counting. mEH activity has been measured by this technique using radio-labelled styrene oxide (Oesch et al., 1971a), benzo[a] pyrene 4, 5-oxide (Schmassmann et al., 1976), and cis-stilbene oxide (Mullin and Hammock, 1980) as substrates. The latter two substrates are preferred because styrene oxide is hydrolyzed by cEH (Gill et al., 1983a), whereas the other two substrates are not (Mullin and Hammock, 1980; Guenthner et al., 1981). Four different articles in Methods in Enzymology present variations on this technique (DePierre et al., 1978; Lu and Levin, 1978; Guenthner et al., 1981; Hammock et al., 1985), and in two of them (DePierre et al., 1978; Guenthner et al., 1981), alternative (non-radiometric) methods for the assay of mEH activity are discussed (see also Seidegård and DePierre, 1983). Determination of cEH activity is likewise most easily performed by radiometric assays, using transstilbene oxide or other trans-substituted styrene oxides as substrates. Technical aspects of cEH assays are detailed in Methods in Enzymology (Hammock et al.,

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1985) and are also discussed at length in a recent review article (Meijer and DePierre, 1988). Measurement of ChEH activity, using thin layer chromatography to separate radio-labelled cholesterol 3, 5, 6-triol from its radiolabelled cholesterol 5, 6-oxide precursor is also detailed in Methods in Enzymology (Hammock et al., 1985). This is a standard method described in many of the publications referenced above (e.g. Sevanian et al., 1980; Levin et al., 1983; Watabe et al., 1983). LTA4H activity is measured by HPLC separation of LTA4 metabolites and quantitation of LTB4 by evaluation of peak height or area, as described in detail in several articles referenced above (e.g. Maycock et al., 1982; Rådmark et al., 1984; McGee and Fitzpatrick, 1985; Evans et al., 1985b; Ohishi et al., 1987). Abbreviations cEH ChEH ChαO and ChβO EET LTA4 or LTB4 LTA4H LTC4 mEH

Cytosolic epoxide hydrolase Cholesterol 5, 6-oxide hydrolase Cholesterol-5, 6α or β-oxide 14, 15-epoxyeicosatetraenoic acid Leukotriene A4 or B4 Leukotriene A4 hydrolase Leukotriene C4 Microsomal epoxide hydrolase

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Index

4-Acetamidotropolene 221 Acetaminophen, see Paracetamol Acetonitrile 377 Acetyl coenzyme A 164, 183, 184 2-Acetylaminofluorene 178, 369, 381, 386 Acetylation 43–4, 163–91, 254 N-Acetylcysteine 314 N-Acetyl-p-benzoquinone imine 318, 325, 331 4-Acetylphenyl sulfate 141 N-Acetyltransferases 163–91, 341 Acivicin 310, 316, 317 ‘Active sulfate’, see PAPS Active site 69, 70, 199–203, 332–3, 372, 378, 389 Acyl CoA synthetases 288 Acyl migration 57, 83, 88 Adenosine 54, 77 Adenosine 3′5′-bisphosphate 135, 136, 143 S-Adenosyl-L-homocysteine 203, 238, 242, 245, 255, 266 S-Adenosyl-L-methionine 194–5, 234, 235, 244, 252, 255 Adrenaline 75, 122, 141, 195, 196, 213, 219, 244, 245 Adrenocortical cells 121 Aflatoxin B1 54, 338, 374 Aflatoxin B1–8, 9-oxide 328 Age 55, 77, 86, 119, 125, 126, 296, 311, 315, 336 Alanine 279 Alcohol, see Ethanol Alcohols 55, 112, 113, 142 Allyl alcohol 74 Allyl isothiocyanate 346 Amino acid conjugation 44, 273–305

