Cytochromes P450: Structure, Function and Mechanism (Taylor & Francis Series in Pharmaceutical Sciences)

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Cytochromes P450

Structure, Function and Mechanism

LEADING EDGE BOOKS IN PHARMACEUTICAL SCIENCES NEW AND FORTHCOMING TITLES International Pharmaceutical Product Registration: Aspects of Quality, Safety and Efficacy (Cartwright & Matthews) 013474974 X Advanced Drug Design and Development: A Medicinal Chemistry Approach (Kourounakis & Rekka) 013336793 2 Pharmaceutical Design and Development: A Molecular Biology Approach (Ramabhadran) 013 553884 X Reverse Transcriptase PCR (Larrick and Siebert) 013 123 118 9 Biopharmaceutics of Orally Administered Drugs (Macheras, Reppas and Dressman) 013 108093 8 Pharmaceutical Coating Technology (Cole, Hogan and Aulton) 013 662891 5 Dielectric Analysis of Pharmaceutical Systems (Craig) 013 210279 X Autonomic Pharmacology (Broadley) 074840 556 9 Photochemical Stability of Drugs and Drug Formulations (Tonnesen) 074840 449 X Potassium Channels and Their Modulators: From Synthesis to Clinical Experience (Evans et al) 074840 557 7 Pharmacokinetic Profiles of Drugs (Labaune) 074840 559 3 Flow Injection Analysis of Pharmaceuticals: Automation in the Laboratory (Martinet-Calatayud) 074840 445 7 Pharmaceutical Experimental Design and Interpretation second edition (Armstrong and James) 074840 436 8 Handbook of Drugs for Tropical Parasitic Infections second edition (Gustafsson,Beerman and Abdi) 07484 0167 9 hbk/07484 0168 7 pbk Biological Interactions of Sulfur Compounds (Mitchell) 0748402446 hbk/07484 0245 4 pbk

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Paracetamol: A Critical Bibliographic Review Review (Prescott) 07484 01369 Zinc Metalloproteases in Health and Disease (Hooper) 07484 442 2 Cytochromes P450 (Lewis) 074840 443 0 1900 Frost Road Suite 101, Bristol PA 19007–1598 USA tel: 1–800 821–8312 fax: 215–785–5515

Rankine Road, Basingstoke, Hants, RG24 8PR, UK tel: +44 (0)1256 813000 fax: +44 (0)1256 479438

Cytochromes P450 Structure, Function and Mechanism

DAVID F.V.LEWIS Molecular Toxicology Group, School of Biological Sciences, University of Surrey, Guildford, Surrey GU2 5XH

UK Taylor & Fancis Ltd, 1 Gunpowder Square, London EC4A 3DE 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.” USA Taylor & Francis Inc., 1900 Frost Road, Suite 101, Bristol, PA 19007 Copyright © Taylor & Francis Ltd 1996 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. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library. ISBN 0-203-48314-6 Master e-book ISBN

ISBN 0-203-79138-X (Adobe eReader Format) ISBN 0-7484-0443-0 (Print Edition) (cloth) Library of Congress Cataloguing Publication data are available Cover design by Jim Wilkie The cover figure shows the crystal structure of cytochrome P450cam, where the ribbon indicates the folding of the polypeptide chain with the camphor substrate at the centre in front of the heme group.

Contents

1

Acknowledgements

ix

Abbreviations

x

Chemical Abbreviations

xii

Foreword by Professor Dennis Parke

xiv

Preface

xvii

Captions for Colour Plates in Chapter 6

xviii

Introduction

1

1.1

Historical background

1

1.2

Distribution

3

1.3

Isolation, purification and characterization of P450s

4

1.4

Spectral and related physicochemical properties

7

1.4.1

Substrate-binding spectra

13

1.4.2

Polarized optical spectroscopy

18

1.4.3

Vibrational spectroscopy

20

1.4.4

Magnetic resonance spectroscopy

26

1.4.5

Mössbauer (MB) spectroscopy

31

1.4.6

EXAFS spectroscopy

33

1.5

Other physical methods

35

1.5.1

X-ray crystallography

35

1.5.2

Redox potentials and their equilibria

36

1.5.3

Other physicochemical and structural studies

39

2

Evolution of the P450 Superfamily

51

2.1

Introduction

51

2.2

The P450 phylogenetic tree and evolutionary aspects

52

vii

2.3

Evolutionary relationships between prokaryotic and eukaryotic P450s

56

2.4

Conclusion

61

2.5

P450 nomenclature

62

The P450 Catalytic Cycle and Oxygenation Mechanism

71

3.1

Introduction

71

3.2

The P450 catalytic cycle

71

3.3

Nature of the oxygenating species

79

3.4

Participation of active site amino acid residues in P450 oxygenations

83

3.5

Thermodynamics of P450 substrate binding and spin-state equilibria

84

3.6

The coupling of redox and spin equilibria

87

3.7

Redox potential and redox interactions in the P450 system

90

3.8

Interactions between redox components

94

3.9

Protein-lipid interactions in the P450 system

96

A proposed mechanistic scheme for the P450 catalytic cycle

98

3

3.10 4

P450 Substrate Specificity and Metabolism

102

4.1

Introduction

102

4.2

Endogenous metabolism

102

4.3

Endogenous steroid hydroxylation by hepatic P450s of families CYP1, CYP2 and CYP3

107

4.4

Exogenous metabolism

116

Induction, Regulation and Inhibition

153

5.1

Introduction

153

5.2

Induction and regulation of P450 genes

157

5.3

Regulatory elements in P450 genes

165

5.4

The toxic consequences of induction

167

5.5

Heterologous expression systems and in vitro models

171

5.6

Inhibition of cytochromes P450

173

Structural Models of P450s and Related Topics

190

6.1

Introduction

190

6.2

P450 modelling

190

6.3

Evaluation of P450-mediated toxicity by the COMPACT approach

262

5

6

viii

6.4

Conclusions and future prospects

266

Bibliography

272

References

273

Index

317

Acknowledgements

There is a fairly substantial number of people who have, knowingly or unknowingly, helped to produce this book. To list those whose names immediately spring to mind may cause offence to others and, therefore, I would like to express my thanks to all who know me, especially those within the School of Biological Sciences at Surrey. However, if I have to name someone who has been the major inspiration behind this work, it would be Professor Dennis Parke. I consider myself extremely fortunate to know and work with Dennis Parke, a true scientist of both vision and intellect, who is able to move freely between many diverse fields in the biological sciences, and to be a recognized international expert in several of them. Such men are rare and, in my opinion, Dennis ranks among the great scientists of our time. Finally, I would like to thank Val Saunders and all of those involved in the preparation of the manuscript for publication, including those at Taylor & Francis, and at the University of Surrey; in particular, the tireless industry of the Inter-Library Loans Section of the George Edwards Library.

Abbreviations

Å CYP D Da DNA e E° ESR eV EXAFS F FAD FMN G H HS INDO J k K LS MAO me MO Ms NADH NADPH PAGE

Ångstrom unit (10−8 cm) Cytochrome P450 Debye (3.33564×10−30 cm) Dalton (1.66×10−24g) Deoxyribonucleic acid Charge on the electron (1.60219×10−19 C) Redox potential (V or mV) Electron spin resonance Electron volt (96.485 kJmole−1) Extended X-ray absorption fine structure Faraday constant (96484.6 C) Flavin adenine dinucleotide Flavin mononucleotide Free energy change Enthalpy change High-spin Intermediate neglect of differential overlap Joule (1 calorie=4.184 Joules) Rate constant Equilibrium constant Low-spin Monoamine oxidase Mass of an electron (9.10953×10−31 kg) Molecular orbital Electron spin angular momentum quantum number Nicotinamide adenine dinucleotide Nicotinamide adenine dinucleotide phosphate Polyacrylamide gel electrophoresis

xi

QSAR R RNA S S SDS T UV

Quantitative structure-activity relationship Gas constant (8.31441 JK−1 mol−1) Ribonucleic acid Spin quantum number Entropy change Sodium dodecyl sulphate Absolute temperature (K) Ultra-violet

Note: Other abbreviations are, in general, referred to in the text; whereas the results of statistical analyses (regression equations) provide values of the correlation coefficient (R), standard error (S) and the variance ratio (F).

Chemical Abbreviations

A C 2-AAF AIA ANF BHA BHT BNF DDD DDT DEHP DEN DHEA DiMeIQx DMN DPEA Glu-P-1 Glu-P-2 IQ LTB4 MC MeA C MEHP MeIQ MeIQx NNK PB PCN PGA

2-amino-9H-pyrido-[2, 3-b]indole 2-acetylaminofluorene allylisopropylacetamide -naphthoflavone butylhydroxyanisole di-t-butylhydroxytoluene ß-naphthoflavone p, p-dichlorodiphenyldichloroethane p, p-dichlorodiphenyltrichloroethane di(2-ethylhexyl) phthalate diethylnitrosamine dehydroepiandrosterone 2-amino-3, 4, 8-trimethylimidazo[4, 5f]quinoxaline dimethylnitrosamine diphenylethylamine 2-amino-6-methyldipyrido[1, 2-a: 3 , 2 -d]imidazole 2-aminodipyrido[1, 2-a: 3 , 2 -d]imidazole 2-amino-3-methylimidazo[4, 5f]quinoline leukotriene B4 3-methylcholanthrene 2-amino-3-methyl-9H-pyrido-[2, 3-b]indole mono(2-ethylhexyl)phthalate 2-amino-3, 8-dimethylimidazo[4, 5f]quinoline 2-amino-3, 8-dimethylimidazo[4, 5f]quinoxaline 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanone phenobarbital pregnenolone-16a-carbonitrile prostaglandin A

xiii

PGE2 PhIP TAO TCDD 1-THC Trp-P-1 Trp-P-2

prostaglandin E2 2-amino-1-methyl-6-phenylimidazo[4, 5-b]pyridine troleandomycin 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin 1-tetrahydrocannabinol 3-amino-1, 4-dimethyl-5H-pyrido[4, 3-b]indole 3-amino-1-methyl-5H-pyrido[4, 3-b]indole

Foreword

The cytochromes P450 are now well established as important integral aspects of biology, their functions extending from hormonal regulation of metabolism to reproduction and evolution; they are also now recognized as having major inputs to medicine, from cancer to diabetes and hepatitis to surgical trauma. Yet they are seldom mentioned in biology textbooks, they are a mystery to most physicians and surgeons and appear to be unknown to those writers of popular science and presenters of TV programmes beguiling viewers with the latest medical marvels of this technological age. So why is this so? Why is this superfamily of enzymes, probably of greater importance to man’s being than even DNA, not regarded with far greater respect and interest by professional scientists and by the general public at large? The reason is probably two-fold. First, these biological entities which were extant some 3500 million years ago, possibly even before the evolution of DNA, and whose existence was appreciated by 18th century scientists, were not finally recognized until some 30 years ago. Secondly, they were a paradox, like God, being one in its functional unity but infinite in its diversity of purpose, and for several decades biologists pondered over whether this was a single entity or a collection of closely related enzymes. The primal biological role of cytochrome P450 is considered to be the detoxication of tissue dioxygen (O2), ultimately to form water. With the advent of oxygen in the earth’s atmosphere and the evolution of aerobic organisms, the toxic properties of singlet oxygen and other reactive oxygen species (ROS) became manifest, resulting in biological dysfunction, tissue damage and death. Consequently, the life-spans of biological species are believed to be determined primarily by oxygen toxicity. A paradox of this is that while cytochrome P450 can detoxicate molecular oxygen by activating it to superoxide anion which is then converted to water by superoxide dismutase and catalase, the superoxide may, alternatively, be converted into hydroxyl radicals which, like singlet oxygen, can wreak severe biological damage, tissue injury and death. Although a major source of these fatally toxic ROS in mammalia are the cytochromes of the mitochondria, this may not have been so in primitive unicellar organisms, where iron and other redox metals could have been responsible. As these primitive simple organisms evolved, a progressive differentiation into photosynthesizing (plants) and parasitic (animals) species occurred, which led to an era of co-evolution, or plant/animal warfare, lasting millions of years, and greatly accelerating the process of evolution. This resulted in the plant species synthesizing protective toxins (phytoalexins) to prevent their own consumption by animal species, and in turn caused the animal species to elaborate defensive measures to detoxicate the phytoalexins and thereby ensure survival. In both of these evolutionary phenomena, namely, phytoalexin biosynthesis, and oxidative detoxication, cytochrome P450 played a major role, evolving into a vast array of closely related, but distinctively different, enzymes, comprising what is known as a ‘multifamily’. Hence, the cytochromes P450 are largely responsible for the phenomenal acceleration of evolution which occurred more than a

xv

thousand million years ago, and have resulted in the earth today being populated with modern life forms instead of the primitive organisms of the primaeval mire. Thus, the problem of one of many enzymes is explained. Initially there was one single enzyme, which activated molecular oxygen to superoxide, this being the first stage in the detoxication of oxygen to water. Then, as more complex molecules were synthesized by the evolving biota, the activated oxygen was incorporated into these new products, building the unpalatable, toxic phytoalexins which protected the primitive plant forms from their predators. Similarly, the activated oxygen of the cytochrome P450 was used to oxidatively metabolize the lipophilic phytoalexins, to make them more susceptible to excretion or oxidative destruction. Hence, it was the versatility of cytochrome P450 to undergo extensive diversification, with consequent differences in the nature of substrate molecules metabolized, and variations in the site at which oxygen is inserted, that enabled biological evolution to occur in the way, and at the pace, that it did. Put ingenuously, it is the cytochromes P450 that are ultimately responsible for man’s being, as they are also responsible for the biosynthesis of the different sex hormones, they are responsible for woman’s existence too. Hence, this vast superfamily of enzymes, the cytochromes P450, comprises several families of enzymes occurring in plants and microorganisms, which contribute to the synthesis of macromolecules vital for their biological survival, such as the phytoalexins and phytosterols; and several families in animal species, concerned in the oxidative detoxication of phytoalexins, drugs, and other xenobiotics. In addition, there are other families also found in animals, which are concerned in the oxidative metabolism of endogenous substrates, such as fatty acids and steroids. Family 1 of the cytochromes P450 (CYP1) appears to have some unique role in reproduction and development, not yet fully elucidated; the polycyclic aromatic hydrocarbons (PAH), and other products of combustion, are its preferred substrates, its genomal regulation is coupled to that of DNA replication, mitosis and growth, and lastly the PAH substrates of CYP1 are structurally very similar to DNA, for which they may have acted as an evolutionary template. This all indicates that cytochrome P450 may have had a role in the evolution of DNA, and that CYP1 continues to have a regulatory role for this core of genetic information. Family 2 is concerned with the oxidative detoxication of animals of phytoalexins, drugs and many man-made chemicals. Similarly Family 3 oxidatively detoxicates large phytoalexins, such as alkaloids; Family 4 oxygenates fatty acids and esters which are resistant to metabolism by -oxidation such as the erucic acid of rapeseed together with phthalate and adipate esters. The remaining families likewise show specificity for different substrates, but all P450s have the same oxygen activating system, enabling these enzymes to insert oxygen into a variety of different compounds. This activation of oxygen is a potentially dangerous phenomenon which could lead to the death of the living organisms if not tightly regulated. In most cases this regulation is accomplished by a redox spin-state mechanism which prevents the P450 from activating oxygen unless it has bound a substrate molecule to act as recipient for the reactive-oxygen insertion; in this way the unlimited production of superoxide anions and other highly toxic ROS is averted. However, no system is perfect, and all P450 oxygenations are accompanied by some release of ROS, a process known as futile cycling; this is considered to play a major role in normal cellular turnover, a consequence of the high cytoxicity of ROS. An exception to this redox spinstate regulation of oxygen activation is seen with cytochrome P4502E1, a cytochrome P450 of Family 2, which is specific for small substrates that are oxygenated with difficulty, such as ether, acetone, ethanol and benzene. It is believed that these substrates ‘stabilize’ the CYP2E1 so that it activates oxygen and that the substrates are subsequently oxygenated by the ROS thus produced. Since ROS are so highly toxic, resulting in oxidative stress, chronic inflammation, malignancy and death, all events which stimulate CYP2E1 activity such as uncontrolled diabetes, fasting, excessive alcohol consumption, exposure to halogenated chemicals and anaesthetics, should consequently be rigorously avoided.

xvi

Another characteristic of the P450 enzymes is their ability to self-regulate, by enzyme stabilization, or by transcriptional or translational regulation of enzyme protein synthesis. When one P450 is highly active because of exposure to a certain substrate, that particular P450 protein is enhanced and, presumably because of the simultaneous futile cycling which occurs, other P450s are destroyed. This is certainly the case with CYP2E1, and high exposure to ethanol can lead to high losses of other CYPs. Enhanced activity of CYP1 and CYP4, by induction of de novo enzyme synthesis, is associated with cytosolic receptors which bind with the specific substrates to effect de-repression of genomal transcription. So, just as this superfamily of enzymes, because of the unique conformational and electronic characteristics of each individual enzyme isoform, exhibits a high degree of substrate specificity, there are likewise for some isoforms corresponding specific regulatory, cytosolic receptors, which exhibit similar, conformation-dependent substrate specificity. This results in a highly versatile, efficient and economical system, whereby biological needs are identified, resources are then mobilized and non-essential structures cannibalized to enhance resources and prevent the dispersion of energies. The requisites for such an operational matrix are: (1) a highly specific recognition system, for the diversity of substrates (enzyme conformation and electronic structure), (2) a directional regulating system for the activation, and substrate-insertion, of the oxygen (active-site conformation, and redox spin-state), and (3) a self-limiting, auto-destruct system for limiting competing enzymes, and for directing resources (electrons) to the P450 of choice. In all of these enzyme features, the conformations and electronic structures of the individual P450s, and especially of their active sites, are of paramount importance. The amino acid sequences of many P450s, of various biological species, have now been established, by molecular biologists, and crystallographers have determined the X-ray crystal structure of a few P450s from bacterial species. This has enabled molecular toxicologists, especially the author of the present volume, to relate chemical toxicity to specific cytochromes P450. This involves the determination of substrate molecular dimensions and electronic structures by molecular orbital calculations and computer graphic interpretation of the data. With the amino acid sequence information and crystallographic data now available, the conformations of the different P450s can be visualized in three-dimensional colour; similarly, interactions with a variety of substrates, with DNA, and other biological molecules can be reproduced in 3-D on a graphics work-station. This is a major development in molecular biology that will ultimately accelerate the progression of molecular toxicology to enable the visualization of toxic, and pathological, processes at the molecular level, and thereby to make predictions concerning the potential toxicity, or safety of a chemical, solely from molecular orbital calculations. In these elegant developments in molecular toxicology, Dr David Lewis, the author of this erudite exposition The Cytochromes P450: Structure, Function and Mechanism, has been one of the foremost pioneers. His numerous studies of the cytochromes P450 have been creative and unique, paving the way to a refreshing new approach to biochemistry and toxicology. This comprehensive and foresighted treatise on the cytochromes P450 is a major work of scholarship and presents a preview of many future developments in the biological sciences. DENNIS V.PARKE June 1995

Preface

Research in the cytochromes P450 superfamily represents a rapid growth area of scientific endeavor. There are now more papers published in this field than in any other area of science. Consequently, a new text on cytochromes P450 appears every few years, containing more recent information and usually covering a proportion of the specific areas of interest. The fundamental importance of these enzymes in toxicology is rarely emphasized sufficiently by authors and editors; so it is hoped that, to some extent, this book will redress that balance. For a subject as vast and detailed as P450, it is impossible to be fully exhaustive in a book of this length, but the reader is directed to reviews and other references where further information may be obtained on those areas where the demands of space have limited the extent of coverage in the text. In writing this book, I have borne in mind the requirements of students in the biological sciences (such as biochemistry, pharmacology and toxicology) together with the needs of research workers in the chemical and pharmaceutical industries engaged in novel compound design and development, those involved with drug and other xenobiotic metabolism, and scientists in the governmental regulatory authorities concerned with the safety evaluation of chemicals. In particular, this volume will cover those aspects of the cytochromes P450 that are relevant to an understanding of their structure, function and mechanisms of action, such as physicochemical methods of characterization, redox interactions, evolution and genetics, experimental and theoretical models, induction, inhibition and metabolism, including pathways of activation and detoxication. Due to the weight of subject material, with over 1000 papers published per annum since 1986, it is only possible to provide an overview of many areas; but key references should direct the interested reader, seeking more in-depth information, to delve further into some of those facets of P450 research which have had to be condensed due to the requirements of space and brevity. D.F.V.LEWIS June 1996

Captions for Colour Plates in Chapter 6 Structural Models of P450s and Related Topics

Figure 6.3 Comparison between the C traces (a) CYP102 (blue) and CYP108 (green) and (b) CYP101 (magenta) and CYP108, using the heme moieties as reference points in the overlays. Figure 6.4 The distribution of the 36 invariant residues between the CYP101, CYP102 and CYP108 sequences shown as they are orientated in the CYP102 structure. Colour coding of amino acid residues is as follows: acidic-red; basic-blue; aromatic-yellow; amide-magenta; hydrophobic-pale blue; cysteine-orange; serine, threonine-blue-green; glycine, proline-green. These colour codes are generally used throughout unless otherwise stated. Figure 6.5 An overlay between the original CYP2B model and that containing phosphorylated serine-103 (coloured by atom type), showing the possible effect of serine phosphorylation on heme binding. The position of the original residues is shown coloured by amino acid type. Figure 6.7 Putative active site region of rat CYP1A1 with the substrate, 7-ethoxyresorufin, orientated for de-ethylation. Aromatic amino acid residues are shown in yellow, whereas the substrate is coloured according to atom type, i.e. carbon-white, oxygen-red, nitrogen-blue, hydrogen-pale blue. Residues mentioned in the text are numbered according to their alignment positions, and this is followed in other figures showing active sites. Figure 6.8 Putative active site region of rat CYP1A2 with the substrate, caffeine, orientated for Ndemethylation. Amino acid residues are labelled and coloured according to type, whereas the substrate is coloured by atom type. Hydrogen bonds are shown as dashed lines and the ribbon indicates folding of the polypeptide chain. Three orientations of caffeine are superimposed in the site, indicating the three Ndemethylations catalyzed by CYP1A2. Figure 6.9 Putative active site region of human CYP1A2 containing the caffeine substrate orientated for N3-demethylation. Colour coding is similar to that shown in Figures 6.7 and 6.8. Figure 6.11 Putative active site region of CYP2A1 showing the substrate, testosterone orientated for 7 hydroxylation (arrowed). Both enzyme and substrate are coloured by atom type. Figure 6.12 Putative active site of CYP2A6 with coumarin orientated for 7-hydroxylation. The heme and substrate and coloured by atom type. Figure 6.14 Putative active site of CYP2B1 showing the substrate androstenedione orientated for 16hydroxylation. The substrate is colour coded by atom type, whereas active site residues are coloured by type, as indicated in the legend to Figure 6.4. Figure 6.15 Close-up of the possible interaction between the FMN domain on reductase and the heme vicinity of CYP2B4. Amino acid residues are coloured by type, whereas the FMN and heme moieties are coloured magenta.

xix

Figure 6.17 Putative active sites of CYP2C3 (a) CYP2C3v (b) with the progesterone substrate orientated for 16 -hydroxylation (a) and 6 -hydroxylation (b). Atoms are coloured by atom type with the site of hydroxylation shown in green. Figure 6.18 A representation of the two substrates, tolbutamide and mephenytoin, overlaid with the putative active site of CYP2C9 where the substrates and active site region are coloured by atom type. Figure 6.20 Putative active site of CYP2D6 showing the substrate, debrisoquine orientated for hydroxylation in the 4-position. The substrate and active site structures are coloured by atom type. Figure 6.21 The substrate, deprenyl, docked in the putative CYP2D6 active site showing how orientation in the heme region can rationalize metabolism of the substrate. Atoms are coloured according to atom type. Figure 6.23 Putative active site of CYP2E1 showing an orientation of the substrate, p-nitrophenol, which rationalizes the known position of metabolism. The substrate and interacting active site residues are coloured according to atom type. Figure 6.24 Putative active site region of CYP3A4 showing the substrate, cyclosporin, (coloured by atom type) oriented for oxygenation in one of the known positions. Amino acid residues are coloured by type. Figure 6.26 The inhibitor, ketoconazole, is shown docked into the putative active site region of CYP3A4, where amino acid residues orientate the inhibitor such that its imidazole ring can ligate the heme iron. Ketoconazole is coloured according to atom type. Figure 6.27 The substrate, granisetron, is shown docked into the putative active site of CYP3A4, and orientated by certain amino acid residues such that N-demethylation of the substrate is favourable. Figure 6.29 Putative active site of CYP4A11 showing the substrate, lauric acid, (coloured by atom type) orientated for end-of-chain hydroxylation. Amino acid residues are coloured by type. Figure 6.30 Putative active site of CYP4F3 showing the substrate, leukotriene B4, docked into the active site where certain amino acid residues orientate the substrate for oxygenation in the known position. Substrate and active site residues are coloured by atom type. Figure 6.31 The endogenous substrate, cholesterol, docked into the putative active site of CYP11A1 showing how amino acid residues orientate the substrate for oxygenation at the known position. The substrate (without hydrogen atoms) is coloured by atom type. Figure 6.32 A possible mode of interaction between adrenodoxin (left) and CYP11A1 (right) showing colour-coded dot surfaces on the ion-pairing residues. Figure 6.33 The endogenous substrate, progesterone, docked into the putative active site of CYP17A1 showing that amino acid residues (coloured by type) can orientate the substrate (coloured by atom type) for oxygenation. Figure 6.34 Active site region of CYP17A1 containing an inhibitor superimposed on the substrate, progesterone, where the amino acid residues are coloured by type and the substrate/inhibitor are coloured by atom type. Figure 6.35 The inhibitor, 4-hydroxyandrostenedione, (coloured by atom type) is shown docked into the putative active site region of CYP19A1, where interactions with key amino acid residues (coloured by type) orientate the inhibitor for heme ligation. Figure 6.36 A possible mode of interaction between putidaredoxin (left) and CYP101 (right) showing colour-coded dot surfaces on the interacting residues. Basic residues are shown in blue, whereas acidic residues are displayed in red, with the C-terminal tryptophan of putidaredoxin high-lighted as a dot surface (magenta). Figure 6.37 The two iron-sulphur redoxins, putidaredoxin and adrenodoxin, superimposed showing their respective regions of interaction with the relevant P450 and reductase displayed as dot surfaces coloured red and purple, respectively.

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Figure 6.38 The heme binding pocket of CYP102 showing the substrate lauric acid orientated for -2 hydroxylation by key active site amino acid residues. Certain distances between substrate atoms and the heme iron are in close agreement with NMR data. Figure 6.39 Active sites of CYP108, CYP101 and CYP102 (viewing from left to right) showing the variation in steric crowding of the heme moiety (magenta) by various amino acid residues (coloured by type). Figure 6.40 Active site region of CYP108 showing a possible orientation of the substrate, -terpineol, with key amino acid residues and the heme moiety which is consistent with the known position of hydroxylation.

1 Introduction

1.1 Historical background The story of how cytochrome P450 was first discovered begins about 40 years ago with the work of Axelrod (1955) and Brodie et al. (1955) who reported that an enzyme system in the liver endoplasmic reticulum was able to metabolize oxidatively certain xenobiotic compounds. Later that year, Hayaishi and co-workers showed that dioxygenase reactions were possible in liver microsomal preparations (Hayaishi et al., 1955) whereas Mason et al. (1955) demonstrated that monooxygenase activity was present in the same system, which utilized NADPH as a reductant. However, the detection of a carbon monoxide (CO) binding pigment in liver microsomes, giving an absorption maximum at 450 nm, was made independently by Garfinkel (1958) and Klingenberg (1958), who are generally accredited with the discovery of P450. Omura and Sato were later able to demonstrate that this pigment was, in fact, a hemoprotein of the b-type cytochrome class (Omura and Sato, 1964a and b) and these co-workers first coined the term cytochrome P450, after the wavelength of the UV absorption maximum (Figure 1.1) in the optical spectrum for the CO complex of the cytochrome pigment (Omura and Sato, 1962). These workers and others subsequently showed that the position of the characteristic Soret band could be shifted by the binding of substrates to the enzyme (Schenkman et al. 1967a and b; Schenkman, 1970) or by treatment with detergent (Omura and Sato, 1962); the latter bringing about a conversion to an inactive solubilized form of the enzyme which produced an absorption maximum at 420 nm in the CO-difference spectrum. The effect of substrate binding on the UV absorption characteristics of cytochrome P450 was of particular importance, as it appeared that different types of substrate elicited differing varieties of spectral change; namely, types I, II and modified type II (Schenkman et al., 1967a and b; Schenkman, 1970; Schenkman et al., 1972). The type I change brought about by apolar substrates involved a blue shift (hypsochromic) in the Soret band, whereas type II substrates gave rise to a red shift (bathochromic) thought to be associated with ligation of the heme iron. Modified type II (or reverse type I) binding gives rise to a UV spectrum which has the appearance of a ‘mirror image’ of the type I spectral change, and this has been attributed to interaction between the substrate and an alternative site on the enzyme. Estabrook and colleagues showed that cytochrome P450 is the terminal oxygenase in the adrenal corticoid system for the C21-hydroxylation of steroids (Estabrook et al., 1963; Cooper et al., 1965) and this role of P450 as the terminal oxidase was subsequently confirmed for the liver microsomal system (Diehl et al., 1969) in the endoplasmic reticulum. The work of Gunsalus and others on bacterial P450cam (Katagiri et al., 1968; Tyson et al., 1972) Kimura and Suzuki on adrenal mitochondrial P450 (Kimura and Suzuki,

2

THE CYTOCHROMES P450

Figure 1.1 Absorption spectrum of P450.CO showing the characteristic Soret peak around 450 nm (reproduced using the data of Estabrook et al. (1963) for P450C21).

1965; Suzuki and Kimura, 1965) and Coon and co-workers on liver microsomal P450 (Lu and Coon, 1968; Lu et al., 1969) established the redox components in these particular electron transport chains as containing an NADH (Fisher and Gaylor, 1982) or NADPH-dependent flavoprotein (Kuby, 1991) which transfers two reducing equivalents (lyanagi, 1987) to cytochrome P450 during two one-electron transfer processes that convert dioxygen to water, with concomitant single oxygen atom insertion into the substrate molecule (Walsh, 1979). The major difference between the liver microsomal system is the absence of any mediating iron-sulphur protein (redoxin), which was found to be required by both the bacterial (putidaredoxin) and adrenal mitochondrial (adrenodoxin) systems (Peterson and Mock, 1979; Hintz and Peterson, 1981; Hintz et al., 1982; Jefcoate, 1986). Additional information was provided by ESR (electron spin resonance) spectroscopy, which suggested that cytochrome P450 is a low-spin ferric hemoprotein (Hashimoto et al., 1962) with a thiol residue as an axial heme ligand (Bayer et al., 1969; Hill et al., 1970a and b). These findings led to explanations for the unusual Soret peak position and its perturbation by substrates and other chemicals (Hanson et al., 1976, 1977; Gunsalus and Sligar, 1978) in terms of charge transfer transitions modulated by the effect of the thiol/ thiolate ligand. Further evidence for an iron-sulphur bond in P450 was obtained from resonance Raman spectra of P450cam (Champion et al., 1982) and finally confirmed as being formed by a covalently bound cysteine residue, following the publication of the X-ray crystal structure of P450cam (Poulos, 1985; Poulos et al., 1985). For further information on the discovery and elucidation of P450, together with additional aspects, the reader is referred to Sato and Omura (1978), Ruckpaul (1978), Schenkman and Kupfer (1982), Ruckpaul and Rein (1984), Ortiz de Montellano (1986a), Archakov and Bachmanova (1990), Omura et al. (1993) and

INTRODUCTION

3

Schenkman and Griem (1993). These accounts provide details of many features of P450 structure and function, with additional references too numerous to be included here; and those who are particularly interested in the role of P450s in drug metabolism are referred to the book by Gibson and Skett (1994), now in its second edition. 1.2 Distribution Although cytochrome P450 (EC 1.14.14.1) was originally discovered in mammalian liver microsomal preparations, it has subsequently been found in many organs and tissues of many other animals, and in some plants, fungi and bacteria. To date, over 230 individual P450s have been characterized according to their protein sequences, and forms of these enzymes appear to be present in every class of biota, including mammalia, birds, fish (Andersson and Förlin, 1992), reptiles, amphibia, insects (Feyereisen, 1993; Cohen and Feyereisen, 1995), plants (Bozak et al., 1990; Durst, 1991; Hallahan et al., 1992), bacteria and fungi (Nelson et al., 1993). The distribution of P450 proteins in mammalian systems (Waterman, 1992) has been investigated in detail and it has been established that they are present mainly in the liver (Paine, 1981) but are also found in the kidney, lung (Smith et al., 1982; Arinc, 1993), gonads, adrenals, brain (Warner et al., 1994), nasal epithelium (Dahl and Hadley, 1991), placenta (Pasanen and Pelkonen, 1989), pancreas, spleen, gastrointestinal tract (Kaminsky and Fasco, 1992) and skin (Mukhtar and Khan, 1989). Further details of the distribution of P450s in various species and in mammalian tissues can be obtained from the recent publication edited by Schenkman and Griem (1993) although it is possible to gain an impression of the ubiquity of P450 proteins from an inspection of the tabulated sequences as catalogued by Nelson and colleagues (Nelson et al., 1993). This compilation provides an update on the new nomenclature of all the P450 sequences obtained thus far, and Table 1.1 has been produced from such information so that the wide distribution of the P450 superfamily in nature can be appreciated; a more detailed version is provided in Chapter 2. Table 1.1 summarizes the various P450 genes/proteins known, according to both the species, the P450 family and subfamily, under which they have been classified. The concentrations of cytochrome P450 in various tissues of mammalia are presented in Table 1.2 (Vainio, 1980) and Table 1.3 shows the relative amounts of P450s in rat liver Table 1.1 P450s present in different species (a to h) (Reference: Nelson et al., 1993) Families

Subfamilies

(a)

Mammalia CYP1 CYP2 CYP3 CYP4 CYP5 CYP7 CYP11 CYP17 CYP19 CYP21 CYP24

2A

2B

2C

4A

4B

4F

11A

11B

2D

2E

2F

2G

2J

2K

4


Families (b)

(c)

(d)

(e) (f) (g) (h)

Subfamilies CYP27 Birds CYP1A2 CYP2H CYP11A Fish CYP1A1 CYP11A CYP17 CYP19 Insects CYP4C, 4D and 4E CYP6A, 6B Molluscs CYP10 Fungi CYP51–57 Plants CYP71–73 Bacteria CYP101–112

(Schenkman and Griem, 1993), whereas Tables 1.4–1.6 show, as an example, the organ distribution of P450 activity towards 7-ethoxy coumarin de-ethylation in the mouse (Ullrich, 1977), in other rodent species (Table 1.5) and various marine organisms (Table 1.6). Clearly, the distribution of these enzymes in biological systems is correctly labelled as being ubiquitous, especially as there are certainly many more P450s present in different species which have yet to be discovered. 1.3 Isolation, purification and characterization of P450s The isolation and purification of various forms of P450 depend on the source of the enzyme, for example, whether it is cytosolic or membrane-bound, and these methods have been extensively described elsewhere (Sato and Omura, 1978; Ruckpaul and Table 1.2 Cytochrome P450 content* in various mammalian tissues (Reference: Vainio, 1980) Mammalian species Organ

Rat

Mouse

Rabbit

Guinea pig

Man

Liver Kidney Lung

0.22–0.92 0.05–0.21 0.035

0.39–1.10 0.40 —

0.81–1.70 0.14–0.36 0.27–0.38

0.43–1.45 0.32 0.07

0.26–1.02 0.03 —

INTRODUCTION

5

Mammalian species Organ

Rat

Mouse

Rabbit

Guinea pig

Man

Intestine 0.02–0.13 0.04 0.07–0.43 0.18 Adrenal gland 0.50 — 1.20 2.0 Testis 0.05–0.10 0.24 0.04 0.078 Skin 0.05 0.022 — — Spleen 0.025 — — — Ovary — — 0.06 — Brain 0.025–0.051 — — — Note: * expressed as nmol/mg microsomal protein—data not available or not known.

— 0.23–0.54 0.005 — — — —

Table 1.3 P450 content in untreated rat liver microsomes (Reference: Funae and Imaoka, 1993) Contenta

P450 family/subfamily

Totala

CYP 1A1 0.001 0.012 CYP 1A2 0.011 CYP 2A1 0.008 0.054 CYP 2A2 0.046 CYP 2B1 0.002 0.019 CYP 2B2 0.017 CYP 2C6 0.88 0.650 CYP 2C11 0.437 CYP 2C12 0.002 CYP 2C13 0.123 CYP 2E1 0.079 0.079 CYP 3A2 0.146 0.146 14.6 CYP 4A1 0.022 0.030 CYP 4A2 0.008 Notes: a Expressed as nmol P450/mg protein; b Percentage of total P450 measured.

Percentageb 1.2 5.4 1.9 65

7.9 3.0

Rein, 1984; Ryan and Levin, 1990; Archakov and Bachmanova, 1990). As far as the mammalian hepatic microsomal forms are concerned, extensive purification procedures are required following the preparation of the microsomal fraction from liver homogenates. The experimental techniques involved in producing microsomal suspensions from tissue homogenates are detailed in a recent publication by Gibson and Skett (1994). This volume also gives descriptions of the various protocols for spectral determination of P450 content, and provides the practical details involved Table 1.4 7-Ethoxycoumarin O-de-ethylase* activity in various organs of the mouse (Reference: Ullrich, 1977) Organ

Activity (nmol/min)

% of total activity

Liver Small intestine Skin

147 15 7

86.5 8.5 4

6


Organ

Activity (nmol/min)

% of total activity

Lung 0.7 Kidney 0.1 Brain 0 Heart 0 Note: * 7-Ethoxycoumarin O-de-ethylation is associated with CYP1 activity.

1 0.1 0 0

Table 1.5 7-Ethoxycoumarin O-de-ethylase activity in the organs of three rodent species (Reference: Vainio, 1980) Organ

Guinea pig

Liver 100 Kidney 0.7 Lung 12.6 Testis 0.3 Spleen 0.3 Adrenal gland 5.3 Duodenum 3.6 Note: Activities are relative to that in the liver expressed as 100%.

Rat

Mouse

100 2.0 5.8 0.5 0.6 5.8 2.4

100 6.7 2.8 0.8 0.2 5.0 3.8

Table 1.6 7-Ethoxycoumarin O-de-ethylase activity in various marine species (Reference: Ullrich, 1977) Species

Organ

Specific activity (nmol/mg protein)

Myoxocephalus scorpius

Liver Intestine Liver Hepatopancreas Intestine Hepatopancreas Hepatopancreas Hepatopancreas

0.21 0.60 0.08 0.04 0.02 0.06 0.11 0.014

Zoarces viviparus Carcinus maenas Eupagurus bernhardus Buccinium undatum Ciona intestinalis

in characterizing P450 substrate binding by UV absorption spectrophotometry; methods for the assay of P450-mediated catalytic activity are also described, together with those relating to other enzymes in the microsomal system. Complete isolation of highly purified P450s from solubilized microsomes is a rather lengthy process, involving a succession of chromatographic, electrophoretic and other procedures which lead to progressively refined preparations (Archakov and Bachmanova, 1990). Separation of distinct bands, corresponding to individual P450 isoforms, using SDS-gel electrophoresis is a common method of characterization according to apparent molecular weight; most P450s lie in the 46–57 kDa region, but other enzymes, such as epoxide hydrolase and flavin monooxygenase can also be present. The techniques of isoelectric focusing and high-performance liquid chromatography using ion-exchange resins have been utilized to produce immunochemically homogenous P450s, which can be characterized according to their spectral and catalytic properties (Ruckpaul and Rein, 1984). Table 1.7 shows the molecular weight values of various P450s estimated from SDS-PAGE measurements compared with sequence data. It can be seen

INTRODUCTION

7

from Table 1.7 and Figure 1.2 that there is, in general, a good agreement between the two methods, especially for the bacterial form, P450cam, although the number of amino acid residues in the sequence also gives a high correlation with molecular weight (Table 1.7 and Figure 1.3). In addition to the determination of molecular weight, the percentage amino acid composition appears to vary somewhat between individual isoforms (Black and Coon, 1986), but only a complete protein (or gene) sequence analysis will show whether a particular P450 is unique. However, there are significant differences in the N-terminal and Cterminal 20–40 residues of various P450s which appear to be quite characteristic (Black and Coon, 1986) and these may reflect variations in substrate specificity (Tsujita and Ichikawa, 1993). Nevertheless, primary sequence homology is an important determinant of P450 classification (Nelson et al., 1993) and a number of P450 families and subfamilies have been investigated for commonalities (Fujii-Kuriyama et al., 1989a; Ouzonis and Melvin, 1991; Degtyarenko, 1992) Table 1.7 Comparison between SDS-PAGE data and molecular weight values for P450s (Reference: Black and Coon, 1986) P450 form

CYP

No. of residues

SDS-PAGE

MWt. (apoprotein)

Rabbit 2 2B4 491 49 500 55 721 Rat b 2B1 491 51 200 55 941 Rat e 2B2 491 51 900 55 924 Rat c 1A1 524 54 300 59 401 Rat d 1A2 513 53 000 58 207 Mouse 1 1A1 521 55 000 58 923 Mouse 3 1A2 513 55 000 58 192 Bovine SCC 11A1 481 51 700 56 406 P.putida CAM 101 412 46 000 46 205 SDS-PAGE=M.Wt. determined by SDS-Polyacrylamide gel electrophoresis Correlations between M.Wt. and no. of residues, and between SDS-PAGE estimates Notes: M.Wt.=113.8 Residue number R=0.9877 (Figure 1.3) (±6.9) M.Wt.=1.08 M.Wt. (PAGE) R=0.9236 (Figure 1.2) (±0.198)

MWt. (holoenzyme) 56 373 56 593 56 576 60 053 58 859 59 575 58 844 57 058 46 857

The difference between the M.Wt. of the holoenzyme and apoprotein is equivalent to that of the heme group. Apart from P450cam, the SDS-PAGE M.Wt. is consistently lower than the true value, probably corresponding to the 20–40 residue transmembrane N-terminal segment. in their protein sequences, as presented in Chapter 6, which contains further details of sequence homology alignments. 1.4 Spectral and related physicochemical properties The cytochromes P450 are hemoproteins, containing a single heme prosthetic group (iron protoporphyrin IX) with the central iron atom ligated to a cysteinyl residue which is situated close to the C-terminus of a single polypeptide chain of apoprotein composed of between 400 and ~ 500 amino acids. To date, only three P450s have been fully characterized by X-ray crystallographic determinations (Poulos, 1985, 1986,

8


Figure 1.2 Relationship (using the data presented in Table 1.7) between MWts of various P450 proteins and those obtained from SDS-PAGE determinations.

Figure 1.3 Correlation (using the data presented in Table 1.7.) between MWts of various P450 proteins and the relevant number of amino acid residues.

1988a and b; Poulos et al., 1985, 1986, 1987; Poulos and Howard, 1987; Raag et al., 1990, 1991, 1993; Raag and Poulos, 1989a and b, 1992; Ravichandran et al., 1993; Boddupalli et al., 1992; Hasemann et al., 1994) although over 230 P450s have been sequenced (Nelson et al., 1993) and many more have been identified by spectral analysis and other physicochemical or biochemical techniques. Many of the particular and different characteristics of various P450s result from interactions between the apoprotein in the active

INTRODUCTION

9

site region (Koymans et al., 1993a and b) and the heme group as this modifies catalytic activity, substrate specificity, redox potential, spin-state equilibria and spectral properties (Sato and Omura, 1978). To fully appreciate this, it is necessary to consider the influence of biologically relevant ligands on the electronic states of iron, and under different field symmetries (Hanzlik, 1976; Murray et al., 1985). Being the sixth element of the first Transition Series, iron contains six electrons in its 3d shell in addition to two 4s electrons in the valence shell (da Silva and Williams, 1991). The chemistry of iron mainly centres around the two oxidation states of ferrous (Fe2+), or Fe(II), and ferric (Fe3+), or Fe(III), brought about by either loss of the 4s2 electrons or an additional 3d electron, giving rise to 3d6 (Fe2+) and 3d5 (Fe3+) configurations, respectively. The ferric electronic state is rather more stable than that of ferrous due to the half-filled 3d shell configuration, which can preferentially adopt the spin-free (high-spin) state. In aqueous solution, therefore, iron(II) compounds readily become oxidized by atmospheric oxygen to form the corresponding iron(III) state, and this is accompanied by a colour change from pale green, for iron(II)hexaaquo, to pale yellow for iron(III)hexa-aquo. The colours of many transition metal compounds arise from the absorption of light in certain regions of the visible spectrum, corresponding to electron transitions within the d shell of the transition metal ion (Cotton and Wilkinson, 1972). Normally, the d orbitals are degenerate in the free gaseous ion of the transition element but, under the influence of the electron field caused by the proximity of coordinated ligands, these energetically equivalent orbitals become split into, usually, two groups, which will facilitate electronic transitions between them. The d orbital splitting pattern depends upon the nature of the ligands (Yamatera, 1958) and their orientation in space with respect to each other and the metal ion, i.e. their symmetry. In the case of iron in hemoproteins, we need only concern ourselves with the octahedral (Oh) ligand field symmetry and its variants, such as the square pyramidal (C4v) case, which arise essentially as distortions from the purely octahedral environment (Figure 1.4). In the majority of hemoproteins, iron can exist in either the ferrous or ferric redox states (and maybe others) as well as at least two possible spin states, commonly referred to as low spin and high spin. For example, ferrous iron can exist in the low-spin state with all six 3d electrons paired (S=0), or in the highspin state with two 3d electrons paired and the remaining four unpaired (S=2). Ferric iron can similarly adopt the low-spin configuration where four 3d electrons are paired and the remaining one unpaired (S=½), or the high-spin state with all five 3d electrons unpaired (S=5/2). Although intermediate spin states may also occur for both redox states, these are not normally found in biological systems. Furthermore, the iron spinstate preferences depend on the nature of the coordinating ligands and the geometry/symmetry of the metalligand complex. In general, however, six-coordinate (Oh) heme iron is low-spin, and five-coordinate (C4v) heme iron adopts the high-spin configuration. The reason for this is, apparently, that for both redox states, the ionic radii of iron are greater for high-spin than for low-spin such that, in the high-spin cases, the iron will preferentially move out of the plane of the porphy-rin ring as the central cavity is too small to accommodate the increased cationic size (da Silva and Williams, 1991). Table 1.8 illustrates this point with reference to the ionic radii of iron(II) and iron(III) in their low- and high-spin states, together with the cavity (or core) size of the porphyrin ring (Shannon and Prewitt, 1970). In five-coordinate square pyramidal (C4v) heme complexes, there is an opportunity for the iron to move out of the plane of the porphyrin ring, and this is clearly observed in the X-ray crystal structures of hemoproteins such as hemoglobin, myoglobin and cytochrome P450, where the iron atom moves about 0.4 Å out of the porphyrin ring plane in the five-coordinate state (Lippard and Berg, 1994; Bertini et al., 1994; Kaim and Schwederski, 1994). In biological systems, iron exists as a thermal equilibrium of the two spin states at physiological temperatures, although this is subject to variation depending on the ligands and other factors (da Silva and Williams, 1991). Hemoproteins, with their porphyrin ring system and axial ligands, appear to have been

10


Figure 1.4 Energy level diagram showing the d orbital splitting under the influence of various ligand field geometries (References: Loew 1983; Makinen and Churg, 1983; Palmer, 1983)

naturally ‘designed’ such that the iron is close to the ‘cross-over’ point between the two spin states and, in cytochrome P450, this position of equilibrium is modulated by substrate binding (Gibson and Tamburini, 1984). These spin state and redox state changes can be observed spectroscopically and techniques such as UV absorption spectrophotometry, can be used to monitor heme geometry and associated iron spin state changes accompanying substrate and oxygen binding to P450s (Guengerich et al., 1976; Coon and White, 1980; White and Coon, 1980). ESR spectroscopy has also been used to investigate spin state and other changes in hemoproteins (Blumberg and Peisach, 1971) although the signal corresponding to the high-spin component can only be readily observable at extremely low temperatures (4K) using liquid helium. In most hemoproteins, the axial ligands to iron may be either both histidine (as in cytochrome b5), histidine and methionine (as in cytochrome c) or cysteine, as in P450. Porphyrin is a good -donor and acceptor tetradentate ligand for iron and, therefore, the heme group acts as a cooperative unit (da Silva and Williams, 1991) which possesses certain properties that are important and relevant to its role in biological systems, such as its ability to accept electrons via - stacking with aromatic rings and the fact that, in the Fe (II) state, it will readily bind oxygen due to its -donor ability. Unfortunately, ferroheme will also have a high affinity for the highly toxic gas carbon monoxide, as well as for the cyanide ion, which are both good

INTRODUCTION

11

-acceptor ligands. In fact, the high affinity of reduced P450 for carbon monoxide, together with the unusual and characteristic spectral properties of the Table 1.8 Ionic radii of iron redox and spin states (Å) (Reference: Shannon and Prewitt, 1970) Fe3+

Fe2+

High-spin 0.64 0.77 Low-spin 0.55 0.61 Note: The diameter of the cavity at the centre of a porphyrin ring system is about 1.22 Å.

