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Investigating the catalytic mechanism of the meta-cleavage product hydrolases Ruzzini, Antonio C. 2013

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 INVESTIGATING THE CATALYTIC MECHANISM OF THE meta-CLEAVAGE PRODUCT HYDROLASES  by Antonio C. Ruzzini  B.Sc. (Honours), Trent University, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Biochemistry and Molecular Biology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  September 2013  ? Antonio C. Ruzzini, 2013   ii ABSTRACT The meta-cleavage product (MCP) hydrolases are members of the ?/?-hydrolase superfamily that utilize a Ser-His-Asp triad to catalyze the hydrolysis of a C-C bond. The catalytic mechanism of the MCP hydrolases is poorly defined and particularly interesting due to a requisite substrate ketonization that precedes hydrolysis. To resolve the catalytic mechanism of the MCP hydrolases, two enzymes were studied: tetrameric BphDLB400 from Burkholderia xenovorans LB400 and dimeric DxnB2 from Sphingomonas witichii RW1. Both efficiently hydrolyze 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoic acid (HOPDA) to produce 2-hydroxypenta-2,4-dienoic acid (HPD) and benzoic acid. A series of experiments established that BphDLB400 uses an histidine-independent nucleophilic mechanism of catalysis and is half-site reactive. Benzoylation of Ser112 was demonstrated by LC ESI/MS/MS analyses and a pre-steady-state kinetic burst of HPD formation indicated the reactivity. While acylation during HOPDA hydrolysis by BphDLB400 occurred on a similar timescale for the WT and H265Q variant, esterase activity was abrogated in the histidine variant. Thus, alternative mechanisms of nucleophile activation are employed for C-C and C-O bond cleavage. A covalent mechanism of catalysis was inferred for DxnB2, however, the turnover of HOPDA was 1:1 with respect to enzyme concentration. A solvent kinetic isotope effect suggested that a proton transfer, and therefore, substrate ketonization determines the rate of acylation in the MCP hydrolases. Substrate ketonization, and therefore acylation, can be indirectly observed as consumption of ESred, an intermediate named for its bathochromically-shifted absorption spectrum. A proton transfer to ESred allowed the assignment of this species to an enzyme-bound HOPDA dianion. An extended Br?nsted analysis revealed a linear   iii correlation between substrate basicity and the rate constant determined for the ketonization reaction. Finally, the MCP hydrolase P-subsite, which contacts the MCP dienoate moiety, was definitively linked to substrate ketonization. In DxnB2 Asn43 and Arg180 variants, ESred formation was found to limit this proton transfer reaction. A substrate-assisted nucleophilic mechanism of catalysis has been proposed for the MCP hydrolases. Therein, the electron-rich dienoate moiety substitutes for the His-Asp pair as the general base for nucleophile activation. Overall, definition of the chemical mechanism of the MCP hydrolases has implications for environmental bioremediation strategies and the rational design of therapeutics.   iv PREFACE Several parts of this thesis have been published in or submitted to refereed journals; an asterisk has been used to indicate shared authorship. Two of these manuscripts are a result of a fruitful collaboration with the laboratory of Jeffrey T. Bolin at Purdue University. In each, the Bolin Lab was responsible for crystallization, X-ray diffraction experiments and the generation and refinement of three-dimensional models of BphDLB400 and DxnB2.  (1) Ruzzini AC*, Ghosh S*, Horsman GP, Foster LJ, Bolin JT, and Eltis LD. 2012. Identification of an acyl-enzyme intermediate in a meta-cleavage product hydrolase reveals the versatility of the catalytic triad. J. Am. Chem. Soc. 134, 4615. For this manuscript, I was responsible for the in vitro characterization of BphDLB400 WT and H265Q, including kinetic and chemical quench experiments, the preparation of samples and analysis of protein MS experiments, and the characterization reactions in H218O. (2) Ruzzini AC, Horsman GP, Eltis LD. 2012. The catalytic serine of meta-cleavage product hydrolases is activated differently for C-O bond cleavage than for C-C bond cleavage. Biochemistry 51, 5831. For this manuscript, I was responsible for the characterization of the rate-limiting step for MCP hydrolases catalysis, the BphDLB400 methanol-dependent esterase activity, the mechanism of nucleophile activation for C-O bond cleavage, and empirically determining the steady-state partitioning of methano- and hydrolysis reaction products.     v  (3) Ruzzini AC*, Bhowmik S*, Yam KC, Ghosh S, Bolin JT, and Eltis LD. 2013. The lid domain of the MCP hydrolase DxnB2 contributes to an increased specificity for recalcitrant PCB metabolites. Biochemistry 52, 5685. For this manuscript, I was responsible for the transient-state kinetic analysis of DxnB2-mediate turnover of recalcitrant 3-Cl HOPDAs, analysis of the enzyme?s oligomeric state and the comparison of seven MCP hydrolases, which lead to discovery of previously unrecognized determinants in enzyme oligomeric state stabilization as well as determinants in substrate specificity.   vi TABLE OF CONTENTS ABSTRACT ............................................................................................................................. ii PREFACE ............................................................................................................................... iv TABLE OF CONTENTS ...................................................................................................... vi LIST OF TABLES ................................................................................................................. ix LIST OF FIGURES ................................................................................................................ x ABBREVIATIONS ............................................................................................................... xii ACKNOWLEDGEMENTS ................................................................................................ xiii Chapter  1: INTRODUCTION .............................................................................................. 1 1.1 Aromatic Compounds .................................................................................................. 1 1.2 Catabolism of aromatic compounds by bacteria .......................................................... 1 1.2.1 The biphenyl (Bph) pathway .................................................................................... 4 1.2.2 The Bph pathway can cometabolize PCBs .............................................................. 5 1.2.3 BphD is a determinant of PCB degradation ............................................................. 7 1.2.4 The MCP hydrolases ................................................................................................ 8 1.2.5 The ?/?-hydrolase superfamily ................................................................................ 9 1.2.6 Structural aspects of the MCP hydrolases .............................................................. 13 1.2.7 Classification of the MCP hydrolases .................................................................... 14 1.3 Mechanistic aspects of MCP hydrolase catalysis ...................................................... 15 1.3.1 Hydrolysis of carbon-carbon and carbon-heteroatom bonds ................................. 16 1.3.2 Substrate ketonization ............................................................................................ 18 1.3.3 ESred: an unidentified catalytic intermediate .......................................................... 19 1.3.4 The importance of ESred and molecular basis of inhibition by PCB metabolites ... 21 1.3.5 The hydrolytic reaction .......................................................................................... 23 1.3.6 Half-site reactivity .................................................................................................. 26 1.4 Aim of this study ........................................................................................................ 27 Chapter  2: MATERIALS AND METHODS ..................................................................... 30 2.1 Chemicals ................................................................................................................... 30 2.1.1 Sourced reagents .................................................................................................... 30 2.1.2 Preparation of 8-F and 5,8-diF DHB ...................................................................... 30 2.1.3 Preparation of 13C-DHB ......................................................................................... 31 2.1.4 Preparation of HOPDAs ......................................................................................... 32 2.2 Mutagenesis, protein production and purification ..................................................... 32 2.2.1 Production and purification of BphDLB400.............................................................. 32 2.2.2 Mutagenesis of DxnB2 ........................................................................................... 32 2.2.3 Production and purification of DxnB2 ................................................................... 33 2.2.4 Protein Analysis ..................................................................................................... 35 2.2.5 Size Exclusion Chromatography Multi-angle Light Scattering (SEC-MALS). ..... 35 2.3 Steady-state kinetic experiments ................................................................................ 36 2.3.1 Standard spectrophotometric assay ........................................................................ 36 2.3.2 Enzyme-substrate complex formation .................................................................... 37 2.3.2.1 Determination of dissociation constants ............................................................. 37 2.4 Transient-state kinetic experiments ........................................................................... 37 2.4.1 Standard protocol for stopped-flow kinetic experiments ....................................... 37 2.4.2 Rapid-quench methods ........................................................................................... 38   vii 2.5 Mass spectrometry of enzyme samples ...................................................................... 39 2.6 Product analysis ......................................................................................................... 39 2.6.1 HPLC analysis of product ratios from nucleophile partitioning ............................ 39 2.6.2 Standard protocol for EI GC/MS analysis ............................................................. 40 2.6.2.1 BphDLB400-mediated hydrolysis in H218O ........................................................... 41 2.7 Characterization of ESred ............................................................................................ 41 2.7.1 NMR of DxnB2 S105A complexes ........................................................................ 41 2.7.2 Sensitivity of ESred to molecular oxygen ............................................................... 42 2.8 Structural comparison of MCP hydrolases ................................................................ 44 Chapter  3: RESULTS .......................................................................................................... 45 3.1 Mechanistic analysis of BphDLB400 ............................................................................ 45 3.1.1 A pre-steady-state kinetic burst of HPD formation from BphDLB400 ..................... 45 3.1.2 Single turnover of HOPDA by BphDLB400 His265Q ............................................. 47 3.1.3 Identification of an acyl-enzyme intermediate in variant and WT enzymes.......... 49 3.1.4 Stoichiometric incorporation of 18O into benzoic acid from reactions in H218O ... 51 3.1.5 Insight into BphDLB400 deacylation from nucleophile partitioning ........................ 55 3.1.6 Proposal of a substrate-assisted mechanism of nucleophile activation .................. 56 3.2 Characterization of BphDLB400 esterase activity .................................................... 57 3.2.1 Methanol-dependent esterase activity .................................................................... 57 3.2.2 Activation of the Serine nucleophile for C-O bond hydrolysis .............................. 59 3.2.3 Biphasic pre-steady-state formation of pNP .......................................................... 61 3.3 Mechanistic analysis of DxnB2 ................................................................................. 63 3.3.1 The oligomeric state of BphDLB400 and DxnB2 are different ................................. 63 3.3.2 Evidence that DxnB2 also utilizes a nucleophilic mechanism of catalysis ........... 64 3.3.2.1 A pre-steady-state kinetic burst of HPD formation ............................................. 64 3.3.2.2 Identification of an acyl-enzyme intermediate in DxnB2 by LC ESI/MS .......... 66 3.3.3 Single turnover of HOPDA by DxnB2 .................................................................. 68 3.3.4 Transient state solvent kinetic isotope effect (SKIE) for HOPDA hydrolysis ....... 70 3.3.5 An extended Br?nsted analysis of substrate ketonization ...................................... 