Mycobacterium tuberculosis - Research Explorer

1

Biochemical and drug targeting studies of

Mycobacterium tuberculosis cholesterol oxidase P450

enzymes

A thesis submitted to the University of Manchester for the degree of Doctor of

Philosophy in the Faculty of Life Sciences.

2015

Cecilia Nwadiuto Amadi

2

Table of contents

Title page 1

Table of contents 2

List of figures 10

List of tables 18

Abbreviations 20

Abstract 24

Declaration and copyright statement 25

Dedication 27

Acknowledgement 28

Chapter 1- Introduction 29

1.1 Tuberculosis: An Update 29

1.1.1 An ‘Ancient and Modern’ disease 29

1.1.2 The Tuberculosis Burden 31

1.1.3 Signs and Symptoms of Tuberculosis 32

1.1.4 Transmission of Tuberculosis: Latent TB Versus Active TB 33

1.2 Mycobacterium tuberculosis: A Description of a Debilitating Human Pathogen

37

1.3 Tuberculosis Treatment: Past, Present and Future 40

1.3.1 The Past: Genesis of Anti-Tubercular Drug Discovery 40

3

1.3.2 The Present: Anti-Tubercular Drugs in Current Use 42

1.3.3. The Future: New Tuberculosis Drug Candidates in Development 48

1.3.4 Anti-Tubercular Drug Resistance: A Cause for Therapeutic Failures 59

1.4 The Cytochrome P450 Systems 60

1.4.1 Structure, Function and Mechanism 60

1.4.2 The P450 Catalytic Cycle 67

1.4.3. Cytochrome P450 Redox Partners 72

1.5 The Mycobacterium tuberculosis Cytochrome P450 Enzymes

77

1.5.1 Discovery of Mtb P450s and the Quest for their Physiological Roles 77

1.5.2 The Cholesterol Oxidase P450 Enzymes 81

1.5.2.1 CYP125A1(Rv3545c):Essential for Mtb Viability and Infectivity

88

1.5.2.2 CYP142A1(Rv3518c): Functional Redundancy 91

1.5.2.3 CYP124A1 (Rv2266): A Methyl-Branched Lipid-Hydroxylase 96

1.5.2.4 Cholesterol Catabolism: A Promising Drug Target in Mycobacterium tuberculosis

100

1.5.3. CYP51B1: The First Member of the CYP51 Family Identified in Prokaryotes

103

1.5.4 CYP121A1: An Essential Gene for Mtb Viability 105

1.5.5 CYP130A1 (Rv1256c): Essential for Virulence in Mtb? 107

1.5.6 CYP126A1 (Rv0778) 108

1.5.7 CYP128A1: An Essential Enzyme with a Role in Hydroxylation of Respiratory Menaquinone

109

1.5.8 Other Partially Characterized P450 Systems in Mycobacterium tuberculosis

111

1.5.9 Azole Antibiotics: Non-Selective Inhibitors of Mtb Cytochrome P450 Enzymes

115

4

1.6 Novel Drug Discovery Approaches 118

1.6.1 Fragment Based Drug Discovery (FBDD): A Novel Approach to Development of New P450 Inhibitor Scaffolds

118

1.6.2 High Throughput Screening (HTS) 123

1.7 Justification of Research 126

1.8 Aims of Research 128

Chapter 2 - Materials and Methods 129

2.1 Materials 129

2.2 Methods 129

2.2.1 Preparation of Plasmid DNA for Expression Constructs 129

2.2.1.1 Source and Description 129

2.2.1.2 Plasmid DNA Purification 130

2.2.2 Generation of Glycerol Stocks of E. coli Transformants 132

2.2.3 Expression Trials for CYP124A1 and CYP142A1 P450s 132

2.2.4 Scale up of the Expression of CYP142A1 134

2.2.5 Scale up of the Expression of CYP124A1 135

2.2.6 Protein Purification for CYP124A1 and CYP142A1 136

2.2.7 Assessment of P450 Concentration and Purity 138

2.2.8 Determination of P450 Extinction Coefficients Using the Pyridine Hemochromagen Method

139

2.2.9 UV-Visible Spectroscopic Studies of Mtb P450s 140

5

2.2.9.1 Binding Assays with Substrates and Inhibitors 140

2.2.9.2 Formation of P450 Carbon Monoxide and Nitric Oxide Adducts

141

2.2.10 Isothermal Titration Calorimetry (ITC) Studies on Mtb P450s 143

2.2.11 Guanidinium Chloride Denaturation of CYP142A1 144

2.2.12 Redox Potentiometry Studies on CYP124A1 and CYP142A1 144

2.2.13 Multi-Angle Laser Light Scattering (MALLS) Studies of Mtb P450s 146

2.2.14 Differential Scanning Calorimetry Analysis of Mtb P450s 147

2.2.15 Electron Paramagnetic Resonance (EPR) Spectroscopy of P450s 148

2.2.16 CYP124A1 Steady-State Kinetics 148

2.2.17 P450 Protein Crystallization and Structure Determination 149

2.2.18 CYP142A1 Nano-ESI Mass Spectrometry 151

Chapter 3 - Biochemical and Biophysical Characterization of P450 CYP142A1: An Example of Functional Redundancy in the Mycobacterium tuberculosis cholesterol oxidases?

153

3.1 Introduction 153

3.2 Results and Discussion 158

3.2.1 Expression and Purification of CYP142A1 158

3.2.2 CYP142A1 Substrate Binding Assays 163

3.2.3 Inhibitor Binding Assays 170

3.2.4 Binding Analysis with CYP142A1 Fragment Hits 178

3.2.5 Binding Analysis with Compounds from CYP121A1 Fragment Elaboration Hits

182

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3.2.6 CYP142A1 Fe(II)-CO Adduct and NO Adduct Formation 187

3.2.7 Determination of an Extinction Coefficient for Mtb CYP142A1 Using the Pyridine Hemochromogen Method

191

3.2.8 Light Scattering (MALLS) Analysis of CYP142A1 194

3.2.9 Electron Paramagnetic Resonance (EPR) Analysis of CYP142A1 199

3.2.9.1 EPR Analysis with Selected CYP142A1 Ligands 199

3.2.9.2 EPR Analysis for CYP142A1 Fragments Hits 202

3.2.9.3 EPR Analysis of CYP142A1 Bound to MEK Compounds 205

3.2.10 Differential Scanning Calorimetry Studies of CYP142A1 207

3.2.11 Guanidinium Chloride Denaturation of CYP142A1 210

3.2.12 Isothermal Titration Calorimetry (ITC) Analysis of CYP142A1 214

3.2.13 Redox Potentiometry of CYP142A1 217

3.2.14 Nanoelectrospray Ionization Mass Spectrometric Analysis of Mtb CYP142A1−Ligand Interactions

223

3.2.14.1 NanoESI Mass Spectra of Ligand-Free CYP142A1 224

3.2.14.2 Interaction of CYP142A1 with DTT 226

3.2.14.3 Interaction of CYP142A1 with Econazole 227

3.2.14.4 Analysis of the Interaction of CYP142A1 with Cholestenone

229

3.2.14.5 Interaction of CYP142A1 with Solvents 232

3.3 Summary 233

Chapter 4 - Biochemical and Biophysical characterization of CYP124A1: A promiscuous enzyme with broad substrate specificity in Mycobacterium tuberculosis

240

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4.2.1 Expression and Purification of CYP124A1 243

4.2.2 Spectroscopic Analysis of CYP124A1 251

4.2.2.1 The UV-Visible Spectrum of CYP124A1 251

4.2.2.2 CYP124A1 Optical Titrations with Substrates 252

4.2.2.3 CYP124A1 Inhibitor Binding Assays 259

4.2.2.4 CYP124A1 Fragment Binding Assays 266

4.2.3 CYP124A1 Heme Iron Coordination by Carbon Monoxide and Nitric Oxide

274

4.2.4 Determination of the CYP124A1 Heme Extinction Coefficient 278

4.2.5 Steady-State Kinetic Analysis for CYP124A1 280

4.2.6 Multiangle Laser Light Scattering (MALLS) Analysis of CYP124A1 286

4.2.7 Thermostability Analysis of CYP124A1 by Differential Scanning Calorimetry

287

4.2.8 Determination of the Heme Iron Redox potentials of Ligand-Free and Ligand-Bound CYP124A1

293

4.2.9 Electron Paramagnetic Resonance (EPR) Analysis of CYP124A1 299

4.2.9.1 EPR Analysis with CYP124A1 Substrates and Azole Inhibitors

299

4.2.9.2 CYP124A1 EPR Analysis with Fragments and MEK Compounds

305

4.3 Summary 309

Chapter 5 - Structural Biology of Ligand-Bound Complexes of the Cholesterol Oxidising P450s CYP142A1 and CYP124A1

315

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5.2.1 X-ray Crystallographic Studies and Structure Determination for CYP142A1 and CYP124A1

317

5.2.1.1 Crystal Structure of the CYP142A1:Cholestenone Complex 321

5.2.1.2 Crystal structure of the CYP124A1:Cholestenone Complex 327

5.2.1.3 A Comparison of Cholestenone-Bound CYP124A1, CYP125A1 and CYP142A1 Structures

335

5.2.1.4 Crystal Structure of the CYP142A1:Econazole Complex 340

5.2.1.5 Crystal Structures of the CYP142A1 in Complex with Fragment-Based Screening Hits

348

5.3 Summary 360

Chapter 6 - Conclusions and Future Directions 365

6.1 Conclusions 365

6.2 Future directions 372

References 375

9

Appendix 396

Word count: 84,129

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List of Figures

Chapter 1

Figure 1.1: World map showing tuberculosis high-burden countries 32

Figure 1.2: Tuberculosis transmission 36

Figure 1.3: An electron micrograph of Mycobacterium tuberculosis 37

Figure 1.4: The Mycobacterium tuberculosis cell wall 39

Figure 1.5: Structures of some selected anti-TB drugs in clinical use for Mtb infections

48

Figure 1.6: Chemical structure of PDKA 52

Figure 1.7: Chemical structures of HT1171 and GL5 53

Figure 1.8: Chemical structures of nitroimidazole compounds 56

Figure 1.9: Chemical structure of Bedaquiline - a diarylquinoline TB drug 57

Figure 1.10: Chemical structure of SQ109 58

Figure 1.11: Chemical structure of BTZ043 59

Figure 1.12: Heme B prosthetic group 62

Figure 1.13: Spectral features for cytochrome P450 and its ferrous–carbon monoxide complex

63

Figure 1.14: Typical topology of a cytochrome P450 67

Figure 1.15: Schematic representation of the d-orbital electron configurations for low- and high-spin ferric heme iron

68

Figure 1.16: A schematic representation of the P450 compound I (oxyferryl radical cation species)

70

Figure 1.17: Schematic representation of the catalytic cycle of a cytochrome P450 enzyme

71

Figure 1.18: Schematic representation of a variety of P450 redox systems and P450 fusion proteins

76

Figure 1.19: Evolutionary analysis of Mtb P450s 78

Figure 1.20: Genetic organization of cholesterol metabolising gene clusters in Rhodococcus sp. RHA1 and Mtb: A comparison

83

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Figure 1.21: The chemical structures of cholesterol and cholest-4-en-3-one

87

Figure 1.22: Cholesterol side chain oxidation reactions 87

Figure 1.23: Structural features of Mtb CYP125A1 in complex with diverse substrates and inhibitor molecules.

90

Figure 1.24: Structural features of CYP142 enzymes from Mtb and M. smegmatis

95

Figure 1.25: CYP124A1 catalyses the -hydroxylation of phytanic acid and other methyl-branched lipids

98

Figure 1.26: Structural features of CYP124A1 from Mtb 99

Figure 1.27: Cholesterol catabolic pathway 100

Figure 1.28: Features of CYP51B1 from Mtb 104

Figure 1.29: CYP121A1 catalyzes the formation of an intramolecular C-C bond between 2 tyrosyl carbon atoms of cyclodityrosine

106

Figure 1.30: Structural features of Mtb CYP121A1 in complex with the substrate (cyclodityrosine) and fluconazole

106

Figure 1.31: Crystal structures of ligand-free and econazole-bound CYP130A1.

108

Figure 1.32: Biosynthesis of the Mtb S881 sulfolipid 111

Figure 1.33: Chemical structures of selected azole antibiotics 117

Figure1.34: A schematic representation of the Fragment Based Drug Discovery (FBDD) Approach

117

Figure 1.35: Application of a FBDD approach to Mtb CYP121. 122

Figure 1.36: A schematic representation of the High Throughput Screening (HTS) approach

125

Figure 1.37: Structures of CYP130A1 with HTS hits (heterocyclic arylamines) bound in the active site

126

Chapter 3

Figure 3.1: Protein purification of the Mtb CYP142A1 from the pET15b/CYP142A1 plasmid

159

12

Figure 3.2: Purification of Mtb CYP142A1 using hydroxyapatite (HA) column chromatography

161

Figure 3.3: Purification of Mtb CYP142A1 using a SuperdexTM S-200 gel filtration column

162

Figure 3.4: Optical titration of CYP142A1 with cholest-4-en-3-one 166

Figure 3.5: Optical titration of CYP142A1 with cholesterol 167

Figure 3.6: Optical binding of CYP142A1 with lanosterol 168

Figure 3.7: CYP142A1 binding titration with econazole 172

Figure 3.8: CYP142A1 binding titration with Miconazole 173

Figure 3.9: CYP142A1 binding titration with clotrimazole 174

Figure 3.10: CYP142A1 binding titration with bifonazole 175

Figure 3.11: CYP142A1 binding titration with sodium cyanide 176

Figure 3.12: Compounds hits from an initial CYP142A1 fragment screen 179

Figure 3.13: CYP142A1 binding titration with NMR170 180



Figure 3.16: Elaborated compounds developed from CYP121A1 fragment hits

184

Figure 3.17: CYP142A1 binding with MEK046 185

Figure 3.18: CYP142A1 binding with MEK065 186

Figure 3.19: UV-visible spectra for gaseous ligand-bound complexes of CYP142A1

191

Figure 3.20: Pyridine hemochromagen spectra for CYP142A1. 193

Figure 3.21: Light scattering (MALLS) data for CYP142A1 in the absence of DTT

196

Figure 3.22: Light scattering (MALLS) data for CYP142A1 in the presence of DTT

197

Figure 3.23: Cysteine residues in CYP142A1 199

Figure 3.24: EPR analysis of CYP142A1 and various ligand complexes. 200

Figure 3.25: EPR analysis of interactions of azole drugs and nitrogen-containing fragments with CYP142A1.

205

13

Figure 3.26: X-band EPR spectra for CYP142A1 in complex with different compounds from the MEK series

207

Figure 3.27: Differential scanning calorimetry analysis of CYP142A1. 209

Figure 3.28: Guanidinium chloride denaturation of ligand-free CYP142A1 213

Figure 3.29: Guanidinium chloride denaturation of cholestenone-bound CYP142A1

214

Figure 3.30: Isothermal titration calorimetric (ITC) binding studies of fragments to CYP142A1

215

Figure 3.31: Redox potentiometry of ligand-free CYP142A1 219

Figure 3.32: Redox potentiometry for cholestenone-bound CYP142A1 221

Figure 3.33: Redox potentiometry for clotrimazole-bound CYP142A1 222

Figure 3.34: NanoESI mass spectra of ligand-free CYP142A1 224

Figure 3.35: NanoESI mass spectrum of 10 μM CYP142A1 with DTT (0-5mM)

226

Figure 3.36: NanoESI mass spectra of 10 μM CYP142A1 with econazole 229

Figure 3.37: NanoESI mass spectra of 10 μM CYP142A1 with cholestenone 231

Figure 3.38: NanoESI mass spectrum of 10 μM CYP142A1 with solvents 232

Chapter 4

Figure 4.1: Nickel affinity chromatography purification of CYP124A1 246

Figure 4.2: Hydroxyapatite (HA) column chromatography purification of CYP124A1

248

Figure 4.3: Purification of Mtb CYP124A1 using a SuperdexTM S-200 gel filtration column

250

Figure 4.4: The UV-visible spectrum for purified, ferric CYP124A1 251

Figure 4.5: Optical titration of CYP124A1 with cholest-4-en-3-one 254

Figure 4.6: Optical titration of CYP124A1 with cholesterol 255

Figure 4.7: Optical titration of CYP124A1 with phytanic acid 256

14

Figure 4.8: Optical titration of CYP124A1 with pristane 256

Figure 4.9: Optical titration of CYP124A1 with geraniol 257

Figure 4.10: Optical titration of CYP124A1 with geranylgeraniol 257

Figure 4.11: Binding of bifonazole to CYP124A1 262

Figure 4.12: Binding of clotrimazole to CYP124A1 262

Figure 4.13: Binding of econazole to CYP124A1 263

Figure 4.14: Binding of miconazole to CYP124A1 264

Figure 4.15: Binding of NMR170 to CYP124A1 266

Figure 4.16: Compounds identified as CYP124A1-specific fragment hits 268



Figure 4.19: Elaborated compounds developed from CYP121A1 fragment hits

272

Figure 4.20: Binding of MEK066 to CYP124A1 273

Figure 4.21: UV-visible absorbance features of CYP124A1 and its carbon monoxide complex

275

Figure 4.22: UV-visible absorbance spectra of CYP124A1 in ferric and ferric-NO bound forms

277

Figure 4.23: The pyridine hemochromogen complex of CYP124A1 280

Figure 4.24: Steady-state kinetic analysis for CYP124A1 using an E. coli redox partner system

283

Figure 4.25: Steady-state kinetic analysis for CYP124A1 using a spinach redox partner system

284

Figure 4.26: MALLS analysis of CYP124A1 286

Figure 4.27: DSC analysis of CYP124A1 in substrate-free and substrate-bound forms

289

Figure 4.28: DSC analysis of CYP124A1 in azole-bound forms 290

Figure 4.29: DSC analysis of CYP124A1 in MEK series-bound forms 291

Figure 4.30: DSC analysis of CYP124A1 in fragment-bound forms 292

Figure 4.31: Redox potentiometry of ligand-free CYP124A1 296

Figure 4.32: Redox potentiometry of phytanic acid-bound CYP124A1 297

15

Figure 4.33: Redox potentiometry of econazole-bound CYP124A1 298

Figure 4.34: EPR spectra of CYP124A1 in ligand-free and sterol-bound forms

303

Figure 4.35: EPR spectra of CYP124A1 in complex with methyl-branched lipids

304

Figure 4.36: EPR spectra of CYP124A1 in complex with azole inhibitors 305

Figure 4.37: EPR analysis of CYP124A1 bound to MEK ligands 308

Figure 4.38: EPR spectra of CYP124A1-specific fragment hits 309

Chapter 5

Figure 5.1: Co-crystals of the CYP142A1:cholestenone complex. 321

Figure 5.2: Overall view of the CYP142A1 cholestenone-bound complex (dimer)

322

Figure 5.3: The CYP142A1 substrate binding channel 324

Figure 5.4: Overview of the CYP142A1-cholestenone binding pocket 325

Figure 5.5: Superimposed structures of ligand-free and cholestenone-bound CYP142A1

326

Figure 5.6: Co-crystals of the CYP124A1:cholestenone complex 327

Figure 5.7: Overall view of the CYP124A1:cholestenone complex 329

Figure 5.8: Superimposed structures of ligand-free and cholestenone-bound forms of CYP124A1

330

Figure 5.9: The CYP124A1 substrate binding channel 331

Figure 5.10: Overview of the CYP124A1-cholestenone binding pocket 332

Figure 5.11: Comparison of cholestenone and phytanic acid binding modes to CYP124A1

334

Figure 5.12: Comparison of CYP125A1, CYP142A1 and CYP124A1 cholestenone binding modes

337

Figure 5.13: The substrate binding channels in the three P450 cholesterol oxidases

338

16

Figure 5.14: Comparison of the cholestenone binding modes for CYP125A1, CYP142A2, CYP142A1 and CYP124A1 with that of the CYP142A2 cholesterol sulfate complex

339

Figure 5.15: Co-crystals of the CYP142A1-econazole complex 342

Figure 5.16: Overall view of CYP142A1 econazole-bound complex 343

Figure 5.17: Overview of the econazole binding pocket in CYP142A1 344

Figure 5.18: Slab view of the CYP142A1 active site channel showing econazole bound to the heme iron

345

Figure 5.19: Superimposed structures of the ligand-free and econazole-bound forms of CYP142A1 showing their secondary structure elements

346

Figure 5.20: Superimposed structures of the CYP125A1 and CYP142A1 econazole complexes

347

Figure 5.21: Superimposed structures of the CYP142A1 and CYP130A1 econazole complexes

348

Figure 5.22: A sample of diamond-shaped CYP142A1 native crystals 350

Figure 5.23: The CYP142A1-NMR623 complex structure 351

Figure 5.24: The CYP142A1-NMR170 complex structure 352

Figure 5.25: Co-crystals of the CYP142A1-NMR491 complex 353

Figure 5.26: The structure of the CYP142A1-NMR491 complex 354

Figure 5.27: Co-crystals of the CYP142A1:1-phenylimidazole complex 354

Figure 5.28: The structure of the CYP142A1:1-phenylimidazole complex 355

Figure 5.29: Superimposed structures of the various CYP142A1-fragment complexes

356

Figure 5.30: Superimposed structures of ligand-free and fragment-bound CYP142A1 enzymes

357

Figure 5.31: Superimposed structures of CYP142A1-fragment complexes with the CYP142A1-econazole complex

358

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Appendix

Figure S1: CYP142A1 (Rv3518c) DNA sequence 398

Figure S2: Synthetic CYP124A1 (Rv2266) gene (codon optimised for E. coli)

400

Figure S3: The CYP142A1 (Rv3518c) gene region in Mycobacterium tuberculosis

401

Figure S4: The CYP124A1 (Rv2266) gene region in Mycobacterium tuberculosis

402

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List of Tables

Chapter 1

Table 1.1: Classical anti-tubercular drugs 43

Table 1.2 : New anti-tubercular drugs in different phases of development 50

Table 1.3: The twenty (20) P450 enzymes in Mycobacterium tuberculosis

80

Chapter 2

Table 2.1 Composition of growth media used for protein production and the amounts of components added per litre of medium

134

Chapter 3

Table 3.1: Typical CYP142A1 purification table 162

Table 3.2: Binding spectral characteristics and Kd values for CYP142A1 ligands

187

Table 3.3: DSC data for the thermal unfolding of CYP142A1 210

Table 3.4: Thermodynamic parameters of CYP142A1-fragment interactions derived from ITC and optical titrations

216

Table 3.5: Redox titration data for CYP142A1 223

Chapter 4

Table 4.1: Binding affinity of CYP124A1 with lipid substrates 253

Table 4.2: Binding affinities for CYP124A1 with azole drug inhibitors 261

Table 4.3: Binding affinity for CYP142A1 fragment hits with Mtb 267

19

cholesterol oxidase P450s

Table 4.4: Binding affinity of CYP124A1 for fragment hits 270

Table 4.5: Binding spectral characteristics of Mtb cholesterol oxidases with CYP121A1 elaborated ligands

273

Table 4.6: Steady-state kinetic parameters for substrate-dependent NADPH oxidation by CYP124A1

285

Table 4.7: DSC data for thermal unfolding of CYP124A1 293

Table 4.8: Redox titration data for CYP124A1 299

Chapter 5

Table 5.1: X-ray data collection and refinement statistics for CYP142A1- and CYP124A1- cholestenone complexes

319

Table 5.2: X-ray data collection and refinement statistics for CYP142A1- econazole/fragment complexes

320

Table 5.3: Dissociation constants for the binding of selected azole drugs to Mtb CYP51B1, CYP121A1, CYP124A1, CYP125A1 and CYP142A1

341

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Abbreviations

°C Degrees Celsius

µ Micro (10-6)

A Absorbance

Å Angstrom (10-10 m)

AIDS Acquired immunodeficiency syndrome

bp Base pair

CO Carbon monoxide

CYP or P450 Cytochrome P450

cYY Cyclo-L-Tyrosine-L-Tyrosine

δ-ALA Delta-aminolevulinic acid

DMSO Dimethyl sulfoxide

DNA Deoxyribonucleic acid

DOTS Direct Observed Therapy Scheme

DSC Differential Scanning Calorimetry

DTT Dithiothreitol

EDTA Ethylenediaminetetraacetic acid

EPR Electron Paramagnetic Resonance

FAD Flavin Adenine Dinucleotide

FDR Ferredoxin reductase

FDX Ferredoxin

FldR/FIdA E. coli flavodoxin reductase/flavodoxin

Fe-S Iron Sulphur

FMN Flavin mononucleotide

g Gram

GdmCl Guanidinium Chloride

HS High-spin

igr Mtb Intracellular growth region

21

INH Isoniazid

ITC Isothermal Titration Calorimetry

IPTG Isopropyl-β-D-1-thiogalactopyranoside

KCl Potassium chloride

Kd Dissociation constant

kDa KiloDalton

kg Kilogram

KPi Potassium phosphate

L Litre

LB Luria-Bertani (Lysogenic Broth) growth medium

LS Low-spin

m Milli (10-3)

M Molar

mce Mammalian cell entry

MALLS Multi Angle Laser Light Scattering

mg Milligram

MDR-TB Multi-drug resistant tuberculosis

MIC Minimum inhibitory concentration

Mtb Mycobacterium tuberculosis

n Nano (10-9)

NAD(H) Nicotinamide adenine dinucleotide (reduced form)

NADP(H) Nicotinamide Adenine Dinucleotide Phosphate (reduced form)

nanoESI-MS Nano-ElectroSpray Ionization-Mass Spectrometry

NHE Normal hydrogen electrode

NMR Nuclear magnetic resonance

nm Nanometre

NO Nitric oxide

OD600 Optical density at 600 nm

22

O/N Overnight

PCR Polymerase Chain Reaction

PEG Polyethylene glycol

P450 Cytochrome P450

P450 BM3 CYP102A1 from Bacillus megaterium

P450cam CYP101A1 from Pseudomonas putida

PCW Periplasmic cell wall

PDIM Phenolphthiocerol-dimycocerosate

Pdr Putidaredoxin reductase

Pdx Putidaredoxin

PIM Phenylimidazole

PMSF Phenylmethanesulfonyl fluoride

PZA Pyrazinamide

RD Region of deletion

RIF Rifampicin

rpm Revolutions per minute

SDS Sodium dodecyl sulphate

SEC Size exclusion chromatography

SOC Super optimal broth with catabolite repression

sp Spinach

TB Tuberculosis

TB Terrific Broth medium

TDM Trehalose-dimycolate

Tm melting temperature

Tet Tetracycline

µg Microgram

µl Microlitre

μM Micromolar

23

UV-Vis UltraViolet-Visible

v/v volume per volume

w/v weight per volume

WHO World Health Organization

XDR-TB Extensively or extremely drug-resistant Mtb

X-rays Electromagnetic radiation with a wavelength in the range of 0.01 to 10 nm

YT Yeast Tryptone medium

ɛ Extinction coefficient

24

Abstract

A thesis submitted to the University of Manchester in 2015, for the degree of Ph.D by Cecilia Nwadiuto Amadi, entitled:

Biochemical and drug targeting studies of Mycobacterium tuberculosis cholesterol oxidase P450 enzymes.

Mycobacterium tuberculosis (Mtb), a deadly pathogen, has scourged mankind for many centuries and has remained a major threat to global world health. Tuberculosis, the disease caused by this bacterium, is a major cause of death in developing nations and there is potential for its re-emergence in developed countries. An alarming rise in cases of multidrug-resistant and extremely-drug resistant tuberculosis (MDR-TB and XDR-TB) that do not respond to the customary first-line antibiotics necessitates the urgent need for development of new anti-TB drugs. Mtb becomes engulfed in human macrophages post infection of the host, but persists in the harsh environment of the human lungs by utilization of host cholesterol as a carbon source. The P450s CYP125A1, CYP142A1 and CYP124A1 are responsible for catalysing the side-chain degradation of cholesterol, which is critical for cholesterol to be used in the Mtb β-oxidation pathway for energy production. This PhD thesis focuses on understanding the structure/mechanism of the Mtb cholesterol 27-oxidases with the aim of facilitating the development of novel inhibitors of these P450s, which are crucial for Mtb to infect the host and to sustain infection. CYP142A1 and CYP124A1 were purified through three chromatographic steps with contaminating proteins successfully removed to give highly pure forms of these enzymes following the final purification step. Spectrophotometric titrations indicate that CYP142A1 and CYP124A1 bind tightly to cholesterol and cholestenone (and also to branched-chain methyl lipids for CYP124A1), highlighting their physiological roles in sterol and fatty acid metabolism, respectively. Binding analyses with a range of azole antibiotics revealed tight binding to bifonazole, clotrimazole, miconazole and econazole, and weak binding to fluconazole. Studies with compounds from a fragment screening library revealed weak binding to fragment hits for the cholesterol oxidases, but much tighter binding to these enzymes was found for ‘elaborated’ hits from a previous fragment screen on the Mtb cyclodipeptide oxidase CYP121A1, indicative of improved ligand potency achieved via ‘fragment merging’ strategies, and of structural similarities between these diverse Mtb P450s. Light scattering data indicate that CYP142A1 exists in dimeric form in solution, but becomes monomeric when treated with DTT; while CYP124A1 is completely monomeric. Crystal structures of CYP142A1 and CYP124A1 in complex with cholestenone, econazole and fragment library hits were determined. CYP142A1 crystal structures with econazole and fragment hits revealed heme coordination via the heterocyclic nitrogen in an azole group, and provide important data towards design of superior inhibitor drugs. The binding of cholestenone within the active site channels of CYP124A1 and CYP142A1 revealed an alignment favourable for C27 hydroxylation of the cholestenone side chain, which supports the physiological roles of CYP142A1 and CYP124A1 (as well as CYP125A1) in host cholesterol catabolism.

25

Declaration

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intellectual property (the “Intellectual Property”) and any reproductions of

copyright works in the thesis, for example graphs and tables (“Reproductions”),

which may be described in this thesis, may not be owned by the author and may be

owned by third parties. Such Intellectual Property and Reproductions cannot and

must not be made available for use without the prior written permission of the

owner(s) of the relevant Intellectual Property and/or Reproductions.

26

iv. Further information on the conditions under which disclosure, publication and

commercialisation of this thesis, the Copyright and any Intellectual Property and/or

Reproductions described in it may take place is available in the University IP Policy

(see http://documents.manchester.ac.uk/DocuInfo.aspx?DocID=487), in any

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University Library’s regulations (see

http://www.manchester.ac.uk/library/aboutus/regulations) and in The University’s

policy on Presentation of Theses.

27

Dedication

This thesis is dedicated to God the father, God the son, God the Holy Spirit. Thank

you Lord for what you have made out of me, this project would not have been

possible without you. It is all about you, Jesus.

28

Acknowledgements I would like to say a big thank you to my supervisor, Professor Andrew Munro for

the opportunity to undertake a PhD in his lab, for his help, support and guidance

throughout the course of my PhD program. Thank you for taking time to teach me

at every step of this work. Special thanks to Dr Kirsty McLean for all that you taught

me in the Laboratory. I would also like to thank Professor David Leys and Dr Colin

Levy for their kind support and guidance through my structural biology work. Dr

Karl Fisher, thank you so much for all the advice and encouragement, they helped

me a lot. Many thanks, Dr Alistair Fielding, for your kind support at the Photon

Science Institute. Furthermore, i appreciate Dr Hazel Girvan, Dr James Belcher, Dr

John Hughes, Dr Binuraj Menon, Marina Golovanova, Michiyo Sakuma and all senior

members of the Munro group for their guidance and help throughout my research

work. To all other members of the Munro group and the entire molecular

enzymology group, I appreciate you all.

My mentor, Professor O.E Orisakwe of the University of Port Harcourt, Nigeria, is

greatly acknowledged ‒ his advice and encouragement stimulated me thus far.

I wish to give a special appreciation to the Nigerian Government and Faculty for the

Future Fellowship from the Schlumberger Foundation for supporting my PhD

studies.

To my family and friends, thanks for your love, support and encouragement. VPA

Manchester church/choir- thanks for being a family away from home. God bless you

all.

29

Chapter 1

Introduction

1.1 Tuberculosis: An Update

1.1.1 An ‘Ancient and Modern’ disease

Tuberculosis (TB) is a disease caused by an intracellular, pathogenic bacterium

known as Mycobacterium tuberculosis (Mtb) and mainly affects the lungs

(pulmonary TB) but can affect other parts of the human body as well (extra-

pulmonary TB) (Russell, 2007). Tuberculosis is an important world health challenge

and has moved up the ladder to become the second leading cause of death globally,

from an infectious, communicable disease, after the human immunodeficiency virus

(HIV). This disease, which has been documented to kill one person every 20

seconds, is predominant in the developing world and has been termed a ‘disease of

poverty’ (Dartois, 2014).

Tuberculosis (TB) remained a mystery disease until the mid-19th century (Chan et

al., 2013). It has scourged mankind for ages and is postulated to have existed more

than 150 million years ago (Daniel, 2006). However, the mystery behind the

tuberculosis disease was unravelled in 1882 when Robert Koch reported the

isolation and cultivation of the causative agent for tuberculosis disease, the

bacterium Mycobacterium tuberculosis (Ho, 2004). Archaeological evidence of early

tuberculosis was found in Egypt and America (Daniel, 2000, Arriaza et al., 1995, Salo

30

et al., 1994). Tuberculosis, also referred to as the ‘white plague’ in early days, was

named by John Bunyan in the 17th century as the ‘captain of all these men of death’

when the disease claimed millions of lives in Europe (Ducati et al., 2006). Other

names include: Consumption, phthisis (meaning ‘wasting away’), scrofula (swollen

glands of the neck), Pott's disease (TB of the bone), and lupus vulgaris (TB of the

skin); but the popular name ‘Tuberculosis’ or ‘TB’ comes from the words 'tubercle

bacillus' a term introduced by Johann L Schönlein to accommodate all the multiple

localizations of this disease. The term ‘tubercle’ refers to a warty outgrowth found

on bones and skin or in the case of TB in the lungs (Das, 2000, Riva, 2014).

Tuberculosis was the major cause of death in Europe and the United States in times

past, although because of the complex forms of symptomatic manifestations, the

disease was often confused with other diseases (Bloom and Murray, 1992). TB

infection resulted in high mortality rates in the past, which were attributed to

absence of drug remedies to tackle the disease. However, the discovery of antibiotic

drugs in the 20th century led to reduction in mortality rates from the disease (Ducati

et al., 2006, Kremer and Besra, 2002).

Even at the start of the 21st century, tuberculosis (TB) resulted in the death of more

than two million people every year, with the highest occurrences of morbidity and

mortality in Sub-Saharan Africa and South East Asia (Kaufmann and McMichael,

2005, Ulrichs and Kaufmann, 2006). To date, TB remains a great threat to global

human health (Johnston et al., 2010). Over two billion people (a third of the world

population) are infected with the latent form of the bacterium and, out of this

population, about 10% will develop active tuberculosis disease in their lifetime.

31

Presently, over two million lives are lost annually due to active tuberculosis

infection (WHO, 2014a, Johnston et al., 2010).

1.1.2 The Tuberculosis Burden

Tuberculosis continues to be one of the deadliest diseases documented in history.

The TB burden can be estimated in terms of incidence (the number of new and

relapse cases of the disease arising in a given time period, usually one year),

prevalence (the number of cases of the disease at a given point in time) and

mortality (the number of deaths caused by TB in a given period of time, usually one

year).

In 2013, the World Health Organization (WHO) documented that about 9 million

people developed the disease and about 1.5 million died from the infection. From

these numbers of deaths, 360 000 patients were HIV positive. The six countries with

the largest number of incident cases in 2013 were India (2.0–2.3 million), China

(0.9–1.1 million), Nigeria (340 000−880 000), Pakistan (370 000−650 000), Indonesia

(410 000−520 000) and South Africa (410 000−520 000) (WHO, 2014a).

The number of TB cases co-infected with HIV was observed to be highest in African

countries. In total, 34% of TB cases were estimated to be co-infected with HIV in

this region, which can be added to the 78% of TB cases among people living with

HIV world-wide. Hence, the African region accounts for about four out of every five

HIV-positive TB cases and TB deaths among patients who were HIV-positive (WHO,

2014a).

32

Figure 1.1: World map showing tuberculosis high-burden countries (green shades). Source: Global Tuberculosis report 2014 (WHO, 2014a).

1.1.3 Signs and Symptoms of Tuberculosis

TB is usually a chronic, slowly progressing disease that often remains undiagnosed

in patients for many years. The disease presents with many symptoms and can

affect many organs, but the most common form in adults is a chronic pulmonary

disease (Young et al., 2008). Typical signs of tuberculosis are chronic or persistent

cough and sputum production (If the disease is at an advanced stage the sputum

will contain blood), fatigue, lack of appetite, weight loss, fever and night sweats.

Infection of other organs causes a wide range of symptoms. Tuberculosis can mimic

many forms of disease and must always be considered if no firm diagnosis has been

made. Although tuberculosis predominantly affects the lungs, it can cause disease

in any organ and must be included within the differential diagnosis of a vast range

of clinical presentations (Lawn and Zumla, 2011). A high index of suspicion for

33

tuberculosis must especially be maintained when caring for patients living with HIV

infection, since risk of tuberculosis is high and diagnosis is problematic (Lawn and

Zumla, 2011).

1.1.4 Transmission of Tuberculosis: Latent TB Versus Active TB

Tuberculosis (TB) is categorised as either ‘Latent TB’ or ‘Active TB’. In Latent TB, the

bacterium is dormant and non-replicating while engulfed in the human

macrophage. The patient is asymptomatic and cannot infect another human. In

Active TB, there is active replication of the bacteria. The patient presents with overt

manifestation of clinical symptoms and the disease at this stage is highly

communicable (Corbett et al., 2003, Kaur et al., 2014). Latent TB with absence of

clinical manifestation comprises about 90% of all cases of tuberculosis infection

(Kondratieva et al., 2014). About 10% of latent infection eventually reactivates to

active TB and becomes contagious. Mechanisms of transition to latency and TB

reactivation are not well understood (Kondratieva et al., 2014).

When Mycobacterium tuberculosis infects the host and reaches its target organs

(the lungs), the bacterium is engulfed by immunological cells (neutrophils and

macrophages) and faces the first line defence mechanism of the host, which

involves the natural and adaptive immune responses. These protective factors

could eradicate the pathogen, but in most cases the bacterium transits to dormancy

or latency, which involves the mobilization of host immune cell populations into

the lung governed by chemokines and cytokines (Kondratieva et al., 2014). This

34

entire process ensures mycobacterial containment at least in the initial stages of

infection (Kondratieva et al., 2014, Monin and Khader, 2014). Latency in

tuberculosis infection is generally accompanied by formation of well-structured

granuloma in the lungs (consisting mainly of leukocytes) which is highly isolated

from the surrounding tissues (Ulrichs and Kaufmann, 2006). Disruption of the

granuloma structure and a rising number of mycobacterial cells result in cavity

formation and active TB disease (Ulrichs and Kaufmann, 2006, Kondratieva et al.,

2014). Research has revealed lung granuloma formation as the hallmark of

pulmonary tuberculosis (Ulrichs and Kaufmann, 2006). Its shape and structure is

characterized by a central necrotic centre enclosed by circular layers of

macrophages, epithelioid cells, multinucleated Langhans giant cells and

lymphocytes (Ulrichs and Kaufmann, 2006, Mariano, 1995). Mtb is contained by a

cellular wall and a fibrotic outer layer, and this stops it from spreading throughout

the host (Ulrichs and Kaufmann, 2006). Latent infection results when the bacterium

is successfully contained within the primary lesion and this is observed as calcified

granulomatous lesions. Transition to active tuberculosis is prevented if this lesion is

maintained or controlled, and this occurs in more than 90% of individuals (Ulrichs

and Kaufmann, 2002, Zhang, 2004).

TB is transmitted through the air when people who have an active Mtb infection

cough, sneeze, or otherwise transmit their saliva through the air (Konstantinos,

2010). Large studies of TB contacts have shown that airborne transmission of Mtb is

promoted by prolonged and close contact with an infectious case, and the key

35

determinant is the amount of time spent sharing room air with a patient who has an

active infection (Richeldi et al., 2004).

Tuberculosis (TB) infection can be established with or without a visible primary

lesion. Such a lesion can be anywhere in the lungs but tends to be sited towards the

base and close to the pleura (Rook et al., 2005). Following exposure to the

bacterium, most humans and animals develop a TH1 cell response and the ability of

peripheral-blood T cells to release interferon gamma in response to secreted

antigens of Mtb is used as a test for exposure (Pathan et al., 2000, Richeldi et al.,

2004). In approximately 90% of infected individuals, this response causes the Mtb

bacilli to remain in the tissues in a latent state, and disease does not occur (Rook et

al., 2005).

In humans, there is a phase of blood-borne spread of approximately 3 weeks after

the bacterium infects initially. The vaccine (Mycobacterium bovis bacillus Calmette–

Guérin (BCG)) might block infection at this stage. A T helper 1 (TH1) cell response

develops rapidly. The infection remains latent in 90-95% of individuals for several

years, but can be reactivated when an individual is immunosuppressed, particularly

through infection with HIV (Rook et al., 2005, Kaur et al., 2014) (Figure 1.2). In

addition, the elderly, malnourished and individuals involved in substance abuse are

at high risk of developing TB (McLean and Munro, 2008). Studies have revealed a

high prevalence of TB and MDR strains in the third world, and co-infection with HIV

is a major problem in sub-Saharan Africa (McLean and Munro, 2008, WHO, 2014a).

36

Progressive disease is characterized by weight loss, toxicity of tumour-necrosis

factor, cavitation and fibrosis, even though interferon- produced by TH1 cells can

decrease the amount of fibrosis. The cavities eventually open into the bronchi,

which allow the transmission of TB by air during coughing or sneezing. Signs and

symptoms of TB infection include: a persistent cough with a duration of more than

3 weeks, bloody sputum, mucus production, night fever and chills, weight loss, loss

of appetite, night sweats, weakness and headache (Kaur et al., 2014).

Figure 1.2: Tuberculosis transmission. Image is adapted from (Kaur et al., 2014).

37

1.2 Mycobacterium tuberculosis: A Description of a Debilitating Human Pathogen

Mtb is a rod-shaped, non-motile, acid fast bacillus that causes tuberculosis infection

in humans. It has been documented that one-third of the world’s population is

already latently infected with Mtb, representing a large potential reservoir for

future reactivation of tuberculosis infection, especially in the dispensation of the

human immunodeficiency virus (HIV) pandemic (Dutta and Karakousis, 2014).

Figure 1.3: An electron micrograph of Mycobacterium tuberculosis. The image shows rod-shaped non motile bacteria. [Taken from http://medimoon.com/2012/07/a-new-weapon-could-show-promising-results-against-resistant-tb/ (Hayat, 2012)].

The architectural structure of the Mtb cell wall is quite distinct from the cell wall

structures of both Gram-negative and Gram-positive bacteria. This complex

structure of the Mtb cell wall accounts for its unusually low permeability to

common antibiotics, and for its pathogenicity and resistance to attack by the host

38

(Brennan, 2003, Alderwick et al., 2007). The Mtb cell wall is composed of two major

segments - the upper and lower segments. Outside the cytoplasmic membrane is

peptidoglycan (PG), which is covalently attached to arabinogalactan (AG) (Brennan,

2003). This covalent attachment consists of a cross-linked network of peptidoglycan

(PG) in which some of the muramic acid residues are substituted with the complex

polysaccharide AG. The arabinogalactan, in turn, is attached to the mycolic acids via

long meromycolate and short α-chains (Brennan, 2003, Alderwick et al., 2007). This

entire complex is termed the ‘cell wall core’ or the mycolylarabinogalactan–

peptidoglycan (mAGP) complex, and is essential for Mtb viability (Alderwick et al.,

2007, Dover et al., 2004). Mycolic acids are the major components of the cell wall

protective barrier and play a major role in the survival, virulence and antibiotic

resistance of Mtb (Barry et al., 1998). Drugs that inhibit mycolic acid biosynthesis,

such as isoniazid, ethambutol and pyrazinamide, are still being used as frontline

anti-TB drugs (Ouellet et al., 2010b, Zhang et al., 2005).

The upper segment consists mainly of complex lipids which are esterified with

multiple methyl-branched long-chain fatty acids, surface and trans-membrane

proteins (Brennan, 2003). These complex lipids include the phenolphthiocerol and

phthiocerol dimycocerosates (PDIMs), sulfatides (SLs), diacyltrehaloses (DATs),

triacyltrehaloses (TATs) and polyacyltrehaloses (PATs) which make up the trehalose

ester families (Ouellet et al., 2010b, Jackson et al., 2007). Unique substances

interspersing the cell wall core and associated with the mycolic acids are the

phosphatidylinositol-containing glycolipids, mainly the lipomannans and the

lipoarabinomannans (Hunter and Brennan, 1990).

39

Collectively, these structures provide a thick and robust layer of lipid on the outer

part of the cell that protects the bacterium against antibiotics, toxic substances and

the host’s immune system (Takayama et al., 2005, Ouellet et al., 2010b).

Figure 1.4: The Mycobacterium tuberculosis cell wall. The image was drawn using Microsoft PowerPoint and adapted from Ouellet et al (Ouellet et al., 2010b).

40

1.3 Tuberculosis Treatment: Past, Present and Future

1.3.1 The Past: Genesis of Anti-Tubercular Drug Discovery

The journey to tuberculosis drug discovery over the years has been a frustrating

one. Before the advent of the first anti-TB drug, tuberculosis was generally

considered ‘hopeless’ and ‘incurable’. Management was based mainly on non-

pharmacological approaches such as the use of herbal dressings, dietary

intervention, climatic remedies such as aero-therapy and heliotherapy as well as

physical measures e.g. bleeding and purging (Riva, 2014, Iseman, 2002). In addition,

tuberculosis was also claimed to have been treated with the legendary ‘royal touch’

(Dossey, 2013). There was a belief that English and French monarchs were endowed

with powers to heal TB-infected individuals by touching them in a ceremonious

ritual, where the quote ‘the King touches you, God cures you’ (‘Le Roy te touché et

Dieu te guérit’ in French) was used to validate the legitimacy of the royals and the

divine source of their healing powers (Riva, 2014). However, these measures gave

little or no solution to the cure of tuberculosis, as the disease continued to claim

many lives. The story of the search for the cure for tuberculosis is well summarised

in an educative documentary novel titled “The Greatest Story Never Told”

(Margulis, 2002).

Furthermore, in the 1930s, a major breakthrough came with the discovery of

sulphonamides and penicillin for tuberculosis treatment (Iseman, 2002).

Sulphonamide was discovered by Gerhard Domagk, assisted by chemists from Bayer

in Germany (Diacon et al., 2012b). In 1944, streptomycin was discovered by Selman

41

Waksman in New Jersey and, in the same year, Jorgen Lehman synthesised para-

aminosalicylic acid (PAS) in Sweden (Iseman, 2002). Having proven streptomycin

and PAS to be effective against tuberculosis, the British Medical Research Council

(MRC), through a randomized clinical trial, found that a combination of the two

drugs had synergistic properties (MRC, 1948, Iseman, 2002). Subsequently, the next

important step forward was the discovery of the anti-tubercular activity of isoniazid

in 1951 (Riva, 2014). This chemical compound (isonicotinyl hydrazine) was studied

and demonstrated independently in three different laboratories (Squibb, Hoffmann

La Roche and Bayer) to have a high level of anti-tuberculosis activity in experimental

animals (Riva, 2014). The compound was first synthesized by two Prague chemists:

Hans Meyer and Josef Mally in 1912, and has proved to be the most potent anti-TB

drug discovered in history, as confirmed from various clinical trials (McDermott,

1969). Further studies revealed that the addition of isoniazid to PAS and

streptomycin (‘triple therapy’) reduced drug resistance and improved the

effectiveness of tuberculosis treatment (Riva, 2014).

Challenging experiences gave physicians insights that TB treatments must be

administered using multiple drugs to prevent emergence of resistance, and that

adherence to prolonged treatments for 24 months or more may be required for a

permanent cure of TB disease (Diacon et al., 2012b). Following the success of the

‘triple therapy’, the next hurdle to cross was the reduction of duration of therapy. In

1961, this was made possible by the replacement of PAS by a more effective drug,

ethambutol, discovered by Lederle laboratories in the United States. The duration

of therapy was then reduced from 24 to 18 months (Doster et al., 1973, Riva, 2014).

42

1.3.2 The Present: Anti-Tubercular Drugs in Current Use

Research into identification of new anti-tuberculosis agents progressed with the

discovery of new compounds, mostly used as second-line drugs. These include:

viomycin, cycloserine, terizidone, kanamycin and amikacin, capreomycin and the

thioamides ethionamide and prothionamide (Diacon et al., 2012b). In the same era,

another ‘wonder’ TB compound (rifampicin) was discovered. Rifampicin was derived

from a chemical modification of ‘rifamycin’, which is a family of compounds

identified from the soil bacterium Streptomyces mediterranei (Riva, 2014).

Between 1970 and 1980, the introduction of rifampicin in tuberculosis therapy

enabled the reduction of therapy duration from 18 to 9 months (Riva, 2014). In the

same period, pyrazinamide was discovered and introduced for TB treatment,

leading to a further reduction of treatment duration to six months when combined

with isoniazid and rifampicin (Diacon et al., 2012b). This was then termed the

‘short-course chemotherapy’ (van Ingen et al., 2011). A 6-month directly observed

treatment short-course (DOTS) based on the three compounds isoniazid, rifampicin

and pyrazinamide was the foundation of tuberculosis treatment strategies world-

wide for about 30 years, and recently this was augmented with ethambutol in view

of increasing rates of isoniazid resistance (Diacon et al., 2012b).

43

Class 1 First-line anti-tubercular drugs Isoniazid Rifampin or Rifampicin

Ethambutol Pyrazinamide

Rifabutin

Class 2 Second-line agents (oral bacteriostatic drugs)

Thioamides Ethionamide Protionamide Cycloserine Terizidone

para-Aminosalicylic acid

Class 3 Second-line agents (injectables/parenterals)

Kanamycin Amikacin

Capreomycin Viomycin

Streptomycin*

Class 4 Second-line bactericidal agents (Fluoroquinolones)

Levofloxacin Moxifloxacin

Ofloxacin Gatifloxacin

Class 5 Third-line agents (Drugs with sparse clinical data)

Clofazimine Linezolid

Amoxicillin/clavulanate Thioacetazone Clarithromycin

Imipenem

Table 1.1: Classical anti-tubercular drugs. (Adapted from (Field et al., 2012, Ahmad and Mokaddas, 2014)). *Can be classified as a first generation anti-tubercular drug.

Isoniazid (INH)

INH is a prodrug and is metabolised to an isonicotinyl-NAD adduct by the bacterial

peroxidase KatG (Mitchison and Davies, 2012). This adduct inhibits InhA (encoded

by inhA gene), which is a major bacterial enzyme in the FAS II (fatty acid

biosynthesis)-dependent production of the cell wall mycolic acid (Mitchison and

Davies, 2012). Resistance often develops by mutations in katG (which encodes a

catalase-peroxidase), but also less commonly in the inhA, ahpC and ndh genes

(Almeida Da Silva and Palomino, 2011, Lee et al., 2001). In Mtb, ahpC has been

44

shown to encode for an enzyme known as alkyl hydroperoxidase reductase and that

is involved in resistance to reactive oxygen and reactive nitrogen intermediates

(Almeida Da Silva and Palomino, 2011). Mutations in the ndh gene, which encodes

an NADH dehydrogenase, was shown to cause defects in the enzyme activity that

generated an increased NADH/NAD+ ratio and co-resistance to isoniazid and

ethionamide (Lee et al., 2001). Isoniazid, though less potent against non-multiplying

cells, nevertheless has shown high bactericidal activity against dividing bacteria,

with a minimal inhibitory concentration (MIC) value of about 0.05 μg/ml (Mitchison

and Davies, 2012). INH causes peripheral neuritis and convulsions, as it

quantitatively depletes vitamin B6 stores in the body when administered at high

dosage (van der Watt et al., 2011, Mitchison and Davies, 2012). In addition,

hepatotoxicity remains a significant concern in patients treated with isoniazid

(Parekh and Schluger, 2013). A lone treatment with isoniazid is given at a dosage of

300 mg daily or 900 mg twice weekly for a period of nine months. However, when

given in combination with rifampicin, the dose changes to 300 mg isoniazid plus 600

mg rifampicin daily for 3 months (Parekh and Schluger, 2013).

Rifampicin

Rifampicin is a potent inhibitor of bacterial DNA-dependent RNA polymerase. It

binds to the rpoB-encoded portion of the bacterial RNA polymerase, hence

inhibiting the formation of new proteins (Mitchison and Davies, 2012, McLean et

al., 2007a, Almeida Da Silva and Palomino, 2011). It is highly bactericidal in its

action against Mtb throughout the course of treatment, with a MIC of 0.5 μg/ml.

However, its therapeutic margin has been shown to be narrow (Mitchison and

45

Davies, 2012). Rifampicin is administered at a dosage of 450-600 mg daily for a

period of 4 months, and side effects include hepatotoxicity, leukopenia and

thrombocytopenia (Mitchison and Davies, 2012, Parekh and Schluger, 2013).

Resistance to rifampicin is mainly due to rpoB mutations of the β subunit of the

RNA polymerase (Field et al., 2012, Uzun et al., 2002). Irrespective of this resistance

due to mutations, about 12-20% of rifampicin-resistant Mtb strains and some MDR

strains remain sensitive to rifabutin (Uzun et al., 2002, Field et al., 2012, Yoshida et

al., 2010, Cavusoglu et al., 2004), which has shown to be a better option for these

MDR-TB categories (Yew and Leung, 2008). Rifabutin is also effective in tuberculosis

patients living with HIV and who are taking protease inhibitors. This is because of its

mild effect on P450 enzyme induction, unlike rifampicin which causes about 40%

enzyme induction (Mitnick et al., 2009, Nuermberger and Mitchison, 2009).

Cytochrome P450 enzymes are responsible for the metabolism of several drugs

(Ramachandran et al., 2013). Rifampicin markedly lowers the blood levels of

protease inhibitors by inducing human cytochrome P450 CYP3A4 activity

significantly. This could result in reduction of antiretroviral activity which could lead

to the development of acquired drug resistance (Vanhove et al., 1996,

Ramachandran et al., 2013).

Pyrazinamide

Pyrazinamide is an essential front-line drug for TB treatment. Pyrazinamide,

alongside isoniazid and rifampicin, forms the bedrock of modern TB chemotherapy

(Zhang et al., 2003). Pyrazinamide, when combined with other anti-TB drugs, helps

46

to shorten the duration of therapy from 9–12 months to 6 months (Zhang et al.,

2003), because it inactivates a population of partially-dormant Mtb in acidic

environments that are not killed by other anti-TB drugs (Zhang et al., 2003).

Pyrazinamide (PZA) is a prodrug that is converted to its active moiety pyrazinoic acid

(POA) by the Mtb amidase encoded by the pncA gene, and resistance to

pyrazinamide develops from the mutation of pncA (Mitchison and Davies, 2012).

The protonated POA (HPOA) formed accumulates under acid conditions within the

bacterium and causes membrane damage (Zhang et al., 2003, McLean et al., 2007a).

Hence, pyrazinamide inactivates Mtb at acid pH (Zhang et al., 2003). This

dependency on pH has accounted for therapeutic failures with pyrazinamide in the

past (Mitchison and Davies, 2012).

Ethambutol

Ethambutol (EMB) is a bacteriostatic agent used for multi-drug-resistant (MDR)

tuberculosis. It also forms part of a cocktail of first-line anti-TB drugs in many

countries. The most common side effect of ethambutol is retro-bulbar optic neuritis

(RON) (Levy et al., 2015). EMB acts on the arabinosyl transferase EmbCAB to inhibit

cell wall synthesis (Almeida Da Silva and Palomino, 2011). Drug resistance to EMB

usually arises through mutations in the embB gene (Mitchison and Davies, 2012).

Fluoroquinolones

Fluoroquinolones have been shown to be highly effective in the treatment of MDR-

TB (Schluger, 2013). They inhibit DNA gyrase (encoded by the gyrA and gyrB genes

47

for the DNA gyrase A and B subunits) and hence block bacterial DNA synthesis (Field

et al., 2012). This stems from the ability of fluoroquinolones to interfere with action

of the A subunit of the bacterial topoisomerase, which is instrumental for

supercoiling DNA (Almeida Da Silva and Palomino, 2011). Resistance to

fluoroquinolones commonly results from mutations in the gyrA gene (Mitchison and

Davies, 2012). Fluoroquinolones with high potency against Mtb include moxifloxacin

(MFX) and gatifloxacin (GTX), which are also chemically related. Levofloxacin has

also been shown to be effective against Mtb, but slightly less active than

moxifloxacin (MFX) and gatifloxacin (Mitchison and Davies, 2012). The

fluoroquinolones as a class of drug are specifically beneficial in the treatment of

MDR-TB, with significant activity against both replicating extracellular and latent

intracellular Mtb (Mitchison and Davies, 2012, Cole and Riccardi, 2011). They are

mycobactericidal and exhibit lower MICs against Mtb than do other first-line drugs

(Donald and Diacon, 2008). The fluoroquinolones have also been suggested to

reduce the treatment duration in drug-susceptible tuberculosis (Rustomjee et al.,

2008, Conde et al., 2009, Dorman et al., 2009, Kwon et al., 2014).

Injectables (Parenterals)

This group of drugs includes streptomycin, the aminoglycosides (kanamycin,

amikacin) and cyclic peptide antibiotics (capreomycin, viomycin). They show high

potency against MDR-TB disease, second to the fluoroquinolones (Mitchison and

Davies, 2012). These drugs inhibit bacterial protein translation by binding to the 16S

rRNA in the ribosomes (Almeida Da Silva and Palomino, 2011, Mitchison and Davies,

2012, Reeves et al., 2015). Since these four drugs bind to a similar location on the

48

ribosome and share the same drug target, cross-resistance is commonly observed

(Reeves et al., 2015, Campbell et al., 2011, Georghiou et al., 2012). Cross-resistance

is linked with mutations on the 16S rRNA (rrs) sequence, and mainly with the

A1401G, C1402T and G1484T mutations (Maus et al., 2005, Jnawali et al., 2013).

The injectables are frequently used as bactericidal second-line drugs (Ahmad and

Mokaddas, 2014).

Figure 1.5: Structures of some selected anti-TB drugs in clinical use for Mtb infections. Structures redrawn with ChemDraw from (McLean et al., 2007a).

1.3.3. The Future: New Tuberculosis Drug Candidates in Development

Novel anti-tubercular drugs with more simple dosing regimens and shorter duration

of administration than the currently available anti-TB drugs are desperately needed

to ensure better patient compliance (Upton et al., 2015). An ideal new anti-TB drug

should be affordable and well tolerated with minimal toxicity, and should also have

a once daily dosing regimen, a short duration of administration, bactericidal activity,

and possess a new mechanism of action with maximal activity against the newly

49

emerged multi-drug (MDR-TB) and extensively drug-resistant (XDR-TB) strains of

Mtb (Upton et al., 2015). In addition, new drugs should be orally available and

should not have interactions with existing anti-tuberculous drugs or anti-retrovirals

in cases where there is co-infection with HIV (Leibert et al., 2014).

After decades of drought, more than 50 years after the last TB drug was discovered,

no new anti-TB drugs were approved for development until 2012 (Upton et al.,

2015). However, the recent re-awakening in TB drug development has yielded two

potent drugs in 2012 and 2013 with new mechanisms of action that have been

approved for the treatment of MDR TB. Thus, there has been a recent resurgence of

TB drug discovery (Upton et al., 2015). These new drugs are bedaquiline (Andries et

al., 2005, Palomino and Martin, 2013, Diacon et al., 2012a, Grosset and Ammerman,

2013), a diarylquinoline inhibitor of Mtb ATP synthase; and delamanid, an anti-

tubercular nitroimidazole (Gler et al., 2012).

50

SN Drug Developers Mechanism Stage

1 Peptide deformylase inhibitors

GSK, TB Alliance Inhibits cell growth Discovery

2 Malate synthase inhibitors

GSK, Rockefeller University, Texas

A&M

Inhibits carbon uptake

Discovery

3 Proteasome inhibitors

Cornell University Inhibits cell maintenance

Discovery

4 Diamine SQ-109 Sequella Inhibits cell wall Biosynthesis

Phase 2

5 Diarylquinolines, e.g. Bedaquiline (TMC207)

Johnson & Johnson ATP depletion and pH Imbalance

Approved for MDR-

TB treatment

6 Nitroimidazoles e.g. Delamanid

Otsuka, Chiron, Novartis,

TB Alliance

Inhibits protein synthesis and cell

wall lipid synthesis

Approved for MDR-

TB treatment

7 Fluoroquinolones (Gatifloxacin

& Moxifloxacin)

NIH, WHO, Bayer, TB Alliance and

others

Inhibits DNA replication

and transcription

Phase 3

8 Benzothiazinones (e.g. BTZ043)

Hans Knöll Institut

Inhibits cell wall biosynthesis

Phase 1

Table 1.2: New anti-tubercular drugs in different phases of development. Table

adapted from (Check, 2007).

Peptide Deformylase Inhibitors

Peptide deformylase (PDF) is an important bacterial metalloenzyme needed for the

maturation of bacterial proteins, and which has been highlighted as a promising

target for new generation antibiotics (Sharma et al., 2009). A large class of PDF

inhibitors has been identified in recent years (Sharma et al., 2009). A number of

them have been tested against Mtb as single agents, as well as in combination with

traditional antibiotics used for TB treatment (e.g. isoniazid and rifampicin). These

PDF inhibitors include: actinonin, BB-3497, hydroxylamine hydrochloride, 1,10-

phenanthroline and galardin (Sharma et al., 2009). Results obtained from the

51

relevant study suggested that PDF inhibitors act synergistically with conventional

anti-tubercular drugs (Sharma et al., 2009).

In another study by Pichota and co-workers, the Mtb PDF was validated as a drug

target and the inhibitor LBK-611 and its analogues showed promising results,

indicating that they could serve as a new class of anti-tubercular agents (Pichota et

al., 2008).