Amino acid N-acyltransferases 288–90 4-Aminoazobenzene 240 2-Amino-4, 5-diphenylthiazole 378 1-Amino-4-nitronaphthylene 140 2-Amino-1, 2, 3, 4,tetrahydronaphthalene 219 4-Aminobenzoic acid 35, 168, 172–8, 292, 295 2-Aminobiphenyl 240 4-Aminobiphenyl 63, 165, 168, 173, 177, 238, 240 2-Aminofluorene 165, 168, 173, 176–82 2-Aminophenol 59–61 4-Aminophenol 87, 118 4-Aminosalicylic acid 42, 168, 173, 176 4-Aminosulfonic acid 168 Amodiaquine 242 Amphetamine 220 Androstene-3, 17-dione 330 Androsterone 60–4 Angiotensins 141 Aniline 77, 118, 140, 168, 235 Aorta 120, 213 APS kinase 109 Apomorphine 238 Arachidonic acid 381, 395 Arginine 279 Aroclor–1254 369, 386 Aromatic amines 44, 55, 164, 168, 170, 180–2, 185 Arylacetic acids 286–7 2-Arylpropionic acid 58 Ascorbic acid 129, 138, 196 L-Asparagine 290 Aspartic acid 292 ATP sulfurylase 109, 129 406

INDEX 407

Atractyloside 117 Azaindole 238 Aziridine carboxylic acid 344 Azothiopurine 259 Benserazide 219 Benzo[a]anthracene 5, 6-oxide 386 Benzidine 165, 168, 182, 238, 240 Benzil 369 Benzo[a]pyrene 329, 383 Benzo[a]pyrene-4, 5-oxide 329, 341, 346, 372, 376, 377, 384, 396 Benzo[c]phenanthrene-5, 6-oxide 372, 386 Benzofuran 369 Benzoic acid 35, 275, 278, 279, 281, 284, 292, 294 Benzylic alcohols 138 1-Benzylimidazole 380 Bile acids 57, 61, 127, 132, 133, 137, 333 Biliary excretion 30, 34, 72, 80–2, 86, 121, 125, 126, 261, 264, 266, 295, 316, 338, 342–6 Bilirubin 59–63, 68, 88, 90, 333 Biphenyl 75, 118, 219 Bipyridyls 238 Bladder 75, 167 Blood-brain barrier 239 Brachymorphic mice 110, 122, 124 Brain 110, 141, 206–8, 215–6, 242, 245 Bromazepam 252 Bromoacetanilide 168, 172 2-Bromooctane 324 α-Bromoisovaleric acid 346 α-Bromoisovalerylurea 45, 323, 324, 331, 343 4-Bromophenol 219 Bromosulfthalein 316, 333, 342, 345 Brompheniramine 282 Buprenorphine 84 Burimamide 242 Butadiene 373 2(3)-t-Butyl-hydroxyanisole 79, 84, 311, 336–8, 345, 369 t-Butylhydroperoxide 317 t-Butyl-4-hydroxytoluene 369 4-tert-Butylaniline 140 Busulfan 344, 347

2, 3-Butanedione 70 Butanephrine 219 Buthionine sulfoximine 310, 311, 313 Caffeine 168 Calcium 75 Carbamazepine 373 Carbidopa 216, 219 Carbon tetrachloride 71, 74 Carboxylic acids 43–4, 55, 273–305 Captopril 252 Cat 114, 121 Catalase 378 Catecholamines 112, 212, 213 Catechol 31, 43, 194, 195, 203, 209, 212, 221 estrogens 209 steroids 212, 221 Catechol-O-methyltransferase 196, 197– 203, 205–18 inhibition 203 Catechols 43, 194, 195, 215–7 Cell cultures 77, 117, 207, 342, 343 Cephalosporin 253 Chalcone oxide 377, 395 Charcoal-broiled diet 126 Chenodeoxycholate 124 Chloramphenicol 59, 64, 69, 84, 86, 138 Chlorpheniramine 282 Chlorate 109, 129 1-Chloro-2, 4-dinitrobenzene 324, 343 2-Chloro-4-nitrophenol 143 1-Chloro-1-phenylethane 324 Cholecystokinin 129, 131, 138, 141 Choleresis 88 Cholestasis 88 Cholesterol 132, 368 Cholesterol 5, 6-oxide hydrolase 384–90, 396 Cholesterol 3-sulfate 130 Chondroitin 130, 136 Choroid plexus 217, 218 Cimetidine 87, 242 Ciprofibrate 386 Ciramadol 87 Clofibric acid 59, 82, 287, 345, 369, 370, 380, 386