71 3.3.6 Probing the electronic nature of ESred by NMR ..................................................... 76 3.3.7 Probing the reactivity of ESred with molecular oxygen .......................................... 78 3.4 Characterization of DxnB2 P-subsite and His255 variants ........................................ 80 3.4.1 Steady-state kinetic analysis of P-subsite and His255 variants ............................. 80 3.4.2 Investigating the ability of P-subsite and His255 variants to accumulate ESred .... 83 3.4.3 Kd Determinations .................................................................................................. 85 3.4.4 Single turnover of HOPDA and 9-Cl HOPDA by P-subsite variants .................... 87 3.4.5 Single turnover of HOPDA and 9-Cl HOPDA by the His255 variants ................. 91 3.5 DxnB2-mediated cleavage of 3-Cl HOPDAs ............................................................ 96 3.5.1 DxnB2:3-Cl HOPDAs complexes do not reveal substrate destabilization ............ 96 3.5.2 DxnB2 is able to access ESred faster than BphDLB400. ............................................ 97 3.5.3 Comparison of the tertiary and quaternary structure of MCP hydrolases .............. 99 Chapter  4: DISCUSSION.................................................................................................. 106 4.1 The chemical mechanism of the MCP hydrolases ................................................... 106 4.1.1 An updated substrate-assisted nucleophilic mechanism of catalysis ................... 107 4.1.2 Evidence for a nucleophilic mechanism............................................................... 109 4.1.2.1 Direct evidence for an acyl-enzyme intermediate ............................................. 110   viii 4.1.2.2 Evidence for an acyl-enzyme intermediate from isotope studies ...................... 112 4.1.2.3 Evidence for an acyl-enzyme from steady-state kinetic approaches................. 113 4.1.3 Evidence for rate-limiting hydrolysis ................................................................... 115 4.1.4 The role of the catalytic histidine ......................................................................... 116 4.1.4.1 The role of the catalytic histidine in the acylation reaction .............................. 117 4.1.4.2 The role of the catalytic histidine in ESred formation ........................................ 119 4.1.4.3 The role of the catalytic histidine in the deacylation reaction........................... 123 4.1.5 Pre-steady-state kinetic behaviour of wild type enzymes and half-site reactivity 123 4.2 Distinct kinetic features of DxnB2 wild type .......................................................... 124 4.2.1 Defining observable kinetic steps in DxnB2 ........................................................ 125 4.2.2 A correlation between substrate basicity and ketonization by DxnB2................. 126 4.2.2.1 Substrate-derived determinants of biphasic kinetics ......................................... 127 4.3 Insight into the identity of ESred ............................................................................... 128 4.3.1 Preliminary experimental investigation of ESred by NMR ................................... 130 4.3.2 EI GC/MS analysis of ESred decay products......................................................... 131 4.4 The role of P-subsite residues in catalysis ............................................................... 132 4.4.1 Kinetic behaviour of P-subsite variants ............................................................... 133 4.4.2 The role of the P-subsite in substrate destabilization and ketonization ............... 134 4.5 Structural determinants in MCP hydrolases ............................................................ 135 4.5.1 Differences in MCP hydrolase structures and activity. ........................................ 135 4.5.2 The DxnB2 lid domain and specificity for 3-Cl HOPDAs .................................. 136 4.6 Concluding Remarks ................................................................................................ 138 APPENDIX I: CHEMICAL PROPERTIES OF SUBSTRATES .................................. 149 APPENDIX II: MASS SPECTROMETRY RESULTS .................................................. 150 APPENDIX III: PRE- OR POST-CATALYTIC EVENTS ............................................ 153 APPENDIX IV: 1H13C HSQC SPECTRUM OF DxnB2 S105A:HOPDA COMPLEX 154 APPENDIX V: STEADY-STATE KINETICS AND EQUILIBRIUM BINDING ....... 155    ix LIST OF TABLES Table 1. List of primers used for the mutagenesis of DxnB2 ................................................ 33 Table 2. Pre-steady-state kinetic burst of HPD formation by BphDLB400 .............................. 47 Table 3. Kinetic data for wild-type BphDLB400 and its His265 variants ................................. 48 Table 4. Incorporation of 18O from H218O into benzoate and HOPDA.................................. 54 Table 5. The effect of alcohol on BphDLB400-mediated HOPDA hydrolysis ......................... 56 Table 6. Methanol-dependent pNPB and HOPDA cleavage ................................................. 59 Table 7. Single turnover kinetic data for pNPB turnover by BphDLB400 and variants ........... 60 Table 8. Pre-steady-state kinetic burst of pNB formation by BphDLB400 ............................... 63 Table 9. Pre-steady-state kinetic burst of HPD formation by DxnB2 .................................... 66 Table 10. Solvent viscosity effects on a single turnover of HOPDA by DxnB2 ................... 70 Table 11. Pre-steady-state SKIE on DxnB2 hydrolysis of HOPDA ...................................... 71 Table 12. Kinetic data for the DxnB2-mediated hydrolysis of substituted HOPDAs............ 72 Table 13. 19F chemical shifts for F-compounds and DxnB2 S105A:F HOPDA complexes. 78 Table 14. The hydrolytic activity of DxnB2 H255 variants ................................................... 81 Table 15. Steady-state kinetic parameters of DxnB2 P-subsite variants................................ 82 Table 16. Kd for DxnB2 S105A/R180M and S105A/R180Q:HOPDA complexes ................ 86 Table 17. Kinetic data for HOPDA and 9-Cl HOPDA turnover by P-subsite variants ......... 91 Table 18. Kinetic data for HOPDA and 9-Cl HOPDA turnover by DxnB2 H255 variants... 95 Table 19. Kinetic values for the DxnB2-mediated hydrolysis of 3-Cl HOPDAs .................. 99 Table 20. HOPDA pKa2 values and extinction coefficients ................................................. 149 Table 21. b-series ion fragment matches to a BphDLB400 peptide ........................................ 150 Table 22. Example of modeling EI GC/MS data to 18O incorporation events ..................... 151 Table 23. Relative abundance ion fragments analyzed by EI GC/MS ................................. 152 Table 24. Transition state stabilization by MCP hydrolase side chains ............................... 159   x LIST OF FIGURES Figure 1. Biologically relevant aromatic compounds. ............................................................. 1 Figure 2. Aerobic and anaerobic aromatic ring degradation. ................................................... 3 Figure 3. The upper Bph pathway. ........................................................................................... 5 Figure 4. The metabolic fate(s) of 3-Cl HOPDAs in soil bacteria. .......................................... 8 Figure 5. The MCP hydrolase reaction. ................................................................................... 9 Figure 6. Examples of ?/?-hydrolase-catalyzed reactions..................................................... 10 Figure 7. Distinct ?/?-hydrolase lids at the common ?6 insertion point. .............................. 11 Figure 8. The BphDLB400 active site. ...................................................................................... 14 Figure 9. Classification of MCP hydrolases. ......................................................................... 15 Figure 10. The mechanism of C-X bond hydrolysis by ?/?-hydrolases. ............................... 17 Figure 11. Representative examples of diketone-like intermediates. .................................... 17 Figure 12. Representative single turnover of HOPDA by BphDLB400. .................................. 19 Figure 13. BphDLB400 variant:HOPDA binary complexes. .................................................... 20 Figure 14. Surface representation of the 3-Cl HOPDA binding mode. ................................. 23 Figure 15. Proposed mechanisms for the MCP hydrolases. .................................................. 23 Figure 16. Mechanistic interpretation from studies of MhpC hydrolysis in H218O. .............. 24 Figure 17. Support for the general base mechanism from studies of BphDLB400 and HsaD. . 25 Figure 18. The two-conformation model of catalysis for BphDLB400. ................................... 27 Figure 19. Pre-steady-state burst of HPD formation by BphDLB400. ...................................... 46 Figure 20. LC ESI/MS analysis of BphDLB400 reacted with HOPDA. ................................... 50 Figure 21. ESI/MS/MS of a modified and an unmodified peptide from BphDLB400. ............ 51 Figure 22. Representative EI/GC/MS analysis of H218O experiments. .................................. 53 Figure 23. The proposed substrate-assisted mechanism of nucleophile activation. .............. 57 Figure 24. Representative single turnover of pNPB by BphDLB400 WT and H265Q. ........... 61 Figure 25. Proposed roles for His265 during the hydrolysis of C-C and C-O bonds. ........... 61 Figure 26. Pre-steady-state burst of pNP by BphDLB400. ....................................................... 63 Figure 27. SEC-MALS analysis of three MCP hydrolases. ................................................... 64 Figure 28. Pre-steady-state burst of HPD formation by DxnB2. ........................................... 65 Figure 29. LC ESI/MS analysis of DxnB2 reacted with HOPDA or 9-Cl HOPDA. ............. 68 Figure 30. Representative single turnover of HOPDA by DxnB2. ........................................ 69 Figure 31. Representative single turnover of HOPDA by DxnB2 in D2O. ........................... 71 Figure 32. Representative single turnover of substituted HOPDAs by DxnB2. .................... 74 Figure 33. Extended Br?nsted relationship of substrate basicity and 1/?2. ............................ 75 Figure 34. 19F NMR spectra of ES complexes. ...................................................................... 77 Figure 35. Mass spectrum of the O2-dependent ESred breakdown product. ........................... 80 Figure 36. DxnB2 S105A/P-subsite variant-substrate complexes ......................................... 84 Figure 37. Titrations of HOPDAs with DxnB2 S105A/R180M and R180Q variants. .......... 86 Figure 38. Representative single turnover of HOPDA by DxnB2 P-subsite variants. .......... 89 Figure 39. Representative single turnover of 9-Cl HOPDA by DxnB2 P-subsite variants ... 90 Figure 40. Representative single turnover of HOPDA by DxnB2 H255 variants. ................ 93 Figure 41. Representative single turnover of 9-Cl HOPDA by DxnB2 H255 variants. ........ 94 Figure 42. DxnB2 S105A:3-Cl HOPDA complexes. ............................................................ 97 Figure 43. Representative single turnover of 3-Cl HOPDAs by DxnB2. .............................. 98 Figure 44. Structure-guided sequence alignment of MCP hydrolases. ................................ 101   xi Figure 45. Structural features of the MCP hydrolase tetramers. .......................................... 102 Figure 46. Superposition of MCP hydrolase lid domains. ................................................... 103 Figure 47. Superposition of DxnB2 S105A:HOPDA complexes. ....................................... 105 Figure 48. An upodated proposal of the MCP hydrolase mechanism. ................................ 108 Figure 49. A Ser112-benzoyl species trapped in a BphDLB400 H265Q crystal. ................... 111 Figure 50. Comparison of BphDLB400 and DxnB2 active sites. ........................................... 118 Figure 51. Comparison of HOPDA binding modes. ............................................................ 121 Figure 52. BphDLB400 P-subsite variability upon substitution of His265. ............................ 122 Figure 53. Kinetic scheme for DxnB2 hydrolysis of HOPDA. ........................................... 126 Figure 54. The HOPDA enol:enolate equilibrium. .............................................................. 149 Figure 55. Visible spectra of post-reaction mixtures of DxnB2 and HOPDA. .................... 153 Figure 56. EI GC/MS demonstrating deuterium exchange into HOPDA. ........................... 153 Figure 57. 1H 13C HSQC spectrum of a DxnB2 S105A:[13C]-HOPDA binary complex.. .. 154 Figure 58. DxnB2 P-subsite steady-state kinetic curves for HOPDA hydrolysis. ............... 156 Figure 59. DxnB2 P-subsite steady-state kinetic curves for 9-Cl HOPDA hydrolysis........ 156 Figure 60. Titrations HOPDAswith DxnB2 N43D/S105A. ................................................. 158    xii ABBREVIATIONS 1/?? ? ? reciprocal relaxation time Abs   absorbance Amp   amplitude AU   absorbance units BPDO   biphenyl dioxygenase (EC 1.14.12.18) CoA   coenzyme A DHB   2,3-dihydroxybiphenyl EI   electron ionization ESI   electrospray ionization FAD   flavin adenine dinucleotide GSH/GSSG  reduced glutathione/oxidized glutathione HEPES  4-(2-hydroxyethyl)-1-piperazineethansulfonic acid HOPDA  2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoic acid HPD   2-hydroxypenta-2,4-dienoic acid HPLC   high performance liquid chromatography KPi   potassium phosphate LC   liquid chromatography MCP   meta-cleavage product MS   mass spectrometery NADH   ?