Transcriptional Regulation of the Human P450 2J2 by Activator -1

by

Nicole Yvonne Marden

A thesis submitted for the degree of Doctor of Philosophy

School of Medical Sciences Faculty of Medicine The University of New South Wales

January 2006 Originality Statement

Originality Statement

I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, or substantial proportions of material which have been accepted for the award of any other degree or diploma at UNSW or any other educational institution, except where due acknowledgement is made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledged in the thesis. I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project’s design and conception or in style, presentation and linguistic expression is acknowledged.

Nicole Yvonne Marden

ii Acknowledgements

Acknowledgements

Firstly, I would like to express my sincere thanks to my supervisor, Professor Michael Murray, for his constant guidance, support and encouragement throughout my PhD. In particular, I must thank Michael for the advice he has given me in terms of the design and analysis of my experimental studies, and for the generous amount of time he has devoted to reviewing this thesis. I would like to extend a special thank you to Dr Gloria Quee and Mrs Eva Fiala-Beer of the Murray Lab for all their help in teaching me experimental techniques, and for their continuous support and advice throughout my PhD. Your friendship and encouragement has been wonderful and has made the long hours in the lab a more pleasurable experience. I must also thank all of the other members of the Murray Lab, past and present, for their friendship and encouragement throughout my studies. I would like to thank Mr Stuart Purvis-Smith from the Molecular and Cytogenetics Unit at the Prince of Wales Hospital for generously providing me with access to a hypoxic incubator to undertake my hypoxic cell culture studies. I would also like to acknowledge Dr Kazuhiko Imakawa and Dr Michael Karin for generously providing various expression plasmids, and Dr Qing-Yu Zhang for generously providing the anti-(rat-CYP2J4) antibody. I would like to thank the members of staff at the Department of Physiology and Pharmacology for their help and interest in my project. A special thanks must go to Ms Rebekah Smith, Ms Sindy Kueh and Dr Kylie Mansfield for their friendship and support. To my fantastic friends Mandy and Beck: thank you for our weekly get-togethers over muffins and coffee. Your friendship has been priceless, and you have helped me to keep things in perspective and keep me smiling. I must also thank you both for your help with proof-reading this thesis. Finally, I would like to thank my wonderful family for their constant love, support and encouragement. Thank you to my Dad for giving me a love of science and learning in general, thank you to Lou for his amazing support and generosity, thank you to my Maxie and thank you to Mark and Nan for their love and encouragement. Most importantly, I would like to thank my wonderful mother, Marlena, and my amazing

iii Acknowledgements

partner, Christian, to whom I dedicate this thesis. You have taught me never to give up, and your unconditional love, patience and encouragement has allowed me to achieve my goals and dreams.

iv Table of Contents

Table of Contents

Originality Statement ii Acknowledgements iii Table of Contents v List of Figures xi List of Tables xiv Abbreviations xv List of Publications and Abstracts xx Abstract xxi

Chapter 1

Introduction 1 1.1 P450 1 1.1.1 Fundamental Aspects 1 1.1.2 Nomenclature of CYP 1 1.1.3 Biochemistry 2 1.2 of Xenobiotics by CYPs 4 1.2.1 CYP1 Family 6 1.2.2 CYP2 Family 7 1.2.3 CYP3 Family 11 1.2.4 CYP4 Family 12 1.3 Metabolism of Endogenous Substances by CYPs 12 1.3.1 CYPs Involved in Steroid Synthesis and Metabolism 14 1.3.2 CYPs Involved in Cholesterol Metabolism and Bile Acid Synthesis 15 1.3.3 CYPs Involved in Vitamin A and Vitamin D Metabolism 17 1.3.4 CYPs Involved in the Metabolism of 19 1.3.4.1 Arachidonic Acid and the Arachidonic Acid Metabolic Cascade 19 1.3.4.2 The Third Pathway in the AA Cascade: Metabolism of AA by CYPs 21

v Table of Contents

1.4 The Z/Z-1 Hydroxylase Pathway of AA Metabolism 22 1.4.1 Biological Activities of 20-HETE 23 1.4.2 Vascular Effects of 20-HETE 24 1.4.2.1 Vasoconstriction and the Regulation of Vascular Tone 24 1.4.2.2 Role for 20-HETE in Vasodilation Pathways 26 1.4.2.3 20-HETE as a Possible Oxygen Sensor 26 1.4.3 Effect of 20-HETE on Ion Transport within the Kidney 27 1.4.4 Mitogenic Actions of 20-HETE 27 1.4.5 Role of 20-HETE in the Pathogenesis of Hypertension 28 1.5 The Pathway of AA Metabolism 30 1.5.1 Biological Activities of EETs 32 1.5.2 Vascular Effects of EETs 32 1.5.2.1 Vasodilatory Effects of EETs within the Vascular System 32 1.5.2.2 EETs Proposed to be the Endothelium-derived Hyperpolarising Factor (EDHF) 34 1.5.2.3 Role of EETs in Reactive Hyperemia 35 1.5.3 Non-vasodilatory Effects of EETs within the Vascular System 36 1.5.3.1 Anti-Inflammatory Effects of EETs 36 1.5.3.2 Anti-migratory Effects of EETs 38 1.5.3.3 Fibrinolytic Effects of EETs 38 1.5.3.4 Mitogenic Properties of EETs 39 1.5.3.5 Effects of EETs on Platelets 40 1.5.3.6 Role of EETs in Protection Against Hypoxia-reoxygenation Injury in Endothelial Cells 40 1.5.4 Non-vascular Effects of EETs 41 1.5.4.1 Effects of EETs on Cardiomyocyte Function and Recovery After Cardiac Ischaemia 42 1.5.4.2 Effect of EETs within the Kidney and Potential Role in Hypertension 43 1.5.4.3 Anti-apoptotic Effects of EETs 45 1.5.4.4 Effects of EETs on the Release of Peptide Hormones 45 1.5.4.5 Effects of EETs in the Lung 46 1.5.4.6 Effects of EETs in the Liver 47 1.5.5 Factors Affecting the Level of EETs within the Body 48 1.6 2J2 (CYP2J2) 49 1.6.1 CYP2J2 Gene and Protein Structure 49 1.6.2 Catalytic Activity of CYP2J2 52

vi Table of Contents

1.6.3 Tissue Distribution of CYP2J2 53 1.7 Biological Significance of CYP2J2 54 1.7.1 Potential Role of CYP2J2 in the Heart and Vasculature 55 1.7.2 Potential Role of CYP2J2 in Other Tissues 59 1.8 Regulation of CYP Gene Expression 62 1.8.1 Regulation of CYP by Liver-enriched Transcription Factors 64 1.8.2 Receptor-mediated Regulation and Induction of CYP Gene Expression 65 1.8.2.1 CYP1A Induction by the Ah Receptor 66 1.8.2.2 Nuclear Receptors Involved in CYP Gene Expression 66 1.8.2.3 CAR-mediated Induction of CYP2B Genes 67 1.8.2.4 PXR-mediated Induction of CYP3A Genes 68 1.8.2.5 PPAR-mediated Induction of CYP4A Genes 69 1.8.2.6 LXR- and FXR-mediated Regulation of CYP7A Gene Expression 70 1.8.3 Down-regulation of CYPs by Inflammatory Mediators 70 1.8.4 Regulation of CYP2J2 Gene Expression 71 1.9 Activator Protein-1 73 1.9.1 AP-1 Components, Dimerisation and DNA Binding 73 1.9.2 Regulation of AP-1 Activity 77 1.9.3 Role of AP-1 in Cellular Physiology and Pathophysiology 79

Chapter 2

Materials and Methods 82 2.1 Materials 82 2.1.1 Reagents and Chemicals 82 2.1.2 Plasmids and Reagents for Molecular Biology 83 2.1.3 Reagents for Cell Culture 83 2.1.4 Reagents for Protein Electrophoresis and Immunoblotting 84 2.2 General Molecular Techniques 84 2.2.1 Preparation of Competent E.coli Cells and Transformation of Plasmids 84 2.2.2 Culture of E.coli Cells and Purification of Plasmids for Transfection 85 2.2.3 DNA Sequencing 87 2.2.4 Electrophoresis and Purification of DNA 88 2.3 Preparation of CYP2J2 Promoter Reporter Constructs 89

vii Table of Contents

2.4 Cell Culture 91 2.4.1 Experimental Conditions 91 2.4.2 Cell Line and Culture Conditions 92 2.4.3 Passaging of Cells 92 2.4.4 Hypoxic Treatment of Cells and Harvesting of Cells for Extraction of Total RNA, Total Cell Lysates and Nuclear Extracts 92 2.4.5 Assessment of Cell Viability 93 2.5 RNA Extraction 94 2.5.1 Experimental Conditions 94 2.5.2 RNA Extraction Procedure 94 2.5.3 Quantitation of RNA by Spectrophotometry 95 2.5.4 Electrophoresis of RNA Samples 95 2.6 Reverse Transcriptase Polymerase Chain Reaction (RT-PCR) 95 2.6.1 Experimental Conditions 95 2.6.2 Semi-quantitative RT-PCR of CYP2J2, c-Fos and c-Jun 96 2.6.3 Competitive RT-PCR for Quantification of CYP2J2 mRNA 98 2.6.3.1 Preparation of a Recombinant CYP2J2 RNA Internal Standard 98 2.6.3.2 Quantitative Competitive RT-PCR for CYP2J2 100 2.7 Protein Analysis 101 2.7.1 Isolation of Total Cell Lysates for Protein Analysis 101 2.7.2 Immunoblotting 102 2.8 Transient Transfection Analysis 103 2.8.1 Transient Transfection of HepG2 Cells 103 2.8.2 Luciferase Reporter Gene Assay 103 2.8.3 E-galactosidase Assay 104 2.9 Electrophoretic Mobility Shift Assay (EMSA) 105 2.9.1 Preparation of Nuclear Extracts 105 2.9.2 Preparation of Double-stranded Probes for use in EMSA 106 2.9.3 EMSA 109

Chapter 3 Regulation of the Expression of CYP2J2, and the AP-1 c-Fos and c-Jun, in Hypoxia and Reoxygenation 110 3.1 Introduction 110 3.2 Viability of HepG2 Cells Following Exposure to Hypoxia 112

viii Table of Contents

3.3 Analysis of CYP2J2 mRNA Levels in HepG2 Cells Following Exposure to Hypoxia and Reoxygenation 114 3.4 Identification of Multiple Potential Binding Sites for the Hypoxia-responsive Transcription Factor AP-1 within the 5’-flanking Region of the CYP2J2 Gene 118 3.5 Analysis of c-Fos and c-Jun Expression in HepG2 Cells During Hypoxia and Reoxygenation 120 3.6 Discussion 122

Chapter 4 Characterisation of an AP-1 Binding Sequence within the CYP2J2 Proximal Promoter that Regulates CYP2J2 Expression in Normoxia and Hypoxia 125 4.1 Introduction 125 4.2 Differential Activation of the CYP2J2 Promoter in HepG2 Cells by AP-1 Proteins 127 4.3 Location of a c-Jun-responsive Region within the Proximal Region of the CYP2J2 5’-flank 128 4.4 Identification of a Functional c-Jun at -56/-63 bp within the CYP2J2 Proximal Promoter 130 4.4.1 Binding of c-Jun to the CYP2J2 Proximal Promoter 130 4.4.2 Analysis of the Binding of Nuclear Protein and Recombinant c-Jun Protein to Sequences Resembling the AP-1 Consensus within the 152 bp c-Jun-responsive Region of the CYP2J2 Promoter 132 4.4.3 The AP-1-like Element at -56 to -63 bp within the CYP2J2 Proximal Promoter is Important in Mediating Transactivation by c-Jun 138 4.5 Effect of Hypoxia on the Binding of c-Jun to the AP-1-like Element at -56 to -63 bp in the CYP2J2 Proximal Promoter 141 4.6 Discussion 144

Chapter 5 Identification and Characterisation of a Second c-Jun Binding Element within the 5’-upstream Region of CYP2J2 that Regulates Expression 148 5.1 Introduction 148

ix Table of Contents

5.2 Identification of c-Jun-responsive Regions Between -152 and -50 bp in the CYP2J2 Proximal Promoter 149 5.3 Identification of c-Jun Binding Sequences within the -152 to -50 bp Region of CYP2J2 151 5.4 The -105 to -95 bp Sequence in CYP2J2 Binds c-Jun and Contributes to Transactivation of CYP2J2 158 5.5 Characterisation of AP-1 Binding to the -105 to -95 bp c-Jun-response Element in the CYP2J2 Promoter 162 5.5.1 Binding of c-Jun to the -105 to -95 bp Element in the CYP2J2 Promoter 162 5.5.2 Effect of Hypoxia on Binding to the -105 to -95 bp c-Jun-response Element 164 5.5.3 Binding of Recombinant c-Jun and c-Fos Proteins to the -105 to -95 bp Element in the CYP2J2 Promoter 165 5.6 The -152 to -122 bp Region of the CYP2J2 Gene is not Involved in c-Jun-dependent Regulation of CYP2J2 Promoter Activity 168 5.7 Discussion 174

Chapter 6

General Discussion 178

References 189

x List of Figures

List of Figures

Figure 1.1 General CYP catalytic reaction cycle 3 Figure 1.2 Steroidogenesis pathway 15 Figure 1.3 Pathways for the metabolism of arachidonic acid 20 Figure 1.4 Role of 20-HETE in the regulation of vascular tone 25 Figure 1.5 Metabolism of arachidonic acid by CYP generates four EET regioisomers; 5,6-EET, 8,9-EET, 11,12-EET and 14,15-EET 31 Figure 1.6 Opposing effects of EETs and 20-HETE on vascular tone 34 Figure 1.7 Anti-inflammatory effects of EETs via inhibition of cytokine-induced NF-NB activation 37 Figure 1.8 Structure of the CYP2J2 gene 50 Figure 1.9 Proposed biological roles of CYP2J2 in the heart and cardiovasculature 56 Figure 1.10 Binding of AP-1 to its DNA consensus sequence 74 Figure 1.11 Regulation of c-fos and c-jun gene transcription by MAP kinases 79

Figure 3.1 Viability of HepG2 cells following exposure to hypoxia for 16 or 40 hr 113 Figure 3.2 Semi-quantitative RT-PCR of CYP2J2 mRNA in HepG2 cells exposed to hypoxia and hypoxia-reoxygenation 114 Figure 3.3 Quantitation of CYP2J2 mRNA in HepG2 cells cultured in hypoxia for 16 hr by competitive RT-PCR 115 Figure 3.4 Quantitation of CYP2J2 mRNA in HepG2 cells exposed to hypoxia for 16 hr followed by reoxygenation for 30 min by competitive RT-PCR 116 Figure 3.5 Quantitation of CYP2J2 mRNA in HepG2 cells cultured in hypoxia for 40 hr by competitive RT-PCR 117 Figure 3.6 Effect of hypoxia and hypoxia-reoxygenation on c-Fos and c-Jun mRNA expression in HepG2 cells 121

xi List of Figures

Figure 3.7 Effect of hypoxia and hypoxia-reoxygenation on CYP2J2, c-Fos and c-Jun protein levels in HepG2 cells 122

Figure 4.1 Nucleotide sequence of the 5’-flanking region of the human CYP2J2 gene 126 Figure 4.2 Differential effects of AP-1 proteins on the CYP2J2 promoter 128 Figure 4.3 Location of a c-Jun-responsive region in the CYP2J2 gene 129 Figure 4.4 EMSAs showing the binding of nuclear protein and recombinant c-Jun protein to the CYP2J2 proximal promoter 131 Figure 4.5 AP-1-like elements within the c-Jun-responsive region of the CYP2J2 promoter 133 Figure 4.6 EMSAs evaluating the binding of c-Jun to the AP-1-like element at -7/+1 in the CYP2J2 promoter 135 Figure 4.7 EMSAs showing the binding of c-Jun to the AP-1-like element at -56/-63 in the CYP2J2 promoter 136 Figure 4.8 Binding of HepG2 nuclear extract to the AP-1 consensus binding motif 138 Figure 4.9 Role of the -7/+1 and -56/-63 bp AP-1-like elements in mediating c-Jun-dependent activation of the CYP2J2 promoter 139 Figure 4.10 Effect of mutating the -7/+1 and -56/-63 bp AP-1-like elements on the basal transcriptional activity of the CYP2J2 promoter 141 Figure 4.11 Differential effect of hypoxia on the binding of c-Jun to the -56/-63 bp AP-1-like element within the CYP2J2 promoter, and the AP-1 consensus binding sequence 143

Figure 5.1 c-Jun-responsive regions between -152 and -50 bp in the CYP2J2 proximal promoter 149 Figure 5.2 Effect of deletions within -152 to -50 bp on basal CYP2J2 promoter activity in HepG2 cells 150 Figure 5.3 Probes and competitors used in EMSAs to identify c-Jun binding sequences within the -152 to -50 bp region of the CYP2J2 proximal promoter 152 Figure 5.4 EMSA analysis of the binding of HepG2 nuclear extract and recombinant c-Jun protein to the -152 to -103 bp CYP2J2 promoter region 155

xii List of Figures

Figure 5.5 EMSA analysis of the binding of HepG2 nuclear extract and recombinant c-Jun protein to the -127 to -79 bp CYP2J2 promoter region 156 Figure 5.6 EMSA analysis of the binding of HepG2 nuclear extract and recombinant c-Jun protein to the -102 to -50 bp CYP2J2 promoter region 157 Figure 5.7 EMSAs reveal the -105 to -95 bp sequence of CYP2J2 is essential for c-Jun binding 159 Figure 5.8 The -56/-63 and -105/-95 bp sequences mediate c-Jun-dependent activation of the CYP2J2 proximal promoter 160 Figure 5.9 The -56/-63 and -105/-95 bp sequences are important in maintaining basal activity of the CYP2J2 proximal promoter in HepG2 cells 161 Figure 5.10 EMSA of the binding of c-Jun to the -105/-95 bp sequence in the CYP2J2 promoter 163 Figure 5.11 EMSA evaluating the effect of hypoxia on the binding of protein to the -105/-95 bp c-Jun binding element 165 Figure 5.12 EMSAs evaluating the binding of c-Jun and c-Fos/c-Jun to the CYP2J2 -105/-95 bp element and the AP-1 consensus binding sequence 167 Figure 5.13 EMSAs evaluating the binding of nuclear proteins and recombinant c-Jun protein within the -152/-122 bp region of the CYP2J2 promoter 170 Figure 5.14 Mutation of the -152/-122 bp region does not affect c-Jun-dependent activation of the CYP2J2 promoter 171 Figure 5.15 Mutation of the -152/-122 bp region does not diminish basal transcriptional activity of the CYP2J2 promoter in HepG2 cells 172 Figure 5.16 Transient transfection analysis of the -152/-122 bp sequence and c-Jun-dependent activation of the CYP2J2 proximal promoter 173

Figure 6.1 Schematic representation of the transcriptional regulation of the CYP2J2 proximal promoter by c-Jun and c-Fos in normoxia and hypoxia 184

xiii List of Tables

List of Tables

Table 1.1 Major human xenobiotic-metabolising CYPs 5 Table 1.2 Human CYPs involved in endogenous metabolism 13 Table 1.3 Biological actions of EETs 33 Table 1.4 CYP2J subfamily enzymes: substrates and tissue distribution 51 Table 1.5 Regioselectivity and stereoselectivity of CYP2J2-mediated metabolism of arachidonic acid into EETs 52 Table 1.6 Possible dimeric combinations between AP-1 subunits, and between AP-1 proteins and other bZIP transcription factors 75

Table 2.1 Sequencing primers 88 Table 2.2 Sequences of oligonucleotides used in site-directed mutagenesis to create CYP2J2 promoter deletion and mutation constructs 91 Table 2.3 Sequences of primers for semi-quantitative and competitive RT-PCR 96 Table 2.4 Sequences of oligonucleotides used in site-directed mutagenesis to create a recombinant CYP2J2 RNA internal standard for competitive RT-PCR 100 Table 2.5 Sequences of double-stranded oligonucleotides used in EMSAs 108

Table 3.1 Location of potential AP-1 response elements within the upstream regulatory region of the CYP2J2 gene 119

xiv Abbreviations

Abbreviations

AA arachidonic acid ADP adenosine 5’-diphosphate Ah aryl hydrocarbon AhRE Ah-responsive element ANP atrial natriuretic peptide AP-1 activator protein-1 APS ammonium persulfate Arnt Ah receptor nuclear translocator protein ATF activating transcription factor ATP adenosine 5’-triphosphate BAEC bovine aortic endothelial cell BARE bile acid-responsive element bp BSA bovine serum albumin bZIP basic region leucine zipper Ca2+ calcium cAMP cyclic adenosine monophosphate CAR constitutive androstanereceptor cDNA complementary DNA C/EBP CCAAT/enhancer binding protein CHOP C/EBP homologous protein 10 Cl- chloride CNC cap ‘n’ collar COX CPF CYP7A1 promoter binding factor CRE cAMP responsive element CREB cAMP responsive element binding protein CYP cytochrome P450 CYP2J2 cytochrome P450 2J2 Da Dalton

xv Abbreviations

DAG diacylglycerol DBD DNA-binding domain DBP albumin D-site binding protein DHET dihydroxyeicosatrienoic acid DMEM Dulbecco’s modified Eagle’s medium DMSO dimethyl sulfoxide dNTP deoxynucleotide triphosphate DR Dahl salt-resistant DS Dahl salt-sensitive DTT dithiothreitol E.coli Escherichia Coli EDHF endothelium-derived hyperpolarising factor EDTA disodium ethylenediaminetetra-acetate EET EGF epidermal growth factor EMSA electrophoretic mobility shift assay eNOS endothelial ERK extracellular-regulated kinase FCS fetal calfserum Fra Fos-related antigen FTF D-fetoprotein transcription factor FXR farnesoid X receptor GH growth hormone HEPES N-(2-hydroxyethyl)piperazine-N’-(2-ethanesulfonic acid) HETE hydroxyeicosatetraenoic acid HIF-1 hypoxia-inducible factor-1 HNF hepatocyte nuclear factor hPAR human pregnane activated receptor hPXR human pregnane X receptor HRE hormone response element hsp90 heat shock protein 90 ICAM-1 intercellular adhesion molecule 1 INB inhibitory factor NB IKK inhibitory factor NB kinase IL-1D interleukin-1D IL-1E interleukin-1E

xvi Abbreviations

IL-2 interleukin-2 IL-6 interleukin-6

IP3 inositol 1,4,5-trisphosphate IPTG isopropyl-E-D-thiogalactopyranoside JAK2-STAT5b Janus kinase 2 – signal transducer and activator of transcription 5b JNK c-Jun N-terminal kinase JT c-Jun-transfected K+ potassium kb kilo bases

KCa large-conductance calcium-activated potassium channel LB Luria-Bertani LBD ligand-binding domain LETF liver-enriched transcription factor LPS lipopolysaccharide LRH-1 liver receptor homologue-1 LXR liver X receptor LXRE LXR response element Maf macrophage-activating factor MAP kinase mitogen-activated protein kinase Meq Marek’s disease virus EcoQ protein Mg2+ magnesium MOPS 3-(N-morpholino)propane sulfonic acid mRNA messenger RNA MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide Na+ sodium NADPH nicotinamide adenine dinucleotide phosphate NFAT nuclear factor of activated T cells NF-E2 nuclear factor-erythroid 2 NF-NB nuclear factor NB NO nitric oxide Nrf NF-E2-related factor Nrl neural retina leucine zipper NS not significant nt nucleotides OD optical density ONPG o-nitrophenyl-E-D-galactopyranoside

xvii Abbreviations

p21SNFT 21-kDa small nuclear factor isolated from T cells PAH polycyclic aromatic hydrocarbon PAI-1 plasminogen activator inhibitor-1 PB phenobarbital PBREM PB-responsive enhancer module PBS phosphate buffered saline PCN pregnenolone 16D-carbonitrile PCR polymerase chainreaction PDGF platelet-derived growth factor

PGE2

PGF2D prostaglandin F2D

PGI2 prostacyclin PKA protein kinase A

PLA2 cytosolic phospholipase A2 PLC phospholipase C PMSF phenylmethanesulfonyl fluoride poly(dI-dC) poly(2’-deoxyinosinic-2’-deoxycytidylic acid) PPARD peroxisome proliferator-activated receptor D PPRE peroxisome proliferator response element PXR pregnane X receptor RAR retinoic acid receptor RT-PCR reverse transcriptase polymerase chain reaction RXR retinoid X receptor SDS sodium dodecyl sulfate SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis S.E.M. standard error SHP-1 small heterodimer partner 1 SHR spontaneously hypertensive rat Sp1 specificity protein 1 SRE serum-response element SRF serum-response factor STAT5 signal transducer and activator of transcription 5 SXR steroid and xenobiotic receptor TALH thick ascending loop of Henle TCF ternary complex factor TE Tris-EDTA buffer TEMED N,N,N’,N’-tetramethylethylenediamine

xviii Abbreviations

TGF-E transforming growth factor-E TNF-D tumour necrosis factor-D TPA 12-O-tetradecanoyl-phorbol-13-acetate t-PA tissue-type plasminogenactivator TRE TPA response element U units Ub ubiquitin UT untransfected VCAM-1 vascular cell adhesion molecule 1 VDR vitamin D receptor VSM vascular smooth muscle WKY Wistar-Kyoto XRE xenobiotic-responsive element

xix List of Publications and Abstracts

List of Publications and Abstracts

Publications

Marden, N. Y., Fiala-Beer, E., Xiang, S.-H., & Murray, M. (2003) Role of activator protein-1 in the down-regulation of the human CYP2J2 gene in hypoxia. Biochemical Journal 373, 669-680.

Marden, N. Y. & Murray, M. (2005) Characterization of a c-Jun-responsive module in the 5’-flank of the human CYP2J2 gene that regulates transactivation. Biochemical Journal 391, 631-640.

Abstracts

Lee, A. C., Marden, N. Y., & Murray, M. (2005) Potential transcriptional control elements in the 5’-upstream region of the human CYP2J2 gene. Australasian Society of Clinical and Experimental Pharmacologists and Toxicologists joint Annual Scientific Meeting with Australasian Pharmaceutical Science Association, Melbourne, Australia, 2005.

Marden, N. Y. & Murray, M. (2004) Characterisation of a c-Jun-responsive module in the 5’-flank of the human CYP2J2 gene that regulates expression. Australian Health and Medical Research Congress, Sydney, Australia, 2004.

Marden, N. Y., Xiang, S.-H., & Murray, M. (2002) Role of AP-1 in the regulation of human cytochrome P450 2J2 in hypoxia. Australian Health and Medical Research Congress, Melbourne, Australia, 2002.

Murray, M., Marden, N. Y., Anggono, V., Xiang, S.-H., & Fiala-Beer, E. (2002) Transcriptional regulation of endobiotic oxidising cytochromes P450. Australian Health and Medical Research Congress, Melbourne, Australia, 2002.

Marden, N. Y., Xiang, S.-H., & Murray, M. (2001) Regulation of human cytochrome P450 2J2 by members of the AP-1 transcription factor family. Australian Society for Medical Research 40th National Scientific Conference, Gold Coast, Australia, 2001.

xx Abstract

Abstract

The cytochrome P450 (CYP) superfamily of enzymes catalyses the oxidative metabolism of lipophilic xenobiotics such as drugs and environmental chemicals, and also plays an essential role in the biosynthesis and metabolism of endogenous compounds such as cholesterol and bile acids, vitamins, steroids, arachidonic acid and eicosanoids. Cytochrome P450 2J2 (CYP2J2) is a recently identified member of the human CYP that is highly expressed in the heart, vasculature, liver and other tissues. CYP2J2 metabolises arachidonic acid (AA) into epoxyeicosatrienoic acids (EETs), which have a number of potent biological activities including cytoprotective, vasodilatory and anti-inflammatory effects. Given its widespread tissue distribution and the biological actions of EETs, CYP2J2 is likely to play an important role in cellular physiology, and altered expression of CYP2J2 may have pathophysiological consequences. Indeed, recent literature studies have indicated that CYP2J2 protein levels are decreased in vascular endothelial cells exposed to hypoxia and reoxygenation, and that maintenance of CYP2J2 expression enhances cell survival. Thus, CYP2J2 expression may be impaired in diseases or conditions associated with decreased oxygen availability, such as ischaemic heart disease, stroke and atherosclerosis, and this may contribute to their pathogenic consequences. Despite its likely importance in human physiology and pathophysiology, very little is known about the regulation of CYP2J2 gene expression. The aim of this study was to investigate the molecular mechanisms that control expression of the CYP2J2 gene. In particular, this study was designed to identify factors that regulate the expression of the CYP2J2 gene in the liver-derived HepG2 cell line during normoxia and hypoxia. A 2.4 kb fragment of the 5’-flanking region of the CYP2J2 gene (corresponding to nucleotides -2341 to +98, relative to the translation start site) was isolated from a human genomic library. Automated searching of the upstream regulatory region of CYP2J2 identified several putative binding sites for the transcription factor activator protein-1 (AP-1). Because AP-1 activity is altered in hypoxia, the possibility that AP-1 may participate in the regulation of CYP2J2 expression in hypoxia was explored. Cell culture studies examined the relationship between the expression of CYP2J2, and the

xxi Abstract

AP-1 genes c-fos and c-jun, in HepG2 cells cultured in normoxia and hypoxia. Down- regulation of CYP2J2 mRNA and protein in hypoxic HepG2 cells was associated with the pronounced up-regulation of c-Fos protein from an undetectable level in normoxic cells; c-Jun protein levels were readily detectable in normoxia, and were also increased in hypoxia. Transient transfection studies revealed distinct effects of Fos and Jun proteins on CYP2J2 promoter activity. While the CYP2J2 promoter was strongly activated by c-Jun, c-Fos was inactive, and also abolished gene transactivation elicited by c-Jun. These results suggest that the constitutively expressed c-Jun is important in the maintenance of CYP2J2 expression in normoxic cells. The up-regulation of c-Fos in hypoxia stimulates the formation of c-Fos/c-Jun heterodimers, which do not support CYP2J2 transcription, leading to gene down-regulation. Experiments with CYP2J2 promoter deletion constructs revealed that the region between -152 to -50 bp relative to the translation start site was crucial for activation of CYP2J2 by c-Jun. Electrophoretic mobility shift assays (EMSAs) and transfection studies identified two distinct elements within this region that were involved in c-Jun-dependent transactivation: an AP-1-like element at -56 to -63 bp, and an atypical AP-1 element at -105 to -95 bp. c-Jun homodimers interacted specifically with both elements. Separate mutagenesis of either element significantly impaired c-Jun-dependent transactivation of CYP2J2, while mutagenesis of both elements eliminated c-Jun-responsiveness. EMSAs established that c-Jun, but not c-Fos, interacted with both elements in normoxic HepG2 cells. Furthermore, mutagenesis of either c-Jun-response element significantly decreased the basal transcriptional activity of the CYP2J2 promoter in HepG2 cells, while mutagenesis of both elements almost completely suppressed basal promoter activity. These findings indicate a pivotal role for c-Jun in the maintenance of CYP2J2 expression in normoxic cells. Transfection studies indicated that c-Fos suppresses c-Jun-dependent activation of CYP2J2 at both the -56/-63 bp and -105/-95 bp c-Jun-response elements. However, c-Fos-dependent inhibition appears to be mediated by distinct mechanisms at these two regulatory elements. While both elements interacted with c-Jun homodimers, only the -105/-95 bp element was able to interact with c-Fos/c-Jun heterodimers. Thus, the up-regulation of c-Fos in hypoxia, and the shift from c-Jun homodimers to c-Fos/c-Jun heterodimers, directly decreased c-Jun binding and transactivation at the -56/-63 bp element. In contrast, up-regulation of c-Fos in hypoxia altered the composition of proteins bound at the -105/-95 bp element from c-Jun to c-Fos/c-Jun. Inhibition of promoter activity occurs because c-Fos/c-Jun heterodimers can occupy, but not transactivate, the CYP2J2 promoter via the -105/-95 bp element.

xxii Abstract

In summary, this thesis provides novel information on the molecular mechanisms that control the differential expression of the human CYP2J2 gene in normoxia and hypoxia. In particular, this study has established that the AP-1 proteins c-Jun and c-Fos play a crucial role in modulating the transcriptional activation of the CYP2J2 promoter in response to cellular stress. Binding of c-Jun to two distinct c-Jun-response elements within the CYP2J2 proximal promoter induces transcriptional activation of the CYP2J2 gene and is essential for maintenance of CYP2J2 expression in normoxic cells. The up-regulation of c-Fos in hypoxia promotes the formation of c-Fos/c-Jun heterodimers, which inhibit transcriptional activation of the CYP2J2 promoter by c-Jun, thus contributing to decreased CYP2J2 expression in hypoxia. Impaired expression of CYP2J2 may contribute to cellular injury in diseases such as atherosclerosis and stroke, and a greater understanding of the mechanisms responsible for mediating altered CYP2J2 expression may eventually lead to therapeutic strategies that manipulate the expression of this important human gene.

xxiii Chapter 1

Chapter 1

Introduction

1.1 Cytochromes P450

1.1.1 Fundamental Aspects The cytochrome P450 (CYP) enzymes are a superfamily of proteins that catalyse the oxidative metabolism of a wide range of exogenous and endogenous substances including drugs, pesticides, alcohols, carcinogens, steroids and fatty acids (Coon et al., 1992). Named for their spectrophotometric absorption maximums at 450 nm when in a reduced and carbon monoxide-bound form (Omura & Sato, 1962), the cytochrome P450 proteins are haemoproteins consisting of a single polypeptide attached to a haem prosthetic group (Lewis, 1996). Cytochrome P450 enzymes have been found in almost all living organisms including fungi, bacteria, plants and animals, and are thought to have originated from a common ancestral gene more than 1.5 billion years ago (Nebert & Gonzalez, 1987). Their widespread existence across the various kingdoms of life highlights a critical role for these enzymes in processes essential to life.

1.1.2 Nomenclature of CYP Enzymes Early study within the field of P450 research was hindered by confusion relating to nomenclature, which arose from the lack of a universal classification system. As researchers around the world began purifying and characterising CYP enzymes, each laboratory essentially developed their own nomenclature (Gonzalez, 1988). Researchers generally assigned trivial names that were based on a variety of

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characteristics such as the source of CYP, the substrate of the , the spectrophotometric properties or even the electrophoretic mobility of the enzyme (Coon et al., 1992). As a result, a single CYP enzyme was often given a multitude of different names. For example, the enzyme called CYP1A1 under the current nomenclature was initially called P-450c, P-4482, P-450BNF-B, P-450 MC-1, P-450MC, P-450 isozyme 6,

P1450, P-450MC1 and PCB P-448-L (Gonzalez, 1988). It became evident that a universal system of nomenclature was needed, however, the standard methods of nomenclature for other enzyme systems, such as those based on the reaction catalysed, were found to be inappropriate for CYPs, because a single CYP enzyme often has a number of substrates, and may even catalyse different chemical reactions (Coon et al., 1992). In the late 1980s, a classification system based on sequence similarity was proposed and adopted (Nebert et al., 1987). Under this system, CYPs from all species, and the genes encoding them, are classified into families (designated by an Arabic number) and subfamilies (designated by a capital letter) on the basis of amino acid sequence identity. CYPs within the same family share a minimum of 40% sequence identity, while those within the same subfamily share greater than 55% sequence identity (Nelson et al., 1996). The individual CYP proteins within a subfamily are then assigned a sequential Arabic number. For example, the enzyme previously given a number of different names including P-450c, P-450BNF-B and P-4482, has now been assigned to family 1, and subfamily A, and is termed CYP1A1 or P4501A1. The gene encoding the enzyme is called CYP1A1. In some cases, a particular P450 enzyme may have a counterpart, or orthologue, in another species. For example, the CYP1A1 gene is ubiquitous in mammals, and is also found in a number of other organisms including fungi, insects, fish and birds (Gonzalez, 1988). On the other hand, some CYPs have been identified in only one species, for example CYP2J2, which is involved in arachidonic acid metabolism in humans (Wu et al., 1996). The CYP superfamily is one of the largest gene families in biology, with more than 480 CYP genes identified to date, giving rise to more than 270 different CYP gene families (Nebert & Russell, 2002). Within humans, 18 different CYP families, comprising of 41 subfamilies, have been identified (Nelson et al., 2004).

1.1.3 Biochemistry Mammalian CYPs are membrane bound enzymes located primarily in the endoplasmic reticulum of liver cells, but also in lower concentrations in mitochondrial, nuclear, and other membranes (Loeper et al., 1990; LaBella, 1991). Within the endoplasmic reticular membrane, CYPs are found in association with cytochrome b5 and the

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nicotinamide adenine dinucleotide phosphate (NADPH)-CYP reductase (Donato & Castell, 2003). The CYPs are the terminal oxidases in this electron transport chain and convert hydrophobic or lipophilic substances into more hydrophilic products via the insertion of an oxygen atom into the substrate molecule (Backes & Kelley, 2003). The general reaction mechanism (summarised in Figure 1.1) begins with substrate binding (1), followed by the reduction of the iron atom within the haem prosthetic group of the CYP enzyme (2). Once in this reduced state, the haem moiety is able to bind molecular oxygen, giving rise to a CYP-dioxygen complex (3). A second reduction reaction activates the bound diatomic oxygen (4), leading to cleavage of the oxygen-oxygen bond and the generation of a water molecule and a reactive iron-oxo species (5). The residual oxygen atom within this oxo complex is subsequently transferred to the bound lipophilic substrate molecule, thus creating a polar centre within the substrate (6). Following oxygen insertion, the product dissociates (7) (Groves & Han, 1995; Coon, 2003). The redox pathways that generate electrons for the two reduction reactions in the CYP reaction cycle vary depending on the CYP system. In mitochondrial and many bacterial systems, electrons are provided by NADH or NADPH via reductases containing FAD as the prosthetic group, whereas in microsomal endoplasmic reticular systems, electrons are generated from NAD(P)H via the FAD- and FMN-containing NADPH-CYP reductase (Porter & Coon, 1991; Lewis, 1996).

ROH RH P450-[Fe3+]

7 1

[ROH]-P450-[Fe3+] [RH]-P450-[Fe3+] e- 6 2

[RH]-P450-[Fe-O]3+ [RH]-P450-[Fe2+]

H2O 5 3 O2 2H+ 3+ - 2+ [RH]-P450-[Fe ]-O2 [RH]-P450-[Fe ]-O2

4

e- 3+ - [RH]-P450-[Fe ]-O2

Figure 1.1 General CYP catalytic reaction cycle Numbers 1 to 7 indicate the various stages in the general CYP catalytic cycle whereby a CYP substrate (RH) is converted into a more polar product (ROH). The redox state of the CYP haem iron (Fe) throughout the catalytic cycle is indicated.

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1.2 Metabolism of Xenobiotics by CYPs

The mammalian CYPs can be divided functionally into two major groups; those primarily involved in the metabolism of foreign substances or xenobiotics, and those primarily involved in the metabolism of endogenous substances (Gonzalez, 1992). CYPs play a critical role in the metabolism of a wide range of xenobiotics including drugs, plant products and environmental chemicals and pollutants (Nebert & Russell, 2002). Most xenobiotics are lipophilic and require chemical modification to make them more hydrophilic and more readily excreted from the body. The biotransformation of xenobiotics into more hydrophilic products often involves phase , and phase ,, metabolism (Donato & Castell, 2003). Phase , metabolism increases the water solubility of lipophilic xenobiotics by the introduction or exposure of a functional polar group within the substance, such as hydroxyl (Vermeulen, 1996). The phase , metabolite may be sufficiently hydrophilic to be directly excreted, or may undergo phase ,, metabolism. Here, the phase , product is metabolised by phase ,, enzymes, which catalyse the conjugation of an endogenous compound such as glucuronic acid, glutathione, and sulfate (Vermeulen, 1996). The final conjugate is extremely hydrophilic and readily excreted from the body via the bile or urine (Gonzalez, 1988). CYPs are the major enzymes involved in phase , metabolism (Vermeulen, 1996). Indeed, it is estimated that CYPs are involved in the metabolism of more than 200 000 chemicals, and greater than 90% of pharmaceuticals (Porter & Coon, 1991; Lewis, 1996). While metabolism of foreign substances by CYPs generally results in their successful detoxification and clearance, in some cases, metabolism by CYPs results in activation and the formation of highly reactive toxic metabolites that bind to DNA and other cellular macromolecules and may initiate cancer (Gonzalez, 1988; Porter & Coon, 1991). Thus, CYPs are considered to be an important factor in the pathogenesis of chemically-mediated tissue injury (Nebert & Gonzalez, 1987; Nebert & Russell, 2002). Human CYPs responsible for the metabolism of pharmaceuticals and the activation of protoxicants are largely concentrated within the CYP families 1-3, and, to a lesser extent, the CYP4 family (Nebert & Russell, 2002). The major human xenobiotic-metabolising CYPs are listed in Table 1.1 and are discussed in sections 1.2.1 – 1.2.3. Xenobiotic-metabolising CYPs are largely expressed within the liver where they play a crucial role in hepatic metabolism, although lower levels have been found in extrahepatic tissues including the lung, kidney, gastrointestinal tract and nasal epithelium (Guengerich, 1995). CYPs expressed within these extrahepatic tissues can

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contribute to overall clearance of drugs from the body (Chang & Kam, 1999). For example, renal CYPs may metabolise compounds transported from the liver and those generated within the kidneys, CYPs found in the lung and nasal tissues are involved in the metabolism of compounds that enter the body through the pulmonary system, and CYPs within the small intestine are important in the metabolism of orally delivered drugs (Guengerich, 1995).

Table 1.1 Major human xenobiotic-metabolising CYPs

Family Subfamily CYP Enzyme Selected Xenobiotic Substrates

CYP1 CYP1A CYP1A1 polycyclic aromatic hydrocarbons (e.g. benzo(a)pyrene)

caffeine, theophylline, polycyclic aromatic hydrocarbons, CYP1A2 heterocyclic amines, aflatoxin B1

CYP1B CYP1B1 polycyclic aromatic hydrocarbons (e.g. benzo(a)pyrene)

CYP2 CYP2A CYP2A6 coumarin, aflatoxin B1, 6-aminochrysene

CYP2A13 coumarin, dimethylaniline

CYP2B CYP2B6 cyclophosphamide, ifosphamide, 6-aminochrysene

CYP2C CYP2C8 taxol, benzo(a)pyrene, benzphetamine

CYP2C9 warfarin, tolbutamide, phenytoin

CYP2C19 S-mephenytoin, omeprazole, diazepam

debrisoquine, sparteine, imipramine, codeine, CYP2D CYP2D6 propranolol

CYP2E CYP2E1 ethanol, benzene, chloroform, carbon tetrachloride

cyclosporin A, erythromycin, nifedipine, midazolam, CYP3 CYP3A CYP3A4 lidocaine

CYP3A5 cyclosporin A, nifedipine

CYP3A7 midazolam, erythromycin

Table is not exhaustive, but lists the well-characterised xenobiotic-metabolising CYPs and some of their substrates.

While some CYPs, such as those involved in steroid metabolism (refer to section 1.3.1) have very distinct substrate specificities, other CYPs, in particular some of the xenobiotic-metabolising CYPs, show broad substrate specificities. Two examples that are noteworthy for the striking diversity in the substrates that they can accommodate are CYP3A4 and CYP2E1 (Guengerich, 1992). CYP3A4 can metabolise a large range of structurally diverse substrates of varying size, including the large antibiotics cyclosporin and erythromycin (Guengerich, 1999), while CYP2E1

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metabolises a range of small molecular weight substrates (Guengerich et al., 1991). Furthermore, some CYPs display overlapping substrate specificities; as such, the one compound may be metabolised by more than one CYP (Smith et al., 1998). In fact, there are only a few examples of compounds that are metabolised exclusively by only one CYP (Donato & Castell, 2003). An interesting and important feature of a number of the xenobiotic-metabolising CYPs is that their expression, which is generally low under normal conditions, is induced by exposure to certain xenobiotics (Dogra et al., 1998). Many inducers are actually substrates for the particular CYP that they induce. In this way, certain xenobiotics are able to enhance their own metabolism, and thus a sophisticated and efficient biological system is revealed, in which enzyme activity is increased only when an enhanced metabolic capacity is required (Denison & Whitlock, 1995). The mechanism of CYP induction is the subject of intense scientific investigation and is discussed further in section 1.8.2. CYP induction has important implications for drug therapy, as it affects CYP-dependent drug metabolism, drug pharmacokinetics, and is the cause of many drug-drug interactions (Waxman, 1999). An important example is the antibiotic , which is both an inducer and substrate of human CYP3A. Ingestion of rifampicin induces the expression of CYP3A enzymes in the liver which facilitates its own metabolism and clearance from the body. However, because the CYP3A enzymes are responsible for the metabolism of many drugs, the increased hepatic content of CYP3A results in increased clearance of other CYP3A substrates. Furthermore, rifampicin also induces the expression of some of the CYP2C enzymes, and increases the clearance of CYP2C substrates. In this way, CYP induction influences the clinical outcome of drug therapy (Nebert & Russell, 2002). As mentioned previously, the human CYPs that metabolise therapeutic drugs and foreign chemicals are found almost exclusively within the CYP1 to CYP4 families (Nebert & Russell, 2002). The important features of these CYPs are discussed below.

1.2.1 CYP1 Family The CYP1 family is one of the most extensively studied CYP families, and contains two subfamilies; CYP1A and CYP1B. Within the CYP1A subfamily, there are two members, CYP1A1 and CYP1A2, both of which can be found in all classes of the animal kingdom (Lewis, 1996). In particular, CYP1A1 is the most highly conserved of the xenobiotic-metabolising enzymes, with its enzymic activities and induction characteristics conserved amongst mammals, chicken and fish (Gonzalez, 1992). Within a given mammalian species, the CYP1A1 and CYP1A2 enzymes are closely related; for example, human CYP1A1 and

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CYP1A2 share approximately 70% sequence identity (Guengerich, 1995). Because land animals possess both CYP1A enzymes, but several species of fish have only CYP1A1, it has been speculated that CYP1A2 may have diverged about 400 million years ago after the colonisation of land, perhaps as a result of exposure to plant toxins (Lewis, 1996). CYP1A1 differs from many of the CYPs involved in xenobiotic metabolism, as it is essentially an extrahepatic enzyme, with only very low levels detected in the human liver (Shimada et al., 1992; Schweikl et al., 1993). CYP1A1 is particularly efficient in the metabolism of polycyclic aromatic hydrocarbons (PAHs; Table 1.1), many of which are procarcinogenic and/or promutagenic after activation by CYP1A1 (Roberts-Thomson et al., 1993; Bauer et al., 1995). Furthermore, the expression of CYP1A1 is induced by PAHs, such as those found in cigarette smoke (Sesardic et al., 1990). The mechanism of induction of CYP1A1 expression has been extensively studied and is discussed in further detail in section 1.8.2.1. Increased expression of CYP1A1 has been detected in the lungs of cigarette smokers (Anttila et al., 1992), and it is thought that there may be a link between the inducibility of human CYP1A1 and susceptibility to lung cancer (Kiyohara et al., 1996). In contrast to CYP1A1, human CYP1A2 is essentially a hepatic enzyme, although low levels of expression have been demonstrated in some extrahepatic tissues such as the brain (Farin & Omiecinski, 1993) and umbilical venous endothelium (Farin et al., 1994). CYP1A2 metabolises a number of drugs including caffeine (Butler et al., 1989; Gu et al., 1992) and theophylline (Table 1.1; Gu et al., 1992). CYP1A2 is known for its role in the metabolic activation of numerous procarcinogens and promutagens such as aflatoxin B1 (Gallagher et al., 1994) and aryl amines and heterocyclic amines (Butler et al., 1989; McManus et al., 1990). The metabolism of the heterocyclic amines by CYP1A2 is of particular interest, as these substances are found in cigarette smoke and charbroiled meat (Sugimura, 1992), and there appears to be some epidemiological evidence for a link between colon cancer and elevated levels of CYP1A2 activity (Lang et al., 1994). The CYP1B subfamily contains a single member, CYP1B1, which is constitutively expressed at a low level in various tissues including the liver, lung, and kidney (Sutter et al., 1994), and metabolises PAHs (Table 1.1; Nebert & Russell, 2002).

1.2.2 CYP2 Family The CYP2 family is the largest of the mammalian CYP families (Nebert & Russell, 2002) and differs from most others in that it contains members that are important in the

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metabolism of both xenobiotics and endobiotics, including arachidonic acid and certain steroids (Waterman, 1996). The human CYP2A subfamily consists of three members; CYP2A6, CYP2A7 and CYP2A13 (Smith et al., 1998). Of these, CYP2A6 has been most extensively studied. CYP2A6 is expressed at highly variable levels within the liver, and generally accounts for only a very small proportion of total hepatic P450 (<1%; Yun et al., 1991). Its expression appears to be linked to that of CYP2B6. The two genes are found in close proximity to each other on 19, and as such their expression may be coordinately regulated (Miles et al., 1989). CYP2A6 is the main enzyme involved in coumarin 7-hydroxylation (Table 1.1; Miles et al., 1990; Yamano et al., 1990; Yun et al., 1991) and 7-ethoxycoumarin O-deethylation (Yun et al., 1991). CYP2A6 also catalyses the activation of the carcinogens aflatoxin B1 and 6-aminochrysene (Table 1.1; Yun et al., 1991), however its importance in contributing to these reactions could be minimal, due to its low level expression in most livers and the fact that these reactions may be carried out more efficiently by other CYPs (Crespi et al., 1991; Yun et al., 1991). CYP2A7 and CYP2A13 are 94% (Yamano et al., 1990) and 93% (Smith et al., 1998) related to CYP2A6. CYP2A7 does not metabolise coumarin (Yamano et al., 1990; Ding et al., 1995), and very little is known about its substrate specificities and patterns of expression. CYP2A13 metabolises coumarin, but with a lower activity than that of CYP2A6, and also catalyses several other reactions including N,N-dimethylaniline N-demethylation (Table 1.1; Guengerich, 2005). The CYP2B subfamily has been studied intensively with the inducibility of rat CYP2B enzymes by barbiturates (Guengerich, 1995; Waxman, 1999). The human CYP2B subfamily consists of a single member, CYP2B6, which has 76% amino acid sequence similarity to rat CYP2B1 (Yamano et al., 1989) and is also inducible by phenobarbital, although to a much lesser extent than CYP2B1 (Waxman, 1999). As with CYP2A6, hepatic expression of CYP2B6 varies significantly between individuals, and is thought to play only a minor role in metabolism of xenobiotics, as it constitutes less than 2% of total CYP expression in the liver (Mimura et al., 1993). Substrates for CYP2B6 include 6-aminochrysene (Mimura et al., 1993) and the anticancer drugs cyclophosphamide and ifosphamide (Table 1.1; Chang et al., 1993). CYP2C is the most complex of the human CYP subfamilies, consisting of 4 functional genes (CYP2C8, CYP2C9, CYP2C18 and CYP2C19), and numerous pseudogenes (Nelson et al., 2004). Together, the human CYP2C enzymes metabolise more than 50% of all frequently prescribed drugs, as well as several important

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endogenous compounds including arachidonic acid and certain steroids (Nebert & Russell, 2002). The human CYP2C enzymes are constitutively expressed within the liver, but show a significant variation in expression levels between individuals (Furuya et al., 1991). They have been found to constitute approximately 20% of total hepatic CYP content (Shimada et al., 1994). CYP2C8 metabolises a variety of substrates (see Table 1.1) including benzphetamine (Wrighton et al., 1987), benzo(a)pyrene (Yun et al., 1992), taxol (Rahman et al., 1994), arachidonic acid (Daikh et al., 1994), retinol and retinoic acid (Leo et al., 1989). CYP2C9 is the major CYP2C protein expressed in most human livers (Läpple et al., 2003), and catalyses the metabolism of arachidonic acid (Daikh et al., 1994), as well as a number of commonly prescribed therapeutics including warfarin (Rettie et al., 1992), tolbutamide and phenytoin (Table 1.1; Veronese et al., 1991). Allelic variants of CYP2C9 that affect metabolism have been identified in various populations (Lee et al., 2002), and are of clinical significance (Aithal et al., 1999; Brandolese et al., 2001). CYP2C18 appears to be a minor member of the CYP2C subfamily; this enzyme exhibits low level hepatic expression (Läpple et al., 2003) and no specific CYP2C18 substrates have been identified (Smith et al., 1998). CYP2C19 is the major enzyme responsible for the metabolism of S-mephenytoin (Table 1.1; Goldstein et al., 1994). It also metabolises a number of other frequently prescribed drugs including diazepam (Jung et al., 1997) and omeprazole (Andersson et al., 1992). A well-studied polymorphism exists for the metabolism of S-mephenytoin, with approximately 3% of Caucasians and nearly 20% of Asians classified as poor metabolisers of S-mephenytoin (Nakamura et al., 1985). A series of defective alleles of the CYP2C19 gene are responsible for the poor metaboliser phenotype (Goldstein & de Morais, 1994). CYP2D6 is the only member of the CYP2D subfamily that is expressed within humans; the two other genes within this subfamily, CYP2D7P and CYP2D8P, are non- functional pseudogenes (Smith et al., 1998). CYP2D6 accounts for less than 5% of total hepatic CYP content (Shimada et al., 1994) but is responsible for the metabolism of approximately 25% of all currently- used therapeutic drugs (Wolf & Smith, 1999). Substrates of CYP2D6 are structurally diverse, but are generally small molecules which contain a basic nitrogen atom that is positively charged at physiological pH (Smith et al., 1998). Drugs metabolised by CYP2D6 include the anti-hypertensive drug debrisoquine (Distlerath & Guengerich, 1984; Distlerath et al., 1985), tricyclic antidepressants including imipramine and

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desipramine (Brosen et al., 1986), analgesics such as codeine (Dayer et al., 1988), E-blockers such as propranolol (Distlerath et al., 1985) and metoprolol (Ellis et al., 1992) and anti-arrhythmics such as sparteine (Table 1.1; Distlerath & Guengerich, 1984; Distlerath et al., 1985). CYP2D6 displays a very well characterised polymorphism within human populations, with individuals classified as poor metabolisers or extensive metabolisers depending on their ability to metabolise CYP2D6 substrates such as debrisoquine (Meyer & Zanger, 1997). Approx 5-10% of Caucasian and 1-2% of Asian and African populations are classified as poor metabolisers (Nakamura et al., 1985; Alvan et al., 1990; Relling et al., 1991); these individuals display a severely compromised ability to metabolise CYP2D6 substrates due to lack of functional CYP2D6 protein that arises from a number of allelic variants in the CYP2D6 gene (Meyer & Zanger, 1997). Extensive metabolisers, on the other hand, produce functional CYP2D6 protein and thus are able to metabolise CYP2D6 substrates. Amongst the extensive metabolisers are the ultra-rapid metabolisers, who contain multiple copies of the functional CYP2D6 gene, presumably due to a duplication event (Johansson et al., 1993). Individuals with the ultra-rapid metaboliser phenotype produce more CYP2D6 protein, and require much higher doses of drugs metabolised by CYP2D6 in order to obtain therapeutic benefit (Smith et al., 1998). The human CYP2E subfamily contains a single gene, CYP2E1 (Gonzalez, 1992). CYP2E1 is toxicologically important because it mediates the metabolic activation of numerous low molecular weight toxins and carcinogens, such as benzene, chloroform and carbon tetrachloride (Table 1.1; Guengerich et al., 1991). It is also one of the pathways by which ethanol is metabolised within the body (Ekström et al., 1989), and oxidises ketone bodies that are produced by gluconeogenesis (Koop & Casazza, 1985). CYP2E1 is expressed in the liver, and has also been detected in brain, lung, skin and placenta (Farin & Omiecinski, 1993; Botto et al., 1994). Whilst hepatic expression is constitutive, CYP2E1 is inducible by many of its substrates, including ethanol (Wrighton et al., 1986; Perrot et al., 1989), and is also induced in physiological states that activate the gluconeogenic pathway such as diabetes and starvation (Dong et al., 1988; Johansson et al., 1990). The regulation of CYP2E1 expression in experimental animals is very complex: rat CYP2E1 is regulated by developmental and hormonal mechanisms, at transcriptional and post-transcriptional levels (Koop & Tierney, 1990). While less is known about the regulation of human CYP2E1, it is thought to involve similar mechanisms to its rodent counterpart (Perrot et al., 1989).

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There exists a large degree of inter-individual variation in hepatic expression of CYP2E1. This variation could be due in part to differential exposure to inducing agents, but may also be due to polymorphisms within the regulatory region of the CYP2E1 gene (Hayashi et al., 1991; Gonzalez, 1992). The human CYP2J subfamily contains one member, CYP2J2, whose regulation is the subject of investigation in this thesis. CYP2J2 is expressed within the liver, but also in a number of extrahepatic tissues including the heart and the lungs (Wu et al., 1996; Zeldin et al., 1996a). While CYP2J2 is primarily involved in metabolism of endogenous arachidonic acid, which will be discussed in detail in section 1.6.2, it is mentioned here as it has also been shown to metabolise certain xenobiotics including ebastine (Hashizume et al., 2002), astemizole (Matsumoto et al., 2002; Matsumoto et al., 2003), diclofenac and bufuralol (Scarborough et al., 1999). While little is known about the CYPs found within the more recently identified human CYP2F, 2R, 2S, 2U and 2W subfamilies, it is possible that they may also contribute to the metabolism of certain xenobiotics (Nebert & Russell, 2002).