CO-P450 complex were instrumental both in the discovery of the enzyme and its name, which derives from the UV absorption maximum of the CO-complex (Sato and Omura, 1978). The axial heme ligands impart a number of properties to the hemoprotein which dictates its biological role, in addition to the means of detection by spectroscopic and other physical techniques. These properties include spin-state and redox equilibria, the ability to stabilize oxygen complexes, and redox potential, which is also modulated by the conformation of the apoprotein (Churg and Warshel, 1986). In the case of P450, many of its unusual properties are due to the cysteinyl fifth ligand which can exist in the thiol (-SH) or thiolate (-S−) state. In the normal resting state of the enzyme, the heme moiety of P450 comprises low-spin iron(III) primarily with bound cysteine as thiolate and (probably) water as the distal sixth ligand (Poulos et al., 1986; Wade, 1990). Being six-coordinate, the heme iron of P450 can be regarded as existing in an octahedral environment which will, in fact, be subject to an axial distortion as the iron-porphyrin bond distances are shorter than those between iron and either axial ligand, as observed in the crystal structure of P450BM3 (CYP102) for example (Ravichandran et al., 1993). Distortions from pure octahedral symmetry give rise to observable spectroscopic effects which enable the degree of axial (or tetragonal) distortion to be calculated. One of the most comprehensive studies of these tetragonal, or Jahn-Teller, distortion effects in hemoproteins has been carried out by Blumberg and Peisach (1971) using ESR spectroscopic measurements. In fact, the ESR spectra of P450s show that they represent a distinct class of hemoprotein in terms of the geometry of the heme iron, as affected by the nature of the apoprotein and its conformation (Blumberg and Peisach, 1971). However, considerable information has also been accumulated from optical spectroscopic techniques, mainly using UV absorption spectrophotometry. In terms of ligand characteristics, the thiolate ion is an extremely weak ligand and a ‘soft’ base, which is also a -donor; whereas, in contrast, water is a medium strength ligand and a ‘hard’ base, and carbon monoxide is a very strong ligand and -acceptor, as is oxygen although slightly less strong than CO (da Silva and Williams, 1991). Thiol ligands are slightly stronger than water, and imidazole (as in histidine) is stronger still, but significantly weaker than carbon monoxide. Other nitrogenous bases such as aniline and, presumably, porphyrin are of a strength which is intermediate between thiol and imidazole (Shimura, 1988). The ligand field strength will determine the degree of splitting of the iron d orbitals and also affect the spinstate equilibrium, such that a strong ligand field will bring about a relatively large d orbital splitting, and thus favour the low-spin state. These facts are summarized in Table 1.9, which shows that water, thiols and imidazole are approximately intermediate between the two extremes of thiolate and carbon monoxide. Consequently, one can calculate that, for example, a change of imidazole to thiolate ligation would be expected to shift the UV absorption band corresponding to a d d transition for a ferroheme-CO complex from about 420 to 450 nm. This is observed in practice when one compares (Hill et al., 1970a) the UV absorption spectra of the CO complexes of hemoglobin (histidine ligand) and P450 (cysteinate ligand). Furthermore, the change from P450 to P420 may be rationalized in terms of a protonation of the cysteine

12


Table 1.9 Some selected ligand field strengths (Reference: Shimura, 1988)

a

data are for Co(III) complexes. Cobalt(III) is d6 as is Iron(II) and it is possible to calculate the corresponding values for iron(II) by multiplying by a factor of 0.965. b estimated value using the data for 2,2 -bipyridine c the value for molecular oxygen, O , is likely to be higher and, in fact, will probably be of the order of the CN– figure 2 Notes: Thiolate (−CH2S–) is a very weak ligand, a soft base and a -donor. In contrast, H2O is a medium-strength ligand and a hard base. Carbon monoxide (CO) is a very strong ligand and a -receptor, as is O2 although slightly less strong than CO. The difference between imidazole and thiolate is about the same as that between thiol and thiolate, and both should produce about a 30 nm shift in the absorption maximum, as is observed when one compares HbCO with P450CO (i.e. shift from 420 450 nm), and P420 with P450 for the CO complex (i.e. a shift from 420 nm to 450 nm). Also, the loss of bound water on substrate binding to P450 will produce a calculated shift in max roughly equal to the observed value.

thiolate ligand to form a thiol (Hill et al., 1970a). However, an alternative explanation for this inactivation of P450 could involve heme ligation by a neighbouring basic residue, such as histidine or arginine (Pratt et al., 1995a). It has been shown that − * transitions in the heme moiety are mainly responsible for the major features of hemoprotein UV spectra and, in P450, the presence of the thiolate ligand makes a contribution to the overall electronic system by mixing sulphur p electrons with the heme molecular orbitals such that an electronic transition corresponding to the Soret absorption maximum at 450 nm in the UV spectrum of the CO adduct is observed (Hanson et al., 1976). It is thought that the unusual intensity of this absorption is due (charge-transfer) transition mixes to a process known as intensity ‘stealing’ whereby the thiolate sulphur with, and borrows intensity from, the porphyrin transition: this appears to be an example of the Fermi splitting of accidentally degenerate electronic states (Lewis, 1986). A comprehensive and detailed analysis of the UV absorption spectra of P450 and its CO complex can be obtained from the definitive work of Hanson et al. (1976, 1977) which provides spectral assignments and orbital energy level diagrams based on polarized electronic absorption spectra of P450cam (CYP101) and iterative extended Hückel MO calculations. More recently, Loew has reviewed the calculations of iron porphyrin optical spectra and electronic states, including those of P450 and other hemoproteins (Loew, 1983). The effect of ligands on the spin states and geometry of hemes has been reviewed by Scheidt and Gouterman (1983) and, in the same monograph, electronic absorption spectra and charge-transfer characteristics of hemoproteins have been extensively tabulated by Makinen and Churg (1983). It is clear that the unusual spectral properties of P450s in the UV/visible region are a direct result of the unique cysteinyl fifth ligand and its effect on the electronic properties of the iron-porphyrin (heme) unit which not

INTRODUCTION

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only give rise to the particular characteristics of P450 optical spectra that have facilitated its study, but also play a fundamental role in the redox and spin state equilibria which are important to the P450 catalytic cycle and oxygenation mechanism (Castro, 1980; Ortiz de Montellano, 1987; Babcock et al., 1992; Hawkins and Dawson, 1992). In particular, the thiolate ligand, being weak field, tends to favour high-spin Fe(II) over low spin, whereas porphyrin itself exerts a fairly strong ligand field, as shown in Table 1.9, such that the latter is close to the cross-over point between the two spin states. Furthermore, the thiolate (or possibly thiol) ligand probably stabilizes the reactive oxygen species, when bound to the heme (Hawkins and Dawson, 1992), so that oxygenation of the substrate can occur, possibly via cleavage of the iron-bound peroxide. 1.4.1 Substrate-binding spectra The UV spectral changes which accompany substrate interactions with P450s have been used to indicate certain binding characteristics, or so-called types of ligand binding (Table 1.10), of which there are three categories; namely, type I, type II and reverse type I (sometimes termed modified type II). These changes, which occur in the UV spectra following ligand-P450 complexation reactions, represent a method for classifying different varieties of P450 active site interactions and also for calcu-lating the modulation of hemoprotein spin state equilibria which can accompany substrate binding (Schenkman and Kupfer, 1982; Schenkman et al., 1981). Apparently, such effects also regulate the redox potential of the P450 involved (Sligar, 1976; Sligar et al., 1979) and, consequently, the rate of metabolism of the substrate (Blanck et al., 1983). In particular, the occurrence of the type I spectral change has been associated with a modulation of the ferric iron spin state equilibrium from low-spin to high-spin, accompanying substrate binding, enabling a calculation of the percentage high-spin content and the equilibrium constant for the process, from which thermodynamic data can be obtained. Specifically, a type I spectral change entails a reduction in the Soret absorption at 420 nm and a concomitant increase in the 390 nm absorption, which may be regarded as a shift in the hemoprotein spin equilibrium from low-spin to high-spin ferric P450. This is because the two spin state forms of native ferric P450 exhibit different Soret absorption maxima: the low-spin form absorbs at around 418 nm (ranging from 416 to 420 nm) whereas the high-spin form absorbs at around 390 nm (ranging from 385 to 394 nm) depending on the type of P450 involved. The isobestic point for the type I spectral change is at around 407 nm (with a range of 406–408 Table 1.10 Substrates of P450 exhibiting various types of spectral change (types I, II and reverse type I) (Reference: Schenkman et al., 1981) Type I substrates Aldrin Allyl neopentyl barbituric acid Allyl butyl barbituric acid Alkanes Aminopyrine Amobarbital d-Amphetamine (low concentration) Aprobarbital Arachidonic acid

2- and 4-Hydroxybiphenyls 2, 2-Hydroxybiphenyl 2-Hydroxydesmethylimipramine Imipramine Isobornyl acetate Isooctane Kelthane Lauric acid Lidocaine (low concentration)

14


Type I substrates Benzpyrene Benzphetamine Biphenyl Butobarbital Caffeine Camphor Carbon tetrachloride Chenodeoxycholic acid Chlordane Chloroethanes Chloroform p-Chloromercuribenzoic acid Chlorpromazine Cocaine Cortisol Cyclobarbital Cyclohexane 1-Cyclohexenyl pyrrolidine 1-Cyclopentyl pyrrolidine Deoxycholic acid Desmethylimipramine Dibutyl sulphide DDT Dieldrin Dihydrosafrole Diphenylhydantoin , -Dipyridyl Endrin Enflurane -Estradiol 5-Ethyl 5-alkyl barbiturates Ethyl morphine Fluoroxene Guthion Halothane Heptabarbital Heptachlor 2-Heptanone Hexane Hexobarbital

Lindane Linoleic acid Linolenic acid Malathion Methionine Methoxychlor Methoxyflurane Methylcyclohexane Methylphenyl sulphide N-Acetylmethionine Naphthalene -Naphthoflavone -Naphthoflavone N-Ethylmaleimide N, N-Dimethylaniline 2-Nonanone Norbenzphetamine Nortriptyline n-Octane n-Octane thiol Oleic acid Oxotremorine Parathion Pentobarbital Phenacetin Phenobarbital Pronethalol Propranolol Prostaglandins Pyrethrins Safrole Secobarbital SKF-525A SKF-8742A Taurochenodeoxycholic acid -Terpineol Testosterone Tetrahydrocannabinol Theophylline Toluene

INTRODUCTION

15

Type I substrates Tremorine Triton X-100 Tryptophan (low concentration) Tween 80 Alkyl and aromatic amides Aminoantipyrine p-Aminophenol d-Amphetamine (high concentration) Aniline p-Anisidine Benzylamine Butylmethyl sulphide p-Chloroamphetamine p-Chloroaniline Cyanide Dapsone Desdimethylimipramine DPEA Acetanilide Agroclavine Aminopyrine d-Amphetamine (high concentration) Barbital Benzyl alcohol Butanol Caffeine Cyclohexene oxide Diallyl barbituric acid 4, 4-Dihydroxybiphenyl Dipyridyl Ethanol Hexobarbital

Vinbarbital Vinyl chloride (R)-Warfarin Zoxazolamine Ethyl isocyanide Imidazole Metyrapone Nicotinamide Nicotine Nitric oxide Pentamethylene sulphide p-Phenetidine Pyridine Pyridine analogs Pyrrolidone SKF-26754A Tryptophan (high concentration) 5-p-Hydroxyphenytoin Isoamyl alcohol Isobutyl alcohol Lidocaine (high concentration) Methanol Phenacetin Phenobarbital 2-Propanol Rotenone Theophylline Tryptophan (intermediate concentration) (S)+(R)-Warfarin (S)-Warfarin

nm) and is itself indicative of an interconversion between two electronic states of the hemoprotein on addition of ligand, which have been confirmed (Kumaki et al., 1978) as corresponding to high- and lowspin ferriheme electronic states. From the differences in absorption and extinction coefficients between the two bands at 390 and 420 nm, the increase in percentage high-spin iron fraction can be determined, together with equilibrium constants for the various processes (Schenkman et al., 1981; Schenkman and Kupfer, 1982) shown in the following scheme:

16


where: S=substrate, LS and HS refer to low-spin and high-spin, respectively, for ferric P450 (Fe3+) with equilibrium constants K1–K4 . This diagram, in fact, forms one face of a ‘cube’ of microequilibria where consideration of the ferrous (Fe2+) equilibria are also included (see for example: Sligar and Murray, 1986). Descriptions of the practical and theoretical aspects of spin equilibria accompanying P450 substrate binding may be obtained from the following: Gibson and Skett, 1994; Tamburini, 1982; Schenkman and Kupfer, 1982; Schenkman et al., 1967a and b, 1972, 1981; Gibson and Tamburini, 1984. The thermodynamic parameters G, H and S which describe the free energy, enthalpy and entropy changes, respectively, for the P450 equilibria outlined above, can be obtained by considering the effect of temperature on the equilibrium constants and, in particular, from construction of van’t Hoff plots for different substrates (Schenkman et al., 1981). Aspects of the thermodynamics of P450 substrate binding will be discussed later in Chapter 3, and the reader is referred to Schenkman and Kupfer (1982) for further details. The fact that ferrous P450 appears to exist predominantly in the high-spin state facilitates a simplification of the thermodynamic ‘cube’ model to the planar scheme outlined above (Sligar, 1976) such that it is possible to derive an expression for the P450 redox potential in terms of spin equilibria as follows: where E0 is the redox potential, R is the gas constant, T is the absolute temperature, F is the Faraday constant, and Ka, b are the equilibrium constants for the ferric high-spin/low-spin and ferrous/ferric high-spin equilibria, respectively. According to the above equation, there should be a linear relationship between the P450 redox potential and spin equilibrium constant, Ka, expressed in the form: ln[(1+Ka)/Ka]. In fact, this appears to be the case for both the bacterial system, P450cam (Sligar and Gunsalus, 1976), and the rat liver microsomal P450LM2 (Sligar et al., 1979), implying that there is a coupling between spin state and redox equilibria in the P450 system, such that a change from low-spin to high-spin which accompanies type I substrate binding also gives rise to a lowering of the P450 redox potential. This would seem to be fundamental for P450 catalytic activity as it implies a relative ease of reduction of ferric P450 by its redox partner to form the ferrous state, which can then bind oxygen and activate it to insert an oxygen atom into the substrate. The fact that the magnitude of the type I spectral change, which occurs on substrate binding, can be related to the rate of metabolism of the substrate (Blanck et al., 1983) seems to support this theory. In contrast to the type I case, substrates which exhibit a type II spectral change appear to act primarily as inhibitors of the enzyme by ligating the heme iron, forming relatively stable, tightly-bound complexes. To some extent, it is possible to make generalizations about the likely nature of these complexes in terms of structural features on the substrates themselves. For example, an inspection of the chemical structures of type II ligands shows that they usually possess atoms with freely accessible non-bonding electrons, such as the nitrogen lone pair of aromatic and aliphatic amines (Table 1.10). The type II spectral change is characterized by a decrease in absorption at around 390–405 nm accompanied by an increase at 425–435 nm, with an isobestic point at about 419 nm (Schenkman et al., 1981). The absorptions corresponding to low- and high-spin iron(III) are, therefore, shifted somewhat in position towards longer wavelengths, in addition to showing a decrease in high-spin content and a concomitant increase in low-spin iron. If the ligands actually bind to the heme of P450, then such changes would be expected as the iron would be

INTRODUCTION

17

preferentially forced to adopt an in-plane conformation by the heme ligand, which favours the low-spin state. Evidence from experimental studies where type II substrates can displace carbon monoxide from the P450-CO adduct provides strong support for this hypothesis (Schenkman et al., 1967a and b, 1972; Omura and Sato, 1962, 1964a and b; Imai and Sato, 1967). The shift to longer wavelengths of the Soret absorption also suggests that there is a direct interaction between the ligand and the heme group, as such shifts tend to follow the pattern exhibited by the change in ligand field strength as summarized in Table 1.9. For example, the absorption maximum for CN– at 445 nm is almost as high as that exhibited by CO at 450 nm, and both are regarded as high field ligands. The third variety of spectral change, that of reverse type I, is essentially a ‘mirror-image’ of the type I case, as can be readily appreciated from an inspection of the UV difference spectra (Schenkman et al., 1981). Consequently, in reverse type I (originally termed modified type II) there is an increased absorption of 420 nm and a decrease at 390 nm. Although this type of spectral change resembles the type II situation, the absorption bands are not shifted and, therefore, it is unlikely that heme ligation by the substrate occurs. Presumably, in this instance, substrate binding favours a spin equilibrium shift in the direction of the low-spin form which implies that the sixth ligand (generally regarded as water) is not displaced, as in type I binding, but is in fact more firmly held. However, an alternative explanation could involve some degree of interaction between the ligand and heme iron, which would involve removal of the water ligand, but still favour the low-spin state. As the majority of reverse type I substrates/ligands (Table 1.10) contain oxygen and/or nitrogen there could be the possibility of hydrogen-bonded interactions between the water sixth ligand and the substrate, or direct heme interaction which is too weak to produce a shift in the absorption maxima. In fact, the reverse type I substrates may bind to a second site in the hydrophobic pocket which is different from that occupied by type I substrates, where the latter tend to displace the sixth ligand as they are, on the whole, more hydrophobic than reverse type II substrates. The position is further complicated because a chemical may be an inhibitor (i.e. type II ligand) of one P450 but a substrate for another, so that it could be both a type I and a type II substrate depending on the nature of the P450 involved. For example, desmethylimipramine is a type I substrate for non-induced P450, but is a type II substrate for phenobarbital-induced P450. A similar situation occurs with caffeine, which is a type I substrate for the PCN-induced form, but a reverse type I substrate for either Arochlor- or MC-induced P450. Furthermore, in some instances, there is a concentration dependency of the substrate type as in amphetamine, lidocaine and tryptophan, where an increase in concentration alters the type of spectral change from type I to type II or reverse type I, as shown in Table 1.10. Table 1.11 Electronic spectral characteristics of P450 and hemoglobin (References: Ruckpaul and Rein, 1984; Lewis, 1986; White and Coon, 1980) Absorption bands (nm) Iron electronic state Fe(III) low-spin 568 568 Fe(III) high-spin 644 645 Fe(II) high-spin —

569 535 535 646 547 540 — 544

535 418 418 540 394 394 542 413

Soret*

Near UV

Hemoprotein

417 360 357 391 — — 408 —

360 P450LM2 P450LM4 — P450LM4 P450SCC — P450LM2

P450cam

P450cam

P450cam

18


Absorption bands (nm) Iron electronic state

Soret*

Near UV

— 542 411 — P450LM4 Fe(II) low-spin — 550 447 363 — 552 451 370 P450LM2 CO — 550 448 — P450LM4 CO 569 540 419 344 Hemoglobin CO 580 552 418 355 P450cam O2 — 558 420 — P450LM O2 577 541 415 344 Hemoglobin O2 * Figure 1.1 shows the appearance of the Soret absorption for the P450C21 CO complex.

Hemoprotein P450cam CO

Another method of producing a spectral change is to increase the pH of the solution, as this will cause the water sixth ligand to be converted into the hydroxide ion (Schenkman et al., 1981). Thus, an increase in the electronegativity of the sixth ligand will also bring about a type I spectral change. However, in reduced Fe (II)CO P450 complexes, increasing the pH may be able to convert the cysteine ligand from thiol to thiolate, which will favour the high-spin form by decreasing the Fe-S bond length, thus assisting out-of-plane movement of the iron and also stabilizing the high spin state via SP- bonding interactions (Lewis, 1986). In fact, it is thought that the inactive form of P450, namely P420, which exhibits a reduced Fe(II)CO absorption maximum at 420 nm, contains cysteine in its protonated thiol state (Hill et al., 1970a). As such, this form exhibits a similar UV spectrum to that of the hemoglobin CO complex, which also absorbs at 420 nm, and there is, moreover, some evidence to suggest that oxygenated P450 has thiol as the fifth ligand, due to similarities in its UV spectrum with that of P420. Table 1.11 summarizes the main absorption bands in the UV/visible spectra of P450s, together with that of hemoglobin as a comparison, such that these spectral effects and differences can be appreciated. 1.4.2 Polarized optical spectroscopy Polarized optical spectroscopy involves the use of polarized light to record optical absorption spectra, and this encompasses the three related techniques of circular dichroism (CD), magnetic circular dichroism (MCD) and optical rotatory dispersion (ORD) spectroscopy. Essentially, circular dichroism relates to the difference in absorption between left- and right-circularly polarized light, which is termed the Cotton effect; magnetic circular dichroism involves the difference between the absorption of left- and right-circularly polarized light induced by an external magnetic field, which is known as the Faraday effect, and optical rotatory dispersion is determined by the rotation of the direction of linear polarization when plane polarized light interacts with matter (Lewis, 1986). Furthermore, an external magnetic field can induce ORD in optically inactive media, and when an ORD curve is recorded through an absorption band, an S-shaped curve results which resembles the first derivative of the absorption peak. Of these three techniques, MCD is probably the most important for the study of hemoproteins, and P450 in particular. However, linearly dichroic polarized optical spectroscopy has also been employed in investigations of hemoproteins, including P450cam, and the use of this technique has been reviewed by Makinen and Churg (1983). Probably the key work on linear dichroism polarized spectroscopy of P450 is

INTRODUCTION

19

that carried out by Hanson and co-workers on P450cam (Hanson et al., 1977) who were able to establish the orientations of electronic transitions in the polarized UV spectrum, and thus utilize molecular orbital calculations by the iterative extended Hückel (IEH) method to compile orbital energy level diagrams for oxy- and carboxy-P450cam. Whereas INDO calculations show that a negatively charged sulphur ligand is required in order to rationalize the UV spectrum of P450.CO (Jung, 1985). The use of polarized optical spectroscopy has been reviewed by Ruckpaul and Rein (1984), Lewis (1986) and in a more recent work by Hawkins and Dawson (1992) which contains further reported studies on P450s, mainly using MCD. The particular interest in MCD spectroscopy of hemoproteins is due to its sensitivity to iron ligation and the accompanying redox and spin state changes (Dawson and Cramer, 1978; Makinen and Churg, 1983). Some of the early CD and MCD work on P450s has been summarized by Coon and White (1980) who report CD data on solubilized liver microsomal P450 and P450cam, and tabulate MCD results for isolated liver microsomal P450s (namely, LM2 and LM4). These data show remarkable similarity with MCD spectra of model iron porphyrin complexes containing thiolate ligands and, furthermore, Dawson and Cramer (1978) have used MCD to disprove the theory that histidine was the fifth ligand in P450cam, in favour of cysteine as the thiolate species. Dawson and co-workers (1982) later showed that the sixth ligand in the resting state of P450cam is, in fact, oxygen-containing (probably water) using a combination of UV, MCD and ESR spectra (Dawson et al., 1982, 1983; Sono et al., 1982; Andersson and Dawson, 1984). A full description of the various stages by which the Dawson group elucidated the heme ligation characteristics of P450, including comparisons with model compounds, can be found in a recent account by Hawkins and Dawson (1992). Low-temperature MCD studies have been carried out by Sharonov and coworkers on P450cam (Greschner et al., 1993) and P450LM2 (Sharonov et al., 1987) which provide evidence for ligand-induced changes in active site structure, including the possibility of alterations in the nature of the proximal ligand accompanying the conversion to P420. Although histidine is suggested (Sharonov et al., 1987) this residue is only present close to the heme in P450cam, whereas arginine would be more likely in microsomal P450s. A basic residue is conserved at two positions upstream of the invariant cysteine in almost all P450s and may be involved in either electron transfer or ion-pairing to a heme propionate (Lewis, 1995a). However, there is another nitrogenous amino acid residue, namely tryptophan, which is highly conserved in many eukaryotic P450s and, in the P450BM3 structure, this residue can form a hydrogen-bonded interaction with the second heme propionate (Lewis, 1995a). Evidence from fluorescence spectroscopy indicates that this tryptophan is indeed relatively close to the heme, as quenching of tryptophan fluorescence is observed (Inouye and Coon, 1985) which also increases following conversion to P420 (Chiang and Coon, 1979). The resulting changes in the CD spectra of both P450LM2 and P450LM4 following treatment with detergent were reported to be indicative of an increase in -helical content of the protein (Chiang and Coon, 1979). Prior to the determination of P450 crystal structures, CD spectroscopy was used to establish the chiral orientation of the prosthetic heme group (Ortiz de Montellano et al., 1983) which was found to be the same as that of hemoglobin. More recently, MCD and other spectroscopic techniques have been used to compare the different heme-thiolate proteins, P450 and chloroperoxidase (Dawson and Sono, 1987), heme-containing oxygenases and peroxidases (Dawson, 1988), and P450 with secondary amine oxidase (Hawkins and Dawson, 1992). Although there are certain similarities between the MCD spectra of P450 and peroxidases, Dawson and coworkers have also highlighted important differences which are suggestive of significant variations in both the hemoproteins, active sites and their mechanisms of oxygen activation (Dawson, 1988; Dawson and Sono, 1987; Dawson et al., 1983). Moreover, a recent MCD study has shown that there are also differences between peroxidases and catalase in terms of their respective oxygenated intermediates (Rodriguez-Maranon et al., 1995). Consequently, it would appear that it is not only the nature of the

20


proximal heme ligand which determines the specific oxygenating activity of hemoproteins (Adachi et al., 1993) but also the particular characteristics of the heme environment that are largely controlled by the apoprotein. 1.4.3 Vibrational spectroscopy Both infra-red (IR) and resonance Raman (RR) spectroscopy have been employed to study cytochromes P450 and related hemoproteins, with RR being the more commonly used, as far as P450 is concerned, because potentially more useful information can be obtained. In particular, vibrational and vibronic transitions associated with the porphyrin ring environment can be investigated by RR spectroscopy (Spiro, 1983; Ruckpaul and Rein, 1984; Lewis, 1986; Hildebrandt, 1992). The region in the immediate vicinity of the heme moiety is well suited to exploration by this technique, especially as the porphyrin ring vibrational modes (Table 1.12) have been extensively characterized, such that their sensitivity to changes in heme ligation, redox- and spin-state can be readily monitored (Hildebrandt, 1992). In contrast, IR spectroscopy is useful for investigating the effects of alteration in heme environment on the stretching frequencies of the distal heme ligand, such as carbon monoxide (CO), dioxygen (O2), etc., in cytochrome P450 and other hemoproteins (Ruckpaul and Rein, 1984; Lewis, 1986). For example, the C−O stretch of the Fe(II)CO complex in P450s varies from 1940 cm−1 in P450cam to around 1950 cm–1 in microsomal P450s (Table 1.13), and this increases to about 1966 cm–1 in the inactive form P420 (Böhm et al., 1979). Table 1.12 Vibrational modes in metal porphyrins (Reference: Lewis, 1986) In-plane vibrations

No. of modes

Out-of-plane vibrations

No. of modes

A1g (Raman) A2g B1g (Raman) B2g (Raman) Eu (Infra-red)

9 8 9 9 18

A1u A2u (Infra-red) B1u B2u (Raman) Eg (Raman)

3 6 5 4 8

Table 1.13 C−O Stretching frequencies of P450-CO complexes (References: Ruckpaul and Rein, 1984; Jung et al., 1992) P450 type

CYP

Frequency (cm−1)

P450cam P450SCC P4501m2 P450rlm

101 11A1 2B4 2B1

1940.6 1953 1949 1948

It is thought that the frequency of the C−O stretching vibration in hemoproteins reflects the degree of distortion from linearity of the Fe−CO grouping (Collman et al., 1976) and crystallographic evidence from the P450camCO complex does indeed show that there is significant non-linearity in the Fe−CO moiety (Raag and Poulos, 1989a) which is at an angle of 166° in the solid state structure. Oxygenated heme complexes also absorb in the infra-red with an O−O stretching frequency indicative of the electronic state of the dioxygen species, being dependent on the O−O bond order and bond energy (Table 1.14 and Figure 1.5). For

INTRODUCTION

Figure 1.5 Linear relationship between oxygen-oxygen stretching frequencies and bond energies for O2, species.

21

and

example, super oxide shows an absorption between 1100 cm–1 and 1150 cm–1, whereas free dioxygen absorbs at about 1555 cm–1, and peroxide has an IR stretching frequency around 850 cm–1 (Lewis, 1986). Resonance Raman spectroscopy may also be used to determine dioxygen stretching bands in hemoproteins and has shown, for example, that under catalytic conditions an O−O stretch occurs at 1141 cm−1 for the oxy form of P450cam (Egawa et al., 1991). In fact, as the O−O bond energy is directly proportional (Figure 1.5) to the vibrational stretching frequency, this suggests that activation of dioxygen by P450 decreases the O−O bond energy and increases the bond length, such that bond cleavage will occur, thus leading to monoxygenation of the substrate. Table 1.14 Dioxygen O−O bond data and vibrational stretching frequencies (Reference: Lewis, 1986) Oxygen species

Bond length (Å)

Bond energy (kJ mole–1)

O−O stretching vibration (cm–1)

O2

1.21 1.33 1.49

497 276 146

1555 1107 850

It is thought that the C−O stretching frequency of hemoprotein-CO complexes may be indicative of effects operating within the heme environment, which could be both electronic and polar, as the C−O stretch is affected by the nature of the proximal heme ligand (Lewis, 1986; Jung et al., 1992) and probably by the hydrogen-bonding possibilities at the distal heme face (Ormos et al., 1988; Jung et al., 1992). In P450cam– CO complexes, the C−O stretching mode shows a dependency on the nature of the bound substrate and, from such studies, it is possible to calculate the angle between the CO bond and the heme normal (Jung et al., 1992). The value of 16º for this angle, obtained from the IR spectrum of the camphor-bound P450camCO complex, is in close agreement with the evidence for X-ray crystallography (Raag and Poulos, 1989a),

22


which appears to be 14° (Table 1.15). Although the oxygen atom of the CO ligand in the P450camCO complex is relatively close to the hydroxyl group of threonine–252 (T252) in the I helix, the interatomic distance of 4.39 Å between the two oxygens (Lewis and Lake, 1995) is too long for any hydrogen bond formation. However, the possibility of free rotation of the T252 side chain can shorten this distance by almost 1Å and, in the case of the dioxygen complex, the longer O−O bond makes ligand-protein hydrogen-bonding fairly likely. This possible scenario could explain the difference in O−O stretching frequencies (Table 1.16) between P450 and other hemoproteins, but an alteration in the electronic distribution of the dioxygen molecule caused by the proximal heme ligand is probably an important factor. A consideration of the relevant vibrational spectra of hemoprotein-CO and -O2 complexes (Nagai et al., 1991; Egawa et al., 1991; Jung et al., 1992), together with Table 1.15 Interatomic distances and vibrational stretching frequencies in hemoglobin, myoglobin and P450cam (References: Lewis, 1986; Egawa et al., 1991; Bangcharoenpaurpong et al., 1986; Dawson et al., 1986; Omura et al., 1993; Jung et al., 1992; Brookhaven Protein Databank) Fe−O distance (Å)

PDB code

Hemoglobin Myoglobin P450cam

1.865, 1.656 1HHO 1.827 1MBO 1.78 (EXAFS) 1.79 (estimated) Fe−C distance (Å) Hemoglobin 2.011, 1.871 1COH Myoglobin 1.924 1MBC P450cam 2.044 3CPP O−O distance (Å) Fe−O−O angle (°) Hemoglobin 1.241, 1.218 1HHO 158.73, 152.69 Myogobin 1.217 1MBO 115.51 P450cam 1.22–1.26 (estimated) 124–136 (estimated) C−O distance (Å) Fe−C−O angle (º) Hemoglobin 1.295, 1.211 1COH 155.37, 152.95 Myoglobin 1.169 1MBC 141.43 P450cam 1.118 3CPP 166.02 O−O stretch (cm–1) C−O stretch (cm–1) Hemoglobin 1107, 1155; 1159 1951 Myoglobin 1103, 1150 1944, 1952 P450cam 1139–1141 1940.6 Note: The estimated values were interpolated from correlations between vibrational stretching frequencies and bond data. Table 1.16 O−O Stretching frequencies in hemoprotein O2 complexes (References: Lewis, 1986; Egawa, 1991; Bangcharoenpaurpong et al., 1986) Hemoprotein

–1 o−o(cm )

Hemoglobin Myoglobin P450cam

1107, 1155 1103, 1150 1140, 1141

INTRODUCTION

Hemoprotein P450cam

23

–1 o−o(cm )

1139 (at room temperature)

Figure 1.6 Correlation between iron-oxygen and oxygen-oxygen stretching frequencies for hemoglobin, P450 and some iron-porphyrin complexes.

their crystallographic data tends to provide a self-consistent picture regarding the correlation between structural studies and stereo-electronic effects in the vicinity of the heme moiety (Hildebrandt, 1992), which is further supported by the work using model iron porphyrins and cobalt porphyrin model complexes (Choi and Spiro, 1983; Spiro, 1983; Collman et al., 1976; Bajdor et al., 1983; Shelnutt, 1983). For example, in addition to the linkage between C−O stretching frequencies and Fe-C−O bond angle in hemoprotein CO complexes (Jung et al., 1992; Nagai et al., 1991) there is also a correlation between Fe−CO and C−O stretching vibrations (Nagai et al., 1991) depending on the type of hemoprotein or model iron porphyrin. It would appear that this may also extend to the respective dioxygen complexes, as can be shown in Figure 1.6, which utilizes the data presented in Table 1.17, obtained from a compilation of vibrational spectra for dioxygen and Fe−O stretching frequencies. Furthermore, it is possible that the variations in these frequencies reflect the bond lengths and bond energies of both O−O and Fe−O, thus providing a means of estimating structural data in heme-dioxygen complexes. Table 1.18 summarizes asymmetric Fe−O stretching frequencies in a number of hemoproteins and model compounds (Egawa et al., 1991). It is thought that an absorption band in this region may be indicative of an Fe(IV)O linkage, although such a vibration frequency was not detected in the RR spectrum of oxygenated P450cam (Egawa et al., 1991) and this Table 1.17 Comparison between dioxygen and iron-oxygen stretching vibrational frequencies for iron porphyrin complexes and hemoproteins (cm−1) (References: Omura et al., 1993; Lewis, 1986; Bangcharoenpaurpong et al., 1986) Iron porphyrin/hemoprotein

O2 stretch

Fe−O stretch (symm.)

Fe tetramesitylporphyrin

1171

522

24


Iron porphyrin/hemoprotein

O2 stretch

Fe tetraphenylporphyrin 1195 Fe meso-tetra(pivalamidophenyl)porphyrin PhS1147 P450cam 1140 Hemoglobin 1107 Note: The correlation between O2 stretch and Fe−O stretch is 0.999 (Figure 1.6)

Fe−O stretch (symm.) 509 536 541 562

Table 1.18 Iron-oxygen stretching vibration frequencies (cm–1) (Reference: Egawa, 1991) Hemoprotein

Fe−O stretch (asymm.)

Ligands

Myoglobin Cytochrome c oxidase Horseradish peroxidase C Horseradish peroxidase A Myeloperoxidase Cytochrome c peroxidase Lactoperoxidase Model porphyrin complexes Peroxide (free)

797 788–790 787 (pH 11.2), 774 (pH 7) 779 782 753 745 780–800 770–850

Histidine Histidine Histidine Histidine Histidine Histidine Histidine Various

was ascribed as being due to the likely transient nature of any type of iron oxene intermediate in the P450 catalytic cycle. Resonance Raman (RR) spectroscopy is an important technique in the study of hemoproteins as the excitation wavelength can be the same as that of the strong electronic absorptions in the UV, such as the Soret absorption band (Lewis, 1986), which enhances the intensity of the vibrational bands associated with the porphyrin ring without affecting those due to the protein. Different regions of the RR spectrum correspond to porphyrin core-size markers, oxidation- and spin-state marker bands, and iron-ligand vibrational modes (Hildebrandt, 1992), all of which can provide useful information for structural investigations of the active sites in P450 complexes, together with those of other hemoproteins. For example, the Ca−N stretching band around 1370 cm−1 is an indicator of the oxidation state of the heme iron, being about 1375 cm−1 for Fe(III) and normally close to 1360 cm−1 in the case of Fe(II), with very little difference between high- and low-spin (Spiro, 1983). For P450 complexes, the oxidation state marker is about 1371 cm−1 for Fe(III) but significantly lower (Ozaki et al., 1978) at around 1341 cm−1 for high-spin Fe(II), which is thought to be due to the effect of thiolate ligation (Hildebrandt, 1992). In the case of the low-spin Fe(II)CO complexes, this oxidation state marker band rises in frequency to between 1365 cm −1 and 1369 cm−1 due to the electron-accepting ability of the axial CO ligand (Ozaki et al., 1978). A number of absorptions in the region 1450 cm−1 to 1650 cm−1, totalling at least eight porphyrin skeletal mode vibrations (Table 1.19), are sensitive to the core size radius of the porphyrin ring and, consequently, can be used as core-size marker bands for the analysis of spin-state and coordination number (Spiro, 1983; Hildebrandt, 1992). These absorption bands all vary in a characteristic manner, which reflects the dimensions of the porphyrin ring core (1.98–2.06 Å), that enable determination of the symmetry (C4v or D4h) of the heme environment, which can vary between 5-coordinate and 6-coordinate, and distinction between Fe(II) and Fe(III) in both high- and low-spin states. Although there are several other vibrational bands of lower frequency in the so-called ‘fingerprint’ region which have been assigned to a number of heme

INTRODUCTION

25

vibrational modes, the iron-ligand absorption bands between 150 cm−1 and 500 cm−1 are important determinants of the heme coordination. In particular, the iron-sulphur stretching vibration at 351 cm−1 indicated the presence of a heme-thiolate moiety in P450cam (Champion et al., 1982), conclu Table 1.19 Resonance Raman marker bands in P450 and other hemoproteins (References: Spiro, 1983; Hildebrandt, 1992; Ruckpaul and Rein, 1984) Fe(III)

Fe(II)

Frequency range (cm −1)

Frequency range (cm– 1)

Comments

B1g

1623–1637

1600–1612

Oxidation and spinstate marker

Eu A2g A1g B1g

1580–1601 1583 1565–1584 1549–1564

1584–1586 – 1556–1564 1532–1534

Spin-state marker

Eu A1g

1548–1550 1485–1502

1521 1462–1466

B2g

1464–1465 A1g

1445 1370–1373

Mode Symmetry Core-size marker bands 37 19 2 11

v38 3

v28 Oxidation state marker

10

4

Spin-state marker Oxidation-state marker Oxidation- and spinstate marker 1341–1344

Increases to 1365 or more in Fe(II)LSCO

sively showing that the heme iron in P450 possessed a cysteinate ligand, prior to the determination of its crystal structure. In the P450BM3 holoenzyme, the Fe-S absorption band has been observed at 346 cm−1 (Munro et al., 1994) which, although close to the value of 350 cm−1 (Miles et al., 1992) for this ligandbinding mode in the P450BM3 hemoprotein domain, may also indicate the effect of reductase domain interactions. Interestingly, the presence of a thiol ligand in P450 has not been observed in RR spectra, which tends to cast some doubt on the likelihood of any mechanistic relevance of the thiol-thiolate equilibrium. Furthermore, the lack of evidence for Fe=O, during the P450 oxygenation reaction, in the RR spectrum of P450cam under catalytic conditions (Egawa et al., 1991), but clear observation of an O−O stretch at 1141 cm −1, provides experimental support for an iron peroxy/superoxy intermediate being the active oxygenating species. Proximal ligand site-directed mutagenesis studies on hemoproteins have shown that the presence of a thiolate ligand is an enhancing factor in O−O bond cleavage (Adachi et al., 1993). Consequently, it is likely that the proximal cysteinate ligand in P450 remains in the thiolate state for catalytic activity as its electron-donating effect, channelled through the heme moiety, is essential for activation of the oxygen ligand and dioxygen bond scission preceding oxygenation of the substrate. Resonance Raman spectroscopy will undoubtedly remain an extremely useful technique for probing the heme environment of P450s, and other hemoproteins, due to the exquisite sensitivity of several RR vibrational modes to changes in the vicinity of the active site.

26


1.4.4 Magnetic resonance spectroscopy Both nuclear magnetic resonance (NMR) spectroscopy and electron spin resonance (ESR) spectroscopy have been employed in the study of the P450 system (Lewis, 1986; Weiner, 1986) and these two techniques are, to some extent, complementary in terms of the possible structural information which can be obtained. Due to the usefulness of ESR in the investigation of Fe(III) spin-state equilibria for P450s and in spin-labelling experiments, this procedure has been used more extensively than NMR in the P450 field. However, one of the advantages of NMR over ESR in the characterization of hemoproteins lies in the fact that the latter is limited to Fe(III) as, being diamagnetic, Fe(II) is ESR-silent. In contrast, NMR can be employed in structural studies of both redox states of P450s, and also in other hemoproteins. 1.4.4.1 NMR spectroscopy Proton (1H) NMR spectroscopy has been used to investigate the effects of substrate and inhibitor binding on the relaxation of solvent protons in the vicinity of the P450 active site (Griffin and Peterson, 1975; Philson et al., 1979; Weiner, 1986). The use of a simplified form of the Solomon-Bloembergen equation enables an estimation of the distances between relaxing protons and the heme iron, due to the influence of the metal centre on proton signals and relaxation times (Weiner, 1986). In fact, 1H NMR has provided considerable evidence for the existence of a solvent water molecule as the distal sixth ligand in various forms of P450 (Lewis, 1986; Weiner, 1986; Hawkins and Dawson, 1992) and this has been confirmed by X-ray crystallographic studies on the substrate-free prokaryotic P450s (Poulos et al., 1986; Ravichandran et al., 1993; Hasemann et al., 1994). The values reported for the distances between protons from the bound water molecule in P450cam as determined by NMR, namely, 2.0–2.9 Å, are in generally close agreement with those observed in the substrate-free crystal structure, where the Fe−O distance is 2.28 Å (Table 1.20). Furthermore, the crystallographic data for the other two bacterial P450s (i.e. P450BM3 and P450terp) indicate similar values, being 2.188 Å and 2.088 Å, respectively (Table 1.20). It is also found that the binding of substrates or inhibitors to P450s will bring about a change in the proton NMR signals indicative of a movement of solvent water away from the heme iron to a distance of up to about 7 Å (Griffin and Peterson, 1975). In addition to solvent water proton relaxation, substrate and inhibitor interactions within the active sites of various P450s can be studied via proton NMR by investigating the effect of the heme iron on relaxation times (Weiner, 1986). It would appear that the distances between aromatic protons of the nitrogenous ligands and iron are consistent with nitrogen coordination to the heme, as shown in Table 1.21. In contrast, the proton signals of P450 substrates tend to indicate greater distances from the heme iron, as would be expected on mechanistic grounds. Recently, an NMR study of lauric acid binding to P450BM3 shows that the -methyl group protons are 5.6 from the iron, whereas the -methylene protons adjacent to the carboxylate moiety are reported to be at a 16.4 distance from the heme (Gibson et al., 1995). In fact, it has been shown that molecular modelling of laurate within the crystal structure of P450BM3 is entirely consistent with the NMR data (Lake and Lewis, 1996). The protons of the protoporphyrin IX system are, moreover, sensitive to the iron redox- and spin-states, thus enabling such changes, which occur during the P450 cycle, to be studied by proton NMR. In fact, this technique can be used to show that Fe(II) in reduced P450 is in the high-spin state prior to dioxygen binding, which could not have been determined by ESR spectroscopy (Hawkins and Dawson, 1992). The presence of thiolate as the fifth proximal heme ligand has been indicated by the shift in methyl group proton resonances on ligation, and from 13C-NMR spectra of CO-bound P450 (Berzinis and Traylor, 1979). P450cam has also been

INTRODUCTION

27

Table 1.20 Heme geometries in P450 crystal structures (References: Poulos et al., 1987; Raag and Poulos, 1989; Ravichandran et al., 1993; Hasemann et al., 1994)

a

Data for substrate-free enzyme.

studied using 15N-NMR to determine the effects of substrate- and putidaredoxin-binding on the N-15 resonances of the cyanide complex (Shiro et al., 1989). The characteristic 15N chemical shift of isotopicallylabelled CN– in these complexes varies in accordance with the electronic environment of the heme, indicating that substrate binding increases the electron density on the iron atom, whereas putidaredoxin binding brings about a decrease in electron spin density, which are thought to reflect the influence of the thiolate ligand on these two stages in the P450cam catalytic cycle. Electron-nuclear double resonance (ENDOR) spectroscopy has been employed to determine the electron-nuclear coupling effects in P450cam

28


(reviewed by Lewis, 1986) but gave conflicting results in two independent studies (Peisach et al., 1979; LoBrutto et al., 1980). However, the supposition that histidine may be the fifth ligand has been discounted, as it is now regarded that the ENDOR coupling observed was due to the porphyrin ring nitrogens (Hawkins and Dawson, 1992). Consequently, ENDOR spectroscopic measurements support the generally-accepted view that cysteine in its anionic form is the fifth proximal ligand in P450. The significant advances in NMR spectroscopic techniques (Roberts, 1993) have facilitated the structural characterization of polypeptides and even small proteins, such as putidaredoxin (Pochapsky and Ye, 1991) and cytochrome b5 (Guiles et al., 1990). In theory, the protein structural conformation in solution could be determined by a combination of various NMR procedures, which enable assignment of Table 1.21 Proton-iron distance ( ) in substrate/inhibitor complexes of various microsomal P450s (Reference: Weiner, 1986) Compound

Methyl protons

Phenyl protons

P450

Xylidine Acetanilide Imidazole 4–Methoxypyridine

2.44–5.35 8.00 (±0.019) H(2):3.9 5.3 (±0.2)

2.3–5.0 7.2 (±0.019) H(4, 5):4.1, 5.9 CH:5.0±(0.5) CH:40 per cent identity between subfamilies within a given family. Although this was originally a somewhat arbitrary distinction, it has been proved to work quite well, with a number of exceptions. For example, proteins of the 2D, 2J Table 2.8 The CYP3 family (References: Nelson et al., 1993; Gonzalez et al., 1985) Species

CYP3A

Rat Human Rabbit Monkey Hamster Mouse Dog Sheep

3A1, 3A2, 3A9 3A3, 3A4, 3A5, 3A7 3A6 3A8 3A10 3a11, 3a13 3A12 3A

Table 2.9 The CYP4 family (References: Nelson et al., 1993; Hardwick et al., 1987; Chen and Hardwick, 1993; Liping and Hardwick, 1993; Kikuta et al., 1993) Species

4A

4B

Human Rabbit Rat Guinea pig Mouse Cockroach Fruit fly

4A9, 4A11 4A4, 4A5, 4A6, 4A7 4A1, 4A2, 4A3, 4A8 4A13 4a10, 4a12

4B1 4B1 4B1

4C

4D

4E

4F 4F2, 4F3 4F1

4C1 4D1

4E1

Table 2.10 Families CYP5, CYP6, CYP7 and CYP10 (References: Nelson et al., 1993; Cohen and Feyereisen, 1995) Species

CYP5

Human House fly Fruit fly Butterfly Rat Rabbit Cow Pond snail

5

CYP6

CYP7

CYP10

7 6A1, 6A3–6, 6C 6A2 6B1 7 7 7 10

EVALUATION OF THE P450 SUPERFAMILY

Table 2.11 The CYP11 family (References: Nelson et al., 1993; Morohashi et al., 1987a and b) Species

CYP11A

CYP11B

Human Cow Pig Rat Chicken Trout Mouse

11A1 11A1 11A1 11A1 11A1 11A1

11B1, 11B2 11B1, 11B2, 11B4 11B1, 11B2, 11B3

11b1, 11b2

Table 2.12 Families CYP17, CYP19, CYP21, CYP24 and CYP27 (References: Nelson et al., 1993; Namiki et al., 1988; Chaplin et al., 1986; Corbin et al., 1988; Means et al., 1989; Chen and Zhou, 1992) Species

CYP17

CYP19

CYP21

Human Cow Pig Chicken Rat Trout Guinea pig Mouse Goldfish Sheep Rabbit

17 17 17 17 17 17 17 17

19

21A2 21A1 21A1

CYP24

27

19 19 19 19 19

CYP27

24

27

21a1 21A1 27

Table 2.13 Fungal and plant CYP families (51 to 73) (References: Nelson et al., 1993; Seghezzi et al., 1992; Kalb et al., 1987; Bozak et al., 1990) Fungal forms

CYP51

Saccharomyces cerevisiae Candida tropicalis

51 51

Candida albicans Candida maltosa

51

Aspergillus niger Neurospora crassa Fusarium oxysporum Nectria haematococca Plant forms

CYP52

CYP53 CYP54 CYP55 CYP56 CYP57 56

52A1, 52A2, 52A6, 52A7, 52A8, 52B1, 52C1 52A3, 52A4, 52A5, 52A9, 52A10, 52A11, 52C2, 52D1 53 54 55 57

CYP71 avocado

CYP72 periwinkle

CYP73 artichoke

67

68


Table 2.14 Bacterial CYP families (CYP101 to CYP112) (Reference: Nelson et al., 1993) Species

CYP family/subfamily

Pseudomonas putida Bacillus megaterium Agrobacterium tumefaciens Streptomyces griseolus Streptomyces spp. Saccharopolyspora erythrea Pseudomonas spp. Bacillus subtillis Anabaena spp. Bradyrhizobium japonicum

101 (cam) 102 (bm-3) 103 105A1, 105B1, 105C1 107A1, 107B1 108 (terp) 109 110 112

106 (bm-1) 104 105D1

111 (lin)

and 2K subfamilies possess generally lower than 40 per cent sequence identity with other CYP2 subfamilies; this is also true for 4C, 4D and 4E subfamilies compared with other CYP4 families, although these all refer to insect forms of the CYP4 family. Furthermore, CYP11A and CYP11B are 34–39 per cent identical but have, nevertheless, been placed in the same family. In fact, these mitochrondrial P450s are sequentially similar to CYP27, which is another P450 present in mitochondria. However, it should be noted that mammalian P450 sequences in the same subfamily are always>55 per cent identical (Nelson et al., 1993). Table 2.15 (see pages 76–77) summarizes the generally accepted timescale for evolution of the P450 superfamily which shows the divergence times calculated from sequence comparisons. This form of phylogenetic analysis has been utilized for the construction of diagrams (Figure 2.5) to illustrate the possible elaboration of the P450 phylogenetic tree. As new P450 sequences are determined, it will be possible to improve the accuracy of such analyses. Table 2.15 Evolution of P450 superfamily and divergence times (References: Nelson and Strobel, 1987; Omura et al., 1993) Divergence time Speciation (Mya) 1400 400

Prokaryoteeukaryote Mammal-fish

300

Mammal-bird

CYP

Evolutionary distance, d

Human-bovinerodent (Human-bovine) (Human-bovine) (Human-bovine)

Mya

Rate**

UEP PAM†

1400

0.9×10−9

5.58

43.16 48.71

271 256

30.52 51.93

240 174

0.8×10−9 0.95×10−9 1.18×10−9 0.64×10−9 1.49×10−9

6.27 63.83 5.25 76.21 4.23 7.87 38.11 3.35 89.63

2.51 1A1 1A2 0.71 19A 17A

85

Sequence % change

21 11A 17A

0.24 0.34 0.36

1.41×10−9 3.54 2.0×10−9 2.50 2.12×10−9 2.36

EVALUATION OF THE P450 SUPERFAMILY

Divergence time Speciation (Mya)

80

75

60

(Mouse-human/ bovine) Mammals

Human-rabbit-rodent (Rabbit-rat) (Rabbit-rat) (Rabbit-human-rodent) (Rabbit-rat) (Human-rabbit/rat) (Human-rodent) (Human-rabbit) (Human-rat) Bovine-porcine

CYP

Evolutionary distance, d

21

0.34

4B 2A 7A 2E 1A1. 2D 2B 19A 21A 11A 4A 1A2 27A 3A 17A 11B

13.33 15.11 17.89 20.22 21.92 22.74 22.39 23.46 24.94 25.17 26.10 26.21 26.83 27.62 32.07 32.74

0.22 0.24

2B 1A1 1A2 2E 2E 1A1 1A2 3A 21A 11A

Sequence % change

69 73 70 69 68 64 63 69 61 67 63 68 64 64 62 60

0.27 0.29 0.31 0.22 0.25 0.24 0.25 0.33 12.40 15.96

Mya

50 52

69

Rate**

UEP PAM†

2.0×10−9

2.54

0.97×10−9 1.04×10−9 1.29×10−9 1.47×10−9 1.60×10−9 1.78×10−9 1.77×10−9 1.71×10−9 2.05×10−9 1.89×10−9 2.06×10−9 1.92×10−9 2.11×10−9 2.17×10−9 2.58×10−9 2.71×10−9

5.15 4.80 3.89 3.41 3.12 2.80 2.83 2.93 2.44 2.65 2.42 2.60 2.37 2.31 1.94 1.84

1.80×10−9 1.93×10−9 2.07×10−9 1.47×10−9 1.67×10−9 1.60×10−9 1.67×10−9 2.20×10−9 1.25×10−9 1.53×10−9

2.78 2.59 2.42 3.41 3.00 3.13 3.00 2.27 4.01 3.28

15.52 16.67 20.89 23.45 25.62 28.53 28.28 27.28 32.85 30.22 33.02 30.72 33.81 34.64 41.24 43.43

14.96 18.32

Hamster-rat/mouse 1A2 11.80 20–17 Rat-mouse 1A1 0.072 1.80×10−9 1A1 0.073 1.83×10−9 2A 4.56 17 1.36×10−9 1A1 6.63 16 2.10×10−9 1A2 6.87 17 2.05×10−9 Key: d=2 where =rate of change (Tajima and Nei, 1984). ** Changes per site per annum. * Unit evolutionary period (time in millions of years for a 1 per cent change in sequence).

13.26 2.78 2.74 3.67 2.38 2.43

4.63 7.13 7.00

70


† Accepted point mutations per 100 residues.

Figure 2.5 A schematic and abbreviated representation of the evolution of the CYP superfamily with an approximate time scale (References: Nelson and Strobel 1987; Gotoh and Fujii-Kuriyama, 1989; Gonzalez and Nebert, 1990; Gonzalez and Gelboin, 1991; Nebert et al., 1991; Degtyarenko and Archakov, 1993; Omura et al., 1993.

3 The P450 Catalytic Cycle and Oxygenation Mechanism

3.1. Introduction For an enzyme system which has been so extensively studied as P450, it is perhaps surprising that there are still many questions about its catalytic cycle and oxygen activation mechanism remaining to be answered. Indeed, even the currently-accepted reaction stages are subject to speculation and controversy: some of which can be focused on the oxene intermediate. This supposedly active oxygenating species remains to be observed in a P450 system and yet would be expected to contain a strong covalent bond, i.e. Fe=O, which should entail stability and an energetically unfavourable cleavage with respect to, say, a single Fe-O linkage that has been postulated to be present at an earlier stage of the catalytic cycle. A second difficulty with the currently in vogue oxenoid mechanism arises from the fact that the iron-oxene intermediate can be expected to be electrophilic in nature whereas, in fact, some P450-mediated reactions would appear to require a nucleophilic oxygenating species. However, there are some advantages to the oxene postulate, as it readily explains single oxygen insertion, particularly for the formation of epoxides and N-oxides, which are otherwise difficult to explain via a peroxide or superoxide intermediate. In contrast, there is considerable evidence for both superoxide and peroxide in P450-mediated oxygenations, including the observation of an O−O stretching vibration by Raman spectroscopy under catalytic conditions (Egawa et al., 1991). On balance, it seems possible that, depending on the reaction conditions, substrate, and type of P450 involved, either mechanism (i.e. oxenoid or peroxide) could occur. It is, therefore, reasonable to discuss both of these mechanisms of oxygen activation and insertion into substrates such that the reader can make up his or her own mind regarding whether either, or both, are possible. 3.2 The P450 catalytic cycle The overall reaction for the majority of cytochrome P450-mediated mono-oxygenations can be represented as follows: where RH represents a substrate which is hydroxylated during the course of the reaction. The above stoichiometry shows that the mono-oxygenation is brought about by a dioxygen molecule, which is cleaved during the catalytic process, to form water and a hydroxylated metabolite, ROH, apparently produced by a

72


single oxygen atom insertion into the substrate. The reaction requires two reducing equivalents, i.e. two electrons and two protons; where the sources of the former are fairly well characterized, but vary according to the location of the P450 concerned. In mitochondria and many bacteria, the electrons are transferred from either NADH or NADPH to an iron-sulphur ferredoxin via an FAD-dependent reductase. However, in microsomal endoplasmic reticular systems, NADPH transfers electrons via an FAD- and FMN-containing flavoprotein, cytochrome P450 oxidoreductase, without the mediation of an Fe2S2 redoxin; although cytochrome b5 may also be the source of the second electron (Backes, 1993). The redox pathways are summarized in Table 3.1, although it should be recognized that there are some variations and exceptions to these generalizations; for example, some bacteria utilize an FAD/FMN-reductase similar to that of microsomal eukaryotic P450s, and can also use NADPH rather than NADH as the source of reducing equivalents. The actual catalytic cycle will vary somewhat, depending on the source of electrons, and the type of substrate undergoing metabolism, but a general simplified representation is shown in Figure 3.1, which depicts a number of intermediate stages. As one proceeds clockwise around the cycle, the intermediates proposed and the detailed mechanisms which operate, become progressively less well understood. The reasons for this mainly lie in the time scale under which some of the conversions take place and the nature of the techniques employed to monitor them. There is, however, much interest and speculation regarding certain features and aspects of the P450 catalytic cycle, including the nature of certain intermediates, particularly the active oxygenating species which carries out the attack on substrate molecules (White and Coon, 1980; Gunsalus and Sligar, 1978; Coon and White, 1980; White, 1991; Hawkins and Dawson, 1992; Gray, 1992; Thomson and Yumike, 1989; Kappus, 1993; Rein and Jung, 1993; Ortiz de Montellano, 1986b, 1989; Akhtar et al., 1994; Lee-Robichaud et al., 1995; Vaz et al., 1991; Coon et al., 1992; Coon and Table 3.1 Electron transport pathways in various P450 systems (Reference: Lewis and Moereels, 1992) Bacterial system:

NADH Putidaredoxin reductase Putidaredoxin P450cam Mitochondrial system: NADPH Adrenodoxin reductase Adrenodoxin P450scc Microsomal system: NADPH Cytochrome P450 oxidoreductase P450 or NADH Cytochrome b5 Cytochrome b5 P450 Notes: Bacteria other than Pseudomonas putia (P450cam) can show variations of the schemes shown above, and the Bacillus megaterium (P450BM3) system resembles the first pathway of the microsomal scheme. In the microsomal system, it is thought that the second pathway is primarily responsible for the delivery of the second electron.

Vaz, 1987; Groves and Watanabe, 1988; White et al., 1984; Ortiz de Montellano and Stearns, 1989). The kinetics and thermodynamics of various stages in the catalytic cycle have been studied in both bacterial and microsomal systems, and show broad similarities suggestive of a common mechanism (Archakov and Bachmanova, 1990; Ruckpaul and Rein, 1984). The camphor mono-oxygenase system from Pseudomonas putida, i.e. P450cam (CYP101), has been extensively studied as it is more readily accessible than microsomal P450s, which are membrane-bound and, consequently, are more difficult to investigate in isolation. However, Table 3.2, which provides a comparison between various kinetic parameters for the two systems, shows that the rate constants for individual stages in the catalytic cycle are roughly equivalent for the liver microsomal system and bacterial P450, camphor mono-oxygenase. The differences which occur

THE P450 CATALYTIC CYCLE

73

The various stages (1–5) whereby a substrate(s) becomes hydroxylated by molecular oxygen through the mediation of cytochrome P450 is shown together with the redox state of the heme iron. Figure 3.1 Reaction cycle and enzymatic intermediates in P450-catalyzed oxygenations (Reference: Lewis, 1992b).

may be partially explained in terms of the variation in redox partners between the two systems, and also due to the fact that microsomal P450s are membrane-bound. There is, furthermore, evidence to suggest that the active site regions are somewhat different (Chapter 6), and this could explain a number of features where there is variation between the two P450 sources. 3.2.1 Stage 1: Substrate binding It is known that in its resting state the enzyme is mainly present in its low-spin ferric form with water occupying the heme pocket, probably with one water molecule ligating the heme iron at the distal face. Substrate binding is rapid, of high affinity, Table 3.2 A comparison between kinetic parameters (rate constants, k) in the P450 catalytic cycle for bacterial and microsomal systems (References: Archakov and Bachmanova, 1990; Ruckpaul and Rein, 1984) Stage

P450cam

Microsomal P450s

1. Substrate binding*

47s−1 (4°C) (4.1×106 M−1 S−1)

2. First reduction

30–33 s−1 (25°C)

3. Oxygen binding 4. Second reduction† Turnover number**

1.7×106 M−1 s−1 (20°C) 1–4 s−1 1900 mm−1

50 s−1 (benzphetamine) (104–105 M−1 s−1) type I substrates 18 s–1 (benzphetamine, LM2) 11.5 s–1 (absence of substrate, LM2) >106 M−1 s−1 (25°C) 2–7 s−1 24 min−1 (benzphetamine, LM2) 28 s−1

>600 s−1 (4°C)

74


Stage

P450cam

Microsomal P450s

s−1

26 – – 30 s–1 ** kcat=number of substrate molecules metabolized to product per molecule of enzyme per unit time (expressed either as s−1 or mi−1). † the second reduction is generally regarded as the rate-determining step and, in microsomes, is probably mediated by cytochrome b5. * this is a high affinity process with KD=1–5 µ M (camphor binding to P450cam). Notes: 1. The catalytic turnover of most microsomal/eukaryotic P450s is low relative to those of prokaryotic P450s, being typically between 0.1 and 10 min−1. However, P450BM3 with its covalently linked reductase has the highest known catalytic turnover of any P450 at 4600 min−1. 2. Most of the kinetic parameters listed above are for the faster phase of a biphasic process. 3. The binding of substrate and of redox components tends to lower the redox potentials of both P450 and redox partners, respectively. 4. The first electron reduction readily gives rise to rapid oxygen binding, followed by the rate-determining second reduction. 5. The first electron reduction is apparently tightly coupled with iron spin-state equilibria and substrate binding.

and is accompanied by spectral changes, readily monitored in the UV, which indicate a modulation in the iron spin-state equilibrium towards the high-spin form (Sligar, 1976) especially in the case of type I substrates. The substrate-bound form of P450cam (CYP101) has been isolated and crystallized, such that its three-dimensional molecular structure can be determined by X-ray crystallography (Chapter 6). The thermodynamics of substrate binding to P450 are consistent with the desolvation of the active site (Griffin and Peterson, 1972) leading to a favourable entropy change ( S>0) and it is generally regarded that the binding of essentially hydrophobic substrates to P450 is largely an entropy-driven process. The relatively stable substrate-bound complex is readily reduced to the Fe(II) state because substrate binding lowers the P450 redox potential by about 100 mV (Ruckpaul et al., 1989), thus facilitating electron transfer from either a flavoprotein reductase or an iron-sulphur redoxin (Sligar and Gunsalus, 1976; Guengerich, 1983; Light and Orme-Johnson, 1981). It is thought that the binding of substrates to P450 brings about a conformational change in the enzyme which triggers interaction with the redox component. However, the solid state three-dimensional structures of substrate-bound and substrate free P450cam(CYP101) are almost identical (Poulos et al., 1985, 1986, 1987). Nevertheless, under the dynamic conditions operating in the biophase, conformational triggering of the redox component interaction brought about by substrate-induced scissoring of two (or more) secondary structures (i.e. either -helices and/or -sheets) could well occur, bearing in mind the conformational flexibility and dynamic aspects of protein structure. Veitch and Williams (1992) have suggested that two approximately parallel -helices in hemoproteins may be subject to a scissoring motion accompanying the binding of substrates. In cytochrome P450, the most obvious choice would be the I and L helices which sandwich the heme group. However, another possibility could involve the I helix and a -sheet close to the binding site as a conserved lysine residue (K314 in P450cam, K349 in P450BM3) at the turn of this -sheet is known to be associated with binding to reductase (Bernhardt et al., 1988). Protein dynamics calculations on P450 models could shed further light on the question of conformational triggering of P450 reduction, but Xray crystallographic studies only show significant thermal motion around threonine–185 (T185) and tyrosine-96 (Y96) in P450cam, which are involved in substrate access and binding, respectively (Poulos et al., 1986).