-nicotinamide adenine dinucleotide NMR   nuclear magnetic resonance PCB   polychlorinated biphenyl PDA   photodiode array pNP   p-nitrophenol pNPB   p-nitrophenyl benzoate SDS-PAGE  sodium dodecyl sulphate polyacrylamide gel electrophoresis SEC-MALS  Size exclusion chromatography multiple angle light scattering SKIE   solvent kinetic isotope effect t1/2   half-life v0   initial reaction velocity vss   steady-state reaction velocity WT   wild type     xiii ACKNOWLEDGEMENTS  To begin, I would like to thank my supervisor Lindsay Eltis for accepting me into his laboratory and providing me with support and guidance throughout my graduate studies.  I must also acknowledge the past and present members of the Eltis lab. I have had the chance to interact with many enthusiastic researchers but I was most inspired working alongside Carola Dresen, Sachi Okamoto, Hiroshi Otani, Nicolas Seghezzi and Rahul Singh. Along with these individuals, Eugene Kuatsjah, Jon Penfield, Joseph Roberts, and many others in the LSI have been good friends in and outside of the lab. Finally, Jennifer Lian was of great assistance during the purification of a number of DxnB2 variants.  Next, I would like to thank my supervisory committee members, Lawrence McIntosh and Michael Murphy. Both have provided helpful suggestions, support and the opportunity to interact with and learn from their own lab members. Specifically, Mark Okon and Jason Grigg were responsible for training and technical assistance with NMR and X-ray crystallography, respectively. It was a great pleasure to work in close proximity to both of these groups as well as the Beatty, Mauk, Mohn and Davies labs. A special thanks is also required for all the helpful discussions and training provided Fred Rosell.  Many thanks to Jeffrey Bolin, Subhangi Ghosh and Shiva Bhowmik. The structural characterization of BphD and DxnB2 were invaluable for the interpretation of kinetic data and experimental design in the latter stages of my studies. To the Davies Lab coffee club, thank you for providing stimulating discussions, academic and beyond. Finally, I would like to thank my family and friends, especially my parents and above everyone else, Marta.    1 Chapter  1: INTRODUCTION 1.1 Aromatic Compounds  Aromatic compounds are widely distributed in nature, possessing diverse chemical structures and indispensible biological functions (Figure 1). For example, simple low molecular weight aromatic compounds are integral components of nucleic acids, proteins and enzyme cofactors. Moreover, the evolution of biosynthetic pathways for complex high molecular weight aromatic compounds reshaped our environment as approximately 30% of organic carbon in the biosphere is now stored as lignin[1]. Indeed, aromatic compounds are the second most abundant class of organic compounds in nature after the carbohydrates.  Figure 1. Biologically relevant aromatic compounds: phenylalanine, adenosine and lignin.  Aromaticity is defined by H?ckel?s rule: an aromatic compound is cyclic, planar, and possesses a conjugated p-orbital system that contains 4n + 2? electrons (where n is a positive integer). Delocalization of these ?-electrons acts to lower the energy of the conjugated system and thereby increase the inherent stability of the aromatic molecule. 1.2 Catabolism of aromatic compounds by bacteria  Bacterial catabolic pathways of aromatic-containing biomass contribute significantly to the global carbon cycle. Aerobic, anaerobic and ?hybrid? catabolic pathways occur in   2 nature and use a shared catabolic logic: activation of the aromatic ring followed by its cleavage[2,3]. For polyaromatic compound degradation, an iterative version of this strategy is employed. In aerobic catabolism, molecular oxygen (O2) serves as a cosubstrate for two separate reactions: O2 is utilized for the activation and cleavage of aromatic rings[4]. Several distinct enzymatic processes have been observed for both the activation and oxygenolysis of aromatic rings. Activation is accomplished by the enzymatic incorporation of oxygen, which typically involves dihydroxylation by a non-heme iron-dependent dioxygenase[5] or a single hydroxylation event by a flavin-dependent monooxygenase[6]. In either case, catecholic compounds that are formed by the hydroxylation reactions are eventually cleaved by either an intra- or extradiol dioxygenase (Figure 2A), which generate ortho and meta-cleavage products (MCPs), respectively[3,7]. The ring-cleavage products are then metabolized by other pathway-specific enzymes and are channeled into central metabolic pathways. The anaerobic catabolism of aromatic compounds has been less extensively studied than the aerobic catabolism[8]. A significant number of aromatic compounds serve as carbon sources in anoxic environments, including aniline, aromatic amino acids, benzoic acids, catechols, hydroquinones, nicotinate, pyrogallol, ?-, ?-, and ?-resorcylate and xylene[2]. Despite the plethora of growth substrates, anaerobic catabolism converges on a relatively smaller number of shared metabolites, as observed in aerobic catabolism. The best-studied pathways involve thioesterification, typically by a CoA ligase, for aromatic ring activation (Figure 2B). Alternatively, CoA-independent pathways exploit the susceptibility of multiply meta-substituted aromatics to an enol-to-keto tautomerization. The relatively few documented shared cyclic metabolites are then subjected to reductive dearomatization before   3 entering primary metabolic pathways as aliphatic acids. These latter reactions include a broad range of enzymatic reactions, many of which are poorly characterized. However, hydrolytic ring-opening is a prevalent strategy.  Figure 2. (A) Aerobic, dioxygenase-catalyzed aromatic ring cleavage. The O2-dependent reactions (solid arrows) and the products of each pathway are shown (dashed arrows indicate from 2 to 5 enzymatic reactions). (B) Shared metabolites that occur in anaerobic aromatic catabolism. Adapted from [2,3].    4 Finally, in addition to the discrete aerobic and anaerobic pathways, some bacteria utilize ?hybrid pathways? to catabolize aromatic compounds. In such pathways, O2 is used in only one of the two critical steps of ring activation and cleavage. A hybrid pathway was first identified and implicated in the degradation of phenylacetic acid in Pseudomonas putida U[9]. Subsequent studies on the benzoyl-CoA oxidizing (Box) pathway enzymes in Azoarcus evansii, BoxABC, demonstrated that benzoyl CoA assimilation proceeds through the formation and hydrolysis of a 2,3-epoxy-benzoyl-CoA rather than a catechol in these hybrid pathways[10]. 1.2.1 The biphenyl (Bph) pathway  The aerobic biphenyl catabolic pathway is a typical meta-cleavage pathway. The microbial capacity to mineralize biphenyl was first described in 1970 when Lunt and Evans demonstrated that it could be utilized as a sole growth substrate by pure cultures of Gram-negative bacteria[11]. This seminal report, reading less than 200 words, also presented three possible biphenyl metabolites: (i) 2,3-dihydroxybiphenyl, (ii) a yellow colored ?-hydroxy-?-phenylmuconic semialdehyde (?max 430 nm), and (iii) phenylpyruvate. Since the pathway?s discovery, all of the enzymatic activities associated with biphenyl utilization have been established and extensively reviewed[12]. While the genes associated with biphenyl degradation can be encoded within a single operon, the Bph pathway has been conceptually divided into upper and lower pathways. The upper Bph pathway generates 2-hydroxypenta-2,4-dienoic acid (HPD) and benzoic acid from biphenyl whereas the lower pathway metabolizes HPD to acetyl-CoA and pyruvate, which can enter the citric acid cycle. Historically, the upper and lower pathways have also been studied from distinct research perspectives. The ability of the upper Bph pathway to   5 transform polychlorinated biphenyls (PCBs; see section 1.2.2) was the impetus for the earliest genetic and biochemical studies[13,14]. In contrast, recent studies on lower pathway enzymes have focused on enzyme engineering for the production of enantiomerically pure antiobiotic precursors[15].  The upper Bph pathway comprises four enzymatic steps that result in the meta-cleavage of one of the biphenyl rings (Figure 3). The first step is catalyzed by biphenyl 2,3-dioxygenase (BPDO), a multicomponent enzyme that includes a two-subunit Rieske non-heme iron dioxygenase (BphAE), a ferredoxin (BphF), and an FAD-containing reductase (BphG)[13,16]. BPDO utilizes O2 and two electrons derived from NADH to transform biphenyl to cis-2,3-dihydro-DHB, which is then reduced to DHB by BphB at the expense of NAD+. The third enzyme, BphC is an Fe(II)-dependent extradiol dioxygenase that incorporates O2 into the catecholic ring of DHB, generating an MCP, 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoic acid (HOPDA). Finally, BphD, an MCP hydrolase, uses water to catalyze the fission of a carbon-carbon (C-C) bond to yield HPD and benzoic acid. Figure 3. The upper Bph pathway. 1.2.2 The Bph pathway can cometabolize PCBs  PCBs are aromatic xenobiotic compounds that had prominent industrial applications in the twentieth century[17]. The chlorination of biphenyl theoretically yields 209 possible congeners. However, commercial mixtures of PCBs typically included from twenty to sixty   6 congeners containing three to six chlorine substitutions. Links between PCB exposure and a multitude of human diseases including cancers, developmental and neurological disorders (reviewed in [18]) ultimately led to bans on their use and production that began in the 1970s. Nevertheless, it is estimated that approximately 10% of all PCBs manufactured, amounting to ~2.8 ? 108 kilograms, are still present in mobile environmental reservoirs[17] and exposure remains a health risk[19].  In general, studies of bacterial PCB degradation have revealed the process to be strain-dependent and limited by the degree and position of chlorination on each PCB congener[14]. Therefore, the degradation patterns observed in bacteria directly reflect the respective substrate specificities of enzymes encoded by the corresponding bph genes. Among the most potent PCB-degrading bacteria, Burkholderia xenovorans LB400 is remarkable not only for its ability to transform certain hexachlorinated congeners[20], but also due to the recruitment of a reductive dehalogenase to its bph operon[21].  The molecular characterization of PCB-degradation in bacteria has focused almost exclusively on the upper Bph pathway?s dioxygenases. As the initiating enzyme, BPDO congener specificity has been thoroughly investigated in a number of homologues[20,22,23], and the reported substrate ranges are often used to classify the corresponding bacterium[24]. Recently, metagenomic analyses of PCB-contaminated soils revealed extensive diversity in the toluene/biphenyl dioxygenases[25], and phylogenetic analyses demonstrated that BPDO could be subgrouped based on sequence similarity to at least two well-characterized enzymes from potent PCB-degrading soil bacteria[26]. More specifically, class I was correlated to bphA from B. xenovorans LB400 and class II to Pseudomonas pseudoalcaligenes KF707. A possible third class was similar to BPDO from Rhodococcus jostii RHA1, although biphenyl   7 may not be the cognate substrate of this enzyme considering that it shares ~40% identity to an alkylbenzene dioxygenase from Rhodococcus strain sp. DK17[27]. Moreover, this strain cannot use biphenyl as a growth substrate[28] and encodes for a ring-cleaving dioxygenase that is identical to a gene annotated as ethylbenzene dioxygenase in R. jostii RHA1[29]. Attempts to increase the range of congeners cometabolized by the Bph pathway have focused on expanding the substrate specificity of BPDO using directed evolution[30-34]. In addition, laboratory evolution of 1,2-dihydroxynapthalene dioxygenase, DoxG, a BphC homologue yielded an enzyme with increased specificity for 3,4-dihydroxybiphenyls, which are often produced during PCB degradation[35]. In spite of the naturally encoded genetic diversity present throughout soil-dwelling bacteria and ongoing in vitro efforts, a number of highly recalcitrant and ?dead-end? metabolites still represent significant bottlenecks to both naturally occurring and potential anthropogenic bioremediation strategies. 1.2.3 BphD is a determinant of PCB degradation The reaction catalyzed by BphD, the final enzyme of the upper Bph pathway, is a common bottleneck during PCB degradation by bacteria. At least three observations indicate that BphD activity can determine the extent and rate PCB cometabolism. First, yellow-coloured metabolites, specifically 3-Cl HOPDAs, accumulated during bacterial growth on 2,4?