1.2.3 CYP3 Family The human CYP3 family is abundantly expressed in liver and plays a critical role in metabolism of xenobiotics, contributing to the biotransformation of more than 40% of all drugs (Lewis, 1996). Erythromycin (Brian et al., 1990), cyclosporin A (Kronbach et al., 1988), nifedipine (Guengerich et al., 1986), midazolam (Kronbach et al., 1989) and lidocaine (Imaoka et al., 1990) are amongst the drugs that are substrates for CYP3 enzymes (Table 1.1). CYP3 members are also important in the metabolism of endogenous compounds such as oestradiol, progesterone, testosterone, cortisol and androstenedione (Waxman et al., 1988; Brian et al., 1990). The human CYP3 family contains three functional members, CYP3A4, CYP3A5 and CYP3A7 (Nelson et al., 2004). CYP3A4 is the most abundantly expressed CYP within the human liver (Guengerich, 1995). It is also expressed at a relatively high level within the gastrointestinal tract, in particular the small intestine, which may be of significance in the metabolism of orally administered drugs (de Waziers et al., 1990). Expression of CYP3A4 is induced by steroids, antifungal drugs, macrolide antibiotics and phenobarbital (Pichard et al., 1990; Kocarek et al., 1995). The mechanism by which this occurs has been the subject of intense investigation and will be presented in greater detail in section 1.8.2.4. CYP3A5 shares 84% amino acid homology to CYP3A4 (Smith et al., 1998). It is expressed in only 10 to 30% of human livers (Aoyama et al., 1989; Wrighton et al.,

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1990), and is present in a higher percentage in children and adolescents compared to adults (Wrighton et al., 1990). CYP3A5 substrates include nifedipine, cyclosporin A and various steroid hormones (Table 1.1; Aoyama et al., 1989; Wrighton et al., 1990). CYP3A7 is expressed in the fetal liver, where it is the major form of P450, constituting between 30 to 50% of total CYP content (Wrighton et al., 1988), and has also been shown to be expressed in the uterine endometrium (Schuetz et al., 1993). CYP3A7 displays similar catalytic activities to CYP3A4 and CYP3A5 (Table 1.1; Guengerich, 2005).

1.2.4 CYP4 Family The CYP4 family consists of 12 enzymes belonging to 6 different subfamilies (Nelson et al., 2004). While primarily known for their role in the metabolism of fatty acids such as arachidonic acid, which will be discussed in section 1.4, various members of the CYP4 family, such as CYP4A11, CYP4B1, CYP4F2 and CYP4F3, have been shown to metabolise certain drugs (Nebert & Russell, 2002).

1.3 Metabolism of Endogenous Substances by CYPs

While CYPs were originally studied for their role in the metabolism of drugs, research over the last few decades has shown that this enzyme system plays an essential role in the biosynthesis and degradation of numerous endogenous compounds. Steroids, bile acids, vitamin D, retinoids and fatty acids are amongst the physiologically important substrates for CYP enzymes (Coon et al., 1992; Nebert & Russell, 2002). In general, the CYPs involved in the metabolism of endogenous substrates belong to gene families that are distinct from the xenobiotic-metabolising CYPs (Waterman, 1996). The CYP2 gene family is a notable exception, containing CYPs that metabolise xenobiotics, as well as endobiotics like arachidonic acid (Waterman, 1996). CYPs that metabolise endogenous substrates display narrow substrate specificities, whereas the xenobiotic-metabolising CYPs generally accommodate a broader range of substrates (Guengerich, 1992). Furthermore, the catalytic activities of the steroid and cholesterol-metabolising CYPs are well conserved across mammalian species. This is thought to be due to the essential role these enzymes play in the synthesis of physiologically important compounds, such that defective genes with

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impaired enzyme activity could be lethal or deleterious to homeostasis (Gonzalez, 1992). On the other hand, many of the xenobiotic-metabolising CYPs show significant variation in their catalytic activities across mammalian species. Another point of difference between the xenobiotic-metabolising CYPs and the endobiotic-metabolising CYPs is that there tends to be less variation in the levels of the latter group of enzymes amongst different individuals. Again, this may be due to their critical role in the biotransformation of endobiotics (Guengerich, 1992). The human CYPs involved in endogenous metabolism are summarised in Table 1.2 and are discussed in further detail below.

Table 1.2 Human CYPs involved in endogenous metabolism

Endogenous Metabolic Pathway CYPs Involved

Steroid Synthesis and Metabolism CYP11A1, CYP11B1, CYP11B2, CYP17A1, CYP19A1, CYP21A2, CYP3A4, CYP3A5

Cholesterol Synthesis CYP51A1

Bile Acid Synthesis CYP7A1, CYP7B1, CYP8B1, CYP27A1, CYP39A1, CYP46A1, CYP3A4

Retinoic Acid Metabolism CYP26A1, CYP26B1

Vitamin D3 Synthesis and Metabolism CYP24A1, CYP27A1, CYP27B1

Arachidonic Acid Metabolism CYP2J2, CYP2C8, CYP2C9, CYP4A11, CYP4F2

Eicosanoid Synthesis and Metabolism CYP4F2, CYP4F3, CYP4F8, CYP5A1, CYP8A1 (e.g. leukotrienes, thromboxane, prostaglandins)

Adapted from Guengerich, 2005 [Tables 10.1 (pg 378-379) and 10.2 (pg 380)].

Briefly, the CYP families involved in steroidogenesis include CYP11A, CYP11B, CYP17, CYP19 and CYP21 (Kagawa & Waterman, 1995). CYP7A1, CYP7B1, CYP8B1, CYP27A1, CYP39A1 and CYP46A1 are involved in the synthesis of bile acids from cholesterol (Nebert & Russell, 2002). CYP5A1 is a thromboxane A

synthase, responsible for converting into thromboxane A2, which is involved in platelet aggregation (Yokoyama et al., 1991; Nebert & Russell, 2002).

CYP8A1, otherwise known as , converts prostaglandin H2 into prostacyclin (Miyata et al., 1994; Nebert & Russell, 2002). CYP24A1, CYP27A1 and

CYP27B1 are involved in vitamin D3 synthesis and metabolism (Wikvall, 2001; Nebert & Russell, 2002), and the CYP26 family is important in retinoic acid hydroxylation (White et al., 1997). Arachidonic acid is metabolised by a number of different CYPs belonging

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to various families, in particular the CYP2 and CYP4 families (Capdevila & Falck, 2002), and CYP51A1 is important in the synthesis of cholesterol (Strömstedt et al., 1996).

1.3.1 CYPs Involved in Steroid Synthesis and Metabolism There are six CYPs, collectively termed the steroid hydroxylases, that are involved in the process of steroidogenesis (the production of steroid hormones from cholesterol). CYPs 11A1, 11B1, 11B2, 17A1, 19A1 and 21A2 convert the precursor cholesterol into steroids with specific physiological functions including glucocorticoids, mineralocorticoids, progestins and sex hormones (Guengerich, 1995; Kagawa & Waterman, 1995). The metabolism of cholesterol into the steroid hormones occurs primarily in the adrenal cortex, the gonads and the placenta, and to a lesser extent within the brain and in adipose tissue (Kagawa & Waterman, 1995). Thus, the specialised steroid hydroxylases are expressed within several extrahepatic tissues (Keeney & Waterman, 1993). While the CYPs are responsible for the biosynthesis of the steroid hormones, their degradation is mediated by several hepatic xenobiotic- metabolising CYPs (Waxman et al., 1991). The general steroidogenic pathway is shown in Figure 1.2. All steroidogenic pathways begin with the conversion of cholesterol into pregnenolone. This reaction occurs within the mitochondrion of the cell, and is catalysed by the mitochondrial cholesterol side chain cleavage cytochrome P450 or CYP11A1 (Hanukoglu, 1992; Keeney & Waterman, 1993). Given the essential role of CYP11A1 in initiating the production of all steroid hormones from their common cholesterol precursor, deficiency in CYP11A1 activity can lead to hypertension, glucocorticoid insufficiency and feminisation (Keeney & Waterman, 1993). The glucocorticoid cortisol and the mineralocorticoid aldosterone are produced by CYP enzymes expressed within the adrenal cortex; CYP11A1, CYP17A1, CYP21A2 and CYP11B1 are expressed within the inner zones of the adrenal cortex and are essential components of the metabolic pathway that produces cortisol, while CYP11A1, CYP21A2, CYP11B1 and CYP11B2 are expressed in the outer zone of the adrenal cortex and are required for the synthesis of aldosterone (Figure 1.2; Keeney & Waterman, 1993; Kawaga & Waterman, 1995). The complement of CYPs within the gonadal steroidogenic tissues gives rise to the production of the sex hormones testosterone and oestrogen. CYP11A1 and CYP17A1 expressed within the steroidogenic cells of the testis (the Leydig cells)

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participate in the metabolic pathway that produces the male sex hormones testosterone and dihydrotestosterone (Figure 1.2; Keeney & Waterman, 1993; Kawaga & Waterman, 1995). CYP11A1 and CYP17A1 expressed within the thecal cells of the ovaries participate in the production of the androgens androstenedione and testosterone, which are subsequently converted into the oestrogens oestrone and oestradiol by CYP19A1 (otherwise known as P450) within the ovarian granulosa cells (Figure 1.2; Kawaga & Waterman, 1995; Guengerich, 1995).

Cholesterol

CYP11A1

Pregnenolone

3EHSD CYP17A1

CYP17A1 Progesterone 17-OH-pregnenolone Dehydroepiandrosterone

CYP21A2 CYP17A1 3EHSD 3EHSD

CYP17A1 Deoxycorticosterone 17-OH-progesterone Androstenedione

CYP11B1 CYP21A2 17EHSD CYP19A1

Corticosterone 11-deoxycortisol Testosterone oestriol

CYP11B2 CYP11B1 5DRED CYP19A1

18-OH-corticosterone Cortisol Dihydrotestosterone oestradiol

CYP11B2

Aldosterone

Figure 1.2 Steroidogenesis pathway 3EHSD=3E-hydroxysteroid dehydrogenase; 17EHSD=17E-hydroxysteroid dehydrogenase; 5DRED=5D-reductase.

1.3.2 CYPs Involved in Cholesterol Metabolism and Bile Acid Synthesis While cholesterol is an essential component of membranes, accumulation of excess cholesterol within the body can lead to cardiovascular disease, and the level of cholesterol within the body is therefore closely regulated (Russell, 1999). The liver is the principal organ of cholesterol metabolism, and cholesterol homeostasis of the entire body is primarily achieved via the coordinate regulation of hepatic cholesterol supply and catabolism pathways. Supply of cholesterol is mediated by two distinct pathways; the endogenous synthesis of cholesterol from acetate precursors, and the uptake of exogenous and endogenous cholesterol from the blood

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via low density lipoprotein receptors. Catabolism of cholesterol involves its conversion into hydrophilic bile acids (Russell & Setchell, 1992). CYP51A1, also known as lanosterol 14D-demethylase, plays an essential role in the biosynthesis of endogenous cholesterol. CYP51A1 catalyses the removal of two methyl groups from the sterol precursor lanosterol within the cholesterol biosynthetic pathway (Strömstedt et al., 1996). Genes encoding enzymes orthologous to human CYP51A1 have been identified in plants, animals and fungi, as well as in the primitive prokaryotic organism Mycobacterium tuberculosis, leading to the suggestion that this CYP may be the common ancestor to all eukaryotic CYPs (Yoshida et al., 2000). CYP enzymes from a number of distinct families including the CYP3, CYP7, CYP8, CYP27, CYP39 and CYP46 families are involved in the conversion of cholesterol into hydrophilic bile acids, which represents the major pathway for the catabolism and elimination of cholesterol from the body (Nebert & Russell, 2002). This cholesterol catabolic pathway consists of at least 14 different enzymes and results in the generation of two major primary bile acids, cholic acid and chenodeoxycholic acid (Russell & Setchell, 1992). There are two pathways for the synthesis of bile acids from cholesterol, both of which involve CYPs. The classical bile acid biosynthetic pathway produces the two primary bile acids, cholic acid and chenodeoxycholic acid, and involves reactions catalysed by CYP7A1 (cholesterol 7D-hydroxylase), CYP8B1 (sterol 12D-hydroxylase) and CYP27A1 (sterol 27-hydroxylase; Riddick et al., 2004). The liver-specific microsomal CYP7A1 catalyses the hydroxylation of the cholesterol molecule at the 7 position of the steroid nucleus, which is the first, and rate-limiting, step in the classic bile acid biosynthetic pathway (Jelinek et al., 1990). In addition to being the rate-limiting step, 7D-hydroxylation by CYP7A1 also serves as the major site of regulation in the bile acid biosynthetic pathway. Regulation of CYP7A1 activity is complex and is reportedly influenced by a number of factors, such as hormones and dietary components including cholesterol, as well as negative feedback by bile acids themselves (Taniguchi et al., 1994). The mechanisms by which cholesterol and bile acids regulate CYP7A1 gene expression are discussed in detail in section 1.8.2.6. The alternate (or acidic) bile acid biosynthetic pathway occurs predominantly within peripheral tissues and macrophages (Riddick et al., 2004) and begins with the oxidation of the steroid side chain of cholesterol to produce oxysterols containing a hydroxyl group at either the 24, 25 or 27 carbon position on the steroid side chain (Russell, 1999). 24S-hydroxycholesterol and 27-hydroxycholesterol are produced, respectively, by the CYP enzymes CYP46A1, a brain-specific cholesterol 24-hydroxylase, and CYP27A1, the widely expressed sterol 27-hydroxylase,

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highlighted above for its role in the classic pathway (Cali & Russell, 1991; Lund et al., 1999). In contrast, 25-hydroxycholesterol is formed from cholesterol by a non-CYP cholesterol 25-hydroxylase (Lund et al., 1998). The cytochrome P450 enzyme CYP7B1 (oxysterol 7D-hydroxylase) converts these oxysterols into 7D-hydroxylated oxysterols that are subsequently funneled into the classical pathway and converted into bile acids in the liver (Russell, 1999; Riddick et al., 2004). This alternate pathway is particularly important in the initiation of cholesterol metabolism in non-hepatic tissues. While many peripheral tissues achieve cholesterol homeostasis by transferring excess intracellular cholesterol to circulating lipoproteins, which are eventually metabolised within the liver, certain tissues, such as the brain, cannot use this pathway (excess cholesterol is unable to pass through the blood brain barrier; Lund et al., 1999). Via the alternate bile acid biosynthetic pathway, excess cholesterol in the brain is converted by CYP46A1 into 24S-hydroxycholesterol, which is considerably more polar than cholesterol, and is more able to pass through the blood brain barrier and be returned to the liver for processing (Björkhem et al., 1997).

1.3.3 CYPs Involved in Vitamin A and Vitamin D Metabolism Various CYP enzymes are important in the metabolism of vitamins A and D. Retinoic acids are derivatives of vitamin A that are key morphogens during the development of vertebrates (Maden, 2000). Retinoic acids exert their biological effects by controlling the transcription of a variety of target genes via interaction with two families of nuclear receptors that behave as ligand-dependent transcription factors, the retinoic acid receptors (RAR D, E and J) and the retinoid X receptors (RXR D, E and J; Chambon, 1996). Retinoic acids are produced from vitamin A by a series of sequential reactions that involve alcohol dehydrogenases or short-chain dehydrogenases, aldehyde dehydrogenases, and several CYP isoforms (Duester, 1996). For example, enzymes from the CYP1 family and CYP3A subfamily have been found to catalyse the oxidation of retinol to retinal, which is the rate-limiting step in the synthesis of retinoic acid (Chen et al., 2000), and CYP1A1 is involved in the oxidation of retinal to retinoic acid (Raner et al., 1996). CYPs are also important in the catabolism of retinoic acid; CYP26A1, a product of the CYP26 gene family, catalyses the 4-hydroxylation of all-trans-retinoic acid (White et al., 1997). Degradation of retinoic acid by CYP26A1-mediated hydroxylation limits the available ligand for the retinoic acid receptors RAR and RXR. Other members of the CYP26 gene family, CYP26B1 and CYP26C1, also appear to be involved in the

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metabolism of retinoic acid or its derivatives, but their precise biological roles remain to be elucidated (Nebert & Russell, 2002).

Vitamin D (vitamin D3) plays an essential role in a variety of important biological processes including bone formation and metabolism, calcium homeostasis, and cellular differentiation (Walters, 1992). Vitamin D3 itself does not initiate any of these biological responses. Rather, vitamin D3 is a precursor that is oxidised to the major active hormonal form of vitamin D, 1D,25-dihydroxyvitamin D3 [1D,25(OH)2D3], by the vitamin D endocrine system, in which various CYPs play a fundamental role (Wikvall, 2001).

1D,25(OH)2D3 exerts its biological effects by acting as a ligand for the vitamin D receptor (VDR), a member of the nuclear hormone receptor superfamily (Haussler et al., 1998). Vitamin D3 is converted into its active hormonal form, 1D,25(OH)2D3, by two sequential hydroxylation reactions that are catalysed by CYPs (Wikvall, 2001). Firstly, vitamin D3 is hydroxylated within the liver to 25-hydroxyvitamin D3 [25(OH)D3], a reaction that is catalysed by the mitochondrial CYP27A1 (Usui et al., 1990; Wikvall, 2001; Nebert & Russell, 2002), which was discussed previously for its key role in bile acid biosynthesis (section 1.3.2). It has been suggested that the microsomal CYP enzymes CYP2D6 and CYP3A4 may also catalyse the 25-hydroxylation of vitamin D3

(Nebert & Russell, 2002). 25-hydroxyvitamin D3 is subsequently hydroxylated in the kidney into 1D,25-dihydroxyvitamin D3 [1D,25(OH)2D3], the active hormonal ligand for the VDR, by CYP27B1 (Monkawa et al., 1997; Wikvall, 2001; Nebert & Russell, 2002). Aside from their essential role in the metabolic activation of vitamin D, CYP enzymes also play a key role in its metabolic deactivation. Specifically, CYP24A1 catalyses the 24-hydroxylation of 1D,25(OH)2D33 into 1D,24,25-(OH) D3; this represents the first step in the major catabolic pathway of 1D,25(OH)2D3, and prevents

1D,25(OH)2D3 from binding to the VDR (Beckman et al., 1996; Nebert & Russell, 2002).

CYP24A1 also acts on the 1D,25(OH)2D3 precursor 25-hydroxyvitamin D3 [25(OH)D3], thus limiting the production of the 1D,25(OH)2D3 VDR ligand (Beckman et al., 1996; Nebert & Russell, 2002).

Due to the essential role of 1D,25(OH)2D3 in calcium homeostasis, its serum levels are tightly controlled by regulating the expression and activity of the CYP enzymes involved in 1D,25(OH)2D3 metabolism (Kato, 1999). Thus, the expression and activity of CYP27B1 is increased by calcitropic hormones such as parathyroid hormone that are released in response to falling serum calcium levels (Walters, 1992; Kato, 1999). Higher expression and activity of CYP27B1 enhances the production of

1D,25(OH)2D3, which serves to increase the intestinal absorption of calcium. On the other hand, high serum phosphate and calcium levels reduce the activity of CYP27B1,

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and in a classic negative feedback loop, 1D,25(OH)2D3 regulates its own production by acting as a negative regulator of CYP27B1, the final enzyme in its biosynthetic pathway (Walters, 1992; Kato, 1999). As a further level of control, the 1D,25(OH)2D3 hormone also induces the expression of the CYP24A1 gene, whose protein product plays a key role in its catabolic breakdown (Chen & DeLuca, 1995).

1.3.4 CYPs Involved in the Metabolism of Arachidonic Acid 1.3.4.1 Arachidonic Acid and the Arachidonic Acid Metabolic Cascade Arachidonic acid (AA) is a 20-carbon fatty acid that plays a fundamental role within the body. AA is a component of most cellular membranes where it serves a recognised structural role within the cell (Makita et al., 1996). In addition to its structural functions, AA may be released from cell membranes and undergo metabolism into a variety of physiologically important metabolites or lipid mediators known as eicosanoids, which serve as mediators in a variety of transmembrane signalling cascades. Eicosanoids may play a role in the pathophysiology of diseases such as , asthma, hypertension, cardiovascular disease, diabetes and cancer (Capdevila et al., 2002). AA is unique amongst other polyunsaturated eicosanoid precursors in that it is present in vivo esterified to hormonally sensitive cell membrane glycerophospholipids and thus is the predominant eicosanoid precursor in mammalian cells (Capdevila et al., 1992; Funk, 2001). Whilst bound to phospholipid, AA cannot undergo metabolism to biologically active eicosanoids (Capdevila & Falck, 2002), however, in response to a variety of biological and mechanical stimuli, phospholipases (e.g. cytosolic phospholipase A2;

PLA2) are activated and facilitate the release of free AA from membrane phospholipids, making it available for oxidative metabolism by three different enzyme systems/pathways (refer to Figure 1.3; Zeldin, 2001). The first two pathways, catalysed by prostaglandin H synthase and lipoxygenase enzymes, have been extensively studied, and the biological significance of their eicosanoid products has been firmly established for quite some time. Prostaglandin H synthases (or ; COXs) metabolise AA to the unstable endoperoxide prostaglandin H2, which is subsequently converted by downstream enzymes into prostaglandins, thromboxane A2 and prostacyclin in a cell specific manner (Figure 1.3; Funk, 2001). Lipoxygenases metabolise AA into several labile regioisomeric allylic hydroperoxy intermediates which serve as precursors for leukotrienes, lipoxins and hydroxyeicosatetraenoic acids (HETEs; Figure 1.3; Brash, 1999).

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Arachidonic acid

COOH

Cyclooxygenase Lipoxygenase

Cytochrome P450 Prostaglandins HETEs Prostacyclin Leukotrienes Thromboxane A2 Lipoxins

Lipoxygenase-like Z/Z-1 Hydroxylase Epoxygenase

COOH COOH COOH

20-HETE 11,12--EET 12-HETE CH2OH HO O 5-HETE 16-HETE 5,6-EET 8-HETE 17-HETE 8,9-EET 9-HETE 18-HETE 11,12-EET 11-HETE 19-HETE 14,15-EET 12-HETE 20-HETE 15-HETE

Figure 1.3 Pathways for the metabolism of arachidonic acid Arachidonic acid is metabolised by cyclooxygenase, lipoxygenase and cytochrome P450 enzymes to yield prostaglandins, prostacyclin, thromboxane A2, lipoxins, leukotrienes, hydroxyeicosatetraenoic acids (HETEs) and epoxyeicosatrienoic acids (EETs).

The third and most recently recognised pathway for AA metabolism is catalysed by CYPs. The first suggestion of a role for CYPs in the metabolism of AA occurred almost thirty years ago (in 1976) when it was demonstrated that CYP inhibitors blocked the AA-induced aggregation of human platelets (Cinti & Feinstein, 1976). In 1981 several groups unequivocally demonstrated that AA was metabolised by CYPs in the liver and in the kidney, and that these CYP-derived eicosanoids were chromatographically distinct from those produced by the COXs and lipoxygenases (Capdevila et al., 1981; Morrison & Pascoe, 1981; Oliw et al., 1981). Subsequent studies demonstrated that the CYP-derived eicosanoids possess potent biological activities and are produced endogenously within a range of rabbit, rat and human organs including the liver, kidney and lung, as well as being detectable in the urine and plasma (Makita et al., 1996; Capdevila et al., 2002). These early studies established that the metabolism of AA by CYPs was an endogenous metabolic pathway and implicated potentially important functional roles in eicosanoid homeostasis for CYPs

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(Capdevila et al., 2002). Subsequent to these early findings, CYP-dependent metabolism of AA and its physiological and pathophysiological significance has become an area of intense research (Capdevila et al., 2002).

1.3.4.2 The Third Pathway in the AA Cascade: Metabolism of AA by CYPs Substantial differences exist between the CYP AA metabolic pathway and the cyclooxygenase and lipoxygenase pathways (Fitzpatrick & Murphy, 1988). COXs and lipoxygenases are dioxygenase enzymes that catalyse the free radical-mediated activation of substrate carbon atoms. On the other hand, CYPs are membrane bound enzymes that catalyse the redox-coupled activation of molecular oxygen and the delivery of one of the oxygen atoms to substrate ground-state carbons (Capdevila et al., 2002). With the exception of 5-lipoxygenase, most of the dioxygenases have minimal requirements other than molecular oxygen (Fitzpatrick & Murphy, 1988). By contrast, CYP-mediated AA metabolism requires molecular oxygen, NADPH/NADP+ or NADH/NAD+, and the flavoprotein NADPH-CYP reductase as cofactors (Fitzpatrick & Murphy, 1988). As with the other enzymes in the AA cascade, CYPs are unable to oxidise AA at a significant rate when it is bound to membrane phospholipids (Capdevila et al., 2002), and thus CYP-mediated biotransformation of AA requires its release from cellular phospholipid pools by the hormonally regulated phospholipases (Capdevila & Falck, 2001). As shown in Figure 1.3, CYPs metabolise AA to three types of eicosanoid products by one or more of the following reactions; (1) allylic and/or bis-allylic oxidation (lipoxygenase-like reaction) This reaction generates several regioisomeric hydroxyeicosatetraenoic acids that contain a cis, trans-conjugated dienol functional group (5-, 8-, 9-, 11-, 12-, and 15- hydroxyeicosatetraenoic acids; HETEs). (2) Z/Z-1-terminal hydroxylations (also called the Z/Z-1 hydroxylase reaction) These reactions catalyse hydroxylations at or near the terminal sp3 carbon to produce

C16 – C 20 alcohols of AA (16-, 17-, 18-, 19-, and 20-hydroxyeicosatetraenoic acids; HETEs). (3) Olefin epoxidation (also called the epoxygenase reaction) This reaction produces four regioisomeric cis-epoxyeicosatrienoic acids (5,6-, 8,9-, 11,12-, and 14,15-epoxyeicosatrienoic acids; EETs). Each of these four EETs can be formed as either the S,R or the R,S enantiomer (Zeldin, 2001; Capdevila & Falck, 2002).

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Studies using either microsomal fractions or purified CYPs demonstrated that the type of reaction catalysed (i.e. allylic oxidation, Z-hydroxylation or olefin epoxidation), the regioselectivity of oxygen insertion (i.e. allylic oxidation at either of the three bis-allylic methylenes, hydroxylation at C16-C 20 or epoxidation at either of the four olefins), and the enantiofacial selectivity of the oxygen insertion (i.e. S,R or R,S) is CYP isoform specific (Capdevila & Falck, 2002). As such, the profile of eicosanoid metabolites produced is dependent on the particular animal species and tissue, as well as sex, age, diet, hormonal status and exposure to xenobiotics, all of which influence the expression of specific CYPs (Capdevila & Falck, 2002). Of the three reactions described above, the Z/Z-1 hydroxylase and epoxygenase pathways have been the most extensively studied. These are the main CYP-catalysed reactions in most tissues, producing, respectively, the major CYP- derived eicosanoids, 20-HETE and EETs (Kroetz & Zeldin, 2002). By contrast, further research is required to establish the role and relevance of CYPs in the in vivo formation of HETEs via the allylic oxidation pathway (Capdevila & Falck, 2001).

1.4 The Z/Z-1 Hydroxylase Pathway of AA Metabolism

The Z/Z-1 hydroxylation of saturated fatty acids is one of the first reactions attributed to the microsomal CYPs (Lu et al., 1969). As a general rule, the rate of fatty acid hydroxylation decreases with increasing carbon chain length. Thus, medium chain fatty acids (C12 to C 16), such as lauric acid (present only at very low levels in mammals) are metabolised significantly faster than AA (Capdevila & Falck, 2002). While microsomal Z/Z-1 hydroxylation of AA occurs in several tissues including the liver, kidney, brain, lung and intestine, it is within the kidney where this reaction is most prevalent, and thought to have its most important functional roles (McGiff & Quilley, 1999; Capdevila & Falck, 2002). 20-HETE is the predominant, and most biologically potent, product of CYP- mediated hydroxylation of AA. In addition to 20-HETE, the Z/Z-1 hydroxylase pathway produces 16-, 17-, 18- and 19- HETEs, but these appear to be minor products in most tissues, and their biological functions are less well defined (Kroetz & Zeldin, 2002). Studies with microsomal fractions and purified CYPs have shown that the CYP4A subfamily is the predominant CYP family that mediates 20-HETE formation in tissues (Capdevila et al., 2000). CYP4As are structurally and functionally conserved

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proteins that are specialised for the metabolism of fatty acids, and display little capacity to metabolise xenobiotics (Capdevila & Falck, 2002). CYP4A expression is regulated by many physiological factors, including age, diet and hormones, and also by pathophysiological states such as diabetes and hypertension (Capdevila & Falck, 2002). As discussed further in section 1.8.2.5, the nuclear peroxisome proliferator- activated receptor alpha (PPARD) plays a key role in regulating the expression of CYP4A genes (Johnson et al., 1996; Waxman, 1999). A number of CYP4A isoforms from several species including rat, mouse, rabbit and human, have been identified, and most have demonstrated the ability to metabolise AA to 20-HETE (Capdevila & Falck, 2002). Four isoforms have been identified in the rat (CYP4A1, 4A2, 4A3 and 4A8), and rabbit (CYP4A4, 4A5, 4A6 and 4A7), and three have been identified in the mouse (CYP4A10, 4A12 and 4A14; Capdevila & Falck, 2002). To date, only one CYP4A isoform with Z/Z-1 hydroxylase activity toward AA, CYP4A11, has been identified in humans (Gainer et al., 2005). CYP4A11 is most related to rat CYP4A8 and mouse CYP4A12 (Capdevila & Falck, 2002) and is expressed in the liver and kidney (Powell et al., 1998). Apart from CYP4A11, human liver and kidney also expresses CYP4F2, which also catalyses AA 20-hydroxylation (Powell et al., 1998). It now appears that CYP4F2 is the major CYP isoform responsible for the production of 20-HETE within the human kidney (Lasker et al., 2000). Additional human CYP4A and 4F isoforms continue to be identified, but to date their functions are unknown (Nebert & Russell, 2002). Other CYPs have demonstrated AA hydroxylase activity. CYPs 1A1 and 1A2 catalyse the oxidation of AA at carbons C16 to C 19 to produce 16-, 17-, 18- and 19- HETEs (Falck et al., 1990), CYP2E1 converts AA to 18- and 19-HETEs (Laethem et al., 1993), and certain members of the CYP2J family, such as rat CYP2J3 (Wu et al., 1997) and mouse CYP2J9 (Qu et al., 2001), also catalyse hydroxylation of AA into HETEs. In addition, several CYP4Fs catalyse prostanoid and leukotriene hydroxylation. Indeed, human CYP4F3 is a specific leukotriene Z-hydroxylase (Capdevila & Falck,

2002), and CYP4F8 is an efficient prostaglandin H2 Z-1 hydroxylase (Bylund et al., 2000).

1.4.1 Biological Activities of 20-HETE The potent biological activities of the products of AA Z/Z-1 hydroxylases, especially 20-HETE itself, has stimulated extensive research into the physiological and pathophysiological roles of the CYP AA Z/Z-1 hydroxylases (Capdevila & Falck, 2002).

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20-HETE has potent vasoactive and mitogenic properties, modulates renal ion transport, and is a second messenger in signal transduction cascades for hormones, such as endothelin-1 and angiotensin ,,, that have key roles in blood pressure regulation via regulation of renal circulation and electrolyte excretion. As such, 20-HETE plays a key role in renal function, and is thought to be involved in the development of hypertension and cardiovascular disease (McGiff & Quilley, 1999; Roman, 2002). The main biological activities of 20-HETE, with emphasis on their renal effects, are discussed below (sections 1.4.2 – 1.4.5).

1.4.2 Vascular Effects of 20-HETE 1.4.2.1 Vasoconstriction and the Regulation of Vascular Tone 20-HETE is produced endogenously within vascular smooth muscle (VSM) cells in response to increases in intracellular calcium (Ca2+; Fleming, 2001), and is a powerful vasoconstrictor of renal (Imig et al., 1996b), cerebral (Gebremedhin et al., 2000), mesenteric (Wang et al., 2001) and skeletal muscular arterioles (Harder et al., 1996). The vasoconstrictor effects of 20-HETE are largely attributed to its ability to inhibit the opening of large-conductance calcium-activated potassium channels (KCa). 2+ Inhibition of KCa channels results in the depolarisation of VSM cells and enhances Ca entry via voltage-sensitive L-type Ca2+ channels; the resultant increase in intracellular Ca2+ ultimately causes vascular contraction (Roman, 2002). The signal transduction mechanism(s) by which 20-HETE blocks the activity of the KCa channel has not been identified (Roman, 2002). Contraction by 20-HETE may also be achieved via the inhibition of the Na+-K+-ATPase (Fleming, 2001). Studies have indicated that 20-HETE plays an important role in the signal transduction pathways mediating the vasoconstrictor response of numerous different vasoactive substances that regulate blood pressure, including angiotensin ,,, endothelin, vasopressin and norepinephrine (Fleming, 2001). Thus, inhibitors of CYPs that generate 20-HETE attenuate the vasoconstrictor responses to these vasoactive agents in rat renal and mesenteric arteries (Roman, 2002). Similarly, 20-HETE has been shown to play a role in blood flow autoregulation, the physiological process in which a constant blood flow to organs is maintained over a wide range of perfusion pressures, with AA Z-hydroxylase inhibitors found to impair autoregulation of renal (Zou et al., 1994a) and cerebral blood flow (Gebremedhin et al., 2000). Constriction of preglomerular arterioles by 20-HETE is a key modulator of renal tubuloglomerular

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feedback (Zou et al., 1994b) which regulates glomerular filtration and ion transport, and consequently, blood pressure (Carroll & McGiff, 2000; Roman, 2002). The mechanism by which 20-HETE contributes to the control of vascular tone is summarised in Figure 1.4.

Stretch Ang,,, NE

phospholipid VASCULAR + + SMOOTH + MUSCLE PLC PLA2 CELL DAG AA CYP Contraction IP3 20-HETE

+ +

Ca2+ K+ - + + - Em 2+ L-type Ca KCa channel

Ca2+ K+

Figure 1.4 Role of 20-HETE in the regulation of vascular tone Membrane stretch and vasoactive agents [e.g. angiotensin ,, (Ang,,), norepinephrine (NE)] activate phospholipase C (PLC) which leads to the release of inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG). IP3 triggers the release of intracellular Ca2+ from the endoplasmic reticulum which induces vascular contraction. 2+ Increased intracellular Ca activates DAG lipase and phospholipase A2 (PLA2) to release arachidonic acid (AA) from membrane phospholipids, which is subsequently converted into 20-HETE by CYP Z-hydroxylases. 20-HETE inhibits the activation of large-conductance calcium-activated potassium channels (KCa ); inhibition of K Ca 2+ channels decreases membrane potential (Em), which increases Ca entry through voltage-sensitive L-type Ca2+ channels and enhances vascular contraction.

Membrane stretch and vasoactive agents such as angiotensin ,, and norepinephrine activate phospholipase C (PLC) in VSM cells and subsequently

increase the synthesis of inositol 1,4,5-trisphosphate (IP3), which triggers the release of Ca2+ from intracellular stores and stimulates vascular contraction. However, the rise in intracellular Ca2+ would also be expected to activate calcium-activated potassium 2+ channels (KCa), leading to membrane hyperpolarisation and decreased Ca influx through voltage-sensitive Ca2+ channels. This would ultimately attenuate the vasoconstrictor response. 20-HETE is thought to regulate vascular tone by buffering or 2+ counteracting the activation of KCa channels after the release of intracellular Ca . As shown in Figure 1.4, increases in intracellular Ca2+ also stimulate the release of AA by 2+ activating Ca -sensitive phospholipase A2 (PLA2) and diacylglycerol (DAG) lipase. The

action of CYP4A and 4F enzymes on AA generates 20-HETE which inhibits KCa

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channels, and opposes the activation of KCa channels that would normally result from 2+ the release of intracellular Ca . Inhibition of KCa channels promotes membrane depolarisation of VSM cells, enhanced entry of Ca2+ through voltage-sensitive L-type Ca2+ channels, and a more intense, prolonged vascular contraction (Roman et al., 2000).

1.4.2.2 Role for 20-HETE in Vasodilation Pathways While 20-HETE has potent vasoconstrictive properties throughout most vascular beds, it is vasodilatory in pulmonary arteries (Birks et al., 1997). The vasodilatory action of 20-HETE within the lung is thought to be dependent on its metabolism by COX to a dilatory substance (Birks et al., 1997), and it has been suggested that 20-HETE may be important in counteracting hypoxia-induced vasoconstriction in the lung (Zhu et al., 2000a). In addition to the pulmonary pathway described above, 20-HETE could play an indirect role in the regulation of vascular tone by nitric oxide (NO), which is an important vasodilatory pathway. The vasoregulatory effect of NO is understood to be mediated, at least in part, by its ability to modulate CYP-dependent 20-HETE formation (Alonso-Galicia et al., 1998b). NO inhibits 20-HETE formation by binding to the haem moiety of CYP4A enzymes; thus, the vasodilatory effects of NO can be partly attributed to its inactivation of CYP4A enzymes and resultant inhibition of 20-HETE formation

(Sun et al., 1998). Indeed, studies have demonstrated that NO activates KCa channels in cerebral and renal VSM cells, which is inhibited by maintaining intracellular 20-HETE levels (Sun et al., 1998). Like NO, carbon monoxide also binds strongly to the haem moiety in CYP enzymes, and evidence is emerging that the vasodilator effects of carbon monoxide may also be mediated, in part, by inhibition of 20-HETE formation (Roman et al., 2000).

1.4.2.3 20-HETE as a Possible Oxygen Sensor In addition to myogenic responses, oxygen-dependent autoregulatory mechanisms contribute to skeletal muscle vascular tone and the regulation of tissue blood flow. A number of studies have suggested that 20-HETE may be an oxygen sensor in the skeletal muscle microcirculation. Thus, the production of 20-HETE within the kidney and vascular tissue is dependent on the local oxygen tension, and inhibitors of 20-HETE formation prevent the vasoconstrictor response to elevated tissue oxygen levels (Harder et al., 1996; Kunert et al., 2001).

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1.4.3 Effect of 20-HETE on Ion Transport within the Kidney In addition to regulating blood pressure via the modulation of renal vascular tone, 20-HETE also influences blood pressure via its effects on ion transport in the kidney. 20-HETE is the predominant renal eicosanoid responsible for the regulation of sodium (Na+) transport and electrolyte excretion (Carroll & McGiff, 2000). In particular, 20-HETE is the major AA metabolite produced within key segments of the nephron involved in ion transport; the proximal tubule and the thick ascending loop of Henle (TALH; Roman et al., 2000). Within the proximal tubule, 20-HETE inhibits Na+ reabsorption by inhibiting Na+-K+-ATPase activity (Roman et al., 2000). Indeed, the inhibitory effects of dopamine, parathyroid hormone and angiotensin ,, on Na+-K++-ATPase activity and Na reabsorption in the proximal tubule are dependent on their ability to stimulate the formation of 20-HETE via the activation of phospholipase A2 (Roman et al., 2000; Roman, 2002). 20-HETE inhibits Na+-K+-ATPase activity by stimulating protein kinase C to phosphorylate the D-subunit of the Na+-K+-ATPase pump (Nowicki et al., 1997). 20-HETE plays a key role in the regulation of chloride (Cl- ) transport by blocking a 70-ps K+ channel in the apical membrane of the TALH, which limits the availability of potassium (K++) for transport via the Na -K+--2Cl cotransporter and reduces Cl - transport (Wang & Lu, 1995; Roman, 2002). Apart from inhibiting Cl- transport, inhibition of the Na+-K+--2Cl cotransporter by 20-HETE also decreases the lumen positive transepithelial potential which is the main force that drives the passive reabsorption of cations including Na+, K+2+, Ca and magnesium (Mg 2+ ) in the TALH (Roman et al., 2000). As such, the inhibitory effects of bradykinin, angiotensin ,, and increased intracellular Ca2+ concentration on Na + transport in the TALH are dependent on 20-HETE (Roman et al., 2000; Roman, 2002).

1.4.4 Mitogenic Actions of 20-HETE Increasing evidence suggests that 20-HETE may contribute to the mitogenic actions of growth factors and vasoactive substances in various cells including VSM cells and renal mesangial cells (Roman et al., 2000). In particular, 20-HETE reportedly increases thymidine incorporation in various different cell types (Roman & Alonso-Galicia, 1999), activates the mitogen-activated protein (MAP) kinase signalling pathway (Roman & Alonso-Galicia, 1999) and stimulates the mitogenic responses to angiotensin ,,, epidermal growth factor (EGF) and norepinephrine in cultured aortic VSM cells (Roman et al., 2000; Roman, 2002). CYP inhibitors attenuate the mitogenic effects of a range of growth-stimulating agents

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such as serum, EGF, phorbol esters and vasopressin in cultured glomerular mesangial cells (Roman & Alonso-Galicia, 1999), and inhibition of 20-HETE formation attenuates EGF-stimulated growth in rat proximal tubular cells (Lin et al., 1995). Based on these studies, 20-HETE is proposed to play a role in angiogenesis and the proliferation of mesangial cells, and may be involved in the pathogenesis of glomerulosclerosis (a mitogenic renal condition) in disease states such as diabetes and hypertension (Roman et al., 2000).

1.4.5 Role of 20-HETE in the Pathogenesis of Hypertension Given the key role of 20-HETE in the regulation of blood pressure via its vasoconstrictive properties within the renal vasculature and its regulation of renal Na+ transport and electrolyte excretion, it is not surprising that 20-HETE is proposed to play a role in the pathophysiology of hypertension (Roman et al., 2000; Roman, 2002). A role for 20-HETE in the development of hypertension was proposed some time ago, and has been intensively investigated in animal models of hypertension. While there is now considerable evidence that the production of 20-HETE within the kidneys is altered in models of hypertension, the consequences of these changes on blood pressure are complicated by the fact that 20-HETE displays both pro- hypertensive and anti-hypertensive properties, depending on its site of production within the kidney. Thus, in the proximal tubule and the TALH, 20-HETE inhibits Na+ reabsorption; this reduces plasma volume and arterial pressure. On the other hand, within the renal vasculature and glomerulus, and also in the peripheral vasculature, 20-HETE is a potent vasoconstrictor that increases vascular tone and blood pressure (Roman, 2002). The spontaneously hypertensive rat (SHR) is the most widely studied model of hypertension with respect to 20-HETE (Kroetz & Zeldin, 2002). SHR rats display a constant increase in blood pressure between 5 and 10 weeks of age, relative to Wistar- Kyoto (WKY) control rats (Capdevila & Falck, 2001). Early studies demonstrated that the formation of 20-HETE is significantly higher in the kidneys of SHR rats compared to controls (Sacerdoti et al., 1988), and that the CYP4A2 gene (whose protein product, CYP4A2, generates 20-HETE from AA), is regulated by sodium chloride and is selectively overexpressed in the kidney of SHR during the developmental phase of hypertension (Iwai & Inagami, 1991). As 20-HETE is a potent renal vasoconstrictor, it is postulated that the increased expression of CYP4A2 and increased production of 20-HETE may contribute to the development of hypertension in SHR by increasing renal vascular resistance (Roman, 2002). Several recent studies support this proposal. In particular, reduced production of 20-HETE by selective inhibition of Z-hydroxylase

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activity in SHR reduces blood pressure (Su et al., 1998), and treatment of SHR with antisense oligonucleotides directed against CYP4A reportedly decreases vascular reactivity and blood pressure (Wang et al., 2001). Another animal model of hypertension that exhibits altered 20-HETE production is the Dahl salt-sensitive (DS) rat, which develops hypertension after 10 to 18 days of consumption of a high salt diet; Dahl salt-resistant (DR) rats remain normotensive (Capdevila & Falck, 2001). In contrast to the situation in SHR, decreased levels of 20-HETE may be involved in the development of hypertension in DS rats. Cl- reabsorption in the TALH of DS rats is elevated compared to control DR rats; this elevated Cl- uptake increases plasma volume and blood pressure (Ito & Roman, 1999). Since 20-HETE is an endogenous inhibitor of Cl- transport in the TALH, it was suggested that altered production of 20-HETE may account for the dysregulation of Cl- transport in DS rats (Roman, 2002). Studies comparing the levels of 20-HETE in kidneys of DS and DR rats found that, although produced at similar levels in the renal cortex, CYP4A expression and 20-HETE production in the outer medulla and TALH is lower in DS rats than in DR controls and other normotensive strains (Ma et al., 1994; Stec et al., 1996). A number of further studies support the hypothesis that a deficiency in 20-HETE contributes to increased Cl- reabsorption and high blood pressure in DS rats. Ito & Roman (1999) demonstrated that exogenous administration of 20-HETE reduces Cl- reabsorption in the TALH of DS rats to normal levels, but does not affect transport of Cl- in the TALH of salt-resistant rats. Administration of fibrate drugs up-regulates renal CYP4A expression in DS rats, which improves renal function and prevents the development of hypertension (Alonso-Galicia et al., 1998a). Furthermore, pharmacological inhibition of the renal medullary formation of 20-HETE produced hypertension in normally salt- resistant Lewis strain rats that were placed on high-salt diets (Stec et al., 1997). In addition to altered 20-HETE production, the up-regulation of renal CYP epoxygenase pathways in response to high salt is defective in DS rats (Makita et al., 1994). This pathway generates the anti-hypertensive EETs (discussed in detail in section 1.5) and is normally induced in response to high salt in normotensive strains of rats (Makita et al., 1994). Thus the impaired ability to increase production of EETs in response to high salt may be another factor that contributes to the development of hypertension in DS rats. Further support for a role for 20-HETE in the development of hypertension was provided by a recent study of mice with a targeted disruption of the Cyp4a14 gene (the murine homologue of rat CYP4A2/4A3). Disruption of this gene was predicted to lower production of 20-HETE, but these animals exhibited a paradoxical increase in renal

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production of 20-HETE and developed spontaneous hypertension. The increase in 20-HETE production was due to a compensatory increase in the expression of Cyp4a12. The enhanced expression of Cyp4a12 in close proximity to the afferent arterioles of the kidney was correlated with increased resistance of the vessels and impaired myogenic responsiveness to increased renal perfusion pressure. It was concluded that the development of hypertension in Cyp4a14 knockout mice may be due to the up-regulation of Cyp4a12 expression and the resultant enhanced renal production of the vasoconstrictor 20-HETE (Holla et al., 2001). Thus, it is clear that the production of 20-HETE within the kidney is altered in a number of models of hypertension, and that manipulation of the production of 20-HETE within these models affects blood pressure. Further studies are required to unravel the complex effects of altered 20-HETE production in the various models. In this respect, studies investigating the catalytic efficiency, regulation and renal localisation of different CYP4A isoforms will be helpful (McGiff & Quilley, 1999). Indeed, it has been shown that the four rat CYP4A isoforms, which differ in their ability to generate 20-HETE, are differentially expressed within various segments of the nephron (Ito et al., 1998).

1.5 The Epoxygenase Pathway of AA Metabolism

The olefinic epoxidation of AA is referred to as the epoxygenase pathway of AA metabolism and generates four regioisomeric cis-epoxyeicosatrienoic acids; 5,6-, 8,9-, 11,12-, and 14,15-epoxyeicosatrienoic acids (EETs; Figure 1.5) that have a number of important biological activities (Zeldin, 2001). In mammals, CYPs are the only enzymes that catalyse the epoxidation of fatty acids to bis-allylic, cis-, and in contrast to the Z/Z-1 hydroxylase pathway by which a number of different fatty acids can be efficiently metabolised, the epoxygenase pathway is essentially selective for AA (Capdevila et al., 2002). CYP-mediated conversion of AA to EETs occurs in numerous tissues including the liver, kidney, lungs, heart, endothelium, brain, adrenal, pituitary, ovaries and skin (Capdevila & Falck, 2001). Studies with purified and microsomal CYPs have indicated that numerous isoforms are able to catalyse EET formation with varying efficiencies. Thus, members of the mammalian CYP1A, CYP2B, CYP2C, CYP2D, CYP2E, CYP2G, CYP2J, CYP2N and CYP4A subfamilies have been reported to catalyse the formation of EETs (Zeldin, 2001; Kroetz & Zeldin, 2002).

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Arachidonic acid COOH

CYP Epoxygenases

5,6-EET 8,9-EET O O COOH COOH

11,12-EET 14,15-EET COOH COOH

O O

Figure 1.5 Metabolism of arachidonic acid by CYP epoxygenases generates four EET regioisomers; 5,6-EET, 8,9-EET, 11,12-EET and 14,15-EET

In humans, EET biosynthesis appears to be predominantly catalysed by CYP2C and CYP2J isoforms (Spector et al., 2004). In any given tissue, multiple CYPs may contribute to EET biosynthesis, and the relative contribution of individual isoforms will depend on their abundance within that tissue as well as their catalytic efficiency (Zeldin, 2001). In human liver and kidney, CYP2C isoforms are probably the major AA epoxygenases (Rifkind et al., 1995; Zeldin et al., 1995a; Zeldin et al., 1996b), while human CYP2J2 appears to be the major enzyme involved in the extrahepatic formation of EETs (Wu et al., 1996; Zeldin et al., 1996a; Zeldin et al., 1997a; Zeldin et al., 1997b). While CYP epoxygenases produce all four regioisomeric EETs from AA (i.e. 5,6-, 8,9-, 11,12- and 14,15- EET), each particular CYP isoform generally displays a characteristic regioselectivity of EET biosynthesis. For example, human CYP2C8 primarily metabolises AA to 14,15-EET and 11,12-EET, while the closely related CYP2C9 produces significant amounts of 14,15-EET, 11,12-EET and 8,9-EET from AA (Daikh et al., 1994). Each regioisomer is produced as a mixture of R,S- and S,R- enantiomers, and, as with regioselectivity, each CYP isoform generally produces EETs in a stereoselective manner. Thus, CYP2C8 predominantly produces 11R,12S-EET, while CYP2C9 predominantly produces the S,R-enantiomer (Daikh et al., 1994). EETs are metabolically unstable, and once formed, are generally rapidly degraded by fatty acid E-oxidation, or are converted into the corresponding dihydroxyeicosatrienoic acids (DHETs) and glutathione-conjugates by

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and glutathione S-, respectively (Capdevila & Falck, 2002). Although DHETs were originally thought to be biologically inactive metabolic byproducts of EETs, they have recently been found to have vasoactive properties (Lu et al., 2001). EETs may also be esterified in membrane glycerophospholipids. The in vivo esterification appears to be unique, amongst other eicosanoids, to EETs, and suggests that cells have the ability to generate EETs independently of oxidative AA metabolism via the hydrolysis of membrane phospholipids. Furthermore, it has been suggested that the incorporation of EETs into the cellular membrane may account for some of the biological activities of EETs (Karara et al., 1991; Capdevila et al., 1992).

1.5.1 Biological Activities of EETs EETs have a wide range of biological activities. In particular, EETs have potent effects within the vascular system, where they are vasodilatory and are proposed to be the putative endothelium-derived hyperpolarising factor (Fleming, 2001; Roman, 2002). EETs are also reported to be anti-inflammatory (Node et al., 1999), fibrinolytic (Node et al., 2001), anti-apoptotic (Chen et al., 2001) and are reported to influence smooth muscle cell migration (Sun et al., 2002). Biological activities of EETs outside the vasculature include effects on renal tubular ion transport, mitogenic activity, and effects on peptide hormone secretion (Roman, 2002). The biological effects of EETs are summarised in Table 1.3 and are discussed in detail in the sections below (sections 1.5.2 – 1.5.4).

1.5.2 Vascular Effects of EETs 1.5.2.1 Vasodilatory Effects of EETs within the Vascular System EETs are produced from AA within numerous different vascular beds including coronary (Campbell et al., 1996), renal (Zou et al., 1994a), cerebral (Gebremedhin et al., 1992) and pulmonary arteries (Zhu et al., 2000b). In contrast to 20-HETE, which is produced in VSM cells, EETs are produced by vascular endothelial cells (Roman et al., 2000). EETs have important physiological effects within the vasculature: in particular, they are potent dilators of renal (Imig et al., 1996a), cerebral (Gebremedhin et al., 1992) and coronary arteries (Campbell et al., 1996). As depicted in Figure 1.6, given the opposing effects of EETs and 20-HETE on arterial tone, it has been suggested that modulation of vascular tone and blood pressure may be partly regulated by both sets of eicosanoids (Fleming, 2001).

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Table 1.3 Biological actions of EETs

Action Reference

Vascular System Vasodilation Fleming, 2001; Roman, 2002 Anti-inflammatory Node et al., 1999 Inhibition of vascular smooth muscle (VSM) cell migration Sun et al., 2002 Stimulation of fibrinolysis Node et al., 2001 Stimulation of endothelial and VSM cell proliferation Fleming et al., 2001a Stimulation of tube formation Munzenmaier & Harder, 2000 Protection against hypoxia-reoxygenation injury Yang et al., 2001 Inhibition of platelet aggregation Fitzpatrick et al., 1986

Heart Inhibition of cardiomyocyte Na+ channels Lee et al., 1999 Modulation of cardiomyocyte Ca2+ channels Xiao et al., 1998; Chen et al., 1999 Modulation of myocardial function following ischaemia Moffat et al., 1993; Wu et al., 1997

Kidney Stimulation of renal mesangial and epithelial cell proliferation Harris et al., 1990; Chen et al., 1998 Modulation of renal ion transport and fluid excretion Roman, 2002 Inhibition of apoptosis Chen et al., 2001

Endocrine System Stimulation of pituitary peptide hormone release (e.g. growth Capdevila et al., 1983; Negro-Vilar et al., 1985; hormone, somatostatin, vasopressin) Snyder et al., 1989 Stimulation of pancreatic hormone release (insulin and glucagon) Falck et al., 1983

Lung Modulation of pulmonary ion transport and airway fluid composition Jacobs & Zeldin, 2001 Modulation of airway smooth muscle tone Zeldin et al., 1995b; Dumoulin et al., 1998

Liver Activation of phosphorylase a and modulation of hepatic Ca2+ Kutsky et al., 1983; Yoshida et al., 1990 metabolism

EETs mediate vasodilation by activation of large-conductance calcium-activated potassium (KCa) channels in VSM cells. This leads to hyperpolarisation of VSMs and vascular relaxation (Figure 1.6; Gebremedhin et al., 1992; Campbell et al., 1996). The mechanism by which EETs alter KCa channel activity is unclear, but may involve EET- mediated ADP-ribosylation of the membrane-associated guanine nucleotide-binding protein GsĮ (Li et al., 1999; Li et al., 2002), and stimulation of the adenylyl cyclase/cyclic AMP (cAMP)/protein kinase A (PKA) signalling pathway (Imig et al., 1999).

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Stretch Ang,,, NE

phospholipid VASCULAR + + SMOOTH + MUSCLE PLC PLA2 CELL DAG AA CYP Contraction IP3 20-HETE

+ + (-)

2+ Ca K+ - (-) + + - Em 2+ (-) (+) L-type Ca KCa channel (+) Ca2+ K+ EETs ENDOTHELIAL CELL

PLA2 CYP AA EETs

Figure 1.6 Opposing effects of EETs and 20-HETE on vascular tone EETs and 20-HETE induce opposing effects on vascular tone via functionally antagonistic effects on large-conductance calcium-activated potassium channels (KCa) in vascular smooth muscle cells. EET- and 20-HETE-mediated effects are indicated by red and black signs, respectively. 20-HETE produced by CYP Z-hydroxylases in vascular smooth muscle cells inhibits KCa channels, which leads to decreased 2+ 2+ membrane potential (Em), activation of L-type Ca channels, increased Ca influx and vascular contraction. In contrast EETs, produced by CYP epoxygenases in endothelial cells, activate KCa channels; this leads to smooth muscle hyperpolarisation, inhibition of L-type Ca2+ channels, decreased Ca 2+ influx and vascular relaxation. Ang,,, angiotensin ,,; NE, norepinephrine; AA, arachidonic acid; PLC, phospholipase C; PLA2, phospholipase A2; IP3, inositol 1,4,5-trisphosphate; DAG, diacylglycerol.

1.5.2.2 EETs Proposed to be the Endothelium-derived Hyperpolarising Factor (EDHF)

Apart from the established vasodilators NO and prostacyclin (PGI2), the endothelium releases another substance, termed the EDHF, that mediates vasodilation (Fisslthaler et al., 1999; Fleming, 2001). There is substantial evidence to suggest that EETs may be the elusive EDHF (Fleming, 2004). The EDHF is actually likely to be several different factors or substances,

because the pharmacological properties of NO/ PGI2-independent endothelium- dependent hyperpolarisation are distinct in different arteries and different species (Fisslthaler et al., 1999; Fleming, 2001).

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While there are several likely EDHFs in different vascular beds and species, there is now considerable evidence that CYP-derived EETs serve as the EDHF in renal and coronary arteries in a number of different species including humans, pigs, rats, rabbits, cows and dogs (Fisslthaler et al., 1999; Fleming, 2004). EETs have the important properties of an EDHF in that they are produced within the endothelium, are potent vasodilators, and mediate vasodilation by hyperpolarising VSM via the activation of KCa channels (Campbell et al., 1996). A proposed role for CYP-derived EETs in mediating the EDHF response was based on early studies showing that CYP inhibitors significantly attenuate the NO/

PGI2-independent hyperpolarisation and relaxation in a number of different vascular preparations (Fleming, 2001). However, the results of these studies were somewhat inconclusive in that the CYP inhibitors were capable of directly affecting the activity of

KCa channels, the proposed cellular target of the EDHF (Alvarez et al., 1992). More selective CYP epoxygenase inhibitors have since been developed, and these specifically attenuate the NO/ PGI2-independent vasodilation of renal arterioles (Imig et al., 2001). The hypothesis that EETs serve as EDHF is further supported by recent studies employing techniques other than the pharmacological inhibition of CYPs. Treatment of porcine coronary artery endothelial cells with the CYP inducers E-naphthoflavone and nifedipine increased CYP2C expression and EET synthesis, and enhanced EDHF activity (Fisslthaler et al., 1999; Fisslthaler et al., 2000). Antisense oligonucleotides directed against human CYP2C8/9 decreased expression of the porcine homologue CYP2C34 in porcine coronary arterial endothelial cells, and also attenuated EDHF activity (Fisslthaler et al., 1999). CYP activity is inhibited by NO, which may explain why EDHF-mediated responses are difficult to detect in the absence of NO synthase inhibition (Fleming, 2001). As healthy endothelial cells produce physiological concentrations of NO that inhibit intrinsic CYP activity, the EET/EDHF pathway may be only of minor importance in normal healthy vessels, but may serve as an important backup vasodilatory mechanism in pathophysiological conditions associated with endothelial dysfunction, in which NO synthesis is impaired (Bauersachs et al., 1996; Fleming, 2001).

1.5.2.3 Role of EETs in Reactive Hyperemia The regulation of cerebral vascular tone is largely dependent on the pressure-induced generation of the vasoconstrictor 20-HETE within cerebral VSM cells, but EETs are also produced within the brain and affect the tone of cerebral arteries (Fleming, 2001). Cerebral EETs are produced by astrocytes and may play a role in reactive or functional hyperemia, the process in which cerebral blood flow is recruited or shunted

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to areas of increased neuronal activity (Alkayed et al., 1997; Fleming, 2001; Roman, 2002). Neuronal function requires oxidative metabolism and reactive hyperemia ensures that active neurons have enhanced cerebral blood flow in order to provide a sufficient level of oxygen (Roman, 2002). The recent demonstration that cultured rat astrocytes express high levels of CYP2C11 and generate EETs from AA, together with the demonstration that EET production by astrocytes is stimulated by the excitatory neurotransmitter glutamate, has led to the hypothesis that astrocyte-derived EETs play a role in reactive hyperemia (Alkayed et al., 1997; Harder et al., 1998). In this model, the astrocytes are stimulated by glutamate released from adjacent active neurons which increases intracellular Ca2+ levels, and stimulates CYP-mediated synthesis and the release of EETs. The EETs diffuse to cerebral VSM cells where they activate KCa channels, elicit vasodilation of cerebral arteries, and increase blood flow to the active areas of the brain (Harder et al., 1998).