75

As threonine-185 is adjacent to another residue (arginine-186) in P450cam, that is also known to show thermal changes on binding, and forms an internal ion-pair with the conserved acidic residue (aspartate-251) which is itself adjacent to the invariant threonine residue (threonine-252 in P450cam) known to participate in the oxygenation mechanism (Atkins and Sligar, 1988; Imai et al., 1989a; Gerber and Sligar, 1992, 1994), it is possible that substrate binding-induced conformational changes distal to the heme face may have a profound relevance to the oxygenation mechanism and catalytic cycle in P450. One can visualize substrate binding to P450 as operating a conformational ‘switch’ by opening and closing the active site during and following substrate access, respectively. The characteristically more open active site of the P450BM3 crystal structure (Ravichandran et al., 1993), which is substrate free, indicates that there may be a significant conformational change on binding substrates which modulates iron spin-state equilibria, redox potential and, ultimately, oxygen binding, activation and insertion into the substrate. It is thought that bound water molecules link charged and/or polar residues in the active site such that they facilitate proton transfer (Gerber and Sligar, 1994), so the binding of substrate is likely to play a part in bringing about key stages in the P450 catalytic cycle. This is due to the proximity of substrate binding residues to those thought to be relevant for oxygen activation and transfer. For example, in P450cam, Val-295, Ile-395 and Thr-185 are close to the bound substrate, camphor (Poulos et al., 1985) and analogous residue positions are present in the P450BM3 structure (Ala-328, Leu-437 and Leu-181, respectively) which are likely to be in close substrate contact (Ravichandran et al., 1993). Although there are some notable differences between the two crystal structures (Lewis, 1995a) and their amino acid sequences, the similarity in tertiary structure suggests that nearby basic residues distal to the heme will be affected by substrate binding, which could modify ion-pairing to the conserved acidic residue adjacent to the invariant threonine and also transmit a conformational change to the conserved lysine residue (Lys-314 in P450cam , Lys-349 in P450BM3) via an intervening -sheet motif. Whatever the mechanism may be, it is clear that substrate binding to P450 will bring about some sort of a change in the enzyme which lowers the iron redox potential as well as causing a shift in the ferric iron spin-state equilibrium towards the high-spin form while, at the same time, desolvating the active site thus giving rise to a favourable entropy change. It is possible that these factors are causally linked and there is some evidence to suggest that they are (Gibson and Tamburini, 1984; Tamburini, 1982; Otsuka, 1970; Sipal et al., 1979; Blanck et al., 1983; Sligar, 1976; Sligar et al., 1976, 1979, 1984; Schwarze et al., 1985). However, following substrate binding, the next stage is a single electron reduction of the P450 via a bound redox partner. 3.2.2 Stage 2: The first reduction A relatively fast step in the cycle is the transfer of the first electron from reductase, or a redoxin; the latter is in itself made up of three separate stages, involving electron transfer from NAD(P)H to reductase, from reductase to redoxin and, finally, from redoxin to P450. In the microsomal system, there is no intermediary redoxin and the reductase contains both FAD and FMN subdomains which are involved in electron transfer directly to P450. In P450cam, electron transfer from NADH to putidaredoxin reductase is rapid ( >600 s−1at 4°C) whereas the subsequent transfer to putidaredoxin and reduction of P450 are significantly slower, being 26 s−1 (at 4°C) and 30–33 s−1 (at 25°C), respectively (Archakov and Bachmanova, 1990). In contrast, the analogous electron transfer chain in microsomal P450s is appreciably slower, especially from NADPH to the flavoprotein reductase (28 s−1) whereas the rate of reduction of P450LM2(CYP2B4) by reductase is increased by the presence of bound substrate, rising from 11.5 s−1 to 18 s−1 in the presence of

76


benzphetamine, for example (Ruckpaul and Rein, 1984). It is possible that the reason for the diminished rate of electron transfer from bacterial to microsomal P450s is because, in the latter, there are two flavin electron transfer components namely FAD and FMN, whereas the bacterial P450 system (P450cam) comprises only the FAD cofactor in putidaredoxin reductase. However, the rate of inter-flavin electron transfer is relatively rapid in the microsomal system, and one can assume that the two flavin rings of reductase are in fairly close proximity (about 4 Å apart) to facilitate an efficient electron-coupling process. If this is true, then the protein environment may be controlling the rate of electron transfer. Apparently, the binding of redox components to P450 lowers the redox potential (Archakov and Bachmanova, 1990) which suggests that there is some cooperation between the binding interaction and electron transport pathway. In the case of P450cam, it is generally thought that at least four basic residues on the P450, proximal to the heme, form ion-paired electrostatic interactions with complementary acidic residues on putidaredoxin (Stayton et al., 1989; Stayton and Sligar, 1990). Presumably, there are similar interactions in the case of reductase binding to microsomal P450s, and some likely candidates have been identified following chemical modification experiments on P450LM2 (Bernhardt et al., 1984, 1987, 1988, 1989a and b). However, there is strong evidence (Munro et al., 1992, 1994) for the role of the conserved tryptophan residue (Trp-96 in P450BM3) in electron transfer from reductase to the heme moiety in microsomal P450s and in P450BM3 (CYP102), where the indole nitrogen atom of this amino acid is only 3 Å away from one of the heme propionates (Lewis, 1995a). Another conserved basic residue in microsomal P450s is a lysine, analogous to Lys-349 in P450BM3 (Lys-314 in P450cam) which appears to be a prime contact point with a complementary acidic residue on reductase (Lewis, 1995a) and, presumably, could also represent a site of interaction between putidaredoxin reductase and P450cam. This possibility has some support from the finding that, in a reconstituted system, P450LM2 is catalytically competent with electron donors obtained from bacterial P450s (Bernhardt and Gunsalus, 1992). 3.2.3 Stage 3: Oxygen binding Following reduction of substrate-bound cytochrome P450, the next step is the rapid binding of molecular dioxygen to the high-spin iron(II)P450-substrate complex. The rate constant of this process is over 106 M−1 s−1 at 25°C (Archakov and Bachmanova, 1990) and the resulting oxyferrous complex is relatively stable, autoxidizing to superoxide at a rate of 0.01 s−1 at 20°C. The presence of a bound substrate increases the stability of the oxygen-bound P450 complex between 12- and 15-fold, and it is possible to isolate the substratebound oxyferrousP450 complex at subzero temperatures. Formation of oxygenated Fe(II)P450 can be observed spectroscopically: the UV difference spectrum of microsomal P450 exhibits absorption maxima at 440, 560 and 590 nm, whereas the bacterial P450cam and mitochondrial P450scc show UV maxima at 418 and 555 nm for the oxygen-bound Fe(II) complexes. The binding of oxygen to P450 results in the ferrous iron returning to the low-spin configuration and, presumably, the iron atom moves back into the plane of the porphyrin ring as is observed in hemoglobin and myoglobin (Lippard and Berg, 1994; Kaim and Schwederski, 1994). It is possible that one of the factors responsible for the high binding affinity of dioxygen for reduced P450 is the spin-spin interaction energy, which will be proportional to the product of the two spin quantum numbers (Tovrog et al., 1976). Molecular dioxygen in the triplet ground state has two unpaired electrons whereas high-spin ferrous P450 has four, so the spin-pairing interaction is expected to be quite significant. This might explain the relevance of the shift from low- to high-spin Fe(III) which accompanies substrate binding to P450. However, it is the Fe(II) state which exhibits high affinity for oxygen (as well as for


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Figure 3.2 The relationship between half wave redox potential (E½) and oxygen affinity (pKO2) for a series of cobalt porphyrin complexes (Carter et al., 1977).

carbon monoxide) and, therefore, other electronic factors pertaining to both the ligand and the heme iron must also be important, such as the electron-deficient nature of dioxygen and the overall negative charge of the Fe(II)heme-cysteinate complex. It is known that heme iron in the Fe(II) state is a good -donor which facilitates the strong binding of -acceptor ligands such as oxygen, carbon monoxide and nitric oxide (da Silva and Williams, 1991). When the iron atom is out-of-plane, as it will be in the high-spin form, one can assume that this affinity for small -donor/ -acceptor ligands is greater (Bertini et al., 1994). It should be remembered that carbon monoxide is also able to bind rapidly to reduced P450 but at a slightly lower rate (k=104–106 M–1 s–1) than, and about a tenth of the affinity of, oxygen. The accompanying spectral change, in particular the intense Soret absorption maximum at around 450 nm, is especially characteristic of the enzyme and was, of course, of prime significance in its discovery (Chapter 1). However, as far as oxygen binding is concerned, the equilibrium constant is proportional to the metal ion redox potential (Figure 3.2) in cobalt complexes with nitrogenous ligands (Carter et al., 1973) and a similar situation may occur in hemoproteins (Addison and Burman, 1985). There is evidence to suggest that the oxyferrous cytochrome P450 complex comprises a significant contribution from the ferric superoxide (Fe3+O2–) canonical form, brought about by the transfer of an electron from iron to dioxygen. Support for this hypothesis comes from the fact that autoxidation of the oxygenated P450 complex produces superoxide and that, in a reconstituted microsomal system, P450 forms two distinct complexes on oxygen binding (Archakov and Bachmanova, 1990). The initial ternary complex exhibits UV absorption maxima at 430 nm and 450 nm, undergoing a slow conversion to a more stable complex, that gives an absorption maximum at 440 nm, which is isolatable at -30°C. The absolute spectrum of Complex II shows the same absorption maxima as that of oxyferrous cytochrome P450cam, namely 420 and 558 nm, with a broad absorption maximum at 442 nm in the UV difference spectrum. These processes can be summarized as follows:

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As is also true for oxyferrous P450cam, the microsomal oxygenated complex autoxidizes biphasically in a first-order process to form, in this instance, peroxide rather than superoxide, although the latter may be a precursor, as the dismutation of superoxide to peroxide is well known (Archakov and Bachmanova, 1990). 3.2.4 Stage 4: The second reduction Apparently, the input of the second reducing equivalent as required by the known stoichiometry of the reaction, is the rate-determining step (Imai et al., 1977) from a consideration of the overall kinetics (Table 3.2). This major pathway for the decomposition of the oxycytochrome P450-substrate complex proceeds with a rate constant of about 3-4 s−1 in P450cam and is slower in microsomes at around 1 s−1 (Archakov and Bachmanova, 1990). In a series of elegant experiments, Gibson and Tamburini demonstrated that cytochrome b5 could mediate in the transfer of the second electron to the oxygenated P450 complex, although its involvement in the first reduction process appears to be unlikely on kinetic grounds (Tamburini and Gibson, 1983; Gibson and Tamburini, 1984). The rate constant for electron transfer from NADH to cytochrome b5 via cytochrome b5 reductase is 30 s−1, whereas that for cytochrome b5 to P450 is between 2 and 7 s−1. The corresponding process for NADPH electron transfer to cytochrome P450 via its reductase proceeds with a rate constant of 1.1 s−1 (Ruckpaul et al., 1989). The addition of a second electron to the oxycytochrome P450 complex gives rise to a species formally expressed as either Fe2+O2– or Fe3+O22–, and these may represent possible canonical structures of the reduced oxygenated P450 complex. Decomposition of this iron peroxy (or superoxy) species can give rise to superoxide, but is diminished by substrate occupancy of the heme site (Archakov and Zhukov, 1989). There is also evidence for the production of hydrogen peroxide in the breakdown of the reduced oxycytochrome P450 complex, especially where substrate oxygenation is unfavourable (Archakov and Zhukov, 1989; Archakov and Bachmanova, 1990). In the absence of substrate, the reduced P450 complex readily forms superoxide with a rate constant of 29 min−1, although dismutation of this species will be evidenced by peroxide production (Ruckpaul et al., 1989). With a bound substrate, the reduced oxycytochrome P450 complex undergoes rapid rearrangement to yield the required oxygenated substrate and water, with a rate constant of around 30 min−1, which varies considerably depending on the substrate and source of P450 (Ruckpaul and Rein, 1984). 3.2.5 Stage 5: Product formation The mechanism by which the oxygenated reduced P450-substrate complex breaks down to form products of the overall reaction is the least well understood process in the P450 catalytic cycle. A number of mechanisms have been proposed for the breakdown of the iron peroxy complex which lead to the formation of the known products (Ortiz de Montellano, 1986b; Kappus, 1993; Rein and Jung, 1993). There is varying support for all of these mechanisms and it is possible that a different one may operate depending on the circumstances, such as type of substrate or P450 isozyme. Most workers favour an iron (V) oxene (Fe=O) intermediate as the oxygenating species in P450 catalysis, although the Fe(V)O species has not been reported in any model porphyrin complexes to date, and this formulation has been challenged (Sawyer,


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1987). One reason for the popularity, however, of the iron oxene (ferryl) intermediate is based on an analogy with peroxidase compound II where a similar species (FeIV=O) is known (Dawson, 1988). However, the Mössbauer spectrum of P450 more closely resembles that of oxyhemoglobin (and indicating the Table 3.3 A comparison between bond lengths, bond energies and O-O stretching frequencies of dioxygen species (Reference: Lewis, 1986) Species

Bond length (Å)

Bond energy (kJ mole–1)

O−O stretching frequency (cm–1)

O−O Dioxygen O−O Superoxide O−O Peroxide

1.21 1.26–1.33 1.49

497 276 146

1555 1107(1145) 850

presence of superoxide) rather than that of a peroxidase, which converts hydrogen peroxide as a substrate into water. The resonance Raman spectrum of oxygenated P450 under catalytic conditions indicates the presence of an iron dioxygen species (O−O stretching vibration frequency 1141 cm−1) which is consistent with the presence of a superoxide complex (Egawa et al., 1991). One can view the formation of superoxide as a possible intermediate stage in the complete activation of dioxygen by the sequential addition of two electrons to produce peroxide, as follows: This process elongates the oxygen-oxygen bond from 1.21 Å, in molecular dioxygen, to 1.49 Å for peroxide via the intermediary superoxide stage, where the bond length may be between 1.26 Å and 1.33 Å (Table 3.3). During this process, where the iron porphyrin system in P450 represents a means of transferring electrons to oxygen in a controlled manner, the oxygen-oxygen bond energy becomes progressively weaker, going from 497 kJ mole–1 for O2, through 276 kJ mole–1 for O , to 146 kJ mole–1 in O . This stretching and concomitant weakening of the dioxygen bond facilitates cleavage of the oxygen species, probably present as an iron peroxy adduct, and subsequent insertion of one oxygen atom into the substrate. The other oxygen atom forms a water molecule by acquiring two protons during the course of the rearrangement. In fact, the oxygen-oxygen bond energy appears to parallel both the bond length and the vibrational stretching frequency of the free dioxygen species (Table 3.3) although one would expect that the coordination to iron in P450 will modify these values. 3.3 Nature of the oxygenating species According to the different mechanisms proposed (Groves and Watanabe, 1988; Sligar et al., 1984; Blake and Coon, 1981; White and Coon, 1980; Coon and White, 1980) the oxygen-oxygen bond is either homolytically or heterolytically cleaved to form a molecule of water and the iron oxene (Fe=O) species similar to that of compound II in peroxidase. Unfortunately, unlike that of peroxidase compound II, this intermediate has not been observed spectroscopically in P450. The iron oxene species in peroxidase, however, has been shown to be relatively stable and exhibits an FeO stretching frequency at around 780 cm −1 in the resonance Raman spectrum (Egawa et al., 1991). One is led to the conclusion, therefore, that the existence of an iron oxene in P450 is either unlikely or too unstable to be observable, possibly due to the trans effect of the thiolate ligand. It is rather curious, however, that such a species is both observable and apparently stable in peroxidase but not in P450; although, in the latter, the presence of substrate would be

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expected to readily accept oxygen from the reactive species, whatever its nature may be. The iron-oxygen bond energy in Fe=O can be expected to be fairly high, possibly as much as 390 kJ mole−1 but more likely to be closer to 270 kJ mole−1 and, consequently, it is surprising that such a widely-quoted intermediate in P450 oxygenations does not appear to have any firm evidence for its existence in the P450 mono-oxygenase system. However, there is some circumstantial evidence for the existence of an iron oxene intermediate in P450 from the work carried out using iodosobenzene (PhIO) as a model system, which has been shown to be capable of carrying out a number of P450-like oxygenations (Okazaki and Guengerich, 1993). There have also been several different model systems, reported in the literature, which can produce aromatic ring hydroxylations and other oxygenations similar to those catalysed by P450, and a number of these have been compared (Ullrich and Staudinger, 1969). The Fenton reagent (Fe(II), H2O2) appears to show some of the characteristics of P450-mediated oxygenation of carbon substrates and organic peracids are also able to reproduce approximately the same reaction products, depending on the nature of the chemical. It is thought that the Fenton reaction generates hydroxyl radicals (OH) whereas a hydroperoxy species has been implicated in other model systems. The isomer ratios of hydroxylation products for simple aromatic (Ullrich et al., 1968) and aliphatic (Frommer et al., 1970) compounds using such model P450 systems, can sometimes be fairly close (Table 3.4) to those found with microsomal preparations, leading to the conclusion that the active oxygen species may differ from one substrate to another, or between different P450 systems. For example, one would expect an electrophilic oxygenating intermediate to carry out aromatic hydroxylations, but a nucleophilic reagent could readily perform proton abstraction of aliphatic groups prior to oxygen insertion (Lewis et al., 1989a). Consequently, one might tend to favour a nucleophilic oxygen species which, under certain circumstances, could give rise to an electrophilic oxygen moiety. Consideration of the P450 reaction cycle and its likely intermediates, especially following the second reduction, suggests that an iron peroxide could fulfil the role of an oxygen nucelophile, whereas possible candidates for an electrophilic oxygenating Table 3.4 A comparison between various P450 model systems in hydroxylation of anisole (References: Ullrich and Staudinger, 1969; Ullrich et al., 1968; Ullrich, 1977) Position of anisole hyd droxylation % Oxygenating system

ortho

meta

para

Reagent/species

P450 Fenton reagent Udenfriend system Hamilton system Ullrich system Trifluoroperoxyacetic acid

11 84 46 66 36 73.7

0 0 22 0 47 0

89 16 32 34 17 26.3

Fe(II), O , 2H+ Fe(II), H202(OHÇ) Fe(II), O2 , EDTA, ascorbate Fe(II) or Fe(III), H2O2, quinol Sn(II), O2, HPO CF3COOOH

species include an iron oxene, an iron hydroperoxide, or an iron hydroxide which may have some of the characteristics of a hydroxyl radical. In fact, an iron oxene radical (Fe=O), resembling compound II of peroxidase, could well represent the optimum electronic configuration for electrophilic attack of certain types of P450 substrates, such as aromatic compounds and tertiary amines (Guengerich, 1993a and b). Electronic structure calculations on some of these species within a thiolate-bound heme environment give theoretical support to these suppositions. Table 3.5 shows the results of molecular orbital calculations by the


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Table 3.5 Results of molecular orbital calculations on P450 heme models (Reference: Lewis, unpublished results, similar values have been reported in Ruckpaul and Rein, 1984)

CNDO/INDO methodology on P450-like heme models with different oxygen species ligating the iron. Table 3.5 shows that the oxygen atom directly adjacent to the heme iron exhibits a partial positive charge, whereas the outer oxygen atom tends to show a negative partial charge. One possible reason why the model reagent systems do not give the same isomeric ratios of hydroxylation products to that shown by P450 may be due to the fact that, in the enzyme system, the

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substrate is likely to be orientated by amino acid residues in the vicinity of the active site, such that certain positions will be preferential for oxygenation. Nevertheless, model oxygenating systems are useful for exploring different oxygen species able to perform substrate oxygenations (Mansuy and Battioni, 1989; Murray and Groves, 1986; Woggon and Matile, 1992) and, currently, the iodosobenzene (PhIO) reagent appears to represent one of the most promising (Blake and Coon, 1989; Sligar et al., 1984). However, the fact that this compound can give rise to oxidation products similar to those obtained by P450 does not necessarily prove that the active oxygenating species in cytochrome P450 oxygenations is an iron oxene. In fact, it could be argued that PhIO bears a closer resemblance to an iron peroxy species (Fe−O−O) rather than the iron oxene (Fe=O), with the oxygen of iodosobenzene being analogous to the outer oxygen of the iron peroxide. PhIO also has the ability to readily produce water, which suggests that either the Fe=O intermediate forms water after (or during) oxygenation of the substrate, or that iodosobenzene is perhaps a better model for the iron peroxy species (as this is thought to produce water prior to the formation of the iron oxene) than the Fe=O species itself. Also, isotopic labelling studies have shown that solvent water oxygen atoms are present in the products of iodosobenzene-driven hydroxylation reactions; this suggests that single oxygen insertion from an iron oxene is not essential (Hawkins and Dawson, 1992). Furthermore, the use of cumene hydroperoxide as a model reagent for P450 oxidations, and its ability to reproduce some of its reaction products, tends to give additional support for an iron (hydro)peroxy intermediate as the oxygenating species in P450 (Blake and Coon, 1981; Vaz et al., 1991; Akhtar et al., 1994; Lee-Robichaud et al., 1995). Such considerations lead one to consider alternative mechanisms for P450-catalyzed oxygenations which involve an iron peroxide (or hydroperoxide) as the active intermediate. However, theoretical calculations of electronic structure using molecular orbital (MO) methodology indicate that the oxygenating species in P450-mediated reactions is likely to be electrophilic in nature (Ruckpaul and Rein, 1984). In particular, it has been reported that HOMO frontier orbital electron densities match the positions of P450-mediated hydroxylations on aromatic rings (Ackland, 1993; Rietjens et al., 1993). Moreover, in the example of toluene, which is metabolized by P450 at different ring positions as well as on the methyl group, the percentage hydroxylation appears to parallel electron density values for both aromatic (Lewis, unpublished results) and methyl hydrogen atoms (Lewis et al., 1995a). As the HOMO (highest occupied MO) is involved in the above examples, it suggests that perhaps the LUMO (lowest unoccupied MO) on an electrophilic oxygenating species may be important, and this finding tends to support the iron oxene mechanism. This is because MO calculations on various heme oxygen model complexes with a thiolate ligand show that the iron oxene complex has a low-lying LUMO and a partial positive charge on oxygen (Loew, 1983). However, the iron superoxide model also exhibits a low LUMO energy, with a partial positive charge on the inner oxygen, and the LUMO of the complex appears to predominate on the superoxide moiety (Table 3.5). Nevertheless, it is likely that steric factors, probably involving key amino acid residues in the P450 active site, will override the often somewhat small variations in substrate electron densities in directing the course of P450–mediated metabolism (Lewis, 1995a). If this is true, it is possible that the orientation of a substrate molecule in the active site may be determined by, for example, - stacking interactions between aromatic rings (Lewis and Lake, 1995). Hence, the electron distribution in a substrate benzene ring may serve to orientate the molecule for metabolism at a given position via an optimum stacking interaction with an aromatic amino acid residue in the P450 active site. As molecular electrostatic potential energy surfaces can sometimes be used to investigate possible interactions between ligands and receptor binding sites, or between substrates and enzyme active centres, a relationship between electronic distribution and molecular orientation within a binding site could explain the finding that, for example, electrostatic potential energy maxima and minima closely match the positions of P450related metabolism in aflatoxin B1 (Lewis, 1994a, 1996b).


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Although the precise nature of the oxygenating species is unknown, the results of MO calculations tend to support the oxenoid mechanism for P450-mediated oxygenations. Using energy calculations, Korzekwa and co-workers have also shown that the site of hydroxylation in P450 substrates is consistent with the attack of an electrophilic ferryl intermediate (Korzekwa et al., 1990; Korzekwa and Jones, 1993). Further experimental support for the participation of an iron oxene is emerging from the studies on deuterium isotope effects in P450 oxidations (Hanzlik et al., 1984; Atkins and Sligar, 1987; Okazaki and Guengerich, 1993). However, a recent review of deuterium isotope effect data by Lu (1992) indicates that the evidence is not conclusively in favour of the oxenoid pathway. Indeed, one of the studies on P450cam has shown that an observed excess water production results from reduction of the oxene (Atkins and Sligar, 1987). This suggests that more than one pathway exists for the breakdown of dioxygen, at least as far as the P450cam system is concerned, with the possibility of a molecule of water being formed from either oxygen atom. It is not clear, however, from the P450cam study whether the substrate can be oxygenated by either the iron peroxide or the iron oxene, and a recent finding has demonstrated the catalytic competence of superoxide and ferrous P450cam which can be interpreted in terms of a peroxy intermediate (Kobayashi et al., 1994). Consequently, it is advisable to regard the nature of the active oxygen species in P450-catalyzed reactions as still being open to question. 3.4 Participation of active site amino acid residues in P450 oxygenations There is emerging evidence for the role of two conserved amino acid residues in the heme environment of P450 as participants in the oxygenation mechanism (Gerber and Sligar, 1992, 1994; Martinis et al., 1989; Imai et al., 1989a; Tuck et al., 1993a and b; Furuya et al., 1989a and b; Shimizu et al., 1994). Site-directed mutagenesis experiments on the P450cam system have shown that modification of Thr-252 to Ser brings about a decrease in camphor hydroxylation, with concomitant increase in peroxide formation (Table 3.6); whereas removal of a hydrogen bond donor/ acceptor amino acid residue at this position, by changing Thr-252 to Val or Ala, progressively switches the major product from 5-hydroxycamphor to peroxide (Imai et al., 1989a). The inference to be drawn from these findings is that Thr-252 in P450cam participates in the oxygenation mechanism by hydrogen bonding with, pre Table 3.6 Effect of site-directed mutagenesis on P450cam activity (Reference: Imai et al., 1989) Mutant

% 5–hydroxy camphor

% H2O2

O2 consumption (min–1)

T252 (Wild type) T252S T252V T252A

96 81 22 6

5 15 45 83

1330 1100 420 1100

sumably, an iron peroxy intermediate. More recently, Gerber and Sligar (1992, 1994) have provided evidence, based on site-directed mutagenesis of both Asp-251 and Thr-252 in P450cam, which indicates that these two highly conserved residues act cooperatively in facilitating scission of the oxygen-oxygen bond, which is probably the key stage in the mechanism leading to the formation of oxygenated products. Based on the crystal structure of P450cam (Poulos et al., 1985, 1987) it is possible to envisage the likely orientation of iron-ligated dioxygen with the outer oxygen pointing towards the Thr-252 hydroxyl group, which could be close enough for hydrogen bond formation. However, a charge relay system of the type

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proposed by Gerber and Sligar (1992, 1994) would necessitate an intervening solvent water molecule between Asp-251 and Thr-252 to enable proton transfer along a hydrogen-bonded conduit to occur. Mutation of Asp-251 to Asn produces a marked decrease in catalytic activity of P450cam, suggesting that the presence of an acidic amino acid residue at this position is important for fully competent oxygenase activity (Gerber and Sligar, 1992, 1994). Furthermore, mutagenesis experiments carried out on two basic amino acid residues (Arg-186 and Lys-178) which are close to Asp-251 in the P450cam active site (and probably interact electrostatically with anionic residues such as Asp-251) provide evidence for an extensive proton relay network in the bacterial enzyme heme environment (Lewis, 1995a) which could have a role in dioxygen bond cleavage (Gerber and Sligar, 1994). Much of this network, with the exception of the intervening water molecule between Asp-251 and Thr-252, can be observed in the P450cam crystal structure, and a similar situation may exist in microsomal P450s as indicated by homology modelling (Chapter 6). Site-directed mutagenesis studies have also been carried out on P450d (CYP1A2) within the putative active site region (Furuya et al., 1989a and b; Tuck et al., 1993a; Shimizu et al., 1994). These experiments also point to an important role for, in this case, Glu-318 and Thr-319, which correspond to Asp-251 and Thr-252, respectively, in P450cam. It is found that mutations to these residues alter the regio-specificity of substrate metabolism in addition to modifying the overall catalytic activities (Furuya et al., 1989a and b). Furthermore, the effect of site-specific changes on the formation of iron-aryl complexes indicates that the position of the distal I helix is shifted in CYP1A2 relative to P450cam(CYP101) suggesting that the P450BM3 (CYP102) crystal structure (Ravichandran et al., 1993) is a better model for microsomal P450s than P450cam (Tuck et al., 1993a). More recently, it has been reported that mutations to Glu-318 and Thr-319 in CYP1A2 may affect the nature of the oxygenating species and appear to be able to enhance the ease of oxygenoxygen bond scission of the bound peroxy species (Shimizu et al., 1994). It should also be noted that, in certain P450s where there is a change from the normally conserved Asp/Glu or Thr, the catalytic activity may be different from that of other P450s which possess the conserved acidic and adjacent threonine residues; some examples of these include allene oxide synthase (Song et al., 1993) and nitric oxide synthase (Degtyarenko and Archakov, 1993). 3.5 Thermodynamics of P450 substrate binding and spin-state equilibria It is generally accepted that there is a coupling between the spin-state and substrate binding equilibria in the P450 system (Gibson and Tamburini, 1984). This may be represented by a four-state model as follows:

where: HS and LS refer to high- and low-spin states, respectively; S refers to the presence of bound substrate, K1−K4 refer to the microequilibrium constants for ferric cytochrome P450; whereas consideration of ferrous P450 states in addition requires a total of eight states, usually represented as a thermodynamic ‘cube’ model (Sligar, 1976). It has been established that substrate binding to P450 modulates the spin-state equilibria usually by bringing about a shift from low-spin ferric to high-spin (Schenkman et al., 1981). Such effects may be investigated by UV difference spectral changes in the Soret region; and temperature dependence studies


85

enable calculation of the relevant thermodynamic quantities, G, H and S. This is achieved as a result of the relationship between equilibrium constant, K, and the Gibbs free energy, G: where: R is the gas constant and T is the absolute temperature; and due to the dependency of G on the enthalpy change, H, and entropy change, S, as follows: where: T is the absolute temperature. Griffin and Peterson (1972) demonstrated the high binding affinity of camphor for P450cam, and calculated the thermodynamic parameters ( G, H and S) for the substrate binding process. The free energy of binding (at 21°C) for the endogenous substrate, camphor, binding to P450cam ( G=−32.2 kJ mole −1) appears to be entirely due to the entropy component (Griffin and Peterson, 1972) which is generally regarded as being caused by the desolvation of bound water molecules in the enzyme active site, although there will be a small effect resulting from the loss of rotational and translational entropy of the substrate on binding (Williams et al., 1991). The crystal structure of substrate-free P450cam shows that there are six water molecules in the active site, one of which ligates the heme iron (Poulos et al., 1986). As these are not present in camphor-bound P450cam, one can conclude that about six water molecules are displaced from the vicinity of the heme in P450cam during substrate binding. A rough calculation (Lewis, unpublished results) indicates that the removal of six water molecules gives rise to an equilibrium binding constant (K=2.43×10– 8) which is close to the experimental value (K=2.21×10−8). However, a calculation of the entropy change brought about by substrate binding suggests that as many as 10 water molecules would be displaced, if one assumes that the entire entropy component is brought about by desolvation (Lewis, unpublished results). For microsomal P450s there are some differences with respect to the bacterial P450cam system, as far as microstate equilibrium thermodynamics are concerned. Although the substrate-free low-spin/high-spin equilibrium constants, K1, appear to be about the same in microsomes as P450cam, there are quite marked variations in the other K values: the substrate-bound spin equilibria (K2) are significantly lower in microsomal P450s than in P450cam, whereas the substrate binding equilibrium constants (K3 and K4) are considerably higher in microsomal P450s. However, substrate binding increases the spin-equilibrium constant irrespective of the source of P450, as shown by comparing the values of K2 with those of K1. It is also found that the equilibria governing substrate binding to low-spin P450 are much greater than for the high-spin state. Table 3.7 shows a comparison between microstate equilibria (K1 to K4) for microsomal P450s and P450cam, whereas Table 3.8 provides the relevant thermodynamic parameters ( G, H and S) for benzphetamine binding to Table 3.7 Substrate-binding and spin-state equilibrium constants (Reference: Ruckpaul et al., 1989) Type of P540

K1

K2

K3 (µ M)

K4 (µ M)

Substrate

P450cam (CYP101) P450LM2 (CYP2B4) P450PB-B (CYP2B1)

0.08 0.08 0.07

15.0 0.4 0.6

9.0 370.0 250.0

0.05 70.0 29.0

Camphor Benzphetamine Benzphetamine

Table 3.8 Thermodynamic parameters for substrate-binding and spin-state equilibrium in P450LM2 (CYP2B4) with benzphetamine as substrate (Reference: Ruckpaul et al., 1989) Equilibrium

constant

K1 K2

0.08 0.39

H (kJ mole−1) −44.3 −38.0

S (Jmole−1 K−1) −130 −121

G(kJ mole−1) −6.3 −2.5

86


Equilibrium

constant

K3 K4

0.37 (mM) 0.07 (mM)

H (kJ mole−1)

S (Jmole−1 K−1)

31.4 26.3

171 171

G(kJ mole−1) −18.8 −23.8

Table 3.9 Thermodynamic parameters for camphor binding and spin-state equilibria in P450cam (CYP101) (Reference: Sligar, 1979) Equilibrium

constant

K1 K2 K3 K4

0.084 15.0 9.0 (µ M) 0.05 (µ M)

H(kcal mole−1)

S(cal mole−1 K−1)

−10.3 −2.5 – –

−30.2 −13.8 – –

G(kcal mole−1) −1.44 1.59 −6.81 −9.84

P450LM2 (CYP2B4), and Table 3.9 gives thermodynamic data for camphor binding to P450cam (CYP101). The relatively large and unfavourable entropy change depicted for the spin-state equilibria reflect a greater ordering of the high-spin form, and this may be due to a conformational change in the P450 structure, particularly in the vicinity of the heme (Otzuka, 1970), which could involve an increased interaction between either individual amino acid residues or secondary structural units: such changes may conceivably influence the binding of redox partners, or ease of reduction. Furthermore, it is apparent that substrate binding to either low-spin or high-spin P450 is an entropy-driven process as, although the enthalpy change is unfavourable, it is more than compensated by a favourable entropy component (Table 3.10). In fact, if one assumes that the entropy change accompanying substrate binding to P450 is mainly composed of the desolvation energy contribution, then the ratio of the S values for benzphetamine and camphor binding to P450LM2 and P450cam, respectively, should Table 3.10 Thermodynamics of camphor binding to P450cam (a) and of benzphetamine binding to P450LM2 (b) (Reference: aGriffin and Peterson, 1972; bRuckpaul et al., 1989) (a)

T(°C)

H (kcal mole−1)

S

−T S (kcal mole −1)

G (kcal mole−1)

4.6 8.7 56.4 −15.7 −7.0 21.0 0 26.2 −7.7 −7.7 (b) T(°C) H (kJ mole−1) S −T S (kJ mole−1) G (kJ mole−1) 20.0 31.4 (±2.1) 171 (±9) −50.1 (±2.6) −18.8 (±2.5) Notes: The magnitude of the entropy component (T S) relative to the enthalpy change ( H) shows that the free energy of binding ( G) is essentially entropic in nature, and it is generally regarded that desolvation of the binding site is the major contribution to the overall entropy change. The volume (and surface area) of the solvent-accessible surface appears to have some relationship to the volume of water molecules displaced on substrate binding to P450, which is likely to be proportional to Sdesolvation. Therefore, physicochemical characteristics of the substrate may provide a means of estimating the magnitude of the entropy change accompanying binding, which is known to be the main factor in P450 binding for both the bacterial system (Griffin and Peterson, 1972) and liver microsomal P450 (Ruckpaul et al., 1989). A comparison between various physico-chemical properties of camphor and benzphetamine indicates that the entropy component of binding free energy is related to the dimensions of the substrate, as shown in Table 3.11.


87

Table 3.11 Comparison between the entropic components on binding to P450 and molecular dimensions for camphor and benzphetamine (Reference: Lewis, unpublished results)

Camphor Benzphetamine

M.Wt.

Volume (Å3)

Surface area (Å2)

−T S (kJ mole −1)

No. of water molecules displaced

152.24 239.38 1.572

151.72 234.36 1.545

158.66 246.97 1.557

−32.2 (21°C) −50.1 (20°C) 1.556

6–8 9–12 1.5–1.57

Note: These parameters may thus facilitate the calculation of binding interaction energies and estimates of the dimensions of various P450 active sites (see text for details).

be roughly equivalent to the ratios of their molecular volumes or surface areas, as the number of water molecules displaced on binding probably depends on the size of the substrate molecules. Inspection of the relevant data presented in Table 3.11 shows that the ratios of these quantities are all about 1.5–1.57, which indicates that estimates of the desolvation entropy component could be made on the basis of substrate physicochemical parameters. However, it should be considered that the active sites of the two types of P450 involved are likely to be topographically different, as are the hydrophobicities of the substrates, and these factors may help to explain some of the differences observed following substrate binding and the subsequent spin-state and redox-state changes. 3.6 The coupling of redox and spin equilibria Based on an analysis of the thermodynamic ‘cube’ model of substrate-binding, spin-state and redox-state equilibria, Sligar (1976) showed that it is possible to reduce the apparently complex series of interactions to a simpler form involving a straightforward coupling between spin and redox equilibria (Schenkman et al., 1981). This reflects the fact that substrate binding to P450 increases the high-spin component (up to 94 per cent in the case of camphor binding to P450cam) with a concomitant lowering of redox potential, which changes from −303 mV to −173 mV when camphor binds to P450cam (Sligar, 1976). One explanation for the simplification of the thermodynamic ‘cube’ model arises from the finding that, even at 4 K, ferrous cytochrome P450 exists almost entirely in the high-spin state, irrespective of substrate binding. In contrast, the temperature dependency of the P450 spin-state equilibria for the ferric state exhibits an increase in highspin content with rise in temperature. Observation of the effect of substrate binding on the P450 spin-state marker bands at 390 nm(HS) and 420 nm(LS) in the Soret region of the UV difference spectrum enables calculation of the modulation of spin-state equilibria on substrate binding (Gibson and Tamburini, 1984; Sligar et al., 1979), whereas redox potential titrations, involving a dye photoreduction technique, facilitate the monitoring of P450 redox potential changes following the addition of substrate (Sligar and Gunsalus, 1974; Sligar et al., 1979). Table 3.12 shows a comparison between spin-state and redox-state equilibria for bacterial P450cam and rat liver microsomal P450 with different substrates. Statistical analysis of these data (Table 3.13) confirms that there is a high correlation (r=0.96) between spin-state and redox equilibria for

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Table 3.12 Spin and redox equilibria for bacterial P450cam and rat liver microsomal P450 (Reference: Gibson, 1986) Spin state

Redox state

P450 type

% high spin

KHS/LS

E0’ (mV)

KFe2+/Fe3+

P450cam P450cam+camphor P450RLM P450RLM+hexobarbital P450RLM+benzphetamine

8 94 10 35 38

0.089 14.4 0.113 0.549 0.607

−303 −173 −300 −237 −225

1.2×105 8.0×102 1.2×105 9.4×103 5.9×103

Table 3.13 Relationships between spin and redox equilibria (data from Table 3.12; reference: Lewis, 1992a) n

s

0.61 E0’+187.1 5 11.76 (±0.11) log KHS/LS= −0.91 log KFe2+/Fe3+ + 3.54 5 0.27 (±0.15) n=number of observations; s=standard error; R=correlation coefficient; F=variance ratio. % high spin=

R

F

0.96

31.9

0.96

31.9

the systems and substrates concerned (Lewis, 1992a) in agreement with the work of Sligar and co-workers (Sligar, 1976; Sligar et al., 1979; Schenkman et al., 1981). Thus, from an application of the Nernst equation to the P450 system, an expression for the observed redox potential, E0 , yields the following:

where: R is the gas constant, T is the absolute temperature, F is the Faraday constant, KFe2+/Fe3+ is the redox equilibrium constant and KHS/LS is the spin equilibrium constant. Therefore, a straight line relationship should be observed between P450 redox potential and high-spin fraction, of negative slope equal to the ratio RT/F and with an intercept corresponding to the redox potential in the absence of the low-spin component (Schenkman et al., 1981). A manipulation of the term in parentheses shows that it may be equated with the reciprocal of the high spin fraction, such that the above expression simplifies to:

and this becomes (for T=298 K) the following: A graph of P450 redox potential against a high spin fraction yields a straight line of high correlation (r=0. 995) for the data shown in Table 3.12 and indicates, on the basis of the intercept, that the high-spin redox potential will be about –74 mV (Figure 3.3). Thus, it can be concluded that redox and spin equilibria are tightly coupled to substrate binding in the P450 system, and that the effect of substrate binding to P450 increases the high-spin content which, in turn, gives rise to a lowering of the iron redox potential (i.e. making it less negative) such that reduction is facilitated. Redox potentials are difficult to measure in the P450 system and, consequently, relatively few have been reported (Lewis, 1986; Ruckpaul et al., 1989). Nevertheless, it appears that, in bacterial and mammalian systems, the hemoprotein exhibits a relatively high negative redox potential of between −300 mV and −400 mV, depending on the source of P450, which diminishes by as much as 130 mV in the presence of bound substrate (Ruckpaul et al., 1989).


89

Notes: Although the redox potential of cobalt porphyrin complexes correloates closely with their oxygen affinities (Carter et al., 1973) it is also of interest to note that a similar relationship exits for hemoproteins comprising myglobins and hemoglobins (Addison and Burman, 1985). Based on an analysis of the oxygen binding affinities (P½O2 values) and iron redox potentials (Eo , mV) of 12 globins these coworkers derived and expression:

which is analogous to that proposed theoretically: that gives a slope value 59 mV, i.e. The log P½O2 for oxygen binding to P450cam can be calculated from the KD value of 0.6 µM, as being—5.921 (based on a P½O2 of 1.2×10−6 Torr). Putting this result into the equation for Eo , derived experimentally, gives a value of—295 mV, which is close to the known redox potential (−303 cV) of P450cam, and well within the limits of experimental error. However, this interesting result does not explain the fact that substrate binding increases oxygen affinity but lowers the redox potential of P450, so there are clearly several factors involved in defining hemoprotein redox potentials. Figure 3.3 The relationship between redox potential (E0’) and percentage high-spin component for several P450 systems (Gibson, 1986; Sligar et al., 1979).

Of additional importance is the finding that the shift in spin-state equilibrium in P450 positively affects both the rate of reduction and the metabolism of substrate (Blanck et al., 1983). For a series of eight benzphetamine analogues, it has been shown that the modulation of spin-state equilibria in phenobarbitalinduced rat liver microsomal P450, following substrate binding, correlates with the reduction rate constant (r=0.81) and, more significantly (r=0.94), with substrate turnover (Blanck et al., 1983) as shown in Figure 3.4. These results indicate that substrate binding probably initiates the entire P450 catalytic cycle where each stage is a natural consequence of the former, and that there is a relationship between the degree of spin-state change and rate of metabolism of substrate, due to a tight coupling between each intervening process.

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Figure 3.4 The relationship between rate of N-demethylation (Vmax) and percentage high-spin shift for a series of benzphetamine analogues (Blanck et al., 1982).

3.7 Redox potential and redox interactions in the P450 system The redox potentials of P450s from a variety of sources indicate that the substrate-free enzyme possesses a relatively high negative redox potential in comparison with other hemoproteins (da Silva and Williams, 1991). Based on a theoretical model proposed by Kassner (1973), where c-type cytochrome redox potentials were rationalized in terms of a local non-polar heme environment, Stellwagen (1978) showed that the iron redox potential in seven hemoproteins (Table 3.14) could be linearly related with the percentage heme exposure to a polar environment (Figure 3.5). Generally, the more buried the heme grouping, the more positive is the iron redox potential, with a relationship of high correlation (r=0.96) between the two quantities given by: Table 3.14 Correlation between Fe2+/Fe3+ redox potential (E0 ) and percentage heme exposure (Reference: Stellwagen, 1978) Hemoprotein

% heme exposure

E0’ (mV)

Cytochrome c2 Cytochrome c Cytochrome c550 Hemoglobin Hemoglobin Myoglobin Cytochrome b5 E0’=-014.94% exposure (±1.95)+343.9 Correlation coefficient: r=0.96

6 4 5 14 20 18 23

320 260 250 113 53 47 20


91

Figure 3.5 The relationship between redox potential (E0 ) and percentage heme exposure for various hemoproteins (Stellwagen, 1978).

E′=−14.94 per cent heme exposure +343.88 n=7; s=37.36; r=0.96; F=46.0 where redox potential (E0’)is expressed in mV. Applying this relation to the P450 system indicates that the percentage heme exposed to an aqueous environment could vary from about 35 per cent to 50 per cent, depending on the presence of substrate and source of P450. In fact, from a consideration of known P450 crystal structures, such a situation does appear to be consistent with this theory, as the overall hydrophobicity of the active sites reflects that of their respective substrates. Furthermore, one would expect that substrate binding should make the P450 redox potential less negative due to occupancy of the heme site, and this is indeed found to be true. As far as rate of reduction is concerned, it has been reported that the reaction rate between hemoproteins and hydrated electrons may be dependent upon a number of factors including molecular mass, number of aromatic residues and net surface charge (Hasinoff, 1985) assuming that the interaction is diffusion controlled. Although cytochrome P450 was one of the hemoproteins included in this study, it is important to consider the particular mode of interaction with the relevant enzyme’s redox partner, as this should have a major bearing on the heme reduction process.

92


Table 3.15 Redox Potentials (mV) in various P450 systems (References: Lewis, 1986; Ruckpaul et al., 1989; Veitch and Williams, 1992) (a) P450cam system (bacterial) E0 (mV) NADH −320 FAD −290 (−320 to −28p5 also reported) Putidaredoxin −240 P450cam −303 O2/ −160 /OH· −100 (b) P450scc system (adrenal cortex mitochondrial) NADPH −324 FAD −290 (−274 has also been reported) Adrenodoxin −270 P450SCC −400 (c) P450RLM system (liver endoplasmic reticulum) NADPH −324 FAD −290 FMN −270 P450LM2 Second reduction NADH Cytochrome b5 reductase Cytochrome b5 O2/H2O2 ‘O’/H2O

−300

O2/H2O

+820

−320 −330 +25 +295 +450

putidaredoxin reductase (−196 when bound to P450cam) (−173 when camphor is bound) (1st electron reduction) (2nd electron reduction) adrenodoxin reductase (−290 has also been reported) (−280 or less when cholesterol is bound) P450 oxidoreductase (−190 to −110 for second reduction depending on redox state) (−237 to −225 on substrate binding)

(+30 to +20 has also been reported) (Fe=O/Fe3+ has been reported to be +1000 to +1500)

As stated previously, the various redox partners in the P450 system vary somewhat, depending on the source of the enzyme, but these all provide a means of delivering the required reducing equivalents in two discrete stages, as demanded by the catalytic cycle (Figure 3.1). The source of electrons in the P450 system is either NADH or NADPH, depending on the type of P450 involved, and reduction is mediated by a flavoprotein reductase, and an iron sulphur redoxin in the case of bacterial P450cam or mitochondrial P450SCC; whereas microsomal P450s tend to be able to utilize cytochrome b5 to mediate transfer of the second electron (Gibson and Tamburini, 1984). The microsomal system does not require a redoxin as the flavoprotein reductase possesses both FMN and FAD, whereas the bacterial P450cam and mitochondrial P450SCC reductases only possess FAD as a cofactor (Degtyarenko and Archakov, 1993). Table 3.15 provides details of the relevant redox potentials of various entities participating in the P450 catalytic cycle, and some of these redox states are represented in Figure 3.6, showing the possible routes of electron transfer for different P450 systems. These redox pathways demonstrate that the electrons may be transferred down a potential gradient from reductant to oxidant

93

Figure 3.6 Electron transport pathways in various P450 systems.


species, between the interacting species and terminating in various oxygen redox couples. Figure 3.6 shows that, for example, the mediation of cytochrome b5 in the microsomal system possibly facilitates reduction of oxygen to hydrogen peroxide, or even to give the oxene intermediate, on the basis of the respective redox potentials. It is also of interest to note that the binding of various key elements in the system serves to promote the transfer of electrons required by lowering the relevant redox potentials of the particular components. For example, the binding of putidaredoxin to P450cam lowers (i.e. becomes less negative) the

94


redox potential of putidaredoxin to enable ease of electron transfer from putidaredoxin reductase, whereas camphor binding to P450cam lowers its redox potential such that reduction via putidaredoxin is facilitated. 3.8 Interactions between redox components Clearly, specific interactions between various redox partners in the different P450 systems cooperate in the transfer of electrons from the reductant species. The means by which redox components in biological systems are able to interact and facilitate electron transfer has been intensely studied (Larsson, 1982; Marcus and Sutin, 1985; Matthew et al., 1983; Simondsen et al., 1982) and a considerable body of information has accumulated from a variety of techniques. The majority of this work, as applied to the P450 system, has been summarized by Schenkman (1993) and Bernhardt (1993), whereas the role of phospholipid as a possible mediating factor in redox component-P450 interactions has been reviewed by Blanck and Ruckpaul (1993). Although much of the evidence points to electrostatic interactions between basic residues on P450 and acidic residues on the appropriate redox partner (redoxin, reductase and/or cytochrome b5) a recent study suggests that the true picture may be rather more complicated (Voznesensky and Schenkman, 1994) at least concerning the reductase-P450 interaction. Apparently, electrostatic repulsions may also play a part in limiting the degree of association between P450 and reductase in the bimolecular complex such that the binding interaction remains reversible. A considerable effort has been expended in determining the key amino acid residues on the various elements in the P450 complex that are involved in intermolecular interactions and electron transfer between redox components in the system. The role of phospholipid in facilitating redox component interactions in the microsomal P450 system has also been extensively studied, including the orientation, association and rotational characteristics of the P450 enzymes in the membrane phospholipid bilayer. The nature of the interactions between cytochrome P450 and its reductase have been investigated by Strobel et al., (1989) and in a number of studies by Bernhardt and colleagues (Bernhardt et al., 1984, 1987, 1988, 1989a and b; Schwarze et al., 1983) where the effect of chemical modification of acidic residues on reductase and basic residues on P450 were evaluated, respectively. Table 3.16 summarizes the results of these and other studies in terms of the likely amino acid residues involved in interactions between P450 and reductase. This comTable 3.16 Amino acid residues involved in ion-pairing and other interactions between redox partners in various P450 systems (References: Stayton et al., 1989; Dailey and Strittmatter, 1979; Sligar et al., 1974, 1991; Davies et al., 1990; Geren et al., 1984; Lambeth et al., 1984; Adamovitch et al., 1989; Wada and Waterman, 1991) Cytochrome b5‡

P450cam

Putidaredoxin*

P450cam

Adrenodoxin†

P450SCC

Heme propionate E48 E44 D60 Heme (E43 K154 K267 K270

R364 R72 K344 R112 F350 Q343)

D58 E65 E67 E72 W106 CO K148

R72 K344 R112 Q343 R364?

E64 E65 E74 D79 D86

K73 K109 K110 K126 K145


Cytochrome b5‡

P450cam

Putidaredoxin*

P450cam

Adrenodoxin†

95

P450SCC

K338 K342 † residues thought to interact with Adx reductase ‡ some of these residues may also bind to b5 reductase * residues also thought to interact with Pdx reductase Notes: 1. The role of the C-terminal trytophan residue (W106) in putidaredoxin (Pdx) has been studied (Davies et al., 1990) and it would appear that this represents a possible mediator for electron transfer from the redoxin to P450cam. As there is a highly conserved tryptophan present in many microsomal P450s, it is thought that, during the course of evolution, this residue became incorporated into P450s to facilitate electron transfer to the heme from reductase. Recent studies have been directed towards establishing the possible relevance of the analogous residue (W96) in P450BM3 (Munro et al., 1992, 1994). 2. It is possible that the mitochondrial P450 system involves a ‘shuttle’ mechanism (Lambeth, 1990) for adrenodoxin (Adx) mediating electron transfer between its reductase and P450SCC (Turko et al., 1989; Hara and Kimura, 1989) which could mean that the same surface residues are associated with ion-pairing interactions between Adx and its reductase, and between P450SCC and Adx. Whether this situation also operates in the bacterial P450cam remains open to further investigation.

pilation clearly demonstrates the relevance of ion-pairing to protein-protein associations within the P450 macromolecular complex. There is clear evidence that basic residues on P450 interact with acidic residues on reductase and on either cytochrome b5 or redoxin (Stayton and Sligar, 1990; Stay ton et al., 1989; Dailey and Strittmatter, 1979; Sligar et al., 1974, 1991; Davies et al., 1990). Moreover, there is also evidence that the interactions between cytochrome b5 and its reductase, and the interactions between redoxins and their reductases, are electrostatic in origin (Geren et al., 1984; Lambeth et al., 1984; Adamovitch et al., 1989; Wada and Waterman, 1992). The identification of the complementary charged residues on redox partners, by site-directed mutagenesis and other experimental techniques, can be used as a guide to facilitate modelling of the tertiary complexes of P450 systems, together with identification of the respective redox centre binding domains (Gerber et al., 1990; Hanukoglu and Gutfinger, 1989). The crystal structure of cytochrome b5 is known but, to date, there are no crystallographically-determined structures of either putidaredoxin, adrenodoxin, or their reductases. However, the crystal structures of various ferredoxins are available and, more recently, the structure of a ferredoxin reductase has been published (Karplus et al., 1991). As the known crystal structures of these proteins are fairly homologous with their analogues in the P450 system, it is relatively straightforward to produce models of the unknown structures via crystal structure templates, as mentioned in Chapter 6. However, in microsomal P450 oxidoreductases, it is necessary to model these based on ferredoxin reductase and flavodoxin, and then combine the two subdomains of the entire reductase structure. The known interaction sites with P450, including evidence from site-directed mutagenesis studies on P450BM3 (Klein and Fulco, 1993) make this process feasible, although it is difficult to link the two subdomains precisely because the topology of the intervening sequence is unknown. Nevertheless, the models of P450–redox partner complexes are in general agreement with known experimental evidence from site-directed mutagenesis and other studies (Gerber et al., 1990; Hanukoglu and Gutfinger, 1989) and enable an exploration of possible electron transfer pathways within the P450 macromolecular assembly (Chapter 6). Furthermore, such studies can indicate the likely orientation of P450 in the endoplasmic reticular membrane, especially when additional experimental data is utilized to aid the modelling process, and it is possible to include a simulated phospholipid bilayer (Damodaran and Merz, 1994) interactively within the model.