-dichlorobiphenyls[14]. Second, this accumulation was strain-independent[36]. Finally, 3-Cl HOPDAs inhibit BphD[37,38]. Steady-state kinetic characterization of several BphD homologues revealed that phenyl-substituents have a relatively modest effect on turnover whereas dienoate-substituents, particularly at C3, have pronounced detrimental effects on catalysis[37-39]. In fact, the inability of Bph pathway hydrolases to metabolize recalcitrant HOPDAs is reflected in the recruitment of a glutathione S-transferase, BphK, which   8 catalyzes a reductive dehalogenation of 3 and 5-Cl HOPDAs at the expense of two molecules of glutathione (GSH) in a subset of bacteria (Figure 4). Nevertheless, many powerful PCB-degraders, such as R. jostii RHA1 cannot produce GSH, underlining the importance of understanding the catalytic limitations of BphD. Along these lines, DxnB2, an MCP hydrolase from Sphingomonas witichii RW1, was recently distinguished for its reactivity towards recalcitrant 3-Cl HOPDAs[39]. In comparison to BphD from B. xenovorans LB400 (BphDLB400) and BphD from Rhodococcus globerulus P6 (BphDP6), DxnB2 was 13- and 83-fold more specific (kcat/KM) for the model PCB metabolite 3-Cl HOPDA or 3.4- and > 2600-fold more specific for the biologically relevant 3,9,11 tri-Cl HOPDA.  Figure 4. The metabolic fate(s) of 3-Cl HOPDAs in soil bacteria. 1.2.4 The MCP hydrolases  MCP hydrolases such as BphD (EC 3.7.1.8) belong to the ?/?-hydrolase superfamily. As outlined above, MCPs are characterized by a 2,6-dioxo-2,4-dienoic acid moiety that results from the extradiol cleavage of catechols (Figure 5). Mirroring the trend in the study of aromatic compound degradation, most identified MCP hydrolases occur in soil-dwelling bacteria. One exception is HsaD, which occurs in a steroid catabolic pathway found in a range of bacteria including the intracellular pathogen Mycobacterium tuberculosis[40,41].   9 BphDLB400 along with 2-hydroxy-6-oxo-nona-2,4-diene-1,9-dioic acid 5,6- hydrolase, MhpC from the phenylproprionate degradation pathway of Escherichia coli W3110, are considered the archetypical MCP hydrolases following a large body of work on these enzymes. Figure 5. The MCP hydrolase reaction. 1.2.5 The ?/?-hydrolase superfamily  The catalytic versatility of the ?/?-hydrolase fold was first highlighted by a structural comparison of five hydrolytic enzymes acting on different substrates[42]. In fact, the ?/?-hydrolases were among the earliest reported examples of divergent enzyme evolution, which constitutes the basis for the classification of enzymes into superfamilies. Since the discovery of the ?/?-hydrolase superfamily, the known chemical reactions have expanded from hydrolyses and acyl transfers involving carbon-heteroatom (C-X) bonds to a number of more interesting reactions including C-C bond cleavage and the cofactor-less activation of O2 (Figure 6).  The ?/?-hydrolase fold consists of an eight-stranded, mostly parallel, ?-sheet that is characterized by a left-handed superhelical twist, and is surrounded by ?-helices[43]. Typically, catalysis by this superfamily originates from a triad of residues that includes a nucleophile, an invariant histidine, and the conjugate base of an acidic amino acid. Reactions are further supported by an ?oxyanion hole,? and inspection of the active site reveals a mirror image relationship between the ?/?-hydrolases and the serine proteases[42]. Interestingly, the ?-helical content of the ?/?-hydrolases is variable. Only the ?C-helix, which is part of a   10 structurally conserved strand-turn-helix motif that carries the catalytic nucleophile is conserved. The nucleophilic residue, an Asp, Cys or Ser, is embedded within a Small-X1-Nu-  Figure 6. Examples of ?/?-hydrolase-catalyzed reactions. The catalytic triad and the proposed catalytic mechanism for each are listed. Adapted from [44].     11 X2-Small consensus sequence located within this structural motif that includes ?5 and ?C, where the ?Small? residues are typically glycines and Xn indicates any amino acid. The nucleophile resides at the tip of this sharp turn called the ?nucleophile elbow?[42], which also provides geometric constraints for the surrounding residues. Specifically, the main chain amide of X2 is positioned to form one half of the ?oxyanion hole.? The invariant catalytic His is carried on a loop, referred to as the Histidine-loop, extending from ?8 and positioning the residue near the conjugate base of an acidic residue (Asp or Glu) that occurs primarily after ?7. Notably, an ?/?-hydrolase with an atypical Ser-His dyad has also been observed[45].  The ability of the ?/?-hydrolase fold to tolerate a large number of distinct insertions has been credited for the wide range of chemistry originating from a relative well-conserved active site[43,44]. Insertions of varying size typically occur at ?3, ?4, ?6, ?7 or ?8. Among the insertions, a flexible cap or lid domain after ?6 is preeminent (Figure 6). For example, the epoxide and MCP hydrolases share similar lid domains yet sequence identity and seemingly subtle structural differences effectively modulate active site geometry and act as Figure 7. Distinct ?/?-hydrolase lids at the common ?6 insertion point. (A) Epoxide hydrolase from Agrobacterium radiobacter AD1 (PDB ID: 1EHY, chain A). (B) BphDLB400 (PDB ID: 2OG1, chain A). (C) Candida antartica lipase B (PDB ID: 1LBS, chain A). The ?/?-hydrolase core is coloured white, the lid domains are distinctly coloured, and the NC-loop (disordered in 1EHY) is coloured yellow. A CB  12 determinants of substrate specificity. Finally, the identity of loops extending from ?/?-hydrolase core to the different lid domains have also been implicated in catalysis[46,47]. The NC-loop, which extends from ?6 connecting the core and N-terminus of the lid domain, is a common example. In keeping with the conserved catalytic machinery, a nucleophilic mechanism of catalysis has been assigned to most ?/?-hydrolases (Figure 6). Early experiments probing catalysis in the ?/?-hydrolases superfamily employed radiolabeled substrates. Initially, acetylation of pancreatic lipase[48,49] and a carbethoxylated cutinase enzyme[50] was demonstrated using highly reactive p-nitrophenyl [3H]-acetate and p-nitrophenyl [1-14C]-acetate, respectively. Later, evidence for covalent catalysis during the turnover of cognate substrates, [3H]-acetylcholine and [3H]-myristoyl-CoA was provided for acetylcholine esterase[51] and myristoyl-ACP thioesterase[52]. Most recently, [14C]-acetylCoA-labeling and mechanism-based inhibition have defined a nucleophilic mechanism for homoserine transacetylases[53,54], and a crystal structure has been reported for an acetlyated deacetylcephalosporin C acetyltransferase[55]. Perhaps the best-characterized catalytic cycle, however, is that of haloalkane dehalogenase for which crystallographic snapshots are available that define a putative reaction coordinate, and include a covalent intermediate[56]. Other ?/?-hydrolase families with inferred nucleophilic mechanisms include C- and N-terminal peptidases[57,58], dienelactone hydrolase[59], the human fatty acid synthase thioesterase domain[60] and enzymes that can moonlight as perhydrolases[61]. Indeed, despite the wealth of indirect evidence for covalent catalysis, direct experimental observation of an acyl-enzyme has only been reported for relatively few members of the ?/?-hydrolase superfamily.   13 In contrast, a general base mechanism of catalysis has been proposed for at least three families of ?/?-hydrolases. General base mechanisms have been proposed for each of two different hydroxynitrile lyases possessing opposite enantioselectivity: paralleled by distinct proton transfer relay systems[62]. More particularly, the proton from the incipient hydroxyl is abstracted by the histidine and transferred to the cyanide leaving group in the R-selective enzyme whereas abstraction and transfer are catalyzed by the serine and histidine, respectively, in the S-selective hydroxynitrile lyases. The catalytic histidine has also been ascribed a role as a general base for the deprotonation of hydroxyl-containing substrates in the cofactor-less dixoygenases[63]. Finally, a general base mechanism for the activation of water rather than the catalytic Serine has been forwarded for the MCP hydrolases[64], as discussed in more detail in section 1.4.4. 1.2.6 Structural aspects of the MCP hydrolases  The MCP hydrolases possess an ?-helical lid domain (residues 146-212, BphDLB400 numbering) inserted between ?6 and ?? of the ?/?-hydrolase core (residues 1-145 and 213-286). The active site occurs at the interface of the two domains, and additional MCP hydrolase-specific residues are borne by the lid domain. The active site can be divided into two distinct subsites separated by the catalytic triad (Ser112-His265-Asp237) and ?oxyanion hole? residues (Gly42 and Met113; Figure 8A). First, the polar (P-) subsite is formed by five strictly conserved residues that contact the substrate?s dienoate moiety: Gly43, Asn51, Asn111, Arg190 and Trp266 (BphDLB400 numbering, Figure 8B). Substitution of each of the last three of these residues in MhpC or Arg190 in BphD lead to their implication in substrate binding and catalysis based on the reduction in the observed steady-state kinetic parameters[65]. The second subsite, named the non-polar (NP-) subsite, houses hydrophobic   14 residues and has been implicated in binding the variable MCP moiety[65,66]. The identities of the residues that form the NP-subsite are not strictly conserved, consistent with their role in determining substrate specificity of distinct enzymes. In BphDLB400, Ile153, Leu156, Phe175, Trp216 and Phe239 are involved in binding the substrate?s phenyl-moiety.  Figure 8. (A) The BphDLB400 active site. The P- and NP-subsite residues are coloured green and blue, respectively. The catalytic triad and ?oxyanion hole? residues are shown in white with the catalytic serine substituted for alanine. (B) The conserved MCP hydrolase P-subsite. The conserved enzyme-substrate interactions are illustrated using the BphDLB400 S112A:HOPDA binary complex (PDB ID: 2PUH). Dashed lines indicated possible H-bonding (distances ? 3.4 ?).  1.2.7 Classification of the MCP hydrolases   Phylogenetic analysis based on amino acid sequences represents a powerful starting point for annotation in the post-genomic era. Such analyses can be particularly informative when functional, sequence and structural data are combined. Accordingly, maximum likelihood analyses of a structure-guided sequence alignment revealed at least three subfamilies of MCP hydrolases (named class I to III; Figure 9, [39]). Overall, this classification echoes the known substrate specificities and oligomeric states of the enzymes included in the analysis. Although the substrate specificity reported for class I enzymes is quite broad, all of these enzymes appear to be tetramers. BphDLB400 has been reported as a A BR190W266N51N111G43W216F175I153 L156L213F239G42M113S112AH265D237  15 tetramer in solution[67], while a tetrahedral tetrameric assembly was either inferred or directly observed from the crystal structures of HsaD[68] or MhpC[69]. In contrast, both class II and III enzymes have been reported as dimers[70,71]. The tight clustering of class II enzymes reflects their preference for alkylbenzene-derived MCPs[67,70,72] whereas the preference of class III enzymes is less well-defined but includes carbazole[71]. The grouping of DxnB2 with the class III MCP hydrolases is consistent with the ability of this enzyme to catalyze the hydrolysis of biphenyl metabolites, specifically 9- and 10-substituted HOPDAs, in vitro[39]. However, the cognate substrate of DxnB2 is unknown. Figure 9. Dendrogram of the best tree obtained from the alignment of 27 MCP hydrolases. The distinct classes and a scale representing the distant of divergence are shown. Numbers at branchpoints represent bootstrap values. Taken from [39].  1.3 Mechanistic aspects of MCP hydrolase catalysis  The MCP hydrolases are interesting due to their purported ability to catalyze both an enol-to-keto substrate tautomerization and hydrolysis of a C-C bond in the absence of organic and inorganic cofactors.    16 1.3.1 Hydrolysis of carbon-carbon and carbon-heteroatom bonds The distinct chemical nature of C-X (where X = N, O or S) and C-C bonds has a profound effect on enzyme-catalyzed hydrolysis in Ser-His-Asp triad containing enzymes. Common obstacles to C-X and C-C bond hydrolysis include activation of the scissile bond and a water molecule, which are performed by a general acid and general base, respectively[44,73]. Covalent catalysis is typically employed for these reactions in the ?/?-hydrolases (Figure 10). For bond activation, the role of the general acid is fulfilled by the ?oxyanion hole,? which stabilizes two tetrahedral oxyanion intermediates that result from nucleophilic attack at a carbonyl center by the catalytic serine or water, respectively. The catalytic His-Asp pair acts as a general base to facilitate the activation of the serine nucleophile for acylation and water for deacylation. The identity of the atom in the leaving group also contributes greatly to the chemical and kinetic mechanism of bond cleavage. The ability of the catalytic histidine to act as a general acid to protonate the leaving group during the collapse of the first tetrahedral oxyanion intermediate facilitates bond cleavage. In contrast to C-X bond cleavage, a leaving carbon atom is typically non-basic, obviating the aforementioned role of the histidine as an acid and necessitating an alternative electron sink for C-C than for C-X bond cleavage. MCP hydrolases and at least two other enzyme families that lack a metal cofactor exploit a diketone or diketone-like functionality for C-C bond hydrolysis (Figure 11). First, the serine nucleophile-dependent and independent ?-ketolases transform substrates that contain a ?-diketone electron sink[74]. A second group of C-C bond hydrolases defined by kynureninase is cofactor-dependent, generating a ?-ketiminium moiety by using pyridoxal 5?