408 INDEX

Clonazepam 168 Clorpromazine 87, 242 Cobalt chloride 311 Codeine 69 Coenzyme A 164, 274, 285 Competition 9–16, 41–9, 294–5 Contraceptive steroids 86, 126, 215 Corpus luteum 211 COS-7 cells 64 Cotinine 237 Coumarin 338 Crigler-Najjar Syndrome 88 Cryopreservation 77 Cumene hydroperoxide 330 2-Cyanoethyl-N-hydroxythioacetamidate 138 Cyclamate 140 Cyclobenzaprine 83, 140 Cyclohexene oxide 369, 377 Cyclohexylamine 168 Cyclopropylcarboxylic acid 283 Cyproheptadine 69, 83 D-Cysteine 111, 124 L-Cysteine 111, 256, 309, 310, 312–4, 317 Cytochrome P-450 8, 240, 314, 367 DDT 85 Deconjugation 31, 81 7-Dehydrocholesterol-5, 6β-oxide 386, 388 Dehydroepiandrosterone 128, 132 Desmethylimipramine 215, 235 Detergent 9, 70, 89 Development 60, 76, 125, 169–70, 205, 210, 245, 296 Diabetes 76 Diazepam 87 Dibromosulfthalein 345 Dibucaine 242 Dibutyryl c-AMP 77, 119 2, 6-Dichlorobenzonitrile 264–6 3, 4-Dichlorobenzyloxyacetic acid 287 1, 2-Dichloro-4-dinitrobenzene 324, 343 2, 6-Dichloro-4-nitrophenol 17, 20, 91, 110, 127, 128, 129, 134, 145, 146 2, 4-Dichlorophenoxyacetic acid 287, 290, 297, 381 Dichlorotetrahydro-isoquinoline 238

Diet 48, 85, 111, 123, 126, 129, 312, 313 Diethyl ether 54, 55, 119 Diethyl maleate 129, 310, 311, 314–6, 325, 343–5 Diethylpyrocarbonate 70 Diethylstilbestrol 60 Diffusional barrier 30–1 Diflunisal 82, 125 Digitoxigenin 59 Digitoxigenin monodigitoxoside 63, 65 Dihydrodiols 367, 371 2, 4-Dihydroxybenzoic acid 198 3, 4-Dihydroxybenzyl alcohol 203 6, 7-Dihydroxy-3, 4-dihydroisoquinolines 204 Dihydroxyindoles 220 3′, 4′-Dihydroxy-2-methylpropiophenone 216, 221 Dihydroxyphenylacetic acid 208 2, 3-Dihydroxypyridine 204 1, 15-Dihydroxy vitamin D 88 Dimaprit 242 4-Dimethylaminophenol 120 Dimethylsulfoxide 169, 342 N, N-Dimethyltryptamine 238 2, 4-Dinitrophenol 54, 110 Diphenhydramine 242, 280 Dipyridyl 240 Disulfiram 338 Dithiothreitol 257, 268 DNA adducts 134, 338 Dobutamine 219 Dog 57, 58, 78, 81, 110, 114, 119, 122, 123, 132, 170 L-Dopa 79, 132, 195, 216–7 Dopamine 128, 131, 132, 141, 195, 213, 216 Double conjugates 44 Ellagic acid 333 Embryo cells 80 Enalapril 35 Enantiomers, see Stereoselectivity Endoplasmic reticulum 47, 52, 70, 71 340, 341, 367, 368 Endorphins 131 Endothelial cells 120, 213