Malate Synthase Inhibitors

The glyoxylate shunt is an anaplerotic bypass of the traditional tricarboxylic acid (or

Krebs) cycle which allows the use of carbon from acetyl-coenzyme A (acetyl CoA)

produced by fatty acid metabolism (Bauza et al., 2014). This bypass mechanism is

available in plants, fungi, and prokaryotes, but is lacking in mammals, which

suggests an interesting drug target that can lead to generation of lead compounds

with minimal human toxicity (Myler and Stacy, 2012, Kondrashov et al., 2006, Bauza

et al., 2014). The shunt utilizes two enzymes: isocitrate lyase (ICL), which converts

isocitrate into glyoxylate and succinate; and malate synthase (GlcB), which converts

glyoxylate into malate using one molecule of acetyl CoA (Krieger et al., 2012, Myler

and Stacy, 2012).

The glyoxylate shunt plays a key role in fatty acid metabolism and virulence in Mtb,

which has proved to be important in the survival of pathogenic organisms that are

involved in chronic infections (Krieger et al., 2012, McKinney et al., 2000). Recent

studies by Krieger et al. showed that a Mtb strain with a dysfunctional glyoxylate

52

shunt was unable to establish infection in a mouse model. This led these

researchers to develop a class of compounds, via a structure-guided approach,

known as phenyl-diketo acids (PDKAs) which specifically inhibit malate synthase

(GlcB). The identification of these PDKA derivatives provides an important

validation of GlcB as an attractive drug target in Mtb (McKinney et al., 2000). The

crystal structures of the complexes of GlcB with PDKA inhibitors have been solved

and these guided optimization of the potency of these compounds (Krieger et al.,

2012). The malate synthase inhibitors are still at the discovery stage and work is on-

going to develop these further as TB therapeutics (Bauza et al., 2014).

Figure 1.6: Chemical structure of PDKA. Image drawn in the enol form using ChemDraw (Krieger et al., 2012).

Proteasome inhibitors

Proteasomes are large protein complexes that are involved in the proteolysis of

cytoplasmic proteins that serve in signalling during adaptation (Yang et al., 2013).

Proteasomes are well conserved and found in eukaryotes, archaea and in some

bacteria (Cheng and Pieters, 2010). Even though proteasomes are present in certain

parasites, Mtb is the only bacterial pathogen known thus far to possess

53

proteasomes (Cheng and Pieters, 2010). The Mycobacterial proteasome plays

important key roles in the bacterium, such as degradation of certain proteins,

survival of nitro-oxidative stress, and in bacterial persistence (Pearce et al., 2008,

Burns et al., 2009, Gandotra et al., 2007). However, the proteasomes in Mtb have a

similar organization to its eukaryotic variants, which makes them less attractive

drugs targets (Cheng and Pieters, 2010).

The drive to use proteasome inhibitors as anti-TB drugs has been dampened due to

the extensive degree of conservation of the mycobacterial proteasome with the

form present in humans. Hence the development of highly selective proteasome

inhibitors that will evade inherent toxicity has become difficult (Cheng and Pieters,

2010). Nevertheless, two new proteasome inhibitors which could kill non-

replicating Mtb were identified (Yang et al., 2013). These are the 1,3,4-oxathiazol-2-

one compounds HT1171 and GL5 (Figure 1.7). These two compounds have been

shown to be both potent and selective in their activity against the Mtb proteasome,

with low inhibitory effects on the human proteasome (Lin et al., 2009).

Figure 1.7: Chemical structures of HT1171 and GL5. Redrawn with ChemDraw from (Yang et al., 2013, Lin et al., 2009).

54

Nitroimidazoles

The nitroimidazoles are a new class of class of anti-tubercular drugs under

development with rather promising prospects (Upton et al., 2015). A wide range of

nitroimidazole subclasses were shown to be potent against members of the Mtb

complex (Mukherjee and Boshoff, 2011). The nitroimidazoles, for example

metronidazole (Flagyl®), have proven effective in the treatment of bacterial and

protozoal infections in recent years and part of their activity results from the

formation of reactive chemical species following bio-reduction of the drugs within

the target pathogen (Upton et al., 2015).

Delamanid, a nitroimidazo-oxazole (also known as OPC-67683), has reached phase

III trials for the treatment of multidrug-resistant tuberculosis, while PA-824, a

nitroimidazo-oxazine, has also entered phase III trials for drug-sensitive and drug-

resistant tuberculosis (Upton et al., 2015). WHO documents that delamanid can

now be used for treatment of adults with MDR-TB (WHO, 2014b). However,

information on the effects of this new drug remains incomplete, since it has only

recently passed through Phase IIb trial and studies for safety and efficacy, and been

approved for MDR-TB. The WHO strongly hence recommends the acceleration of

Phase III trials in order to generate a more wholesome evidence base to inform

future policy on delamanid (WHO, 2014b). Delamanid, on bio-reduction within Mtb,

blocks the biosynthesis of mycolic acid, which is a major component of the bacterial

cell wall (Matsumoto et al., 2006, Skripconoka et al., 2013). Recent studies have

documented that it is efficacious against Mtb in vitro and in mice, with potency

both as a single drug and as part of drug multi-therapy for MDR-TB in a 2-month

55

research trial that evaluated bacterial CFUs (colony forming units) in serial sputum

samples (Diacon et al., 2011, Gler et al., 2012).

Research has shown the nitroimdazole drug PA-824 to have activity against both

replicating and hypoxic non-replicating Mtb (Somasundaram et al., 2013, Stover et

al., 2000, Singh et al., 2008). It has also been documented that PA-824 shows potent

bactericidal and sterilising activity against active TB infection in mice (Lenaerts et

al., 2005, Tyagi et al., 2005) and guinea pigs (Lenaerts et al., 2005, Dutta et al., 2013,

Garcia-Contreras et al., 2010), and also enormous early bactericidal activity against

tuberculosis disease in humans (Gler et al., 2012). PA-824 is a pro-drug that

becomes activated by nitro-reduction to one or more active compounds (Dutta and

Karakousis, 2014). Apart from inhibiting keto-mycolic acid and protein synthesis,

PA-824 also destroys Mtb through a new mechanism which involves generation of

intracellular nitric oxide (Singh et al., 2008).

TBA-354, a pyridine-containing biaryl compound, was shown to have exceptional

activity against chronic murine tuberculosis and good bioavailability in preliminary

studies carried out in rodents (Upton et al., 2015). Although TBA-354 has a narrow

spectrum of activity, it has bactericidal activity against both replicating and non-

replicating Mtb in vitro, with near-identical potency to that of delamanid and higher

potency than PA-824 (Upton et al., 2015).

56

Figure 1.8: Chemical structures of nitroimidazole compounds. Figures drawn with ChemDraw (Mukherjee and Boshoff, 2011).

Diarylquinolines

Diarylquinolines, though related to the quinolones, do not inhibit the Mtb DNA

gyrase (Leibert et al., 2014). Rather, they link to the trans-membrane domain of the

adenosine triphosphate (ATP) synthase, which inhibits the mycobacterial conversion

of adenosine diphosphate (ADP) into ATP by interrupting trans-membrane and

central stalk rotation of the proton pump. This mechanism of action for this class of

anti-tuberculous drugs is new and a notable example is bedaquiline (Leibert et al.,

2014, Andries et al., 2005). Bedaquiline is more selective for the mycobacterial ATP

synthase than that from mammals. It was shown to be effective against both non-

57

resistant and MDR strains of Mtb and other mycobacterial species (Andries et al.,

2005, Haagsma et al., 2009).

Bedaquiline is orally bioavailable with a half-life of 43–64 hours in plasma and 28–

92 hours in tissues (including lung and spleen) (Andries et al., 2005). While

bedaquiline is a promising drug and marks a breakthrough in the quest for novel

anti-tuberculous drugs, its position in TB chemotherapy remains unclear pending a

full Phase III trial and the development of other upcoming new TB drugs (Leibert et

al., 2014). Nevertheless, bedaquiline was granted accelerated approval in December

2012 by the United States Food and Drug Administration (WHO, 2013).

Figure 1.9: Chemical structure of Bedaquiline - a diarylquinoline TB drug. Figure drawn with ChemDraw (WHO, 2013).

58

Diamine SQ109

Diamine SQ109 is a 1,2-ethylenediamine compound and is an analogue of

ethambutol (Kwon et al., 2014). It inhibits protein synthesis by targeting the mycolic

acid transporter MmpL3 in Mtb and was shown to be potent against both drug-

susceptible and drug-resistant Mtb (Sacksteder et al., 2012). It also shows activity

against other mycobacteria (M. bovis and M. bovis BCG) and kills Mtb inside

macrophages with similar efficiency to isoniazid, but with superior activity to

ethambutol (Sacksteder et al., 2012, Jia et al., 2005). Diamine SQ109, which is

currently in phase II clinical trials, acts synergistically when combined together with

bedaquiline (Reddy et al., 2010, Reddy et al., 2012, Sacksteder et al., 2012).

Figure 1.10: Chemical structure of SQ109: Figure drawn using ChemDraw (Onajole et al., 2010).

Benzothiazinones (BTZ043) BTZ043, a nitro-aromatic compound, inhibits the synthesis of decaprenylphospho-

arabinose, which is a precursor of the arabinans in the cell wall of Mtb (Kwon et al.,

2014). It is highly efficacious against drug-susceptible TB, MDR-TB and XDR-TB

(Pasca et al., 2010). It was shown to possess additive effects when combined with

59

rifampin, isoniazid, ethambutol, TMC207, PA-824, moxifloxacin, meropenem (with

or without clavulanate) and SQ109; and synergistic effects with bedaquiline

(Lechartier et al., 2012). Studies showed that BTZ043 displays nanomolar

bactericidal activity both in vitro and in ex vivo models of tuberculosis (Makarov et

al., 2014).

Figure 1.11: Chemical structure of BTZ043: Figure drawn using ChemDraw (Makarov et al., 2014).

1.3.4 Anti-Tubercular Drug Resistance: A Cause for Therapeutic Failures

The increased spread of TB worldwide has been propelled by the emergence of Mtb

strains that have become insensitive to the conventional antibiotics used for TB

treatment (McLean and Munro, 2008, Kaur et al., 2014). Many of these drugs have

been in existence for over 50 years and new therapeutic measures are urgently

needed to replace the older, ineffective drugs (McLean and Munro, 2008). The

slow-replicating nature of Mtb and its ability to persist and remain dormant within

the human macrophage for a long period of time necessitates prolonged therapy

60

duration of anti-TB drugs. This is to ensure complete clearance of the bacterium

from the human body and the restoration of the immune system (McLean and

Munro, 2008).

Presently, the standard therapy regimen lasts between 6–12 months and involves

the administration of four anti-tubercular drugs: namely isoniazid, rifampicin,

pyrazinamide and either streptomycin or ethambutol. However, this long duration

of treatment often leads to non-compliance on the part of the patient, which

contributes to the development of resistant strains of the bacterium (Kaur et al.,

2014, McLean and Munro, 2008). Co-infection with HIV has also been documented

to add to the growing problem of resistance and mutations in the Mtb genome

have worsened the problem of treatment failure in humans and have resulted in

various types of drug resistance (McLean and Munro, 2008). These include multi-

drug resistance (MDR), which signifies Mtb strains resistant to at least two of the

front line drugs (isoniazid and rifampicin), single-drug resistance (SDR), and

extensive drug resistance (XDR). XDR Mtb strains are resistant to isoniazid,

rifampicin, any one of the quinolone antibiotics, and to at least one of the second

line anti-TB drugs kanamycin, capreomycin and amikacin (Kaur et al., 2014).

1.4 The Cytochrome P450 Systems

1.4.1 Structure, Function and Mechanism

The cytochromes P450 (CYPs or P450s) are a large, ubiquitous family of enzymes

found in most organisms. They are found in all biological kingdoms, ranging from

61

archaea, bacteria and fungi through to plants and mammals, with over 14,000 P450

genomic sequences discovered so far (Hrycay and Bandiera, 2012, Cederbaum,

2014). The P450s were first identified in the mammalian liver endoplasmic

reticulum as unusual membranous pigments (Munro et al., 2007b). The highest

levels of P450s in mammals are found in the microsomes of the liver, but they also

are present in microsomes from other organs including kidney, small intestine,

lungs, adrenal cortex, skin, brain, testis, placenta and others (Cederbaum, 2014).

Mitochondria from liver and endocrine tissue also contain P450s. Most P450s

consist of about 400–500 amino acids with a size (molecular mass) of about 50 kDa

(Cederbaum, 2014).They are cysteine thiolate-ligated heme b-binding proteins

(Figure 1.12) and have a diagnostic absorption peak at approximately 450 nm

(hence ‘pigment 450’ or ‘P450’) when the reduced heme iron binds to carbon

monoxide (McLean et al., 2012). This 450 nm peak shifts to about 420 nm (P420) on

protonation of the cysteine thiolate to thiol. Protonation of the thiolate ligand (i.e.

P420 formation in the Fe2+–CO complex) results in enzyme inactivation, but is

reversible in some P450s. (Perera et al., 2003, McLean et al., 2010). The electron

donating property of the cysteine thiolate ligand is crucial for P450 catalysis (Munro

et al., 2007b).

62

Figure 1.12: The heme B prosthetic group. The iron (Fe) atom in the centre is shown in red, bonded to four pyrrole nitrogen atoms shown in blue. Figure was drawn with ChemDraw (Mills, 2006)

63

Figure 1.13: Spectral features for cytochrome P450 and its ferrous–carbon monoxide complex. A typical absorption spectrum for a P450 enzyme (CYP142A1 from Mtb, ∼5 µM enzyme) is shown in its oxidised (ferric) state (black solid line, Amax = 418 nm) and in its dithionite-reduced Fe(II)–CO complex (red solid line, Amax = 450 nm and 420 nm). The major absorption (Soret) band shift to ∼450 nm in the CO complex is a characteristic signature of the cysteinate-coordinated heme iron of P450 enzymes. A small shoulder originating from the inactive (thiol-coordinated) P420 form of CYP142 is seen at ∼420 nm in the spectrum of the Fe(II)–CO complex.

The cytochrome P450 enzymes are involved in the catalysis of different types of

reactions mediated through redox chemistry via their heme prosthetic group (an

iron–porphyrin complex). Among the most common reactions are hydroxylations

and other oxidations of organic substrates (Ortiz de Montellano, 2005). A notable

example is the sequential oxidations of the cholesterol side-chain at the C27

position by the P450s CYP125A1 and CYP142A1 in Mtb, forming first the terminal

alcohol moiety, then the aldehyde, and finally the acid (Garcia-Fernandez et al.,

2013).

64

The most common P450 reaction is mono-oxygenation, in which one oxygen atom

of molecular oxygen is inserted into an organic substrate (which may be compounds

central to metabolic processes, or exogenous/xenobiotic molecules), while the

second oxygen atom is reduced to water. This reaction is supported by two

electrons supplied to the P450 by NAD(P)H, with electron transfer mediated by

redox partner flavoproteins and iron-sulfur proteins (Hrycay and Bandiera, 2012,

Ouellet et al., 2010b). The general mono-oxygenation reaction of P450 enzymes can

be represented by the following equation:

RH + O2 + 2H+ + 2e- → ROH + H2O

This P450 reaction has also been referred to as a mixed function oxidase reaction –

reflecting that both the electron donor and the substrate become oxidised in a

typical P450 reaction (Cederbaum, 2014). RH is the organic substrate and ROH is the

oxidised product. The reducing equivalents (2e-) are supplied either by NADH or

NADPH and delivered by redox partner(s) (Hrycay and Bandiera, 2012). The first of

the two electrons (delivered at distinct points in the catalytic cycle) is used to

reduce ferric P450 heme iron into its ferrous state, ready to bind a dioxygen

molecule (McLean et al., 2005). Studies with P450cam (a camphor hydroxylase) and

P450 BM3 (a fatty acid hydroxylase) revealed that substrate binding, which serves

as a control mechanism, is required for efficient electron transfer to the heme iron

(Daff et al., 1997, McLean et al., 2005). A second electron transfer reaction to the

65

ferrous-oxy P450 heme precedes protonation events that produce a reactive iron-

oxo species that catalyses substrate oxidation.

Reactions including reduction, epoxidation, desaturation, ester cleavage, ring

expansion and dehydration are also typical of P450 enzymes. P450s in plants play

key roles in the synthesis of lignins and alkaloids (Cederbaum, 2014). Human P450s

are involved in reactions such as xenobiotic (e.g. drug) metabolism and in the

synthesis of essential endogenous compounds (e.g. steroids) (Balding et al., 2008,

McLean et al., 2012). A notable illustration is the P450 enzyme CYP3A4, a highly

expressed P450 in humans which is involved in the metabolism of about half of all

drugs that enter the body (Balding et al., 2008). On the other hand, prokaryotic

P450s play key roles in pathways for utilization of exogenous compounds (e.g.

cholesterol) as carbon sources, production of antibiotics, and also make

contributions to pathogen biochemistry (Balding et al., 2008). The eukaryotic

cytochrome P450 enzymes are membrane-bound enzymes, linked to their

respective membranes by an N-terminal hydrophobic anchor region and interacting

with membranous redox partner enzymes. These enzymes supply electrons needed

to reduce the heme-bound dioxygen to form a reactive iron-oxo species used to

convert substrate to mono-oxygenated product, and to reduce the second oxygen

atom to water (McLean et al., 2005). However, their prokaryotic relatives are

cytosolic, soluble enzymes that lack a membrane “anchor” region, and interact with

soluble redox partner proteins. The prokaryotic P450s have numerous functions

that are different from those of the eukaryotic P450 enzymes (McLean et al., 2010).

66

A typical bacterial P450 redox system (known as a class I system) involves a

NAD(P)H-specific FAD-containing reductase and an iron-sulfur protein (or

ferredoxin) that shuttles electrons from the reductase to the P450, and the P450

itself as the terminal oxidase (McLean et al., 2007b). Early bacterial P450s studied

(notably P450cam) were shown to be involved in oxidation reactions, such as the

utilization of camphor as a carbon source for growth in Pseudomonas putida

(Poulos, 2003). Characterization of a wider range of microbial P450s has highlighted

several other important physiological roles, including oxidation of polyketides in

biosynthetic pathways; specific examples including hydroxylation reactions in the

synthesis of erythromycin, and epoxidations in the synthesis of the anticancer

agents epothilones C and D in different microbes (Ogura et al., 2004, McLean et al.,

2010).

P450s generally share low amino acid sequence identity, with P450s sharing ≥40%

identity classified in the same P450 “family” (within the cytochrome P450

“superfamily”) and usually exhibiting similar substrate specificity. Despite wide

variations in amino acid sequence identity, The P450s exhibit strong similarities in

their overall structure, with a number of highly conserved secondary structural

elements. The regions which vary most in the P450 structures are those associated

with diverse ligand binding and catalytic reactions. Variations in P450s structure are

also important to enable distinct redox partner interactions and to allow P450

structural flexibility (Munro et al., 2007b, McLean et al., 2012). The majority of

P450s have been shown, through structural studies, to possess relatively

67

hydrophobic active site cavities for substrate binding – which reflects the nature of

the substrates themselves (Kim et al., 2005, Munro et al., 2007b).

Figure 1.14: Typical topology of a cytochrome P450: A representation of the high resolution crystal structure (1.63 Å) of the P. putida camphor hydroxylase P450cam (CYP101A1) is shown (PDB code 2CPP). P450 enzymes are primarily comprised of α-helices (red cylinders) with heme (purple sticks) in the centre. The β-sheets are shown as yellow arrows and the loops are represented as green strings. Significant structural elements include the long I helix, which runs above the distal face of the heme and consists of several amino acid residues important for catalysis. The α-helices are well conserved in the P450s.

1.4.2 The P450 Catalytic Cycle

The P450s contain ferric heme iron in their resting state, and usually undergo shifts

in the spin-state equilibrium of the heme iron from predominantly low-spin (S =

68

1/2) to partially or predominantly high-spin (S = 5/2) form on substrate binding

(McLean et al., 2007a).

Figure 1.15: Schematic representation of the d-orbital electron configurations for low- and high-spin ferric heme iron. In the low-spin state (resting state), the electronic configuration has all the 5 electrons in the lower energy orbitals (t2g) with total spin = ½. The high-spin configuration shows 3 electrons occupying the lower energy orbitals and 2 electrons occupying the higher energy (eg) orbitals with total spin = 5/2. ΔOct represents the energy difference between the two energy levels. Substrate binding usually causes a low- to high-spin shift of the ferric heme iron.

The binding of substrates to P450 enzymes often results in an elevation of the heme

iron reduction potential by about 130–140 mV, i.e. the ferric heme iron potential

becomes more positive. This reaction favours first electron transfer to the ferric

heme iron from the redox partner, reducing it to the ferrous form (McLean et al.,

2005, Ouellet et al., 2010b, Munro et al., 2002). The second electron is transferred

to the heme iron after molecular oxygen binds to the ferrous iron, producing a

ferric-peroxy species. This intermediate product is protonated (to ferric

hydroperoxy) and a subsequent protonation leads to the cleavage of the O-O bond

and to water molecule formation with the formation of a highly reactive oxyferryl

radical cation species (compound I), which is known to be a powerful oxidant in

P450 catalysis (Munro et al., 2007b, McLean et al., 2005, Ouellet et al., 2010b).

Attack of this highly reactive species on the bound substrate leads to its mono-

69

oxygenation and to the formation of an oxidised product (Ortiz de Montellano,

2005, Anderson and Chapman, 2005, McLean et al., 2005).

Research has also revealed that P450 mono-oxygenase reactions can be driven by

H2O2 or organic peroxides (known as the ‘peroxide shunt’ mechanism), thus

bypassing the role of NAD(P)H, molecular oxygen and redox partner(s) (McLean et

al., 2005). This H2O2 or organic peroxide-driven process can result in the direct

conversion of a ferric P450 to the ferric-hydroperoxo (Compound 0) intermediate

(Fe3+-OOH). Further protonation of this intermediate leads to loss of a water

molecule and to the generation of the reactive ferryl-oxo Compound I species

(FeIV=O porphyrin radical cation) that oxidises the substrate (Ortiz de Montellano,

2005, Denisov et al., 2005, Guengerich and Munro, 2013). The Bacillus subtilis

P450BS, for example, and its homologue P450SP from Sphingomonas

paucimobilis utilise H2O2 as an oxidant to catalyse the - and -hydroxylation of

fatty acids. The crystal structure of the B. subtilis enzyme (CYP152A1) has been

determined, providing important insights into the mechanism of the reaction and

highlighting the major residues (Arg242, Leu17, Leu70, Val74, Leu78, Phe79, Val170,

Phe173, Ala246, Phe289 and Phe292) responsible for binding the fatty acid substrate

and for catalysis (Lee et al., 2003).

70

Figure 1.16: A schematic representation of the P450 compound I (oxyferryl radical cation species). The general structure of the P450 compound I shows the four nitrogen ligands of the porphyrin macrocycle, the ferryl (FeIV) state of the heme iron, the radical cation of the porphyrin, and the cysteine thiolate ligand (Hrycay and Bandiera, 2012).

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Figure 1.17: Schematic representation of the catalytic cycle of a cytochrome P450 enzyme. The catalytic metabolism of organic substrates by P450s involves a series of oxidation, reduction and protonation steps. First, the substrate (RH) binds close to the ferric heme iron of the P450 enzyme (displacing a water ligand), and this generates a more positive heme iron potential (in the high-spin form) which favours electron transfer from the redox partner to reduce the heme iron to the ferrous state. The binding of dioxygen to the ferrous heme iron leads to the formation of ferric superoxy species which is reduced and protonated to form a ferric hydroperoxy species (compound 0). The further protonation and subsequent dehydration of the compound 0 species forms a ferryl-oxo intermediate which is known to be a highly reactive oxidant (compound I), and this species catalyses oxygenation (often hydroxylation) of the substrate. The dissociation of the hydroxylated product (R–OH) from the enzyme restores the heme iron to the initial, resting ferric low-spin state. The oxy intermediates in the cycle can collapse via non-productive pathways to produce either superoxide (from the ferric superoxy form), hydrogen peroxide (from compound 0) or water (from compound I). The formation of compound 0 can be productively driven by mixing H2O2 (or organic peroxides) with ferric, substrate-bound P450. This reaction is often not very efficient. The non-productive pathways may occur if electron delivery from redox partners or if protonation events are not timely, or if substrate is improperly positioned for oxidative attack, or is completely absent. Image is adapted from (Munro et al., 2007b).

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1.4.3. Cytochrome P450 Redox Partners

Most P450s interact with one or more redox partners to acquire their reducing

equivalents (electrons). Important studies on P450 systems, particularly those from

bacterial species, have revealed an extensive diversity in the machinery used to

deliver electrons to the P450s (McLean et al., 2005).

The progression of the P450 reaction cycle depicted in Figure 1.17 requires

consecutive delivery of two electrons to its heme iron at distinct points in its

catalytic cycle (Munro et al., 2007b, Ortiz de Montellano, 2005). These electrons are

provided by the nicotinamide adenine dinucleotide cofactors (NAD(P)H) and their

delivery to the heme iron is mediated by accessory redox partner proteins. Two

primary types of redox systems are involved in shuttling electrons to the heme iron

(Ortiz de Montellano, 2005). The first category, often called class I, employs

ferredoxins (iron–sulfur cluster containing enzymes) or sometimes flavodoxins

(FMN-containing proteins) in conjunction with a separate reductase protein that

uses flavin adenine dinucleotide (FAD) as its cofactor and accepts electrons from

NAD(P)H (Ouellet et al., 2010b). The class I systems are employed by organisms with

soluble P450 enzymes, such as bacteria, but they are also found in higher

organisms, where their reductase and P450 components are associated with the

mitochondrial membrane and involved in reactions associated with steroid

synthesis (Ouellet et al., 2010b, Munro et al., 2007b). Typical prokaryotic class I

redox systems use ferredoxin reductase and ferredoxin (Fdx) partners proteins

(Munro et al., 2007b, Guengerich and Munro, 2013).

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The second major type of redox system, often called class II (eukaryotic), is

exclusively membrane bound (i.e. P450 and redox partner are bound by an N-

terminal membrane anchor). The reductase (cytochrome P450 reductase or CPR)

possesses two different flavin-binding domains within a single polypeptide (McLean

et al., 2005). The mammalian CPRs are typical of the class II reductase system and

these diflavin reductases, like the class I enzymes, contain a FAD-binding domain

that is structurally similar to the microbial ferredoxin reductases. However, in class

II systems these modules are fused to an FMN-binding (flavodoxin-like) domain,

forming a single CPR redox partner protein (Ouellet et al., 2010b). The CPRs have a

N-terminal membrane anchor on their FMN-binding domain. There are also

examples of P450 enzymes, such as CYP102A1 (P450 BM3) from Bacillus

megaterium, that contain a CPR-like domain fused to the P450 domain (Munro et

al., 2007b, McLean et al., 2005, Ortiz de Montellano, 2005). However, in this case

the P450 and CPR components lack membrane anchors and BM3 is a soluble

enzyme. The systems supporting the Mtb P450s will be representatives of the Class

I system, since there is no CPR encoded in the Mtb genome (Munro et al., 2007b).

Bacterial flavodoxins and ferredoxins are important redox proteins that play key

roles in P450 metabolic pathways (McLean et al., 2005). Ferredoxins are functional

one-electron carriers which bind iron–sulfur cofactors with 2Fe-2S, 3Fe-4S or 4Fe-4S

clusters. The iron ions in the ferredoxins co-ordinate to sulfur atoms from cysteine

residues in the protein and from inorganic sulfide (McLean et al., 2005). Flavodoxins

can carry either one or two electrons and bind an FMN cofactor non-covalently.

74

However, they are frequently isolated in their single-electron reduced semiquinone

form and typically act as single electron donors/acceptors, mainly by electron

transfer from their 2-electron reduced hydroquinone form (McLean et al., 2005).

Studies have revealed that flavodoxins act as surrogates for ferredoxins under

conditions of cellular iron limitation, and hence they are believed to support

functions of selected bacterial P450s (McLean et al., 2005). The catalytic function of

bovine P450c17 (CYP17) was shown to be supported by the E. coli flavodoxin

reductase/flavodoxin system (Jenkins et al., 1997). In a physiologically relevant

example, the cineole metabolizing P450cin (CYP176A1) from Citrobacter braakii was

shown to interact with the host flavodoxin cindoxin, the product of a gene located

adjacent to the P450 on the bacterial chromosome (Hawkes et al., 2002).

The P450cam system utilizes a 2Fe-2S cluster ferredoxin (putidaredoxin) to reduce

the P450, but studies have revealed other microbial P450s that also exploit 3Fe-4S

ferredoxins and probably 4Fe-4S ferredoxins for the electron relay mechanism

(McLean et al., 2005). Supporting examples include the xenobiotic-transforming

P450soy from Streptomyces griseus which uses a 3Fe-4S ferredoxin (Trower et al.,

1992), and a 4Fe-4S ferredoxin shown to support catalytic activity in the fatty acid

oxidising P450 BioI system (Lawson et al., 2004).

It has been documented that flavodoxins are not present in the Mtb genome, but

ferredoxins and ferredoxin-like proteins do exist in the genome. Notable examples

of Mtb P450 redox partners are the ferredoxin products of the Rv1786 gene

(adjacent to the gene for the P450 CYP143A1) and the Rv0764c gene (adjacent to

75

the gene encoding CYP51B1) (McLean et al., 2005). CYP51B1’s 3Fe-4S ferredoxin

partner (Fdx) has been characterised and shown to support the catalytic properties

of this enzyme (Bellamine et al., 1999, McLean et al., 2005).

Research in the last decade has identified several unique types of P450 redox

systems that do not fall into the typical Class I/II categories, many of which have

been discovered via characterization of P450 enzyme systems identified from

genome sequencing studies (Guengerich and Munro, 2013). Some selected

examples are depicted in Figure 1.18.

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Figure 1.18: Schematic representation of a variety of P450 redox systems and P450 fusion proteins. Selected P450 enzymes and their redox partner systems are shown. 1) Fatty acid hydroxylase. For example, P450–CPR fusions of the BM3 type (Munro et al., 2007a). 2) glycosyltransferases (of the GTB-type superfamily) fused to P450s (Breton et al., 2006). 3) Short-chain reductases (SDR) fused to P450s. These enzymes catalyze a wide range of activities including the metabolism of steroids, cofactors, carbohydrates, lipids, aromatic compounds, and amino acids, and act in redox sensing (Kallberg et al., 2010). 4) Esterases fused to P450s: These are esterases and lipases which includes fungal lipases, cholinesterases etc. (Hang et al., 2012). 5) P450s fused to medium chain reductases (MDR) (Persson et al., 2008). 6) P450s fused to Dolichyl-phosphate-mannose-protein mannosyltransferases (PMT_2) (Haft et al., 2012). 7) An ammonium transporter fused to a P450 found in Saccharomyces cerevisiae (Marini et al., 1994). 8) P450s fused to membrane bound O-acyl transferase (MBOAT). A conserved histidine is suggested to be an important active site residue in such enzymes (Hofmann, 2000). 9) P450s fused to a histidine phosphatase (HP) domain, as found in phosphatase proteins that containing a His residue which is phosphorylated during the course of their reaction (Rigden, 2008). (10) SPFH-like superfamily protein fused to P450s, where SPFH indicates stomatin, prohibitin, flotillin and HflK/C type proteins that are often associated with lipid rafts (Tanaka and Tsukihara, 2012).

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1.5 The Mycobacterium tuberculosis Cytochrome P450 Enzymes

1.5.1 Discovery of Mtb P450s and the Quest for their

Physiological Roles Genomic sequencing of Mtb by Cole and his group in 1998 revealed a large number

(20) of cytochrome P450 enzymes, which is unusual for a bacterium of that genomic

size (Cole et al., 1998). This number of P450 enzymes in a 4.4 Mb Mtb H37Rv

genome reveals an ~200-fold greater CYP gene “density” than in the human

genome (3 Gb) which contains 57 CYPs (McLean et al., 2006a, Brueggemeier et al.,

2005, Ouellet et al., 2010b, McLean et al., 2010). Unusually, another bacterium of

similar genomic size to Mtb, E. coli, has no P450s in its genome, which makes this

unusual occurrence even more interesting (McLean et al., 2006a, Hudson et al.,

2012a). Nevertheless, this large number of P450s in Mtb indicates strongly that they

have important physiological functions, which recent studies are beginning to

unravel (McLean et al., 2006a). The Mtb P450s exhibit low evolutionary

relationships to one another with no define groupings (McLean et al., 2006a,

Hudson et al., 2012a) (Figure 1.19).

In recent years, characterization of the Mtb P450 enzymes has revealed novel and

exciting functions, and has led to the recognition of their potential for exploitation

as novel therapeutic targets (McLean et al., 2010). Table 1.3 summarizes the gene

essentiality of the twenty (20) Mtb P450 enzymes along with data indicating their

level of biochemical and structural characterization.

78

Figure 1.19: Evolutionary analysis of Mtb P450s. The Mtb P450s show limited evolutionary relationships to one another, with no distinct groupings, as demonstrated by the evolutionary tree shown. The Mtb P450s are in red text while other selected members of the P450 superfamily are in black. MonD and NanP play key roles in synthesis of the polyether ionophore antibiotics monensin (in Streptomyces cinnamonensis) and nanchangmycin (in Streptomyces nanchangensis). The P450 isoforms that currently present the most attractive anti-tubercular drug targets are indicated and these include the cholesterol oxidase enzymes. Adapted from (McLean et al., 2006a, Hudson et al., 2012a).

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S/N Mycobacterium tuberculosis P450 enzymes

Essentiality in Mtb

Level of Characterization

Crystal structures

References

1 CYP51B1 (Rv0764c) Non-essential gene. Involved in host steroid inactivation?

Expressed and purified; binds avidly to azole drugs; unstable Fe-CO complex

Ligand-free and fluconazole complexes solved

Mclean et al., 2007a, Bellamine et al., 1999, Podust et al., 2001.

2 CYP121A1 (Rv2276) Essential for bacterial viability.

Expressed and purified; cyclodipeptide (cYY) oxidase

Ligand-free, fluconazole and cyclodityrosine complexes solved

Mclean et al., 2002a, 2007a, 2010, Leys et al.,2003.

3 CYP123A1 (Rv0766c) Function unknown. Upregulated at high temperatures.

Partially characterized

No crystal structure

Stewart et al., 2002, Mclean et al., 2007a.

4 CYP124A1 (Rv2266) Important in viability and infectivity. Expressed in dormancy.

Expressed and purified; binds cholesterol/ cholestenone and methyl branched lipids

Ligand-free, cholestenone and phytanic acid-bound structures solved

Johnston et a., 2010, Mougous et al., 2006, Mclean et al., 2010, Sasseti et al., 2003.

5 CYP125A1 (Rv3545c) Important in viability and infectivity. Expressed in dormancy.

Expressed and purified. Oxidises cholesterol and cholestenone

Ligand-free, cholestenone, econazole, and androstenedione structures solved

Capyk et al., 2009, Mclean et al., 2010,Chang et al.,2009.

6 CYP126A1 (Rv0778) Possible operon with enzymes in purine synthesis.

Expressed and purified

Ligand-freea and ketoconazole complexes solved

Unpublisheda.

7 CYP128A1 (Rv2268c) Required for optimal growth of Mtb

Not expressed, but expected to be a menaquinone hydroxylase


Sassetti et al., 2003, Mclean et al., 2006a.

8 CYP130A1 (Rv1256c) Non-essential gene. Absent in M. bovis BCG vaccine strain.

Expressed and purified, binds inhibitors but substrate unknown

Ligand-free, econazole and heterocyclic arylamine bound structures

Ouellet et al., 2008, Podust et al., 2009.

9 CYP132A1 (Rv1394c) Role in Mtb virulence?



Recchi et al., 2003, Mclean et al., 2007a.

10 CYP135A1 Non-essential Not No crystal Sassetti et al.,

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gene. characterized structure 2003, Mclean etal., 2010.

11 CYP135B1 (Rv0568) Non-essential gene. Expressed in dormancy.

Not characterized


Sassetti et al., 2003, Mclean etal., 2010.

12 CYP136A1 (Rv3059) Non-essential gene. Expressed in dormancy.

Not characterized


Sassetti et al., 2003, Mclean etal., 2010.

13 CYP137A1 (Rv3685c) Non-essential gene.

Not characterized


Sassetti et al., 2003.

14 CYP138A1 (Rv0136c) Virulence role. Upregulated at high temperatures.



Stewart et al., 2002, Mclean et al., 2007a.

15 CYP139A1 (Rv1666c) Non-essential gene.



Mclean et al., 2010, Sasseti et al., 2003.

16 CYP140A1 (Rv1880c) Non-essential gene. Expressed in dormancy.

Not characterized


Sassetti et al., 2003.

17 CYP141A1 (Rv3121) Function unknown. Absent in M. bovis BCG vaccine strain.



Mclean et al., 2007a.

18 CYP142A1 (Rv3518c) Role in viability and infectivity. Expressed in dormancy.

Expressed and purified. Oxidises cholesterol and cholestenone

Ligand-free, cholestenone and econazole-bound structures solved

Driscoll et al., 2010. Mclean et al., 2010, Sasseti et al., 2003.

19 CYP143A1(Rv1785c) Non-essential gene.


Ligand-free structure solvedb

Sassetti et al., 2003, Unpublishedb.

20 CYP144A1 (Rv1777) Possible role in virulence?


Ligand-free structure solvedc

Tailleux et al., 2008, Driscoll et al., 2011, Unpublishedc.

Table 1.3: The twenty (20) P450 enzymes in Mycobacterium tuberculosis (Adapted from Mclean et al., 2010). aMclean et al.; bSwami et al.; cDriscoll et al. (unpublished data).

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1.5.2 The Cholesterol Oxidase P450 Enzymes

With the growing line of evidence that Mtb lacks the machinery to synthesize

sterols, paradoxically, the gene for CYP51B1 was found to be encoded in the Mtb

genome. CYP51B1 is a P450 enzyme that can catalyze the 14α-demethylation of

lanosterol, which is an important step in cholesterol biosynthesis in eukaryotes to

give the 8,14-diene product (Bellamine et al., 1999). CYP51 enzymes, though well

conserved across the actinobacteria, are apparently non-essential for in vitro cell

viability of these organisms, unlike the eukaryotic CYP51s. The physiological role of

the CYP51B1 enzyme in Mtb and other bacteria remains unknown (Lamb et al.,

2002).

Interestingly, a study carried out by Van der Geize et al. identified a cluster of genes

that are involved in steroid metabolism in Rhodococcus jostii (Van der Geize et al.,

2007). This organism is a soil bacterium that metabolizes a large range of organic

compounds (mainly hydrophobic). It is also a mycolic acid-producing bacterium

within the order Actinomycetales, which includes the mycobacteria (Gurtler et al.,

2004, Van der Geize et al., 2007). Furthermore, genomic sequencing revealed that

approximately 60% of the 3,999 genes of Mtb H37Rv are conserved in R. jostii

RHA1, and these include many genes with unknown functions (McLeod et al., 2006).

Hence these data suggests that rhodococci could form part of a useful model for

many mycobacterial processes (Van der Geize et al., 2007).

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The large steroid metabolism-associated gene cluster identified in R. jostii was

initially discovered by bioinformatics analyses of genes that were expressed during

growth on cholesterol, and this cluster was also shown to encode the Mtb P450

enzymes CYP142A1 and CYP125A1 (Van der Geize et al., 2007, Driscoll et al., 2010)

(Figure 1.20). Organic compounds such as cholesterol occur widely in plants (albeit

in small amounts), animals and some micro-organisms, and potentially comprise an

essential source of energy for bacteria and fungi that live and feed on dead organic

matter, especially actinomycetes that utilize hydrophobic substrates (Van der Geize

et al., 2007). The sterol genes identified in R. jostii and another Rhodococcus sp.

were shown to be involved in sterol uptake and in sterol side-chain and ring

degradation, and are conserved in related pathogenic actinobacteria, such as Mtb,

M. bovis and M. avium. These pathogens appear to have retained the capacity for

cholesterol metabolism, which they use to enable infectivity and survival in their

hosts (Van der Geize et al., 2007).

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Figure 1.20: Genetic organization of cholesterol metabolising gene clusters in Rhodococcus sp. RHA1 and Mtb: A comparison. The image above shows genes from Rhodococcus sp. RHA1 and Mtb which are colour-coded according to designated functions: cholesterol uptake: purple-colour; side-chain degradation: red-colour; cleavage of cholesterol rings A and B: blue colour; degradation of the propionate moiety of 9,17-dioxo-1,2,3,4,10,19-hexanorandrostan-5-oic acid (DOHNAA: an intermediate in the breakdown pathway of cholesterol rings A and B): orange colour; degradation of rings C and D: green colour. White arrows represent genes for which no reciprocal homologue is present. Image adapted from (Van der Geize et al., 2007).

Research has highlighted the importance of cholesterol in Mtb in latent and chronic

infection. Cholesterol serves as a major source of carbon while the pathogen is

engulfed in the human macrophages of the lungs and it also enhances bacterial

entry into the macrophage. This subsequently enhances infectivity and persistence

of the bacterium within the host (Chang et al., 2009, McLean et al., 2012, Johnston

et al., 2012).

Two major P450 enzymes, CYP125A1 and CYP142A1, were demonstrated to play

key roles in host cholesterol catabolism by Mtb. Another P450 gene, CYP124A1, is

located next to a sulfotransferase (Sft3, Rv2267c) on the Mtb H37Rv genome, which

84

in turn is adjacent to the Rv2268 gene encoding the P450 CYP128. CYP128 is

thought to hydroxylate menaquinone, which is then sulfated by Stf3 (Holsclaw et

al., 2008).

CYP124A1 was also identified to metabolise cholesterol, although expression of

CYP124A1 is not induced by cholesterol (McLean et al., 2012, Johnston et al., 2009,

Ouellet et al., 2010b). These three P450 isoforms were shown to catalyse the C-27

(or omega)-hydroxylation of the hydrocarbon side chain of cholesterol to produce

27-hydroxycholesterol, and then to further oxidise the substrate through to the

carboxylic acid via an aldehyde (Figure 1.22) (Johnston et al., 2012, McLean et al.,

2012). This important reaction subsequently enables the β-oxidation of the

cholesterol side chain (McLean et al., 2012). The acid moiety generated from the

reaction then undergoes trans-esterification with coenzyme-A (CoA) in an ATP-

dependent step that prepares the acyl-CoA for β-oxidation. Subsequently, three

rounds of β-oxidation produce one molecule of acetyl-CoA and two propionyl-CoA

equivalents (Johnston et al., 2012, Driscoll et al., 2010).

The major enzyme implicated in the cholesterol side chain degradation is

CYP125A1, while CYP142A1 plays a complementary role with CYP125A1 in certain

mycobacterial strains (Thomas et al., 2011, McLean et al., 2012). This cholesterol

catabolic pathway is conserved in the soil-dwelling relative of Mtb, M. smegmatis,

in which both CYP125A3 and CYP142A2 serve as paralogous enzymes (Frank et al.,

2014, Garcia-Fernandez et al., 2013). These two enzymes share approximately 70%

sequence identity with their Mtb orthologues and activity towards the substrate 4-

85

cholesten-3-one (Frank et al., 2014). Although some mycobacterial variants, such as

the CDC1551 strain of Mtb and M. bovis BCG, have lost a functional CYP142A1 gene,

they still retain the CYP125A1 gene (Klansek et al., 1995, Ouellet et al., 2010a, Capyk

et al., 2009, Frank et al., 2014). Recent studies have revealed that CYP142A1, but

not CYP124A1, fully restores the growth phenotype for a CYP125A1 gene deletion

strain of Mtb (Brzostek et al., 2007). Recent studies in the Mtb vaccine strain M.

bovis BCG revealed CYP125A1 to be up-regulated 7.1-fold during growth in the

presence of cholesterol (McLean et al., 2010). A ∆CYP125 M. bovis BCG strain

showed no growth on cholesterol and accumulated 4-cholesten-3-one during

growth in the presence of cholesterol (McLean et al., 2010).

In Mtb, CYP125A1 is found in a gene cluster known as the intracellular growth

operon (igr), which is essential for the survival of the bacterium in macrophages

(McLean et al., 2012). Even though CYP124A1 and CYP125A1 are closely related and

show remarkable sequence identity (40.7%) over 428 residues, CYP124A1 also

metabolises a wide range of branched chain fatty acids and isoprenoids, while

CYP125A1 has no such activity (Johnston et al., 2009, Johnston et al., 2010, Driscoll

et al., 2010). CYP142A1 is far less related to CYP125A1 (with a sequence identity of

28.0% identity over 397 residues) than is CYP124A1. The comparable sequence

identity between CYP142A1 and CYP124A1 is 35.5% over 392 residues (Driscoll et

al., 2010).

The key steps required for cholesterol utilization by Mtb can be categorised into

four major phases: (1) cholesterol uptake into the Mtb cell, (2) its oxidation to

86

cholest-4-en-3-one, (3) degradation of its side chain and (4) breakdown of the

steroid ring (Ouellet et al., 2011, Johnston et al., 2012). The uptake of cholesterol

into the Mtb cell is carried out by the MCE (mammalian cell entry) proteins and

then cholesterol is oxidized to cholest-4-en-3-one (cholestenone) either by a

cholesterol oxidase (ChoD) or by the 3β-hydroxysteroid dehydrogenase (3β-HSD)

(Johnston et al., 2012, Brzostek et al., 2007, Ouellet et al., 2010a, Ouellet et al.,

2011). The important P450-mediated reaction of cholesterol oxidation is required

for the activation of the side-chain for entry into β-oxidation, and subsequent

steroid ring degradation (Johnston et al., 2012). However, it is not very clear

whether there is a strict order of the cholesterol catabolic pathway in Mtb (Ouellet

et al., 2011). This has been supported by evidence from the literature on

rhodococcal sterol catabolism, which postulated that intermediates of ring and side

chain degradation can be intertwined between the two pathway routes (Rosloniec

et al., 2009). Studies have also shown that in Mtb the blockage of the cholesterol

side chain degradation resulted in the accumulation of cholest-4-en-3-one as a key

metabolite which is bacteriostatic or toxic to the bacterium (Ouellet et al., 2010a).

This suggests that the ring-degrading enzymes (e.g. KsaAB and HsaA-C) act more

efficiently after the side chain has been cleaved off (Ouellet et al., 2011).

The chemical structures of cholesterol and cholest-4-en-3- one are shown in Figure

1.21. The important functions of these cholesterol-hydroxylating P450s for Mtb

infection, persistence and survival makes them promising drug targets and a

potential focus for anti-TB drug development (McLean et al., 2012, Johnston et al.,

2012).

87

Figure 1.21: The chemical structures of cholesterol (A) and cholest-4-en-3-one (B). Figures were drawn using ChemDraw (Mills, 2006).

Figure 1.22: Cholesterol side chain oxidation reactions. The figure shows a schematic illustration of the CYP125/CYP142/CYP124-mediated hydroxylation of cholesterol at the C-27 position to 27-hydroxycholesterol, and the subsequent conversion into cholestenoic acid via an intermediate with an aldehyde moiety (McLean et al., 2012). Image was drawn using ChemDraw (Mills, 2006).

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1.5.2.1 CYP125A1 (Rv3545c): Essential for Mtb Viability and Infectivity

CYP125A1 has been documented to be the most important P450 drug target in Mtb

(McLean et al., 2010). It is the major P450 enzyme that catalyses the 27-hydroxyl-

ation of the cholesterol and cholestenone side chain, enabling cholesterol/one

degradation for energy generation (Ouellet et al., 2010a, Capyk et al., 2009).

CYP125A1 initially sparked an interest as an Mtb drug target when a gene cluster

was identified in the Rhodococcus sp. RHA1 Strain by Van der Geize and co-workers

in 2007 (Van der Geize et al., 2007). This gene operon was shown to be involved in

cholesterol catabolism, with several of these genes conserved in Mtb (including

CYP125A1 and CYP142A1 in Mtb) (Van der Geize et al., 2007, McLean and Munro,

2008). In Mtb, CYP125A1 is located in the igr operon, which is essential for Mtb

survival in human macrophages (Chang et al., 2009, Garcia-Fernandez et al., 2013).

The crystal structure of the ligand-free CYP125A1 enzyme has been solved at a

resolution of 1.4 Å, revealing a ‘letterbox’-like hydrophobic active site entry cavity

which narrows in a funnel-like manner towards the heme centre (McLean et al.,

2009). In addition, the structures of CYP125A1 in complex with androstenedione,

econazole and cholestenone have also been determined to resolutions of 2.0, 2.2

and 1.58 Å, respectively (McLean et al., 2009). For the econazole and

androstenedione structures, the molecules bound within the letterbox cavity, with

neither of the two compounds able to penetrate the funnel-shaped access tunnel to

the heme (the closest distances to the heme iron were at 12.9 Å and 9.3 Å for

89

androstenedione and econazole, respectively) (McLean et al., 2009).

Androstenedione lacks the aliphatic side chain found in cholesterol and hence

exhibited a binding mode that is not consistent with P450 oxidation (McLean et al.,

2009). This as a result of the narrow nature of the active site, which clearly prevents

the steroid molecule from reaching the heme iron directly (McLean et al., 2009).

However, the androstenedione steroid ring nucleus does occupy the space that

would naturally be occupied by the cholesterol steroid nucleus. Minimal

conformational changes in the P450 were observed upon the binding of

androstenedione and econazole ligands to CYP125A1 (McLean et al., 2009).

The crystal structures of CYP125A1 in complex with sterols reveal substantial

lipophilic interactions with the steroid ring nucleus which help to fix the substrate’s

binding mode and to position the aliphatic side-chain at the correct distance from

the heme iron for its C27-oxidation (Johnston et al., 2012). Cholest-4-en-3-one

(cholestenone) binding to CYP125A1 resulted in conformational changes which are

largely due to repositioning of the N-terminal portion of the I-helix and the H-helix

to envelop the substrate in the active site (Ouellet et al., 2010a). Furthermore, the

N-terminal portion of the I-helix bends to make hydrophobic contacts with the

cholestenone molecule. These conformational changes produced a r.m.s.d of 0.78 Å

between the substrate-free and substrate-bound structures (Ouellet et al., 2010a).

90

Figure 1.23: Structural features of Mtb CYP125A1 in complex with diverse substrates and inhibitor molecules. A: solvent-accessible surface of CYP125A1 (PDB code 2XN8). The colour coding shows the helices in cyan, the sheets in magenta and the loops in pink. The arrow indicates the narrow access site entry for CYP125A1 which can be readily identified by the direct view onto the heme cofactor in red spacefill (McLean et al., 2009). B: cholestenone-bound CYP125A1 (PDB code 2X5W). Cholest-4-en-3-one aligns in the active-site channel with the aliphatic side chain facing the distal surface of the heme cofactor at the C27 position (Ouellet et al., 2010a). C: androstenedione-bound CYP125A1 (PDB code 3IW1). The ligand is in blue-coloured sticks. The image shows a binding mode for androstenedione which is not compatible with P450 oxidation due to absence of the alkyl side chain found in cholestenone (McLean et al., 2009). D: econazole-bound CYP125A1 (PDB code 3IW2). The ligand is in magenta-coloured sticks. The picture shows the binding mode for econazole, which is prevented from migration into the active site by steric constraints due to active site narrowing near the heme, with no heme ligation occurring (McLean et al., 2009). Structures were drawn using PyMol (DeLano, 2002).

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1.5.2.2 CYP142A1 (Rv3518c): Functional Redundancy

Recent studies revealed a functional redundancy in cholesterol oxidation capacity in

the Mtb H37Rv strain (but not in M. bovis BCG) and further highlighted that a

compensatory mechanism can overcome absence of CYP125A1 in this bacterium

(following inactivation of the CYP125A1 gene) to aid growth on cholesterol (Capyk

et al., 2009). Hence it is worth noting that another P450 gene (CYP142A1, Rv3518c)

is located in the cholesterol gene cluster of Mtb. (Driscoll et al., 2010). In research

carried out by Capyk and co-workers, it was evident that though M. bovis BCG grew

on cholesterol, the M. bovis BCG CYP125 deletion strain did not show any growth,

unless complemented by another CYP125 gene copy (Capyk et al., 2009). However,

the CYP125 deletion strain of Mtb H37Rv still grew on cholesterol, suggesting a

compensatory mechanism in cholesterol 27-hydroxylase activity in this strain

Driscoll et al., 2010). Studies showed that a ∆CYP125 Mtb CDC1551 strain

accumulated cholest-4-en-3-one that was toxic to the cells, suggesting that, in Mtb

CDC1551, CYP125 plays a key role in detoxification of cholest-4-en-3-one by 27-

hydroxylation, leading to its degradation (Capyk et al., 2009). To probe further into

the mystery behind the inability of the M. bovis BCG ∆CYP125 strain to grow on

cholesterol (and the ability of the ∆CYP125A1 Mtb H37Rv strains to grow on

cholesterol), Driscoll et al. noted that Mtb H37Rv had another CYP gene in the

cholesterol operon (but outside the igr region) and hence suggested that CYP142A1

(the Rv3518c gene product) might also metabolize cholesterol/cholest-4-en-3-one,

or catabolize products from these molecules. Results generated from the studies

identified CYP142A1 as a second cholesterol 27-hydroxylase and a possible

92

complementary enzyme in the absence of CYP125A1 (Driscoll et al., 2010). A recent

study has suggested that CYP142A1 may play a key role during Mtb infection by

making extra reservoirs of esterified intracellular cholesterol in the host more

accessible to the pathogen than would ordinarily be available for energy generation

(Frank et al., 2014). The results from this study demonstrated that CYP142A2 from

M. smegmatis (the soil-dwelling counterpart of Mtb) metabolises cholesteryl sulfate

and cholesteryl propionate, whereas CYP125 enzymes metabolise cholesteryl

sulfate at a much slower rate and do not significantly oxidize cholesteryl propionate

(Frank et al., 2014). The crystal structure of CYP142A2 complexed with cholesteryl

sulfate revealed the substrate in a conformation similar to that of the 4-cholesten-

3-one-bound structure solved previously, with the bulky sulfate group projecting

out toward the solvent (Garcia-Fernandez et al., 2013, Frank et al., 2014) (Fig. 4A).

The crystal structure of the ligand-free CYP142A1 has also been solved at a

resolution of 1.6 Å, with the overall fold highly similar to that of Mtb CYP124A1 and

CYP125A1, which also hydroxylate cholesterol and cholestenone (Driscoll et al.,

2010). The active site topologies of CYP142A1 and CYP125A1 are similar, with a

letterbox-shaped entry-exit channel formed by the FG-loop, the BC-loop and the I-

helix N-terminal region. Their active site channel is lined by mainly hydrophobic

residues which curve upwards away from the heme cofactor (Driscoll et al., 2010).

In contrast, the CYP124A1 entry-exit channel is located perpendicular to the heme

and is formed by the FG- and BC-loops in addition to the β1-β2-loop domain

(Driscoll et al., 2010). This may be consistent with the recent suggestion that

CYP142A1 does not oxidize fatty acids or methyl branched lipids, but rather is

involved in the metabolism of the cholesterol side chain (McLean et al., 2010).

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In Mtb, CYP142A1 appears to serve only (or mainly) as a “back-up” for CYP125A1 in

the side-chain degradation of host cholesterol, while in a ∆CYP125-∆CYP142 mutant

of M. smegmatis, the cholesterol metabolizing property is retained. These findings

suggest the presence of another source of redundancy within the M. smegmatis

genome (Garcia-Fernandez et al., 2013). Studies had earlier revealed that an

unusually high number of CYPs (e.g. 20 CYPs in Mtb H37Rv, 29 CYPs in Rhodococcus

jostii RHA1 and 40 CYPs in M. smegmatis) are found in actinobacterial genomes,

compared to Escherichia coli which has a similar genomic size with Mtb, for

example, and yet possesses no P450 enzymes (Hudson et al., 2012a, Garcia-

Fernandez et al., 2013, Ouellet et al., 2010a). In line with the above facts, M.

smegmatis possesses more CYPs than Mtb H37Rv (Garcia-Fernandez et al., 2013).

Hence an alternative cholesterol catabolic pathway or enzyme encoded within the

M. smegmatis genome could be responsible and one possible suggestion is the

cytochrome P450 encoded by the MSMEG_4829 gene (CYP189A1) (Garcia-

Fernandez et al., 2013). CYP189A1 was shown to be upregulated in both wild-type

and mutant strains of M. smegmatis grown on cholesterol, but it does not have an

orthologue in Mtb (Garcia-Fernandez et al., 2013). However, studies are ongoing to

further characterize this enzyme and the putative role it plays in cholesterol

metabolism. This research could give more insights into the differences in the

cholesterol catabolic pathways between Mtb and M. smegmatis (Garcia-Fernandez

et al., 2013).

CYP142A1 exists in a low-spin resting state with a Soret peak at 418 nm, and hence

differs from CYP125A1 which is typically isolated in an extensively high-spin state

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(Soret peak at 393 nm). This high-spin state appear to result from the side chain

conformation of the Val-267 in CYP125A1 and the effect that this has on the water

molecule which serves as the sixth axial ligand to the CYP125A1 heme iron (McLean

et al., 2009, Driscoll et al., 2010).

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Figure 1.24: Structural features of CYP142 enzymes from Mtb and M. smegmatis. A: solvent-accessible surface of Mtb CYP142A1 (PDB code 2XKR). B: solvent-accessible surface of M. smegmatis CYP142A2 (PDB code 3ZBY). The colour coding for A and B shows the helices in cyan, the sheets in magenta and the loops in pink. The arrow indicates the narrow access site entry for the CYP142 enzymes which can be readily identified by the direct view onto the heme cofactor in red spacefill (Frank et al., 2014, Garcia-Fernandez et al., 2013, Driscoll et al., 2010). C: Cholestenone-bound CYP142A2 (PDB code 2YOO). Cholestenone binds close to the heme with the C-27 methyl group displacing the distal water from the heme iron. This binding mode favours cholesterol/cholestenone oxidation at the C-27 position to initiate side chain metabolism for energy generation (Garcia-Fernandez et al., 2013). D: Cholesterol sulfate-bound CYP142A2 (PDB code 4TRI), with substrate binding mode similar to that for the cholestenone-bound structure (Frank et al., 2014). Structures were drawn using PyMol (DeLano, 2002).

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1.5.2.3 CYP124A1 (Rv2266): A Methyl-Branched Lipid-

Hydroxylase

CYP124 P450s are found in an array of microorganisms, including actinobacteria,

some proteobacteria as well as pathogenic and non-pathogenic mycobacterial

species (Ouellet et al., 2010b, Johnston et al., 2009). This wide occurrence of

CYP124 genes is indicative of an important physiological role (Johnston et al., 2009).

Mtb CYP124A1 is found in a gene cluster containing a sulfotransferase (Sft3 encode

by Rv2267c) that plays a key role in a 3'-phosphoadenosine 5'-phosphosulfate

(PAPS)-dependent sulfation of menaquinone MK-9 DH-2 at the omega-position

(Johnston et al., 2009, Holsclaw et al., 2008, Mougous et al., 2006). The gene

location for CYP124A1 is in the same chromosomal region as those for CYP128A1

(Rv2268c) and CYP121A1 (Rv2276) (McLean et al., 2010). The sulfated product of

menaquinone MK-9 DH-2 (referred to as “S881”) is found in the cell membrane of

Mtb, where it was shown to be involved in virulence in Mtb-infected mice

(Mougous et al., 2006). Recent work has shown that CYP124A1 metabolises a wide

range of substrates with similar chemical structures to menaquinone MK-9 DH-2

(i.e. compounds with repeating methyl branching) (Johnston et al., 2009). The

CYP124A1 reaction involves a hydroxylation reaction at the omega position, with a

significant preference for methyl-branched lipids (Johnston et al., 2009). CYP124A1

binds a range of methyl branched fatty acids (e.g. phytanic acid) with high affinity

and catalyses their hydroxylation at the omega-position (Figure 1.25) (Johnston et

al., 2009). Comparing with the other 20 Mtb P450s, it is evident that CYP125A1 has

the closest similarity to CYP124A1 (38%), followed by CYP126A1 (34%).

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The crystal structures of Mtb CYP124A1 have been solved in both the ligand-free

and phytanic acid-bound forms (Figure 1.26) (Johnston et al., 2009). The crystal

structure of CYP124A1 in complex with phytanic acid helps reveal the mechanism

by which the active site enables an unfavourable regioselectivity of substrate

oxidation (Johnston et al., 2009). A small hydrophobic space located near the heme

cofactor binds one of the terminal methyl groups such that the other is aligned

close to the heme iron for oxidation (Johnston et al., 2009). As described above,

recent studies have identified CYP124A1, CYP142A1 and CYP125A1 as involved in

the successive oxidations of the aliphatic side chain of cholesterol/cholest-4-en-3-

one at the C-27 position to the alcohol, aldehyde and then the carboxylic acid

(Johnston et al., 2010) (Figure 1.22). Recent studies by Johnston et al have revealed

that all the three P450 isoforms oxidize cholest-4-en-3-one side chain completely to

the carboxylic acid, which is a prerequisite for entry into the β-oxidation pathway

(Johnston et al., 2010). However, out of three isoforms, CYP125A1 is the most

efficient catalyst, followed by CYP142A1 and lastly CYP124A1 (Johnston et al.,

2010).

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Figure 1.25: CYP124A1 catalyses the -hydroxylation of phytanic acid and other methyl-branched lipids. Image redrawn from (McLean et al., 2010) using chemdraw (Mills, 2006).