-phosphate (PLP)[75]. MCP hydrolases are thought to catalyze a substrate enol-to-keto   17 tautomerization prior to C-C bond cleavage[76]. Common to all these enzymatic reactions is the generation of an electron sink that is capable of stabilizing a leaving carbanion by charge delocalization. In contrast, a mechanistic proposals for orotidine 5?-monophosphate decarboxylase invoke protonation of during[77] or after[78] C-C bond fission. Figure 10. The mechanism of C-X bond hydrolysis in Ser-His-Asp containing ?/?-hydrolases. The first (Td1) and second (Td2) tetrahedral oxyanion intermediates and acyl-enzyme are drawn. Taken from [79].   Figure 11. Representative examples of proposed diketone- or diketone-like intermediates during C-C bond hydrolysis by (A) ?-ketolases, (B) kynuerninase and (C) the MCP hydrolases.     18 1.3.2 Substrate ketonization  The mechanism of substrate activation by the MCP hydrolases is poorly understood. The generation of an ?,?-unsaturated ?-diketone (Figure 11C) is dependent on protonation of the dienoate moiety at C5. Evidence for an enol-to-keto tautomerization was initially derived from product analyses of reactions performed in 2H2O, demonstrating that MhpC catalyzed the stereospecific incorporation of deuterium into the H-5E position of HPD[80]. This stereospecific deuterium incorporation was confirmed in studies of the BphDLB400-mediated hydrolysis of HOPDA[81]. Finally, the ability of BphDLB400 to catalyze the tautomerization of HPD to (E)-2-oxo-3-pentenoate was also indicative of this prerequisite electron-sink generating enol-to-keto tautomerase activity[66,81].  Although the mechanism is not understood, three residues of the strictly conserved P-subsite have been implicated in substrate binding and/or activation[65]. Notably, the conserved arginine (MhpC Arg188 and BphDLB400 Arg190) was ascribed a role in substrate destabilization. Furthermore, a hypothetical role for the catalytic histidine in mediating a proton transfer from an incoming MCP enol, specifically from the C2-hydroxyl to C5, has been forwarded[66,81]. This mechanistic proposal was based largely on two observations. First, the proximity of His265 to the C2-oxo group of the substrate or product in BphDLB400 S112A:HOPDA or S112C:HPD binary complexes. Second, BphD H265A and BphD S112A/H265A did not accumulate an intermediate with a bathochromically-shifted spectrum, which is thought to represent a destabilized form of the substrate at the BphDLB400 active site (see next section). In spite of the proposal, the protonation state of the substrate as it is bound is not well established. For HOPDA, the pKa for the enol:enolate equilibrium (pKa2) is close to physiological pH (see Appendix I for the known chemical properties of MCPs). Moreover,   19 the visible spectrum of the red-shifted intermediate is also more consistent with an enolate or dianionic species (?max 434 nm in solution) rather than an enol form of the substrate (?max 345 nm) arguing against a simple enol-to-keto tautomerization of substrate. 1.3.3 ESred: an unidentified catalytic intermediate  Single turnover kinetic analysis of the BphDLB400-catalyzed hydrolysis of HOPDA by stopped-flow revealed the existence of an ES complex with a bathochromically-shifted absorption spectrum (?max 473 and 492 nm; Figure 12A). The intermediate was named ESred, and its disappearance was mirrored by the production of HPD (Figure 12B), which can be monitored at 270 nm[81]. Subsequent studies revealed that similar red-shifted species with extended lifetimes accumulated in catalytic serine-to-alanine variants of BphDLB400 and HsaD, implying that the active site causes substrate destabilization[41,66]. Figure 12. Representative stopped-flow experiment illustrating a single turnover of 4 ?M HOPDA by 8.1 ?M BphDLB400 at 3.2 ?C. (A) Changes to the visible spectrum revealing ESred. (B) Data showing that ESred decay (492 nm) is commensurate with HPD production (270 nm). An additional post-catalytic process is visible at 270 nm, HPD ketonization.  Crystallographic characterization of ES complexes in BphDLB400 variants with catalytic triad substitutions exposed significantly different HOPDA binding conformations   20 (Figure 13). Specifically, restrained refinement of HOPDA to the electron density in a 1.6 ? resolution structure of BphDLB400 S112A revealed a non-planar binding mode that included a gauche+ torsion angle around the C4-C5 bond[66]. Single crystal absorption spectra were used to correlate the crystallographic data to the aqueous form of ESred, and therefore, the binding mode was considered to represent a mimic of the catalytic intermediate. The large deviation of the torsion angle around the C4-C5 bond from planarity, resulted in the HOPDA molecule being modeled as a (E)-2,6-dioxo-6-phenylhexa-3-enoate. Similar deviations from planarity were also observed in crystalline HsaD S114A:MCP complexes[68]. In contrast to the S112A variants, the binary BphDLB400 S112A/H265A:HOPDA complex revealed a binding mode in which C1 through C6 of the dienoate moiety were coplanar. This result was consistent with the lack of spectral perturbation observed in solution and the assignment of the free substrate in acetone in the trans-transoid configuration[37,82]. Figure 13. Ball-and-stick diagrams of the BphDLB400 variant:HOPDA binary complexes highlighting the differences in binding conformation and P-subsite interactions. (A) BphDLB400 S112A:HOPDA (PDB ID: 2PUH) and (B) BphDLB400 S112A/H265A:HOPDA (PDB ID: 2PUJ). Polar interactions (? 3.4 ?) are indicated using dashed lines.  The crystalline models imply that the catalytic serine variants are capable of substrate activation: generation of the 2,6-diketo intermediate. Thus, the structural data are apparently consistent with the observation that BphDLB400 S112A but not H265A is capable of R190N51W266H265N111S112AG43R190N51W266H265AN111S112AG43A B  21 catalyzing the ketonization of HPD, albeit at approximately 18% of the WT activity[66]. However, the models are contradictory to Woodward?s rules, which dictate that the ?,?-unsaturated ?-diketone should possess a hypsochromically-shifted spectra relative to the HOPDA enolate dianion. Moreover, ESred formation occurs at essentially an equal rate in BphDLB400 WT and S112A[66]. Accordingly, double bond strain has been suggested as a source of the spectral perturbation[83]. In this proposal, a reduction, but not complete disruption, of the p-orbital overlap would raise the ground state energy of the substrate thereby lowering the energy of the visible electronic transition. A second hypothesis that is concurrent with the crystalline models, suggests that polarization of ?-electrons in the ketonized substrate could also account for the spectral change. Polarization of similar ?/?-unsaturated systems in liver alcohol dehydrogenase and enoyl-CoA hydratase explained a 60 and 90 nm red-shift, respectively[84,85]. Despite these proposals, the identity of ESred remains unknown in the MCP hydrolases. 1.3.4 The importance of ESred and molecular basis of inhibition by PCB metabolites The mechanism of inhibition by 3-Cl HOPDA has been studied extensively in BphDLB400 using both kinetic and biophysical approaches. Steady-state kinetic studies revealed that 3-Cl HOPDA is a competitive inhibitor of HOPDA hydrolysis (Kic = 0.57 ?M), and that it is ultimately hydrolyzed 470 to 730 times slower than HOPDA[37,86]. The crystal structures of the catalytically inactive BphDLB400 S112A variant in complex with HOPDA and 3-Cl HOPDA also revealed major differences in the substrate-binding modes[86]. Instead of the non-planar conformation observed for HOPDA, the 3-Cl HOPDA molecule was bound in a planar conformation that was reminiscent but distinct from the planar binding mode of HOPDA to the S112A/H265A variant. For 3-Cl HOPDA, stabilization of a planar   22 binding mode was attributed to specific interactions between the chlorine substituent and the side chains of Leu156, Phe175, Phe239 and Met171. These residues form a hydrophobic pocket that accommodates the chlorine substituent (Figure 14). Additionally, unlike all other MCP hydrolase:MCP structures, the carboxylate of 3-Cl HOPDA does not interact with Arg190. Further comparison of available structural data implied that conformational changes would be required to alleviate the observed steric inhibition and reach the productive non-planar binding mode observed for HOPDA. Much like the BphDLB400 H265A variants, ESred was not observed during the hydrolysis of 3-Cl HOPDA and a similar species did not accumulate in complex with BphDLB400 S112A in solution[86]. Steady-state kinetic analyses showed that BphDLB400 preferentially hydrolyzed a series of 3-substituted HOPDAs in the order H > F > Cl > CH3. Stopped-flow studies monitoring the single turnover of 3,10-diF HOPDA by BphDLB400 revealed a small population of ESred, reinforcing the importance of this intermediate to catalysis. Taken together, the structural and kinetic data suggest that large C3-substituents impede formation of ESred, which can ultimately become rate-limiting to catalysis in the case of 3-Cl HOPDA hydrolysis by BphDLB400.   23 Figure 14. Surface representation of the 3-Cl HOPDA binding mode to BphDLB400 S112A. The hydrophobic pocket that accommodates the chlorine substituent is shown (PDB ID: 2RHT).  1.3.5 The hydrolytic reaction  The bulk of the mechanistic studies on the MCP hydrolases have attempted to define the mechanism of the hydrolytic reaction. Despite the wealth of data spanning two decades, no experimental evidence has been reported that satisfactorily delineates the catalytic mechanism of the MCP hydrolases. In fact, both a general base and a nucleophilic mechanism have been proposed (Figure 15), with the former having gained the greatest favour in the literature[87]. Figure 15. Proposed mechanisms for the MCP hydrolases. M171L156 G43F239F175G43M171L156F175F239rotate x 90o  24 Early work on the MCP hydrolase XylF from Pseudomonase putida, which cleaves 2-hydroxymuconic semialdehyde, revealed a chemical inactivation profile resembling that of a typical serine hydrolases[72]. In spite of these experiments, there have been several failed attempts to trap an acyl-enzyme intermediate in MhpC, stimulating support for a general base mechanism of catalysis in the MCP hydrolases[64,83]. Two observations have propelled support for the general base mechanism[64]: (i) a second 18O incorporation event into 3-6% of the succinate product of MhpC-catalyzed hydrolysis of 2-hydroxy-6-oxo-nona-2,4-dienoic acid in H218O, and (ii) an enzyme-catalyzed isotope exchange at the carbonyl of a non-hydrolyzable substrate analog. These data were interpreted as evidence for the reversible formation of a gem-diolate intermediate (Figure 16). Precedence for the purported gem-diolate was derived from studies on HIV-1 protease, an aspartic protease that utilizes a general base mechanism[88]. Figure 16. Mechanistic interpretation from studies of MhpC hydrolysis in H218O. The observation of a 13C NMR signal with a chemical shift that matched a calculated value for a gem-diolate species that was hydrogen bonded to a serine residue has been reported as an empirical demonstration of the intermediate and an alternative catalytic role for the catalytic serine[89]. More specifically, a broad signal at 128 ppm was detected during steady-state turnover of (1E:10S) of [13C6]-HOPDA by three variant enzymes, BphDLB400 S112A and H265A as well as by MhpC H114A (Figure 17A). These catalytically impaired BphDLB400 variants were used to ostensibly increase the likelihood of observing catalytic   25 intermediates. The MhpC H114A substitution affords the slow turnover of HOPDA by MhpC, a reaction that is not catalyzed by the WT enzyme. Structural characterization of crystalline HsaD S114A:MCP complexes were also used to strengthen the support for a general base mechanism in the MCP hydrolases. In particular, a conserved active site water molecule was designated as the nucleophilic water. The basis of this assignment were angles measured between the bound MCP carbonyls and the water molecule, which deviated from 10 to 20? from the B?rgi-Dunitz angle of attack on a carbonyl center[41]. Nevertheless, the alleged nucleophilic water was > 5 ? away from the catalytic histidine, and was at hydrogen bonding distances to an asparagine side chain and a ketone group on the substrate, ultimately raising questions about the possible mechanism of nucleophile activation. Figure 17. (A) 13C NMR spectra of the putative gem-diolate intermediate observed using either (i) BphDLB400 H265A, (ii) BphDLB400 S112A, (iii) MhpC H114A or (iv) the spectrum of the substrate free in solution. Taken from [89]. (B) Proposed catalytic water in the binary complex of HsaD S114A:4,9 DHSA (a cholesterol-derived metabolite, PDB ID: 2WUF).  Finally, more indirect evidence for a general base mechanism was founded on a Hammett analysis of the BphDLB400-catlayzed hydrolysis of para-substituted HOPDAs revealing a negative correlation, ? = ? 0.71, between electron withdrawing para-substituents 134 132 130 128 126 12413C (ppm)(i)(ii)(iii)(iv)A BN244catalytic waterH269N54R192G45L115S114A  26 and kcat[90,91]. This interpretation was largely based on the contrast between these results and the positive correlation observed in serine proteases[92]. Interestingly, Hammett analysis of BphDLB400 esterase activity using p-substituted p-nitrophenyl benzoates was also interpreted as evidence for a general base mechanism[93]. For the p-substituted pNPBs, the Hammet plot was non-linear, a feature that was attributed to a change in the rate-limiting step from ester bond cleavage to product release for substrates with ? ? 