INDEX 409

Ephedrine 138 Epichlorohydrin 373, 374 Epinephrine, see Adrenaline Epoxides 44, 325–30, 365–404 Epoxide hydrolase 8, 44, 344, 365–404 cis-Epoxymethylstearate 387 1, 2-Epoxy-3-(p-nitrophenoxy) propane 325 Erythrocytes 214–5, 258, 259, 310, 311, 315, 336, 392 Estradiol 60, 64, 141 Estriol 63 Estrone 60, 63, 141 Ethacrynic acid 242, 311, 325, 346, 347 Ethanol 21, 53, 54, 76, 110, 117, 119, 137, 313, 314, 338 di-(2-Ethylhexyl)-phthalate 381 Ethinyloestradiol 88, 119, 123, 125, 221 Ethionine 110, 117, 255, 317 7-Ethoxycoumarin 73, 75, 116, 117 Ethyl acetate 377 Ethylene dibromide 323, 348 Ethylmorphine 317 Ethylene glycol 114 Etiocholanolone 64 Etoposide 71 Extrahepatic conjugation 32–5, 61, 71, 81, 83–4, 121 Eye 218 Fasting 54, 75, 77, 85, 111, 116, 119, 126, 312–5 Fatty acid epoxides 382 Fenofibrate 380 Fenoldopam 123 Fenoterol 58, 79, 84 Fetus 86, 126 Fibroblasts 70 First pass 49, 82, 112, 114, 119, 123, 126 Flavones 204, 370, 377 7-Fluorenone 370 9-Fluorenone oxime 134 5-Fluoronorepinephrine 201 Folic acid 169 Follitropine 130 Fructose 54, 75, 110 Furosemide 59, 69

D-Galactosamine 54, 129 Genetic polymorphism 214, 258–9, 368 Gentisamide 16, 22, 23, 26, 73, 115 Genotoxicity 348 Geranyl epoxide 375 Gastrines 131 Gilbert syndrome 88, 127 Glucocorticoid 138 Glucose 54, 75, 79 β-Glucuronidase 75 Glucuronidation 9–34, 43, 44, 47, 51–105, 286, 294–5, 299 Glutamic acid 279, 283 L-Glutamic acid 309 Glutamine 280, 286, 290 γ-Glutamylcysteine synthesis 309, 311, 315 γ-Glutamyltranspeptidase 309, 316, 317, 341, 348 Glutathione 28, 43, 48, 134, 256, 258, 309– 17, 318, 332, 342, 343, 345, 374, 384 analogues 332 biosynthesis 309–11, 343 degradation 309–11 distribution 309–11 Glutathione conjugates 28–30, 44, 254, 307–63, 373, 374, 391 Glutathione transferases 8, 318–22, 334– 42, 346, 374, 384 membrane bound 340–1 Glutathione synthetase 309 Glycerol 43 Glycine 43, 278, 282, 286, 290, 292, 293, 295, 296, 309 Glycine conjugation 35, 273–305 Glycocholic acid 278 Glycosaminoglycans 130–1 Glycoproteins 130–1 Glycosylation 43 Gunn rat 61, 62, 88 Haematin 333 Halothane 54 Harman 386 Harmaline 169 Harmine 169 Harmol 11, 16, 20, 22, 23, 32, 71–3, 84–8, 110, 111, 115, 116, 118, 119, 121, 126