1.5.3 Non-vasodilatory Effects of EETs within the Vascular System 1.5.3.1 Anti-inflammatory Effects of EETs EETs, especially the 11,12-EET enantiomer, have been demonstrated to possess potent anti-inflammatory effects in endothelial cells (Node et al., 1999). Endothelial cells play a key role in the initiation of the vascular inflammatory process and in the pathogenesis of atherosclerosis (Ross, 1999; Campbell, 2000). Upon stimulation with the pro-inflammatory macrophage- and leukocyte-derived cytokines tumour necrosis factor-Į (TNF- Į) and interleukin-1Į (IL-1Į), endothelial cells express vascular cell adhesion molecule 1 (VCAM-1), intercellular adhesion molecule 1 (ICAM-1) and E-selectin (Campbell, 2000). These proteins facilitate the recruitment and adhesion of inflammatory cells such as mononuclear cells (Node et al., 1999). The excessive adhesion of inflammatory cells to the vascular wall narrows the vascular lumen and restricts blood flow, and represents a critical stage in the development of atherosclerosis (Ross, 1999). Node et al. (1999) has demonstrated that low, physiologically relevant (nanomolar), concentrations of EETs inhibit or prevent the cytokine-induced expression of cell adhesion molecules in cultured human endothelial cells. Pretreatment of human endothelial cells with nanomolar concentrations of 11,12-EET, 8,9-EET and 5,6-EET (but not 14,15-EET) attenuated TNF-Į-induced VCAM-1 expression. Further to these in vitro experiments, Node et al. (1999) demonstrated anti-inflammatory effects of EETs in an in vivo model of TNF-Į-induced cellular adhesion: VCAM-1 expression and

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mononuclear cell adhesion to the mouse carotid artery was inhibited by intra-arterial infusion of 11,12-EET. The anti-inflammatory effects of EETs are independent of their membrane-

hyperpolarising effects, as two different selective KCa channel inhibitors prevented EET- induced vasodilation, but did not affect 11,12-EET-mediated inhibition of cytokine- induced VCAM-1 expression (Node et al., 1999). Rather, EETs exerted their anti- inflammatory effect by inhibiting activation of the pro-inflammatory transcription factor nuclear factor NB (NF-NB) which up-regulates VCAM-1, ICAM-1 and E-selectin expression (May & Ghosh, 1998; Node et al., 1999; Campbell, 2000). The cytokine- induced activation of NF-NB involves the phosphorylation, and subsequent degradation, of inhibitory factor NB (INB), the cytosolic inhibitor of the NF-NB transcription factor (May & Ghosh, 1998; Campbell, 2000). 11,12-EET inhibits the activity of the INB kinase that phosphorylates INB (Figure 1.7; Node et al., 1999).

phospholipid

TNF-D (+) PLA2 CYP IL-1D AA EETs (-)

P P Degradation INB

VCAM-1 VCAM-1, NF-NB IKK INB ICAM-1 ICAM-1, E-selectin E-selectin mRNA NF-NB ADP ATP

ENDOTHELIAL CELL

Figure 1.7 Anti-inflammatory effects of EETs via inhibition of cytokine-induced NF-NB activation Upon stimulation with pro-inflammatory cytokines such as tumour necrosis factor-D (TNF-D) and interleukin-1D (IL-1D), inhibitory factor NB kinase (IKK) is activated and phosphorylates inhibitory factor NB (INB). Phosphorylation of INB stimulates its ubiquitination and degradation by the 26S proteosome, and enables the unbound nuclear factor NB (NF-NB) to translocate into the nucleus and activate transcription of pro-inflammatory genes such as those encoding the cell adhesion molecules vascular cell adhesion molecule 1 (VCAM-1), intercellular adhesion molecule 1 (ICAM-1) and E-selectin. EETs prevent NF-NB activation and the expression of pro-inflammatory genes by inhibiting cytokine-induced activation of IKK. AA, arachidonic acid; PLA2, phospholipase A2; ATP, adenosine 5’-triphosphate; ADP, adenosine 5’-diphosphate.

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Further support for EETs as anti-inflammatory mediators comes from the demonstration that 11,12-EET attenuates the pyretic response to lipopolysaccharide (LPS) injection (Kozak et al., 2000) and interleukin-1E (IL-1E; Nakashima et al., 2001). While further studies are clearly required, it has been suggested that CYP- derived EETs may prove useful in treating vascular inflammatory disorders such as atherosclerosis, and possibly also non-vascular inflammatory diseases (Node et al., 1999).

1.5.3.2 Anti-migratory Effects of EETs Proliferation and migration of VSM cells is a key feature of atherosclerosis and vascular proliferative diseases (Spiecker & Liao, 2005). Sun et al. (2002) demonstrated that 11,12-EET significantly inhibited the migration of rat aortic smooth muscle cells in response to platelet-derived growth factor (PDGF) and fetal calf serum (FCS). 14,15- EET and 5,6-EET also exhibited significant, but smaller, inhibitory effects on PDGF- stimulated smooth muscle migration, whereas 8,9-EET was ineffective (Sun et al., 2002). The anti-migratory effects of 11,12-EET were independent of its membrane hyperpolarisation properties, as treatment of cells with a KCa channel blocker did not prevent the modulation of cell migration by EETs (Sun et al., 2002). Rather, EETs may inhibit smooth muscle cell migration by activating the cAMP/PKA signalling pathway that participates in the regulation of smooth muscle cell migration (Itoh et al., 2001; Sun et al., 2002). As the migration of VSM cells into the intima of vessels is a key stage in the development of numerous vascular proliferative diseases, it is suggested that 11,12- EET may prove to be of therapeutic benefit in reducing the risk of vascular disease by preventing VSM cell migration (Sun et al., 2002).

1.5.3.3 Fibrinolytic Effects of EETs A key feature of acute coronary syndromes is the rupture of atherosclerotic plaques with ensuing vascular thrombosis and occlusion (Libby, 1995). The process of vascular thrombosis is regulated, in part, by the enzyme tissue- type plasminogen activator (t-PA), and its inhibitor, PAI-1 (Spiecker & Liao, 2005). EETs, in particular 11,12-EET, have fibrinolytic properties that are mediated by increasing the expression and activity of t-PA protein in vascular endothelial cells; expression of the t-PA inhibitor PAI-1 is unchanged (Node et al., 2001). The fibrinolytic effects of EETs are not related to their membrane hyperpolarisation effects. Treatment of cells with a selective KCa channel blocker did

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not affect the 11,12-EET-mediated increase in t-PA expression. Rather, the increase in t-PA protein levels by 11-12-EET was due to increased activation of the cAMP- responsive t-PA gene promoter. Thus, it was shown that treatment of endothelial cells with 11,12-EET increased intracellular cAMP levels and the activity of the t-PA promoter. The mechanism by which EETs increase cAMP levels was suggested to be mediated by the activation of the guanine nucleotide-binding protein GĮs; EETs were found to increase GĮs activity, and activation of GĮs stimulates the adenylyl cyclase/PKA/cAMP pathway and increases the activity of cAMP-dependent gene promoters (Node et al., 2001). The demonstration that EETs mediate the activation of the fibrinolytic mediator t-PA suggests that these eicosanoids may play a role the regulation of fibrinolysis in the vascular system. In particular, EETs may lower the risks of cardiovascular thrombosis and occlusion by increasing t-PA levels and enhancing fibrinolytic activity within the vasculature (Node et al., 2001).

1.5.3.4 Mitogenic Properties of EETs EETs are potent mitogens. EETs, especially the 11,12-EET and 14,15-EET enantiomers, increase growth factor-stimulated proliferation of a range of cell types, including vascular endothelial and smooth muscle cells (Munzenmaier & Harder, 2000; Fleming et al., 2001a), and renal epithelial and mesangial cells (Harris et al., 1990; Chen et al., 1998). Although the precise mechanism by which EETs mediate cellular proliferation remains to be elucidated, EETs activate intracellular protein kinases that regulate cellular proliferation, including tyrosine kinases and the MAP kinase family. In particular, EETs activate the extracellular-regulated kinases 1 and 2 (ERK1/2), which are MAP kinases that mediate the proliferation of vascular cells (Fleming et al., 2001a), and the mitogenic effects of 14,15-EET in epithelial cells appear to be mediated by activation of the Src family of tyrosine kinases (Chen et al., 1998). The demonstration that EETs are potent vascular mitogens has led to the suggestion that they may play a role in angiogenesis. In support of this suggestion are the results of Munzenmaier & Harder (2000), showing that the co-culture of EET- producing astrocytes with brain microvascular endothelial cells increased cell growth and the formation of endothelial tubes that are precursors of capillaries. Thus, EETs may influence capillary density and the long term regulation of cerebral blood flow. Furthermore, this study demonstrated that CYP epoxygenase inhibitors prevented the promotion of endothelial tube formation by astrocytes and provided the first direct evidence for a link between EETs and angiogenesis (Munzenmaier & Harder, 2000).

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Other investigators have since provided further evidence of a role for CYP-derived EETs in the promotion of angiogenesis (Medhora et al., 2003).

1.5.3.5 Effects of EETs on Platelets The activation of platelets with thrombin and platelet activating factor stimulates the release of EETs, which inhibit platelet aggregation in response to vascular injury and AA (Fitzpatrick et al., 1986; Roman, 2002). The structural relationship of AA to EETs leads to potential competition for eicosanoid biotransformation enzymes. Thromboxane is the principal product of AA metabolism by COX in platelets, and stimulates platelet activation (Fitzpatrick & Murphy, 1988). 8,9-EET and 14R,15S-EET inhibit thromboxane synthesis and therefore platelet aggregation, by competing with AA for metabolism by COX (Fitzpatrick et al., 1986; Roman, 2002).

1.5.3.6 Role of EETs in Protection Against Hypoxia-reoxygenation Injury in Endothelial Cells Hypoxia occurs when the blood supply to a tissue or organ is decreased. Hypoxia, and the subsequent reoxygenation, is a key feature of ischaemic injury that occurs in a number of pathophysiological conditions such as atherosclerosis, ischaemic heart disease, stroke, cancer, chronic lung disease, organ transplantation and chemical- mediated cellular damage (Fan et al., 1999; Semenza, 1999). The role of vascular endothelial cells in cardiovascular physiology and pathophysiology is well established, and endothelial dysfunction arising from hypoxia- reoxygenation is a critical event in the pathophysiology of nearly all types of ischaemia- reperfusion injury (Lefer & Lefer, 1993). Endothelial dysfunction is associated with an increase in the generation of oxygen-derived free radicals, such as hydrogen peroxide and superoxide anions, that occurs within minutes of reperfusion, and is characterised by a decrease in the bioavailability of NO, most likely due to the destruction of NO by reactive oxygen species (Lefer & Lefer, 1993; Fleming, 2004). The production of NO is one of the most important functions of endothelial cells; NO, like EETs, has important vasodilatory, anti-thrombotic and anti-inflammatory effects that play a key role in maintaining vascular homeostasis (Yang et al., 2001). As such, the maintenance or improvement of endothelial function following myocardial ischaemia-reperfusion injury has been shown to improve cardiac function (Yang et al., 2001), and the therapeutic nitrate drugs, which are a source of NO to the vasculature, are commonly used for the treatment of ischaemic heart disease (Abrams, 1996).

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In addition to endothelial dysfunction and reduced NO bioavailability, ischaemia-reperfusion is associated with infiltration of leukocytes and platelets, as well as reactive oxygen species-mediated lipid peroxidation, which may also contribute to vascular injury (Spiecker & Liao, 2005). EETs have been reported recently to protect endothelial cells against hypoxia- reoxygenation injury. Yang et al. (2001) showed that pretreatment of bovine aortic endothelial cells with 11,12-EET significantly reduced cell death associated with hypoxia and reoxygenation. 14,15-EET and 11,12-DHET were also effective, although to a lesser extent than 11,12-EET. Protection against hypoxia and reoxygenation by EETs was suggested to be mediated via their anti-inflammatory and anti-oxidative properties (Yang et al., 2001). An association between EETs and vascular ischaemic injury had also been reported previously by Rosolowsky et al. (1990), in a study using a canine model of concentric coronary artery stenosis. This model is characterised by endothelial damage, vasoconstriction, infiltration of leukocytes into the media of the coronary vessel, and adherence of platelets to the vessel wall. Vasoconstriction and the accumulation of platelets contribute to spontaneous decreases and increases in coronary blood flow (known as cyclic flow variation). It was demonstrated that stenosed coronary vessels produced higher levels of EETs than normal vessels, and that EETs induced relaxation of coronary artery rings. Thus, higher levels of EETs may be produced within the vasculature under ischaemic conditions, and EETs may influence the development of vascular disease (Rosolowsky et al., 1990).

In summary, similar to the endogenous vasodilator NO, EETs mediate a number of important non-vasodilatory protective effects within the vascular wall and are proposed to play an important role in vascular homeostasis and vascular disease (Node et al., 1999; Node et al., 2001; Sun et al., 2002).

1.5.4 Non-vascular Effects of EETs EETs are produced in many tissues outside the vasculature. In particular, EETs are synthesised within the heart, liver, kidney, lungs and pancreas, and are known to possess potent biological activities within these tissues (Table 1.3). Some of the important biological activities of EETs outside the vascular system are discussed below.

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1.5.4.1 Effects of EETs on Cardiomyocyte Function and Recovery After Cardiac Ischaemia Substantial amounts of EETs are synthesised within human and rat cardiac myocytes by the CYP2J subfamily (Wu et al., 1996; Wu et al., 1997), and significant quantities of EETs, and their hydrated products, DHETs, are incorporated into the membrane phospholipids of cardiac myocytes (Lee et al., 1999; Roman, 2002). In response to hypoxia, and agonists that activate phospholipases, incorporated EETs are released and influence cardiac function (Roman, 2002). In particular, EETs formed within cardiac myocytes affect intracellular Ca2+ concentration, activity of ion channels, the contractile function of the heart, and modulate the recovery of the heart during ischaemia (Roman, 2002). 8,9-EET has been shown to induce a hyperpolarised shift in the steady-state membrane potential of cardiomyocytes by inhibition of cardiac Na+ channels (Lee et al., 1999). In general, inhibition of Na+ channels offers cardioprotection during ischaemia- reperfusion (Seubert et al., 2004). EETs also affect the activity of cardiac L-type Ca2+ channels, which play an important role in regulating the strength and rate of cardiac contractions. In particular, the influx of Ca2+through L-type Ca 2+ channels stimulates the release of Ca 2+ from the sarcoplasmic reticulum, which initiates and regulates the force of cardiomuscular contraction (Chen et al., 1999). EETs inhibit the opening of cardiac L-type Ca2+ channels, an effect partially mediated by an increase in the rate of channel inactivation (Chen et al., 1999). Because inhibition of cardiac L-type Ca2+ channels reduces the force and rate of contraction of cardiac tissue, it has been suggested that the release of EETs during cardiac ischaemia would reduce contractile force and decrease oxygen utilisation, and thus may provide a protective effect. Protection may also be achieved by preventing the development of dangerous arrhythmias associated with activation of Ca2+ channels (Chen et al., 1999). Consistent with this hypothesis is the study by Wu et al. (1997), which demonstrated that 11,12-EET significantly improved the functional recovery of the heart following prolonged global cardiac ischaemia in an isolated-perfused rat heart model. By contrast, other investigators have reported opposite effects of EETs on cardiac L-type Ca2+ channels and cardiac recovery following an ischaemic event. Xiao et al. (1998) reported that 11,12-EET stimulates cardiac L-type Ca2+ channel activity and increases Ca2+ current in rat ventricular myocytes, and that CYP inhibitors suppress the Ca2+ current through these channels. Similarly, Moffat et al. (1993) found that 5,6-EET and 11,12-EET increase intracellular Ca2+ concentration and contraction in guinea pig cardiac myocytes. This study reported that EETs caused a delay in the

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recovery of myocardial function in guinea pig hearts following ischaemia and reperfusion (Moffat et al., 1993). The discrepancies in the reported effect of EETs on cardiac L-type channels and recovery following an ischaemic event may be due to species differences, variation in the severity of ischaemia, and differences in the cellular metabolic state (Wu et al., 1997; Chen et al., 1999). In summary, ischaemia stimulates the release of EETs from cardiac myocytes. Current data suggests that the released EETs affect cardiac Na+2+ and Ca channels, which subsequently affect contractile force and heart rate during ischaemia. Additional studies are required to establish whether EETs improve or diminish the recovery of cardiac function following ischaemia (Roman, 2002).

1.5.4.2 Effect of EETs within the Kidney and Potential Role in Hypertension EETs are endogenously produced within the human and rat kidney and regulate renal function (Zeldin, 2001). While CYPs from several subfamilies are able to catalyse the formation of EETs, the CYP2C subfamily are likely to represent the major renal CYP epoxygenases (Zeldin et al., 1995a); the CYP2J subfamily may also have a role (Wu et al., 1996; Yu et al., 2000). As discussed previously, EETs have potent effects on vascular tone (section 1.5.2.1). As in most vascular systems, EETs are generally vasodilatory within the kidney. Specifically, 11,12-EET, 14,15-EET and 8,9-EET are vasodilatory within the renal circulation, but 5,6-EET is a renal vasoconstrictor (Imig et al., 1996a). This effect of 5,6-EET is blocked by inhibitors of COX, suggesting that vasoconstriction is dependent on further metabolism by COX enzymes (Imig et al., 1996a). EETs also have mitogenic effects within the kidney, and may be intracellular mediators for growth factors and other vasoactive compounds (Roman & Alonso- Galicia, 1999). In particular, EETs stimulate cellular proliferation in renal glomerular mesangial cells (Harris et al., 1990) and renal epithelial cells (Chen et al., 1998). Apart from their effects on renal vascular tone and cellular proliferation, EETs regulate renal ion transport and modulate fluid and electrolyte excretion (Roman, 2002). EETs activate the Na+/H+ exchanger in cultured rat glomerular mesangial cells, which implicates EETs in the regulation of glomerular filtration rate (Harris et al., 1990; Roman et al., 2000). EETs inhibit Na+ transport within the proximal tubule by inhibiting the translocation of the Na+/H+ exchanger to the apical membrane (Roman, 2002), and may be second messengers in the natriuretic actions of angiotensin ,, within the proximal tubule (Roman, 2002). EETs also regulate ion transport within the collecting duct, where they inhibit Na+ reabsorption and K+ secretion (Ma et al., 1999). Thus,

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EETs mediate anti-hypertensive effects by inhibition of Na++ transport, increasing Na and water excretion, and lowering plasma volume and blood pressure (Roman, 2002). In view of their effects on renal vascular tone, and fluid and electrolyte transport, it has been suggested that EETs may play a role in the pathogenesis of hypertension (Roman, 2002). While the majority of studies have focused primarily on the role of 20-HETE in hypertension (section 1.4.5), others have found that renal EET production is altered in certain animal models of hypertension. For example, it has been demonstrated that excess dietary salt increases EET production in some rat strains, and that rat renal epoxygenases are induced by high dietary salt (Makita et al., 1994). Because EETs inhibit renal sodium reabsorption, the up-regulation of renal epoxygenases and increased EET production may be an efficient adaptive mechanism to prevent salt retention and hypertension in response to increased salt intake. In support of this contention are experiments in Sprague-Dawley rats that demonstrated that inhibition of the salt-induced up-regulation of renal epoxygenases results in the development of salt-dependent hypertension (Makita et al., 1994). Also supportive of an anti-hypertensive role for EETs are studies in the Dahl salt-sensitive rat model which suggest that the development of hypertension in DS rats may be associated with the inability of this strain to increase renal EET production. In particular, EET production increased in normotensive DR rats in response to elevations in salt intake, whereas the DS rats remained unresponsive to high salt (Makita et al., 1994). In contrast to these findings, other investigators have reported an association between increased EET production and the development of hypertension. For example, the urinary excretion of EETs is higher in the developmental phase of hypertension in the SHR compared to normotensive WKY controls. Increased expression of a renal CYP2J isoform in SHRs could be responsible for the increased production of EETs (Yu et al., 2000). Similarly, in women with pregnancy-induced hypertension, the urinary excretion of epoxygenase metabolites is significantly increased (Catella et al., 1990). While these observations may indicate that increased EET formation contributes to the development of hypertension, it is also possible that increased EET formation in these systems may be a consequence, rather than a cause, of hypertension. Given that the EETs are largely anti-hypertensive (they induce vasodilation and inhibit Na+ reabsorption), the increase in EET production in SHR and pregnancy-induced hypertension may be a compensatory mechanism against increased blood pressure (Yu et al., 2000). This suggestion is supported by the study of Pomposiello et al. (2001), which reported that renal EET-mediated vasodilation was increased in the SHRs.

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Thus, in summary, production of EETs is clearly altered in hypertension, but further research is now required to firmly establish the role of EETs in the pathogenesis of hypertension.

1.5.4.3 Anti-apoptotic Effects of EETs Apoptosis, or programmed cell death, plays a central role in the pathogenesis of diseases and conditions including cancer and heart disease (Kroetz & Zeldin, 2002). Recent experimental evidence suggests that EETs, especially 14,15-EET, may inhibit apoptosis. In a study by Chen et al. (2001) using a renal proximal tubule-like epithelial cell line, the exogenous application of 14,15-EET inhibited apoptosis induced by a diverse range of stimuli, including serum withdrawal, etoposide, hydrogen peroxide, and excess AA. Stable transfection of a bacterial CYP AA epoxygenase that generated 14,15-EET also inhibited apoptosis in response to pro-apoptotic stimuli (Chen et al., 2001). The mechanism by which 14,15-EET exerts its anti-apoptotic effects could involve activation of the phosphatidylinositol-3 kinase-Akt signalling pathway (Chen et al., 2001). Phosphatidylinositol-3 kinase is activated by a wide range of stimuli such as hormones, cytokines and growth factors, and has a role in growth factor-mediated cell survival and the prevention of apoptosis in different types of cells (Toker & Cantley, 1997; Chen et al., 2001). The downstream target of phosphatidylinositol-3 kinase in the growth factor-mediated cytoprotective signalling pathway is the serine/threonine protein kinase B called Akt (Toker & Cantley, 1997). It was demonstrated that 14,15-EET significantly activates both phosphatidylinositol-3 kinase and Akt, and that the anti- apoptotic effects of 14,15-EET were prevented with the use of specific inhibitors of phosphatidylinositol-3 kinase. Thus, it appears that 14,15 EET inhibits apoptosis by activation of the anti-apoptotic phosphatidylinositol-3 kinase-Akt signalling pathway (Chen et al., 2001).

1.5.4.4 Effects of EETs on the Release of Peptide Hormones EETs are produced within mammalian endocrine tissues and play a role in mediating the release of certain peptide hormones. For example, EETs are produced within the pituitary gland and potentiate the release of growth hormone, vasopressin, oxytocin, somatostatin and luteinizing hormone from the pituitary by stimulating Ca2+ entry and release (Capdevila et al., 1983; Negro-Vilar et al., 1985; Snyder et al., 1989; Roman, 2002). EETs are also produced endogenously by the human and rat pancreas (Zeldin et al., 1997a), and stimulate the release of insulin and glucagon from isolated islet

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cells, leading to the suggestion that they may modulate glucose homeostasis (Falck et al., 1983; Zeldin et al., 1997a). The effects of EETs on insulin and glucagon release occurred at low concentrations and were regioselective; 5,6-EET stimulated insulin secretion, but did not affect the release of glucagon, while 8,9-EET, 11,12-EET and 14,15-EET stimulated glucagon secretion but did not affect the secretion of insulin (Falck et al., 1983). The mechanism by which EETs alter insulin and glucagon secretion from islets is unclear, but could be related to increased intracellular Ca2+ concentration, which has a role in the secretion of insulin and glucose (Roman, 2002). Because EETs have the capacity to increase intracellular Ca2+ levels in many different cell types, EET-stimulated insulin and glucagon release may occur through a Ca2+- dependent mechanism (Zeldin et al., 1997a; Roman, 2002).

1.5.4.5 Effects of EETs in the Lung The COX and lipoxygenase metabolites of AA have long been recognised to play an important role in pulmonary function (Roman, 2002). COX metabolites have various well established pulmonary effects. For example, prostaglandin E2 (PGE2), which is produced within the airway epithelium, is a potent bronchodilator and has anti- inflammatory properties. PGI2, a major product of pulmonary arterial endothelial cells, is a pulmonary vasodilator and bronchodilator and inhibits platelet aggregation, while prostaglandin F2Į (PGF2Į) and thromboxane are pulmonary vasoconstrictors and bronchoconstrictors and enhance platelet aggregation (Jacobs & Zeldin, 2001). Leukotrienes, the products of the action of lipoxygenase on AA, play a significant role in the increased mucous secretion, airway inflammation, bronchoconstriction and vascular permeability seen in asthma (Jacobs & Zeldin, 2001). While less is known about the role of CYP metabolites of AA within the lung, increasing evidence suggests that 20-HETE and the EETs may also have an important role in critical pulmonary functions (Jacobs & Zeldin, 2001). Several CYP epoxygenases are expressed within different cell types throughout the lung in various species (Zeldin et al., 1996a; Zhu et al., 2000b; Hukkanen et al., 2002). Moreover, microsomal fractions from rabbit and guinea pig lung convert AA to EETs (Knickle & Bend, 1994; Zeldin et al., 1995b), and substantial amounts of EETs have also been detected in human and rat lung tissue (Zeldin et al., 1996a). A number of investigators have reported various effects of EETs within the lung. For example, EETs have been found to affect pulmonary vascular tone. As outlined in section 1.5.2.1, EETs are generally vasodilatory within renal, cerebral, coronary and other systemic arteries, and consistent with these observations, 5,6-EET stimulated the vasodilation of isolated perfused rabbit lungs that had been preconstricted with a

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thromboxane mimetic (Tan et al., 1997). On the other hand, Zhu et al. (2000b) has reported that all regioisomers of EETs effect vasoconstriction in isolated pressurised rabbit pulmonary arteries. Thus, EETs may regulate pulmonary vascular tone, although the nature of these effects of EETs may be dependent on factors such as species and the preexisting vascular tension (Jacobs & Zeldin, 2001). EETs are involved in the regulation of ion transport within the lung. CYP epoxygenases are abundantly expressed within airway epithelial cells (Zeldin et al., 1996a), and EETs have been found to inhibit Ca2+-sensitive Cl - currents in tracheal epithelial cells. Thus, EETs decrease Cl- transport and fluid secretion by the airway epithelium, and may regulate the volume and composition of fluids lining the airways (Jacobs & Zeldin, 2001; Roman, 2002). Finally, EETs probably modulate airway tone. CYP epoxygenases are expressed within bronchial smooth muscle cells (Zeldin et al., 1996a), and EETs regulate the membrane potential of bronchial smooth muscle cells. For example, 5,6-EET and 11,12-EET activate airway smooth muscle large-conductance Ca2+- activated K+ channels and hyperpolarise and relax airway smooth muscle cells (Dumoulin et al., 1998). Furthermore, 5,6-EET and 8,9-EET relaxed guinea pig bronchi that had been preconstricted with histamine (Zeldin et al., 1995b). In summary, EETs appear to play a role in the regulation of pulmonary vascular tone, bronchial smooth muscle tone, and airway epithelial ion transport, and may be important mediators of pulmonary function under both normal physiological conditions, as well as pathophysiological conditions (Jacobs & Zeldin, 2001).

1.5.4.6 Effects of EETs in the Liver The liver has the highest total content of CYPs compared with other organs (Sacerdoti et al., 2003) and EETs are endogenously produced by hepatic CYP epoxygenases. Indeed, EETs are the principal products of AA metabolism in rat (Capdevila et al., 1990) and human liver microsomes (Zeldin et al., 1996b). CYP2C8 appears to be the primary hepatic CYP responsible for the production of EETs, although other family 2 CYPs, such as CYP2C9 and CYP2J2, are also expressed in the liver and may contribute to hepatic EET formation (Rifkind et al., 1995; Wu et al., 1996; Zeldin et al., 1996b; Enayetallah et al., 2004). There is limited information on the role on EETs within the liver. For example, Yoshida et al. (1990) demonstrated that EETs activate phosphorylase a and increase intracellular Ca2+ concentration in isolated rat hepatocytes. That study reported that inhibition of EET formation prevented the increase in intracellular Ca2+ concentration in vasopressin-stimulated hepatocytes, and proposed that EETs may influence

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vasopressin-induced glycogenolysis in the liver via activation of phosphorylase a and stimulation of increased intracellular Ca2+ (Yoshida et al., 1990). Similarly, Kutsky et al. (1983) reported that EETs modulate calcium metabolism in rat liver microsomes. Despite limited information on the role of EETs within the liver, their importance in other tissues suggests that they may regulate hepatic homeostasis (Zeldin et al., 1996b; Sacerdoti et al., 2003). Indeed, they may regulate hepatic vascular tone and the action of peptide hormones within the liver (Zeldin et al., 1996b). Furthermore, the regiochemistry of EETs in human plasma is similar to that in liver, which implies that large quantities of EETs produced by the liver could be secreted into the circulation (Zeldin et al., 1996b). Finally, in a rat model of liver cirrhosis, the hepatic production of EETs is decreased (Sacerdoti et al., 2003). Although the significance of this finding is uncertain, it may be indicative of a role for EETs in the pathophysiology of hepatic diseases and conditions (Sacerdoti et al., 2003).

1.5.5 Factors Affecting the Level of EETs within the Body The total amount of EETs within a given tissue is influenced by a number of factors including EET biosynthesis by CYP epoxygenases, EET storage via esterification to membrane phospholipids, and EET metabolism by enzymes such as epoxide hydrolases and phase ,, enzymes such as the glutathione S-transferases (Qu et al., 1998). The biosynthesis of EETs by CYP epoxygenases is altered by factors that modulate expression and/or activity of CYPs, including exposure to xenobiotics (Fisslthaler et al., 1999; Fisslthaler et al., 2000), nutritional factors such as fasting and salt intake (Makita et al., 1994; Qu et al., 1998), hormonal stimuli (Roman, 2002), physiological conditions such as pregnancy (Catella et al., 1990), and pathophysiological conditions such as hypertension (Yu et al., 2000). EET biosynthesis may also be affected by genetic polymorphisms of CYPs (Fleming, 2001; Jacobs & Zeldin, 2001). CYP genes are highly polymorphic which can predispose to altered drug and chemical elimination in certain individuals (Ingelman- Sundberg, 2001). There is some information about polymorphisms of CYP2C and CYP2J that oxidise AA to EETs. Polymorphisms in these genes have been reported within the coding and/or promoter regions and may affect EET synthesis (Dai et al., 2001; King et al., 2002). For example, the CYP2C8*3 variant has a decreased capacity to synthesise EETs (Dai et al., 2001). It is conceivable that polymorphisms in CYP epoxygenase genes may be associated with increased susceptibility to certain diseases and conditions (Fleming, 2001; Jacobs & Zeldin, 2001; Zeldin, 2001).

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Because EETs are important in normal cellular physiology, and altered EET production may contribute to the pathophysiology of certain diseases, information on the tissue-specificity and regulation of CYP epoxygenases is important (Zeldin, 2001; Kroetz & Zeldin, 2002). The work detailed in this thesis identified some of the transcriptional mechanisms that regulate expression of the important human CYP epoxygenase CYP2J2.

1.6 Cytochrome P450 2J2 (CYP2J2)

The recently described CYP2J subfamily comprises a number of enzymes with a wide tissue distribution that catalyse AA oxidation (Table 1.4; Ma et al., 1998; Scarborough et al., 1999). Human CYP2J2 is a recently identified member of this subfamily that has been demonstrated to actively catalyse the formation of EETs from AA (Wu et al., 1996). CYP2J2 is the only member of the human CYP2J subfamily (Wu et al., 1996; King et al., 2002), whereas CYP2J enzymes in other species include rabbit CYP2J1 (Ichihara et al., 1985; Kikuta et al., 1991), rat CYP2J3 (Wu et al., 1997), CYP2J4 (Zhang et al., 1997) and CYP2J10 (Scarborough et al., 1999), and mouse CYP2J5 (Ma et al., 1999), CYP2J6 (Ma et al., 2002), CYP2J7 (Scarborough et al., 1999; Nelson et al., 2004), CYP2J8 (Scarborough et al., 1999; Nelson et al., 2004), CYP2J9 (Qu et al., 2001), CYP2J11 (Nelson et al., 2004), CYP2J12 (Nelson et al., 2004) and CYP2J13 (Nelson et al., 2004).

1.6.1 CYP2J2 Gene and Protein Structure The entire CYP2J2 gene has been cloned and sequenced (King et al., 2002). As shown in Figure 1.8, the CYP2J2 gene is approximately 40.3 kilo bases (kb) in length, and like all previously characterised CYP2 genes, consists of nine exons and eight introns (King et al., 2002). The exons vary in length from 139 to 522 base pairs (bp), and exons 8 and 9 encode the putative haem-binding region (King et al., 2002). Introns (non-coding sequences of nucleotides) vary in length from 393 bp to 10.4 kb. The CYP2J2 gene contains an approximate 6000 bp 5’-flanking region and an approximate 1000 bp 3’-untranslated region (King et al., 2002). The CYP2J2 cDNA was found to be 1876 nucleotides in length and encodes a protein of 502 amino acids with a molecular mass of 57653 Daltons (Da; Wu et al.,

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1996). Comparison of the amino acid sequence of the CYP2J2 protein with that of other CYPs shows that CYP2J2 shares between 19 to 30% sequence identity with proteins belonging to the CYP1, CYP3, CYP4, CYP5 and CYP6 families, and between 40 to 46% identity with CYP2 family proteins (Wu et al., 1996). Comparison of the CYP2J2 amino acid sequence with other members of the CYP2J subfamily has found that CYP2J2 is 80% identical to rabbit CYP2J1 (Wu et al., 1996), 73% identical to rat CYP2J3 (Wu et al., 1997), 76% identical to rat CYP2J4 (Zhang et al., 1997), 69% identical to mouse CYP2J5 (Ma et al., 1999), 76% identical to mouse CYP2J6 (Scarborough et al., 1999; Ma et al., 2002) and 72% identical to mouse CYP2J9 (Qu et al., 2001).

a6000 10436 3625393 1647 1857 2727 3767 7138 a1000 5’ 1 2 3 4 5 6 7 8 9 3’ 220 163 150 161 177 142 188 139 522

Figure 1.8 Structure of the CYP2J2 gene The CYP2J2 gene is approximately 40.3 kb and contains nine exons and eight introns. Exons are represented as numbered grey boxes, and the length of each exon is given below in bp. Intron lengths (bp) are indicated in blue. The CYP2J2 gene contains an approximate 6000 bp 5’-flanking region and an approximate 1000 bp 3’-untranslated region, which are indicated in red and green, respectively.

The CYP2J2 gene has been mapped to the short arm of human chromosome 1 (Ma et al., 1998). Interestingly, although CYP genes are widely spread throughout the entire genome (Nelson et al., 1996), CYP2J2 appears to be closely linked to the CYP4B1 gene on chromosome 1, and CYP4A11, the human orthologue of mouse Cyp4a10, has also been mapped to chromosome 1 (Nelson et al., 1996; Ma et al., 1998). Similarly, the mouse Cyp2j5 and Cyp2j6 genes have been mapped to a region of chromosome 4 that is in close proximity to the mouse Cyp4a10 and Cyp4a12 genes (Nelson et al., 1996; Ma et al., 1998). Like CYP2J members, the CYP4A subfamily actively catalyses AA oxidation to biologically active eicosanoids (section 1.4). The close proximity of the CYP2J genes to the CYP4A genes in both human and mouse suggests that these genes may form part of a cassette of CYP genes that are involved in the generation of biologically active fatty acids (Ma et al., 1998).

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Table 1.4 CYP2J subfamily enzymes: substrates and tissue distribution

Enzyme Known Tissue Reference Substrates Distribution

Rabbit CYP2J1 benzphetamine small intestine Ichihara et al., 1985; Kikuta et al., 1991; aminopyrine Koike et al., 1997 N,N-dimethylaniline

Human CYP2J2 arachidonic acid heart Wu et al., 1996; Zeldin et al., 1996a; liver Zeldin et al., 1997a; Zeldin et al., 1997b; vasculature Node et al., 1999; Scarborough et al., astemizole lung 1999; Gu et al., 2000; Moran et al., 2000; ebastine GIT Fleming, 2001; Hashizume et al., 2002; diclofenac kidney Matsumoto et al., 2002; Matsumoto et al., bufuralol pancreas 2003; Enayetallah et al., 2004 pituitary fetal liver

Rat CYP2J3 arachidonic acid heart Zeldin et al., 1996a; Wu et al., 1997; benzphetamine liver Zeldin et al., 1997a; Zeldin et al., 1997b; lung Scarborough et al., 1999 GIT kidney pancreas

Rat CYP2J4 arachidonic acid small intestine Zhang et al., 1997; Zhang et al., 1998 testosterone liver progesterone olfactory mucosa retinal

Rat CYP2J10 unknown unknown Scarborough et al., 1999

Mouse CYP2J5 arachidonic acid kidney Ma et al., 1999; Scarborough et al., 1999 testosterone liver diclofenac bufuralol

Mouse CYP2J6 benzphetamine small intestine* Ma et al., 2002 liver* kidney* colon* brain* heart* lung*

Mouse CYP2J7 unknown unknown Scarborough et al., 1999; Nelson et al., 2004

Mouse CYP2J8 unknown unknown Scarborough et al., 1999; Nelson et al., 2004

Mouse CYP2J9 arachidonic acid brain Qu et al., 2001 linoleic acid kidney*

Mouse CYP2J11 unknown unknown Nelson et al., 2004

Mouse CYP2J12 unknown Unknown Nelson et al., 2004

Mouse CYP2J13 unknown Unknown Nelson et al., 2004

* Expression demonstrated at the level of mRNA rather than protein. GIT, gastrointestinal tract.

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1.6.2 Catalytic Activity of CYP2J2 Studies have established that CYP2J2 is primarily an AA epoxygenase: CYP2J2 recombinant protein catalyses the NADPH-dependent oxidation of AA to all four regioisomeric EETs as the main products, with 19-HETE a minor reaction product (Wu et al., 1996; Scarborough et al., 1999). CYP2J2-mediated EET synthesis is regioselective, with epoxidation occurring preferentially at the 14,15-olefinic bonds (37% of total EETs produced), and less often at the 8,9-, 5,6- and 11,12-bonds (24, 21 and 18% of total EETs produced, respectively; Table 1.5; Wu et al., 1996). Production of 14,15-EET by CYP2J2 was highly stereoselective, with the R,S enantiomer accounting for 76% of total 14,15-EET produced. On the other hand, epoxidation at the 8,9- and 11,12-olefinic bonds occurred in a non-stereoselective manner (Table 1.5). The stereoselectivity of CYP2J2-mediated 5,6-EET production could not be investigated due to the rapid hydration of 5,6-EET to 5,6-DHET and the G-lactone of 5,6-DHET (Wu et al., 1996). The regioselectivity and stereoselectivity of CYP2J2 is different from that of other known human AA epoxygenases including CYP2C8 and CYP2C9 (Daikh et al., 1994; Wu et al., 1996).

Table 1.5 Regioselectivity and stereoselectivity of CYP2J2-mediated metabolism of arachidonic acid into EETs

Regioisomer Proportion of Stereoselectivity Total EETs R,S enantiomer S,R enantiomer 5,6-EET 21% Not determined Not determined 8,9-EET 24% 47% 53% 11,12-EET 18% 49% 51% 14,15-EET 37% 76% 24%

Adapted from Wu et al., 1996 (Table 1, pg 3465).

In addition to AA oxidation, CYP2J2 also oxidises other unsaturated fatty acids, including linoleic acid and eicosapentaenoic acid (Table 1.4; Moran et al., 2000; Fleming, 2001). Metabolism of linoleic acid and eicosapentaenoic acid by CYP2J2 generates epoxyoctadecenoic acids and epoxyeicosaquatraenoic acids, both of which have been shown to possess biological activities (Moran et al., 2000; Fleming, 2001). While primarily a fatty acid epoxygenase, as shown in Table 1.4, CYP2J2 displays a limited capacity to metabolise certain xenobiotics, including astemizole (Matsumoto et al., 2002; Matsumoto et al., 2003), ebastine (Hashizume et al., 2002), diclofenac and bufuralol (Scarborough et al., 1999).

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Other members of the CYP2J subfamily actively metabolise AA, but the eicosanoids generated by these enzymes are distinct from those formed by CYP2J2. For example, rat CYP2J3 metabolises AA to 14,15-EET, 11,12-EET, 8,9-EET and 19-HETE and is therefore both an AA epoxygenase and Z-1 hydroxylase (Wu et al., 1997). Similarly, rat CYP2J4 generates both EETs and 19-HETE from AA (Zhang et al., 1997). Mouse CYP2J5 generates EETs and midchain HETEs, in particular 11-HETE and 15-HETE, as the principal products of AA metabolism (Ma et al., 1999), and mouse CYP2J9 is predominantly an AA Z-1 hydroxylase, generating 19-HETE as the principal reaction product, with EETs and other HETEs as more minor reaction products (Qu et al., 2001).

1.6.3 Tissue Distribution of CYP2J2 Most previously characterised mammalian CYPs are primarily expressed within the liver, and display a much lower level of expression in extrahepatic tissues (Guengerich, 1995; Scarborough et al., 1999). CYP2J2 differs from many mammalian CYPs in this regard: while exhibiting significant constitutive expression in the liver, CYP2J2 is highly and constitutively expressed in a wide range of extrahepatic tissues (Table 1.4; Wu et al., 1996; Enayetallah et al., 2004). In particular, CYP2J2 is highly expressed within the heart (Wu et al., 1996; Enayetallah et al., 2004). To date, CYP2J2 is the only isoform that has been found to be predominantly expressed in this tissue (Wu et al., 1996). Significant expression of CYP2J2 has also been demonstrated in the lung (Zeldin et al., 1996a), vasculature (Node et al., 1999; Enayetallah et al., 2004), gastrointestinal tract (Zeldin et al., 1997b; Enayetallah et al., 2004), kidney (Wu et al., 1996; Enayetallah et al., 2004), pancreas (Zeldin et al., 1997a; Enayetallah et al., 2004), adrenal and pituitary glands (Enayetallah et al., 2004), and also within the fetal liver (Gu et al., 2000). CYP2J2 expression within these tissues has often been localised to specific cell types. Within the heart, CYP2J2 is primarily expressed in cardiomyocytes and in coronary arterial endothelial cells (Wu et al., 1996; Fisslthaler et al., 1999; Node et al., 1999; Fisslthaler et al., 2000; Enayetallah et al., 2004). Expression of CYP2J2 within the lungs is predominantly localised to ciliated airway epithelial cells, but is also significant in nonciliated airway epithelial cells, bronchial smooth muscle cells, pulmonary vascular smooth muscle cells, pulmonary vascular endothelial cells and alveolar macrophages (Zeldin et al., 1996a). CYP2J2 exhibits cell specific expression throughout the entire gastrointestinal tract including the oesophagus, stomach, duodenum, jejunum, ileum and colon (Zeldin et al., 1997b). In the oesophagus,

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CYP2J2 expression is primarily localised to the squamous epithelium and autonomic ganglion cells, but is also evident in oesophageal smooth muscle and vascular endothelial cells (Zeldin et al., 1997b). In the stomach, CYP2J2 is most prominently expressed in the chief and parietal cells of the gastric glands, and in the nerve cells of the autonomic ganglia (Zeldin et al., 1997b; Enayetallah et al., 2004). In the small intestine, CYP2J2 is primarily expressed in the autonomic ganglion cells and in the absorptive villi epithelial cells, and is also expressed, to a lesser extent, in the muscularis smooth muscle cells, mucous-producing goblet cells, and vascular endothelial cells (Zeldin et al., 1997b). Finally, within the colon, CYP2J2 expression is most prominent within the autonomic ganglion cells and surface columnar epithelial cells, but is also evident in goblet cells, muscularis smooth muscle cells and vascular endothelial cells (Zeldin et al., 1997b). Within the liver, CYP2J2 is expressed in hepatocytes (Enayetallah et al., 2004), and within the kidney, CYP2J2 expression has been localised to the proximal tubules, distal tubules and collecting ducts of the nephron (Enayetallah et al., 2004). CYP2J2 expression in the pancreas is highly localised to the cells in the islets of Langerhans, but is also detected at a much lower level in the pancreatic exocrine and vascular smooth muscle cells (Zeldin et al., 1997a; Enayetallah et al., 2004). Other members of the CYP2J subfamily also exhibit significant extrahepatic expression in other species (Table 1.4). Rat CYP2J3 exhibits a very similar tissue distribution to that of CYP2J2, with high expression in the heart and liver (Wu et al., 1997), as well as significant expression in the lungs (Zeldin et al., 1996a; Wu et al., 1997), gastrointestinal tract (Wu et al., 1997; Zeldin et al., 1997b), kidney (Wu et al., 1997), and pancreas (Zeldin et al., 1997a). Rat CYP2J4 is primarily expressed within the small intestine and the olfactory mucosa, and to a lesser extent in the liver (Zhang et al., 1997). Likewise, rabbit CYP2J1 appears to be primarily expressed in the small intestine (Kikuta et al., 1991; Koike et al., 1997). Mouse CYP2J5 is predominantly expressed within the kidney, but also at lower levels in the liver (Ma et al., 1999), and mouse CYP2J9 is primarily expressed within the brain (Qu et al., 2001).

1.7 Biological Significance of CYP2J2

As was discussed in detail previously (sections 1.5.1 – 1.5.4), EETs exhibit a range of potent biological activities in a variety of tissues, and current evidence suggests that

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CYP2J2 is the predominant AA epoxygenase responsible for the generation of EETs in extrahepatic tissues (Wu et al., 1996; Zeldin et al., 1996a; Zeldin et al., 1997a; Zeldin et al., 1997b). The widespread tissue distribution of CYP2J2 and its localisation to particular cell types in which EETs have significant biological actions suggests that CYP2J2 plays an important functional role within the body. Some of the potential physiological roles of CYP2J2 in the various tissues in which it is expressed are discussed in sections 1.7.1-1.7.2.

1.7.1 Potential Role of CYP2J2 in the Heart and Vasculature CYP2J2 is highly expressed within the heart and vasculature and is proposed to play an important role within these tissues (Figure 1.9; Wu et al., 1996; Node et al., 1999). CYP2J2 is abundantly expressed within cardiomyocytes and EETs are produced endogenously within the human heart (Wu et al., 1996; Enayetallah et al., 2004). Significantly, the enantiomeric distribution of EETs produced within the heart is in accordance with that produced by recombinant CYP2J2-mediated metabolism of AA, suggesting that CYP2J2 is the predominant epoxygenase in the heart (Wu et al., 1996). The influence of EETs on cardiac function suggests an important physiological role within the cardiovascular system for CYP2J2. As discussed in section 1.5.4.1, EETs modulate the activity of Na+2+ and Ca ion channels in cardiomyocytes, thereby affecting heart rate and the contractile force of the myocardium (Chen et al., 1999; Lee et al., 1999). Importantly, EET biosynthesis is enhanced in damaged coronary arteries and during cardiac ischaemia and reperfusion (Rosolowsky et al., 1990; Seubert et al., 2004), and EETs have been shown to improve the recovery of the myocardium following ischaemia-reperfusion injury (Wu et al., 1997). Thus, EETs, and CYP epoxygenases such as CYP2J2, may play an important cardioprotective role (Figure 1.9; Seubert et al., 2004; Spiecker & Liao, 2005). Understanding of the role of CYP2J2 in cardiac function has been enhanced by a recent study in which transgenic mice with cardiomyocyte-specific over-expression of CYP2J2 were generated (Seubert et al., 2004). These CYP2J2 transgenic mice were found to have normal heart anatomy and basal myocardial contractile function, but importantly, exhibited significantly better recovery of myocardial function following ischaemia and reperfusion compared with wild type mice (Seubert et al., 2004). Analysis of epoxygenase activity and fatty acid metabolism indicated that cardiomyocytes from the CYP2J2 transgenic mice had significantly higher AA epoxygenase activity and released more of the stable hydrated EET metabolites 8,9-DHET, 11,12-DHET and 14,15-DHET, than cardiomyocytes from wild type mice (Seubert et al., 2004). In this model, perfusion with physiologically relevant

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concentrations of 11,12-EET improved the postischaemic recovery of wild type hearts. Furthermore, it was reported that the inclusion of an epoxygenase inhibitor in this model completely abolished the improvement in postischaemic recovery in CYP2J2 transgenic mice. These results indicate that CYP2J2 plays a role in cardioprotection following ischaemia, and suggests that the cardioprotective effects of CYP2J2 are mediated by EETs (Seubert et al., 2004).

K+ hyperpolarisation vasodilation VASCULAR SMOOTH MUSCLE CELL K Ca smooth muscle cell migration + - K +

- - NF-NB activation inflammation phospholipid vasoprotection CYP2J2 + + AA EETs t-PA expression fibrinolysis

- - HR injury cell death

ENDOTHELIAL CELL

Na+ + phospholipid K

CYP2J2 - cardioprotection AA EETs heart rate Ca2+ - contractility

CARDIOMYOCYTE

L-type Ca2+ Channel Ca2+

Figure 1.9 Proposed biological roles of CYP2J2 in the heart and cardiovasculature CYP2J2 is abundantly expressed within the cardiovascular system and is proposed to play an important protective role in both the vascular endothelium and in cardiomyocytes via the generation of EETs. Protective effects of EETs within the cardiovascular system include; induction of vasodilation via the activation of large-conductance calcium- activated potassium channels (KCa) channels; inhibition of vascular smooth muscle cell migration; anti-inflammatory activity via inhibition of the pro-inflammatory transcription factor nuclear factor NB (NF-NB); stimulation of fibrinolysis via the activation of tissue- type plasminogen activator (t-PA) expression; protection against hypoxia-reoxygenation (HR) injury; and modulation of heart rate and myocardial contractile force via inhibition of cardiac Na+2+ and L-type Ca channels.

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Apart from being expressed in cardiac myocytes, CYP2J2 is also abundantly expressed in the cardiovasculature; in particular, CYP2J2 is localised to the endothelial cells of small and large human coronary arteries (Node et al., 1999). As discussed previously (sections 1.5.2 – 1.5.3), EETs have a number of important effects within the vascular system. In particular, EETs are vasodilatory and mediate anti-inflammatory, anti-migratory, anti-thrombotic, anti-oxidant and anti-apoptotic effects within the vascular wall (Node et al., 1999; Node et al., 2001; Yang et al., 2001; Sun et al., 2002; Spiecker & Liao, 2005). Thus, the abundant expression of CYP2J2 within the cardiovasculature suggests that this enzyme may play an important role in regulating vascular homeostasis (Figure 1.9; Spiecker & Liao, 2005). In addition to CYP2J2, the alternate CYP epoxygenases CYP2C8 and CYP2C9 have been identified in vascular endothelial cells and may contribute to the vascular effects of EETs (Fisslthaler et al., 1999; Fisslthaler et al., 2000). For example, a series of elegant experiments employing inducers and antisense oligonucleotides to enhance and attenuate CYP2C expression have indicated that the EDHF in porcine coronary arteries is a metabolite of the porcine homologue of human CYP2C8/9 (Fisslthaler et al., 1999; Fisslthaler et al., 2000). The EDHF response in human coronary arteries could be mediated by a CYP2C isoform, but evidence that CYP2J2 is not involved has not yet been presented. While CYP2C subfamily members have been associated with the EDHF response in the coronary vasculature, a number of the non-vasodilatory responses of EETs within the vasculature have been associated with CYP2J2 expression and activity. As discussed previously (section 1.5.3.1), 11,12-EET exhibits potent anti- inflammatory effects in vascular endothelial cells by inhibiting the pro-inflammatory transcription factor NF-NB and preventing cytokine-induced adhesion of leukocytes to the vascular wall (Node et al. 1999). Similar anti-inflammatory effects were achieved by over-expressing CYP2J2 in vascular endothelial cells (Node et al., 1999). Indirect support for the importance of CYP2J2 in mediating anti-inflammatory effects in the vascular wall has been provided by an in vitro study showing that the CYP2C9 epoxygenase is a significant source of reactive oxygen species in coronary arteries, and is, in fact pro-inflammatory in vascular endothelial cells, despite generating 11,12-EET. Fleming et al. (2001b) demonstrated, in accordance with the findings of Node et al. (1999), that the exogenous application of 11,12-EET inhibited cytokine- induced activation of NF-NB and cell adhesion molecule expression in vascular endothelial cells. However, while over-expression of CYP2J2 previously produced an anti-inflammatory effect similar to that seen with exogenous application of 11,12-EET (Node et al., 1999), the over-expression of CYP2C9 enhanced NF-NB activity and was

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pro-inflammatory (Fleming et al., 2001b). Although over-expression of CYP2C9 increased 11,12-EET formation, it also stimulated the production of reactive oxygen species within the vascular wall, which activated the redox-sensitive NF-NB and increased endothelial cell adhesion molecule expression (Fleming et al., 2001b). Thus, in contrast to CYP2J2, the anti-inflammatory effect of 11,12-EET derived from CYP2C9 is overshadowed by the generation of pro-inflammatory reactive oxygen species (Fleming et al., 2001b). The formation of superoxide anions, hydroxyl radicals and hydrogen peroxide during the CYP reaction cycle has been previously documented, and appears to be isoform-specific. Thus, like CYP2C9, CYP2E1 activity generates significant levels of reactive oxygen species (Davydov, 2001), whereas CYP2J2 is inactive in this regard (Yang et al., 2001). Differences in the generation of reactive oxygen species by different CYPs could be due to the efficiency of coupling of substrate and NADPH oxidation (Davydov, 2001; Fleming, 2001). Aside from anti-inflammatory actions, EETs, especially 11,12-EET, possess fibrinolytic properties in vascular endothelial cells (Node et al., 2001), anti-migratory effects on vascular smooth muscle cells (Sun et al., 2002), and protect the endothelium against hypoxia-reoxygenation injury (Yang et al., 2001). Importantly, these vascular effects of 11,12-EET are similarly achieved by over-expression of CYP2J2 (Figure 1.9). Thus, over-expression of CYP2J2 in vascular endothelial cells mimicked the effect of 11,12-EET on expression of the fibrinolytic protein t-PA (Node et al., 2001), while over- expression of CYP2J2 in vascular smooth muscle cells inhibited FCS- and PDGF- induced cell migration in a similar fashion to that achieved with the application of exogenous 11,12-EET (Sun et al., 2002). This suggests that CYP2J2 may be an important modulator of fibrinolytic activity and smooth muscle cell migration in the vascular wall. In bovine aortic endothelial cells exposed to hypoxia-reoxygenation, the expression of CYP2J2 immunoreactive protein was decreased (Yang et al., 2001). Significantly, over-expression of CYP2J2 reduced the generation of reactive oxygen species during hypoxia-reoxygenation, and prevented endothelial cell death. Thus, CYP2J2 may be protective in the endothelium, and down-regulation of CYP2J2 during hypoxia-reoxygenation may contribute to endothelial injury (Yang et al., 2001). Vascular inflammation and thrombosis, smooth muscle cell migration, and endothelial injury are key features of cardiovascular diseases such as atherosclerosis; thus the demonstration that CYP2J2 can modulate these processes implicates an important role for CYP2J2 in the regulation of vascular function under normal and pathophysiological conditions, and suggests that alterations in CYP2J2 expression and/or activity may be associated with altered risk of developing cardiovascular disease (Yang et al., 2001; Spiecker & Liao, 2005).

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The importance of CYP2J2 in vascular function and cardiovascular disease has been strengthened by a recent study of the impact of CYP2J2 polymorphism on coronary artery disease (Spiecker et al., 2004). Several polymorphisms were identified within the CYP2J2 gene. In particular, nine polymorphisms were detected within the coding region of the CYP2J2 gene, four of which result in amino acid substitution and decreased AA oxidation, and one polymorphism was detected in the CYP2J2 promoter region (King et al., 2002). The promoter polymorphism was found within a region resembling the consensus sequence for the specificity protein 1 (Sp1) transcription factor, and, unlike the coding sequence polymorphisms that affect CYP2J2 function, the promoter polymorphism occurs at a relatively high frequency of 8-16% of the population, depending on racial/ethnic background (King et al., 2002). The CYP2J2 promoter polymorphism, in which a guanidine is replaced by a thymidine, decreased Sp1 binding and CYP2J2 promoter activity in transfected cells (Spiecker et al., 2004). Given the relative frequency of this polymorphism, and its potential functional importance to CYP2J2 expression, an epidemiological study was conducted to determine the frequency of the CYP2J2 promoter polymorphism in a cohort of subjects with coronary artery disease as well as control subjects without any angiographic evidence of coronary artery disease. The promoter polymorphism occurred at a higher frequency in patients with coronary artery disease compared with control subjects, and the plasma concentrations of stable EET metabolites of CYP2J2 were significantly lower in subjects with the CYP2J2 promoter polymorphism (Spiecker et al., 2004). While further studies are required, the data from this investigation supports previous experimental evidence of a protective role for CYP2J2-derived EETs in the cardiovascular system, and supports the hypothesis that altered CYP2J2 expression/activity is associated with altered susceptibility to cardiovascular disease (Spiecker et al., 2004).