0. While this interpretation was ambiguous, the failure to observe a pre-steady-state kinetic burst of product formation from pNPB was cited to strengthen the proposal of a general base mechanism for the MCP hydrolase in vitro esterase activity. 1.3.6 Half-site reactivity  Half-site reactivity can be classified as an extreme case of negative cooperativity in which the maximal yield of product formation from a single enzymatic turnover amounts to only half of the apparently equivalent active sites[94]. Differences in substrate binding between protomers in higher order quaternary structures or structural non-equivalence attributed to oligomeric subunit organization are among the explanations for this phenomenon. In the MCP hydrolases, half-site reactivity was first proposed based on half-site occupancy of a crystalline MhpC:inhibitor complex[69]. Similar observations have been reported for two HsaD S114A:MCP complexes. Nevertheless, determination of dissociation constants to the S114A variant suggested stoichiometric binding in solution[41]. Unfortunately, crystallographic symmetry of all BphDLB400:MCP complexes requires equivalence across all active sites, limiting comparisons of active site occupancy as in MhpC and HsaD[66,81,86]. In BphDLB400, half-site reactivity could account for the biphasic decay of ESred during a single turnover of HOPDA[81]. A two-conformation model of catalysis was   27 initially proposed based on the mirrored kinetic data correlating ESred decay to product formation (Figure 18). In contrast to the paradigm, the relatively large amplitudes associated with both transient kinetic phases at 492 and 270 nm contradict the proposal of reversible (3-6%) gem-diolate formation. Instead, the two-conformation model suggests that the MCP hydrolases utilize a nucleophilic mechanism of catalysis in which deacylation at one site is linked to acylation at another. Figure 18. The two-conformation model of catalysis for BphDLB400.  1.4.5 Summary of mechanistic studies on the MCP hydrolases Overall, the balance of the data collected using BphDLB400, MhpC and HsaD has been used to challenge the accepted paradigm for serine hydrolases. A general base mechanism is not outside of the scope of the ?/?-hydrolase superfamily and is consistent with the mechanisms proposed for other C-C hydrolases that act on a diketone functionality. In contrast, the two- conformation model, the geometric arrangement of the ?nucleophile elbow?, and the comparative nucleophilic strength of alkoxides versus hydroxides[95,96] raise questions about the reported mechanism. Accordingly, the chemical mechanism of the MCP hydrolases has not been definitively established and mechanistic studies have been focused only on class I enzymes. 1.4 Aim of this study  Overall, the goal of this thesis was to investigate the kinetic and chemical mechanism of the MCP hydrolases. At the outset of these studies, a general base mechanism of catalysis   28 had been proposed as the paradigm for the MCP hydrolases[87]. Thus, the first goal of this work was to establish the hydrolytic mechanism of this enzyme family. Stopped-flow and chemical quench experiments followed by mass spectrometry were employed to directly detect catalytically relevant reaction intermediates. Both BphDLB400 WT and an H265Q variant were analyzed, and the crystal structures resulting from HOPDA soaking experiments in the H265Q and S112A/H265Q variants are included in the interpretation of the results. These experiments were complemented with indirect studies, which included product analyses of reactions in H218O by EI GC/MS and solvent partitioning experiments by HPLC.  The second objective of the study was to determine the mechanism of ester hydrolysis by the MCP hydrolases. Stopped-flow was utilized to perform pre-steady-state kinetic analysis of the BphDLB400 WT and variant-mediated hydrolysis of pNPB. Nucleophile partitioning experiments were also used to compare the C-O and C-C bond cleavage reactions.  The third goal of this thesis was to investigate the poorly understood mechanism of substrate ketonization. For these studies, DxnB2 from S. witichii RW1 was used based on the relatively simple kinetic behaviour observed during preliminary pre-steady-state characterization, namely stoichiometric turnover of HOPDA rather than the half-site reactivity observed for BphDLB400. Again, stopped-flow was employed to investigate the mechanism of ketonization by monitoring the single turnover of a series of substituted HOPDAs and performing an extended Br?nsted analysis thereof. Solvent viscosity and isotope effects were also measured in order to substantiate and provide additional evidence for the assignment of observable transient kinetic phases. 19F NMR spectra of free and bound   29 fluorinated substrates were also recorded in order to attempt to assign the electronic nature of ESred.  Next, the role of the catalytic histidine and two conserved P-subsite residues in substrate binding, destabilization and ketonization were assessed. DxnB2 Asn43, Arg180 and His256 variants were prepared and studied using steady- and transient-state kinetic approaches in order to assess their ability to accumulate ESred during the hydrolysis of HOPDA and 9-Cl HOPDA.   The final goal of this work was to investigate the reactivity of DxnB2 towards recalcitrant 3-Cl HOPDAs, which is higher than previously characterized BphD enzymes. Single turnover studies by stopped-flow were performed in order to assess the rate-limiting step for the turnover of 3-Cl HOPDA, a model PCB metabolite, and 3,9,11 tri-Cl HOPDA, a metabolite that can be found in nature. A structural comparison of DxnB2 and other MCP hydrolases was also conducted in order to identify potential determinants in catalysis. The crystal structures of the DxnB2 S105A: binary complexes are included in the discussion of these results.     30 Chapter  2: MATERIALS AND METHODS 2.1 Chemicals 2.1.1 Sourced reagents  Chlorinated DHBs, including 4-Cl-DHB, 6-Cl-DHB, 2?-Cl-DHB and 3?-Cl-DHB were gifts from Victor Snieckus (Department of Chemistry, Queen?s University, Kingston, ON). All other chemicals were of analytical grade and used without further purification unless stated otherwise. Preparations of BphAELB400, BphFLB400 and BphG from Pandoraea pnomenusa B-356 (BphGB356) were purified by Leticia G?mez-Gil according to published protocols[16,22,97]. An E. coli OverExpress C41(DE3) strain harbouring a pET14b vector carrying bphBB356 was provided for protein production by Michel Sylvestre (INRS ? Institut Armand Frappier, Point-Claire, QC). His6-BphB356 was purified as previously described[98] except that glycerol was omitted from the purification buffers. 2.1.2 Preparation of 8-F and 5,8-diF DHB  2?-F- and 2?,6-diF-DHB was generated from whole cell biotransformation of 2,2?-difluorobiphenyl using P. pnomenusa B-356 in the presence of 3-chlorocatechol. A 1 L culture of P. pnomenusa B-356 was grown on M9[99] + Goodies[100] supplemented with 100 mg biphenyl was grown until an OD600 nm of 0.5. These cells were then harvested and washed three times with phosphate buffer saline (PBS, pH 7.3) in an attempt to remove residual biphenyl. The harvested cells were divided into two flasks containing 500 mL PBS + 8 mg 2,2?-diF-biphenyl + 8 mg 3-chlorocatechol, an inhibitor of BphC. These flasks were then incubated at 30 ?C for 90 minutes, shaking at 200 rpm. The two samples were then pooled, acidified to pH 2.5, centrifuged and the supernatant was filtered to remove any insoluble 2,2?-difluorobiphenyl. The filtrate was extracted with three 200 mL volumes of   31 ethyl acetate, dried with MgSO4, and concentrated by rotary evaporation. Dry samples from each biotransformation were stored under nitrogen gas in an Mbraun Labmaster glovebox (Stratham, NH, USA) until further purification. A Waters 2695 HPLC system (Waters Corp., Milford, MA) was used for the purification of DHBs. The dry samples were dissolved in a solution of 0.5% formic acid (v/v), 50% methanol (v/v). Precipitate was removed from the resuspension by centrifugation and filtration (0.45 ?m).. The mixtures were injected on to a 250 x 10 mm Luna 5 ?m C18(2) column (Phenomenex, Torrence, CA), equilibrated at a 50:50 ratio of solvent A and B, and operating at a flow rate of 3.5 mL/min. Solvent A was 0.5% formic acid (v/v) and solvent B was methanol. Isocratic elution of 2?-F- and 2?,6-diF-DHB was achieved after 27.4 and 25.5 minutes, respectively. Pure fractions were collected, pooled, and partially dried under a stream of N2 gas to remove excess methanol. The identity of each DHB was confirmed by performing EI GC/MS. Each resulting fluorinated DHB solution, which typically amounted to 10 mL post-purification, was diluted to 200 mL and the pH of the solution was adjusted to 7.5 with 1 M NaOH prior to HOPDA preparation. 2.1.3 Preparation of 13C-DHB Uniformly labeled 13C-DHB was prepared from the enzymatic transformation of 13C-biphenyl (Sigma). A 40 mL reaction mixture containing 5 mg BphAELB400, 9 mg His6-BphFLB400, 10 mg BphGB356, 20 mg His6-BphB356, and 190 ?M NADH in KPi (I = 0.05 M), pH 7.5, was used to generate 13C-DHB. The reaction was complete in 1 hour at room temperature with constant stirring. The solution was then passed through a 0.45 ?m filter, acidified to pH 2.5 with HCl, extracted with three 20 mL volumes of ethyl acetate, dried with   32 MgSO4, and concentrated by rotary evaporation. The 13C-DHB, Rt = 32 min, was purified by HPLC, as described for the fluorinated DHBs. 2.1.4 Preparation of HOPDAs  HOPDAs were generated enzymatically using recombinantly expressed and purified BphC from B. xenovorans LB400 and the appropriate DHB as previously described[37]. Briefly, ~20 mg of DHB was solubilized in 1 mL of ethanol and added to a 500 mL volume of potassium phosphate buffer (I = 0.1 M), pH 7.5 containing 1 to 2 mg of BphC. For the fluorinated DHBs, 2 to 4 mg were transformed in a 200 mL volume. Quantitative conversion was monitored spectrophotometrically by following the increase in absorbance at the ?max specific to HOPDA enolate dianion that was being generated (see Appendix I). Upon complete conversion, the HOPDA solution was acidified to pH 2.5 using HCl and extracted with three 0.4 volumes of ethyl acetate. The organic extract was then dried over anhydrous MgSO4, concentrated by rotary evaporation, and ultimately dried using a stream of nitrogen gas. The purity of each sample was inspected by HPLC before use. The pKa of 5,8-diF HOPDA was determined to be 5.0 ? 0.2 by titration of an unbuffered solution, as previously described[37]. 2.2 Mutagenesis, protein production and purification 2.2.1 Production and purification of BphDLB400  The yield from the purification of recombinant BphDLB400 was improved two-fold by using E. coli Rosetta(DE3)pLysS cells (EMD4Biosciences) as a host strain at 30 ?C. Otherwise, BphDLB400 was purified as previously described[101]. 2.2.2 Mutagenesis of DxnB2 BphDLB400 and DxnB2 were mutated using a modified Quikchange oligonucleotide-  33 directed mutagenesis method (Stratagene, La Jolla, CA). Briefly, a single, 5?-phosphorylated mutagenic primer was used to amplify the plasmids encoding each gene. For BphDLB400, a broad host range plasmid, pSS184, carrying either bphD or bphD S112A was used as a template with the following primer, 5?-PO4-CTCCAAGTGCGGCCAGTGGGCGCAATGG, affording a His to Gln codon substitution. For DxnB2, the pEMBL18-based plasmids, pEMDXN1 carrying the wild type sequence or pEMDXN105A carrying a codon substitution for the S105A variant were used as templates. A complete list of mutagenic DxnB2 primers can be found in Table 1. After amplification, the PCR template was digested with DpnI, and the single stranded mutated amplicons were transformed and propagated in NovaBlue cells (EMD4Biosciences). The nucleotide sequences of plasmids carrying the mutated genes were confirmed using an ABI 373 Stretch DNA sequencer (Applied Biosystems, Foster City, CA) and a BigDye v3.1 Terminator kit. Table 1. List of primers used for the mutagenesis of DxnB2 Mutants  Primer sequence N43A GGCCGGTCCGCCTTCGCCGACAATTTTCCG N43D GGCCGGTCCGACTTCGCCGACAATTTTCCG N43H GGCCGGTCCCACTTCGCCGACAATTTTCCG S105A CCTGATCGGCAACGCCATGGGCGGGACG R180K GATATCGTCCATTACAAGCACGAGGCGTCGCTG R180M GATATCGTCCATTACATGCACGAGGCGTCGCTG R180Q GATATCGTCCATTACCAGCACGAGGCGTCGCTG H255A CCGAACTGCGGCGCTTGGGTGATGATCGAA H255E CCGAACTGCGGCGAATGGGTGATGATCGAA H255Q CCGAACTGCGGCCAGTGGGTGATGATCGAA  2.2.3 Production and purification of DxnB2  DxnB2 was produced in E. coli DH5?, as previously described, but purified using a modified version of the published protocol[39]. Buffers were made using water with a   34 conductivity of > 17 M? purified from a Barnstead NANOpure UV apparatus (Barnsted International, Dubuque, IA). Buffer A was 20 mM HEPES, pH 7.5 and Buffer B was 20 mM HEPES, 500 mM NaCl, pH 7.5. Typically, a cell pellet obtained from 2 litres of culture (~8 g, wet weight) was resuspended to a final volume of 40 mL in Buffer A supplemented with 1 mM CaCl2, 1 mM MgCl2, and 1 mg DNase I (Roche Diagnostics, Laval, QC). The cells were lysed by three successive passages through an Emulsi Flex-C5 homogenizer (Avestin Inc, Ottawa, ON) operating between 10 and 15 kPSI. Cell debris was removed by ultracentrifugation at 50000g for 45 min at 4 ?C. The resulting supernatant was passed through a 0.45 ?m filter before being loaded onto a 28 mL bed volume anion exchange Source 15Q column (GE Healthcare), which was equilibrated at 7% Buffer B and operating at 15 mL/min using an Akta Explorer or Purifier 100 (GE Healthcare). Proteins were eluted using a linear gradient from 7 to 40% B over 15 column volumes. Fractions containing DxnB2, which centered near 130 mM NaCl, were pooled and concentrated using an Amicon stirred cell concentrator fitted with an YM30 membrane (Millipore, Billerica, MA). The concentrated sample was then brought to 1 M (NH4)2SO4 at pH 7.5 and stirred for 12 hours at 4 ?C. The resulting precipitate was washed with three 25 mL volumes of Buffer A supplemented with 1 M (NH4)2SO4 before dissolution in 20 mM TAPS, pH 8.5. Purified preparations (> 99% homogeneity based on SDS-PAGE) were then exchanged into potassium phosphate buffer (KPi; I = 0.1 M), pH 7.5, concentrated to > 20 mg/mL, flash frozen in liquid nitrogen, and stored at -80 ?C until further use. The yield obtained from each purification was ~80 mg/L.  