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3.9 Protein-lipid interactions in the P450 system There is considerable evidence to support the view that the membrane plays an important role in the microsomal P450 system (Archakov and Bachmanova, 1990; Arinc et al., 1991; Ruckpaul and Rein, 1984; Ingelman-Sundberg, 1986; Schenkman and Griem, 1993). In fact, it would appear that there is a multifaceted role of the membrane phospholipid in both facilitating and moderating the interaction between the various redox partners, and in providing a vehicle for the ingress of substrates to the P450 access channel and active site. The N-terminal segment of between 20 and 40 residues in microsomal P450s, the N-terminus of the P450 oxidoreductase, and the C-terminus of cytochrome b5, are all thought to span the phospholipid bilayer and serve to anchor these elements of the redox system within the membrane, such that their binding kinetics can be modulated by protein-lipid interactions. These stretches of peptide, which probably restrict the motion of the various components of the P450 system, possibly also interact with each other when complexation occurs, with the formation of macromolecular assemblies of several P450s clustered around a central oxidoreductase (Greinert et al., 1982) with cytochrome b5 units able to bind also, as required by the stoichiometry of the electron transfer reaction. The fact that the lengths of the P450 N-termini vary from about 20 to 40 residues, has led to the suggestion that they may comprise either a single helical stretch of 20 residues or a helix-turn-helix motif for the longer segments (Ozols, 1989). Apparently, an -helix of 20 amino acid residues is about the same length as the width of a phospholipid bilayer, thus providing some support for this hypothesis. Tryptic cleavage of the N-termini of microsomal P450s leads to either a loss or reduction in their catalytic activity, as does association of the redox components in a membrane-free system. This diminution of enzymatic activity following separation from the membrane phospholipid has discouraged crystallographic studies on microsomal P450s, although the current emergence of expression and reconstituted systems have led to a renewed interest in such endeavours. Rotational diffusion experiments using an electron spin resonance (ESR) technique (Schwarz and Pirrwitz, 1990) provides evidence of the membrane topology of microsomal P450 on the basis of the observed correlation time, , of 18 µ s. Calculations of the corresponding diffusion coefficient, D, using the expression =1/6D, suggest that six individual P450 units associate as a cluster on the microsomal membrane, and rotate about the six-fold axis of symmetry parallel to the plane of the membrane. In such an orientation, the N-terminal helical segments of each P450 unit in the hexamer could associate as a membrane-spanning agreggate which serves to anchor the P450 cluster in the phospholipid bilayer and, thus, explains the relatively slow rate of rotation of the hexameric P450 cluster. From a consideration of the known crystal structures of prokaryotic P450s, the overall shape of the enzyme, which resembles a triangular prism, would favour the formation of hexameric units on purely geometric grounds. In fact, the requirement of a 60° apical angle for such a formation is fulfilled in both P450cam and P450BM3. Consequently, the saturation-transfer ESR experiments provide some evidence for the retention of overall tertiary structure between prokaryotic and eukaryotic P450s. Supportive information from freeze-fracture scanning electron micrographs of intramembrane protein particles indicates that aggregation of P450 membrane-anchoring N-termini occurs, with an overall diameter of the helical bundle being between 3 and 5 nm (30–50 Å); which agrees favourably with the 3.5 nm (35 Å) estimate for the diameter of a parallel, symmetrical 6-helical structure (Schwarz et al., 1990). More recent studies on the rotation of an adrenocorticoid mitochrondrial P450, namely P45011A1, in proteoliposomes using time-resolved delayed fluorescence depolarization anisotropy (Schwarz et al., 1993) are consistent with the hexameric model of membrane-bound P450s embedded in the phospholipid bilayer to a depth of about 1 nm (10 Å) with an overall diameter of 11.4 nm (114 Å). Furthermore, di-iodofluorescein iodoacetamide labelling of cysteine residues in bovine P45011A1 has indicated a region of the structure which is exposed to the membrane


97

surface and accessible to adrenodoxin (Chernogolov et al., 1994). This appears to be a hinge region corresponding to a stretch of polypeptide between the H and I helices, which is longer than that found in the prokaryotic P450 crystal structures, and probably existing as a surface loop in eukaryotic P450s, that could play a role in some form of membrane interaction, and may also be associated with redox partner binding. Additional evidence of protein-lipid interactions in a microsomal P450 system has been provided by rotational diffusion studies on P4501A1 performed with and without the N-terminal 30 residue segment (Ohta et al., 1994). Using flash-induced absorption anisotropy of the P450-CO complex, the rotation of P4501A1 about an axis perpendular to the membrane showed a reduced relaxation time from 1101 µ s to 1020 µ s when the N-terminal peptide was removed. The results of these experiments suggest that, in addition to the N-terminal region, other segments of protein may also be involved in membrane binding. In fact, sequence alignment of eukaryotic P450s with the prokaryotic P450s of known structure does indeed show a number of peptide sections as likely candidates for interactions with membrane phospholipid (Lewis, 1995a). It is interesting to note that the emerging model for the membrane topology of eukaryotic P450s is consistent with the multicomponent cluster representation described by Anton Stier about 20 years ago (Stier, 1976). In this model, approximately six P450 monomeric units cluster around a central reductase molecule as a macromolecular assembly embedded within the microsomal membrane. In such a description there is a striking analogy with the bacteriorhodopsin structure generally accepted as a template for Gprotein-coupled receptors, which contains seven transmembrane helices. Presumably, the hexameric P450reductase complex may also comprise a total of seven helical segments able to span the microsomal membrane phospholipid bilayer. However, the complex probably has a dynamic nature, reflecting the fluidity (Denner and Kaiser, 1982; Funk et al., 1982) of the membrane matrix, where the P450 reductase is able to move and interact with individual P450 monomers in turn by an association which accompanies electron transfer. The situation in mitochondrial P450 systems is likely to be somewhat different from that of microsomal P450s, with the membrane-free adrenodoxin molecule acting as a mobile electron shuttle between its reductase and the mitochondrial P450 cluster (Lambeth, 1990). Nevertheless, the overall membrane topology (Brown and Black, 1989; Black, 1992; Uvarov et al., 1994) of eukaryotic P450s is likely to conform in general, with the model proposed by Nelson and Strobel (1988) on the basis of sequence alignment of over 30 P450s, which shows their essential similarity with the prokaryotic P450cam structure, and where the heme moiety may lie in an orientation parallel with the membrane surface. As the N-terminal anchor varies in length between different P450s, according to the position of tryptic cleavage, it is possible that the N-terminus could be composed of a helix-turn-helix motif (Nelson and Strobel, 1988) for sequences of about 40 residues in length, as opposed to a single N-terminal helix for shorter sequences of roughly 20 residues. It can be shown that a helix of about 20 amino acid residues long is able to span a phospholipid bilayer and, based on the hydrophobicity of P450 N-termini, it is likely that the preferred conformation of such sequences will be -helical. The possibility that part of the P450 N-terminal sequence may represent a signal peptide has been explored by site-directed mutagenesis (Szczesna-Skorupa et al., 1988) and an analysis of various membrane-protein topologies has been reported on the basis of membrane integration of modified microsomal P450 in a cell-free system (Szczesna-Skorupa and Kemper, 1991). Clearly, the phospholipid bilayer has an important function in eukaryotic P450s, as evidenced by its influence on substrate binding (Ebel et al., 1978; Sipal et al., 1979) where substrate lipophilicity plays a role in microsomal P450 binding (Al-Gailany et al., 1978). Apparently, the structural properties of phospholipid molecules are of relevance to modulation of the hemoprotein spin-state equilibria in microsomal P450 (Ruckpaul et al., 1982) and may also mediate conformational changes in P450 that are associated with

98


substrate binding interactions (Omata et al., 1986, 1987). Apparently, the competition between different P450s for reductase is dependent on the presence of substrate (Cawley et al., 1995) which suggests that the substrate-induced conformational change in P450 triggers the interaction with its redox partner. Furthermore, it has been shown that electron transfer between adrenodoxin and P45011A1 (CYP11A1) is facilitated by the binding characteristics of the membrane phospholipid, cardiolipin (Pember et al., 1983). In fact, the lipid-P450 interactions may be of a synergistic nature, as it has been reported that the presence of microsomal P450 increases the rate of phospholipid tranverse diffusion, commonly referred to as a ‘flipflop’ mechanism (Barsukov et al., 1982) which could have a relevance to substrate access to the heme site of membrane-bound P450s. The rate-determining process in the P450 catalytic cycle appears to be the second reduction (Imai et al., 1977). It is possible to show (Archakov and Bachmanova, 1990) that the velocity of this step is given by: reduction rate=KakET [P450] [Fpt] where: Ka=binding constant between the flavoprotein reductase (Fpt) of concentration [Fpt] and P450 of concentration [P450]; and, kET=rate of electron transfer between Fpt and P450. The values for the concentrations of P450 and reductase in microsomal preparations have been estimated as 4×10–3 M–1 and 2×10–4 M–1, respectively; whereas the association constant, Ka, has been recorded to be 0.05×10−6 M in negatively-charged vesicles and 0.5×10–6 M in neutral vesicles (Archakov and Bachmanova, 1990). The rates of P450 reduction by reductase are biphasic with rate constants of 2 s−1 (fast phase) and 0.3 s−1 (slow phase) in reconstituted systems whereas, in liver microsomes, values of 11.5 s−1 (no substrate) and 18 s−1 (+benzphetamine) have been recorded (Archakov and Bachmanova, 1990). Using these values, it is possible to calculate kET, the rate of electron transfer, to be between 2.875 and 4.5×1014 s −1. This is consistent with an electron jump of between 5 and 6 Å (based on k =h/m 2), which agrees ET e closely with the distance between FMN and heme propionate in flavocytochrome b2 at 5.659 Å, and the calculated wavelength of a 2 s electron (5.54 Å). However, an electron jump from FMN directly to the heme iron would require a change in the dielectric constant of the medium, and it is likely that the binding of substrate will facilitate this by desolvation of the heme environment. The fact that the values for the P450cam (bacterial) system are somewhat lower (3–4 s–1) than the microsomal P450 case (1–10 s−1) suggests that the role of phospholipid may be to facilitate electron transfer between redox components, whereas the broader range of values in the microsomal systems could reflect the variation in substrate hydrophobicity or their molecular volumes. Clearly, however, the relative concentrations of the various redox components, and their association constants, will play a major role in the overall rate of the reduction stages in the P450 cycle. 3.10 A proposed mechanistic scheme for the P450 catalytic cycle A consideration of the overall thermodynamics of the P450 catalytic cycle (Table 3.17) indicates that the reaction is energetically feasible provided that the required activation energy is overcome. The major component of this energy requirement is Table 3.17 Thermodynamic inventory of the P450 mono-oxygenase reaction (References: Ruckpaul and Rein, 1984; Ruckpaul et al., 1989) Chemical process H+

NADPH + O2(g) 2O(g)

H2+NADP

G (kJ mole–1) 19.3 460.5


Chemical process

99

G (kJ mole–1)

H2(g)+O(g) H2O(1) −470.1 RH(1) + O(g) ROH(1) −393.9 Overall G = −384.2 RH = a hydrocarbon substrate, e.g. cyclohexane Notes: The reaction is, therefore, thermodynamically favourable, but rates are generally quite slow (0.1–10 min–1 in mammalian P450s) due to the relatively high activation energy required for the reaction to proceed. Apart from the high dissociation energy of dioxygen, the other energetically unfavourable reaction is the cleavage of the C-H bond in the substrate (~418 kJ mole–1). Without the presence of an enzyme, the activation energy of the reaction can be between ~418–460 kJ mole–1 but, in the case of P450, this reduces to ~38–71 kJ mole– 1 which implies that the enzyme facilitates the cleavage of high energy bonds in both oxygen and substrate due to binding to the heme and protein, respectively. P450 achieves this by facilitating the fission of the dioxygen bond by the addition of two reducing equivalents to the heme-thiolate oxygen complex.

that represented by the bond energy of the dioxygen double bond (460.5 kJ mole–1). Consequently, the enzyme will need to reduce this high activation energy by lowering the O−O bond energy. Figure 3.7 shows the electronic states, orbital energies and other relevant bond data for various dioxygen species, such as superoxide and peroxide, where it can be appreciated that addition of two electrons sequentially to molecular oxygen both increases its reactivity and weakens the oxygen-oxygen bond. The energy requirement to produce superoxide, and then peroxide, is significantly less than that of an oxygen-oxygen double bond, and it is therefore feasible that P450 facilitates these conversions via the twostage input of electrons and utilization of the unique properties of the heme-thiolate combination in a protein environment which enables proton transfer and hydrogen-bond interactions between the dioxygen species and active site amino acid residues. It is, therefore, unnecessary for the enzyme to employ such a large amount of free energy to generate energetically unfavourable species, such as free oxygen atoms or heptavalent oxygen radicals, when more feasible reactive intermediates, like superoxide and peroxide, could readily perform the necessary reactions with substrates and have, furthermore, been observed in many P450 systems. Based on the accumulating evidence for an iron peroxy species as the active oxygenating species in P450 reactions, it is possible to propose a scheme (Figure 3.8) for the mechanistic cycle of a typical P450 monooxygenase reaction, which is consistent with some of the recently-emerging data on P450 catalysis (Pratt et al., 1995b; Lee-Robichaud et al., 1995; Akhtar et al., 1994; Kobayashi et al., 1994; Gerber and Sligar, 1994). In this scheme, ferric P450 predominantly in the low-spin state, with a coordinated water molecule in the resting state, will be converted to high-spin ferric on substrate binding, which is consistent with the displacement of active site water molecules including the distal ligand. The subsequent lowering of P450 redox potential and conformational change, which accompanies substrate binding, brings about the first reduction from a bound redox partner (i.e. reductase or a redoxin). The resultant high-spin ferrous P450 possesses a high affinity for oxygen which can then bind via a spin-pairing interaction, causing the iron to become low-spin. Although this intermediate can be regarded as an equilibrium between two canonical forms (FeIIO2 FeIIIO2–) the oxygen species does not appear able to oxygenate the substrate. However, the second reduction which forms a potentially more reactive entity (FeIIO2– FeIIIO22–) could either generate superoxide or peroxide, if uncoupling occurs, or give rise to oxygenated substrate and water provided that there is an available source of protons. The possibility of a distal charge relay with hydrogen-bonded interactions between various active site amino acid residues and, perhaps, intervening water molecules makes the production of water and

100


Note: Moleculardioxygen (2 O2) can be converted to singlet oxygen (1 O2) via the intermediate stage of superoxide (O2−·) but the direct route is spin-forbidden. Thus, if P450 convets triplet ground state dioxygen to superoxide, it provides a means of activating the oxygen molecule such that the singlet state O2 (or O2−/2) can be formed, which will then be able to react with organic substrates that are generally singlet. Molecular dioxygen will normally be unreactive towards organic substrates due to the fact that the reaction is spin-forbidden, as shown abvoe. Figure 3.7 Electronic states of oxygen species and related data (References: Hanzlik, 1976; Bonnett, 1981).

hydroxylated substrate feasible via either an iron hydroperoxy species or iron-coordinated hydrogen peroxide. The latter could also effect other types of P450 oxygenations, if one assumes that there will be a charge separation between the two oxygen atoms due to iron ligation, such that the peroxy species can act either as a nucleophile or electrophile. As there is a possibility of proton exchange with solvent water for either the hydroperoxide or hydrogen peroxide intermediate, deuterium isotope effects would be expected in accordance with the experimental observations. Following oxygenation of substrate, the other oxygen atom can readily ligate the ferric P450 as coordinated water, thus returning the enzyme to its original resting state. Although aliphatic hydroxylation is the example shown in Figure 3.8, it is also possible to modify the mechanism to accommodate aromatic hydroxylation, epoxidation and N- or S-oxygenations. Furthermore, this scheme can be readily adapted to show that a concerted mechanism is feasible, whereas the formation of the unusual high-valent or high-energy states (FevO2−, FeIVO− or FeIII


101

Notes:It is possbile that the iron peroxy intermediate is stabilized by hydrogen bonding with the conserved distal threonine residue, with likely cooperation from the adjacent conserved acidic residue for conducting protons from the environment to the active oxygen species. In P450cam, it is thought that cooperation between Thr 252 and Asp 251 chnnels protons into the heme vicinity, possibly via ion-pairing between Arg 186 and Asp 251, but other basic protonated residues in the heme pocket may also be involved (Gerber and Sligar, 1994). This mechanism could be varied to give rise to the iron oxene intermediate if desired, but it is not essential. For further details of iron-oxygen electronic states, the reader is referred to an ex cellent discussion by Harcourt (1977). For aromiatic and N/S oxygenation, the uncleophilic substrate attacks the iron peroxide species in an analogous fashion to the carbanion shown above. The hydroxide then acts as a base by removing a proton from the intermediate, thus forming the known metabolite and water. In this variant, two protons from the solvent and/or protein are required to from the active oxygen species thus allowing more scoope for protein control of the reaction. Figure 3.8 Mechanistic cycle for aliphatic hydroxylation (RCH3

RCH2OH).

4 P450 Substrate Specificity and Metabolism

4.1 Introduction Over 60 different varieties of enzymic reactions are known to be catalyzed by P450s, of which over 300 genetically distinct isoforms have now been sequenced (Nelson et al., 1993), that are capable of metabolizing hundreds of thousands of different chemicals (Porter and Coon, 1991). The classification system employed for P450s (Nelson et al., 1993) does not necessarily equate precisely with types of substrates metabolized, but there are some correlations which will be outlined in this chapter. As the means of classifying P450s is based on their amino acid sequence homologies, the types of reactions catalyzed do not form any obviously recognizable pattern with respect to the particular P450 isoforms, although there is a fair degree of discrimination in terms of substrate specificities. Although many aspects of P450 metabolism have been studied in rodents and other laboratory animals, there is an understandable interest in human P450s, which are currently being investigated to an everincreasing extent, especially as it is now possible to express human (and other) P450s in reconstituted systems using recombinant DNA technology (Chapter 5). This chapter will, therefore, focus primarily on those species which have been most extensively studied and where major interest exists, i.e. mammalia, especially rodents and man. Endogenous substrate metabolism, however, has also been investigated in other species, including unicellular organisms, and this will be considered first. 4.2 Endogenous metabolism Although cytochromes P450 of families CYP1, CYP2 and CYP3 are able to metabolize some endogenous chemicals, such as steroids, these families appear to be primarily associated with the metabolism of exogenous compounds. There is, however, increasing evidence for an endogenous role in some cases, for example: CYP1A2, CYP2A, CYP2D, CYP2E and CYP3A. Nevertheless, even where an endogenous function can be demonstrated, there is an overwhelming number of foreign com-pounds that are known to be metabolized by P450s of these three families (LaBella, 1991). In contrast, members of the other P450 families, especially those in mammalia, have been shown to possess specific metabolic functions. For example, enzymes of the CYP4 family are responsible for the end-chain hydroxylation of long-chain fatty acids, particularly lauric acid, whereas some CYP4 isozymes are associated with the -hydroxylation of certain prostaglandins and leukotrienes (Kupfer, 1980). The remaining P450 families present in mammalia

P450 SUBSTRATE SPECIFICITY AND METABOLISM

103

are involved in the biosynthesis of steroid hormones, and their nomenclature is derived from the various positions in the steroid nucleus where metabolism occurs (Nelson et al., 1993). Therefore, CYP7 mediates the hydroxylation of cholesterol at the 7 -position, whereas CYP11B1 brings about 11 -hydroxylation of progesterone. CYP17 and CYP21 catalyze the 17 -and 21-hydroxylations of progesterone, respectively, and CYP19 facilitates the aromatization of androgens to estrogens by the initial step of hydroxylation at the 19position (Chen et al., 1989). The particular P450s involved in endogenous metabolism, which have had their amino acid sequences determined, are listed in Table 4.1, together with their species and catalytic functions. In addition to the mammalian P450s mentioned above, this table summarizes those which have been found in other species such as insects, fungi, plants and bacteria. In some cases, xenobiotic metabolism has also been demonstrated in these non-mammalian P450s and the reader is referred to a number of relatively recent reviews for further information, namely, Schenkman (1992), Schenkman and Griem (1993), Fulco (1991) and various chapters in volumes 3, 4 and 6 of the book series Frontiers in Biotransformation, edited by Ruckpaul and Rein, and listed in the Bibliography. In mammalia, the cytochromes P450 involved in steroidogenesis are present in the adrenal cortex. These enzymes mediate the formation of pregnenolone, progesterone, corticosterone, testosterone, estradiol and other steroids, using cholesterol as a precursor. The various pathways and the P450s concerned are illustrated in Figure 4.1 where it can be seen that pathways involving CYP11 occur in the mitochondria, whereas those associated with CYP17, CYP19 and CYP21 take place on the endoplasmic reticular membrane (Takemori and Kominami, 1984; Gibson and Skett, 1994). It should be noted that some steroids may be metabolized in various positions by a variety of different P450s. For example, in a relatively recent review by Martucci and Fishman (1993), the species differences in estrogen metabo Table 4.1 Endogenous metabolism (References: Nelson et al., 1993; Schenkman and Griem, 1993) CYP family/ subfamily Species 4A1 4A2 4A3 4A4 4A5 4A6 4A7 4A8 4A9

Substrate metabolism

Rat (liver) Rat (kidney) Rat Rabbit (lung)

-hydroxylation of lauric acid -hydroxylation of lauric acid -hydroxylation of lauric acid -hydroxylation of PGE2 and other prostaglandins (El, A1, A2, D1 and F2 ) Rabbit kidney -hydroxylation of lauric acid Rabbit (kidney and liver) -1 hydroxylation of lauric acid Rabbit (kidney and liver) -2 hydroxylation of PGA Rat -hydroxylation of lauric acid Human -hydroxylation of lauric acid

CYP family/ subfamily

Species


4A10 4A11

Mouse Human (kidney)

4B1

Human, rat, rabbit (lung)

4C1

Cockroach

-hydroxylation of lauric acid - and -1 hydroxylation of lauric acid - and -1 hydroxylation of lauric acid Fatty acid hydroxylation

104



Species

4D1

Fruit fly


Unknown but probably lauric acid - and -1 hydroxylation 4E1 Fruit fly Unknown but probably lauric acid - and -1 hydroxylation 4F1 Rat -hydroxylation of lauric acid 4F2 Human Unknown 4F3 Human -hydroxylation of LTB4 5 Human Thromboxane synthase 6A1 House fly Unknown 6A2 Fruit fly Unknown 6B1 Butterfly Xanthotoxin (8-methoxypsoralen) Note: Insect forms of P450 (which have not been sequenced) are associated with steroid and juvenile hormone synthesis and metabolism of cholesterol to ecdysone, which can be further metabolized by hydroxylation at the 20, 2, 22 and 25 positions (Feyereisen, 1993), and of methyl farnesoate epoxidation to form the sesquiterpenoid juvenile hormone. Sex pheromone biosynthesis from monoterpenes may also be catalyzed by P450 enzymes, as well as the biosynthesis of defensive chemicals, in addition to fatty acid metabolism summarized above. CYP7 Human, rat, rabbit 7 -hydroxylation of cholesterol CYP11A1 Human, rat, etc. Side chain cleavage of cholesterol (C20-C22) to pregnenolone CYP11B1 Human, rat, etc. 11 -hydroxylation of progesterone CYP11B2 Human, rat, etc. Aldocorticoid synthase CYP 17 Human, rat, etc. 17 -hydroxylation of progesterone (and pregnenolone) CYP 19 Human, rat, etc. Androgen aromatization of androstenedione (to estrone) and testosterone (to estradiol) CYP21 Human, mouse 21-hydroxylation of pregesterone CYP27 Human, rat, rabbit 25-, 26-, 27-hydroxylation of cholesterol metabolites, bile acids, vitamin D 25-hydroxylase CYP51 Yeasts Lanosterol 14 -demethylation CYP52 Yeasts Alkane -hydroxylation Fatty acid -hydroxylation CYP53 Fungi Benzoic acid p-hydroxylation CYP54 Fungi Unknown CYP55 Fungi Nitrate/nitrite reductase CYP56 Yeast Dityrosine synthase CYP57 Fungi Pisatin demethylation (phytoalexin) CYP71 Avocado Unknown CYP72 Periwinkle Geraniol 10-hydroxylation CYP73 Artichoke Cinnamic acid 4-hydroxylation



Species

105


Note: Fungal and plant forms not sequenced also display other catalytic activities of relevance to endogenous and, possibly, exogenous metabolism. CYP family /subfamily

Species


CYP101 CYP102 CYP103 CYP104 CYP105

Pseudomonas putida Bacillus megaterium Agrobacterium tumefaciens A. tumefaciens Streptomyces griseolus

Camphor 5-exo hydroxylation -2 fatty acid hydroxylation Unknown Unknown Sulfomethuron* and other sulphonylurea methyl hydroxylation Unknown 6-deoxyerythronolide hydroxylation -terpineol 4-methyl hydroxylation Unknown Unknown Linalool 8-methyl hydroxylation Unknown

CYP106 B. megaterium CYP107 Saccharopolyspora erythrea CYP108 Pseudomonas putida (spp.) CYP109 Bacillus subtilis CYP110 Anabaena spp. CYP111 P. putida (incognita) CYP112 Bradyrhizobium japonicum * not an endogenous substrate (sulphonylurea herbicide) Note: In addition to the activities shown above, there are many other catalytic reactions of bacterial P450s not sequenced to date, which are associated with the metabolism of other substrates.

lism by P450s is described, together with the pathophysiological roles of estrogens and their metabolites. Although both mitochondrial and endoplasmic reticular adrenocorticoid P450s are membrane-bound, the N-terminal sequences of the former contain generally more polar amino acid residues (Omura, 1993) whereas the latter have highly hydrophobic N-termini resembling those of the hepatic microsomal P450s, such as CYP2B (Takemori and Kominami, 1984). As mentioned previously, the electron transfer systems are also different: the mitochondrial system utilizes an iron-sulphur protein, adrenodoxin, which ‘shuttles’ between its FAD-dependent NADPH-adrenodoxin reductase; whereas the endoplasmic reticulum system requires NADPH-cytochrome P450 reductase for electron transfer which, as in hepatic microsomes, contains both FMN and FAD moieties for electron transfer. It is interesting to note that, in some cases, steroidal substrates which are intermediate stages in these biosynthetic pathways for the production of corticoids and andogens, are required to move from the mitochondria to the endoplasmic reticulum and vice versa. There are, almost certainly, important regulatory control rationales under-lying such steroidogenic systems (Zimniak and Waxman, 1993). In addition to the hydroxylations which lead to steroid hormones, cytochromes P450 also catalyze the formation of bile acids from cholesterol. These conversions, however, occur in the liver and involve 7 hydroxylation (CYP7), 27-hydroxylation (CYP27) and 12 -hydroxylation (CYP12?) forming, eventually, cholic acid and chenodeoxycholic acid (Okuda et al., 1993). In addition to being a biosynthetic pathway for a group of functionally important steroids, these P450-mediated reactions also represent the most significant route of cholesterol degradation and elimination in mammalia. Furthermore, hydroxylation of vitamin D3 and some of its hydroxy derivatives are also catalyzed by CYP27, although these hydroxylations appear to occur at positions 25 and 26 on the steroid-like nucleus (Hollis and Gray, 1993). Apparently, this enzyme (or

106


Figure 4.1 Steroid hormone biosynthetic pathways in the adrenal cortex.

isoforms of it) is able to mediate in hydroxylations at analogous positions in certain C27 steroids; vitamin D3 hydroxylase is present in both hepatic and renal mitochondria, whereas the steroid 27-hydroxylase (sometimes referred to as 26-hydroxylase) is a microsomal form. Presumably, amino acid sequence


107

comparison between these isozymes will lead to a clarification of the apparent ambiguity regarding the classification of both mitochondrial and microsomal forms to the same CYP family. 4.3 Endogenous steroid hydroxylation by hepatic P450s of families CYP1, CYP2 and CYP3 Endogenous steroids, such as testosterone, are hydroxylated in various positions by liver microsomal P450s normally associated with xenobiotic metabolism (Schenkman, 1992). There is some speculation regarding any possible physiological significance in these findings as they are relatively minor, occur in positions generally distinct from those relating to biosynthesis, and may be incidental to the primary functions of those P450s (Zimniak and Waxman, 1993). However, it is thought that isoenzymes of the CYP3 family are evolutionarily distant relative to those of the CYP2 family, which could have evolved as a response to plant toxins; whereas there is evidence for an endogenous role for CYP3 in the 6 -hydroxylation of the bile acid, lithocholic acid (Zimniak and Waxman, 1993). It is possible, based on the specificity of various members of the CYP2 family for steroid hydroxylation, that some of these enzymes have been latterly adapted for steroid metabolism, following their original exogenous roles (Zimniak and Waxman, 1993). One of the recurrent features of the P450s is their apparently simple genetic modification to alter substrate specificity, and this may explain their ubiquity in living systems for the metabolism of a multitude of structurallydiverse chemicals. Taking, as an example, the metabolism of testosterone by rat liver microsomal P450s, it can be appreciated that enzymes of families CYP1, CYP2 and CYP3 all catalyze hydroxylations in a number of different positions (Schenkman, 1992; Waxman, 1988; Funae and Imaoka, 1993). These are summarized in Table 4.2, which also provides additional information regarding nomenclature and induction of these isoenzymes. Table 4.3 gives an indication of the control levels of constitutive and inducible P450s in rat liver, together with their relative inducibility by typical inducing agents. The corresponding levels of human orthologues are to some extent different from those found in the rat, and are subject to considerable variation due to lifestyle and pathophysiological states (George and Farrell, 1991; Watkins, 1990, 1992a and b; Wolff and Strecker, 1992; Renton and Knickle, 1990), thus making comparisons somewhat difficult. However, it appears that the CYP3 family accounts for up to 60 per cent of the human liver P450 complement (Gonzalez and Gelboin, 1994), with much lower levels for CYP2B (0–2 per cent) and CYP2A ( 1 per cent), and some variability in CYP1A2, CYP2C, CYP2D and CYP2E; the levels of which are subject to, for example, the individual’s medication, disease state, age, genetic polymorphism (CYP2D6 and CYP2C19) and alcohol intake, diabetes, and fasting (CYP2E). CYP1A1 is not expressed in human liver but is present extrahepatically, such as in the lung: in contrast CYP1A2 has only been detected significantly in the Table 4.2 Rat hepatic P450s, nomenclature, inducers, testosterone metabolism and sex differences (References: Ryan and Levin, 1990; Schenkman et al., 1989; Nelson et al., 1993; Soucek and Gut, 1992; Schenkman and Griem, 1993; Omura et al., 1993) Nomenclature CYP

Guengerich Funae

Ryan

Inducer

I/C Site of testosterone metabolism

Sex specificity

1A1 1A2

BNF-B ISF-G

P450C P450d

TCDD, BNF 3MC, ANF

I C

None Female dominant

MC-5 MC-1

6 6

108


Nomenclature CYP

Guengerich Funae

Ryan

Inducer

I/C Site of testosterone metabolism

2A1 UT-F IF-3 P450a PCB, 3MC C 7 *, 6 2A2 — UT-4 P450a2 — C 15 *, 6 , 7 , 16 , (7 ) 2B1 PB-B PB-4 P450b PB I 16 , 17, 16 2B2 PB-D PB-5 P450e PB C 16 , 16 2C6 PB-C PB-2 P450k PB C 2 , 16 , 17 2C7 — UT-16 P450f ETOH C 16 2C11 UT-A UT-2 P450h — C 16 *, 2 *, 17, 6 2C12 UT-I IF-2 P450i — C 15 *, (15 ) 2C13 — UT-5 P450g — C 6 , 15 , 16 , 7 2D1 UT-H UT-7 (P450db1) — C 6 2E1 — DM P450j ETOH, ACE C — 3A1 PCN-B — P450p PCN I 6 * 3A2 PCN-E PB-1 (P450pcn2) PCN C 6 ,2 4A1 — — (P450la ) CLOF C — I=Inducible, C=Constitutive; TCDD =2, 3, 7, 8-tetrachlorodibenzo-p-dioxin; BNF- -naphthoflavone; ANF= -naphtoflavone; 3MC=3-methylcholanthrene; PCB=polychlorinated biphenyls; PB=phenobarbital; ETOH=ethanol; ACE=acetone; PCN=pregnenolone 16 -carbonitrile; CLOF=clofibrate. * Almost all (over 85 per cent) of the indicated metabolite is produced by this form. Minor metabolites of testosterone produced are shown in parentheses.

Sex specificity Female dominant Male Male dominant Male dominant None Female dominant Male Female Male Male dominant Female dominant Male dominant Male None

Table 4.3 Sex differences and control levels of rat liver P450s (Reference: Schenkman and Griem, 1993) CYP

Sex specificity

Inducibility

Control levels*

%†

2C11 3A2 2A2 2C13 2C22 2B1 2B2 3A1 2D1 2C12 2A1 2C7 2E1 1A2 2C6 1A1

Male specific Male specific Male specific Male specific Male specific Male dominant Male dominant Male dominant Male dominant Female specific Female dominant Female dominant Female dominant Female dominant No sex difference No sex difference

Non-inducible 3–fold induction by PB Non-inducible Non-inducible Non-inducible 40–fold induction by PB 12–fold induction by PB 12–fold induction by PB Non-inducible Non-inducible 2–fold induction by PB Non-inducible 3–fold induction by ACE 20–fold induction by 3MC, BNF 2–fold induction by PB 50–fold induction by 3MC, BNF

0.3–1.2 0.088–0.39 0.01–0.026 0.086–0.171 not known 0.001–0.08 0.004–0.07 < 0.015 not known 20 k 2C2 Rabbit 9 ~20k 11B1 Human 9 ~4.8 k 2C3 Rabbit 9 >25 k 11B2 Human 9 ~5.3 k 2C5 Rabbit 9 >20k 17A Bovine 8 ~6.8 k 2C11 Rat 9 ~35k 17A Human 8 6659 2C12 Rat 9 >35k 19A Human 10 >70 k 2D2 Rat 9 4036 21A Bovine 10 3457 2D3 Rat 9 4371 21A2 Human 10 ~3230 2D4 Rat 9 4678 21a1 Mouse 10 ~3060 2D5 Rat 9 4567 * The number of introns will be (obviously) one less than the number of exons and these tend to be of variable length from one gene family to another but there are overall pattern similarities, i.e. the gene organization is generally conserved. Table 5.4 Chromosomal locations of mammalian P450 genes (Reference: Omura et al., 1993, Matsunaga et al., 1990b) Human

Mouse

CYP

Chromosome

1A1 1A2 2A 2B 2C 2D 2E

15q22-q24 15 19q13.1–13.2 19q13.1–13.2 10q24. 1–24.3 22q11.2-qter 10

CYP

Chromosome

1a2 2a

9R 7

2c 2d 2e

19 15 7

INDUCTION, REGULATION AND INHIBITION

Human CYP

159

Mouse Chromosome

2F 19q13.1–13.2 3A 7q21.3–q22.1 4B 1p12-p34 11A 15q23–24 11B 8q21–22 17A 10 19 15q21.1–21.3 21 6p21.3 27 2q33-qter P450 gene clusters on different human and mouse chromosomes Human P450 genes Chromosome 15 1A1, 1A2, 11A and 19 Chromosome 19 2A, 2B and 2F Chromosome 6 21A Chromosome 22 2D Chromosome 10 2C, 2E and 17A

CYP

Chromosome

3 4a 11a

6 4 9

19 21 27

9 17 1

Mouse Chromosome 9 Chromosome 7 (2A and 2E) Chromosome 17 Chromosome 15 —

Figure 5.1 Stages in P450 gene expression.

Transcription of the P450 gene gives rise to a specific mRNA which requires further processing to remove (i.e. splice) the non-coding regions (introns) to produce an active mRNA capable of translation to form the appropriate P450 protein, as shown in Figure 5.1. The possibility of alternative initiation and splicing sites in P450 genes giving rise to different mRNAs in particular cell types may account for the altered levels of P450 proteins in different tissues and organs as the mRNAs in various cells possess the same coding sequence but different untranslated 5’ leaders such that initiation occurs at altered rates; indeed, similar situations operate in the Table 5.5 Mechanisms of P450 induction (References: Porter and Coon, 1991; Gibson and Skett, 1994) Induction mechanism P450s known to be induced/regulated Gene transcription mRNA processing mRNA stabilization

1A1, 1A2; 2B1, 2B2; 2C7, 2C11, 2C12; 2D9; 2E1; 2H1, 2H2; 3A1, 3A2, 3A6; 4A1; 11A1, 11B1; 17A1;21A1 1A2 1A1; 2B1, 2B2; 2C12; 2E1; 2H1, 2H2; 3A1, 3A2, 3A6; 11A1

160


Induction mechanism P450s known to be induced/regulated Translation 2E1 Enzyme stabilization 2E1; 3A1, 3A2, 3A6

Figure 5.2 Simplified structure of a P450 gene.

organ/tissue specific expression of other proteins (Breitbart et al., 1987). The different mechanisms of enzyme induction can operate at any of the various stages in the expression and regulation of the gene, as indicated in the scheme presented in Figure 5.1. It is thought that P450 inducers may act at any one of these stages, as shown in Table 5.5. However, it appears that some induction mechanisms (e.g. transcriptional activation) are more common than others (Waxman and Azaroff, 1992). There are a number of different regulatory elements on the P450 gene which will modulate expression of the protein via transcriptional activation or repression, depending on the function of that element and the nature of the binding factor (Omura et al., 1993). The arrangement of these various elements is shown schematically in Figure 5.2, but there will be significant variation from one P450 to another, and the binding of promotors or repressers will depend on several factors, including cell type, sex, age (Horbach et al., 1992) and developmental status of the organism (Hines et al., 1994). This means that the activity of P450s expressed in heterologous systems may not necessarily equate with the situation prevailing in a specific organ of a particular animal. The binding of some positive or negative regulatory factors may be able to alter the geometry of that section of DNA, i.e. the response element, which could increase the curvature of the DNA and, consequently, lead to the interaction between normally distant segments (Hines et al., 1994). This model (Figure 5.3) would explain how binding to one section of the 5’ -flanking region might modulate effects elsewhere on the regulatory portion of the gene (Nebert, 1989b) and there is more recent evidence showing that this occurs in other systems (Carey, 1994; Kerppola and Curran, 1995; Nordheim, 1994; Ptashne and Gann, 1990). Although these various regulatory elements generally lie upstream of the transcription start site, it is also known that response elements can be located within the first intron (Hines et al., 1994). It is possible, therefore, that regulation away from the 5’-flanking region could be associated with P450 induction involving mRNA stabilization because such introns will be transcribed prior to further processing of the mRNA, which involves excision of these segments. There are clearly many regulatory factors and response elements with which they interact, that can have either an activating or repressing influence. For example, the binding of TCDD to the cytosolic Ah receptor leads to transcriptional activation of the CYP1A gene, following translocation to the nucleus via an association with the Arnt factor (Reyes et al., 1992; Burbach et al., 1992; Hankinson, 1995). Furthermore, other xenobiotic (XRE) and glucocorticoid/estrogen (GRE, ERE) response elements exist in different P450 genes (Kemper, 1993) which could be related to induction by such agents. There is also the possibility of PB-type induction occurring via ligand binding to a repressor (Waxman and Azaroff, 1992) or some other protein which is subsequently released; and this process is termed genomal depression. The discovery of several response elements on the same gene thus explains the possible induction of the relevant P450 by different inducing agents, and the effect of promoters on the basal transcription element (BTE). Cytokines, such as the interleukins, appear to be able to modulate (Clark et al., 1995) the levels of P450s (as well as


161

Figure 5.3 A model for receptor-mediated transcriptional activation (Adapted from Nebert, 1989b).

those of other proteins) and, although the mechanisms are not well understood, it is possible that these proteins may be mediating interactions between upstream regulatory elements. In addition to the aforementioned factors, the levels of P450s will be affected by genetic polymorphism and allelic variants within the coding regions of the gene although, sometimes, mutations can occur in the non-coding regions (Heim and Meyer, 1991). This can result in poorly-expressed P450s or those showing impaired catalytic activity due to the loss of a particular function, such as the ability to bind heme effectively, or from alternative splicing (Ding et al., 1995). In some cases, even an entire P450 gene can be

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deleted from a gene locus (Meyer, 1991). Genetic polymorphism has been extensively studied in the CYP2D subfamily (Mikus et al., 1994) but is also known to exist in the CYP2C subfamily (Price-Evans, 1993), and in the CYP1 family (Gonzalez and Gelboin, 1994). It is likely that further evidence of polymorphism in other human (and animal) P450 genes will be discovered in due course. Induction has an effect on, and appears to be related to, the turnover rates of P450s (Correia, 1993). It is thought that the binding of some substrates to the P450 protein can increase the half-life of the enzyme itself by blocking degradation via the protein kinase-mediated phosphorylation pathway, which utilizes cAMP (Jansson, 1993). Judging by the structural differences between inducing agents of P450s, it would appear that the mechanisms of induction of P450s vary, although there are also some similarities, as has been noted previously (Table 5.5). For example, CYP1 inducers (Table 5.6) all bind to the Ah receptor and possess significant similarity in their molecular structures, namely, relative planarity coupled with a certain degree of rectangularity, exemplified by that of TCDD (Landers and Bunce, 1992). Such similarity is not as apparent in CYP2B inducers (Table 5.7), i.e. PB-type inducing agents (Waxman and Azaroff, 1992; Nims et al., 1993), although most of these agents possess non-planar geometries and V-shaped molecular conformations, sometimes also having an electronegative atom (such as oxygen or nitrogen) in a Table 5.6 Inducers of CYP1A isozymes (References: Okey, 1990; Gibson and Skett, 1994; Liu et al., 1993b) Polyaromatic hydrocarbons (e.g. 3MC, BP, BA and DBA) Phenothiazines -Naphthoflavone (and other flavones) Plant indoles (e.g. indole-3-carbinol and indole-3-acetonitrile) Indolocarbazoles Ellipticine TCDD (and other halogenated dibenzo-p-dioxins) Halogenated dibenzofurans Polychlorobiphenyls (specifically those with planar molecules) Polybromobiphenyls (specifically those with planar molecules) 2-Acetylaminofluorene Cigarette smoke Crude petroleum Benzocoumarins Key: 3MC=3-methylcholanthrene BP=benzo(a)pyrene BA=benz(a)anthracene DBA=dibenz(a, h)anthracene TCDD=2, 3, 7, 8-tetrachlorodibenzo-p-dioxin Table 5.7 Inducers of CYP2B isozymes (References: Okey, 1990; Gibson and Skett, 1994) Phenobarbital (and related barbiturates) Phenytoin (and other hydantoins) DDT l, 4-Bis[2-(3, 5-dichloropyridyloxy)]benzene Octachlorostyrene


163

trans-Stilbene oxide Pentamethylbenzene Polychlorobiphenyls (with non-planar molecules) Dieldrin DDT=p, p-Dichlorodiphenyltrichloroethane

similar position in the molecule, which is only apparent when the structures are superimposed in threedimensions (Lewis et al., 1987), especially with an electron-density coded van der Waals surface. CYP2E inducers (Table 5.8) are characterized by relatively small, structurally-diverse, molecules (Lewis et al., 1994a) and some of these may be able to stabilize the protein against degradation by binding to the serine residue normally associated with phosphorylation, i.e. a post-translational modification. Such agents would require at least one electronegative atom in order to hydrogen bond with serine and, therefore, ethanol, acetone and diethyl ether represent typical examples of CYP2E inducers. Chemicals which are able to induce CYP3A (Table 5.9) include endogenous and synthetic steroids (Okey, 1990; Waxman et al., 1985), leading to the view Table 5.8 Inducers of CYP2E isozymes (References: Okey, 1990; Gibson and Skett, 1994) Ethanol Benzene Isopropanol Imidazole Acetone Diethylether

Trichloroethane Pyrazole Isoniazid Diabetic state Fasting state

Table 5.9 Inducers of CYP3A isozymes (Reference: Okey, 1990) Pregnenolone 16 -carbonitrile Dexamethasone Spironolactone Erythromycin Rifampicin Triacetyloleandomycin Clotrimazole (also an inhibitor) Ketoconazole (also an inhibitor) Endogenous glucocorticoids (e.g. cortisone, hydrocortisone) Phenobarbital

that a DNA-binding steroid hormone receptor, such as the glucocorticoid receptor, may mediate induction of CYP3, although there is also evidence for the role of a microsomal protein (Wright and Paine, 1994). As steroid receptor response elements (GRE and ERE) are present in the upstream 5’ -flanking region of the CYP3 genes (Kemper, 1993), there would appear to be some evidence to support the hypothesis of steroid receptor mediation. However, there is stronger evidence for the involvement of another receptor protein of the steroid hormone superfamily in the induction of CYP4.

164


Induction of CYP4 (Table 5.10 for typical inducers) appears to be associated with peroxisomal proliferation (Gibson, 1992a; Hard wick, 1991), which is a well-characterized mechanism for carcinogenesis in rodents (Lake, 1995). A receptor protein, which is recognized as being a member of the steroid hormone receptor superfamily, and termed the peroxisome proliferator-activated receptor (ppar), has been identified (Issemann and Green, 1990) as a mediator of both CYP4 induction and peroxisome proliferation; and it is likely that ppar-binding response elements, which regulate the two events, are present in the regulatory regions of the relevant genes, i.e. those of the CYP4 proteins (Aldridge et al., 1995; Kimura et al., 1989) and peroxisome proliferation enzymes. It is thought, however, that peroxisome proliferators could actually displace an endogenous ligand from the ppar, rather than bind to it themselves, in order to effect peroxisome proliferating activity and CYP4 induction (Lake, 1995). This would, therefore, constitute a different mechanism for induction than that of CYP1, for example, where it is known that the binding affinity of ligands to the Ah receptor correlates closely with induction of CYP1 (Denomme et al., 1985; Golas et al., 1990). It is known that induction of P450s generally increases their half-lives (Correia, 1991) as shown in Table 5.11. The turnover of proteins is dependent on the nature of the N-terminal amino acid to some extent and, methionine which is a common N-terminal residue in most P450s, appears to be one example which gives rise to longer protein half-lives. However, it has been established that a cAMP-regulated phosphorylation-dependent mechanism exists for P450 turnover which involves conversion to P420, followed by denaturation, loss of heme and subsequent degradation of the protein (Jansson et al., 1990). Clearly, there are a number of different modes of P450 induction, the precise molecular mechanisms of which have yet to be fully elucidated and understood. It is possible, moreover, for more than one mecha Table 5.10 Inducers of CYP4A isozymes (References: Okey, 1990; Gibson and Skett, 1994; Lake and Lewis, 1996; Lake, 1995) Clofibrate Ciprofibrate Methyl clofenapate Nafenopin DEHP (Di-2-ethylhexyl phthalate) MEHP (Mono-2-ethylhexyl phthalate) Other peroxisome proliferators DHEA (dihydroepiandrosterone) Table 5.11 Half-lives of P450s (t in h) (Reference: Correia, 1991) CYP

Basal

Induced

Inducer

1A1 1A2 2A1 2B1, 2 2C6 2C11 3A1 3A2

15 10 14 19 20 20 – 12

20, 38 12, 22 20, 21 37, 20 35 32 14 27, 15

PB, arochor 1254 PB, NF PB, NF PB, NF PB PB DEX PB, NF


CYP PB=phenobarbital NF= -naphthoflavone DEX=dexamethasone Half-lives of proteins (t½ in h) Cytochrome b5 Catalase CYP2C11 Cytochrome P450 reductase Epoxide hydrolase

Basal

Induced

165

Inducer

84 60 20 29 19

nism to operate simultaneously, even for the same P450 and inducing agent, although it is usual for one type of inducer to manifest a particular mode of induction. 5.3 Regulatory elements in P450 genes The regulatory regions of P450 genes (Figure 5.2) contain both similarities and differences which, in theory, should explain the variations in regulation between P450s in different species, sexes and tissues. For example, the steroidogenic P450 genes (C7P11A, 11B, 17, 19 and 21) all possess response elements (CREs) in their regulatory regions which are associated with cAMP-mediated regulation via a specific binding protein (Kemper, 1993), and it is known that all five of these P450s have their levels regulated by cAMP. The 5 flanking regions of all P450 genes possess a TATA box, usually between 20 and 40 base pairs (bp) upstream of the transcription initiation site, with a basal transcription element (BTE) about 20 or more base pairs upstream of the TATA box. The BTE region normally contains a CAAT box (for liver-specific P450s) and a GC box, whereas a specific 17 bp sequence (commonly referred to as a ‘Barbie’ box) is present in the BTE region of both CYP2B1 and CYP2E2 genes, together with those of CYP102 and CYP106, all of which are inducible by phenobarbital (Padmanaban and Nirodi, 1994; Fulco, 1991). Tissue-specific elements, such as those thought to bind hepatic nuclear factors, can occur between 40 and 200 bp upstream of the RNA transcription site but are common at between −80 and −120 bp of the transcription start site (Kemper, 1993). Further upstream of the tissue specific factor regulatory elements, but occasionally overlapping with them, are frequently found a number of glucocorticoid response elements (GREs), which could explain the fact that several P450s are induced by glucocorticoids. In the C7P1A1 gene, however, these GREs lie within the first intron (Hines et al., 1994) and, unlike other P450s, the first exon is not transcribed in CYP1A. The region between the promoter site (BTE and TATA) and about -200 bp can also contain CREs, in steroidogenic P450s or xenobiotic response elements (XREs) for P450s of families 2 and 3, although the CYP1 genes have their XREs between roughly–500 and–3500 bp of the initiation site (Hines et al., 1994). The position and number of these response elements and other regulatory elements can vary from one P450 family (or subfamily) to another; sometimes there is even a variation between genes in the same subfamily, and this could relate to different levels of expression moderated by sex, species and tissue type. Probably the most extensively-studied regulatory region is that of the CYP1A1 gene (Kemper, 1993). Apparently, there are a total of five XREs (at -3586, -1233, -1089, -1010 and -537 bp in the rat CYP1A1), of which the two XREs between -1000 and -1100 bp probably constitute the preferred Ah receptor binding site. All five XREs contain a core sequence CACGC as part of a 17 bp homologous sequence (Omura et al.,

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1993; Hines et al., 1994). Nebert has proposed a model for the regulation of CYP1A1 which involves a curved conformational change in the regulatory region that occurs on Ah receptor binding, thus bringing together linearly distant elements (Nebert, 1989b). It is known that the binding of some receptor proteins to DNA can induce a conformational change which produces a curve in the DNA, although it is known that the crystal structures of synthetic poly-GC DNAs naturally exhibit a distinct curvature in their overall conformation. Transcriptional activation of CYP1A1 via the ligand-bound Ah receptor has been well established (Nebert, 1989a; Nebert and Gonzalez, 1985; Nebert et al., 1990; Lin et al., 1991; FujiiKuriyama et al., 1989b). It appears that the binding of a potent inducer, such as TCDD, displaces the 90 kDa heat shock protein (HSP90) from the cytosolic Ah receptor which can then bind another protein, known as the Ah receptor-nuclear translocation factor (Arnt), that facilitates translocation of the complex to the nucleus (Hankinson, 1995), whereupon binding to the regulatory region of CYP1A occurs and, possibly, preferentially at the XRE1 and XRE2 sites (Omura et al., 1993). The subsequent conformational change in the DNA, which accompanies binding of the heterodimer Ah/ Arnt complex, probably leads to partial unwinding (Richard-Foy, 1994) of the double helix, thus exposing the DNA bases for transcription by RNA polymerase II at the initiation site, following release of the represser protein (Hankinson, 1995) bound to the basal transcription element (BTE). For this to occur, it requires significant curvature of the 5’ -flanking region to trigger release of the BTE-binding protein (BTEB), although it is possible that any conformational change, which occurs on binding of the heterodimeric enhancement factor to the XRE, could be transmitted downstream to the BTE (-43 bp), and there is recent evidence for electron conduction in DNA, although the possible role of this is as yet unknown. The BTEB sequence indicates that the potential DNA-binding motif is likely to be of the ‘zinc-finger’ type (Fujii-Kuriyama et al., 1992). This helix-turn-helix structure is a common DNA-binding motif found in other proteins which interact with DNA, as the helices are able to fit into the major groove of the double helix. It is also possible to predict the likelihood that a section of DNA forms part of a major groove and, for example, the 17 bp consensus sequence representing the XRE contains a common core (CAGCG) which is thought to lie in a major groove of the double helix (Kemper, 1993). Presumably, ligand binding to a specific receptor (e.g. the Ah receptor) modifies the conformation of the protein such that a favourable DNA interaction is likely between the response element and DNA-binding region. Although the DNA-binding motifs and ligand-binding domains on such receptors are sequentially distant, it is likely that ligand-induced folding of the polypeptide chain will bring these into relatively close proximity, as has been postulated for the human estrogen receptor (Lewis et al., 1995e). Apparently, there are some differences between the regulatory regions of 1A1 and 1A2 genes, which may relate to the known variations in expression of the two P450s (Kemper, 1993). For example, there are more HNF-REs in the 1A2 gene than are present in that of 1A1 (which only contains one) and, conversely, 1A2 possesses only one AhRE in its 5’ -flanking region, whereas 1A1 has five (Hines et al., 1994). This would imply that 1A2 should be more readily expressed in liver than 1A1, which is in fact true; and 1A1 would be expected to be more easily inducible by 1A inducers than 1A2, and this is also known to be the actual situation found in vivo (Gonzalez and Gelboin, 1994; Omura et al., 1993). Consequently, it would appear that there is much useful information to be gleaned from an analysis of the regulatory regions of P450 genes. For example, a particular consensus nucleotide sequence has been noted for both 2B1/2 and CYP102 regulatory regions, all of which are inducible by phenobarbital (Kemper, 1993). In the 2B1 and 2B2 genes, this so-called ‘Barbie’-box motif is close to the BTE, as it is slightly upstream of this element, and there may be some form of coordinate regulation mechanism operating via these associated regions (Padmanaban and Nirodi, 1994).