410 INDEX

Heart 141, 212, 213 Heparine 131, 136 Hepatocyte(s) 47, 55, 76–8, 110, 117–9, 178, 182, 312, 313, 342–4, 383 HepG2 cell line 77 Heptanethiol 258 Hexachlorobenzene 261 Hexachlorobutadiene 323, 341, 348 Hippuric acid 275, 283 Histamine 168, 235, 241–2, 279 Homocysteine 255, 256 Homovanillic acid 209 Hydralazine 164, 167, 168, 170, 179, 181 Hydrocortisone 137, 169 Hydroxamic acids 44, 55 25-Hydroxy vitamin D3 133 N-Hydroxy-2-acetylaminofluorene 115, 126, 134, 138, 176 7-Hydroxycoumarin 22, 71, 73, 116, 117, 119 3-Hydroxybenzo(a)pyrene 63 4-Hydroxybiphenyl 59, 63, 64, 76, 117, 118, 129 6-Hydroxydopamine 207, 215 p-Hydroxyephedrine 220 2-Hydroxyestradiol 210, 211 4-Hydroxyestradiol 210, 211 5-Hydroxy-indoleacetic acid 31 4-Hydroxynon-2-enal 325 8-Hydroxyquinoline 204 Hydroxylamines 44, 55, 112, 168 E-1-Hydroxynortryptiline 57 3, 4-Hydroxyphenylalanine 141 4-Hydroxypropranolol 58, 114, 220 1′-Hydroxysafrole 127, 134 17β-Hydroxysteroid 63, 64 3α-Hydroxysteroid 64 3β-Hydroxysteriod 137 5-Hydroxytryptamine 31, 168 Hyodeoxycholic acid 55, 64 Hyperbilirubinaemia 61 Hypoxia 75 Iminocholestanol 388, 389 Imipramine 238 Imodazole 241 Indol-3-ylacetic acids 280

Indole 3-carbinol 369 Indoxylsulfate 141 Induction 48, 59, 74, 77, 84, 168–9, 255, 295–6, 336–8, 369, 370, 380, 381 Inhibitors 67–8, 87, 127–9, 168–9, 216, 221, 238, 242–5, 266–7, 310, 333–4, 346–7, 369–70, 377, 378, 388, 394–5 Intestine 33–4, 60, 61, 65, 67, 78, 79, 82–4, 111, 119, 120, 123, 141, 169, 216, 254, 291 Insulin 11, 77 2-Iodooctane 342 Iodothyronine 126 Iopanoic acid 86 Isoetharine 219 Isoniazid 168, 172, 176, 177, 179, 181 Isoprenaline 123, 213, 219 Isopropoxyacetic acid 283 Isoquinoline 370 Isotretinoin 82 Kainic acid 207 Ketocholestanol 386, 388 Kidney 31, 34–5, 60, 61, 79, 80, 84, 85, 110, 120, 121, 127, 141, 169, 218, 254, 255, 258, 264–6, 290–3, 311, 313–6 Kinetics 5–39, 45–6, 74, 80–3, 114, 126, 172–4, 314 β-Lactam antibiotics 219 Levophanol 220 Levophed 219 Leukotriene A3 395 Leukotriene A4 341 Leukotriene A4 hydrolase 390–5 Leukotriene A5 395 Leukotriene B4 391 Leukotriene biosynthesis 347–8 Leukotriene C4 333 Linopleic acid 334 Lipid peroxidation 71, 382, 385 Lipid solubility 46, 47, 55, 72, 76, 110, 113, 121, 142, 252 Liposomes 312 Lipoxin A 382 Lithocholic acid 64, 88