The purification scheme for DxnB2 S105A/H255E differed from the protocol outlined above since this variant was produced at lower levels and was recalcitrant to the   35 standard ammonium sulfate precipitation procedure. Instead, enriched DxnB2 S105A/H255E samples (post-Source 15Q) were brought to 1 M (NH4)2SO4, 20 M HEPES, pH 7.5, filtered, and loaded onto a 28 mL bed volume Source 15PHE column (GE Healthcare) for separation based primarily on hydrophobicity. The column was equilibrated with 1 M (NH4)2SO4, 20 M HEPES, pH 7.5 and was operated at 15 mL/min. Proteins were eluted using a gradient from 1 to 0 M (NH4)2SO4, in 20 mM HEPES, pH 7.5 over 30 column volumes. DxnB2 eluted near 60 mM (NH4)2SO4. The purified DxnB2 S105A/H255E enzyme was then buffer exchanged, concentrated, frozen and stored as described above. 2.2.4 Protein Analysis  The concentration of purified enzyme was routinely determined using spectrophotometric analysis of samples prior to all experimentation using the molar absorptivity of BphDLB400, ?280 nm = 55.4 mM-1cm-1[81], and DxnB2, ?280 nm = 36.2 mM-1cm-1. These values were calculated from the absorbance at 280 nm of samples whose concentrations had been determined by amino acid analysis at the Advanced Protein Technology Centre of the Hospital for Sick Children (Toronto, ON, Canada). It was assumed that single or double amino acid substitutions did not significantly perturb the absorptivity of the enzymes. 2.2.5 Size Exclusion Chromatography Multi-angle Light Scattering (SEC-MALS). SEC-MALS experiments were performed using an AKTA Purifier (GE Healthcare) fitted with Wyatt Technology Corporation SEC column and guard column (WTC-030S5 and WTC-030S5G). The protein concentrations used were between 1.5 and 2.5 mg/ml, prepared in 20 mM HEPES, pH 7.5. Multi-angle light scattering and refractive index was measured using the miniDAWN TREOS and Optilab T-rEX detectors (Wyatt Technology Corporation,   36 Santa Barbara, CA). Calibration was performed using bovine serum albumin. 2.3 Steady-state kinetic experiments 2.3.1 Standard spectrophotometric assay  A Cary60 or Cary5000 spectrophotometer equipped with a thermostatted cuvette holder (Agilent Technologies Canada Inc., Mississauga, ON) was utilized to monitor the generation or hydrolysis of HOPDAs. DHB ring-cleavage by BphC was followed by monitoring an increase in absorbance at the appropriate wavelength corresponding to the ?max of the HOPDA enolate chromophore (Appendix I). The hydro- or alcoholytic activity of BphDLB400 and DxnB2 were measured by monitoring the decay in the absorbance attributed to the HOPDA enolate chromophore. Alternatively esterase activity was monitored at 400 nm, following the appearance of para-nitrophenyl (pNP) from the cleavage of para-nitrophenyl benzoate (pNPB). Unless otherwise noted, kinetic measurements were performed in KPi (I = 0.1 M), pH 7.5, and the reaction systems were maintained at 25.0 ? 0.5 ?C. All reactions were initiated by the addition of a 10 ?L aliquot of enzyme prepared at an appropriate concentration. The reaction progress curves were deemed to be linear for at least 1 minute by fitting a series of 12 s intervals without the observation of significant (> 10%) decay in the measured initial rate. For higher concentrations of HOPDAs equal to or exceeding 50 ?M, a 2 mm pathlength cuvette was employed. Initial velocities (v0) were determined from the slopes of the linear progress curves using the regression tool within the Cary WinUV Kinetics application. The Michaelis-Menten equation or one accounting for substrate inhibition was fit to the data using the dynamic least squares fitting option in LEONORA[102].   37 2.3.2 Enzyme-substrate complex formation  Unless otherwise indicated, enzyme-substrate (ES) complexes were formed by mixing 40 ?M DxnB2 S105A with 10 ?M HOPDA in KPi (I = 0.1 M), pH 7.5 at 25.0 ? 0.5 ?C. Electronic absorption spectra of the substrates alone or in complex were recorded using the Scan application of a Cary5000 spectrophotometer. 2.3.2.1 Determination of dissociation constants  Dissociation constants for DxnB2 S105A P-subsite:substrate complexes were determined by measuring concentration dependent changes to the visible spectra of HOPDAs in KPi (I = 0.1 M), pH 7.5 at 25.0 ? 0.5 ?C. Specifically, 4 ?M HOPDA or 3 ?M 9-Cl HOPDA was titrated with a concentrated solution of enzyme. Titrations into 400 ?L of the substrate did not exceed 2% (v/v). The dissociation constants were then evaluated by fitting either a quadratic or hyperbolic binding equation to the data using the non-linear curve fitting were conducted in R[103]. 2.4 Transient-state kinetic experiments 2.4.1 Standard protocol for stopped-flow kinetic experiments  An SX.18MV stopped-flow reaction analyzer (Applied Photophysics Ltd., Leatherhead, UK) was used to perform transient-state kinetic measurements. The temperature of the optical cell and drive syringe chamber was maintained at 25 ?C using a circulating water. The concentration of reactants and specific buffer conditions are stated in the Results section as they depended on the aim of each individual experiment. Multiple wavelength data from 180 to 750 nm were collected using the system?s photodiode array (PDA) detector directly coupled to the Xe light source (the practical range was from 250 nm to 750 nm). The settings for the PDA detector were set manually as follows: integration period 1.0 ms, scan   38 period 0.921 ms, offset 0.1 V, gain 153. A monochromator (4.96 nm/mm bandpass open to 0.5 mm; ~2.5 nm bandpass) was employed to collect single wavelength data. Rate constants from stopped-flow experiments are reported using distinct nomenclature: (i) k when the process has been unambiguously assigned or (ii) 1/? (reciprocal relaxation time) when the assignment of processes were provisional or supported by this work alone. 2.4.2 Rapid-quench methods  Acyl-enzymes were prepared using the sequential mixing option of an SX.18MV stopped-flow reaction analyzer. The temperature of the drive syringe chamber and optical cell were held at 25 ?C by a circulating water bath. Samples analyzed by mass spectrometry originated from reactions of 4 ?M enzyme and 20 ?M HOPDA or 9-Cl HOPDA in KPi (I = 0.1 M), pH 7.5 before quenching with acetic acid (5% final v/v). The system was manually calibrated so that each quenched reaction volume totaled 400 ?L: 220 ?L of the reaction mixture was quenched with 180 ?L of 11.1% acetic acid (v/v) originating from a second set of stopped-flow syringes. Quenched samples, which originated from approximately 10-15 shots from the stopped-flow syringes (4 to 6 mL total volume), were collected on ice, measured to be at pH 2.3, and then concentrated 10-fold at 4 ?C. Samples were either flash frozen in liquid N2 and stored at -20 ?C until whole protein LC ESI/MS analysis or incubated with 1:50 pepsin (w/w) for 1 h at 37 ?C. The digests were halted by flash freezing with liquid N2, and the resulting peptide mixtures were stored at -20 ?C until further analysis. Samples from the reaction of BphDLB400 WT with HOPDA were allowed to age for 0.2, 1 and 10 seconds prior to quenching, while the H265Q variant reaction was quenched after 0.6 seconds. For DxnB2, the reactions with 9-Cl HOPDA were allowed to proceed for 1 second before quenching in the stopped-flow apparatus whereas the reaction with HOPDA   39 was quenched manually after 10 seconds. 2.5 Mass spectrometry of enzyme samples  Whole protein samples were analyzed using LC ESI-MS. Quenched reaction mixtures were loaded on a C18 precolumn (Dionex) and buffer salts were washed from the samples before elution in 80% acetonitrile, 0.1% formic acid (v/v). A Q-TOF hybrid instrument (QSTAR, Pulsar i, Applied Biosystems, Foster City, CA) was used to analyze the eluted samples. Estimates of the amount of modified BphDLB400 were based on comparing the observed peak areas calculated from Gaussian peak fitting function in Origin 8.0 (OriginLab Corporation, Northampton, MA).   Pepsin-digested peptides were analyzed by LC ESI MS/MS and identified as previously described[104], except that pepsin specificity was used to limit the MASCOT database search, benzoylated serine residues were allowed as a variable modification, and the fragments were searched against the BphDLB400 sequence. 2.6 Product analysis 2.6.1 HPLC analysis of product ratios from nucleophile partitioning Reactions were analyzed using a Waters 2695 HPLC system (Waters Corp., Milford, MA) equipped with a Hewlett-Packard ODS Hypersil C18 column (5 ?m pore size, 125 mm ? 4 mm) operating at a flow rate of 1 mL min-1 and equilibrated at 10% solvent B (methanol) in solvent A (0.5% H3PO4 v/v, in H2O). Reactions were quenched with H3PO4 (0.5%, v/v) and mixtures passed through a 0.45 ?m filter before injection. Product ratios or percentages of acid product were calculated by measuring the amount of benzoate produced in each reaction relative to controls performed without alcohol. Control reactions in potassium phosphate (I = 0.1 M, pH 7.5) with or without 0.2% acetone (v/v) were considered to be 100% hydrolytic,   40 and the amount of benzoate was quantified by integrating the eluted peak volume using Empower 3 software (Waters Corp.). All reactions analyzed using HPLC were performed in triplicate: the mean and standard deviation are reported. Alcoholysis reactions were performed in 400 ?L of potassium phosphate (I = 0.1 M) pH 7.5 at 25 ? C containing 40 ?M HOPDA and were initiated by adding BphDLB400 to a final concentration of 4 nM. The completeness of the reactions was judged spectrophotometrically before quenching. Samples were eluted using the following solvent gradients: (i) 10 to 30% B from 0 to 20 min, (ii) 30 to 60% B from 20 to 30 min, and (iii) 60 to 100% B from 30 to 40 min. Product elution times were as follows: 5.5 min for HPD, 13.7 for benzoate, 25.5 for methyl benzoate, 29.9 for ethyl benzoate, and 32.4 for propyl benzoate. The retention times of the alcoholytic products matched those of purchased standards.  Experiments that aimed to assess the methanol dependence of BphDLB400-mediated hydrolysis of HOPDA and pNPB were conducted via elution using similar gradients: (i) 10 to 30% B from 0 to 20 min, (ii) 30 to 60% B from 20 to 30 min, and (iii) 60 to 100% B from 30 to 32 min. Because of the relatively low solubility of the ester, reaction volumes were increased to 1 mL and mixtures contained 4 ?M pNPB. Moreover, 500 ?L was injected to compensate for the reduced substrate and/or product concentration. 2.6.2 Standard protocol for EI GC/MS analysis All samples analyzed by EI GC/MS were solubilized in pyridine and derivatized using an equal volume of BSTFA + TMCS (99:1) for at least 10 min. EI GC/MS was carried out using an HP 6890 series GC system fitted with an HP 5973 mass selective detector and an HP 5 ms column (30 m ? 250 ?m). The operating conditions were: TGC (injector) 280 ?C; TMS (ion source) 230 ?C; oven program time T0 min 80 ?C, T60 min 290 ?C, T0 min 80 ?C   41 (heating rate 5 ?C min-1). 2.6.2.1 BphDLB400-mediated hydrolysis in H218O For solvent exchange experiments, 20 ?M BphDLB400 was incubated with 200 ?M benzoate for 300 min or used to cleave 200 ?M HOPDA over 20 min in 50 ?L of KPi (I = 0.1 M), pH 7.5, in 92% H218O (v/v). Benzoate samples were prepared in acetone, applied to microfuge tubes, and allowed to evaporate to dryness before incubation in 18O-buffer with or without enzyme. Quantified samples of HOPDA in KPi (I = 0.1 M), pH 7.5, were lyophilized before resuspension in H218O with or without enzyme. In some instances, HOPDA was pre-incubated with the 18O-buffer for either 5 or 20 min prior to the addition of enzyme. The enzyme was prepared in natural abundance buffer. This buffer was also added to samples that did not contain enzyme in order to account for the dilution of H218O to 92% (v/v). After incubation (20 or 300 min), samples were acidified to pH 2.3 with acetic acid (5% final v/v) and extracted with 2 volumes (100 ?L) of ethyl acetate. The organic fraction was dried under vacuum. The average distribution of fragment ions observed in H2O was used to estimate the amount of 18O incorporation by fitting the data in H218O to two distinct models accounting for (i) a single 18O incorporation event, and (ii) low level of incorporation of a second 18O equivalent into benzoate. The root-mean-square error is reported for each fit. The relative abundance of fragment ions is reported as the percentage of a single species and the error between two replicated is reported as a standard deviation. 