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The possible mechanisms of induction of the bacterial P450, CYP102, by phenobarbital and structurallyrelated compounds, have been studied by Fulco and coworkers (Fulco, 1991; Fulco and Ruettinger, 1987; Ruettinger et al., 1989) who have shown that the presence of both positive and negative regulatory elements within the same gene are likely to modulate expression of the protein. This may explain why it is found that many P450 inducers ultimately inhibit enzyme activity at sufficiently high dose (Okey, 1990). Furthermore, the finding that the appropriate response elements are present in the regulatory regions of other inducible P450s appears to rationalize the known inducing agents for CYP3A and CYP4A, for example. In fact, two peroxisome proliferator receptor response elements (PPREs) have been shown to be present at about -4300 bp upstream of the transcriptional initiation site on the 4A1 gene (Aldridge et al., 1995), whereas there are several steroid hormone receptor response elements (three EREs, one PRE and GRE) between about -200 bp and about -500 bp (and flanked by HNF-REs) on the CYP3A4 gene (Hashimoto et al., 1993). It is also found that coordinate regulation of all four steroidogenic P450 genes present in the adrenal gland is associated with cAMP via specific response elements (CREs) which are able to bind a receptor protein, known as the cAMP responsive element binding protein (CREB), and there is evidence for the involvement of PKC in cAMP-mediated induction of these P450s (Kemper, 1993). Although the induction of P450 enzyme activity is generally regarded as involving an increase in the concentration of that particular P450, it should be noted that the mRNA levels of the relevant P450 are also known to increase, following treatment by an inducing agent, in a time-dependent manner which parallels the increase in P450 protein levels, as has been shown in 3A induction by rifampicin, for example (Potenza et al., 1989). However, it is found that, in general, mRNA levels remain fairly elevated for some time after induction as the inducing agent often stabilizes the mRNA from degradation. Consequently, there is often a link between these two mechanisms of induction, namely, transcriptional activation and mRNA stabilization, as can be appreciated from Table 5.5. 5.4 The toxic consequences of induction If the inducing agent is a poor substrate, (e.g. TCDD or clofibrate) but a potent inducer of a P450, there may be toxic consequences and/or down-regulation of other P450s. For example, there could be DNA damage resulting from either reactive metabolites (e.g. benzo(a)pyrene-7,8-diol-9,10-epoxide) or from the generation of other reactive intermediates, such as ROS (e.g. O2–., OH, 1O2, H2O2, etc.) which can cause miscoding, mutagenesis and carcinogenesis (Kappus, 1993). Induction may also alter the promotional stages by giving rise to genomal derepression, and lead to poor DNA repair (Parke and loannides, 1990a; loannides and Parke, 1993). Repeated dosage of an inducer can overwhelm DNA repair mechanisms and the natural defence systems (Parke and loannides, 1994) against ROS (i.e. SOD, catalase and GSH), and the reactive intermediates may be sufficiently long-lived to cause irreversible DNA damage, especially if they are poor substrates for the detoxifying enzymes (loannides et al., 1995); for example, benzo(a)pyrene-7,8diol-9,10-epoxide is not a good substrate of epoxide hydrolase (EH). Furthermore, the reactive species may be generated in a localized tissue; for example, -lyase cleavage of haloalkene cysteine conjugates occurs in the kidney, giving rise to nephrotoxicity and carcinogenicity (Dekant et al., 1989, 1990; Koob and Dekant, 1991). This may explain the finding that some agents only cause cancer in certain organs, but variation in the regulation of P450s depending on the tissue type could also be important. For example, Table 5.12 shows that CYP1 can be induced in various human cells and tissues, whereas extra-hepatic induction of CYP1 and CYP2B in rodents is summarized in Table 5.13.

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There are many examples where the level of induction is related to the degree of toxic effect produced, and the measured degree of P450 enzyme induction in experimental animals forms the basis of a procedure known as ENACT (loannides et al., 1995) which was developed from the earlier work of Hollebone (1986). In general, therefore, potent induction can be regarded as generally deleterious, since it is associated with cell growth and proliferation via activation of the PKC cascade, and may lead to forms of toxicity associated with the production of oxygen radicals and other ROS (reactive oxygen species) (Parke, 1987a, 1994), including carcinogenicity. The precise nature and degree of toxic effect produced depends on a number of factors, including the species (Boobis et al., 1990), strain (i.e. genetics), pathophysiological status, exposure level or dosage of inducer, and the type of P450 involved. For example, it is known that the DBA/2 strain of mouse is not responsive to CYP1 induction by TCDD and other potent inducers, whereas C57BL/6 represents a responsive mouse strain (Okey, 1990). Presumably, this is due to variation in the ability to bind the Ah receptor in the regulatory region of the CYP1 gene, or caused by variability in the Ah receptor itself. Just as there are species differences in metabolism, it is also found that the nature of the species is relevant to the manifestation of toxic effects (Parke and loannides, 1990a). In particular, it is known that Table 5.12 Induction of CYP1 in human cells and tissues (Reference: Okey, 1990) Cell/tissue (a) In vivo Placenta Placenta Placenta Lung Epidermis (b) In cell culture Fetal liver cells Lymphoblastoid cells Epidermal cells Esophageal epithelium Squamous cell carcinoma Epidermis Hair follicle Hepatoma cell line Breast carcinoma cell line Thymic epithelial cells Peripheral blood lymphocytes * PCBs=polychlorobiphenyls; 3MC=3-methylcholanthrene; † TCDD=2,3,7,8-tetrachlorodibenzo-p-dioxin

Inducer Cigarette smoke PCBs* Dibenzofurans Phenobarbital Coal tar Benz(a)anthracene (BA) Benz(a)anthracene (BA) BA, 3MC BA, 3MC †TCDD, BA BA Coal tar †TCDD †TCDD, BA †TCDD BA

small rodents, especially the mouse, are more susceptible to the potentially toxic effects caused by P450 inducers, such as those associated with the production of oxygen radicals and other reactive oxygen species (ROS). This is due to the fact that metabolic rate, oxygen consumption and level of DNA damage all


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increase with decrease in body weight (Parke, 1987a; Martin and Palumbi, 1993). The ability of the organism to counteract the effects of oxygen radicals and protect against their formation is also an important factor in the likelihood of overt toxicity (Parke and loannides, 1994). Table 5.13 Induction of P450s in extrahepatic tissue (Reference: Okey, 1990) CYP Rat

Mouse

1A 1A1

Lung, ovary, testis, skin, Zimbal gland Lung, kidney, colon, spleen, small and large intestine

Lung, kidney, mammary gland, prostate Lung, kidney, prostate, duodenum, adrenal, Zimbal gland, pulmonary epithelium, alveoli, Clara cells, olfactory epithelium

1A2 2B 2B1 Lung, adrenal, testis, small intestine 2B2 Adrenal, testis Notes: Inducers of 1A: MC, TCDD, NF Inducer of2B: PB

Lung, kidney, spleen, small and large intestine Lung, bronchiolar epithelium

There are natural defence mechanisms which have been developed by the higher animals for dealing with oxygen radical toxicity and these include: superoxide dismutase (SOD), catalase and glutathione (GSH), together with vitamins E and C, which are quite effective reducing agents and antioxidants (Parke and loannides, 1994). Small rodents rapidly deplete their glutathione (Lorenz et al., 1984) following exposure to some toxic agents, whereas man is able to conserve glutathione levels by utilizing epoxide hydrolase (EH) for the detoxication of epoxides produced during the metabolism of certain compounds (Parke and loannides, 1990a). Consequently, the toxic effects of chemical agents which produce ROS by redox cycling, such as BHA and BHT, may vary between small rodents and man (Parke, 1987a). Due to the association between induction of CYP2E and fasting or restricted diet, and the ability of glutathione and other radical scavengers/antioxidants present in certain foodstuffs to detoxify ROS, the nutritional status of the subject will have an important bearing on the manifestation of toxicity following chemical exposure (Parke and loannides, 1994; Parke, 1994). Indeed, it is thought that the ability of the host organism to defend itself against the potentially deleterious ROS may have a relevance to ageing and life span (Orr and Sohal, 1994) whereas the varying anti-oxidant effects of dietary fatty acid composition have been linked with tumour development (De Vries and Van Noorden, 1992). There is substantial evidence, therefore, for the production of oxygen radicals and other activated oxygen species following induction of several forms of P450, particularly, CYP2E and CYP4A (Parke et al., 1990). These ROS are able to give rise to a number of different forms of toxicity which can be associated with lipid peroxidation, inflammation and DNA damage, depending on the nature of the active oxygen species (Kappus, 1993). There is, also, considerable evidence for a link between CYP1A induction and toxicity, being associated with cell proliferation via initiation of the PKC cascade (Enan and Matsumura, 1995) and activation of EGF, which leads, ultimately, to carcinogenesis (Parke et al., 1990; loannides and Parke, 1993). However, although induction of the PB-inducible forms, such as CYP2B (Table 5.14), might be generally regarded as beneficial, as it should enhance the ease of detoxifying metabolism, there are examples (Liu and Wells, 1994) where CYP2B induction can be potentially hazardous, especially if the inducing agent is a poor substrate, such that futile cycling can occur (Parke, 1994). In fact, it is possible that induction of CYP1 by

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TCDD may also produce ROS by futile cycling, as TCDD (Huff et al., 1994) is not a good substrate because of the blocking effect of the four Table 5.14 Variation in phenobarbital induction of rat hepatic P450s (References: Waxman and Azaroff, 1992; Gibson and Skett, 1994) CYP

Fold-induction

Comments

2A1 2B1 2B2 2C6 2C7 2C11 3A1 3A2

2.4 50–100 20 2–4 20 2 10 10

Also inducible by 1A1 inducers Major PB-inducible form Shows 97% homology with 2B1 Induction has been demonstrated in hepatoma cells Shows developmental regulation mRNA induced but not protein Also induced by PCN and some other synthetic steroids (Table 5.9) Shows male-specific expression

chlorine atoms in the molecule (loannides and Parke, 1993). However, in general, P450 substrates induce their own metabolism, that is they are inducers of the particular isoform which can metabolize them, although the metabolites of some P450 substrates may actually be more toxic and reactive than the parent compound; the metabolic activation of benzo(a)pyrene to the 7, 8-diol-9, 10-epoxide by CYP1 is an important example of this (loannides and Parke, 1993). Due to the fact that there is an elaborate biological defence mechanism for protection against the potential damage caused by oxygen radical production, especially in the higher mammals, it is possible that some of the toxic effects observed in small rodent species, following high dose levels of known P450 inducers, are not physiologically relevant to man (Parke, 1987a; Parke and loannides, 1994). Recent studies have shown, for example, that the expression and tissue specificities of human P450s are different from those in experiment rodents (Gonzalez and Gelboin, 1994). The variation in expression of CYP1A1 and CYP1A2 between rat and man is a case in point and, furthermore, peroxisome proliferation may not represent a significant hazard to man (Lake, 1995). However, the known hepatotoxicity of paracetamol can be attributed to the effects of redox cycling (Parke, 1987a) via the quinoneimine metabolite to produce superoxide radicals (Kappus, 1993), and similar processes may operate for the antioxidants BHA and BHT. Moreover, the induction of CYP2E and subsequent oxidative burst of ROS, following exposure to ether and other small molecular weight anesthetics, may represent a possible cause of post-surgical trauma (Liu et al., 1993a), especially when combined with pre-operative starvation. The numerous toxic effects of free radicals and their association with a variety of disease states and other conditions has been reviewed (Kehrer, 1993) and it would appear that a specific marker for the production of hydroxyl radicals (OH˙), for example, is the formation of 8-hydroxyguanosine. The hydroxyl radical, which can be formed from either superoxide or hydrogen peroxide, is able to attack DNA preferentially at the C-8 position on guanine, and the subsequent formation of the 8-hydroxy derivative gives rise to translation errors leading to carcinogenicity (Kehrer, 1993). Although the complete picture for the role of ROS in signal transduction is exceedingly complex and insufficiently understood, it has been shown that there is a good correlation between 8-hydroxyguanosine formation and carcinogenesis (Kehrer, 1993). Consequently, the involvement of the P450 system and its regulation by internal and external factors are likely to be of considerable relevance to the manifestation of toxic effects in both animals and man (Park and Kitteringham, 1988, 1990). Table 5.15 summarizes the association between various types of P450 induction and toxicity,


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of which CYP1A1 induction appears to have been most closely linked with genotoxicity, for example (Kopponen et al., 1994), although Table 5.15 Mechanisms of toxicity associated with P450 induction (Reference: Parke et al., 1990) CYP Mechanisms of potential toxicity 1A 2B 2E 4A

Binding to Ah receptor, activation of transcription by genomal derepression, initiation of PKC cascade Redox cycling and ROS generation Futile cycling and ROS generation Peroxisome proliferation and peroxide generation

there is also strong evidence for the role of CYP2E induction in toxicity (Terelius et al., 1993). Recent findings indicating an association between the effects of chemical carcinogens, such as aflatoxin B1 (Aguilar et al., 1993), and mutations in the p53 tumour suppressor (Cho et al., 1994) which appear to be related to cancer susceptibility (Shields et al., 1993), show that the molecular mechanisms governing tumorigenesis may be synergistic in nature. 5.5 Heterologous expression systems and in vitro models Following the recent developments in molecular biology and recombinant DNA biotechnology, it has become possible to express mammalian P450s (Gonzalez and Korzekwa, 1995) in other cell systems (Estabrook et al., 1991) such as those from bacteria, yeast and isolated animal cells; and some examples of these so-called heterologous expression systems are summarized in Table 5.16. The relationship between native and heterologous expression, together with the various processes involved, is presented in Figure 5.4, where it can be seen that an important intermediate step in the procedure is the determination of the complementary DNA (cDNA) sequence for the relevant P450 gene (Doehmer and Griem, 1993). Yeast appears to be an important host organism for the production of recombinant P450s (Sakaki et al., 1987, 1990; Imai, 1988; Ohkawa et al., 1990; Kedzie et al., 1991b; Yabusaki and Table 5.16 P450s expressed in heterologous systems (Reference: Doehmer and Griem, 1993) Cell system

CYP

Species

Cell system

E. coli

2E1 2G1 2A7 17A1 2C3 4A 1A1 1A1 1A1 1A1 1A2 1A2

Human Human Human Bovine Rabbit Rabbit Rat Mouse Human Rabbit Mouse Rat

Mammalian cells (a) Monkey

Yeast

(b) Hamster

CYP

Species

17A1 1A1 1A2

Bovine Mouse Mouse

2B1 1A1 1A2 1A2 2E1 1A1 1A2

Rat Rat Rat Human Human Human Mouse

172


Cell system

CYP

Species

1A2 2B1 2B2 2E1 3A4 17A1 21B1

Rabbit Rat Rat Rabbit Human Bovine Bovine

Cell system

CYP

Species

1A1

Mouse

(c) Rat

1A1 1A2 2E1

Mouse Mouse Human

(d) Mouse

1A2 2E1 2B1 1A1 1A2 1A1 1A2 1A2 2A6 2B7 2C8 2C9 2D6 2E1 2F1 4B1 3A3 3A4 3A5

Mouse Human Rat Mouse Human Mouse Mouse Human Human Human Human Human Human Human Human Human Human Human Human

(e) Human


173

Figure 5.4 Native and heterologous expression of P450 proteins (Reference: Doehmer and Griem, 1993).

Ohkawa, 1991; Winkler and Wiseman, 1992; Wiseman, 1993) and the bacterium Escherichia coli has also been used as an expression vector (Gillam et al., 1994; Sandhu et al., 1994). Moreover, a number of animal cell systems have been established that are capable of expressing human and other mammalian P450s which exhibit enzyme activities that parallel the in vivo situation (Miles and Wolf, 1991; Estabrook et al., 1991; Langenbach et al., 1992; Doehmer and Griem, 1993; Gonzalez et al., 1991). The maintenance of P450 activities in hepatocytes (Saad et al., 1994) and other cell cultures represents a convenient in vitro technique for the study of metabolism (Paine, 1990) and such systems may be used for the screening of development compounds and potentially toxic chemicals and carcinogens (Miles and Wolf, 1991). In vitro forecasting of potentially adverse drug interactions may also be carried out using human liver microsomal test systems (Dayer, 1990a and b; Dayer et al., 1989, 1992). One of the disadvantages of in vitro systems, however, is the generally rapid loss of enzymatic activity, although there appears to be considerable species variation and a number of modifications, such as co-culture and alteration of the culture medium, have been employed to increase the viability of, for example, hepatocyte cultures of mammalian P450s. For further details, the interested reader is referred to in an excellent review by Schwarz and Wiebel (1993). Various methods for the utilization of P450s, particularly human isoforms, in the early screening of novel pharmaceuticals are regularly employed by the drug industry, and it would appear that there have been major advances recently, particularly in heterologous expression systems (Gonzalez and Korzekwa, 1995). 5.6 Inhibition of cytochromes P450 There are many known inhibitors of various P450s, some of which are specific for individual isoforms, and there are several different mechanisms by which a P450 enzyme may be inhibited, such that it is possible to

174


classify inhibitors into particular categories. Inhibitors may be metal ions (e.g. Co2+), small molecules (e.g. CO and NO), heme iron ligands (e.g. ketoconazole and metyrapone), terminal alkenes or alkynes (e.g. gestodene) and other mechanism-based inhibitors (e.g. chloramphenicol), some of which may be of a competitive nature, and many P450 antibodies are capable of inhibiting the interaction between redox components in the P450 system. There have been a number of reviews describing the various types of P450 inhibitors, including those which exhibit some degree of specificity towards individual isoforms, and the reader is referred to the following for further details: Testa and Jenner, 1981; Ortiz de Montellano and Reich, 1986; Kapetanovic, 1990; Murray and Reidy, 1990; Testa, 1990; Murray, 1992; Correia and Ortiz de Montellano, 1993; Gibson and Skett, 1994; Halpert et al., 1994; Karuzina and Archakov, 1994; Halpert, 1995. The following sections describe the various types of P450 inhibitors, although there are some examples which can be regarded as of a mixed type. 5.6.1 Competitive inhibitors A P450 inhibitor is described as being competitive if it is able to bind reversibly to either the heme iron, or elsewhere in the active site, and thus compete with either oxygen or a particular substrate for binding to the P450 concerned. (In practice, however, the term competitive inhibition is usually restricted to that of substrate metabolism.) For example, carbon monoxide (CO) appears to be a specific inhibitor of P450s in general, but it is not specific for any particular isoform (Testa and Jenner, 1981). This is because it competes with molecular oxygen for ligation of the heme iron (in the Fe2+ state) and, although it is used as an identifier of P450, indeed, the wavelength of the absorption maximum of the CO complex (Table 5.17) led to the ‘450’ portion of the enzymes’ name, carbon monoxide is also able to bind to the heme iron of other hemoproteins, such as hemoglobin and myoglobin (Chapter 1). However, carbon monoxide exhibits lower affinity for P450 than does oxygen, and there are variations in its affinity between different isoenzymes (Testa and Jenner, 1981). Other small molecular species, such as nitric oxide (NO), cyanide (CN¯ ) and isocyanides (e.g. EtNC), are able to inhibit P450 in a similar way to CO by competing with O2 for the heme (Hill et al., 1970a). It is likely that the orientation of these small molecular ligands differs from that of oxygen due to the variations in bonding between the different complexes and because of possible interactions with one or more amino acid residues, such as the invariant threonine, within the P450 heme pocket (Chapter 1). All of these small molecular ligands are relatively strong -acceptors and give rise to a bathochromic shift in the wavelength (Table 5.18) of the Soret maximum in the UV, which is indicative of a modulation in the iron spin state equilibrium in favour of the low-spin form (Ortiz de Montellano and Reich, 1986). The CO-complex of P450cam (CYP101) has been characterized by X-ray crystallography to a 1.9 Å resolution (Raag and Poulos, 1989). Interestingly, the CO ligand assumes a torsional angle which is non-perpendicular with respect to the plane of the porphyrin ring, in contrast to the situation encountered in other hemethiolate CO complexes, where there is a strong tendency to bind perpendicularly (Testa and Jenner, 1981). The likely explanation for the observed CO ligand bond angle in P450camCO probably lies in the possibility of a hydrogen bonding interaction between the CO oxygen atom and the hydroxyl side chain of Thr-252, and it may be that dioxygen binds to P450 in an analogous fashion, with the hydrogen-bonded interaction with Thr–252 probably playing a key role in the catalytic mechanism (Chapter 3). In addition to the small molecular ligands (CO and NO) which compete with oxygen for the P450 heme iron, there is a large number of organic P450 inhibitors that are able to ligate the heme moiety but also mimic P450 substrates, sometimes in a specific manner (Testa and Jenner, 1981). These organic ligands all


175

contain an electronegative atom (N or O) which coordinates to the heme iron via donation of a lone pair of electrons, and there is often some back-bonding from iron to augment the strength of the coordinate bond as a synergic effect. Heterocyclic groupings, such as imidazole, triazole, pyridine, quinoline, furan and benzofuran, are particularly good heme ligands due to their ability to form p -d bonded interactions with iron and, in P450, the proximal thiolate ligand from the invariant cysteine residue probably pushes electron density onto the heme iron (Hawkins and Dawson, 1992). This effect favours imidazole (and other related heterocycles mentioned previously) as ligands in P450 inhibitors such as ketoconazole, for example, where there is also likely to be an interaction between other groupings on the inhibitor and certain amino acid residues in the P450 active site (Correia and Ortiz de Montellano, 1993). Although imidazole and triazole are stronger ligands than pyridine, metyrapone (which contains two pyridine rings) is an example of a competitive, non-specific, P450 inhibitor (Testa and Jenner, 1981). The crystal structures of P450cam with several bound inhibitors including metyrapone and phenylimidazole, have been determined to a 2.2 Å resolution (Poulos, 1988). In the crystal structure of metyrapone-bound P450cam, one pyridine nitrogen ligates the heme, whereas the second forms a hydrogen bond with Tyr-96, which is the same residue involved in hydrogen bonding to the endogenous substrate, camphor (Poulos et al., 1987). However, metyrapone and phenylimidazole also inhibit the phenobarbitalinducible microsomal P450s, and it is possible that their mode of interaction is different from that with P450cam, as there will be a number of Table 5.17 Characteristics of purified drug-metabolizing P450s from rat, rabbit and mouse (References: Omura et al., 1993; Nelson et al., 1993) CYP Species

(nm.) COadduct

Fe3+ spin state Apparent M.Wt. (kDa)

No. of amino acid Typical inducers residues

1A1

Rat

447

L

56

524

1A1 1A1 1A2

Mouse Rabbit Rat

449 448 447

L H

55 56 54

524 518 513

1A2 2A1 2A2 2A4 2A5 2B1 2B2 2B4 2B5 2B9 2C2 2C3 2C4 2C5 2C6

Mouse Rat Rat (M) Mouse (F) Mouse (M) Rat Rat Rabbit (lung) Rabbit Mouse Rabbit Rabbit Rabbit Rabbit Rat

448 451 452 451 451 450 451 451 451

L L H H L L L L

451 450

L>H L

55 48 48 48 48 52 52 49 49 54 50 51

450 450

L L

48 50

513 492 492 494 494 491 491 491 491 491 490 490 487 487 490

max

MC, TCDD, PCB, NF MC TCDD MC, PCB, isosafrole MC, isosafrole

PB PB PB PB

176


CYP Species

(nm.) COadduct

Fe3+ spin state Apparent M.Wt. (kDa)

No. of amino acid Typical inducers residues

2C7

448

H>L

490

Rat

max

2C11 2C12 2C13 2C14 2D1 2D2 2D9 2E1

Rat (M) Rat (F) Rat (M) Rabbit Rat Rat (neonate) Mouse Rat

2E1 2E1

Mouse Rabbit

451 448 449 451 448 448 449 452

51 L>H L L L L

452

H

50 51 50 50 52 52 50 51

500 490 490 490 504 500 504 493

H

51 51

493 493

Non-inducible Non-inducible Ethanol, isoniazid Ethanol, isoniazid

2E2 2J1

Rabbit 452 L>H 52 493 Rabbit 56 501 (intestine) 3A2 Rat 449 L 51 504 PCN, TAO 3A6 Rabbit 449 L 52 501 TAO 4A1 Rat 452 L 52 511 Clofibrate 4A4 Rabbit 450 L 52 506 Progesterone 4A5 Rabbit L 55 511 Clofibrate 4A6 Rabbit L+H 53 510 Clofibrate 4A7 Rabbit L+H 53 511 Clofibrate 4B1 Rabbit 449 56 506 PB Notes: L=low-spin, H=high-spin; M, F=male, female dominance/specificity; PB=phenobarbital ; PCN=pregnenolone 16 -carbonitrile; MC=3-methylcholanthrene; PCB=polychlorobiphenyls; TCDD=2, 3, 7, 8tetrachorodibenzo-p-dioxin; NF= naphthoflavone; TAO=triacetyloleandomycin. Table 5.18 Soret absorption maxima of oxidized (Fe3+) and reduced (Fe2+) microsomal P450 showing the effect of small molecular inhibitors (Reference: Hill et al., 1970a) Inhibitor Fe2+

max

(nm.)

Inhibitor Fe3+

max

(nm.)

none 412 none 416 CO 450 NO 432 EtNC 429, 454* EtNC 430 CN− 432† CN− 436 * Value produced at high pH. † Anionic ligands, such as cyanide (CN−) exhibit relatively low affinity for reduced P450 due to the fact that the iron bears an overall net electronic charge of −1 when Fe2+.


177

differences between the active sites of these enzymes, including the fact that Tyr-96 is not conserved in an analogous position in the protein sequences of microsomal PB-induced P450s (Lewis, 1995a). In addition to metyrapone, proadifen (SKF-525A) is an inhibitor of the phenobarbital-inducible forms (together with some steroidogenic P450s) and its mode of inhibition is thought to be somewhat different. Apparently, SKF-525A is N-deethylated by the P450 concerned and is converted to an N-oxide (or a nitroso derivative) which then inhibits the enzyme via heme ligation (Testa and Jenner, 1981; Murray, 1992). It is, therefore, an example of a mechanism-based reversible inhibitor. The crystal structures of both SKF-525A and metyrapone have been determined and exhibit the characteristic V-shaped or ‘butterfly-wing’ conformation shown by substrates of PB-inducible P450s (Rossi et al., 1987). In addition to molecular shape similarities, there are other common structural features (Lewis et al., 1987) between these P450 inhibitors and typical substrates of the same enzymes, which point to certain commonalities in their interactions with key active site residues (Chapter 6). Knowledge of the structural specificities of known P450 substrates, and a consequent inference of the relevant active site features, can assist in the design of specific inhibitors (Halpert, 1995). Therefore, a specific P450 inhibitor must be complementary to the active site topography in addition to possessing a hemebinding moiety, or a functionality capable to interacting with a particular amino acid in the binding pocket; sometimes this will be an irreversible interaction, such as that encountered with chloramphenicol (Miller and Halpert, 1986; Halpert et al., 1988; Bories and Cravedi, 1994). An example of a reversible inhibitor which has complementarity with key amino acid residues in the appropriate P450 active site is that of the specific 2D6 inhibitor, quinidine (Halpert et al., 1994). In addition to its quinoline nucleus, which ligates the heme via the nitrogen atom, this 2D6 inhibitor also contains a second nitrogen in a quinuclidine ring system which is capable of becoming protonated at physiological pH, and can thus form an ion-paired interaction with a key aspartate residue in the 2D6 active site (Lewis, 1995a). Furthermore, quinidine possesses a hydroxyl group which could enter into a hydrogen-bonded interaction with a complementary amino acid residue (probably serine) within the vicinity of the heme pocket (Lewis, 1995a). Interestingly, the enantiomeric antipode of quinidine, namely quinine, is a specific inhibitor of the rat orthologue, 2D1, and a comparison between the protein sequences of 2D6 and 2D1 shows a conservative change from serine to threonine in the vicinity of the putative active site (Lewis, 1995a). Molecular modelling of the two isomeric inhibitors within the relevant P450s indicates that the specificity of these compounds can be explained in terms of hydrogen bonding possibilities between the inhibitors and the key amino acid residue change, serine to threonine, within the distal I helix close to the heme (Lewis, 1995a). The azole series of P450 inhibitors, which includes the structurally-related compounds ketoconazole, itraconazole, miconazole and clotrimazole, for example, are able to inhibit a number of different P450s, including fungal forms of the enzyme, and are, consequently, useful in the treatment of fungal infections (Correia and Ortiz de Montellano, 1993). Inhibition of some key pathways in steroid metabolism, such as the aromatization of androgens, by certain azole derivatives (including ketoconazole) has also led to the development of these P450 inhibitors as therapeutic agents for the treatment of some forms of cancer (Correia and Ortiz de Montellano, 1993). The search for specific inhibitors of various P450s can have important therapeutic consequences, therefore, not only in the design of novel agents which can be employed as antifungal or antineoplastic therapies, but also in the development of diagnostic probes for P450–mediated metabolism which may be useful for the screening of new pharmaceutical discoveries. As the hepatic microsomal P450s play such a key role in the Phase I metabolism of foreign compounds, an early indication of the likely routes of metabolism in man can be of potential benefit to novel drug development programmes. Table 5.19 shows some examples of specific P450 inhibitors for a number of different forms of the enzyme. It can be seen that such inhibitors may be either mechanism-based (see section 5.6.2) or competitive heme ligands, but

178


both types will also combine substrate mimicry in the rest of the molecular structure to enable specific binding to the relevant active site residues. However, Table 5.19 Specific inhibitors of P450s (References: Murray and Reidy, 1990; Halpert et al., 1989, 1994; Correia and Ortiz de Montellano, 1993; Laughton and Neidle, 1990; Vickery and Kellis, 1983; Van den Bossche, 1988) CYP

Specific inhibitor

Other inhibitors

1A1 1A2 2A6 2B1 2C9 2D1 2D6 2E1 3A4 4A1 11A1 11B1 17A1

1-ethynyl pyrene Furafylline Diethyldithiocarbamate Secobarbital Sulfaphenazole Quinine Quinidine Disulfiram Gestodene 11-undecynoic acid 22-amino-23, 24-bisnor-5-cholen-3 -ol Metyrapone (1S, 2S, 3S, 5R)-(+)-isopinocamphenyl 4-pyridylacetic acid ester 19A1 4-hydroxyandrostenedione 21 21, 21-difluoroprogesterone 51 Itraconazole 101 Metyrapone

-naphthoflavone 9-hydroxy ellipiticine Metyrapone Proadifen (SKF-525A) — — — Diallyl sulfide Ketoconazole 11-dodecynoic acid (20R)-20-phenyl-5-pregnene-3 , 20-diol — 4-cyclohexyl-2-methyl-2(4-pyridyl) propanoate Vorozole, aminoglutethimide — Ketoconazole Phenylimidazole

Table 5.20 Iron porphyrin complexes with axial nitrogen ligands: physicochemical and structural data# Ligand

pKa

log Kexpected

log Kobs

log Kdecrease

Bond angle (°) Bond length N−Fe−N (Å) Fe−N

Benzimidazole 5.46 3.9 2.7 1.2 175.0 27.9 4.7 2.6 2.1 170.38 Methylimidazo le 26.0 2.8 1.0 1.8 174.06 Methylpyridin e Dimethylamin 10.64 4.0 2.7 1.3 176.68 e Isopropylamin 10.63 4.0 1.5 2.5 170.63 e t-Butylamine 10.55 4.0 0.5 3.5 162.19 Trimethylamin 9.8 3.8 0.0 3.8 178.37 e Notes: log Kdecrease=–0.16 (±0.02) Bond angle N−Fe−N+29.44† R=0.97 (excluding Trimethylamine*)

2.076 2.064

2.104

2.042 2.042 2.053 2.069


Ligand

pKa

log Kexpected

log Kobs

log Kdecrease

179

Bond angle (°) Bond length N−Fe−N (Å) Fe−N

pKa=ligand basicity K=equilibrium (stability) constant for formation of iron porphyrin complex (Byfield et al., 1993) log Kdecrease=log Kexpected–log Kobserved log Kexpected values were calculated from ligand pKa data using the relationship :log K=a.pKa+b (Byfield et al., 1993) * As trimethylamine is a symmetrical tertiary amine, this ligand shows an increase in Fe−N distance rather than a decrease in the axial N−Fe−N bond angle, thus justifying its position as an outlier. † A plot of this relationship is shown in Figure 5.5. # Pratt and Lewis, unpublished results.

from a therapeutic point of view, reversible inhibition may be more desirable than the irreversible inhibition generally associated with many mechanism-based inhibitors, as some of the P450s involved have important physiological functions. Although the azole series of compounds possess inhibitory activity towards a variety of P450s, it is found that the change from imidazole to triazole increases the specificity for the fungal form P45014DM (CYP51) which catalyzes the C14demethylation of lanosterol (Correia and Ortiz de Montellano, 1993). The H2antagonist cimetidine is another example of an imidazole derivative which exhibits P450 inhibitory activity; however, the furan analogue, ranitidine, does not appear to do so (Murray, 1992), and this is probably due to steric hindrance of the iron ligation by the bulky 2-substituent on the furan ring. It is found that, in a series of cyclic nitrogenous iron porphyrin ligands, an increase in sterically-hindering groups is paralleled by a decrease in the observed association constant, Kobs. The steric effect of bulky groups flanking the ring nitrogen atom gives rise to both an increase in the Fe-N distance and in the N-Fe-N bond angle between the ligand nitrogens and iron porphyrin moiety: these geometric parameters (Table 5.20) correlate closely (r=0.97) with the difference between the Kobs values relative to that expected on the grounds of ligand pKa (Pratt and Lewis, unpublished results) as shown in Figure 5.5. The typical Fe-N distance in both model compounds and in the P450cam-bound nitrogenous inhibitor complexes is about 2.0 Å (including the iron-porphyrin nitrogen distances) and this is similar to the results obtained from molecular modelling studies on related iron porphyrin complexes, as shown in Table 5.20. In fact, the movement out of perpendicular shown by sterically-hindered iron-porphyrin ligands is mirrored in the P450cam-inhibitor crystal structures (Poulos, 1988). Furthermore, the ligating nitrogen electron density is also important to achieve a high binding affinity with iron porphyrins and heme complexes, whereas the orientation of the coordinated ligand represents a compromise between optimal p –d orbital overlap and steric interactions. It would appear that both electronic and molecular structural factors operate in describing the inhibitory potencies of nitrogen heterocyclic P450 ligands, such as imidazoles, triazoles, benzimidazoles, pyridines and quinolines, and for aliphatic amines (Lewis, 1995b). Moreover, these effects can explain the variations in activity of oxygen-based P450 inhibitors such as ethers, ketones, quinones, phenols, coumarins (Lewis et al., 1994b,c) and alcohols (Lewis, 1987). However, the potency differences of methylenedioxybenzenes, that are able to inhibit P450 via formation of the carbene which subsequently ligates the heme, can be rationalized solely on the basis of molecular planarity (Lewis et al., 1995b). However, these mechanism-based inhibitors are like SKF-525A and orphenadrine, competitive inhibitors. It should be noted that the type of spectral change associated with substrate binding provides an indication of the likelihood of heme ligation, as this form of interaction is generally accompanied by a type II spectral change (Chapter 1). Consequently, many P450 inhibitors which act as heme ligands will exhibit type II spectra, indicative of a modulation in the iron spin-state equilibria in favour of the low-spin form

180


Figure 5.5 Correlation between binding affinity difference and axial ligand bond angle in iron porphyrins.

(Schenkman et al., 1981). However, there are some inhibitors which show mixed types of spectral change, for example SKF-525A; the probable reason for this could be due to its mechanism of inhibition. The related compound, orphenadrine, undergoes the following sequence (Murray, 1992) of metabolic transformations :

and it is possible that the structurally-analogous SKF-525A operates via a similar mechanism, although this would involve N-deethylation rather than N-demethylation. Figure 5.6 shows the above scheme in more detail, together with a comparison between the structures of SKF-525A and orphenadrine. The formation of the nitroso compound, which presumably will ligate the heme iron in an analogous fashion to nitric oxide, may involve the transient production of a nitroxide intermediate. Earlier studies on simpler amines have demonstrated that the sequence of events outlined in Figure 5.6 is a likely mechanistic route for the production of nitroso metabolites (Testa and Jenner, 1981). As stated previously, the electronegative ligand atoms (i.e. N or O) do not necessarily have to be present in an aromatic ring system to give rise to inhibition, so aliphatic amines and alcohols will also be capable of acting as P450 inhibitors to varying degrees. In a series of primary aliphatic amines binding to microsomal P450, it has been found that the affinity constants correlate with several structural parameters (Lewis,


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Figure 5.6 Metabolic transformations in orphenadrine (Reference: Testa and Jenner, 1981).

1995b). Apparently, aliphatic amines exhibit both high and low affinity binding characteristics towards phenobarbital-induced rat hepatic microsomal P450 (Jefcoate et al., 1969). The relevant binding affinities increase with increasing length of the aliphatic chain until a point is reached where the binding levels off, and then commences to diminish after about n = 8 (where n is the number of carbon atoms in the chain). It can be shown that the strength of this binding is dependent on hydrophobicity, as determined by log P, the electronic charge on the nitrogen atom and by the pKa of the amine (Lewis, 1996a). These findings suggest that there may be binding interactions to (a) the heme iron, (b) a nearby carboxylate group from an acidic amino acid residue, and (c) complementary hydrophobic residues within the P450 active site, which applies a size constraint to substrate binding. The high and low affinity binding characteristics may, therefore, involve interactions between the amine and either the heme iron or the conserved acidic residue one position upstream from the invariant threonine, with hydrophobic interactions governing the association of the alkyl chain to the protein in the region of the active site (Lewis, 1996a). If one assumes that the major PB-inducible form of P450 is involved in the binding interactions, then the limitation on length of the alkyl moiety is similar to that observed in a homologous series of 2, 4-dichlorophenoxy N-alkyl N-methylethylamines, which exhibit a peak activity at the N-butyl homologue for metabolism by CYP2B1 (Roffey, 1993). It is possible that the interaction between SKF-525A and the phenobarbital inducible P450 isoforms is similar to the aforementioned cases, and molecular modelling indicates that this is likely to be the case (Chapter 6). Aliphatic primary alcohols also act as competitive P450-inhibitors but, in this case, it would appear that the p-hydroxylation of aniline is inhibited (Testa and Jenner, 1981), thus indicating that CYP2E1 is the relevant enzyme involved. Quantitative structure-activity relationships generated for a series of short chain alcohols show that inhibition of CYP2E-mediated aniline p-hydroxylation can be explained in terms of molecular length and electronic features in the molecule (Lewis, 1987). Nevertheless, heterocyclic compounds are stronger ligands than their aliphatic counterparts, as evinced by pyridines, imidazoles, ellipticines, benzimidazoles, furans, benzofurans and triazole P450 inhibitors. Quinolines and related heterocyclics containing an exocyclic protonable nitrogen at between 5 and 7 Å distance from the quinoline ligating nitrogen (e.g. quinidine), and at an optimum orientation, tend to possess specific inhibitory activity towards CYP2D6 and some of the other orthologues in this subfamily (Smith and Jones, 1992). The definitive work, which combines molecular modelling with inhibition studies, is that of Strobl and colleagues (Strobl et al., 1993). As mentioned previously, specific inhibitors of P450s, such as quinidine for 2D6, are particularly important in determining which P450 isoform is involved in a metabolic pathway of, say, a novel development compound. This identification is especially relevant for the assessment of the

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relative importance of P450 isoforms associated with the ‘poor-metabolizer’ phenotypes, i.e. 2D6 (Otton et al., 1988) and 2C19. For imidazole and benzimidazole inhibitors, a combination of molecular and electronic structural descriptors is able to rationalize the variations in potency for limited numbers of analogues. It appears that both whole molecular properties, such as E(LUMO) and dipole moment, and electronic factors associated with the imidazole nucleus, such as individual net atomic charges and electrostatic potential energy minima, give good correlations with inhibitory activity (Lewis, 1995b). Cyclic and aromatic ethers are also able to inhibit P450s and furafylline, with its iron-ligating furan ring system, is an example of a specific inhibitor of CYP1A2 (Table 5.19 ). Aromatic compounds containing the benzoflavone ring system, such as α- and β-naphthoflavone, also inhibit CYP1 (Testa and Jenner, 1981) but these do not show the specificity of furafylline, as this ligand was designed from the structure of caffeine, which is a specific CYP1A2 substrate. Certain aromatic amines are also able to act as P450 inhibitors, possibly via heme ligation, and aniline is an example which exhibits a type II spectral change indicative of the change from high- to low-spin iron that is associated with heme iron ligation. Fuhr and co-workers have reported a QSAR and molecular modelling study on a number of quinolone derivatives which act as inhibitors of CYP1A2 (Fuhr et al., 1993). This important work highlights the powerful combination of electrostatic potential energy contour surface mapping, molecular fitting between substrates and inhibitors, and multivariate analysis of molecular and electronic properties. In this investigation, where the experimental work on CYP1A2 inhibition was also carried out in tandem with the theoretical studies, a good correlation was found between a key atom electron density, electrostatic potential energy minima and molecular volume (Fuhr et al., 1993). It is possible, therefore, with the current ‘state-of-the-art’ in structural modelling technology, to rationalize the specific inhibition of P450s in terms of molecular features of the relevant compounds. 5.6.2 Irreversible and mechanism-based inhibitors Mechanism-based inhibitors are those types which are converted into an active form by the enzyme concerned, and are then able to bind (usually irreversibly) with either the heme moiety or to the protein, normally within the active site region (Testa, 1990). Some notable examples of the latter have already been mentioned, for example, SKF-525A and chloramphenicol. However, there are other mechanism-based inhibitors which act as suicide substrates via heme ligation, and the methylene dioxybenzenes are examples of this type (Testa and Jenner, 1981) as has been noted in the preceding section. Another class of suicide substrate inhibitors are the haloalkanes, notably small molecular weight chloroalkanes, such as chloroform and tetrachloromethane; these form carbene intermediates that are capable of heme ligation in a similar manner to the methylene dioxybenzenes (Testa and Jenner, 1981). An important class of mechanism-based inhibitors which bind irreversibly to the heme and thus inactivate the P450 are certain terminal alkenes and alkynes which comprise a number of specific P450 inhibitors, such as gestodene (Guengerich, 1990c), secobarbital, allylisopropylacetamide and ethinylestradiol (Murray, 1992). The action of these irreversible inhibitors has been reviewed by Ortiz de Montellano (1987), and by Ortiz de Montellano and Reich (1986), whereas suicide substrates in general have been reviewed recently (Karuzina and Archakov, 1994). Selective P450 inhibitors have been reviewed by Murray and Ryan (1990), Halpert (1995) and by Halpert et al., (1994); whereas Correia and Ortiz de Montellano have produced a useful review on the therapeutic uses of P450 inhibitors (Correia and Ortiz de Montellano, 1993).


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It is generally accepted that the terminal alkene and alkyne P450 inhibitors exert their activity by an initial oxygenation step to form a ketene (or oxetane) which then reacts with the porphyrin skeleton of the heme moiety (Correia and Ortiz de Montellano, 1993). Such reagents are, therefore, non-competitive inhibitors as they are associated with deactivation and destruction of the heme. It should be noted that, in the cases of specific inhibitors, the essentially non-reactive part of the molecule is analogous to the structure of a known substrate of that enzyme, for example, 11-undecynoic acid and lauric acid in the case of P4504A1 (Table 4.19). In fact, it can be regarded as a general characteristic of specific inhibitors (Figure 5.7) to be complementary with the relevant P450 active site, in addition to possessing a heme binding group. There are several other classes of compounds which give rise to an irreversible reaction that is similar to the heme alkylation processes carried out by terminal alkenes and alkynes. For example, phenyldiazene (PhN = NH) and other diazenes, phenyl hydrazine (PhNHNH2) and other hydrazines, aminobenzotriazoles, alkyl-dihydropyridines and dialkyldihydroquinolines all react with the prosthetic heme group of P450 to give rise to covalent heme adducts (Ortiz de Montellano and Reich, 1986). The X-ray crystal structure of the catalytically inactive complex, resulting from the reaction between phenyldiazene and cytochrome P450cam, has been determined and refined to a 1.9 Å resolution (Raag et al., 1990). The complex shows that the phenyl group of phenyldiazene is able to form a covalent bond with the heme iron, following the loss of nitrogen, and it is thought that subsequent migration of the phenyl group onto one of the porphyrin nitrogens will give rise to an N-phenyl protoporphyrin IX derivative. Extensive studies on the nature of these porphyrin adducts has provided an important insight into the active site topology of various P450 isozymes (Ortiz de Montellano, 1987; Swanson et al., 1991, 1992; Ortiz de Montellano and Graham-Lorence, 1993; Tuck et al., 1992, 1993b). Apparently, the relative proportions of N-phenyl protoporphyrin IX regioisomers produced from the ferricyanide-induced oxidative breakdown of the phenyl-heme complex provides an indication of the active site topographies of different P450s (Ortiz de Montellano and Graham-Lorence, 1993). A compilation of the results from the active site studies of Ortiz de Montellano and co-workers is presented in Table 5.21, where it can be seen that there are marked differences between P450 isozymes, even within the same subfamily. Following the X-ray crystallographic determinations of the three prokaryotic P450s (CYP102, CYP101 and CYP108) studied by the above heme adduct rearrangement technique, there is an opportunity to investigate whether there is, in general, broad agreement between the inhibition product ratios and the actual crystal structure geometries. If the X-ray data for these bacterial P450s corroborate with the inhibition studies, then the latter may be used to assess the validity of other P450 models generated from these crystal structures. This will be discussed in the following chapter, which is concerned with P450 models. In addition to P450 inhibitors which bind covalently with either the heme or active site, there are other chemical agents (Table 5.22) that are able to react specifically with certain types of amino acid residues such as lysine and cysteine. It has been demonstrated by Bernhardt and colleagues that certain exposed lysine residues on the surface of microsomal CYP2B4 are important for reductase binding (Bernhardt, 1993; Bernhardt et al., 1984, 1988, 1989). Consequently, irreversible chemical modification of such basic residues may inhibit P450 catalysis by blocking the interaction between a P450 and its redox partner(s). This type of information is important for elucidating the mode of electron transfer and redox interactions between reductase (or other redox partners) and P450, together with defining the surface topography of the enzymes in the microsomal membrane (Chapter 3). Another variety of inhibitors which can be more specific for individual P450 isozymes, but also relate to surface characteristics, are antibodies, as some of these may be designed to block redox partner interactions (Reik et al., 1982; De Lemos-Chiarandini et al., 1987; Edwards et al., 1991, 1993, 1994, 1995; Murray et al., 1993). Antibodies raised to recognize a particular surface epitope on a P450 may also either prevent

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Figure 5.7 Structures of some specific P450 inhibitors.

redox partner binding, if the recognition site is sufficiently close to a reductase/redoxin interaction point, or obstruct the access of a substrate to the heme, depending on the position and orientation of the epitope


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within the P450 sequence. In both instances, the binding of a specific antibody will act in an inhibitory fashion although, in this case, the inhibition can be reversible. However, the main use of specific antibodies for P450 isozymes lies in their ability to recognize individual isoforms (Reik et al., 1982) and thus aid immunoquantification of the respective P450 levels in a particular sample under investigation. For example, specific P450 antibodies have been employed to immunoquantify the amounts and variations of human hepatic microsomal P450s in Japanese and Caucasian groups (Shimada et al., 1994). 5.6.3 Therapeutic and agrochemical uses of P450 inhibitors Table 5.18 shows a number of specific P450 inhibitors together with the relevant isoform involved, and the structures of some of these are given in Figure 5.7. As certain P450-mediated reactions are physiologically relevant, particularly those associated with steroid metabolism, agents have been designed to inhibit some of these steroidogenic pathways, and have been found to possess therapeutic value in the control of a number of disease states, fungal infections and other conditions (Coulson et al., 1984; Covey, 1988; Berg and Plempel, 1988; Guengerich, 1990b; Correia and Ortiz de Montellano, 1993). A major use of some of these P450 inhibitors lies in the treatment of various forms of cancer, such as post-menopausal breast cancer, prostate cancer and related disease states, such as benign prostatic hyperplasia (Correia and Ortiz de Montellano, 1993). There is also some potential utility for P450 inhibitors in the treatment of hyperaldosteronism, hypertension, and for the alleviation of various conditions associated with

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Table 5.21 Ratios of N-phenylprotoporphyrin IX regioisomers produced via ferricyanide-induced phenyl group migration of the phenyl-iron P450 complexes formed by the reaction of various P450s with phenyldiazene*

* Modified and augmented from Ortiz de Montellano and Graham-Lorence, 1993.

abnormalities in arachidonic and retinoic acid metabolism (Correia and Ortiz de Montellano, 1993). In particular, aromatase (CYP19) inhibitors have been employed in the treatment of breast cancer, whereas anti-prostatic agents include compounds active against both CYP17 and CYP19. The various therapeutic applications of P450 inhibitors have been summarized recently in an excellent review article (Correia and Ortiz de Montellano, 1993), and the reader is directed to this work for further details. In the agrochemical area, P450 inhibitors have been shown to be effective fungicides, particularly with respect to the marked inhibitory properties of certain azole derivatives (Berg and Plempel, 1988) on the biosynthetic pathway of ergosterol from lanosterol, which involves a C-14 demethylation step that is catalyzed by a fungal form of the enzyme, namely, CYP51. Other areas of potential use for P450 inhibitors


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Table 5.22 Chemical modification of amino acid residues by some specific reagents

*EDC=1-ethyl-3[3’ -dimethylaminopropyl]carbodiimide (Strobel et al., 1989) MNT=2-methoxy-5-nitropone (Bernhardt et al., 1988) FITC=fluorescein isothiocyanate (Bernhardt et al., 1984) DIFIA=diiodofluorescein iodoacetamide (Chernogolov et al., 1994)

include plant growth regulation (Coulson et al., 1984) and in the development of novel herbicides, antimicrobial agents and fungicides (Durst, 1991; Durst and Benveniste, 1993). 5.6.4 Metal ions as P450 inhibitors and iron redox state preferences Metal ions are also able to inhibit P450, but these are, in general, not specific for any particular isoform (Testa and Jenner, 1981). However, there has been some recent evidence for possible isoenzyme specificity as, for example, lead nitrate appears to preferentially inhibit CYP1A2 (Degawa et al., 1993), whereas cadmium and mercury salts have been shown to inhibit CYP2E but not CYP3A (Alexidis et al., 1994). A mechanism has been proposed which rationalizes the way in which certain metal ions, such as Co2+, may act as inhibitors of P450 (Testa and Jenner, 1981). Apparently, cobaltous ions are able to both inhibit heme biosynthesis and increase the rate of heme degradation, and similar effects have been observed for

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gold salts, in addition to a number of other transition metal and heavy metal ions. However, cadmium seems to act in a different manner, by promoting the conversion of P450 to the inactive form, P420, and this may involve binding to either thiol or carboxylic acid groups, thus affecting the conformation of the protein, presumably in the vicinity of the heme binding site. An obvious candidate for such an interaction is the invariant cysteine which ligates the heme, although the conserved acidic residue (aspartate or glutamate) one position upstream of the invariant threonine, distal to the heme, is another possibility for binding cadmium ions. In the crystal structure of P450cam (CYP101) there is a recognizable cationic binding site close to the heme pocket, and it is known that bound potassium ions, for example, can increase substrate binding affinities, possibly by affecting the conformation of the B’ helix to facilitate displacement of the sixth ligand (Raag and Poulos, 1992). Perhaps there are cationic binding sites in other P450s which may be responsible for metal ion-induced alteration in substrate binding characteristics by affecting the localized protein conformation in the active site region. Whether such metal binding sites exist in P450s other than P450cam, however, has not been demonstrated to date. The fact that Cd2+ and Hg2+ administration preferentially affects CYP2E, indicates that the high redox potentials of the metal/metal ion couple may cause some form of interference with the iron redox or spin-state equilibrium in CYP2E, as this form is closely associated with the generation of oxygen free radicals, primarily due to its preference for the high-spin ferric state. Clearly, there are several possible explanations for the effect of metal ions on P450 and it could be that more than one is in operation simultaneously. Although there is likely to be a dose dependency on the type of mechanism by which metal ions inhibit P450s, it may be possible to rationalize their effects in terms of some combination of redox potential and ionic radius compared with those of either Fe2+ or Fe3+. In fact, it is known that some inhibitors exhibit a preference for either of the two iron redox states (Testa and Jenner, 1981). For example, coordination of a strong ligand to ferric iron of P450 in its resting state, or displacement of a weak ligand, shifts the P450 spin-state equilibrium in favour of the low-spin form, thus giving rise to a type II binding spectrum. (Note that most substrates for the enzyme exhibit type I binding spectra.) This alters the P450 redox potential making it more difficult for reductase to bring about the first electron transfer, as the iron redox potential of the P450 will be too high (Chapter 3). This factor, in addition to occupancy of the distal (sixth) ligand site, brings about an inhibition of the P450 (Ortiz de Montellano and Reich, 1986). It is found that cyanide (CN−) binds preferentially to P450 in its ferric (Fe3+) resting state as opposed to the reduced ferrous P450, and this is probably due to the fact that there is no electrostatic repulsion between the cyanide ligand and the heme iron, which will have a formal charge of zero due to the balance of three singly negatively-charged ligands (comprised from the thiolate -S− of cysteine and two negative charges from two of the porphyrin ring nitrogen atoms). This is in contrast with the case of reduced ferrous (Fe2+) P450 where the overall charge on the heme will be -1, thus disfavouring the binding of anionic ligands such as CN−. Furthermore, this effect also explains the stronger binding of CN− to ferric myoglobin as the heme iron will possess an overall charge of +1 due to the fifth (proximal) ligand being imidazole, rather than thiolate as it is in P450. Moreover, the binding of ionic ligands would be expected to be somewhat unfavourable in an essentially lipophilic (hydrophobic) environment, such as that encountered in the P450 binding site. Carbon monoxide, however, binds exclusively to reduced (Fe2 +) P450 and this is probably due to the change in electronic distribution of the iron caused by the addition of an electron to iron(III), with the unique role of the thiolate ligand being an important factor in the overall bonding description and orbital occupancy. Consequently, carbon monoxide (CO) acts both as a -donor and a good -acceptor ligand, with the additional electron (i.e. ferrous as opposed to ferric) being involved in back-donation to the CO ligand (Chapter 1). A similar effect probably operates in the binding of dioxygen to P450, as this also favours the


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iron(II) state with spin-spin coupling likely to be involved in the interaction between the two high-spin species (i.e. triplet oxygen and high-spin ferrous). In conclusion, it can be appreciated that a considerable number and variety of agents are able to induce or inhibit various P450s, either in general or selectively. It should also be considered, however, that at sufficiently high doses, inducers will also act in an inhibitory manner, frequently towards the same P450 isozyme; for example, benzocoumarins have been shown to be both inducers and inhibitors of CYP1 (Liu et al., 1993b). It is also known that inhibitors are able to induce P450s, depending on the concentration, as is found with SKF-525A (Bornheim et al., 1983).