1.7.2 Potential Role of CYP2J2 in Other Tissues CYP2J2 expression has been demonstrated in a number of tissues outside the cardiovascular system in which EETs have important biological activities. CYP2J2 is abundantly expressed in the lung, and substantial amounts of EETs have been detected in human lung tissue (Zeldin et al., 1996a). Significantly, the regioselectivity of endogenous EETs produced within the human lung is similar to that produced by recombinant CYP2J2-mediated metabolism of AA, suggesting that CYP2J2 is one of the key epoxygenases responsible for the generation of EETs within the lung (Zeldin et al., 1996a). The pulmonary expression of other CYP enzymes has been previously documented (Hukkanen et al., 2002), but the distribution of CYP2J2

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within the human lung appears to be unique compared to other CYPs (Zeldin et al., 1996a). In particular, CYP2J2 is significantly expressed within a variety of different cell types throughout the lung where EETs have important biological functions. For example, CYP2J2 is expressed in airway epithelial cells and bronchial smooth muscle cells (Zeldin et al., 1996a), in which EETs respectively modulate Cl- secretion (Jacobs & Zeldin, 2001; Roman, 2002) and bronchial smooth muscle tone (Jacobs & Zeldin, 2001). CYP2J2 is also localised within pulmonary vascular smooth muscle and endothelial cells (Zeldin et al., 1996a) where EETs may modulate pulmonary vascular tone (Tan et al., 1997) and mediate anti-inflammatory and cytoprotective effects similar to that observed in the cardiovasculature (Jacobs & Zeldin, 2001). Thus, via the production of pulmonary EETs, CYP2J2 may influence normal lung functions such as airway fluid production, bronchial tone and pulmonary vascular tone, and alterations in CYP2J2 expression and/or activity may be associated with altered susceptibility to pulmonary injury and disease (Zeldin et al., 1996a; Jacobs & Zeldin, 2001). EETs are produced endogenously within the human gastrointestinal tract, and CYP2J2 is abundantly expressed from the oesophagus to the colon (Zeldin et al., 1997b). Although further studies are required, the cellular localisation of CYP2J2 within the gastrointestinal tract may reflect important functional roles for CYP2J2-derived EETs in gastrointestinal physiology (Zeldin et al., 1997b). For example, the expression of CYP2J2 within the intestinal vascular endothelium, together with the demonstrated effects of EETs on vascular tone within the intestine (Proctor et al., 1987) suggests that CYP2J2 may participate in the regulation of intestinal vascular tone (Zeldin et al., 1997b). Given the role of EETs in controlling fluid and electrolyte transport in extraintestinal tissues such as the kidney (Roman, 2002) and the lung (Jacobs & Zeldin, 2001), the localisation of CYP2J2 within the absorptive epithelial cells in the colon and small intestine may reflect a role for CYP2J2 and its eicosanoid metabolites in fluid and electrolyte transport in the gut (Zeldin et al., 1997b), while the expression of CYP2J2 in the nerve cells of the autonomic ganglia, combined with the demonstrated effects of EETs in stimulating the release of neuropeptides (Ojeda et al., 1989), suggests a role for CYP2J2-derived EETs in mediating the release of intestinal neuropeptides and regulating intestinal motility (Zeldin et al., 1997b). EETs are produced endogenously within the human pancreas and the total amount of EETs recovered from different human pancreatic specimens has been found to be highly correlated with the amount of CYP2J2 immunoreactive protein in the particular pancreatic specimen, suggesting that CYP2J2 is one of the predominant epoxygenases present in the pancreas (Zeldin et al., 1997a). Moreover, while studies have demonstrated the expression of other CYPs within exocrine acinar pancreatic

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cells (Murray et al., 1988), CYP2J2 expression within the pancreas is highly localised to the endocrine cells of the islets of Langerhans (Zeldin et al., 1997a). These cells release insulin and/or glucagon in response to various stimuli, and play a critical role in regulating glucose homeostasis (Zeldin et al., 1997a). The localisation of CYP2J2 to these insulin- and glucagon- releasing islet cells, combined with the demonstration that low concentrations of EETs potently stimulate the release of insulin and glucagon from isolated rat pancreatic islets (section 1.5.4.4; Falck et al., 1983), suggests that CYP2J2 and its eicosanoid products may be involved in stimulus-secretion coupling within the pancreas, which contributes to glucose homeostasis (Zeldin et al., 1997a). In view of this, it has been hypothesised that alterations in CYP2J2 expression and/or activity may be associated with the development of diabetes mellitus. To this effect, the frequency of a particular CYP2J2 polymorphism, the allelic variant CYP2J2*6, was examined in a Caucasian population with type 1 or type 2 diabetes mellitus. The CYP2J2*6 variant encodes an enzyme with lower capacity to metabolise AA, but the frequency of this polymorphism within the study population was very low, and was not different between patients with type 1 or type 2 diabetes and control subjects. Further studies are required to determine whether any of the other CYP2J2 polymorphisms are associated with diabetes (Pucci et al., 2003). Aside from the pancreas, CYP2J2 is expressed in another endocrine tissue: the pituitary gland (Enayetallah et al., 2004). As EETs have been shown to stimulate the release of hormones from the pituitary such as growth hormone (Snyder et al., 1989) and vasopressin (Negro-Vilar et al., 1985), the expression of CYP2J2 in the pituitary may reflect a role for CYP2J2-derived EETs in the modulation of pituitary hormone release (Enayetallah et al., 2004). EETs are endogenous constituents of the human kidney, and, as outlined in section 1.5.4.2, are thought to play a role in the regulation of renal function via the modulation of fluid and electrolyte excretion and renal vascular tone (Zeldin, 2001; Roman, 2002). The renal effects of EETs are generally anti-hypertensive, and altered renal EET production may contribute to the pathogenesis of hypertension (Makita et al., 1994; Yu et al., 2000; Roman, 2002). CYP2C subfamily isoforms are considered to be the primary epoxygenases that mediate renal EET formation (Zeldin et al., 1995a), but CYP2J2 is also expressed within the kidney and may contribute to renal function (Wu et al., 1996; Scarborough et al., 1999; Enayetallah et al., 2004). Immunohistochemical analysis has shown that renal CYP2J2 expression is localised to the proximal tubules and collecting ducts (Enayetallah et al., 2004), both of which are regions of the nephron in which EETs affect fluid and electrolyte transport (refer to section 1.5.4.2). Thus, CYP2J2-derived EETs may modulate renal ion transport, and may play a role in the

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pathogenesis of hypertension (Scarborough et al., 1999). Indeed, the decreased renal expression of rat CYP2J isoform(s) in angiotensin ,,-infused rats fed a high-salt diet may contribute to the development of hypertension in these animals (Zhao et al., 2003), and studies with CYP2J2 transgenic animals would greatly improve our current limited understanding of the role of CYP2J2-derived EETs in kidney function and blood pressure regulation (Scarborough et al., 1999). Finally, CYP2J2 may play an important functional role in the liver. The hepatic production of EETs is significant (Zeldin et al., 1996b), and EETs may play a role in vasopressin-stimulated glycogenolysis in hepatocytes (Yoshida et al., 1990). They may also modulate hepatic calcium homeostasis (Kutsky et al., 1983), and may be secreted into the circulation to have effects in extrahepatic tissues (Zeldin et al., 1996b). As in the kidney, CYP2C isoforms appear to be the primary hepatic epoxygenases (Rifkind et al., 1995; Zeldin et al., 1996b), but CYP2J2 is also significantly expressed within hepatocytes, and thus may contribute to the effects of EETs in liver physiology and pathophysiology (Wu et al., 1996; Enayetallah et al., 2004).

In summary, the widespread expression of CYP2J2 in tissues in which EETs are produced endogenously and exert important biological effects implicates important roles for this enzyme in normal cellular physiology. Alterations in CYP2J2 expression and/or activity may also be involved in the pathophysiology of various human diseases and conditions. In view of this, it is of key interest to study factors influencing CYP2J2 expression and activity, such as genetic polymorphisms and regulation of CYP2J2 gene expression. Some progress has been made in characterising CYP2J2 genetic polymorphisms, but little is known about the molecular mechanisms regulating CYP2J2 gene expression. The aim of this thesis was to identify factors important in regulating CYP2J2 gene expression; in particular, to identify molecular mechanisms which influence transcriptional activity of the CYP2J2 gene.

1.8 Regulation of CYP Gene Expression

Regulation of CYP genes is complex, with many factors found to influence their expression in tissues and during development (Gonzalez, 1988; Morgan et al., 1998; Waxman, 1999). Certain CYPs in some species display sexual dimorphism i.e. significant gender differences in their expression (Gonzalez, 1988; Wiwi & Waxman,

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2004). CYP gene expression is known to be regulated by inflammatory cytokines and endogenous hormones (e.g. sex steroids, glucocorticoids and growth hormone; Morgan et al., 1998; Waxman, 1999). Furthermore, certain CYP genes are expressed at low levels under normal conditions, but are rapidly induced upon exposure to xenobiotics such as drugs, chemicals and environmental pollutants (Denison & Whitlock, 1995; Dogra et al., 1998). Xenobiotic inducers are often substrates for the induced CYP and thus act to enhance their own metabolism. In this way, xenobiotic induction of CYP expression is an efficient biological mechanism to control metabolic capacity (Denison & Whitlock, 1995; Williams et al., 2005). Control of transcription initiation is a key regulatory mechanism used for determining whether a particular gene will be expressed and how much mRNA and protein is produced (Cartharius et al., 2005). Indeed, regulation of CYP expression is primarily achieved via modulation of the transcriptional activity of CYP genes (Gonzalez, 1988; Okey, 1990; Denison & Whitlock, 1995; Waxman, 1999; Akiyama & Gonzalez, 2003), although post-transcriptional, translational and post-translational mechanisms of regulation have been reported in certain instances (Gonzalez, 1988; Okey, 1990; Porter & Coon, 1991). An example is the ethanol-inducible CYP2E1, which is regulated by xenobiotics and pathophysiological factors by transcriptional and post-transcriptional mechanisms, such as mRNA and protein stabilisation (Song et al., 1987; Gonzalez, 1988; Roberts et al., 1995; Donato & Castell, 2003). The transcriptional activity of genes is regulated by the interaction of transcription factor proteins with their specific cis-acting regulatory elements, which are primarily located in the upstream regulatory (promoter or enhancer) region of the gene (Gonzalez & Lee, 1996; Cartharius et al., 2005). A large number of different transcription factors are present in any given cell; indeed the may encode as many as 3000 different transcription factors, and more than 1400 have been identified (Cartharius et al., 2005). The binding of transcription factors within the regulatory region of genes may enhance or diminish the rate of transcription initiation by RNA polymerases. Transcriptional regulation of genes is complex and often involves the actions of a number of different transcription factors within gene promoters (Gonzalez & Lee, 1996; Akiyama & Gonzalez, 2003). The molecular mechanisms governing the transcriptional regulation of CYP genes are poorly understood in many cases (Akiyama & Gonzalez, 2003). However, considerable progress has been made in understanding some of the mechanisms involved in regulating the constitutive and tissue-specific expression of certain hepatic CYPs. Furthermore, the mechanism of induction of certain xenobiotic- and endobiotic- metabolising CYPs by nuclear receptors, a particular class of transcription factors, has

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been well-studied. The down-regulation of many CYP isoforms by inflammatory mediators has also been investigated. Key features of these mechanisms of regulation of CYP gene expression are discussed below.

1.8.1 Regulation of CYP Genes by Liver-enriched Transcription Factors As most mammalian CYPs are primarily expressed within the liver (Guengerich, 1995; Akiyama & Gonzalez, 2003), the tissue-specific expression of hepatic CYP genes has been best-studied, with less information on mechanisms controlling extrahepatic CYP expression (Gonzalez & Lee, 1996; Scarborough et al., 1999). The liver-enriched transcription factors (LETFs) appear to be important in the liver-specific expression of CYP genes (Gonzalez & Lee, 1996; Akiyama & Gonzalez, 2003; Wiwi & Waxman, 2004). LETFs include the hepatocyte nuclear factors (HNFs) HNF1Į, HNF1ȕ, HNF3, HNF4Į, and HNF6, albumin D-site binding protein (DBP) and several CCAAT/enhancer binding proteins (C/EBPs; Gonzalez & Lee, 1996; Akiyama & Gonzalez, 2003). The promoters of many liver-specific genes, including CYPs, contain binding sites for LETFs, which do not appear to be under the control of endogenous ligands. Thus, LETFs are emerging as key regulators of constitutive and tissue-specific expression of numerous hepatic genes including CYPs (Akiyama & Gonzalez, 2003). As many CYP genes contain multiple LETF binding sites within their 5’-regulatory regions, liver-specific CYP expression is likely to reflect the coordinated effects of multiple LETFs (Gonzalez & Lee, 1996; Akiyama & Gonzalez, 2003). For example, the rat CYP2C11 gene promoter is strongly activated by the LETFs HNF1Į and HNF3E (Park & Waxman, 2001), and the rat CYP2C12 gene is regulated, in part, by the synergistic actions of HNF3ȕ and HNF6 (Delesque-Touchard et al., 2000). Amongst the LETFs, HNF4Į is a particularly important regulator of CYP gene expression. HNF4Į is a member of the nuclear receptor superfamily that is expressed in the liver, intestine, pancreas and kidney, and is classified as an orphan receptor, as no endogenous or exogenous ligand has yet been identified (Sladek et al., 1990; Akiyama & Gonzalez, 2003). HNF4Į binds as a homodimer to consensus sequences containing direct repeats of the hexamer AGGTCA separated by one base pair (DR1; Sladek et al., 1990), as well as similar sequences such as the HPF-1 motif present in a number of CYP gene promoters (Akiyama & Gonzalez, 2003). CYP genes found to be activated by HNF4Į include rabbit CYP2C genes (Chen et al., 1994), rat CYP3A genes (Ogino et al., 1999), and the mouse Cyp2a4 gene (Yokomori et al., 1997).

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The expression of the sexually dimorphic rat CYP2C11 gene in male, but not female, rat liver is dependent on the intermittent pulses of plasma growth hormone (GH) that is characteristic of adult male rats (Wiwi & Waxman, 2004). While only partially understood, substantial evidence indicates that the sexually dimorphic effects of pulsatile GH on hepatic CYP2C11 gene expression are mediated via the Janus kinase 2 – signal transducer and activator of transcription 5b (JAK2-STAT5b) signalling pathway (Waxman & Chang, 2005). STAT5b is efficiently activated by the male pulsatile GH pattern and binds to target elements within the promoters of male-specific hepatic CYPs (Waxman et al., 1995; Wiwi & Waxman, 2004). Through the use of STAT5b-null male mice, STAT5b has been demonstrated to play a critical role in sex- specific CYP expression (Davey et al., 1999), and LETFs, especially HNF4Į, appear to cooperate with STAT5b in the GH-regulated expression of sex-specific CYPs (Wiwi & Waxman, 2004). In addition to regulation of CYPs in other mammalian species, HNF4Į is also important in the regulation of several human CYPs. Using antisense strategies targeting HNF4Į in human hepatocytes, HNF4Į was identified as an important regulator of human CYP3A4, CYP3A5 and CYP2A6 genes (Jover et al., 2001). Other studies have also implicated HNF4Į in the regulation of the human CYPs 7A1 and 8B1, that are involved in conversion of cholesterol to bile acids (Stroup & Chiang, 2000; Zhang & Chiang, 2001). Furthermore, as HNF4Į has been identified as a key regulator of the expression of other LETFs (Wiwi & Waxman, 2004), HNF4Į may also be indirectly involved in the regulation of a broader range of CYPs by controlling the expression of their LETF regulators (Akiyama & Gonzalez, 2003). CYPs that are regulated by LETFs other than HNF4Į include CYP2E1, CYP1A2 and CYP27 genes which are activated by HNF1Į, CYP2A4 and CYP2A5 genes that are regulated by DBP, and CYP2B1, CYP2B10 and CYP2D5 which are regulated by C/EBP members (Akiyama & Gonzalez, 2003).

1.8.2 Receptor-mediated Regulation and Induction of CYP Gene Expression A number of the major classes of CYP genes are selectively regulated by certain ligand-activated receptors (Honkakoski & Negishi, 2000). These regulatory mechanisms are discussed below in sections 1.8.2.1 – 1.8.2.6.

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1.8.2.1 CYP1A Induction by the Ah Receptor The ligand-dependent regulation of the CYP1A genes by the aryl hydrocarbon (Ah) receptor has been intensively investigated and is quite well understood (Waxman, 1999). The Ah receptor, a member of the Per-Arnt-Sim family of basic helix-loop-helix transcription factors, normally resides within the cytosol of the cell in an inactive form bound to the molecular chaperone heat shock protein 90 (hsp90; Hankinson, 1995; Whitlock, 1999). Hsp90 is thought to mask a nuclear localisation signal within the receptor and maintain it in a high affinity ligand-binding conformation (Dogra et al., 1998; Williams et al., 2005). A number of high affinity ligands for the Ah receptor have been identified and include a variety of toxic hydrophobic chemicals that are planar and aromatic in structure. These include polycyclic aromatic hydrocarbons (PAHs), such as benzo(a)pyrene and 3-methylchlolanthrene, that can be found in cigarette smoke, halogenated aromatic hydrocarbons such as polychlorinated dibenzo-p-dioxins, biphenyls and dibenzofurans, and other chemicals (Hankinson, 1995; Williams et al., 2005). Upon ligand binding, the Ah receptor is activated and hsp90 is released. The activated ligand-bound Ah receptor is translocated to the nucleus, where it heterodimerises with the nuclear factor Arnt (Ah receptor nuclear translocator protein). Dimerisation with Arnt allows the Ah receptor to bind to specific target sequences, known as xenobiotic-responsive elements (XREs) or Ah-responsive elements (AhREs), that are located in the 5’-regulatory regions of CYP1A and other Ah receptor-inducible genes, and activate transcription (Hankinson, 1995; Waxman, 1999; Whitlock, 1999; Williams et al., 2005). This process is well conserved across different species and accounts for the induction of CYP1A gene expression by PAHs in many cell types (Waxman, 1999). The CYP1B1 promoter also contains an XRE and is regulated by the Ah receptor in a similar fashion to CYPs 1A (Smith et al., 1998; Williams et al., 2005).

1.8.2.2 Nuclear Receptors Involved in CYP Gene Expression In contrast to the induction of the CYP1 genes by the Ah receptor, the regulation of other CYP families by certain xenobiotics and endogenous molecules involves members of the nuclear receptor superfamily (Waxman, 1999). Nuclear receptors comprise a large family of transcription factors that transform endogenous and exogenous stimuli (e.g. hormones, drugs and environmental compounds) into cellular responses (Honkakoski & Negishi, 2000; Wang & LeCluyse, 2003). By regulating expression of a wide range of target genes, nuclear receptors play a crucial role in development and adult physiology, and also in the body’s defense system against xenobiotics (Akiyama & Gonzalez, 2003; Wang & LeCluyse, 2003).

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Members of the nuclear receptor superfamily include the steroid, thyroid, retinoid, and vitamin D receptors, as well as the co-called ‘orphan receptors’, for which endogenous ligands or activators were unknown at the time of discovery (Honkakoski & Negishi, 2000; Akiyama & Gonzalez, 2003; Wang & LeCluyse, 2003). Nuclear receptors generally have a conserved modular protein structure that consists of a highly conserved DNA-binding domain (DBD), and a less conserved ligand-binding domain (LBD) at their carboxy-terminal that is responsible for ligand binding and ligand-dependent transactivation (Wang & LeCluyse, 2003). Via their DBD, nuclear receptors bind to specific sequences within the promoter regions of target genes known as hormone response elements (HREs) or xenobiotic response elements (XREs; Wang & LeCluyse, 2003). These nuclear receptor response elements are generally composed of two core half-sites of the consensus hexamer AGGTCA that are present as inverted, everted or direct repeats with a 3- to 6 bp spacing. The receptors generally bind to these elements as homo- or heterodimers (Honkakoski & Negishi, 2000; Wang & LeCluyse, 2003; Eloranta & Kullak-Ublick, 2005). Steroid hormone receptors bind as homodimers, while many other hormone and orphan receptors, including those important in CYP gene regulation, bind as heterodimers with the retinoid X receptor (RXR; Honkakoski & Negishi, 2000; Wang & LeCluyse, 2003). The general model of nuclear receptor-mediated activation of gene expression involves the binding of ligand to the LBD, which results in a change in receptor conformation that leads to the dissociation of corepressor proteins, association with accessory proteins such as transcriptional coactivators, and transcriptional activation of the target gene (Akiyama & Gonzalez, 2003; Eloranta & Kullak-Ublick, 2005). Induction of particular CYP gene subfamilies by xenobiotics, as well as regulation by hormones and other endogenous mediators, involves various members of the nuclear receptor superfamily, including the constitutive androstane receptor, the pregnane X receptor, the peroxisome proliferator-activated receptor, the liver X receptor and the farnesoid X receptor (Waxman, 1999; Honkakoski & Negishi, 2000; Wang & LeCluyse, 2003).

1.8.2.3 CAR-mediated Induction of CYP2B Genes The induction of CYP2B genes, including human CYP2B6, by phenobarbital (PB) and many other “PB-like” lipophilic chemicals has been widely studied and has recently been shown to be mediated by the constitutive androstane receptor (CAR; Honkakoski et al., 1998; Sueyoshi et al., 1999). CAR induces CYP2B gene expression by binding as a heterodimeric complex with RXR to specific sequences known as PB-responsive

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enhancer modules (PBREMs) that have been identified in rat, mouse and human CYP2B genes (Trottier et al., 1995; Honkakoski et al., 1998; Sueyoshi et al., 1999). CAR is a constitutively active receptor that is able to activate PBREM- containing reporter genes in the absence of PB ligands (Honkakoski et al., 1998). Because of its constitutive activity, the subcellular localisation of CAR is likely to be a crucial determinant of its activity, and several theories have been proposed to account for PB-dependent activation of CYP2B expression by CAR (Honkakoski & Negishi, 2000; Williams et al., 2005). In particular, the transcriptional activity of CAR has been found to be inhibited by the binding of the endogenous steroidal ligands, androstanol (5Į-androstan-3Į-ol) and androstenol (5Į-androst-16-en-3Į-ol; Forman et al., 1998), and it has been suggested that PB and other structurally diverse so-called PB-like chemicals may relieve the inhibitory binding of androstanes to CAR, and thus stimulate its translocation from the cytoplasm to the nucleus, where it can bind to PBREMs and activate transcription (Sueyoshi et al., 1999; Waxman, 1999). However, supraphysiological concentrations of androstanes are required to suppress CAR- mediated gene expression, so that the true endogenous inhibitory ligand for CAR may be a different steroid (Honkakoski & Negishi, 2000; Williams et al., 2005). Other studies have implicated a phosphorylation-dependent regulation of CAR activity by PB. Thus, it has been suggested that PB may stimulate nuclear translocation of CAR and CYP2B gene transcription by altering the phosphorylation status of the receptor (Kawamoto et al., 1999; Honkakoski & Negishi, 2000; Williams et al., 2005).

1.8.2.4 PXR-mediated Induction of CYP3A Genes The well known, but seemingly paradoxical induction of members of the CYP3A subfamily by both glucocorticoids such as dexamethasone, and anti-glucocorticoids such as pregnenolone 16Į-carbonitrile (PCN), as well as a broad range of drugs and environmental chemicals, is mediated by the pregnane X receptor (PXR; Waxman, 1999; Honkakoski & Negishi, 2000; Williams et al., 2005). In a species-specific manner, CYP3A inducers bind and activate PXR (or the human homologue that is variously known as hPXR, hPAR, human pregnane activated receptor, and SXR, steroid and xenobiotic receptor; Bertilsson et al., 1998; Kliewer et al., 1998; Lehmann et al., 1998). Ligand-activated PXR-RXR heterodimers bind and transactivate specific elements composed of nuclear receptor half-site consensus or consensus-like sequences within the promoters of mammalian CYP3A genes including human CYP3A4 (Bertilsson et al., 1998; Kliewer et al., 1998; Lehmann et al., 1998; Goodwin et al., 1999).

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The species-specific pattern of induction observed with CYP3A genes reflects the different ligand binding specificities of the PXR receptor in different species. This accounts for the efficient induction of human and rabbit, but not rat and mouse, CYP3A genes by the antibiotic rifampicin. In contrast, potent induction of rat CYP3A, but not human or rabbit CYP3A, by the anti-glucocorticoid PCN, reflects the activation of rat, but not human or rabbit, PXR by PCN (Jones et al., 2000). The differences in the ligand activation profile of PXR receptors from different species arises from relatively low amino acid conservation in their LBDs (Jones et al., 2000).

1.8.2.5 PPAR-mediated Induction of CYP4A Genes The renal and hepatic expression of CYP4A genes from a number of mammalian species is induced by a range of xenobiotics including lipid-lowering fibrate drugs, phthalate esters used in the medical and chemical industries, as well as various other chemicals and environmental pollutants (Waxman, 1999; Honkakoski & Negishi, 2000; Williams et al., 2005). These chemical inducers of CYP4A expression are known as peroxisome proliferators, as they induce hepatic peroxisomal enzymes and increase the size and number of peroxisomes in liver (Waxman, 1999). The induction of CYP4A enzymes and peroxisomal enzymes by peroxisome proliferators is mediated by the peroxisome proliferator-activated receptor alpha (PPARĮ; Johnson et al., 1996; Waxman, 1999), which is also activated by fatty acids and eicosanoid metabolites (Wang & LeCluyse, 2003). PPARĮ is constitutively located within the nucleus, and activates gene transcription by binding as a heterodimer with RXR to specific sequences, referred to as peroxisome proliferator response elements (PPREs), in the upstream regulatory region of target genes (Waxman, 1999; Wang & LeCluyse, 2003). The binding of peroxisome proliferators or endogenous fatty acid ligands to PPARĮ induces a conformational change in the receptor that facilitates its interaction with transcriptional coactivators (Wang & LeCluyse, 2003; Williams et al., 2005). Induction of CYP4A genes by peroxisome proliferators has been demonstrated in rats and rabbits (Aldridge et al., 1995; Johnson et al., 1996). On the other hand, the hepatic peroxisome proliferator response is weak in humans (Waxman, 1999), with no induction of CYP4A isoforms reported thus far (Wang & LeCluyse, 2003). This may be due to the significantly reduced expression, and sensitivity to activation by peroxisome proliferators, of human PPARĮ compared to rat PPARĮ (Keller et al., 1997; Palmer et al., 1998), and the existence of an inhibitory variant of PPARD in humans (Gervois et al., 1999).

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1.8.2.6 LXR- and FXR-mediated Regulation of CYP7A Gene Expression The expression of CYP7A1, which catalyses the first and rate-limiting step in the catabolism of cholesterol to bile acids (section 1.3.2), is modulated by cholesterol and bile acids acting at the orphan liver X receptor (LXR) and farnesoid X receptor (FXR; Waxman, 1999; Honkakoski & Negishi, 2000; Edwards et al., 2002). LXR is activated by several oxysterol metabolites of cholesterol (Lehmann et al., 1997), and up-regulates the rat CYP7A1 gene in response to excess cholesterol by binding to an LXR response element (LXRE) in the rodent CYP7A1 gene (Lehmann et al., 1997). Up-regulation of CYP7A1 in rats by high cholesterol diets is an adaptive mechanism facilitating the conversion of excess cholesterol into bile acids (Jelinek et al., 1990; Lehmann et al., 1997). The critical role of LXR in cholesterol homeostasis was confirmed by the demonstration that LXR null mice are unable to up-regulate Cyp7a1 in response to high levels of dietary cholesterol (Peet et al., 1998). There may be species specificity in the regulation of CYP7A1 by LXR. Thus, the rat and mouse CYP7A1 genes contain LXR response elements, but the human CYP7A1 gene lacks an LXR binding sequence and is not regulated by oxysterols or LXR (Chiang et al., 2001; Agellon et al., 2002; Edwards et al., 2002). The farnesoid X receptor (FXR), so-named due to its activation by supraphysiological levels of farnesol in rats (Forman et al., 1995), binds, and is activated by, physiologically relevant concentrations of bile acids such as chenodeoxycholic acid (Makishima et al., 1999). In a negative feedback mechanism, bile acids suppress CYP7A1 via its bile acid-responsive element (BARE; Chiang & Stroup, 1994). The BARE of CYP7A1 contains a binding site for the nuclear receptor liver receptor homologue-1 (LRH-1; also known as CYP7A1 promoter binding factor; CPF; and Į-fetoprotein transcription factor; FTF), which activates transcription of the CYP7A1 gene and is critical for its liver-specific expression (Nitta et al., 1999; Lu et al., 2000). While FXR suppresses CYP7A1 promoter activity and mediates the down- regulatory effects of bile acids (Makishima et al., 1999), FXR does not interact directly with the CYP7A1 promoter (Riddick et al., 2004). Rather, suppression by FXR is indirect and is thought to involve FXR-mediated up-regulation of the nuclear receptor small heterodimer partner 1 (SHP-1), which interferes with transcriptional activation within the BARE of CYP7A1 by LRH-1 (Goodwin et al., 2000; Lu et al., 2000).

1.8.3 Down-regulation of CYPs by Inflammatory Mediators In humans and animals, infections and inflammatory stimuli have long been recognised to cause changes in the activity and expression of various CYP isoforms in the liver and extrahepatic tissues (Morgan, 2001).

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While certain CYP isoforms are unaffected, or induced by inflammatory stimuli (e.g. CYP4A subfamily; Sewer et al., 1996), in most cases CYP activity and expression is suppressed (Morgan, 2001). For example, induction of CYP1 enzymes by PAHs is inhibited by TNF-D or by transforming growth factor-ȕ (TGF-ȕ; Muntané-Relat et al., 1995; Döhr et al., 1997). Interleukin-6 (IL-6) inhibits PB-induced expression of CYP2B in rat hepatocytes (Clark et al., 1996) and rifampicin-induced expression of CYP3A in human hepatocytes (Muntané-Relat et al., 1995). The expression of both CYP2C11 and CYP3A in rat hepatocytes and liver is down-regulated by TNF-D (Nadin et al., 1995) and interleukin-2 (IL-2; Tinel et al., 1999); CYP2C11 is also suppressed by administration of IL-1 or the bacterial endotoxin LPS (Iber et al., 2000). There have been numerous additional studies that have documented similar effects of cytokines and interleukins that are activated during infection and inflammation, on the expression of both basal and xenobiotic-inducible hepatic CYPs (Waxman, 1999; Morgan, 2001). The suppression of CYPs by cytokines and other inflammatory stimuli appears to occur primarily at the level of transcription (Waxman, 1999; Morgan, 2001). In most cases the mechanisms of transcriptional suppression are poorly understood, although greater understanding of the molecular mechanisms by which some isoforms are down-regulated have emerged (Morgan, 2001). Studies have indicated a role for the cytokine-activated transcription factor NF-NB in the down-regulation of PAH-induced CYP1A1 and CYP1A2 expression by cytokines (Tian et al., 1999; Ke et al., 2001). Activation of NF-NB results in a physical interaction and mutual functional repression between NF-NB and the Ah receptor that is required for CYP1A induction; this decreases cellular Ah receptor availability and diminishes induction (Tian et al., 1999; Ke et al., 2001). The IL-2-stimulated down-regulation of CYP2C11 and CYP3A in rat hepatocytes could be linked to the up-regulation of the proto-oncogene transcription factor c-myc (Tinel et al., 1999); the suppression of CYP2C11 may also involve NF-NB, with Iber et al. (2000) identifying a low-affinity binding site for NF-NB in the CYP2C11 promoter that is required for IL-1 or LPS-mediated suppression of CYP2C11 promoter activity.

1.8.4 Regulation of CYP2J2 Gene Expression Very little is known about the regulation of CYP2J2 or other members of the CYP2J subfamily. Preliminary data suggests that some CYP2J isoforms may be developmentally regulated (Scarborough et al., 1999). For example, mouse Cyp2j5 appears to be regulated in an age-related manner in the kidney and liver; CYP2J5

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protein is present in the kidney before birth, is optimal at 2 to 4 weeks of age, declines at 8 to 10 weeks of age, and then reaches adult levels by 16 weeks of age (Ma et al., 1999). On the other hand, CYP2J5 protein is not expressed in liver before birth, appears for the first time at 1 week of age, remains relatively constant between weeks 1 to 10, and reaches adult levels by 16 weeks (Ma et al., 1999). The expression of Cyp2j9 within the mouse brain also follows an age-related pattern of expression (Qu et al., 2001). Further work is required to determine whether postnatal exposure to environmental or endogenous agents contributes to the age-related changes in CYP2J expression (Scarborough et al., 1999). CYP2J expression may be altered in certain animal models of disease; e.g. renal expression of CYP2J2 immunoreactive protein is increased in SHR compared to control rats (Yu et al., 2000). There is also evidence to suggest that CYP2J expression may be influenced by sex hormones, with the renal expression of CYP2J5 protein demonstrated to be higher in male mice than in female mice after puberty (Ma et al., 2004). As discussed previously, CYP2J2 is constitutively expressed in liver, but is also highly and constitutively expressed in a number of extrahepatic tissues (section 1.6.3; Wu et al., 1996; Zeldin et al., 1996a; Zeldin et al., 1997a; Zeldin et al., 1997b; Enayetallah et al., 2004). The strong extrahepatic expression of CYP2J enzymes, including CYP2J2, suggests that the liver-enriched transcription factors that are important in constitutive expression of many other CYPs (see section 1.8.1) may not regulate CYPs 2J. In keeping with their pattern of strong, constitutive expression, current data shows that CYP2J members are not readily induced by well known CYP inducers (Scarborough et al., 1999). In particular, Zeldin et al. (1996a and 1997b) and Wu et al. (1997) demonstrated that treatment of animals with ȕ-naphthoflavone, phenobarbital, clofibrate or acetone induced, respectively, the hepatic expression of CYP1A, CYP2B, CYP4A, and CYP2E1, but did not affect the hepatic or extrahepatic expression of CYP2J3, the rat homologue of CYP2J2. Therefore nuclear receptors that regulate xenobiotic-inducible CYP expression may not be important in CYP2J2 gene regulation. Prior to the work presented in this thesis, the only data available on the regulation of CYP2J2 expression was a report by Yang et al. (2001) showing that the levels of CYP2J2 immunoreactive protein (presumably the bovine CYP2J orthologue) were decreased in bovine aortic endothelial cells (BAECs) exposed to hypoxia and reoxygenation. In that study, the authors found that CYP2J2 expression was decreased in human coronary arterial endothelial cells exposed to hypoxia and reoxygenation (Yang et al., 2001). Maintenance of CYP2J expression by transfection of BAECs with CYP2J2 cDNA, or the exogenous application of CYP2J2-derived EETs, significantly

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reduced endothelial cell injury arising from hypoxia-reoxygenation (Yang et al., 2001). In view of the vasoprotective properties of CYP2J2 and its eicosanoid metabolites, the loss of CYP2J2 protein could contribute to endothelial cell dysfunction in ischaemia- reperfusion injury, and possibly the pathogenesis of cardiovascular diseases such as atherosclerosis (Yang et al., 2001). However, a molecular mechanism for impaired CYP2J expression was not provided. Altered gene expression in hypoxia and hypoxia-reoxygenation involves the transcription factors hypoxia-inducible factor-1 (HIF-1), activator protein-1 (AP-1) and NF-NB (Semenza, 1996; Guillemin & Krasnow, 1997; Schmedtje et al., 1997; Faller, 1999; Michiels et al., 2001). The experimental work presented in this thesis investigated the role of AP-1 in regulating the expression of the human CYP2J2 gene in normoxia and hypoxia.

1.9 Activator Protein-1

AP-1 is an important transcription factor that plays a key role in the regulation of eukaryotic gene expression (Angel & Karin, 1991). AP-1 is activated by a wide range of external stimuli and plays a role in cellular proliferation, differentiation, tumourigenesis and adaptation to external stresses by regulating the expression of numerous mammalian target genes in a variety of tissues and cell types (Angel & Karin, 1991; Chinenov & Kerppola, 2001; Shaulian & Karin, 2001).

1.9.1 AP-1 Components, Dimerisation and DNA Binding AP-1 is actually a multiprotein transcription factor, composed of members of the Jun and Fos family of proteins (Angel & Karin, 1991; Ryseck & Bravo, 1991). Jun proteins include c-Jun, JunB and JunD, with c-Jun being the most widely studied member of this family (Bohmann et al., 1987; Ryder et al., 1988; Sassone-Corsi et al., 1988; Hirai et al., 1989; Ryder et al., 1989; Angel & Karin, 1991; Ryseck & Bravo, 1991). c-Fos was the first of the Fos family of proteins to be identified, and is also the most widely studied (Chiu et al., 1988; Rauscher et al., 1988; Angel & Karin, 1991; Ryseck & Bravo, 1991); other Fos proteins include FosB and the Fos-related antigens, Fra-1 and Fra-2 (Cohen et al., 1989; Zerial et al., 1989; Nishina et al., 1990; Angel & Karin, 1991; Ryseck & Bravo, 1991).

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Jun and Fos proteins belong to the basic region leucine zipper (bZIP) family of proteins that contain a highly conserved basic region involved in DNA binding, and an adjacent domain consisting of a heptad repeat of leucine residues, known as a leucine zipper, that facilitates dimerisation to other bZIP proteins (Kouzarides & Ziff, 1988; Chinenov & Kerppola, 2001; Vogt, 2001). Dimerisation is a requirement for the binding of bZIP proteins to DNA, bringing the basic amino acid sequence of each protein subunit into close proximity and providing a contiguous DNA contact surface (Alber, 1992; Abate et al., 1993; Chinenov & Kerppola, 2001). The leucine zipper of a particular bZIP protein dimerises with the leucine zipper of other bZIP proteins in a specific fashion that is controlled by the non-leucine residues of the zipper (Smeal et al., 1989; Alber, 1992; Vogt, 2001). As shown in Table 1.6, the three Jun family members can bind to DNA as homodimers, or as heterodimers with other Jun or Fos proteins (Nakabeppu et al., 1988; Angel & Karin, 1991; Chinenov & Kerppola, 2001). On the other hand, Fos proteins do not form stable homodimers or heterodimers with other Fos proteins, and heterodimerisation with Jun proteins is essential for DNA binding (Chiu et al., 1988; Halazonetis et al., 1988; Smeal et al., 1989; Angel & Karin, 1991; Chinenov & Kerppola, 2001). The AP-1 transcription factor complex binds with high affinity to the 7 bp DNA consensus sequence TGAG/CTCA that is found within the regulatory region of numerous genes (Figure 1.10; Nakabeppu et al., 1988; Angel & Karin, 1991; Chinenov & Kerppola, 2001).

Fos Jun Jun Jun

TGAG/CTCA OR TGAG/CTCA 7 bp AP-1 consensus 7 bp AP-1 consensus

target gene

Figure 1.10 Binding of AP-1 to its DNA consensus sequence AP-1 (Jun/Jun or Jun/Fos dimers) modulates transcription by binding to the 7 bp DNA consensus sequence TGAG/CTCA located within the regulatory region of target genes.

This sequence is often referred to as the 12-O-tetradecanoyl-phorbol-13- acetate (TPA) response element (TRE), based on early studies identifying AP-1 as a TPA-inducible transcription factor that mediates transcriptional induction of the metallothionein and other genes (Angel et al., 1987; Lee et al., 1987b). The binding of the dimeric AP-1 complex to its specific target sequence influences the basal and stimulated expression of numerous target genes (Angel & Karin, 1991). While AP-1

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binding generally facilitates transcriptional activation and increased expression of the target gene, transcriptional repression and down-regulation of target gene expression by AP-1 has also been reported (Bushel et al., 1995; Schreiber et al., 1999).

Table 1.6 Possible dimeric combinations between AP-1 subunits, and between AP-1 proteins and other bZIP transcription factors

bZIP c-Jun JunB JunD c-Fos FosB Fra-1 Fra-2 Family

c-Jun AP-1 Yes Yes Yes Yes Yes Yes Yes JunB AP-1 Yes Yes Yes Yes Yes Yes Yes JunD AP-1 Yes Yes Yes Yes Yes Yes Yes c-Fos AP-1 Yes Yes Yes No No No No FosB AP-1 Yes Yes Yes No No No No Fra-1 AP-1 Yes Yes Yes No No No No Fra-2 AP-1 Yes Yes Yes No No No No ATF-2 ATF/CREB Yes Yes ATF-3 ATF/CREB Yes Yes Yes ATF-4 ATF/CREB Yes Yes Yes Yes ATF-a ATF/CREB Yes Yes Yes B-ATF ATF/CREB Yes Yes Nrf-1 CNC Yes Yes Yes Nrf-2 CNC Yes Yes cMaf Maf Yes Yes No Yes MafB Maf No Yes MafA Maf Yes Yes MafK Maf Yes Yes MafF Maf Yes Yes MafG Maf Yes Yes Nrl Maf Yes Yes C/EBPE C/EBP Yes Yes CHOP C/EBP Yes Yes Yes Meq bZIP Yes p21SNFT bZIP Yes Yes

Adapted from Herdegen & Leah, 1998 (Table 1, pg 377) and Chinenov & Kerppola, 2001 (Table 1, pg 2440). bZIP, basic region leucine zipper; AP-1, activator protein-1; ATF, activating transcription factor; CREB, cAMP responsive element binding protein; CNC, cap ’n’ collar; Nrf, NF-E2 (nuclear factor-erythroid 2)-related factor; Maf, macrophage-activating factor; Nrl, neural retina leucine zipper; C/EBP, CCAAT/enhancer binding protein; CHOP, C/EBP homologous protein 10; Meq, Marek’s disease virus EcoQ protein; p21SNFT, 21-kDa small nuclear factor isolated from T cells.

AP-1 complexes, composed of different dimeric combinations of Jun and Fos proteins, differ in their DNA binding and transcriptional activities (Angel & Karin, 1991; Ryseck & Bravo, 1991; Abate et al., 1993; Karin, 1995). For example, c-Jun homodimers exhibit strong DNA binding and transactivation activity, while JunB and JunD homodimers have a lower DNA binding affinity and are poor transactivators

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(Angel & Karin, 1991; Ryseck & Bravo, 1991; Herdegen & Leah, 1998). Heterodimerisation with JunB substantially lowers the capacity of c-Jun to bind and transactivate AP-1-responsive promoters (Chiu et al., 1989; Schütte et al., 1989; Ryseck & Bravo, 1991; Deng & Karin, 1993). On the other hand, c-Fos/c-Jun heterodimers are generally more potent than c-Jun homodimers as transactivators at AP-1-responsive promoters, due to increased stability and binding to the AP-1 consensus sequence (Halazonetis et al., 1988; Sassone-Corsi et al., 1988; Hirai et al., 1989; Allegretto et al., 1990; Ryseck & Bravo, 1991). Heterodimers between Jun and other Fos family proteins also have greater DNA binding affinity (Ryseck & Bravo, 1991), although transactivation potential is variable (Suzuki et al., 1991; Karin, 1995). The different binding and transactivation potential of the various AP-1 complexes, combined with the differential expression of AP-1 subunit proteins in response to extracellular stimuli, is a major mechanism by which AP-1 activity and the expression of AP-1-responsive genes is regulated (Mechta-Grigoriou et al., 2001; Shaulian & Karin, 2001). While the principal partners of Jun within the AP-1 complex are the Fos and Fos-related antigens (Fras) as discussed above (Vogt, 2001), it is now recognised that Jun and Fos proteins can also form stable dimers with certain members of other bZIP transcription factor families including the activating transcription factor (ATF)/cAMP responsive element binding protein (CREB) and macrophage-activating factor (Maf)/cap ‘n’ collar (CNC) families of bZIP proteins (Chinenov & Kerppola, 2001; Vogt, 2001). For example, c-Jun forms stable dimers with ATF-2, ATF-3 and ATF-4 (Hai & Curran, 1991), and c-Fos heterodimerises with MafB, MafK, MafF and MafG (Kataoka et al., 1994a; Kataoka et al., 1995). Thus, the potential for different dimeric Jun- and Fos-containing complexes in cells now exceeds 50 (Table 1.6; Chinenov & Kerppola, 2001). Dimerisation of Jun and Fos proteins with other bZIP proteins results in diversification of DNA binding specificity (Chinenov & Kerppola, 2001). Thus, while Jun-Jun and Jun-Fos dimers bind with high affinity to the TRE/AP-1 consensus sequence, dimers formed between Fos or Jun and the ATF proteins bind preferentially to the consensus sequence for the ATF/CREB proteins, named the cAMP responsive element (CRE; TGACGTCA), which differs from the AP-1 consensus by only one nucleotide (Hai & Curran, 1991; Shaulian & Karin, 2001). Maf homodimers recognise a binding sequence consisting of TGCtgaC half-sites (capital letters indicate the most conserved base pairs; Kataoka et al., 1994b; Kerppola & Curran, 1994), and Fos-Maf heterodimers have been found to preferentially interact with sites composed of juxtaposed AP-1 (TGAC) and Maf (TGCtgaC) half-sites (Kerppola & Curran, 1994).

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In addition to their interactions with other bZIP proteins, Fos and Jun can interact and bind to DNA in association with non-bZIP transcription factors such as nuclear factor of activated T cells (NFAT; Jain et al., 1993; McCaffrey et al., 1993), NF-NB (Stein et al., 1993; Yang et al., 1999) and Smad proteins (Liberati et al., 1999); interaction with these other DNA binding proteins can direct Fos and Jun proteins to promoter sequences that only slightly resemble the AP-1 consensus motif (van Dam & Castellazzi, 2001). In summary, selective dimerisation between Fos and Jun proteins, dimerisation with other bZIP proteins, and interaction with non-bZIP transcription factors alters the DNA binding site specificity of AP-1 proteins and is thought to provide flexibility in their potential to regulate target genes (Chinenov & Kerppola, 2001; van Dam & Castellazzi, 2001; Vogt, 2001).

1.9.2 Regulation of AP-1 Activity Unlike other response elements, such as those that mediate induction by glucocorticoid hormones, the AP-1/TRE element has basal activity in the absence of inducers like TPA, and regulates basal and stimulated expression of target genes (Angel et al., 1987; Lee et al., 1987a; Lee et al., 1987b; Angel & Karin, 1991). c-Jun is constitutively expressed in various cells, which has led to the suggestion that c-Jun homodimers may be the pre-existing AP-1 proteins in resting cells (De Cesare et al., 1995; Mechta- Grigoriou et al., 2001; Minet et al., 2001). The activity of AP-1 is induced by a diverse range of extracellular stimuli including the tumour promoter TPA and other phorbol esters, growth factors, cytokines, and neurotransmitters, as well as cellular stresses such as oxidative stress, hypoxia and UV irradiation (Angel & Karin, 1991; Karin, 1995; Rupec & Baeuerle, 1995; Wisdom, 1999; Shaulian & Karin, 2001). The regulation of AP-1 activity by extracellular stimuli is primarily achieved via modulation of the abundance and activity of the Fos and Jun proteins. The abundance of AP-1 proteins is primarily regulated via the transcriptional regulation of the fos and jun genes, while post-translational modification influences the DNA binding and transactivation capacity of both pre-existing and newly synthesised AP-1 proteins (Angel & Karin, 1991; Karin, 1995; Karin et al., 1997). The MAP kinases, which include the extracellular-regulated kinase (ERK), the c-Jun N-terminal kinase (JNK), and the p38 MAP kinase, are variously activated by stimuli known to induce AP-1 activity, and subsequently modulate both the expression and activity of the AP-1 subunits (Karin, 1995; Whitmarsh & Davis, 1996). ERKs are

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rapidly activated by growth factors and phorbol esters, while the JNK and p38 MAP kinases are activated by inflammatory cytokines and cellular stresses such as osmotic shock, heat shock and UV irradiation (Karin, 1995; Whitmarsh & Davis, 1996; Ip & Davis, 1998). Amongst the Fos and Jun members, the regulation of c-Fos and c-Jun expression and activity by the MAP kinases has been best studied. The ERK MAP kinases have been found to be important in mediating the rapid induction of c-fos expression in response to extracellular stimuli such as growth factors (Whitmarsh & Davis, 1996; Karin et al., 1997). The c-fos gene promoter contains a cis regulatory element called the serum-response element (SRE) that is crucial in mediating induction by ERKs (Gille et al., 1995; Karin et al., 1997). The SRE binds the serum-response factor (SRF), which subsequently recruits proteins known as ternary complex factors (TCFs). In response to mitogenic stimuli, Elk-1, one of several TCFs, is rapidly phosphorylated by the ERK MAP kinases (Gille et al., 1995; Karin, 1995; Whitmarsh & Davis, 1996). The phosphorylation of Elk-1 facilitates its association with the SRF and SRE and enhances its ability to activate transcription, thereby increasing c-fos expression (Figure 1.11; Gille et al., 1995; Karin, 1995; Whitmarsh & Davis, 1996). The SRE also mediates the induction of c-fos in response to stimuli such as UV irradiation and cytokines in a manner that is dependent on the JNK and p38 MAP kinases rather than the ERKs (Cavigelli et al., 1995; Whitmarsh et al., 1995; Raingeaud et al., 1996). The activation of JNK and p38 MAP kinases by these stimuli leads to the phosphorylation of Elk-1, and a subsequent increase in c-fos gene transcription (Figure 1.11; Cavigelli et al., 1995; Whitmarsh et al., 1995; Raingeaud et al., 1996; Karin et al., 1997). Amongst the MAP kinases, the JNK family is important in mediating an increase in the activity of pre-existing c-Jun subunits as well as increasing the abundance of c-Jun subunits (Karin, 1995; Karin et al., 1997). The c-jun gene promoter contains a sequence that resembles the TRE/AP-1 consensus, known as the c-Jun TRE, which plays a key role in the induction of c-jun by extracellular stimuli (Angel et al., 1988; van Dam et al., 1993; Karin et al., 1997). This c-Jun TRE is efficiently recognised by c-Jun/ATF-2 heterodimers (van Dam et al., 1993; Whitmarsh & Davis, 1996). The activation of JNK by external stimuli such as UV irradiation, cytokines and growth factors stimulates the JNK-dependent phosphorylation of serine residues within the transactivation domain of the c-Jun protein, increasing its ability to activate transcription (Dérijard et al., 1994; Whitmarsh & Davis, 1996). JNK also increases the activity of ATF-2 via a phosphorylation dependent mechanism (Gupta et al., 1995). In an auto- regulatory fashion, increased transactivation capacity of c-Jun, and also ATF-2,

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stimulates increased expression of the c-jun gene (Figure 1.11; Karin, 1995; Mechta- Grigoriou et al., 2001). Thus, the activation of JNK by external stimuli leads to an increase in both the activity and abundance of the c-Jun protein (Karin, 1995).

extracellular stimulus extracellular stimulus

ERK, JNK, p38 JNK

P P P P P P P TCF SRF c-Jun Elk-1 ATF-2

SRE c-Jun TRE

c-fos gene c-jun gene

Figure 1.11 Regulation of c-fos and c-jun gene transcription by MAP kinases Extracellular stimuli (e.g. growth factors, cytokines and cellular stress) variously activate the ERK, JNK and p38 mitogen-activated protein (MAP) kinases. ERK, JNK and p38 phosphorylate the ternary complex factor (TCF) Elk-1, which increases its transactivation potential and enhances its association with the serum-response element (SRE) in the c-fos gene promoter and its associated transcription factor, the serum- response factor (SRF), thus leading to enhanced c-fos expression. The c-jun gene promoter contains an important AP-1-like regulatory element, known as the c-Jun TRE (TPA response element) which binds c-Jun/ATF-2 heterodimers. JNK-dependent phosphorylation of c-Jun and ATF-2 enhances their transcriptional activities and leads to increased expression of c-jun. ERK, extracellular-regulated kinase; JNK, c-Jun N-terminal kinase.

1.9.3 Role of AP-1 in Cellular Physiology and Pathophysiology Given the activation of AP-1 and its subsequent modulation of downstream target genes by a wide range of stimuli, AP-1 is proposed to play a role in a number of physiological and pathophysiological processes (Angel & Karin, 1991; Wisdom, 1999; Shaulian & Karin, 2001; van Dam & Castellazzi, 2001). There is an abundance of evidence to indicate that the activation of AP-1 by growth factors and tumour promoters plays an important role in proliferation, differentiation and oncogenic transformation of various cell types (Angel & Karin, 1991; Leppä & Bohmann, 1999). For example, various AP-1 components, including c-Jun, are essential for embryonal development (van Dam & Castellazzi, 2001), and the proliferative response to growth factors has been associated with AP-1-dependent transcriptional activation of cyclin D1, which plays a key role in cell cycle progression (Wisdom et al., 1999; Mechta-Grigoriou et al., 2001).

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Aside from its role in cell proliferation and differentiation, AP-1 is implicated in the regulation of apoptosis (Karin et al., 1997; Shaulian & Karin, 2001). Whether AP-1 activity stimulates or protects against apoptosis appears to be cell type and stimulus- specific (Leppä & Bohmann, 1999; Shaulian & Karin, 2001). Thus, AP-1 is important in protecting cells from UV-induced apoptosis, with fibroblasts harboring a null mutation in either the c-fos or c-jun gene displaying increased apoptosis following UV irradiation (Schreiber et al., 1995; Wisdom et al., 1999). The mechanism by which AP-1 prevents UV-induced apoptosis is unclear, but c-Jun may repress activation of the tumour suppressor gene p53, which allows growth arrested cells to re-enter the cell cycle (Schreiber et al., 1999; Shaulian & Karin, 2001). c-Jun may also be protective against apoptosis induced by the inflammatory mediator TNFD (Wisdom et al., 1999). In contrast to these studies identifying an anti-apoptotic role for AP-1, a number of other studies have identified a pro-apoptotic role for AP-1, particularly within the nervous system (Shaulian & Karin, 2001). For example, over-expression of c-Fos enhances neuronal cell death (Smeyne et al., 1993). The over-expression of c-Jun in sympathetic neurons induced apoptosis, while a dominant negative c-Jun mutant prevented apoptosis induced by the withdrawal of nerve growth factor (Ham et al., 1995). Furthermore, the accumulation of c-Jun has been demonstrated in neurons undergoing hypoxia-induced apoptosis (Dragunow et al., 1993). These divergent effects of AP-1 on apoptosis are likely to be due to the differential activation of AP-1 subunits in response to different stimuli as well as interaction with other critical regulatory proteins (Leppä & Bohmann, 1999; Mechta-Grigoriou et al., 2001; Shaulian & Karin, 2001). In addition to UV irradiation, AP-1 is activated by other stress stimuli including hypoxia and oxidative stress, as well as by pro-inflammatory cytokines that are released in response to tissue injury, infection and trauma such as TNFD, IL-1 and IL-6; thus AP-1 is thought to play an important role in the adaptive response of the body to these stresses (Hattori et al., 1993; Rupec & Baeuerle, 1995; Wisdom, 1999). The activation of AP-1 by pro-inflammatory cytokines, and the demonstration that AP-1 target genes include the collagenase and IL-2 genes (Angel et al., 1987; Muegge et al., 1989), provided early indications that AP-1 is important in inflammation and the innate immune response (Shaulian & Karin, 2001). A particularly important response to injury and inflammation is activation of the acute phase response, in which the altered expression of acute phase proteins is a cellular response to prevent further tissue damage, destroy any infective organisms, and activate cellular repair processes (Baumann & Gauldie, 1994). AP-1 proteins are activated during the hepatic acute phase response and may interact with AP-1 binding elements within genes that encode

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acute phase proteins; thus, AP-1 is proposed to play a critical role in the regulation of the acute phase response to injury and infection (Hattori et al., 1993). AP-1 is activated in hypoxia and controls the expression of genes that regulate the cellular adaptation to a low oxygen environment (Norris & Millhorn, 1995; Rupec & Baeuerle, 1995; Mishra et al., 1998; Faller, 1999; Michiels et al., 2001). Thus, AP-1 activation is implicated in pathophysiological conditions that are characterised by low oxygen availability, such as ischaemic injury and tumourigenesis. Considered together, the AP-1 transcription factor complex is activated by a wide range of stimuli, and via regulation of numerous target genes, modulates cellular differentiation and proliferation, apoptosis, and the cellular response to injury, inflammation and ischaemia. The number of AP-1-responsive genes that are involved in these processes continues to expand. As discussed previously, the CYP2J2 protein has important functions in cellular physiology and pathophysiology, even though little is known about the regulation of the gene. A study conducted by Yang et al. (2001) suggested that CYP2J2 may be differentially expressed in hypoxia, and a number of potential AP-1 binding sites within the regulatory region of the CYP2J2 gene were identified in preliminary work underlying the present project. Given the well documented activation of AP-1 in hypoxia, the involvement of AP-1 in the regulation of CYP2J2 gene expression was explored. The results presented in this thesis establish a role for the AP-1 proteins c-Fos and c-Jun in the transcriptional control of CYP2J2 in normoxia and hypoxia.

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Chapter 2

Materials and Methods

2.1 Materials

2.1.1 Reagents and Chemicals All general chemicals and reagents were of analytical grade and were purchased from local suppliers (Sigma-Aldrich, Castle Hill, NSW, Australia; Astral Scientific, Gymea, NSW, Australia; Bacto Laboratories, Liverpool, NSW, Australia; Merck, Kilsyth, Vic., Australia). All solutions used for cell culture and RNA extraction were prepared using sterile water purchased from Baxter Healthcare (Old Toongabbie, NSW, Australia). All other buffers and solutions were prepared using MilliQ water. Tryptone, yeast extract and agar were purchased from Bacto Laboratories. Choramphenicol, bovine serum albumin (BSA), o-nitrophenyl-E-D-galactopyranoside (ONPG), Folins reagent, ethidium bromide, N-lauroylsarcosine, isoamyl alcohol, 2-mercaptoethanol, formaldehyde, thiazolyl blue (MTT) and dimethyl sulfoxide (DMSO) were obtained from Sigma-Aldrich. Phosphate buffered saline (PBS), guanidinium thiocyanate, 3-(N-morpholino)propane sulfonic acid (MOPS), acrylamide and bis-acrylamide were obtained from Amresco (Astral Scientific). Isopropyl-E-D-thiogalactopyranoside (IPTG) and ammonium persulfate (APS) were purchased from Promega (Annandale, NSW, Australia). Ampicillin, X-Gal and N,N,N’,N’-tetramethylethylenediamine (TEMED) were obtained from Progen Industries (Darra, QLD, Australia). Consumables (e.g. pipettor tips, microcentrifuge tubes, PCR tubes) were purchased from local suppliers (Interpath Services, Caringbah, NSW, Australia; Sarstedt, Technology Park, SA, Australia) and were autoclaved prior to use.