INDEX 411

Liver 6–32, 35, 54, 55, 60, 71, 81–3, 110, 114–7, 164, 170, 197, 342–4, 254, 291– 3, 310–4, 317, 334, 335, 338 zonal heterogeneity 6–28, 73, 74, 114– 6, 342, 343 Lorazepam 71, 82, 84–7 Lung 60, 80, 83, 120, 122, 141, 169, 266, 313 Lutropine 130 β-Lyase(s) 253–4, 261–4, 265, 267, 268, 349 Lymphocytes 70 Mammary glands 211 Melphalan 323, 324 Menadione 317 Merallutide 242 2-Mercaptoethanol 252, 253, 257 6-Mercaptopurine 253, 258, 259, 268 3-Mercaptotyramine 204 Mercapturic acids 253, 261, 264, 265, 341, 344, 346, 348 Metanephrine 244 Methane sulfinic acid 263 Methinin 238, 244, 245 Methionine sulfoximine 255 L-Methionine 111, 195, 312, 317 L-Methionine S-adenosyltransferase 194 Methotrexate 281 4-Methoxybiphenyl 77, 118 3-Methoxy-4-hydroxyphenylethylene glycol 114 N-Methylaniline 140 Methylation 43, 44 N-Methylation 233–50 O-Methylation 112, 193–232 S-Methylation 251–72 N-Methyl-4-aminoazobenzene 322, 338 Methyl-bis-2-chloraniline 168 1-Methyl-4-phenyltetrahydropyridine 239 3-Methylcholanthrene 59, 73, 74, 79, 85, 116, 118, 120, 255, 340, 369, 381 α-Methyldopa 114, 123, 126, 219 3-O-Methyldopa 216 N-Methylhistamine 242 4-Methylpyrazole 54 Methylpyridinium 234

Methylsulfide 258 Methylsulfones 266 Methylsulfoxides 266 Methylthio metabolites 252 Methylthio shunt 261–3 3-Methyotoxy-4-hydroxyphenylethanol 209 N-Methyltransferases 233–50 O-Methyltransferases 193–232 S-Methyltransferases 255–9 4-Methylumbelliferone 30, 34, 35, 59, 60, 63, 64, 71, 74, 78, 84, 91, 116, 124 4-Methylumbelliferylsulfate 141, 116 Metiamide 242 Metyrapone 87, 370 Mineralocorticoid 138 Monodigitoxoside 59 Morphine 31, 34, 57, 59, 60, 61, 63, 64, 65, 69, 71, 72, 79, 81, 84, 86, 88, 90, 91, 125, 134, 220 Multiple indicator dilution technique 21, 74 Myelosuppression 259 Nafenopin 380 Nalorphine 88, 220 Naloxone 84 Naphthalene oxide 372 β-Naphthoflavone 59, 340, 369 1-Naphthol 34, 35, 59, 60–8, 69, 71, 77, 80, 84–6, 91, 109, 110, 119, 120, 121 1-Naphthylacetic acid 69, 286, 295 2-Naphthylacetic acid 280, 286 1-Naphthylamine 63, 168 2-Naphthylamine 89, 140, 141, 165, 168 Neoplasms 86, 205, 211, 212, 240, 336, 338, 339, 383 Nicotine 237 Nitrazepam 168 4-Nitroaniline 168, 172 4-Nitroanisole 77 4-Nitrobenzyl chloride 322, 324 3-Nitrocatechol 221 4-Nitroisovanillin 204 4-Nitro-estra-1 ,5(10)trien-17β-ol 129 4-Nitrophenol 35, 50–65, 71–3, 77, 84, 85, 91, 110, 116–8, 120, 126

412 INDEX

4-Nitrophenyl sulfate 111, 141, 144 1-Nitropyrene 329 1-Nitropyrene-4, 5-oxide 318, 332 1-Nitropyridine-N-oxide 341 4-Nitropyridine-N-oxide 324 Noradrenaline 131, 195, 213–6, 243, 344 Nordihydroguaiaretic acid 378 Norharman 386 Novobiocin 68, 88 Octene 1, 2-oxide 386 Octopamine 244 Oestrone 116 Ontogeny 60 Ophthalmic acid 333 Ornithine 279 OR-462, 217 Ovary 167, 210 Ouabain 138 Oxidative stress 317 Oxaprozin 82 Oxazepam 58, 69, 82, 85–7 β-Oxidation 282 N-Oxidation 42, 44 Oxiranes 368 Oxygen 117 PAPS 48, 109–12, 117, 120, 124, 126, 128, 129, 135, 136 analogues 129 availability 126 carrier 110 synthesis 109–12, 124, 128 Paracetamol 11, 22, 28, 32, 54, 55, 57, 59, 71, 77, 79, 81–7, 91, 253, 110, 111, 115– 20, 123–7, 313–8, 325, 344, 346 Paraoxon 75, 117, 168, 169, 178 Parkinson’s disease 216–7, 239 Penicillamine 253 Pentachloronitrobenzene 261 Pentachlorophenol 91, 110, 127, 128, 129, 134, 145, 169 Pentachlorothioanisole 261–6 Peptide(s) 131, 164 hormones 164 Perfusion