6 Structural Models of P450s and Related Topics

6.1 Introduction There is a need for models of some kind in science, to aid in the conceptualization and understanding of what may be a continuum of complex processes describing an overall effect: the P450 field is no exception. In general, the previous chapters have been associated with experimental findings related to various aspects of P450 biophysical chemistry and metabolism. To some extent, it is hoped that the modelling of a number of key aspects of P450 structure and function could facilitate progress in our knowledge of the P450 system, and rationalize much of the biochemical data in terms of the fundamental interactions between the molecules involved. This final chapter of the book tries to illustrate how it is possible to utilize a number of recent technological advances in order to obtain a description of P450-mediated pathways at the molecular level. Molecular modelling has now become commonplace in the chemical and biological sciences due to an increased awareness of the potential utility of its various structural techniques, and to the recent advances in computer hardware technology and software development (Balbes et al., 1994). Novel chemical entities are now being developed routinely with the aid of molecular modelling systems in both academia and the pharmaceutical, agrochemical and other fine chemical industries. The various features of most molecular modelling software packages usually interface closely with the data produced from crystallographic and NMR spectroscopic studies, such that the visualization and manipulation of three-dimensional structures in real time is now a facile procedure (Lewis, 1994a). However, the construction of protein and enzyme models remains somewhat problematic in the absence of a sufficiently close crystallographic analogue, although NMR spectroscopy (Roberts, 1993) is now becoming a viable alternative as far as the structural studies of relatively small proteins are concerned. 6.2 P450 modelling In theory, there should be some form of structural explanation for the various phenomena associated with P450-mediated reactions, and modelling represents a possible means of achieving such molecular structural rationalizations for many aspects of the P450 system. Consequently, one might hope that molecular modelling could help to explain, for example, why certain chemicals are specific substrates for individual P450 isozymes, and whether there are any common 3-D structural characteristics which could facilitate the

STRUCTURAL MODELS OF P450S AND RELATED TOPICS

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prediction of P450-mediated metabolism using computational techniques. To some extent, there has been progress made in this area, and this has been achieved via the generation of structural models for both P450 substrate templates and of the enzymes themselves (see Table 6.1). Furthermore, some idea of the way in which the various components in the P450 macromolecular complex may interact can be obtained from structural modelling of these redox partners and it is also possible, to some extent at least, to obtain information about ligand-receptor binding interactions (Lewis et al., 1995e; Lewis and Lake, 1993; Lewis et al., 1994a) which could have relevance to the mode of P450 induction. In the absence of robust models of the relevant biomolecular structures, however, the various techniques of QSAR analysis remain popular procedures not only in the P450 field (Wolff et al., 1993), but also in other areas of biochemistry, toxicology and pharmacology for the rationalization of potency differences in series of structurally-related congeners (Lewis et al., 1995f; Lewis and Parke, 1995; Lewis, 1990; 1992a and b; 1994a; 1995b; Benigni and Giuliani, 1994). Due to the virtually exponential rise in the number of published protein sequences as opposed to the relatively small increase in number of protein crystal structure determinations, there is a general need for methods to enable the construction of protein models with a reasonable degree of accuracy in order to investigate biological processes at a mechanistic level. Although a large number (over 400) of P450s have had their cDNA sequences determined, there are at present only three† P450 crystal structures available, and all of these are from prokaryotic sources (Poulos et al., 1987; Ravichandran et al., 1993; Hasemann et al., 1994). In spite of this potential difficulty, many groups worldwide have investigated ways of modelling P450 active sites or, in some cases, the entire enzymes themselves, using X-ray crystal structures of the bacterial P450s, namely, P450cam (CYP101), P450BM3 (CYP102) and P450terp (CYP108). 6.2.1 Physical methods Protein crystallography has made an enormous impact on our understanding of the structure and function of enzymes, particularly in connection with their mechanisms of action. However, it is a popular misconception that the X-ray crystal structure of an enzyme or protein represents the ultimate reality for defining its physiologically relevant state. In fact, this is not the case, as proteins will be subject to dynamic motion within the structure itself due to thermal effects, and also from interactions with water molecules and the phospholipid milieu as well, if the protein is memTable 6.1 A Summary of various types of P450 models based on P450cam or substrate templates P450 Active Site

Whole enzyme

Substrate overlay

Molecular template

1A

Zvelebil et al., 1991 Lewis et al., 1994a Lewis 1995b

Lewis et al., 1986 Yagi and Jerina, 1982

Koymans, 1992 Tuck et al., 1993

2B

2C

Lewis et al., 1987 Lewis and Moereels, 1992 Lewis et al., 1987 Schwarze et al., 1988 Lewis and Moereels, 1992

Korzekwa and Jones, 1993

† A fourth prokaryotic P450 crystal structure has recently been published (Cupp-Vickery and Poulos, 1995) which is of P450eryF (CYP107).

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P450 Active Site

Whole enzyme

2D

Lewis, 1995b Lewis, 1995b

2E 3A 11A 17A 19A 51A 105

Koymans et al., 1993b Ackland, 1993 Lewis, 1987 Lewis and Moereels, 1992 Ackland et al., 1996 Lewis and Moereels, 1992

Lewis et al., 1994a Lewis, 1995b Ferenzy and Morris, 1989

Substrate overlay

Molecular template

Smith and Jones, 1992 Islam et al., 1991 Koymans et al., 1992 Strobl et al., 1993 Koymans et al., 1993b Ackland et al., 1996

Vijayakumar and Salerno, 1992 Laughton et al., 1990 Graham-Lorence et al., 1990 Laughton et al., 1993 Lewis and Moereels, 1992 Lewis, 1995b Ishida et al., 1988 Morris and Richards, 1991 Braatz et al., 1994

brane bound. There may also be interactions between small molecules and, moreover, with other proteins which may result in the formation of a complex assembly embedded within the phospholipid bilayer. However, the determination of a three-dimensional structure by X-ray crystallography represents an excellent starting point for investigations of the molecular mechanisms of proteins and enzymes: the cytochromes P450 are no exception in this respect. The crystal structure depicts a composite X-ray ‘image’ of the protein molecule in the solid state, usually containing water molecules resulting from the aqueous environment in which the crystallization process occurred (Creighton, 1993). The accuracy of this model, generated within the time frame of the experiment, will be determined by the resolution of the structure and a number of other factors pertinent to the crystallographic technique, including the interpretation of its results. Nuclear magnetic resonance (NMR) spectroscopy is becoming increasingly important as a complementary procedure to X-ray crystallography for the study of protein structures, and this can give information relating to a number of solution phase conformations corresponding with likely threedimensional structures experienced by the protein in an essentially aqueous environment. At present, however, NMR techniques are limited to relatively small proteins due to the difficulties involved in resolution and interpretation of the complex number of signals obtained from the NMR experiment. Nevertheless, methods are being developed for the generation of 3-D protein structures directly from a analysis of the high-resolution NMR data, and the reader is referred to a recent publication edited by Roberts (1993). Together with the visualization of NMR-derived conformations, molecular mechanics and molecular dynamics simulations of protein structures are now quite feasible with the current state of computer software and hardware technology. However, for relatively large proteins, X-ray crystallography remains the recognized technique for structure determination and, if one can assume (Black, 1993) that proteins from the same family bear a generally conserved tertiary fold, successful homology modelling of eukaryotic P450s from one or other of the three prokaryotic P450 crystal structures should be reasonably achievable (Hasemann et al., 1995).


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6.2.2 P450 sequence alignment The starting point for the generation of a protein or enzyme model is the production of an alignment between the sequence of the crystal structure template and that of the protein or enzyme which is to be constructed from it. In P450s it would appear that, of the three crystal structures determined so far, the P450BM3 (CYP102) hemoprotein domain probably represents the best overall template for producing models of mammalian P450s as it is generally of greater sequence homology than the other two prokaryotic structures, although there may be some localized variations in homology between them. It is also known that, in common with microsomal P450s, P450BM3 utilizes an NADPH-dependent P450 oxidoreductase as a redox partner, whereas the other two bacterial forms of known 3-D structure, namely P450cam and P450terp, employ iron-sulphur redoxins similar to those of mitochondrial P450s (Asperger and Kleber, 1991). This may, in fact, reflect the prokaryotic origins of these eukaryotic forms and certain eukaryotic cell organelles (Yang et al., 1985). Consequently, most of the sequence alignments listed herein have been based on P450BM3, although it is possible to map the P450cam (and/or P450terp) sequences onto these, as is shown in Figure 6.1, which also demonstrates that, by and large, the P450BM3 sequence is more appropriate (Lewis, 1995a). Although it is possible to derive sequence alignments of proteins across the P450 superfamily, it has been found that alignments within individual families and subfamilies is preferred (Lewis and Lake, 1995) as these appear to provide the more useful P450 model endpoints. However, an example of such an alignment between several P450s from different families and subfamilies is presented in Figure 6.1, as this illustrates certain similarities and differences so that comparisons can be made (Lewis, 1995a). It can be useful to include the P450cam and P450terp sequences together with P450BM3 in alignments between prokaryotic and eukaryotic P450 sequences, as the former are sometimes helpful in enabling decisions to be taken regarding the best overall fit within some regions where there is low homology (Hasemann et al., 1995). This can be particularly applicable to the N-terminal half of the sequence alignment where conservation of primary structure is normally low. It should be noted that the P450cam sequence is about 40 residues shorter than that of P450BM3, whereas the latter is roughly of equivalent length to the microsomal P450s, if one excludes the 20 or so residues which constitute the N-terminal membrane anchor. Moreover, the known substrates of P450BM3, which are long-chain carboxylic acids (Fulco, 1991), are very similar to those of the CYP4 family and, consequently, relatively high homology is found between P450BM3 and P4504A sequences, leading to what one might expect to be relatively valid models (Lake and Lewis, 1996). In fact, having constructed homology models, it is important to assess their overall geometric accuracy in relation to the ranges of bond lengths and angles allowable in known proteins. Based on the author’s own experience, it is recommended that all workers in this field employ some means of checking the validity of a protein model derived either from homology with a crystal structure, or by other means. The ProCheck package (which can be obtained from Oxford Molecular Ltd., Oxford, UK) is probably the most comprehensive system for assessing how well a new model conforms to protein stereochemical quality (Morris et al., 1992). This software comprises a number of complementary procedures for evaluating protein structures and identifies regions of the model which may require further refinement. There are many algorithms available to facilitate the alignment of protein sequences, and the GCG (Genetics Computer Group, Inc., Madison, Wisconsin) suite of programs, for example, contains a variety of software tools which enable sequence alignment to be carried out; although, in many cases, some visual editing is recommended because there can be specific patterns or motifs within sequences that may not be easily recognizable by purely computational methods. However, the combination of the CGEMA and

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VGAP packages developed at Janssen Pharmaceutica (Moereels et al., 1990) is able to reproduce fairly satisfactorily some of the sequence alignments which have been based primarily on additional experimental information regarding conserved positions and motifs (Lewis and Moereels, 1992). In P450 sequence alignments, however, there is relatively low homology between proteins from different families and, therefore, it is necessary to combine information from several experimental findings, together with additional indicators of protein substructural preferences (Lewis, 1995a; Schulz, 1988; Sternberg, 1986). Table 6.2 shows a comparison between sequence homologies for both P450BM3 and P450cam against a number of P450s from different families and subfamilies, based on the alignment presented in Figure 6.1, which shows that P450BM3 displays a higher Table 6.2 Sequence homology (%) between P450BM3 (CYP102), P450cam (CYP101) and a number of mammalian microsomal P450s CYP102 (461) Homology

CYP101 (414) Similarity

Homology

Similarity

(502) CYP3A4 27.1 52.1 16.7 45.9 (506) CYP4A4 26.7 54.4 17.4 49.0 (519) CYP4A11 26.0 53.6 11.6 36.2 (492) CYP2E1 24.9 53.6 17.6 47.8 (524) CYP1A1 24.9 54.2 14.5 42.3 (513) CYP1A2 24.3 50.5 16.4 45.7 (494) CYP2A6 22.6 54.0 18.6 46.6 (491) CYP2B1 22.3 52.1 17.6 49.5 (497) CYP2D6 20.8 51.2 19.3 50.2 (490) CYP2C9 20.2 51.6 17.4 48.8 (414) CYP101 19.3 48.8 — — Notes: The length of each sequence is given in parentheses. The per cent (%) homology columns relate to sequence identity whereas the per cent similarity columns include sequence homology plus identity. It should be noted that the eukaryotic P450s all possess additional N-terminal membrane anchor sequences of about 20 residues or more. If these are excluded (as has been done in the alignment in Figure 6.1) then the overall homologies will increase. The above values were calculated via the CGEMA suite of programs, and the kind assistance of Dr Luc Koymans from the Janssen Research Foundation is gratefully acknowledged.

overall homology with microsomal P450s than does P450cam. Moreover, it should be appreciated that, whereas the P450BM3 sequence is roughly similar in length to the eukaryotic P450s, the P450cam sequence is about 40 residues shorter with several large gaps in its alignment (Lewis, 1995a). The P450terp sequence is slightly ‘better’ than that of P450cam in terms of sequence homology with eukaryotic P450s, but is still approximately 20 residues shorter than that of P450BM3. The alignments presented here take into account a number of pieces of information derived from different sources which, when combined, tend to give rise to a largely self-consistent alignment that can be employed with some degree of confidence in model building (Lewis, 1995a). The various factors used to produce these alignments include the following: 1 Site-directed mutagenesis data relating to residues known to be involved in sub-strated binding, oxygen activation or electrostatic interactions with redox partners (Lewis, 1995a). Some of these are outlined in Figure 6.2 and are also referred to in the discussion of individual P450s modelled from P450BM3.

Figure 6.1 Alignment between CYP101, CYP102 and a number of microsomal P450s from families 1, 2, 3 and 4. References to amino acid sequences are as follows: CYP101—Unger et al., 1986; CYP102—Ruettinger et al., 1989; CYP4A4—Matsubara et al., 1987; CYP4A11—Palmer et al., 1993; CYP3A4 -Beaune et al., 1986; Bork et al., 1989; CYP2A6—Yamano et al., 1989a; CYP2B1—Suwa et al., 1987; CYP2C9—Kimura et al., 1987; CYP2D6—Gonzalez et al., 1988b; CYP2E1—Khani et al., 1987; CYP1A1—Sogawa et al., 1984; CYP1A2—Kawajiri et al., 1984.

STRUCTURAL MODELS OF P450S AND RELATED TOPICS 195

2 Experimental information on key basic residues associated with ion-paired interactions with redox partners, such as reductase, redoxin and cytochrome b5. Chemical modification of surface lysine residues in P450LM2 (CYP2B4), for example, identifies a number of key lysines which appear to be

196


important for electrostatic interactions with acidic residues on NADPH-dependent P450 oxidoreductase (Bernhardt et al., 1984, 1988, 1989; Bernhardt, 1993). 3 Antibody recognition sequences which indicate whether certain stretches of polypeptide are likely to reside on the surface of the enzyme and, consequently, are able to be recognized by specific antibodies (De Lemos-Chiarandini et al., 1987; Edwards et al., 1991, 1993, 1995).


197

4 Retention of secondary structural and super-secondary structural motifs with reard to turns between helices and -sheets. It is known that certain amino acids exhibit preferences for helices, sheets or turns, whereas the pattern of repeating hydrophobic residues varies between helices and sheets (Creighton, 1993; Eisenberg et al., 1984; Ptitsyn and Finkelstein, 1983). Such information, based on observations from proteins of known structure, greatly facilitates sequence alignment, especially when homology is fairly low, as is the case with P450s (Lewis, 1995a). However, it is found that there are

198


several motifs which tend to be conserved, by and large, throughout the superfamily and these are also of assistance in aligning the sequences. In fact, there is a 10–residue signature motif common to all P450s, but not encountered in any other proteins, which includes the invariant cysteine that ligates the heme iron. This consensus sequence is:


199

where x can be any residue, although there are preferences in some cases; for example, the first residue after F is either S, G, N or H, whereas the residue after C is either L, I, V, M, F, A or P (see Table 6.3 for amino acid codes). The first G can be D in some cases, whereas the last G can be also either A or D. Moreover, the R can also be either H, P or T, although R is by far the most common.

200


It is interesting to note that this signature motif is not entirely conserved in nitric oxide synthase (NOS) even though this enzyme bears some common characteristics to P450s (Degtyarenko and Archakov, 1993). It is possible, therefore, that NOS is not part of the P450 superfamily although it does show some similarities including, for example, a UV absorption maximum of the CO complex close to 450 nm, which is indicative of a heme-thiolate protein. 5 Substrate recognition sites (SRSs) are regions (Gotoh, 1992) where the homology tends to be rather low due to the fact that they will contain specific amino acid residue changes in order that the relevant active sites will be complementary with the substrates of different P450s (Lewis, 1995a). However,


201

202



203

204



Table 6.3 The 20 naturally-occurring amino acids (Reference: Creighton, 1993) Amino acid

Three-letter code

Single-letter code

Type of side chain

Arginine Lysine Aspartate

Arg Lys Asp

R K D

Basic Basic Acidic

Glutamate Asparagine Glutamine Serine Threonine Cysteine Histidine Tryptophan Tyrosine

Glu Asn Gin Ser Thr Cys His Trp Tyr

E N Q s T C H W Y

Acidic Polar Polar Polar Polar Weakly polar Weakly basic Aromatic, polar Aromatic, polar

205


Methionine Alanine Proline Glycine

Met Ala Pro Gly

M A P G

Hydrophobic Aliphatic, hydrophobic Aliphatic, hydrophobic Neutral Figure 6.2 Site-directed mutagenesis and crystallographic studies on P450s. (Dashed lines indicate allelic variants)

206

even in these areas some similarities emerge and the results of site-directed mutagenesis experiments have assisted in defining the appropriate complementary residues pertaining to substrate binding and recognition. For example, several P450s have been mutated in SRS1 (Figure 6.2) and it is likely that the residue(s) in contact with substrates will be hydrophobic. SRS2 contains the GERL motif in the CYP11 family, and this is known to be associated with steroid recognition (Gotoh et al., 1985). In P450BM3, the LI81 residue in this region (helix F) points directly towards the heme and this appears to be of


207

particular importance in substrate binding and recognition, as P450s showing a specificity towards aromatic substrates all possess phenylalanine at this position, whereas others tend to have aliphatic hydrophobic amino acid residues, such as I or L (Figure 6.1). There are similar findings for the other SRSs which will be discussed elsewhere in this chapter. 6 Substrate specificities of different P450s facilitate alignment decisions in the regions of low conservation associated with the putative active sites, and tend to produce some degree of selfconsistency in the homology modelling based on such alignments (Lewis, 1995a). This procedure appears to be quite effective both at a general level, for rationalizing broad substrate preferences, and within families and subfamilies where subtle changes appear to be responsible for small, but crucial, alterations in substrate regio- and stereo-specificity (Lewis, 1995a). 6.2.3 General features of P450 crystal structures and sequences At this point it is useful to consider the crystal structures of P450s and their sequences, in conjunction with alignments between the prokaryotic isozymes and eukaryotic forms. It is generally accepted that the P450s all possess a largely conserved tertiary structural core of amino acid residues (Lewis, 1995a) which represents the key structural and functional elements characteristic of P450 isozymes, and inspection of the three bacterial P450 crystal structures (Hasemann et al., 1995) tends to confirm this, although there are some differences which reflect the biological source and environment, including redox and membrane interactions (Lewis, 1995a). Consequently, sequence alignment and visual comparison between the crystal structures can highlight similarities and differences, such that it is possible to investigate the extent to which primary sequence homology is reflected in the conservation of secondary and tertiary structure (Lewis, 1995a). Many of the regions of peptide which comprise the P450 structural core are situated within the Cterminal portion of the sequence from the I helix onwards, particularly as far as the L helix, and this contains a number of highly conserved motifs, including the 10–residue P450 signature mentioned previously. The N-terminal region tends to exhibit lower homology overall than the C-terminal, although there are some stretches of relatively high conservation for either functional or structural reasons (Lewis, 1995a). Figure 6.3 shows a comparison between the C tracings of the three prokaryotic P450 crystal structures, and an alignment of these sequences indicates that the primary and secondary structures remain essentially conserved between these three bacterial forms (Hasemann et al., 1995). In fact, 36 amino acids show identity for the three sequences, and Figure 6.4 indicates the spatial distribution of these residues within the P450BM3 (CYP102) structure. It is likely that a reasonable proportion of these residues will be conserved across the P450 superfamily, especially those which are specifically relevant to the catalytic functionality of the enzymes such as the binding of heme, redox partners, for oxygen activation, and for folding of the polypeptide chain. However, it is important to show how a description of the key elements of the crystal structures may be integrated with the primary sequences of other P450s, and this will now be described, by starting at the N-terminus and progressing along the sequences and structures. The reader is referred to the alignment presented as Figure 6.1 for an appreciation of the various aspects under discussion, whereas Tables 6.4, 6.5 and 6.6 show the secondary structural elements in the three bacterial crystal structures. In eukaryotic P450s, the first 20 or more residues (not shown in the alignment presented as Figure 6.1) at the N-terminus tend to be generally hydrophobic in character, and it is thought that this segment represents a transmembrane anchor for the protein, which may also have a role in inter-P450 and reductase interactions (Finch and Stier, 1991). It is likely that this part of the sequence is essentially helical in nature, possibly

208


comprising two helices separated by a turn, which could enable the N-terminus to loop in and out of the phospholipid bilayer (Bernhardt et al., Table 6.4 Substructural motifs within the P450cam structure (Reference: Lewis, 1995a) Helix

Residue range

A B BND BPR C D E F G H I J K 310 L

G37-Q46 R67-D77 D77-F81 P89-Y96 P106-K126 L127-Q145 N149-L169 D173-T185 T1192-K214 D218-N225 T234-S267 S267-Q276 R280-F292 K324-N328 G359-I378

Sheet (1) (2) 5 (1) 5 (2) 5 (3) 2 (1) 2 (2) 3 (1) 3 (2) 4 (1) 4 (2) 1

1

Residue range

Turn

Residue range

D52-C58 G60-T66 G146-F150 I395-V405 S382-S397 G226-V228 G230-I233 V295-L301 G315-M323 Y305-H308 V310-L312

T1 T2 T3 T4 T5 T6 T7 T8 T9 T10 T11 T12 T13 T14 T15 T16

P15-V18 D27-N30 N33-A36 E47-V50 C58-G61 F98-S102 D104-Q108 P170-D173 V228-R231 R277-I281 F307-V310 N328-E331 C334-H337 T348-G351 G353-L356 A384-A387

Table 6.5 Substructural motifs within the P450BM3 structure (Reference: Lewis, 1995a) Helix

Residue range

A A1 B B’ C B1 C1 D E D1 F G H I J J’ K

N16-L20 P25-G37 S54-C62 S72-G83 E93-L104 P105-S108 Q109-K113 G114-R132 P142-F158 N163-R167 P172-Q189 A197-S226 L233-N239 D250-K282 N283-L298 S304-Q310 L311-W325

Sheet (1) 1 (2) 1 (3) 1 (4) 1 (5) 2 (1) 2 (2) 3 (1) 3 (2) 3 (3) 4 (1) 4 (2)

1

Residue range E38-A44 R47-S53 G350-L356 P329-A335 R66-N70 E337-L341 E344-E348 H138-V141 F444-K451 H420-E424 D432-T436 T438-E442


Helix

Residue range

K’ E1 F1 L

I357-H361 R375-F379 N381-I385 G402-K419

Sheet

209

Residue range

1989; Sakaguchi et al., 1994). This N-terminal membrane anchor ends with a short sequence of basic residues, prior to an alternating polyproline motif (P x P x P) which is fairly well conserved between both eukaryotic and prokaryotic P450s, although P450terp (CYP108) appears to possess two alternating alanine residues instead of the usual prolines at this point (Hasemann et al., 1994). Various conserved motifs in P450 sequences, discussed in this section, are summarized in Table 6.7. The folding pattern of the peptide is different between P450cam (CYP101) and P450BM3 (CYP102) in this section and this is reflected in their sequences, with the latter being a closer match to the eukaryotic P450s (Lewis, 1995a). The region from the polyproline motif to the highly conserved tryptophan (W96 in P450BM3) is regarded as representative of a Rossman domain (Tretiakov et al., 1989), and this consists of three -helices (A, B and B’) and three strands of a -sheet ( 1) with the conserved tryptophan being present close to the start of the C helix (Ravichandran et al., 1993). The section which immediately follows the polyproline motif in P450BM3 (CYP102) tends to be a short stretch of 310 helix, and this appears to ‘match up’ with a complementary strand at the end of helix F, about 170 residues downstream. Both of these regions are recognized by antibodies in CYP2B1 (De Lemos-Chiarandini et al., 1987) and, therefore, may be associated with either membrane interactions and/or substrate recognition and access (Lewis, 1995a). Although there is some contact between these surface portions in the P450BM3 crystal structure via both polar and hydrophobic residues, the fact that there are complementary conserved segments recognizable by antibodies suggests that this region lies on the Table 6.6 Secondary structural elements in CYP108 (Reference: Hasemann et al., 1994) Helices A A3 A B B B3 C C C3 D E E D3 F G

-sheets E9-I16 Q19-A22 D24-E37 H57-K65 Q82-I91 L102-S104 P108-L118 P123-I126 R127-L129 E130-L145 F153-D156 Y160-L170 E174-D176 E177-Q185 A208-R231

(1) (2) 1 (3) 1 (4) 1 (5) 2 (1) 2 (2) 3 (1) 3 (2) 3 (3) 4 (1) 4 (2) 5 (1) 5 (2) 1 1

P39-A43 P50-T55 G335-S341 F317-L322 L69-S71 T325-V327 Q330-I332 G149-F153 V421-A428 K397-S404 R408-V410 G417-N420 S245-L247 N250-I252

210


Helices

-sheets

H V238-N244 I D254-R285 J P287-S295 E3 P297-L299 K I300-T311 K Y342-N346 L Q380-L394 F3 L395-K397 3 denotes 310 helix

surface in the microsomal P450s (Lewis, 1995a). This section of peptide leads into the A helix which, although well conserved between P450BM3 and P450terp, does not exhibit much homology across other members of the superfamily, apart from a number of exceptions; the pattern of hydrophobic residues is, however, a good indicator of a helical conformation and these do show conservation, being repeated every four or five residues (Lewis, 1995a). The A helix terminates in a ‘helix-breaker’ residue which is usually glycine, but can also be proline in some cases (Figure 6.1). This conserved residue marks the start of the first strand of a -sheet ( 1) which terminates with a highly conserved glycine, after a sequence of seven or eight intervening residues that is punctuated by a conserved pattern of hydrophobic side chains (Figure 6.1). The first strand of 1 is also recognized by antibodies in the CYP2B1 enzyme and, therefore, as is the case with the bacterial crystal structures, this segment is likely to be conserved as a surface region across the superfamily (Lewis, 1995a). Following the glycine which marks the turn into the second strand of 1, an arginine residue (R47) is present in P450BM3 which is thought to be involved in ion-pairing to the long-chain carboxylic acid substrates of this enzyme (Lake and Lewis, 1996; Graham-Lorence et al., 1994). The equivalent region in the eukaryotic P450s shows a sequence of hydrophobic residues at the end of this second strand of -sheet, and a conserved glycine (or proline) indicates the start of the B helix which contains a recognizable motif, Table 6.7 Motifs in P450 sequences Residue motifs

Position in structure

PxPxPxP G 7 or 8 residues G

At end of N-terminal anchor At first strand of 1

KEAL WKxxRRxS GERL R K 7 or more residues D G x D/E T KxxEE ExxR FSxGxRxCFG

At end of B helix In C helix In F helix (of P45011 family) In G and H helices In I helix In J helix In K helix At start of L helix (P450 signature motif)

The region containing these motifs comprises aRossman domain


Residue motifs

211

Position in structure

ELD At first strand of 4 x=any residue. Notes: Most of the helical segments are highly conserved between the three bacterial P450 crystal structures, in terms of their spatial orientation, with four pairs of helices in contact, namely, D and E, F and G, I and L, J and K helices. Of these, helices D, E, I and L are regarded as core helices whereas the E, F, G and I helices form a Greek key helical bundle. The 10-residue signature motif, which contains the invariant cysteine, shows some degree of variation but the changes are usually conservative, and this is generally the case for the other motifs listed above.

KEAL, in some of the microsomal forms (present as KEAC in P450BM3 and REAY in P450cam) which is usually preceded by one or two hydrophobic residues (Figure 6.1). The conserved basic residue (i.e. Lys or Arg) could be involved in redox partner interactions, as it has been shown that this is one of the four points of electrostatic interaction between P450cam and either putidaredoxin or cytochrome b5 (Stayton et al., 1989; Stayton and Sligar, 1990). The following region, which comprises the B’ helix is not well conserved between the various sequences and is variable in both length and, consequently, conformation in the prokaryotic crystal structures, being unusually long in P450terp (Hasemann et al., 1994). This region represents the first of the substrate recognition sites, SRS1, and is thought to reflect the variability in substrate specificities between different P450s. However, the usual ‘helical’ pattern of hydrophobic residues is generally retained and this assists sequence alignment, with P450BM3 representing the overall best match with most eukaryotic forms (Figure 6.1). P450terp has an additional segment, not present in either P450cam or P450BM3, which does not map favourably with the eukaryotic P450 sequences; whereas a glycine at the end of the B’ helix in P450BM3 is fairly well conserved within the microsomal enzymes’ sequences (Figure 6.1). In fact, the G x G motif at the end of this helix appears to show a commonality between P450BM3 and several of the eukaryotic isoforms (Lewis, 1995a). This is followed by a short stretch of hydrophobic residues which are likely to be involved in substrate binding as has been indicated by site-directed mutagenesis experiments (Figure 6.2); most of these match up with F87 in P450BM3 and this points directly over the heme in the crystal structure (Ravichandran et al., 1993). This phenylalanine is followed by a threonine and serine in P450BM3, which is identical with the corresponding sequences of P450cam and P450terp. The start of the C helix is marked by a conserved glycine (or proline) although P450BM3 shows a glutamate at this point (Figure 6.1). The C helix constitutes a highly conserved section which is likely to be surface-exposed for interaction with redox partners (Lewis, 1995a). It contains a number of basic residues, one of which forms an ion-pair with one of the heme propionates in all three bacterial crystal structures, and this is usually an arginine in most P450s, although it is conserved as histidine (H100) in P450BM3. The corresponding arginine (R112) in P450cam is the second of the four basic residues which appear to be associated with electrostatic ion-paired interactions with either putidaredoxin or cytochrome b5 (Stayton et al., 1989; Stayton and Sligar, 1990) and, therefore, may be evolutionarily conserved for redox interactions within other P450s. This basic residue lies four positions downstream from a highly conserved tryptophan (W96 in P450BM3) which is though to be involved in electron transfer between reductase and the heme moiety (Munro et al., 1992; Lewis, 1995a). In P450BM3, these substructures are in close contact, with the nitrogen of the tryptophan indole ring being only about 3 Å from one of the heme propionates, indicative of a possible hydrogen-bonded electron conduit to transfer charge from reductase to P450 (Lewis, 1995a). Tryptophan-96 is a surface residue and could, therefore, readily interact with the electron transfer flavin component of a bound reductase. The crystal structure of flavocytochrome b2 (Xia and Mathews, 1990)

212


seems to represent a close analogue of this type of interaction, since this protein contains both heme- and flavin-binding domains with, in this case, a tyrosine residue bridging the two prosthetic groups via a hydrogen-bonded interaction. However, a crystal structure of the complex between cytochrome c peroxidase and cytochrome c shows a tryptophan grouping intervening between the heme moieties of the two redox-coupled hemoproteins (Pelletier and Kraut, 1992). It is likely, therefore, that at least one conserved aromatic residue is involved in electron transfer to P450 from its redox partner(s), possibly via - stacking with another delocalized system on the redox partner. A 10-residue stretch following the conversed tryptophan in CYP2B1 is recognized by antibodies (De Lemos-Chiarandini et al., 1987) and contains the serine phosphorylation site which forms part of the protein kinase A substrate recognition motif, R R x S, where the first arginine is the highly conserved heme-binding residue mentioned previously. Molecular modelling of CYP2B1 at this point, by phosphorylating the serine residue, shows that the phosphate moiety will interfere with the arginine-heme propionate ion-pairing, such that the heme binding interaction with the arginine is likely to be weakened by preferential ion-pairing to the phosphorylated serine (Figure 6.5). Presumably, this provides some explanation for the mechanism of P450 degradation via conversion to the inactive form, P420, and subsequent loss of heme (Jansson, 1993; Jansson et al., 1990). It has been shown that cytochrome b5 inhibits serine phosphorylation (Epstein et al., 1989) by binding to this region, presumably with the conserved arginine, and the enzyme models show that this serine lies on the surface in the CYP2 family, although the position is conserved as threonine in CYP4, and as serine in P4503, but shifted one residue downstream; whereas, in the CYP1 family, there is a serine at a further six positions downstream (Figure 6.1). The C helix is fragmented into more than one stretch of helix in both P450BM3 and P450terp, but there is close correspondence between the two, with the final helical segment terminating in a lysine (K113 in P450BM3) which is largely conserved in other P450s, apart from those of family CYP1. In the CYP2B4 orthologue, however, it has been demonstrated that this conserved lysine residue is involved in the binding of reductase to P450 via electrostatic interactions (Bernhardt et al., 1988; 1989) and, consequently, this position is probably relevant to the binding of reductase in other P450s. A conserved glycine (or proline) indicates the start of the D helix which is fairly hydrophilic in character, although the usual repeating pattern of hydrophobic residues is observed (Figure 6.1). This helix is in contact with the E helix in the bacterial crystal structures, and these comprise half of the P450 common core of four helices, which is completed by the highly conserved helix I and helix L. The first strand of the 3 sheet, containing a fairly well-conserved acidic residue (El40 in P450BM3) leads into the start of the E helix, which is itself part of a Greek key helical bundle (Poulos, 1986) consisting of the four helices E, F, G and I, that may constitute an evolutionarily conserved globin fold. The pattern of hydrophobic residues in the E helix is clearly recognizable and includes a highly conserved (almost invariant) isoleucine residue (I153 in P450BM3) that seems to pair with another hydrophobic side-chain in helix I (L262 in P450BM3). In fact, it appears that hydrophobic interactions are a common feature of interhelical contacts in the internally facing sections of several P450 helices, of which helices I and L are the most striking examples (Lewis, 1995a). Helix E terminates in a phenylalanine residue (F158 in P450BM3) which is well conserved in most P450s, and this leads into a sequence of residues linking up with the F helix. Although this portion does not appear in either P450cam or P450terp, it is present in P450BM3 with the ‘gap’ being filled by the D1 helix (Ravichandran et al., 1993). A conserved RF motif, which is common between P450BM3 and most of the eukaryotic P450s, precedes the short stretch of D1 helix and this seems to pair with complementary residues in helix D (e.g. D121 in P450BM3). The F helix which follows D1 is variable in sequence between P450s and constitutes SRS2, as it is thought to be involved in substrate interactions (Gotoh, 1992). Sequence alignment of this region is facilitated by the conserved pattern of hydrophobic residues, however, and site-directed mutagenesis


213

Fig. 6.3a

Fig. 6.3b

(Lindberg and Negishi, 1989) has shown that the position corresponding to L181 in P450BM3 is important for defining substrate specificity in the CYP2A subfamily (Lewis and Lake, 1995). Table 6.8 summarizes the results of site-directed mutagenesis in various P450s. In P450BM3, the L181 residue points directly into the heme binding pocket and it can be shown that long-chain fatty acid substrates are likely to form hydrophobic contacts with the complementary side chain of this leucine (Lake and Lewis, 1996). This sequence position, which appears to be conserved as a hydrophobic residue, exhibits a change to phenylalanine in those P450s that exhibit specificity for aromatic substrates, some of which may possess planar molecular geometries (Lewis, 1995a). In P450s of the 11A and 11B subfamilies, a leucine residue at this position denotes the final residue of the steroid recognition motif, GERL (Gotoh et al., 1985), and this may be relevant to the specificity of isozymes of the CYP11 family for endogenous steroid substrates. In the P450cam crystal structure, a methionine residue (M184) at an analogous position is close to the bound substrate, camphor, and a comparison between the substrate-bound and substrate-free crystal structures indicates that there is thermal fluctuation within this region which is consistent with substrate access (Poulos et al., 1986). However, in the P450terp crystal structure, a segment of sequence following the F helix is so subject to disorder that the electron density map could not be interpreted for residues 191 to 207 (Hasemann et al., 1994). As the P450cam structure shows a shortened F helix and the sequence alignment indicates a large gap with respect to the other P450s, the P450BM3 structure remains the only template structure for modelling this region effectively and the second half of the F helix, in particular, shows a number of interesting changes which point to possible substrate interactions and recognition sites in

214


Fig. 6.4

Fig. 6.5

Fig. 6.7

eukaryotic P450s (Lewis, 1995a). The region comprising the end of the F helix to the start of the G helix is recognized by antibodies in CYP2B1 and, therefore, probably lies on the surface of the protein, which is in agreement with evidence shown in the prokaryotic P450 crystal structures. A lack of homology in most of the intervening residues between these two helices (i.e. F and G) may reflect the difference in environments for prokaryotic and eukaryotic microsomal P450s, although a possible role for this segment in substrate recognition cannot be ruled out.


215

Fig. 6.8

Fig. 6.9

Fig. 6.11

Helix G contains a number of fairly well conserved acidic and basic residues which, together with albeit relatively few hydrophobic markers, enables alignment with the bacterial crystal structure sequences (Figure 6.1). Towards the end of this helix there is a pattern of repeating basic residues which may have two functions, namely, binding to reductase and ion-pairing to an invariant aspartate at the start of helix H (Lewis, 1995a). The ion-pair, exemplified by R223 and D232 in P450BM3, is also present in the other two prokaryotic P450 crystal structures and is, therefore, likely to be of structural importance. In CYP2B4, a

216


Fig. 6.12

Fig. 6.14

Fig. 6.15

lysine residue one position upstream of that corresponding to R223 has been linked with reductase binding using chemical modification experiments (Bernhardt et al., 1988, 1989), and there is a striking conservation of two or three basic amino acids in this region across the P450 superfamily, which suggests that it is a point of contact for both redox partner interaction and for retention of the tertiary fold (Lewis, 1995a). In P450BM3, the second of these two basic residues (K224) forms an ion-pair with an aspartate (D251) at the start of helix I, and there is general conservation of this pattern in other P450s, although the position may


217

Fig. 6.17a

Fig. 6.17b

Fig. 6.18

shift by one residue in some cases. As the I helix leads directly to the distal heme face where oxygen activation and single oxygen atom insertion into substrates occurs, and the basic residues at the end of the G helix may be associated with redox interactions, it is tempting to speculate that the binding of a redox partner at this point may initiate either electron and/or proton transfer via a hydrogen-bonded conduit along the I helix itself (Lewis, 1995a).

218


Fig. 6.20

Fig. 6.21

Fig. 6.23

The H helix is well conserved, especially between the prokaryotic crystal structure sequences, and the pattern of hydrophobic residues together with the invariant aspartate map closely onto the eukaryotic P450 sequences (Figure 6.1). Antibodies raised against CYP1 are able to discriminate between CYP1A1 and CYP1A2 within this H helix region and the variable loop which follows it (Edwards et al., 1993), so it is likely that this stretch of sequence is on the enzyme surface (Murray et al., 1993) and could, therefore, provide a means of distinguishing between different P450s via specific antibody binding. The fact that the


219

Fig. 6.24

Fig. 6.26

Fig. 6.27

interaction with antibodies within this region inhibits P450 activity (Edwards et al., 1993) gives some support to the view that there may be key redox partner interactions or substrate recognition sites along this so-called ‘hinge’ between the two major domains of the P450 structure. The I helix, which is preceded by a conserved hydrophobic residue (L249 in P450BM3), is the longest segment of secondary structure and stretches the entire width of the enzyme in the bacterial crystal structures (Figure 6.3). This highly conserved, generally hydrophobic helix is distal to the heme and contains a

220


Fig. 6.29

Fig. 6.30

Fig. 6.31

substrate recognition site prior to the proposed oxygen binding pocket (Poulos, 1986) and invariant threonine (T268 in P450BM3). The conserved motif G x D/E T ensures a straightforward alignment of P450 sequences in this region, and this is followed by a group of hydrophobic residues after a cluster of threonines and serines that are conserved between P450BM3 and eukaryotic P450s (Figure 6.1). The hydrophobic region towards the end of the I helix is in contact with a complementary portion of the L helix, which is at the proximal heme face. The orientation of these two helices is suggestive of a possible


221

Fig. 6.32

Fig. 6.33

Fig. 6.34

scissoring conformational change (Veitch and Williams, 1992), which could occur following substrate binding and may be associated with the initiation of reductase interaction leading to electron transfer and oxygenation. It is likely that two strands of -sheet ( 2) which link the putative substrate binding site with a known point of interaction (Bernhardt et al., 1988, 1989) with reductase (corresponding to K349 in P450BM3) may also undergo a conformational scissoring motion with the I helix when a substrate binds, and this could trigger reduction of the heme iron.

222


Fig. 6.35

Fig. 6.36

Fig. 6.37

Site-directed mutagenesis of several residues in different P450s within the I helix have provided strong indications of the important roles of some of these in both substrate binding and oxygen activation (Figure 6.2). In fact, in addition to the invariant threonine (T268 in P450BM3) the previous residue, which is usually either glutamate or aspartate, appears to act in concert with the threonine in shuttling protons to oxygen when it is bound to the heme iron, and it is possible that ion-pairing to basic residues within the heme pocket may facilitate this process (Gerber and Sligar, 1992, 1994). It is thought that the sequence of


223

Fig. 6.38

Fig. 6.39

Fig. 6.40

threonine and serine residues following the conserved acidic residue (E267 in P450BM3) could be associated with hydrogen-bonded interactions between a number of bound water molecules which may be present along a channel within one corner of the substrate binding pocket (Raag and Poulos, 1992). Several water molecules are observed in this region of the P450 crystal structures and appear to form a hydrogen bonded

224


channel, which could represent a conduit for the transfer of protons from the aqueous environment to the distal heme face (Raag and Poulos, 1992). At the end of the I helix, a highly conserved proline signifies the start of the J helix which contains a probable ion pair made from K289 and E292 in P450BM3 that forms part of a motif, K x x E E. The system is not conserved in either P450cam or P450terp, however and the helix is shorter in these latter two structures. Between the J and K helices there is a stretch of sequence in the eukaryotic P450s which is absent in both P450cam and P450terp, but present and fairly well conserved in P450BM3 as the J’ helix. This usually starts with a proline and, together with the first half of the K helix, contains an antibody recognition sequence in CYP3A (Leeder et al., 1994). The K helix (which is in contact with helix J) contains a conserved tyrosine residue (Y313 in P450BM3) that is absent in the P450cam and P450terp sequences, and a pattern of hydrophobic amino acids leads into another highly conserved motif, E x x R, which exhibits ion pairing between the oppositely charged glutamate (E320 in P450BM3) and arginine (R323 in P450BM3) residues. It is not known, however, whether this motif is purely structural or has a relevance to redox partner binding. The end of the helix K leads into a turn that represents a likely substrate recognition site (SRS5) as it is within the heme pocket of each prokaryotic P450 crystal structure, and contains a conserved hydrophobic residue (A328 in P450BM3). Site-directed mutagenesis studies (Figure 6.2) have identified key residues within this region that appear to be relevant to substrate interactions and a conserved proline (P329 in P450BM3) aids alignment with most eukaryotic P450s, although this is not present in either P450cam or P450terp. Three sections of -sheet connect this proline to a highly conserved basic residue, usually lysine (K349 in P450BM3) that has been shown to bind reductase in CYP2B4 (Bernhardt et al., 1984, 1988, 1989). The first of these -strands is associated with substrate binding, as indicated by site-directed mutagenesis, and usually ends with a basic residue which is largely conserved between prokaryotic and eukaryotic P450s, but is absent in P450BM3 (Lewis, 1995a). In the other two bacterial P450 crystal structures, this residue is an arginine (R299 in P450cam) which ion-pairs with the second of the two heme propionates (bonded to pyrrole ring D). The following two strands of the 2 sheet include a number of relatively well conserved residues that facilitate sequence alignment and a motif, comprising D T x x x G, is a common feature (Figure 6.1). Another motif, P K G, signifies the end of the second strand of 2 and the start of another strand of 1 which is characterized by a number of hydrophobic residues. The relevance of the central lysine (K349 in P450BM3) to this motif (which is on a surface-exposed turn in the bacterial P450 crystal structures) for interaction with either reductase or another redox partner has been mentioned previously and there is compelling evidence that a basic residue at this point is essential for reductase binding (Bernhardt, 1993). After the -strand, a short helical segment that is present in all three P450 crystal structures ends in a generally conserved basic residue followed by an acidic residue, with an aromatic amino acid four positions downstream (Figure 6.1). This region is an antibody recognition site in CYP2B1 (De Lemos-Chiarandini et al., 1987) and, presumably, lies on the surface in all P450s. Another motif follows this segment which comprises the sequence F x P E R F, and is well conserved between P450BM3 and the eukaryotic sequences (Lewis, 1995a). The basic residue of this motif in the short E1 helix may be a redox partner binding point, although the two oppositely charged residues (E377 and R378 in P450BM3) form an ion-pair and could, therefore, be of a structural nature. A second short piece of helix, F1 in P450BM3, follows E1 and although this is not very well conserved between the P450 sequences, it includes a lysine residue which binds putidaredoxin and cytochrome b5 in P450cam, namely, K344 (Stay ton et al., 1989; Stay ton and Sligar, 1990). A variable length of peptide sequence, which is better defined in the P450BM3 structure with respect to the eukaryotic P450s than in the other two bacterial P450s, leads into the highly conserved 10-residue P450


225

signature motif beginning with an invariant phenylalanine (F393 in P450BM3). This P450 signature sequence contains two well conserved glycines (G396 and G402 in P450BM3) together with the invariant cysteine (C400 in P450BM3) which is essential to P450 catalytic activity and ligates the heme iron via a thiolate functionality (Hawkins and Dawson, 1992). The following residue tends to be hydrophobic (1401 in P450BM3) and this leads into the L helix which begins with the second highly conserved glycine (Figure 6.1). The invariant phenylalanine at the start of the 10-residue signature appears to form a stacking interaction with the porphyrin ring system of the heme moiety and it is possible that this phenylalanine could be involved in electron transfer from a redox partner to the heme (Yasukochi et al., 1994). A similar interaction has been observed between cytochrome c peroxidase and cytochrome c (Beratan et al., 1992; Pelletier and Kraut, 1992) where a tryptophan residue appears to form an electrontunnelling pathway between the two heme structures. There is also a conserved basic residue in this region (R398 in P450BM3) which may be associated with either an electrostatic interaction with a heme propionate and/or with a redox partner (Lewis, 1995a). Table 6.8 Site-directed mutagenesis studies on various P450s* CYP

Residues modified

References to mutagenesis experiments

Analogous residues in CYP 102

1A2

Glu318, T319‡

Glu267, T268

2A4, 5

Alall7, Leu209, Met365

2B1

I1e114, Thr302, Val363 and 367, Gly478

2C1–5

Val113, Thr301, Ile359, Ser364

2D1, 6

Asp301, Ile380

2E1 11B1 19A1

Thr303 Phe66, Ser126 Glu302, Ala307, Pro308

21A1

Val281, Cys428

51A1 101A1

Gly310 Asp251, Thr252

102A1

Arg47, Phe87, Trp96

Shimizu et al., 1994; Furuya et al., 1989a and b; Tuck et al., 1993a; Hiroya et al., 1992 Lindberg et al., 1992; Lindberg and Negishi, 1989; Iwasaki et al., 1993; Juvonen et al., 1991 Aoyama et al., 1989; Kedzie et al., 199la; Halpert and He, 1993; He et al., 1994 Hsu et al., 1993; Straub et al., 1991; Imai et al., 1989; Imai and Nakamura, 1988, 1989; Kronbach and Johnson, 1991; Kronbach et al., 1989, 1991 Ellis et al., 1994; Matsunaga et al., 1990a Fukuda et al., 1993 Mathew et al., 1990 Graham-Lorence et al., 1991; Zhou et al., 1991 Wu et al., 1991; Wu and Chung, 1991 Ishida et al., 1988 Gerber and Sligar, 1992, 1994; Martinis et al., 1984; Atkins and Sligar, 1989; Stayton and Sligar, 1989; Davies and Sligar, 1992; Imai et al., 1989a Graham-Lorence et al., 1994; Munro et al., 1994

Phe87, Leu18l, Ala330

Phe87, Thr268, Ala328, Ser332, Leu439 Phe87, Thr268, Trp325, Phe331

Thr260, Thr327 Thr268 Tyr51, Glu109 Thr260, Gly265, His266 Ile254, Cys400 Thr260 Glu267, Thr268

Arg47, Phe87, Trp96

226

CYP


Residues modified

References to mutagenesis experiments

Analogous residues in CYP 102

* The majority of key positions where mutations have been carried out in the CYP2 family are shown in Figure 6.2 and mapped onto the alignment with CYP102 and CYP101 (Figure 6.1). ‡ A number of other distal and proximal residue positions were also mutated (Furuya et al., 1989a and b).

In P450cam, the conserved basic residue (H355) could constitute a central grouping in a charge relay system linking the heme propionate with a serine residue (S83 in P450cam) which may represent a hydrogenbonded electron conduit as a network of hydrogen bonds (Lewis, 1995a) extends from the heme propionate to the conserved surface lysine (K314 in P450cam) shown to be analogous with a site of electrostatic pairing to reductase (Bernhardt et al., 1988, 1989) mentioned previously. The distance between the heme and this lysine residue (25.4 Å) is virtually identical (25.2 Å) to that of FAD and a surface aspartate (D123) in the crystal structure of ferredoxin reductase (Lewis, 1995a), which suggests that ion-pairing between P450 and reductase orientates the electron transfer groupings in the two structures such that they are in close proximity, thus facilitating electron transfer from reductase to P450. The second half of the L helix contains a number of hydrophobic residues which are in contact with a complementary stretch in the I helix, and these two helices ‘sandwich’ the heme on the proximal and distal faces, respectively. Prior to this hydrophobic section of the L helix is a conserved glutamate (E409 in P450BM3) which could assist in the attraction of water molecules into the hydrophobic channel that links the heme environment with the enzyme surface. In the P450cam crystal structure, an analogous glutamate (E366) has been shown to participate in a hydrogen-bonded assembly of water molecules which extend from the invariant distal threonine (T252 in P450cam) to the external aqueous environment (Raag and Poulos, 1992). There is a basic residue two positions upstream of this glutamate in both P450cam and P450terp, but it is absent in P450BM3, indicating that it is probably involved in redox partner interactions in the former bacterial P450s. In P450cam, this residue (R364) has been shown to occupy a position that can both bind putidaredoxin or cytochrome b5 (Stayton et al., 1989; Stayton and Sligar, 1990), and a similar basic amino acid is present at an analogous point in some of the aligned eukaryotic P450 sequences (Figure 6.1). Following the L helix, the P450 crystal structures contain a number of -sheet strands that are fairly well conserved between the bacterial forms but the overall alignment with eukaryotic P450 sequences is impeded by a lack of homology except from a small number of moderately conserved regions. In general, the P450BM3 template displays the most homology with the microsomal P450s along this final stretch of the protein sequences. In particular, the first strand of -sheet ( 3) following the L helix contains a conserved alternating pattern of hydrophobic residues and this is largely mirrored in a similar short -strand ( 4) which is preceded by a fairly well conserved motif, ELD, where the first of the two acidic residues is at position 430 in P450BM3 (Figure 6.1). Between the two strands of the 4-sheet, a turn residue lies within the heme pocket in all bacterial P450 crystal structures (L437 in P450BM3) and is usually hydrophobic in character. This stretch corresponds to the final substrate recognition sequence (SRS6), and site-directed mutagenesis (Figure 6.2) has shown that it probably contacts substrates in other P450s as well as in the prokaryotic crystal structures. The final strand of -sheet, 3(2), exhibits an alternating pattern of hydrophobic amino acids in many of the P450s that is characteristic of this type of secondary structure and alignment is facilitated by a fairly well-conserved pair of prolines eight residues apart, with intervening repeats of basic and hydrophobic amino acids (Figure 6.1). Consequently, even in this poorly conserved C-terminal portion from strand 3(3) onwards, it is possible to produce a fairly good alignment of the P450 sequences.


227

A recurring theme in this description of how the P450 amino acid sequences may be aligned is that, of the three bacterial crystal structures, P450BM3 represents the best overall template for modelling other forms of the enzyme, especially eukaryotic P450s. The following section summarizes in general terms how it is possible to construct models of eukaryotic P450s from the P450BM3 crystal structure; the results of P450 modelling using the relevant sequence alignments will be discussed in section 6.2.5. 6.2.4 Recommended procedure for homology modelling of P450s The following series of steps can be recommended as a procedure for deriving models of P450s from bacterial crystal structures. 1 Produce a satisfactory alignment between the relevant sequences. 2 Begin at the N-terminus of the crystal structure and work downstream, changing each amino acid residue as required by the alignment. 3 Inspect each change visually to check for any bad contacts which may be produced and note where a change in torsional angle of a residue could give rise to a more favourable interaction, which may be electrostatic, hydrogen-bonded or hydrophobic in character. 4 Delete any residues required by the alignment; this can be carried out concurrently with (2) above, but one should ensure that any secondary structural element will not become grossly distorted by the deletions. 5 Include the additional residues required by the alignment using a loop search routine to extract best fits of homologous peptides from the protein databank. In P450s constructed from P450BM3, this rarely involves more than one or two residues in each ‘loop’. 6 Check the ‘raw’ structure for unfavourable steric contacts. Some packages have a torsional scan option which will relieve steric bumps automatically, but this may not necessarily give the best results. It should be noted, moreover, that a full torsional scan on a large protein can take some time to execute. A known substrate could also be positioned in the putative active site to investigate whether there is any complementarity between groupings on the substrate and residues within the active site region. 7 Energy minimize the entire structure using molecular mechanics. If the unfavourable steric interactions have been modified previously, localized ‘annealing’ is not usually necessary. In the Sybyl (Tripos Associates, St. Louis, MO) software suite, it is possible to monitor the geometry optimization visually, as bonds can be colour-coded by energy. This can be very helpful to ensure that there are no local ‘hot’ areas of high energy where the geometry is unfavourable. The final structure should show a total minimized energy of the order of –1000 kcal.mole−1 (Table 6.9 gives some examples) and the criteria used to determine optimization of the geometry should be that the change in energy per iteration is less than 0.1 kcal.mole−1 and that the change in rms force (or energy derivative) is less than 0.01 kcal.mole −1. Å−1 (Lewis, 1995a). However, some texts recommend that these latter criteria may be double the values mentioned above, for example, Hirst (1990). Table 6.9 Minimized energies* of P450s modelled from P450BM3(CYP102) CYP species

Energy (kcal.mole−1)

Substrate

1A1 rat 1A1 trout

−1086.889 −1125.610

None Ethoxyresorufin

228


CYP species

Energy (kcal.mole−1)

1A2 rat −1058.396 1A2 human −920.268 2A1 rat −1332.946 2A4 mouse −1361.018 2A5 mouse −1342.871 2A6 human −1582.917 2B1 rat −1049.516 2B4 rabbit −1125.856 2B6 human −1080.373 2C9 human −1102.285 2C19 human −1204.931 2D6 human −976.713 2E1 human −1252.090 2E1 rat −1066.169 3A4 human −1260.057 4A11 human −970.404 4A4 rabbit −886.654 4F3 human −962.533 11A1 bovine −829.582 11A1 bovine −1065.453 17A1 human −1053.295 17A1 human −1143.564 19 A 1 human −1129.483 102 bacillus megaterium −1206.412 * Final minimum energy of the optimized geometry.