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2.1.2 Plasmids and Reagents for Molecular Biology The pGL3basic luciferase reporter vector, pCMV-E-galactosidase expression vector, pGEM-T Easy vector, Steady-Glo¥ Luciferase Assay System, E-galactosidase Enzyme Assay System with Reporter Lysis Buffer, RiboMAX¥ Large Scale RNA Production System and recombinant human c-Jun protein were purchased from Promega. Active Motif recombinant human c-Fos protein was from Bioscientific (Gymea, NSW, Australia). ABI Prism BigDye¥ Terminator Ready Reaction Mix and [32P]dCTP were obtained from PerkinElmer (Rowville, Vic., Australia). FuGENE¥ 6 transfection reagent and poly(2’-deoxyinosinic-2’-deoxycytidylic acid) [poly(dI-dC)] were purchased from Roche Diagnostics (Castle Hill, NSW, Australia). The Stratagene QuikChange¥ Site- Directed Mutagenesis Kit and Pfu Turbo polymerase were obtained from Integrated Sciences (Willoughby, NSW, Australia). The Epicentre MasterPure¥ Complete DNA and RNA Purification Kit was purchased from Astral Scientific. Expression plasmids for murine c-Fos and c-Jun were kindly provided by Dr. Kazuhiko Imakawa (University of Tokyo, Tokyo, Japan). Expression plasmids for human JunB, JunD, Fra-1 and Fra-2 were kindly provided by Dr. Michael Karin (University of California, San Diego, CA, U.S.A.). The Plasmid Midi Kit, QIAquick Gel Extraction Kit and OneStep RT-PCR Kit were purchased from Qiagen (Doncaster, Vic., Australia). The Quantum Prep Plasmid Miniprep Kit was purchased from Bio-Rad (Regents Park, NSW, Australia). The Megaprime¥ DNA Labelling System and ProbeQuant¥ G-50 Micro Columns were purchased from Amersham Biosciences (Castle Hill, NSW, Australia). T4 DNA , RQ1 RNase-free DNase, Recombinant RNasin Ribonuclease Inhibitor, Taq DNA polymerase and deoxynucleotide triphosphates (dNTPs) were purchased from Promega. Restriction enzymes were from Roche Diagnostics unless otherwise specified and oligonucleotides were synthesised by Geneworks (Adelaide, SA, Australia).

2.1.3 Reagents for Cell Culture The HepG2 human hepatoma cell line was purchased from American Type Culture Collection (A.T.C.C.; Manassas, VA, U.S.A.). Cell culture media, L-glutamine, penicillin/streptomycin antibiotic mix and fetal calf serum were purchased from Thermo Trace (Noble Park, Vic., Australia). Cells were grown in Nunc sterile 6-well plates, 25 cm3 and 75 cm3 flasks purchased from Medos (Gladesville, NSW, Australia).

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2.1.4 Reagents for Protein Electrophoresis and Immunoblotting Polyclonal rabbit anti-c-Fos and anti-c-Jun antibodies, polyclonal goat anti-ubiquitin (Ub) antibody and monoclonal mouse anti-E-actin antibody were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, U.S.A.). Polyclonal rabbit anti-(rat CYP2J4) antibody, which is cross-reactive with human CYP2J2, was generously provided by Dr. Qing-Yu Zhang (Wadsworth Centre, New York State Department of Health, New York, NY, U.S.A.). Secondary anti-(rabbit IgG) and anti-(mouse IgG) antibodies, and protease inhibitors were purchased from Sigma-Aldrich. Tween 20, ECL Western blotting detection reagents and autoradiography film were from Amersham Biosciences. Schleicher and Schuell Protran nitrocellulose transfer membrane was purchased from Medos and Fermentas Prestained Protein Molecular Weight Markers were purchased from Progen Industries.

2.2 General Molecular Techniques

2.2.1 Preparation of Competent E.coli Cells and Transformation of Plasmids All plasmids used in this study (i.e. CYP2J2 promoter reporter constructs and expression plasmids for the various AP-1 protein subunits) were transformed into competent E.coli (strain JM109) cells for the purposes of storage, propagation and purification for downstream applications. Competent cells were prepared from E.coli JM109 cells (Promega) according to the following protocol. Luria-Bertani (LB) medium (10 g/L tryptone, 5 g/L yeast extract, 5 g/L sodium chloride, pH 7.0) was inoculated with a single colony of E.coli JM109 and grown overnight at 37qC with shaking. The following morning, 100 mL of LB medium was inoculated with 1 mL of the overnight starter culture, and allowed to grow at 37qC with shaking until the OD590 of the culture reached 0.5. The culture was then cooled on ice for 10 min and the cell pellet was collected by centrifugation of the culture at 2500g for 5 min at 4qC. The pellet was gently resuspended in 20 mL ice-cold CaCl2 (50 mM), re-pelleted by centrifugation at 2500g for 5 min at 4qC and then resuspended in 4 mL ice-cold CaCl2 (50 mM) and 1.26 mL 50% (v/v) glycerol. This competent cell preparation was aliquoted, frozen slowly at -20qC, and then stored at -80qC.

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For transformation of plasmids, 100 PL of competent E.coli JM109 cells were thawed on ice and gently combined with plasmid DNA (1 to 10 PL) in pre-chilled sterile PCR tubes. The plasmid-cell mixture was incubated on ice for 20 min, subjected to heat shock at 42qC for 50 sec and then incubated on ice for a further 2 min. Sterile 10 mL tubes containing 900 PL of SOC medium (2 g/100 mL tryptone, 0.5 g/100 mL yeast extract, 10 mM sodium chloride, 2.5 mM potassium chloride, 10 mM magnesium chloride, 10 mM magnesium sulphate and 20 mM glucose) were inoculated with the plasmid-cell mixture and incubated for 90 min at 37qC with shaking. The cell pellet was collected by centrifugation, resuspended in a small volume of SOC medium (<100 ȝL) and spread onto LB-agar plates (10 g/L tryptone, 5 g/L yeast extract, 5 g/L sodium chloride, 15 g/L agar, pH 7.0) supplemented with the antibiotic ampicillin (100 Pg/mL) for selection of successfully transformed cells (all plasmids used carried an ampicillin gene for selection). Plates were incubated overnight at 37qC. Single colonies were picked from transformation plates, inoculated into 3 mL of LB medium supplemented with 100 ȝg/mL ampicillin (LB-Amp) and cultured overnight at 37qC with shaking. Glycerol stocks for long-term storage of the plasmid-transformed cells were made by combining 0.5 mL of overnight culture with 0.5 mL sterile 50% (v/v) glycerol; this glycerol mixture was aliquoted and stored at -80qC. The overnight culture remaining after glycerol stock preparation was used to confirm the identity of the transformed plasmid; plasmid DNA was isolated and purified from E.coli JM109 cells using the Quantum Prep Plasmid Miniprep Kit (described in section 2.2.2) and confirmed by DNA sequencing or digestion with suitable restriction enzymes.

2.2.2 Culture of E.coli Cells and Purification of Plasmids for Transfection Plasmid DNA was isolated from transformed E.coli cultures using either the Bio-Rad Quantum Prep Plasmid Miniprep Kit or the Qiagen Plasmid Midi Kit. The Miniprep Kit was used in accordance with the manufacturer’s instructions to extract plasmid DNA on a small scale for the purpose of sequencing and restriction enzyme digestion. Briefly, plasmid-transformed E.coli JM109 cells were inoculated into 2 to 3 mL of LB-Amp, and the culture was grown overnight at 37qC with shaking. The overnight culture (1 to 2 mL) was centrifuged at 15800g for 30 sec, the supernatant discarded and the cell pellet resuspended in 200 PL of Cell Resuspension Solution. Cells were lysed by gentle mixing with Cell Lysis Solution (250 PL), and then the lysis reaction was stopped by the addition of Neutralisation Solution (250 PL). The cellular debris was pelleted by centrifugation at 15800g for 5 min and the plasmid-containing

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supernatant was mixed with 200 PL of Quantum Prep Matrix. Supernatant-Matrix mixtures were applied to Quantum Prep Kit Spin Filters, and centrifugation at 15800g for 30 sec facilitated binding of plasmid DNA to the Spin Filter. The bound plasmid DNA was washed twice with 500 ȝL Wash Buffer, and then eluted in 100 PL sterile water by centrifugation of the Spin Filter at 15800g for 1 min. Concentration of plasmid DNA was measured by spectrophotometry (Cary 300 Bio spectrophotometer, Varian) at a wavelength of 260 nm. Concentrations were calculated from the assumption that 50 ȝg/mL of double-stranded DNA and 30 ȝg/mL of single-stranded DNA has an absorbance of 1. To obtain high yields of plasmid DNA of suitable purity for transient transfection, the Qiagen Plasmid Midi Kit was used. Glycerol stocks of plasmid-transformed E.coli JM109 cells were streaked onto LB-agar plates supplemented with ampicillin and grown overnight at 37qC. Single colonies were picked and inoculated into 2 mL LB-Amp. Starter cultures were grown at 37qC with shaking for 8 hr and subsequently inoculated into 25 mL of LB-Amp at a dilution of 1:500. These cultures were then grown overnight at 37qC with vigorous shaking. The protocol for the culture of transformed E.coli JM109 cells harboring the low-copy c-Fos and c-Jun expression plasmids was modified to maximise plasmid yield. For these plasmids, 2 mL LB-Amp starter cultures were inoculated, incubated overnight at 37qC with shaking, and subsequently used to inoculate 100 mL LB-Amp at a dilution of 1: 166. Cultures were incubated at 37qC with vigorous shaking until the culture reached an OD600 of 0.5 to 0.6 (approximately 3.5 hr); at this point chloramphenicol was added to the culture at a final concentration of 170 Pg/mL, and the cultures were incubated at 37qC with shaking for a further 16-20 hr. Both the high- and low-copy plasmids were purified from culture using the Qiagen Plasmid Midi Kit according to the manufacturer’s instructions. Cultures were centrifuged at 6000g for 15 min at 4qC to collect the bacterial cells. The supernatant was discarded and the bacterial pellet was resuspended in 4 mL Buffer P1 (50 mM Tris-Cl, pH 8.0, 10 mM EDTA, 100 Pg/mL RNase A). Buffer P2 (4 mL; 200 mM NaOH, 1% (w/v) SDS) was added to lyse the cells, and the lysis reaction was stopped after 5 min with the addition of 4 mL Buffer P3 (3 M potassium acetate, pH 5.5). Samples were transferred to QIAfilter cartridges and incubated at room temperature for 10 min so that proteins, genomic DNA and detergent could form a layer above the sample. Following incubation, the sample was passed though the QIAfilter; the plasmid DNA collected as the filtrate whereas the protein, genomic DNA and detergent mixture was retained by the filter. The lysate was applied to a QIAGEN-tip that had been pre- equilibrated with Buffer QBT (4 mL; 750 mM NaCl, 50 mM MOPS, pH 7.0, 15% (v/v)

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isopropanol, 0.15% (v/v) Triton X-100), and allowed to pass through the resin by gravity flow, leaving the plasmid DNA bound to the resin. Contaminants were removed from the bound plasmid preparation by the application and passage of two consecutive volumes of Buffer QC (10 mL; 1 M NaCl, 50 mM MOPS, pH 7.0, 15% (v/v) isopropanol), and then the plasmid DNA was eluted from the resin in 5 mL Buffer QF (1.25 M NaCl, 50 mM Tris-Cl, pH 8.5, 15% (v/v) isopropanol). The plasmid DNA was precipitated by the addition of 0.7 volumes of isopropanol, and pelleted by centrifugation at 15000g for 30 min at 4qC. Plasmid DNA was washed with 2 mL 70% ethanol, and re-pelleted by centrifugation at 15000g for a further 10 min. Following air- drying for 10 min, the plasmid DNA pellet was resuspended in 200 ȝL Tris-disodium ethylenediaminetetra-acetate (EDTA) buffer (TE; 10mM Tris-Cl, pH 8.0, 1 mM EDTA, pH 8.0), and stored at -20qC until used in transient transfections. The concentration of plasmid DNA preparations were measured spectrophotometrically at 260 nm.

2.2.3 DNA Sequencing Sequencing of DNA was performed using the ABI Prism BigDye¥ Terminator Ready Reaction Mix (PerkinElmer) and sequencing primers shown in Table 2.1. A reaction mixture containing 8 ȝL of the BigDye Terminator Ready Reaction Mix, 1 ȝg of plasmid DNA, and 5 pmoles of sequencing primer was prepared in a final volume of 20 ȝL. Reactions were amplified by PCR in a thermal cycler (GeneAmp PCR system 2400, PerkinElmer) using a cycling program that consisted of initial denaturation at 96qC for 3 min, followed by 25 cycles of denaturation at 96qC for 10 sec, primer annealing at 50qC for 5 sec, and extension at 60qC for 4 min. Following amplification, an ethanol/potassium acetate precipitation was performed to remove excess unincorporated BigDye terminators prior to sequencing. Amplification reactions were mixed with 50 ȝL of 95% (v/v) ethanol and 2 ȝL 5M potassium acetate (pH 8.0) and allowed to incubate for 15 min at room temperature. The precipitated DNA was pelleted by centrifugation at 20800g for 20 min, and then washed with 250 ȝL 70% (v/v) ethanol. DNA was re-pelleted after washing by a further centrifugation at 20800g for 5 min, and then dried by heating at 90qC for 1 min. Sequencing of purified DNA samples were performed by the Automated DNA Analysis Facility, University of New South Wales. The Genomatix MatInspector Professional consensus sequence identification program (www.genomatix.de/matinspector.html; Quandt et al., 1995) was used to detect potential transcription factor consensus recognition sites within the CYP2J2 upstream regulatory region.

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Table 2.1 Sequencing primers

Oligonucleotide Sense/ Location Sequence (5’ – 3’) Name Antisense

CYP2J2 Sequencing Primers Sp1 sense CYP2J2 (-1989) TGGAACCTCTATCTCAT Sp7 antisense CYP2J2 (-1769) CAAATGAAGGCTACAGT Flank 13 antisense CYP2J2 (-1706) CAGATCAACGTGACAGAA Sp5 sense CYP2J2 (-1515) TTCTGTGGGGTTTCTCA Flank 14 antisense CYP2J2 (-1395) GCTCATTCCCGTATCAG Sp3 antisense CYP2J2 (-1155) GGTCCAGAGAGAAATACTA Flank 10 sense CYP2J2 (-968) TGTAGTCCCAGCTACATG Flank 12 antisense CYP2J2 (-861) GACAGAGTCTTGCTCTGT Flank 11 sense CYP2J2 (-449) TTCCTCAACAGCAAGATG Sp4 antisense CYP2J2 (-400) CAAAATTATGAAGGAGAAG Sp2 antisense CYP2J2 (-229) AGAGCGCCACAGACATT -2161 sense CYP2J2 (-181) TGGCCTTTTCTGAGACCGGTGCGTG T7 sense CYP2J2 (+42) AGTGGTCCATCCTCGGAC Sequencing Primers Flanking Vector Cloning Sites RVprimer3 sense pGL3basic CTAGCAAAATAGGCTGTCCC GLprimer2 antisense pGL3basic CTTTATGTTTTTGGCGTCTTCCA M13 sense pGEM-T GTTTTCCCAGTCACGAC

CYP2J2 sequencing primers were synthesised by Geneworks. Primer locations within the CYP2J2 gene are numbered relative to the translation start site (+1). The sequencing primers binding within the pGL3basic (RVprimer3 and GLprimer2) and pGEM-T (M13) vectors were purchased from Promega.

2.2.4 Electrophoresis and Purification of DNA Where necessary, DNA samples were combined with a small volume (2 - 5 ȝL) of 6X DNA loading buffer (0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cyanol, 50% (v/v) glycerol) and electrophoresed on 1% agarose gels containing 0.1 ȝg/mL ethidium bromide. Electrophoresis was carried out in TBE buffer (90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA) at 110V for 50 min, and DNA was visualised under UV light on a transilluminator (Gel Doc 2000, Bio-Rad). For purification following restriction digestion, DNA fragments were separated by electrophoresis at 100V for 1 – 2 hr on a 2% low-melting agarose gel in TAE buffer (40 mM Tris-acetate, 1mM EDTA) and visualised under UV light. Agarose slices containing the required DNA fragment were excised from the gel with a sterile scalpel blade, and DNA was purified using the QIAquick Gel Extraction kit (Qiagen) in accordance with the manufacturer’s instructions. Briefly, the gel slice was weighed and placed into a sterile 1.5 mL microcentrifuge tube. Three volumes of Buffer QG were added, and the gel slice was melted by incubation at 50qC for 10 min with intermittent vortexing. Once the agarose slice was completely melted, the sample was mixed with one volume of isopropanol and loaded onto a QIAquick spin column. DNA present

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within the sample was bound to the silica gel membrane of the column by centrifugation at 15800g for 1 min. After the application of Buffer QG (0.5 mL), the column was centrifuged at 15800g for 1 min to remove remaining traces of agarose. Buffer PE (0.75 mL) was then applied to the column, and after 5 min the column was centrifuged at 15800g for 1 min. The Buffer PE washing step was repeated, and then the bound and purified DNA was eluted from the column by incubation with 50 PL Buffer EB (10 mM Tris-HCl, pH 8.5) followed by centrifugation at 15800g for 1 min. Purified DNA was quantified by measuring absorbance at 260 nm and stored at -20qC until used in downstream applications.

2.3 Preparation of CYP2J2 Promoter Reporter Constructs

An approximate 2.4 kb fragment of the CYP2J2 gene consisting of 2341 bp of the sequence upstream of the translation start site (i.e. 2341 bp of the 5’-regulatory region) and 98 bp of the sequence downstream of the translation start site (i.e. 98 bp of exon 1) was isolated from a human genomic library using the Clontech Human GenomeWalker¥ Kit (Progen Industries) and subcloned into the pGL3basic luciferase reporter vector to generate the plasmid 2J2 (-2341/+98). The 5’-truncated constructs 2J2 (-1894/+98), 2J2 (-1228/+98), 2J2 (-574/+98) and 2J2 (-152/+98) were generated by restriction enzyme digestion and re-ligation from 2J2 (-2341/+98) using available sites within the CYP2J2 5’-flanking sequence. 2J2 (-2341/+98) was digested, respectively, with EcoRV to create 2J2 (-1894/+98), PvuII to create 2J2 (-1228/+98), NheI to create 2J2 (-574/+98) and SmaI to create 2J2 (-152/+98). The 2.4 kb fragment of CYP2J2 and the 4 deletion constructs were gifts from Dr. Shi-Hua Xiang of this laboratory. All other constructs were prepared by the author. The CYP2J2 promoter construct 2J2 (-152/+98) contains a SmaI/XmaI recognition sequence at -152 bp. Thus, to prepare the shorter CYP2J2 promoter deletion constructs, 2J2 (-122/+98), 2J2 (-82/+98) and 2J2 (-49/+98), SmaI/XmaI recognition sites were introduced at appropriate positions within 2J2 (-152/+98) using the Stratagene QuikChange¥ Site-Directed Mutagenesis Kit in accordance with the manufacturer’s instructions. Briefly, a complementary pair of primers were designed to introduce SmaI/XmaI recognition sequences in 2J2 (-152/+98) [Table 2.2; Geneworks]. PCR reaction mixes containing 50 ng plasmid template, 150 ng each of the appropriate sense and antisense oligonucleotide, 1X Pfu Turbo reaction buffer, 200 ȝM dNTP mix

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and 2.5U Pfu Turbo polymerase were prepared in 0.2 mL PCR tubes, and cycling was carried out in a thermal cycler (GeneAmp PCR system 2400, PerkinElmer). An initial cycle of denaturation at 95qC for 30 sec was followed by 18 cycles of denaturation at 95qC for 30 sec, annealing at 55qC for 1 min and extension at 68qC for 13 min. Following PCR, the amplification reactions were incubated with 10U of DpnI (Promega) at 37qC for 1 hr to digest the parental methylated DNA. The DpnI-digested reactions were further incubated with 3U T4 DNA ligase at room temperature for 1 hr to facilitate ligation of the newly synthesised plasmid DNA. 7 ȝL of this final reaction mixture was transformed into competent E.coli JM109 cells; colonies resulting from the transformation were picked and grown at 37qC overnight in LB-Amp medium, and then plasmid preparations were obtained using the Quantum Prep Plasmid Miniprep Kit as outlined previously (section 2.2.2). The successful introduction of SmaI/XmaI recognition sites at either -122, -82 or -49 bp within 2J2 (-152/+98) was confirmed by sequencing using Promega’s pGL3basic sequencing primers RVprimer3 (sense) and GLprimer2 (antisense) as outlined in section 2.2.3. Incubation of the resultant plasmids (approximately 1 ȝg) with 5U of XmaI at 37qC for 1 hr facilitated excision of the CYP2J2 promoter sequence from -152 to -123 bp, -152 to -83 bp, or -152 to -50 bp within 2J2 (-152/+98) to create, respectively, 2J2 (-122/+98), 2J2 (-82/+98) and 2J2 (-49/+98). The digested plasmid preparations were electrophoresed (1% low-melt agarose TAE gel; 70V for 1 to 2 hr) to separate the excised sequence from the larger fragment containing the residual promoter – plasmid sequence. Plasmid DNA fragments were excised from agarose gels using a sterile scalpel blade, purified using the QIAquick Gel Extraction Kit outlined previously (section 2.2.4) and re-ligated using T4 DNA ligase. The deleted and re-ligated plasmids were transformed and confirmed by sequencing using the RVprimer3 and GLprimer2 sequencing primers. Mutation constructs were similarly prepared using the QuikChange¥ Site- Directed Mutagenesis Kit. Complementary primer pairs incorporating mutated nucleotides were designed and synthesised as shown in Table 2.2. The mutations were incorporated into the template plasmid [either 2J2 (-152/+98) or 2J2 (-122/+98)] by PCR using reaction and cycling conditions identical to those described above for the preparation of deletion constructs. Following the PCR, the parental methylated template DNA was digested with DpnI, and the newly mutated plasmid was ligated with T4 DNA ligase as described above. The mutated plasmids were transformed into competent E.coli JM109 cells, and colonies picked and grown overnight for propagation. The plasmid DNA was then purified using the Quantum Prep Plasmid

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Miniprep Kit, and sequenced to confirm the successful incorporation of mutated nucleotides using the RVprimer3 and GLprimer2 sequencing primers. Purified and concentrated preparations of the CYP2J2 promoter reporter constructs for use in transient transfection into HepG2 cells were prepared using the Qiagen Plasmid Midi Kit as described in section 2.2.2.

Table 2.2 Sequences of oligonucleotides used in site-directed mutagenesis to create CYP2J2 promoter deletion and mutation constructs

Oligonucleotide Sense/ Sequence (5’ – 3’) Name Antisense

For Creation of CYP2J2 Mutation Constructs 2J2-mt-(-7/+1) sense AGCAGGAGGACGTAAAAAAAATGCTCGCGGCGAT 2J2-mt-(-7/+1) antisense ATCGCCGCGAGCATTTTTTTTACGTCCTCCTGCT 2J2-mt-(-56/-63) sense GAGGCGGGGCGGGAAAAAAAACCTGCTGGGACCG 2J2-mt-(-56/-63) antisense CGGTCCCAGCAGGTTTTTTTTCCCGCCCCGCCTC 2J2-mt-(-148/-142) sense GCGTGCTAGCCCGCTTAAAAAGCGCCTGGCATC 2J2-mt-(-148/-142) antisense GATGCCAGGCGCTTTTTAAGCGGGCTAGCACGC 2J2-mt-(-141/-134) sense AGCCCGGGAATCCTTTTAAAAGCATCTTCGCAGG 2J2-mt-(-141/-134) antisense CCTGCGAAGATGCTTTTAAAAGGATTCCCGGGCT 2J2-mt-(-133/-126) sense AATCCAGCGCCTGAAAAAAAAGCAGGGTGCTGCG 2J2-mt-(-133/-126) antisense CGCAGCACCCTGCTTTTTTTTCAGGCGCTGGATT 2J2-mt-(-106/-96) sense GGTGCTGCGAAGGGGGAAAAAAAAAAGCGGGGCACGGCTGG 2J2-mt-(-106/-96) antisense CCAGCCGTGCCCCGCTTTTTTTTTTCCCCCTTCGCAGCACC For Creation of CYP2J2 Deletion Constructs 2J2-mt-(-122/Sma,) sense CTGGCATCTTCGCACCCGGGTGCTGCGAAGG 2J2-mt-(-122/Sma,) antisense CCTTCGCAGCACCCGGGTGCGAAGATGCCAG 2J2-mt-(-82/Sma,) sense CGGGGCACGGCTCCCGGGAGCGAGGCG 2J2-mt-(-82/Sma,) antisense CGCCTCGCTCCCGGGAGCCGTGCCCCG 2J2-mt-(-49/Sma,) sense CCGTCGCCTGCTCCCGGGACCGCCGCC 2J2-mt-(-49/Sma,) antisense GGCGGCGGTCCCGGGAGCAGGCGACGG

Mutated or introduced nucleotides are underlined

2.4 Cell Culture

2.4.1 Experimental Conditions All manipulations were performed in a laminar flow hood (Biological Safety Cabinet Class II, Email Air Handling) which was sterilised by UV light for 20 min and wiped with 70% ethanol prior to use. All solutions were pre-heated to 37qC and the vessels were sprayed with 70% ethanol before being placed in the hood. Flasks and plates used to grow cells were sterile and of tissue culture grade.

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2.4.2 Cell Line and Culture Conditions The HepG2 cell line was cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal calf serum (heat-inactivated at 56qC for 30 min), 1% L-glutamine, 1% penicillin/streptomycin antibiotic mix, 20 mM N-(2-hydroxyethyl) piperazine-N’-(2-ethanesulfonic acid) [HEPES] and 26 mM sodium hydrogen bicarbonate (complete DMEM). Cells were cultured at 37qC in a humidified incubator using a mixture of 95% air and 5% CO2 and were passaged twice weekly as described in section 2.4.3. Cells at passage 4 were used for all experiments.

2.4.3 Passaging of Cells Cells were passaged when they reached approximately 70% confluence. At this point, media was removed by aspiration with a sterile pasteur pipette and vacuum line, the cell monolayer was washed once with 10 mL phosphate buffered saline (PBS; 137 mM NaCl, 2 mM KCl, 10 mM phosphate buffer, pH 7.4) and subsequently detached from the vessel surface by incubation at 37qC (in 95% air and 5% CO2) with 2 mL trypsin (0.4%) for 9 min. Following trypsinisation, cells were immediately resuspended in 12 mL complete DMEM to inactivate trypsin and prevent cell membrane disruption. The cells were then sub-cultured at a ratio of 1:7.5 by transferring 4 mL aliquots of resuspended cells into new flasks with fresh complete DMEM (30 mL).

2.4.4 Hypoxic Treatment of Cells and Harvesting of Cells for Extraction of Total RNA, Total Cell Lysates and Nuclear Extracts For isolation of RNA, cell lysates and nuclear extracts, HepG2 cells were seeded in 75 cm35 flasks at a density of 2x10 /mL for 48 hr prior to hypoxic exposure. For hypoxic incubations, flasks were transferred to an oxygen-regulated incubator containing a mixture of 94% N2, 5% CO2 and 1% O2 for 16 hr or 40 hr. Control (normoxic) flasks were cultured for the same time period in 21% O2 (95% air and 5% CO2). For reoxygenation, flasks were removed from 1% O2 and returned to 21% O2 for a further 30 min. Following the defined treatments, cells were harvested for preparation of total RNA, total cell lysates and nuclear extracts as follows. Media was removed by aspiration, the cell monolayer was washed with 10 mL PBS and detached by incubation with 2 mL trypsin (0.4%). The detached cells were resuspended in 12 mL complete DMEM and the cell pellet was collected by centrifugation at 330g for 5 min at 4qC. To remove remaining traces of media and trypsin, the resultant cell pellet was resuspended in 10 mL ice-cold PBS, centrifuged again at 330g for 5 min at 4qC and the

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supernatant discarded. Cell pellets were then processed according to the protocols for extraction of total RNA (section 2.5.2), total cell lysates (section 2.7.1) or nuclear protein (section 2.9.1).

2.4.5 Assessment of Cell Viability The MTT assay was used to assess cell viability following hypoxic exposure. Thiazolyl blue [MTT; 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide] is taken up and reduced within the mitochondria of living cells (Mosmann, 1983). The dark blue products of MTT reduction were dissolved in DMSO (1 mL) and their absorbance measured at 540 nm. For MTT cell viability assays, HepG2 cells were seeded (in triplicate) at 2 x 105 cells/mL in 6-well plates. Following a 48 hr incubation under normal oxygen conditions

(21% O2; 95% air and 5% CO2), cells were transferred to an oxygen-regulated incubator and subjected to hypoxia for 16 or 40 hr (94% N2, 5% CO2 and 1% O2).

Normoxic controls were incubated at 21% O2 for 16 hr or 40 hr. MTT (625 Pg/250 PL) was added, and cells were incubated for 2 hr at 37qC. The cell supernatant was aspirated and DMSO (1 mL) was added to each well. Plates were shaken vigorously at room temperature for 30 min and the absorbance at 540 nm was measured. The absorbance of cells cultured in normoxia (21% O2) was taken to represent

100% viability; the viability of cells cultured in hypoxia (1% O2) was expressed as a percentage of normoxic control. In addition to assaying for cell viability immediately following hypoxic exposure, viability was determined after allowing a period of 3-6 days of recovery from hypoxic exposure. For these experiments, cells were seeded at 2 x 105 cells/mL in 75 cm3 flasks and incubated at 21% O22for 48 hr prior to exposure to 1% O for 16 hr and 40 hr.

Normoxic controls were incubated at 21% O2 for 16 hr and 40 hr. At the end of this time, cells were trypsinised, resuspended in fresh complete DMEM and counted (Brightline haemocytometer, Sigma). Cells were seeded at 2 x 105 cells/mL in 6-well plates and incubated for 3-6 days at 21% O2 to enable recovery from treatments. Cells exposed to hypoxia for sub-toxic periods grew normally during this period, while cells exposed to hypoxia for toxic periods exhibited slow growth and poor recovery. After the 3-6 day recovery period, plates were removed from the incubator and viability was assessed by MTT assay as described above.

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2.5 RNA Extraction

2.5.1 Experimental Conditions To ensure an RNase free environment, all glassware used for the preparation of RNA extraction solutions was baked overnight at 190qC. Sterile (Baxter) water was used in the preparation of all solutions and all consumables (e.g. pipettor tips) were autoclaved at 121qC for 20 min prior to use. Unless otherwise stated, all procedures were carried out on ice.

2.5.2 RNA Extraction Procedure Total RNA was extracted from cultured HepG2 cells using the acid guanidinium thiocyanate/phenol method of Chomczynski & Sacchi (1987). Following the defined treatments, HepG2 cells were harvested as described in section 2.4.4 and the resultant cell pellets were immediately resuspended in 3 mL ice-cold solution D (4 M guanidinium thiocyanate, 25 mM sodium citrate, pH 7.0, 0.5% (w/v) N-lauroylsarcosine, 0.1 M 2-mercaptoethanol). The DNA was sheared by five passes through a 23 gauge needle. The resultant suspension was treated with 0.3 mL 2 M sodium acetate, pH 4, 3.0 mL water-saturated phenol and 0.6 mL of chloroform-isoamyl alcohol mixture (49:1 v:v). After the addition of each component the samples were mixed gently by inversion, and after the addition of the final component, the samples were mixed vigorously by hand for 10 sec and placed on ice for 15 min. Samples were centrifuged at 12000g for 25 min at 4qC to facilitate separation of RNA into the aqueous phase, DNA into the interphase, and proteins into the phenolic phase. The upper aqueous layer containing the RNA was transferred to a fresh tube and combined with 8 mL ice-cold 100% ethanol, mixed gently, and stored overnight at -80qC to precipitate the RNA. The following morning, samples were centrifuged at 12000g for 25 min at 4qC and the RNA pellet was then resuspended in 0.3 mL ice-cold solution D. This sample was transferred to a 1.5 mL microcentrifuge tube, mixed with 0.6 mL 100% ethanol and the RNA was re-precipitated overnight at -80qC. The RNA was collected by centrifugation at 20800g for 15 min at 4qC. The supernatant was removed and the cell pellet was washed by resuspension in 0.3 mL ice-cold 70% ethanol. After centrifugation at 20800g for 10 min at 4qC, the supernatant was discarded and the RNA pellet was air-dried at room temperature for 20 – 25 min. The dried pellet was resuspended in 50 PL sterile water and an aliquot removed for

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quantitation and electrophoresis (to assess RNA quality). RNA samples were then aliquoted and stored at -80qC until required in experiments.

2.5.3 Quantitation of RNA by Spectrophotometry Extracted RNA was quantified by spectrophotometry at 260 nm (40 Pg/mL of RNA has an absorbance of 1 at 260 nm). Samples were also read at 280 nm, and the ratio of the absorbances at 260 nm to 280 nm used as an indication of RNA purity (high quality preparations of RNA generally have an A260/A280 ratio of 1.6 – 2.0, whereas samples containing impurities give a lower ratio). Only high quality RNA preparations were used for RT-PCR.

2.5.4 Electrophoresis of RNA Samples To confirm integrity before use in RT-PCR, an aliquot of each extracted RNA sample was electrophoresed in a denaturing 1% agarose gel containing 5 mL of 10X MOPS buffer [0.2 M 3-(N-morpholino)propane sulfonic acid (MOPS), 80 mM sodium acetate, 0.01 M EDTA, pH 7.0] and 2.5 mL formaldehyde. For preparation prior to electrophoresis, 5 ȝg of RNA was diluted in water to a final volume of 10 ȝL and combined with 2 PL 10X MOPS and 4 PL formaldehyde. Following denaturation of RNA by heating at 65qC for 10 min, samples were cooled on ice and mixed with 2 PL loading dye (0.25% bromophenol blue, 25% sucrose) and 1 PL 1 μg/μL ethidium bromide. Samples were loaded onto the gel, electrophoresed in 1X MOPS (20 mM MOPS, 8 mM sodium acetate, 1 mM EDTA) at 90V for 1.5 hr and the RNA visualised under UV light (Gel Doc 2000 system, Bio-Rad). Samples of high quality, non-degraded RNA exhibited two distinct bands on the gel, corresponding to the most abundant cellular RNA species: 18S and 28S ribosomal RNA, whilst samples containing degraded RNA appeared as smears. Only high quality, non-degraded samples were used for RT-PCR.

2.6 Reverse Transcriptase Polymerase Chain Reaction (RT-PCR)

2.6.1 Experimental Conditions The extreme sensitivity of PCR amplification to even very small quantities of foreign DNA necessitated a number of precautions:

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1. Manipulations relating to RT-PCR, such as aliquoting of RNA and reagents, and preparation of reaction mastermixes, were performed in a laminar flow hood in a different laboratory. 2. Plugged pipettor tips (Art barrier tips, Interpath Services) were used throughout. The aerosol barrier prevents cross-contamination of samples during the dispensing of solutions. 3. Suitable controls were included during RT-PCR; template and RT negative controls were included for every RT-PCR experiment. The template negative control contained no template RNA (water was substituted instead) and was used to detect possible contamination of the reagents used in the RT-PCR. The RT negative control did not undergo reverse transcription and was used to detect contamination of the RNA with genomic DNA.

2.6.2 Semi-quantitative RT-PCR of CYP2J2, c-Fos and c-Jun Semi-quantitative RT-PCR was performed by co-amplification of CYP2J2, c-Fos or c-Jun with E-actin (internal control) using the primer sequences listed in Table 2.3.

Table 2.3 Sequences of primers for semi-quantitative and competitive RT-PCR Oligonucleotide Sense/ Sequence (5’ – 3’) Antisense

CYP2J2 forward sense GGACCCCACCAAACTCTCTTCAGCAA CYP2J2 reverse antisense ATAAAGCAGAGCCCATCGCAGAGTTG c-Fos forward sense ATGTTCTCGGGCTTCAACGCAGA c-Fos reverse antisense CAGTGACCGTGGGAATGAAGTTGG c-Jun forward* sense CATGAGGAACCGCATTGCCGC c-Jun reverse* antisense TAGCATGAGTTGGCACCCACTG E-actin forward** sense ACGGGGTCACCCACACTGTGC E-actin reverse** antisense CTAGAAGCATTTGCGGTGGAC

* The c-Jun primers are based on those previously reported by Chung et al. (1996). ** The E-actin primers are those previously reported by Lederer et al. (1996). The c-Fos, c-Jun and E-actin primers were used in semi-quantitative RT-PCR; the CYP2J2 primers were used in both semi-quantitative and competitive RT-PCR.

The CYP2J2-forward and -reverse primers correspond to nucleotides 724 to 749 and 953 to 978, respectively, within the coding sequence of the CYP2J2 gene, giving an amplification product of 254 bp. c-Fos-forward and -reverse primers corresponded to nucleotides 4 to 26 and 173 to 196, respectively, giving an amplification product of 193 bp. The c-Jun-forward and -reverse primers are based on those reported previously by Chung et al. (1996), and correspond to nucleotides 777 to

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797 and 951 to 972, respectively, to give a product of 196 bp. The E-actin-forward and - reverse primers were reported previously by Lederer et al. (1996), and correspond to nucleotides 511 to 531 and 1149 to 1169, respectively, giving an amplification product of 659 bp. Reverse transcription of mRNA into cDNA and subsequent amplification of the cDNA by PCR was performed in a single tube using the Qiagen OneStep RT-PCR Kit. Prior to RT-PCR, RNA samples were treated with DNase to digest any genomic DNA potentially present within the sample and thus prevent amplification of contaminating genomic DNA. Total RNA samples (0.5 Pg) were incubated with 1 PL (1U) RQ1 RNase-free DNase in a final volume of 10 μL for 10 min at 37qC, followed by an incubation at 70qC for 5 min to inactivate the DNase. While the DNase reactions were proceeding, a mastermix containing all RT-PCR reaction components (other than the template RNA) was prepared to minimise errors associated with pipetting individual components. The DNase-treated RNA samples were cooled on ice, and 40 PL of the mastermix was added to give reactions with a final volume of 50 μL that contained

1X OneStep RT-PCR Buffer (with 2.5 mM MgCl2); 400 PM each of dATP, dCTP, dGTP, dTTP; 10U Recombinant RNasin Ribonuclease Inhibitor; 0.1 to 1 PM of the appropriate primers (as outlined below); and 2 μL OneStep RT-PCR Enzyme Mix (containing an optimised concentration of Omniscript and Sensiscript Reverse Transcriptases and HotStarTaq DNA polymerase). Reactions were mixed thoroughly and amplified by cycling in a GeneAmp PCR system 2400 thermal cycler (PerkinElmer). An initial cycle of 50qC for 30 min to facilitate reverse transcription, and then 95qC for 15 min to facilitate inactivation of reverse transcriptases and activation of DNA polymerase, was followed by cycles of denaturation at 94qC for 20 sec, annealing for 20 sec at the temperatures specified below, and extension at 72qC for 30 sec. Reactions were concluded by a final extension at 72qC for 10 min. The annealing temperature, primer concentration and cycle number for each co-amplification reaction was optimised to ensure that reactions were within the exponential phase of amplification. For co-amplification of CYP2J2 and E-actin, annealing temperature was 55qC, primer concentrations were 0.6 PM and 0.1 PM for CYP2J2 and E-actin primers respectively, and 28 cycles of amplification were used. For co-amplification of c-Fos and E-actin, annealing temperature was 58qC, primer concentrations were 1 PM and 0.2 PM for c-Fos and E-actin primers respectively, and 26 cycles of amplification were used. For c-Jun and E-actin co-amplification, annealing temperature was 58qC, primer concentrations were 1 PM and 0.2 PM for c-Jun and E-actin primers respectively, and 25 cycles of amplification were used.

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Following cycling, RT-PCR reactions (30 μL) were mixed with 5 μL DNA loading dye and electrophoresed with TBE buffer (90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA) on 2% agarose gels containing 0.1 μg/mL ethidium bromide. A 100 bp DNA size ladder (Progen Industries) was included for estimation of the size of RT-PCR products. After electrophoresis at 110V for 50 min, the RT-PCR products were visualised under UV light (Gel Doc 2000 transilluminator, Bio-Rad), and their intensities determined densitometrically using Quantity One software (Bio-Rad). The relative intensity of the CYP2J2, c-Fos and c-Jun amplification products were normalised to that of the ȕ-actin control.

2.6.3 Competitive RT-PCR for Quantification of CYP2J2 mRNA 2.6.3.1 Preparation of a Recombinant CYP2J2 RNA Internal Standard Competitive RT-PCR was used to supplement the semi-quantitative RT-PCR data obtained for CYP2J2. Briefly, competitive RT-PCR involves spiking the amplification reaction with varying amounts of an internal RNA standard, which is constructed to contain the same primer binding sites as the target RNA, but modified from the target RNA sequence so that their amplification products are distinguishable from one another, usually by size. The target and standard RNAs compete for amplification by the same primer, with the target RNA amplification product decreasing as the amount of spiked standard RNA increases. The reaction in which the target and standard amplification products are equal indicates that the initial concentrations of target and standard RNA were also equivalent, thus enabling the quantity of target RNA to be calculated (Vanden Heuvel et al., 1993; Zimmermann & Mannhalter, 1996). A 197 bp recombinant RNA template [(2J2-197)-RNA] for use as an internal standard in competitive RT-PCR was generated by Mrs Eva Fiala-Beer of this laboratory as follows. PCR amplification of CYP2J2 cDNA with the CYP2J2-forward and -reverse primers used for semi-quantitative RT-PCR (Table 2.3) was used to generate a CYP2J2 amplification product of 254 bp corresponding to nucleotides 724 to 978. The PCR reaction contained 1 μg of CYP2J2 cDNA template, 0.2 μM each of CYP2J2-forward and -reverse primer, 1X Pfu Turbo reaction buffer, 200 μM dNTP mix and 3U Pfu Turbo polymerase in a final reaction volume of 50 μL. Amplification was performed on a GeneAmp PCR system 2400 thermal cycler (PerkinElmer) using an initial cycle of denaturation at 95qC for 3 min, followed by 28 cycles of denaturation at 95qC for 30 sec, primer annealing at 55qC for 1 min, and extension at 68qC for 2 min. The amplification reaction was concluded with a final extension at 68qC for 7 min. The resultant 254 bp CYP2J2 PCR product was purified using Microcon PCR columns

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(Millipore, North Ryde, NSW, Australia) according to the manufacturer’s instructions, and then A-tailed to produce overhanging adenosine nucleotides for cloning into the pGEM-T Easy vector (Promega). For A-tailing, a reaction mix containing 200 ng of the purified 254 bp CYP2J2 PCR product, 200 μM dATP, 2.5 mM MgCl2, 1X Taq buffer and 5U Taq DNA polymerase in a final volume of 10 μL was incubated at 70qC for 30 min. The A-tailed 254 bp CYP2J2 PCR product was cloned into the pGEM-T Easy vector using a ligation reaction that contained a 3:1 molar ratio of insert to vector, 1X Rapid Ligation Buffer (Promega) and 3U T4 DNA ligase. The ligation reaction was incubated at room temperature for 60 min, transformed into E.coli JM109 cells, and transformed colonies harboring the cloned (2J2-254)-pGEM plasmid were identified by blue/white screening of transformants grown on LB-agar plates supplemented with isopropyl-E-D-thiogalactopyranoside (IPTG; 0.5 mM) and X-Gal (0.05 mg/mL). Colonies were selected, grown overnight, and the (2J2-254)-pGEM plasmid DNA was extracted and confirmed by sequencing using the pGEM-T vector sequencing primer M13 (Promega; sequence shown in Table 2.1). Two Hindȱȱȱ recognition sites separated by 57 bp were created within (2J2-254)-pGEM by site-directed mutagenesis according to the protocol described in section 2.3. The sequences of the complementary primer pairs used to introduce the two Hindȱȱȱ recognition sites are listed in Table 2.4, and their successful incorporation to create the plasmid 2J2-254-Hindȱȱȱ-pGEM was confirmed by sequencing (M13 primer). 2J2-254-Hindȱȱȱ-pGEM (10 μg) was incubated with 20U Hindȱȱȱ at 37qC for 90 min to release a 57 bp fragment from within the CYP2J2 DNA insert. The larger CYP2J2-plasmid fragment was separated from the smaller 57 bp fragment by gel electrophoresis, purified using the QIAquick Gel Extraction Kit and re-ligated with T4 DNA ligase to generate the plasmid (2J2-197)-pGEM. This plasmid contained the shortened recombinant 197 bp CYP2J2 DNA insert downstream of the pGEM-T Easy T7 promoter. Recombinant (2J2-197)-RNA was produced by in vitro transcription of (2J2-197)-pGEM using the RiboMAX¥ Large Scale RNA Production System and T7 polymerase according to the manufacturer’s instructions. Briefly, the (2J2-197)-pGEM plasmid (12 Pg) was linearised by digestion with Salȱ (20U) and then purified by gel electrophoresis and extraction using the QIAquick Gel Extraction Kit. The linearised plasmid (1 μg) was combined with 1X T7 Transcription Buffer, 7.5 mM each of dATP, dCTP, dGTP and dUTP and 10 μL T7 enzyme mix in a final volume of 100 μL, and incubated at 37qC for 5 hr. RQ1 RNase-free DNase (1U) was added and the reactions incubated at 37qC for a further 15 min to digest the DNA template. The transcribed (2J2-197)-RNA was subsequently purified using the Epicentre MasterPure¥ Complete

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DNA and RNA Purification Kit according to the following protocol supplied by the manufacturer. The DNase-treated in vitro transcription reaction was combined with 100 μL 2X T and C Lysis Solution and mixed by vortexing for 5 sec. Following the addition of 100 μL MPC Protein Precipitation Reagent, mixing by vortexing for 10 sec, and incubation on ice for 3 min, samples were centrifuged at 10000g for 10 min at room temperature. The supernatant containing the RNA was transferred to a fresh microcentrifuge tube, mixed with 300 μL isopropanol to precipitate the RNA, and then centrifuged at 20800g for 10 min at 4qC. The supernatant was removed, and the resultant RNA pellet was washed with 200 μL 70% ethanol, and then re-pelleted by further centrifugation at 20800g for 10 min at 4qC. The wash procedure was repeated, the ethanol was carefully removed, and the (2J2-197)-RNA pellet was resuspended in 10 to 35 μL sterile water. (2J2-197)-RNA was quantitated by measuring absorbance at 260 nm (Cary 300 Bio spectrophotometer, Varian), and stored in aliquots at -80qC until used in competitive RT-PCR.

Table 2.4 Sequences of oligonucleotides used in site-directed mutagenesis to create a recombinant CYP2J2 RNA internal standard for competitive RT-PCR

Oligonucleotide Sense/ Location Sequence (5’ – 3’) Antisense

2J2-mt-Hind,,, 1 sense 746-768 GCAACTGGAAAAAAGCTTGAAATTG 2J2-mt-Hind,,, 1 antisense 768-746 CAATTTCAAGCTTTTTTCCAGTTGC 2J2-mt-Hind,,, 2 sense 806-835 GGAATCCTGCAGAAAGCTTAAGAGACTTTATTG 2J2-mt-Hind,,, 2 antisense 835-806 CAATAAAGTCTCTTAAGCTTTCTGCAGGATTCC

Hind,,, sites created within oligonucleotides by site-directed mutagenesis are underlined, and mutated nucleotides are highlighted in bold. Primer locations within the coding sequence of the CYP2J2 gene are numbered relative to the translation start site (1).

2.6.3.2 Quantitative Competitive RT-PCR for CYP2J2 For quantification of CYP2J2 mRNA, various amounts (0.1 pg to 100 pg) of the in vitro transcribed recombinant (2J2-197)-RNA was added as an internal competitive standard to a fixed quantity (0.5Pg) of RNA extracted from HepG2 cells. Competitive RT-PCR was performed using the CYP2J2-forward and -reverse primers that were used for semi-quantitative RT-PCR of CYP2J2 (sequences listed in Table 2.3). These primers amplified both the native target CYP2J2 mRNA and the recombinant (2J2-197)-RNA standard to produce amplification products of 254 bp and 197 bp respectively. The RNA mixture was incubated with RQ1 RNase-free DNase (1U) for 10 min at 37qC,

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cooled on ice, and then combined with RT-PCR mastermix to give 50 μL reactions containing 1X OneStep RT-PCR Buffer (with 2.5 mM MgCl2); 0.6 μM CYP2J2-forward and -reverse primer; 400 μM each of dATP, dCTP, dGTP and dTTP; 10U Recombinant RNasin Ribonuclease Inhibitor and 2 μL OneStep RT-PCR Enzyme Mix. Reactions were mixed and subjected to cycling in a GeneAmp PCR system 2400 thermal cycler (PerkinElmer), using conditions that were identical to those described for the semi- quantitative CYP2J2 RT-PCR. For separation and quantitation of RT-PCR products, RT-PCR reactions (30 μL) were mixed with 5 μL DNA loading dye and electrophoresed on 2.5% agarose gels containing 0.1 μg/mL ethidium bromide in TBE buffer (90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA). Following electrophoresis at 100V for 100 min, the amplification products from the CYP2J2 target mRNA and standard competitor were visualised under UV light (Gel Doc 2000 transilluminator, Bio-Rad), and their relative intensities were determined by densitometric analysis using Quantity One software (Bio-Rad). For quantitation of the amount of CYP2J2 mRNA present within each sample, the ratio of the signals derived from the standard and target was plotted against the amount of standard competitor RNA included in the reaction.

2.7 Protein Analysis

2.7.1 Isolation of Total Cell Lysates for Protein Analysis Following treatment, HepG2 cells were harvested as described in section 2.4.4 and the resultant cell pellets were resuspended in 1 mL ice-cold PBS and transferred to clean microcentrifuge tubes. Cells were pelleted by centrifugation at 20800g for 3 min at 4qC and lysed by resuspension in 300 PL of sample buffer [400 mM Tris-HCl, 3% (w/v) sodium dodecyl sulfate (SDS), 10% (v/v) 2-mercaptoethanol, 20% (v/v) glycerol and 0.002% (w/v) bromophenol blue]. The suspension was passed through a 23 gauge needle 10 times to shear the DNA and separate interacting DNA and protein, and then the samples were heated at 100qC for 5 min to facilitate protein denaturing. Cellular debris was pelleted by centrifugation at 20800g for 3 min at 4qC, and the supernatant containing total cellular protein (total cell lysate) was aliquoted and stored at –20qC until used in immunoblotting.

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2.7.2 Immunoblotting Immunoblotting was used to examine the abundance of CYP2J2, c-Fos and c-Jun protein in total cell lysates extracted from HepG2 cells. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) was performed according to the method of Laemmli (1970). Equal volumes of protein samples (total cell lysates) were resolved by electrophoresis on 10% SDS-polyacrylamide gels [10% (v/v) acrylamide (29:1 acrylamide:bis), 0.75 M Tris-HCl, pH 8.8, 0.1% (w/v) SDS, 0.05% (w/v) ammonium persulfate (APS), 0.1% (v/v) N,N,N’,N’-tetramethylethylenediamine (TEMED)] in Tris-glycine buffer (24 mM Tris, 192 mM glycine, 0.1% (w/v) SDS). Following electrophoresis at 120V for 1.5 to 2.5 hr, separated proteins were electrophoretically transferred from the gel onto nitrocellulose in transfer buffer (25 mM Tris, 192 mM glycine, 20% (v/v) methanol) at 100V for 1 hr. For detection of proteins, membranes were blocked for 1 hr at room temperature in Tris-buffered saline (50 mM Tris-HCl, pH 7.4, 200 mM NaCl and 0.05% (v/v) Tween-20) containing 5% (w/v) non-fat dried milk, and then incubated for 2 hr at room temperature with primary antibodies diluted as follows in the blocking solution; polyclonal anti-(rat CYP2J4) [16 Pg/mL; kindly provided by Dr. Qing-Yu Zhang], anti- c-Fos (0.75 Pg/mL; Santa Cruz) or anti- c-Jun (0.4 Pg/mL; Santa Cruz). Immunoblotting for E-actin (1 Pg/mL anti-actin in blocking solution; Santa Cruz) on parallel nitrocellulose filters was performed to correct for protein variation between samples. After incubation with primary antibodies, the membranes were washed 5 times in Tris-buffered saline and incubated for 1 hr at room temperature in secondary anti-(rabbit IgG) or anti-(mouse IgG) antibody conjugated with horseradish peroxidase. The anti-(rabbit IgG) antibody was diluted in 5% blocking solution at 1:1000, 1:1500, and 1:2000 dilutions for CYP2J2, c-Fos and c-Jun blots respectively, while anti-(mouse IgG) antibody was used at a dilution of 1:500 in 5% blocking solution for the detection of E-actin. After incubation with secondary antibody, the membranes were washed five times in Tris-buffered saline and the proteins visualised by autoradiography following incubation with ECL Western blotting detection reagents according to the manufacturer’s instructions (Amersham Biosciences). Bacterial-expressed CYP2J2 (E.coli DH5D cells), recombinant human c-Jun protein (Promega) and nuclear extract from phorbol-ester-treated Jurkat cells (Santa Cruz) were included in immunoblots as standards for CYP2J2, c-Jun and c-Fos, respectively.

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2.8 Transient Transfection Analysis

2.8.1 Transient Transfection of HepG2 Cells HepG2 cells were seeded at a density of 6 x 105 cells per well in 6-well plates and incubated for 24 hr prior to transfection. Transfections were carried out using the FuGENE¥ 6 transfection reagent at a ratio of 3:2 (FuGENE¥ 6 reagent: total DNA transfected) in accordance with the manufacturer’s instructions. Cells were co-transfected with 1 Pg/well CYP2J2 promoter constructs and 0.5 Pg/well pCMV-E-galactosidase expression plasmid to control for transfection efficiency. Mammalian expression plasmids encoding AP-1 proteins were added at 0.5 Pg/well, and total DNA in each transfection was equalised with salmon sperm DNA. The plasmid DNA mixture for each transfection was incubated with the FuGENE¥ 6 transfection reagent (diluted to a volume of 100 μL in serum-free DMEM) for 20 min at room temperature and then added to each well. Plates were gently swirled to ensure an even distribution of the transfection mixture. The cells were incubated with the transfection mixture for 24 hr, after which time the medium was replaced with fresh complete DMEM and the cells incubated for a further 48 hr before harvesting. For harvest, the media was aspirated, the cells were washed in PBS (2 mL) and detached from the surface of the plates by incubation with trypsin (0.3 mL) at 37qC for 6 min. Following trypsinisation, cells were resuspended in 2 mL DMEM and transferred to centrifuge tubes. Samples were centrifuged for 5 min at 360g, the supernatant was aspirated and cell pellets were resuspended in 0.5 mL of fresh complete DMEM and stored at -80qC until assayed for luciferase and E-galactosidase activity. Luciferase and ȕ-galactosidase assays were performed as outlined in sections 2.8.2 and 2.8.3. Luciferase activity was normalised to ȕ-galactosidase activity to control for variations in transfection efficiency. All transfections within each experiment were performed in duplicate, and each experiment was performed on at least three separate occasions. Data from transfections is expressed as means ± standard error (S.E.M.) relative to a specified promoter construct, and differences between samples were detected using a Student’s t-test, with P<0.05 considered to be statistically significant.

2.8.2 Luciferase Reporter Gene Assay A series of CYP2J2 promoter constructs was prepared in which the CYP2J2 5’-flanking region was inserted into the pGL3basic luciferase reporter gene vector (section 2.3). The pGL3basic reporter vector encodes the firefly (Photinus pyralis) luciferase protein,

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but lacks a functional promoter or enhancer necessary for transcription of the firefly gene and production of firefly luciferase protein. To identify gene regulatory elements important in the control of CYP2J2 gene transcription, promoter reporter constructs were transfected into HepG2 cells as described in section 2.8.1, and luciferase activity was measured using the Steady-Glo¥ Luciferase Assay System (Promega). The luciferase assay was performed as follows. Transfected cells were harvested as outlined in section 2.8.1, and 10 – 50 ȝL of the thoroughly resuspended cell sample was diluted in complete DMEM to a final volume of 100ȝL. Under reduced light conditions, an equal volume of the Steady-Glo reagent (100 PL) was added, the samples were mixed by vortexing briefly and then incubated for 5 min at room temperature to ensure complete cell lysis. Samples were transferred to scintillation vials, and luminescence was quantified in a scintillation counter (1900 TR, Packard).

2.8.3 ȕ-galactosidase Assay As outlined in section 2.8.1, a pCMV-ȕ-galactosidase plasmid was included in all transfections to control for variation in transfection efficiency. Thus, aliquots of cells from all transfections were assayed for ȕ-galactosidase activity (E-galactosidase Enzyme Assay System with Reporter Lysis Buffer; Promega). Here, the cell lysate is incubated with Assay 2X Buffer containing the ȕ-galactosidase substrate o-nitrophenyl-ȕ-D-galactopyranoside (ONPG). The resultant o-nitrophenol was detected spectrophotometrically at 420 nm. To prepare cell lysates, a 300 ȝL aliquot of the cell sample harvested from each transfection was centrifuged at 8000g for 3 min at room temperature and the supernatant was discarded. Traces of media in the resultant cell pellet were removed by resuspension of the cell pellet in 500 ȝL PBS and re-centrifugation at 8000g for 3 min at room temperature. Cell pellets were resuspended in 165 ȝL 1X Reporter Lysis Buffer (prepared by dilution of 5X Reporter Lysis Buffer in water), incubated for 15 min at room temperature, and then vigorously mixed by vortexing for 10 seconds. Cellular debris was pelleted by centrifugation at 8000g for 3 min at room temperature and the supernatant (cell lysate; 150 ȝL) was transferred to a fresh microcentrifuge tube and stored at -80qC until the ȕ-galactosidase assay was conducted. ȕ-galactosidase assay was performed according to the protocol described by Promega for the ȕ-galactosidase Enzyme Assay System with Reporter Lysis Buffer. Cell lysates were allowed to thaw at room temperature and an equal volume (150 ȝL) of Assay 2X Buffer (200 mM sodium phosphate buffer, pH 7.3, 2 mM MgCl2, 100 mM

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ȕ-mercaptoethanol, 1.33 mg/mL ONPG) was added. Samples were mixed, and incubated at 37qC for 15 to 30 min. Reactions were stopped by the addition of 500 ȝL 1M sodium carbonate, and the absorbance at 420 nm was measured immediately (Cary 300 Bio spectrophotometer, Varian). Product formation was determined with reference to a E-galactosidase standard curve (constructed using 0.0 to 6.0 x 10-3 units of E-galactosidase).