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Figure 1.26: Structural features of CYP124A1 from Mtb. A: Cartoon representation of ligand-free Mtb CYP124A1 (PDB code 2WM5). Heme is in pink, iron is coloured brown and the secondary structural elements are in grey. B: the solvent-accessible surface of ligand-free Mtb CYP142A1 (PDB code 2WM5). The colour coding shows the helices in red, the sheets in yellow and the loops in green. The arrow indicates the narrow access site entry for CYP124A1, which is readily identified by the direct view onto the heme cofactor in pink spacefill with brown heme iron (Johnston et al., 2009). C: Phytanic acid-bound Mtb CYP124A1 (PDB code 2WM4). Phytanic acid binds near the heme with its C27 methyl group close to the heme iron, leading to hydroxylation of the substrate at this position (Johnston et al., 2009). D: Active site of phytanic acid-bound CYP124A1 from Mtb, showing the ligand in yellow sticks and in transparent spacefill (PDB code 2WM4) (Johnston et al., 2009). Structures were drawn using PyMol (DeLano, 2002).

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1.5.2.4 Cholesterol Catabolism: A Promising Drug Target in Mycobacterium tuberculosis

Figure 1.27: Cholesterol catabolic pathway. The aliphatic tail of cholesterol is metabolized to the acid by the cholesterol oxidizing P450s and channelled to β-oxidation for energy generation. Sterol ring degration occurs simultaneously with carbon atoms derived from the ring nucleus converted to CO2 via the tricarboxylic acid cycle (TCA), whereas the propionyl-CoA produced from the degradation of the side chain is channelled into mycobacterial lipids, including the virulence factor PDIM. Figure adapted from Ouellet et al., 2011. Even though Mtb, like most bacteria, does not synthesize its own sterols, studies

have revealed that cholesterol is essential for infection of macrophages and for

survival in the latent phase of infection (Ouellet et al., 2011). The inability of Mtb to

make sterols is due to the absence of squalene monooxygenase and oxidosqualene

cyclase, enzymes essential for sterol biosynthesis. Mtb, however, persists in the

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harsh environment of macrophages in the human lung by utilization of host

cholesterol as a sole source of carbon and energy (Ouellet et al., 2011). Studies have

revealed that high levels of cholesterol in the diet can increase the bio-load of Mtb

in the human lung significantly, which could overwhelm the host defence

mechanisms leading to impaired immunity to the bacterium (Koul et al., 2004, Kim

et al., 2010, Schafer et al., 2009, Martens et al., 2008). Cholesterol is essential for

the phagocytosis of Mtb into macrophages, and Mtb enters the phagocytic cells via

cholesterol-rich membrane microdomains. Furthermore, the bacterium utilises

cholesterol to maintain the host protein coronin 1 (TACO or tryptophan-aspartate-

containing coat protein) on Mtb-infected phagosomes, and this inhibits

phagosome–lysosome fusion (Gatfield and Pieters, 2000, Munoz et al., 2009).

The mechanism by which Mtb takes up host cholesterol is poorly understood.

However, an ABC-like transport system, mce4, that is involved in cholesterol import

into the bacterium was discovered recently (Pandey and Sassetti, 2008). The growth

of mce4 gene deletion strains of Mtb was impaired when cholesterol was utilized as

the primary source of carbon (Pandey and Sassetti, 2008, Ouellet et al., 2011). Using

radio-labelled cholesterol derivatives, it was unambiguously shown that cholesterol

is metabolised by Mtb, with the carbon atoms from the sterol backbone and the

aliphatic side chain channelled to energy generation and lipid synthesis, respectively

(Figure 1.27)(Pandey and Sassetti, 2008, Ouellet et al., 2011). Furthermore, Yang et

al. demonstrated that cholesterol degradation in Mtb elevates the average mass of

the lipid virulence factor phthiocerol dimycocerosate (PDIM), which results from the

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higher metabolic flux of propionate derived from cholesterol breakdown (Yang et

al., 2009).

In addition to the above, the blockage of cholesterol import in mce4 gene deletion

strains of Mtb significantly reduced Mtb virulence in both activated macrophages

and in a mouse model of infection, further validating the important role of

cholesterol metabolism in chronic infection (Pandey and Sassetti, 2008). Screening

of a transposon mutant library revealed another locus, igr, which is important for

Mtb growth in activated macrophages (Chang et al., 2007). The igr gene locus

comprises six genes, including a cytochrome P450 (CYP125A1), two acyl-CoA

dehydrogenases (fadE28 and fadE29), two conserved hypothetical proteins

(Rv3541c and Rv3542c) and a putative lipid carrier protein (ltp2) (Ouellet et al.,

2011).

Inactivation of the whole igr operon in a subsequent study revealed a pronounced

effect on growth on cholesterol alone or in combination with glycerol, signifying

some level of cell intoxication by cholesterol or its metabolites (Chang et al., 2009).

The mce4 operon is also found in many other actinobacterial species, including

Rhodococcus jostii, M. smegmatis and M. bovis BCG, and these bacteria have also

been shown to utilize cholesterol for growth (Van der Geize et al., 2007, Mohn et

al., 2008, Av-Gay and Sobouti, 2000). Hence, research to date indicates that

cholesterol utilisation by the Mtb bacterium is important for bacterial infectivity

and persistence, and the cholesterol degradation pathway could provide important

therapeutic targets for the development of new anti-tubercular agents (Ouellet et

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al., 2011). As described above, it seems highly likely that targeting cholesterol

catabolism (and the P450 enzymes involved) in persistent/latent bacteria could hold

the key to a major breakthrough in anti-TB therapy (McLean et al., 2010).

1.5.3. CYP51B1: The First Member of the CYP51 Family Identified in Prokaryotes

The 14α-sterol demethylases (CYP51s) are eukaryotic cytochromes P450 that

catalyse the removal of the 14α-methyl group from lanosterol, 24-methylene

dihydrolanosterol and obtusifoliol in the manufacture of cholesterol, ergosterol and

phytosterols in animals, fungi and plants, respectively (Noshiro et al., 1997, Monk et

al., 2014). Sterol 14α-demethylases have been discovered in plants (e.g. Sorghum

bicolor) and in a large number of eukaryotes, and by the year 2000 the orthologous

nature of a CYP51-like gene from Mtb to the eukaryotic CYP51s was confirmed,

signalling the first potential sterol demethylase in a bacterial genome (Aoyama et

al., 1998, Bellamine et al., 1999, McLean et al., 2010). This was a major

breakthrough given that the fungal CYP51 enzymes (e.g. in Aspergillus spp. and in

Candida albicans) are important drug target enzymes known to be inactivated by

azole-based drugs such as econazole, clotrimazole and ketoconazole (McLean et al.,

2010).

The sterol 14α-demethylase (CYP51) family is the only CYP gene family of P450s

which is widely distributed in different biological kingdoms, being found in animals,

plants, fungi, yeast, lower eukaryotes and bacteria (Noshiro et al., 1997, Lepesheva

and Waterman, 2004, Monk et al., 2014) and is widely considered to be the most

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genetically ancient member of the P450 superfamily (Aoyama et al., 1996, Nelson,

1999, Monk et al., 2014). Most forms of CYP51 are membrane-bound proteins,

which have complicated structural studies of this type of protein using X-ray

crystallography (Podust et al., 2001). However, the full length structure of the

CYP51 from Saccharomyces cerevisiae, an integral membrane protein, has been

successfully solved, revealing a single transmembrane helix at the N-terminal region

of the P450. This has given further insight into how single-transmembrane helices

orient cytochrome P450 enzymes at the bilayer surface (Monk et al., 2014). Crystal

structures of other CYP51 enzymes include those for the soluble CYP51B1 enzyme

from Mtb in complex with azole inhibitors (Podust et al., 2001); and N-terminal

(membrane anchor) truncated enzymes from humans (Strushkevich et al., 2010).

Figure 1.28: Features of CYP51B1 from Mtb. The left panel shows a cartoon representation of fluconazole-bound Mtb CYP51B1 (PDB code 1EA1). The heme is in red sticks with iron in brown, and the secondary structural elements are in a wheat colour. The fluconazole ligand is shown in spacefill with carbon atoms in cyan. The right panel shows a close-up of the fluconazole binding site – illustrating the coordination of the heme iron by a triazole ring nitrogen from the inhibitor (Podust et al., 2001). Structures were drawn using PyMol (DeLano, 2002).

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1.5.4 CYP121A1: An Essential Gene for Mtb Viability

The gene encoding the Mtb P450 CYP121A1 (rv2276 or CYP121A1), was

documented to be important for Mtb viability (McLean et al., 2008). This important

discovery was made following the demonstration that the CYP121A1 gene could not

be deleted from the Mtb H37Rv genome unless a second copy of the gene had been

inserted elsewhere on the chromosome (McLean et al., 2010). This also suggests

that the product of the CYP121A1 reaction could play important physiological roles

for the bacterium (McLean et al., 2008, McLean et al., 2010). CYP121A1 has

received a remarkable amount of interest among the twenty P450 enzymes found

in Mtb H37Rv and a lot of work has been done on this particular enzyme (Dumas et

al., 2014). This results from its importance for the viability of Mtb (in vitro and likely

in the human host) and also its ability to catalyse a carbon–carbon coupling

reaction, which is unusual for a P450 enzyme (Dumas et al., 2014). CYP121A1

catalyses the oxidative crosslinking of the tyrosyl side chains of the cyclic dipeptide

cyclo-L-Tyr-L-Tyr [or cYY] to produce the metabolite mycocyclosin (mcyc) (Figure

1.28). This reaction leads to a cyclization process where two tyrosine aromatic rings

are covalently joined through a carbon-carbon bond in the ortho-position with

respect to each tyrosine hydroxyl group (Belin et al., 2009). Crystal structures of

CYP121A1 in complex with the substrate (cYY) and fluconazole (as well as mutant

forms of CYP121A1) have been solved (McLean et al., 2008, Seward et al., 2006,

Belin et al., 2009). The fluconazole-bound structure revealed a new mode of azole

drug binding in which the heterocyclic nitrogen of the triazole ring did not

coordinate directly to the heme iron, but instead ligated the ferric iron via a water

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molecule that remained as the sixth ligand to the P450 heme iron (Seward et al.,

2006).

Figure 1.29: CYP121A1 catalyses the formation of an intramolecular C-C bond between 2 tyrosyl carbon atoms of cyclodityrosine. Image redrawn from (McLean et al., 2010) using ChemDraw (Mills, 2006).

Figure 1.30: Structural features of Mtb CYP121A1 in complex with the substrate (cyclodityrosine) and fluconazole. A: cyclodityrosine (cYY)-bound CYP121A1 (PDB code 3G5H). The ligand is in stick representation, with carbons in yellow and other atoms in standard colours. The heme is in red sticks with pyrrole nitrogens in blue, and the heme iron in brown (Belin et al., 2009) B: Fluconazole-bound CYP121A1 (PDB code 2IJ7). The ligand is again in stick representation, with carbons in green and other atoms in standard colours. Heme is represented as described in panel A (Seward et al., 2006). Structures were drawn using PyMol (DeLano, 2002).

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1.5.5 CYP130A1 (Rv1256c): Essential for Virulence in Mtb?

Comparative genome analysis shows that the Mtb CYP130A1 and CYP141A1 genes

are absent in the virulent M. bovis strain and from its avirulent counterpart M. bovis

BCG, indicating that they are likely not important for Mtb growth per se, but could

be essential for Mtb virulence and pathogenicity in humans (Ouellet et al., 2008,

McLean et al., 2007a). Rv1256c (CYP130A1), the gene encoding the P450 CYP130A1

in Mtb, was shown to be part of an operon along with the gene Rv1258c that

encodes for a tetracycline/aminoglycoside resistance TAP[2]-like efflux pump (Ainsa

et al., 1998).

Though an orphan P450 (in terms of a known catalytic function), work by Ortiz de

Montellano’s group produced the crystal structure of the ligand-free and econazole-

bound CYP130A1 at resolutions of 1.46 Å and 3.0 Å, respectively (McLean et al.,

2010, Ouellet et al., 2008). Interestingly, the ligand-free CYP130A1 crystallized as a

monomer in the “open” conformation, while the econazole-bound form crystallized

as a dimer in a “closed” conformation. Changes in the conformation of the

secondary structural elements resulted in a reorganization of the BC-loop, F- and G-

helices to accommodate the binding of econazole in the active site, which also

affected the shape of the protein surface (Ouellet et al., 2008). Econazole ligates the

heme iron (with a coordination bond length of 2.75 Å) via the lone pair electrons of

its imidazole nitrogen atom (Ouellet et al., 2008).

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Figure 1.31: Crystal structures of ligand-free and econazole-bound CYP130A1. A: Features of the econazole-bound CYP130A1 dimer, revealing a “closed” conformation (PDB code 2UVN). Secondary structure elements are in wheat colour, heme is in red and econazole is in green sticks and spheres. B: monomeric features of the ligand-free CYP130A1 revealing an “open” conformation (PDB code 2UUQ) (Ouellet et al., 2008). Structures were drawn using PyMol (DeLano, 2002).

1.5.6 CYP126A1 (Rv0778)

CYP126A1 (encoded by Mtb H37Rv gene rv0778) is located adjacent to important

Mtb genes that encode enzymes involved in purine synthesis, and CYP126A1 is also

part of a gene cluster encoding the adenylosuccinate lyase, PurB (Lew et al., 2011).

However, CYP126A1 also shares about 35% identity with the Mtb cholesterol

hydroxylases CYP124A1 and CYP125A1, and it is highly conserved across both

pathogenic and non-pathogenic strains of actinobacteria, which suggests important

physiological roles that are still unknown (Ouellet et al., 2010b, Hudson et al.,

2014). Recent studies to predict the functional role of CYP126A1 have involved the

use of a novel fragment based screening approach, which is a method for

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identifying and designing small-molecule ligands as chemical tools and leads for

drug development (Anand et al., 2011, Hudson et al., 2014). Interestingly, the

fragment screening hit rate against CYP126A1 (14%) was significantly higher than

those observed for Mtb CYP121A1 and CYP125A1 (which were only 4% and 1%,

respectively) (Hudson et al., 2012b, Hudson et al., 2013).

1.5.7 CYP128A1: An Essential Enzyme with a Role in

Hydroxylation of Respiratory Menaquinone Mtb synthesizes a wide range of lipids, some of which are involved in immune

response modulation within infected individuals, while others envelop the organism

within a thick and resilient physical barrier (Gokhale et al., 2007, Brennan and Crick,

2007, Schelle and Bertozzi, 2006, Onwueme et al., 2005). Research carried out by

Bertozzi’s group identified several sulfotransferases within the Mtb genome (Schelle

and Bertozzi, 2006, Mougous et al., 2002). S881 is a sulfated metabolite that was

found to be located on the outer cell wall of the bacterium, where it acts as a

negative regulator of virulence in the mouse model of tuberculosis (Mougous et al.,

2006, ten Bokum et al., 2008). It was also revealed that the biosynthesis of S881 is

dependent on the PAPS (3′-phosphoadenosine 5′-phosphosulfate)-dependent

sulfotransferase Stf3 (encoded by Rv2267c) (Mougous et al., 2006, Holsclaw et al.,

2008), which is located in a three gene operon containing CYP128A1 (Holsclaw et

al., 2008, Cole et al., 1998). Subsequently, Holsclaw and co-workers elucidated the

structure of S881 (Figure 1.31), revealing it to be a menaquinone-like molecule (MK-

9 DH-2) containing a linear chain consisting of nine isoprenoid units appended to a

naphthoquinone ring (Ouellet et al., 2010b). Menaquinone is a molecule that was

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shown to be the major quinol electron carrier in Mtb respiration (McLean et al.,

2010). This then led to a biosynthetic scheme which suggested that CYP128A1 first

hydroxylates the terminal isoprene unit of MK-9 (DH-2) and hence activates it for

nucleophilic PAPS-dependent sulfation (Ouellet et al., 2010b). Sulfation of

menaquinone is suggested to play a key role in Mtb virulence regulation (Holsclaw

et al., 2008) (Figure 1.31).

Attempts to produce CYP128A1 in a soluble form to enable its characterization have

proven unsuccessful to date, even though high levels of its expression have been

reported in E. coli (Ouellet et al., 2010b, McLean et al., 2010). Due to the

hydrophobic nature of its quinol substrate, it was suggested that CYP128A1 could

be associated with the membrane to access this molecule, and hence detergent

solubilization of CYP128A1 may be helpful to enable biochemical characterization of

this P450 and to confirm its physiological role (McLean et al., 2010).

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Figure 1.32: Biosynthesis of the Mtb S881 sulfolipid. CYP128A1 is postulated to

hydroxylate the terminal -position of MK-9 (DH2). This is the primary step in its metabolism, and is followed by a PAPS-dependent sulfation by the Stf3 enzyme to form the S881 sulfolipid. Image redrawn from (Ouellet et al., 2010b) using ChemDraw (Mills, 2006).

1.5.8 Other Partially Characterized P450 Systems in Mycobacterium tuberculosis

CYP139A1

The CYP139A1 gene is located in a cluster with polyketide synthase (pks) genes and

with a component of the prospective polyketide exporting membrane protein

system (encoded by Rv1668c), suggesting a role for this cytochrome P450 in

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polyketide metabolism. However, the structure and function of this P450 remain

unknown (McLean et al., 2010).

CYP132A1

CYP132A1 has significant protein sequence similarity to members of a well

characterized cytochrome P450 family (the CYP4 fatty acid oxidases). This similarity

has helped researchers to postulate CYP132A1’s probable function. Presently, there

is no documentation of the characterization of CYP132A1, even though the

CYP132A1 gene was reported as a major target of gene induction for the AraC-type

transcriptional regulator (product of gene Rv1395), which is encoded by a gene

adjacent to CYP132A1 on the chromosome. This suggests a physiological role for

CYP132A1 in bacterial virulence (Recchi et al., 2003). Nevertheless, CYP132A1’s

function and any role in Mtb infectivity or pathogenesis remain to be defined

(McLean et al., 2010).

CYP123A1

CYP123A1 (encoded by the Mtb H37Rv Rv0766c gene) is documented to be highly

conserved among actinobacteria and proteobacteria, and the CYP123A1 gene was

shown to be up-regulated in Mtb at high temperatures (Stewart et al., 2002).

However, other studies also showed CYP123A1 to be down-regulated in a PhoPR

two-component system mutant strain of Mtb (Walters et al., 2006). The PhoPR two-

component system is essential for Mtb virulence. It is a positive transcriptional

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regulator of genes associated with the synthesis of components in the Mtb cell

envelope (Walters et al., 2006). It is postulated that when Mtb encounters the

harsh environment of the human lung, PhoPR plays an important role in cell

envelope remodelling and also enables Mtb to switch to alternative metabolic

pathways involved in lipid metabolism or synthesis. This enables the bacterium to

successfully persist and survive in the infected host (Walters et al., 2006).

CYP138A1 and CYP141A1

Research has shown that the Mtb CYP141A1 and CYP138A1 genes are up-regulated

following a brief (about 2 h) exposure to lung surfactant, indicating a possible role

for these P450s during the early stages of infection (Schwab et al., 2009).

Furthermore, the CYP141A1 and fprA (Rv3106) genes are found within the Mtb

genome in a location adjacent to a gene cluster involved in molybdopterin cofactor

biosynthesis (Ouellet et al., 2010b). Molybdopterin is required by enzymes involved

in anaerobic metabolism and also for activity of nitrate reductase (Dubnau et al.,

2005). The CYP138A1 gene is also up-regulated during heat shock response at high

temperatures (Stewart et al., 2002). The CYP141A1 protein is a Mtb P450 enzyme

predicted to have important function(s) in the bacterium (McLean et al., 2007a).

The gene sequence of CYP141A1 was used by Darban-Sarokhalil and co-workers as

the basis for design of a PCR-based technique for the rapid detection of Mtb from

respiratory specimens (Darban-Sarokhalil et al., 2011, Rabiee-Faradonbeh et al.,

2014).

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The CYP141A1 and CYP130A1 genes are absent from the Mtb vaccine strain M.

bovis BCG, and are located in “regions of deletion” (also known as “regions of

difference”; RDs 13 and 12, respectively) in the BCG strain, which are considered to

eliminate key genes involved in Mtb virulence (McLean et al., 2010). The likely

important roles played by CYP130A1 and CYP141A1 in Mtb virulence is thus a major

drive for their enzymatic characterization (McLean et al., 2010).

Other Mycobacterium tuberculosis P450s

Relatively little information is currently available for the remaining Mtb P450

enzymes. CYP136A1 is suggested to be a distant relative of sterol demethylase

(CYP51) enzymes, but such functional assignment and any physiological role

remains unclear (Ouellet et al., 2010b). The CYP137A1 gene is found in a cluster

close to one of the essential WhiB-like transcriptional regulatory proteins, WhiB4.

These are proteins that act in response to cellular redox changes or metabolic shifts

(Soliveri et al., 2000). CYP139A1 is predicted to be eukaryotic-like (i.e. CYP139A1

may be membrane-associated with a potential N-terminal membrane anchor) and

is conserved among pathogenic mycobacterial strains, but is not found in non-

pathogenic strains. The CYP139A1 gene is chromosomally adjacent to a cluster of

polyketide synthase genes (pks17, pks9 and pks11 being the closest) and is also

located adjacent to a gene encoding a predicted macrolide transport protein

(Rv1667c), suggesting a possible role for CYP139A1 in oxidative tailoring of a

nascent polyketide prior to its export from the cell (McLean and Munro, 2008). The

pks gene pks17 alone was reported to be essential in Mtb from data generated by

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screening a transposon-mutant library (Sassetti et al., 2003). The proteins encoded

by genes pks17 and pks8 were shown to play key roles in the biosynthesis of

methyl-branched unsaturated fatty acids (Dubey et al., 2003). The Mtb genes

CYP143A1 and CYP144A1 are each highly conserved across many pathogenic and

non-pathogenic strains of actinobacteria (Ouellet et al., 2010b). The CYP144A1

gene, along with CYP125A1 and CYP132A1, was identified by transcriptional

profiling to be expressed specifically in Mtb-infected human dendritic cells after 18

hours (Tailleux et al., 2008), opening up an additional connection between virulence

and Mtb P450 enzymes (Ouellet et al., 2010b). CYP143A1 is located in the same

genomic region as CYP144A1, but little is known about their physiological functions.

The CYP143A1 gene is located immediately adjacent to (on the opposite strand) the

ferredoxin encoded by Rv1786, suggesting that these are redox partner proteins.

1.5.9 Azole Antibiotics: Non-Selective Inhibitors of Mtb Cytochrome P450 Enzymes

The druggability of Mtb cytochrome P450 enzymes is validated by the high level of

anti-mycobacterial activity exhibited by azole anti-fungal drugs (Hudson et al.,

2012a). These are compounds with a five-membered nitrogen heterocyclic ring

containing at least one other non-carbon atom which could be nitrogen, sulphur or

oxygen (Figure 1.33). Azole antibiotics were originally developed as antifungals

targeting CYP51, but azole drugs such as econazole and clotrimazole were shown to

be potent against most of the Mtb P450 enzymes tested. For example, CYP121A1,

which is important for Mtb viability, exhibits a very high affinity for some of the

azole drugs, with Kd values in the nanomolar range (McLean et al., 2008). Studies

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have also shown that the azole antifungal econazole is capable of clearing both

active and latent Mtb infection from infected mice (Ahmad et al., 2006a, Munro et

al., 2013, Byrne et al., 2007, Ahmad et al., 2006c, Ahmad et al., 2006b).

To evaluate the effects of azole inhibitors against Mtb H37Rv, the minimal

inhibitory concentration (MIC) data were determined (Mclean et al., 2008). The

data revealed that econazole (MIC = 8 µg/ml) and miconazole (8 µg/ml) were most

potent, followed by clotrimazole (11 µg/ml and lastly ketoconazole (16 µg/ml). For

M. smegmatis, MIC data obtained were: econazole (MIC <0.1 µg/ml), miconazole

(1.25 µg/ml), clotrimazole (0.1 µg/ml), and ketoconazole (20 µg/ml). Even though

the MIC values for Mtb H37Rv were higher than those obtained previously for M.

smegmatis (Mclean et al., 2002b), the data revealed that the azoles possessed

activity against Mtb.

The azole compounds inhibit P450 catalysis by reversibly ligating the ferric heme

iron via a heterocyclic nitrogen atom (on an imidazole or triazole group) in the sixth

distal ligand position, to give a type-II spectral shift (Soret shift to a longer

wavelength) (Ortiz de Montellano, 2005, Odds et al., 2003). This binding displaces

the weakly bound water ligand on the 6th axial position and hence also prevents

substrate access to the active site (Hudson et al., 2012a). As discussed earlier, the

crystal structures of some Mtb P450 enzymes in complex with azole inhibitors have

been determined.

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Due to their poor bioavailability and promiscuous affinity for human P450s (which

can result in high toxicity levels and drug-drug interactions when co-administered

with other drugs), the use of azole antibiotics in TB treatment remains questionable

(Odds et al., 2003, Perea et al., 2004, Como and Dismukes, 1994). In addition,

mutations in Mtb have led to the emergence of resistance to azole drugs, which

results from up-regulation of a transmembrane transporter protein believed to act

as an efflux pump for azoles and other compounds (Milano et al., 2009).

Understanding molecular interactions of azoles in P450 active sites provides a route

for designing more specific azole-based inhibitors and for rationalising (and

avoiding) development of drug resistance (Podust et al., 2001, Podust et al., 2009).

Other routes to making Mtb P450-specific inhibitors as novel TB drugs are also

attractive options – given their important functions in the physiology and infectivity

of the bacterium (Hudson et al., 2012a).

Figure 1.33: Chemical structures of selected azole antibiotics: Figures were drawn using ChemDraw (Mills, 2006). Coordination of the P450 heme iron occurs from an imidazole nitrogen atom in the case of each of these azole drugs.

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1.6 Novel Drug Discovery Approaches

1.6.1 Fragment Based Drug Discovery (FBDD): A Novel Approach to Development of New P450 Inhibitor Scaffolds

The desperate need for the development of new TB drugs has become a global

problem. The majority of anti-tubercular drugs currently in use have been in

existence for over 40 years and have been rendered ineffective due to the

emergence of multi-drug (MDR-TB) and extensively drug-resistant (XDR-TB) strains

of Mtb (McLean et al., 2010, Hudson et al., 2012a). The WHO has declared this

situation a “global emergency” and only now, for the first time in many decades, are

there newly developed compounds undergoing clinical and preclinical trials to

tackle this public health problem (Check, 2007).

Fragment-based drug discovery (FBDD) has been introduced in the last decade as a

promising tool for drug discovery and has caused a new awakening in the process of

drug design, with many FBDD compounds being put into clinical trials or approved

for use in recent years (Wang et al., 2015, Erlanson, 2012). The origin of the

concepts behind FBDD can be traced back to pioneering work of two renowned

scientists, Jencks and Ariens, about 30 years ago (Jencks, 1981, Rees et al., 2004,

Erlanson, 2012). It was only in the mid-1990s that experimental techniques became

adequately sensitive and rapid for the concept to be become feasible (Erlanson,

2012). Jencks and Ariens demonstrated that drug-like molecules can be regarded as

the combination of two or more distinct binding epitopes (or fragments) (Rees et

al., 2004). The screening for these discrete binding moieties using small ligand

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molecules (fragments) generated what is known as the ‘fragment approach’ (Rees

et al., 2004). In this approach, the starting points are very small, weak-binding

molecules (fragments) that are about half the size of typical drugs with molecular

weights between 120-250 Da or 8-18 non-hydrogen atoms (Erlanson, 2012, Rees et

al., 2004). These fragments are then chemically elaborated to create drug leads

(Figure 1.34 and 1.35) (Erlanson, 2012).

There are three major methods for the elaboration of initial fragment hits (Scott et

al., 2012). 1) fragment growing, in which fragments are chemically grown (guided by

data from structural biology that shows their target binding modes) into new

unexplored spaces within/around their binding sites; 2) fragment linking, where two

or more fragments that bind closely within the active site are covalently linked to

one another (while trying to retain their individual binding modes); and 3) fragment

merging, in which the components of fragments that overlap in the binding pocket

are fused together to generate more potent inhibitors (Hudson et al., 2013).

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Figure 1.34: A schematic representation of the Fragment Based Drug Discovery (FBDD) approach. Fragments are screened against targets of interest, leading to identification of “hits”, which are further linked/grown/merged and optimised to create lead compounds for drug development. Image was drawn using Microsoft PowerPoint, and adapted from (Erlanson, 2012).

Another widely used current method of drug discovery is high-throughput screening

(HTS), in which very large numbers (up to millions) of compounds are collected and

screened against a target of interest (Erlanson, 2012). HTS has been used

successfully in development of many drugs, but has also proved ineffective in many

cases over the years for a number of reasons. These include the fact that the

compounds are expensive to purchase, maintain and screen; and since a set of even

hundreds of thousands or millions of compounds in HTS may not allow for efficient

exploration of chemical space, and hence may even achieve a lower hit rate than

FBDD (Figure 1.36) (Erlanson, 2012). In contrast to the traditional high-throughput

screening, the FBDD has remarkable advantages, such as efficiently covering

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chemical space (i.e. a relatively small fragment library, usually containing 102 to 103

fragments, can occupy a significantly larger proportion of chemical space compared

with the approximately 105-106 larger molecules (Mw 300–500 Da) as would be

used in a high-throughput screen (HTS) (Hudson et al., 2014, Scott et al., 2012).

Furthermore, FBDD generates a higher hit rate than the HTS, and these fragment

hits must make reasonably strong interactions with the target in order to bind with

sufficient affinity for detection by biophysical methods (e.g. NMR- and calorimetry-

based methods), leading to higher ligand specificity detection (Wang et al., 2015,

Hudson et al., 2014). The magnitude of these interactions is expressed by hits

having high or low ligand efficiency (where ligand efficiency (LE) equals the negative

∆G of binding divided by the number of non-hydrogen atoms (NHA) in the

fragment) (Hopkins et al., 2004, Hudson et al., 2014).

X-ray crystallography plays a key role in both approaches in that it provides precise

three-dimensional detail on the molecules’ binding modes, and hence guides their

subsequent elaboration and validation (Murray and Blundell, 2010). Structure-

based drug design is used in FBDD to increase potency and selectivity, whilst

maintaining drug-like properties (Murray and Blundell, 2010).

FBDD has been applied to some of the 20 Mtb CYPs; however CYP121A1 represents

the first successful P450 enzyme to be successfully studied via fragment-based

approaches (Hudson et al., 2012b). An initial fragment-screening process involving

thermal shift and NMR spectroscopy (for detection of fragment binding), and X-ray

crystallography (to pinpoint fragment binding positions in CYP121A1) identified four

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fragments that bound within the CYP121A1 active site, including one that bound in

two overlapping orientations. A direct fragment–fragment merging process was

carried out, leading to the discovery of a novel type-II aminoquinoline inhibitor with

high ligand efficiency and affinity (Figure 1.35) (Hudson et al., 2012b).

Figure 1.35: Application of a fragment-based drug discovery approach to Mtb CYP121. A: Identical CYP121 fragments (a non-heme-coordinating triazolylphenol fragment) binding in overlapping orientations (PDB code 4G47) (Hudson et al., 2013). B: The fragments from A chemically “merged”, producing a molecule with slightly lower affinity than the parent fragments (PDB code 4G2G). C: Further optimization of the merged fragment with the addition of a primary amine group gives a compound with improved affinity and ligand efficiency (PDB code 4KTF) (Hudson et al., 2013). For all figures, the parent fragments/merged/optimised hits are shown in sticks with carbon atoms in yellow, oxygen atoms in red and nitrogen atoms in blue. The heme is shown in atom coloured sticks with carbon atoms in magenta. Structures were drawn using PyMol (DeLano, 2002).

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1.6.2 High Throughput Screening (HTS)

High-throughput screening is a large scale approach to drug discovery. It was first

used in the pharmaceutical industry in the early 1990’s, but has become widely

used in other industries with improved technologies allowing the screening of the

screening of million-compound libraries within 2-3 months (Flotow, 2014). The data

generated from HTS have consistently increased over the years and several drugs

initially identified from HTS approaches are currently in therapeutic use for different

ailments (Macarron et al., 2011).

The rate for HTS success is generally estimated to be ~50% which could b attributed

to is short fall of not allowing efficient exploration of chemical space (Figue 1.36)

(Fox et al., 2006, Bleicher et al., 2003, Erlanson, 2012). However, it has been

documented that other lead discovery strategies (such as fragment screening,

structure-based design or virtual screening) have shown higher success rates than

HTS. Nevertheless, no hit identification method is 100% fruitful. The methods all

have their advantages and disadvantages, but when coupled with good integration

of hit identification approaches the chances of success can be maximised (Fox et al.,

2006). An important advantage of HTS over other screening approaches is that HTS

can be applied to a broader range of targets and/or ligands (Macarron et al., 2011).

Unfortunately, many difficult targets, especially those with inaccessible binding

pockets, have shown low success rates using HTS (Macarron et al., 2011). Another

advantage of HTS is its inherent ability to identify compounds that modulate

biological activity (for example, cell viability, protein translocation or second

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messenger pathway monitoring) without the need for prior knowledge of the mode

of action or the characteristics of the drug target (Gao et al., 2010).

Examples of drugs identified via HTS include maraviroc (Selzentry/Celsentri from

Pfizer), an anti-retroviral drug. The journey to maraviroc discovery began with the

screening of the Pfizer library (~500,000 compounds) in 1997 and ended with the

US Food and Drug Administration (FDA) approval of maraviroc in 2007 (Macarron et

al., 2011). The HTS done using the CC-chemokine receptor 5 (CCR5; also known as

MIP1) and a radio-ligand binding assay resulted in the identification of a weak

agonist hit that had no cellular antiviral activity, but gave a good starting point for

an extensive structure–activity relationship (SAR) study (Lemm et al., 2010).

Another example is the discovery of Hepatitis C virus NS5A inhibitors. The HTS

approach here was used to identify compounds that inhibited hepatitis C virus

replication and further optimization of these hits resulted in highly effective clinical

candidates (Lemm et al., 2010, Gao et al., 2010).

HTS screening on Mtb CYP130A1 identified some specific heme-coordinating,

inhibitor-like molecules with no substrate-like molecules discovered so far (Figure

1.36). These inhibitor-like molecules are mostly heterocyclic arylamines which have

been shown to cause toxicity issues. This property has, to date, deterred their

exploitation as antibiotics (Podust et al., 2009, Kim and Guengerich, 2005).

Furthermore, HTS has also been carried out on other Mtb P450s, including

CYP121A1 and CYP126A1, with some of the crystal structures solved (Kirsty Mclean

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et al., University of Manchester, unpublished data). The aim here is to find potent

and highly selective inhibitors of Mtb P450 enzymes in addition to identifying

substrates for the orphan P450s which would help to define their mechanisms and

physiological roles.

Figure 1.36: A schematic representation of the High Throughput Screening (HTS) approach. Large numbers of compounds (many of them quite bulky) are screened against the target of interest, leading to identification of hit compounds. In most cases HTS hits do not make optimal interactions with the target, and further elaboration of these hits is needed to produce potent and selective drugs. The image was drawn using Microsoft PowerPoint and adapted from (Erlanson, 2012).

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Figure 1.37: Structures of CYP130A1 with HTS hits (heterocyclic arylamines) bound in the active site. A: two molecules of “compound 4” (1-(p-tolyl)-1H-benzo[d]imidazol-5-amine) are shown with carbon atoms in yellow and pink in stick representation (PDB code 2WHF). B: compound 2 (5-amino-2-(4-((4-aminophenyl)thio)phenyl)isoindoline-1,3-dione) is shown in sticks with carbon atoms in cyan (PDB code 2WH8). In both cases, the protein backbone is shown as purple-blue ribbon and the heme is shown with carbon atoms in red (Podust et al., 2009). Structures were drawn using PyMol (DeLano, 2002).

1.7 Justification of Research

Research on tuberculosis has been boosted by the determination of the Mtb H37Rv

genome sequence in 1998 (Cole et al., 1998), which provided a major steps towards

identifying new drug targets and in understanding the complex biology of the Mtb

bacterium (Driscoll et al., 2010). The genome sequence of Mtb H37Rv revealed a

large number of cytochrome P450 enzymes. There are 20 CYP genes in Mtb,

suggesting important physiological roles (McLean et al., 2006b). In addition, Mtb

was also shown to encode the first prokaryotic 14α-sterol demethylase (CYP51B1),

which was characterized in the Munro group (Dunford et al., 2007, McLean et al.,

2006b). An important question was raised as to whether the azole drugs (e.g.

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clotrimazole, fluconazole and miconazole, which are known anti-fungals targeting

the CYP51s) could also have activity against Mtb (Driscoll et al., 2010, McLean et al.,

2006b). The results obtained were positive, suggesting the possibility that one or

more of the Mtb P450s might be novel drug targets in the bacterium (McLean et al.,

2007a, McLean and Munro, 2008, McLean et al., 2002b).

Research in recent years has led to the expression and structural/biochemical

characterization for a number of the Mtb P450 enzymes, and this has been

complemented by genetic studies that have highlighted absolute or conditional CYP

gene essentiality in some cases. In particular, previously unexpected functions in

sterol demethylation (CYP51B1), cholesterol oxidation (CYPs 124A1, 125A1, 142A1),

novel secondary metabolite synthesis (CYP121A1) and probably in menaquinone

oxidation (CYP128A1) have emerged, all suggesting important cellular functions for

the Mtb P450s that further highlight their potential as targets for therapeutic

intervention (McLean and Munro, 2008, Munro et al., 2003, Ouellet et al., 2010b).

Recent studies have also led to some exciting results with respect to the use of

fragment based screening for targeting the Mtb P450s, with work at Manchester

and at the University of Cambridge producing novel inhibitors of the cyclodipeptide

(cyclo-L-Tyr-L-Tyr) oxidase CYP121A1 (Hudson et al., 2012b, Hudson et al., 2013). In

the case of CYP142A1 (and also for CYPs 125A1/124A1), there is strong evidence

that inhibiting their activities could be important in the killing of latent Mtb - where

metabolism of host cholesterol is essential for the bacterium to survive in the host

(Pandey and Sassetti, 2008, Chang et al., 2009, Yam et al., 2009, Ouellet et al., 2011,

Munoz et al., 2009) (also see section 1.5.2.4). The efficient inhibition of CYP124A1,

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CYP125A1 and CYP142A1 would completely prevent cholesterol/cholestenone

oxidation by Mtb and likely prevent the bacterium infecting the host macrophage.

Hence, there is a compelling need for this study, in which I have researched further

into the structure, function and drug targeting of the cholesterol oxidases in the

human pathogen Mycobacterium tuberculosis.

1.8 Aims of Research

In this PhD thesis, my work has focused on structure-guided approaches for drug

targeting and selective inhibition of the Mtb cholesterol oxidases.

The key objectives for this thesis include:

Production and biochemical/biophysical characterization of the Mtb

cholesterol oxidases CYP124A1 and CYP142A1.

Determination of crystal structures of the cholesterol oxidases in complex

with substrate and inhibitor molecules in order to rationalise their binding

modes and to analyse molecular determinants of inhibitor binding.

Identification and evaluation of specific inhibitors for the Mtb cholesterol

oxidases using fragment-based screening approaches in collaboration with

researchers at the University of Cambridge.

Provision of novel data on the biochemical and biophysical properties of the

Mtb cholesterol oxidase enzymes.

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Chapter 2

Materials and Methods

2.1 Materials

E. coli growth media were obtained from ForMedium (Norfolk, UK). Competent

cells of NovaBlue and C41 (DE3) were from Novagen (UK). Protein marker,

restriction endonucleases and other DNA modifying enzymes were obtained from

New England Biolabs (Herts, UK) and were used according to the manufacturer’s

specifications and with the commercial buffer system supplied. NADPH was

obtained from Europa Bioproducts Ltd (Cambridge, UK). All other antibiotics and

other chemicals were either from MP Biomedicals (Cambridge, UK) or Sigma-Aldrich

(Poole, UK) and were of the highest grade available, unless otherwise stated.

2.2 Methods

2.2.1 Preparation of Plasmid DNA for Expression Constructs

2.2.1.1 Source and Description

CYP142A1 (Gene Rv3518)

The CYP142A1 (gene Rv3518 from the Mtb H37Rv genome) from a Mycobacterium

tuberculosis H37Rv cosmid DNA library (from Dr. Roland Brosch, Institut Pasteur,

Paris) was transformed into E. coli Novablue competent cells (Novagen, Darmstadt,

Germany).

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The Rv3518 gene was expressed using the previously cloned Rv3518 construct in

the pET15b plasmid vector (Novagen), with the gene having been cloned from a

Mtb H37Rv chromosomal cosmid library (BAC clone Rv416) using the forward

primer, upstream (CYP142-NdeF), 5’-

GGAGGATCCATATGACTGAAGCTCCGGACGTGG-3’, and reverse primer, downstream

(CYP142-BamHIR), 5’-CGTTCGGGATCCCTCAGCCCAGCGGCGTGAAC-3’. The letters

underlined in the upstream primer indicate an engineered NdeI restriction-cloning

site, including the initiation codon ATG (bold). The underlined letters in the

downstream primer indicate a BamHI restriction-cloning site, with the stop codon in

bold (Driscoll et al., 2010).

CYP124A1 (Gene Rv2266)

The CYP124A1 (Rv2266) gene was synthesized by Genscript (Piscataway, USA)

following codon optimization for expression in E. coli, and was cloned into pET47b

using the restriction enzymes sites BamHI and HindIII by the supplier.

2.2.1.2 Plasmid DNA Purification

Plasmid DNA for both CYP142A1 and CYP124A1 were purified using standard

protocols. Competent cells of Novablue strains of E. coli were first transformed with

the plasmid DNA and transformants were grown in culture. Subsequently, plasmids

were purified from these cultures using a QIAGEN miniprep kit and the supplier’s

protocol (Qiagen, Manchester UK).

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Ultra-competent E. coli Novablue cells (Novagen, UK) were taken from a -80oC

freezer and used to inoculate 5 ml of LB (Luria Bertani) medium containing a

selective antibiotic specific for Novablue cells (tetracycline at 12.5 g/ml). This

culture was grown at 37oC at 190 rpm overnight. Working cultures were then grown

from the starter culture to an optical density of 0.4 (OD600 = 0.4) and cells were

harvested by centrifugation in a microfuge tube for 30 s and then resuspended in

50 mM calcium chloride twice (i.e. re-centrifuged and resuspended in 0.5 mL of

fresh 50 mM CaCl2), and then left on ice for 60 minutes. 1 μl of the

CYP142A1/pET15b plasmid (40.2 µg/ml) or the CYP124A1/pET47b (84.1 µg/ml) was

added and the mixture left on ice for another 60 minutes before being subjected to

a heat shock treatment at 42oC for one minute, and then returned to ice for

another 2 minutes. Sterile SOC medium (0.8 ml) was then added and the mixture

left to incubate at 37oC for 60 minutes in a shaking incubator. 100 μl of each E. coli

strain were then plated onto LB agar plates containing 50 μg/ml carbenicillin (for

CYP142A1/pET15b plasmid) or 30 μg/ml kanamycin (for CYP124A1/pET47b) and

incubated overnight at 37oC. The following morning, a single colony was picked

from each plate and used to inoculate LB starter cultures, and these cells were

grown overnight in 5 ml LB medium containing 50 μg/ml carbenicillin (for

CYP142A1) or 30 μg/ml kanamycin (for CYP124A1) at 37oC with agitation at 190

RPM. DNA was extracted using a QIAGEN miniprep kit and protocol (Qiagen,

Manchester UK). The concentration of plasmid DNA used was determined using a

NanoDrop 2000 instrument (Thermo Scientific, Wilmington USA) and the stock

stored in a labelled, sterile Eppendorf tube at -80oC. The constructs used were all

checked for correct CYP gene insertion using 0.8% agarose Tris-Acetate EDTA (TAE)

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gel electrophoresis with ethidium bromide (0.2 µg/ml), with the DNA resolved by

running the gel at 90 V and by analysis of both undigested plasmid DNA, and

plasmids digested with BamHI and NdeI. To verify that the entire gene was correctly

inserted, a sample was also sent to Source Bioscience (Nottingham, UK) for

sequencing with the T7 primers: T7F - TAA TAC GAC TCA CTA TAG GG and T7R - GCT

AGT TAT TGC TCA GCG G. These data confirmed that the correct genes were

present.

2.2.2 Generation of Glycerol Stocks of E. coli transformants Glycerol stocks of transformant E. coli cells were prepared in order to preserve the

plasmid DNA by storage in expression strains of E. coli. In these cases, 5 ml cultures

of transformed E. coli cells (C41 (DE3)) were grown overnight in medium containing

relevant antibiotics specific for the plasmids. The next day, a freshly re-inoculated

culture in the same medium was grown until an OD600 = 0.4-0.6. Subsequently, 800

µl of cell culture and 200 µl of sterilized 80% glycerol were mixed gently in an

Eppendorf tube and the transformant cell sample was stored at -80°C until

required.

2.2.3 Expression Trials for CYP124A1 and CYP142A1 P450s

E. coli transformant expression trials for the CYP142A1 and CYP124A1 genes were

carried out in 50 ml volumes of 2YT medium supplemented with ampicillin (50

µg/ml) or kanamycin (30 μg/ml) (for CYPs 142A1 and 124A1, respectively) in 250 ml

flasks using various media (Luria-Bertani (LB), 2x yeast tryptone (2YT) and terrific

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broth (TB)) with different IPTG concentrations (0.1-1.0 mM) and expression

temperatures (18-25°C), both with/without addition of the heme precursor -

aminolevulinic acid (0.1-0.5 mM δ-ALA), using different E. coli expression hosts

(HMS174 (DE3), C41 (DE3), BL21 (DE3) and Rosetta (DE3) Novagen). The CYP142A1

gene was expressed under control of a T7 RNA polymerase/promoter system in

pET15b while CYP124A1 gene was expressed in pET47b using different DE3 lysogen

E. coli strains. Transformed cells were grown at 37°C overnight in LB medium (5 ml)

supplemented with antibiotics selective for each gene, and this culture was used as

an inoculum for 250 ml 2YT medium with the same additives.

In the 250 ml flasks, cells were grown at 37°C for 4 hours, and the growth

temperature was then reduced to 18-25°C prior to a further 4-48 hours growth

following IPTG (0.1-1.0 mM) addition to induce target gene expression, and the

addition of 0.1-0.5 mM δ-ALA at an OD600 of 0.6–0.8. Subsequently, 1 ml samples

were collected and centrifuged at 13,000 rpm for 10 min and 4°C on a benchtop

microfuge (MicrofugeR 22R centrifuge, Beckman Coulter) and the supernatant was

then discarded and the pellet resuspended in 200 μl 50 mM KPi, 250 mM KCl, 10%

glycerol, pH 8 buffer. 12 μl of the normalised cell resuspension was heated with 6 μl

of 3x SDS sample buffer at 95°C for 3 minutes and analysed by SDS-polyacrylamide

gel electrophoresis (SDS-PAGE). Normalised loading volumes for SDS-PAGE analysis

were calculated using the worksheet provided in the pET System Manual

(Novagen). 10 μl of protein marker, broad range (2-212 kDa) (New England Biolabs,

Hitchin UK) was run alongside the samples to determine the relevant P450 protein

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band mass, and (unless stated otherwise) the same protein marker was used in all

SDS-PAGE gels.

Medium type Composition Amount added

LB 10 g/l Tryptone 5 g/l Yeast Extract 10 g/l NaCl pH 7.0

25 g/l (premixed)

TB 12 g/lTryptone 24 g/l Yeast extract 4 ml glycerol 72 mM K2HPO4, 17 mM KH2PO4, pH 7.0

36 g/l (premixed)

2YT 16 g/l Bacto Tryptone 10 g/l Bacto Yeast Extract 5 g/l NaCl pH to 7.0 with 5N NaOH

31 g/l (premixed)

SOC 20 g/l Tryptone 5 g/l Yeast Extract 10 mM NaCl 2.5 mM KCl 10 mM MgCl2 10 mM MgSO4 20 mM glucose pH 7.0

800 μl premixed SOC added to final volume of 500 μl re-suspended cells.

Table 2.1 Composition of growth media used for protein production and the amounts of components added per litre of medium. All media were sterilised by autoclaving prior to use.

2.2.4 Scale up of the Expression of CYP142A1

From the protein expression trials carried out, the E. coli strain C41 (DE3) and

growth in 2YT medium was observed to produce the greatest proportion and

quantity of soluble CYP142A1 protein. Hence, further expression (to enable P450

purification) was carried out on a larger scale using 2 litre flasks containing 600 ml

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of 2YT medium. The E. coli strain C41 (DE3) was transformed (as described in

section 2.2.1.2) with the pET15b-CYP142A1 plasmid (hereafter referred to as

pCYP142A1). Single colonies were taken and inoculated into 5 ml LB medium

containing 50 μg/ml carbenicillin and cultures were grown for six hours. 0.5 ml of

the starter culture was then inoculated into 200 ml LB containing the same

antibiotics and culture was continued at 37°C. For protein expression, the

transformed cells were grown in 24 x 2 litre flask cultures in 2xYT medium. Each 2

litre flask contained 600 ml of growth medium supplemented with ampicillin (50

µg/ml), and was inoculated with 6 ml of bacteria from an overnight culture in the

same medium. Cells were then grown at 37°C with agitation (200 rpm) to an OD600

of 0.4; the temperature was then reduced to 22°C and the cultures were grown to

an OD600 of 0.8. Expression was induced by addition of IPTG (0.1 mM) and culture

was continued for a further 20–24 h. The cells were harvested by centrifugation at

6000 g for 10 min at 4°C and stored at -80°C until required.

2.2.5 Scale up of the Expression of CYP124A1

CYP124A1 was expressed using a similar protocol to that for CYP142A1, with some

minor differences. The C41 (DE3) E. coli strain and 2YT medium combination was

found to be best for protein expression. E. coli C41 (DE3) was transformed (as

described in section 2.2.1.2) with the pET47b-CYP124A1 plasmid (hereafter referred

to as pCYP124A1).

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Each 2 litre flask contained 600 ml of growth medium supplemented with

kanamycin (30 µg/ml), and was inoculated with 6 ml of bacteria from an overnight

culture in the same medium. Cells were then grown at 37°C with agitation (210

rpm) to an OD600 of 0.3; the temperature was then reduced to 25°C and the

cultures were grown to an OD600 of 0.6. Expression was induced by addition of IPTG

(0.5 mM) with addition of 0.5 mM δ-ALA heme precursor. Bacterial cell culture was

continued for a further 30-36 h. The cells were harvested by centrifugation at 6000

g for 15 min at 4°C and the cell pellet stored at -80°C until required.

2.2.6 Protein Purification for CYP124A1 and CYP142A1

The same purification protocol was used for both CYP142A1/CYP124A1. Cell pellets

from approximately 15 litres of culture were resuspended in approximately 400 ml

ice cold 50 mM potassium phosphate (KPi, pH 8.0) containing 250 mM KCl and 10%

glycerol (resuspension buffer). DNase (10 mg/ml from a standardized vial

containing 2,000 Kunitz units of DNase I), lysozyme (10 mg/ml) and 2.5 mM MgCl2

were added, and eight complete EDTA-free protease inhibitor tablet (Roche) were

added while stirring on a magnetic stirrer.

Ultrasonication was carried out with ~15 cycles of 20 seconds on ice with 1 minute

rest periods, using a Bandelin Sonopuls sonicator at 45% full power. The lysate was

centrifuged at 40,000 g for 1 h at 4°C to pellet insoluble material. The supernatant

was retained and loaded onto a nickel-nitrilotriacetic acid (Ni-NTA) column (Qiagen)

for protein purification. The Ni-NTA column was pre-equilibrated with 50 mM KPi

buffer (pH 8.0) containing 250 mM KCl and 10% glycerol (binding buffer), and the

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supernatant sample allowed to flow through the column under gravity. Flow-

through from the column was collected and analysed by SDS-PAGE. The column was

washed with 10 column volumes of resuspension buffer plus 5 mM imidazole.

Successive washes were then performed with loading buffer containing imidazole

at 20 mM, 40 mM, 60 mM and finally 100 mM. The eluent after each wash was

collected and analysed spectrally (between 240 nm and 800 nm, analysing for P450

heme signals) using a Cary 50 UV-visible scanning spectrophotometer (Varian, UK).

CYP142A1 purity was also established using its Reinheitszahl (Rz) value — i.e. the

ratio of absorbance at the heme Soret peak to the protein absorbance at 280 nm

(A418/A280). Fractions containing CYP142A1 (mainly from the 40-100 mM imidazole

fractions) were pooled, concentrated by ultrafiltration using a Vivaspin 20 ml

ultrafiltration device (Sartorius, 30 kDa MWCO) by centrifugation at 3500 rpm using

a bench top centrifuge (MicrofugeR 22R centrifuge, as before), and then dialysed

into 15 mM KPi (pH 6.5) to remove imidazole (which can act as a ligand to the P450

heme iron).

The pooled samples from the Ni-NTA column were further concentrated to ~5 ml by

ultrafiltration as described above. Further purification was then done using a

hydroxyapatite (HA) column on an AKTA purifier. The purification was done using a

linear gradient of 15 mM KPi, pH 6.5 (Buffer A) and 500 mM KPi pH 6.5 (Buffer B)

over a range of 350 to 500 ml. Fractions from the HA column were analysed by SDS-

PAGE using 12% precast gels (Expedeon, Cambridge UK) and by UV–Vis

spectroscopic analysis. The purest fractions from the HA column purification

containing P450s with A418/A280 ratios >1.0 (for CYP124A1) and >1.5 (for CYP142A1)

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were pooled. These were then concentrated to ~50 μl by ultrafiltration using a

Vivaspin 2 ml ultrafiltration device (Sartorius, 30 kDa MWCO) at ∼1500 g, and then

subjected to a final purification step on a SuperdexTM S-200 gel (120 ml S200

16/600GL, GE Healthcare) gel filtration column on an AKTA purifier using 10 mM

Tris plus 150 mM NaCl, pH 7.2 as the loading buffer. For CYP142A1 purification, this

buffer was supplemented with 1 mM DTT (dithiothreitol, Melford, Ipswich UK).

Fractions were analysed by SDS-PAGE as described in section 2.2.6. Purity was

confirmed by a single band present on an SDS-polyacrylamide gel and by UV-visible

spectroscopy, where an A418/A280 ratio ≥2 (for CYP142A1) and ≥1.3 (for CYP124A1)

correlated with highly pure enzyme.

2.2.7 Assessment of P450 Concentration and Purity

Protein purity was assessed by SDS-PAGE on 12% pre-cast SDS-PAGE acrylamide

gels (Biorad, UK) run at 350 V for 20 minutes according to the manufacturer’s

protocol. CYP142A1 purity was also verified using its Rz value (A418/A280). Protein

samples (20 μl) were mixed with 20 μl of 2x SDS protein sample buffer/loading

buffer (40% glycerol, 240 mM Tris/HCl pH 6.8, 8% SDS, 0.04% bromophenol blue,

5% beta-mercaptoethanol). The mixture was heated at 95-100°C for 5 minutes to

denature proteins prior to SDS-PAGE. For each well on the gels, 10 μl of protein-

buffer mixture was loaded and the gel cassette placed in the electrophoresis tank.

The samples were then run in 0.1% SDS, 1x running buffer (containing 28.8 g

glycine, 6.04 g Tris base, 2 g SDS, and deionized, distilled water [ddH2O] to a final

volume of 2 litres). Electrophoresis was performed at 350 V for 20 minutes. After

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the completion of electrophoresis, the SDS-PAGE gel was removed from the tank

and a gel image picture taken with a BioRad Bioimaging instrument (BioRad, UK).

2.2.8 Determination of P450 Extinction Coefficients using the Pyridine Hemochromagen Method

Determination of the Soret extinction coefficient for CYP142A1 was done by the

pyridine hemochromogen method (Berry and Trumpower, 1987). UV–visible

absorbance spectra were recorded for CYP142A1/CYP124A1 solutions (using 5.5

μM enzyme for CYP142A1/5.8 μM enzyme for CYP124A1) in a 1 ml quartz cuvette,

and then 500 µl was removed and retained for later use. An equal volume (500 µl)

of a 40% pyridine stock solution (containing 0.8 mM potassium ferricyanide and 200

mM NaOH) was added and the hemochrome spectrum taken afterwards.

Further spectral changes were recorded after addition of a few grains of sodium

dithionite and until no further spectral changes occurred and peak formation

between 550-570 nm was consistent. The CYP142A1/124A1 concentrations were

calculated using the difference in absorbance at 557 nm between the oxidized and

reduced spectra and by employing a difference extinction coefficient at this

wavelength of ∆A557 = 23.98 mM−1 cm−1.

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2.2.9 UV-Visible Spectroscopic Studies of Mtb P450s

2.2.9.1 Binding Assays with Substrates and Inhibitors

All ligand binding (for potential substrates and inhibitors) assays were performed by

spectrophotometric titration at 25oC in 100 mM potassium phosphate (KPi), 100

mM KCl buffer, (pH 7.5 for CYP142A1 and pH 7.0 for CYP124A1) using a Cary UV-

visible scanning spectrophotometer (Varian UK) and a 1 cm path length quartz

cuvette, recording spectra between 250 and 800 nm, and typically using a protein

concentration between 2-10 μM. Imidazole and cyanide stocks were made up in the

same buffer, while other azole drugs (clotrimazole, econazole, fluconazole,

bifonazole, ketoconazole, miconazole, and voriconazole) were prepared in

dimethylsulfoxide (DMSO). Cholesterol and cholestenone solutions were made up

in 45% 2-hydroxypropyl-β-cyclodextrin (HPCD, in water), fatty acids solutions were

made in ethanol, while a lanosterol solution was made up in a 9:1 ethanol:HPCD

mixture. Ligands were added in small volumes (typically 0.1-0.5 µl aliquots) from

concentrated stock solutions to the protein in a 1 ml final volume. Spectral

measurements were taken before ligand addition, and following addition of each

aliquot of ligand. The Kd values for each ligand were determined by plotting the

optical change induced after each addition of ligand against the relevant ligand

concentration, and by fitting the data using either the Morrison equation (Equation

1), a standard hyberbolic function (Equation 2) or the Hill equation (equation 3)

using Origin software (OriginLab, Northampton, MA). Equation 1 provides robust

fitting of binding data for tight binding ligands, accounting for the concentration of

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protein in cases where the Kd value is not substantially greater than the protein

concentration used.

Aobs = (Amax/(2Et) × ((S+Et+Kd)-((S+Et+Kd)2-(4EtS))0.5) (Equation 1)

In Equation 1 (the Morrison equation), Aobs is the observed absorbance change at

ligand concentration S; Amax is the absorbance change at ligand saturation; Et is the

enzyme concentration, and Kd is the dissociation constant for the enzyme-ligand

complex.

Aobs = (Amax*S/(Kd+S)) (Equation 2)

In Equation 2 (the standard hyperbolic function, essentially the Michaelis-Menten

function adapted for ligand binding), Aobs is the observed absorbance change at

ligand concentration S, Amax is the maximal absorbance change observed at ligand

saturation, and Kd is dissociation constant for the binding of the ligand (the

substrate concentration at which Aobs = 0.5 x Amax).

Aobs = (Amax × Sn)/(Kn+Sn) (Equation 3)

In equation 3 (the Hill equation), Aobs is the observed absorbance change at ligand

concentration S, Amax is the absorbance change at ligand saturation, K is the

apparent dissociation constant, and n is the Hill coefficient, a value describing the

apparent extent of cooperativity observed in ligand binding.

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2.2.9.2 Formation of P450 Carbon Monoxide and Nitric Oxide Adducts

Conversion of the enzyme Fe(II)-CO complex from the “native” (cysteine thiolate-

coordinated) form with a Soret band maximum close to 450 nm (P450) to the

“inactive” form with a maximum near 420 nm (P420, with cysteine thiol

coordination) was monitored spectrophotometrically in the presence and absence

of cholesterol. This was done using a Cary 50 UV visible spectrophotometer under

anaerobic conditions in a glove box (Belle Technology, Weymouth, UK) for ferrous

CO-bound enzymes (McLean et al., 2008, Quaroni et al., 2004).

Solutions of CYP142A1/CYP124A1 (2-6 µM) were prepared in 100 mM KPi

containing 100 mM KCl (pH 7.5 for CYP142A1 and pH 7.0 for CYP124A1). UV/Visible

absorption spectra were recorded on a Cary UV-50 Bio UV/Visible scanning

spectrophotometer using a sealed 1 cm pathlength cuvette with anaerobic buffer at

25°C. Spectra were first recorded for the oxidized species, and subsequently

CYP142A1 (4.4 µM) and CYP124A1 (3.8 µM) were reduced using a small amount of

sodium dithionite and bubbled briefly with carbon monoxide to form the P450-CO

complex. Spectra were then recorded every 10 s for several hours to observe any

changes relating to the P450 “collapse” into the P420 form due to cysteine thiolate

protonation. The stability of the enzyme-CO complex was also examined in the

same way in the presence of cholestenone (1 μM sterol).

Nitric oxide (NO) complexes of CYP142A1 and CYP124A1 were obtained by brief

bubbling of the buffered oxidized enzyme solution with NO gas. This experiment

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was performed with caution to avoid excessive addition of NO given its ability to

form nitrous acid in solution which would lead to protein denaturation. Spectra

were recorded as described above.

2.2.10 Isothermal Titration Calorimetry (ITC) Studies on Mtb

P450s

Experiments were performed using a VP-ITC calorimeter equipped with the control

and data acquisition/analysis software ORIGIN 7 (MICROCAL Inc., Northampton,

MA). Solutions of the protein and fragments used for ITC titrations were prepared

in 100 mM KPi, 100 mM KCl, pH 7.5 containing 5% DMSO. The protein solution

(CYP142A1 at 62 µM) was placed in the calorimetric cell and titrated with different

concentrations of the fragments in the titration syringe. Twenty injections of 15 l

aliquots of fragments (at different concentrations) were run at 300 seconds

intervals. The titration syringe was continuously stirred at 310 rpm, and the

temperature of the calorimetric cell maintained at 25oC. Injecting the ligand into

buffer alone was also carried out as a reference titration and the resulting heat of

dilution subtracted from the protein-fragment titration. Data were generated by

fitting to a single binding model. ∆G was then determined by the relationship:

∆G=∆H-T∆S (Equation 4)

Where T is the temperature of the experiment in Kelvin (oK = oC + 273.15), ∆G is

Gibbs free energy of reaction, ∆H is the change in enthalpy of the system and ∆S is

the change in entropy of the system.