Substrate None None Coumarin Testosterone Coumarin Coumarin None None None None Omeprazole None p-nitrophenol p-nitrophenol None Lauric acid PGE2 LTB4 None Cholesterol None Progesterone Androstenedione Lauric acid

8 Validate the authenticity of the model using ProCheck (Oxford Molecular Ltd., Oxford, UK) or other software packages for assessing the stereochemical quality of protein models. It can also be an interesting exercise to run the original crystal structure template through such a system, in addition to the model generated from it (see Table 6.10 for an example). Some workers in this field routinely perform energy minimization on the protein crystal structure, prior to homology modelling, to relax the solid state geometry. 9 The model may need further refinement if there are any non-allowed regions of torsional space and/or unacceptable substructures, such as D-amino acid residues, cis-prolines and non-planar geometries of peptide links or aromatic ring systems. 10 Following refinement, a further minimization of the structure is recommended, and one could also consider the inclusion of a substrate (or inhibitor) docked into the putative binding site. It is possible to obtain an estimate of the substrate binding energy from the difference between the minimum energies of the enzyme-substrate complex and the substrate-free enzyme using the equation


229

Table 6.10 Results of ProCheck analysis on the stereochemical quality of P450 model structures (References: Morris et al., 1992 (ProCheck technique); Lewis and Lake, 1995 (CYP2A6 model)) CYP102 (Crystal Structure)

CYP2A6 (Model)

CYP2A6 (Refined)

Notes: The favourable regions correspond to the conformations of the peptide, side chain angles, and , andother torsional angles and overall geometric characteristics of chemical bonds in the protein structure. The above table represents a brief summary of the entire output information derived from a ProCheck analysis.

However, this is only an approximation as it represents the change in internal energy without considering a number of other factors such as the entropy changes due to desolvation. It is possible to make estimations of the other contributions to the binding energy using, for example, a multicomponent equation derived by Williams and co-workers (Williams et al., 1991). The desolvation entropy change, which is likely to be a major contribution to the overall observed binding energy, can be estimated from the relative molecular mass of the substrate as this is proportional to the number of water molecules displaced from the active site when the substrate binds. For example, the binding energy for camphor binding to P450cam (CYP101) can be calculated, using a modification of the Williams’ equation, to be -31.931 kJ.mole–1 (Lewis, unpublished results) which is in close agreement with the experimental value of -32.217 kJ.mole-1 (Griffin and Peterson, 1972). Alternatively, one could carry out interactive docking energy calculations which facilitate optimization of the orientation of the substrate within the putative enzyme active site. Such interactive docking studies can be achieved within Sybyl, and involve automatic calculation of electrostatic and steric interaction energies as the substrate is manipulated interactively relative to the enzyme active site. In order to perform such investigations successfully, however, it is necessary to add hydrogen atoms to the relevant structures and also to calculate the atomic charges. Finally, molecular dynamics simulations (see, for example Paulsen and Ornstein, 1992, 1994, 1995) could be carried out on the enzyme-substrate complex which may include consideration of the local dielectric constant, for example, and other factors relating to the likely environment medium encountered by the enzyme. 6.2.5 Eukaryotic P450s modelled from P450BM 3 (CYP102) The comparisons between P450BM3 and P450cam crystal structures have been described previously (Lewis, 1995a). In terms of sequence alignments, the former should be regarded as being the preferred starting point for producing models of other P450s. However, the lack of a bound substrate in the P450BM3 structure does present something of a drawback, compared with P450cam, and there is a marked ‘openness’ in P450BM3 which is not reflected in P450cam (Ravichandran et al., 1993; Lewis, 1995a). A further difficulty is the lack of an obvious basic residue to form the second ion-paired interaction with one of the heme propionates of P450BM3 (Lewis, 1995a). Apart from these potential problems, it can be generally assumed that models derived from P450BM3 should represent relatively close analogues of the actual eukaryotic P450s. In fact, it has been found that every P450 model produced so far from P450BM3 appears to rationalize known substrate specificity, and preliminary findings will now be summarized for each isoform constructed, roughly in order of their classification. As these models have been produced by alignments within each family or subfamily, there may be some small variations from the global alignment shown in Figure 6.1. Residue

230


Figure 6.6 Alignment between CYP102 and members of the CYP1 family.

positions are referred to on the basis of the P450BM3 alignment rather than in the actual enzyme concerned, however. 6.2.5.1 The CYP1 family 1. CYP1A1. The putative active site of rat CYP1A1 generated from a modelling alignment with P450BM3 (see Figure 6.6 for this alignment) is shown in Figure 6.7, with the known substrate, ethoxyresorufin, orientated for O-de-ethylation (Lewis et al., 1995a). The planar poly aromatic substrate occupies a complementary ‘slot’ in the heme pocket made up from a number of approximately coplanar aromatic amino acid residues (F88, F181, F266 and Y437) of which two, F181 and Y437, lying about 6.2 Å apart, appear to ‘sandwich’ the essentially planar substrate via - stacking interactions between co-planar aromatic rings (Figure 6.7). In addition, there are several polar amino acid residues, within the putative active site region, that can enter into hydrogen-bonded interactions with certain electronegative atoms on the ethoxyresorufin substrate. In particular, amino acids T185, T78, T87 and N82 are able to serve as potential hydrogen bond donor/acceptor residues, of which the former can readily donate a hydrogen bond to the ketonic oxygen of the resorufin nucleus (Figure 6.7). The key interations of the two aromatic residues, F181 and Y437, together with the T185 side chain, serve to orientate the ethoxyresorufin substrate for O-de-ethylation, as the methylene portion of the ethoxy group is positioned above the heme iron such that oxygenation can occur and lead to dealkylation, which is the experimentally observed reaction catalyzed primarily by this enzyme (Burke et al., 1985). It is not clear whether N82 would form a hydrogen bond with the nitrogen atom of the resorufin nucleus, as the phenyl ring of F88 tends to sterically hinder such an interaction. The other two potential hydrogen bond donors, T78 and T87, are positionally too high and too low, respectively, to contact the substrate at this point (Figure 6.7). However, it is possible to reorientate the ethoxyresorufin substrate such that S178 could perform a similar function to T185 in the original orientation, thus enabling either T78 or T185 to donate a hydrogen bond to the substrate’s central ring nitrogen. It is interesting to speculate whether the close structural analogy between the resorufin nucleus and a flavin ring system indicates a possible endogenous role of either of the CYP1 isozymes in flavin metabolism. A flavin nucleus, i.e. the isoalloxazine ring system, could readily fit the putative active site of CYP1A1 (or 1A2) in a similar manner to ethoxyresorufin (or methoxyresorufin in the case of CYP1A2) and, furthermore, the additional carbonyl group on the C ring of isoalloxazine would be in an optimum position for accepting a hydrogen bond from T78. In the putative active site of CYP1A1, F327 may restrict the chain length of the alkoxy substituent such that the ethoxy congener is the preferred member of the alkoxyresorufin series (Lewis et al., 1995c). Another two phenylalanine residues, F88 and F266, could facilitate the binding of polyaromatic


231

hydrocarbon (PAH) substrates, such as benzo(a)pyrene, and those PAHs containing more extensively delocalized -systems; although there will be dimensional constraints on such substrates due to the

232


topography of the active site (Figure 6.7). In fact, it is possible that at least two aromatic amino acid


233

residues may operate as an access channel for PAHs, and other structurally related substrates which can differentiate between their molecular shape characteristics. Our group has reported such a situation using a CYP1A1 model derived from P450cam (Lewis et al., 1994d) and, in the current P450BM3 model, it would appear that Y213 and F259 may serve this function. The latter of these two amino acids is particularly interesting in this respect as it changes to an asparagine residue in CYP1A2, thus providing some rationale for the preference of the 1A2 isozyme for polyaromatic amine and polyaromatic amide substrates. In fact, both benzo(a)pyrene and its 7, 8-diol are able to fit the CYP1A1 active site in the current model, such that the co-planar phenylalanine and tyrosine residues (F181, F266 and Y437) orientate the pyrene ring system for oxygenation in the known positions, which are 7, 8-epoxidation in benzo(a)pyrene and 9, 10epoxidation for the 7, 8-diol. The latter oxygenation is specifically catalyzed by CYP1A1, and the enzyme model (Lewis and Lake, 1996) shows that T87 can form a hydrogen bond with one of the hydroxyl groups of the substrate to assist in orientation of the molecule such that it will be preferentially epoxidized with the known stereospecificity (Conney et al., 1994). It is possible that N82 could form a hydrogen bond with the second hydroxyl moiety of the 7, 8-diol to enable precise oxygen attack at the 9, 10-anti position that would give rise to the ultimate carcinogen (Conney et al., 1994). Although there is no apparent role for T185 in orientation of the benzo(a)pyrene substrate for metabolism by CYP1A1 as there is for ethoxyresorufin, the model for this enzyme is able to explain the experimentally observed activating metabolism of the structurally related proximate carcinogen 15, 16-dihydro-11methylcyclopenta[a] phenanthren-17-one, with reference to a key interaction between T185 and the ketonic function of the substrate (Boyd et al., 1995). Although this compound is metabolized at several positions in the molecule, the formation of the 3, 4-diol represents the metabolically activating reaction catalyzed by CYP1A1. It is possible to orientate the substrate in several ways within the putative P4501A1 active site, all of which present the molecule to the heme iron for oxygenation to occur in the known positions (Boyd et al., 1995). The situation where 3, 4-oxygenation can take place would necessitate the substrate to be positioned such that the carbonyl group accepts a hydrogen bond from T185; whereas, in another orientation, involving donation of a hydrogen bond from T87 to the ketonic function of the cyclopentaphenanthrenone derivative, the 11–methyl group on the substrate could be hydroxylated (Boyd et al., 1995). However in an alternative position which involves hydrogen bonding to the ketone via T78 such that oxygenation can occur in the 1, 2-position, it is found that orientation of the cyclopentaphenanthrenone substrate for 6, 7-oxygenation gives rise to unfavourable steric interactions without any possibility of hydrogen bonding and, furthermore, with less favourable - stacking between aromatic rings. This observation would appear to explain why there are no metabolites found for this substrate corresponding to oxygenation at the 6, 7–bond (Boyd et al., 1995). Site-directed mutagenesis studies in the 2A and 2B subfamilies (Table 6.8) have identified residues corresponding to T87, F181 and Y437 as being associated with substrate interactions but, as far as the CYP1 family is concerned, only the CYP1A2 isozyme has been investigated using this technique (Furuya et al., 1989a and b). In this case, however, only proximal and distal heme mutations have been produced, so further studies would have to be performed in order to establish whether the previously mentioned residues are, in fact, involved in substrate binding. It is difficult to visualize how proximal heme mutants can possibly affect substrate binding, but they may alter the (largely hydrophobic) contacts between I and L helices which maintain the position of the heme and overall topography of the heme binding site, including those regions which may be involved in substrate-protein interactions. Additionally, mutations carried out along the L helix, which is proximal to the heme, may affect the binding of redox partners and this could, therefore, alter the catalytic activity of the enzyme in general.

234


In fact, the natural mutant form of CYP1A1, which has been identified in certain individuals where there is an association with lung cancer, shows a single residue change at position 405 in the alignment and this corresponds to the start of the L helix (Kawajiri et al., 1992). However, it is not established whether this rare allelic variant in human CYP1A1 has a direct causal relationship with the incidence of lung cancer in man and it is not easy to see how such a conservative change (I V), which is associated with this polymorphic mutation, could have any significant effect on the catalytic properties of the P450. Perhaps the I462V mutant brings about an uncoupling of P450 oxygenation leading to an increase in ROS, or by moderating an alteration of the PAH-oxygenase activity that leads to an increase in activation pathways relative to those which are generally detoxifying in nature. It is possible that even this conservative change could have a subsequent effect on the electron delocalization within the heme environment, as the site-directed mutagenesis data on CYP1A2 proximal to the heme do indicate that the catalytic activity of the enzyme can be altered by apparently minor changes in this region (Furuya et al., 1989a and b). One explanation may involve the effect of proximal heme variations on the geometry of the cysteinate ligand which could, in turn, alter the ability of the enzyme to stabilize oxygenated intermediates in the catalytic cycle. Alternatively, the I462V mutation may affect the hydrogen-bonded conduit of water molecules thought to play a role in proton coupling (Raag and Poulos, 1992). 2. CYP1A2. Although this isozyme is highly homologous with CYP1A1, there are several notable amino acid changes between the two putative active sites which may explain the known differences in metabolism and substrate preferences. For example, CYP1A2 displays specificity towards methoxyresorufin as opposed to the ethoxy homologue which is more specific for CYP1A1 (Burke et al., 1985). In particular, T185 in CYP1A1 is a valine in CYP1A2, whereas V184 in CYP1A1 becomes asparagine in P4501A2. This change at two adjacent residues in the F helix, which lies above the heme, causes a reorientation of the resorufin substrate such that it is difficult for the ethoxy congener to become de-ethylated by CYP1A2, whereas methoxyresorufin can be favourably orientated in CYP1A2 for O-demethylation (Lewis et al., 1995c). Furthermore, F259 in CYP1A1 changes to an asparagine in CYP1A2 and this residue in the I helix may be involved in controlling the access of substrates to the active site, thus explaining the preference of CYP1A2 for aromatic amines and amides, as opposed to the polyaromatic hydrocarbon (PAH) substrates of CYP1A1, where F259 appears to form an access channel for planar PAHs by adopting a - stacked coplanar conformation with F212. Caffeine is another known substrate of CYP1A2, and this is metabolized by the enzyme at three positions, namely, N1-, N3- and N7-demethylation (Berthou et al., 1992). In order to explain the known sites of caffeine metabolism mediated by CYP1A2, it is clear that the substrate will have to adopt at least three possible orientations within the active site; all of which should involve particular interactions with key amino acid residues in this region. Moreover, N-demethylation in the N3-position may be associated with a preferred interaction in the CYP1A2 active site as the N3–demethylated derivative is the major metabolite (Berthou et al., 1992). In fact, the putative CYP1A2 active site constructed from P450BM3 is able to show how all three caffeine metabolites can be formed and why N-demethylation in the 3-position is preferred (Ayalogu et al., 1995). Figure 6.8 shows that caffeine is able to adopt three superimposable positions in the active site of CYP1A2 by forming two hydrogen-bonded interactions with both N82 and T87, which each donate a hydrogen bond to either the two carbonyl oxygen atoms, or to one of these and the N9 nitrogen atom in the caffeine molecule. In each case, therefore, a different N-methyl group of the substrate will be positioned close to the heme iron for oxygenation. The orientation which would correspond to N3-demethylation gives not only the best - stacked overlap between the aromatic rings of the substrate and those of F181 and


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Y437, but also presents the N9 nitrogen towards E267, which could either form a hydrogen bond, if the latter is protonated, or an ion-pair if the former basic nitrogen is protonated instead (Ayalogu et al., 1995). It should be noted that residues in positions analogous to F181, Y437 and T87 have been shown to be important for substrate binding in other P450s, by using site-directed mutagenesis (Table 6.8). In CYP1A2, however, only a number of positions distal and proximal to the heme face have been examined by this technique (Shimizu et al., 1988, 1989, 1994; Tuck et al., 1993a; Furuya et al., 1989a and b). Two independent studies (Shimizu et al., 1994; Tuck et al., 1993a) have confirmed the importance of the distal charge-relay system comprising the conserved acidic residue E267 (E318 in CYP1A2) and the invariant threonine T268 (T319 in CYP1A2) that are crucial to substrate oxygenations (Gerber and Sligar, 1992; 1994). Earlier site-directed mutagenesis experiments focused on the effects of these and other distal (and proximal) residues on substrate regio-specificity in acetanilide hydroxylation (Furuya et al., 1989a) and catalytic activities towards benzphetamine and 7-ethoxycoumarin metabolism (Furuya et al., 1989b). It is not clear, however, from these early studies, whether the mutated residues are important for substrate binding or simply modulate the catalytic activity of the enzyme. In the human CYP1A2 orthologue, there is a change in the putative active site region relative to the rat form which could rationalize, to some extent, the clear preference of this enzyme to N-demethylate caffeine at the 3-position, as opposed to the other possibilities mentioned previously. The modelling alignment shows that N82 in rat CYP1A2 changes to aspartate in the human isoform, and it is possible that T78, in addition to T87, will constitute the two hydrogen bond donor residues in this case. Apparently, this difference indicates that the caffeine substrate will show a more favourable interaction with these (and other) putative active site residues, when orientated for N3-demethylation as shown in Figure 6.9. The CYP1A2 model also explains the variations in metabolism of the four isomers of diaminotoluene (Lewis and Lake, 1996). It is known that 2, 4-diaminotoluene is metabolized via N-hydroxylation at the 4amino position rather than the 2-position, whereas the 2, 5-diaminotoluene is N-hydroxylated at a higher rate; the other two isomers are only weakly metabolized. It is possible to superimpose all four diaminotoluenes onto caffeine in the CYP1A2 active site where the 2, 5-isomer fits most closely with the caffeine molecule, as the former can adopt two orientations which superimpose the methyl group of the toluene with either N1- and N3-methyl groups in caffeine, so that the 5-amino group of 2, 5-diaminotoluene will be hydroxylated preferentially and the 2-amino group can donate a hydrogen bond to T78. Similarly, 2, 4-diaminotoluene will N-hydroxylate at the 4-amino group if its 2-amino function is able to donate a hydrogen bond to T87. When the other two diaminotoluenes are superimposed onto this template, it can be shown that each amino group is sterically hindered either by an adjacent methyl or another amine function and, consequently, hydrogen-bonded interactions are not favourable with any of the putative active site residues; oxygenation at the free amino group is also sterically hindered by ortho substituents in a similar fashion. It appears, therefore, that the CYP1A2 model agrees closely with known experimental data on substrate metabolism; moreover, it can be shown that the specific 1A2 inhibitor, furafylline, will also fit the putative active site as does the specific substrate Glu-P-1 (Lewis and Lake, 1996). Additional supporting evidence for the use of P450BM3 as a template for this enzyme has been reported from active site studies using the phenyldiazene inhibitor (Tuck et al., 1992) where the heme environments of CYP1A2 and P450BM3 appear to show some commonalities. Furthermore, antibodies raised to recognize and differentiate between CYP1A2 and CYP1A1 have been designed to complement a region of polypeptide between the H and I helices (Edwards et al., 1993, 1994, 1995). The CYP1A2 and CYP1A1 models show that this stretch of peptide is on the enzyme surface and lies close to one of the likely points of interaction with reductase, which may explain the inhibitory effect of antibody binding on enzyme activity.

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6.2.5.2 The CYP2 family 1. CYP2A subfamily. Of the three rat orthologues, CYP2A1 has been modelled from P450BM3 and it appears that the putative active site is able to accommodate the known substrates of this isoform, where specific amino acid residues can orientate the relevant structures for metabolism at the experimentally observed positions (Lewis and Lake, 1995). From the modelling alignment between the CYP2A sub-family and P450BM3, shown in Figure 6.10, amino acid residues which can be expected to occupy the active site region of CYP2A1 include N181, N78 and Q74, together with a number of essentially hydrophobic amino acid side chains. The combination of the two asparagine residues, N181 and N78 in particular, is able to orientate both coumarin for oxygenation in the 3-position, and NNK for hydroxylation at the -carbon adjacent to the N-nitroso moiety. This is due to the fact that the oxygen atoms on these two substrates can accept hydrogen bonds from the two amide groups of these amino acid residues in the CYP2A1 active site. Furthermore, testosterone is hydroxylated in the 7 -position by CYP2A1 and this can be explained in terms of hydrogen-bonded interactions between N181 and Q74 amide side chains and the two oxygen atoms in the steroid substrate, as shown in Figure 6.11. The two mouse orthologues, CYP2A4 and CYP2A5, are highly homologous, differing in only 11 amino acid residues out of 494 (Squires and Negishi, 1988). It has been established that three of these residue positions are especially important for determining the different substrate specificities of the two isozymes, via site-directed mutagenesis experiments (Lindberg and Negishi, 1989; Juvonen et al., 1991; Lindberg et al., 1992; Iwasaki et al., 1993). All three of these positions lie within the putative active site regions of the two CYP2A models (Lewis and Lake, 1995) and one in particular, L181F, appears to be in close contact with the relevant substrates, either coumarin or testosterone. A mutation from leucine to phenylalanine at position 181 (209 in 2A4, 5) is sufficient to alter the enzyme specificity from testosterone 15 -hydroxylase to coumarin 7-hydroxylase activity (Lindberg and Negishi, 1989; Poulos, 1989) and the model of CYP2A5 shows that a phenylalanine at this position will be able to form a - stacking interaction with coumarin so that the substrate can be orientated, with the aid of T184 which hydrogen bonds to the ketone moiety, for hydroxylation at the 7-position (Lewis and Lake, 1995). The two other residues, L327 and A87, are also relatively close to the testosterone substrate, in particular, in the CYP2A4 model, where there is complementarity between these hydrophobic residues and similar regions of the testosterone molecule. These amino acid residues become methionine and valine, respectively, in CYP2A5 and these two changes, although less important then the L181F change, restrict the size of the heme pocket which will, consequently, favour the occupancy of coumarin rather than testosterone (Lewis and Lake, 1995). Both of these substrates, however, contain a carbonyl group which appears to be able to hydrogen bond with T184, and this residue is common to both isozymes, thus indicating a possible role for this residue in controlling substrate specificity and regioselectivity. The major human orthologue, CYP2A6, bears similarity with CYP2A5, especially in possessing a phenylalanine residue at position 181, and it is, therefore, perhaps not surprising that CYP2A6 also hydroxylates coumarin at the 7-position (Yamano et al., 1990). The model of this enzyme (Figure 6.12) clearly demonstrates how the preferred substrate, coumarin, readily occupies the putative active site where there is considerable complementarity between the substrate and certain key residues (Lewis and Lake, 1995). In addition to F181, the side chains of T184 and H437 appear to be in close proximity to the coumarin substrate, where the two aromatic amino acid residues adopt a coplanar geometry by - stacking interactions with the benzene ring of coumarin (Figure 6.12). As in the other CYP2A orthologues, T184 in CYP2A6 can donate a hydrogen bond to the carbonyl oxygen atom of the substrate which, in the case of coumarin, will orientate the molecule for 7-hydroxylation (Lewis and Lake, 1995). CYP2A6 can also


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Figure 6.10 Alignment between CYP102 and members of the CYP2A subfamily.

metabolize aflatoxin B1 and it is possible to show how this substrate can be positioned in the putative active site, which explains the known CYP2A6-mediated metabolism. In fact, it appears that the same amino acid residues are involved in binding aflatoxin B1 as there are for coumarin, and modelling shows that there is some degree of structural similarity between the two molecules; in fact, visual inspection reveals that aflatoxin B1 contains the coumarin nucleus as a substructure of the molecule.

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A second human form of CYP2A, namely CYP2A7, does not appear to be able to incorporate heme readily, and a comparison between the sequences of the two human orthologues shows that one of the conserved basic amino acid residues, likely to be necessary for ion pairing with one of the heme propionates, is not conserved in CYP2A7. It is possible that this fact may explain the apparent lack of full functionality of the 2A7 isoform (Lewis and Lake, 1995). Members of the CYP2A subfamily differ from other microsomal P450s in not possessing a conserved tryptophan residue analogous to W96 in P450BM3.


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Figure 6.13 Alignment between CYP102 and members of the CYP2B subfamily.

However, Y84 may fulfil the same role in 2A as W96 does in other P450s, because this residue appears to be close enough to the heme moiety for either - stacking or hydrogen-bonding to the relevant propionate group. It would be interesting, therefore, to ascertain the possible role of Y84 using site-directed mutagenesis and other techniques. Nevertheless, many aspects of the 2A subfamily can be rationalized from the models generated from P450BM3, as has been reported previously (Lewis and Lake, 1995). 2. CYP2B subfamily. As with the CYP2A subfamily, models of CYP2B indicate a close agreement with site-directed mutagenesis studies, and it is also possible to show how substrates of these isozymes are able to occupy their putative active sites such that the known positions of metabolism can be explained. For example, it has been demonstrated that single point mutations at certain specific amino acid residue positions in CYP2B1 will affect the regio- and stereo-specificity of steroid metabolism (Halpert and He, 1993; He et al., 1994). In particular, the change from valine to leucine at position 332 in the alignment shown in Figure 6.13 (corresponding to 367 in CYP2B1) results in only 16 -hydroxylation of androstenedione, as opposed to both 16 - and 16 -hydroxylation of this substrate in the wild type (He et al., 1994). The model of CYP2B1, with androstenedione docked in the putative active site, indicates that the Aring ketonic function of the substrate may form a hydrogen bond with a serine residue in position 69, and that the V L change at 332 is likely to bring about a reorientation of the substrate molecule which will cause the 16 -hydrogen to become preferentially positioned directly above the heme iron such that only this site will become hydroxylated (Figure 6.14). The change of valine to leucine involves the addition of a CH2– group to the side chain, which is sufficient to bring about the alteration in substrate position that leads to the known stereo-selectivity in metabolism mentioned above. In fact, the other major rat orthologue, P4502B2, shows this V L difference which could, therefore, explain the known variation in stereospecificity of steroid metabolism between CYP2B1 and CYP2B2 (He et al., 1993). There are other differences in metabolism between these two highly homologous proteins (differing in only 14 amino acids) which may be rationalized in terms of amino acid changes in their active sites (Aoyama et al., 1989; Kedzie et al., 199la; Halpert and He, 1993; He et al., 1994). Allelic variants of CYP2B1 and CYP2B2 have been explored via site-directed mutagenesis in order to establish which amino acid residue positions are particularly important determinants of steroid substrate regiospecificity in metabolism (Kedzie et al., 1991a; Aoyama et al., 1989). These studies have demonstrated that a further two residues are relevant to variations in androgenic steroid metabolism, namely, I87 (position 114 in 2B1 and 2B2) and G439 (position 478 in 2B1 and 2B2). Both of these amino acids lie in the putative active site of the CYP2B1 model (Lewis, 1995a), and it can be shown that modification at either position will affect the orientation of both androstenedione and testosterone substrates, thus explaining the experimentally observed change from 16 - to 15 -hydroxylation. Furthermore, another two amino acid residue positions that are sensitive to steroid metabolism regiospecificity (He et al., 1994) are also present within the putative active site of CYP2B1 (Lewis, 1995a). These are F181 (206 in 2B1) and V328 (363 in 2B1) which confer testosterone 7 and 15 -hydroxylating activity, respectively, in this enzyme following mutation to small aliphatic sidechain hydrophobic amino acid residues (He et al., 1994).

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Exogenous substrate metabolism can also be explained in terms of key amino acid residues in the putative CYP2B1 active site. Many known CYP2B1 substrates, including: phenobarbital, pentoxyresorufin, benzphetamine, nicotine, chloramphenicol, DDT, androstenedione and testosterone, can be orientated in the


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CYP2B1 binding site for metabolism in the known positions, and inhibitors such as: SKF-525A, metyrapone and secobarbital will also fit the CYP2B1 model. For both substrates and inhibitors, there is complementarity between key amino acid residue positions and certain structural features of the substrates

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and inhibitors themselves, and some of these amino acids have been mentioned previously in this section. For example, the specific substrate pentoxyresorufin can occupy the CYP2B1 site by accepting a hydrogen bond from S69 which, with other key residues, orientates the substrate for O-depentylation (Lewis et al., 1995c). The pentyl group appears to be of the optimum length to lie in an essentially hydrophobic channel containing L327 and V328, of which the latter has been shown previously to be relevant to steroid regioselectivity. The N-pentyl congener in a series of 2, 4-dichlorophenoxy N-alkyl N-methyl-ethylamines (Roffey, 1993) will fit the CYP2B1 active site in a similar manner, and it is also found that chloramphenicol can form a hydrogen bond between S69 and its p-nitro group to position the molecule for metabolism at the dichloromethyl ketone functionality. It is interesting to note that the distance between the nitro group and site of metabolism in chloramphenicol is 11.03 Å whereas, in androstenedione, the A-ring ketonic function is 11.10 Å from the 16 -hydrogen; these two substrates are readily superimposible in the putative CYP2B1 active site, conforming to their respective positions of metabolism. The equivalent distance in pentoxyresorufin is somewhat shorter at 10.20 Å, but it is possible to dock this substrate in an alternative conformation, where it can form a hydrogen bond with S185 instead of S69, and F181 could interact via stacking with the resorufin ring system (Lewis et al., 1996a). Both phenobarbital, and its inhibitory analogue secobarbital, may also enter into hydrogen-bonded interactions with S185 and could moreover stack with F181, although F263 is another possibility. Furthermore, another known CYP2B substrate, DDT, can be orientated in the 2B1 active site by assuming that there may be interactions between its two phenyl rings and those of F181 and F263 in such a way that its position of metabolism lies above the heme iron. The CYP2B inhibitor, SKF–525A, also contains two phenyl rings in an analogous conformation to DDT (Rossi et al., 1987) which could, therefore, fit into the 2B1 active site in a similar way. Chloramphenicol is another mechanism-based inhibitor of this isozyme (Halpert et al., 1988) but, in this case, the reactive metabolite forms an imine by covalent binding to an active site lysine residue (Miller and Halpert, 1986). The model of CYP2B1 indicates that such an interaction is possible with either K82 or K436, of which the latter seems most likely. In addition to substrate and inhibitor interactions, the models of CYP2B1 and the major rabbit orthologue CYP2B4, appear to be in agreement with likely redox partner binding sites, antibody recognition sites, and serine phosphorylation sites (Lewis, 1995a). Apparently, the binding of cytochrome b5 blocks serine phosphorylation (Epstein et al., 1989) which suggests that the relevant interacting sites are in relatively close proximity. This is clearly shown in the CYP2B1 (or 2B4) model, as S103 (S128 in 2B1 and 2B4) is sufficiently near R100 (which is also a surface residue) to be inhibited from phosphorylation if cytochrome b5 interacts with R100 (R125 in 2B1 and 2B4). As this basic residue also forms an ion-pair with one of the heme propionates, phosphorylation of the nearby serine–103 will bring about a movement in R100 (Figure 6.5) which will tend to ion-pair with the phosphate moiety, thus diminishing the heme binding affinity of the apoprotein. If one assumes that cytochrome b5 is likely to bind to microsomal P450s in a similar way to its interaction with P450cam (CYP101) especially as there is a general conservation of basic residues at these positions, then R100 will be involved in the binding of cytochrome b5 to CYP2B enzymes (Lewis, 1995a). Other surface regions of the CYP2B1 model correlate with known antibody recognition sites (DeLemosChiarandini et al., 1987) whereas the same is true for the CYP2B4 model, as this also equates with epitope mapping studies in the 2B4 isozyme (Kolesanova et al., 1994) which have been used, moreover, to ascertain the probable membrane-binding regions (Uvarov et al., 1994). The likely surface residue positions for reductase interaction with CYP2B4 have been investigated by Bernhardt and co-workers (Bernhardt et al., 1984, 1987, 1988, 1989). All of these basic residues are present as surface sites in the CYP2B4 structure


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and, furthermore, it is possible to show via molecular modelling how the FMN and FAD domains (Black and Coon, 1982; Porter and Kasper, 1985, 1986; Porter et al., 1990) of NADPH-dependent P450 oxidoreductase could bind to CYP2B4 such that electron transfer to the heme can occur (Figure 6.15). Details of the reductase modelling studies will be published elsewhere due to the space limitations of the current work; however, these involve construction of the FMN and FAD domains from flavodoxin and ferredoxin reductase, respectively (Lewis, unpublished data). 3. CYP2C subfamily. There has been a number of site-directed mutagenesis studies conducted on various isozymes in this subfamily which tend to support the models produced (Lewis, 1995a). For example, the residue at position 87 (113 in the 2C subfamily) seems to be important for determining substrate specificity (Straub et al., 1993a and b; Kronbach et al., 1991, 1989). This hydrophobic residue is valine in both 2C9 and 2C19, and the ethyl group of either phenytoin or mephenytoin will be in contact with the aliphatic side chain of this amino acid if these substrates are orientated for hydroxylation at the known site of metabolism mediated by these isozymes (Lewis, 1995a). The (S)-isomer of warfarin can also fit the 2C9 active site in an orientation suited for 7-hydroxylation (Rettie et al., 1992) with its methyl group in close hydrophobia contact with V87. This residue lies in a region generally regarded to be one of the substrate recognition sites (SRS1) for P450s of the 2 family (Gotoh, 1992). However, this so-called hypervariable region is strikingly well-conserved between P450BM3 and members of the CYP2C subfamily with a 7-residue motif of G x G x x x S, where the first two residues after the second glycine are hydrophobic, usually isoleucine and valine respectively (Figure 6.16). This may not be so surprising when one considers that, in common with P450BM3, some CYP2C isozymes can hydroxylate long chain carboxylic acids (see Chapter 4 for further details). The importance of various amino acid residues in this putative substrate binding site region have been extensively investigated by both site-specific mutagenesis (Kronbach et al., 1989, 1991; Straub et al., 1993a and b) and by antibody recognition studies (Kronbach and Johnson, 1991). These findings point to at least three amino acid positions (including the one mentioned previously) being relevant to progesterone 21hydroxylase and lauric acid -1 hydroxylase activities in 2C1, 2C2, 2C4 and 2C5 isozymes, whereas antibody binding to this region brings about inhibition of catalytic activity in 2C5 (Kronbach and Johnson, 1991). In the CYP2C models, the stretch of peptide corresponding to the epitope recognized by the 2C5 specific monoclonal antibody is a surface loop between the putative substrate binding site and the conserved region of C helix which contains a likely reductase interaction point. Another region of the putative CYP2C binding site which has been studied by site-directed mutagenesis (Hsu et al., 1993; Kaminsky et al., 1993; Richardson and Johnson, 1994) corresponds to a second substrate recognition site close to the conserved proline, P329 (P362 in the CYP2C subfamily). Apparently, an allelic variant of CYP2C3 contains a conservative amino acid change S331T (S364T in CYP2C3), two residue positions downstream from the aforementioned proline, which confers progesterone 6 -hydroxylase activity in the variant 2C3v (Hsu et al., 1993). As the wild-type enzyme normally possesses progesterone 16 hydroxylase activity, it would seem that a single residue change brings about a considerable alteration in substrate orientation in the binding site. In fact, the CYP2C model can rationalize this dramatic change in enzyme regiospecificity in terms of hydrogen bonding between the side chain of this residue position and the two carbonyl groups at either end of the progesterone molecule (Lewis, 1995a). In 2C3, the serine-331 hydroxyl group can donate a hydrogen bond to the A-ring carbonyl oxygen of progesterone such that the 16 -hydrogen is positioned directly above the heme iron (Figure 6.17). However, if this serine is changed to threonine, the hydrogen-bonded interaction is sterically hindered by the -methyl group of threonine, but progesterone can still form a hydrogen bond to the threonine hydroxyl via its D-ring carbonyl substituent; this interaction will cause the substrate to reorientate for 6 -hydroxylation (Lewis, 1995a). As this residue

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Figure 6.16 Alignment between CYP102 and members of the CYP2C subfamily.

position is occupied by asparagine in CYP2C4 and 2C5, it is possible that such a change may explain the fact that progesterone is hydroxylated at the 21–position in these isozymes, as the steroid substrate would have to alter its orientation in the site accordingly. Position 331 is occupied by serine in both 2C9 and 2C19, two of the major human CYP2C isoforms, and it is found that many known substrates of these enzymes can be orientated for oxygenation at the experimentally observed positions by forming hydrogen bond interactions with S331 and key electronegative hydrogen bond acceptor atoms (usually oxygen) in the relevant substrate molecules (Lewis,


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1995a). For example, both tolbutamide and mephenytoin will fit the 2C9 active site by hydrogen bonding between ketonic oxygen atoms on the substrates and S331, such that their respective positions of metabolism lie above the heme iron (Figure 6.18). In this instance, the distances between the hydrogen bond acceptor atoms and the sites of metabolism are both 7.86 Å for these two substrates. Tienilic acid will also fit the putative 2C9 site in a similar fashion with its ether oxygen atom able to accept a hydrogen bond from S331, being at a distance of 7.99 Å from the site of oxygenation on the thiophene ring (Lopez-Garcia et al., 1993). In this case, the chlorine atom ortho to the oxyethanoic acid side chain is in hydrophobic contact with another residue known to be present in the active site, namely, V87. The n-butyl side chain of tolbutamide appears to contact V87 similarly (Lewis, 1995a) whereas, in mephenytoin, this hydrophobic contact is achieved by either the ethyl or N-methyl group, as two orientations are possible. With (S)-warfarin, it is the ring carbonyl oxygen that hydrogen bonds with S331 to enable hydroxylation at the 7-position on its coumarin nucleus. The other carbonyl group can form a second

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hydrogen bond with S69, and the methyl group adjacent to this ketonic function is in hydrophobic contact with V87. Furthermore, the phenyl ring of warfarin could enter into a - stacking interaction with F74. Returning to tienilic acid, one of the carboxylate oxygen atoms can also form a hydrogen bond with S69 in a similar fashion to the ketone oxygen of warfarin, mentioned previously. However, as far as progesterone is concerned, the distance between the A-ring carbonyl and 16 -hydrogen is 10.74 Å, which is over 2 Å greater than the analogous distance in other substrates. Nevertheless, the oxygen atom of S331 lies 9.43 Å from the iron-oxene oxygen atom in the putative active site of 2C9, so it is likely that there is a range of possible distances between the relevant atoms on 2C substrates which will enable hydrogen-bonded interactions to occur with S331. From the substrates considered so far, this range is 7.8 to 10.8 Å which agrees closely with the 2C9 substrate template model of Smith and Jones (Jones et al., 1993; Smith and Jones, 1992). The specific inhibitor, sulfaphenazole is also able to fit the putative active site of 2C9 in a similar manner to its substrate analogue, tolbutamide, although it is also possible for the inhibitor to ligate the heme iron in an alternative orientation, where its amino group donates a hydrogen bond to S331. Moreover, it can be shown that omeprazole will readily occupy the 2C19 active site, at an orientation consistent with its known position of metabolism (Andersson et al., 1993) by forming a hydrogen bond between S331 and the sulphone oxygen, with one of its methoxy groups in hydrophobic contact to V87. These findings clearly demonstrate that the CYP2C models are in good agreement with experimental observations. 4. CYP2D subfamily. The putative 2D active site acidic residue, corresponding to D260 in the alignment shown in Figure 6.19 (D301 in the CYP2D subfamily) which is thought to be associated with the binding of basic protonated substrates to these isozymes, has been shown by site-directed mutagenesis to be critical to 2D6 substrate interactions (Ellis et al., 1994). It should be noted that many other P450s possess an acidic residue at this position in the I helix, but all of these also have a second acidic amino acid side chain seven residues downstream, which is probably involved in the catalytic oxygenation mechanism (Gerber and Sligar, 1994). Isozymes of the CYP2D subfamily, however, have a hydrophobic amino acid (usually valine) instead of this conserved acidic residue next to the invariant threonine. Presumably, the reason for this is to ensure substrate orientation in the active site such that the nitrogenous 2D substrates will be metabolized at a certain position in the molecule some distance removed from the protonated nitrogen. The striking similarity between this stretch of I helix in the 2D subfamily and that of the third transmembrane helix in the tryptaminergic receptors and in some of the other G-protein-coupled receptors, which contain a highly conserved aspartate for the binding of biogenic amines and their antagonists, leads one to speculate on the possible evolutionary origin of the 2D subfamily for the detoxication of plant toxins containing protonatable nitrogen functions. Typical 2D6 substrates, such as debrisoquine, are able to fit the putative active site of the CYP2D6 model as shown in Figure 6.20, where the carboxylate side chain of D260 can readily form ion-paired interactions

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Figure 6.19 Alignment between CYP102 and members of the CYP2D subfamily.

with the basic nitrogen of these substrates, that are protonated as physiological pH, giving rise to relatively low Km values (Smith and Jones, 1992). Presumably, the electrostatic forces between substrate and enzyme constitute a major contribution to the overall binding energy, thus facilitating significant catalytic activity at


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low substrate concentrations. Many 2D6 substrates also possess an aromatic ring, in addition to the basic nitrogen, and it is possible that there may be a - stacking interaction between F87 and planar aromatic groups on the substrates. Several groups have reported substrate template models for 2D6 (Strobl et al., 1993; Islam et al., 1991; Koymans et al., 1992) whereas Koymans and co-workers have constructed an

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active site model of 2D6 by homology with P450cam (Koymans et al., 1993b). The model (Figure 6.20) of the entire 3D structure of P4502D6 generated from P450BM3 (Lewis, 1995a) generally agrees with Koymans’ active site model, and particularly as far as the I helical region is concerned. However, there are several other regions of the putative active sites of 2D6 where differences between the models can be observed which are due to alignment variations, and the 2D subfamily sequences have been aligned with that of P450BM3 (Figure 6.19) to give an example of one possibility. Unfortunately, there have been not as many site-directed mutagenesis studies reported on the 2D subfamily as, for example, with enzymes of the 2A, 2B and 2C subfamilies, but position 332 (380 in 2D1) has been mutated from leucine to phenylalanine in the rat orthologue, CYP2D1, in order to simulate the situation in an allelic variant of this enzyme (Matsunaga et al., 1990a). This position corresponds to methionine in 2D6 and is present at another region of the putative active site from the aspartate mentioned previously, but close to the alignment position occupied by serine in 2C9 (S331) which was discussed in the preceding section. It is not surprising, therefore, that alteration of a leucine to phenylalanine at this point will affect the rate of bufuralol metabolism in 2D1, but not that of debrisoquine, as the former substrate molecule is likely to occupy this region (i.e. L331) of the 2D1 active site, whereas the latter substrate, debrisoquine, being of a smaller size, does not appear to interact significantly with any amino acid residues in the vicinity. Genetic polymorphism in CYP2D6 has been associated with certain allelic variants in the enzyme itself, and some of these result in either reduced catalytic activity or an inactive form of the enzyme (Meyer, 1991; Price-Evans, 1993). One interesting example is the 2D6-F allele which gives rise to ‘poor-metabolizer’ status, as the 2D6 isoform in this case is completely devoid of debrisoquine 4-hydroxylase activity. This mutant allele has been related to a non-conservative change in the coding region of the 2D6 gene which will bring about a replacement of a glycine residue, at position 212, by glutamic acid. In the alignment (Figure 6.19) used to generate the current 2D6 model, this residue change occurs at position 180, which is one position upstream of that responsible for the change in substrate specificity in the 2A subfamily (Lindberg and Negishi, 1989). This amino acid residue position will correspond to a region of the F helix (Lewis, 1995a) which lies across the upper portion of the putative active site and, consequently, is likely to affect substrate binding. The presence of an acidic amino acid residue at this point could bring about a considerable reorientation of a basic nitrogenous substrate in the 2D6 active site and, in the case of debrisoquine, metabolism would be extremely difficult if the substrate were to bind preferentially in this region, which is likely to be a substrate access channel. However, three positions downstream of this site there is another glutamate which is invariant in all of the 2D subfamily proteins, together with a third glutamate as the next residue in 2D6, although this is not well-conserved in the other 2D isozymes (Figure 6.19). It is possible that this latter position may represent an alternative anionic binding site for relatively large 2D6 substrates, such as tropisetron (Fischer et al., 1994), as it is difficult to explain the enzyme specificity of these chemicals in terms of the generally


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accepted substrate template model (Islam et al., 1991). Furthermore, at the other end of the scale with reference to size of substrates, the MAO inhibitor deprenyl has been shown to possess 2D6 specificity (Grace et al., 1994) despite the fact that its protonated nitrogen is only about 2.5 Å from the known site of metabolism. Modelling of deprenyl in the current 2D6 model (Figure 6.21) indicates that the proton bound to the positively charged nitrogen is roughly 3.0 Å away from the position of oxygenation but, nevertheless, the substrate can form an ion-pair with D260 due to the conformational flexibility of the aspartate side chain. The distance between one of the aspartate carboxylate oxygen atoms and the iron oxene is 7.4 Å in the energy-minimized 2D6 model, whereas the corresponding distance of the proton bonded to the basic nitrogen in debrisoquine is 4.4 Å from the site of metabolism in this substrate (Figure 6.20). In addition to substrates, the specific 2D6 inhibitor, quinidine (Boobis et al., 1990; Smith, 1991) readily fits the putative active site of 2D6 where its quinoline nitrogen can ligate the heme iron, and the protonated nitrogen atom on the quinuclidine ring will form an ion-pair with D260 (Lewis, 1995a). Moreover, a hydroxyl group on the inhibitor could form a hydrogen bond with S263 (position 304 in 2D6). Interestingly, the enantiomer of this inhibitor, namely quinine, is specific for the rat orthologue 2D1, which has a conservative change of S T at the 263 position (Figure 6.19). It is possible to show that the alteration in chirality between the two inhibitors will enable the hydroxyl group of quinine to hydrogen bond with T263 in 2D1 (Lewis, 1995a). Consequently, the CYP2D models show satisfactory correlations with known substrate specificity, together with other information obtained from site-directed mutagenesis studies and consideration of allelic variants. It is hoped that further investigations will rationalize in molecular terms a number of other interesting observations within this important P450 subfamily, such as the finding that the mouse orthologue, 2D9, can hydroxylate testosterone in the 16 -position (Funae and Imaoka, 1993) but is unable to metabolize debrisoquine (Paine, 1991). 5. CYP2E subfamily. Site-directed mutagenesis studies on rabbit CYP2E1 (Figure 6.22) show that replacement of the conserved distal threonine, T268 (position 303 in CYP2E1) with serine altered the spinstate equilibrium in favour of the low-spin form, and also modified the regioselectivity of fatty acid hydroxylation (Fukuda et al., 1993). These findings point to the possible involvement of this threonine residue in substrate interactions, although the removal of the -methyl group by the change T S will probably also allow a water molecule to ligate the heme in the resting state of the 2E1 enzyme, such that its catalytic activity may be affected. Interestingly, replacement of the conserved threonine by valine did not markedly affect the spin-state equilibrium in 2E1 (Fukuda et al., 1993), presumably because such a change would not necessarily lead to a less-hindered heme environment. The known substrate specificity of CYP2E1 isozymes for relatively small-sized molecules suggests that the active site will be conformationally restricted by various amino acid residues, and the human CYP2E1 model (Figure 6.23) supports this, as there are several bulky amino acid side chains in the putative heme environment, including four phenylalanine residues, F78, F88, F181 and F263, which appear to form two pairs of - stacked phenyl rings, respectively. Moreover, there are a further four bulky aliphatic hydrophobic residues in the active site, namely, 1436, 1438, L327 and V328, which will also constrain the pocket so that small-sized hydrophobic substrates, such as benzene, will be preferred. Furthermore, the specific marker substrate, pnitrophenol, can readily fit the putative CYP2E1 active site via interactions with two complementary amino acid residues which orientate the molecule for the known position of hydroxylation (Figure 6.23). The phenolic hydroxyl group of this substrate can form a hydrogen-bonded interaction with T266 (position 301 in CYP2E1) whereas the p-nitro moiety could interact with R82, which may also be involved in the binding of other polar or fatty acid substrates via hydrogen bonding or electrostatic interactions. In fact, many of the known CYP2E substrates, such as ethanol, acetone, DMN and chlorzoxazone, can fit the putative 2E active site, where a number of key amino acid residues (mentioned previously) orientate

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each substrate for metabolism at the preferred position. Moreover, there are several differences between rat, mouse and human CYP2E1 (Figure 6.22) which may explain the experimentally observed species differences in metabolism, such as the formation of butadiene epoxides (Csanady et al., 1992; Duescher and Elfarra, 1994; Bond et al., 1995; Melnick and Kohn, 1995), because even conservative changes in the P450 active site can have a marked effect on substrate metabolic profiles, as have been reported in orthologous P450s from other families and subfamilies. 6.2.5.3 The CYP3 family In contrast to the CYP2E model, that of CYP3A4 (Figure 6.24) displays a highly unrestricted active site, in keeping with the known structural diversity of CYP3A (Namkung et al., 1988) substrates, some of which possess relatively large-sized molecules (Smith and Jones, 1992). However, it is important to show that substrates ranging in size from simple steroids to macrocyclic antibiotics, such as cyclosporin, are able to both fit into the putative 3A4 binding site, and become orientated via interactions with certain amino acid residues, such that they will be metabolized in the known positions. The model of the human isozyme, CYP3A4, does indeed demonstrate that diverse substrates can readily occupy the heme pocket for oxygenation at the experimentally observed sites (Lewis and Lake, 1996c). For example, cyclosporin (Kronbach et al., 1988) is able to bind within the putative active site of CYP3A4 by forming a number of hydrogen-bonded and hydrophobic interactions with several complementary amino acid residues (Figure 6.24). In particular, the macrolide substrate forms two hydrogen bonds with N74, a residue which appears to be involved in binding many other 3A4 substrates, and one of the amide groups of cyclosporin can be orientated parallel to the phenyl ring of F72; this residue also seems to be important for the binding interactions with substrates containing at least one aromatic ring (see Figure 6.25 for the 3A alignment). Testosterone can also fit the 3A4 active site, in an orientation which will enable 6 -hydroxylation to occur, with its D-ring hydroxyl group forming a hydrogen bond to N74, whereas the A-ring carbonyl oxygen atom can accept a hydrogen bond from S271. The steroidal analogue gestodene, a specific inhibitor for CYP3A4, can be orientated within the 3A4 active site with its D-ring ethynyl moiety above the heme iron, by forming a hydrogen bond with N74 via its A ring carbonyl group; this position would then block the oxygen access channel close to the conserved distal threonine, T268, by the occupancy of the inhibitor’s ethyl substituent which lies between the C and D rings of the steroid nucleus. The weaker and less-specific inhibitor, ketoconazole, may also bind in the CYP3A4 active site via interactions with N74 and F72, such that its imidazole ring nitrogen atom ligates the heme (Figure 6.26). Other substrates of diverse structure which can, nevertheless, fit the 3A4 active site in an orientation for oxygenation at the known positions include: omeprazole, tamoxifen (Wiseman and Lewis, 1996) and granisetron, where the N74 and F72 pair appear to form complementary binding interactions with each of these substrates (Figure 6.27). Although there have been no site-directed mutagenesis studies reported for this P450 family, the relatively high amino acid sequence homology between P450BM3 and the CYP3A subfamily (Figure 6.25) tends to instill a reasonable degree of confidence in the models produced, especially as many CYP3A substrates fit the enzyme active site irrespective of their structural complexity or molecular size; and the orientations of these substrates appear to be constrained by certain amino acid residues that position the relevant molecules for metabolism at the experimentally observed positions.


253

Figure 6.22 Alignment between CYP102 and members of the CYP2E subfamily.

6.2.5.4 The CYP4 family As long chain carboxylic acids are substrates of both P450BM3 and most of the enzymes in this family, it is

254


perhaps not surprising that it is relatively straightforward to show that typical substrates, such as lauric acid, can bind to the putative actives site of, for example, the human orthogue, CYP4A11 (Lewis, 1995a). There is considerable sequence homology between P450BM3 and most members of the CYP4 family, particularly those of the CYP4A subfamily (Figure 6.28). However, a small number of significant changes are able to modify the substrate specificity of CYP4A subfamily isozymes from that of long chain fatty acids to prostaglandins, in the case of CYP4A4 (Lewis, 1995a). In CYP4A11, lauric acid can occupy the putative active site, orientated for either - or - 1 hydroxylation, by forming an ion-pair between its carboxylate head group and the side chain of R188 (Lake and Lewis, 1996). A number of complementary hydrophobic amino acid residues line the heme pocket in the CYP4A11 model (Figure 6.29) and appear to ‘mould’ the conformation of the dodecanoate substrate to produce an optimum fit (Lake and Lewis, 1996). The active site phenylalanine residue F87 in P450BM3, which ensures that only -2 hydroxylation of long chain fatty acids occurs, becomes leucine in the CYP4A subfamily and this change allows - and -1 hydroxylation of the aliphatic chains of their carboxylic acid substrates (Lake and Lewis, 1996). In addition, MEHP will occupy the CYP4A11 active site such that end-of-chain hydroxylation can occur; the specific inhibitor, 11-undecynoic acid, being a substrate analogue of lauric acid, can also fit the enzyme active site in a similar way. Furthermore, the substrate preference of the rabbit orthologue CYP4A4 for prostaglandin E2 can be explained by the model of this isozyme, where several key amino acid changes enable hydrogen bonding to complementary groups on the substrate (Lake and Lewis, 1996). Finally, the model of CYP4F3 (Figure 6.30) can be shown to accommodate the specific endogenous substrate, leukotriene B4, where a relatively small number of key amino acid residue changes give rise to complementarity with groupings on the substrate molecule. 6.2.5.5 The CYP11 family As this mitochondrial P450 family does not utilize an NADPH-dependent cytochrome P450 oxidoreductase as a redox partner, it might be expected that P450BM3 would not be a particularly appropriate template for modelling these isozymes. However, this does not appear to be the case, as the model of CYP11A1 (Figure 6.31), generated from P450BM3, agrees closely with known experimental data on substrate metabolism, inhibitor binding, interaction with adrenodoxin, and the postulated shuttle mechanism governing electron transfer from adrenodoxin reductase (Lambeth, 1990). One similarity between the substrates of P450BM3 and the CYP11 family is that they contain a system of aliphatic carbon atoms of approximately the same length (if one compares the structures of cholesterol and lauric acid, for example) with hydrophilic head groups at roughly the same distance from the respective sites of metabolism (Lake and Lewis, 1996). It may be, therefore, that enzymes of the CYP11 and CYP4 families and P450BM3 are evolutionarily related with respect to the metabolism of C12 substrates. The endogenous substrate, cholesterol, can fit the putative active site of CYP11A1 via hydrogen bond formation between the A-ring hydroxyl group of the substrate and the complementary amino acid residue, T330, which is present in the heme pocket of the bovine CYP11A1 model (Figure 6.31). There are also several hydrophobic aliphatic amino acid residues which appear to be relevant for defining substrate


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specificity, including L87, L181, V329, A263 and L437. The combination of these side chains appears to orientate the cholesterol molecule for oxygenation at the 20, 22 carbon-carbon bond, which is cleaved during the metabolism of this substrate. An intermediate stage in the conversion of cholesterol to pregnenolone involves the formation of (R)-22-hydroxycholesterol, and this intermediate compound could bind in a slightly different manner to cholesterol via a hydrogen bond to S328. The specific inhibitor, (20R)-20-phenyl-5-pregnene-3 , 20-diol, will also dock into the CYP11A1 site by heme ligation involving the exocyclic hydroxyl group and hydrogen bonding between the A-ring hydroxyl and T330. The phenyl

256


ring of this inhibitor tends to obstruct the oxygen access channel to the heme iron in this orientation, which may represent a possible mode of inhibition by this compound. It is also possible to show how the redox partner, adrenodoxin, is able to interact with the bovine CYP11A1 model via electrostatic binding between the two protein surfaces (Figure 6.32). As the relevant complementary acidic and basic residues have been identified (Table 6.11) in both adrenodoxin and CYP11A1, interactive docking between the two models indicates a likely mode of binding which leads to electron transfer. A possible way in which these oppositely-charged residues may interact in the adrenodoxin-CYP11A1 complex in shown in Figure 6.32, and this bears close similarity with the putidaredoxin-P450cam binary complex, which is described in a following section.