2.9 Electrophoretic Mobility Shift Assay (EMSA)

2.9.1 Preparation of Nuclear Extracts Nuclear extracts were prepared from HepG2 cells using the method outlined below that was adapted from Schreiber et al. (1989). All solutions used for nuclear protein extraction were stored at 4qC and protease inhibitors, dithiothreitol (DTT), phenylmethanesulfonyl fluoride (PMSF), Na3VO4, NaF, spermine and spermidine were added to buffers just prior to the commencement of the extraction procedure. All procedures were carried out on ice or at 4qC unless otherwise stated. Nuclear extracts were prepared from HepG2 cultured under normoxic (16 hr at 21% O2) and hypoxic conditions (16 hr at 1% O2; section 2.4.4), and also from untransfected HepG2 cells or HepG2 cells that had been transfected with a c-Jun expression plasmid (0.5 μg/6 x 105 cells) according to the protocol described in section 2.8.1. Following the defined treatments, cells were harvested as outlined in section 2.4.4, and the resultant cell pellets were washed by resuspension in 1 mL ice-cold PBS and transferred to clean microcentrifuge tubes. Cells were pelleted by centrifugation at 15800g for 15 sec, the supernatant was removed, and the cell pellet was resuspended in 400 PL of buffer containing 10 mM HEPES (pH 7.9), 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM

DTT, 0.5 mM PMSF, 0.4 mM Na3VO4, 1 mM NaF, 0.15 mM spermine, 0.5 mM spermidine, 2 Pg/mL aprotinin, 1 Pg/mL pepstatin and 1 Pg/mL leupeptin. Cells were allowed to swell on ice for 15 min, after which time 0.6% (v/v) Nonidet NP-40 (Fluka) was added, and the samples then vortexed vigorously for 10 sec. Following centrifugation at 15800g for 30 sec, the supernatant was discarded and the resultant nuclear pellets were resuspended in 50 PL of buffer containing 20 mM HEPES (pH 7.9), 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 1 mM PMSF, 0.4 mM

Na3VO4, 1 mM NaF, 4 Pg/mL aprotinin, 1 Pg/mL pepstatin and 10 Pg/mL leupeptin. The tubes were rocked at 4qC for 15 min and then the nuclear membranes were pelleted by

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centrifugation at 15800g for 5 min. The supernatant (nuclear extract) was separated into aliquots, frozen in liquid N2 and stored at –80qC until used in EMSAs. The protein concentration of nuclear extracts was determined by a Lowry assay adapted from the Lowry method using bovine serum albumin (BSA) as a standard (Lowry et al., 1951). Briefly, 1 to 20 μg of BSA was aliquoted into microcentrifuge tubes and diluted to a final volume of 200 μL with water. Likewise, 1 μL and 3 μL aliquots of each nuclear protein sample were diluted to 200 μL with water. Lowry solution

(185 mM Na2CO3, 98 mM NaOH, 0.4 mM CuSO4, 0.87 mM Na tartrate; 1 mL) was added to the nuclear protein samples and BSA standards, and samples were mixed by vortexing and incubated for 10 min at room temperature. Folins reagent (100 μL of 50% solution in water) was added, the samples were vortexed and incubated for a further 30 min at room temperature, and then the absorbance at 750 nm was measured (Cary 300 Bio spectrophotometer, Varian). Protein was quantified with reference to BSA. The absorbance at 280 nm of the BSA stock (1 μg/μL) from which the BSA standards were prepared was measured, and a correction factor was determined by comparison to the theoretical absorbance at 280 nm of 1 μg/μL BSA (0.66).

2.9.2 Preparation of Double-stranded Probes for use in EMSA The sequences of complementary sense and antisense oligonucleotides used as labelled probes or unlabelled competitors in EMSAs are listed in Table 2.5. These oligonucleotides generally contained a 3’-GGG overhang for the purpose of labelling with 32P-radiolabelled dCTP. The oligonucleotides containing the consensus AP-1 recognition element were based on those described by Stratagene and contained 5’-CTAG (sense) and -GATC (antisense) overhangs for labelling. The oligonucleotides containing the STAT5 recognition sequence from the ȕ-casein promoter were based on those reported by Gebert et al. (1997), with the antisense oligonucleotide containing a 5’-G overhang for labelling. For annealing of complementary oligonucleotides, equal amounts (0.2 nmoles) of complementary sense and antisense oligonucleotides were combined with 1X Multicore restriction enzyme buffer (Promega) in a final volume of 40 μL. Samples were heated to 100qC for 10 min to disrupt any secondary coiling of the oligonucleotides that may otherwise impede proper annealing, and were then annealed by slow cooling overnight. To confirm annealing, a 5 μL sample of each annealing reaction was separated by PAGE alongside a 2 μL aliquot of the appropriate sense or antisense single-stranded oligonucleotide as follows. The annealing reactions and oligonucleotide samples were combined with DNA loading dye (2 μL) and

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electrophoresed on a 20% polyacrylamide gel [20% (v/v) acrylamide (29:1 acrylamide:bis), 90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA, 0.06% (v/v) APS, 0.06% (v/v) TEMED] in TBE buffer (90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA) at 100V for 40 min. The gels were stained with ethidium bromide and the DNA fragments were visualised under UV light. Successfully annealed oligonucleotide probes did not migrate as far on gels as non-annealed probes. Annealed probes were end-labelled with [32 P]dCTP using the Megaprime¥ DNA Labelling System. Labelling reactions contained 20 pmoles of annealed probe,

1X reaction buffer (containing Tris-HCl, pH 7.5, 2-mercaptoethanol and MgCl2), 2 PL each of dATP, dGTP and dTTP supplied with the kit, 3 PL (30 μCi) of [32P]dCTP and 1U Klenow DNA polymerase, and were incubated at room temperature for 90 min. To confirm probe labelling, an aliquot (1 PL) of the labelling reaction was spotted onto PEI Cellulose F strips (Merck), suspended in a small volume of 0.75 M potassium dihydrogen orthophosphate to allow the separation of unincorporated 32P-radiolabelled dCTP and labelled probe, and visualised by autoradiography. Following confirmation of labelling, unincorporated radioactivity was removed by purification through ProbeQuant¥ G-50 Micro Columns (Amersham Biosciences) according to the manufacturer’s instructions. Briefly, columns were vortexed to resuspend the matrix and then centrifuged at 735g for 1 min to equilibrate the column. The labelling reaction was applied to the column and centrifuged at 735g for 2 min. The labelled probe passed through the column and was collected in a microcentrifuge tube, whilst the unincorporated radioactivity remained trapped in the column and was discarded. Radiolabelled probes were kept at 4qC until used in EMSA. In addition to small oligonucleotide probes, a larger double-stranded probe consisting of 167 bp of the CYP2J2 promoter spanning -152 to +15 bp (relative to the translation start site) was used in some EMSAs. To prepare this fragment (2J2/167), the CYP2J2 reporter construct 2J2 (-152/+98) was digested with Nheȱ and Banȱȱ restriction enzymes (37qC for 90 min), and the released 167 bp fragment was separated from the other digested plasmid fragments by electrophoresis on a 2% low-melting agarose gel. Digested fragments were visualised under UV light, gel slices containing the 2J2/167 fragment were excised, and the 2J2/167 fragment was purified from agarose using the QIAquick Gel Extraction Kit. The purified 2J2/167 fragment was quantified from the absorbance at 260 nm, and labelled as outlined above for the oligonucleotide probes.

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Table 2.5 Sequences of double-stranded oligonucleotides used in EMSAs

Oligonucleotide S/AS Sequence (5’ – 3’)

2J2-(-7/+1) S GAGGACGTCTGAGCCATGCTCGCggg 2J2-(-7/+1) AS GCGAGCATGGCTCAGACGTCCTCggg 2J2-mt-(-7/+1)*** S GAGGACGTAAAAAAAATGCTCGCggg 2J2-mt-(-7/+1)*** AS GCGAGCATTTTTTTTACGTCCTCggg 2J2-(-56/-63) S CGGGGCGGGGACCGTCGCCTGCTGGGggg 2J2-(-56/-63) AS CCCAGCAGGCGACGGTCCCCGCCCCGggg 2J2-mt-(-56/-63)*** S CGGGGCGGGAAAAAAAACCTGCTGGGggg 2J2-mt-(-56/-63)*** AS CCCAGCAGGTTTTTTTTCCCGCCCCGggg 2J2-A (-152/-103)* S ctagCCCGGGAATCCAGCGCCTGGCATCTTCGCA GGGTGCTGCGAAGGGGCGGG 2J2-A (-152/-103)* AS gatcCCCGCCCCTTCGCAGCACCCTGCGAAGATG CCAGGCGCTGGATTCCCGGG 2J2-B (-127/-79) S TCGCAGGGTGCTGCGAAGGGGCGGGCTGGGAG GCGGGGCACGGCTGGGAggg 2J2-B (-127/-79) AS TCCCAGCCGTGCCCCGCCTCCCAGCCCGCCCC TTCGCAGCACCCTGCGAggg 2J2-C (-102/-50) S CTGGGAGGCGGGGCACGGCTGGGAGCGAGGC GGGGCGGGGACCGTCGCCTGCTggg 2J2-C (-102/-50) AS AGCAGGCGACGGTCCCCGCCCCGCCTCGCTCC CAGCCGTGCCCCGCCTCCCAGggg 2J2-(-152/-122) S CCCGGGAATCCAGCGCCTGGCATCTTCGCAggg 2J2-(-152/-122) AS TGCGAAGATGCCAGGCGCTGGATTCCCGGGggg -152/-128 (probe 1) S CCCGGGAATCCAGCGCCTGGCATCTggg -152/-128 (probe 1) AS AGATGCCAGGCGCTGGATTCCCGGGggg -137/-119 (probe 2) S CTGGCATCTTCGCAGGGTggg -137/-119 (probe 2) AS ACCCTGCGAAGATGCCAGggg -127/-106 (probe 3) S TCGCAGGGTGCTGCGAAGGGGCggg -127/-106 (probe 3) AS GCCCCTTCGCAGCACCCTGCGAggg -114/-97 (probe 4) S CGAAGGGGCGGGCTGGGAggg -114/-97 (probe 4) AS TCCCAGCCCGCCCCTTCGggg -105/-86 (probe 5) S GGGCTGGGAGGCGGGGCACGggg -105/-86 (probe 5) AS CGTGCCCCGCCTCCCAGCCCggg -105/-86; mt –105/-96 (probe mt 5)*** S AAAAAAAAAAGCGGGGCACGggg -105/-86; mt –105/-96 (probe mt 5)*** AS CGTGCCCCGCTTTTTTTTTTggg -94/-77 (probe 6) S CGGGGCACGGCTGGGAGCggg -94/-77 (probe 6) AS GCTCCCAGCCGTGCCCCGggg -85/-69 (probe 7) S GCTGGGAGCGAGGCGGGggg -85/-69 (probe 7) AS CCCGCCTCGCTCCCAGCggg -77/-60 (probe 8) S CGAGGCGGGGCGGGGACCggg -77/-60 (probe 8) AS GGTCCCCGCCCCGCCTCGggg -68/-48 (probe 9) S GCGGGGACCGTCGCCTGCTGGggg -68/-48 (probe 9) AS CCAGCAGGCGACGGTCCCCGCggg -157/-138 (probe 10) S GTCCTCCCGGGAATCCAGCGggg -157/-138 (probe 10) AS CGCTGGATTCCCGGGAGGACggg -147/-128 (probe 11) S GAATCCAGCGCCTGGCATCTggg -147/-128 (probe 11) AS AGATGCCAGGCGCTGGATTCggg AP-1 consensus* S ctagTGATGAGTCAGCCGGATC AP-1 consensus* AS gatcGATCCGGCTGACTCATCA Stat5 E-casein promoter S GGACTTCTTGGAATTAAGGGA Stat5 E-casein promoter** AS gTCCCTTAATTCCAAGAAGTCC

All oligonucleotides have a 3’-ggg overhang for labelling with [32P]dCTP with exceptions as follows; * the AP-1 consensus and 2J2-A (-152/-103) oligonucleotides have a 5’-ctag overhang on the sense (S) strand and a 5’-gatc overhang on the antisense (AS) strand; ** the Stat5 antisense (AS) oligonucleotide has a 5’-g overhang. The AP-1 consensus sequence in the sense (S) strand of the AP-1 consensus oligonucleotide, the STAT5 element in the Stat5 E-casein promoter oligonucleotides, the AP-1-like element in the antisense (AS) strand of 2J2-(-56/-63) probe, and the c-Jun binding site at -105/-95 bp in the sense strand of probe 5 (-105/-86) are highlighted in bold.*** Mutated nucleotides are underlined. 108 Chapter 2

2.9.3 EMSA Radiolabelled probes (10 – 200 fmoles) were incubated with 5-30 Pg of nuclear protein fractions for 20 min at room temperature and 10 min at 4qC in a buffer containing

50 mM NaCl, 10 mM Tris-HCL (pH 7.5), 1 mM MgCl2, 0.5 mM EDTA (pH 8.0), 0.5 mM DTT, 4% glycerol and 1Pg poly(2’-deoxyinosinic-2’-deoxycytidylic acid) [poly(dI-dC)]. For reactions with recombinant proteins, 0.6 μg recombinant human c-Jun, or a mixture consisting of 0.15 μg of recombinant human c-Fos and 0.15 μg of recombinant human c-Jun, were incubated as outlined for nuclear extracts, except that the incubation buffer was adjusted to contain 0.2 μg poly(dI-dC). Loading dye (2 PL) was added to the reactions, and protein-DNA complexes were resolved by electrophoresis on a 5% polyacrylamide gel (4.96% acrylamide, 0.06% bis-acrylamide, 90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA, 2.5% glycerol, 0.04% APS, 0.1% TEMED) in TBE buffer (90 mM Tris-HCl, 90 mM boric acid, 2.5 mM EDTA) at 100V for 1.5 to 2.5 hr at 4qC. Following electrophoresis, gels were dried (model 583 Gel Dryer and HydroTech¥ Vacuum Pump, Bio-Rad), and protein-DNA interactions were visualised by autoradiography. For competition experiments, 200-fold excess unlabelled probe was included in binding reactions, with the STAT5 consensus sequence from the E-casein promoter (described by Gebert et al., 1997) used as a negative control. For supershift experiments, nuclear or recombinant protein was incubated with rabbit polyclonal c-Jun or c-Fos antibodies (2 Pg; Santa Cruz) for 1 hr at 4qC prior to the binding reaction. An anti-ubiquitin (Ub) antibody (Santa Cruz) was used as a negative control in supershift experiments.

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Chapter 3

Regulation of the Expression of CYP2J2, and the AP-1 Proteins c-Fos and c-Jun, in Hypoxia and Reoxygenation

3.1 Introduction

Hypoxia is defined as a decreased oxygen tension in tissues and occurs when the supply of blood is decreased or occluded. Reoxygenation occurs when the blood flow is restored. Hypoxia is a feature of normal physiological processes such as embryogenesis, but also occurs in numerous pathological processes or conditions such as ischaemic heart disease, atherosclerosis, stroke, chronic lung disease and tumour growth (Semenza, 1999). The cellular expression of certain genes is altered in hypoxia as part of an adaptive mechanism aimed at enhancing cell survival (Bunn & Poyton, 1996; Semenza, 1996). Reprogramming of gene expression has been established as an important mechanism by which cells respond and adapt to harmful environmental changes or toxic substances (Tacchini et al., 2002). Genes known to be up-regulated in hypoxia include those encoding erythropoietin, which stimulates the production of red blood cells, vascular endothelial growth factor, a protein that is important in the formation of new blood vessels (angiogenesis), , which stimulates respiration, and genes encoding key glycolytic enzymes such as lactate dehydrogenase and pyruvate kinase, which support the production of ATP under anaerobic conditions (Bunn & Poyton, 1996; Semenza, 1996; Guillemin & Krasnow, 1997). The enhanced expression of these genes supports energy production and promotes improved oxygen transport and delivery, and thus enhances survival of cells in an oxygen-deficient environment (Semenza, 1996; Guillemin & Krasnow, 1997).

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Significant progress has been made in understanding the mechanisms by which gene expression is regulated in hypoxia, with the identification of several transcription factors that are activated in hypoxia, in particular HIF-1 and AP-1. These transcription factors have been found to play an important role in mediating the changes in gene expression that occur in hypoxia (Norris & Millhorn, 1995; Semenza, 1996; Mishra et al., 1998; Faller, 1999; Semenza, 1999; Michiels et al., 2001). HIF-1 plays a critical role in the increased expression of erythropoietin, vascular endothelial growth factor and the genes involved in glycolytic metabolism (Semenza, 1996; Guillemin & Krasnow, 1997; Semenza, 1998; Faller, 1999). Similarly, AP-1 also participates in the hypoxic regulation of vascular endothelial growth factor, basic fibroblast growth factor, collagenase IV, platelet-derived growth factor-B, endothelin-1 and thrombospondin-1 (Damert et al., 1997; Faller, 1999; Lee & Corry, 1999; Minet et al., 2001). These proteins have potent vasoconstricting, chemoattractant, mitogenic or angiogenic activities and are involved in tissue remodelling that occurs in chronic hypoxia (Faller, 1999; Minet et al., 2001). Some investigators have indicated that the redox-sensitive transcription factor NF-NB is also activated in hypoxia (Koong et al., 1994) and may mediate altered regulation of cyclooxygenase-2 under low oxygen conditions (Schmedtje et al., 1997). By contrast, other studies have proposed that NF-NB is not activated by hypoxia, but rather by the reoxygenation process, which is associated with increased formation of reactive oxygen species (Rupec & Baeuerle, 1995). Hypoxia can also decrease the expression of certain genes. For example, endothelial nitric oxide synthase (eNOS) is down-regulated in hypoxia, and the resultant decrease in nitric oxide availability may contribute to the pathogenesis of certain diseases that have a component of hypoxia (Liao et al., 1995; Phelan & Faller, 1996; Giraldez et al., 1997; Faller, 1999). At the commencement of this research project, very little was known about the regulation of CYP2J2, other than a study by Yang et al. (2001) which indicated that CYP2J2 protein expression was reduced in endothelial cells subjected to hypoxia and reoxygenation. However, the molecular mechanism by which CYP2J2 expression is reduced in hypoxia was not investigated (Yang et al., 2001). The present thesis investigated CYP2J2 regulation in the human liver-derived HepG2 cell line. The HepG2 cell line was chosen as an appropriate system in which to study CYP2J2 gene regulation for a number of reasons. First, CYP2J2 mRNA and protein is expressed in both human liver tissue (Wu et al., 1996; Enayetallah et al., 2004), and HepG2 cells (preliminary findings). Therefore, this liver-derived cell line contains the transcriptional machinery necessary for controlling the CYP2J2 promoter. Second, the HepG2 cell line is easy to maintain and readily transfectable, and has also been used successfully as a

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model to study the regulation of certain other CYP genes (Fukuda et al., 1992; Ibeanu & Goldstein, 1995; Morel & Barouki, 1998; Goodwin et al., 1999; Makishima et al., 1999; Nitta et al., 1999; Sueyoshi et al., 1999; Zhang & Chiang, 2001). This chapter investigates the effect of exposure to low oxygen on CYP2J2 mRNA and protein expression in HepG2 cells, and identifies the AP-1 transcription factor as a potential regulator of CYP2J2 expression. The molecular mechanisms by which the AP-1 proteins c-Fos and c-Jun regulate the expression of CYP2J2 under normal and decreased oxygen conditions is explored in detail in subsequent chapters.

3.2 Viability of HepG2 Cells Following Exposure to Hypoxia

In this chapter, the expression of CYP2J2 was examined in HepG2 cells exposed to hypoxia (1% O2) for a period of 16 hr and 40 hr. Preliminary experiments assessed the viability of HepG2 cells in hypoxia. Using the MTT reduction assay, cell viability was assessed immediately after hypoxic exposure, and also after allowing a period of recovery from hypoxic exposure. In these experiments, cells were first cultured in hypoxia for 16 or 40 hr, and then under normal oxygen conditions (21% O2) for 3 to 6 days before MTT assay. Control cultures were incubated in normoxia (21% O2) at each time point, and the viability of cells cultured in hypoxia was expressed as a percentage of the suitable normoxic control (Figure 3.1). Exposure of cells to hypoxia for 16 hr decreased viability to 74.5% of normoxic control (P<0.0001), but viability was restored after 3 days of recovery (Figure 3.1 A). On the other hand, exposure to 40 hr of hypoxia produced a much greater effect, decreasing cell viability to 39.4% of control, which remained low even after 6 days recovery at normal oxygen (Figure 3.1 B; P<0.0001 at each time point). Thus, the viability of HepG2 cells was relatively unimpaired by exposure to hypoxia for 16 hr, but significantly affected by exposure to low oxygen for the longer period of 40 hr.

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80 * Normoxia (16 hr) 60 Hypoxia (16 hr) 40 % Cell Viability % Cell 20

0 0 days 3 days 4 days 6 days recovery recovery recovery recovery

B 120

100

80 Normoxia (40 hr) 60 * Hypoxia (40 hr) * 40 * * % Cell Viability % Cell 20

0 0 days 3 days 4 days 6 days recovery recovery recovery recovery

Figure 3.1 Viability of HepG2 cells following exposure to hypoxia for 16 or 40 hr A. Viability of HepG2 cells following exposure to hypoxia for 16 hr. HepG2 cells were cultured for 16 hr in hypoxia (1% O22 ) or normoxia (21% O ). Cell viability was assessed by MTT assay immediately following treatment (0 days recovery), or after allowing 3 to 6 days recovery in normoxic culture conditions (21% O2), as outlined in Methods. Results shown are means ± S.E.M., (n=3), with viability expressed as a percentage of normoxic control. *Significantly different from normoxic control (P<0.0001). B. Viability of HepG2 cells following exposure to hypoxia for 40 hr. HepG2 cells were cultured for 40 hr in hypoxia (1% O2) or normoxia (21% O2), and cell viability assessed by MTT assay immediately following treatment (0 days recovery), or after allowing 3 to 6 days recovery in normoxic culture conditions (21% O2), as outlined in Methods. Results shown are means ± S.E.M., (n=3), with viability expressed as a percentage of normoxic control. *Significantly different from normoxic control (P<0.0001).

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3.3 Analysis of CYP2J2 mRNA Levels in HepG2 Cells Following Exposure to Hypoxia and Reoxygenation

To assess the effect of hypoxia on CYP2J2 mRNA levels, RNA was extracted from HepG2 cells exposed to hypoxia or hypoxia followed by reoxygenation, and semi-quantitative RT-PCR of CYP2J2 and ȕ-actin (internal control) was performed. Exposure to hypoxia for 16 hr did not affect expression of the ȕ-actin reference gene, but caused a pronounced decrease in CYP2J2 mRNA expression (Figure 3.2 A). In contrast, CYP2J2 mRNA levels were rapidly restored to control levels in HepG2 cells that were exposed to hypoxia for 16 hr and then a 30 min period of reoxygenation (Figure 3.2 B). CYP2J2 mRNA expression remained markedly suppressed after exposure to a longer period of hypoxia (40 hr; Figure 3.2 C).

A B C

CYP2J2

ȕ-actin

normoxia hypoxia normoxia hypoxia/ normoxia hypoxia (16 hr) (16 hr) (16 hr 30 min) reoxygenation (40 hr) (40 hr) (16 hr/30 min)

Figure 3.2 Semi-quantitative RT-PCR of CYP2J2 mRNA in HepG2 cells exposed to hypoxia and hypoxia-reoxygenation HepG2 cells were cultured in hypoxia (1% O2) for 16 hr (A), hypoxia for 16 hr followed by reoxygenation (21% O2) for 30 min (B), or hypoxia for 40 hr (C); controls were cultured in normoxia (21% O2) for each time period. Following treatment, cells were harvested and total RNA was extracted. CYP2J2 and ȕ-actin were amplified by semi-quantitative RT-PCR as described in Methods, and the amplification products visualised under UV after electrophoresis on 2% agarose gels.

To confirm the trend from semi-quantitative RT-PCR approaches, CYP2J2 mRNA levels were quantitated using a competitive RT-PCR assay. (2J2-197)-RNA, the 197 bp recombinant CYP2J2 RNA internal standard, was generated as described in section 2.6.3.1, and added at varying amounts (0.1 to 100 pg) to fixed quantities of RNA (0.5 μg) extracted from differently treated HepG2 cells. CYP2J2 target and standard RNA were amplified by RT-PCR, and separated by electrophoresis on agarose gels (Figures 3.3 A to 3.5 A). The intensity of the CYP2J2 target and standard amplification products were measured by densitometric analysis, and the amount of standard RNA introduced into each reaction was plotted against the ratio of the amplification products on a logarithmic scale (Figures 3.3 B to 3.5 B).

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Target (254 bp) Normoxia (16 hr) Standard (197 bp)

Target (254 bp) Hypoxia (16 hr) Standard (197 bp)

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1

0.8 Normoxia (16 hr) 0.6 g total RNA)

ȝ Hypoxia (16 hr) 0.4 CYP2J2 mRNA mRNA CYP2J2 * (pg/0.5 0.2

0 Treatment

Figure 3.3 Quantitation of CYP2J2 mRNA in HepG2 cells cultured in hypoxia for 16 hr by competitive RT-PCR A. RNA was isolated from HepG2 cells that had been cultured for 16 hr in normoxia (21% O2; upper panel) or hypoxia (1% O2; lower panel). Known quantities of (2J2-197)-RNA standard were added to 0.5 μg total cellular RNA, and competitive RT-PCR was performed as described in Methods. B. Typical relationship between the relative intensities of CYP2J2 target RNA (254 bp) and (2J2-197) standard RNA (197 bp). C. Effect of exposure to hypoxia for 16 hr on CYP2J2 mRNA expression in HepG2 cells. Results shown are pg of CYP2J2 mRNA per 0.5 ȝg total RNA (means ± S.E.M., n=3). *Significantly different from CYP2J2 mRNA levels in normoxic controls (P<0.0001).

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0.6 Normoxia (16 hr 30 min) g total RNA) total g

ȝ Hypoxia (16 hr)/ 0.4 reoxygenation (30 min) CYP2J2 mRNA mRNA CYP2J2

(pg/0.5 0.2

0 Treatment

Figure 3.4 Quantitation of CYP2J2 mRNA in HepG2 cells exposed to hypoxia for 16 hr followed by reoxygenation for 30 min by competitive RT-PCR A. RNA was isolated from HepG2 cells that had been cultured for 16 hr 30 min in normoxia (21% O2; upper panel) or 16 hr of hypoxia (1% O2), followed by 30 min reoxygenation (lower panel). Known quantities of (2J2-197)-RNA standard were added to 0.5 μg total cellular RNA, and competitive RT-PCR was performed as described in Methods. B. Typical relationship between the relative intensities of CYP2J2 target RNA (254 bp) and (2J2-197) standard RNA (197 bp). C. Effect of hypoxia- reoxygenation on CYP2J2 mRNA expression in HepG2 cells. Results shown are pg of CYP2J2 mRNA per 0.5 ȝg total RNA (means ± S.E.M., n=3).

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Target (254 bp) Hypoxia (40 hr) Standard (197 bp)

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RNA standard (pg) C 1.2

1

0.8 Normoxia (40 hr) 0.6 g total RNA) total g

ȝ Hypoxia (40 hr) 0.4 CYP2J2 mRNA (pg/0.5 0.2 * 0 Treatment

Figure 3.5 Quantitation of CYP2J2 mRNA in HepG2 cells cultured in hypoxia for 40 hr by competitive RT-PCR A. RNA was isolated from HepG2 cells that had been cultured for 40 hr in normoxia (21% O2; upper panel) or hypoxia (1% O2; lower panel). Known quantities of (2J2-197)- RNA standard were added to 0.5 μg total cellular RNA, and competitive RT-PCR was performed as described in Methods. B. Typical relationship between the relative intensities of CYP2J2 target RNA (254 bp) and (2J2-197) standard RNA (197 bp). C. Effect of exposure to hypoxia for 40 hr on CYP2J2 mRNA expression in HepG2 cells. Results shown are pg of CYP2J2 mRNA per 0.5 ȝg total RNA (means ± S.E.M., n=3). *Significantly different from CYP2J2 mRNA levels in normoxic controls (P<0.0001).

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Findings from competitive RT-PCR assays confirmed the data from semi-quantitative RT-PCR analysis. Thus, CYP2J2 mRNA was decreased (to 25% of control) in HepG2 cells exposed to hypoxia for 16 hr [0.26 ± 0.02 pg CYP2J2 mRNA/0.5 μg total RNA compared with 1.06 ± 0.07 pg CYP2J2 mRNA/0.5 μg of total RNA for control (P<0.0001); Figure 3.3 C]. Exposure of HepG2 cells to 30 min of reoxygenation following the 16 hr hypoxic incubation restored CYP2J2 mRNA levels to control levels [0.81 ± 0.04 pg CYP2J2 mRNA/0.5 μg total RNA compared with 0.74 ± 0.10 pg CYP2J2 mRNA/0.5 μg total RNA for control (not significant; NS); Figure 3.4 C]. In HepG2 cells exposed to hypoxia for 40 hr, CYP2J2 mRNA levels were reduced to 15% that of control [0.14 ± 0.02 pg CYP2J2 mRNA/0.5 Pg total RNA compared with 0.89 ± 0.16 pg CYP2J2 mRNA/0.5 Pg total RNA for control (P<0.0001); Figure 3.5 C]. These findings demonstrate that CYP2J2 mRNA was significantly decreased in HepG2 cells cultured in hypoxia, but was rapidly restored to control levels upon reoxygenation. While the longer 40 hr hypoxic incubation period produced a somewhat more pronounced decrease in CYP2J2 expression than the 16 hr incubation, it also decreased cell viability significantly (Figure 3.1); most studies that have investigated the effect of hypoxia on gene expression have not generally exceeded 24 hr of treatment (Bandyopadhyay et al., 1995; Norris & Millhorn, 1995; Schmedtje et al., 1997; Mishra et al., 1998; Lee & Corry, 1999; Minet et al., 2001). For these reasons, the 16 hr incubation period was chosen for subsequent experiments.

3.4 Identification of Multiple Potential Binding Sites for the Hypoxia-responsive Transcription Factor AP-1 within the 5’-flanking Region of the CYP2J2 Gene

Prior to the commencement of this thesis, an approximate 2.4 kb fragment of the 5’-flanking region of the CYP2J2 gene was isolated from a human genomic library to facilitate study of the regulation of the gene. The Genomatix MatInspector Professional consensus sequence identification program (Quandt et al., 1995) was used to identify potential transcription factor binding sites within the 2.4 kb CYP2J2 upstream regulatory region. Using high-stringency searching parameters, a number of sequences with high homology to the AP-1 consensus binding element (TGAG/CTCA; Angel & Karin, 1991) were identified within the CYP2J2 5’-flanking region (Table 3.1). Given the

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down-regulation of CYP2J2 mRNA levels in HepG2 cells exposed to hypoxia (Figures 3.2 – 3.5) and similar reports in endothelial cells (Yang et al., 2001), the presence of multiple putative binding sites for the hypoxia-inducible transcription factor AP-1 within the CYP2J2 upstream region was of potential significance and was investigated further.

Table 3.1 Location of potential AP-1 response elements within the upstream regulatory region of the CYP2J2 gene

AP-1-like Match to DNA Strand - Location Sequence Consensus Sense or Antisense TGACTTC 5/7 Sense (+) -2301 to -2295 TGACCGA 5/7 Sense (+) -2135 to -2129 TGACTTG 5/7 Antisense (-) -1911 to -1917 TGACATT 4/7 Sense (+) -1842 to -1836 TGACAGA 5/7 Antisense (-) -1716 to -1722 TGAGTTA 6/7 Sense (+) -1688 to -1682 TAACTCA 6/7 Antisense (-) -1682 to -1688 TGACTCA 7/7 Sense (+) -1681 to -1675 TGAGTCA 7/7 Antisense (-) -1675 to -1681 TGACTTC 5/7 Antisense (-) -1479 to -1485 TGACGTC 4/7 Sense (+) -1420 to -1414 TCAGTCA 6/7 Sense (+) -1326 to -1320 TGACTGA 6/7 Antisense (-) -1320 to -1326 TGACTAG 5/7 Sense (+) -1177 to -1171 TGACCAT 4/7 Antisense (-) -1054 to -1060 TGACCAA 5/7 Sense (+) -1036 to -1030 TGACTCT 6/7 Sense (+) -817 to -811 TGACAAA 5/7 Antisense (-) -397 to -403 CGACGGT 3/7 Antisense (-) -56 to -63 TGAGCCA 6/7 Sense (+) -7 to +1

Potential AP-1 response elements within the CYP2J2 promoter were identified using the Genomatix MatInspector Professional consensus sequence identification program (Quandt et al., 1995). The listed AP-1-like sequences were identified using high-stringency searching parameters (a matrix similarity score of greater than 0.85), with the exception of the sequence at -56 to -63 bp, which was identified under low-stringency parameters (matrix similarity score of 0.70). Nucleotides matching that of the AP-1 consensus binding sequence (TGAG/CTCA) are highlighted in bold, and the location of potential elements is given relative to the translation start site (+1).

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3.5 Analysis of c-Fos and c-Jun Expression in HepG2 Cells During Hypoxia and Reoxygenation

Because potential binding sites for the transcription factor AP-1 were identified within the CYP2J2 5’-flank, the relationship between CYP2J2 expression and that of the AP-1 proteins c-Fos and c-Jun, was explored in HepG2 cells exposed to hypoxia. CYP2J2 mRNA was previously demonstrated to be down-regulated in hypoxia but restored by a 30 min period of reoxygenation (Figure 3.2). Measurement of c-Fos and c-Jun mRNA levels by semi-quantitative RT-PCR indicated that both genes were up-regulated several-fold in hypoxic cells (P<0.005), but were restored to control by a 30 min period of reoxygenation in a similar fashion to that observed with CYP2J2 (Figure 3.6). Thus, the expression of CYP2J2 was inversely related to the increased expression of AP-1; CYP2J2 was down-regulated in hypoxia and c-Fos and c-Jun were increased, but gene expression was normalised during reoxygenation. Protein expression was analysed by immunoblotting of total cell lysates. Hypoxia decreased CYP2J2 immunoreactive protein in HepG2 cells and increased the expression of c-Jun protein (Figure 3.7 A). c-Fos protein was essentially undetectable in cell lysates extracted from normoxic cells, but was strongly up-regulated in hypoxic HepG2 cells (Figure 3.7 A). c-Fos and c-Jun protein levels were normalised in cells after hypoxia-reoxygenation, although CYP2J2 protein remained suppressed (Figure 3.7 B).

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1 n *

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0.2 normoxia hypoxia mRNA levels relative to to relative levels mRNA 0 c-Fos c-Jun

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c-Fos ȕ 0.4 c-Jun Normoxia 0.3 Hypoxia / ȕ-actin / reoxygenation 0.2

0.1 normoxia hypoxia/ reoxygenation to relative levels mRNA 0 c-Fos c-Jun

Figure 3.6 Effect of hypoxia and hypoxia-reoxygenation on c-Fos and c-Jun mRNA expression in HepG2 cells A. Semi-quantitative amplification of c-Fos, c-Jun and ȕ-actin (control) mRNA in HepG2 cells cultured for 16 hr in normoxia (21% O2) or hypoxia (1% O2). RT-PCR was performed as described in Methods, and amplification products were visualised under UV following electrophoresis on 2% agarose gels. B. Densitometric analysis of the expression of c-Fos and c-Jun mRNA relative to ȕ-actin in HepG2 cells cultured for 16 hr in normoxia (21% O2) or hypoxia (1% O2). C. Semi-quantitative amplification of c-Fos, c-Jun and ȕ-actin mRNA in HepG2 cells cultured for 16 hr 30 min in normoxia (21% O2) or hypoxia (16 hr at 1% O2), followed by 30 min reoxygenation (21% O2). D. Densitometric analysis of c-Fos and c-Jun mRNA relative to ȕ-actin in HepG2 cells cultured for 16 hr 30 min in normoxia (21% O2) or hypoxia (16 hr at 1% O2), followed by 30 min reoxygenation (21% O2). Results shown in B and D represent means ± S.E.M., (n=3). *Significantly different from normoxic controls (P<0.005).

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CYP2J2

c-Fos

c-Jun

ȕ-actin standard normoxia hypoxia B

CYP2J2

c-Fos

c-Jun

ȕ-actin standard normoxia hypoxia/ reoxygenation

Figure 3.7 Effect of hypoxia and hypoxia-reoxygenation on CYP2J2, c-Fos and c-Jun protein levels in HepG2 cells A. Immunoblot analysis of CYP2J2, c-Fos and c-Jun protein expression in total cell lysates prepared from HepG2 cells that were cultured for 16 hr in normoxia (21% O2) or hypoxia (1% O2). B. Immunoblot analysis of CYP2J2, c-Fos and c-Jun protein expression in total cell lysates prepared from HepG2 cells cultured for 16 hr 30 min in normoxia (21% O2) or hypoxia (16 hr at 1% O2), followed by 30 min reoxygenation (21% O2). Proteins were separated by electrophoresis on 10% polyacrylamide gels and immunoblotted as described in Methods. Equivalent protein loading between lanes was confirmed by immunoblotting with an anti-actin antibody. Lysate from E.coli cells expressing recombinant CYP2J2 protein was loaded as a standard for CYP2J2. Lysate from phorbol-ester-treated Jurkat cells, and recombinant human c-Jun protein, were loaded as standards for c-Fos and c-Jun respectively.

3.6 Discussion

The results presented in this chapter demonstrate the down-regulation of CYP2J2 mRNA and protein in HepG2 cells exposed to hypoxia. Hypoxia activates the transcription factor AP-1, which regulates the differential expression of several genes in hypoxia (Norris & Millhorn, 1995; Rupec & Baeuerle, 1995; Mishra et al., 1998; Faller, 1999; Lee & Corry, 1999; Michiels et al., 2001; Minet et al., 2001). The identification of a number of sequences within the 5’-flanking region of the CYP2J2 gene that are closely related to the AP-1 consensus binding element (Table 3.1) implicated AP-1 in CYP2J2 regulation in hypoxia. Parallel measurements of

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CYP2J2, and the major components of the AP-1 complex, c-Fos and c-Jun, indicated that the down-regulation of CYP2J2 in hypoxic HepG2 cells was associated with the increased expression of c-Fos and c-Jun mRNA and protein. The activation of AP-1 in HepG2 cells cultured under low oxygen conditions is consistent with previous studies by Bae et al. (1998) and Minet et al. (2001), and has also been reported in numerous other cell types, including human umbilical vein endothelial cells and bovine aortic endothelial cells (Bandyopadhyay et al., 1995), HeLa cells (Rupec & Baeuerle, 1995; Müller et al., 1997), SiHa cells (Ausserer et al., 1994), and MCF-7 cells (Lee & Corry, 1999). While the activation of AP-1 in hypoxia has been clearly demonstrated in the literature, the effect of hypoxia followed by a period of reoxygenation on the expression and activity of AP-1 is controversial. There are reports that AP-1 activity remains elevated following reoxygenation, while other studies have found that the mRNAs corresponding to certain AP-1 subunits are increased in hypoxia and decreased by reoxygenation (Yao et al., 1994; Rupec & Baeuerle, 1995). Similarly, AP-1 has been reported to be activated in ischaemia-reperfusion (Brand et al., 1992; Morooka et al., 1995), or to be activated in ischaemia, normalised during the early reoxygenation phase, and then elevated by prolonged reoxygenation (Tacchini et al., 1999). Results presented in this chapter indicated that the expression of c-Fos and c-Jun in HepG2 cells exposed to hypoxia followed by reoxygenation was not significantly different from normoxic control, which is consistent with the rapid restitution, at both the mRNA and protein level, of these immediate early genes that control the acute-phase response to external stresses. Consistent with the apparent relationship to AP-1 subunit expression, CYP2J2 mRNA levels in HepG2 cells returned to control levels after hypoxia-reoxygenation. CYP2J2 protein, on the other hand, remained suppressed after hypoxia-reoxygenation; this is consistent with the results of Yang et al. (2001), who reported the down-regulation of CYP2J protein in hypoxia followed by 4 hr reoxygenation. A number of studies have documented rapid increases in CYP mRNAs, but delayed synthesis of the corresponding proteins following in vivo exposure to foreign compounds. For example, in a study of rats administered the industrial chemical pyridine, maximal induction of hepatic CYP1A1 mRNA was observed at 9 hr post-treatment, whereas maximal induction of CYP1A1 protein was not observed until 24 hr after treatment; a similar pattern was also observed for the induction of renal CYP1A1 mRNA and protein (Iba et al., 1999). Thus, the time required for the restoration of CYP2J protein could well be longer than that required for normalisation of the mRNA, which would explain the discrepancy between CYP2J2 mRNA and protein levels in hypoxia-reoxygenation.

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Decreased expression of CYP2J2 under low oxygen conditions is likely to be of pathophysiological significance. As outlined previously (section 1.5.1 – 1.5.4), EETs, the cellular products of metabolism of AA by CYP2J2, have a number of cytoprotective physiological actions including vasodilatory, anti-inflammatory and anti-apoptotic effects (Node et al., 1999; Chen et al., 2001; Fleming, 2001; Yang et al., 2001; Roman, 2002; Spiecker & Liao, 2005). As such, the decreased expression of CYP2J2, and the resultant decline in cytoprotective EETs, may contribute to the cellular damage associated with pathogenic conditions characterised by tissue hypoxia, including ischaemic heart disease and stroke (Yang et al., 2001). Thus, maintenance of CYP2J2 in cells exposed to low oxygen conditions, via transfection of CYP2J2 cDNA, or treatment with CYP2J2-derived EETs, limited cellular injury and enhanced survival (Yang et al., 2001). Interestingly, the expression of eNOS is also decreased in endothelial cells cultured in a low oxygen environment (Liao et al., 1995; Phelan & Faller, 1996; Giraldez et al., 1997; Faller, 1999). NO and EETs share several protective properties such as potent vasodilatory and anti-inflammatory properties. Similar to the situation with CYP2J2, the decreased expression of eNOS may be an important event that contributes to the pathogenesis of ischaemic injury (Liao et al., 1995; Giraldez et al., 1997; Faller, 1999; Yang et al., 2001). In summary, findings presented in this chapter have suggested an apparent relationship between the down-regulation of CYP2J2 in hypoxia and the up-regulation of the AP-1 subunits c-Fos and c-Jun. The role of AP-1 in the regulation of CYP2J2 expression under normal and reduced oxygen conditions was further investigated in experiments presented in the following chapters.

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Chapter 4

Characterisation of an AP-1 Binding Sequence within the CYP2J2 Proximal Promoter that Regulates CYP2J2 Expression in Normoxia and Hypoxia

4.1 Introduction

Results presented in Chapter 3 showed that the expression of CYP2J2 mRNA and protein is significantly decreased in hypoxic HepG2 cells, and provided preliminary evidence for a role for AP-1 in this setting. Thus, the down-regulation of CYP2J2 in hypoxic HepG2 cells was inversely associated with the up-regulation of the AP-1 proteins c-Fos and c-Jun (Figures 3.2 to 3.7). Numerous potential AP-1 binding sequences were identified within the 5’-regulatory region of the CYP2J2 gene (Table 3.1). Experiments presented in this chapter were designed to investigate the role of AP-1 in the regulation of CYP2J2 under normoxic and hypoxic conditions. As described in Chapter 3, a 2.4 kb fragment of the 5’-flanking region of the human CYP2J2 gene was isolated from a human genomic library to facilitate a molecular analysis of CYP2J2 gene regulation. This 2.4 kb fragment of the CYP2J2 gene corresponded to nucleotides -2341 to +98 relative to the translation start site (Figure 4.1), and contained the major transcription start site 26 bp upstream of the translation start site (King et al., 2002; GenBank accession number AF272142). Our sequence was also submitted to GenBank and has the accession number AF039089.

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Figure 4.1 Nucleotide sequence of the 5’-flanking region of the human CYP2J2 gene An approximate 2.4 kb fragment of the 5’-flanking region of the human CYP2J2 gene, corresponding to nucleotides -2341 to +98 bp (relative to the translation start site), was isolated from a human genomic library. This fragment was cloned into the pGL3basic reporter vector to facilitate a molecular analysis of the transcriptional regulation of the CYP2J2 gene. Potential AP-1 transcription factor binding sites identified using the Genomatix MatInspector Professional consensus sequence identification software are labelled and marked by arrows (the 3’ or 5’ direction of arrows indicates binding sites on the sense and antisense strands, respectively). The potential CCAAT box and Sp1 binding sequences identified within the proximal promoter region are also highlighted. The major transcription start site identified by King et al. (2002) is marked with an asterisk, and the translation start site (+1) is highlighted in bold. The restriction enzyme recognition sites used to create the CYP2J2 promoter deletion constructs 2J2 (-1894/+98), 2J2 (-1228/+98), 2J2 (-574/+98) and 2J2 (-152/+98) are also indicated.

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The Genomatix MatInspector Professional consensus sequence identification program (Quandt et al., 1995) was used to identify potential transcription factor binding sites within the CYP2J2 5’-flank. Consistent with the sequence reported subsequently by King et al. (2002), no canonical TATA box was identified within the CYP2J2 proximal promoter, although a potential CCAAT box, as well as several GC-rich regions, which are known to bind the ubiquitously expressed transcription factor Sp1 (Lania et al., 1997), were present (Figure 4.1). Importantly, as outlined previously (Chapter 3), a number of sequences with high homology to the AP-1 consensus binding motif were identified within the CYP2J2 5’-flanking region (Table 3.1, Figure 4.1).

4.2 Differential Activation of the CYP2J2 Promoter in HepG2 Cells by AP-1 Proteins

The apparent inverse association between CYP2J2 and AP-1 expression in hypoxia (Chapter 3) prompted a molecular analysis of the effects of AP-1 proteins on transcriptional activity of the CYP2J2 gene. The 2.4 kb 5’-flanking sequence of the CYP2J2 gene (Figure 4.1) was cloned into the pGL3basic luciferase reporter vector to produce 2J2 (-2341/+98). To test the intrinsic AP-1-responsiveness of the CYP2J2 promoter, transient transfection studies were performed with the reporter construct 2J2 (-2341/+98) in HepG2 cells, with a range of expression plasmids encoding members of the AP-1 complex. c-Jun was found to be a strong inducer of CYP2J2 promoter activity, activating 2J2 (-2341/+98) to approximately 7-fold that of untransfected control (Figure 4.2). The related proteins JunB and JunD also stimulated CYP2J2 promoter activity, but to a lesser extent than c-Jun (Figure 4.2). In contrast, co-transfection of Fos family genes (c-Fos, Fra-1 and Fra-2) did not stimulate reporter activity. Furthermore, with the exception of the combination of JunB and Fra-1, Fos proteins abolished the activation of the reporter construct 2J2 (-2341/+98) produced by Jun proteins. Thus, the data presented in Figure 4.2 indicates that the CYP2J2 promoter is differentially regulated by AP-1 proteins; CYP2J2 promoter activity is strongly enhanced by c-Jun, but this induction is abolished by the Fos proteins.

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2J2 (-2341/+98) 8 6 4 2 0 -2 Fold Difference Fold -4 -6 JunB JunD Fra-1 Fra-2 c-Jun c-Fos JunB + JunD + JunB Fra-1 +Fra-1 JunB +Fra-1 JunD +Fra-2 JunB +Fra-2 JunD c-Jun + JunB c-Jun + JunD c-Jun Fra-1 + c-Jun Fra-1 + c-Jun Fra-2 c-Fos + JunB + c-Fos JunD + c-Fos c-Fos + c-Jun + c-Fos

Figure 4.2 Differential effects of AP-1 proteins on the CYP2J2 promoter A large fragment of the CYP2J2 gene containing 2341 bp of the upstream regulatory region, relative to the translation start site (+1), was cloned into the pGL3basic reporter vector to produce the CYP2J2 promoter construct 2J2 (-2341/+98). 2J2(-2341/+98) was co-transfected (1 μg per well) into HepG2 cells with combinations of expression plasmids encoding AP-1 proteins (0.5 μg per well). A pCMV-ȕ-galactosidase expression plasmid was included in each well to control for transfection efficiency (0.5 μg per well). Cells were harvested and assayed for luciferase and ȕ-galactosidase activity as described in Methods. Luciferase activity was normalised to ȕ-galactosidase activity, with results presented as fold differences relative to the activity of 2J2 (-2341/+98) in the absence of co-transfected AP-1 proteins (set to 1-fold). Results shown are means ± S.E.M. for three independent experiments.

4.3 Location of a c-Jun-responsive Region within the Proximal Region of the CYP2J2 5’-flank

A series of promoter deletion constructs were prepared by 5’-truncation of 2J2 (-2341/+98) and used in transfection studies to identify the region of the CYP2J2 gene responsible for mediating the divergent regulation by c-Jun and c-Fos. The activity of CYP2J2 promoter constructs in HepG2 cells was assessed following co-transfection with c-Fos and c-Jun expression plasmids. Consistent with the results presented in Figure 4.2, the full-length construct 2J2 (-2341/+98) was strongly activated by over-expression of c-Jun (to 8.2 ± 1.5-fold of control; Figure 4.3), while over-expression of c-Fos did not enhance reporter activity (1.2 ± 0.4-fold of control) and abolished induction by c-Jun (1.1 ± 0.4-fold of control). A similar pattern of transactivation was seen with the deletion constructs 2J2 (-1894/+98), 2J2 (-1228/+98) and 2J2 (-574/+98); all of these deletion constructs

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exhibited c-Jun-responsiveness and inhibition of c-Jun-dependent activation by c-Fos (Figure 4.3). The smallest deletion construct 2J2 (-152/+98) also remained strongly inducible by c-Jun (to 4.1 ± 0.3-fold of control). Furthermore, the c-Jun-dependent activation of this construct was suppressed by c-Fos (Figure 4.3). These results indicate that transactivation of CYP2J2 by c-Jun, and repression by c-Fos, is mediated by element(s) in the proximal promoter region, within 152 bp of the CYP2J2 start codon.

-2341 +1 +98 2J2 (-2341/+98)

-1894 2J2 (-1894/+98)

-1228 2J2 (-1228/+98)

-574 construct alone 2J2 (-574/+98) + c-Fos AP-1-like element -152 + c-Jun 2J2 (-152/+98) + c-Fos/c-Jun

024681012 Fold Difference

Figure 4.3 Location of a c-Jun-responsive region in the CYP2J2 gene Identification of a c-Jun-responsive region in the CYP2J2 gene using 2J2 (-2341/+98) and the CYP2J2 promoter deletion constructs 2J2 (-1894/+98), 2J2 (-1228/+98), 2J2 (-574/+98) and 2J2 (-152/+98), which were prepared by 5’-truncation of 2J2 (-2341/+98). Potential AP-1 response elements located within the constructs are indicated by black boxes. HepG2 cells were co-transfected with CYP2J2 reporter constructs (1 ȝg per well) and expression plasmids encoding the AP-1 proteins c-Fos and c-Jun (0.5 ȝg per well) as indicated. A ȕ-galactosidase expression plasmid was included in each well to control for transfection efficiency (0.5 ȝg per well). Following transfection, cells were harvested, and luciferase and ȕ-galactosidase activity was measured as outlined in Methods. Results represent luciferase activity corrected for ȕ-galactosidase activity, and are expressed as fold differences relative to the activity of the construct in the absence of co-transfected protein (set to 1-fold). Data shown are means ± S.E.M. for three independent experiments.

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4.4 Identification of a Functional c-Jun Binding Site at -56/-63 bp within the CYP2J2 Proximal Promoter

4.4.1 Binding of c-Jun to the CYP2J2 Proximal Promoter Given the results obtained from transient transfection studies indicating that the 152 bp sequence immediately upstream of the CYP2J2 start codon is important in mediating c-Jun-responsiveness (Figure 4.3), the binding of c-Jun within this region was evaluated in a series of electrophoretic mobility shift assays (EMSAs). A 167 bp double-stranded fragment (2J2/167) corresponding to nucleotides (nt) -152 to +15 of the CYP2J2 promoter (Figure 4.4 A) was generated from the deletion construct 2J2 (-152/+98) by digestion with the restriction enzymes Nheȱ and Banȱȱ as outlined in Methods. This fragment was labelled with [32P]dCTP and used as a probe in EMSAs with nuclear protein fractions prepared from untransfected HepG2 cells or HepG2 cells that had been transfected with a c-Jun expression plasmid (Figure 4.4 B and C). Two shifted complexes, representing distinct DNA-protein interactions, were observed upon incubation of the labelled 2J2/167 probe with nuclear protein fractions from untransfected HepG2 cells (shifts 1 and 2; Figure 4.4 B, lane 2). Both shifted complexes were competed by a 200-fold excess of unlabelled 2J2/167 probe (self), but not by an unrelated probe containing the STAT5 element from the ȕ-casein promoter (stat; Figure 4.4 B, lanes 3 and 4), thus confirming the specific nature of the two distinct DNA-protein interactions observed with the 2J2/167 probe. To evaluate the presence of c-Jun within the protein complexes bound to the 2J2/167 probe, supershift experiments were performed using antibodies directed against c-Jun, or the unrelated protein ubiquitin, which was included as a negative control. As shown in Figure 4.4 B, antibodies directed against c-Jun, but not ubiquitin, enhanced the apparent amount of a supershifted complex (lanes 5 and 6), indicating that c-Jun is part of the protein complex bound to the CYP2J2 proximal promoter in HepG2 cells.

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A 2J2/167

-152 CCCGGGAATCCAGCGCCTGGCATCTTCGCAGGGTGCTGCGAAGGGGCGGGCTGGGAGGCGGGGCACGGCTGGGAGCGAGGCGG GGCGGGGACCGTCGCCTGCTGGGACCGCCGCCTGCTTGGACCGCAGAAGAGCAGGAGGACGTCTGAGCCATGCTCGCGGCGATG +15

B Competitor - - self stat - - C Antibody - - - - Jun Ub Anti-c-Jun - - - +

Protein - UT UT UT UT UT Protein - UT JT JT supershift supershift

shift 3 shift 1 shift 1

shift 2 shift 2

1 2 3 4 5 6 1 2 3 4

Probe 2J2/167 Probe 2J2/167

D Competitor - - AP-1 rec. c-Jun - + +

shift

1 2 3

Probe 2J2/167

Figure 4.4 EMSAs showing the binding of nuclear protein and recombinant c-Jun protein to the CYP2J2 proximal promoter A. Sequence of 2J2/167, a 167 bp probe corresponding to -152 to +15 bp of the CYP2J2 gene (relative to the translation start site). 2J2/167 was end-labelled with [32P]dCTP and used in EMSAs with nuclear protein fractions prepared from untransfected (UT; B, C) and c-Jun-transfected (JT; C) HepG2 cells, and with recombinant c-Jun protein (D). For competition experiments, 200-fold excess of unlabelled 2J2/167 probe (self), stat probe (STAT5 element from the ȕ-casein promoter) or AP-1 probe (containing the AP-1 consensus binding sequence) was included in the binding reaction, and for EMSAs with antibodies, nuclear proteins were pre-incubated with antibodies to c-Jun (Jun) or ubiquitin (Ub) prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Shifted and supershifted complexes are indicated by arrows.

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Incubation of the 2J2/167 probe with nuclear protein fractions prepared from c-Jun-transfected HepG2 cells produced a more intense signal at shift 1, and also resulted in the appearance of an additional shifted complex (shift 3) with slower mobility compared to shifts 1 and 2 (Figure 4.4 C, lanes 2 and 3). In contrast, the signal intensity of shift 2 was not altered with the use of nuclear protein from c-Jun-transfected HepG2 cells (lanes 2 and 3). These results suggested that c-Jun is present within the DNA-protein complex giving rise to shifts 1 and 3, but that interaction with a protein other than c-Jun produces shift 2. Pre-incubation of the c-Jun-transfected HepG2 nuclear extract with anti-c-Jun antibody resulted in the appearance of a prominent supershifted complex, and a parallel decrease in signal intensity at shifts 1 and 3, confirming the presence of c-Jun in the DNA-protein complexes producing shifts 1 and 3 (Figure 4.4 C, lane 4). To confirm that c-Jun binds directly to the CYP2J2 proximal promoter, additional EMSAs were performed using the 2J2/167 probe and recombinant human c-Jun protein. As seen in Figure 4.4 D, a prominent shifted complex was observed upon incubation of 2J2/167 with recombinant c-Jun (lane 2), which was successfully competed by 200-fold excess of unlabelled probe that contained the AP-1 consensus binding sequence (AP-1; lane 3). Thus, EMSAs presented in Figure 4.4 indicate that c-Jun protein binds within the 152 bp sequence of CYP2J2 immediately upstream of the translation start site. Consistent with the strong activation of CYP2J2 proximal promoter constructs observed in co-transfection studies with c-Jun, binding within this c-Jun-responsive region was enhanced upon transfection with c-Jun. Furthermore, EMSAs using recombinant c-Jun protein demonstrated the capacity of c-Jun homodimers to interact directly with the CYP2J2 proximal promoter.

4.4.2 Analysis of the Binding of Nuclear Protein and Recombinant c-Jun Protein to Sequences Resembling the AP-1 Consensus within the 152 bp c-Jun-responsive Region of the CYP2J2 Promoter Results presented in sections 4.3 and 4.4.1 indicate that the 152 bp sequence of the CYP2J2 promoter immediately upstream of the start of the gene binds c-Jun and mediates transactivation of the CYP2J2 promoter by c-Jun. Using the Genomatix MatInspector software, two sequences resembling the AP-1 consensus binding motif were identified within the 152 bp c-Jun-responsive region of CYP2J2. The first AP-1-like element, located at -7 to +1 bp on the positive strand, is highly homologous to the AP-1 consensus sequence, while the second

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AP-1-like element, located at -56 to -63 bp on the negative strand, is less related to the AP-1 consensus and was only identified by low-stringency searching (Figure 4.5). The ability of these AP-1-like elements, located within the c-Jun-responsive region of the CYP2J2 promoter, to interact with c-Jun, was assessed in a series of EMSA experiments. 32P-labelled oligonucleotide probes containing the AP-1-like element at -7 to +1 bp [2J2-(-7/+1) probe] or the AP-1-like element at -56 to -63 bp [2J2-(-56/-63) probe] were prepared (Figures 4.6 A and 4.7 A, respectively), and incubated with nuclear protein fractions extracted from untransfected and c-Jun-transfected HepG2 cells; EMSA reactions with recombinant c-Jun protein were also performed to directly evaluate binding to c-Jun.

-7/+1 bp -152 TGAGCCA +1 (+) A -152 TGGCAGC +1 (-) -56/-63 bp

B TGAG/CTCA

Figure 4.5 AP-1-like elements within the c-Jun-responsive region of the CYP2J2 promoter A. Sequences of AP-1-like binding elements within the c-Jun-responsive region (-152/+1) of the CYP2J2 promoter. Nucleotides sharing homology with the AP-1 consensus binding sequence (B) are indicated in bold. Potential AP-1 binding sites were identified by the Genomatix MatInspector consensus sequence identification software. The AP-1-like element spanning -7/+1 bp on the positive strand of CYP2J2 (+) is highly homologous to the AP-1 consensus binding sequence, while the AP-1-like element spanning -56/-63 bp on the negative strand of CYP2J2 (-) deviates further from the consensus sequence. Nucleotide numbers are given relative to the translation start site (+1).