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2.2.11 Guanidinium Chloride Denaturation of CYP142A1

Aromatic amino acid (mainly tryptophan) fluorescence measurements of pure

CYP142A1 (5 µM) was carried out by incubating the P450 with increasing

concentrations of guanidinium chloride (GdmCl, 0-6 M) for 30 minutes (i.e. 30 min

incubation period at each [GdmCl]) in the presence and absence of cholestenone.

Fluorescence was measured using a fluorescence spectrophotometer (Varian Cary

Eclipse instrument) at a temperature at 25oC in 100 mM KPi, 100 mM KCl, pH 7.5.

To measure the tryptophan fluorescence, the excitation wavelength was 280 nm

and emission data were collected from 290 to 470 nm. Slit widths for excitation and

emission monochromators were set at 5 nm each. The signal produced by a buffer

solution containing the applied concentrations of GdmHCl without the enzyme was

subtracted from the data sets containing P450 protein, to eliminate background

emission.

The spectra collected were analysed using OriginLab software. The observed

fluorescence changes were plotted against the applied GdmCl concentration. Data

were fitted using a sigmoidal function to determine the midpoint GdmCl

concentration required for 50% loss of protein tertiary structure.

2.2.12 Redox Potentiometry Studies on CYP124A1 and CYP142A1

Redox potentiometry was carried out in a Belle Technology glove box (Weymouth,

UK) under anaerobic conditions in a nitrogen atmosphere (<2 ppm oxygen). Oxygen

was removed from buffers and solutions by bubbling with O2-free nitrogen prior to

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the experiments. CYP142A1/CYP124A1 enzymes (approx. 700 µM) were applied to

a BioRad PD-10 (GE Healthcare) desalting column in the anaerobic box, pre-

equilibrated with degassed 100 mM KPi, 200 mM KCl, 10% glycerol, pH 7.5 (titration

buffer) to remove oxygen. Titrations were carried out electrochemically in a 5 ml

final volume reaction containing 5-8 µM enzyme using sodium dithionite as the

reductant and potassium ferricyanide as the oxidant in the presence of redox

mediators (2 µM phenazine methosulfate [PMS], 7 μM hydroxynaphthoquinone

[HNQ], 0.3 μM methyl viologen [MV] and 1 μM benzyl viologen [BV]). The enzyme

sample was then left to equilibrate for ~10 minutes after each addition of reductant

before the spectral readings were taken along with the corresponding reduction

potential. Where applicable, ligands were added into the reaction at a saturating

concentration, but avoiding excess additions that might cause protein precipitation

and turbidity. This experiment was performed at 25°C according to the method of

Dutton (Dutton, 1978). Changes in absorbance at the heme Soret peak were plotted

against applied potential, corrected for the potential of the electrode used

(Ag/AgCl) against the standard hydrogen electrode (SHE) as E0 = +220 mV. All data

were fitted using the Nernst equation for the midpoint potential determination for

the heme Fe3+/Fe2+ transition. Data manipulation and analysis were performed

using Origin software (OriginLab, Northampton MA, USA).

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𝑨 = (𝑨𝒂𝒃𝒔 + 𝑩𝒂𝒃𝒔 ∗ ((𝑬𝟎−𝑬)/𝑹𝑻𝑭))/(𝟏 + 𝟏𝟎((𝑬𝟎−𝑬)/𝑹𝑻𝑭)) (Equation 5)

Nernst equation for a 1-electron redox reaction: A – absorption of the analyte

observed at a given potential, Aabs – absorbance of oxidised P450, Babs – absorbance

of reduced P450, E0 –standard midpoint redox potential of the P450 heme iron

FeIII/FeII transition (or other relevant redox process), E – applied electrode potential

corrected against the normal hydrogen electrode (NHE), RTF – a compound

parameter of the universal gas constant (R), absolute temperature of the system (T)

and the Faraday constant (F).

2.2.13 Multi-Angle Laser Light Scattering (MALLS) Studies of Mtb P450s

MALLS analysis was carried out for CYP142A1 and CYP124A1 to estimate the

molecular weight and homogeneity of the protein. Light scattering data were

collected using a DAWN HELEOS-II laser photometer (laser wavelength 658 nm,

Wyatt, USA) and an Optilab rEX refractometer (Wyatt, USA) with a QELS dynamic

light scattering attachment, following an integrated Superdex 200 gel filtration step

(24 ml S200 10/300GL, GE Healthcare) at a flow rate of approximately 0.75 ml/min.

0.2 ml of 2.5 mg P450 enzyme was run in 10 mM Tris, 150 mM NaCl, pH 7.2. For

CYP142A1, the experiment was done with buffers containing no salt, with 150 mM

NaCl and with 300 mM NaCl, and in the presence and absence of DTT (1 mM). Data

were collected using a k5 cell type and a laser wavelength of 690 nm. Light

147

scattering intensity and eluant refractive index (concentration) were analysed using

ASTRA v5.3.4.13 software to give a weight-averaged molecular mass (MW). MALLS

experiments were carried out by Mrs Marjorie Howard at the Biomolecular

Interactions Facility in the Faculty of Life Sciences, University of Manchester, UK.

2.2.14 Differential Scanning Calorimetry Analysis of Mtb P450s

Differential scanning calorimetry (DSC) studies were performed on a Microcal VP-

DSC calorimeter (MicroCal Inc., Amherst MA, USA). Protein samples of CYP142A1

and CYP124A1 (8 µM) in 100 mM NaCl, 10 mM KPi (pH 7.2) were used for the

experiment for both ligand-free and ligand-bound samples. The buffer (filtered and

degassed) was used to baseline the equipment prior to the actual experiment. The

thermal transitions occurring during the unfolding of the P450 proteins were

recorded between 20-90oC at a 90oC/hour scan rate (10 min pre-scan thermostat).

Data were collected in repeated scans until a consistent reading was obtained.

Data collected were fitted using DSC OriginLab software (Microcal) to determine

melting temperatures or transition midpoints (Tm values), and enthalpy or

calorimetric heat change (ΔH) and van’t Hoff heat change (ΔHvH) values for protein

unfolding transitions.

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2.2.15 Electron Paramagnetic Resonance (EPR) Spectroscopy of P450s

EPR data for the ligand-free and ligand-bound forms of CYP142A1/CYP124A1 were

obtained using a Bruker ELEXSYS E500/E580 EPR spectrometer (Bruker GmbH,

Rheinstetten, Germany) fitted with an ESR-9 liquid helium flow cryostat (Oxford

Instruments). Spectra were recorded at 10 K with a microwave power of 2.08 mW

and modulation amplitude of 1 mT. Protein samples (200 μM) were prepared in 100

mM KPi, 100 mM KCl, pH 7.5 and ligand concentrations added were at least 10x

their Kd value plus 200 M. The final ligand concentration added was typically 450

µM dissolved in 1.7 µl solvent. The software packages supplied with the EPR

instrument were used to generate the g-values for all samples. EPR spectra were

collected by Dr. Stephen Rigby, Dr. Karl Fisher and Dr. Kirsty McLean (University of

Manchester).

2.2.16 CYP124A1 Steady-State Kinetics

Steady-state kinetic assays were carried out on a Cary UV-50 spectrophotometer

(Agilent) at 340 nm. An electron transport system was set up using CYP124A1,

spinach ferredoxin (spFDX), and E. coli flavodoxin reductase (E. coli FLDR) in the

ratio 1:10:2 (CYP:spFLD:FLDR) i.e. 200 nM CYP:2 µM spFDX:400 nM E. coli FLDR

(used for the spinach system). Another set of experiments was also performed

using CYP124A1, E. coli flavodoxin (E. coli FLD) and E. coli FLDR in the same ratio

(used for the E. coli system). This assay was performed in 1 ml assay buffer

containing a 200 nM final concentration of enzyme with varied substrate

concentrations (0-50 µM) at 25°C. Enzyme rate constants for substrate-induced

149

NADPH oxidation were determined in triplicate at each substrate (cholestenone,

phytanic acid, geraniol, farnesol, geranyl geraniol, and 15 methylpalmitic acid)

concentration. Substrate-dependent consumption of NADPH was followed by the

rate of change in absorbance at 340 nm (340 = 6.21 mM-1 cm-1) and was

monitored over 5 minutes. The data generated were plotted against the relevant

substrate concentration and fitted using the Michaelis-Menten function with Origin

software.

2.2.17 P450 Protein Crystallization and Structure Determination

Following their final size exclusion purification step, CYP142A1/CYP124A1 were

immediately buffer exchanged into 10 mM Tris, pH 7.2, using a 10 DG disposable

chromatography column (Bio-Rad), and concentrated by ultrafiltration to the

concentration required for crystallography. Crystallization trials for CYP142A1 were

carried out using varying concentrations of highly purified CYP142A1 ranging from

10-30 mg/ml in 10 mM Tris pH 7.2 for both ligand-free and ligand-bound enzyme.

The initial screening of crystallization conditions was performed by using a

Molecular Dimensions screening kit (including MorpheusTM, Clear Strategy Screen

ITM, Clear Strategy Screen IITM, PACT premierTM, and JCSG-plusTM) and a Mosquito

nanolitre pipetting robot (TTP Labtech, Melbourn UK) operating with 96-well plates,

and a sitting drop vapour diffusion protocol. Experiments were set up using 400 nl

(200 nl protein drops plus 200 nl drops of mother liquor) sitting drops and trays

were sealed with crystal clear tape (Manco® Inc. Ohio, USA) and stored in a cold

150

room at 4°C. Crystals appeared within 3-14 days from several conditions. Diffraction

quality crystals were obtained by further optimization of the initial conditions and

by micro-seeding using the protocol described by (D'Arcy et al., 2007).

For native CYP142A1, the crystals giving the best diffraction were formed under the

following conditions: 15 mg/ml protein in 0.1 M sodium acetate at a pH range of

4.7–5.0, with 0.1 M potassium thiocyanate, 8-10% PEG 200 (v/v), and 8-12% PEG

550MME (v/v). For econazole-bound CYP142A1 (15 mg/ml protein supplemented

with 1.3 mM econazole), crystals grew from 0.2 M potassium thiocyanate, 0.1 M

bis-Tris propane, pH 6.5, 20% PEG 3350. Crystals of the cholestenone-bound

CYP142A1 (15 mg/ml enzyme supplemented with 1 mM cholestenone) grew from

2.4 M ammonium sulfate, 0.1 M sodium acetate, pH 5.5. For cholestenone-bound

CYP124A1 (10 mg/ml enzyme supplemented with 1 mM cholestenone), crystals

grew from 0.3 M magnesium formate dehydrate, 0.1 M bis-Tris propane, pH 5.5.

For NMR491- and 1-phenylimidazole-bound CYP142A1 (25 mg/ml enzyme

supplemented with 2 mM ligand), crystals grew from 0.2 M magnesium chloride,

0.1 M sodium chloride (pH 5.3-5.6), 6-12% PEG 20K, 8-10% PEG 550 MME. The

NMR170-bound and NMR623-bound CYP142A1 structures were solved by back-

soaking native crystals in 24% PEG 550 MME, 0.1 M sodium acetate at pH 4.5, 0.1 M

potassium thiocyanate.

Crystals were flash-cooled in liquid nitrogen with Paratone N as cryoprotectant.

Diffraction data were collected at the Diamond Light Source (Oxford, UK) by Dr.

Colin Levy (University of Manchester). The data were scaled and integrated using

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the Xia 2 package (Kabsch, 1993). Structures were solved by molecular replacement

(McCoy et al., 2007) with the previously solved CYP142A1 crystal structure as a

search model (PDB 2XKR). The structures were built using COOT (Emsley and

Cowtan, 2004) in conjunction with MOLPROBITY (Davis et al., 2007) and refined

using Phenix (Adams et al., 2010) to resolutions of 1.70 Å (NMR170-bound

CYP142A1), 1.91 Å (NMR491-bound CYP142A1), 2.0 Å (NMR623-bound CYP142A1),

1.34 Å (1PIM-bound CYP142A1), 2.09Å (cholestenone-bound CYP142A1), 2.12 Å

(econazole-bound CYP142A1) and 2.54 Å (cholestenone-bound CYP124A1).

Diagrams and models in this thesis were made using PyMol™ (DeLano, 2002)

molecular graphics software using CCP4 mesh files and PDB files.

2.2.18 CYP142A1 Nano-ESI Mass Spectrometry

Protein stock solutions (20 μM) were prepared by dilution of purified CYP142A1

(638 μM) in 200 mM ammonium acetate buffer, pH 7.0. Samples were buffer

exchanged by size exclusion chromatography using Micro Biospin 6 columns,

molecular weight cut-off 6 kDa (BioRad, Hemel Hempstead, UK). Azole compounds

and MEK ligands were prepared as stock solutions in d6-DMSO at 0.4-2 mM

concentrations. Cholestenone (0.4-5 mM) stock solutions were prepared in

absolute ethanol and DTT (2-20 mM) was dissolved directly in 200 mM ammonium

acetate buffer, pH 7.0. Ligand-protein samples were prepared by diluting protein

stocks (10 μl) and ligand stocks (0.5 μl) with ammonium acetate buffer (9.5 μl) to

give final concentrations of 10 μM CYP142A1, 10-125 μM ligand and 2.5% v/v d6-

DMSO or ethanol. Samples containing DTT were prepared by diluting protein stocks

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(10 μl) with stocks of DTT (10 μl) to give a final concentration of 10 μM CYP142A1

and 1-10 mM DTT. Mass spectra were recorded on a Synapt HDMS instrument

(Waters UK Ltd., Manchester, UK). Capillaries for nano-ESI were purchased from

ThermoFisher (Hemel Hempstead, UK). Capillary tips were cut under a stereo

microscope to give inner diameters of 1−5 μm and then loaded with 2.5 μl of

sample solutions. Given below are the general instrumental conditions used to

acquire the reported spectra. However, parameters were recorded and varied over

the course of each experiment to observe the strength of protein-ligand complexes

under different ionising strengths. All measurements were carried out in a positive

ion mode with ion source temperature of 20oC. A capillary voltage of 1.5 kV, cone

voltage of 40 V and extraction cone voltage of 4.8 V was applied to perform

nanoESI. All reported spectra were collected with a trap collision energy 12-30 V,

transfer collision energy 12-30 V, IMS pressure 5.02 × 10−1 mbar, TOF analyser

pressure 1.17 × 10−6 mbar. External calibration of the spectra was achieved using

caesium iodide at 100 mg ml−1 in water. Data acquisition and processing were

performed using Micromass MassLynx v4.1. Mass differences resulting from ligand

binding were calculated from the unbound protein peak internal to each spectrum.

The unbound protein peak was compared to the relevant 2.5% v/v d6-DMSO or

2.5% v/v ethanol control spectra for consistency. Mass differences were divided by

the molecular weight of the ligand to calculate binding stoichiometry.

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Chapter 3

Biochemical and Biophysical Characterization of P450 CYP142A1: An Example of Functional Redundancy in the

Mycobacterium tuberculosis Cholesterol Oxidases? 3.1 Introduction

Cholesterol is important in all facets of life and plays a central role in many

physiological processes (McLean et al., 2012). Its homoeostasis is essential for brain

and central nervous system function (McLean et al., 2012). Cholesterol and related

sterols are found everywhere throughout the environment (Garcia-Fernandez

(Garcia-Fernandez et al., 2013). They are present in cytoplasmic membranes and

play key roles as precursors of vitamin D, the bile acids and all the sterol hormones

(Garcia-Fernandez et al., 2013). Cholesterol is an essential molecule with many

roles involving cytochrome P450 enzymes (McLean et al., 2012).

In Mtb, cholesterol functions as an important source of energy during both latent

and chronic infection, and recent data have revealed that Mtb uses P450s to initiate

breakdown of host cholesterol for this purpose (Ouellet et al., 2011, Johnston et al.,

2010). The mycobacterial metabolism of cholesterol goes through two reaction

stages: firstly sterol side-chain activation/degradation and secondly steroid ring

opening (van der Geize and Dijkhuizen, 2004). Three key Mtb P450s are involved in

the first reaction and these are CYP125A1, CYP142A1 and CYP124A1 (hereafter,

referred to as ‘cholesterol oxidases’) (Johnston et al., 2010). These cholesterol

oxidases sequentially metabolise the cholesterol side chain at the C27 position to

154

the carboxylic acid state; first to an alcohol, then to an aldehyde and finally to the

acid moiety (Driscoll et al., 2010). CYP125A1 is the enzyme that plays the major

role in this cholesterol side chain degradation in Mtb CDC1551, but not in the Mtb

H37Rv strain (Johnston et al., 2010). Functional redundancy in cholesterol oxidation

capacity has been previously reported in Mtb H37Rv (but not in M. bovis BCG),

indicating that the H37Rv strain possesses compensatory enzyme(s) that allow it to

survive in the absence of a functional CYP125A1 (Johnston et al., 2010, Capyk et al.,

2009). However, studies by Johnston et al. revealed that a compensatory role in

cholesterol side chain oxidation was played by CYP142A1 through genetic

complementation of a CDC1551 ∆cyp125A1 strain by introduction of the Mtb

CYP142A1 gene (Johnston et al., 2010). Differences in the expression profiles of the

three cholesterol oxidase enzymes in the wild-type Mtb H37Rv strain revealed that,

though all three Mtb strains can oxidize cholesterol, only CYP142A1 (but not

CYP124A1) can complement the defect associated with the absence of CYP125A1

(Johnston et al., 2010). This indicates that CYP142A1 provides a functionally

redundant cholesterol side chain catabolic activity that can fully compensate for

loss or absence of CYP125A1 activity in the H37Rv ∆cyp125A1 strain (Johnston et

al., 2010). To further validate this hypothesis, CYP142A1 and CYP125A1 were shown

to be part of a gene cluster involved in lipid catabolism and suggested to play a

major role in the metabolism of host lipids, including cholesterol (Johnston et al.,

2010, Ouellet et al., 2010b).

In view of the gene location of CYP142A1 and its role in functional redundancy, a

vast amount of knowledge is still needed to unravel the role played by this enzyme

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in the pathogenicity and survival in the human macrophage during infection and

latency. This chapter hence aims to validate CYP142A1as a cholesterol oxidase using

series of biophysical/biochemical techniques and to evaluate its molecular

interaction with azole/novel inhibitors identified via fragment based screening.

Furthermore, recent studies by Manca et al., 1999 also revealed Mtb CDC1551 to

be less virulent but more immunogenic than the H37Rv strain, in that post-infection

it induces a rapid and vigorous cytokine response which may explain the high

frequency and large size of purified protein derivative (PPD) responses following

exposure to patients infected with the CDC1551 strain (Manca et al., 1999). A

similar differential response was induced on exposure to lipids from the two Mtb

strains. The CDC1551 strain of Mtb differs from the H37Rv strain in having a

mutation that inactivates CYP142 enzyme production. Thus, it is possible that

differences in lipid metabolism due to the presence or absence of CYP142 activity

result in different sterol/lipid profiles that influence the strength of the host

immune response.

The crystal structure of CYP142A1 has previously been solved (Driscoll et al., 2010).

However, in a later chapter of this thesis, the crystal structure of CYP142A1 in

complex with cholestenone will be reported. These studies will provide additional

insights into the cholesterol catabolic role of this enzyme. The results in this chapter

provide a broad analysis of the properties of CYP142A1, employing both biophysical

and biochemical techniques to further characterize an enzyme with a potentially

functionally redundant role in cholesterol catabolism in Mtb H37Rv (i.e. an ability to

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compensate for deficiencies in CYP125A1 activity and to enable continued Mtb

growth/survival using cholesterol as a carbon source). The data presented here

include studies to investigate the binding affinities of CYP142A1 to a variety of

substrates and inhibitors. Specific inhibitors of CYP142A1 could be used as chemical

tools to reveal how this enzyme relates to Mtb infection, growth and persistence in

the human host. They may also have potential as novel antibiotics that target

CYP142A1 and inhibit Mtb utilization of host cholesterol. Such compounds would

obviously be of even greater potency if they were also active against the other

major Mtb cholesterol 27-oxidase, CYP125A1.

Hence, CYP142A1 interactions with compound hits from fragment-based screening

studies (in collaboration with researchers from the University of Cambridge) were

also analysed, in order to allow for identification of small chemical ligands that bind

specifically to the active site of CYP142A1.

The UV-visible spectrum of a ligand-bound cytochrome P450 enzyme usually

provides a simple and accurate method for the determination of the nature of the

interaction between the ligand and the P450 (Locuson et al., 2007). The binding of

azole inhibitors and other nitrogen heterocycles to P450s usually involves

displacement of a water molecule (weakly ligated as an axial ligand to the heme

iron) by a basic nitrogen on the heterocyclic ring of the inhibitor (McLean et al.,

2002b). This leads to a shift in the Soret absorption maximum spectrum of the P450

to a longer wavelength, also known as a type II shift or a red shift. On the other

hand, the binding of substrates, in addition to displacing off the water molecule on

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the sixth axial position on the heme, switches the ferric heme iron from being hexa-

cordinated to penta-cordinated. This leads to a shift in the Soret absorption

maximum spectrum of the P450 to a shorter wavelength, also known as a type I

shift or a blue shift.

The structure of CYP142A1 was previously published as a monomer (Driscoll et al.,

2010). However, results from this work revealed that CYP142A1 can dimerize in

solution, and that the dimerization can be disrupted by the reducing agent DTT

(dithiothreitol). Analysis of the stability of CYP142A1 was also crucial for this work,

because this enzyme would be required in its most stable form for crystallographic

studies. Hence, results from stability assays on CYP142A1 are also presented in this

chapter.

The overall aims of the work in this chapter were to provide a detailed study of the

properties of CYP142A1, including analysis of its binding to substrates and

inhibitors, including fragment compounds with a view to developing novel

inhibitors. Further aims were to gain a detailed understanding of the

thermodynamic, hydrodynamic and spectroscopic properties of this P450, including

its aggregation state in solution. In particular, studies were aimed at defining the

coordination state of its heme iron and how this property, along with the overall

thermal stability of CYP142A1, is influenced by ligand binding.

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3.2 Results and Discussion 3.2.1 Expression and purification of CYP142A1 Preliminary CYP142A1 expression trials were investigated under different cell

growth and induction conditions, and these conditions were optimised as described

in the Materials and Methods (section 2.2.3-2.2.5). The results obtained showed

that more soluble protein was produced from growth in 2YT medium using C41

(DE3)/pET15b-CYP142A1 than was the case in other media and using other E. coli

strains. Furthermore, CYP142A1 was purified to homogeneity via three

chromatographic steps, as described in the Materials and Methods (section 2.2.6).

The first step used was affinity chromatography using a Ni-NTA column, followed by

affinity for Hydroxyapatite (HA) and a final “polishing step” using size exclusion

chromatography. The mechanisms of these chromatographic techniques are

described in more detail in the next chapter.

For nickel affinity purification, CYP142A1 crude cell extract was loaded onto a Ni-

NTA column, and red (heme-containing) protein was seen to bind. Following

loading of the sample and washing the column with the loading buffer, CYP142A1

was eluted with 40 mM, 60 mM and then 100 mM imidazole washes in 50 mM KPi

buffer (pH 8.0), containing 250 mM KCl and 10% glycerol. A large CYP142A1 band

was seen by SDS-PAGE analysis following the wash at 40 mM imidazole, and this

was confirmed by UV-visible spectral results, which showed the highest intensity

Soret peak of the P450 heme at a wavelength of 424 nm, signifying the presence of

substantial amounts of a P450 protein. P450s in their low-spin ferric state typically

have a Soret absorption maximum at ~418 nm, and thus the peak at 424 is likely a

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consequence of imidazole being bound to CYP142A1 in the distal position on the

heme iron at this stage. The apparent molecular mass of CYP142A1 (based on

comparisons with standards) from SDS-PAGE analysis is ∼46 kDa (Figure 3.1),

consistent with the predicted mass from the amino acid sequence at 46.6 kDa,

including the N-terminal hexahistidine tag region.

Figure 3.1: Protein purification of the Mtb CYP142A1 from the pET15b/CYP142A1 plasmid. SDS PAGE analysis (10% polyacrylamide gel) shows molecular weight markers (Lane 1, NEB Broad range marker bands labelled in kDa), Lysate (total protein) (lane 2), and flow-through from the column (lane 3). Lanes 4-8 show CYP142A1 eluted from the column with increasing concentrations of imidazole in the wash buffer.

The largest amount of CYP142A1 was clearly obtained using the 40 mM and 60 mM

imidazole washes, whereas there appeared to be lower amounts of hemoprotein in

the sample collected at 100 mM imidazole. The relative CYP142A1 protein purity

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improved with increased concentration of imidazole. This was confirmed through

observation of intense bands of CYP142A1 along with only minor bands of other

protein contaminants. Samples isolated from Ni-NTA chromatography were

exchanged into 15 mM KPi, pH 6.5 (Buffer A) and loaded onto a HA column pre-

equilibrated in the same buffer. Protein was eluted using a gradient of 15 mM to

500 mM KPi (pH 6.5) buffer and samples were retained for SDS-PAGE analysis. From

the SDS-PAGE results (Figure 3.2), single bands of CYP142A1 were seen after the HA

purification along with only very minor bands of other protein contaminants.

Samples from the HA purification step had an Rz (Reinheitszahl) purity ratio value of

A417/A280 ≥1.5. Hence, the next step was to subject CYP142A1 to a further

purification using gel filtration (size exclusion) chromatography in order to obtain

highly purified enzyme for crystallographic and other analyses.

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Figure 3.2: Purification of Mtb CYP142A1 using hydroxyapatite (HA) column chromatography. SDS-PAGE analysis (10% polyacrylamide gel) shows molecular weight markers (lane 1, NEB Broad range marker, bands labelled in kDa) and purified CYP142A1 as a near-single band (lanes 2-9) and close to the predicted molecular weight of ∼46 kDa (for CYP142A1 plus its His-tag) when compared to the molecular weight markers. Other proteins eluted along with CYP142A1 are seen as faint contaminant bands by SDS-PAGE.

From the HA column chromatography, purer samples were pooled together,

concentrated to ~50 µl by ultrafiltration and loaded onto an S-200 gel filtration

column, which served as a final “polishing” step for CYP142A1 purification. This was

carried out on an AKTA purifier using 10 mM Tris plus 150 mM NaCl, 1 mM DTT, pH

7.2 as the loading buffer. Fractions collected from the gel filtration column were

analysed using SDS-PAGE. The final step resulted in a purer enzyme, depicted by

single bands of CYP142A1 (Figure 3.3) close to the predicted molecular weight of

∼46 kDa (CYP142A1 plus His-tag) on an SDS-PAGE gel, and with an improved Rz of

A418/A280 ≥ 2.0.

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Figure 3.3: Purification of Mtb CYP142A1 using a SuperdexTM S-200 gel filtration column. SDS-PAGE analysis shows molecular weight markers (lane 1, NEB Broad range marker, bands labelled in kDa) and pure CYP142A1 as a single band (lanes 2-7) and close to the predicted molecular weight of ∼46 kDa (CYP142A1 plus His-Tag) when compared to the molecular weight markers.

PURIFICATION STAGE

TOTAL PROTEIN (mg)

TOTAL P450 (mg)

P450/PROTEIN (A418/A280)

STEPWISE PURIFICATION FOLD

OVERALL PURIFICATION FOLD

STEPWISE YIELD %

OVERALL YIELD %

Lysate 76415.00 692.13 0.01 1.00 1.00 100 100

Ni-NTA 660.15 319.39 0.48 48.00 48.00 46.15 46.15

HA Purification 126.68 74.51 0.59 1.23 59.00 23.33 10.77

Gel filtration 67.55 43.29 0.64 1.08 64.00 58.10 6.25

Table 3.1: Typical CYP142A1 purification table.

Table 3.1 above shows a typical purification scheme from the three purification

steps carried out for isolation of CYP142A1. There was a decrease in the amount of

the total protein (which includes all contaminating proteins) and an increase in the

relative amount of P450 protein, signifying an increase in the purity of CYP142A1

during the purification process. The stepwise purification of CYP142A1 after passing

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through the nickel column was about 48-fold, which highlights an extensive

removal of contaminating proteins at this stage. For the HA and the gel filtration

steps, the stepwise purification fold was lower (1.23- and 1.08-fold, respectively)

and this further confirms the relative effectiveness of the nickel column

chromatographic step in the purification of His-tagged recombinant proteins. The

overall purification was 64-fold and resulted in a single protein band by SDS-PAGE

analysis. Assuming near-complete CYP142A1 purification at this stage, this suggests

that CYP142A1 did not comprise more than ~1.6% of the total protein content in

the E. coli expression cells used. The overall yield decreased from one purification

step to the next and this suggests significant losses in the total amount of

CYP142A1 recovered. However, since calculations of CYP142A1 recovery were

based on heme absorbance measurements, it should also be borne in mind that

other cofactor-binding (e.g. heme, flavin iron-sulfur cluster) proteins in E. coli may

also contribute to the absorption used to quantify CYP142A1, and thus the overall

yield of CYP142A1 may actually be rather greater than the estimate given in Table

3.1.

3.2.2 CYP142A1 Substrate Binding Assays

Studies have highlighted that, due to the genetic location of CYP142A1 in Mtb, this

P450 could be involved in the metabolism of cholesterol while the pathogen is

engulfed in human macrophages in the latent phase of infection (Van der Geize et

al., 2007). Interactions of CYP142A1 with cholesterol (i.e. productive, type I binding

spectra) were reported previously (Driscoll et al., 2010). Hence, with these data in

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hand, we examined the spectral properties of CYP142A1 on binding to selected

sterols, and specifically cholestenone, cholesterol and lanosterol. CYP142A1

displays UV–visible spectral characteristics typical of low-spin ferric P450 enzymes,

with its Soret peak at 418 nm in the oxidized, substrate-free form. CYP142A1

undergoes shifts in heme iron spin-state equilibrium from a predominantly low-spin

(S = 1/2) to a partially/predominantly high-spin (S = 5/2) form on binding selected

sterol substrates and analogues, with these molecules evidently binding to the P450

in the active site close to the heme, and inducing the displacement of the distal H2O

ligand to the heme iron.

For the substrates analysed in this study (cholesterol, cholestenone and lanosterol),

binding of the sterol substrates gives a distinctive high-spin (type I shift), with the

Soret band moving from 418 towards ~393 nm in the substrate-bound form. Figure

3.4A shows an optical titration of CYP142A1 (3.1 μM) with cholestenone, displaying

the type I Soret shift as cholestenone binds, and with a Soret isosbestic point at 406

nm. The inset shows an overlaid set of difference spectra derived from the

cholestenone titration of CYP142A1. Figure 3.4B shows a plot of the induced

spectral change versus the steroid concentration, fitted to generate a Kd value of

0.22 ± 0.02 μM.

Figure 3.5 A shows an optical titration of CYP142A1 (3.4 μM) with cholesterol.

Cholesterol induced similar spectral changes to cholestenone and bound CYP142A1

with an apparent Kd of 0.25 ± 0.01 μM. The inset shows an overlaid set of difference

spectra derived from binding CYP142A1 with cholesterol. Figure 3.5B shows a fit of

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the induced absorption change versus cholesterol concentration. Due to the low

solubility of cholesterol, the conversion to the high-spin state was not as extensive

when compared to that for cholestenone. As the optical titration continued with

higher concentrations of cholesterol, some reconversion of heme absorption

towards the low-spin state was also observed.

Figure 3.6 shows an optical titration of CYP142A1 (3.5 μM) with lanosterol.

Lanosterol is a substrate for the Mtb sterol demethylase CYP51B1, and for other

eukaryotic lanosterol demethylases - which catalyse a three-step reaction of

oxidative removal of the 14-α-methyl group from lanosterol to form ergosterol

(Lepesheva et al., 2008). The data here also show tight binding of lanosterol to

CYP142A1 but relatively weaker as compared to cholestenone and cholesterol with

a Kd value of 7.50 ± 0.99 μM.

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Figure 3.4: Optical titration of CYP142A1 with cholest-4-en-3-one. Panel A shows

absolute spectra recorded during a titration of CYP142A1 (3.6 μM) with

cholestenone. The Soret peak shifts from 418 to 393 nm as the high-spin ferric

heme iron form accumulates. The inset shows overlaid difference spectra from the

optical titration. Panel B shows cholestenone-induced absorption change plotted

versus cholestenone concentration, with data fitted using the Morrison equation

(Equation 1) to produce a Kd value of 0.22 ± 0.02 μM.

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Figure 3.5: Optical titration of CYP142A1 with cholesterol. Panel A shows UV-visible absorption spectra from a titration of cholesterol with CYP142A1 (3.6 μM). Some reconversion from high-spin towards low-spin heme is observed at higher concentrations of cholesterol. The inset shows difference spectra from the titration with peak and trough values at 387 nm and 420 nm. Panel B shows a fit of cholesterol-induced absorption change (ΔA387 minus ΔA420, reflecting the peak and trough values in the difference spectra, and computed by subtracting the spectrum for cholesterol-free CYP142A1 from each of the spectra for the cholesterol-bound forms, and by plotting the absorbance difference data versus [cholesterol] added, with data fitted using equation 1 (see main text) to generate an apparent Kd value of 0.25 ± 0.01 μM for cholesterol binding to CYP142A1. The solid line indicates the data fit up to 1.2 μM cholesterol, with the black dots indicating the projected fit beyond these points using the Morrison equation (Equation 1).

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Figure 3.6: Optical binding of CYP142A1 with lanosterol. Panel A shows UV visible

absorption spectra from a titration of lanosterol with CYP142A1 (3.7 μM). The Soret

band shifts towards high spin (HS) and from 418 to 393 nm as the HS ferric heme

iron form accumulates. The formation of the HS substrate-bound form was not

extensive. The inset shows overlaid difference spectra from the optical titration.

Panel B shows lanosterol-induced absorption change plotted versus lanosterol

concentration, with data fitted using the Morrison equation (Equation 1) to

produce a Kd value of 7.50 ± 0.99 μM.

P450s exist in large numbers in eukaryotes, but are often considered to be

relatively rare in bacteria. When found in bacteria, they function frequently as

components of dispensable catabolic pathways for the breakdown of unusual

carbon sources, e.g. in the case of P450cam (Poulos et al., 1987, Peterson et al.,

1992). However, the more recent genome sequencing projects for Mtb have

highlighted that this pathogen encodes a large numbers of P450s, with 20 different

P450 isoforms encoded within the bacterial genome in the case of Mtb H37Rv (Cole

et al., 1998). The location of CYP142A1 in a large gene cluster conserved from the

Rhodococcus sp. strain RHA1 to Mtb highlights an important physiological role in

cholesterol metabolism that is shared in the two organisms (Van der Geize et al.,

2007, McLean and Munro, 2008).

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Studies have demonstrated that cholesterol, which is a major source of carbon for

Mtb during infection, also plays a crucial role in aiding the entry of the pathogen

into human macrophages (McLean et al., 2012, Chang et al., 2009). The P450

enzymes implicated in the early stages of Mtb cholesterol metabolism are

CYP125A1, CYP142A1 and CYP124A1 (Johnston et al., 2010). These P450 isoforms

have also been demonstrated to perform hydroxylation at the cholesterol C27

position and also further sequential oxidations at the same position (for both

cholesterol and cholestenone) via an aldehyde product to form the carboxylic acid

moiety, which prepares the cholesterol side chain for β-oxidation (Ouellet et al.,

2010a, McLean et al., 2009, Capyk et al., 2009, Driscoll et al., 2010). Studies have

also shown that steroid ring metabolism cannot proceed when the sterol has a full-

length aliphatic side chain, and thus these P450s perform a crucial role in initiating

cholesterol breakdown. Capyk et al. carried out a study to compare the growth of

CYP125 gene deletion strains and the wild-type forms of M. bovis BCG and Mtb

H37Rv on cholesterol. While M. bovis BCG grew on cholesterol and cholestenone,

the CYP125 deletion strain failed to grow (Capyk et al., 2009). For Mtb H37Rv,

however, the ∆CYP125A1 strain grew on cholesterol, suggesting a compensatory

mechanism involved in the metabolism of cholesterol in Mtb H37Rv. Recent studies

have also suggested that while CYP125A1 is the major enzyme involved in this side

chain hydroxylation, CYP142A1 could also function as compensatory enzyme

(McLean et al., 2012). In this study, the spectral changes on binding of cholesterol,

cholest-4-en-3-one (cholestenone) and lanosterol to this P450 enzyme were

investigated. Cholestenone and cholesterol exhibited tight binding to CYP142A1,

with Kd values of 0.22 ± 0.02 μM and 0.25 ± 0.01 μM, respectively, and a Soret

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maximum shift towards a shorter wavelength, consistent with displacement of a

water ligand from the heme iron to form a 5-coordinate high-spin form of the ferric

heme iron. These data further suggest these sterols to be the major substrates for

CYP142A1, which is consistent with results presented in previous studies that

indicate CYP142A1 is a cholesterol hydroxylase. Lanosterol, a major substrate for

CYP51 enzymes (including the Mtb CYP51B1 isoform), was also used in this study to

investigate the behaviour of other sterols with CYP142A1. This also showed a

relatively low Kd value (7.50 ± 0.99 μM) for binding to CYP142A1, thus signifying

that this structurally diverse sterol retains capacity to bind to CYP142A1, albeit with

a Kd value ~30-fold weaker than those for cholesterol and cholestenone. Moreover,

the binding mode for lanosterol results in a much less extensive shift towards the

high-spin state, indicating that while CYP142A1 can bind to all three of these

sterols, it has a clear preference for cholesterol/cholestenone over lansoterol.

3.2.3 Inhibitor Binding Assays CYP142A1 binds to a range of imidazole and triazole antifungals, inducing a type II

(Soret red shift). In this study, on binding CYP142A1 with the azole inhibitors, the

Soret band shifted from 418 ± 1 nm (inhibitor-free) to ~423 ± 1 nm (azole-

saturated). Highest affinity was observed for bifonazole, clotrimazole, miconazole

and econazole (Kd values of 0.55 ± 0.10 μM, 1.14 ± 0.10 μM, 1.42 ± 0.16 μM, and

2.28 ± 0.19 μM, respectively) (Figure 3.7-3.11) (Table 3.2).

For these four tight-binding azoles, the computed spectral changes associated with

azole ligation occurred almost linearly with increasing azole concentrations in the

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lower concentration range, and then sharply reached a plateau, indicative of tight,

near-stoichiometric binding to the P450. Optical titrations with fluconazole

revealed weaker binding of this azole to CYP142A1, with a Kd value of 309.1 ± 36.3

μM. Ketoconazole bound to CYP142A1 with a higher affinity than fluconazole, with

a Kd value of 11.95 ± 0.64 μM.

The binding of imidazole (283.3 ± 6.4 µM) to CYP142A1 was considerably weaker

than to the other azoles (apart from fluconazole), indicating that the high affinity of

the azole antifungals for CYP142A1 is determined primarily by favourable

interactions between the bulky, polycyclic azole antifungals and the hydrophobic

residues in the largely apolar active site of CYP142A1, rather than being driven

mainly by ligation of the azole group to the ferric heme iron. No significant optical

changes were observed on titration of CYP142A1 with the more polar voriconazole

and the bulky itraconazole, suggesting that these drugs bind much more weakly to

CYP142A1 than do the other azole drugs, or that they cannot penetrate the P450’s

active site. Kd values were determined from fits using the Morrison equation

(equation 1) for tighter binding azoles, and using a standard hyperbolic function

(Michaelis-Menten, equation 2) for weaker binding azoles.

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Figure 3.7: CYP142A1 binding titration with econazole. Panel A shows accumulated absorption and absorbance difference spectra induced by the addition of econazole to oxidized CYP142A1 (5.9 μM), with difference spectra generated by subtraction of the starting (azole-free) CYP142A1 absolute spectrum from those collected following addition of 0.6-14 μM econazole. The Soret absorption maximum shifted to 423 nm in the absolute spectrum for the econazole-saturated enzyme. In the difference spectra (inset), minima and maxima resulting from azole addition are located at approximately 411 nm and 431 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A431 minus ∆A411) for binding of econazole to CYP142A1 versus the relevant econazole concentrations. Fitting the data using the Morrison equation (equation 1) generates a Kd value of 2.28 ± 0.19 μM for the drug.

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Figure 3.8: CYP142A1 binding titration with miconazole. Panel A shows collected absorption and difference spectra induced by the addition of miconazole to oxidized CYP142A1 (4.0 μM), with difference spectra generated by subtraction of the starting (azole-free) CYP142A1 absolute spectrum from those collected following addition of 1.2-24.1 μM miconazole. The Soret absorption maximum shifts from 418 to 423 nm in the absolute spectrum on miconazole saturation. In the difference spectra, minima and maxima resulting from azole addition are located approximately at 432 nm and 413 nm. Panel B shows a fit of absorption change (∆A432 minus ∆A413) for binding of miconazole to CYP142A1 versus the relevant miconazole concentrations. Fitting the data using the Morrison equation (equation 1) generates a Kd value of 1.42 ± 0.16 μM for the drug.

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Figure 3.9: CYP142A1 binding titration with clotrimazole. Panel A shows collected absorption and difference spectra induced by addition of clotrimazole to oxidized CYP142A1 (5.8 μM), with difference spectra generated by subtraction of the azole-free CYP142A1 spectrum from those collected after addition of 1.5-21.8 μM clotrimazole. The azole drug complex is near-fully formed by ~21.8 μM, with the Soret maximum at 423 nm. In the difference spectra (inset), minima and maxima resulting from azole addition are at ~412 nm and 431 nm. Panel B shows a plot of maximal shift in absorption data (∆A431 minus ∆A412) for CYP142A1 binding to clotrimazole CYP142A1 versus the relevant [clotrimazole]. Fitting using equation 1 gives a Kd value of 1.14 ± 0.10 µM for the drug.

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Figure 3.10: CYP142A1 binding titration with bifonazole. Binding of bifonazole to the P450 heme produces a shift in the UV-visible absorption spectrum, due to replacement of the sixth water ligand to the ferric heme iron by the azole moiety. Panel A shows a collection of absorption and difference spectra induced by the binding of the azole antifungal drug bifonazole to oxidized CYP142A1 (4.0 μM), with difference spectra generated by subtraction of the starting (azole-free) CYP142A1 absolute spectrum from those collected following addition of 0.8-12.1 μM bifonazole. The Soret absorption maximum shifted from 418 to 423 nm in the absolute spectrum. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from azole addition are located at approximately 415 nm and 436 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A436 minus ∆A415) for binding of bifonazole to CYP142A1 versus the relevant bifonazole concentrations. Fitting the data using the Morrison equation (equation 1) generates a Kd value of 0.55 ± 0.10 μM for the drug.

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Figure 3.11: CYP142A1 binding titration with sodium cyanide. Panel A shows collected absorption and difference spectra induced by the addition of cyanide to oxidized CYP142A1 (3.2 μM), with difference spectra generated by subtraction of the starting (cyanide-free) CYP142A1 absolute spectrum from those collected following addition of 2.7-46.2 mM cyanide. The Soret absorption maximum shifts from 418 to 433 nm in the absolute spectrum on cyanide saturation. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from azole addition are located at approximately 415 nm and 441 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A441 minus ∆A415) for binding of cyanide to CYP142A1 versus the relevant cyanide concentrations. Fitting the data using the Hill function (equation 3) generates an apparent Kd value of 25.29 ± 0.29 mM for cyanide binding.

In view of the importance of cholesterol metabolism in Mtb infection and

persistence in human macrophages, targeting this pathway could provide the much

needed route for the development of new therapeutic agents. Azole antibiotics are

generally used as antifungal drugs and the effect of these drugs results from the

inhibition of the sterol biosynthesis pathway at the step catalysed by the CYP51

enzyme – a lanosterol demethylase (Ouellet et al., 2011, McLean et al., 2002b).

Azole antibiotics such as econazole were shown to have antimycobacterial activities

against both latent Mtb and multidrug-resistant strains (Ahmad et al., 2006b,

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Ahmad et al., 2006c) and this discovery suggests one or more cytochrome P450

enzymes are likely drug targets.

In this study, CYP142A1 exhibited tight-binding characteristics to a number of azole

drugs, with Kd values in the μM range. This high affinity of azole drugs for CYP142A1

suggests that the enzyme could be a valuable drug target in Mtb. This is consistent

with other azole drug/Mtb P450 interactions that have been characterised to date,

including those with CYP121A1 (McLean et al., 2002b), CYP144A1 (Driscoll et al.,

2011) and CYP130A1 (Ouellet et al., 2008). The tightest binding azole drug

interactions obtained for CYP142A1 were with bifonazole, clotrimazole, miconazole

and econazole, while the weakest detectable binding was with fluconazole. The

order of potency of the azole drugs (see MIC data in section 1.5.9) correlated with

their Kd values for binding to the Mtb CYP142A1 enzyme (Table 3.2), suggesting

that this P450 is a valuable drug target.

The cyanide ion acts as an ionic ligand and binds preferentially to the ferric iron (III)

form of P450s. In the resting state of the enzyme, the 3+ charge of the ferric heme

iron (Fe3+) matches that of the three negative ions provided by two of the porphyrin

nitrogens and the thiolate ligand, but in the reduced state of the enzyme the

negative charges exceed the two positive charges of the Fe2+ species. Hence

cyanide binds more readily, but still weakly, to the ferrous species of cytochrome

P450 enzymes (Correia and Ortiz de Montellano, 2005).

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From the titration carried out in this study, a sigmoidal cyanide titration plot was

obtained, and data were best fitted using the Hill equation (Eq. 3), with an apparent

Hill coefficient of 8.68 ± 0.70. While it is not possible to infer too much from the Hill

coefficient itself, these data do suggest that some form of binding “cooperativity”

occurs with cyanide, and an apparent binding constant (Kd or KH) of 25.29 ± 0.29

mM was obtained, signifying very weak binding to CYP142A1. The sigmoidal curve

obtained could result from multiple binding sites for cyanide on CYP142A1.

Possibly, the binding of sodium ions in the CYP142A1 active site ultimately shields

repulsive interactions with the cyanide anion, enabling it to ligate the heme iron.

This model might explain the unusual sigmoidal dependence of heme ligation on

the concentration of sodium cyanide.

3.2.4 Binding Analysis with CYP142A1 Fragment Hits

Fragment based drug discovery is a novel approach for developing small-molecule

ligands as chemical tools and leads for drug development (Scott et al., 2012). It

involves a structure-based design and synthesis of specific and potent ligands from

weak-binding low molecular weight ligand (fragment) molecules with molecular

weights less than ~250 Da (Hudson et al., 2014). An initial fragment screening (using

fluorescence-based protein stability thermal shift and heme binding assays)

performed on CYP142A1 using a fragment library of 720 fragments generated six

fragment hits. These hits are named: NMR170, NMR540, NMR623, NMR089,

NMR491 and NMR099. Hit validation studies were then carried out using UV-Vis

spectroscopy, electron paramagnetic resonance (EPR) spectroscopy, Isothermal

titration calorimetry (ITC) and X-ray crystallography. Selected results from the UV-

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Vis spectroscopy studies are shown below. UV-Vis spectroscopy revealed a

substrate-like type I optical shift for fragment NMR099, and inhibitor-like type II

shifts for the five others – indicating direct interaction with the heme iron.

Dissociation constants for the fragments ranged from NMR491 (0.68 µM) to

NMR099 (6.8 mM) (Table 3.2).

NMR540, NMR623, NMR089, NMR491 and NMR099 were observed to bind weakly

to CYP142A1, which is expected for fragments at the first stage of the screening

process. The aim of the fragment based drug discovery approach is to develop

highly potent inhibitors from small ligands which exhibit low affinity for the drug

targets, but bind in distinct parts of the enzyme active site (Hudson et al., 2012b).

These initial fragments hits can then be developed further via structure-based

fragment elaboration and optimization to generate inhibitors with high affinity and

potency for the enzyme target (Hudson et al., 2013, Hudson et al., 2012b).

Figure 3.12: Compounds hits from an initial CYP142A1 fragment screen

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Figure 3.13: CYP142A1 binding titration with NMR170. Panel A shows collected absorption and difference spectra induced by the addition of NMR170 to oxidized CYP142A1 (5.0 μM), with difference spectra generated by subtraction of the starting (azole-free) CYP142A1 absolute spectrum from those collected following additions of NMR170. The Soret absorption maximum shifts from 418 to 421 nm in the absolute spectrum on NMR170 saturation. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from azole addition are located at approximately 411 nm and 431 nm. Panel B shows the plot of the maximal shifts in absorption (∆A431 minus ∆A411) for binding of NMR170 to CYP142A1 versus the relevant ligand concentrations. Fitting the data using the Hill function (equation 3) generates an apparent Kd value of 1.87 ± 0.07 µM, where n = 1.28 ± 0.56.

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Figure 3.14: CYP142A1 binding titration with NMR540. Panel A shows collected absorption and difference spectra induced by the addition of NMR540 to oxidized CYP142A1 (3.8 μM), with difference spectra generated by subtraction of the starting (azole-free) CYP142A1 absolute spectrum from those collected following sequential additions of NMR540. The Soret absorption maximum shifts from 418 to 420 nm in the absolute spectrum on NMR540 saturation. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from azole addition are located at approximately 411 nm and 432 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A432 minus ∆A411) for binding of NMR540 to CYP142A1 versus the relevant NMR540 concentrations. Fitting the data using the hyperbolic function (equation 2) generates a Kd value of 107.46 ± 7.35 µM for NMR540.

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Figure 3.15: CYP142A1 binding titration with NMR623. Panel A shows collected absorption and difference spectra induced by the addition of NMR623 to oxidized CYP142A1 (4.2 μM), with difference spectra generated by subtraction of the starting (azole-free) CYP142A1 absolute spectrum from those collected following addition of NMR623. The Soret absorption maximum shifts from 418 to 420 nm in the absolute spectrum on NMR623 saturation. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from NMR623 azole addition are located at approximately 413 nm and 433 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A433 minus ∆A413) for binding of NMR623 to CYP142A1 versus the relevant NMR623 concentrations. Fitting the data using a hyperbolic function (equation 2) generates an apparent Kd value of 232.41 ± 11.51 µM.

3.2.5 Binding Analysis with Compounds from CYP121A1 Fragment Elaboration Hits

The first successful application of fragment-based approach to the Mtb P450

enzymes was achieved with CYP121A1 (Hudson et al., 2012b, Hudson et al., 2013).

A preliminary fragment-screening process involving thermal shift and NMR

spectroscopy generated four fragments which were shown to bind within the

CYP121A1 active site. One of these bound in two overlapping binding modes

(mimicking the binding of the cyclic dipeptide substrate), with others interacting in

both heme-binding and non-heme binding modes (Hudson et al., 2012b, Hudson et

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al., 2013). A fragment–fragment merging approach was used to develop these

further and this led to the discovery of a novel type-II aminoquinoline inhibitor with

high ligand efficiency (LE, where LE = -∆G of binding/number of non-hydrogen

atoms (NHA) in the ligand) and about four times higher affinity than the natural

CYP121A1 substrate cYY (Hudson et al., 2012b). This novel inhibitor was also shown

to be the highest affinity ligand developed using fragment-based approaches

against any cytochrome P450 enzyme (Hudson et al., 2013). Further studies

involving structural biology and synthetic chemistry have continued around this

CYP121A1-specific inhibitor and these have led to the development of other ligands

that bind to other Mtb P450s. A number of these ligands were shown to bind the

cholesterol oxidases CYP124A1, 125A1 and 124A1. These are named MEKs 046,

047, 065, 066, 076, 077 and 050f2. Their structures are shown in Figure 3.15. Out of

the seven compounds, three appeared to stand out in terms of affinity for these

cholesterol oxidases, and these are MEKs 047, 065 and 066. MEK046 showed a type

II spectral shift (inhibitor-like), while MEK065 and MEK066 showed type I shifts

(substrate-like) with partial conversion (~25% conversion) to a high-spin state. The

Kd values ranged from 3.24 µM (MEK046) to 35.30 µM (MEK066). MEKs 047, 050f2,

076 and 077 gave weak (about 1 nm shift) or no spectral shifts for these enzymes at

all.

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Figure 3.16: Elaborated compounds developed from CYP121A1 fragment hits.

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Figure 3.17: CYP142A1 binding with MEK046. Panel A shows collected absorption and difference spectra induced by the addition of MEK046 to oxidized CYP142A1 (5.2 μM), with difference spectra generated by subtraction of the starting (ligand-free) CYP142A1 absolute spectrum from those collected following addition of MEK046. The Soret absorption maximum shifts from 418 to 421 nm in the absolute spectrum on MEK046 saturation. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from MEK046 addition are located at approximately 411 nm and 441 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A432 minus ∆A411) for binding of MEK046 to CYP142A1 versus the relevant MEK046 concentrations. Fitting the data using the Morrison equation (Equation 1) generates a Kd value of 1.00 ± 0.09 µM.

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Figure 3.18: CYP142A1 binding with MEK065. Panel A shows collected absorption and difference spectra induced by the addition of MEK065 to oxidized CYP142A1 (5.2 μM), with difference spectra generated by subtraction of the starting (ligand-free) CYP142A1 absolute spectrum from those collected following addition of MEK065. The Soret absorption maximum shifts from 418 to 405 nm in the absolute spectrum on MEK065 saturation. In the difference spectra (inset), minima (trough) and maxima (peak) resulting from MEK065 addition are located at approximately 420 nm and 387 nm. Panel B shows the plot of the maximal shifts in absorption data (∆A387 minus ∆A420) for binding of MEK065 to CYP142A1 versus the relevant ligand concentrations. Fitting the data using the Morrison equation (equation 1) generates a Kd value of 3.13 ± 0.10 µM.

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S/N Ligand Kd Value (µM) Type of shift

1 Cholestenone 0.22 ± 0.02 I

2 Cholesterol 0.25 ± 0.01 I

3 Lanosterol 7.50 ± 0.99 I

4 Econazole 2.28 ± 0.19 II

5 Miconazole 1.42 ± 0.16 II

6 Clotrimazole 1.14 ± 0.10 II

7 Bifonazole 0.55 ± 0.10 II

8 Ketoconazole 11.95 ± 0.64 II

9 Fluconazole 309.2 ± 36.3 II

10 Imidazole 283.3 ± 6.4 II

11 1-Phenylimidazole 23.20 ± 3.84 II

12 Sodium cyanide 25290 ± 290 II

13 NMR170 1.87 ± 0.07 II

14 NMR491 0.68 ± 0.14 II

15 NMR540 107.46 ± 7.35 II

16 NMR623 232.41 ± 11.51 II

17 NMR089 377.4 ± 9.3 II

18 NMR099 7214 ± 332 I

19 MEK046 1.00 ± 0.09 II

20 MEK065 3.13 ± 0.10 I

21 MEK066 35.30 ± 3.80 I

Table 3.2: Binding spectral characteristics and Kd values for CYP142A1 ligands. Kd values were determined as described in the Materials and Methods (section 2.2.9).

3.2.6 CYP142A1 Fe(II)-CO Adduct and NO-Adduct Formation

Formation of a carbon monoxide adduct is a diagnostic signature for cytochrome

P450 enzymes (Driscoll et al., 2010). These enzymes exhibit a characteristic shift in

the Soret peak to approximately 450 nm (ferrous-CO complex) on reduction and

binding with carbon monoxide (Omura and Sato, 1964). This characteristic Soret

shift is as a result of the retention of the thiolate proximal ligand to the heme iron

(Cys339 in CYP142A1) (Driscoll et al., 2010), while a shift to ~420 nm is indicative of

the protonation of the thiolate ligand, resulting in the formation of a cysteine thiol

as the proximal ligand. In this P420 state, the enzyme is inactive (McLean et al.,

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2008, Omura and Sato, 1964). A study carried out by Driscoll et al. to check the

effect of pH on the stability of the CYP142A1 CO-adduct revealed that the P450

form of CYP142A1 is most stable at pH 7 and highly unstable at pH values less than

6 and greater than 8, which is marked by the formation of the P420 form (Driscoll et

al., 2010).

In this study, the UV-Vis absorption spectrum for carbon monoxide binding to

CYP142A1 was recorded between 250 and 700 nm. It shows features typical of a

heme-containing protein. In its resting state, CYP142A1 exhibited a UV-Vis

spectrum characteristic of Fe(III) (ferric) P450s in their low-spin, hexacoordinated

state, with a Soret peak at 418 nm and α and β bands at 564 and 532 nm,

respectively. Upon reduction with dithionite and after bubbling with CO, the Soret

peak shifted to 450 nm, as expected for a P450 Fe(II)-CO complex (Omura and Sato,

1964). This complex was stable for several minutes in the absence and presence of

the substrate cholestenone (Figure 3.19) and did not convert to a P420 complex,

which absorbs maximally at around 420 nm and arises from protonation of the

cysteine thiolate to a thiol form. A final P420:P450 peak height ratio of

approximately 1:3 was obtained. The same ratio was observed in the presence and

absence of the natural substrate cholestenone (Figure 3.19).

Previous studies carried out on the Mtb P450 CYP51B1 revealed the propensity of

the enzyme to convert from the thiolate to the thiol proximal cysteinate ligand on

reduction of the heme iron. However, this process was found to be retarded in the

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presence of a substrate analogue (estriol) (Driscoll et al., 2011, McLean et al.,

2006b, Aoyama et al., 1998). Previously, it was postulated that the P420 Fe2+-CO

species reflects an ‘inactivated’ form of the enzyme (i.e. that ferric form of the

same enzyme preparation may have thiol coordination and be catalytically inactive)

but this has been counteracted by follow-up studies reported for CYP121A1 and the

P450epoK enzyme from Sorangium cellulosum (Dunford et al., 2007, Ogura et al.,

2004). These results revealed that the P450-CO adduct in the P450epoK enzyme is

restored to the P450 state from the P420 form on binding with its epothilone

substrate. In addition, CYP121A1 can also be reversibly converted between the

P420/P450 forms by titrating the reduced/CO-bound forms in the pH range from

6.1 to 10.5 (Dunford et al., 2007). Hence, these results are indicative of the

reversibility of the P450/P420 transition and also the relevance of substrate binding

in the stabilization of the catalytically active thiolate-coordinated heme state

(Driscoll et al., 2011).

At the early stage of Mtb infection, nitric oxide (.NO) generated by host

macrophages inhibits heme-containing terminal cytochrome oxidases, inactivates

iron/sulfur proteins, and mobilises entry into the latent phase (Ouellet et al., 2009).

A study carried out by Ouellet et al. revealed that nitric oxide binds tightly to

CYP125A1 and CYP142A1 at sub-micromolar concentration, but binds with lower

affinity to CYP130A1 and CYP51A1 (Ouellet et al., 2009). However, the ferrous NO-

P450 adducts formed with CYP125A1 and CYP142A1 decomposed back to their

ferric P450 resting forms within minutes of exposure to oxygen, while the ferrous

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CYP130A1 and CYP51B1 adducts remained bound almost irreversibly to NO (Ouellet

et al., 2009). This study suggested that, at physiological concentrations of

approximately 1 M, nitric oxide could inhibit the activity of CYP130A1 and

CYP51A1, whereas the cholesterol hydroxylases CYP125A1 and CYP142A1 are more

resistant (Ouellet et al., 2009).

In this study, the formation of a ferric CYP142A1-nitric oxide adduct was monitored

over a period of 30 minutes. The brief bubbling of nitric oxide gas into solutions of

ferric CYP142A1 prepared under strictly anaerobic conditions resulted in the

formation of nitrosyl complexes characterized by Soret, α and β bands located at

433, 572, and 541 nm, respectively. This complex, however, remained stable over a

period of 30 minutes without significant decomposition, which further agrees with

the study carried out by Ouellet et al. (Ouellet et al., 2009). The NO-bound

CYP142A1 spectral features (Soret at 433 nm with strong alpha and beta band

development) are typical of those for other characterized Mtb P450s, e.g.

CYP130A1, CYP51B1 and the cholesterol 27-hydroxylase CYP125A1 (Ouellet et al.,

2010b, McLean et al., 2009).

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Figure 3.19: UV-visible spectra for gaseous ligand-bound complexes of CYP142A1. A) UV-visible spectral features of substrate-free CYP142A1 (5.6 µM) in the ferric (black), sodium dithionite-reduced ferrous (red), and ferrous-CO bound (green) forms. B) Spectral properties for CYP142A1 (5.1 µM) are shown for the ferric (oxidized) form with a Soret maximum at 418 nm (black), the sodium dithionite-reduced form with a maximum at 414 nm (red), the CYP142A1-cholestenone complex after reduction with sodium dithionite with a peak at 414 nm (green), and the ferrous-CO CYP142A1 complex with a major peak at 449 nm and a minor peak at 422 nm (blue), likely reflecting a small proportion of the P420 state. C) UV-visible

absorbance spectra of CYP142A1 (3.3 M) in the resting ferric (black), and ferric-NO bound (red) forms. The data reveal Soret, α and β bands at 433, 572 and 541 nm, respectively.

3.2.7 Determination of an Extinction Coefficient for Mtb CYP142A1 Using the Pyridine Hemochromogen Method

The extinction coefficient indicates how much light a protein (or other molecule)

absorbs at a certain wavelength. The CYP142A1 extinction coefficient was

determined using the pyridine hemochromogen method (Berry and Trumpower,

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1987). Figure 3.20 shows spectroscopic features for oxidized CYP142A1 along with

the spectra for the oxidized pyridine hemochrome form after reaction with sodium

dithionite. The heme concentration was calculated from the difference in

absorbance generated by subtraction of the oxidized hemochrome spectrum from

that of the reduced hemochrome (see Materials and Methods section 2.2.8). The

hemoprotein concentration was determined using ∆555 = 23.98 mM-1 cm-1 to give

an extinction coefficient of 92 mM-1 cm-1 for CYP142A1 in its oxidized state at the

heme Soret peak (418 nm).

This method was recently used to determine the heme Soret coefficient of

CYP144A1 (ε420.5 = 100 mM−1 cm−1) (Driscoll et al., 2010), and to determine ε416.5 =

110 mM−1 cm−1 for CYP121A1, and ε419 = 134 mM−1 cm−1 for CYP51B1 (McLean et al.,

2006b). A coefficient of ε417 = 115 mM−1 cm−1 was also reported for the P450cam

(CYP101A1) camphor hydroxylase from Pseudomonas putida (Dawson et al., 1982).