257


6.2.5.6 The CYP17 family The known endogenous steroidal substrate for this enzyme, progesterone, can readily fit into the putative active site of the CYP17A1 model for 17 -hydroxylation by forming hydrogen bonds to T77 and Y177 from

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its A-ring hydroxyl and 17substituted ketone groupings, respectively (Figure 6.33). There are also several hydrophobic residues in the putative binding site of 17A1 which also contact complementary substructures of the substrate molecule. Furthermore, one member of a class of known CYP17A inhibitors, namely, cyclohexyl esters of pyridyl-substituted carboxylic acids (Laughton and Neidle, 1990), is able to occupy the


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enzyme’s active site in an analogous fashion to that of the substrate, where the ester carbonyl moiety can form a hydrogen bond to Y177, with the pyridyl nitrogen atom in position for ligating the heme iron. This inhibitor is superimposable onto the progesterone substrate in the CYP17A1 active site, with its cyclohexyl group overlaying the C ring of progesterone (Figure 6.34). The conformation of the inhibitor used in the docking study (shown in Figure 6.34) is essentially the same as that reported from the crystallographic and modelling investigations on these inhibitory agents (Laughton and Neidle, 1990). 6.2.5.7 CYP19 family The model of CYP19A1 (aromatase) is able to show how one of the known inhibitors, 4hydroxyandrostenedione, and the specific substrates, androstenedione and testosterone, can occupy the putative active site of this enzyme by forming hydrogen bonds with key amino acid residues within the heme environment. For example, the D-ring carbonyl (or hydroxyl group) on the steroids can accept a hydrogen bond from S72, and S437 may be involved in hydrogen bonding with the A-ring carbonyl oxygen of the substrates (Figure 6.35). Site-directed mutagenesis experiments have focused on the region close to the invariant threonine distal to the heme (Zhou et al., 1991; Kadohama et al., 1992; Graham-Lorence et al., 1990, 1991, 1994), and these studies indicate that the I helix could represent a point of contact for the orientation of substrates. Possibly, there is more than one orientation of substrates for this enzyme, which can lead to the formation of the aromatized product, as there are intermediate stages in the overall reaction which, presumably, would require some movement of the substrate within the active site. 6.2.5.8 The CYP101 family The crystal structure of P450cam (CYP101), until recently has been the only template available for modelling P450 isozymes, and details of this structure have been reported by Poulos and co-workers, including the effects of various substrates, ligands and inhibitors (Poulos et al., 1985, 1986, 1987a and b; Poulos, 1985, 1988a and b, 1991; Poulos and Raag, 1992; Raag and Poulos, 1989, 1992; Raag et al., 1990, 1991, 1993). It has also been shown that cytochrome b5 is able to bind to P450cam in a similar manner to the natural redox partner, putidaredoxin (Stayton et al., 1989; Stayton and Sligar, 1990; Sligar et al., 1991); whereas, it would appear that the C-terminal amino acid of putidaredoxin (a tryptophan residue) is involved in electron transfer to the P450 concerned (Davies et al., 1990; Davies and Sligar, 1992; Sligar et al., 1974). A model has been proposed for the mode of interaction between putidaredoxin and P450cam which could lead to electron transfer from the iron-sulphur protein to the bacteral P450 enzyme (Baldwin et al., 1991) and it is possible to demonstrate that both cytochrome b5 and putidaredoxin contain anionic surface groups which can ion-

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pair with a quartet of complementary basic residues (Table 6.11) at the proximal heme face of P450cam, as shown in Figure 6.36. Interestingly, the sets of negatively charged acidic residues on putidaredoxin associated with P450cam and putidaredoxin reductase binding are on different surface regions of the redoxin model (Figure 6.37) and there is considerable analogy with those utilized for similar purposes in adrenodoxin, as mentioned previously (Table 6.11). The various features of P450cam that are of structural and functional importance have been compared with those in the more recent prokaryotic crystal structure, P450BM3 (Lewis, 1995a) and the reader is referred to this publication for further details. 6.2.5.9 The CYP102 family There are now two crystal structures (pdb codes:- 2hpd and 2bmh) available for the hemoprotein domain of P450BM3 (CYP102A1), although only the original model (2hpd) has been described in the literature (Ravichandran et al., 1993). Table 6.12 Table 6.11 Interactions between P450s and their redox partners (References: Stayton et al., 1989; Adamovitch, et al., 1989; Sligar et al., 1991; Geren et al., 1984; Dailey and Strittmatter, 1979; Lambeth et al., 1984) (a) Likely residues involved in redox partner interactions with P450cam and P450scc Pdx CAM b5 Adx 11A1 D36 … K344 … E44 D36 … K315, 319 D12 … R364 … heme CO2– D12 … K114 D40 … R72 … E48 E42 … K63 W97 CO … R112 … D60 D94 … K97 D98 … K245 (b) Likely residues involved in interactions between P450 redox partners and their respective reductases Pdx Adx b5 D59 (58) E59 (60) E37 E67 (65) D67 (72) E43 E69 (67) E69 (74) E44 E72 (72) D72 (79) E48 Notes: The references given above relate to reported identification of certain residues previously published, which are either in broad or exact agreement with those listed in the tables above. The Pdx and Adx residues involved in P450 interactions have been largely determined by interactive molecular modelling docking studies and are unpublished to date. The numbering of residues in CYP11A1 refers to their alignment positions relative to P450BM3. The numbering of P450cam and cytochrome b5 refer to their actual positions in their respective sequences. The numbering of residues in Pdx and Adx refers to their alignment positions relative to ferredoxin (pdb code : 3fxc) with the actual positions in their respective sequences given in parentheses. Pdx=Putidaredoxin Adx=Adrenodoxin b5=cytochrome b5 CAM=P450cam(CYP101) 11A1=CYP11A1(P450scc) ¼ denotes ion-pairine interactions


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Table 6.12 Key residues conserved or modified between CYP102 and CYP101 CYP102

CYP101

G46d

G60e

Comments

Inter-helical turn K59 R72a Ion-pairing to reductase/redoxin F87d P100 F87 blocks -hydroxylation of substrate in CYP102 W96C Q108 Electron transfer from reductase to heme in CYP102 a H100 R112 Ion-pairing to reductase/redoxin, heme and/or b5 K113 K126b Ion-pairing to reductase K187 R186e Ion-pairing to D251 in CYP101 R223 R211b Ion-pairing to reductase in CYP102 d b,e K224 R212 Reductase interaction and ion-pairing to D233/D218 D232d D218e Ion-pairing to K224/R212 E267f D251e, f Ion-pairing to R186 in CYP101, involved in catalytic mechanism T268 T252f Involved in oxygenation process and mechanism d e E320 E287 Ion-pairing to R323/R290 R323d R290e Ion-pairing to E320/E287 and redoxin or b5 A328 V295e Hydrophobic interactions with substrate K349b K314b Ion-pairing to reductase in CYP102 d e R398 H355 Ion-pairing to reductase/redoxin and heme C400d C357e Ligation to the heme L437 I395e Hydrophobic interactions with substrate Note: References tabulating functional or structural roles of the residues listed above are as follows: a Stay ton and Sligar, 1990; Stay ton et al., 1989 b Bernhardt et al., 1984, 1987, 1988, 1989 c Baldwin et al., 1991 d Ravichandran et al., 1993 e Poulos et al., 1985, 1986, 1987 f Gerber and Sligar, 1994, 1992; Imai et al., 1989

provides a comparison between certain elements of the P450cam (pdb code:- 2cpp) and P450BM3 crystal structures, whereas Tables 6.4 and 6.5 list secondary structural motifs for these two crystallographic models. Although there are several features of commonality between P450cam and P450BM3, the latter structure is clearly the more suitable template for eukaryotic P450 modelling, as has been described previously (Lewis, 1995a). However, it is unfortunate that neither of the two available crystal structures of P450BM3 contains a bound substrate so consequently it is not entirely clear whether there is any significant difference between the substrate-bound and substrate-free conformations. Information from site-directed mutagenesis and NMR spectroscopic studies have provided some useful criteria to enable a description of the likely mode of substrate binding. For example, the roles of R47 and F87 in substrate interactions have been identified from mutagenesis experiments (Graham-Lorence et al., 1994) where it has been shown that R47 probably ion-pairs with the carboxylate head group of the long chain fatty acid substrate, e.g. lauric acid, and F87 sterically hinders the - and -1 carbon atoms of the substrate from oxygenation, such that only -2 hydroxylation is possible. More recently, NMR spectroscopic measurements have indicated the probable distances between the extremities of the lauric acid substrate molecule and the heme iron (Gibson et al., 1995). Molecular modelling of the interaction between

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lauric acid and P450BM3 (Lake and Lewis, 1996) is in very close agreement with the results of the NMR studies based on the paramagnetic shifts of protons in the vicinity of the heme iron (Figure 6.38). The methylene protons are found to be about 16.25 Å distance from the iron, whereas the NMR data reports this as 16.4 Å (Gibson et al., 1995). The -methyl protons, on the other hand, lie at around 5.55 Å from the heme iron in the model, which is in good agreement with the NMR value of 5.6 Å. Consequently, the energyminimized structure of P450BM3 with bound substrate represents an extremely good fit with the available experimental information. 6.2.5.10 The CYP108 family The crystal structure of P450terp (CYP108) has been published recently (Hasemann et al., 1994) and the overall disposition of the model (pdb code: Icpt) appears to confirm the view that the tertiary fold of P450s is largely conserved, at least between prokaryotic forms. In fact, there is significant conservation between the protein sequences of CYP101, CYP102 and CYP108, including 36 invariant amino acid residues, and it is possible to compare the C tracings of these three bacterial crystal structures (Figure 6.3). The secondary structural elements also show broad commonality and Table 6.6 lists these for the P450terp structure. Moreover, the active site regions of these enzymes display good agreement with the topology studies of Swanson and co-workers (Swanson et al., 1991) which were published before the CYP102 and CYP108 crystal structures were available. Figure 6.39 shows a comparison between the three active sites for these prokaryotic forms, where it can be seen that the presence of certain amino acid residues close to the heme moiety are responsible for sterically hindering migration of the phenyl group, in line with the results (Table 6.13) of the phenyl diazene inhibition studies reported by the Ortiz de Montellano group (Swanson et al., 1991; Tuck et al., 1992). Although there is no substrate present in the P450terp crystal structure, it is possible to demonstrate the mode of interaction between -terpineol and the active site of this enzyme which agrees with the known position of oxygenation. Modelling indicates that S101 is able to form a hydrogen bond with the substrate’s hydroxyl group, and there are several hydrophobic amino acid residues within the heme pocket which appear to orientate the substrate for hydroxylation of the cyclo-hexenyl methyl substituent, as shown in Figure 6.40. At present, no site-directed mutagenesis studies have been reported on CYP108, and the lack of an important segment of polypeptide between the F and G helices gives rise to some uncertainty regarding the potential utility of this structure as a possible molecular template for other P450 isozymes. However, the current P450BM3 -derived models all demonstrate a satisfactory agreement with experimental data on the relevant eukaryotic forms, thus indicating that this is the preferred template for constructing models of eukaryotic P450s (Lewis, 1995a). 6.3 Evaluation of P450±mediated toxicity by the COMPACT approach A knowledge of the P450 substrate specificity and the structural characteristics of these different classes of substrates (and inducers) has facilitated the development of


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Table 6.13 Comparison between ratios of N-phenyl protoporphyrin IX regioisomers and active site residues in P450s Rations of regioisomersa

Residues in corresponding vicinityb

CYP

NB

NA

Nc

ND

B

A

C

1A1 (rat) 1A2 (rat) 2B1 (rat) 2B2 (rat) 2B4 (rabbit) 2B10 (mouse) 2B11 (dog) 2E1 (rat) 102 (BM3) 101 (CAM) 108 (TERP)

0 2 0 0 0 0 0 0 2 0 0

2 9 2 3 3 3 4 1 10 0 0

0 2 0 0 2 1 1 0 2 1 0

1 3 3 2 3 3 2 2 1 4 1

AGF AGF AGT AGT AGT AGT AGT AGR AGH VGG AGH I helix

DTI ETV ETS ETG ETT ETS ETT ETT ETT DTV DTT

T, A T, A I, A I, A I, A I, A V, A I, A F, A T, L T, A B helix and

D T, V T, V I, V I, A I, I I, I V, L I, V F, A V V sheet 1

a

Swanson et al., 1991; Tuck et al., 1992 From crystal structures and sequence alignments Note: Designation of the pyrrole rings in the heme group; b

a method for the prediction of P450-mediated toxicity, known as COMPACT (Lewis, 1992a, b and d; Lewis et al., 1990a and b; 1993a and b, 1994a and d; loannides et al., 1993, 1994, 1995). This technique utilizes molecular modelling procedures to calculate the electronic structures and spatial dimensions of molecules in order to assess the likelihood that the chemicals concerned may act as substrates (or inducers) of various P450 isozymes and, consequently, enables determination of the possible toxicological or metabolic consequences of exposure to such compounds. It has been possible to determine the structural criteria which enable various chemicals to act as substrates and/or inducers of CYP1 (Lewis et al., 1986b) and the combination of molecular planarity with electronic activation energy is discrimi-natory for CYP1 specificity relative to that of other P450s (Lewis et al., 1987, 1989a and b; Parke et al., 1988a, 1990a,b and c; 199la and b). Furthermore, it can be shown that the magnitude of CYP1 induction is largely determined by molecular shape (Parke et al., 1986) whereas the calculated electronic activation energy correlates with both

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mutagenicity (see Maron and Ames (1983) for a description of the Ames test for bacterial mutagenicity) and carcinogenicity (Lewis, 1995b; Lewis et al., 1995f, 1996b). Computer-optimized molecular parametric analysis of chemical toxicity (COMPACT) represents a relatively straightforward method for the prediction of carcinogenicity (and possibly other forms of toxicity) in diverse series of chemicals where P450s are likely to be involved, and validation of this system against rodent carcinogenicity has produced concordances of between 70 and 90 per cent (Lewis et al., 1995d; Brown et al., 1994; Lewis, 1994c; Lewis et al., 1993). It has been found that the inclusion of additional structural parameters can provide a means of distinguishing CYP2E substrates and inducers from those specific for other P450s, thus enabling an overall improvement in the predictive power of the COMPACT technique in identifying P450-mediated carcinogenicity (Lewis et al., 1995d). Many known rodent and human carcinogens are correctly identified by this method, which also gives predictions of carcinogenicity subsequently found to produce an over 70 per cent accuracy (Lewis et al., 1995d), and it is also possible to differentiate between Table 6.14 Structure activity data for 14 structurally diverse P4501 inducers (Reference to induction data: loannides and Parke, 1993) Compound

Area/depth (Å)

E (eV)

µ (D.)

Log induction potential

1. Benzanthracene 12.019 7.3951 0.043 3.2553 2. Benzpyrene 12.023 6.8058 0.033 3.1761 3. 3-Methylcholanthrene 8.130 7.2547 0.815 2.6021 4. -Naphthoflavone 13.821 7.9668 4.168 2.7782 5. -Napthylamine 7.392 8.0543 1.623 1.8751 6. Acridine Orange 7.574 6.7297 0.601 1.5185 7. 4-Aminobiphenyl 8.166 8.3128 1.692 1.3979 8. Safrole 1.949 9.0495 0.744 1.3424 9. 2-Aminoanthracene 9.141 7.1113 1.720 1.2041 10. 2-Aminofluorene 5.197 8.0134 1.773 1.2041 11. -Naphthylamine 4.773 7.9269 1.547 1.0000 12. o-Toluidine 3.275 9.0332 1.563 0.9031 13. Benoxaprofen 2.609 8.1023 4.513 0.3010 14. Cimetidine 2.168 8.6993 6.762 −0.0969 Log induction†=0.10Area/depth+0.58 E–0.27µ –5.17 potential (±0.01) (±0.19) (±0.05) n=14; s=0.315; R=0.96; F=42.0 E(HOMO), E(LUMO)=energies of the highest occupied and lowest empty molecular orbitals E=E(LUMO)–E(HOMO) µ =dipole moment Area/depth=the ratio of the product of molecular length and width, relative to molecular depth † see Figure 6.41 for a plot of this relationship

structural pairs using certain molecular descriptors (Lewis et al., 1994a; loannides et al., 1993). Quantitative Structure-Activity Relationships (QSARs) on many series of structurally related chemicals provide considerable support for the COMPACT philos-ophy towards P450-related activity and toxicity (Lewis, 1992a and b, 1995b; Lewis et al., 1994a; Lewis and Parke, 1995). It can also be demonstrated that


265

Figure 6.41 Graph showing the agreement between observed values of the logarithm of induction potential, for CYP1 induction by a structurally diverse group of 14 CYP1 inducers, and calculated values of the logarithm of CYP1 induction potential. The relevant data are presented in Table 6.14.

COMPACT parameters are able to describe quantitatively the induction potential of diverse groups of CYP1 inducers, as shown in Table 6.14 and Figure 6.41. Furthermore, Hansch and co-workers have reported that the hydrophobic parameter, log P, is an important factor for rationalizing potency differences in series of P450 substrates (Hansch and Zhang, 1993; Debnath et al., 1992) and it is possible to include this factor in carcinogenicity prediction, together with the use of structural alerts for direct-acting effects not associated with P450 pathways (Lewis, 1994c; Lewis et al., 1995d). The QSAR studies which exhibit satisfactory correlations between molecular and electronic structural parameters and both P450-related activity and toxicity, are summarized in Table 6.15 which represent the results of a large number of investigations carried out in the author’s laboratory (Lewis, 1995b). To give an example for how the use of three COMPACT parameters can classify a number of known carcinogens in terms of their possible induction of (or activation by) CYP1 and CYP2E, Table 6.16 lists the relevant information for 10 structurally diverse chemicals. Figure 6.42 presents a two-dimensional COMPACT plot (of area/ depth2 versus E) for these compounds which classifies 8 of the 10 chemicals as possessing CYP1 specificity: this is in full agreement with known experimental find-ings (Lewis et al., 1994a). Concerning the two compounds not activated via CYP1, the small molecular diameters of these chemicals suggests that they are able to act as substrates of CYP2E, and their relatively low activation energies ( E values) indicate that they may give rise to reactive intermediates which could be either carcinogenic or show some other form of toxicity. Although all of the 10 chemicals are known carcinogens, it is not possible to predict their likely organ of toxicity, however, using the COMPACT technique. In order to achieve this, it would be necessary to investigate the various levels of different P450s in a number of organs, and assess the potential for phase II conjugation (Ghauri et al., 1992). Moreover, the possibility of other mechanisms of toxicity, which may be organ specific, would need to be considered.

266


6.4 Conclusions and future prospects Cytochrome P450 is one of the most extensively studied enzyme systems. The number of research publications reported annually exceeds 1000 and is increasing yearly, thus reflecting the considerable scientific interest in the enzyme superfamily, especially its pivotal role in the toxicology of foreign compounds (Connors, 1978; Coon et al., 1980; Alexander and Goff, 1982; Guengerich, 1991b, 1992c). By combin-ing the considerable amount of accumulated experimental data on compound Table 6.15 QSARs within P540 substrates, inducers and inhibitors (Reference: Lewis, 1995b) (A) Cytochromes P4501 Compounds

Equation

n

s

R

F

Key to biological activity

log EROD=0.29E(HOMO) +0.80 1/w+2.22 (±0.12) (±0.26) log EROD=0.51E(HOMO)– 0.17 a/d2–8.02Q3H+9.29 (±0.10) (±0.04) (±2.14) 2. Benzimidazoles pI50=0.07 a/d + 0.01µ 2+2.33 (±0.01) (±0.003) 3. Methylene dioxybenzenes pI50=0.23 a/d–5.39 (±0.03) pI50=1.22 a/d2–5.68 (±0.14) 4. Resorufins log EROD=0.08SN+5.17 (±0.01) 5. Ellipticines pI50=–0.26 a/d2 + 4.07 1/w – 3.54 (±0.11) (±1.12) 6. EROD=22.15 1/w –19.92 Cyclopentaphenanthrenones (±6.45) 7. Polychlorobiphenyls pEC50=0.40 a/d2+3.19 (±0.08) EROD=0.49 a/d2+3.87 (±0.12) 8. TCDDS log Ah=0.25 a/d2+447.8Q6H –3.73 (±0.05) (±85.12) log A=5.53 E + 46.57 8 – 234.5 (±1.01) (±4.53)

14

0.313

0.75

7.2

EROD=Ethoxyresorufin O-deethylase activity

14

0.220

0.87

10.7

EROD=Ethoxyresorufin O-deethylase activity

8

0.127

0.98

53.0

8

0.268

0.93

40.9

I50=Inhibition of AHH activity I50=EROD inhibition of uM

8

0.208

0.96

71.7

I50=EROD inhibition in µ M

8

0.094

0.96

80.0

16

0.492

0.71

6.7

EROD=Ethoxyresorufin O-deethylase activity I50=4S protein binding affinity

5

3.71

0.89

11.8

15

0.650

0.82

25.9

10

0.777

0.83

18.0

6

0.181

0.96

16.3

6

0.256

0.99

54.0

9. DCDDS

8

0.550 0.87

19.3 EC50=Ah receptor binding affinity

7

0.692 0.88

16.6 EROD=Ethoxyresorufin O-deethylase activity

1. Aromatic amines

pEC50=−3.94E(HOMO)–24. 87 (±0.90) EROD=10.32Q7L+5.09 (±2.53)

EROD=Ethoxyresorufin O-deethylase activity EC 50=Ah receptor binding affinity EROD=Ethoxyresorufin O-deethylase activity Ah=Ah receptor binding affinity A=Relative potency of AHH induction


10. Coumarins 11. Chrysenes

12. Polybromobiphenyls

13. Dichlorobiphenyls

14. Polyaromatic hydrocarbons 15. Benoxaprofen analogs

16. Various

pI50=5.64E(HOMO)+48.05 (±0.88) log EROD=33.65Q12L+4. 18µ 2 –0.19 (±16.90) (±1.78) log GluP1=32.70Q12–0.12 +4.50 (±7.98) (±0.04) log MC=0.43 a/d2 +0.76µ –3. 56 (±0.12) (±0.21) A=2.72Q6+1.96E(HOMO) +0.01SN+6.43 (±0.25) (±0.27) (±0. 003) log A=1.591ogP–2.29µ 2 – 10.05 (±0.66) (±0.98) pLD50=–0.16SN5 –1.74E (HOMO) –17.22 (±0.04) (±0.33) logke=–0.34 E+4.44 (±0.05)

267

23

0.482 0.82

9

0.175 0.80

10

0.264 0.89

13.9 GluP1=Glu-P-1 activation

14

1.076 0.81

10.6 MC=Rate of metabolism by 3MC-induced microsomes

11

0.086 0.98

61.1 A=Ratio of rates of metabolism

6

0.74

3.7

16

0.124 0.83

14.5 LD50=Lethal dose

26

0.730 0.82

49.9 ke=electron uptake rate constant

0.84

41.6 I50=Inhibition of AHH in rat liver microsomes 5.5 EROD=Ethoxyresorufin O-deethylase activity

A=Carcinogenic potency

Key: E(HOMO)=Energy of the highest occupied molecular orbital (MO) SN=Nucleophilic superdelocalizability E(LUMO)=Energy of the lowest unoccupied MO QN=Electronic charge on atom N E=E(LUMO)–E(HOMO) QNH = HOMO electron density on atom N 1/w=Molecular length/width QNL=LUMO electron density on atom N a/d=Molecular area/depth n=Number of observations a/d2=Molecular area/depth2 s=Standard error µ =Dipole moment R=Correlation coefficient =Polarizability F=Variance ratio logP=Logarithm of the octan-1-o1/water partition coefficient Error limits are given in parentheses (B) Cytochrome P4502E Compounds

Equation

n

s

R

F


1. Alcohols

pIC50=−13.3QcL+6.42 (±1.4) pIC50=0.45 length–4.04 (±0.09)

20

0.337

0.92

93.0

22

0.636

0.74

23.6

IC50=50% Inhibition of aniline phydroxylation IC50=50% Inhibition of aniline phydroxylation

268


(B) Cytochrome P4502E Compounds

Equation

n

s

R

F


2. Nitriles

pKi=−1.25 / E+0.95 (±0.32) log Rate=–0.006 +0.28 (±0.001) pLD50=–0.03 +0.43 (±0.01) log P=0.49 –0.50 (±0.07) pLD50=4.79 E –67.47 (±0.80) pLD50=54.83Q1H+3.16 (±13.32) pLD50=8.80 (±1.88)

13

0.282

0.76

15.3

20

0.046

0.79

30.5

Ki=Ethanol inhibition constant of nitrile metabolism Rate=Ratio of metabolic rates

16

0.199

0.87

42.1

LD50=Lethal dose for 50% kill

18

0.380

0.86

45.1

P=Octanol/water partition coefficient

6

0.334

0.95

35.7

LD50=Carcinogenicity

31

0.602

0.61

16.9

LD50=Carcinogenicity

5

0.087

0.94

15.2

LD50=Rat lethal dose in mg/kg

3. Halothanes 4. Nitrosamines

5. Pyrazines

Key: IC50=Concentration required for 50% inhibition of aniline p-hydroxylation Ki=Ethanol inhibition constant of nitrile metabolism Rate=Ratio of metabolic rates following pretreatment with ethanol with respect to glucose (fold increase in metabolism of ethanol) LD50=Lethal dose for 50% kill logP=Octanol/water partition coefficient =Molar polarizability Length=Molecular length in Å Other symbols defined previously (C) Cytochrome P4502B Compounds

n

s

R

1. Aliphatic amines K1=196.3QNH –268.8 (±29.9)

6

1.670

0.98 83.6

K2=495.9QNH–677.4 (±81.8)

5

2.630

0.96 36.7

2. Benzphetamines 3. Alkylbenzenes

Equation

F

pK1=1.67 log P–0.17(log P)2 –2. 8 0.267 0.98 73.8 15 (±0.26) (±0.05) pK2=1.52 log P–0.15(log P)2–2. 7 0.094 0.99 306.4 68 (±0.13) (±0.02) log P=0.26 –2.55 14 0.488 0.91 60.4 (±0.03) K2=1.74SE–50.44 E–1+1.61 10 0.645 0.93 23.9 (±0.26) (±33.79) K2=1.56SE–2.58 10 0.692 0.91 39.5 (±0.25)

Key to biological activity K1=High affinity binding constant to rat liver microsomal P450 K2=High affinity binding constant to rat liver microsomal P450 K1=defined above K2=defined above P=Octanol/water partition coefficient K2=Spin state equilibrium constant K2=Spin state equilibrium constant


269

(C) Cytochrome P4502B Compounds

Equation

n

s

R

4. Phenytoins

p

20

0.364

0.70 17.6

20

0.291

0.75 23.1

=0.03 –4.62 (±0.007) p =0.03 –4.32 (±0.006)

F

Key to biological activity =teratogenicity in limbic bud development =teratogenicity in CNS development

Key: K2=Spin state equilibrium constant SE=Electrophilic superdelocalizability Other symbols defined previously (D) Cytochrome P4503 Compounds

Equation

1. Steroids

n

s

R

F


a/d2 –0.37

log A=0.23 E+6.75 14 0.115 0.92 32.3 A=% Increase in ethylmorphine ( ±0.05) ( ±0.05) N-demethylase activity 2. Imidazoles –log MIC=9.12Q3+0.40 6 0.058 0.97 73.7 MIC=Minimum inhibitory (±1.06) concentration pI50=0.58 a/d2 –1.14E(LUMO)+2. 7 0.084 0.95 18.6 I50=50% Inhibition of P450 45 (±0.10) (±0.41) Key: A=Ethylmorphine N-demethylase activity MIC=Minimum concentration for inhibition of C-14 demethylation of lanostesol I50=Inhibition (50%) of yeast cytochrome P450 Other symbols defined previously (E) Cytochrome P4501 Compounds 1. Benzanthracenes

Equation

n

s

log M=13.38E(LUMO)+ 2.26 14 0.272 (±2.71) 2. Chrysenes log M=49.16Q12H+7.56 6 0.210 (±12.91) 3. Aromatic amines log M=3.83Q6 –0.08a/d+0.65E 14 0.301 (HOMO)+10.95 (+1.85) (±0.03) (±0. 15) 4. Benzonitrofurans log M=1.92E(HOMO)+26.60 6 1.048 (±0.55) log R=–7.09E(LUMO)+4.72 6 0.474 (±2.57) 5. Heterocyclic amines log M=–0.97 E+14.53 20 0.8 (±0.14) Key: E(LUMO)=Energy of the lowest unoccupied molecular orbital (MO) E(HOMO)=Energy of the highest occupied molecular orbital MO

R

F


0.82

24.4

M=Mutagenicity

0.89

14.51

M=Mutagenicity

0.81

6.14

M=Mutagenicity

0.87

12.3

M=Mutagenicity

0.81

7.6

R=Reversion frequency

0.86

49.4

Salmonella mutagenicity (TA 100)

270


(E) Cytochrome P4501 Compounds Equation E=E(LUMO)–E(HOMO) Q12H=Electron density in the HOMO of atom 12 Q6=Electron charge density on atom 6 a/d=Molecular area/molecular depth n=Number of observations s=Standard error R=Correlation coefficient F=Variance ratio Error ranges are shown in parentheses.

n

s

R

F


Table 6.16 Toxic chemicals with known P450 specificityf (Reference: Lewis et al., 1994a) Compound

P450 involvec d Toxic effect

Area/depth2

E (eV.) Diameter (Å)

1. Benzo(a)pyrene CYP1 Lung carcinogen 12.0 9.3 2. -naphthylamine CYP1 Bladder carcinogen 7.9 12.5 3. TCDD CYP1 Liver carcinogen 7.9 12.7 4. Benzidine CYP1 Bladder carcinogen 7.9 12.7 5. Dimethylaminoazobenzene CYP1 Liver carcinogen 3.8 10.6 6. Diethylstilbestrol CYP1 Uterus carcinogen 4.8 13.0 7. Aflatoxin B1 CYP1 Stomach carcinogen 3.2 11.9 8. Benzene CYP2E Hemoreticular carcinogen 5.4 17.9 9. Halothane CYP2E Immunotoxic carcinogen 2.0 10.8 10. Dimethylnitrosamine CYP2E Liver carcinogen 2.9 14.5 TCDD=2, 3, 7, 8-tetrachlorodibenzo-p-dioxin † Figure 6.42 shows a plot of these data which indicates how CYP1 specificity can be evaluated.

7.6 6.4 7.4 7.4 7.4 7.9 7.7 5.4 5.6 5.1

metabolism with crystallographic details of the various prokaryotic forms and site-directed mutagenesis studies, it is possible to construct models for eukaryotic P450s which are self-consistent with the known facts currently available on substrate specificity (Lewis, 1995a and b). Consequently, this rationalization of P450 characteristics at the molecular level provides a means for the prediction of the route of novel compound metabolism which involves P450-mediated pathways. Although it will be important to produce the crystal structure of at least one eukaryotic P450 to confirm the validity of bacterial P450-derived models, the current procedures described previously may prove to be adequate for evaluating the potential toxicity and metabolic fate of development compounds in both animals and man. Furthermore, there could be many biotechnological applications of P450 research, especially from the use of structural models of the relevant enzymes to derive genetically engineered P450 mutants expressed in heterologous systems (Fowler et al., 1994). For example, the use of bioreactors involving P450s expressed in yeast or bacteria could be made to produce therapeutically important chemicals that are difficult to synthesize by conventional means. Also, there may be applications in the agricultural industry for the generation of novel insecticides produced from genetically-engineered variants of plant P450s which would normally give rise to pyrethroids, or other natural pesticides, biosynthetically. Moreover, it is likely that there could be biotechnological applications in the production of flavours and essences derived from plant or animal sources, whereas it is possible that there are many potential uses of P450 biotechnology in fruit


271

Figure 6.42 COMPACT plot of 10 carcinogenic chemicals based on the data presented in Table 6.16. The points are numbered according to their order in Table 6.16 and the curve indicates the division between CYP1 specificity (above the line) and that for other P450s.

ripening, enhancement of root or stem stability, phytoalexin production associated with the bruising of plants, and in the area of bioremediation. It may be possible to design novel P450 mutant forms which could be used in the treatment or removal of chemical spillage or waste products from industrial processes. However, because of the normally rather low catalytic rate of most P450 enzymes, it is probably essential to derive chimeric systems comprising the P450 and a fused reductase, although P450BM3 may prove to be a useful starting point for such applications, as this is catalytically competent with a high catalytic turnover (Fulco, 1991). Finally, inorganic catalysts utilizing zeolite-caged iron complexes have been shown to possess the capacity to mimic P450 reactions thus indicating potential industrial applications of model P450like systems (Parton et al., 1994). With so many possibilities envisaged, it is likely that there will be an exciting future for the progress of P450 research, in which structural modelling will undoubtedly play a major role.

Bibliography

Sato, R. and Omura, T., 1976, Cytochrome P–450, New York, Academic Press. Ruckpaul, K. and Rein, H. (Eds), 1984, Cytochrome P–450, Berlin: Akademie-Verlag. Ortiz de Montellano, P.R. (Ed.), 1986, Cytochrome P–450, 2nd edition, New York: Plenum. Schenkman, J.B. and Griem, H. (Eds), 1993, Cytochrome P450, Berlin: Springer-Verlag. Omura, T., Ishimura, Y. and Fujii-Kuriyama, Y. (Eds), 1993, Cytochrome P–450 (2nd edition), Tokyo: Kodansha. Guengerich, F.P. (Ed.), 1982, Mammalian Cytochromes P–450, Boca Raton, Florida: CRC Press. Archakov, A.I. and Bachmanova, G.I., 1990, Cytochrome P–450 and Active Oxygen, London: Taylor & Francis. Gibson, G.G. and Skett, P., 1994, Introduction to Drug Metabolism (2nd edition) London: Chapman and Hall. Schenkman, J.B. and Kupfer, D. (Eds), 1982, Hepatic Cytochrome P–450 Mono-Oxygenase System, Oxford: Pergamon. Ioannides, C. (Ed) 1996, Cytochromes P450: Metabolic and Toxicological Aspects, Boca Raton Florida:CRC Press. The following multi-volume book series contains a considerable amount of useful background information on P450 and related subjects: Frontiers in Biotransformation, edited by K.Ruckpaul and H.Rein, Akademie Verlag, Berlin. Volume 1 Basis and Mechanisms of Regulation of Cytochrome P–450, 1989. Volume 2 Principles, Mechanisms and Biological Consequences of Induction, 1990. Volume 3 Molecular Mechanisms of Adrenal Steroidogenesis and Aspects of Regulation and Application, 1990. Volume 4 Microbial and Plant Cytochromes P–450: Biochemical Characteristics, Genetic Engineering and Practical Implications, 1991. Volume 5 Membrane Organization and Phospholipid Interaction of Cytochrome P–450, 1991. Volume 6 Cytochrome P–450 Dependent Biotransformation of Endogenous Substrates, 1991. Volume 7 Relationships between Structure and Function of Cytochrome P–450—Experiments, Calculations, Models, 1992. Volume 8 Medicinal Implications in Cytochrome P–450 catalyzed Biotransformations, 1993. Volume 9 Regulation and Control of Complex Biological Processes by Biotransformation, 1994.

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Index

absorption maximum 1 see also Soret absorption spectra, spectroscopy 11, 14 see also IR, UV, ESR, EXAFS, NMR, etc. acetaminophen see paracetamol acetone 128, 159, 172 2-acetylaminofluorene (2-AAF) 138 ACTH (adrenocorticotrophic hormone) 171 activation of carcinogens 131 of oxygen 21, 28, 37, 83 active site(s) 92–4 CYP1A1 see Figure 6.7, plate section CYP1A2 see Figures 6.8 and 6.9, plate section CYP2A1 see Figure 6.11, plate section CYP2A6 see Figure 6.12, plate section CYP2B1 see Figure 6.14, plate section CYP2C3 see Figure 6.17, plate section CYP2C9 see Figure 6.18, plate section CYP2D6 see Figures 6.20 and 6.21, plate section CYP2E1 see Figure 6.23, plate section CYP3A4 see Figures 6.24, 6.26 and 6.27, plate section CYP4A11 see Figure 6.29, plate section CYP4F3 see Figure 6.30, plate section CYP11A1 see Figure 6.31, plate section CYP17A1 see Figures 6.33 and 6.34, plate section CYP19A1 see Figure 6.35, plate section adrenaline 128 adrenodoxin 2, 118, see also Figure 6.32, plate section aflatoxin B1 92, 138, 151, 164, 188 Ah (aryl hydrocarbon) receptor 66, 138–40, 170, 171, 177, 180, 182, 184 AIA (allylisopropylacetamide) 152, 201 aldehyde dehydrogenase 171 aldrin 152

alignment of sequences 9, 55, 59, 212–40, 214–224, 243– 5, 250–1, 253–5, 263–5, 268–9, 270–1, 273–5 aliphatic alcohols 199 amines 47, 153, 198 hydrocarbons 67 ketones 47, 198, 46 alkoxyresorufins 140 alkyl benzenes 153 allelic variants 67, 127, 141, 159, 266 allene oxide synthase 94 amino acid(s) see also individual amino acids codes 227 composition of P450s 7 residues in P450s 7 specific reagents for 205 4-aminobiphenyl 138 6-aminochrysene 154, 164 2-aminofluorene 165 amphetamine 18 androgens 47 androstenedione 122–6 aniline(s) 45, 153, 159, 200 antibodies 141, 170, 204 arachidonic acid 206 archaebacteria 55, 61, 63 arginine 12, 20, 83 arochlor (1254) 18, 129 aromatase 116 see also CYP19 aromatic amines 45 hydrocarbons 45 aryl hydrocarbon receptor nuclear translocator (ARNT) 139, 170, 177 asparagine 164 317

318

INDEX

aspartate, aspartic acid 83, 93, 157 bacteria 55, 60–6 bacterial P450s 61–6 P450cam 2, 7, 14, 20–3, 29, 30, 34–50, 61, 69, 82–7, 92–7, 104, 105, 107, 191, 201, 210, 277 P450BM3 12, 28–30, 38, 43, 61–5, 69, 83–5, 93, 106, 107, 128, 155, 210–40, 277–9 P450terp 29, 38, 210, 279 system(s) 62, 63, 80, 84 bacteriorhodopsin 108 basal transcription element (BTE) 177, 181–3 bathochromic shift 1, 18 benzene(s) 45, 47, 153, 159 benzidine 138 benzimidazole 200 benzoflavone 200 benzofuran 191 benzo(a)pyrene 67, 138 diol 164 diolepoxide 184 benzphetamine(s) 84, 95, 100, 109, 152 BHA, BHT (butylated hydroxy anisole, toluene) 186 bile acids 118 bilirubin 129 NF see -naphthoflavone bromotoluene 153 buta-1, 3-diene 128, 160 caffeine 18, 140, 200 cambrian 58 cAMP (cyclic adenosine monophosphate) 181, 183 camphor 40, 43, 46, 47, 50, 69, 94, 191 carcinogenicity 138, 142, 149–50, 154, 160–4, 184–8, 290 catalase 21, 186 catalytic cycle 14, 34, 36, 37, 43–5, 80, 81 turnover 64, 82, 100 cDNA (complementary DNA) 55, 67 characterization of P450s 4–8, 192–3 charge-transfer (electron transfer) 2, 13 chenodeoxycholic acid 118 chloramphenicol 152, 201 chlordane 152 chloroalkanes 201 chlorobenzene 153 chloroform 201 chlorzoxazone 159, 162 cholesterol 67, 69, 116, 171

chromosomal locations 65, 144, 175 cimetidine 197 circular dichroism (CD) spectroscopy 19–21 clofibrate 184 clotrimazole 195 CO-complex 1, 12, 13, 18–24, 30, 35, 43, 67, 69, 191, 207 difference spectrum 1, 2 stretching vibrations 21–4 codeine 163 coevolution 58, 67 COMPACT 138, 279–83, F6.42 complement of P450s 120, 122, 156 corticosterone 116 coumarin(s) 58, 125, 145, 150, 198 coupling of spin and redox equilibria 97–100 CPO (chloroperoxidase) 35, 37 crystallography 3, 9 see also X-ray crystallography crystal structures 11, 12, 21, 29 see also bacterial P450s c-terminus 7, 64 cyclcophosphamide 152, 154 cyclosporin 164 cysteine 3, 9, 11–14, 20, 32–3, 50, 64, 85, 107, 191, 203 conjugation 161, 184 cytochrome b5 11, 30, 50, 80, 87, 104–6, 170 cytochrome P420, 1, 12, 19, 34 cytochromes P450 (CYP) 4, 65, 68, 116–18 CYP1 58, 66, 115, 120, 135–43, 170, 171, 177, 182, 184 CYP1A1 58, 69, 120, 129, 171 CYP1A2 58, 69, 115, 120, 129, 171 CYP2 58, 70, 120, 143–62 CYP2A 58, 70, 115, 120, 125, 126, 130, 144–51, 172 CYP2B 70, 118, 120, 123, 128, 151–4, 169, 178, 183, 184 CYP2C 70, 120, 123, 127, 129, 154–6, 178 CYP2D 58, 66, 70, 71, 115, 120, 127, 129, 156–9, 171, 178, 194 CYP2E 58, 70, 115, 120, 128, 159–62, 170, 179, 186 CYP2F 70, 162 CYP2J 70, 71, 162 CYP2K 70, 74, 162 CYP3 71, 115, 120, 126, 163–4, 179, 183 CYP3A 12, 71, 123, 126, 169 CYP3A4 43, 71, 164 CYP3A7 71, 127 CYP4 66, 71, 74, 116, 180, 183 CYP5 72, 116–18

INDEX

CYP6 72, 116–18 CYP7 72, 116–18 CYP10 72 CYP11 72, 116–18, 181 CYP11A 72, 74, 107, 171 CYP11B 72, 74, 116 CYP17 72, 116–18, 171, 181, 206 CYP19 72, 116–18, 171, 181, 206 CYP21 72, 116–18, 181 CYP24 72, 116–18 CYP27 72, 74, 116–18 CYP51 73, 116–18, 197, 206 CYP101–112 74, 116–18, 181, 183 cytokines 177 d orbital(s) 9 splitting 9, 12, 31 DDT (p, p-dichlorodiphenyltrichlorethane) 172 debrisoquine 156, 159 DEN (diethylnitrosamine) 147, 151 desmethylimipramine 18 desolvation 94–6 detergent 1, 21 deuterium isotope effects 45, 92, 113, 153 devonian 58 dexamethasone 126, 163, 172 DHEA (dehydroepiandrosterone) sulphate 163 diabetic state 128 diallyl sulphide 162 2, 4-dichlorophenoxy N-alkyl N-methylethylamines 152, 199 diethyl ether 159 dimethyl benzanthracene 138 diolepoxides 138 dioxygen 2, 21, 22, 36, 44, 85, 88, 110 see also oxygen discovery of P450 1 distribution of P450s 3, 4, 5, 6 disulphiram 162 DMN (dimethylnitrosamine) 147, 159 dopamine 128 EGF (epidermal growth factor) 139, 186 electron transfer, transport 2, 64, 80, 84 electronic spectroscopy 1, 6, 11–14 see also UV spectroscopy ENACT 184 endogeneous metabolism, substrates 115–30

319

regulation 172, 175 steroid metabolism via CYP1-3 120–30 endoplasmic reticulum 1, 2, 80 ENDOR (electron-nuclear double resonance) spectroscopy 30 endrin 152 entropy change(s) 95 epoxides 138, 162, 186 epoxide hydrolase (EH) 7, 162, 184, 186 ergosterol 206 erythromycin 163 ESR (electron spin resonance) spectroscopy 3, 11, 31–4, 37, 107 estradiol 116, 129, 171 estrogen(s) 116, 127, 171 estrogen receptor 183 ethanol 128, 159, 172 ethers 198 7-ethoxy coumarin 4 ethyl morphine 164 ethynyl estradiol 164, 171, 201 1-ethynyl pyrene 140 eubacteria 61, 63 eukaryotes 56, 60–6 eukaryotic P450s 56–66, 80 evolution 55–67, 75, 76 EXAFS (extended X-ray absorption fine structure) spectroscopy 36–8 exogenous metabolism, substrates 130–67 expression systems 189 heterologous 173, 188–90 FAD (flavin adenine dinucleotide) 40, 61, 80, 84, 104 fatty acids 67 Fenton reagent 89 feprazone 152 Fermi splitting 13 ferredoxin 64, 80 ferric, ferrous 9–11, 32–3, 43–4 see also iron ferriheme, ferroheme 3, 9–14 flash photolysis 43 flavin 84 monooxygenase 7 flavocytochrome b2 109 flavone(s) 58 flavoprotein 2, 65 reductase 31, 80, 82–4 see also Figure 6.15, plate section

320

INDEX

FMN (flavin mononucleotide) 31, 61, 80, 84, 104, 109 FSH (follicle stimulating hormone) 171 Furafylline 140, 200 Furan 191 g tensor, values 31–3 genes, P450 3, 55, 64–74, 174–84 expression, regulation 169–176 structure 65, 174, 176, 177 genetic polymorphism 66–7, 120, 143, 156, 159, 170, 178 gestodene 201 globins 56, 64, 66 glucocorticoid(s), receptor 126, 143, 163, 180 gluconeogenesis 160 glutathione (GSH) 160, 162, 184 transferase 171

inhibition 18, 48–9, 190–208 inhibitors competitive 190–200 irreversible, mechanism-based 200–5 therapeutic and agrochemical uses 205 iodosobenzene 89, 91 ipomeanol 165 IR (infra-ed) spectroscopy 21–5 iron ionic radii 10, 11, T1.8 porphyrin complexes 9, 14, 196, 204, 280 isobestic point 14 isolation of P450s 4–7 isosafrole 71 itraconazole 195 Jahn-Teller distortion, effect 12, 31

half-lives of P450s 181 haloalkanes, haloalkenes 162 heme ligand, ligation 1, 9–14, 191 hemoglobin 11, 19, 21, 24, 32, 34, 38, 60, 85, 87 hemoproteins 1, 3, 9, 11–4, 20, 24, 25, 26, 31, 34, 39, 56, 66, 83, 86 heptachlor 153 high-spin state 9–11, 14, 16–9, 30, 32–5, 40, 42, 43, 85 histidine 11–12, 20, 30, 32 historical background 1–3 HOMO (highest occupied molecular orbital) 45 homology of P450 sequences 7, 55, 67, 71, 74, 212–42 Horseradish peroxidase (HRP) 34 HPLC (high-performance liquid chromatography) 7 Hückel MO claculations 14, 20 human P450s 123 hydroperoxide 90 hydrophobicity 18, 44, 47, 97, 108, 153 hydroxyl radical(s) 89 8-hydroxy guanosine 187 hypsochromic shift 1 I helix 23, 83 imidazole 12, 50, 159, 191 immunochemical homogeneity 7 inducers CYP1A 178, 281, 282 CYP2B 179 CYP2E 179 CYP3A 179 CYP4A 180 induction 66, 174–81, 184–8

ketoconazole 191 ketone bodies 128 lanosterol 197 lauric acid, laurate 30, 67, 116, 201 leucine 125 leukotrienes 116 LH (lutenizing hormone) 171 lidocaine 18 ligand field strength 12–13, 18 symmetries 9, 13, 34 octahedral 9–10, 35 rhombic 33, 35 square pyramidal 9–11 tetragonal 12, 31, 33 ligands -acceptor 11, 85, 191 -donor 11, 85 ligation 9–19 lignin peroxidase 50 lipid interactions 106–9 peroxidation 141, 186 lithocholic acid 120 log P (P=octanol/water partition coefficient) 47 low-spin state 3, 10–12, 14, 16–9, 31–5, 85 LUMO (lowest unoccupied molecular orbital) 46 -lyase 161, 184 lysine(s) 83, 85, 93, 152, 203

INDEX

magnetic resonance spectroscopy 28–34 see also ESR and NMR mammalian P450s 48–9, 56–9, 68 MC (3-methylcholanthrene) 18, 67, 129, 281 MCD (magnetic circular dichroism) spectroscopy 19–21, 37–8 MDBs (methylene dioxybenzenes) 198, 201 mechanism oxygenation 14, 44, 103–11 MEHP (mono-2-ethyl hexyl phthalate) 165 membrane binding 4, 64, 81, 106 menthofuran 165 mephenytoin 155 mercaptide 47 metabolism via P450s 48–9, 50, 58–67 metal ions as inhibitors 206–8 methionine 11 8-methoxy psoralen 165 metmyoglobin 33 metoprolol 159 metyrapone 152, 191 miconazole 195 microsomes, microsomal 1, 5–7, 67 P450s, 5, 20, 45–7, 63, 67 mitochondria 60–1, 67 mitochondrial system 2, 74, 85, 108 ML-236B 165 model complexes, systems 47, 89 molecular dynamics (MD) 43 molecular modelling of P450s 210, 211, 240–79 molecular orbital (MO) calculations 35, 36, 90–2 Hückel method 14, 20 CNDO/INDO method(s) 20, 90 Mössbauer (MB) spectroscopy 34–6 mRNA 67, 175 stabilization 170, 177, 184 mutagenicity, mutation 60, 138 myoglobin 11, 33, 85, T1.15 NAD(H), NADP(H) 2, 40, 61, 80, 84, 87, 104 NADPH menadione oxidoreductase 171 -naphthoflavone ( NF) 67, 129 -naphthylamine 138, 162 nicotine 154 nifedipine 163 -nitrophenol 153, 159, 161 1-nitropyrene 164 nitrosamines 147, 161

321

NMR (nuclear magnetic resonance) spectroscopy 28–30, 209, 212 NNK (4-methylnitrosamino-1-(3-pyridyl)-1butanone) 151 nomenclature of P450s 3, 4, 55, 67–77 NOS (nitric oxide synthase) 63–4, 94 N-terminus 7, 64, 106–8 omeprazole 140 optical spectroscopy 1, 9–21, 38 polarized 14, 19–21 ORD (optical rotatory dispersion) 20 orphenadrine 198, 199 oxene 50, 79, 87–92 oxygen see also dioxygen atmospheric 55–61, 66 binding affinity 11, 32, 43, 85 bond stretch 22–25, 28, 79, 88 complexes 12 electronic states 111 insertion mechanism 2 transfer, activation 17, 37, 83, 88 oxygenase 1 oxygenating species 88–92 P420 1, 12, 19–20, 34, 170, 180, 206, 233 P450LM2 (CYP2B4) 17, 20, 69, 84 LM4 20, 69 paracetamol 161 pathophysiological state 120, 129 PCBs (Polychlorobiphenyls) 141, 152 PCN (pregnenolone 16 -carbonitrile) 18, 126, 163, 172 PCR (polymerase chain reaction) 173 pentoxy resorufin 152, 154 peroxidase 21, 31, 87 peroxide 14, 22, 44, 79, 87, 104, 110 peroxisome proliferator(s) 180 activated receptor (ppar) 66, 180, 183 peroxy species 91–3, 110 phenacetin 161 phenobarbital 67, 126, 128, 143, 151, 154, 163, 172, 181, 183 phenol(s) 45, 153 phenylalanine 125 phenylbutazone 152 phenyldiazene 201, 204 phenylhydrazine 201 phenytoin 152, 154 phospholipid 64, 104–9, 170, 174 phosphorylation 170, 233

322

INDEX

site see Figure 6.5, plate section phylogenetic tree 56–60, 63, 66, 74, 75 physical properties of P450s 9–50 polyaromatic hydrocarbons (PAHs) 129, 135, 138 poor-metabolizer (PM) phenotype 129, 155, 157, 171 porphyrin ring, ligand 11–14, 35–6, 38, 47, 97 vibrational modes 21–2, 26 post-translational modification 170 potassium ions 43 pravastatin 165 pregnenoline 116 proadifen (SKF–525A) 152, 194, 198, 199 ProCheck 242 progesterone 116, 122, 171 prokaryotes 55, 66 propanediol 128 prostaglandins 116 proteins, P450 3 protein kinase A (PKA) 170, 178 C(PKC) 139, 184 protein stabilization 170 protoporphyrin IX 9, 30 PTH (parathyroid hormone) 171 pulegone 165 purification of P450s 4, 173 putidaredoxin 2, 30, 40, 43, 64, 84, 104 see also Figure 6.36, plate section pyridine 160, 193 QSARs (quantitative structure-activity relationships) 45– 7, 152, 161, 164, 210, 281, 282, 284–8 quenching of tryptophan fluorescence 21 quinidine 157, 163, 194 quinine 157, 194 quinoline(s) 58, 191 quinolones 141 recombinant DNA technology 172 redox components 2, 61–5, 84, 102–6 equilibria, states 12, 14, 17, 34, 39, 97, 98 interactions 100–6, 276 potentials 39–42, 45, 46, 64, 83–4, 86, 97–104 redoxin(s) 2, 61–4, 82, 104–6 reductase 28, 64, 84, 104–6, 172 reduction of P450, 84, 87, 100 rate(s) 102, 109 regulation

of P450s 66, 174 response elements 176, 181 resonance Raman (RR) spectroscopy 3, 21, 26–8, 38, 88 retinioc acid 206 rhombicity 31–3, T1.23 rifampicin 163, 172 ROS (reactive oxygen species) 66, 128, 154, 160, 184–7 rotational diffusion 107 safrole(s) 58 SDS-PAGE (sodium dodecyl sulphate-polyacrylamide gel electrophoresis) 7, 8, 69, 173 secobarbital 152, 201 senecionine 164 sequences of P450s, 3, 7, 55, 67, 71, 210, 214–224, 226, 232, 243–5, 250–1, 252–5, 259–62, 263–5, 268–9, 270– 1, 273–5, 278 serine 157 serotonin 128 sex differences mouse P450s 125 rat P450s 121, 122, 124 signal peptide 108 site-directed mutagenesis 50, 92, 93, 125, 127, 151, 155, 159, 226, 235 skatole 162 SKF-525A see proadifen SOD (superoxide dismutase) 160, 184, 186 Solomon-Bloembergen equation 28 Soret peak, absorption 1, 2, 14, 19, 26, 97, 191, 194 sparteine 159 spectral properties of P450s 9–38 spin-orbit coupling 33 spin-state equilibria 9–19, 26–8, 33–4, 38, 40, 42, 83–4, 94–8, 99–100, 108 effect of pH 42 effect of pressure 42 effect of temperature 42 spironolactone 163 steroidogenic P450s 2, 64, 67, 71, 116, 119 substrates 14–18, 48–9, 67 CYP1A 137 CYP2A 145 CYP2B 145 CYP2C 146, 147 CYP2D 148 CYP2E 149–50 CYP2F 145 CYP3A 158

INDEX

substrate binding 81–4 spectra 14–19, 82 sulfaphenazole 156 sulphur 33, 35–7, 61 superoxide, superoxy anion 22, 44, 79, 85–7, 91, 110 symmetry of heme environment 10–12, 26 TAO (troleandomycin) 163 TCDD (2,3,7,8-tetrachlorodibenzo-p-dioxin) 129, 138, 177, 182, 184 testosterone 116, 121, 122–8, 145–6, 150, 159 tetrachloromethane 201 thalidomide 154 therapeutic applications of inhibitors 205 thermodynamics of substrate binding 94–7, 110 thiolate ligand, ligation 3, 12, 20, 26, 30, 33, 35, 82 threonine 23, 43–4, 64, 83, 92, 157, 191 tienilic acid 155 tolbutamide 156 toluene(s) 45, 91, 153 toxicity and CYP1 induction 184, 187 transverse diffusion 109 traizole 191 trichloroethene 159 tris-(2,3-dibromopropyl) phosphate 164 tryptophan 18, 21, 64, 84 type I, II spectra 1, 14–19, 42, 198 substrates 15–16, 153 tyrosine 32, 191 UDP-glucuronosyl transferase 171 11-undecynoic acid 201 UV (ultra-violet) absorption spectroscopy 1, 6, 12–19, 86, F1.1 vibrational spectroscopy 21–8 frequencies 22, 24–27 modes 22 vitamin A 129 vitamin C 186 vitamin D3 118 vitamin E 186 warfarin 155 water 2, 12, 18–20, 29, 42–4, 56, 93–7, 113 xanthotoxin (8–methoxy psoralen) 165 X-ray crystallography 33, 38–9, 82

323

crystal structures of P450s 3, 9, 24, 29, 37, 39, 203, 210, 229, 230, 231 see also Figures 6.3 and 6.4, plate section P450cam see Figure 6.39, plate section P450BM3 see Figure 6.38, plate section P450terp see Figure 6.40, plate section

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