Incubation of the 2J2-(-7/+1) probe with nuclear protein fractions from untransfected HepG2 cells produced a shifted complex of low intensity (Figure 4.6 B, lane 1), which was more pronounced in nuclear extracts from c-Jun-transfected cells (lane 2). This shifted complex was effectively competed by 200-fold excess of unlabelled 2J2-(-7/+1) probe (self; lane 3), but not by excess of the unrelated probe containing the STAT5 element from the ȕ-casein promoter (stat; lane 4), thus confirming the specific nature of the DNA-protein interaction. The signal of greater mobility seen in EMSAs represented a non-specific interaction as it was unaffected by excess of the unlabelled self probe. The presence of c-Jun, but not c-Fos, in the specific DNA-protein complex was demonstrated by disruption (or block-shift) of the retarded complex by an anti-c-Jun antibody (lane 5), but not by an anti-c-Fos antibody (lane 6). The complex was unaffected by an antibody directed against the unrelated

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protein ubiquitin, included routinely as a negative control in antibody supershift experiments (lane 7). The 2J2-mt-(-7/+1) probe was created by mutagenesis of the AP-1-like sequence spanning -7 to +1 bp (Figure 4.6 A) and was used in EMSAs to assess the importance of the -7 to +1 bp element in DNA-protein complex formation. As shown in Figure 4.6 C, the 2J2-mt-(-7/+1) probe produced a substantially weaker interaction with nuclear extracts from untransfected and c-Jun-transfected cells compared to that with the wild type probe. The ability of the -7 to +1 bp sequence to directly bind c-Jun was confirmed in EMSAs with recombinant c-Jun protein; a retarded complex was observed upon incubation of recombinant c-Jun protein with the wild type 2J2-(-7/+1) probe (Figure 4.6 D, lane 2), but not with the 2J2-mt-(-7/+1) probe (lane 4). Similar EMSAs were conducted to assess the capacity of c-Jun to interact with the second AP-1-like element identified within the c-Jun-responsive region of CYP2J2 (Figure 4.7 A). As indicated previously, the second AP-1-like element, located at -56 to -63 bp within the CYP2J2 5’-flank, is less related to the AP-1 consensus than the -7 to +1 bp sequence, and was only identified by the MatInspector software using low- stringency searching parameters. Despite this, incubation of the 2J2-(-56/-63) probe with nuclear protein fractions from untransfected HepG2 cells produced a specific retarded complex of stronger intensity than that observed with the 2J2-(-7/+1) probe (Figure 4.7 B, lane 2). The specificity of the DNA-protein interaction represented by this shifted complex was demonstrated by competition with 200-fold excess of unlabelled 2J2-(-56/-63) probe (self; lane 4), but not by 200-fold excess of the unrelated STAT5 element probe (stat; lane 5); the lower running signal was not affected by the inclusion of unlabelled competitor and thus represented a non-specific interaction. The intensity of the specific shifted complex observed with nuclear extracts from untransfected HepG2 cells (lane 2) was increased in nuclear extracts from c-Jun-transfected HepG2 cells (lane 3), and was block-shifted by an antibody directed against c-Jun (lane 7), but not by antibodies directed against c-Fos (lane 6) or ubiquitin (lane 8). Thus, c-Jun, but not c-Fos, is present in the protein complex bound to this probe.

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A GAGGACGTCTGAGCCATGCTCGC GAGGACGTAAAAAAAATGCTCGC 2J2-(-7/+1)probe 2J2-mt-(-7/+1) probe

B Competitor - - self stat - - - Antibody - - - - Jun Fos Ub Protein UT JT UT UT UT UT UT

shift

1 2 3 4 5 6 7

Probe 2J2-(-7/+1)

C Protein UT JT UT JTD Protein - Jun - Jun

shift shift

1 2 3 4 1 2 3 4

Probe 2J2- 2J2-mt- Probe 2J2- 2J2-mt- (-7/+1) (-7/+1) (-7/+1) (-7/+1)

Figure 4.6 EMSAs evaluating the binding of c-Jun to the AP-1-like element at -7/+1 in the CYP2J2 promoter A. Sequences of probes used in EMSAs to evaluate the binding of c-Jun to the -7/+1 bp AP-1-like element within the CYP2J2 promoter. 2J2-(-7/+1) incorporates the wild type -7/+1 bp element (bolded); 2J2-mt-(-7/+1) is mutated at the -7/+1 bp element as indicated by asterisks. Probes were labelled with [32P]dCTP and incubated with nuclear protein fractions from untransfected (UT) and c-Jun-transfected (JT) HepG2 cells (B, C), and with recombinant c-Jun protein (Jun; D) as described in Methods. For competition experiments, 200-fold excess unlabelled 2J2-(-7/+1) probe (self) or stat probe (containing the STAT5 element from the ȕ-casein promoter) was included in the binding reaction, and in antibody experiments, nuclear protein was pre-incubated for 1 hr with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows.

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A CCCAGCAGGCGACGGTCCCCGCCCCG CCCAGCAGGTTTTTTTTCCCGCCCCG 2J2-(-56/-63)probe 2J2-mt-(-56/-63) probe

B Competitor - - - self stat - - - Antibody - - - - - Fos Jun Ub Protein - UT JT UT UT UT UT UT

shift

1 2 3 4 5 6 7 8

Probe 2J2-(-56/-63)

C Protein UT JT UT JTD Protein - Jun - Jun

shift shift

1 2 3 4 1 2 3 4

Probe 2J2- 2J2-mt- Probe 2J2- 2J2-mt- (-56/-63) (-56/-63) (-56/-63) (-56/-63)

Figure 4.7 EMSAs showing the binding of c-Jun to the AP-1-like element at -56/-63 in the CYP2J2 promoter A. Sequences of probes used in EMSAs to evaluate the binding of c-Jun to the -56/-63 bp AP-1-like element within the CYP2J2 promoter. 2J2-(-56/-63) incorporates the wild type -56/-63 bp element (bolded); 2J2-mt-(-56/-63) is mutated at the -56/-63 bp element as indicated by asterisks. Probes were labelled with [32P]dCTP and incubated with nuclear protein fractions from untransfected (UT) and c-Jun-transfected (JT) HepG2 cells (B, C), and with recombinant c-Jun protein (Jun; D) as described in Methods. For competition experiments, 200-fold excess unlabelled 2J2-(-56/-63) probe (self) or stat probe (containing the STAT5 element from the ȕ-casein promoter) was included in the binding reaction, and in antibody experiments, nuclear protein was pre-incubated for 1 hr with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows.

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The importance of the -56 to -63 bp AP-1-like element in mediating DNA-protein interaction was assessed specifically in EMSAs with the 2J2-mt-(-56/-63) probe, in which the -56 to -63 bp sequence is mutated (Figure 4.7 A). As shown in Figure 4.7 C, the 2J2-mt-(-56/-63) probe produced only a very weak interaction with nuclear extracts from untransfected and c-Jun-transfected HepG2 cells compared to that of the wild type probe, which indicates that the -56 to -63 bp AP-1-like element is critical for protein binding (Figure 4.7 C). To confirm the ability of c-Jun to interact directly with the -56 to -63 bp AP-1-like element, EMSAs were performed with recombinant c-Jun protein. As shown in Figure 4.7 D, a prominent shifted complex was observed upon incubation of the 2J2-(-56/-63) probe with recombinant c-Jun (lane 2). In contrast, no interaction was apparent with the mutant probe (lane 4). Comparative EMSAs were also conducted with a probe containing the AP-1 consensus binding sequence (Figure 4.8 A). Incubation of the AP-1 consensus probe with HepG2 nuclear extracts produced a shifted complex of very similar mobility to that seen with the 2J2-(-56/-63 probe) in Figure 4.7 (Figure 4.8 B, lane 2). The signal observed in nuclear protein fractions from untransfected HepG2 cells (lane 2) was more intense in fractions from c-Jun-transfected cells (lane 3), while the specificity of the shifted complex was demonstrated by competition of binding by 200-fold excess unlabelled AP-1 probe (self; lane 4), but not by 200-fold excess of the unrelated STAT5 element probe (stat; lane 5). Similar to the results obtained with the probes containing the AP-1-like elements present within the CYP2J2 proximal promoter, the complex formed between the AP-1 consensus probe and HepG2 nuclear extracts was disrupted by incubation with c-Jun antibody (lane 6), but not by incubation with anti-c-Fos (lane 7) or anti-ubiquitin (lane 8) antibodies. Thus, c-Jun, but not c-Fos, forms the protein complex bound to the AP-1 consensus element in untransfected HepG2 cells.

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A TGATGAGTCAGCCGGATC AP-1 consensus probe

B Competitor - - - self stat - - - Antibody - - - - - Jun Fos Ub Protein - UT JT UT UT UT UT UT

shift

1 2 3 4 5 6 7 8

Probe AP-1 consensus

Figure 4.8 Binding of HepG2 nuclear extract to the AP-1 consensus binding motif A. Sequence of the AP-1 consensus probe used in EMSAs, with the AP-1 consensus binding motif highlighted in bold. B. c-Jun, but not c-Fos, forms part of the protein complex bound to the AP-1 consensus element in untransfected HepG2 cells. The AP-1 consensus probe was labelled with [32P]dCTP and incubated with nuclear protein fractions from untransfected (UT) and c-Jun-transfected (JT) HepG2 cells as described in Methods. Competition experiments were performed by the inclusion of 200-fold excess unlabelled AP-1 consensus probe (self) or stat probe (containing the STAT5 element from the ȕ-casein promoter) in the binding reaction, and antibody experiments were performed by pre-incubation of nuclear protein with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) for 1 hr prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Shifted complexes are indicated by arrows.

4.4.3 The AP-1-like Element at -56 to -63 bp within the CYP2J2 Proximal Promoter is Important in Mediating Transactivation by c-Jun EMSAs shown in Figures 4.6 and 4.7 indicated that both AP-1-like elements located within the c-Jun-responsive region of the CYP2J2 proximal promoter are capable of binding c-Jun. To determine whether these elements play a functional role in c-Jun-dependent transactivation of the CYP2J2 promoter, CYP2J2 reporter constructs were prepared in which the -7 to +1 bp or -56 to -63 bp elements were mutated or deleted. The effect of these mutations or deletions on c-Jun-dependent promoter activity was assessed in transfection studies.

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As shown in Figure 4.9, activation by c-Jun of the construct 2J2 (-152/+98; mt -7/+1), in which the -7 to +1 bp sequence is mutated, was not different to that of the wild type construct 2J2 (-152/+98). This finding indicates that the AP-1-like element at -7 to +1 bp does not mediate c-Jun-dependent transactivation of the CYP2J2 promoter. Deletion of the -152 to -50 bp region generated the reporter construct 2J2 (-49/+98), which contained the -7 to +1 bp sequence, but not the sequence at -56 to -63 bp. This construct was not inducible by c-Jun (Figure 4.9), confirming the lack of involvement of the -7 to +1 bp sequence and suggesting an important role for the -56 to -63 bp element in c-Jun-dependent activation.

-152 -56/-63 -7/+1 +98 2J2 (-152/+98)

-152 -56/-63 -7/+1 +98 2J2 (-152/+98; mt -7/+1) mutated - c-Jun -49 -7/+1 +98 + c-Jun 2J2 (-49/+98) * -152 -56/-63 -7/+1 +98 2J2 (-152/+98; mt -56/-63) mutated * -1012345

-56/-63 AP-1-like site Fold Difference -56/-63 AP-1-like site, mutated -7/+1 AP-1-like site -7/+1 AP-1-like site, mutated

Figure 4.9 Role of the -7/+1 and -56/-63 bp AP-1-like elements in mediating c-Jun-dependent activation of the CYP2J2 promoter The CYP2J2 promoter deletion construct 2J2 (-49/+98) was prepared by 5’-truncation of 2J2 (-152/+98), while the mutagenised constructs 2J2 (-152/+98; mt -7/+1) and 2J2 (-152/+98; mt -56/-63) were prepared by site-directed mutagenesis of the AP-1-like elements at -7/+1 bp and -56/-63 bp, respectively. CYP2J2 reporter constructs were transfected (1 ȝg per well) into HepG2 cells with a ȕ-galactosidase expression plasmid (0.5 ȝg per well; to control for transfection efficiency). A c-Jun expression plasmid was co-transfected at 0.5 ȝg per well as indicated. Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity. The normalised activity of reporter constructs in the absence of c-Jun was set to 1-fold, and the activity of constructs in the presence of c-Jun was expressed as fold differences relative to activity in the absence of c-Jun. Results shown are means ± S.E.M. for at least three independent experiments. *Significantly different from 2J2 (-152/+98) in the presence of c-Jun (P<0.0001).

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To directly assess the role of the -56 to -63 bp element, the mutagenised construct 2J2 (-152/+98; mt -56/-63) was prepared (Figure 4.9). Activation of 2J2(-152/+98; mt -56/-63) by c-Jun was decreased (2.4 ± 0.2-fold of control) relative to the wild type construct 2J2 (-152/+98) [3.9 ± 0.2-fold of control; P<0.0001]. Thus, the -56 to -63 bp AP-1-like element plays an important role in mediating activation of CYP2J2 by c-Jun. However, while producing a significant reduction in c-Jun-dependent activity, mutagenesis of the -56 to -63 bp element did not result in the complete elimination in c-Jun-responsiveness that was observed with the deletion construct 2J2 (-49/+98), suggesting that an additional element between nt -152 to -50 may contribute to c-Jun-responsiveness. Analysis of the basal transcriptional activity of the CYP2J2 mutation and deletion constructs described above generated results that were consistent with those obtained for c-Jun-dependent activation. Thus, as shown in Figure 4.10, mutagenesis of the -7 to +1 bp sequence did not diminish basal activation of the CYP2J2 promoter in HepG2 cells. In contrast, basal promoter activity was significantly impaired by mutagenesis of the -56 to -63 bp AP-1-like element [the basal transcriptional activity of 2J2 (-152/+98; mt -56/-63) was 4.6 ± 0.4-fold that of the pGL3basic vector backbone, compared to 6.0 ± 0.4-fold for the wild type construct 2J2 (-152/+98); Figure 4.10; P<0.01]. Mirroring the pattern observed for c-Jun-dependent activation, deletion of the -152 to -50 bp sequence produced a more pronounced effect, with the construct 2J2 (-49/+98) displaying a basal promoter activity only 1.7 ± 0.2-fold higher than the promoterless pGL3basic backbone, compared with 6.0 ± 0.4-fold for 2J2 (-152/+98) [Figure 4.10; P<0.0001]. These results are consistent with a role for c-Jun in maintaining CYP2J2 gene expression in cells by acting at c-Jun-response elements within the -152 to -50 bp region of the CYP2J2 promoter.

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pGL3basic

-152 -56/-63 -7/+1 +98 2J2 (-152/+98)

-152 -56/-63 -7/+1 mt +98 2J2 (-152/+98; mt -7/+1)

-49 -7/+1 +98 2J2 (-49/+98) ** -152 -56/-63 mt -7/+1 +98 2J2 (-152/+98; mt -56/-63) *

02468 -56/-63 AP-1-like site Fold Difference -56/-63 AP-1-like site, mutated -7/+1 AP-1-like site -7/+1 AP-1-like site, mutated

Figure 4.10 Effect of mutating the -7/+1 and -56/-63 bp AP-1-like elements on the basal transcriptional activity of the CYP2J2 promoter The CYP2J2 reporter constructs 2J2 (-152/+98), 2J2 (-152/+98; mt -7/+1), 2J2 (-49/+98) and 2J2 (-152/+98; mt -56/-63), or the empty pGL3basic vector, were co- transfected (1 ȝg per well) into HepG2 cells with a ȕ-galactosidase expression plasmid (0.5 ȝg per well; to control for transfection efficiency). Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity and presented as fold differences relative to the normalised activity of the empty pGL3basic vector (set to 1-fold). Results shown are means ± S.E.M. for at least three independent experiments. *Significantly different from 2J2 (-152/+98) [P<0.01], **significantly different from 2J2 (-152/+98) [P<0.0001].

4.5 Effect of Hypoxia on the Binding of c-Jun to the AP-1-like Element at -56 to -63 bp in the CYP2J2 Proximal Promoter

To assess the effect of hypoxia on protein binding to the -56/-63 AP-1-like element within the CYP2J2 promoter, EMSA studies were conducted using the labelled 2J2-(-56/-63) probe and nuclear extracts prepared from HepG2 cells that had been cultured under hypoxic and normoxic conditions. Incubation of probe 2J2-(-56/-63) with nuclear protein fractions from normoxic cells produced a shifted complex (Figure 4.11 A, lane 2), that was competed by 200-fold excess of unlabelled 2J2-(-56/-63) probe (self; lane 3), but not by 200-fold excess of the unrelated probe containing the STAT5 element from the ȕ-casein promoter (stat; lane 4). This retarded complex was block-shifted by an anti-c-Jun antibody (lane 6), but not by anti-c-Fos

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(lane 5) or anti-ubiquitin (lane 7) antibodies. Thus, c-Jun, but not c-Fos, binds to the -56 to -63 bp element under normal oxygen conditions. In hypoxia, the intensity of the shifted complex was markedly decreased (lane 8), and was apparently unaffected by either of the antibodies directed against c-Fos or c-Jun (lanes 9 and 10), indicating that the binding of c-Jun to the -56/-63 AP-1-like element was decreased in nuclear protein fractions from hypoxic cells. Comparative studies with an AP-1 consensus probe evaluated the effect of hypoxia on the binding of proteins to the AP-1 consensus binding element (Figure 4.11 B). Similar to the results observed for the 2J2-(-56/-63) probe, the intensity of the shifted complex observed with the AP-1 consensus probe and nuclear protein fractions from normoxic cells (lane 1) was diminished by an anti-c-Jun antibody (lane 2), but not by anti-c-Fos or anti-ubiquitin antibodies (lanes 3 and 4, respectively). However, in contrast to the results observed with the 2J2-(-56/-63) probe, the intensity of the shifted complex produced by the AP-1 consensus probe was more pronounced in nuclear extracts from hypoxic cells (lane 5), and both c-Jun and c-Fos antibodies (lanes 6 and 7), but not the ubiquitin antibody (lane 8) supershifted the complex. These results indicate that the protein complex bound to the AP-1 consensus element under normal oxygen conditions contains c-Jun, while both c-Jun and c-Fos are present in the complex that binds to the AP-1 consensus under low oxygen conditions. Thus, the -56 to -63 bp AP-1-like element in the CYP2J2 promoter exhibits a different pattern of binding in hypoxic conditions compared to the consensus AP-1 binding element.

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A Competitor - - self stat ------Antibody - - - - Fos Jun Ub - Fos Jun Protein - N N N N N N H H H

shift

1 2 3 4 5 6 7 8 9 10

Probe 2J2-(-56/-63)

B Antibody - Jun Fos Ub - Jun Fos Ub Protein N N N N H H H H supershift

shift

1 2 3 4 5 6 7 8

Probe AP-1 consensus

Figure 4.11 Differential effect of hypoxia on the binding of c-Jun to the -56/-63 bp AP-1-like element within the CYP2J2 promoter, and the AP-1 consensus binding sequence A. Hypoxia diminishes the binding of c-Jun to the -56/-63 bp element within the CYP2J2 promoter. B. In hypoxia, the binding of c-Jun and c-Fos to the AP-1 consensus binding sequence is increased. 32P-labelled double-stranded probes incorporating (A) the -56/-63 bp AP-1-like element within the upstream region of CYP2J2 [(2J2-(-56/-63)], and (B) the AP-1 consensus binding motif (AP-1 consensus) were incubated with nuclear protein fractions prepared from HepG2 cells cultured for 16 hr in normoxia (21% O2; N) or hypoxia (1% O2; H) as described in Methods. For competition experiments, 200-fold excess of unlabelled 2J2-(-56/-63) probe (self), or stat probe (containing the STAT5 element from the ȕ-casein promoter) was included in the binding reaction; for experiments with antibodies, nuclear protein was pre- incubated for 1 hr with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis and visualised by autoradiography. Shifted and supershifted complexes are indicated by arrows.

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4.6 Discussion

Results presented in Chapter 3 demonstrated that the expression of CYP2J2 is down- regulated in HepG2 cells cultured under hypoxic conditions. A number of potential AP-1 binding motifs were identified within the 5’-upstream regulatory region of the CYP2J2 gene, and the down-regulation of CYP2J2 in hypoxia was associated with the up-regulation of AP-1 proteins, including c-Fos. These preliminary studies prompted a molecular analysis of the involvement of AP-1 proteins in the differential regulation of CYP2J2 expression under conditions of normal and decreased oxygen tension. Results presented in this chapter demonstrate that c-Jun is important in the transcriptional regulation of CYP2J2 and indicate that c-Fos is involved in the down-regulation of CYP2J2 in hypoxia. To investigate the molecular regulation of the CYP2J2 gene, the 5’-flanking region was cloned and its transactivation by AP-1 gene products was studied by transient transfection analysis. As outlined in Chapter 1 (section 1.9.1), the AP-1 complex exists as homodimers between Jun proteins or heterodimers between members of the Fos and Jun families (Angel & Karin, 1991). Importantly, AP-1 complexes composed of different dimeric combinations of Jun and Fos proteins may exhibit distinct DNA binding affinities and different transcriptional activities, which gives rise to a high degree of flexibility in its gene regulatory potential (Angel & Karin, 1991; Ryseck & Bravo, 1991; Kerppola & Curran, 1993; Shaulian & Karin, 2001; van Dam & Castellazzi, 2001). Transient transfection analysis indicated divergent regulatory effects of different AP-1 dimers on CYP2J2 gene transcription. The CYP2J2 promoter was found to be strongly activated by c-Jun, whereas c-Fos did not stimulate CYP2J2 promoter activity and abolished the activation of the promoter elicited by c-Jun alone. Thus, the transcription of the CYP2J2 gene is influenced by the composition of AP-1 complexes present within the cell, which is determined by the differential expression of specific family members (Angel & Karin, 1991; Karin, 1995; Shaulian & Karin, 2001). It is well established that c-Jun is expressed constitutively in resting cells and that c-Jun homodimers constitute the pre-existing AP-1 complexes present within unstimulated cells (Lafyatis et al., 1990; De Cesare et al., 1995; Karin et al., 1997; Mishra et al., 1998; Mechta-Grigoriou et al., 2001; Minet et al., 2001). In contrast to c-Jun, c-Fos is either absent or expressed at very low levels in resting cells, but is up-regulated in response to a range of exogenous stimuli. Accordingly, rapid changes in the composition of AP-1 complexes from c-Jun homodimers to c-Jun/c-Fos heterodimers occurs in stimulated cells (Karin, 1995; Karin et al., 1997; Mishra et al.,

144 Chapter 4

1998; Minet et al., 2001). Consistent with these reports, the findings in Chapter 3 demonstrated the basal expression of c-Jun in HepG2 cells cultured under normal oxygen conditions. On the other hand, c-Fos was not expressed in normoxic HepG2 cells, but was strongly induced in response to hypoxia (Figures 3.6 to 3.7). Thus, exposure of HepG2 cells to hypoxia results in a shift in the composition of AP-1 complexes from c-Jun homodimers to c-Fos/c-Jun heterodimers. Taken together, these results suggest that c-Jun is involved in the maintenance of CYP2J2 expression in normoxic cells, and that down-regulation of CYP2J2 in hypoxia is mediated by c-Fos up-regulation and heterodimerisation with c-Jun. Transfection studies indicated that c-Jun-dependent induction of CYP2J2 promoter activity was also suppressed by the Fos-related proteins Fra-1 and Fra-2. It is feasible that Fra-1 and Fra-2, which have been shown to be expressed in liver and other cells (Yoshioka et al., 1995; Wild et al., 1998; Shih et al., 2000), may contribute with c-Fos to the suppression of CYP2J2 in hypoxia. Although c-Fos usually potentiates the activity of AP-1-regulated promoters (Sassone-Corsi et al., 1988; Hirai et al., 1989), there have been several reports of antagonism of c-Jun-dependent gene activation. For example, Kovacic-Milivojevic & Gardner (1992) described the activation of the human atrial natriuretic peptide (ANP) promoter by c-Jun and its inhibition by over-expression of c-Fos. Similarly, De Cesare et al. (1995) reported that c-Fos inhibits c-Jun-dependent activation of the human urokinase gene promoter. Thus, it appears that CYP2J2 is another human gene whose expression is positively regulated by c-Jun and negatively regulated by c-Fos. Transient transfection studies using CYP2J2 promoter deletion constructs identified the 152 bp sequence immediately upstream of the start of the CYP2J2 coding region as important for transactivation by c-Jun and repression by c-Fos; EMSAs confirmed the binding of c-Jun within this region. Sequence analysis identified two AP-1-like sequences within this 152 bp c-Jun-responsive region of CYP2J2: one on the positive strand located at -7 to +1 bp relative to the translation start site, and the other on the negative strand located at -56 to -63 bp. EMSAs established the capacity of both sites to bind c-Jun protein, albeit with different affinities, with the -56 to -63 bp element exhibiting stronger interactions with both nuclear protein fractions and recombinant c-Jun protein than those observed with the -7 to +1 bp element. Consistent with its apparently weaker ability to bind to c-Jun, site-directed mutagenesis of the -7 to +1 bp AP-1-like element did not affect basal or c-Jun-dependent activity of the CYP2J2 promoter in transient transfections. Thus, this element does not play a functional role in c-Jun-mediated activation of CYP2J2. On the other hand, the -56 to -63 bp element has an important functional role in c-Jun-dependent activation of

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the CYP2J2 promoter. Mutagenesis of this element decreased the activation of CYP2J2 by c-Jun in transient transfections. Moreover, mutagenesis at -56/-63 bp also reduced the basal transcriptional activity of the CYP2J2 promoter in HepG2 cells. This finding strongly suggests that the -56 to -63 bp element has a role in the basal expression of CYP2J2 in normoxia. EMSAs conducted using nuclear protein fractions extracted from normoxic and hypoxic HepG2 cells indicated that binding to the -56/-63 bp c-Jun-response element within the CYP2J2 promoter is altered in hypoxia. c-Jun, but not c-Fos, was identified in the protein complex bound to the element in normoxic HepG2 cells. This is consistent with the expression of c-Jun, but not c-Fos, in normoxic HepG2 cells (De Cesare et al., 1995). c-Fos was up-regulated in hypoxic HepG2 cells, which facilitated the formation of c-Fos/c-Jun heterodimers and increased the binding of proteins to an AP-1 consensus sequence in EMSAs. By contrast, c-Fos/c-Jun heterodimers diminished the binding of c-Jun to the -56/-63 bp AP-1-like element in the CYP2J2 promoter. It is noteworthy that complexes composed of different dimeric combinations of AP-1 proteins have been shown to bind differently to AP-1 elements within gene promoters (Ryseck & Bravo, 1991; Kerppola & Curran, 1993) and to induce different patterns of DNA bending at and around AP-1 sites (Kerppola & Curran, 1993). Importantly, AP-1 sites are often part of complex regulatory elements that contain binding sites for multiple different transcription factors, and it has been suggested that constraints on the DNA curvature at a particular AP-1 site, determined by other proteins binding to overlapping or adjacent sites, may determine the particular AP-1 dimers that can bind to a given gene element (Kerppola & Curran, 1993). Considerations of this type may account for the apparent ability of c-Jun, but not c-Fos/c-Jun, to interact with the -56/-63 bp AP-1-like element within the CYP2J2 promoter and contribute to the differential regulation of CYP2J2. Indeed, a similar mechanism apparently mediates the differential regulation of the human urokinase gene promoter by c-Jun and c-Fos. De Cesare et al. (1995) found that urokinase promoter activity is positively regulated by c-Jun and ATF-2, and inhibited by c-Fos. Thus, c-Jun/ATF-2 heterodimers activated transcription by binding to an AP-1-like element in the upstream enhancer region of the gene, whereas c-Fos/c-Jun heterodimers interacted poorly with this element. It was proposed that the inhibitory effect of c-Fos on c-Jun-dependent urokinase promoter activity arises from sequestration of c-Jun from the active pool of c-Jun/ATF-2 heterodimers to produce transcriptionally ineffective c-Fos/c-Jun heterodimers (De Cesare et al., 1995). In summary, the findings in this chapter have established an important role for the AP-1 proteins c-Jun and c-Fos in the regulation of CYP2J2 expression in normoxia

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and hypoxia, and have provided a molecular basis for the association of hypoxic up-regulation of c-Fos and down-regulation of CYP2J2 (from Chapter 3). c-Jun, which is expressed endogenously within HepG2 and other cells, was found to be a strong inducer of CYP2J2 promoter activity. In contrast, c-Fos, which is absent in resting cells but rapidly up-regulated in response to stress stimuli such as hypoxia, suppressed c-Jun-dependent activation of CYP2J2. Thus, it emerges that c-Jun homodimers stimulate CYP2J2 promoter activity and support basal expression of CYP2J2 in normoxia. In hypoxia, the formation of c-Fos/c-Jun heterodimers does not support CYP2J2 transcription. Regulation by AP-1 was found to be partially dependent on the -56/-63 bp element within the CYP2J2 proximal promoter. This element interacted strongly with c-Jun in normoxic cells, but displayed diminished binding to c-Jun following the up-regulation of c-Fos in hypoxia. While mutagenesis of the -56/-63 bp AP-1-like element significantly decreased activation of the CYP2J2 promoter by c-Jun, induction was not abolished completely. In contrast, deletion of the upstream region between -152 and -50 bp completely eliminated c-Jun-responsiveness, which implicates another element within this region that is involved in regulation of CYP2J2 by c-Jun. Chapter 5 describes the identification and characterisation of the additional element that participates with the -56/-63 bp site in c-Jun-dependent regulation of the CYP2J2 promoter.

147 Chapter 5

Chapter 5

Identification and Characterisation of a Second c-Jun Binding Element within the 5’-upstream Region of CYP2J2 that Regulates Expression

5.1 Introduction

Results presented in the previous two chapters have indicated that the AP-1 transcription factor is involved in the regulation of hepatic CYP2J2 expression in normoxia and hypoxia. Down-regulation of CYP2J2 in hypoxic HepG2 cells is associated with up-regulation of the AP-1 protein c-Fos. In transient transfection studies, the CYP2J2 promoter was strongly activated by c-Jun; this effect was abolished by c-Fos. Thus, the constitutively expressed c-Jun maintains CYP2J2 expression in resting HepG2 cells. In hypoxia, the up-regulation of c-Fos and formation of c-Fos/c-Jun heterodimers does not support CYP2J2 transcription. Using a series of CYP2J2 deletion-reporter constructs, a c-Jun-responsive region was identified between -152 and -50 bp relative to the translation start site. An AP-1-like element at -56 to -63 bp was important in mediating c-Jun-dependent transactivation of CYP2J2. The finding that mutagenesis of the -56/-63 bp element only partially impaired c-Jun-dependent activation, whereas deletion of the -152 to -50 bp region completely abolished activation by c-Jun, implicated a second AP-1-responsive element in CYP2J2 regulation. However, a search of the -152 to -50 bp region of the CYP2J2 promoter did not reveal additional AP-1-like binding sequences. Thus, it was considered that an atypical or cryptic AP-1 site that does not resemble the AP-1 consensus binding sequence must mediate c-Jun-dependent transactivation. Results presented in this chapter describe

148 Chapter 5

the identification and characterisation of a second c-Jun-responsive element within the CYP2J2 proximal promoter.

5.2 Identification of c-Jun-responsive Regions Between -152 and -50 bp in the CYP2J2 Proximal Promoter

Deletion of the -152 to -50 bp sequence rendered the CYP2J2 promoter completely unresponsive to c-Jun [2J2 (-49/+98); Figure 5.1; P<0.0001]. Consistent with a role for c-Jun in maintaining expression of CYP2J2 in resting cells, the basal promoter activity of 2J2 (-49/+98) in HepG2 cells was only 1.6 ± 0.1-fold higher than that of the promoterless pGL3basic vector backbone, compared with 6.3 ± 0.5-fold for 2J2 (-152/+98) [Figure 5.2; P<0.0001].

-152 +1 +98 2J2 (-152/+98) -122 +1 +98 2J2 (-122/+98) * - c-Jun -82 +1 +98 2J2 (-82/+98) + c-Jun ** -49+1 +98 -56/-63 c-Jun site 2J2 (-49/+98) *** -1012345 Fold Difference

Figure 5.1 c-Jun-responsive regions between -152 and -50 bp in the CYP2J2 proximal promoter The CYP2J2 promoter deletion constructs 2J2 (-122/+98), 2J2 (-82/+98) and 2J2 (-49/+98) were prepared by 5’-truncation of 2J2 (-152/+98) as described in Methods. The location of the previously identified c-Jun binding site at -56/-63 bp is indicated. CYP2J2 reporter constructs were co-transfected (1 ȝg per well) into HepG2 cells with a ȕ-galactosidase expression plasmid (0.5 ȝg per well; to control for transfection efficiency). A c-Jun expression plasmid was co-transfected at 0.5 ȝg per well as indicated. Following transfection, cells were harvested, and assayed for luciferase and ȕ-galactosidase activity as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity. The normalised activity of reporter constructs in the absence of c-Jun was set to 1-fold, and the activity of constructs in the presence of c-Jun was expressed as fold differences relative to activity in the absence of c-Jun. Results shown are means ± S.E.M. for at least three independent experiments. *Significantly different from 2J2 (-152/+98) in the presence of c-Jun (P=0.0001), **significantly different from 2J2 (-122/+98) in the presence of c-Jun (P<0.0001), ***significantly different from 2J2 (-82/+98) in the presence of c-Jun (P<0.0001).

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pGL3basic

-152 +1 +98 2J2 (-152/+98)

-122 +1 +98 2J2 (-122/+98) * -82 +1 +98 2J2 (-82/+98) ** , # -49+1 +98 -56/-63 c-Jun site 2J2 (-49/+98) ** , +

0246810 Fold Difference

Figure 5.2 Effect of deletions within -152 to -50 bp on basal CYP2J2 promoter activity in HepG2 cells The CYP2J2 reporter constructs 2J2 (-152/+98), 2J2 (-122/+98), 2J2 (-82/+98) and 2J2 (-49/+98), or the empty pGL3basic vector, were co-transfected (1 ȝg per well) into HepG2 cells with a ȕ-galactosidase expression plasmid (0.5 ȝg per well; to control for transfection efficiency). Following transfection, cells were harvested, and assayed for luciferase and ȕ-galactosidase activity as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity and presented as fold of the normalised activity of the empty pGL3basic vector (set to 1-fold). Results shown are means ± S.E.M. for at least three independent experiments. *Significantly different from 2J2 (-152/+98) [P<0.05], **significantly different from 2J2 (-152/+98) [P<0.0001], #significantly different from 2J2 (-122/+98) [P<0.0001], +significantly different from 2J2 (-82/+98) [P<0.02].

To locate the additional c-Jun-responsive element(s) within the -152/-50 bp region, the deletion constructs 2J2 (-122/+98) and 2J2 (-82/+98) were prepared by 5’-truncation from 2J2 (-152/+98), and transfected into HepG2 cells. As shown in Figure 5.1, deletion of the region between -152 to -122 bp decreased c-Jun-dependent transactivation to 3.2 ± 0.2-fold of control, compared with 4.2 ± 0.1-fold for 2J2 (-152/+98) [P=0.0001], but did not diminish basal promoter activity (Figure 5.2). In fact, 2J2 (-122/+98) displayed a higher basal activity than 2J2 (-152/+98) [Figure 5.2; P<0.05]. In contrast, deletion of the promoter sequence between -122 to -82 bp caused a pronounced loss of both c-Jun-dependent activation (to 1.8 ± 0.2-fold of control; Figure 5.1; P<0.0001), and basal transcriptional activity [to 2.7 ± 0.2-fold that of the empty pGL3basic vector backbone, compared to 6.3 ± 0.5-fold for 2J2 (-152/+98); Figure 5.2; P<0.0001]. These results indicate that, in addition to the previously identified -56/-63 bp element, the CYP2J2 proximal promoter contains an important c-Jun-responsive element between -122 and -82 bp. From these studies, the -152 to -122 bp region may also contain a third element that participates in regulation by c-Jun.

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5.3 Identification of c-Jun Binding Sequences within the -152 to -50 bp Region of CYP2J2

Although transfection studies clearly indicated the presence of additional c-Jun-response elements within the -152 to -50 bp region of the CYP2J2 gene, other than the site at -56/-63 bp (Figure 5.1), these sites were not identified by high- and low- stringency automated screening. Thus, a series of EMSAs were conducted to evaluate the binding of c-Jun within the -152/-50 bp region. Double-stranded oligonucleotides corresponding to three large overlapping regions of the -152 to -50 bp sequence (2J2-A, spanning -152 to -103 bp; 2J2-B, spanning -127 to -79 bp; and 2J2-C, spanning -102 to -50 bp; Figure 5.3) were labelled with [32P]dCTP and used as probes in EMSAs with nuclear protein fractions prepared from c-Jun-transfected HepG2 cells, and with recombinant c-Jun protein. In order to identify specific sequences important for c-Jun binding, competition experiments were performed using unlabelled double- stranded oligonucleotides corresponding to smaller overlapping sequences within the -152 to -50 bp region (probes 1 to 9, Figure 5.3). The results of these EMSA studies are shown in Figures 5.4 to 5.6. Incubation of 2J2-A (-152/-103 bp) with nuclear protein fractions from c-Jun-transfected HepG2 cells generated a shifted complex (Figure 5.4 B, lane 1) that decreased in intensity with the inclusion of 200-fold excess of unlabelled self probe (lane 2), but not by excess of the unrelated probe containing the STAT5 element from the ȕ-casein promoter (stat; lane 3). The signal intensity of the shifted complex was also diminished by the inclusion of 200-fold excess of the unlabelled probes 5 (-105/-86 bp; lane 6), 4 (-114/-97 bp; lane 8), and, to a lesser extent, 1 (-152/-128 bp; lane 4), but not by probes 3 (-127/-106 bp; lane 5), or 2 (-137/-119 bp; lane 7). Probe 2J2-A (-152/-103 bp) elicited a significant binding reaction with recombinant c-Jun protein (Figure 5.4 C, lane 1), which was diminished by excess unlabelled self probe but not by excess of the unrelated STAT5 element probe (lanes 2 and 3). Competition experiments with the smaller unlabelled probes gave similar results to those observed with nuclear protein fractions. Thus, the interaction between 2J2-A (-152/-103 bp) and recombinant c-Jun protein was decreased by 200-fold excesses of probes 5 (-105/-86 bp; lane 6) and 4 (-114/-97 bp; lane 8), and also, to a lesser extent, probe 1 (-152/-128 bp; lane 4), whereas probes 3 (-127/-106 bp; lane 5) and 2 (-137/-119 bp; lane 7) did not compete.

151 Chapter 5 t P]dCTP. The double- 32 , were end-labelled with [ (-102/-50), which span three large overlapping regions (-102/-50), CYP2J2 proximal promoter CYP2J2 stranded oligonucleotides -152/-128 (probe 1), -137/-119 (probe 2), -127/-106 3), -114/-97 (probe 4), -105/-86 (probe 5), -94/-77 6), -85/-69 7), -77/-60 8) and -68/-48 9), which span smaller overlapping sequences within -152/-50 bp, were used as unlabelled competitors. The c-Jun binding site a The probes 2J2-A (-152/-103), 2J2-B (-127/-79) and 2J2-C 2J2-B The probes 2J2-A (-152/-103), within the -152/-50 bp c-Jun-responsive region of Figure 5.3 Probes and competitors used in EMSAs to identify c-Jun binding sequences within the -152 to -50 bp region of the -56 to -63 bp (identified in Chapter 4) is highlighted. The results of EMSAs with these probes are shown 5.6. Figures 5.4 to

152 Chapter 5

The probe 2J2-B, which spans -127 to -79 bp of the CYP2J2 promoter (Figure 5.5 A), formed a number of strong interactions with nuclear extracts from c-Jun-transfected HepG2 cells (Figure 5.5 B, lane 1), that were found to represent specific DNA-protein interactions because they were competed by excess unlabelled self probe (Figure 5.5 B, lane 2) but not by the unrelated STAT5 element probe (stat; lane 3). These specific DNA-protein interactions were competed by the inclusion of 200-fold excess of the unlabelled smaller overlapping probe 5 (-105/-86 bp; lane 5), and to a lesser extent, probe 4 (-114/-97 bp; lane 8). On the other hand, excess of the smaller overlapping probes 3 (-127/-106 bp; lane 4), 7 (-85/-69 bp; lane 6), 2 (-137/-119 bp; lane 7) or 6 (-94/-77 bp; lane 9) did not inhibit complex formation. Probe 2J2-B (-127/-79 bp) also interacted strongly with recombinant c-Jun protein (Figure 5.5 C, lane 1). The interaction was confirmed to be specific by competition of complex formation with 200-fold excess of unlabelled self probe (lane 2), but not by the unrelated STAT5 element probe (stat; lane 3). Following the same pattern as that seen with nuclear protein, the binding of 2J2-B (-127/-79 bp) to recombinant c-Jun protein was effectively inhibited by 200-fold excess of the unlabelled smaller probes 5 (-105/-86 bp; lane 5) and 4 (-114/-97; lane 8), but not by probes 3 (-127/-106 bp; lane 4), 7 (-85/-69 bp; lane 6), 2 (-137/-119 bp; lane 7) or 6 (-94/-77; lane 9). The probe 2J2-C (-102/-50 bp) generated retarded complexes upon incubation with nuclear protein fractions from c-Jun-transfected HepG2 cells (Figure 5.6 B, lane 1). These complexes were effectively competed by 200-fold excess of unlabelled self probe (lane 2), but not by similar excess of the unrelated STAT5 element probe (stat; lane 3), thus indicating specificity. The inclusion of excess of the unlabelled smaller overlapping probes 5 (-105/-86 bp; lane 4), 9 (-68/-48 bp; lane 6), 4 (-114/-97 bp; lane 7) and 8 (-77/-60 bp; lane 9) diminished complex formation to varying extents, while the inclusion of excess of the unlabelled probes 7 (-85/-69 bp; lane 5) or 6 (-94/-77 bp; lane 8) did not markedly affect complex formation. Incubation of 2J2-C (-102/-50 bp) with recombinant c-Jun protein produced a strong specific binding interaction (Figure 5.6 C, lane 1), as demonstrated by competition with 200-fold excess of unlabelled self probe (lane 2) but not the unrelated STAT5 element probe (stat; lane 3). Consistent with the results of EMSA experiments with nuclear protein, the binding of probe 2J2-C (-102/-50 bp) to recombinant c-Jun protein was decreased by 200-fold excess of the unlabelled smaller overlapping probes 5 (-105/-86 bp; lane 4), 9 (-68/-48 bp; lane 6), 4 (-114/-97 bp; lane 7) and, to a lesser extent, probe 8 (-77/-60 bp; lane 9), but not by excess of probes 7 (-85/-69 bp; lane 5) or 6 (-94/-77 bp; lane 8). The combination of the two most effective competitors (probes 5 and 9) proved

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to be particularly efficient in inhibiting the binding of probe 2J2-C (-102/-50 bp) to HepG2 nuclear extract and recombinant c-Jun protein (Figures 5.6 B and C, lane 10). Taken together, the results of EMSAs shown in Figures 5.4 to 5.6 indicate that, within the -152 to -50 bp c-Jun-responsive region of the CYP2J2 gene, the sequences between -114 to -86 bp and -77 to -48 bp, and possibly also the -152 to -128 bp sequence, are capable of interacting with c-Jun. Binding within the -77 to -48 bp region was attributed to the c-Jun/AP-1-like element at -56 to -63 bp characterised in Chapter 4. The -152 to -128 bp and -114 to -86 bp sequences lie, respectively, within the -152 to -122 bp and -122 to -82 bp regions that were found in transfection studies to have potential roles in c-Jun-dependent activation (Figure 5.1). Their possible involvement in CYP2J2 regulation by c-Jun was further evaluated by EMSAs and transfection experiments.

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-105/-86 (5) -114/-97 (4) -127/-106 (3) -137/-119 (2) -152/-128 (1) A -152 CCCGGGAATCCAGCGCCTGGCATCTTCGCAGGGTGCTGCGAAGGGGCGGG -103 2J2-A

B Competitor - self stat 1 3 5 2 4 Protein JT JT JT JT JT JT JT JT

shift

1 2 3 4 5 6 7 8

Probe 2J2-A (-152/-103)

C Competitor - self stat 1 3 5 2 4 Protein Jun Jun Jun Jun Jun Jun Jun Jun

shift

1 2 3 4 5 6 7 8

Probe 2J2-A (-152/-103)

Figure 5.4 EMSA analysis of the binding of HepG2 nuclear extract and recombinant c-Jun protein to the -152 to -103 bp CYP2J2 promoter region A. Sequences of probes used in EMSAs to evaluate protein binding within the -152 to -103 bp region of the CYP2J2 promoter. 2J2-A corresponds to -152 to -103 bp of the CYP2J2 promoter and was labelled with [32P]dCTP. Probes corresponding to smaller overlapping sequences of CYP2J2 [-152/-128 (probe 1), -137/-119 (probe 2), -127/-106 (probe 3), -114/-97 (probe 4), and -105/-86 (probe 5)] were used in EMSAs as unlabelled competitors. The labelled 2J2-A probe was incubated with nuclear protein fractions from c-Jun-transfected HepG2 cells (JT; B), and with recombinant c-Jun protein (Jun; C) as described in Methods. 200-fold excess of unlabelled 2J2-A probe (self), stat probe (STAT5 element from the ȕ-casein promoter), or the smaller overlapping probes 1-5 were included in the binding reactions as indicated. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Shifted complexes are indicated by arrows.

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-85/-69 (7) -94/-77 (6) -105/-86 (5) -114/-97 (4) -127/-106 (3) -137/-119 (2) A -127 TCGCAGGGTGCTGCGAAGGGGCGGGCTGGGAGGCGGGGCACGGCTGGGA -79 2J2-B

B Competitor - self stat 3 5 7 2 4 6 Protein JT JT JT JT JT JT JT JT JT

shift shift

shift

shift

1 2 3 4 5 6 7 8 9

Probe 2J2-B (-127/-79)

C Competitor - self stat 3 5 7 2 4 6 Protein Jun Jun Jun Jun Jun Jun Jun Jun Jun

shift

1 2 3 4 5 6 7 8 9

Probe 2J2-B (-127/-79)

Figure 5.5 EMSA analysis of the binding of HepG2 nuclear extract and recombinant c-Jun protein to the -127 to -79 bp CYP2J2 promoter region A. Sequences of probes used in EMSAs to evaluate protein binding within the -127 to -79 bp region of the CYP2J2 promoter. 2J2-B corresponds to the -127 to -79 bp sequence of CYP2J2 and was labelled with [32P]dCTP. The probes spanning smaller overlapping sequences of CYP2J2 [-137/-119 (probe 2), -127/-106 (probe 3), -114/-97 (probe 4), -105/-86 (probe 5), -94/-77 (probe 6) and -85/-69 (probe 7)] were included in EMSAs as unlabelled competitors. The labelled 2J2-B probe was incubated with nuclear extracts from c-Jun-transfected HepG2 cells (JT; B), and with recombinant c-Jun protein (Jun; C) as outlined in Methods. 200-fold excess of unlabelled 2J2-B probe (self), stat probe (STAT5 element from the ȕ-casein promoter), or the smaller overlapping probes 2-7 were included in the binding reactions as indicated. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows. 156 Chapter 5

-68/-48 (9) -77/-60 (8) -85/-69 (7) -94/-77 (6) -105/-86 (5) -114/-97 (4) A -102 CTGGGAGGCGGGGCACGGCTGGGAGCGAGGCGGGGCGGGGACCGTCGCCTGCT -50

2J2-C

B Competitor - self stat 5 7 9 4 6 8 5+9 Protein JT JT JT JT JT JT JT JT JT JT

shift shift

shift shift

1 2 3 4 5 6 7 8 9 10 Probe 2J2-C (-102/-50)

C Competitor - self stat 5 7 9 4 6 8 5+9 Protein Jun Jun Jun Jun Jun Jun Jun Jun Jun Jun

shift

1 2 3 4 5 6 7 8 9 10

Probe 2J2-C (-102/-50)

Figure 5.6 EMSA analysis of the binding of HepG2 nuclear extract and recombinant c-Jun protein to the -102 to -50 bp CYP2J2 promoter region A. Sequences of probes used in EMSAs to evaluate protein binding within the -102 to -50 bp region of the CYP2J2 promoter. 2J2-C corresponds to the -102 to -50 bp sequence of CYP2J2 and was labelled with [32P]dCTP. The c-Jun binding element at -56 to -63 bp is underlined. Probes spanning smaller overlapping sequences of CYP2J2 [-114/-97 (probe 4), -105/-86 (probe 5), -94/-77 (probe 6), -85/-69 (probe 7), -77/-60 (probe 8) and -68/-48 (probe 9)] were included in EMSAs as unlabelled competitors. The labelled 2J2-C probe was incubated with nuclear extract from c-Jun-transfected HepG2 cells (JT; B), and with recombinant c-Jun protein (Jun; C) as described in Methods. 200-fold excess of unlabelled 2J2-C probe (self), stat probe (STAT5 element from the ȕ-casein promoter), or the smaller overlapping probes 4-9 were included in the binding reactions as indicated. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows. 157 Chapter 5

5.4 The -105 to -95 bp Sequence in CYP2J2 Binds c-Jun and Contributes to Transactivation of CYP2J2

From Figure 5.1, deletion of the CYP2J2 gene between -122 and -82 bp significantly decreased c-Jun-dependent activation of CYP2J2, indicating the presence of an important c-Jun-response element within this region. These results were supported by EMSAs that showed the specific interaction of nuclear extract and recombinant c-Jun protein with probe 2J2-B (-127/-79 bp), which incorporates the -122/-82 bp sequence (Figure 5.5). To further investigate the binding of c-Jun to the -122/-82 bp region of CYP2J2, probe 5 (-105/-86 bp) was labelled and used directly in EMSA experiments. This probe, which spans the -105 to -86 bp region of the CYP2J2 gene (Figure 5.7 A), was chosen because it was an extremely efficient competitor of the binding of c-Jun to probe 2J2-B (-127/-79 bp; Figure 5.5). As shown in Figure 5.7, a pronounced shifted complex was observed upon incubation of labelled probe 5 (-105/-86 bp) with both HepG2 nuclear protein and recombinant c-Jun protein (lanes 1 in Figures 5.7 B and C, respectively). Complex formation was competed by 200-fold excess of unlabelled self probe, but not by similar excess of the unrelated STAT5 probe (Figures 5.7 B and C, lanes 2 and 3), indicating that c-Jun binds in a specific manner to the -105 to -86 bp region of CYP2J2. The binding of nuclear protein and recombinant c-Jun protein to the -105 to -86 bp CYP2J2 promoter sequence was inhibited by the inclusion of 200-fold excess of the overlapping probe 4, which spans -114 to -97 bp, but not by similar excess of the overlapping probe 6, which spans -94 to -77 bp (Figures 5.7 B and C, lanes 4 and 5 respectively). Thus, within the -105 to -86 bp region of CYP2J2, the sequence at -105 to -95 bp is critical for c-Jun binding. The functional role of the -105 to -95 bp sequence in mediating c-Jun-dependent transactivation of the CYP2J2 promoter was tested in transfection analysis. As shown in Figure 5.8, the wild type promoter construct 2J2 (-152/+98) was activated strongly by c-Jun (to 3.7 ± 0.2-fold of control). Mutagenesis of the previously identified c-Jun binding site at -56 to -63 bp decreased c-Jun-responsiveness to 3.0 ± 0.5-fold of control (P<0.02); mutagenesis of the -105 to -95 bp sequence also significantly decreased c-Jun-dependent activation of reporter activity (to 1.5 ± 0.2-fold of control; Figure 5.8; P<0.0001), demonstrating the functional role of this site. Thus, both the -56/-63 bp and -105/-95 bp elements mediate transactivation of CYP2J2 by c-Jun.

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-105 -95 ********** A GGGCTGGGAGGCGGGGCACG -105/-86 (probe 5)

CGAAGGGGCGGGCTGGGAGGCGGGGCACGGCTGGGAGC -114/-97 (probe 4) -94/-77 (probe 6)

B Competitor - self stat 4 6 C - self stat 4 6 Protein UT UT UT UT UT Jun Jun Jun Jun Jun

shift shift

1 2 3 4 5 1 2 3 4 5 Probe -105/-86 (probe 5) -105/-86 (probe 5)

Figure 5.7 EMSAs reveal the -105 to -95 bp sequence of CYP2J2 is essential for c-Jun binding A. Sequences of probes used in EMSAs to evaluate the binding of c-Jun within the -114 to -86 bp region of the CYP2J2 promoter. Probes 4, 5 and 6 span -114 to -97 bp, -105 to -86 bp and -94 to -77 bp, respectively, of CYP2J2. Probe 5 (-105/-86 bp) was end-labelled with [32P]dCTP and incubated with nuclear protein fractions from untransfected HepG2 cells (UT; B), and with recombinant c-Jun protein (Jun; C) as outlined in Methods. 200-fold excess of unlabelled probe 5 (self), stat probe (STAT5 element from the ȕ-casein promoter), or the overlapping probes 4 (-114/-97 bp) or 6 (-94/-77 bp) were included in the binding reactions as indicated. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows. Nucleotides required for interaction with c-Jun (-105 to -95 bp) are highlighted in bold and marked by asterisks (A).

To evaluate the role of c-Fos at the two c-Jun-response elements, co-transfection studies with c-Jun and c-Fos were performed. Results presented in Chapter 4 indicated that c-Jun-dependent activation of the CYP2J2 proximal promoter is suppressed by c-Fos [the activity of the wild type proximal promoter construct 2J2 (-152/+98) was decreased from 4.1 ± 0.3-fold of control with c-Jun to 1.1 ± 0.1-fold of control with c-Fos/c-Jun; Figure 4.3]. The residual c-Jun-dependent activity observed with the two mutated constructs 2J2 (-152/+98; mt -56/-63) and 2J2 (-152/+98; mt -106/-96) was also suppressed by co-transfection of c-Fos. Thus, c-Fos decreased c-Jun-dependent activation of 2J2 (-152/+98; mt -56/-63) from 3.0 ± 0.5-fold of control to -0.1 ± 0.03-fold of control. Similarly, the activity of the construct 2J2 (-152/+98;

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mt -106/-96) was decreased from 1.5 ± 0.2-fold of control with c-Jun to -1.4 ± 0.2-fold with c-Fos/c-Jun. These results indicated that separate mutagenesis of either c-Jun-response element did not completely eliminate c-Jun-dependent activation, or c-Fos-mediated suppression, of the CYP2J2 promoter. Instead, mutagenesis of both elements rendered the promoter unresponsive to AP-1 [construct 2J2 (-152/+98; mt -56/-63, mt -106/-96); Figure 5.8]. Thus, both the -56/-63 bp and -105/-95 bp elements contribute to c-Jun-dependent activation and c-Fos-dependent inhibition of the CYP2J2 promoter.

-152 -105/-95 -56/-63 +1 +98 2J2 (-152/+98)

-152 -105/-95 -56/-63 +1 +98 2J2 (-152/+98; mt -56/-63) mutated * - c-Jun -152 -105/-95 -56/-63 +1 +98 + c-Jun 2J2 (-152/+98; mt -106/-96) mutated ** -152 -105/-95 -56/-63 +1 +98 2J2 (-152/+98; mt -56/-63, , # , + mutated mutated mt -106/-96) ** 01234 -56/-63 c-Jun binding site Fold Difference -56/-63 c-Jun binding site, mutated -105/-95 c-Jun binding site -105/-95 c-Jun binding site, mutated

Figure 5.8 The -56/-63 and -105/-95 bp sequences mediate c-Jun-dependent activation of the CYP2J2 proximal promoter The CYP2J2 promoter constructs 2J2 (-152/+98; mt -56/-63) and 2J2 (-152/+98; mt -106/-96) contain mutations, respectively, in the -56 to -63 bp and -105 to -95 bp c-Jun binding sequences. The construct 2J2 (-152/+98; mt -56/-63, mt -106/-96) contains mutations in both the -56 to -63 bp and -105 to -95 bp sequences. These constructs were prepared from the wild type construct 2J2 (-152/+98) by site-directed mutagenesis as outlined in Methods. HepG2 cells were co-transfected with CYP2J2 reporter constructs (1 ȝg per well) and a c-Jun expression plasmid (0.5 ȝg per well) as indicated. A ȕ-galactosidase expression plasmid (0.5 ȝg per well) was co-transfected to control for transfection efficiency. Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity. The normalised activity of reporter constructs in the absence of c-Jun was set to 1-fold, and the activity of constructs in the presence of c-Jun was expressed as fold differences relative to activity in the absence of c-Jun. Results shown are means ± S.E.M. for three independent experiments. *Significantly different from the wild type construct 2J2 (-152/+98) in the presence of c-Jun (P<0.02), **significantly different from the wild type construct 2J2 (-152/+98) in the presence of c-Jun (P<0.0001), #significantly different from the mutated construct 2J2 (-152/+98; mt -56/-63) in the presence of c-Jun (P<0.0001), +significantly different from the mutated construct 2J2 (-152/+98; mt -106/-96) in the presence of c-Jun (P<0.005).