P450cin (CYP176A1), an enzyme that degrades cineole from Citrobacter braakii, had

its extinction coefficient determined at 150 mM−1 cm−1 using the same method

(Hawkes et al., 2002).

In recent studies involving Mtb P450 enzymes (CYP51B1, CYP125A1, CYP124A1 and

CYP130A1) (Bellamine et al., 1999, Ouellet et al., 2009), the method of extinction

coefficient determination used was that of Omura and Sato and based on the

development of Soret absorption of the Fe2+-CO complex at, or near, 450 nm

(Omura and Sato, 1964). However, there are some limitations with this method, at

least with respect to CYP51B1 (where the thiolate-coordinated P450 form

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consistently collapses to the thiolate-coordinated P420 species with Soret

maximum at ∼420 nm) and CYP125A1 (where a mixture of P450 and P420 species is

formed in the Fe2+-CO complex) (Bellamine et al., 1999, McLean et al., 2002a,

Aoyama et al., 1998, Dunford et al., 2007).

Figure 3.20: Pyridine hemochromagen spectra for CYP142A1. The main panel shows spectra for oxidized, substrate-free CYP142A1 (5.6 μM, black line), the oxidized pyridine hemochrome form following reaction with pyridine (red line), and the reduced pyridine hemochrome following reduction with sodium dithionite (green line). The inset shows details of these spectra. An extinction coefficient of ε418 = 92 mM-1 cm-1 for oxidised, substrate-free CYP142A1 was calculated from the difference between oxidized and reduced pyridine hemochrome spectra, using Δε555 = 23.98 mM-1 cm-1.

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3.2.8 Light Scattering (MALLS) Analysis of CYP142A1

In order to examine the aggregation state of the pure CYP142A1 enzyme, size

exclusion chromatography (SEC) coupled to MALLS (multiangle laser light

scattering) analysis was performed. A pure sample of CYP142A1 was resolved by

SEC on a Superdex 200 gel filtration column before passing through MALLS and

refractive index (RI) detectors.

One of the most important parameters for characterizing macromolecules such as

proteins, polysaccharides, oligonucleotides, and antibodies is their molecular

weight and/or molecular weight distribution. Different techniques have been used

to investigate protein aggregation. These techniques include: size exclusion

chromatography (SEC) (Carpenter et al., 1999), native gel electrophoresis, analytical

ultracentrifugation, circular dichroism, fluorescence spectroscopy, Fourier

transform/IR spectroscopy, UV spectroscopy and light blockage tests, and visual

inspection (Arakawa and Kita, 2000, Charman et al., 1993, Zuo et al., 2003, Borchert

et al., 1986, Wang, 1999). Among these different techniques, SEC is a simple and

fast method for determination of the molecular weight of a protein based on its

elution profile (Ye, 2006). Light scattering studies were performed on CYP142A1 to

determine its molecular weight and homogeneity. This was carried out as a crucial

step leading to crystallographic studies. Studies were done with buffers containing

no salt, 150 mM NaCl and 300 mM NaCl, both in the presence and absence of 1 mM

DTT (dithiothreitol), as described in the Materials and Methods section (section

2.2.13).

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In the absence of DTT and NaCl, MALLS data show one peak with a higher than

expected molecular weight prediction of 61 kDa, and eluting earlier than expected

from the column at between ~12.2-13.8 ml. This high molecular weight and broad

elution profile suggests a mixture of monomer and dimer (Figure 3.21A). Results

with 150 mM NaCl in the buffer showed two peaks, with the smaller peak giving a

molecular weight of 94.2 kDa (dimer) and the larger peak giving a molecular weight

of 45.9 kDa (monomer) (Figure 3.21B). A similar result was obtained when the salt

concentration was increased to 300 mM, with protein eluting much later at about

13.0-15.8 ml of buffer (Figure 3.20C).

When treated with DTT (1 mM), CYP142A1 was completely monomeric in both the

presence and absence of salt. The apparent molecular weight was close to the

predicted mass of 46.6 kDa from the CYP142A1 amino acid sequence. However, a

higher apparent molecular weight species was observed in the absence of salt

(Figure 3.22 A-D). These results indicate that DTT plays a major role in the

elimination of the dimeric species, likely through reduction of a disulfide bridge

across two CYP142A1 monomers. Elution volume was also delayed with increased

salt concentration, similar to the phenomenon observed in the absence of DTT

(Figure 3.21).

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Figure 3.21: Light scattering (MALLS) data for CYP142A1 in the absence of DTT. A: CYP142A1 in the absence of DTT and NaCl. Data shows one peak with a predicted molecular weight of 61.0 kDa that is higher than the true monomer MWt. CYP142A1 elutes early at about 12.2-13.8 ml buffer volume. B: CYP142A1 with 150 mM NaCl, minus DTT. The sample shows two peaks with the smaller peak giving a MWt of 94.2 kDa (dimer) and the larger peak giving a MWt of 45.9 kDa (monomer). The protein elutes between about 12.5-15.8 ml buffer. C: CYP142A1 with 300 mM NaCl, minus DTT. This sample shows two peaks with the smaller peak having a MWt of 94.1 kDa (dimer) and the larger peak a MWt of 46.4 kDa (monomer). The protein elutes between 13.0-15.8 ml buffer. D: Superimposed data for the three conditions. Green (No NaCl, minus DTT), Black (150 mM NaCl, minus DTT), Red (300 mM NaCl, minus DTT).

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Figure 3.22: Light scattering (MALLS) data for CYP142A1 in the presence of DTT. A: CYP142A1 in the presence of 1 mM DTT, minus NaCl. Data indicate a molecular weight (47.8 kDa) higher than that predicted for the monomer, and early elution at ~12.2-13.2 ml of buffer. B: CYP142A1 with 150 mM NaCl, plus DTT. This sample has a MWt of 44.5 kDa and protein elutes at ~14.4-15.8 ml buffer. C: CYP142A1 with 300 mM NaCl, plus DTT. This sample has a MWt of 44.9 kDa and protein elutes at ~15-16 ml buffer. D: Superimposed data for the three conditions. Green (No NaCl, plus DTT), Black (150 mM NaCl, plus DTT), Red (300 mM NaCl, plus DTT). CYP142A1 was completely monomeric in all three conditions.

The dimerization properties and activities of some enzymes, transcription factors,

sensor proteins and transcription factor modulators are determined by redox-

sensitive cysteine residues and disulfide (S-S) bonds in their subunits, which

perform redox-sensing roles to modulate the protein/enzyme functions (Hu et al.,

2011, Ramjeesingh et al., 1999). Disulfide (S−S) bond reducing agents or thiol-

reducing reagents (e.g. DTT and β-mercaptoethanol) may break the disulfide bond

between cysteine residues to modify the protein/enzyme function via

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conformational or other changes (Hu et al., 2011, Ramjeesingh et al., 1999). These

compounds reduce and break the disulfide (S−S) bonds and leave their cysteine

residues in a reduced (SH) state (Hu et al., 2011).

Studies have revealed that some Mtb P450s, like CYP121A1 and CYP125A1, exist as

monomeric species (Driscoll et al., 2011) while others appear to be dimers, e.g. Mtb

CYP130A1 (Ouellet et al., 2008). The Bacillus megaterium flavocytochrome P450

BM3 (CYP102A1, BM3) enzyme is also dimeric. In BM3, a soluble P450:P450

reductase (CPR) fusion protein dimerizes to enable electron exchange between

monomers to activate fatty acid hydroxylation (Neeli et al., 2005, Kitazume et al.,

2007). Oligomers of a P450 were also reported in the case of CYP3A4 (Davydov et

al., 1999). In the present study, however, the tendency of CYP142A1 to dimerize in

solution is likely to be as a result of the presence of intermolecular disulphide bonds

between cysteine residues on the protein surface of CYP142A1. As shown in Figure

3.23, analysis of the crystal structure of CYP142A1 reveals Cys296 and Cys316 to be

exposed on the P450 surface. It is thus likely that disulfide bond(s) can form

between pairs of these residues on different monomers to form dimeric species.

Cleavage of the disulfide bond(s) on DTT treatment then results in the formation of

monomeric species.

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Figure 3.23: Cysteine residues in CYP142A1. A CYP142A1 transparent surface representation is shown, with cysteine residues (in yellow spacefill) shown on the surface protein surface and within the macromolecule. Two cysteine residues are clearly present on the surface of CYP142A1 (Cys296 and Cys316), while most are buried inside the molecule. It is likely that partial dimerization of CYP142A1 occurs through disulfide bonds between these cysteines (from different monomers) and that DTT treatment breaks the disulfide bonds to restore the monomeric state.

3.2.9 Electron Paramagnetic Resonance (EPR) Analysis of CYP142A1

3.2.9.1 EPR analysis with selected CYP142A1 ligands

EPR is a technique used to study molecules (e.g. proteins) with unpaired electrons,

and can provide important information about the state of ferric heme (and other

EPR active cofactors) as well as e.g. providing details of the interactions of

substrates and inhibitors with heme and other cofactors in proteins and enzymes

(Andersson and Barra, 2002). EPR spectra were recorded (as described in Materials

and Methods, section 2.2.15) to probe CYP142A1 heme iron coordination and the

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effects of CYP142A1 binding to various ligands. Continuous wave X-band EPR data

were collected at 10 K for both ligand-free and ligand-bound forms of CYP142A1,

and revealed a characteristic rhombic signal for ferric P450s. Figure 3.24 shows

overlaid X-band EPR spectra for native, cholestenone-, econazole- and DMSO-

bound forms of CYP142A1.

Figure 3.24: EPR analysis of CYP142A1 and various ligand complexes. Panel A. EPR features for CYP142A1 in the major region for detection of high-spin heme iron. Negligible high-spin features were detected for the ligand-free, DMSO-bound or econazole-bound forms of CYP142A1. In the cholestenone substrate-bound form, high-spin features were detected at gz = 7.95 and gy = 3.61. Panel B. Low-spin ferric heme EPR spectra for the same species as in panel A. The ligand-free CYP142A1 has a low-spin rhombic trio at g = 2.40/2.23/1.92, typical of a cysteine thiolate/water coordinated P450. A minor species with gz = 2.48/gx = 1.90 suggests a proportion of a species with a stronger (e.g. nitrogen) distal ligand to the heme iron. The cholestenone-bound CYP142A1 has a similar spectrum to ligand-free P450. The econazole-bound CYP142A1 shows a single set of EPR values at g = 2.47/2.25/1.90, consistent with coordination of the heme iron by an econazole nitrogen atom. The DMSO/CYP142A1 sample shows (in addition to the ligand-free type species at 2.40/2.24/1.92) two other species at 2.42/2.24/1.91 and at 2.47/2.24/1.89. While the gz = 2.47 species may be the same one as observed in the ligand-free enzyme, it is also possible that these species result from the interactions of DMSO oxygen/sulfur atoms with the CYP142A1 heme iron.

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The EPR spectrum of ligand-free CYP142A1 displays features attributable to a S =

1/2 LS ferric heme iron with a thiolate-proximal ligand to the iron and a distal water

ligating the heme (Figure 3.23). The g-values observed for the ligand-free CYP142A1

are g = 2.40/2.23/1.92, with a minor species at 2.48/2.23/1.90. The latter species

may result from an altered coordination state of the distal water ligand, or possibly

from the interaction of a stronger ligand (e.g. a distal nitrogen) in a small

proportion of the enzyme. These data are consistent with a low-spin P450 enzyme

and are similar to EPR spectra previously reported for the Mtb cholesterol 27-

hydroxylase CYP125A1 (2.40/2.25/1.94) and the cyclodipeptide oxidase CYP121A1

(2.47/2.25/1.91), both of which show LS EPR signals (McLean et al., 2008, McLean

et al., 2009). In the case of CYP121A1, there was no evidence for distal nitrogen

coordination, and thus the gz = 2.47 value may instead be consistent with a

particular organisation of water molecules around the 6th ligand water molecule.

The heterogeneity observed in the ligand-free CYP142A1 spectrum might also be

consistent with that observed with CYP125A1, which was suggested to arise from

different conformers of the P450 with altered angles of the axial ligands (McLean et

al., 2009). Even though CYP125A1 is extensively high-spin at ambient temperature,

the heme iron is almost completely low-spin at the cryogenic temperatures (10K)

required for heme EPR (McLean et al., 2009).

In its complex with cholestenone, CYP142A1 has a similar set of LS g-values to those

seen for the ligand-free form (2.40/2.24/1.92 and a minor signal at 2.47/2.24/1.90),

but also a small signal indicative of high-spin heme iron with gz/gy values at

7.95/3.61. There is clearly the retention of a small amount of high-spin heme iron in

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the cholestenone-bound form, despite the very low temperature used (10 K),

suggesting that the binding mode of the steroid is one in which the axial water

ligand is effectively displaced by the substrate and a proportion of high-spin heme

iron is maintained even at cryogenic temperatures. In its complex with econazole,

CYP142A1 shows a homogeneous set of g-values (2.47/2.25/1.90) indicative of the

replacement of the water ligand by the azole nitrogen. Interestingly, this contrasts

with data generated for econazole-bound Mtb CYP144A1, in which there was

heterogeneity, with g-values at 2.62 (minor)/2.45 (major), 2.26 and 1.89 (Driscoll et

al., 2011), suggesting a more complete coordination of the CYP142A1 heme iron by

econazole. Other recent studies showed near-identical g-values for the fluconazole

complex of Mtb CYP51B1 (2.45, 2.26, 1.90) with that for the CYP121A1-fluconazole

complex (McLean et al., 2008, McLean and Munro, 2008).

The addition of the solvent DMSO produced a perturbation of the CYP142A1 LS EPR

spectrum, with two sets of g-values being near-identical to those for the ligand-free

CYP142A1, but a third set (2.42/2.24/1.91) being distinct. These data may indicate

that the DMSO affects the solvent environment around the heme iron, or could

possibly reflect direct interactions with DMSO solvent oxygen and/or sulfur atoms

with the heme iron (Kuper et al., 2012).

3.2.9.2 EPR analysis for CYP142A1 fragments hits

EPR analysis also carried out on CYP142A1 complexes with fragments generated

from fragment based screening studies (Figure 3.25). These fragments are NMR170,

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NMR491, NMR623, NMR540, and NMR089. This study is a follow-up to the UV-Vis

spectroscopy analyses presented earlier (section 3.2.4) to study the interactions of

these compounds with the P450. These fragments are imidazoles or substituted

imidazoles (NMR’s 540, 089, 491 and 623), or pyridine ring-containing (NMR540).

They thus have potential to interact with the CYP142A1 heme iron via nitrogen

atom interactions (and potentially either directly with the heme iron or via a water

molecule on the sixth axial position, as observed for fluconazole with CYP121A1)

(Seward et al., 2006). EPR data generated for CYP142A1 binding to these fragments

were compared with those for their interactions with three different azole

inhibitors (clotrimazole, 1-PIM and 2-PIM). From results obtained, heterogeneity of

low-spin EPR spectral features was observed for most of the fragments, with low-

spin EPR spectra observed in all cases. CYP142A1 in complex with 1-PIM gives a

homogeneous set of g-values (gz = 2.47, gy = 2.25, gx = 1.89) indicative of the

replacement of the water ligand by the imidazole nitrogen. NMR491, NMR089,

NMR540, NMR170, 2-PIM and clotrimazole all showed splitting of the g-values for

their CYP142A1 complexes, suggesting heterogeneity and the presence of 2-3 low-

spin species with different distal coordination states of the heme iron. For example,

CYP142A1 in complex with 2-PIM exhibits three sets of g-values at 2.51/2.24/1.88

and 2.47/2.24/1.89 (major) and at 2.40/2.24/1.90. The final form may be similar to

the ligand-free form (possibly with 2-PIM influencing the environment of the distal

water ligand), while the other two (major) species likely originate from the

coordination of the heme iron by the imidazole group, possibly with altered

geometries of Fe-azole bonding, or (for the gz = 2.47 species) a species where the 2-

PIM interacts indirectly with the CYP142A1 heme iron via the distal water that

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remains as the heme 6th ligand, as was revealed in a recent study on the structure

of the CYP121A1-fluconazole complex (Seward et al., 2006). Among the other

azoles and fragments tested, the pyridine-containing NMR170 clearly coordinates

to the CYP142A1 heme iron in two states, with g-values of 2.54/2.25/1.87 and

2.47/2.24/1.89, while 1-phenylimidazole gives a relatively homogeneous heme-

coordinated species at 2.47/2.25/1.89. The substituted imidazole NMR623 also

gives a relatively homogeneous EPR spectrum, with the main species having g-

values of 2.44/2.25/1.90, and with a minor species at 2.57/2.25/1.85.

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Figure 3.25: EPR analysis of interactions of azole drugs and nitrogen-containing fragments with CYP142A1. X-band EPR spectra are shown for CYP142A1 (200 μM) in its ligand-free form and in complex with fragments and selected azole drugs. The g-values are labelled on each spectrum. Heterogeneous low-spin heme iron signals were obtained on binding many of the azoles and fragments, arising from either direct ligation of a heterocyclic nitrogen in the fragment/ligand to the heme iron (potentially with different geometries of coordination), or through their making indirect interactions with the heme iron via the (6th) water ligand that remains on the heme iron following ligand addition.

3.2.9.3 EPR analysis of CYP142A1 bound to MEK compounds

From the UV visible spectroscopy, MEK065, MEK046 and MEK066 showed

substantial spectral shifts and these were selected for further analysis using the EPR

technique. In addition to ligand binding investigation, EPR is also used to probe

heterogeneity in spin state of the ferric heme iron. EPR analysis of complexes of

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CYP142A1 with MEK065 and MEK066 (which were reported earlier in this thesis to

produce type I spectral shift in UV-visible spectroscopy titration) revealed some

heterogeneity, with a mixture of low-spin and high-spin species observed in X-band

EPR studies (Figure 3.26). The MEK065 complex with CYP142A1 gave

heterogeneous low-spin g-values at 2.51/2.24/1.91 (minor) 2.45/2.24/1.91 (minor)

and 2.40/2.24/1.93 (major). However, the CYP142A1/MEK066 complex exhibited

much less heterogeneity in the low-spin spectrum, with g-values of 2.45/2.24/1.90

(minor) and 2.40/2.24/1.93 (major). In the high-spin spectral region, there were

small signals for both the MEK065 (gz = 8.01, gy = 3.54) and the MEK066 complexes

(gz = 7.92, gy = 3.42). The high-spin EPR features are consistent with binding data

from UV-visible spectroscopy, where an ~2 nm Soret shift towards a high-spin

species were observed for the binding of both MEK065 and MEK066 to CYP142A1

(section 3.2.5). The retention of a high-spin component at 10 K is again likely due to

the displacement of the axial water ligand on the heme iron by these molecules in

at least a proportion of the CYP142A1 enzyme, as also observed with cholestenone

in section 3.2.9.1 above.

With MEK046, a type II binder, the spectrum has two major components at (i)

2.62/2.25/1.84 and (ii) 2.47/2.25/1.89. These spectra likely result from the direct

interaction of the heterocyclic imidazole nitrogen of this compound with the heme

iron (i) and possibly from an indirect interaction of the drug with a retained distal

water (ii). Consistent with ligation of the heme iron, there was no significant high-

spin EPR signal for the CYP142A1/MEK046 complex. In the case of the MEK065 and

066 compounds, these contain thiophene and furan rings in the same position as

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the imidazole group in MEK046, and these are likely to be less effective ligands for

the P450 heme iron. DMSO was used as a solvent for these ligands, and data for its

mixture with CYP142A1 revealed some perturbation of the EPR spectrum with g-

values of 2.46/2.24/1.91 and 2.40/2.24/1.93. The former set of g-values likely

reflects the interaction of the DMSO sulfur atom with the heme iron (which may

also be observed to a lesser extent in the other complexes), while the latter set is

consistent with the resting state of CYP142A1 with thiolate/water ligands to the

ferric heme iron.

Figure 3.26: X-band EPR spectra for CYP142A1 in complex with different compounds from the MEK series. The MEK compounds originated as ligands for CYP121A1, but also show affinity for other Mtb P450s. The g-values for different low-spin and high-spin X-band EPR features are labelled on each spectrum. Panel A: High-spin ferric heme EPR features. Panel B: Low-spin ferric heme EPR features. There is clear evidence for the distal coordination of the CYP142A1 heme iron by the imidazole-containing MEK046, whereas both MEK065 and 066 also clearly bind to CYP142A1 and induce formation of a proportion of high-spin heme iron in the enzyme.

3.2.10 Differential Scanning Calorimetry studies of CYP142A1

Differential scanning calorimetry (DSC) is a thermodynamic tool often used for

probing thermal denaturation of proteins (Goyal et al., 2014, Gill et al., 2010). It is

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commonly used in the pharmaceutical industry to study thermal stability of

proteins, including analysis of their overall conformation, phase transitions, and

domain folding properties (Arthur et al., 2015, Johnson, 2013). DSC is a technique

that measures heat capacity as a function of temperature, and protein unfolding

transitions (Arthur et al., 2015). Parameters in the DSC thermogram, such as the

transition midpoint (which is also referred to as the ‘melting temperature’, Tm), can

be used to investigate the thermal stability of proteins under various conditions

(Cederbaum, 2014). The Tm is an important indicator of thermostability and it is

postulated that the higher the Tm, the more thermodynamically stable is a protein

(Bruylants et al., 2005).

Furthermore, the Tm of a protein is an important parameter that can reveal other

useful properties of the molecule, such as its propensity for successful

crystallization for high-resolution structural studies (Dupeux et al., 2011). A recent

study revealed that samples with Tm values of 318 K or higher crystallized in 49% of

cases, while the success rate of crystallization declined rapidly for samples with

lower Tm values, with only about 23% of samples with a Tm below 316 K producing

crystals (Dupeux et al., 2011). Hence conditions, or stabilising additives, that elevate

Tm are important tools utilized to optimise success rates in crystallization

experiments (Dupeux et al., 2011, Ericsson et al., 2006).

Thermal stability of CYP142A1 was studied by DSC, which was used to monitor heat

absorption and conformational unfolding. DSC was performed as detailed in the

Materials and Methods (section 2.2.14) on the substrate-free, cholestenone-bound

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and clotrimazole-bound forms of the enzyme. The results showed that CYP142A1

displays single unfolding transitions for both the ligand-free and ligand-bound

forms of the enzyme. Cholestenone increased the CYP142A1 Tm value by

approximately 5 oC, while clotrimazole increased the Tm value by approximately 3

oC. This is indicative of stabilization (albeit to a small extent) of the P450 when

complexed with these ligands.

Figure 3.27: Differential scanning calorimetry analysis of CYP142A1. The black line is for the raw data and the red line is for the data fit, showing the protein unfolding transition events illustrated by changes in heat capacity with applied temperature The thermal unfolding profile is shown for a sample of 8 μM CYP142A1 in 10 mM KPi, 100 mM NaCl, pH7.2 buffer. The data were baselined and concentration corrected, and data were fitted using a non-2-state function using Microcal software (OriginLab). A: Ligand-free CYP142A1 with a Tm of 53.29 ± 0.06 oC. B: Cholestenone-bound CYP142A1 with a Tm of 58.43 ± 0.06 oC. C: Clotrimazole-bound CYP142A1 with a Tm of 56.06 ± 0.06 oC.

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Tm (oC) ∆H (cal mol-1) ΔHvH (cal mol-1)

Ligand-free CYP142A1 53.29 ± 0.06 1.26 x 105 ± 1.77 x 103 9.44 x 104 ± 1.65 x 103

CYP142A1 + cholestenone 58.43 ± 0.06 1.47 x 105 ± 2.22 x 103 1.05 x 105 ± 1.97 x 103

CYP142A1 + clotrimazole 56.06 ± 0.06 7.45 x 104 ± 1.65 x 103 1.45 x 105 ± 3.84 x 103

Table 3.3: DSC data for the thermal unfolding of CYP142A1. The thermal transition midpoints (Tm values), calorimetric enthalpy (ΔH) and Van’t Hoff enthalpy (ΔHvH) are shown for the single transition events observed for ligand-free CYP142A1 and for its complexes with cholestenone substrate and clotrimazole inhibitor.

DSC measures enthalpy (∆H) of unfolding as a result of thermal denaturation of

macromolecules (Gill et al., 2010). Calorimetric enthalpy (∆Hcal) refers to the total

integrated zone below the thermogram peak (or apparent area under the peak),

which indicates total heat energy absorbed by the sample in the experiment

(Holdgate, 2009). The van’t Hoff enthalpy (ΔHvH) is an independent measurement

of the enthalpy of the transition associated with the model of the experiment (Gill

et al., 2010) in the sense that, if ΔHvH is equal to ∆Hcal, the transition occurs in a

two-state mode. This was not the case with CYP142A1, in which the transition

occurred in a single-state mode for the ligand-free and ligand-bound enzymes.

3.2.11 Guanidinium chloride denaturation of CYP142A1

The folded conformation adopted by a protein is completely determined by its

amino acid sequence (Almeida Da Silva and Palomino, 2011). A protein is

synthesised as a linear unfolded polypeptide form, but rapidly folds into its

biologically active structure (Silva-Lucca et al., 2013). Protein unfolding can be

investigated in vitro using chemical denaturants or chaotropic agents, as well as by

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using pH or temperature variations. Chaotropic agents such as guanidinium chloride

(GdmCl) or urea are frequently used to study protein unfolding/refolding, even

though the unfolding mechanism remains unclear. These studies are commonly

followed by methods including far UV-CD (to follow secondary structural change)

and protein fluorescence assays (to measure changes in protein tertiary structure)

(Almeida Da Silva and Palomino, 2011). Intrinsic fluorescence emission provides a

sensitive and effective tool to characterize proteins by monitoring tryptophan

residues that are very sensitive to the polarity of the environment. The wavelength

at the emission peak (λmax) and the fluorescence intensity can be used to study

protein unfolding/refolding and to determine conformational changes (Eftink,

1998).

In this study, experiments were carried out to investigate CYP142A1

unfolding/denaturation using increasing concentrations of GdmCl in the presence

and absence of the substrate cholestenone, followed using fluorescence assays.

Aromatic amino acid fluorescence (with excitation at ~280 nm, close to the

absorbance peak for tryptophan) provides emission spectra reporting on protein

tertiary structure. Structural perturbations induced by GdmCl were monitored by

following fluorescence changes from aromatic amino acids (mainly tryptophans) in

the emission spectra (in the range from ~300-450 nm). During GdmCl-dependent

denaturation of CYP142A1, changes in both the fluorescence emission intensity and

the max of fluorescence were observed. CYP142A1 (5 µM) was incubated with

increasing concentrations of GdmCl (0-6 M) for 30 minutes in both the absence and

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presence of cholestenone (60 µM). Experiments were performed as described in

the Materials and Methods (section 2.2.11).

The effects of GdmCl on the fluorescence emission spectra of CYP142A1 are shown

in Figure 3.28. The intrinsic fluorescence emission spectrum of resting, substrate-

free CYP142A1 has a maximum (λmax) at 330 nm, which is red shifted to 358 nm and

accompanied by a pronounced increase in fluorescence intensity as the

concentration of GdmCl is increased. This indicates protein unfolding and the

exposure of tryptophan residues to a more polar environment. To better examine

the conformational changes induced by GdmCl on CYP142A1 in the absence and

presence of cholestenone, the percentage of unfolded CYP142A1 (taken as the ratio

of fluorescence emission near the maximum for the unfolded enzyme at 358 nm,

divided by the emission near the maximum for the native enzyme at 333 nm – i.e.

F358/F333) was plotted against the relevant GdmCl concentration and the data fitted

using a sigmoidal (Hill) function. From the fits, the midpoint values of 1.72 ± 0.05 M

GdmCl (plus cholestenone) and 1.77 ± 0.04 M GdmHCl (minus cholestenone) were

determined (Figure 3.28).

The sigmoidal curves suggest that cooperative conformational changes are induced

during GdmHCl unfolding. These data, however, showed that cholestenone had no

significant tertiary structural stabilization effect on CYP142A1 against the

denaturant. In the absence of cholestenone (Figure 3.28), the fluorescence intensity

decreased with increasing concentrations of GdmCl until ~1.75 M, and then

increased at higher concentrations of the denaturant. However, in the presence of

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cholestenone (Figure 3.29), the fluorescence intensity increased continuously with

increasing GdmCl concentrations, suggesting that some conformational changes (or

other structural reorganization of the enzyme) are induced by binding of the

substrate. In both cases, there was a shift of the fluorescence emission maximum

(λmax) to a longer wavelength (red shift) as the proteins unfolded.

Figure 3.28: Guanidinium chloride denaturation of ligand-free CYP142A1. Panel A. Intrinsic fluorescence emission spectra recorded after incubation of CYP142A1 (5

M) with increasing concentrations of GdmCl (0-6 M) for 30 minutes in the absence of cholestenone. The emission spectral maximum is red shifted from 330 to 358 nm with increasing concentrations of GdmCl and as the protein unfolds. Panel B. shows a data plot for the percentage unfolded CYP142A1 (from the ratio of fluorescence values at 358 nm and 333 nm – F358/F333) as the concentration of GdmCl is increased. The data were fitted using the Hill function to give a midpoint value of 1.77 ± 0.04 M GdmCl (n = 2.73 ± 0.15).

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Figure 3.29: Guanidinium chloride denaturation of cholestenone-bound CYP142A1. Panel A. Intrinsic fluorescence emission spectra recorded after

incubation of CYP142A1 (5 M) with increasing concentrations of GdmCl (0-6 M) for

30 minutes following the addition of cholestenone (60 M) to the P450. The spectra become red shifted from 331 nm to 358 nm with increasing concentrations of GdmCl and as the protein unfolds. Panel B shows a data plot for the percentage unfolded CYP142A1 (from the ratio of fluorescence values at 358 nm and 331 nm – F358/F331) as the concentration of GdmCl is increased. The data were fitted using the Hill function to give a midpoint value of 1.72 ± 0.05 M GdmCl (n = 3.32 ± 0.27).

3.2.12 Isothermal Titration Calorimetry (ITC) analysis of

CYP142A1

ITC is an important technique used to study protein-ligand binding, and is also

being increasingly used to study protein-protein interactions (Velazquez-Campoy et

al., 2004). When two proteins bind or when a ligand binds to a protein, there are

changes in the thermodynamic parameters (∆G, ∆H, ∆S), which can be detected by

highly sensitive calorimetric methods such as ITC (Velazquez-Campoy et al., 2004).

The thermodynamics of association are measured by the stoichiometry of the

interaction (n), the association constant (Ka), the free energy (ΔG), enthalpy (ΔH),

entropy (ΔS), and heat capacity of binding (ΔC) (Brautigam, 2015).

215

Figure 3.30: Isothermal titration calorimetric (ITC) binding studies of fragments to

CYP142A1. The calorimetric enthalpy changes (upper panels) and the resulting binding

isotherms (lower panels) are shown for titrations of CYP142A1 with NMR491 (A), 1-

phenylimidazole (B), NMR170 (C), NMR623 (D), and NMR540 (E). The data were best

fitted using a one-step binding model. The binding parameters obtained are detailed in

Table 3.4.

Binding of fragments to CYP142A1 was investigated by ITC to probe the

thermodynamics of protein-fragment interactions. The experiment was performed

as described in the Materials and Methods (section 2.2.10). Out of all the fragments

tested in the study, the experiment proved successful with five of the fragment hits

and the data for these are presented in Figure 3.30. The data were best fitted using

216

a one-step binding model. The thermodynamic parameters derived from the

analysis are summarised in Table 3.4. Binding of the fragments to CYP142A1

revealed an exothermic driven process, as evidenced by the large and negative ∆H

and ∆G values. The Kd value obtained from ITC for the binding of NMR491 (26.65

M) was much higher than that obtained from UV-visible spectroscopic binding

analysis (0.68 M), which is consistent with a previous comparative ligand-binding

study carried out on CYP130A1 (Ouellet et al., 2008). However, the ITC Kd values

obtained for NMR170, NMR540, NMR623 and 1-PIM closely match the

spectroscopic dissociation constant values, and thus the Kd value for NMR491 is the

only outlier Kd from the two techniques used. Though the values derived from ITC

are all slightly higher, the trend of ligand affinity for CYP142A1 is similar for both

techniques and (apart from NMR491) the values appear to converge better as the

ligand affinity gets weaker.

Thermodynamic Parameters Fragments

Ka * 104(M

-1) Kd

cal (1/Ka)

(µM) Kd

optical

(µM) ∆H (kcal/mol/K)

∆Sb

(cal/mol/K)

∆Ga

(kcal/mol)

N

NMR491 3.75 ± 0.31 26.65 0.68 -9.86 ± 0.91 -12.13 -6.24 0.40

NMR170 21.53 ± 2.68 4.64 1.87 3.50 ± 0.13 12.33 -7.27 0.44

NMR540 0.47 ± 0.11 213.36 175.82 -0.96 ± 0.31 13.57 -5.01 2.20

NMR623 0.37 ± 0.07 272.63 253.77 -1.12 ± 0.21 12.56 -4.86 4.31

1-Phenylimidazole 3.92 ± 0.65 25.46 23.20 -11.17 -16.46 -6.26 0.41

Table 3.4: Thermodynamic parameters for CYP142A1-fragment interactions derived from ITC and optical titrations. a∆G = ∆H-T∆S; b∆S = (∆H-∆G)/T.

217

3.2.13 Redox potentiometry of CYP142A1

The cytochrome P450 enzymes contain ferric heme iron in their resting states, and

a change in spin-state equilibrium of the heme iron from predominantly low-spin (S

= 1/2) to predominantly high-spin (S = 5/2) occurs on substrate binding in most

cases (McLean et al., 2007a). This is as a result of the displacement of the 6th

(water) ligand to the heme iron (Driscoll et al., 2011). Substrate binding to P450

enzymes in this way mostly leads to an elevation of the heme iron reduction

potential by about 130-140 mV, meaning that the ferric heme iron potential

becomes more positive. This positive shift in potential facilitates/accelerates heme

iron reduction by the NAD(P)H-dependent redox partner enzymes as a

consequence of a greater driving force for electron transfer to the heme iron (Daff

et al., 1997).

This is an important regulatory process, allowing efficient heme reduction only

when the P450 is substrate-bound, and avoiding production of damaging oxygen

radicals (H2O2 and superoxide). Figure 3.31 shows spectral data from a redox

titration of CYP142A1 in the ligand-free form. On reduction of the heme iron from

Fe3+ to Fe2+, there is a decrease in heme absorption with a blue shift from 418 nm to

~405 nm, and the merging of the α and β bands at longer wavelength.

Previous redox potentiometry studies performed on ligand-free Mtb CYP51B1

showed a Soret shift to a longer wavelength from 419 nm to 423 nm, and with a

significant feature developing at 558.5 nm for the reduced enzyme (McLean et al.,

218

2006b). CYP144A1 also exhibited similar features with a Soret shift from 420.5 nm

to 425 nm (Driscoll et al., 2011). The spectral changes revealed by CYP144A1 and

CYP51B1 on heme reduction are indicative of the heme iron becoming thiol-

coordinated in the ferrous state (McLean et al., 2006b, Driscoll et al., 2011).

However, in contrast to the above results, redox titration of CYP121A1 revealed a

shift in the Soret band from 416.5 nm to 407 nm on heme reduction and similar

results were obtained from P450 BM3 and P450cam enzymes, indicating the

retention of heme thiolate coordination in the ferrous enzyme (McLean et al., 2008,

Daff et al., 1997, Sligar and Gunsalus, 1976). The spectral changes observed for

CYP142A1 on heme reduction with dithionite are thus consistent with the heme

iron retention of cysteine thiolate coordination in the ferrous enzyme (Figure

3.31A).

Figure 3.31 panel B shows a fit of the CYP142A1 heme absorbance at 418 nm versus

applied potential data, using the Nernst equation. This produces a CYP142A1 midpoint

potential of −394 ± 4 mV versus the normal hydrogen electrode (NHE). This value is

somewhat more negative than the values determined for some other Mtb P450s e.g.

CYP144A1 (−355 ± 5 mV), CYP51B1 (−375 ± 5 mV) and CYP125A1 (−303 ± 5 mV).

However, the midpoint potential obtained for ligand-free CYP142A1 (−394 ± 4 mV) is

consistent with the values determined for CYP121A1 and for some of its mutants, which

were all found to be more negative than -400 mV (McLean et al., 2008).

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Figure 3.31: Redox potentiometry of ligand-free CYP142A1. The main panel shows spectra from a redox titration of ligand-free CYP142A1 (∼8.5 μM). The arrows indicate the direction of absorption changes occurring during the reductive phase of the titration, as the ferric heme iron is reduced to the ferrous form. The ferric heme Soret band at 418 nm (black solid line) decreases in intensity and shifts to 405 nm (red solid line) in the ferrous state. In the visible region, an absorption band develops at 550 nm on heme reduction with a merging of the α and β bands. The Inset shows a plot of heme absorbance change at 418 nm against the applied potential (versus the normal hydrogen electrode, NHE), and with the data fitted using the Nernst equation to produce a midpoint potential of −394 ± 4 mV for the heme iron Fe3+/Fe2+ transition.

Figure 3.32 (main panel) shows spectral data from a redox titration of CYP142A1

complexed with cholestenone. On reduction of the heme iron from Fe3+ to Fe2+,

there is a decrease in heme absorption with a red shift from 393 nm to 405 nm. The

Figure 3.32 inset shows a plot of the heme absorbance at 393 nm versus applied

potential, with data fitted using the Nernst equation, producing a CYP142A1

midpoint potential of −150 ± 4 mV versus NHE. When compared to the midpoint

220

potential for the ligand-free CYP142A1 (−394 ± 4 mV), the large potential change

(~240 mV) between the low-spin (substrate-free, SF) and high-spin (substrate-

bound, SB) forms reflects a major change in heme iron spin-state on cholestenone

binding and a tightly regulated redox system. Studies have revealed that the redox

potential for CYP51B1 is −375 ± 5 mV in the ligand-free form and this is elevated to

−225 ± 8 mV for the estriol-bound form (giving a difference of 150 mV), in which

the estriol-bound protein becomes extensively high-spin (McLean et al., 2006b).

The ligand-free cholesterol hydroxylase CYP125A1 has a more positive heme

potential of −303 ± 5 mV, consistent with the more HS nature of this Mtb P450

(McLean et al., 2009). In addition, the midpoint redox potential of the P450 BM3

heme iron is −427 mV in the ligand-free form elevating to −289 mV when

complexed with the substrate arachidonic acid, giving a difference of 138 mV (Ost

et al., 2001). The much larger potential change (~250 mV) between the low-spin

(SF) and high-spin (SB) forms of CYP142A1 is, however, unusual and the precise

reasons for this magnitude of change are unclear.

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Figure 3.32: Redox potentiometry for cholestenone-bound CYP142A1. The main panel shows spectra from a redox titration of Mtb CYP142A1 (∼5.4 μM) in complex with cholestenone. The arrows indicate the direction of absorption changes occurring during the reductive phase of the titration, as the ferric heme iron is reduced to the ferrous form. The heme Soret band at 393 nm (ferric, solid black line) decreases in intensity and shifts to 405 nm (ferrous, red solid line). In the visible region, an absorption band develops at 550 nm on heme reduction. The inset shows a plot of heme absorbance change at 393 nm against the applied potential (versus the normal hydrogen electrode, NHE), and with the data fitted using the Nernst equation to produce a midpoint potential of −150 ± 4 mV for the heme iron Fe3+/Fe2+ transition.

Figure 3.33 (main panel) shows spectral data from a redox titration of CYP142A1

complexed with an azole inhibitor, clotrimazole. On reduction of the heme iron

from Fe3+ to Fe2+, there is an increase in heme Soret absorption with a red shift

from 424 nm to 428 nm. The Figure 3.33 inset shows a plot of the heme absorbance

at 424 nm versus applied potential, with data fitted using the Nernst equation,

producing a CYP142A1 midpoint potential of −360 ± 4 mV versus NHE. When

222

compared to the midpoint potential for the ligand-free (−394 ± 4 mV), there is a

rather small heme iron potential change (~34 mV) between the low-spin ligand-free

and low-spin clotrimazole-bound forms. A significant spectral change is observed in

the Q-band region, which is marked by increased absorbance intensity and the clear

development of two distinct absorption bands at 529 nm and 559 nm. This is

indicative of distal nitrogen ligation to the reduced CYP142A1 heme iron.

Figure 3.33: Redox potentiometry for clotrimazole-bound CYP142A1. The main panel shows spectra from a redox titration of Mtb CYP142A1 in complex with clotrimazole (∼7 μM). The arrows indicate the direction of absorption change occurring during the reductive phase of the titration in regions of the spectrum at which major changes occur. The heme Soret band at 424 nm (ferric, black solid line) increases in intensity and shifts to 428 nm (ferrous, red solid line). The α (532 nm) and β (558 nm) bands showed a marked increase in absorbance intensity on heme reduction. The inset shows a plot of heme absorbance change at 424 nm against the applied potential (versus the normal hydrogen electrode, NHE), and with the data fitted using the Nernst equation to produce a midpoint potential of −360 ± 4 mV for the heme iron Fe3+/Fe2+ transition

223

Midpoint potential (mV vs NHE)

Type of shift

Ligand-free CYP142A1 −394 ± 4 Blue

CYP142A1 + cholestenone −150 ± 4 Red

CYP142A1 + clotrimazole −360 ± 4 Red

Table 3.5: Redox potential data for CYP142A1. The midpoint potential values and the type of Soret shifts are indicated for the ligand-free CYP142A1 and for its complexes with cholestenone and clotrimazole.

3.2.14 Nanoelectrospray Ionization Mass Spectrometric

Analysis of Mtb CYP142A1−Ligand Interactions Nanoelectrospray Ionization Mass Spectrometry (nanoESI) is a non-destructive tool

used to generate positively or negatively charged molecular protein ions that are

transferred into a given mass spectrometer (Brugger, 2014). NanoESI was first used

to analyse molecules of low molecular weight (Whitehouse et al., 1985) and then,

over time, its uses were extended to macromolecules such as oligonucleotides and

proteins (Fenn et al., 1989). A unique characteristic of nanoESI is its ability to

produce multiple charged ions, an attribute that is important for protein analysis

(Brugger, 2014). Among many important advantages of the nanoESI assay are its

sensitivity, selectivity, and the fact that experiments do not require immobilization

or labelling, and allow simultaneous measurements of multiple binding equilibria,

and require only minute (microgram) quantities of protein samples (El-Hawiet et al.,

2012). NanoESI is one of the softest ionization tools in current use and enables the

analysis of non-covalent molecular interactions (Benesch and Robinson, 2006).

In this work, nanoESI was used to characterize the oligomerization state of

CYP142A1 and to probe the binding stoichiometries and interactions of CYP142A1

224

in its ligand-free state and in complex with its substrate cholestenone, with the

azole inhibitor econazole and with DTT. All experiments were performed as

described in the Materials and Methods (section 2.2.19).

3.2.14.1 NanoESI mass spectra of ligand-free CYP142A1

Figure 3.34: NanoESI mass spectra of ligand-free CYP142A1. Samples contained 10 and 20 μM CYP142A1 in 200 mM ammonium acetate (pH 7.0). Peaks in the range m/z 3200 to 4400 were assigned to monomer (A) and in the range m/z 4800 to 7000 to dimer (B). The mass spectra recorded reveal two species, the major one at 47080 ± 6 Da (monomer) and the minor species at 94412 ± 48 Da (dimer, CYP142A1 at 20 µM). For CYP142A1 at 10 µM, the major species is at 47072 ± 6 Da (monomer) and the minor species is at 94109 ± 8 Da (dimer).

Some characterized Mtb P450s, such as CYP121A1, were shown to be dimeric in

solution (Duffell et al., 2013) and this is consistent with the findings derived from

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CYP142A1 characterization, as observed from the light scattering studies (section

3.2.8). Another example of an Mtb P450 that has shown evidence of dimerization is

CYP130A1. In recent studies, CYP130A1 was crystallized in the ligand-free form as a

monomer and in an econazole-bound form as a dimer. The ligand-bound “closed”

form of CYP130A1 also formed a dimer in solution (Ouellet et al., 2008). Hence, in

order to further probe the oligomerization state of CYP142A1, nanoESI mass

spectra were recorded at two different concentrations (10 µM and 20 µM) of the

protein (Figure 3.34). The signals corresponding to the monomer persisted at the

different CYP142A1 concentrations, while the weaker features in the range m/z

4800−7000 at 10 M CYP142A1 (corresponding to the dimer) increased as the

protein concentration was elevated to 20 M. In contrast, two other Mtb P450

enzymes, CYP125A1 and CYP126A1, were also investigated under the same

experimental conditions as those for which CYP121A1 was studied, and both were

exclusively monomeric (Duffell et al., 2013). The mass spectrum of CYP142A1 at 20

M has major peaks in the m/z range from 3200 to 4200, assigned to the

monomeric CYP142A1, with a molecular weight of 47080 ± 6 Da, close to the value

computed from the amino acid sequence. Weaker intensity peaks, in the range

from m/z 4800 to 6800, were assigned to dimeric CYP142A1 with a molecular

weight of 94412 ± 48 Da (Figure 3.34). The proportion of dimeric CYP142A1

increased considerably when the protein concentration was raised from 10 to 20

M, with the percentage of the dimer estimated to be ~33% that of the monomer

at 20 M CYP142A1.

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3.2.14.2 Interaction of CYP142A1 with DTT

Figure 3.35: NanoESI mass spectrum of 10 μM CYP142A1 with DTT (0-5 mM). 10 μM CYP142A1 was prepared in 200 mM ammonium acetate (pH 7.0) and mixed with 0 mM, 1 mM, and 5 mM DTT. Assigned CYP142A1 peaks for the three different concentrations of DTT are for monomer: (A) m/z 3200 to 4400; and for dimer: (B) m/z 4600 to 6000. The presence of CYP142A1 dimer was observed in all samples, but decreased considerably with the introduction of DTT. From the light scattering (MALLS) data (Figure 3.20A), there was evidence of

dimerization of CYP142A1. However, the dimer content was diminished by treating

with DTT. To investigate this further, nanoESI was used to study the influence of

DTT (at 1 mM and 5 mM) on CYP142A1 dimerization. From the results generated

(Figure 3.35), the dimerization of CYP142A1 decreased significantly on addition of

DTT, but there was little difference between the effects produced by 1 mM DTT and

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5 mM DTT. However, the reduction in the percentage of CYP142A1 dimer present

was from approximately 13% dimer in absence of DTT to 7% dimer at 5 mM DTT.

3.2.14.3 Interaction of CYP142A1 with Econazole

The interactions of CYP142A1 with two different concentrations (10 and 50 µM) of

a known azole inhibitor (econazole) were also investigated. CYP142A1 was directly

mixed with econazole in the form of stock solutions in 100% DMSO. The influence

of DMSO (2.5%) on the quality of the spectra was also analysed (Figure 3.36). The

dimerization of CYP142A1 was almost negligible when complexed with econazole,

but a significant proportion of dimer was evident in the presence of DMSO, possibly

indicating that changes in structural conformation which favour the dissociation of

the dimer occur in the presence of econazole. In addition, it is worth noting that the

presence of DMSO alone did not have any obvious impact on the measured masses.

Econazole complexes with CYP142A1 by ligating to the P450 heme iron via an

imidazole nitrogen atom on the molecule. The nanoESI MS data revealed that

econazole bound to CYP142A1 in a proportion of the molecules. There was also

some influence on the aggregation state of the enzyme, as the proportion of

CYP142A1 dimer was diminished to some extent in the samples to which econazole

was added (Figures 3.36). The molecular weight of econazole is ~382 Da and hence

a deduction of this value from 47436 ± 2 Da (for example, in the case of the 50 M

econazole complex) gives a figure approximately that of the molecular weight of

the ligand-free CYP142A1 (~47054 Da). This finding is consistent between the data

sets obtained using two different concentrations of econazole. The recorded

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spectra for the monomeric CYP142A1 ligand complex revealed a splitting of the

peaks associated with both unbound and econazole-bound monomeric forms.

Signals associated with CYP142A1 dimer are almost absent for the econazole-bound

CYP142A1 (with more dimer seen in the DMSO-only sample). This observation is

consistent with the results obtained for CYP121A1 in recent studies, where stronger

binding azoles such as clotrimazole or miconazole cause dissociation of the dimers

present (Duffell et al., 2013).

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Figure 3.36: NanoESI mass spectra of 10 μM CYP142A1 with econazole. 10 μM CYP142A1 was mixed with 2.5% DMSO, and either 10 μM or 50 μM econazole in 200 mM ammonium acetate (pH 7). Peaks in the range m/z = 42700 to 5400 were assigned to monomer. Red dots indicate the unbound monomers and green dots represent econazole-bound monomers. Dimeric features are almost absent for the econazole-bound CYP142A1. At 50 µM econazole, the ligand-free monomer has a Mw = 47046 ± 1 Da and the ligand-bound monomer a Mw = 47436 ± 2 Da. The difference in mass (390 Da) is consistent with that of econazole (382 Da), within the error range of the experiment. At 10 µM econazole, the ligand-free monomer has a Mw = 47058 ± 38 Da and the ligand-bound monomer a Mw = 47489 ± 24 Da. In the presence of DMSO only (at 2.5% v/v), the ligand-free monomer has a Mw = 47058 ± 16 Da, similar to that for ligand-free CYP142A1.

3.2.14.4 Analysis of the Interaction of CYP142A1 with Cholestenone.

Cholestenone and cholesterol are substrates for CYP142A1 and Mtb utilises these

substrates as a major source of energy during chronic and latent infection (Ouellet

et al., 2011). CYP142A1 (alongside CYP125A1 and CYP124A1) catalyses the C27

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hydroxylation of the cholestenone/cholesterol side chain via a three step reaction,

first to an alcohol, then to an aldehyde and finally to the carboxylic acid moiety

(Johnston et al., 2010). Binding assays for CYP142A1 with cholestenone reveal a

heme iron high-spin shift associated with a Soret band shift to lower wavelength (a

‘blue shift’). This shift in the heme iron spin-state is accompanied by the substrate-

dependent displacement of the axial water ligand on the heme iron.

In this experiment 0-125 μM cholestenone was added to a 10 μM solution of

CYP142A1. Data revealed mainly monomeric species (ligand-free and cholestenone-

bound) with a proportion of dimeric peaks. Peaks in the range from m/z 3400 to

4400 correspond to the monomer. The molecular weight of cholestenone is ~385

Da and hence a deduction of this value from 47424 ± 40 Da (for the 125 M

cholestenone-bound monomer form), for example, gives a figure approximating to

the molecular weight of the ligand-free CYP142A1 (~47036 ± 3 Da). This finding is

consistent for the data sets obtained at three different concentrations of

cholestenone. The spectra for CYP142A1 at each cholestenone concentration

revealed a splitting of the peaks into unbound and cholestenone-bound monomer

forms. Weak peaks representing dimeric species are evident with each of the

cholestenone-bound CYP142A1 forms, and with the cholestenone-free sample that

contains only ethanol (used as solvent for cholestenone). The results obtained here

for CYP142A1 points to important differences with recent studies carried out using

CYP121A1, and where both monomeric and dimeric CYP121A1 azole ligand-bound

peaks were observed (Duffell et al., 2013).

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Figure 3.37: NanoESI mass spectra of 10 μM CYP142A1 with cholestenone. 10 μM CYP142A1 was mixed with 0-125 μM cholestenone (dissolved in 2.5% ethanol) and in 200 mM ammonium acetate (pH 7). Peaks in the range m/z 3400−4400 were assigned to monomer, where red dots indicate the unbound monomers and green dots represent cholestenone-bound monomers. At 125 µM cholestenone, the ligand-free monomer has a Mw = 47036 ± 3 Da and the ligand-bound monomer a Mw = 47424 ± 40 Da. At 50 µM cholestenone, the ligand-free monomer has a Mw = 47004 ± 50 Da and the ligand-bound monomer a Mw = 47438 ± 50 Da. At 10 µM cholestenone, the ligand-free monomer has a Mw = 47060 ± 37 Da and the ligand-bound monomer a Mw = 47476 ± 8 Da. In the CYP142A1 sample containing ethanol (2.5%), the CYP142A1 monomer has a Mw = 47058 ± 16 Da.

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3.2.14.5 Interaction of CYP142A1 with Solvents.

Figure 3.38: NanoESI mass spectrum of 10 μM CYP142A1 with solvents. 10 μM CYP142A1 was mixed with 200 mM ammonium acetate (pH 7.0), and with buffer plus 1% ethanol and 2.5% DMSO. Assigned peaks for DMSO: monomer (A) m/z 4200 to 5400; dimer (B) m/z 6200 to 7400; for ethanol: monomer (A) m/z 3400 to 4400; dimer (B) m/z 4400 to 5400; and for buffer alone: monomer (A) m/z 3400 to 4400; dimer (B) m/z 4800 to 5800. The presence of dimer was observed for all solvents but was minimal with DMSO as compared with buffer and ethanol. In order to evaluate the effects of selected solvents (DMSO and ethanol, used to

solubilise various ligands during the course of experiments done in this thesis),

nanoESI experiments were carried out to investigate whether interactions with the

solvents alone influenced the propensity of CYP142A1 to undergo changes in

aggregation state. It was found that CYP142A1 dimers formed in the presence of

both DMSO (2.5% v/v) and ethanol (1% v/v), but that the largest proportion of

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dimer was seen in ethanol, while lower (and quite similar) amounts of dimer were

present in the CYP142A1 samples containing buffer alone (200 mM ammonium

acetate, pH 7), or buffer containing DMSO. Thus, addition of ethanol appears to

have an effect in promoting (to a small extent) the formation of CYP142A1 dimers.

3.3 Summary

CYP142A1, a cholesterol 27-oxidase enzyme that likely plays a compensatory role

with CYP125A1 in host cholesterol/cholestenone oxidation, was expressed using an

E. coli expression system and purified to homogeneity via three chromatographic

steps. Results obtained from the expression and purification trials showed that

CYP142A1 was expressed best in 2YT medium after 24 hours of culture in C41 (DE3)

transformant cells. Purification of CYP142A1 was quite efficient, as most of the

contaminants were eliminated in the initial purification step (using a Ni-NTA

column), while remaining protein contaminants were successfully removed on

passing the material collected from the affinity purification stage through first a

hydroxyapatite column, and then by using size exclusion (gel filtration)

chromatography. The effectiveness of the purification regime was confirmed based

on the analyses of eluates from these columns using SDS-PAGE and UV-visible

spectrophotometric analyses, with CYP142A1 appearing as single bands on SDS-

PAGE gels, and with an A418/A280 (heme/protein or Rz) ratio ≥ 1.9.

The extinction coefficient for the low-spin ferric heme in the resting state of

CYP142A1 was determined using the pyridine hemochromogen method (Berry and

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Trumpower, 1987). This experiment resulted in the determination of an absorbance

coefficient of ε418 = 92 mM−1 cm−1 at the Soret peak for the oxidised enzyme.

Carbon monoxide binding to CYP142A1 revealed features typical of a heme-

containing P450 protein. A CYP142A1 CO-adduct was formed with a Soret

maximum at ~450 nm, as expected for a P450 Fe(II)-CO complex with retention of

cysteine thiolate proximal ligation to the heme iron (Omura and Sato, 1964). The

CYP142A1 ferrous-CO complex was stable for several minutes in both the absence

and the presence of the substrate cholestenone, and did not convert to a P420

complex (which absorbs at ~420 nm and arises from protonation of the cysteine

thiolate to a thiol form). Thus, CYP142A1 forms a stable ferrous-CO adduct that

does not require substrate for the retention of its native (thiolate-ligated) state.

The Mtb genome sequence revealed a large number (20) of cytochrome P450

enzymes (Cole et al., 1998), some of which participate in cholesterol metabolism

(Ouellet et al., 2011). CYP125A1, CYP142A1 and CYP124A1 can all initiate oxidative

degradation of the cholesterol side chain, a critical first step in its breakdown that

ultimately enables energy generation via the β-oxidation pathway (Ouellet et al.,

2011). Optical titrations with cholestenone, cholesterol and lanosterol show that

CYP142A1 binds tightly to each of the first two sterols (and more weakly to

lanosterol) consistent with a physiological role for CYP142A1 in

cholesterol/cholestenone metabolism. CYP142A1 also binds tightly to a range of

azole antifungal drugs, and some of these azoles have been shown to clear Mtb

infection in mice, with econazole being the most effective among drugs tested

(Ahmad et al., 2006c). Many of these azoles were also shown to bind tightly to

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other Mtb P450 enzymes, and to be effective in preventing the growth of Mtb and

other mycobacterial cells in vitro (McLean et al., 2002b, McLean et al., 2008). The

order of potency of the azole drugs against Mtb H37Rv (econazole and miconazole

at 8 g ml-1, clotrimazole at 11 g ml-1 and ketoconazole at 16 g ml-1) (section

1.5.9) correlates quite well with their Kd values for binding to the Mtb CYP142A1

enzyme (2.28 M, 1.42 M, 1.14 M and 11.95 M, respectively) (Table 3.2),

conforming that the CYP142A1 has high affinity for the most effective anti-Mtb

azoles, and lower affinity for the one (ketoconazole) with the weakest MIC. It also

has a weak Kd value for fluconazole (Kd = 309 M); a drug known to be ineffective

against Mtb. These data are consistent with CYP142A1 being an important azole

drug target in Mtb, presumably through the crucial role that it plays in host

cholesterol/cholestenone metabolism.

Preliminary fragment based screening with CYP142A1 conducted at the University

of Cambridge generated 6 hits from a fragment chemical library (named NMR089,

099, 170, 491, 540 and 623), and the interactions of these molecules with the P450

were studied further using biochemical and biophysical techniques. Validation of

CYP142A1 ligand binding to these compounds was done using UV-visible

spectroscopy, electron paramagnetic resonance (EPR) spectroscopy, isothermal

titration calorimetry (ITC) and crystallography (further results are given in chapter

4). UV-visible spectroscopy revealed a substrate-like type I optical shift for fragment

NMR099, and inhibitor-like type II shifts for the five others – indicating direct

interaction with heme iron. Dissociation constants (Kd values) for the fragments

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ranged from NMR491 (0.68 µM) to NMR099 (7.2 mM). ITC confirmed compound

binding in the same rank order as seen from the optical binding studies.

EPR was done to probe CYP142A1 heme coordination and

substrate/inhibitor/fragment binding. X-band EPR data at 10 K for ligand-free and

fragment-bound forms of CYP142A1 revealed characteristic rhombic signals for

ferric P450s. Heterogeneous low-spin heme iron signals were obtained on binding

azole inhibitors/fragments, arising from either direct ligation of a heterocyclic

nitrogen in the fragment/ligand to the heme iron, or through their making indirect

interactions with heme iron via a (6th) water ligand that remains on the iron

following ligand addition. A mixture of low-spin and high-spin features was

obtained for cholestenone binding, indicating a displacement of the water ligand in

the sixth axial position. EPR provides important confirmatory information for

binding of these molecules to CYP142A1. Data from differential scanning

calorimetry (DSC) also revealed that the binding of cholestenone increased the

CYP142A1 Tm value by approximately 5 oC, while clotrimazole increased the Tm

value by approximately 3 oC. This is indicative of the stabilization of the P450 when

complexed with these ligands.

Redox potentiometry studies were done for CYP142A1 in cholestenone-bound,

clotrimazole-bound and ligand-free forms. Many P450s have low-spin ferric heme

iron in their resting state, but convert to high-spin ferric heme iron on binding

substrates, with loss of the 6th (water) ligand to the heme iron causing heme iron

(3d orbital) electronic reorganization (Ouellet et al., 2010b). This causes a large shift

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in heme potential, enabling electron transfer to heme iron from NAD(P)H-

dependent redox partners (Daff et al., 1997) .

This is an important regulatory process, allowing heme iron reduction to occur

efficiently only when substrate is bound, avoiding production of damaging oxygen

radicals. For substrate-free CYP142A1, reduction of heme iron induces a change in

the absorbance maximum of the ferric heme Soret band from 418 nm to ~405 nm

(with a midpoint potential of -394 ± 4 mV vs NHE). For the cholestenone substrate-

bound form, the Soret shift on heme reduction is from 393 nm to ~405 nm (with a

midpoint potential of -150 ± 4 mV vs NHE). For the clotrimazole-bound CYP142A1,

the Soret band shifted from 424 nm to ~428 nm with a midpoint potential of -360 ±

4 mV. In these experiments, anaerobic reductive titration of CYP142A1 forms was

undertaken, and heme absorbance changes at wavelengths reflecting large changes

between the oxidised and reduced forms of CYP142A1 were plotted versus applied

potential, and data were fitted using the Nernst function in order to determine

midpoint potentials for the CYP142A1 heme Fe3+/Fe2+ couples in the substrate-free

and cholestenone-bound forms of CYP142A1 as -394 mV and -150 mV (vs NHE),

respectively. The large potential change (~244 mV) between the low-spin (SF) and

high-spin (SB) forms reflects at least in part a major change in heme iron spin-state

on cholestenone binding and is indicative of a tightly regulated redox system. The

particularly large change in potential may also indicate a further level of regulation

– e.g. through structural changes in the heme environment that occur on substrate

binding. Re-oxidation of the reduced P450 samples demonstrated reversibility and

also that CYP142A1 retains its cysteine thiolate ligand to the heme iron on return to

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the ferric state. Light scattering (MALLS) data revealed CYP142A1 to have a

substantial component of dimeric P450 in the absence of the reducing agent DTT.

On addition of DTT, the P450 became almost completely monomeric. This

treatment enabled CYP142A1 to be crystallized in the ligand-free and ligand-bound

forms (see results in chapter 5). The oligomerization of CYP142A1 was further

investigated with nanoESI mass spectrometry. With this technique, the proportion

of CYP142A1 dimer was found to increase as the concentration of CYP142A1 was

elevated from 10 µM to 20 µM. Under these conditions, treatment with different

DTT concentrations diminished the proportion of dimer present, but did not cause a

complete conversion to the monomeric state. In contrast, the CYP142A1 complex

with econazole showed complete conversion to a monomeric state. This was not

the case for cholestenone, and a proportion of the dimeric form was retained in the

presence of the substrate. It appears likely that the binding of econazole favours

dissociation of the CYP142A1 dimer.