liver 9–16, 30, 71–6, 111, 114–7, 283, 312, 314–7, 342–4, 347, 384 kidney 31, 79, 120, 299, 344 lung 345, 120 Peroxisome(s) 369, 380, 381, 382, 386 Phenacetin 28, 32, 85, 115 Phenelzine 168 4-Phenetidine 168 Phenobarbital 54, 57, 59, 73, 74, 79, 82, 84, 85, 116–8, 295, 311, 317, 338, 340, 343, 345, 369, 381, 386 Phenol 43, 83, 85, 91, 121, 126, 141–4, 219 Phenols 55, 112, 118, 120, 129, 194 Phenolphthalein 34, 69, 77, 78 3-Phenoxybenzoic acid 281, 291, 294 Phenozocine 220 2-Phenylacetamidopenicillanic 219 Phenylacetic acid 279–81, 294, 295 trans-4-Phenyl-3-buten-2-one 325 1-Phenytethanol 138 Phenylethanolamine 235 β-Phenylethylamine 140 5-Phenyl-5-p-hydroxyphenyl hydantoin 71, 86 2-Phenylpropionic acid 58, 286 Phenyl sulfate 144, 145 Phenyltetrahydropyridine 238 4-Phenyl-1, 2, 3, 6-tetrahydropyridine 239 4-Phenylpyridine 238 Phenytoin 59, 179, 181, 373 Phorbol ester 340 Phorone 311, 313 Phosphate conjugation 43 3′-Phosphoadenylyl sulfate, see PAPS Pig 121 Piperine 79 Placenta 167, 209 Platelets 110, 215, 336 Potyethylene glycols 116 Polyglutamates 281 Polymorphism 170–8, 180, 181, 214 Potassium cyanide 54, 75 Pregnancy 76, 86, 126 Pregnanediol 60 Pregnenolone-16α-carbonrtrile 59 Preneoplastic foci 86, 134, 338 Probenicid 87

INDEX 413

Probucol 380 Procainamide 168, 172, 176, 179, 181 Procaine 242 Pro-opiomelanocortine 130 Propaehlor 253, 264, 266 Propargylglycine 312 Propofol 82 Propranolol 58, 138, 220 β-Propylstyrene oxide 375 Protein binding 20, 31, 47, 82, 117, 120, 124, 311, 342 Pyrazole 313 Pyridine 140, 204, 234, 237 Pyrogallol 204, 221 Pyrone 204 Quaternary ammonium glucuronides 84 Quercetin 333 Quinacrine 242 Rabbit 121 Ranitidine 87 Regioselectivity 371, 372 Reproductive system 209–12 Reserpine 215 Resorufin 117, 118 Retinol 138 Retinoic acid 88 Rhesus monkey 57 Rifampicin 85, 126 Rimiterol 219 RO 4–4602 221 Route of administration 48–9 Saccharine 140 Salbutamol 114, 126 Salicylamide 11, 16, 17, 20, 22–4, 26, 33, 34, 69, 82, 87, 110, 114, 119, 122, 123, 126, 127, 169 Salicylic acid 31, 35, 82, 86, 112, 292, 294, 295, 297, 314 Salicyluric acid, 31, 297, 299 Sarcosine 282 Selenium deficiency 314 Serine 279, 283