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Consistent with a role for c-Jun in maintenance of the basal expression of CYP2J2 in resting cells, separate mutagenesis of either the -56/-63 bp or -105/-95 bp c-Jun binding elements significantly decreased the basal activity of the CYP2J2 promoter in HepG2 cells (Figure 5.9). Mutagenesis of both elements produced the most pronounced effect, reducing the basal activity of the CYP2J2 promoter to only 2.9 ± 0.1-fold that of the empty pGL3basic backbone, compared with 6.6 ± 0.4-fold for the wild type construct 2J2 (-152/+98) [Figure 5.9; P<0.0005]. Thus, the -56/-63 bp and -105/-95 bp c-Jun-response elements are essential for maintenance of the basal transcriptional activity of the CYP2J2 gene.

pGL3basic -152 -105/-95 -56/-63 +1 +98 2J2 (-152/+98)

-152 -105/-95 -56/-63 mt +1 +98 2J2 (-152/+98; mt -56/-63) * -152 -105/-95 mt -56/-63 +1 +98 2J2 (-152/+98; mt -106/-96) ** -152 -105/-95 mt -56/-63 mt +1 +98 2J2 (-152/+98; mt -56/-63, , # , + mt -106/-96) **

02468 -56/-63 c-Jun binding site Fold Difference -56/-63 c-Jun binding site, mutated -105/-95 c-Jun binding site -105/-95 c-Jun binding site, mutated

Figure 5.9 The -56/-63 and -105/-95 bp sequences are important in maintaining basal activity of the CYP2J2 proximal promoter in HepG2 cells The wild type CYP2J2 promoter construct 2J2 (-152/+98), the mutagenised constructs 2J2 (-152/+98; mt -56/-63), 2J2 (-152/+98; mt -106/-96) and 2J2 (-152/+98; mt -56/-63, mt -106/-96), or the empty pGL3basic vector, were co-transfected (1 ȝg per well) into HepG2 cells with a ȕ-galactosidase expression plasmid (0.5 ȝg per well; to control for transfection efficiency). Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity and presented as fold differences relative to the normalised activity of the empty pGL3basic vector (set to 1-fold). Results shown are means ± S.E.M. for at least three independent experiments. *Significantly different from 2J2 (-152/+98) [P<0.002], **significantly different from 2J2 (-152/+98) [P<0.0005], #significantly different from 2J2 (-152/+98; mt -56/-63) [P<0.005], +significantly different from 2J2 (-152/+98; mt -106/-96) [P<0.01].

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5.5 Characterisation of AP-1 Binding to the -105 to -95 bp c-Jun-response Element in the CYP2J2 Promoter

5.5.1 Binding of c-Jun to the -105 to -95 bp Element in the CYP2J2 Promoter EMSAs were conducted to further evaluate the binding of c-Jun to the -105 to -95 bp sequence. Probe 5 (-105/-86 bp; Figure 5.10 A), which incorporates the -105 to 95 bp sequence of CYP2J2, generated a shifted complex with nuclear protein fractions from untransfected HepG2 cells, as shown in Figure 5.10 B (lane 1). A more intense signal was observed with nuclear protein fractions from c-Jun-transfected HepG2 cells (lane 2). Complex formation was competed by 200-fold excess of unlabelled self probe (lane 3), but not by excess of the unrelated STAT5 element probe (stat; lane 4), thus confirming specificity. In antibody supershift experiments, the retarded complex was disrupted by an anti-c-Jun antibody (lane 6), but not by an anti-c-Fos antibody (lane 5), or an antibody directed against ubiquitin (lane 7). Thus c-Jun, but not c-Fos, was present in the protein complex bound to probe 5 (-105/-86 bp). The importance of the -105 to -95 bp element in binding c-Jun was evaluated directly in EMSAs using the probe mt 5, in which the -105 to -95 bp sequence was mutated (Figure 5.10 A). The binding interaction of probe mt 5 with nuclear extracts from untransfected and c-Jun-transfected HepG2 cells was greatly decreased compared to that with the wild type probe (Figure 5.10 C). Probe 5, containing the wild type -105/-95 bp sequence, also interacted strongly with recombinant c-Jun protein (Figure 5.10 D, lane 2), whereas the mutant sequence did not interact with c-Jun (lane 4).

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A GGGCTGGGAGGCGGGGCACG AAAAAAAAAAGCGGGGCACG -105/-86 (probe 5) -105/-86; mt -105/-96 (probe mt 5)

B Competitor - - self stat - - - Antibody - - - - Fos Jun Ub Protein UT JT UT UT UT UT UT

shift

1 2 3 4 5 6 7

Probe probe 5

C Protein UT JT UT JTD Protein - Jun - Jun

shift shift

1 2 3 4 1 2 3 4

Probe probe 5 probe Probe probe 5 probe mt 5 mt 5

Figure 5.10 EMSA of the binding of c-Jun to the -105/-95 bp sequence in the CYP2J2 promoter A. Sequences of probes used in EMSAs to evaluate the binding of c-Jun to the -105/-95 bp sequence within the CYP2J2 promoter. Probe 5 incorporates the wild type -105/-95 bp element (bolded); probe mt 5 is mutated at the -105/-95 bp element as indicated by asterisks. Probes were labelled with [32P]dCTP and incubated with nuclear protein fractions from untransfected (UT) and c-Jun-transfected (JT) HepG2 cells (B, C), and with recombinant c-Jun protein (Jun; D) as described in Methods. For competition experiments, 200-fold excess unlabelled probe 5 (self) or stat probe (containing the STAT5 element from the ȕ-casein promoter) was included in the binding reaction, and in antibody experiments, nuclear protein was pre-incubated with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) for 1 hr prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows.

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5.5.2 Effect of Hypoxia on Binding to the -105 to -95 bp c-Jun-response Element Results presented in the previous chapter demonstrated that exposure of HepG2 cells to hypoxia diminishes the binding of c-Jun to the -56 to -63 bp AP-1-like element in the CYP2J2 gene promoter (Figure 4.11). A similar approach was taken to assess the effect of hypoxia on the binding of protein to the -105 to -95 bp c-Jun-response element. As shown in Figure 5.11, incubation of probe 5 (harbouring the -105 to -95 bp sequence) with nuclear protein fractions from normoxic HepG2 cells generated a specific shifted complex (lane 1) that was effectively competed by 200-fold excess of unlabelled self probe (lane 2), but not by excess of the unrelated STAT5 element probe (lane 3). This DNA-protein complex was block-shifted by an anti-c-Jun antibody (lane 5), but was not affected by antibodies directed against c-Fos (lane 4) or ubiquitin (lane 6), indicating that c-Jun, but not c-Fos, binds to the -105 to -95 bp element in normoxic HepG2 cells. In contrast, a shifted complex was observed with nuclear protein fractions from hypoxic HepG2 cells (lane 7), that was disrupted by antibodies directed against both c-Jun (lane 9) and c-Fos (lane 8); the anti-ubiquitin antibody did not disrupt the complex (lane 10). These results indicate that exposure of HepG2 cells to hypoxia results in a change in the composition of proteins bound at the -105 to -95 bp element from c-Jun to c-Fos/c-Jun, and contrast those obtained for the -56 to -63 bp element (Chapter 4). The latter element interacted with c-Jun homodimers, but not c-Fos/c-Jun heterodimers, and therefore binding of c-Jun was decreased in hypoxic conditions in which c-Fos/c-Jun heterodimers predominated (Figure 4.11). Thus, hypoxia induces different effects at the two c-Jun-response elements within the CYP2J2 promoter. Whereas the binding of c-Jun to the -56 to -63 bp AP-1-like element was decreased, hypoxic exposure altered the proteins bound at the -105 to -95 bp element from c-Jun to both c-Jun and c-Fos.

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Competitor - self stat ------Antibody - - - Fos Jun Ub - Fos Jun Ub Protein N N N N N N H H H H

shift

1 2 3 4 5 6 7 8 9 10

Probe -105/-86 (probe 5)

Figure 5.11 EMSA evaluating the effect of hypoxia on the binding of protein to the -105/-95 bp c-Jun binding element [32P]dCTP-labelled probe 5, which incorporates the -105/-95 bp c-Jun binding sequence, was incubated with nuclear protein fractions prepared from HepG2 cells cultured for 16 hr in normoxia (21% O2; N) or hypoxia (1% O2; H) as described in Methods. Competition experiments were performed by the inclusion of 200-fold excess of unlabelled probe 5 (self) or stat probe (containing the STAT5 element from the ȕ-casein promoter) in the binding reaction, and antibody experiments were performed by pre-incubation of nuclear protein with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) for 1 hr prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Specific shifted complexes are indicated by arrows.

5.5.3 Binding of Recombinant c-Jun and c-Fos Proteins to the -105 to -95 bp Element in the CYP2J2 Promoter To confirm the capacity of the -105 to -95 bp CYP2J2 promoter element to directly bind both c-Jun homodimers and c-Fos/c-Jun heterodimers, EMSAs were conducted with probe 5 (-105/-86 bp) and recombinant human proteins (Figure 5.12). A shifted complex was observed with recombinant c-Jun protein (Figure 5.12 A, lane 1). Competition by 200-fold excess of unlabelled self probe (lane 5), but not the unrelated STAT5 element probe (stat; lane 6) demonstrated specificity. As anticipated, the complex was disrupted by an anti-c-Jun antibody (lane 2), but not by antibodies directed against c-Fos (lane 3) or ubiquitin (lane 4). No retarded complex was observed upon incubation of the labelled probe with recombinant c-Fos protein alone (Figure 5.12 B, lane 1), which is consistent with the requirement of Fos proteins to dimerise with Jun proteins for DNA binding (Halazonetis et al., 1988; Angel & Karin,

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1991). On the other hand, a shifted complex was observed with the c-Fos/c-Jun protein combination (Figure 5.12 B, lane 2), which was disrupted by antibodies directed against c-Jun and c-Fos, but not ubiquitin (Figure 5.12 B, lanes 3–5). These results confirm that the -105 to -95 bp element is able to interact with c-Fos/c-Jun heterodimers as well as c-Jun homodimers. The specific nature of the interaction with c-Fos/c-Jun heterodimers was indicated by competition with excess unlabelled self probe, but not with excess of the unrelated STAT5 element probe (Figure 5.12 B, lanes 6 and 7 respectively). Also shown in Figure 5.12 (lower panel C and D) are the results of control experiments conducted in parallel with a probe containing the consensus AP-1 binding element. As expected, c-Fos alone did not bind to the consensus probe (Figure 5.12 D, lane 1), but shifted complexes were observed with c-Jun alone (Figure 5.12 C, lane 1), and with the c-Fos/c-Jun combination (Figure 5.12 D, lane 2). Antibody supershift experiments (Figure 5.12 C, lanes 2-4 and Figure 5.12 D, lanes 3-5) and unlabelled competition experiments (Figure 5.12 C, lanes 5-6 and Figure 5.12 D, lanes 6-7) confirmed the specificity of the interactions between the AP-1 consensus sequence and c-Jun homodimers and c-Fos/c-Jun heterodimers.

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A B Competitor - - - - - self stat Competitor - - - - self stat Antibody - - Jun Fos Ub - - Antibody - Jun Fos Ub - - rec c-Jun - + + + + + + rec c-Jun + + + + + + rec c-Fos + + + + + + +

shift shift

1 2 3 4 5 6 1 2 3 4 5 6 7

Probe -105/-86 (probe 5) -105/-86 (probe 5)

C D Competitor - - - - - self stat Competitor - - - - self stat Antibody - - Jun Fos Ub - -

Antibody - Jun Fos Ub - - rec c-Jun - + + + + + + rec c-Jun + + + + + + rec c-Fos + + + + + + +

shift shift

1 2 3 4 5 6 1 2 3 4 5 6 7

Probe AP-1 consensus AP-1 consensus

Figure 5.12 EMSAs evaluating the binding of c-Jun and c-Fos/c-Jun to the CYP2J2 -105/-95 bp element and the AP-1 consensus binding sequence Double-stranded probes containing the CYP2J2 -105/-95 bp element (probe 5; A and B) and the AP-1 consensus binding sequence (C and D) were labelled with [32P]dCTP and incubated with recombinant (rec) c-Jun protein alone (A and C), or a combination of recombinant c-Jun and recombinant c-Fos protein (B and D) as described in Methods. In competition experiments, binding reactions included 200-fold excess unlabelled probe 5 (self; A and B), AP-1 consensus probe (self; C and D), or stat probe (STAT5 element from the ȕ-casein promoter). For reactions with antibodies, recombinant proteins were pre-incubated with antibodies directed against c-Jun (Jun), c-Fos (Fos) or ubiquitin (Ub) for 1 hr prior to the addition of labelled probe. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Shifted complexes are indicated by arrows.

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5.6 The -152 to -122 bp Region of the CYP2J2 Gene is not Involved in c-Jun-dependent Regulation of CYP2J2 Promoter Activity

From studies presented in this chapter so far, the role of the -152 to -122 bp region of the CYP2J2 gene in regulation of expression by c-Jun remained equivocal. Deletion of the promoter region of CYP2J2 between -152 and 122 bp decreased activation by c-Jun, suggesting that the -152 to -122 bp region may contain an element that participates in the regulation of CYP2J2 by c-Jun (Figure 5.1). On the other hand, deletion of this region did not impair basal transcriptional activity of the CYP2J2 promoter (Figure 5.2). Furthermore, preservation of the -152 to -122 bp region, but mutation of both the -56 to -63 bp and -105 to -95 bp c-Jun-response elements, rendered the CYP2J2 promoter completely unresponsive to c-Jun (Figure 5.8); this demonstrated the importance of the -56 to -63 bp and -105 to -95 bp elements in c-Jun-dependent activation, and argued against a role for the -152 to -122 bp region in c-Jun-responsiveness. In EMSA experiments, the probe 2J2-A (-152/-103 bp), which incorporates the -152 to -122 bp region, produced interactions with nuclear extract from c-Jun-transfected HepG2 cells (Figure 5.4 B), and recombinant c-Jun protein (Figure 5.4 C), that were partially diminished by excess of the unlabelled probe 1, which corresponds to the -152 to -128 bp sequence of CYP2J2. To pursue whether the -152 to -122 bp region contained a c-Jun-responsive sequence, a series of additional EMSAs and transfections were performed. A probe corresponding to the -152 to -122 bp region of the CYP2J2 gene was designed and labelled with [32P]dCTP for use in EMSAs (Figure 5.13 A). The overlapping probes 1 (-152/-128 bp), 2 (-137/-119 bp) and 3 (-127/-106 bp) used previously (section 5.3) were included in binding reactions as unlabelled competitors (Figure 5.13 A). Additional overlapping probes spanning -157 to -138 bp (probe 10) and -147 to -128 bp (probe 11) were designed and also tested for their ability to compete binding (Figure 5.13 A). Incubation of the -152/-122 bp probe with nuclear protein fractions isolated from c-Jun-transfected HepG2 cells produced several weak shifted complexes (Figure 5.13 B, lane 1) that were diminished by excess unlabelled self probe (lane 2), but not by excess of the unrelated STAT5 element probe (lane 3). Competition experiments in which excess of the unlabelled overlapping probes 1, 2, 3, 10 and 11 were incorporated into the binding reaction did not provide clear indications

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that any region within the -152 to -122 bp sequence is involved in protein interaction (lanes 4 to 8). EMSAs were also performed with the -152/-122 bp probe and recombinant c-Jun protein, and also proved to be inconclusive (Figure 5.13 C). A single shifted complex was observed upon incubation of the -152/-122 bp probe with recombinant c-Jun protein (lane 1). However, it was unclear from competition experiments whether this shifted complex was a specific or non-specific DNA-protein interaction. Inclusion of excess unlabelled self probe did not alter the intensity of the shifted complex, but appeared to affect its mobility (lane 2). A similar finding was noted with the inclusion of an excess of unlabelled probe 1 (-152/-128 bp; lane 4). The inclusion of excess of the unrelated STAT5 element probe (stat; lane 3), or excess of the unlabelled overlapping probes 3 (-127/-106 bp; lane 5), 2 (-137/-119 bp; lane 6), 10 (-157/-138 bp; lane 7) or 11 (-147/-128 bp; lane 8) did not affect the intensity or mobility of the shifted complex. Thus, the EMSA studies in Figure 5.13 failed to identify a specific region within the -152 to -122 bp sequence that mediates interactions with HepG2 nuclear protein or recombinant c-Jun protein. Transfection studies were more effective than EMSAs in evaluating the involvement of the -152 to -122 bp region in regulation by c-Jun. CYP2J2 reporter constructs containing consecutive 8 bp mutations within the -152 to -122 bp region were prepared from the wild type construct 2J2 (-152/+98) by site-directed mutagenesis. As shown in Figure 5.14, all three mutated constructs [2J2 (-152/+98; mt -148/-142), 2J2 (-152/+98; mt -141/-134) and 2J2 (-152/+98; mt -133/-126)] displayed similar c-Jun-responsiveness to that of the wild type construct 2J2 (-152/+98). Similarly, these mutations did not impair basal transcriptional activity of the CYP2J2 promoter [the constructs 2J2 (-152/+98; mt -148/-142), 2J2 (-152/+98; mt -141/-134) and 2J2 (-152/+98; mt -133/-126) displayed respective basal activities of 7.8 ± 0.2-fold, 6.5 ± 0.3-fold, and 8.4 ± 0.8-fold that of pGL3basic, compared with 5.9 ± 0.01-fold for the wild type construct 2J2 (-152/+98); Figure 5.15]. These findings are consistent with the lack of a significant role for the -152/-122 bp region in basal and c-Jun-mediated regulation of the gene.

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-127/-106 (3) -137/-119 (2) -147/-128 (11) -152/-128 (1) -157/-138 (10) A -152 CCCGGGAATCCAGCGCCTGGCATCTTCGCA -122

2J2-(-152/-122)

B Competitor - self stat 1 3 2 10 11 Protein JT JT JT JT JT JT JT JT

shift shift

shift

1 2 3 4 5 6 7 8

Probe 2J2-(-152/-122)

C Competitor - self stat 1 3 2 10 11 Protein Jun Jun Jun Jun Jun Jun Jun Jun

shift

1 2 3 4 5 6 7 8

Probe 2J2-(-152/-122)

Figure 5.13 EMSAs evaluating the binding of nuclear proteins and recombinant c-Jun protein within the -152/-122 bp region of the CYP2J2 promoter A. Sequences of probes used in EMSAs to evaluate protein binding within the -152 to -122 bp region of the CYP2J2 promoter. 2J2-(-152/-122) spans the -152 to -122 bp sequence of CYP2J2 and was labelled with [32P]dCTP. Probes spanning overlapping sequences [-157/-138 (10), -152/-128 (1), -147/-128 (11), -137/-119 (2) and -127/-106 (3)] were included in EMSA reactions as unlabelled competitors. Labelled 2J2-(-152/-122) was incubated with nuclear protein fractions from c-Jun-transfected HepG2 cells (JT; B), and with recombinant c-Jun protein (Jun; C) as described in Methods. 200-fold excess of unlabelled 2J2-(-152/-122) probe (self), stat probe (STAT5 element from the ȕ-casein promoter), or the overlapping probes 1, 2, 3, 10 and 11, were included in the binding reactions as indicated. DNA-protein complexes were resolved by polyacrylamide gel electrophoresis, and visualised by autoradiography. Shifted complexes are indicated by arrows.

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-152 -122 +1 +98 2J2 (-152/+98)

-148/-142 +1 +98 2J2 (-152/+98; mt -148/-142) - c-Jun -141/-134 +1 +98 + c-Jun 2J2 (-152/+98; mt -141/-134)

-133/-126 +1 +98 2J2 (-152/+98; mt -133/-126)

0123456 mutated sequence Fold Dif f erence -56/-63 c-Jun binding site -105/-95 c-Jun binding site

Figure 5.14 Mutation of the -152/-122 bp region does not affect c-Jun-dependent activation of the CYP2J2 promoter The CYP2J2 promoter constructs 2J2 (-152/+98; mt -148/-142), 2J2 (-152/+98; mt -141/-134) and 2J2 (-152/+98; mt -133/-126) contain mutations in the nucleotides spanning -148 to -142 bp, -141 to -134 bp, and -133 to -126 bp, respectively. These mutagenised constructs were prepared by site-directed mutagenesis of the wild type construct 2J2 (-152/+98) as outlined in Methods. The position of the -56/-63 bp and -105/-95 bp c-Jun binding elements are highlighted. HepG2 cells were co-transfected with CYP2J2 reporter constructs (1 ȝg per well) and a c-Jun expression plasmid (0.5 ȝg per well) as indicated. A ȕ-galactosidase expression plasmid (0.5 ȝg per well) was co-transfected to control for transfection efficiency. Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity. The normalised activity of reporter constructs in the absence of c-Jun was set to 1-fold, and the activity of constructs in the presence of c-Jun was expressed as fold differences relative to activity in the absence of c-Jun. Results shown are means ± S.E.M. for three independent experiments.

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pGL3basic

-152 -122 +1 +98 2J2 (-152/+98)

-148/-142 +1 +98 2J2 (-152/+98; mt -148/-142) * -141/-134 +1 +98 2J2 (-152/+98; mt -141/-134)

-133/-126 +1 +98 2J2 (-152/+98; mt -133/-126) **

0246810 mutated sequence Fold Difference -56/-63 c-Jun binding site -105/-95 c-Jun binding site

Figure 5.15 Mutation of the -152/-122 bp region does not diminish basal transcriptional activity of the CYP2J2 promoter in HepG2 cells The CYP2J2 promoter constructs 2J2 (-152/+98), 2J2 (-152/+98; mt -148/-142), 2J2 (-152/+98; mt -141/-134), 2J2 (-152/+98; mt -133/-126), or the empty pGL3basic vector, were co-transfected (1 ȝg per well) into HepG2 cells with a ȕ-galactosidase expression plasmid (0.5 ȝg per well; to control for transfection efficiency). Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity and presented as fold differences relative to the normalised activity of the empty pGL3basic vector (set to 1-fold). Results shown are means ± S.E.M. for three independent experiments. *Significantly different from the wild type construct 2J2 (-152/+98) [P<0.01], **significantly different from the wild type construct 2J2 (-152/+98) [P<0.005].

Finally, further reporter constructs, spanning -122 to +98 bp of the CYP2J2 gene, in which the -56 to -63 bp element, the -105 to -95 bp element, or both elements, were mutated, were also prepared and tested for their ability to respond to c-Jun (Figure 5.16). In particular, the activation of these constructs by c-Jun was compared to that of similarly mutated constructs in which the -152 to -122 bp sequence was intact. As shown in Figure 5.16, the activities of the mutated constructs which lacked the -152 to -122 bp region [i.e. 2J2 (-122/+98; mt -56/-63), 2J2 (-122/+98; mt -106/-96), and 2J2 (-122/+98; mt -56/-63, mt -106/-96)] were not different to the corresponding constructs containing the -152 to -122 bp sequence [i.e. 2J2 (-152/+98; mt -56/-63), 2J2 (-152/+98; mt -106/-96) and 2J2 (-152/+98; mt -56/-63, mt -106/-96)]. Thus, the apparently lower activity of 2J2 (-122/+98), relative to 2J2 (-152/+98) in initial experiments [Figure 5.1], was not supported by more detailed evaluations with mutagenised and truncated constructs.

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-105/-95 -56/-63 +1 +98 -152 2J2 (-152/+98) -122 2J2 (-122/+98) * -105/-95 -56/-63 mt +1 +98 -152 2J2 (-152/+98; mt -56/-63) -122 2J2 (-122/+98; mt -56/-63)

-105/-95 mt -56/-63 +1 +98 -152 2J2 (-152/+98; mt -106/-96) -122 2J2 (-122/+98; mt -106/-96)

-105/-95 mt -56/-63 mt +1 +98 -152 2J2 (-152/+98; mt -56/-63, mt -106/-96) - c-Jun + c-Jun -122 2J2 (-122/+98; mt -56/-63, mt -106/-96)

012345 -56/-63 c-Jun binding site Fold Difference -56/-63 c-Jun binding site, mutated -105/-95 c-Jun binding site -105/-95 c-Jun binding site, mutated

Figure 5.16 Transient transfection analysis of the -152/-122 bp sequence and c-Jun-dependent activation of the CYP2J2 proximal promoter The wild type CYP2J2 promoter constructs 2J2 (-152/+98) and 2J2 (-122/+98) span -152 to +98 bp and -122 to +98 bp of CYP2J2, respectively. The mutagenised constructs 2J2 (-152/+98; mt -56/-63), 2J2 (-152/+98; mt -106/-96) and 2J2 (-152/+98; mt -56/-63, mt -106/-96) span -152 to +98 bp of CYP2J2 and are mutated, respectively, at the -56 to -63 bp c-Jun binding element, the -105 to -95 bp c-Jun binding element, and at both elements. The mutagenised constructs 2J2 (-122/+98; mt -56/-63), 2J2 (-122/+98; mt -106/-96) and 2J2 (-122/+98; mt -56/-63, mt -106/-96) are similarly mutated, but are lacking the -152 to -122 bp region. HepG2 cells were co-transfected with CYP2J2 reporter constructs (1 ȝg per well) and a c-Jun expression plasmid (0.5 ȝg per well) as indicated. A ȕ-galactosidase expression plasmid (0.5 ȝg per well) was co-transfected to control for transfection efficiency. Following transfection, cells were harvested, and luciferase and ȕ-galactosidase assays were performed as outlined in Methods. Luciferase activity was normalised to ȕ-galactosidase activity. The normalised activity of reporter constructs in the absence of c-Jun was set to 1-fold, and the activity of constructs in the presence of c-Jun was expressed as fold differences relative to activity in the absence of c-Jun. Results shown are means ± S.E.M. for at least three independent experiments. *Significantly different from 2J2 (-152/+98) in the presence of c-Jun (P<0.001).

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5.7 Discussion

Results presented in Chapter 4 indicated that the proximal promoter region of CYP2J2 between -152 to -50 bp was essential for regulation by c-Jun. An AP-1-like element at -56 to -63 bp within this region was involved in c-Jun binding and in c-Jun-dependent activation of CYP2J2. However, mutagenesis of this element only partially decreased activation by c-Jun, which suggested that an additional element within this region contributes to c-Jun-responsiveness. Results in Chapter 5 characterised a second c-Jun binding site at -105 to -95 bp that has a major role in the activation of CYP2J2 by c-Jun. From extensive EMSA studies, the -105 to -95 bp sequence was found to be involved in direct binding to c-Jun. Mutagenesis of this sequence impaired c-Jun binding and significantly decreased both c-Jun-dependent transactivation and basal transcriptional activity of the CYP2J2 promoter. Analysis of the -105 to -95 bp sequence of CYP2J2 revealed little homology to the AP-1 consensus binding sequence. This region contains the sequence TGGGAGG, which shares only three of the seven nucleotides constituting the AP-1 consensus binding motif (TGAG/CTCA; Angel et al., 1987; Angel & Karin, 1991). Thus, the -105 to -95 bp sequence of CYP2J2 is an atypical AP-1 binding element. In this respect, it is noteworthy that numerous genes that are regulated by Fos and Jun proteins do not contain consensus AP-1 recognition sequences within their promoter regions, and a wide range of different sequence motifs have been shown to modulate AP-1-dependent gene transcription (Ryseck & Bravo, 1991; Pestell et al., 1994; Chinenov & Kerppola, 2001). The interaction of AP-1 proteins with other transcription factors has also been demonstrated, and interactions of this type may facilitate the binding of AP-1 components to promoter sequences that only slightly resemble the AP-1 consensus motif (Chinenov & Kerppola, 2001; van Dam & Castellazzi, 2001). c-Jun, in particular, has been shown to interact with several different transcription factors including GATA-2 (Kawana et al., 1995), Ets family members (Bassuk & Leiden, 1995; Behre et al., 1999), NF-NB (Yang et al., 1999), NFAT (McCaffrey et al., 1993), Smad family proteins (Liberati et al., 1999) and Sp1 (Kardassis et al., 1999) to activate transcription of target genes. Thus, it is possible that c-Jun activates transcription of the CYP2J2 gene in concert with other transcription factors that bind in close proximity to the -105 to -95 bp c-Jun binding motif. Analysis of the promoter sequence surrounding the -105 to -95 bp element detected several potential Sp1 binding motifs that overlap (-110/-100 bp and -98/-88 bp), or flank (-76/-66 bp and -71/-61 bp), the -105 to -95 bp c-Jun binding site. Significantly, a recent study indicated that Sp1 positively regulates

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CYP2J2 gene transcription via the -76/-66 bp sequence (Spiecker et al., 2004). Thus, it appears that c-Jun and Sp1 bind to adjacent elements within the CYP2J2 proximal promoter, and raises the possibility that these two transcription factors interact to regulate CYP2J2 gene expression. Indeed, interaction between c-Jun and Sp1 is an important regulatory mechanism for a number of different gene promoters (Kardassis et al., 1999; Chen & Chang, 2000; Wu et al., 2003; Chen & Feener, 2004). In accord with findings relating to the -56/-63 bp c-Jun-response element (Chapter 4), EMSA analysis established that c-Jun, but not c-Fos, was present in the protein complex bound to the -105/-95 bp sequence in normoxic HepG2 cells. Again, this finding is consistent with the basal expression of c-Jun, but not c-Fos, in unstimulated HepG2 cells (De Cesare et al., 1995), and with the proposed role of c-Jun in the maintenance of CYP2J2 expression. Because c-Fos abolishes the activation of the CYP2J2 promoter by c-Jun and contributes to the down-regulation of CYP2J2 in hypoxia, the interaction of the -105/-95 bp element with c-Fos was examined. Co-transfection studies with c-Fos and c-Jun expression plasmids established that c-Fos exerts its suppressive effects at both the -56/-63 bp and -105/-95 bp c-Jun-response elements, as mutagenesis of either site alone did not eliminate c-Fos-mediated inhibition. However, mutagenesis of both elements rendered the CYP2J2 promoter unresponsive to AP-1. Both c-Jun and c-Fos were present in the protein complex bound to the -105/-95 bp element in EMSAs with nuclear extracts prepared from hypoxic HepG2 cells, and EMSAs with recombinant proteins confirmed the capacity of the -105/-95 bp element to interact specifically with both c-Jun homodimers and c-Fos/c-Jun heterodimers. These results contrast those observed for the -56/-63 bp element, which did not interact with c-Fos/c-Jun heterodimers (Chapter 4). Taken together, these findings suggest that the inhibitory effect of c-Fos on the expression of CYP2J2 involves divergent actions at the two c-Jun-response elements within the c-Jun-responsive module of the promoter. At the -56 to -63 bp element, the up-regulation of c-Fos in hypoxia and the resultant shift from c-Jun homodimers to c-Fos/c-Jun heterodimers leads to diminished binding of, and transactivation by, c-Jun. On the other hand, the up-regulation of c-Fos in hypoxia alters the composition of the AP-1 complex bound at the -105 to -95 bp element from c-Jun to c-Fos/c-Jun. The resultant inhibitory effect on transactivation of CYP2J2 may arise from the ability of the c-Fos/c-Jun heterodimer to bind to the -105 to -95 bp element, but not facilitate activation. As noted previously, Fos and Jun proteins are known to regulate transcription through cooperation with other transcription factors, and AP-1 binding sites are often part of complex regulatory elements that contain binding sites for many transcription

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factors (Kerppola & Curran, 1993). Complexes composed of different AP-1 proteins have been shown to induce different DNA bending patterns at AP-1 sites, which is likely to alter the interactions between transcription factors bound to flanking sequences, and may account for the different transcriptional activity of different AP-1 dimers (Kerppola & Curran, 1993). Binding of c-Jun to the -105 to -95 bp element under basal conditions may facilitate interplay with other important transcription factors and stimulate transcriptional activation. On the other hand, the binding of c-Fos/c-Jun heterodimers at the -105 to -95 bp element under conditions in which c-Fos is highly expressed, such as hypoxia, may promote a conformation of the CYP2J2 promoter that prevents the optimal interaction of transcription factors. An analogous situation has been reported for the osteocalcin gene promoter, in which transactivation by the vitamin D receptor is impaired by the binding of c-Fos/c-Jun heterodimers at an AP-1 element adjacent to the vitamin D receptor response element (Schüle et al., 1990). As mentioned previously, the CYP2J2 promoter region surrounding the -105 to -95 bp c-Jun binding element contains several sequences that are highly homologous to the Sp1 consensus recognition element, one of which has been recently reported to bind Sp1 and positively activate transcription of CYP2J2 (Spiecker et al., 2004). It seems reasonable then to speculate that c-Jun and Sp1 may cooperate to positively activate transcription of the CYP2J2 gene, and that this cooperation may be impaired by c-Fos/c-Jun heterodimerisation. Indeed, positive regulation by c-Jun and Sp1, and inhibition by c-Fos, has been demonstrated for the monoamine oxidase B gene promoter (Wong et al., 2002). Initial experiments presented in this chapter suggested that the -152/-122 bp region of CYP2J2 may also contain an element that participates with the -56/-63 bp and -105/-95 bp elements in regulation by c-Jun. However EMSA analysis did not provide evidence of significant binding interactions between the -152 to -122 bp region and c-Jun, and transfection studies with further mutagenised constructs showed that the -152 to -122 bp region was not important for basal transcriptional activity or c-Jun-dependent transactivation of CYP2J2. Moreover, mutagenesis of both the -56/-63 bp and -105/-95 bp elements completely abolished c-Jun-responsiveness. In summary, this chapter has established that the CYP2J2 promoter contains a c-Jun-responsive module consisting of two distinct c-Jun-response elements, an AP-1-like element at -56/-63 bp and an atypical AP-1 site at -105/-95 bp, that positively regulate expression of the gene. The binding of c-Jun to these two response elements stimulates transcriptional activation of the CYP2J2 gene and is important for maintaining expression of CYP2J2 in resting cells. By apparently distinct mechanisms, c-Fos inhibits or interferes with transcriptional activation by c-Jun at these two critical

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regulatory elements, thus contributing to the down-regulation of CYP2J2 expression in hypoxia.

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Chapter 6

General Discussion

The human CYP gene superfamily encodes enzymes that play a fundamental role in the oxidative metabolism of lipophilic compounds from both exogenous and endogenous sources (Nebert & Russell, 2002). CYP enzymes catalyse the phase I metabolism of hydrophobic xenobiotics, such as drugs, plant products and environmental chemicals and carcinogens, into polar products that are more readily excreted from the body (Gonzalez, 1988; Vermeulen, 1996; Nebert & Russell, 2002). While generally leading to detoxification and increased clearance, CYP-mediated metabolism of certain xenobiotics can result in bioactivation and formation of toxic metabolites. This is considered to be an important factor in the pathogenesis of chemically-associated tissue injury, and in the increased risk of certain cancers and birth defects (Nebert & Gonzalez, 1987; Gonzalez, 1988; Nebert & Russell, 2002). Aside from their role in the metabolism of foreign chemicals, CYP enzymes participate in the biosynthesis and metabolism of numerous essential endogenous compounds including steroid hormones, cholesterol and bile acids, fat-soluble vitamins, arachidonic acid and eicosanoids (Nebert & Russell, 2002). The recently identified CYP2J2 is an arachidonic acid epoxygenase that catalyses the oxidative metabolism of arachidonic acid (AA) into epoxyeicosatrienoic acids (EETs; Wu et al., 1996). These eicosanoids have a number of potent biological activities in the various different cells and tissues in which they are formed. For example, EETs have been found to modulate the activity of Na+ and Ca2+ ion channels in cardiomyocytes (Chen et al., 1999; Lee et al., 1999) and improve the recovery of the myocardium following ischaemia and reperfusion (Wu et al., 1997). EETs are potent vasodilators of coronary, renal and cerebral arteries (Roman, 2002), and are proposed to be the elusive endothelium-derived hyperpolarising factor (EDHF) that mediates

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vasodilation in these vascular beds (Fisslthaler et al., 1999; Fleming, 2004). In addition to their vasodilatory actions, EETs mediate other protective effects within the vascular system which include inhibition of vascular inflammation (Node et al., 1999) and smooth muscle cell migration (Sun et al., 2002), activation of the vascular fibrinolytic pathway (Node et al., 2001), and protection against hypoxia-reoxygenation-induced cell death (Yang et al., 2001), and are therefore proposed to play an important role in vascular homeostasis and protection against cardiovascular diseases such as atherosclerosis (Spiecker & Liao, 2005). EETs also have potent biological effects outside the vasculature which include the modulation of renal fluid and electrolyte transport, mitogenic activity, and effects on peptide hormone secretion (Roman, 2002). Thus, EETs regulate a number of fundamental processes within the body, and while further research is required, it is likely that altered cellular production of EETs contributes to the pathophysiology of certain diseases and conditions such as cardiovascular disorders and hypertension (Yang et al., 2001; Kroetz & Zeldin, 2002; Roman, 2002; Spiecker & Liao, 2005). CYP2J2 is highly expressed in many of the tissues in which EETs mediate important activities, including the heart and vasculature (Wu et al., 1996; Node et al., 1999), liver (Wu et al., 1996; Enayetallah et al., 2004), lungs (Zeldin et al., 1996a), gastrointestinal tract (Zeldin et al., 1997b; Enayetallah et al., 2004), kidney (Wu et al., 1996; Enayetallah et al., 2004), and pancreas (Zeldin et al., 1997a; Enayetallah et al., 2004). Thus, CYP2J2 is likely to be important in cellular physiology, and alterations in the expression or activity of CYP2J2 may contribute to the development of certain human diseases and pathological conditions. For example, a study by Yang et al. (2001) found that CYP2J2 protein was decreased in vascular endothelial cells that had been subjected to hypoxia and reoxygenation. Transfection of cells with CYP2J2 cDNA, or direct treatment with EETs, prior to culture at low oxygen tension, markedly enhanced cell survival (Yang et al., 2001). Given that endothelial injury arising from ischaemia and reperfusion is a critical event in the development of atherosclerosis and ischaemic heart disease, it is proposed that reduced CYP2J2 protein expression and the subsequently lower production of cytoprotective EETs may contribute to the pathogenesis of cardiovascular disorders (Yang et al., 2001). Lending support to this hypothesis is a recent study that found that increased risk of coronary artery disease was related to a functional polymorphism within the CYP2J2 gene that decreased CYP2J2 promoter activity and EET biosynthesis (Spiecker et al., 2004). Despite its likely importance in cellular physiology and pathophysiology, little is known about the mechanisms that regulate human CYP2J2 gene expression. The work presented in this thesis is the first major study of the molecular mechanisms controlling

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CYP2J2 gene expression. Specifically, this study has demonstrated that CYP2J2 gene expression is impaired in hypoxia, and has identified factors that regulate the differential expression of CYP2J2 in normal and reduced oxygen conditions. Previous findings by Yang et al. (2001) demonstrated that the endothelial expression of CYP2J2 protein was decreased by hypoxia and reoxygenation. This study was the first to suggest that the CYP2J2 gene may be differentially regulated in response to altered oxygen levels, but did not offer any insight into the underlying molecular mechanism. Using this earlier study as a starting point, the expression of CYP2J2 was investigated in HepG2 cells cultured in normoxia and hypoxia. Consistent with findings in endothelial cells, CYP2J2 mRNA and protein levels were decreased in HepG2 cells following exposure to hypoxia, indicating that down-regulation of CYP2J2 occurs at a pre-translational level. Several redox-responsive transcription factors, such as AP-1, play a key role in mediating the changes in gene transcription and expression that occur in hypoxia (Norris & Millhorn, 1995; Michiels et al., 2001; Minet et al., 2001). Significantly, a number of potential AP-1 binding sites were identified within the 5’-flanking region of the CYP2J2 gene, leading to the hypothesis that the AP-1 transcription factor may be involved in the pre-translational down-regulation of CYP2J2 in HepG2 cells cultured at low oxygen. AP-1 proteins from the Jun family form homodimers that appear to predominate in resting cells (Angel & Karin, 1991; Minet et al., 2001). In response to external stimuli such as hypoxia, the expression of Fos proteins is induced; this leads to the formation of heterodimers between Jun and Fos proteins (Karin, 1995; Karin et al., 1997; Mishra et al., 1998; Minet et al., 2001), which may exhibit distinct binding affinities and transcriptional activities to Jun homodimers (Angel & Karin, 1991; Ryseck & Bravo, 1991; Kerppola & Curran, 1993). Thus, the AP-1 system can regulate the basal transcription of genes, and can also rapidly modulate the expression of target genes in response to stress and other stimuli (Angel & Karin, 1991). By modulating target gene expression in response to a wide range of extracellular stimuli, AP-1 influences critical processes such as cellular proliferation, differentiation, apoptosis and adaptation to cellular stress (Angel & Karin, 1991; Mechta-Grigoriou et al., 2001; Shaulian & Karin, 2001). c-Jun has been identified as the major component of the AP-1 complex in numerous cells, and c-Fos is its best characterised partner (Karin et al., 1997; Vogt, 2001). In transient transfection and cell culture experiments, an important role was established for the major AP-1 proteins, c-Fos and c-Jun, in the regulation of CYP2J2 in normoxia and hypoxia. The down-regulation of CYP2J2 mRNA was closely associated with pronounced up-regulation of c-Fos protein. c-Jun strongly enhanced

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transcriptional activity of the CYP2J2 promoter, whereas c-Fos was inactive and also abolished the activation of CYP2J2 elicited by c-Jun alone. Thus, CYP2J2 promoter activity is modulated by the composition of AP-1 complexes present within the cell, which is a reflection of the differential expression of subunit proteins in response to the cellular environment (Angel & Karin, 1991; Karin, 1995; Shaulian & Karin, 2001). These results implicate the constitutively expressed c-Jun in the maintenance of CYP2J2 expression in normoxic cells. Down-regulation of CYP2J2 in hypoxia occurs in response to c-Fos up-regulation and the appearance of c-Fos/c-Jun heterodimers which are transcriptionally inactive towards the CYP2J2 gene promoter. Consistent with the relationship between CYP2J2 and c-Fos expression, a short period of reoxygenation following hypoxic exposure returned c-Fos protein levels to those observed in normoxic cells, and CYP2J2 mRNA to control levels. CYP2J2 protein levels remained suppressed after hypoxia-reoxygenation, which is consistent with the earlier study by Yang et al. (2001) conducted in endothelial cells exposed to hypoxia- reoxygenation, and is probably due to the time required for synthesis of new CYP2J2 protein. This phenomenon is not unique to CYP2J2; although CYP mRNAs respond rapidly to various stimuli, changes in CYP protein expression are delayed. For example, CYP1A1 mRNA has been shown to be maximally induced within the liver at 9 hr after treatment with the chemical pyridine, whereas maximal induction of CYP1A1 protein was not apparent until 24 hr after treatment (Iba et al., 1999). The c-fos gene is highly responsive to a range of different stimuli (Wisdom, 1999). In addition to hypoxia, c-fos gene transcription and expression is activated in response to growth factors and cytokines, as well as other forms of cellular stress (Angel & Karin, 1991; Wisdom, 1999). Thus, it is feasible that CYP2J2 may be down- regulated in response to a variety of other external stimuli that up-regulate c-fos and alter the cellular composition of the AP-1 complex. Indeed, preliminary experiments undertaken by other members of this laboratory have indicated that treatment of HepG2 cells with the pro-inflammatory cytokine IL-6, or with the nitric-oxide-releasing agent sodium nitroprusside, also induces c-fos expression and down-regulates CYP2J2. While further studies are now required, these preliminary findings suggest that CYP2J2 down-regulation may be a more general response to exogenous cellular stresses. This is an area which warrants further investigation given the likely importance of CYP2J2 and EETs in cellular physiology. Another area that requires further attention is the mechanism by which stress stimuli up-regulate c-fos and suppress CYP2J2 expression. The MAP kinase signalling pathway is recognised to play a key role in the regulation of AP-1 activity (Karin, 1995; Whitmarsh & Davis, 1996). In particular, the ERK, JNK and p38 MAP kinases are variously activated by a

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range of extracellular stimuli, which modulates the expression and activity of the AP-1 subunit proteins (Karin, 1995; Whitmarsh & Davis, 1996). Preliminary experiments conducted in this laboratory have found that MAP kinase inhibitors modulate c-fos and CYP2J2 expression in sodium nitroprusside-treated HepG2 cells. c-Fos, acting in conjunction with c-Jun, has generally been found to potentiate the activity of AP-1-responsive gene promoters (Sassone-Corsi et al., 1988; Hirai et al., 1989). This is probably due to the increased stability and binding efficiency of c-Fos/c-Jun heterodimers for the AP-1 consensus compared to that of c-Jun homodimers (Halazonetis et al., 1988; Ryseck & Bravo, 1991). However, there are also several studies that report the antagonism of c-Jun-dependent gene activation by c-Fos; this situation is similar to that observed in the present studies of CYP2J2 gene regulation. Indeed, Kovacic-Milivojevic & Gardner (1992) demonstrated that the human atrial natriuretic peptide gene promoter is strongly activated by c-Jun, and that over- expression of c-Fos inhibits c-Jun-dependent promoter activity. c-Fos also inhibited c-Jun-dependent activation of the human urokinase gene promoter (De Cesare et al., 1995). Thus, CYP2J2 appears to be one of several human genes whose expression is regulated positively by c-Jun and negatively by c-Fos. The work presented in this thesis supports the growing number of studies documenting the importance of the intracellular concentration and differential dimerisation of AP-1 proteins on the transcription of AP-1-responsive genes (Suzuki et al., 1991; Kerppola & Curran, 1993). The CYP2J2 5’-flank between -152 to -50 bp relative to the translation start site was crucial for c-Jun-dependent transactivation of CYP2J2. EMSA and transfection studies identified two distinct gene elements that were involved in transactivation: an AP-1-like element at -56 to -63 bp, and an atypical c-Jun binding element at -105 to -95 bp. Both elements interacted strongly with c-Jun homodimers, and separate mutagenesis of either element significantly impaired activation by c-Jun, while mutagenesis of both elements completely abolished c-Jun-responsiveness. Furthermore, c-Jun interacted with both elements in HepG2 cells cultured under normoxia. Mutagenesis of either element impaired basal transcriptional activity of the CYP2J2 promoter in HepG2 cells, and mutagenesis of both elements almost completely suppressed basal promoter activity. These findings are consistent with an essential role for c-Jun in the maintenance of CYP2J2 expression in normoxia via interaction with the -56/-63 bp and -105/-95 bp promoter elements. The -105/-95 bp site may be the more important of the two elements, as mutagenesis of this sequence produced a more pronounced decrease in basal and c-Jun-inducible promoter activity than after mutagenesis of the -56/-63 bp element. The -105/-95 bp sequence is an atypical AP-1 element that shares limited homology with

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the AP-1 consensus binding sequence. Indeed, this sequence was not detected during automated screening of the CYP2J2 promoter region with the transcription factor identification program MatInspector, and only came to attention during extensive EMSA analysis. Regulation of AP-1-responsive gene promoters by sequences that deviate significantly from the AP-1 consensus is not uncommon. Indeed, many genes regulated by Fos and Jun proteins do not contain consensus AP-1 binding sequences within their regulatory regions, and an increasing number of different sequence motifs have been shown to mediate AP-1-dependent gene transcription (Ryseck & Bravo, 1991; Pestell et al., 1994; Chinenov & Kerppola, 2001). AP-1 proteins interact with many different transcription factors, and such interactions may facilitate the binding of AP-1 proteins to regulatory sequences that deviate significantly from the AP-1 consensus binding motif (Chinenov & Kerppola, 2001; van Dam & Castellazzi, 2001). Thus, it is possible that interactions between c-Jun and another transcription factor bound to a nearby regulatory element within the CYP2J2 promoter may augment binding to the -105/-95 bp sequence. GATA proteins (Kawana et al., 1995), Ets family proteins (Behre et al., 1999), NFAT (McCaffrey et al., 1993), NF-NB (Yang et al., 1999), Smad proteins (Liberati et al., 1999) and Sp1 (Kardassis et al., 1999) are among the various transcription factors that have been shown to interact with c-Jun to regulate gene transcription. The CYP2J2 promoter contains several potential Sp1 binding sites that overlap or flank the -105/-95 bp c-Jun binding site. A recent report that the putative Sp1 binding sequence at -76/-66 bp may mediate Sp1-dependent transactivation of CYP2J2 is of interest (Spiecker et al., 2004). Many different human gene promoters have been found to be regulated by interactions between c-Jun and Sp1, including the human vimentin (Wu et al., 2003), 12(S)-lipoxygenase (Chen & Chang, 2000) and p21WAF1/Cip1 (Kardassis et al., 1999) gene promoters. Thus, the close proximity of functional c-Jun and Sp1 binding elements within the CYP2J2 promoter raises the possibility that these two transcription factors may also interact to regulate CYP2J2 gene expression. Co-transfection studies established that c-Fos suppresses c-Jun-dependent activation at both the -56/-63 bp and -105/-95 bp c-Jun-response elements. However, EMSAs revealed important differences in the capacity of c-Fos to interact with these sites. c-Jun, but not c-Fos, was present in the protein complex bound to both elements in HepG2 cells cultured under normal oxygen conditions, which is consistent with the basal expression of c-Jun, but not c-Fos, in unstimulated HepG2 cells (De Cesare et al., 1995), and with c-Jun-dependent activation of the CYP2J2 promoter under basal conditions. Exposure of HepG2 cells to hypoxia stimulated a change in the composition of the complex bound at the -105/-95 bp element from c-Jun to both c-Jun and c-Fos.

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Experiments with recombinant proteins confirmed the capacity of this element to interact with both c-Jun homodimers and c-Fos/c-Jun heterodimers. In contrast, the -56/-63 bp element did not interact with c-Fos/c-Jun heterodimers, and the up-regulation of c-Fos in hypoxic HepG2 cells diminished the binding of c-Jun to this region. These findings suggest that c-Fos suppresses CYP2J2 gene expression via divergent actions at the two c-Jun-response elements within the CYP2J2 proximal promoter (Figure 6.1). Hypoxic up-regulation of c-Fos and the resultant shift from c-Jun homodimers to c-Fos/c-Jun heterodimers results in diminished binding of c-Jun to the -56/-63 bp element, thereby decreasing c-Jun-dependent promoter activity and CYP2J2 gene transcription (Figure 6.1 A and B).

A NORMOXIA

Sp1 ? Sp1 ?

RNA ++++ ? c-Jun c-Jun ? c-Jun c-Jun Pol.

-105/-95 bp -56/-63 bp

B HYPOXIA

c-Fos Sp1 ? Sp1 ? c-Jun

RNA + c-Fos ? c-Jun ? Pol.

-105/-95 bp -56/-63 bp

Figure 6.1 Schematic representation of the transcriptional regulation of the CYP2J2 proximal promoter by c-Jun and c-Fos in normoxia and hypoxia A. In normoxia, the binding of c-Jun homodimers to the -56/-63 bp and -105/-95 bp elements stimulates transcriptional activation of the CYP2J2 promoter. Interactions between c-Jun and Sp1 at putative Sp1 binding sites flanking the -105/-95 bp element may by important for activation of the CYP2J2 promoter and are included in the schematic. B. c-Fos inhibits or interferes with transcriptional activation by c-Jun at the two promoter elements; thus, the up-regulation of c-Fos in hypoxia and the formation of c-Fos/c-Jun heterodimers leads to diminished transactivation of the CYP2J2 promoter. Question marks indicate unidentified transcription factors, and transcriptional activity is indicated by red arrows. RNA Pol.=RNA polymerase.

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A similar scenario is apparently responsible for the divergent regulation of the human urokinase gene promoter by c-Jun and c-Fos. While urokinase promoter activity was positively regulated by the binding of c-Jun/ATF-2 heterodimers to an AP-1-like element within the upstream enhancer region of the gene, c-Fos inhibited c-Jun-dependent promoter activity, and EMSA studies revealed that c-Fos/c-Jun heterodimers interacted very poorly with the AP-1 response element (De Cesare et al., 1995). These authors proposed that the inhibitory effects of c-Fos on urokinase promoter activity was due to sequestration of c-Jun, which limited its availability for dimerisation with ATF-2 (De Cesare et al., 1995). It appears likely that up-regulation of c-Fos in hypoxia decreases the availability of c-Jun for binding at the -56/-63 bp element and inhibits transcription of the CYP2J2 gene. By contrast, a different mechanism may be responsible for c-Fos-dependent suppression of CYP2J2 promoter activity at the -105/-95 bp c-Jun-response element. The inhibition of CYP2J2 promoter activity presumably reflects the ability of c-Fos/c-Jun heterodimers to bind to the -105/-95 bp element, but not facilitate activation. Numerous reports have documented differences in transcriptional activation by different combinations of AP-1 proteins at a particular AP-1 response element (Angel & Karin, 1991; Kerppola & Curran, 1993; Mechta-Grigoriou et al., 2001). Because AP-1 binding sites are frequently found in complex regulatory regions, and Fos and Jun proteins cooperate with other transcription factors to regulate gene expression, the promoter context is probably an important determinant of the transcriptional activity of Fos and Jun proteins (Kerppola & Curran, 1993). Transcriptional activation requires the bending or looping of DNA to enable transcription factors bound at distinct regulatory elements to interact with each other and with RNA polymerase and its associated factors at the transcription initiation site (Kerppola & Curran, 1991; Kerppola & Curran, 1993). Different combinations of AP-1 proteins induce different patterns of bending at AP-1 sites, which may influence transcription factor interactions at flanking sequences and with the transcription initiation complex (Kerppola & Curran, 1993). Alternate bending by different AP-1 dimers may also inhibit or promote the binding of proteins to overlapping or adjacent regulatory elements (Kerppola & Curran, 1993). Considerations of this type may account for the different transcriptional activity of different AP-1 complexes and contribute to their regulatory specificity (Kerppola & Curran, 1993). Significantly, c-Jun homodimers and c-Fos/c-Jun heterodimers have been demonstrated to induce DNA bends that are directed in opposite directions (Kerppola & Curran, 1991; Kerppola & Curran, 1993). The binding of c-Jun to the -105/-95 bp sequence under basal conditions may facilitate interaction with other transcription factors bound to nearby

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regulatory elements and promote transcriptional activation (Figure 6.1 A). In contrast, the binding of c-Fos/c-Jun heterodimers to the -105/-95 bp element under conditions in which c-Fos is highly expressed, such as hypoxia, may induce a conformational change in the CYP2J2 promoter that is suboptimal for transcriptional activation (Figure 6.1 B). Similar mechanisms may regulate the transcriptional activity of other gene promoters. For example, interaction between c-Jun and the glucocorticoid receptor has been found to generate a stable complex that strongly enhances transcriptional activation of the murine proliferin gene promoter. In contrast, transcriptional repression is observed in the presence of both c-Jun and c-Fos, which may reflect a conformational change due to c-Fos/c-Jun heterodimers that prevents transactivation (Diamond et al., 1990). The potential Sp1 binding sites adjacent to the -105/-95 bp c-Jun-response element may be significant for CYP2J2 transcription. Given that interactions between c-Jun and Sp1 are important for the positive regulation of numerous different gene promoters (Kardassis et al., 1999; Chen & Chang, 2000; Wu et al., 2003), it is tempting to speculate that c-Jun and Sp1 may similarly activate the CYP2J2 gene (Figure 6.1 A). It is also possible that up-regulation of c-Fos in hypoxia impairs this cooperation and diminishes transcriptional activation (Figure 6.1 B). Again, a similar situation has been reported in which the human monoamine oxidase B gene promoter is activated by c-Jun and Sp1, but inhibited by c-Fos (Wong et al., 2002). Further attention is now warranted as to whether such regulatory mechanisms account for the divergent regulation of CYP2J2 by c-Jun and c-Fos at the two promoter elements. The present studies were undertaken in the HepG2 hepatocarcinoma-derived cell line, which is an appropriate cellular environment. CYP2J2 mRNA and protein are readily detected in the liver (Wu et al., 1996; Enayetallah et al., 2004) and in HepG2 cells. In addition, HepG2 cells are readily transfectable and have been successfully used to study the regulation of certain other CYP genes (Ibeanu & Goldstein, 1995; Goodwin et al., 1999; Nitta et al., 1999; Zhang & Chiang, 2001). Considered together, the HepG2 cell line is a suitable system in which to study CYP2J2 gene regulation. Results presented in this thesis have established that c-Jun and c-Fos play an important role in controlling the expression of the CYP2J2 gene in normoxic and hypoxic HepG2 cells. Because these AP-1 proteins are widely expressed in many different cell types (Wisdom, 1999), it seems plausible that the CYP2J2 gene may be similarly regulated in the other cell types in which it is expressed. Indeed, the similar effect of hypoxia on the expression of CYP2J2 in endothelial (Yang et al., 2001) and HepG2 cells is suggestive of a common regulatory mechanism. However, it remains to

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be established whether common regulatory pathways exist in the range of cells and tissues that express CYP2J2. It is noteworthy that other Jun and Fos proteins were capable of modulating CYP2J2 promoter activity in transient transfection studies in a similar fashion to c-Jun and c-Fos. Thus, both JunB and JunD stimulated CYP2J2 promoter activity, although to a much smaller extent than c-Jun, and c-Fos also suppressed the actions of these Jun family members. Furthermore, with the exception of the combination of JunB and Fra-1, the other Fos family proteins (Fra-1 and Fra-2) also suppressed CYP2J2 activation by c-Jun and the other Jun proteins. These findings suggest that the CYP2J2 promoter may be generally activated by Jun proteins and suppressed by Fos proteins. While the relative expression of c-Jun and c-Fos plays a key role in controlling CYP2J2 expression in HepG2 cells, it is possible that different subsets of Jun and Fos proteins may regulate CYP2J2 expression in other cellular environments. Indeed, the specific AP-1 proteins expressed in a particular cell may be an important determinant of CYP2J2 expression in response to stress stimuli. In summary, the findings described in this thesis have improved the understanding of the molecular mechanisms involved in regulating expression of the human CYP2J2 gene. In particular, this study has identified the AP-1 transcription factor as a key modulator of the transcriptional activity of the CYP2J2 gene, and shown that the major AP-1 proteins c-Jun and c-Fos play a crucial role in regulating the differential hepatic expression of CYP2J2 in normoxia and hypoxia. The switch between c-Jun homodimers and c-Jun/c-Fos heterodimers at the -56/-63 bp and -105/-95 bp elements in the CYP2J2 upstream sequence enables the rapid modulation of gene expression in situations of cellular stress. The altered regulation of CYP2J2 in cells may have major physiological consequences. CYP2J2-derived fatty acid epoxides are now recognised to play a fundamental role in numerous cellular processes. Therefore, changes in the cellular production of EETs, due to impaired expression of CYP2J2, is likely to have important physiological and pathophysiological consequences. Decreased expression of CYP2J2 in hypoxia, and the resultant lower levels of cellular EETs, may contribute to cellular injury seen in pathophysiological diseases such as cardiovascular disease, stroke, cancer and tissue injury. Indeed, maintenance of CYP2J2 expression helps to protect cells against hypoxic injury (Yang et al., 2001), and modulation of CYP2J2 expression may prove to be an attractive therapeutic target. An understanding of the factors that regulate CYP2J2 expression is therefore essential for the development of strategies - pharmacological or otherwise - to favourably manipulate the cellular expression of CYP2J2. To date, very limited information has appeared on the regulation of CYP2J2 expression. This thesis contains

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novel information on the molecular mechanisms that control CYP2J2 gene expression, leading to further avenues of investigation that could facilitate a greater understanding of the regulation of this important human gene.

188 References

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