Protein stability studies were also done on substrate-free and cholestenone-bound

CYP142A1 using guanidinium chloride (GdmCl) as a denaturant. Protein

denaturation by GdmCl was monitored using tryptophan fluorescence.

Fluorescence (with excitation at ~280 nm) provides emission spectra that report on

the protein tertiary structure, with structural perturbations evident from changes in

the fluorescence intensity from aromatic amino acids (mainly tryptophans) in the

P450, and from changes in the emission maximum in the range from ~340 nm

upwards. CYP142A1 was progressively denatured with increasing [GdmCl].

Cholestenone binding had little effect in stabilizing CYP142A1 to denaturation by

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GdmCl, perhaps indicating that dissociation of the substrate occurs at a low

concentration of GdmCl. However, the stabilising effects of the substrate are

evident from DSC studies, where an ~5 oC increase in Tm with cholestenone was

observed.

Collectively, results in this chapter provide a substantial body of work describing

fundamental biochemical, spectroscopic and thermodynamic properties of Mtb

CYP142A1. In results presented in Chapter 4, the catalytic and biophysical

properties of CYP124A1, the third member of the cholesterol oxidase family, will be

discussed.

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Chapter 4

Biochemical and Biophysical characterization of CYP124A1: A

promiscuous enzyme with broad substrate specificity in

Mycobacterium tuberculosis

4.1 Introduction

Similarly to CYP142A1, CYP124A1 can also oxidize the aliphatic side chain of

cholesterol/cholestenone to the carboxylic acid state by sequential metabolism to

the alcohol, then to the aldehyde, and ultimately to the acid (Johnston et al., 2010).

However, in addition to this function, CYP124A1 has also been shown to possess

broad substrate specificity, including activity towards branched chain lipids

(Johnston et al., 2009). To date there is no evidence for the essentiality of the

CYP124A1 gene in Mtb, and a Mtb H37Rv CYP124A1 transposon mutant can grow in

vitro. However, given the relevance of this P450 to sterol and other lipid oxidation

reactions in Mtb, it is suspected that that CYP124A1 may be important for growth

of the bacterium in the host macrophage.

Mycobacterium tuberculosis, the human pathogen that causes tuberculosis,

generates sulfated metabolites associated with virulence. However, these

metabolites have remained poorly characterized for several years (Mougous et al.,

2006). One of these metabolites is a methyl-branched lipid known as S881 which

was shown to be associated with the cell wall of Mtb and also involved in the

negative modulation of virulence in Mtb infected mouse models (Johnston et al.,

2009, Holsclaw et al., 2008). Studies have also highlighted that such sulfated

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metabolites function as signalling molecules between pathogenic bacteria and their

hosts (Holsclaw et al., 2008).

CYP124A1 (Rv2266) is located in the same gene region as CYP128A1 and CYP121A1.

One of the genes in this cluster (Sft3, Rv2267c) encodes a sulfotransferase enzyme

(Johnston et al., 2009, McLean et al., 2010). This sulfotransferase was shown to

catalyze the 3’-phosphoadenosine-5’-phosphosulfate (PAPS)-dependent sulfation of

menaquinone MK-9 DH-2 (a compound with repeated methyl branching) at the -

position (Holsclaw et al., 2008, Mougous et al., 2006). Sulfotransferases are also

known to mediate the transfer of a sulfuryl group from PAPS to a carbohydrate, or

to a tyrosine residue within a protein (Mougous et al., 2002). CYP128A1 (the

product of Rv2268c) is thought to hydroxylate the -position of menaquinone MK-9

DH-2 before its Sft3-dependent sulfation to the S881product, which is involved in

virulence in Mtb (Holsclaw et al., 2008, Mougous et al., 2006, Johnston et al., 2009).

Menaquinone MK-9 DH-2 is the major quinol electron carrier of Mtb and thus its

sulfation may be a process by which its respiratory function is regulated (Holsclaw

et al., 2008).

It is postulated that the utilization of host cholesterol by CYP125A1 (and CYP142A1)

enables Mtb persistence and survival in the cholesterol-rich macrophage (Ouellet et

al., 2010a, Johnston et al., 2010, Pandey and Sassetti, 2008). Hence a drug that can

inhibit the cholesterol catabolic pathway could be an important candidate for

eliminating non-replicating, latent Mtb. CYP124A1, in addition to being a methyl-

branched lipid -hydroxylase, was also shown by Johnston et al. to oxidize

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cholesterol at the C27 position (Johnston et al., 2009, Johnston et al., 2010). Studies

have shown that CYP124A1 and CYP142A1 possess the same sterol 27-oxidase

activity and can complement the growth defect of a ∆CYP125 Mtb strain on

cholesterol. Thus, these duo may present additional promising secondary drug

targets against latent Mtb (Johnston et al., 2010). CYP124 genes are conserved

across a wide spectrum of organisms ranging from pathogenic to non-pathogenic

mycobacterial species, actinobacteria and some proteobacteria, suggesting multiple

and important physiological and catalytic roles for this enzyme (Ouellet et al., 2008,

Johnston et al., 2009).

In this chapter, results from the biochemical and biophysical characterization of

CYP124A1 are presented, with the aim of providing further insights into the

substrate specificity of this enzyme and defining which substrates have greatest

affinity for the P450. Binding assays for CYP124A1 were carried out using a wide

range of substrates include cholesterol, cholestenone, fatty acids and other long-

alkyl-chain lipids. These studies were done to validate its substrate selectivity and

to characterize its hydroxylase and oxidase activities with diverse substrates.

Studies were also done to analyse the interactions of CYP124A1 with selected azole

inhibitors and other compounds emanating from fragment based screening studies;

the latter providing the basis for a future fragment merging/linking programme by

which CYP124A1-specific inhibitors could be generated. Results from these

experiments are aimed at providing further insights into the druggability of this

enzyme and its potential as a novel drug target for existing azole drugs and for

novel types of inhibitors. Further aims of the research in this chapter include the

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spectroscopic analysis of CYP124A1 to provide further understanding of the modes

of binding of substrates and inhibitors with the P450, and the determination of the

influence of substrate binding on the heme iron potential in CYP124A1 to establish

how effective substrate binding is in elevating the heme iron potential to enable

reduction of CYP124A1 and progression of the catalytic cycle.

4.2 Results and Discussion

4.2.1 Expression and Purification of CYP124A1

The CYP124A1 (Rv2266) gene was codon-optimised and synthesised by Genscript

(Piscataway, USA), and cloned into the expression vector pET47b using BamHI and

HindIII restriction sites. The pET47b vector contains an N-terminal His6-tag, which

allows recombinant proteins to be purified by nickel affinity column

chromatography. Gene expression is under the control of a T7lac promoter. The

pET47b vector carries the gene for kanamycin resistance, which allows for antibiotic

selection of cells carrying the plasmid.

Expression trials for the CYP124A1/pET47b construct was done using various

different conditions, as detailed in the Materials and Methods (section 2.2.3) and

these included different E. coli strains, growth media, IPTG induction conditions,

inclusion or absence of ∆ALA post-induction of CYP124A1 expression, and various

growth temperature conditions. These conditions were optimized and the best

condition for CYP124A1 expression was chosen. Hence, CYP124A1 was best

expressed using E. coli C41 (DE3) cells grown in 2YT medium supplemented with

kanamcyin (30 μg/ml). The other E. coli strains that were tested were the DE3

244

lysogens BL21 (DE3), Rosetta 2 (DE3), C41 (DE3) and HMS174 (DE3). The DE3

lysogen strains encode the T7 RNA polymerase gene needed for recombinant

protein expression from the T7 promoter on the plasmid. This gene is under the

control of the lacUV5 promoter, and expression is induced by the addition of IPTG.

The construct CYP124A1/pET47b was transformed into these three E. coli DE3

strains and CYP124A1 expression trials were conducted. Addition of IPTG

(isopropyl-β,D-thiogalactopyranoside) regulates the lacUV5 promoter which

controls expression of the T7 polymerase gene, specifically by binding to the

inhibitory Lac repressor protein and displacing it from its DNA target, enabling the

RNA polymerase to bind. IPTG enables the expression of the recombinant protein

by acting as a stable analogue of lactose to bind and displace the Lac repressor from

its operator binding site (Studier and Moffatt, 1986, Wang et al., 1989).

Following growth of CYP124A1-producing transformant cells, the cells were

collected and disrupted, and the P450 was purified to homogeneity via three

chromatographic steps using the same protocol as for the CYP142A1 purification, as

detailed in the Materials and Methods (section 2.2.6). The first step was affinity

chromatography using a Ni-NTA (nickel) column, followed by a further affinity step

using a hydroxyapatite (HA) column, and then a final polishing step using size

exclusion chromatography.

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A) Nickel Column Purification

A DNA sequence specifying a string of six to nine histidine residues is frequently

used in plasmid expression vectors to enable purification of recombinant proteins.

This results in the expression of recombinant proteins with a 6xHis or other poly-

His tag fused to the protein’s N- or C-terminus (Smith et al., 1988). Immobilized

Metal-Affinity Chromatography (IMAC) (e.g. nickel column chromatography) is an

effective separation technique that is utilized in the purification of proteins both

with natural surface-exposed histidine residues and for recombinant proteins with

engineered histidine tags or other histidine clusters (Gaberc-Porekar and Menart,

2001).

This purification technique uses covalently bound chelating compounds or metal

ions immobilized on solid chromatographic media to serve as affinity ligands for

target proteins, making use of coordinative binding to relevant amino acid residues

exposed on the protein surface (Gaberc-Porekar and Menart, 2001). Recombinant

proteins with engineered histidine-tags can usually be purified easily because the

string of histidine residues binds to several types of immobilized metal ions,

including nickel, cobalt and copper, under specific buffer conditions. Subsequently,

the fraction or protein of interest is eluted by e.g. varying the pH of the washing

buffer, or by addition of high concentrations of imidazole (Block et al., 2009).

CYP124A1 purification with nickel column chromatography was successful, with

most of the contaminant proteins being removed with the flow-through. Purified

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CYP124A1 samples were eluted with 40 mM imidazole as described above, and

were pooled together and subjected to the next purification step using

hydroxyapatite column chromatography.

Figure 4.1: Nickel affinity chromatography purification of CYP124A1. The image shows a 12% (w/v) SDS-PAGE gel showing (from left to right): Protein marker (PM) - NEB protein ladder with bands labelled in kDa (10-250 kDa), Flow-through from the column (FT), and Buffer wash (BF). Lanes 4-9 show CYP124A1 eluted from the column with increasing concentrations of imidazole in the wash buffer.

B) Hydroxyapatite (HA) Column Purification

Hydroxyapatite chromatography is also referred to as a “pseudo-affinity

chromatography’’ or "mixed-mode" ion exchange, since the strength and selectivity

of the interaction depends on the structural features of both the chromatographic

material and the retained substance (Hilbrig and Freitag, 2012). Hydroxyapatite

(HA), a calcium and phosphate based inorganic material, has the stoichiometric

formula of Ca10(PO4)6(OH)2 (Hilbrig and Freitag, 2012).

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The mechanism of HA chromatography is complex, because it involves non-specific

interactions between positively charged calcium ions and negatively charged

phosphate ions (on the stationary phase HA resin) with negatively charged carboxyl

groups and positively charged amino groups in the protein. A buffer with increasing

phosphate concentration is typically used for elution of the protein of interest.

Hence, while acidic proteins bind through C (calcium)-sites, basic proteins bind

through P (phosphate)-sites on the hydroxyapatite (Cummings et al., 2009). With

HA chromatography, most weakly bound proteins are purified with low ionic

strength buffers with low phosphate concentrations as low as 1 mM. More strongly

adsorbed or HA-bound proteins are often eluted with higher concentrations of

phosphate, or even using NaCl or KCl salts (Cummings et al., 2009).

CYP124A1 was purified using HA column chromatography immediately after the

nickel column chromatography, and this resulted in the elimination of most of the

contaminating proteins left after the nickel affinity step. Samples isolated from

nickel chromatography were exchanged into 15 mM KPi, pH 7.0 (Buffer A) and

loaded onto a HA column pre-equilibrated in the same buffer. Protein was eluted

using a gradient of 15 mM to 500 mM KPi (pH 7.0) buffer. Purer fractions of eluates

from the HA column (as evidenced from SDS PAGE analysis) were then pooled

together and subjected to a final polishing step using size exclusion

chromatography.

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Figure 4.2: Hydroxyapatite (HA) column chromatography purification of CYP124A1. A 12% (w/v) SDS-PAGE gel is shown, with lanes (from left to right) containing: Protein marker (lane 1; NEB protein ladder bands labelled in kDa (10-250 kDa)), Lanes 2-5 show CYP124A1 eluted from the column with increasing potassium phosphate buffer strength. CYP124A1 bands shown are close to the predicted molecular weight of ∼50.52 kDa (for CYP124A1 plus the His-tag) when compared to the molecular weight markers. Other proteins eluted along with CYP124A1 are seen as faint contaminant bands on SDS-PAGE.

C) Size exclusion or Gel filtration column chromatography

Size exclusion or gel filtration chromatography is a technique used for the

separation of molecules, e.g. proteins and peptides, based on their size (Duong-Ly

and Gabelli, 2014). Furthermore, gel filtration chromatography can also be used to

resolve oligomeric forms of proteins and to exchange the buffer of a sample for a

different one, often referred to as a desalting column in the latter case (Duong-Ly

and Gabelli, 2014).

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The matrix of the gel filtration column consists of porous beads, and the size of the

bead pores defines the size of macromolecules that may be separated (Bollag,

1994). Molecules that that are too large to enter the bead pores are “excluded,”

and thus separate out from the column first, while small molecules, which can enter

the sieves in the matrix of the stationary phase, elute much later. This is because

large molecules that do not enter the bead pores have a smaller volume to pass

through, and they are the first molecules to elute from the column (Bollag, 1994,

Duong-Ly and Gabelli, 2014).

Immediately after the HA column purification, CYP124A1 was subjected to a final

polishing step using gel filtration chromatography. In this case, highly pure protein

was resolved, as shown on the SDS-PAGE gel depicted below (Figure 4.3).

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Figure 4.3: Purification of Mtb CYP124A1 using a SuperdexTM S-200 gel filtration column. SDS-PAGE analysis shows molecular weight markers (lane 1, NEB protein ladder with bands labelled in kDa (10-250 kDa) and pure CYP124A1 as a single band in lanes 2-3) and close to the predicted molecular weight of ∼50.52 kDa (CYP124A1 plus His-Tag) when compared to the molecular weight marker.

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4.2.2 Spectroscopic analysis of CYP124A1

4.2.2.1 The UV-visible spectrum of CYP124A1

Figure 4.4: The UV-visible spectrum for purified, ferric CYP124A1. CYP124A1 (5.3

M) has a low-spin Soret peak at 418 nm, and a high-spin shoulder at ~395 nm

(arrowed). The alpha and beta bands are at ~570 and ~538 nm, with a high-spin

charge transfer band (cysteine thiolate-to-high-spin ferric heme iron) at ~650 nm.

Studies carried out recently by Johnston et al. revealed that the purified CYP124A1

exhibited typical spectral features of a ferric P450, but with about 70% of the heme

iron in a low-spin state and with a Soret peak at 418 nm and a proportion of high-

spin heme, as evidenced by a shoulder at 395 nm (Johnston et al., 2010). This is

consistent with the findings in this study. CYP124A1 was found to be mixed-spin,

with a low-spin Soret peak at 418 nm and a weaker high-spin shoulder at ~395 nm,

as depicted with black arrows in Figure 4.4. The mixed-spin heme iron feature was

observed to be perturbed in favour of the high-spin species if CYP124A1 was

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continuously frozen and thawed. It is speculated that this phenomenon may be due

to structural perturbations induced and their effects on the high-spin/low-spin

equilibrium that is likely mediated by the displacement and replacement of the 6th

water ligand to the ferric heme iron. In recent studies, Mtb CYP125A1 was reported

to be extensively high-spin in its native, resting state (McLean et al., 2009). Based

on evolutionary relationships, there is a quite close similarity between CYP124A1

and CYP125A1 (40.7% identity over 428 residues) (Driscoll et al., 2010) and

structural similarities in the heme environment could account for the similarity in

their spin-state equilibrium. However, this is in contrast to the spectral features

displayed by CYP142A1, in which the ferric heme is extensively low-spin (Johnston

et al., 2010).

4.2.2.2 CYP124A1 optical titrations with substrates

On the basis of the genetic location of CYP124A1 in an operon containing a

sulfotransferase (Stf3) involved in the sulfation of the respiratory menaquinone MK-

9, it was speculated that CYP124A1 could be involved in the metabolism of

substrates with a similar structure (Johnston et al., 2009). Johnston et al. also

demonstrated that CYP124A1 is among the three key P450 enzymes involved in

sequential oxidation reactions of cholesterol side chain at the C27 position

(Johnston et al., 2010).

In this study, CYP124A1 was tested for binding with cholesterol, cholestenone and a

series of methyl-branched lipids. The results show that CYP124A1 binds tightly to

each of these substrates, giving further insights into the likely functionality of

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CYP124A1 in sterol metabolism, as well as in metabolism of other (including

methyl-branched) lipids. The high affinity of CYP124A1 for a wide range of

substrates could provide some explanation as to its relatively low catalytic

efficiency as a sterol 27-hydroxylase compared with CYP125A1 and CYP142A1

(Johnston et al., 2010, Hudson et al., 2012a). That is, it is feasible that CYP124A1

has evolved towards the oxidation of substrates distinct from sterols, but still

retains relatively low activity towards 27-oxidation of sterols. The results of

substrate binding assays with CYP124A1 are summarized (in the order of their

decreasing affinities) in Table 4.1 below.

S/N Substrate Kd (μM) Type of Shift/Fitting Function

1 Cholesterol 0.27 ± 0.01 I (Hill equation)

2 Cholestenone 0.50 ± 0.01 I (Hill equation)

3 Farnesol 0.89 ± 0.05 I (Hill equation)

4 Phytanic Acid 1.67 ± 0.39 I (Quadratic equation)

5 15-Methylpalmitic acid 1.71 ± 0.07 I (Hill equation)

6 Geraniol 2.85 ± 0.59 I (quadratic equation)

7 Geranylgeraniol 3.76 ± 0.05 I (Hill equation)

8 Phytane 844 ± 23 I (Hill equation)

9 Pristane 3179 ± 482 I (Hyperbolic function)

10 Menaquinone-4 - -

Table 4.1: Binding affinity of CYP124A1 with lipid substrates. The table shows Kd values and the types of spin-state shifts induced on binding to sterols and methyl-branched substrates. All molecules tested (other than menaquinone-4) induced substrate-like type I (high-spin) heme iron shifts. S/N indicates substrate number.

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Figure 4.5: Optical titration of CYP124A1 with cholest-4-en-3-one. Panel A shows

absolute spectra recorded during titration of CYP142A1 (3.4 μM) with cholest-4-en-

3-one. The Soret peak shifts from 418 to 393 nm as the high-spin ferric heme iron

form accumulates. The inset shows overlaid difference spectra from the optical

titration. Panel B shows the cholest-4-en-3-one-induced absorption change plotted

versus cholest-4-en-3-one concentration, with data fitted using the Hill equation to

give a Kd value of 0.50 ± 0.01 μM, n = 1.51 ± 0.02.

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Figure 4.6: Optical titration of CYP124A1 with cholesterol. Panel A shows UV-

visible absorption spectra from a titration of cholesterol with CYP142A1 (3.9 μM).

Reconversion from high-spin towards low-spin heme was not observed at higher

concentrations of cholesterol, but was observed with CYP142A1. The inset shows

difference spectra from the titration with peak and trough values at 387 nm and

422 nm. Panel B shows a fit of cholesterol-induced absorption change (ΔA387 minus

ΔA422, reflecting the peak and trough values in the difference spectra computed by

subtracting the spectrum for cholesterol-free CYP124A1 from each of the spectra

for the cholesterol-bound forms) versus [cholesterol] added, with data fitted using

equation 3 (the Hill equation) to generate an apparent Kd value of 0.27 ± 0.01 μM, n

= 1.54 ± 0.01.

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Figure 4.7: Optical titration of CYP124A1 with phytanic acid. Panel A shows UV visible absorption spectra from a titration of phytanic acid with CYP124A1 (~3.6 μM). The inset shows difference spectra from the titration with peak and trough values at 387 nm and 422 nm. Panel B shows a fit of phytanic acid-induced absorption change (ΔA387 minus ΔA422, reflecting the peak and trough values in the difference spectra computed by subtracting the spectrum for phytanic acid-free CYP124A1 from each of the spectra for the phytanic acid-bound forms) versus [phytanic acid] added, with data fitted using equation 1 (the quadratic equation) to generate an apparent Kd value of 1.67 ± 0.39 µM for phytanic acid binding to CYP124A1.

Figure 4.8: Optical titration of CYP124A1 with pristane. Panel A shows UV-visible absorption spectra from a titration of cholesterol with CYP124A1 (2.7 μM). The inset shows difference spectra from the titration with peak and trough values at 387 nm and 420 nm. Panel B shows a plot of pristane-induced absorption change (ΔA387 minus ΔA422, reflecting the peak and trough in the spectra) against [pristane], with data fitted using equation 2 (the hyperbolic function) to generate an apparent Kd value of 3.18 ± 0.48 mM for pristane binding to CYP124A1.

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Figure 4.9: Optical titration of CYP124A1 with geraniol. Panel A shows absolute spectra from a titration of geraniol with CYP124A1 (3.5 μM). The inset shows difference spectra from the titration with peak and trough values at 390 nm and 424 nm. Panel B shows a plot of geraniol-induced absorption change (ΔA390 minus ΔA424) against [geraniol] with data fitted using equation 1 (the quadratic equation) to generate a Kd value of 2.85 ± 0.59 µM for geraniol binding to CYP124A1.

Figure 4.10: Optical titration of CYP124A1 with geranylgeraniol. Panel A shows UV-visible absorption spectra from a titration of geranylgeraniol with CYP124A1 (5.6 μM). The inset shows difference spectra from the titration with peak and trough values at 387 nm and 421 nm. Panel B shows a plot of geranylgeraniol-induced absorption change (ΔA387 minus ΔA421, reflecting the peak and trough values in the difference spectra, computed by subtracting the spectrum for geranylgeraniol-free CYP124A1 from each of the spectra for the geranylgeraniol-bound forms) versus [geranylgeraniol] added, with data fitted using equation 3 (the Hill equation) to generate a Kd value of 3.76 ± 0.05 µM for geranylgeraniol binding to CYP124A1, n = 2.49 ± 0.08.

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Substrate binding to cytochrome P450 enzymes typically generates an absorbance

shift of the Soret peak from a higher wavelength to a lower wavelength (i.e. a shift

from ferric low-spin to high-spin). This is also referred to as a ‘blue-shift’ and is

associated with the displacement of the 6th ligand water molecule coordinated to

the heme iron (Denisov et al., 2005).

The binding of cholestenone to CYP124A1 resulted in a near-full conversion of the

heme iron to high-spin. However, the addition of cholesterol to CYP124A1 gave a

less extensive conversion to high-spin, although no reconversion or reversal of the

spectral shift towards low-spin was observed. The difference in extent of shift might

be associated with small differences in solubility of these compounds in aqueous

buffer. Both compounds are only sparingly soluble in water. Binding of cholesterol

to CYP142A1 and CYP125A1 was shown to give a partial conversion towards high-

spin (Ouellet et al., 2010a, Driscoll et al., 2010, McLean et al., 2009, Capyk et al.,

2009). An earlier study by Johnston et al. documented a partial reversal of the high-

spin spectral shift on binding CYP124A1 with cholesterol, and hence the affinity of

CYP124A1 for cholesterol could not be determined accurately (Johnston et al.,

2010). However, in this study, no reconversion back towards low-spin was observed

in the cholesterol titration with CYP124A1, and hence it was possible to determine

the affinity of CYP124A1 for cholesterol.

The Kd values of CYP124A1 for cholestenone, cholesterol and other lipids were

obtained from fitting UV-visible spectral titration data (Figures 4.5 - 4.10). The

binding curves were best fitted using sigmoidal or Michaelis-Menten/quadratic

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equations (see Materials and Methods, section 2.2.9.1) depending on the nature of

the dependence of the induced absorption change versus substrate concentration.

The apparent Kd values are listed in Table 4.1 in order of their affinities for

CYP124A1. From the results obtained, CYP124A1 displayed highest affinity for

cholesterol (Kd = 0.27 ± 0.01 µM) and cholestenone (Kd = 0.50 ± 0.01 µM), and the

weakest affinity for pristane (Kd = 3.18 ± 0.48 mM), while menaquinone-4 showed

no detectable binding. The data collected indicate a preference of CYP124A1 for

cholestenone and cholesterol over the methyl branched lipids. It is tempting at this

point to say that cholestenone and cholesterol could really be the natural

substrates for this enzyme, rather than the methyl branched lipids. In addition to

cholestenone, near-full conversion of CYP124A1 to the high-spin state was also

observed with some of the other lipids tested, irrespective of their weaker Kd

values. These lipids include phytanic acid, 15-methyl palmitic acid, farnesol,

geranylgeraniol and phytane, and these data suggest that these lipids bind close to

the heme iron and should be good substrates for CYP124A1, irrespective of their

weaker binding affinities compared to the sterol substrates. However, it should be

noted that all of the non-sterol substrates tested that did induce CYP124A1 high-

spin heme iron development (other than pristane [Kd = 3179 M] and phytane [Kd =

844 M]) have Kd values of <5 M.

4.2.2.3 CYP124A1 inhibitor binding assays

Like many characterized cytochrome P450 enzymes, CYP124A1 binds tightly to a

range of azole inhibitor drugs, some of which are known to bind to other Mtb P450s

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and to clear Mtb infection in mice (Ahmad et al., 2006c, Balding et al., 2008). These

azole compounds ligate the heme iron via their heterocyclic (usually imidazole or

triazole) nitrogen and this produces characteristic type II Soret spectral shifts (also

known as ‘red shifts’), usually leading to formation of a peak between 425 and 435

nm and a broad trough at about 390–410 nm in absorption difference spectra,

indicative of azole coordination to the heme iron through a nitrogen atom

(Jefcoate, 1978).

CYP124A1 was assayed with a range of azole inhibitors and the results are

summarised in Table 4.2. Unexpectedly, bifonazole bound as a Type I ligand and

showed the highest affinity towards CYP124A1 among the drugs tested (Kd = 0.19 ±

0.02 µM), with approximately 80% conversion to the high-spin state. Previous work

showed that its binding induced a near-complete high-spin conversion (Johnston et

al., 2009) and these findings are consistent with the results derived from this study.

The other azole drugs assayed exhibited type II shifts with CYP124A1, with

voriconazole showing the weakest affinity. Voriconazole also exhibits weak binding

affinity for CYP121A1 and CYP144A1 (Driscoll et al., 2011, McLean et al., 2002b).

Econazole and miconazole exhibited an odd binding spectrum that seems to show

that the heme initially goes high-spin, before converting to a new inhibitor-ligated

low-spin species. For posaconazole, no detectable heme perturbation was

observed, possibly due to this very large drug (molecular mass = 700.8 g/mol) being

unable to access the CYP124A1 active site. For fluconazole, no spectral shift was

again observed. Fluconazole is relatively water soluble compared to the other

clinically used azoles. The same is true for voriconazole — which also displayed

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weak binding to CYP124A1 (Kd = 959 ± 47 µM). The order of potency of the azole

drugs (section 1.5.9) correlated with their Kd values for binding to the Mtb

CYP124A1 enzyme (see Table 4.2), suggesting this P450 to be an important drug

target.

I/N Inhibitor Kd (μM) MIC value for Mtb H37Rv (µg ml-1) (Mclean et al., 2008)

Type of Shift

1 Bifonazole 0.19 ± 0.02 - I

2 Clotrimazole 4.76 ± 0.13 11.0 II

3 Econazole 18.6 ± 0.5 8.0 II

4 Miconazole 19.6 ± 0.3 8.0 II

5 Ketoconazole 70.9 ± 3.7 16.0 II

6 1-Phenylimidazole 335 ± 13 - II

7 Voriconazole 959 ± 47 - II

8 Fluconazole - - ND

9 Posaconazole - - ND

Table 4.2: Binding affinities for CYP124A1 with azole drug inhibitors. Kd values for CYP124A1 ligand binding and the resulting heme iron absorbance changes (type I or type II) are shown. ND = no spectral change detected. I/N indicates inhibitor number.

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Figure 4.11: Binding of bifonazole to CYP124A1. Panel A shows UV-visible absorption spectra from a titration of bifonazole with CYP124A1 (3.35 μM). The inset shows difference spectra from the titration with peak and trough values at 390 nm and 423 nm. Panel B shows a fit of bifonazole-induced absorption change (ΔA390 minus ΔA423, reflecting the peak and trough values in difference spectra computed by subtracting the spectrum for bifonazole-free CYP124A1 from each of the spectra for the bifonazole-bound forms) versus [bifonazole] added, with data fitted using equation 1 to generate an apparent Kd value of 0.19 ± 0.02 µM for bifonazole binding to CYP124A1.

Figure 4.12: Binding of clotrimazole to CYP124A1. Panel A shows UV-visible absorption spectra from a clotrimazole binding assay with CYP124A1 (2.7 μM). The inset shows difference spectra from the titration with peak and trough values at 431 nm and 392 nm. Panel B shows a fit of clotrimazole-induced absorption change (ΔA431 minus ΔA392, reflecting the peak and trough values in the difference spectra computed by subtracting the spectrum for clotrimazole-free CYP124A1 from each of the spectra for the clotrimazole-bound forms) versus [clotrimazole] added, with data fitted using equation 3 (the Hill equation) to generate an apparent Kd value of 4.76 ± 0.13 µM for clotrimazole binding to CYP124A1, n= 2.11 + 0.11.

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Figure 4.13: Binding of econazole to CYP124A1. Panel A shows UV visible absorption spectra from a titration of econazole with CYP124A1 (3.8 μM). The inset shows difference spectra from the titration with peak and trough values at 428 nm and 389 nm. Panel B shows a fit of cholesterol-induced absorption change (ΔA428 minus ΔA389, reflecting the peak and trough values in difference spectra computed by subtracting the spectrum for econazole-free CYP124A1 from each of the spectra for the econazole-bound forms) versus [econazole] added, with data fitted using equation 3 (the Hill equation) to generate an apparent Kd value of 18.56 ± 0.50 µM for econazole binding to CYP124A1, n = 3.53 + 0.31.

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Figure 4.14: Binding of miconazole to CYP124A1. Panel A shows UV-visible absorption spectra from a titration of miconazole with CYP124A1 (3.8 μM). Inset shows difference spectra from the titration with peak and trough values at 429 nm and 390 nm. Panel B shows a fit of miconazole-induced absorption change (ΔA429 minus ΔA390, reflecting the peak and trough values in the difference spectra computed by subtracting the spectrum for miconazole-free CYP124A1 from each of the spectra for the miconazole bound forms) versus [Miconazole] added, with data fitted using equation 3 (the Hill equation) to generate an apparent Kd value of 19.57 ± 0.30 µM for miconazole binding to CYP124A1, n = 3.70 + 0.19.

Azole-based compounds were shown to be potent inhibitors of fungal CYP51s and

of fungal growth. In addition, they also demonstrate inhibitory effects on the

growth of Mtb, M. smegmatis and Streptomyces strains (McLean et al., 2002b,

McLean et al., 2008). High affinity for a series of azole drugs has also been reported

for a number of characterized Mtb P450s, with econazole binding very tightly to

CYP121A1 (Kd = 24 ± 6 nM) and also being shown to clear Mtb infection in a mouse

model (Ahmad et al., 2005, Ahmad et al., 2006a).

Table 4.2 shows the Kd values for a range of azole drugs used clinically, as well as for

the smaller azole 1-phenylimidazole (1-PIM); while Figures 4.11 – 4.14 show the

spectral data for the binding of a range of azoles to CYP124A1. Spectral changes

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consistent with coordination of the CYP124A1 heme iron by the azoles (type II Soret

shifts to ∼422 nm) were observed for all drugs with the exception of bifonazole,

which instead showed a type I (substrate-like) shift and was also the tightest binder.

Other drugs (including clotrimazole with a Kd at 4.76 ± 0.13 μM, econazole at 18.56

± 0.50 μM and miconazole at 19.57 ± 0.30 μM) bound relatively tightly compared to

other azoles tested. However, the affinities for these azoles are not as high as those

for other Mtb cytochrome P450s studied, such as CYP121A1 (McLean et al., 2008);

CYP125A1 (McLean et al., 2009), CYP142A1 and CYP144A1 (Driscoll et al., 2011,

Driscoll et al., 2010).

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4.2.2.4 CYP124A1 fragment binding assays

A) Analysis of CYP124A1 interactions with CYP142A1-specific

fragment hits

Figure 4.15: Binding of NMR170 to CYP124A1. Panel A shows UV-visible absorption spectra from a binding titration of NMR170 with CYP124A1 (3.3 μM). The inset shows difference spectra from the titration with peak and trough values at 422 nm and 390 nm. Panel B shows a fit of NMR170-induced absorption change (ΔA422 minus ΔA390, reflecting the peak and trough values in difference spectra computed by subtracting the spectrum for NMR170-free CYP124A1 from each of the spectra for the NMR170 bound forms) versus [NMR170] added, with data fitted using equation 1 (the quadratic equation) to generate a Kd value of 19.57 ± 0.30 µM for NMR170 binding to CYP124A1.

CYP124A1 was also assayed against fragments generated from fragment-based

screening studies conducted at the University of Cambridge. Fragments are small,

weak-binding molecules with low molecular weight (typically <250 Da) (Murray and

Blundell, 2010). Fragment-based drug design is a relatively new technique that

involves developing small molecule ligands as chemical tools and leads for drug

development (Scott et al., 2012). Initial fragment screening at Cambridge generated

6 specific hits for CYP142A1 (Table 4.3) and 4 specific hits for CYP124A1 (Table 4.4).

Interestingly, the CYP142A1-specific hits were found to bind all the three

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cholesterol oxidases (CYP125A1, CYP124A1 and CYP142A1), indicating a good start

point for the development of specific inhibitors that are selective for each of these

three P450 isoforms and which could completely block host cholesterol utilization

by Mtb. Table 4.3 below compares the relative affinities of these fragments for the

three Mtb P450 isoforms.

Fragment CYP142A1 (Kd, µM)

Type of

shift

CYP124A1 (Kd, µM)

Type of shift

CYP125A1 (Kd, µM)

Type of shift

NMR491 0.68 ± 0.14 II 72.3 ± 1.9 II 4325 ± 285

II

NMR170 1.87 ± 0.07 II 43.3 ± 1.2

II 127 ± 5

II

NMR 540 176 ± 61 II 340 ± 220

II 3730 ± 380

II

NMR623 254 ± 47 II 2920 ± 250

II ND ND

NMR089 377 ± 10 II 3990 ± 160

II ND ND

NMR099 7215 ± 330 I 12100 ± 330

II ND ND

Table 4.3: Binding affinity for CYP142A1 fragment hits with Mtb cholesterol oxidase P450s. The data show Kd values determined from optical titrations, and the heme Soret absorption shifts induced in each case. All fragments (except NMR099 with CYP142A1) demonstrate type II binding, this being indicative of interactions between the fragments and the heme iron of the P450s. ND = no binding detected. These compounds showed highest affinity for CYP142A1 and weakest affinity for

CYP125A1. NMR491, NMR170 and NMR540 were observed to bind to all of these

cholesterol oxidase P450s. This is a desirable characteristic, since we aim to develop

highly potent and selective inhibitors which can completely block host cholesterol

utilization by Mtb.

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B) Studies with CYP124A1-specific fragment hits

Four fragments were generated specifically for CYP124A1. These are named:

NMR415, NMR115, NMR356 and NMR515 (Figure 4.16). The relevant spectral data

and binding titration analyses are detailed in Table 4.4. The tightest binder was

NMR415 and the weakest binding was seen with NMR356. No spectral shifts were

observed with NMR515. The three fragments that showed weak binding gave type

II spectral shifts, indicative of direct coordination to the heme iron by the small

chemical ligands.

Figure 4.16: Compounds identified as CYP124A1-specific fragment hits.

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Figure 4.17: Binding of NMR115 to CYP124A1. Panel A shows UV visible absorption spectra from a titration of NMR115 with CYP124A1 (~4.35 μM). The inset shows difference spectra from the titration with peak and trough values at 430 nm and 394 nm. Panel B shows a fit of NMR115-induced absorption change (ΔA430 minus ΔA394, reflecting the peak and trough values in the difference spectra, computed by subtracting the spectrum for NMR115-free CYP124A1 from each of the spectra for the NMR115-bound forms) versus [NMR115] added, with data fitted using equation 3 (the Hill equation) to generate a Kd value of 716.3 ± 14.5 µM for NMR115 binding to CYP124A1, n = 2.45 ± 0.11.

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Figure 4.18: Binding of NMR415 to CYP124A1. Panel A shows UV-visible absorption spectra from a titration of NMR415 with CYP124A1 (3.3 μM). The inset shows difference spectra from the titration with peak and trough values at 430 nm and 394 nm. Panel B shows a fit of NMR415-induced absorption change (ΔA430 minus ΔA394, reflecting the peak and trough values in difference spectra, computed by subtracting the spectrum for NMR415-free CYP124A1 from each of the spectra for the NMR415-bound forms) versus [NMR415] added, with data fitted using equation 3 (the Hill equation) to generate a Kd value of 282.0 ± 8.5 µM for NMR415 binding to CYP124A1, n= 1.76 + 0.10.

Fragment Kd (µM) Spectral shift

NMR415 282.0 ± 8.5 II

NMR115 716.3 ± 14.5 II

NMR356 4440 ± 2240 II

NMR515 ND No spectral shift

Table 4.4: Binding affinity of CYP124A1 for fragment hits. The Kd values were determined by optical titration, as described in the Materials and Methods (section 2.2.9.1). Inhibitor-like (type II) Soret shifts were observed for the binding of NMR415, 115 and 356. In the case of NMR356, only minor Soret changes were observed at high ligand concentrations, resulting in weak affinity for CYP124A1 and with a large error on the Kd value due to points collected being in the near-linear portion of the binding curve. No binding to CYP124A1 was detected (ND) in the case of fragment NMR515.

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C) Binding analysis with compounds from CYP121A1 fragment elaborated hits.

The first successful fragment-based approach to targeting Mtb cytochrome P450

enzymes was achieved with CYP121A1 (the gene for which is essential for Mtb

viability) (Hudson et al., 2013). Initial fragment screening for CYP121A generated 26

hits, giving a 46% validated hit rate (Hudson et al., 2012b). A parallel fragment

screen was also carried out against the Mtb CYP125A1 cholesterol/cholestenone

C27 monooxygenase, which produced a total of nine hits (Hudson et al., 2012b).

Surprisingly, only a single hit common to both CYPs was observed, suggesting a high

level of isoform selectivity, even at the fragment-screening level (Hudson et al.,

2012b).

The CYP121A1 specific hits generated were further validated and elaborated (via a

combination of synthetic chemistry and using data from structural biology studies)

to generate more potent compounds with higher affinity for CYP121A1 (Hudson et

al., 2012b). A fragment–fragment merging approach was used to generate these

compounds with higher affinity and selectivity, and this provided an excellent

scaffold for development of further type II CYP121A1 inhibitors. The findings from

this work lay the groundwork for the application of fragment-based drug design to

other members of the cytochrome P450 superfamily. CYP121A1 inhibitors

generated from this study were also assayed against the other Mtb P450s

CYP142A1, CYP125A1 and CYP124A1 in my PhD work, and interestingly they were

found to bind with differing affinities. These compounds are shown in Figure 4.19

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and Table 4.5 compares the relative affinities of these compounds for CYP142A1,

CYP125A1 and CYP124A1.

Figure 4.19: Elaborated compounds developed from CYP121A1 fragment hits. Compounds were synthesised by Madeline Kavanagh (University of Cambridge), based on the binding position of fragments determined from CYP121A1/fragment complex structures, and through a synthetic chemistry approach to “elaborate” initial hits such that they should bind tighter and more specifically to CYP121A1.

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Figure 4.20: Binding of MEK066 to CYP124A1. Panel A shows UV-visible absorption spectra from a titration of MEK066 with CYP124A1 (3.2 μM). The inset shows difference spectra from the titration with peak and trough values at 423 nm and 388 nm. Panel B shows a fit of MEK066-induced absorption change (ΔA423 minus ΔA388, reflecting the peak and trough values in the difference spectra, computed by subtracting the spectrum for MEK066-free CYP124A1 from each of the spectra for the MEK066-bound forms) versus [MEK066] added, with data fitted using equation 3 (the Hill equation) to generate a Kd value of 293.1 ± 8.9 µM for MEK066 binding to CYP124A1, n= 2.44 ± 0.14.

Ligand CYP142A1 (Kd, µM)

Type of shift

CYP124A1 (Kd, µM)

Type of shift

CYP125A1 (Kd, µM)

Type of shift

MEK046 3.24 ± 0.07

II 38.0 ± 2.6 II 86.4 ± 5.1 II

MEK065 5.62 ± 0.09

I 89.6 ± 6.3 Rev I ND ND

MEK066 35.3 ± 3.8 I 293.1 ± 8.9

Rev I ND ND

MEK076 1 nm shift I ND ND ND ND



MEK050f2 1 nm shift I ND ND ND ND

Table 4.5: Binding spectral characteristics of Mtb cholesterol oxidases with CYP121A1 elaborated ligands. Kd values were determined as described in the Materials and Methods (section 2.2.9.1). For fragments MEK076, 077, 047 and 050f2, minor type I (high-spin) shifts were observed only with CYP142A1. ND = no binding detected by P450 heme optical titration.

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From the results obtained, CYP142A1 bound tightest to the three MEK compounds

for which Kd values could be determined, while CYP125A1 bound weakest and

showed measurable affinity only for MEK046. CYP142A1 gave type I spectral shifts

(substrate-like) with MEK065 and MEK066, while CYP124A1 showed a reverse type I

spectral shift with these compounds, with the high-spin heme signal at 395 nm

diminished and the low-spin peak at 418 nm increased. MEK046 gave an inhibitor-

like (Soret red shift) with both CYP124A1 and CYP142A1. Since CYP125A1 exists in

the high-spin form, it was not possible to observe the spin shift particularly if the

compound were to give a type I shift; hence only MEK046 gave an observable type

II spectral shift (inhibitor-like). MEK076, MEK077, MEK047 and MEK050F2 showed

either very weak binding (with small type I shifts of ~1 nm) in the case of CYP142A1,

or no binding at all that could be detected optically for the CYP124A1 and

CYP125A1 enzymes.

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4.2.3 CYP124A1 heme iron coordination by carbon monoxide and nitric oxide

Figure 4.21: UV-visible absorbance features of CYP124A1 and its carbon monoxide complex. Absorbance spectra are shown for the resting (ferric) CYP124A1 (3.5 μM, black line), for the sodium dithionite-reduced form with a maximum at 414 nm (red line) and for the CYP124A1 ferrous-CO form at 450 nm (blue line). A shoulder at 423 nm in the Fe2+-CO complex is indicative of a proportion of the enzyme in a form where cysteine thiol (rather than thiolate) is the proximal ligand to the heme iron. After 15 minutes incubation, the P420 form becomes more prominent (magenta dashed line), indicating some instability of the thiolate ligand to protonation in the CYP124A1 ferrous-CO complex. The Soret peak for the resting (ferric) form of CYP124A1 is at 418 nm, consistent

with that for some other characterized Mtb P450s, e.g. CYP142A1 (Driscoll et al.,

2010). However, CYP124A1 is purified in a mixed-spin form with a shoulder at 395

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nm. For comparison, other Mtb P450 enzymes have their low-spin Soret peaks at

416.5 nm (CYP121A1) (McLean et al., 2002a), 419 nm (the sterol 14α-demethylase

CYP51B1) (McLean et al., 2006b, Bellamine et al., 1999), and 418 nm (CYP130A1)

(Ouellet et al., 2008).

A fundamental signature of P450 enzymes is their ability to form a ferrous-CO

adduct on binding of carbon monoxide to ferrous heme iron, with a Soret shift to an

absorbance maximum near 450 nm for the cysteine thiolate coordinated enzyme

(McLean et al., 2009). For CYP124A1, the Fe2+-CO complex spectrum gave a

maximum at 450 nm immediately after the Fe2+-CO complex was formed, with a

shoulder at 423 nm. However, after 15 minutes the 423 nm (P420) species

increased in intensity while the 450 nm (P450) species decreased. The formation of

the P420 form results from protonation of the proximal cysteinate ligand to a thiol.

In previous studies, similar phenomena have been observed whereby the

equilibrium between P450 and P420 changes over periods of minutes. This was

observed previously with Mtb CYP125A1, and also with CYP51B1, where rapid P450

collapse to P420 could be retarded by addition of estriol substrate (McLean et al.,

2009, McLean et al., 2006b).

The P420 Fe2+-CO species was commonly referred to as an “inactivated” form of a

cytochrome P450 enzyme, but further studies on CYP121A1, CYP142A1 and the

P450epoK enzyme from Sorangium cellulosum have shown the P450/P420

equilibrium to be reversible. In CYP142A1, the P420 form could be reconverted into

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P450 by addition of the substrate cholest-4-en-3-one (Ogura et al., 2004, Dunford

et al., 2007, Driscoll et al., 2010).

Figure 4.22: UV-visible absorbance spectra of CYP124A1 in ferric and ferric-NO

bound forms. The resting ferric (black line, 4.8 M), and the ferric-NO bound (red line) forms of CYP124A1 are shown. For the ferric-NO bound form, the Soret, α and β bands are at 432, 573 and 541 nm, respectively.

From the NO-bound CYP124A1 (Figure 4.22) spectral features (Soret band at 433

nm with strong alpha and beta band development at 573 nm and 541 nm,

respectively) are typical of those seen for other Mtb P450s, e.g. CYP130A1,

CYP144A1, CYP125A1 and CYP142A1 (Ouellet et al., 2009, McLean et al., 2009). The

Soret peak increased in intensity on formation of the NO-adduct, due in the main to

the presence of a proportion of high-spin ferric heme iron in the resting state, and

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the conversion to a homogeneous species in the ferric-NO complex (which is likely

formally ferrous-NO+). Similar spectral features were observed for the NO complex

of CYP125A1, which is mainly high-spin in its ferric resting state (McLean et al.,

2009).

Nitric oxide was documented to be an inhibitor of Mtb cytochrome P450 enzymes

(Ouellet et al., 2009). Nitric oxide radical produced by host macrophages inhibit

heme-containing terminal oxidases, inactivate iron-sulfur proteins, and enhance

bacterial entry into latency during the preliminary growth/infection stage of Mtb

(Ouellet et al., 2009).

4.2.4 Determination of the CYP124A1 heme extinction coefficient

CYP124A1 hemoprotein concentration was estimated using the pyridine

hemochromogen method. A sample of ferric CYP124A1 (subsequently determined

as 4.9 μM) was mixed with pyridine and subsequently reduced using sodium

dithionite, as detailed in the Materials and Methods (section 2.2.8). Using the

method of Berry and Trumpower (Berry and Trumpower, 1987), the extinction

coefficient was calculated as ε418 = 110 mM−1 cm−1 for the oxidized, ligand-free

CYP124A1 enzyme. This coefficient takes into consideration that CYP124A1 is

purified in a partially high-spin state, and that the high-spin proportion is quite

consistent. Thus, the coefficient at 418 nm estimates the additional contribution

that the high-spin component would make if CYP124A1 was essentially completely

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low-spin. The extinction coefficients of some other Mtb P450 enzymes have

determined using the pyridine hemochromogen method. These include CYP144A1

at ε420.5 = 100 mM−1 cm−1 (Driscoll et al., 2011), CYP121A1 and CYP51B1 at ε416.5 =

110 mM−1 cm−1 and ε419 = 134 mM−1 cm−1, respectively (McLean et al., 2006b). An

extinction coefficient of ε417 = 115 mM−1 cm−1 was also determined for P450cam

(CYP101A1), a camphor hydroxylase from Pseudomonas putida (Dawson et al.,

1982). The data obtained for CYP124A1 are comparable with the extinction

coefficients of these previously characterized P450s.

The extinction coefficient for CYP124A1 was previously determined by Johnston et

al. to be 91 mM-1 cm-1, using the method developed by Omura and Sato and which

is based on the development of Soret absorption of the Fe2+-CO complex at, or

near, 450 nm (Omura and Sato, 1964). However, as shown in Figure 4.21 above, this

method has limitations with respect to the thiolate-coordinated CYP124A1 P450

form progressively collapsing into the thiol-coordinated P420 species with a Soret

maximum at ∼423 nm. This phenomenon was also observed with CYP51B1, albeit it

occurring more rapidly than with CYP124A1 (Bellamine et al., 1999, Aoyama et al.,

1998). However, the binding of estriol to CYP51B1 did retard considerably the rate

of P450-to-P420 collapse in CYP51B1 (McLean et al., 2006b). In the case of

CYP125A1, a mixture of P450 and P420 species is formed in the Fe2+–CO complex,

and a relatively stable equilibrium is formed (McLean et al., 2009).

280

Figure 4.23: The pyridine hemochromogen complex of CYP124A1. The UV-visible

absorption spectrum is shown for CYP124A1 in the ferric state (black, 4.9 M). Further spectra are shown for the formation of the pyridine complex of CYP124A1 (red), and on reduction to the pyridine hemochromogen complex by sodium dithionite (green). The inset shows a magnification of the 480-650 nm region, highlighting the spectral features of the pyridine hemochromogen complex which absorbs maximally at 556 nm. The extinction coefficient was calculated as ε418 = 110 mM−1 cm−1 for the oxidized, ligand-free CYP124A1 enzyme.

4.2.5 Steady-state kinetic analysis for CYP124A1

Most cytochrome P450 enzymes interact with one or more redox partners to

acquire their reducing equivalents (electrons) (McLean et al., 2005) and the

progression of the P450 catalytic cycle requires consecutive delivery of two

electrons to its heme iron at distinct points in the cycle (Munro et al., 2007b, Ortiz

de Montellano, 2005, Munro et al., 2007a). These electrons are provided by the

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nicotinamide adenine dinucleotide cofactors (NAD(P)H) and their delivery to the

heme iron is mediated by accessory redox partner proteins.

In view of the high affinity of CYP124A1 for a wide variety of substrates, steady-

state kinetic analysis was performed to investigate the apparent rate of substrate

turnover by reconstituting the P450 with spinach ferredoxin (spFDX) and E. coli

flavodoxin reductase (E. coli FLDR) redox partners, hereafter referred to as the

‘spinach system’ in this work. Another set of experiments was also performed

reconstituting CYP124A1 with E. coli flavodoxin (E. coli FLD) and E. coli FLDR

(referred to as the E. coli system in this work). Experiments were carried out as

detailed in the Materials and Methods (section 2.2.17). Substrate-dependent

NADPH oxidation was monitored on additions of varying amounts of the substrate

cholestenone and of the methyl-branched lipids (15-methylpalmitic acid, phytanic

acid, farnesol, geraniol, and geranylgeraniol). These redox systems were shown to

support cholesterol/cholest-4-en-3-one oxidation by Mtb CYP125A1 and CYP142A1

(Driscoll et al., 2010, McLean et al., 2009), However, a comparison of the two

systems was needed to ascertain which system supports the faster substrate-

dependent NADPH oxidation by CYP124A1.

Figure 4.24 shows hyperbolic dependence of NADPH oxidation rate on substrate

concentration for a 1:10:2 (200 nM CYP: 2 M E. coli FLD: 400 nM E. coli FLDR)

reaction mixture, while Figure 4.25 shows hyperbolic dependence of NADPH

oxidation rate on substrate concentration for a 1:10:2 (200 nM CYP: 2 M spFDX:

400 nM E. coli FLDR) mixture. Table 4.6 shows a summary of the kinetic data

282

derived from these experiments. The spinach system produced higher kcat values

(40-122 min-1) relative to the E. coli system, suggestive of a higher catalytic

efficiency with the spinach ferredoxin present. The highest kcat values were

observed for phytanic acid 122.6 ± 5.4 min-1) and cholestenone (120.3 ± 1.7 min-1)

substrates with the spinach system. Phytanic acid showed the highest kcat values

with both redox partner systems, suggesting that this is a good substrate for

CYP124A1. With the spinach system, geraniol showed the lowest Km (0.46 ± 0.10

µM), but also the lowest kcat (40.4 ± 1.5 min-1); while geranylgeraniol showed the

lowest Km (0.33 ± 0.09 µM) and a modest kcat (28.3 ± 1.2 min-1) with the E. coli

system. However, the spinach system failed to function efficiently with

geranylgeraniol. The reason for this is unknown.

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Figure 4.24: Steady-state kinetic analysis for CYP124A1 using an E. coli redox partner system. The redox partner system used was E. coli flavodoxin (E. coli FLD) and E. coli flavodoxin reductase (E. coli FLDR). Kinetic data were collected for turnover studies conducted with CYP124A1 (200 nM final concentration) and with different substrates, as described in the Materials and Methods (section 2.2.17)

using the E. coli redox system. Reactions were initiated by the addition of 200 M NADPH. Data points were collected in triplicate, with error bars showing the S.E.M. Data were fitted using the Michaelis-Menten equation. (A) Phytanic acid, (B) Cholestenone, (C) Farnesol, (D) Geraniol. The kcat and Km values determined are shown in Table 4.6.

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Figure 4.25: Steady-state kinetic analysis for CYP124A1 using the spinach redox system. The redox partner system used was spinach ferredoxin (spFDX) and E. coli flavodoxin reductase (E. coli FLDR). Kinetic data were collected for turnover studies conducted with CYP124A1 (200 nM final concentration) and with different substrates, as described in the Materials and Methods (section 2.2.17) using the

spinach redox system. Reactions were initiated by the addition of 200 M NADPH. Data points were collected in triplicate, with error bars showing the S.E.M. Data were fitted using the Michaelis-Menten equation. (A) Phytanic acid, (B) Cholestenone, (C) Farnesol, (D) Geraniol. The kcat and Km values determined are shown in Table 4.6.

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Substrate E. coli system Spinach system

kcat (min-1) Km (µM) kcat (min-1) Km (µM)

15-Methyl Palmitic Acid 28.6 ± 1.6 1.33 ± 0.39 81.2 ± 9.1 3.30 ± 1.37

Cholestenone 24.0 ± 1.85 1.78 ± 0.70 120.3 ± 1.75 2.68 ± 0.18

Farnesol 26.6 ± 1.1 0.57 ± 0.13 42.1 ± 2.0 0.92 ± 0.25

Geraniol 27.6 ± 2.6 1.65 ± 0.77 40.4 ± 1.5 0.46 ± 0.10

Phytanic Acid 40.0 ± 2.8 2.45 ± 0.73 122.6 ± 5.4 2.67 ± 0.48

Geranyl geraniol 28.3 ± 1.2 0.33 ± 0.09 ND ND

Table 4.6: Steady-state kinetic parameters for substrate-dependent NADPH oxidation by CYP124A1. Turnover studies were done using either the spinach (E. coli FLDR and spinach ferredoxin) or the E. coli (E. coli FLDR and FLD) redox partner systems, as described in the Materials and Methods (section 2.2.17). ND indicates “not determined” – due to negligible geranylgeraniol-dependent stimulation of NADPH oxidation in this case.

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4.2.6 Multiangle Laser Light Scattering (MALLS) analysis of

CYP124A1

Figure 4.26: MALLS analysis of CYP124A1. The figure shows the MALLS profile for

pure, intact CYP124A1 resolved by size exclusion chromatography using a Superdex

200 gel filtration column, with elution in a single peak at 16 ml and a molecular

mass across the single peak determined as 50.37 kDa, close to the predicted mass

of 50.52 kDa from the CYP124A1 amino acid sequence. These data are consistent

with CYP124A1 being a monomeric enzyme.

To investigate the oligomerization state of CYP124A1, MALLS (multiangle laser light

scattering) analysis was carried out. CYP124A1 is clearly a single monomeric

species, and these data are consistent with light scattering studies performed on

CYP125A1 and CYP121A1, both of which appear monomeric in solution (Driscoll et

al., 2010). However, these data contrast with those for CYP142A1, shown in chapter

3 of this thesis. While CYP142A1 appeared dimeric in solution, the dimer formation

287

completely disappeared when the enzyme was treated with DTT. The homogeneity

of CYP124A1 is crucial for its successful crystallization, and successful crystallization

of CYP124A1 forms a major part of the fragment based approach, which requires

structural elucidation of fragment-bound P450 complexes in order to identify

fragment binding modes and to drive fragment merging, growing and elaboration

for the subsequent development of specific and potent inhibitors of the Mtb P450s.

Certain Mtb P450s were shown to crystallize as dimers, for example CYP130A1

(Ouellet et al., 2008). The molecular weight of CYP124A1 estimated from MALLS is

50.37 kDa, which is close to the predicted mass of 50.52 kDa from the CYP124A1

amino acid sequence and consistent with an entirely monomeric species with little

propensity to form dimers in solution.

4.2.7 Thermostability analysis of CYP124A1 by Differential

Scanning Calorimetry DSC is a technique that is used to monitor thermally-induced conformational

changes of proteins by measuring the amount of heat absorbed (or released)

associated with such changes (Privalov and Dragan, 2007, Johnson, 2013). DSC

analysis of CYP124A1 was performed on the ligand-free and ligand-bound

(substrate- and inhibitor-bound) forms of the enzyme to investigate their

thermodynamic properties on unfolding. Experiments were carried out as described

in the Materials and Methods (section 2.2.14).

288

The results (Figure 4.27) obtained revealed minor effects of ligand binding on the

thermodynamic properties (unfolding midpoint temperatures, or Tm values) of

CYP124A1. A summary of the thermodynamic parameters derived from the DSC

experiments is given in Table 4.7. The bifonazole and NMR115-bound CYP124A1

showed two unfolding transitions (Tm values) at 60.31 °C (Tm2, minor) and 53.68 °C

(Tm1, major) for bifonazole and (Tm values) at 54.38 °C (Tm2, major) and 60.43 °C

(Tm1, minor) for NMR115. However, the type II inhibitors (econazole, miconazole,

MEK046 and NMR415) produced a considerable increase in Tm by a value of ~3 to 6

oC, indicating a much more stabilizing effect induced by these type II azoles. The

significant effect on protein stability induced by binding of miconazole may result

from its decreasing the conformational flexibility of CYP124A1 and favouring a

single major conformation, including its influence in providing a new (nitrogen) 6th

ligand to the heme iron. On the other hand, the substrate cholestenone increased

the Tm of CYP124A1 by ~1.4 oC (to 54.31 ± 0.05 oC); while phytanic acid (a methyl

branched lipid) increased the Tm by ~2 oC (to 54.91 ± 0.08 oC), indicating a less

extensive extent of thermal stabilization mediated by binding of substrates to

CYP124A1.

289

Figure 4.27: DSC analysis of CYP124A1 in substrate-free and substrate-bound forms. DSC experiments are shown for CYP124A1 (8 µM) in the absence of ligand (A), and when bound to cholestenone (30 µM) (B), and phytanic acid (30 µM) (C). Data were collected for 8 μM protein samples using a reference sample containing identical buffer and ligand. Final traces were baseline and concentration corrected, and fitted to a non-2-State equation. The unfolding transition midpoints (Tm values), enthalpy (ΔH) and Van’t Hoff enthalpy (ΔHvH) are summarised in Table 4.7.

290

Figure 4.28: DSC analysis of CYP124A1 in azole-bound forms. DSC experiments are shown for CYP124A1 (8 µM) when bound to miconazole (300 µM) (A), econazole (300 µM) (B), clotrimazole (120 µM) (C) and bifonazole (90 µM) (D). Data were collected for 8 μM protein samples using a reference sample containing identical buffer and ligand. Final traces were baseline and concentration corrected, and fitted to a non-2-State equation. The unfolding transition midpoints (Tm values), enthalpy (ΔH) and Van’t Hoff enthalpy (ΔHvH) are summarised in Table 4.7.

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Figure 4.29: DSC analysis of CYP124A1 in MEK compound series-bound forms. DSC experiments are shown for CYP124A1 (8 µM) when bound to MEK046 (301 µM) (A), MEK065 (336 µM) (B), and MEK066 (686 µM) (C). Data were collected for 8 μM protein samples using a reference sample containing identical buffer and ligand. Final traces were baseline and concentration corrected, and fitted to a non-2-State equation. The unfolding transition midpoints (Tm values), enthalpy (ΔH) and Van’t Hoff enthalpy (ΔHvH) are summarised in Table 4.7.

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Figure 4.30: DSC analysis of CYP124A1 in fragment-bound forms. DSC experiments are shown for CYP124A1 (8 µM) when bound to NMR115 (2 mM) (A), NMR415 (1 mM) (B), and MEK066 (50 µM) (C). Data were collected for 8 μM protein samples using a reference sample containing identical buffer and ligand. Final traces were baseline and concentration corrected, and fitted to a non-2-State equation. The unfolding transition midpoints (Tm values), enthalpy (ΔH) and Van’t Hoff enthalpy (ΔHvH) are summarised in Table 4.7.