Sex differences 48, 84, 85, 110, 114, 126, 205, 206, 245, 255, 279, 280, 291–3, 322, 342 Sheep 126 SKF 525A 87, 267 Skin 60, 70, 80, 121 Smoking 85, 126 Smooth muscle 213 Spinaceamine 242 Spironolactone 59 Spleen 141, 167 Stereoselectivity 44, 45, 57, 58, 82, 90, 114, 138, 195, 282, 286, 324, 328, 329, 333, 334, 344, 346, 371, 372, 375, 388, 392, 394 Steroids 87, 132, 135 cis-Stilbene oxide 368, 396 trans-Stilbene oxide 64, 336, 338, 345, 369, 381, 382, 383, 386, 392, 395 Streptozotocin 77, 116, 119, 127, 133, 338, 343 Stucture-activity 57, 65–7, 113, 145, 167, 203, 204, 244, 254, 281–7, 334, 368, 375, 393 Styrene 325, 373 Styrene oxide 328, 342, 344, 346, 368, 372, 375, 376, 380, 384, 386, 392, 396 Suicide inhibitors 176 Sulfadiazine 168 Sulfamethazine 168, 172, 174 Sulfanilamide 178 Sulfapyridine 168 Sulfasalazine 168, 180 Sulfate 71, 73, 76, 111, 112, 115, 117, 119, 123, 124, 126 Sulfate depletion 121–4, 126 Sulfation 9–28, 30, 31, 32, 35, 43, 44, 47, 57, 71–8, 81, 85, 86, 87, 107–61 Sulfolipids 130 Sulfonamide(s) 178, 180 Sulfotransferases 8, 24, 43, 46, 134–46 Sulfodimethoxine 83 7-Sulfonyloxymethyl, 12-hydroxymethylbenzo[a]anthracene 323 Synephrine 244 Taurine 43, 48, 281–3, 286, 287, 294, 295

414 INDEX

Testosterone 60–5, 69 Tetrahydrofuran 377 2-Thioacetanilide 253, 268 6-Thioguanine 259 Thiols 255, 259 Thiomethyl shunt 253, 254 Thiophenols 55, 261 Thiopurine 252, 256 Thiopyrimidine 252 2-Thiouracil 256, 260 Thyroid 138 Thyroid hormones 117 Thyrotropine 130 Thyroxine 87, 133 Tiadenol 380 Tiaramide 114 Tissue distribution 60–1, 167–8, 178, 197, 206–8, 241, 243, 256, 263, 290–3, 334– 6, 340, 341, 350, 367, 379, 386, 392 Tocainide 82, 85 Toluidine 140 Trauma 86, 127 Triamterene 126 2, 4, 5-Trichlorophenoxyacetic acid 287, 290 Trichloropropene oxide 369, 374, 377, 383, 386, 388 Tridecylamine 140 Trimethylene oxide 369 Trimoprostil 283 Tripelennamine 83 Triphenylacetic acid 68 7, 7, 7,-Triphenylheptanoic acid 68 Triptamine 235, 236 Tropolone 204, 217, 267 Tyramine 141, 168, 220 Tyrosine methylester 141, 143–4 U–0521 221 UDP 62, 63, 68, 70 N-acetylglucosamine 68, 70 hexanolamine Sepharose 4B 62, 63 UDP glucose 53, 54 dehydrogenase 53, 54 pyrophosphorylase 53 UDP glucuronic acid 52, 53–5, 67, 70, 79, 84, 91

carrier 67, 70 dehydrogenase 53, 54 pyrophosphorylase 53 UDP glucuronosyltransferases 8, 43, 46, 58–60, 64, 65, 69, 70–1 active site 69, 70 cloning 64, 65 latency 70–1 purification 62–4 UTP 54 Urinary excretion 81 Uterus 167, 209, 210, 218 Valproic acid 82 Vas deferens 212 Vernolepin 204 Vinyl chloride 373 Vitamin A deficiency 346 Xamoterol 71–3, 81, 87, 123 Zomepirac 87 Zymosan 169