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CYP124A1 sample Tm (oC) ∆H ( cal mol-1) ∆HvH ( cal mol-1)

Ligand-free CYP124A1

52.89 ± 0.04 1.30 x 105 ± 9.91 x 102 6.95 x 104 ± 6.58 x 102

CYP124A1+ cholestenone

54.31 ± 0.05 1.08 x 105 ± 1.37 x 103 9.72 x 104 ± 1.54 x 103

CYP124A1 + phytanic acid

54.91 ± 0.08 7.87 x 104 ± 1.44 x 103 8.14 x 104 ± 1.86 x 103

CYP124A1 + econazole

58.62 ± 0.05 5.69 x 104 ± 1.06 x 103 1.54 x 105 ± 3.59 x 103

CYP124A1 + Miconazole

56.29 ± 0.06 9.64 x 104 ± 1.8 x 103 1.13 x 105 ± 2.61 x 103

CYP124A1 + clotrimazole

55.18 ± 0.06 1.26 x 105 ± 2.27 x 103 1.17 x 105 ± 2.63 x 103

CYP124A1 + bifonazole

a) 53.68 ± 0.23 b) 60.31 ± 0.17

a) 3.24 x 105 ± 1.52 x 104

b) 1.02 x 105 ± 1.45 x 104 a) 6.82 x 104 ± 1.48 x 103 b) 1.13 x 105 ± 7.44 x 103

CYP124A1 + MEK046 57.24 0.04 1.01 x 105 ± 1.51 x 103 1.62 x 105 ± 3.04 x 103

CYP124A1 + MEK065 54.34± 0.02 1.77 x 105 ± 1.14 x 103 1.02 x 105 ± 8.12 x 102

CYP124A1 + MEK066 52.86 ± 0.03 1.32 x 105 ± 1.14 x 103 1.2 x 105 ± 1.29 x 103

CYP124A1 + NMR115

a) 60.43± 0.23 b) 54.38±0.12

a) 3.05 x 104 ± 3.98 x 103

b) 6.04 x 104 ± 4.07 x 103 a) 1.48 x 105 ± 1.49 x 104

b) 1.13 x 105 ± 5.47 x 103

CYP124A1 + NMR415

57.88± 0.05 8.87 x 104 ± 1.83 x 103 1.57 x 105 ± 4.03 x 103

Table 4.7: DSC data for thermal unfolding of CYP124A1. The thermal transition midpoints (Tm values), calorimetric enthalpy (ΔH) and Van’t Hoff enthalpy (ΔHvH) are shown for the individual thermal transition events observed in ligand-free CYP124A1 and various ligand-bound forms of CYP124A1.

4.2.8 Determination of the heme iron redox potentials of

ligand-free and ligand-bound CYP124A1

Redox titrations were carried out for ligand-free, phytanic acid-bound and

econazole-bound forms of CYP124A1 in order to obtain the midpoint potentials for

the heme iron Fe3+/Fe2+ couples (versus NHE). A redox titration with ligand-free

CYP124A1 revealed a shift in the Soret peak from 419 nm (ferric) to 410 nm

(ferrous), with a decrease in intensity of absorption at the reduced peak. Previous

work on CYP121A1 showed a similar shift from 416.5 nm to 407 nm on heme

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reduction, indicative of the retention of heme thiolate coordination in the ferrous

(reduced) enzyme (McLean et al., 2008).

The inset to Figure 4.28 shows a fit of the redox titration absorbance versus

potential data using the Nernst equation, producing a ligand-free CYP124A1

midpoint potential of -337 ± 4 mV versus NHE. This value is rather more positive

than that for CYP121A1 (−467 ± 5 mV) (McLean et al., 2008), but more negative

than the value obtained for CYP125A1 (−303 ± 5 mV) (McLean et al., 2009),

although this P450 has substantial high-spin content in its resting state. CYP124A1's

potential is in the same range as that for CYP142A1 (−394 ± 4 mV) (see chapter 3 of

this thesis) and P450 BM3 (−427 ± 4 mV) (Ost et al., 2001).

Phytanic acid binding to CYP124A1 generates a substantial shift in the ferric heme

iron spin-state equilibrium toward high-spin (Figure 4.29), and such shifts in spin-

state equilibrium are often accompanied with elevation of the heme iron potential;

i.e. the heme iron develops a more positive potential and becomes easier to reduce

(Bui et al., 2012, Sligar and Gunsalus, 1976). The binding of phytanic acid to

CYP124A1 (Figure 4.29) induced a substantial increase in the heme potential from -

337 ± 4 mV to -230 ± 5 mV, a difference of 107 mV. This difference in potential

between the substrate-free and substrate-bound forms of CYP124A1 is consistent

with the extent of potential change observed for other P450 enzymes that have

been analyzed in their substrate-free and substrate-bound from. These include

CYP51B1 (150 mV positive shift for the Fe3+/Fe2+ couple) (McLean et al., 2006b), and

P450 BM3 (138 mV) (Ost et al., 2001). In prokaryotic P450 enzymes, an electronic

reorganization (in the ferric iron 3d orbitals, from low- to high-spin) is associated

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with substrate binding, resulting in a positive shift in heme iron potential (typically

by ∼140 mV) which enables/enhances heme iron reduction by the NAD(P)H-

dependent redox partners, and where E°’ for NAD(P)H/NAD(P)+ couple is -320 mV

(Daff et al., 1997). The redox titration of phytanic acid-bound CYP124A1 revealed a

shift in the Soret peak from ~393 nm to ~412 nm, with a decrease in intensity of

absorption at the ferrous Soret peak and the development of a strong feature at

548 nm in the Q-band region of the reduced enzyme. These features are consistent

with the retention of the heme thiolate in the ferrous state of the enzyme (McLean

et al., 2006b).

Spectroelectrochemical titrations with econazole-bound CYP124A1 generated a

shift in the Soret peak from 422 nm to 429 nm with a large increase in the intensity

of absorption at the ferrous Soret peak, and with the development of strong split

α/β bands features at ~561 nm and 530 nm, respectively in the reduced enzyme.

These features are indicative of nitrogen ligation of the CYP124A1 heme iron from

an econazole imidazole nitrogen atom in both the oxidized and reduced forms of

the complex. The redox potential for the econazole-bound CYP124A1 (-318 ± 4 mV)

was more negative than that for the substrate-bound enzyme (at -230 ± 5 mV), but

more positive than that for the ligand-free CYP124A1 (at -337 ± 4 mV). This is the

first report of the determination of the redox potential for a Mtb P450 enzyme in

complex with an azole inhibitor drug.

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Figure 4.31: Redox potentiometry of ligand-free CYP124A1. The main panel shows spectra from a redox titration of Mtb CYP124A1 (∼7.5 μM), as described in the Materials and Methods (section 2.2.12). The arrows indicate the direction of absorption changes occurring during the reductive phase of the titration, as the ferric heme iron is reduced to the ferrous form. The heme Soret band at 419 nm (ferric) decreases in intensity and shifts to 410 nm (ferrous), and an absorption band develops at ∼316 nm as the titration nears completion – due to accumulation of dithionite. In the visible region, an absorption band develops at 545 nm on heme reduction. Arrows indicate directions of absorbance change during reduction in the Soret and Q-band regions. The inset shows a plot of heme absorbance change at 419 nm against the applied potential (versus the normal hydrogen electrode, NHE), and with the data fitted using the Nernst equation to produce a midpoint potential of −337 ± 4 mV for the heme iron Fe3+/Fe2+ transition.

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Figure 4.32: Redox potentiometry of phytanic acid-bound CYP124A1. The main panel shows spectral data from a redox titration of CYP124A1 (6.2 μM) bound to phytanic acid (~15 μM). Phytanic acid binding induces an absorbance shift of the ferric low-spin heme iron to the ferric high-spin form at ~393 nm (black spectrum with highest intensity at 393 nm). The progressive addition of sodium dithionite reductant leads to decreases in intensity of the Soret band at 393 nm with a peak shift to a longer wavelength (~412 nm). Arrows indicate directions of absorbance change during reduction in the Soret and Q-band regions. The inset presents a plot of heme absorbance change at 393 nm against applied potential along with the structure of phytanic acid. The data were fitted using the Nernst equation to produce a midpoint potential of -230 ± 5 mV for the Fe3+/Fe2+ transition of the phytanic acid-bound CYP124A1 heme iron.

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Figure 4.33: Redox potentiometry of econazole-bound CYP124A1. The main panel shows spectral data from the redox titration of CYP124A1 (7.5 μM) bound to econazole (~90 μM). Econazole binding induces an absorbance shift of the Soret peak to a longer wavelength (from 419 nm to 422 nm). The progressive addition of sodium dithionite reductant leads to increases in intensity of the Soret band with a further red shift of the peak to 429 nm. Increases in absorbance at ~330 nm are indicative of gradual dithionite accumulation towards the end of the titration. Arrows indicate directions of absorbance change during reduction in the Soret and Q-band regions (at 561 nm and 530 nm). The inset shows a plot of heme absorbance change at 429 nm against applied potential. The data were fitted using the Nernst equation to produce a midpoint potential of -318 ± 4 mV for the Fe3+/Fe2+ transition of the econazole-bound CYP124A1 heme iron. The structure of econazole is also shown in the inset.

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CYP124A1 sample Midpoint potential vs NHE (mV)

Type of Soret shift

Ligand-free CYP124A1 -337 ± 4 Blue

CYP124A1 + phytanic acid -230 ± 5 Red

CYP124A1 + econazole -318 ± 4 Red

Table 4.8: Redox titration data for CYP124A1. The table shows the midpoint redox potential values for the CYP124A1 heme iron Fe3+/Fe2+ transition in ligand-free and substrate-/inhibitor-bound forms, and the type of Soret shifts occurring during heme iron reduction.

4.2.9 Electron Paramagnetic Resonance (EPR) analysis of CYP124A1

4.2.9.1 EPR analysis with CYP124A1 substrates and azole

inhibitors.

Electron paramagnetic resonance (EPR) is a useful technique for studying transition

metal ions in biological systems (Lowe, 1992). It can also be used to study

aggregation state and heme coordination of metalloproteins. The effects of binding

substrate- and inhibitor-like molecules on the heme coordination of CYP124A1

were investigated. Figure 4.34 shows the EPR spectra for CYP124A1 in the ligand-

free form and in complex with cholesterol, cholestenone and HPCD. Continuous-

wave (CW) X-band EPR data for the various ferric forms of CYP142A1 show the

characteristic rhombic resonance pattern of P450 enzymes, consistent with data

derived from previous studies (Driscoll et al., 2011, McLean et al., 2008).

The ligand-free CYP124A1 EPR spectrum revealed a mixture of low-spin and high-

spin ferric species, consistent with data obtained from UV-visible spectroscopy,

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where the enzyme displayed heterogeneous high-spin and low-spin features. The

low-spin species has g-values of gz = 2.39, gy = 2.24, and gx = 1.97 while the high-spin

species produced g-values of 7.98/3.59/1.70. The addition of the substrates

cholestenone and cholesterol produced minor changes to the EPR spectra, with

retention of both high-spin and low-spin features. There is an apparent splitting at

the low-spin gy feature, possibly due to some heterogeneity of the low-spin state. In

addition, there is clearly an increased proportion of high-spin heme iron relative to

the low-spin component, and consistent with observations from UV-visible

spectroscopy in which the substrates cholestenone and cholesterol induced a

substantial increase in the proportion of pentacoordinated (high-spin) heme iron.

This increase in the proportion of high-spin heme iron is indicative of substrate

binding and subsequent removal of the water molecule from the heme iron. The

EPR spectra for HPCD (hydroxypropyl-β-cyclodextrin), the solvent for cholestenone

and cholesterol, was also analyzed. This produced a mixture of low-spin and high-

spin features. However, the amount of high-spin was visibly less that obtained using

the substrates, indicating that the increased high-spin features resulting from

cholestenone and cholesterol was mainly due to the substrates themselves, and not

due to the solvent.

However, the spectra for geranyl geraniol- (GG), geraniol-, 15-methyl palmitic acid-

(15-MPA) and phytanic acid-bound forms of CYP124A1 (Figure 4.35), despite these

lipids being good type I substrates for CYP124A1, produced EPR spectra dominated

by low-spin heme iron, and with minimal high-spin features. This may result from

the different binding modes of these substrates. For instance, these methyl

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branched-lipids may bind more distant from the heme iron compared to

cholestenone, and this could impact greatly on the retention of high-spin heme iron

under the conditions of EPR data collection at 10 K. However, farnesol, phytane and

pristane produced an increase in the proportion of high-spin heme iron relative to

the low-spin component. Form UV-Vis spectroscopy, phytane and pristane showed

low affinity for CYP124A1 as compared to other fatty acids tested; however, this

was not reproduced under the conditions of EPR data collection at 10 K. Addition of

the solvent ethanol (at a final concentration of 1%) perturbs the EPR spectrum and

gives almost complete low-spin signals with g-values of 2.39/2.24/1.93, similar to

the data obtained for geraniol and 15-methyl palmitic acid. This could indicate that

ethanol solvent may have minimal effects on the environment around the heme

iron and the result obtained is due to the effect of the substrates and not the

solvent. EPR studies by Lipscomb on P450cam reported that the signal at g = 1.97 is

unique for a substrate-bound enzyme in which the substrate displaces the water

molecule coordinating the heme iron (Lipscomb, 1980). This postulation is

consistent with data obtained in this study, which revealed CYP124A1 showing a g =

1.97 signal when in complex with cholestenone, cholesterol and some branched-

chain fatty acids. The 1.97/1.98 signal, though present in all the spectra (including

for native CYP124A1 and the solvent control) was found to increase in proportion

with those enzyme-substrate complexes with greater high-spin content. This was

evident for the spectra obtained for cholestenone-, cholesterol-, pristane-, phytane-

, farnesol- and phytanic acid-bound forms of CYP124A1. The proportion of the

1.97/1.98 signal, however, decreased in the enzyme-inhibitor complexes and in

CYP124A1 complexes with substrates that showed more low-spin features.

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For the complexes of CYP124A1 with the azole inhibitors bifonazole, clotrimazole,

econazole and miconazole (Figure 4.36), the high-spin features as seen in the

purified CYP124A1 enzyme have almost completely disappeared in their EPR

spectra. For miconazole-bound CYP124A1, a new rhombic, low-spin species

dominated the EPR spectrum with g-values at 2.38/2.23/1.93, and with a small gz

shoulder at 2.44. Econazole and bifonazole gave similar signals at 2.39/2.23/1.94

and 2.38/2.24/1.93, and with small gz shoulders at 2.43 and 2.45, respectively.

These data are indicative of a novel coordination state in CYP124A1 when bound to

these azole inhibitors. Studies with the CYP121A1-fluconazole complex revealed a

unique feature where fluconazole was shown to coordinate the ferric heme iron by

formation of a direct hydrogen bond to the aqua sixth heme ligand, and with low-

spin g-values at 2.45/2.26/1.90 (Seward et al., 2006). These data are consistent with

those obtained for the major species in the CYP124A1-clotrimazole complex

(2.40/2.25/1.92), in which a gz shoulder at ~2.43 is also seen. The gz ~2.40 signals in

the CYP124A1/azole complexes may result from a proportion of the CYP124A1

enzyme in which water is retained as 6th ligand and/or azole is not bound. The gz

~2.44 signals obtained for econazole, miconazole, bifonazole and clotrimazole may

result from the azole drug interaction with a retained water molecule coordinating

the heme iron. Given that direct azole nitrogen ligation of P450 heme iron is often

associated with an EPR spectrum with gz ~2.50, it may be the case here that each of

these azoles ligate predominantly to the heme iron indirectly via the retained water

6th ligand.

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The EPR spectrum obtained with DMSO solvent (1%) and CYP124A1 was similar to

that obtained with the ligand-free enzyme and buffer. Thus, the spectral changes

produced by the azole inhibitors were clearly due to the influence of the ligands

themselves, and not due to the solvent.

Figure 4.34: EPR spectra for CYP124A1 in ligand-free and sterol-bound forms. X-band CW EPR spectra are shown for purified, ligand-free CYP124A1 and for the P450 in complex with cholesterol, cholestenone and HPCD. The low-spin and high-spin g-values are marked in all cases. Data were collected as described in the Materials and Methods (section 2.2.15).

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Figure 4.35: EPR spectra for CYP124A1 in complex with methyl-branched lipids. X-band continuous wave EPR spectra are shown for purified, ligand-free CYP124A1 and for the P450 in complex with various methyl-branched lipids. The low-spin and high-spin g-values are marked in all cases. Data were collected as described in the Materials and Methods (section 2.2.15).

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Figure 4.36: EPR spectra for CYP124A1 in complex with azole inhibitors. X-band continuous wave EPR spectra are shown for purified, ligand-free CYP124A1 and for the P450 in complex with azole inhibitors. The low-spin and high-spin g-values are marked in all cases. Data were collected as described in the Materials and Methods (section 2.2.15).

4.2.9.2 CYP124A1 EPR analysis with Fragments and MEK

compounds

Further EPR spectra were collected for CYP124A1 in complex with compounds

generated originally from a CYP121A1 fragment based screening study (MEK

compounds MEK046, 065 and 066) (Figure 4.37) and with CYP124A1-specific

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fragment hits (NMR115, NMR356, NMR415 and NMR515) (Figure 4.38). MEK065-

and MEK066-bound CYP124A1 EPR spectra revealed a mixture of low-spin and high-

spin species, similar to that seen with ligand-free CYP124A1, but with a slightly

lower proportion of the high-spin component. This is consistent with the data

obtained from UV-visible spectroscopy, where these compounds showed a reverse-

type I (R-I) spectral shift, converting high-spin ferric heme iron towards low-spin.

The EPR spectrum for MEK065 in complex with CYP124A1 gave ferric low-spin g-

values at 2.38/2.24/1.93, with minor high-spin features at gz = 8.13 and gy = 3.45.

The CYP124A1/MEK066 complex exhibited a similar spectrum to that for MEK065,

again with minor HS features.

The EPR spectrum for the CYP124A1/MEK046 was distinct from those for the

MEK065/066 complexes. In this case there was a splitting of the gz values, with

components at 2.50/2.25/1.89 (major) and 2.43/2.25/1.91. The gz = 2.50

component is likely due to nitrogen ligation of the heme iron from the MEK046

compound – probably from its pendant imidazole group (see Figure 4.19). Typical gz

values for histidine- or imidazole-coordinated P450s lie in the range from 2.65-2.5

(Dawson et al., 1982). DMSO was used as solvent for these ligands and an EPR

spectrum was also collected for CYP124A1 with DMSO. There was minimal

perturbation of the EPR spectrum in this case, and the g-values were similar to

those for the ligand-free CYP124A1 with pronounced high-spin features with g-

values of gz = 8.0 and gy = 3.57, and a low-spin species with g-values of

2.39/2.23/1.93.

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The EPR data for CYP124A1-fragment complexes revealed significant heterogeneity

in spin states. The spectrum for CYP124A1-NMR515 complex revealed a mixture of

low-spin and high-spin species. Two low-spin species are observed for the NMR515

and NMR415 complexes with CYP124A1. The NMR515-CYP124A1 species has

heterogeneous low-spin features with g-values at 2.39/2.23/1.93 (major) and

2.44/2.23/2.00 (minor); while the NMR415-CYP124A1 species also shows

heterogeneous low-spin features with g-values at 2.38/2.24/1.90 (major) and

2.47/2.24/1.93 (minor). As with the azole drugs, these data likely reflect the

interactions of ligand nitrogens with the heme iron (directly or indirectly). While

NMR415 showed the highest affinity (among the fragment series) for CYP124A1

from UV-Vis spectroscopic titrations, NMR515 showed no detectable spectral shift

in UV-Vis titrations with this P450; but showed clear perturbation of the CYP124A1

EPR spectrum at 10 K. In the case of both NMR415 and NMR515, it is postulated

that the low-spin species with the higher gz value reflect the direct interaction of

the pyridine and pyrazole nitrogens, respectively, with the CYP124A1 heme iron.

The low-spin species with the lower gz value instead likely involve indirect

interactions of the drugs via a water molecule retained as the 6th ligand on the

CYP124A1 heme iron. The CYP124A1 complex with NMR356 showed almost

complete low-spin features with no splitting of the gz species, similar to the data

obtained for the CYP124A1-clotrimazole complex. It is possible here that this

reflects the direct coordination of the heme iron by the isoquinoline nitrogen in

NMR356. The NMR115-CYP124A1 complex showed almost complete low-spin

features with the major species at 2.39/2.21/1.93 as well with a minor species seen

as shoulder at gz = 2.46. Again, the former (major) species may reflect indirect (via

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H2O) heme iron coordination, while the latter (minor) species may result from

direct coordination of the heme iron by the NMR115 pyrazole nitrogen.

Figure 4.37: EPR analysis of CYP124A1 bound to MEK ligands. The figure shows both the high-spin and low-spin heme iron regions of the X-band EPR spectrum, with small high-spin features seen for the MEK065/066 complexes – consistent with these drugs causing a reverse type I (R-I) shift in UV-visible titration studies. The heme-coordinating MEK046 removes the entire high-spin component. DMSO solvent alone shows retention of the high-spin component seen in ligand-free CYP124A1, consistent with DMSO not having a major influence on the CYP124A1 spectrum in its own right. Low-spin features for the MEK065/066 complexes are similar to those for ligand-free CYP124A1, as is the spectrum for the DMSO-bound form. The MEK046 spectrum shows a large signal at 2.50/2.25/1.89 – consistent with direct coordination of heme iron via a MEK046 imidazole nitrogen atom. Data were collected as described in the Materials and Methods (section 2.2.15).

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Figure 4.38: EPR spectra of CYP124A1 bound to specific fragment hits. X-band continuous wave EPR spectra are shown for purified, ligand-free CYP124A1 and for the P450 in complex with selected fragment hits. The low-spin and high-spin g-values are marked in all cases. Data were collected as described in the Materials and Methods (section 2.2.15).

4.3 Summary

The identification of novel sulfated metabolites associated with virulence in

Mycobacterium tuberculosis lipid extracts has driven research towards

identification and characterization of enzymes responsible for the sulfation of these

molecules (Holsclaw et al., 2008). The location of CYP124A1 (rv2266) within the

genome of Mtb is adjacent to a region encoding both CYP128A1 (the product of

gene rv2268c, that hydroxylates menaquinone) and Rv2267c (product of rv2267c,

the sulfotransferase that generates the sulfated menaquinone by transfer of a

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sulfate group to the hydroxylated menaquinone). As such, characterization of

CYP124A1 may shed more light on the roles of these enzymes in the metabolism of

methyl-branched lipids and other sterols (Johnston et al., 2009). Further, the genes

CYP121A1 (rv2276) and rv2275 are located close by and play important roles in the

synthesis of a cyclic dipeptide (cYY) and its oxidative crosslinking. Thus, this region

of the Mtb genome is particularly important with respect to oxidative catalysis

involving P450 enzymes (Belin et al., 2009, Vetting et al., 2010).

In this chapter, I report the expression, biophysical and biochemical

characterization of CYP124A1. Optimal protein expression was achieved using the

C41 (DE3) E. coli strain with transformant cells grown in 2YT medium, and

CYP124A1 purification to homogeneity was done via three chromatographic steps,

similarly to the method used for CYP142A1. Optical titrations revealed a high

affinity for cholesterol, cholestenone and methyl branched lipids, further validating

its role as a cholesterol oxidase and a methyl-branched lipid hydroxylase. This wide

spectrum of substrate specificity is indicative of important physiological and

catalytic roles in Mtb. CYP124A1 also demonstrated affinity for a range of azole

inhibitors, some of which have been shown to clear Mtb infection in a mouse model

(Ahmad et al., 2006c). The order of potency of the azole inhibitors (section 1.5.9)

for binding to the Mtb CYP124A1 enzyme (Table 4.2) showed a similar pattern to

that observed with CYP142A1 with clotrimazole (Kd = 4.78 M, MIC = 11 g ml-1 for

Mtb H37Rv) having the tightest Kd among drugs with validated MIC values; followed

by econazole and miconazole (Kd = 18.6 and 19.6 M, respectively; with MIC = 8 g

ml-1 for Mtb H37Rv in both cases) and ketoconazole (Kd = 71 M, MIC = 16 g ml-1

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for Mtb H37Rv ). The general affinity of binding of these azole drugs to CYP124A1 is

weaker than seen for CYP142A1, but the pattern of affinity continues to resemble

that of CYP142A1 with fluconazole binding substantially more weakly than these

other azoles to CYP124A1, to the extent that neglible heme absorbance shift is

observed with fluconazole. The structurally related triazole voriconazole also binds

very weakly to CYP124A1 (Kd = 959 M). However, bifonazole binds much tigher (Kd

= 0.19 M), albeit it causing a type I (substrate-like) shift rather than inhibiting

CYP124A1 by heme iron coordination. Thus, CYPs 142A1 and 124A1 show a similar

pattern of affinity for various azoles with validated MIC values for Mtb H37Rv, likely

consistent with their similar active site architecture that allows

cholesterol/cholestenone binding in both cases.

One of the aims of the fragment based screening approach used in our studies of

Mtb P450 enzymes is to identify commonalities among the cholesterol oxidase

P450 enzymes which could be exploited to develop potent and specific inhibitors

that recognize the CYP124A1, CYP125A1 and CYP142A1 P450 isoforms. These

inhibitors potentially could block completely the utilization of host cholesterol for

energy generation by Mtb during infection. Interestingly, in this work, fragments

that bind each of the three P450 isoforms were identified, and these could be

considered a starting point for fragment merging/linking/growing to produce more

potent inhibitors.

A typical spectrum for CYP124A1 is that of a low-spin, hexa-coordinated P450

enzyme with its Soret peak at 418 nm in its native resting state, but with a shoulder

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at ~394 nm indicative of the presence of a proportion of a high-spin species. This

proportion of high-spin was taken into account when the extinction coefficient was

determined for CYP124A1. Formation of a ferrous-CO adduct is a characteristic

property for P450 enzymes, and this adduct has an absorption maximum close to

450 nm – providing the basis for the nomenclature of this enzyme superfamily

(Omura and Sato, 1964). CYP124A1 formed a CO-adduct at 450 nm (thiolate-

coordinated P450) which was moderately unstable and partially collapsed towards

the P420 form (thiol-coordinated) after a period of several minutes, indicating the

sensitivity of the proximal cysteinate ligand to protonation in the ferrous-CO state.

Steady-state kinetics for CYP124A1 were analysed using a series of substrates, and

the rate constants for substrate-dependent NADPH oxidation were determined

using two sets of redox partners, referred to as the ‘E. coli’ (E. coli flavodoxin

reductase and flavodoxin) and ‘Spinach’ (E. coli flavodoxin reductase and spinach

ferredoxin) systems in this project. The kinetics of NADPH oxidation were faster

using the spinach system, with phytanic acid showing the highest kcat values from

both redox partner systems, suggesting that this should be the best substrate

tested for CYP124A1.

Light scattering analysis showed CYP124A1 to be essentially completely monomeric

in solution. This is an important and desirable property for its successful

crystallization. Stability studies were also performed with DSC, in order to probe the

robustness of the enzyme and its stabilization by the binding of ligands/substrates.

The Tm value for ligand-free CYP124A1 was determined to be 52.9 oC. The type II

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inhibitors (econazole, miconazole, MEK046 and NMR415) produced a considerable

increase in Tm by values of ~3 to 6 oC, indicating a considerable stabilizing effect

induced by the binding of these type II azoles. However, the substrates

cholestenone and phytanic acid showed lower stabilizing effects on CYP124A1, with

Tm increases of ~2 oC.

The midpoint heme iron potential for the ligand-free CYP124A1 P450 enzyme was

found to be quite negative (-337 ± 4 mV), and the Soret band was seen to shift to

shorter wavelength on heme iron reduction (a blue shift, and consistent with

retention of the cysteine thiolate ligation). The CYP124A1 redox potential was

elevated by ~107 mV on binding the substrate phytanic acid, a result consistent

with that observed for a number of other P450 enzymes on binding their substrates

(Ost et al., 2001, McLean et al., 2006b). This elevation in potential enables the P450

to accept electrons from NAD(P)H-dependent redox partners. The redox potential

for the econazole-bound CYP124A1 (-318 ± 4 mV) was more negative than that for

the substrate-bound enzyme, but more positive than that for the ligand-free

enzyme, despite the fact that econazole-bound CYP124A1 has a nitrogen ligand in

the distal position on the heme iron, and that this is not displaced on heme iron

reduction.

EPR analysis confirmed heterogeneity in the CYP124A1 heme iron spin-state, as also

observed by UV-visible spectroscopy. The ligand-free CYP124A1 enzyme displayed

mixed-spin character at both cryogenic temperatures (needed for EPR) and at

ambient temperature (for UV-visible spectroscopy). Interestingly, the binding of

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some methyl-branched lipid substrates (geranyl geraniol (GG), geraniol, 15-methyl

palmitic acid (15-MPA) and phytanic acid) diminished the high-spin content of

CYP124A1 in EPR analysis (despite increasing high-spin content by UV-visible

spectroscopy at ambient temperature). In contrast, EPR analysis of the

CYP124A1/cholestenone and cholesterol complexes showed some enhancement of

the high-spin content, suggesting different binding modes of these substrates to

CYP124A1. However, retention of a substantial high-spin component in substrate-

bound P450s by EPR can vary considerably between different P450s with different

substrates, due primarily to the fact that these studies are typically done at ~10 K

and since the P450 high-spin/low-spin equilibrium is dependent on temperature.

With the azole inhibitors econazole, clotrimazole and miconazole, CYP124A1 high-

spin content was essentially completely removed, and a new low-spin EPR

spectrum appeared that was consistent with distal nitrogen ligation of the heme

iron, as also inferred from binding studies by UV-visible spectroscopy at ambient

temperature. This is also consistent with the data obtained with the CYP124A1-

specific fragments hits NMR115, NMR356 and NMR415.

Collectively, these studies on CYP124A1 have provided novel data on its

thermodynamic, spectroscopic and catalytic features, including quantification of

the productive interactions of this P450 with a variety of substrates, inhibitors and

ligands identified from fragment screening studies. In the next chapter, this work is

extended into analysis of the structural properties of both CYP124A1 and CYP142A1

enzymes in complex with novel substrates and other ligands.

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Chapter 5

Structural biology of ligand-bound complexes of the

cholesterol oxidising P450s CYP142A1 and CYP124A1

5.1 Introduction

Structural biology has emerged as an important tool for the design of new

therapeutics (Malito et al., 2015). A combination of synthetic chemistry and

structural biology of CYP121A1 has played a major role in the fragment based drug

design of potent inhibitors against this enzyme (Hudson et al., 2012b) and this

approach has been extended to other Mtb P450 isoforms. The use of X-ray

crystallography for determining protein structure, for understanding protein

function and for structure-guided design of small-molecule drugs is well

documented, and includes several significant success stories with a large number of

protein structures now available in the protein data bank (PDB) (Malito et al., 2015,

Scapin, 2013). However, despite significant technical advances, the generation of

high-quality crystals for protein X-ray crystallography undoubtedly remains the

major bottleneck in structure determination (Malito et al., 2015).

From the previous chapters in this thesis, CYP124A1 and CYP142A1 were shown to

bind cholesterol/cholestenone (and methyl-branched lipids for CYP124A1) with

high affinities, consistent with their role in sterol metabolism and (for CYP124A1)

oxidation of branched chain lipids. Data presented in chapters 3 and 4 demonstrate

the purification of both of these P450s, leading to the production of highly pure

P450 forms suitable for crystallization both in their ligand-free and ligand-bound

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states. In this chapter, additional data was sought from structural biology (X-ray

crystallography) to provide further insights into the three dimensional structures of

these P450s and how they adapt on binding substrate and inhibitors. These data

would be crucial in ongoing efforts to generate effective inhibitors of the Mtb

cholesterol/cholestenone metabolising P450s using a fragment based approach. For

this reason, crystallographic trials were undertaken using cholestenone as well as

the tight-binding azole inhibitor econazole and a variety of fragment molecules.

Both cholestenone and cholesterol are excellent substrates for CYP124A1 and

CYP142A1, and these molecules are considered to be the natural substrates used in

vivo by both CYP142A1 and CYP125A1, as well as being good substrates for

CYP124A1 (Johnston et al., 2010). The pathway for Mtb-dependent catabolism of

cholesterol/cholestenone is shown in Figure 1.27 in Chapter 1 (Ouellet et al., 2011).

For those Mtb P450s implicated in cholesterol oxidation, crystals structures are

available for CYP124A1, CYP125A1 and CYP142A1. The crystal structures of

CYP125A1 in complex with cholestenone (Ouellet et al., 2010a) and econazole

(McLean et al., 2009) were determined. In the case of CYP124A1, both ligand-free

and phytanic acid-bound forms have been solved previously by Johnston et al.

(Johnston et al., 2009) but, prior to the start of this project, the structure of the

complex with cholest-4-en-3-one (cholestenone) was not resolved. In the case of

CYP142A1, the ligand-free structure was resolved in our group. However, efforts to

obtain a ligand-bound form of the enzyme were unsuccessful, due to the binding of

a PEG molecule present in the mother liquor (Driscoll et al., 2010).

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Structural studies at both the amino acid sequence and tertiary structure level have

provided important insights into the common function and evolutionary origins of

these Mtb cholesterol-oxidase P450 enzymes. It is likely that the differences in

substrate specificity between these enzymes are a consequence of the disparity in

the shape and chemical composition of their respective active sites (Driscoll et al.,

2010). A superimposition of CYP142A1 with the cholestenone-bound CYP125A1

complex (PDB code 2X5W) (Ouellet et al., 2010a) reveals that the active site of

CYP142A1 could provide sufficient space for a cholesterol-like substrate, oriented in

a similar position as that observed in CYP125A1 (Driscoll et al., 2010). However, it

was suggested that it was unlikely for cholesterol to bind to CYP124A1 in the same

orientation, due to the non-complementary shape of the entry of the active site

channel. In this chapter, work is done to explore the structural properties of

different ligand- and substrate-bound complexes of CYP124A1 and CYP142A1.

These studies are aimed at providing a more in-depth understanding of substrate

binding in the Mtb P450s that are implicated in cholesterol oxidation.

5.2 Results and Discussion 5.2.1 X-ray Crystallographic Studies and Structure

Determination for CYP142A1 and CYP124A1 CYP142A1 and CYP124A1 were purified to homogeneity via three chromatographic

steps as described in the Materials and Methods (sections 2.2.6). These steps were

necessary for successful crystallization and structure determination. In this study,

the crystal structures of CYP142A1 in complex with cholestenone, econazole and

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fragment hits were determined, as well as that of CYP124A1 in complex with

cholestenone.

The X-ray data for the complexes were scaled and integrated using the Xia 2

package (Kabsch, 1993). Structures were solved by molecular replacement (McCoy

et al., 2007) with the previously solved CYP142A1 crystal structure as a search

model (PDB 2XKR). The structures were built using COOT (Emsley and Cowtan,

2004) in conjunction with MOLPROBITY (Davis et al., 2007) and refined using Phenix

(Adams et al., 2010). The data collection and refinement statistics are shown in

Table 5.1 and 5.2.

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Data collection CYP142A1:

cholestenone

CYP124A1:

cholestenone

Space group P21212 P212121

Cell dimensions

a, b, c (Å) 65.81 128.13

146.60

46.96 81.35

148.21

Resolution (Å) 58.7-2.09

(2.12-2.09)

74.11-2.65

(2.72-2.65)

Rmeas 0.224 (1.464) 0.163 (0.984)

I/σI 6.0 (1.3) 11.9 (2.2)

Completeness (%) 99.89 (100) 99.86 (97.6)

Redundancy 6.6 7.1

Refinement

No. of reflections 74167 16309

Rwork/Rfree 0.216/0.246

(0.314/0.347)

0.218/0.257

(0.343/0.375)

No. atoms 6811 3510

B-factors (Å2) 28.45 40.2

R.m.s. deviations

Bond lengths (Å) 0.03 0.013

Bond angles (°) 0.802 1.543

Table 5.1: X-ray data collection and refinement statistics for CYP142A1- and CYP124A1- cholestenone complexes

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Data collection CYP142A1: econazole CYP142A1:NMR623 CYP142A1:NMR170 CYP142A1:NMR491 CYP142A1:1-

phenylimidazole

Space group C222 P212121 P212121 P212121 P212121

Cell dimensions

a, b, c (Å) 117.90 185.22 57.58 55.71 65.77 131.81 55.52 65.72 130.57 56.04 65.74 129.44 55.62 65.62

129.06

Resolution (Å) 30.77-2.12

(2.15-2.12)

22.4-2.00

(2.059-2.00)

65.285-1.70

(1.73-1.70)

58.64-1.91

(1.94-1.91)

51.12 – 1.34

(1.355-1.34)

Rmeas 0.085 (0.670) 0.062 (0.701) 0.057 (0.756) 0.102 (0.748) 0.099 (0.753)

I/Σi 11.4 (2.1) 21.6 (3.4) 22.3 (2.7) 14.1 (2.7) 10.5 (2.4)

Completeness (%) 99.89 (100) 99.86 (97.6) 99.89 (100) 99.86 (97.6) 99.89 (100)

Redundancy 3.4 8.9 8.6 6.6 5.9

Refinement

No. of reflections 67981 33457 53352 37809 105716

Rwork/Rfree 0.1926/0.2246

(0.2633/0.2905)

0.181/0.221

(0.233/0.289)

0.169/0.199

(0.253/0.311)

0.169/0.196

(0.265/0.337)

0.188/0.207

(0.276/0.288)

No. atoms 7582 3405 3597 8054 3672

B-factors (Å2) 29.281 38.85 25.61 14.059 13.35

R.m.s. deviations

Bond lengths (Å) 0.008 0.006 0.007 0.008 0.007

Bond angles (°) 1.089 0.982 1.055 1.066 1.158

Table 5.2: X-ray data collection and refinement statistics for CYP142A1- econazole/fragment complexes

321

5.2.1.1 Crystal structure of the CYP142A1:Cholestenone complex

Figure 5.1: Co-crystals of the CYP142A1:cholestenone complex. Crystals were

obtained from 2.4 M ammonium sulfate, 0.1 M sodium acetate, pH 5.5, as

described in the Materials and Methods (section 2.2.18). The crystal structure was

solved to a resolution of 2.09 Å.

Studies have documented that cholesterol/cholestenone serve as major sources of

carbon and energy during both latent and chronic phases of tuberculosis infection

(Pandey and Sassetti, 2008, Miner et al., 2009, Munoz-Elias and McKinney, 2006,

Brzostek et al., 2009). CYP142A1 binds tightly to cholestenone/cholesterol

producing a blue or type I shift (i.e. a shift in the Soret maximum to a shorter

wavelength) (see section 3.2.2). This spectral behaviour is associated with the

displacement of the loosely coordinated distal H2O ligand to the heme iron upon

ligand binding, switching the ferric heme iron from six-coordinated to five-

coordinated.

Diffraction-quality co-crystals of the CYP142A1:cholestenone complex were

obtained as described in the Materials and Methods (section 2.2.18). The X-ray

322

crystal structure of the CYP142A1: cholestenone complex was determined to a

resolution of 2.09 Å. The asymmetric unit contains a CYP142A1 dimer with

cholestenone molecules (Figure 5.2) positioned in an orientation relative to the

heme which is similar to that of the previously published cholestenone structures in

complex with CYP125A1 (Ouellet et al., 2010a) (PDB code 2X5W), CYP142A2

(Garcia-Fernandez et al., 2013) (PDB code 2YOO), and CYP142A2 in complex with

cholesterol sulfate (Frank et al., 2014) (PDB code 4TRI) (see Figures 5.3 and 5.4).

Figure 5.2: Overall view of the CYP142A1 cholestenone-bound complex (dimer). The crystal structure of CYP142A1 is shown in complex with cholestenone, with the H-, G- and I-helices coloured in blue, green and pink, respectively. Cholestenone is highlighted in cyan and yellow spacefill with the heme prosthetic groups in red spacefill. Cholestenone is bound within the active site tunnel with the aliphatic side-chain

facing the distal surface of the heme cofactor, and the 3-keto group pointing

towards the protein surface (Figure 5.3). The active site cavity becomes more

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narrow directly above the heme (i.e. at the catalytic site), to tightly accommodate

the aliphatic side-chain of cholestenone. Cholestenone bound in the active site

makes contact with a series of hydrophobic and polar amino acid residues, as

depicted in Figure 5.4.

It has been documented that the crystal structures of CYP125A1 reveal an active

site entirely enclosed within the protein interior (McLean et al., 2009). However,

this is in contrast to the active site of the cholestenone-bound CYP142A1, where

the carbonyl/keto group of cholestenone remains exposed to the bulk solvent

(Figure 5.3). This difference in topology could explain compensatory or additional

roles linked to CYP142A1 with respect to this P450 metabolizing a pool of sterol

derivatives that are inaccessible to CYP125A1 (Frank et al., 2014). CYP142A1

positions cholestenone such that the aliphatic side chain -terminal carbon is in

close proximity to the heme iron, an alignment favourable for C27-hydroxylation.

324

Figure 5.3: The CYP142A1 substrate binding channel. A view of the cholest-4-en-3-one-bound CYP142A1 is shown, cut by a plane through the substrate binding channel. The H-, G- and I-helices are coloured in blue, green, and pink, respectively. Cholestenone is highlighted in yellow sticks with the heme in red sticks.

325

Figure 5.4: Overview of the CYP142A1-cholestenone binding pocket. Key residues contacting the cholestenone substrate are shown in atom coloured sticks. The heme cofactor is shown with red sticks. For clarity, main chain atoms have been removed. The distances between the C26/C27 terminal carbons and the heme iron are approximately 5.8 Å and 4.3 Å, respectively.

Two terminal methyl groups of the alkyl side chain are positioned at 4.3 Å and 5.8 Å

from the heme iron centre, and interact with Ile76 and Ile65. The alkyl side chain is

positioned well for CYP142A1-mediated oxidation, which is a critical step for

cholestenone ring degradation and for subsequent β-oxidation reactions to

catabolise the steroid (Ouellet et al., 2011). The keto group of cholestenone resides

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at the more hydrophilic end of the substrate binding tunnel and is surrounded by

the residues Met176, Pro179, Thr175 and Gly69.

Figure 5.5: Superimposed structures of ligand-free and cholestenone-bound CYP142A1. The structures are shown with the common P450 secondary structure elements labelled. The protein backbone is depicted by coloured ribbon, with cholestenone in green spacefill and the heme in red sticks. The I-, G- and H-helices (grey for ligand-free and blue for cholestenone-bound forms) undergo conformational change upon substrate binding.

A protein conformational change is observed upon cholestenone binding (Figure

5.5) and includes repositioning of the I-, G- and H-helices to accommodate the

substrate in the active site and to establish hydrophobic contacts with the ligand.

The substrate-induced reorganization of secondary structure elements is largely

confined to the regions typically involved in P450 ligand binding (Poulos and

Johnson, 2005).

327

Substrate-bound CYP142A1 reveals a more open conformation than the substrate-

free form, with the G-, H- and I-helices positioned further from the protein core to

accommodate the substrate molecule. This is consistent with similar structural

reorganizations observed for cholestenone-bound CYP142A2 (Garcia-Fernandez et

al., 2013) and cholestenone-bound CYP125A1 (Ouellet et al., 2010a).

5.2.1.2 Crystal structure of the CYP124A1:Cholestenone complex

CYP124A1 oxidizes methyl-branched chain lipids (Johnston et al., 2009).

Cholesterol/cholestenone has a terminal methyl-branched side chain similar to the

methylated fatty acid substrates of CYP124A1, and a superimposition of the

substrate-bound CYP124A1 structure with CYP142A1 reveals a similar active site

conformation (Driscoll et al., 2010).

Figure 5.6: Co-crystals of the CYP124A1:cholestenone complex. Crystals were

obtained from 0.3 M magnesium formate dihydrate, 0.1 M bis-Tris propane, pH 5.5

as described in the Materials and Methods (section 2.2.18). The crystal structure

was solved to a resolution of 2.54 Å.

328

Diffraction-quality co-crystals of cholestenone bound-CYP124A1 where generated

as described in the Materials and Methods (section 2.2.18). The X-ray crystal

structure of the CYP124A1–cholestenone complex was determined to a resolution

of 2.54 Å. The crystal structure of the CYP124A1:cholestenone complex provides

important insight into how the enzyme catalyses -hydroxylation. A cholestenone

ligand is observed bound to protein in a similar binding orientation as that

previously observed for phytanic acid in complex with CYP124A1 (PDB code 2WM4)

(Johnston et al., 2009) (Figure 5.11). However, this means that CYP124A1 displays a

slightly different binding mode for cholestenone relative to the other cholestenone-

Mtb P450 complexes reported (Figure 5.14). The cholestenone-bound Mtb

CYP124A1 structure reveals a more ‘open’ conformation than the substrate-free

enzyme, which is due to the repositioning of the F-, G-, D- and I-helices (Figure 5.8).

329

Figure 5.7: Overall view of the CYP124A1:cholestenone complex. The crystal structure of CYP124A1 is shown in complex with cholestenone, with the F-, G- and I-helices coloured in yellow, pink, and blue respectively. Cholestenone is highlighted in green spacefill with the heme in red spacefill.

330

Figure 5.8: Superimposed structures of ligand-free and cholestenone-bound forms of CYP124A1. The structures are shown with the common P450 secondary structure elements labelled. The protein backbone is depicted by coloured ribbons, with the cholestenone in cyan spacefill and the heme in red sticks. The pink (ligand-free) and grey (cholestenone-bound) structural elements undergo conformational change upon substrate binding.

331

Figure 5.9: The CYP124A1 substrate binding channel. A view of the CYP124A1:cholestenone complex active site region is shown, cut by a plane through the substrate-binding channel. Cholestenone is represented with green sticks with the heme in red spheres. The F-, G- and I-helices are highlighted in blue, yellow, and pink respectively.

332

Figure 5.10: Overview of the CYP124A1-cholestenone binding pocket. Key residues contacting the cholestenone moiety are shown in atom coloured sticks, the heme cofactor is shown with red sticks and pyrrole nitrogens in blue. The cholestenone is in green with its keto group in red. For clarity, main chain atoms have been removed. The distances between the C26/C27 terminal carbons and the heme iron are approximately 6.3 Å and 4.1 Å, respectively.

The active site of the cholestenone-bound CYP124A1 reveals a channel enclosed

within the protein interior, consistent with that previously documented for

CYP125A1 (McLean et al., 2009). This could be attributed to the evolutionary

relationships between CYP124A1 and CYP125A1; two P450s that share relatively

high sequence identity (40.7%) identity over 428 residues (Driscoll et al., 2010).

333

Cholestenone-binding generates a conformational reorganization of those

secondary structural elements typically involved in the formation of the active site

and substrate-recognition region in P450s. This reorganization is typified by a ‘kink’

or ‘bend’ in the F-, G- and I -helices to enclose the substrate in the active site

(Figure 5.8).

The terminal methyl groups of cholestenone are at 4.1 Å and 6.3 Å away from the

high-spin heme iron, and thus appropriately positioned for -oxidation (Figure

5.10). The reorganization of the secondary structure elements on substrate binding

provides many hydrophobic interactions with the substrate. The I-helix positions

several hydrophobic residues to bind the cholestenone (Leu285, Leu286, Ile284,

Val288, Thr293, Thr294 and Ala297), while the movement of the F-helix relocates

Ile219 and Leu220 to bind the substrate, and Phe231 and Phe234 from the G-helix

also bind cholestenone at its carbonyl group.

Figure 5.11 shows overlaid structures of the cholestenone-bound and phytanic acid-

bound forms of CYP124A1. The distances between the heme iron and the carbon

atoms of the substrates’ branched methyl groups are 3.90 Å and 6.16 Å

(cholestenone) and 3.82 Å and 5.88 Å (phytanic acid), and this proximity is

consistent with the closer methyl groups (in each case) being oxidised by the

reactive iron-oxo (compound I) species in CYP124A1. These observed binding

modes of the substrates are also consistent with the -regiospecificity observed

previously by Johnston et al. (Johnston et al., 2009). The CYP124A1 active site is

relatively narrow, indicative of only one substrate binding conformation and hence

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consistent with a single cholestenone metabolite. Similar to the previously

published data on the CYP124A1-phytanic acid bound structure (Johnston et al.,

2009), the binding mode for cholestenone in the CYP124A1 active site is highly

suggestive of regio-specific hydroxylation of the substrate.

Figure 5.11: Comparison of cholestenone and phytanic acid binding modes to CYP124A1. The image shows an overlay of the cholestenone- and phytanic acid-bound forms of CYP124A1 (PDB code 2WM4) (Johnston et al., 2009). Phytanic acid binds closest to the heme with the terminal carbons at approximately 3.8 Å and 5.9 Å from the heme. The terminal carbons for cholestenone are at 4.1 Å and 6.3 Å from the heme. A comparison of the secondary structure elements reveals minimal differences in their conformations. The inset shows a magnified image of the active site, revealing a similar CYP124A1 binding mode for both cholestenone and phytanic acid.

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5.2.1.3 A comparison of cholestenone-bound CYP124A1, CYP125A1 and CYP142A1 structures

It has been documented that cholesterol/cholestenone is an important source of

energy and carbon for Mtb during both chronic and latent infection (Pandey and

Sassetti, 2008, Chang et al., 2009, Johnston et al., 2010, Yam et al., 2009). A major

stage in the cholesterol degradation pathway is the three-step oxidation of the

cholesterol aliphatic side chain to form the carboxylic acid by CYP125A1 and

CYP142A1. This product is subsequently degraded from both ends of the molecule,

with the involvement of the β-oxidation pathway in the degradation of cholesterol

side chain for energy generation (Ouellet et al., 2011, McLean et al., 2010). All three

P450s (CYPs 124A1, 125A1 and 142A1) can oxidize cholesterol/cholestenone.

However, only the CYP142A1 gene (not CYP124A1) was able complement the defect

associated with the deletion of the CYP125A1 gene from the Mtb CDC1551 strain in

studies in which CYP124A1 and CYP142A1 genes were independently integrated

into the deletion strain under the control of the strong, constitutive hsp60 gene

promoter (Johnston et al., 2010).

A superimposed structure of the three P450 isoforms reveals that CYP125A1 and

CYP142A1 bind cholestenone in a very similar orientation, but that CYP124A1

binding to cholestenone is slightly different (Figure 5.12). Figures 5.13 and 5.14

show comparative binding modes for cholestenone in CYPs 124A1, 125A1 and

142A1 (Figure 5.13) and for cholestenone (and cholesteryl sulfate) binding in

CYP124A1, 125A1, 142A1 and 142A2 (Figure 5.14). In Figure 5.14, the

Mycobacterium smegmatis mc(2) CYP142A2 complexes with both cholestenone and

336

cholesterol sulfate are shown. The data in both Figures demonstrate a different

binding mode for cholestenone in the CYP124A1 enzyme compared to the

CYP125A1 and CYP142A1/A2 P450 enzymes.

As the terminal carbon provides a pro-chiral centre for cholestenone, oxidation of

one methyl group will lead to the 27(S) as opposed to the 27(R) stereochemistry for

the other. The crystal structures suggest that the CYP142A1 complex with

cholestenone is in the pro-S-form, while the CYP124A1 cholestenone complex is in a

pro-R-form. The CYP125A1:cholestenone complex structure places both methyl

groups relatively close to the heme iron, making it more difficult to confidently

predict product stereochemistry. Studies have shown that CYP142A1-driven

cholestenone oxidation results in generation of the 26/27(S)-product, whereas

CYP125A1 produces the opposite stereochemistry (Johnston et al., 2010).

337

Figure 5.12: Comparison of CYP125A1, CYP142A1 and CYP124A1 cholestenone binding modes. CYP125A1 and its bound cholestenone are in green (PDB code 2X5W), while the CYP142A1 and CYP124A1 complexes are in magenta and yellow, respectively. Oxidation of one of the two terminal methyl carbons on the pro-chiral centre will generate an R- or S-product, depending on which methyl group is oxidized. CYP125A1 and CYP142A1 bind cholestenone in a similar orientation, while CYP124A1 binds cholestenone in a different orientation.

338

Figure 5.13: The substrate binding channels in the three P450 cholesterol oxidases. Views are given representing overlays of the three cholestenone ligands (coloured in green sticks (CYP125A1), purple sticks (CYP142A1) and yellow sticks (CYP124A1)) in the active site channels of CYP124A1, CYP142A1 and CYP125A1 (coloured the same way). While CYP125A1 and CYP142A1 exhibit a very similar overall shape, and as a consequence have very similar conformations for the bound ligand, the CYP124A1 active site channel displays a marked kink, accommodated by a more bent conformation of the bound cholestenone.

339

Figure 5.14: Comparison of the cholestenone binding modes for CYP125A1, CYP142A2, CYP142A1 and CYP124A1 with that of the CYP142A2 cholesterol sulfate complex. Structural data for the cholestenone complexes are from CYP125A1 (green, PDB code 2X5W), Mycobacterium smegmatis mc(2) CYP142A2 (pink, PDB code 2YOO), CYP142A1 (white) and CYP124A1 (yellow). The cholesterol sulfate complex of CYP142A2 is shown in blue (PDB code 4TRI). CYP124A1 displays a different binding mode for cholestenone relative to the other complexes.

340

5.2.1.4 Crystal structure of the CYP142A1: Econazole complex

Imidazole and triazole-based drugs are potent cytochrome P450 inhibitors that are

widely used as antifungals, and also possess strong anti-mycobacterial activity

(Seward et al., 2006).

Econazole is an antifungal drug with a potent activity against both latent and

multidrug-resistant forms of tuberculosis, and has been shown to clear Mtb

infection in mouse models (Ahmad et al., 2006c, Ahmad et al., 2006a). Mtb

cytochrome P450 enzymes, including CYP142A1, are therefore possible therapeutic

targets for the azole-based antifungal antibiotics. CYP142A1 exhibits high affinity

for a number of antifungal drugs, including econazole, miconazole, clotrimazole,

and bifonazole (see Table 5.1). However, the azole binding affinities for CYP142A1

are typically lower than those observed for two other structurally characterized

Mtb P450 enzymes: CYP121A1 (McLean et al., 2002b, McLean et al., 2002a) and

CYP51B1 (Guardiola-Diaz et al., 2001, Bellamine et al., 1999, McLean et al., 2002b)

(see Table 5.1). The crystal structures of some Mtb P450s in complex with

econazole have been solved, including the heterocyclic arylamine-binding

CYP130A1 (Ouellet et al., 2008) and CYP125A1 (McLean et al., 2009).

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Azole antibiotics CYP51B1 (µM)

CYP121A1 (µM)

CYP124A1 (µM)

CYP125A1 (µM)

CYP142A1 (µM)

MIC value µg ml-1

Econazole 2.4 ± 0.8 0.024 ± 0.006 18.6 ± 0.5 11.7 ± 0.7 2.3 ± 0.2 8.0

Miconazole ND 0.136 ± 0.021 19.6 ± 0.3 4.6 ± 0.4 1.4 ± 0.2 8.0

Clotrimazole 0.18 ± 0.02 0.073 ± 0.008 4.76 ± 0.13 5.3 ± 0.6 1.1 ± 0.1 11.0

Bifonazole NA NA 0.19 ± 0.02 NA 0.6 ± 0.1 NA

Ketoconazole 5.9 ± 2.7 3.44 ± 0.31 70.9 ± 3.7 27.1 ± 0.9 12.0 ± 0.6 16.0

Fluconazole 5.8 ± 0.1 8.61 ± 0.21 ND 43.1 ± 3.8 309.2 ± 36.3 NA

Voriconazole ND 16.3 ± 2.11 959 ± 47 ND ND NA

Itraconazole ND NA NA 30.2 ± 4.3 ND NA

4-Phenylimidazole ND 32.3 ± 2.2 72.3 ± 1.9 216 ± 5 0.7 ± 0.1 NA

1-Phenylimidazole NA NA 335 ± 13 NA 23.2 ± 3.8 NA

Table 5.3: Dissociation constants for the binding of selected azole drugs to Mtb CYP51A1, CYP121A1, CYP124A1, CYP125A1 and CYP142A1. Comparative data for the binding of azoles to the Mtb CYP121A1 enzyme and MIC data are from (McLean et al., 2008), CYP125A1 from (McLean et al., 2009), CYP51B1 from (McLean et al., 2002b), and CYP142A1 and CYP124A1 from Chapters 3 and 4 of this thesis. ND indicates that a Kd value could not be determined due to lack of any significant heme spectral perturbation induced on binding of the relevant azole to the particular Mtb P450. NA indicates data not available (i.e. no significant spectral shift was induced in these cases). Mtb P450 gene essentially data are fully detailed in Table 1.3 (chapter 1). For the P450s enzymes tabulated here, CYP121A1 is considered essential for Mtb growth, while CYP125A1 is essential for infection of the host and CYP142A1 catalyses the same reaction and may compensate for loss of CYP125A1 activity. CYP51B1 and CYP124A1 appear to be non-essential for Mtb growth in vitro.

UV-Visible spectroscopic titrations indicate that econazole binds CYP142A1 through

direct ligation of the heme iron via an imidazole nitrogen atom, producing a type II

absorbance shift to a longer wavelength (a Soret red shift). Formation of a low-spin

complex involving an indirect iron-azole nitrogen coordination through a water

molecule was observed for the CYP121A1-fluconazole complex, consistent with a

less extensive Soret type II shift observed in optical titrations with this ligand

(Seward et al., 2006).

To explore the binding mode of a potent azole inhibitor with CYP142A1 and in

efforts towards identifying effective inhibitors of the cholesterol metabolising Mtb

P450s, we crystallised the CYP142A1:econazole complex, revealing that econazole

342

ligates CYP142A1 directly via a heterocyclic nitrogen. A comparison with the

CYP130:econazole complex reveals a very similar binding mode (Figures 5.16 and

5.21).

Econazole-bound CYP142A1 co-crystals (Figure 5.15) were obtained from obtained

from 0.2 M potassium thiocyanate, 0.1 M bis-Tris propane, pH 6.5, 20% PEG 3350,

as described in the Materials and Methods (section 2.2.18). The crystal structure of

the CYP142A1:econazole complex was solved to a resolution of 2.12 Å.

Figure 5.15: Co-crystals of the CYP142A1-econazole complex. Crystals were

obtained from 0.2 M potassium thiocyanate, 0.1 M bis-Tris propane, pH 6.5, 20%

PEG 3350 as described in the Materials and Methods (section 2.2.18). The crystal

structure was solved to 2.12 Å.

343

Figure 5.16: Overall view of CYP142A1 econazole-bound complex. The crystal structure of CYP142A1 in complex with econazole is shown, with the H-, G- and I-helices coloured in orange, blue, and pink, respectively. Econazole is highlighted in green-coloured sticks and heme is in red sticks. Panel A shows an overall view of the protein structure and panel B shows an enlarged image of the active site, with the distance between the heme iron and the heterocyclic nitrogen at approximately 2.3 Å.

Econazole binds to CYP142A1 through a set of predominantly hydrophobic

interactions, in addition to the coordination (at 2.28 Å) between the heme iron and

the azole nitrogen lone pair of electrons. Econazole binding induces a

reorganization of the secondary structure elements, with the P450 adopting a more

open conformation with a kink in the I-helix to accommodate the econazole

molecule (see Figure 5.19). The ligand forms hydrophobic contacts with the amino

acid side chains of Val160, Val381, Ser164, Leu163, Phe380, Ile229, Met74, Pro276,

Val277, Ile277, Leu226, Glu223, Ile65, Ile76, Met280, Thr234 and Thr235.

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Figure 5.17: Overview of the econazole binding pocket in CYP142A1. Amino acid residues in contact with the econazole inhibitor ligand are shown in atom coloured sticks (blue carbons, chlorines in green and the ether-linked oxygen in red), while the heme cofactor backbone is shown in red sticks with the heme iron in orange.

345

Figure 5.18: Slab view of the CYP142A1 active site channel showing econazole bound to the heme iron. The image shows the surface of the CYP142A1 active site cavity in grey, the heme prosthetic group in red spacefill, water molecules in magenta and econazole with cyan carbons and coordination to the CYP142A1 heme iron via an imidazole nitrogen.

346

Figure 5.19: Superimposed structures of the ligand-free and econazole-bound forms of CYP142A1 showing their secondary structure elements. Several secondary structure elements in CYP142A1 undergo conformational changes upon econazole inhibitor binding. The secondary structure of the ligand-free enzyme is shown in green while the econazole-bound enzyme is in grey. Econazole is depicted in spacefill with carbons in purple, and heme is in red sticks with pyrrole nitrogens in blue. Structural changes associated with the binding of econazole are particularly clear in this view for the G-, H- and I-helix regions.

The crystal structure of CYP125A1 in complex with econazole was previously

published by (McLean et al., 2009). A comparison of the binding modes of

econazole to CYP125A1 and CYP142A1 reveals large differences. In CYP125A1,

econazole was prevented from binding directly to the heme by steric constraints

attributable to the funnel shape of the active site near the heme (McLean et al.,

2009) (Figure 5.20). In contrast to the CYP125A1-econazole complex (where

econazole was positioned at ~9.3 Å from the heme), the econazole molecule bound

347

to CYP142A1 ligates the heme iron directly with a distance of only 2.3 Å between

the heme iron and the heterocyclic nitrogen ligand.

Figure 5.20: Superimposed structures of the CYP125A1 and CYP142A1 econazole

complexes. The image shows the binding modes of econazole to CYP125A1 (PDB

code 3IW2) and CYP142A1. The CYP125A1-econazole structure is shown in blue,

while the CYP142A1-econazole structure is in violet. Econazole ligates to the

CYP142A1 heme iron directly via a heterocyclic nitrogen, while CYP125A1 binds

econazole in a position distant from the heme.

348

Figure 5.21: Superimposed structures of the CYP142A1 and CYP130A1 econazole

complexes. The image shows the binding modes of econazole to CYP130A1 (PDB

code 2UVN) (Ouellet et al., 2008) and to CYP142A1. The CYP130A1 structure is

shown in wheat colour while the CYP142A1 structure is in white. Econazole ligates

both the CYP142A1 and CYP130A1 heme iron directly via a heterocyclic nitrogen.

The CYP130A1-econazole complex crystallized as a dimer, whereas the CYP142A1-

econazole complex crystallized as a monomer. A comparison of their econazole

binding modes reveals very similar orientations.

5.2.1.5 Crystal structures of CYP142A1 in complex with fragment- based screening hits

As part of this research project, novel inhibitors of CYP142A1 were sought using a

fragment screening approach. Central to the success of such a project is the

identification of small ligands that bind to the P450 target (using thermal shift and

high-throughput NMR methods), followed by the determination of the binding

349

modes of compound “hits” – done here using X-ray crystallographic methods

(Fischer and Hubbard, 2009). The identification of the binding modes of small

ligands in the target proteins then enable strategies such as fragment linking,

merging or chemical elaboration to be done to generate next-phase molecules with

improved affinity for the protein. Ligand-free crystals for CYP142A1 were obtained

as described in the Materials and Methods (section 2.2.18). These crystals were

used to soak small molecule ligands that were identified from fragment-based

screening with CYP142A1 (done with collaborators at the University of Cambridge),

to determine the respective binding modes and to enable further elaboration of

these ligands. A crystal soaking strategy was considered the best option, since most

of these ligands possessed weak affinity for CYP142A1 (see section 3.2.4).

Diffraction quality ligand-free crystals of CYP142A1 were obtained from 0.1 M

sodium acetate in a pH range of 4.7–5.0 with 0.1 M potassium thiocyanate, 8-10%

PEG 200 (v/v), and 8-12% PEG 550MME (v/v), using 15 mg/ml enzyme as described

in the Materials and Methods (section 2.2.18). However, this resulted in the

generation of crystals with PEG200 (from the mother liquor) bound to the heme

cofactor. This problem was overcome by back-soaking native crystals in 24% PEG

550 MME, 0.1 M sodium acetate (pH 4.5), 0.1 M potassium thiocyanate.

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Figure 5.22: A sample of diamond-shaped CYP142A1 native crystals. Crystals were obtained from 0.1 M sodium acetate across a pH range of 4.7–5.0 with 0.1 M potassium thiocyanate, 8-10% PEG 200 (v/v), and 8-12% PEG 550MME (v/v) using 15 mg/ml enzyme.

CYP142A1 crystals were back-soaked for 24 hours in 24% PEG 550 MME, 0.1 M

sodium acetate (pH 4.5), 0.1 M potassium thiocyanate, and then supplemented

with fragment hits at concentrations ranging from 1-4 mM. From the six fragments

tested, only two fragments led to ligand-bound structures, possibly due to factors

such as the low affinity of the compounds at this early stage in the fragment

screening process, or their binding to multiple positions in the protein, or their

inability to access the active site of the crystallized P450. However, crystals of

CYP142A1 complexes with NMR170 and NMR623 were obtained, and these were

solved to resolutions of 1.70 Å (NMR170-bound CYP142A1) and 2.0 Å (NMR623-

bound CYP142A1).

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Figure 5.23: The CYP142A1-NMR623 complex structure. Two NMR623 ligands are bound near to the CYP142A1 heme. One (lower) molecule ligates the heme iron directly via an imidazole/pyrazole nitrogen, while the second (upper) molecule binds closer to the protein surface. NMR623 is shown in atom coloured sticks (carbons in cyan and bromine in brown) and transparent spheres, and the heme is in red with pyrrole nitrogens in blue and the heme iron in orange. The inset shows the NMR623 chemical structure.

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Figure 5.24: The CYP142A1-NMR170 complex structure. Two NMR170 ligands are bound close to the heme in the active site. One NMR170 molecule ligates the heme directly via a pyridine nitrogen atom, while the second molecule binds further up the active site. NMR170 is depicted in yellow sticks (with the nitrogen in blue) and transparent spheres; heme is in pink (with pyrrole nitrogens in blue and the iron in orange) and the I-helix is shown in light purple. The inset shows the NMR170 chemical structure.

In view of the failure to obtain CYP142A1 complexes by soaking pre-formed P450

crystals with four of the fragments, these remaining fragments were subjected to

co-crystallization trials with CYP142A1, which yielded two additional complexes

with the fragments NMR491 and 1-phenylimidazole.

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Figure 5.25: Co-crystals of the CYP142A1-NMR491 complex. Crystals were obtained from 0.2 M magnesium chloride, 0.1 M sodium chloride (pH 5.3-5.6), 6-12% PEG 20K, 8-10% PEG 550 MME, as described in the Materials and Methods (section 2.2.18). The crystal structure of the complex was solved to a resolution of 1.91 Å.

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Figure 5.26: The structure of the CYP142A1-NMR491 complex. A single NMR491 ligand is bound to the heme iron via an imidazole nitrogen. NMR491 is depicted in wheat-coloured stick and transparent spheres, with imidazole nitrogens in blue. The heme is in purple with pyrrole nitrogens in blue and heme iron in orange, and the CYP142A1 I-helix is in yellow. The inset shows the NMR491 chemical structure.

Figure 5.27: Co-crystals of the CYP142A1:1-phenylimidazole complex. Crystals were obtained from 0.2 M magnesium chloride, 0.1 M Na acetate (pH 5.3-5.6), 6-12% PEG 20K, 8-14% PEG 550 MME, as described in the Materials and Methods (section 2.2.18). The crystal structure was solved to a resolution of 1.34 Å.

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Figure 5.28: The structure of the CYP142A1:1-phenylimidazole complex. The structure shows that one molecule of 1-phenylimidazole is bound to the heme, with coordination of the iron by an imidazole nitrogen. The 1-phenylimidazole is depicted in magenta-coloured stick and transparent spheres, with nitrogen atoms in blue. The heme is in red with pyrrole nitrogens in blue and the heme iron as an orange sphere. The I-helix is shown in green.

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Figure 5.29: Superimposed structures of the various CYP142A1-fragment complexes. The figure shows 1-phenylimidazole (1-PIM) in magenta-coloured sticks; NMR170 in yellow sticks, NMR491 in green sticks and NMR623 in cyan sticks (two molecules of the heme-coordinating NMR623 molecule are shown in this figure, with the second molecule distant from the heme). The heme is in red with pyrrole nitrogens in blue and heme iron as an orange sphere. The image reveals the binding modes of the four fragments in the CYP142A1 active site, and presents a good starting point for fragment merging/linking/growing via synthetic chemistry to produce more potent CYP142A1 inhibitors.

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Figure 5.30: Superimposed structures of ligand-free and fragment-bound CYP142A1 enzymes. The figure shows the protein backbone with the native structure in grey, the 1-phenylimidazole (1-PIM) complex in magenta; the NMR170 complex in yellow, the NMR491 complex in green and the NMR623 complex in cyan. The heme is shown in red sticks with pyrrole nitrogens in blue and heme iron as an orange sphere. No significant reorganization of the secondary structural elements was observed on binding of these fragments to CYP142A1, despite all these molecules ligating to the heme iron.

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Figure 5.31: Superimposed structures of CYP142A1-fragment complexes with the CYP142A1 econazole complex. The figure shows 1-phenylimidazole (1-PIM) in magenta-coloured sticks; NMR170 in yellow sticks, NMR491 in green, NMR623 in cyan, econazole in orange sticks (with chlorine atoms in green and oxygen in red) and the CYP142A1 heme backbone in red with pyrrole nitrogens in blue and heme iron in orange. Fragments (all containing either imidazole or pyridine groups that coordinate the CYP142A1 heme iron in the distal position) show generally similar binding modes to econazole (a much larger azole drug and clinically used CYP51 inhibitor), with only NMR170 occupying a different position in the active site in the second of its two binding modes.

The crystal structures of the CYP142A1 P450 in complex with the fragments

NMR623 (4-bromo-1H-imidazole), NMR170 (4-benzylpyridine), NMR491 (4-phenyl

Imidazole) and 1-phenylimidazole revealed that all fragments ligate the heme iron

in the distal position via coordination through a nitrogen lone pair of electrons. In

addition, NMR623 and NMR170 (see Figures 5.23 and 5.24) revealed two molecules

binding in the active site, with one directly ligating the heme in each case and the

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other binding in the active site further from the heme (for NMR170), or at the

protein surface (for NMR623). This is possibly a consequence of the high

concentrations of fragment ligands used during the soaking experiments. However,

NMR491 and 1-phenylimidazole (both obtained through co-crysallisation, see

Figures 5.26 and 5.28) show only one molecule binding for each compound and

with a similar binding mode observed for the two compounds (involving a direct

coordination of the heme iron). While the second binding mode for the NMR623

fragment is at a superficial site on the P450, the second molecule on NMR170 binds

in the active site cavity in a position away from the heme iron. This is a useful

finding in terms of developing a fragment linking/merging/building strategy, since

the binding modes of the other fragment ligands are clustered around the

CYP142A1 heme (see below). In previous fragment screening studies with Mtb

CYP121A1, a phenolic fragment was found to bind in two adjacent positions in the

active site (Hudson et al., 2012b). These positions mapped closely onto the space

occupied by the cyclic dipeptide substrate cyclo-L-tyr-L-tyr, and these data also

point to how fragment screening can be used to probe physiological roles in

enzymes. No significant conformational reorganization in the CYP142A1

polypeptide chain was associated with the binding of the fragments to the P450

(Figure 5.30).

Even though a relatively large gap is observed between the two molecules of

NMR170 bound in the CYP142A1 complex (Figure 5.24), further studies using

synthetic chemistry could still generate second generation molecules that bridge

this gap using an appropriate linker in order to generate more potent inhibitors and

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tighter binding ligands. Overlapping fragments were found in the heme-binding

active site position, and these provide a rational start point for fragment

elaboration involving a linking/merging/growing strategy via synthetic chemistry to

produce specific inhibitors as probes for CYP142A1 function both in vitro and in

vivo. As mentioned above, this chemical elaboration strategy has previously been

successfully applied to CYP121A1 (Hudson et al., 2012b, Hudson et al., 2014).

Interestingly, these fragments were found to bind all the three cholesterol oxidases

(CYP125A1, CYP124A1 and CYP142A1) (see Tables 4.2 and 4.3) , indicating a good

start point for the development of potent inhibitors that are effective against each

of these three cholesterol oxidising Mtb P450s, and hence could completely block

host cholesterol utilization by Mtb.

An overlay of the structures of CYP142A1-fragment complexes (bound to small

azole- and pyridine-based inhibitors) with the CYP142A1-econazole complex (a

much larger azole inhibitor) (Figure 5.31) reveals a similar binding mode and

proximity to the heme iron for each of these molecules, and provides important

insights into the selective inhibition and druggability of this enzyme that could be

achieved by the further development of the fragment screening strategy.

5.3 Summary

CYP125A1, CYP142A1 and CYP124A1 are involved in the C27 oxidation of the

aliphatic side chain of cholesterol/cholestenone, a critical step leading to

cholesterol utilization for energy generation during both latent and chronic

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infection in the human host (Ouellet et al., 2011, Johnston et al., 2010). In this

chapter, the crystal structures of various CYP124A1 and CYP142A1 ligand

complexes were determined, including structures of complexes with the substrate

(cholestenone, with CYP124A1) and with azole (and pyridine) inhibitors, including

econazole (with CYP142A1), a drug shown to be effective in clearing Mtb infections

in a mouse model (Ahmad et al., 2006c). Cholestenone was found to bind to

CYP142A1 in an orientation that is similar to that seen in other, previously resolved

mycobacterial P450 structures (i.e. the CYP125A1 and CYP142A2 complexes)

(Ouellet et al., 2010a, Frank et al., 2014, Garcia-Fernandez et al., 2013). However,

the crystal structure of CYP124A1 bound to cholestenone revealed a different

cholestenone binding orientation, due to its narrow active site channel. It was

previously postulated by Driscoll et al. that it may be unlikely for

cholesterol/cholestenone to bind to CYP124A1 in the same orientation as seen in

the other cholesterol-metabolizing P450 enzymes, due to the distinct shape of the

entry of the active site channel (Driscoll et al., 2010). The cholestenone molecules

bound within the active sites of CYP124A1 and CYP142A1 make contacts with

hydrophobic residues. A superimposition of the substrate-free and substrate-bound

structures revealed conformational changes in the secondary structure elements,

mainly in the I-, G- and F-helices, in order to accommodate the substrate in the

active sites of these enzymes. Cholestenone binds to CYP124A1 and CYP142A1 with

its aliphatic tail positioned directly above the heme in an orientation favourable for

C27 hydroxylation, while the keto group projects outwards to the solvent-accessible

surface. A comparison of the active site channels of the three different types of Mtb

cholesterol hydroxylases revealed that the substrate-binding tunnel for CYP142A1

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provides more space as it approaches towards the protein surface when compared

to CYP125A1 and CYP124A1. This indicates the possibility of binding multiple

cholesterol derivatives, and suggests that CYP142A1 enzymes may play an essential

role during Mtb infection by providing access to more diverse cholesterol

derivatives that otherwise would not be available to the pathogen, as postulated by

Frank et al. (Frank et al., 2014).

Structural data presented in this chapter validate further the biophysical binding

experiments, nanoESI-MS and EPR data. From the nano ESI-MS data, the dimeric

features of CYP142A1 were almost completely eliminated on incubation with DTT

and econazole, but retained with cholestenone. These data are comparable to the

results derived from the structural studies which showed econazole to bind

CYP142A1, while cholestenone binds CYP142A1 in a dimer and CYP124A1 binds

cholestenone in a monomer.

Azole antibiotics are known to be rather non-selective, but potent inhibitors of Mtb

P450 enzymes, and some of these azoles have been shown to clear Mtb infection in

mice (Ahmad et al., 2006c, Ahmad et al., 2006a). The crystal structure of CYP142A1

in complex with econazole revealed a direct heme iron coordination via a

heterocyclic (imidazole) nitrogen, consistent with the canonical mechanism of P450

enzyme inhibition by azole drugs. An interesting observation was that the crystal

structure of the CYP125A1:econazole complex showed that the drug could not

penetrate deep enough into the active site to interact with the heme as a result of

the narrowing of its funnel-shaped access channel to the heme, and instead bound

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in a mode in which the imidazole moiety was orientated away from the active site

(McLean et al., 2009). These data point to interesting differences in the structural

organization of the CYP142A1 and CYP125A1 enzymes that might help to

understand subtle differences in their catalytic properties.

Initial fragment based screening identified six fragments hits for CYP142A1.

Interestingly, these fragments also bind the other two Mtb cholesterol oxidase

P450s (CYP124A1 and CYP125A1) albeit with lower affinity (see Tables 4.2 and 4.3).

Crystal structures for four of the fragments were resolved in complex with

CYP142A1 in this study, and these are NMR491, NMR623, NMR170 and 1-

phenylimidazole (1-PIM). For NMR170 and NMR623, two molecules were found to

bind in different positions to CYP142A1 (one ligating the heme iron in each case),

while single NMR491 and 1-PIM molecules were bound coordinated to the

CYP142A1 heme iron. It is worth noting that each of these fragments are heme-

binders and ligate the heme iron directly via a heterocyclic nitrogen in an imidazole

or pyridine ring; as also observed for the much larger econazole inhibitor. The

crystallographic data for CYP142A1 and CYP124A1 in complex with substrates and

inhibitors are confirmatory of ligand-binding data presented in Chapters 3 and 4. In

the case of cholestenone, this substrate induces a low- to high-spin shift (type I) in

the ferric heme iron in both these P450s, consistent with a binding mode in which

the 6th water ligand to the heme iron is displaced. This is confirmed from the

structural data for the cholestenone complexes in both cases. Similarly, econazole

coordinates the heme iron to induce a red (long wavelength) shift of the Soret

maximum in both CYP124A1 and CYP142A1, which is consistent with an imidazole

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nitrogen displacing the 6th water ligand. This phenomenon is observed clearly in the

crystal structure of the CYP142A1/econazole complex. Similarly, CYP142A1 heme

iron distal coordination by the imidazole or pyridine nitrogens of the fragment

molecules NMR170, NMR491, NMR623 and 1-phenylimidazole is inferred from UV-

visible and EPR studies, and is again confirmed in the crystal structures of the

CYP142A1 complexes with these molecules. Binding data for the various substrates

and inhibitors (including fragments) used in structural studies are presented in

Table 3.2 and Tables 4.1-4.3 in Chapters 3 and 4 of this thesis. The fragment-based

approach used in this work aims at developing potent and selective inhibitors for

important drug targets such as the Mtb P450 enzymes (Hudson et al., 2012b,

Hudson et al., 2014). With the binding modes of these fragments determined, a

door is opened for further chemical elaboration via merging/linking/growing

strategies in order to develop novel inhibitors that could completely block host

cholesterol utilization/metabolism by Mycobacterium tuberculosis.

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Chapter 6

Conclusions and Future Directions

6.1 Conclusions

This PhD research provides a comprehensive study of the biochemical, biophysical

and structural properties of the cholesterol oxidase P450 enzymes in Mycobacterium

tuberculosis.

A cholesterol catabolism gene cluster was discovered recently in the genome of the

Mtb-related actinobacterium Rhodococcus jostti RHA1 (Van der Geize et al., 2007).

Interestingly, many of these genes were also found to be conserved in Mtb, among

which are genes that code for proteins involved in both cholesterol uptake and

degradation (including CYP125A1 and CYP142A1), suggesting that Mtb can utilize

cholesterol for growth during infection (Ouellet et al., 2011). The

cholesterol/branched chain fatty acid binding CYP124A1, on the other hand, is

located in the same gene operon as CYP128A1 and CYP121A1. One of the other

genes in this cluster (Sft3, Rv2267c) encodes a sulfotransferase enzyme involved in

the sulfation of menaquinone MK-9 DH-2 at the -position (Johnston et al., 2009).

The hydroxylation of menaquinone at the -position by CYP128A1 enables the Sft3-

catalysed sulfation at this position. Interestingly CYP124A1 was found to metabolize

substrates with chemical structures similar to menaquinone and these are

substrates with repeated methyl branching, which includes

cholesterol/cholestenone (Johnston et al., 2010, Johnston et al., 2009).

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These P450 isoforms (CYPs 125A1/142A1/124A1) have been shown to be involved

in the C27 hydroxylation of cholesterol side chain subsequent to cholesterol ring

degradation necessary for energy generation, bacterial survival and infectivity in

Mtb (Ouellet et al., 2011, Johnston et al., 2010). In the absence of these P450

enzymes, the cholesterol metabolite cholestenone accumulates and exerts a

bacteriostatic effect (Ouellet et al., 2011). CYP125A1 is the primary enzyme

involved in cholesterol catabolism, while CYP142A1 functions as a compensatory

enzyme to the cholesterol 27-oxidase CYP125A1 in some Mtb strains. CYP124A1 is

found in both pathogenic and non-pathogenic mycobacterial species and shows a

broad lipid substrate specificity. These P450 enzymes were demonstrated to be

promising drug targets for new generations of anti-tuberculosis drugs. Targeting

these three enzymes and the cholesterol degradation pathway in Mtb could

provide a much needed new route to killing the bacterium.

CYP142A1 may play a “redundant” role in Mtb cholesterol metabolism, where it can

serve as a back-up enzyme for CYP125A1 (Johnston et al., 2010). However, it may

also catalyse sterol metabolism reactions that are complementary to those

performed by CYP125A1. Even though all three of the P450s can oxidize the

cholesterol side chain, only CYP142A1, but not CYP124A1, can substitute for

CYP125A1 when its gene is inactivated in Mtb (Johnston et al., 2010). From my

work, the crystal structures of CYP124A1 and CYP142A1 in complex with

cholestenone revealed a close proximity of the aliphatic side chain of the sterol to

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the heme iron centre, consistent with their role as cholesterol/cholestenone C27

oxidases.

CYP124A1 and CYP142A1 were successfully purified via three chromatographic

steps, with CYP142A1 being predominantly low-spin and CYP124A1 purified in a

mixed-spin form. Binding assays carried out on the three P450 isoforms revealed

tight binding to sterols, and to branched-chain lipids (for CYP124A1) consistent with

their proposed physiological roles. Studies have revealed azole antifungals to be

potent inhibitors of mycobacterial P450 enzymes, with econazole shown to clear

Mtb infection in mice (Ahmad et al., 2006a). Affinity for selected azole antibiotics

and novel inhibitors from fragment-based screening was also established for the

cholesterol-metabolising P450s, consistent with other previously characterized Mtb

P450s (McLean et al., 2002b, Seward et al., 2006, Hudson et al., 2014). Specific

inhibitors of the cholesterol oxidase P450 enzymes could be used as chemical tools

to reveal how the activities of these enzymes relate to Mtb infection, growth and

persistence in the human host.

CYP142A1 and CYP124A1 exhibit fundamental characteristics of P450 enzymes with

respect to cysteine thiolate coordination of the heme iron and affinity for molecules

such as carbon monoxide (CO) and nitric oxide (NO). The characteristic signature for

P450 enzymes is that they display a shift in the Soret peak to approximately 450 nm

(in the ferrous-CO complex) on heme iron reduction and binding with carbon

monoxide. This “P450” spectrum is a result of the retention of the thiolate proximal

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ligand to the heme iron when CO binds trans to the cysteinate ligand (Omura and

Sato, 1964). The carbon monoxide adduct for CYP142A1 was stable for over 30

minutes in both the absence and the presence of the substrate cholestenone and

did not convert to a P420 (cysteine thiol-coordinated) complex, which absorbs

maximally at around 420 nm. However, this was in contrast to CYP124A1 where the

423 nm (P420) species gradually increased in intensity while the 450 nm (P450)

species continuously decreased in intensity over 15 minutes.

Electron paramagnetic resonance (EPR) spectroscopy was done to probe CYP142A1

and CYP124A1 heme coordination and ligand binding. X-band EPR data collected at

10 K for ligand-free and ligand-bound forms of the enzymes revealed a

characteristic rhombic signal for ferric, thiolate-coordinated P450s, consistent with

conclusions made from UV-visible absorbance spectra. Heterogeneous low-spin

heme iron signals were obtained on binding fragments and azole inhibitors to the

P450s, arising from either direct ligation of a heterocyclic nitrogen in the

fragment/ligand to heme iron, or through their making indirect interactions with

heme iron via a retained water ligand coordinating the heme iron in the 6th

position. EPR data for CYP142A1/CYP124A1 complexed with relevant substrates

demonstrated a heterogeneous mixture of high-spin and low-spin features,

consistent with data obtained from previously characterized P450s (McLean et al.,

2008, Driscoll et al., 2011). The high-spin species result from the displacement of

the water ligand coordinating to the heme iron in the 6th axial position following the

binding of the substrate. EPR provides important confirmatory information for the

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binding of inhibitor and substrate molecules to CYP142A1 and CYP124A1, in

addition to providing insights into their modes of binding.

The structure of ligand-free CYP142A1 was previously published as a monomer

(Driscoll et al., 2010). However, results from light scattering analysis revealed that

CYP142A1 can dimerize in solution, and this dimerization could be disrupted with

the reducing agent DTT (dithiothreitol). Appropriately treated CYP124A1 was found

to be completely monomeric in solution, appearing as a single species through

MALLS analysis with an apparent molecular weight of 50.37 kDa, close to the

predicted mass of 50.52 kDa from the CYP124A1 amino acid sequence. The stability

parameters of a protein give important indications of its probability for successful

crystallization to enable high-resolution structural studies (Dupeux et al., 2011).

Studies have revealed that samples with Tm values of 318 K (45 oC) or higher

crystallized in 49% of cases, while the success rate of crystallization declined rapidly

for samples with lower Tm values (Dupeux et al., 2011). Thermal stability studies of

CYP142A1 and CYP124A1 generated Tm values of 53.29 ± 0.06 oC and 52.89 ± 0.04

oC, respectively, which can be taken as positive indications for successful

crystallizability of these P450s.

The heme iron mid-point redox potential values for CYP142A1 and CYP124A1 were

consistent with values previously reported for other characterized P450 enzymes

(McLean et al., 2008, Ost et al., 2001, Driscoll et al., 2011). Ligand-free CYP142A1

has a more negative mid-point potential (−394 ± 4 mV) than the ligand-free

CYP124A1 (−337 ± 4 mV). Substrate binding to P450 enzymes usually induces

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significant shifts in heme iron spin-state equilibrium toward the high-spin ferric

state, and such shifts in spin-state equilibrium are typically associated with the

heme iron developing a more positive potential and becoming easier to reduce

(Daff et al., 1997, Bui et al., 2012). Cholestenone binding to CYP142A1 produced a

large heme iron potential change (~244 mV) between the low-spin (substrate-free)

and high-spin (substrate-bound) forms, indicative of a tightly regulated redox

system. The extent of substrate-induced potential change is rather greater than

observed for other P450s (e.g. ~140 mV for P450 BM3 and P450cam), and might

reflect the impact of changes to heme environment or structural arrangements in

addition to the change in heme iron spin-state on substrate binding to CYP142A1.

The CYP124A1 redox potential was elevated by ~107 mV on binding the substrate

phytanic acid, a result consistent with that observed for a number of other P450

enzymes on substrate binding (Ost et al., 2001, McLean et al., 2006b). Azole binding

to CYP124A1 and CYP142A1 resulted in a more heme iron negative potential (−360

± 4 mV for the CYP142A1-clotrimazole complex and −318 ± 4 mV CYP124A1-

econazole complex) than found for their respective substrate-bound forms,

although these are slightly more positive potentials than found in the ligand-free

forms of the enzymes.

Structural data for the CYP142A1 and CYP124A1 enzymes revealed active site

architecture that favours the binding of a sterol molecule. Significant reorganization

of various secondary structural elements were observed on substrate- and

inhibitor-binding, and this was most evident at the I-, G- and H-helices. The

cholestenone complex of CYP142A1 showed that the positioning of the

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cholestenone molecule within the substrate binding channel is similar to that

observed in the crystal structures of other mycobacterial P450 enzymes that have

been solved in complex with cholestenone (Frank et al., 2014, Garcia-Fernandez et

al., 2013). Cholestenone binds in an orientation that favours C27-hydroxylation,

consistent with recent data reporting the catalytic properties of the enzyme

(Johnston et al., 2010, Ouellet et al., 2010a, Ouellet et al., 2011). A superimposed

structure of the three Mtb cholesterol oxidase P450 isoforms reveals that

CYP125A1 and CYP142A1 bind cholestenone in a very similar orientation, but that

CYP124A1 binding to cholestenone is slightly different, resulting from its distinctive

active site cavity. Crystal structures of CYP142A1 in complex with econazole and

fragment hits revealed direct heme ligation by azole nitrogens, and in the case of

the fragments NMR170 and NMR623 two molecules were bound to the P450, with

one ligating the heme iron and the other bound more distant from the heme. The

binding modes of CYP142A1-specific hits were also determined in my studies. By

combining structural biology with synthetic chemistry, the “linking” of such

molecules that bind to different regions of the CYP142A1 active site could enable

production of highly specific P450 ligands to be used as chemical probes for the

Mtb cholesterol oxidase enzymes and/or as leads for new drug development for TB.

The results presented in this thesis are mainly for two of the cholesterol

metabolizing P450 enzymes in Mtb (CYP124A1 and CYP142A1) and more

importantly these enzymes are among the 20 P450s that are potential drug targets.

The biochemical, biophysical and structural data presented provide important clues

about their in vivo function. The substrate specificity data presented here, together

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with the molecular interactions of novel inhibitors from fragment based screening

and the crystal structures of the various ligand-bound complexes, provide a scaffold

for design and development of specific inhibitors for the Mtb cholesterol oxidases

and of other Mtb P450 enzymes.

In conclusion, these studies have provided fundamental new data on the

spectroscopic, ligand-binding, kinetic, thermodynamic, hydrodynamic and

structural properties of two Mtb P450s pivotal to the metabolism of sterols (and

branched chain lipids in the case of CYP124A1). Substantial new data are presented

in all these areas, with novel structural data being particularly important in

understanding how CYP142A1 and CYP124A1 interact with the substrate

cholestenone, and (for CYP142A1) how various inhibitors interact with this P450

and how a fragment based screening approach could be useful in the production of

new inhibitors and probes for the function of CYP142A1 (and of the related Mtb

cholesterol oxidases CYP125A1 and CYP124A1). Such an approach will complement

ongoing fragment screening work on Mtb CYP121A1 and CYP144A1 and could lead

to development of novel therapeutics against Mtb.

6.2 Future directions

Work done in this thesis led to the identification of small chemicals (“fragments”)

that bind to the CYP124A1 and CYP142A1 P450s. Fragments are generally weak

binding ligands with low affinity for their targets; although the recognition of their

binding is frequently due to their occupancy of a specific binding “pocket” in the

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target enzyme (Hudson et al., 2014). In this study, the design/identification of new

inhibitors common to the three Mtb cholesterol oxidases was undertaken using

fragment based screening approaches, and in collaboration with researchers at the

University of Cambridge. Fragments hits were validated against the cholesterol

oxidases and crystal structures of fragments in complex with CYP142A1 were

determined for some of the fragment “hits”. Further progress in this area could be

made by using fragment linking/merging/growing strategies to improve the potency

and affinities of next-generation molecules developed from the fragment hits.

Through synergistic studies using protein crystallography to define binding modes

for fragments and synthetic chemistry to link neighbouring fragments; potent

inhibitors should be sought and their ability to inhibit CYPs 124A1/125A1/CYP142A1

determined, along with their effects on growth of mycobacterial strains.

Binding assays were also done for CYP124A1-specific fragment hits. However,

crystallisation trials with CYP124A1 and these fragments were not successful,

possibly due to low affinity of the fragment hits. Future work on this enzyme should

progress studies to identify the binding modes of fragment hits by X-ray

crystallography (using both co-crystallisation and crystal-soaking approaches). Once

the CYP124A1 binding modes for sufficient numbers of fragments have been

determined, the same approaches detailed above for CYP142A1 could be followed

until highly specific and potent inhibitors are developed. These should be tested at

each stage for their ability to bind to CYP124A1 (and the other Mtb cholesterol

oxidases) and to inhibit enzymatic activity (Kd and Ki values), as well as for their anti-

Mtb activity (MIC values), with the aim being to generate new, potent drugs that

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are effective against drug-resistant strains of Mtb. For CYP125A1, while structural

data are available, protein crystal packing results in occlusion of the substrate

binding site – impairing the soaking of inhibitory fragments into the active site, and

thus preventing determination of their binding modes. Protein engineering on

CYP125A1 should enable its crystallization in a form with a more open active site

that will enable fragments (various hits have already been identified) to be soaked

into the active site. Co-crystallization of CYP125A1 with fragments should also be

done to expedite identification of fragment binding modes and to progress

CYP125A1 inhibitor development. The overall aim is to build novel types of P450

isoform-specific inhibitors that inactivate CYP142A1, 124A1 and 125A1, and that

can effectively inhibit host cholesterol utilization and human infection by Mtb.

375

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396

Appendix

ATGGGCAGCAGCCATCATCATCATCATCACAGCAGCGGCCTGGTGCCGCGCGGCAGCCATAT

GACTGAAGCTCCGGACGTGGATCTGGCCGACGGCAACTTCTACGCCAGCCGCGAGGCGCGGG

CCGCGTACCGGTGGATGCGGGCCAACCAACCGGTGTTCCGCGATCGCAACGGCCTGGCGGCC

GCGTCGACGTACCAGGCGGTGATCGACGCCGAACGTCAACCCGAGCTGTTCTCCAACGCCGG

CGGCATCCGCCCCGACCAGCCCGCCCTGCCGATGATGATCGACATGGACGATCCCGCACATC

TGTTGCGGCGCAAGCTGGTTAACGCCGGCTTCACCCGCAAGCGGGTGAAGGACAAGGAGGCG

TCGATTGCCGCGCTGTGTGACACCCTGATCGACGCCGTGTGCGAACGCGGCGAGTGTGACTT

CGTGCGGGACCTGGCCGCGCCGCTACCGATGGCGGTGATCGGCGACATGCTCGGGGTGCGTC

CAGAGCAGCGGGACATGTTCTTGCGGTGGTCCGACGATCTGGTGACATTCCTCAGTTCGCAT

GTGTCTCAAGAGGATTTCCAGATCACCATGGACGCCTTCGCGGCCTACAACGACTTCACCCG

GGCCACCATTGCGGCACGGCGAGCGGACCCCACCGACGACCTGGTCAGCGTGCTGGTGAGTT

CCGAAGTTGACGGCGAGCGGCTAAGCGACGACGAGCTGGTCATGGAGACGCTGCTGATCCTG

ATCGGCGGCGACGAGACCACGCGGCATACCTTGAGCGGTGGTACCGAGCAGCTGCTGCGCAA

CCGTGACCAGTGGGACCTGCTGCAGCGCGACCCGTCGTTGCTGCCCGGGGCCATCGAGGAGA

TGCTACGTTGGACCGCCCCGGTAAAGAACATGTGCCGGGTGTTGACCGCGGATACCGAGTTT

CACGGCACGGCGTTGTGTGCCGGCGAGAAGATGATGCTGCTCTTCGAGTCGGCGAACTTCGA

CGAGGCGGTTTTCTGTGAACCGGAAAAGTTTGATGTTCAGCGAAATCCAAACAGCCACTTGG

CGTTTGGCTTCGGCACGCATTTCTGCCTGGGCAATCAGCTGGCCCGGTTGGAGCTGTCGTTG

ATGACGGAACGGGTGTTGCGGCGGCTACCCGACCTGCGGTTGGTCGCCGATGACTCCGTGTT

GCCGCTGCGGCCGGCGAACTTTGTCAGCGGCCTGGAATCCATGCCGGTGGTGTTCACGCCGA

GCCCGCCGCTGGGCTGAGGGATCC

GGCTGCTAACAAAGCCCGAAAGGAAGCTGAGTTGGCTGCTGCCACCGCTGAGCAATAACTAG

CATAACCCCTTGGGGCCTCTAAACGGGTCTTGAGGGGTTTTTTGCTGAAAGGAGGAACTATA

TCCGGATATCCCGCAAGAGGCCCGGCAGTACCGGCATAACCAAGCCTATGCCTACAGCATCC

AGGGTGACGGTGCCGAGGATGACGATGAGCGCATTGTTAGATTTCATACACGGTGCCTGACT

GCGTTAGCAATTTAACTGTGATAAACTACCGCATTAAAGCTTATCGATGATAAGCTGTCAAA

CATGAGAATTCTTGAAGACGAAAGGGCCTCGTGATACGCCTATTTTTATAGGTTAATGTCAT

GATAATAATGGTTTCTTAGACGTCAGGTGGCACTTTTCGGGGAAATGTGCGCGGAACCCCTA

TTTGTTTATTTTTCTAAATACATTCAAATATGTATCCGCTCATGAGACAATAACCCTGATAA

ATGCTTCAATAATATTGAAAAAGGAAGAGTATGAGTATTCAACATTTCCGTGTCGCCCTTAT

TCCCTTTTTTGCGGCATTTTGCCTTCCTGTTTTTGCTCACCCAGAAACGCTGGTGAAAGTAA

AAGATGCTGAAGATCAGTTGGGTGCACGAGTGGGTTACATCGAACTGGATCTCAACAGCGGT

AAGATCCTTGAGAGTTTTCGCCCCGAAGAACGTTTTCCAATGATGAGCACTTTTAAAGTTCT

GCTATGTGGCGCGGTATTATCCCGTGTTGACGCCGGGCAAGAGCAACTCGGTCGCCGCATAC

ACTATTCTCAGAATGACTTGGTTGAGTACTCACCAGTCACAGAAAAGCATCTTACGGATGGC

ATGACAGTAAGAGAATTATGCAGTGCTGCCATAACCATGAGTGATAACACTGCGGCCAACTT

ACTTCTGACAACGATCGGAGGACCGAAGGAGCTAACCGCTTTTTTGCACAACATGGGGGATC

ATGTAACTCGCCTTGATCGTTGGGAACCGGAGCTGAATGAAGCCATACCAAACGACGAGCGT

GACACCACGATGCCTGCAGCAATGGCAACAACGTTGCGCAAACTATTAACTGGCGAACTACT

TACTCTAGCTTCCCGGCAACAATTAATAGACTGGATGGAGGCGGATAAAGTTGCAGGACCAC

TTCTGCGCTCGGCCCTTCCGGCTGGCTGGTTTATTGCTGATAAATCTGGAGCCGGTGAGCGT

GGGTCTCGCGGTATCATTGCAGCACTGGGGCCAGATGGTAAGCCCTCCCGTATCGTAGTTAT

CTACACGACGGGGAGTCAGGCAACTATGGATGAACGAAATAGACAGATCGCTGAGATAGGTG

CCTCACTGATTAAGCATTGGTAACTGTCAGACCAAGTTTACTCATATATACTTTAGATTGAT

TTAAAACTTCATTTTTAATTTAAAAGGATCTAGGTGAAGATCCTTTTTGATAATCTCATGAC

CAAAATCCCTTAACGTGAGTTTTCGTTCCACTGAGCGTCAGACCCCGTAGAAAAGATCAAAG

GATCTTCTTGAGATCCTTTTTTTCTGCGCGTAATCTGCTGCTTGCAAACAAAAAAACCACCG

CTACCAGCGGTGGTTTGTTTGCCGGATCAAGAGCTACCAACTCTTTTTCCGAAGGTAACTGG

CTTCAGCAGAGCGCAGATACCAAATACTGTCCTTCTAGTGTAGCCGTAGTTAGGCCACCACT

TCAAGAACTCTGTAGCACCGCCTACATACCTCGCTCTGCTAATCCTGTTACCAGTGGCTGCT

GCCAGTGGCGATAAGTCGTGTCTTACCGGGTTGGACTCAAGACGATAGTTACCGGATAAGGC

GCAGCGGTCGGGCTGAACGGGGGGTTCGTGCACACAGCCCAGCTTGGAGCGAACGACCTACA

397

CCGAACTGAGATACCTACAGCGTGAGCTATGAGAAAGCGCCACGCTTCCCGAAGGGAGAAAG

GCGGACAGGTATCCGGTAAGCGGCAGGGTCGGAACAGGAGAGCGCACGAGGGAGCTTCCAGG

GGGAAACGCCTGGTATCTTTATAGTCCTGTCGGGTTTCGCCACCTCTGACTTGAGCGTCGAT

TTTTGTGATGCTCGTCAGGGGGGCGGAGCCTATGGAAAAACGCCAGCAACGCGGCCTTTTTA

CGGTTCCTGGCCTTTTGCTGGCCTTTTGCTCACATGTTCTTTCCTGCGTTATCCCCTGATTC

TGTGGATAACCGTATTACCGCCTTTGAGTGAGCTGATACCGCTCGCCGCAGCCGAACGACCG

AGCGCAGCGAGTCAGTGAGCGAGGAAGCGGAAGAGCGCCTGATGCGGTATTTTCTCCTTACG

CATCTGTGCGGTATTTCACACCGCATATATGGTGCACTCTCAGTACAATCTGCTCTGATGCC

GCATAGTTAAGCCAGTATACACTCCGCTATCGCTACGTGACTGGGTCATGGCTGCGCCCCGA

CACCCGCCAACACCCGCTGACGCGCCCTGACGGGCTTGTCTGCTCCCGGCATCCGCTTACAG

ACAAGCTGTGACCGTCTCCGGGAGCTGCATGTGTCAGAGGTTTTCACCGTCATCACCGAAAC

GCGCGAGGCAGCTGCGGTAAAGCTCATCAGCGTGGTCGTGAAGCGATTCACAGATGTCTGCC

TGTTCATCCGCGTCCAGCTCGTTGAGTTTCTCCAGAAGCGTTAATGTCTGGCTTCTGATAAA

GCGGGCCATGTTAAGGGCGGTTTTTTCCTGTTTGGTCACTGATGCCTCCGTGTAAGGGGGAT

TTCTGTTCATGGGGGTAATGATACCGATGAAACGAGAGAGGATGCTCACGATACGGGTTACT

GATGATGAACATGCCCGGTTACTGGAACGTTGTGAGGGTAAACAACTGGCGGTATGGATGCG

GCGGGACCAGAGAAAAATCACTCAGGGTCAATGCCAGCGCTTCGTTAATACAGATGTAGGTG

TTCCACAGGGTAGCCAGCAGCATCCTGCGATGCAGATCCGGAACATAATGGTGCAGGGCGCT

GACTTCCGCGTTTCCAGACTTTACGAAACACGGAAACCGAAGACCATTCATGTTGTTGCTCA

GGTCGCAGACGTTTTGCAGCAGCAGTCGCTTCACGTTCGCTCGCGTATCGGTGATTCATTCT

GCTAACCAGTAAGGCAACCCCGCCAGCCTAGCCGGGTCCTCAACGACAGGAGCACGATCATG

CGCACCCGTGGCCAGGACCCAACGCTGCCCGAGATGCGCCGCGTGCGGCTGCTGGAGATGGC

GGACGCGATGGATATGTTCTGCCAAGGGTTGGTTTGCGCATTCACAGTTCTCCGCAAGAATT

GATTGGCTCCAATTCTTGGAGTGGTGAATCCGTTAGCGAGGTGCCGCCGGCTTCCATTCAGG

TCGAGGTGGCCCGGCTCCATGCACCGCGACGCAACGCGGGGAGGCAGACAAGGTATAGGGCG

GCGCCTACAATCCATGCCAACCCGTTCCATGTGCTCGCCGAGGCGGCATAAATCGCCGTGAC

GATCAGCGGTCCAGTGATCGAAGTTAGGCTGGTAAGAGCCGCGAGCGATCCTTGAAGCTGTC

CCTGATGGTCGTCATCTACCTGCCTGGACAGCATGGCCTGCAACGCGGGCATCCCGATGCCG

CCGGAAGCGAGAAGAATCATAATGGGGAAGGCCATCCAGCCTCGCGTCGCGAACGCCAGCAA

GACGTAGCCCAGCGCGTCGGCCGCCATGCCGGCGATAATGGCCTGCTTCTCGCCGAAACGTT

TGGTGGCGGGACCAGTGACGAAGGCTTGAGCGAGGGCGTGCAAGATTCCGAATACCGCAAGC

GACAGGCCGATCATCGTCGCGCTCCAGCGAAAGCGGTCCTCGCCGAAAATGACCCAGAGCGC

TGCCGGCACCTGTCCTACGAGTTGCATGATAAAGAAGACAGTCATAAGTGCGGCGACGATAG

TCATGCCCCGCGCCCACCGGAAGGAGCTGACTGGGTTGAAGGCTCTCAAGGGCATCGGTCGA

GATCCCGGTGCCTAATGAGTGAGCTAACTTACATTAATTGCGTTGCGCTCACTGCCCGCTTT

CCAGTCGGGAAACCTGTCGTGCCAGCTGCATTAATGAATCGGCCAACGCGCGGGGAGAGGCG

GTTTGCGTATTGGGCGCCAGGGTGGTTTTTCTTTTCACCAGTGAGACGGGCAACAGCTGATT

GCCCTTCACCGCCTGGCCCTGAGAGAGTTGCAGCAAGCGGTCCACGCTGGTTTGCCCCAGCA

GGCGAAAATCCTGTTTGATGGTGGTTAACGGCGGGATATAACATGAGCTGTCTTCGGTATCG

TCGTATCCCACTACCGAGATATCCGCACCAACGCGCAGCCCGGACTCGGTAATGGCGCGCAT

TGCGCCCAGCGCCATCTGATCGTTGGCAACCAGCATCGCAGTGGGAACGATGCCCTCATTCA

GCATTTGCATGGTTTGTTGAAAACCGGACATGGCACTCCAGTCGCCTTCCCGTTCCGCTATC

GGCTGAATTTGATTGCGAGTGAGATATTTATGCCAGCCAGCCAGACGCAGACGCGCCGAGAC

AGAACTTAATGGGCCCGCTAACAGCGCGATTTGCTGGTGACCCAATGCGACCAGATGCTCCA

CGCCCAGTCGCGTACCGTCTTCATGGGAGAAAATAATACTGTTGATGGGTGTCTGGTCAGAG

ACATCAAGAAATAACGCCGGAACATTAGTGCAGGCAGCTTCCACAGCAATGGCATCCTGGTC

ATCCAGCGGATAGTTAATGATCAGCCCACTGACGCGTTGCGCGAGAAGATTGTGCACCGCCG

CTTTACAGGCTTCGACGCCGCTTCGTTCTACCATCGACACCACCACGCTGGCACCCAGTTGA

TCGGCGCGAGATTTAATCGCCGCGACAATTTGCGACGGCGCGTGCAGGGCCAGACTGGAGGT

GGCAACGCCAATCAGCAACGACTGTTTGCCCGCCAGTTGTTGTGCCACGCGGTTGGGAATGT

AATTCAGCTCCGCCATCGCCGCTTCCACTTTTTCCCGCGTTTTCGCAGAAACGTGGCTGGCC

TGGTTCACCACGCGGGAAACGGTCTGATAAGAGACACCGGCATACTCTGCGACATCGTATAA

CGTTACTGGTTTCACATTCACCACCCTGAATTGACTCTCTTCCGGGCGCTATCATGCCATAC

CGCGAAAGGTTTTGCGCCATTCGATGGTGTCCGGGATCTCGACGCTCTCCCTTATGCGACTC

CTGCATTAGGAAGCAGCCCAGTAGTAGGTTGAGGCCGTTGAGCACCGCCGCCGCAAGGAATG

GTGCATGCAAGGAGATGGCGCCCAACAGTCCCCCGGCCACGGGGCCTGCCACCATACCCACG

398

CCGAAACAAGCGCTCATGAGCCCGAAGTGGCGAGCCCGATCTTCCCCATCGGTGATGTCGGC

GATATAGGCGCCAGCAACCGCACCTGTGGCGCCGGTGATGCCGGCCACGATGCGTCCGGCGT

AGAGGATCGAGATCTCGATCCCGCGAAATTAATACGACTCACTATAGGGGAATTGTGAGCGG

ATAACAATTCCCCTCTAGAAATAATTTTGTTTAACTTTAAGAAGGAGATATACC

Figure S1: CYP142A1 (Rv3518c) DNA sequence. The CYP142A1 gene was cloned into the plasmid vector pET15b using NdeI/BamHI restriction enzyme sites to create the CYP142A1/p15b plasmid (6898 bases). NdeI (CATATG) and BamHI (GGATCC) PCR-engineered restriction sites are underlined. The start and stop codons are shaded in red.

CATATGGCACATCACCACCACCATCACTCCGCGGCTCTTGAAGTCCTCTTTCAGGGACCCGG

GTACCAGGATCCGATGGGTCTGAATACCGCAATTGCAACCCGTGTTAATGGTACACCGCCTC

CGGAAGTTCCGATTGCAGATATTGAACTGGGTAGCCTGGATTTTTGGGCACTGGATGATGAT

GTTCGTGATGGTGCATTTGCAACCCTGCGTCGTGAAGCACCGATTAGCTTTTGGCCGACCAT

TGAACTGCCTGGTTTTGTTGCAGGTAATGGTCATTGGGCACTGACCAAATATGATGATGTTT

TTTATGCAAGCCGTCACCCGGATATCTTTAGCAGCTATCCGAATATTACCATCAATGATCAG

ACACCGGAACTGGCAGAATATTTTGGTAGCATGATTGTTCTGGATGATCCGCGTCATCAGCG

TCTGCGTAGCATTGTTAGCCGTGCATTTACCCCGAAAGTTGTTGCACGTATTGAAGCAGCAG

TTCGTGATCGTGCACATCGTCTGGTTAGCAGCATGATTGCAAATAATCCGGATCGTCAGGCA

GATCTGGTTAGCGAACTGGCAGGTCCGCTGCCGCTGCAGATTATTTGTGATATGATGGGTAT

TCCGAAAGCCGATCATCAGCGTATTTTTCATTGGACCAATGTGATTCTGGGTTTTGGTGATC

CGGATCTGGCAACCGATTTTGATGAATTTATGCAGGTTAGCGCAGATATTGGTGCATATGCC

ACCGCACTGGCCGAAGATCGTCGTGTTAACCATCATGATGATCTGACCAGCAGCCTGGTTGA

AGCCGAAGTTGATGGTGAACGTCTGAGCAGCCGTGAAATTGCCAGCTTTTTTATCCTGCTGG

TTGTTGCCGGTAATGAAACCACCCGTAATGCAATTACCCATGGTGTTCTGGCACTGAGCCGT

TATCCGGAACAGCGTGATCGTTGGTGGTCAGATTTTGATGGTCTGGCACCGACCGCAGTTGA

AGAAATTGTTCGTTGGGCAAGTCCGGTTGTTTATATGCGTCGTACCCTGACCCAGGATATCG

AACTGCGTGGCACCAAAATGGCAGCCGGTGATAAAGTTAGCCTGTGGTATTGTAGCGCAAAT

CGTGATGAAAGCAAATTTGCAGATCCGTGGACCTTTGATCTGGCACGTAATCCGAATCCGCA

TCTGGGCTTTGGTGGTGGTGGTGCACATTTTTGTCTGGGTGCAAATCTGGCACGTCGTGAAA

TTCGTGTTGCATTTGATGAACTGCGTCGTCAGATGCCGGATGTTGTTGCAACCGAAGAACCG

GCACGTCTGCTGAGCCAGTTTATTCATGGTATTAAAACCCTGCCGGTTACCTGGTCATAATA

AGCTTGCGGCCGCAGAGCTCGCTCTGGTGCCACGCGGTAGTAAAGAAACCGCTGCTGCTAAA

TTCGAACGCCAGCACATGGACAGCTCTACTTCTGCTGCTCTCGAGGCTTAATTAACCTAGGC

TGCTAAACAAAGCCCGAAAGGAAGCTGAGTTGGCTGCTGCCACCGCTGAGCAATAACTAGCA

TAACCCCTTGGGGCCTCTAAACGGGTCTTGAGGGGTTTTTTGCTGAAAGGAGGAACTATATC

CGGATVATGGCGAATGGGACGCGCCCTGTAGCGGCGCATTAAGCGCGGCGGGTGTGGTGGTT

ACGCGCAGCGTGACCGCTACACTTGCCAGCGCCCTAGCGCCCGCTCCTTTCGCTTTCTTCCC

TTCCTTTCTCGCCACGTTCGCCGGCTTTCCCCGTCAAGCTCTAAATCGGGGGCTCCCTTTAG

GGTTCCGATTTAGTGCTTTACGGCACCTCGACCCCAAAAAACTTGATTAGGGTGATGGTTCA

CGTAGTGGGCCATCGCCCTGATAGACGGTTTTTCGCCCTTTGACGTTGGAGTCCACGTTCTT

TAATAGTGGACTCTTGTTCCAAACTGGAACAACACTCAACCCTATCTCGGTCTATTCTTTTG

ATTTATAAGGGATTTTGCCGATTTCGGCCTATTGGTTAAAAAATGAGCTGATTTAACAAAAA

TTTAACGCGAATTTTAACAAAATATTAACGTTTACAATTTCAGGTGGCACTTTTCGGGGAAA

TGTGCGCGGAACCCCTATTTGTTTATTTTTCTAAATACATTCAAATATGTATCCGCTCATGA

399

ATTAATTCTTAGAAAAACTCATCGAGCATCAAATGAAACTGCAATTTATTCATATCAGGATT

ATCAATACCATATTTTTGAAAAAGCCGTTTCTGTAATGAAGGAGAAAACTCACCGAGGCAGT

TCCATAGGATGGCAAGATCCTGGTATCGGTCTGCGATTCCGACTCGTCCAACATCAATACAA

CCTATTAATTTCCCCTCGTCAAAAATAAGGTTATCAAGTGAGAAATCACCATGAGTGACGAC

TGAATCCGGTGAGAATGGCAAAAGTTTATGCATTTCTTTCCAGACTTGTTCAACAGGCCAGC

CATTACGCTCGTCATCAAAATCACTCGCATCAACCAAACCGTTATTCATTCGTGATTGCGCC

TGAGCGAGACGAAATACGCGATCACTGTTAAAAGGACAATTACAAACAGGAATCGAATGCAA

CCGGCGCAGGAACACTGCCAGCGCATCAACAATATTTTCACCTGAATCAGGATATTCTTCTA

ATACCTGGAATGCTGTTTTGCCGGGGATCGCAGTGGTGAGTAACCATGCATCATCAGGAGTA

CGGATAAAATGCTTGATGGTCGGAAGAGGCATAAATTCCGTCAGCCAGTTTAGTCTGACCAT

CTCATCTGTAACATCATTGGCAACGCTACCTTTGCCATGTTTCAGAAACAACTCTGGCGCAT

CGGGCTTCCCATACAATCGATAGATTGTCGCACCTGATTGCCCGACATTATCGCGAGCCCAT

TTATACCCATATAAATCAGCATCCATGTTGGAATTTAATCGCGGCCTAGAGCAAGACGTTTC

CCGTTGAATATGGCTCATAACACCCCTTGTATTACTGTTTATGTAAGCAGACAGTTTTATTG

TTCATGACCAAAATCCCTTAACGTGAGTTTTCGTTCCACTGAGCGTCAGACCCCGTAGAAAA

GATCAAAGGATCTTCTTGAGATCCTTTTTTTCTGCGCGTAATCTGCTGCTTGCAAACAAAAA

AACCACCGCTACCAGCGGTGGTTTGTTTGCCGGATCAAGAGCTACCAACTCTTTTTCCGAAG

GTAACTGGCTTCAGCAGAGCGCAGATACCAAATACTGTCCTTCTAGTGTAGCCGTAGTTAGG

CCACCACTTCAAGAACTCTGTAGCACCGCCTACATACCTCGCTCTGCTAATCCTGTTACCAG

TGGCTGCTGCCAGTGGCGATAAGTCGTGTCTTACCGGGTTGGACTCAAGACGATAGTTACCG

GATAAGGCGCAGCGGTCGGGCTGAACGGGGGGTTCGTGCACACAGCCCAGCTTGGAGCGAAC

GACCTACACCGAACTGAGATACCTACAGCGTGAGCTATGAGAAAGCGCCACGCTTCCCGAAG

GGAGAAAGGCGGACAGGTATCCGGTAAGCGGCAGGGTCGGAACAGGAGAGCGCACGAGGGAG

CTTCCAGGGGGAAACGCCTGGTATCTTTATAGTCCTGTCGGGTTTCGCCACCTCTGACTTGA

GCGTCGATTTTTGTGATGCTCGTCAGGGGGGCGGAGCCTATGGAAAAACGCCAGCAACGCGG

CCTTTTTACGGTTCCTGGCCTTTTGCTGGCCTTTTGCTCACATGTTCTTTCCTGCGTTATCC

CCTGATTCTGTGGATAACCGTATTACCGCCTTTGAGTGAGCTGATACCGCTCGCCGCAGCCG

AACGACCGAGCGCAGCGAGTCAGTGAGCGAGGAAGCGGAAGAGCGCCTGATGCGGTATTTTC

TCCTTACGCATCTGTGCGGTATTTCACACCGCATATATGGTGCACTCTCAGTACAATCTGCT

CTGATGCCGCATAGTTAAGCCAGTATACACTCCGCTATCGCTACGTGACTGGGTCATGGCTG

CGCCCCGACACCCGCCAACACCCGCTGACGCGCCCTGACGGGCTTGTCTGCTCCCGGCATCC

GCTTACAGACAAGCTGTGACCGTCTCCGGGAGCTGCATGTGTCAGAGGTTTTCACCGTCATC

ACCGAAACGCGCGAGGCAGCTGCGGTAAAGCTCATCAGCGTGGTCGTGAAGCGATTCACAGA

TGTCTGCCTGTTCATCCGCGTCCAGCTCGTTGAGTTTCTCCAGAAGCGTTAATGTCTGGCTT

CTGATAAAGCGGGCCATGTTAAGGGCGGTTTTTTCCTGTTTGGTCACTGATGCCTCCGTGTA

AGGGGGATTTCTGTTCATGGGGGTAATGATACCGATGAAACGAGAGAGGATGCTCACGATAC

GGGTTACTGATGATGAACATGCCCGGTTACTGGAACGTTGTGAGGGTAAACAACTGGCGGTA

TGGATGCGGCGGGACCAGAGAAAAATCACTCAGGGTCAATGCCAGCGCTTCGTTAATACAGA

TGTAGGTGTTCCACAGGGTAGCCAGCAGCATCCTGCGATGCAGATCCGGAACATAATGGTGC

AGGGCGCTGACTTCCGCGTTTCCAGACTTTACGAAACACGGAAACCGAAGACCATTCATGTT

GTTGCTCAGGTCGCAGACGTTTTGCAGCAGCAGTCGCTTCACGTTCGCTCGCGTATCGGTGA

TTCATTCTGCTAACCAGTAAGGCAACCCCGCCAGCCTAGCCGGGTCCTCAACGACAGGAGCA

CGATCATGCTAGTCATGCCCCGCGCCCACCGGAAGGAGCTGACTGGGTTGAAGGCTCTCAAG

GGCATCGGTCGAGATCCCGGTGCCTAATGAGTGAGCTAACTTACATTAATTGCGTTGCGCTC

ACTGCCCGCTTTCCAGTCGGGAAACCTGTCGTGCCAGCTGCATTAATGAATCGGCCAACGCG

CGGGGAGAGGCGGTTTGCGTATTGGGCGCCAGGGTGGTTTTTCTTTTCACCAGTGAGACGGG

CAACAGCTGATTGCCCTTCACCGCCTGGCCCTGAGAGAGTTGCAGCAAGCGGTCCACGCTGG

TTTGCCCCAGCAGGCGAAAATCCTGTTTGATGGTGGTTAACGGCGGGATATAACATGAGCTG

400

TCTTCGGTATCGTCGTATCCCACTACCGAGATGTCCGCACCAACGCGCAGCCCGGACTCGGT

AATGGCGCGCATTGCGCCCAGCGCCATCTGATCGTTGGCAACCAGCATCGCAGTGGGAACGA

TGCCCTCATTCAGCATTTGCATGGTTTGTTGAAAACCGGACATGGCACTCCAGTCGCCTTCC

CGTTCCGCTATCGGCTGAATTTGATTGCGAGTGAGATATTTATGCCAGCCAGCCAGACGCAG

ACGCGCCGAGACAGAACTTAATGGGCCCGCTAACAGCGCGATTTGCTGGTGACCCAATGCGA

CCAGATGCTCCACGCCCAGTCGCGTACCGTCTTCATGGGAGAAAATAATACTGTTGATGGGT

GTCTGGTCAGAGACATCAAGAAATAACGCCGGAACATTAGTGCAGGCAGCTTCCACAGCAAT

GGCATCCTGGTCATCCAGCGGATAGTTAATGATCAGCCCACTGACGCGTTGCGCGAGAAGAT

TGTGCACCGCCGCTTTACAGGCTTCGACGCCGCTTCGTTCTACCATCGACACCACCACGCTG

GCACCCAGTTGATCGGCGCGAGATTTAATCGCCGCGACAATTTGCGACGGCGCGTGCAGGGC

CAGACTGGAGGTGGCAACGCCAATCAGCAACGACTGTTTGCCCGCCAGTTGTTGTGCCACGC

GGTTGGGAATGTAATTCAGCTCCGCCATCGCCGCTTCCACTTTTTCCCGCGTTTTCGCAGAA

ACGTGGCTGGCCTGGTTCACCACGCGGGAAACGGTCTGATAAGAGACACCGGCATACTCTGC

GACATCGTATAACGTTACTGGTTTCACATTCACCACCCTGAATTGACTCTCTTCCGGGCGCT

ATCATGCCATACCGCGAAAGGTTTTGCGCCATTCGATGGTGTCCGGGATCTCGACGCTCTCC

CTTATGCGACTCCTGCATTAGGAAGCAGCCCAGTAGTAGGTTGAGGCCGTTGAGCACCGCCG

CCGCAAGGAATGGTGCATGCAAGGAGATGGCGCCCAACAGTCCCCCGGCCACGGGGCCTGCC

ACCATACCCACGCCGAAACAAGCGCTCATGAGCCCGAAGTGGCGAGCCCGATCTTCCCCATC

GGTGATGTCGGCGATATAGGCGCCAGCAACCGCACCTGTGGCGCCGGTGATGCCGGCCACGA

TGCGTCCGGCGTAGAGGATCGAGATCGATCTCGATCCCGCGAAATTAATACGACTCACTATA

GGGGAATTGTGAGCGGATAACAATTCCCCTCTAGAAATAATTTTGTTTAACTTTAAGAAGGA

GATATA

Figure S2: Synthetic CYP124A1 (Rv2266) gene (codon optimised for E. coli). The CYP124A1 gene was cloned into plasmid vector pET47b using BamHI (GGATCC) and HindIII (AAGCTT) restriction sites. The CYP124A1 gene has an N-terminal His-tag with a HRV-3C cleavage site (yellow). BamHI (GGATCC) and HindIII (AAGCTT) restriction sites are underlined. The start is coloured in green while the stop codon is coloured red. Figure shows sequence for the entire plasmid.

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Figure S3: The CYP142A1 (Rv3518c) gene region in Mycobacterium tuberculosis. The CYP142A1 gene is located in a gene cluster involved in metabolism of a variety of structurally unrelated compounds, including steroids and fatty acids. The surrounding genes include PE-PGRS57 (Rv3514), a member of the Mtb PE (N-terminal Pro(P)-Glu(E) sequence) family whose functions remain largely unknown; echA19 (Rv3516), a potential enoyl-CoA hydratase; Rv3520c, a potential coenzyme F420-binding oxidoreductase; ltp4 (Rv3522), a putative lipid transfer protein; and fadD19 (Rv3515c) that is a predicted fatty acid-CoA-synthase. The genes Rv3517, Rv3519 and Rv3521, are conserved hypotheticals with unknown function. The image was generated using Tuberculist (http://www.tuberculist.epfl.ch).

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Figure S4: The CYP124A1 (Rv2266) gene region in Mycobacterium tuberculosis. The CYP124A1 gene is located among a gene cluster that also contains a menaquinone sulfotransferase (Sft3, Rv2267c) and the P450 CYP128A1 which is involved in the hydroxylation of menaquinone MK-9 at the omega-position, leading to its sulfation by the Rv2267c gene product. Other flanking genes include Rv2262c which encodes a conserved hypothetical protein possibly involved in lipid metabolism; and Rv2264c which encodes a conserved hypothetical Pro-rich protein that has a highly Pro-, Thr-rich C-terminus and which is predicted to be an outer membrane protein. Its function remains unknown. The lppN (Rv2270) gene is a predicted lipoprotein. Rv2263 is a putative NADP(H)-dependent oxidoreductase. Rv2265 encodes a likely conserved integral membrane protein. Rv2267c encodes a conserved hypothetical protein of unknown function, as does Rv2269c. The image was generated using tuberculist (http://www.tuberculist.epfl.ch).

Mycobacterium tuberculosis - Research Explorer

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