2.7 Characterization of ESred 2.7.1 NMR of DxnB2 S105A complexes NMR spectra were recorded at 25 ?C using either a Bruker Avance 500 or 600 MHz   42 spectrometer equipped with cryogenic probes. 19F chemical shifts were referenced to a 10% solution of trifluoroacetic acid (TFA). Binary complexes of 400 ?M DxnB2 S105A with either 200 ?M 8-F or 5,8-diF HOPDA were prepared in KPi (I = 0.1 M), pH 7.5. 19F NMR spectra of standards, including 200 ?M 8-F HOPDA, 200 ?M 5,8-diF HOPDA, and saturated solutions of 2-fluorophenylpropanone and 2-flourophenylprop-2-eneone in KPi (I = 0.1 M), pH 7.5, were also recorded. 1H 13C HSQC spectra were recorded for 13C-HOPDA and a DxnB2 S105A:13C-HOPDA binary complex (1 mM S105A to 500 ?M 13C-HOPDA) in KPi (I = 0.1 M), pH 7.5. For comparison, an HSQC spectrum of 5 mM unlabeled HOPDA in acetone-d6 was also recorded. The spectra were processed and analyzed using Bruker Topspin 3.0 software. 2.7.2 Sensitivity of ESred to molecular oxygen The half-life (t1/2) of the ESred species resulting from the DxnB2 S105A:HOPDA binary complex was measured spectrophotometrically at 25 ?C in KPi (I = 0.1 M), pH 7.5 at each of three concentrations of O2. First, a sample was prepared under ambient conditions, 280 ?M. Second, a solution was prepared at 650 ?M O2 by bubbling oxygen into a pre-formed solution of ESred. For the samples containing excess O2, the concentration dissolved into each system was measured using a Clark-type Oxygen electrode (Hansatech Instrument Ltd., Norfolk, UK). Finally, oxygen-free samples were prepared using degassed buffers that were equilibrated in an Mbraun Labmaster glovebox operating at < 5 ppm O2. Each 1 mL sample was sealed in a screw cap cuvette topped with a silicone rubber septum. Visible spectra were recorded at 5-20 minute intervals, and a t1/2 for each sample was determined by fitting a single exponential decay to the data using Origin 8.0. Product analysis of ESred samples were performed by comparing the DxnB2   43 S105A:9-Cl HOPDA decay products to those from the non-enzymatic process. EI GC/MS was performed as described for the 18O-exchange experiments.     44 2.8 Structural comparison of MCP hydrolases Superposition of MCP hydrolase structures was performed using the STAMP algorithm implemented in Multiseq, a plugin available in VMD[105-107]. Tools available in COOT[108] were also routinely used for superposition and manual inspection of the aligned proteins. The common names, biological sources and PDB codes of the aligned enzymes are: BphDLB400 from B. xenovorans LB400 (2OG1, chain A)[81], BphDRHA1 from Rhodococcus jostii RHA1 (1C4X)[109], CarC from Janthinobacterium sp. strain J3 (1J1I)[110], CumD from Pseudomonas fluorescens IP01 (1IUP)[70], DxnB2 from Sphingomonas witichii RW1 (coordinates provided by Shiva Bhowmik and Jeffrey T. Bolin, Purdue University, IN, USA), HsaD (2VF2, chain A)[68], and MhpC (1U2E, chain B)[69].     45 Chapter  3: RESULTS 3.1 Mechanistic analysis of BphDLB400  At the outset of these studies a general base mechanism of catalysis by the MCP hydrolases was prevailing in the literature[87]. In spite of the apparently defined reaction mechanism, continued efforts to more precisely define the chemical mechanism of the MCP hydrolases stimulated the generation of a BphDLB400 H265Q variant. In collaboration with the Bolin Lab at Purdue University, BphDLB400 H265Q was crystallized and incubated with HOPDA, yielding a covalent Ser112-benzoyl adduct. The observation of an acyl-enzyme in BphDLB400 H265Q (PDB ID: 3V1N) represented the first experimental demonstration of a possible reaction intermediate on the MCP hydrolase reaction pathway. These results contradicted previous attempts to trap or identify an acyl-enzyme intermediate during MCP hydrolase catalysis in solution[64,83], and prompted further study.  3.1.1 A pre-steady-state kinetic burst of HPD formation from BphDLB400  A previously proposed two-conformation model of catalysis predicts the existence of a pre-steady-state burst of HPD formation during the hydrolysis HOPDA[81], which is also in line with a covalent mechanism of catalysis in which kacylation > kdeacylation. Thus, to further investigate the chemical and kinetic mechanism of the MCP hydrolases, pre-steady-state kinetic studies of the turnover of HOPDA by BphDLB400 were performed using stopped-flow spectrophotometry. Multiple turnover reactions were performed using 1, 2 or 4 ?M BphDLB400 and 20 ?M HOPDA in KPi (I = 0.1 M), pH 7.5 at 25 ?C. The reactions were monitored at 270 nm and the resulting pre-steady-state kinetic burst (Figure 19) was ascribed to HPD formation, as previously described[111]. The values derived from fitting the spectrophotometric measurements with a burst equation are summarized in Table 2. The pre-  46 steady burst rate constant, kburst ~55 s-1, is in agreement with that observed, 1/?2 ~50 s-1, and provisionally assigned to acylation under single turnover conditions[81]. Interestingly, the magnitude of the burst accounted for only half of the enzyme used in each assay, consistent with the two-conformation model and the proposal that BphDLB400 and other MCP hydrolases are half-site reactive enzymes. Figure 19. Representative stopped-flow experiments demonstrating a pre-steady-state burst of HPD formation by BphDLB400 in KPi (I = 0.1 M), pH 7.5 at 25 ?C. Enzyme concentrations are indicated. The data has been normalized to account for the combined starting absorbance of the enzyme and substrate.  The measured steady-state rate constants, kss, were slightly slower than the previously determined kcat values of 4.2[37] or 6.5 s-1[81], and the burst equation could not account for data beyond 0.25 s at higher enzyme concentrations. This slight divergence from linearity in the kinetic traces can be explained by substrate depletion, which was ~20% after 0.3 s at 4 ?M enzyme. Concomitant product accumulation and inhibition (Kic HPD ~80 ?M, Kiu HPD ~120 ?M and Kic benzoate ~160 ?M[81]) as well as a combination of non-enzymatic and enzymatic ketonization of HPD, which is associated with a decay of the signal at 270 nm are likely to contribute to the apparent reduction in the observed kss. This is supported by the decrease in kss with increasing enzyme concentration. Thus, while the experiment allowed the   47 definition of a pre-steady-state burst of product formation, it did not afford an accurate determination of kcat. Table 2. Pre-steady-state kinetic parameters derived from monitoring HPD formationa [E] (?M) kburst (s-1) Ampburst (?A270nm) [P]burst (?M) vss (?A270nm s-1) kss (s-1) 1 52 ? 5 0.0098 ? 0.0007 0.51 ? 0.04 0.081 ? 0.007 4.2 ? 0.4 2 55 ? 5 0.0199 ? 0.0008 1.04 ? 0.04 0.150 ? 0.003 3.91 ? 0.08 4 62 ? 1 0.0376 ? 0.0007 1.96 ? 0.04 0.243 ? 0.004 3.16 ? 0.05 a?270 nm HPD = 19.7 mM-1cm-1[111] 3.1.2 Single turnover of HOPDA by BphDLB400 His265Q  The measured half-site reactivity helps to account for the biphasic HPD formation observed during the single turnover of HOPDA by BphDLB400 WT[81]. However, hydrolysis was readily observed in the H265A variant[66]. Moreover, the aforementioned crystalline acyl-enzyme was observed in the H265Q variant[112]. In contrast, mutation of the catalytic His in serine hydrolases results in the abrogation of enzyme activity[73]. His265 was also proposed to play a critical role in substrate tautomerization in BphDLB400[66,81]. Thus, to complement the crystallographic studies, single turnover experiments were performed to evaluate the kinetic behaviour of the H265Q variant and the ability of the S112A/H265Q variant to accumulate an electronically perturbed intermediate.  In BphDLB400 WT, ESred is rapidly formed, exceeding the detection limit of the stopped-flow instrument, 1/?1 > 500 s-1. Next, ESred decays in a biphasic manner, which is mirrored by an increase in absorbance at 270 nm, ascribed to HPD formation (Table 3). Under identical conditions, the reaction of 8.1 ?M H265Q and 4 ?M HOPDA in KPi (I = 0.1 M), pH 7.5 at 25 ?C, could also be described by three discrete transient kinetic phases. Instead of ESred formation, a small hyper- and hypsochromic shift to 430 nm was observed during a single catalytic cycle of H265Q, monitored using a PDA detector. The hyper- and 4   48 nm hypsochromically-shifted spectrum was consistent with the presence of a dianionic enolate substrate at the active site. This ES species then decayed in two phases. The first decay phase, 1/?2, proceeded at 10.3 s-1, approximately 5-fold slower than that of ESred decay in the WT and 8-fold faster than the enolate decay observed in the H265A variant. This phase was provisionally assigned to acylation based on monitoring HPD formation at 270 nm, for which a rate constant of 10.8 s-1 was determined. The second phase of enolate decay in H265Q, 1/?3 of 0.022 s-1 was similar to an observable relaxation at 270 nm, occurring at 0.08 s-1. Interestingly, in the His265 variants, the slowest processes observed under single turnover conditions do not match the kobs values measured in the presence of excess substrate, which in the case of H265Q is reduced by 106-fold (Table 3). Table 3. Kinetic data for wild-type BphDLB400 and its His265 variantsa Enzyme ? (nm) 1/?1 (s-1) [%Amp] 1/?2 (s-1) [%Amp] 1/?3 (s-1) [%Amp]a kcat (s-1)b WTc 270   50 ? 10 [89] 8.3 ? 0.7 [11] 6.5  492 ~500  54 ? 3 [64] 5.8 ? 0.9 [32]  H265Ad 434 78 [82] 1.3 [18]    0.0058 H265Q 270   10.8 ? 0.6 [28] 0.08 ? 0.02 [72]   434 130 ? 10 [16] 10.3 ? 0.1 [21] 0.022 ? 0.001 [64] 3 ? 10-5 aValues in square brackets indicate the % change of the total signal amplitude associated with each rate constant bkcat for H265Q was determined from the turnover of 50 ?M HOPDA with 2 or 4 ?M enzyme c,dData was taken from [81] and [66], respectively.  The assignment of a dianonic species at the active site of the H265Q variant was supported by the observation of triphasic ES complex formation between HOPDA and the S112A/H265Q variant (1/?1 = 114 ? 8 s-1, 1/?2 = 32 ? 2 s-1 and 1/?3 = 0.78 ? 0.02 s-1). The resulting stable S112A/H265Q:HOPDA complex was also hyperchromically shifted by ~125% (?434 nm = 32.1 ? 0.2 mM-1cm-1) and possessed a half-life of 31 hours. Curiously, the hyperchromic shift accounted for only half of that expected for the complete conversion to an all enolate sample (?434 nm = 40.1 mM-1cm-1). Incubation of HOPDA with a two-fold excess   49 of S112A/H265Q in Na-CHES (I = 0.1 M), pH 9.5 at 25 ?C, shifted the enolate band to 432 nm without a significant change in absorptivity. 3.1.3 Identification of an acyl-enzyme intermediate in variant and WT enzymes  The observation of pre-steady-state kinetic burst and HPD production from the single turnover of HOPDA by BphDLB400 H265Q with a measured rate constant comparable to the WT enzyme stimulated further investigation of catalytically relevant intermediates. Specifically, chemical quench experiments were performed during turnover of 20 ?M HOPDA by 4 ?M WT or the H265Q variant in KPi (I = 0.1 M), pH 7.5 at 25 ?C. The quench times were selected based on the observed kcat value for the WT or after 1/?2 for the H265Q variant. LC ESI/MS analyses of the acetic acid quenched reactions (5% v/v final) indicated that ~40% of the H265Q variant (Figure 20A) was benzoylated (M+104) after 600 ms while ~45% of the WT was benzoylated after 200 ms (Figure 20B). The signals for the unmodified WT and H265Q variant were observed at 31898 and 31889 m/z, respectively, which corresponded exactly to the theoretical values for BphDLB400 enzymes missing their N-terminal methionine. In WT-catalyzed reactions, the proportion of benzoylated enzyme dropped to ~30% after 1 s (Figure 20C), and no acylated enzyme was detected after 10 s (Figure 20D). The later obviated the possibility of a non-enzymatic acid-catalyzed process leading to the sub-stoichiometric acylation. Indeed, the observed decay of the M+104 species in WT BphDLB400 was consistent with the kss measured in kinetic burst experiments under similar conditions. Interestingly, the rapid acylation of BphDLB400 H265Q indicated that this process, and accordingly, nucleophile activation was His265-independent, and allowed for the assignment of 1/?2 and 1/?3 to acylation events.   50 Figure 20. Whole protein LC ESI/MS analysis of BphDLB400 reacted with HOPDA. Reactions contained 4 ?M enzyme and 20 ?M HOPDA in KPi (I = 0.1 M), pH 7.5 at 25 ?C. (A) An H265Q-catalyzed reaction was quenched after 600 ms. The WT-catalyzed reactions were quenched after (B) 200 ms, (C) 1 s and (D) 10 s. The additional peak observed at M+115 is consistent with a non-covalent WT:HPD adduct and the M+217 peak is consistent with either a non-covalent WT:HOPDA complex or a benzoylated enzyme:HPD complex.   Peptide matching from LC ESI/MS/MS of pepsin-digested reaction mixtures was used to umabiguously identify the site of enzyme modification. The benzoylation of Ser112 was confirmed from a hydrophobic peptide identified in samples from WT and H265Q reactions with HOPDA that were allowed to age 200 and 600 ms, respectively, before quenching with 5% acetic acid (v/v). The peptide spanned residues 102-120 (DIDRAHLVGNSMGGATALNF), containing several missed pepsin cleavage sites. Peptide fragment ions, matched in both the WT and H265Q samples, indicated modification of Ser112. In total, 15 ion fragments from the WT reaction mixture and 6 ion fragments from the H265Q sample were matched to b-series ions that could be explained only by   51 benzoylation at Ser112 (Figure 21, Table 23 in Appendix II). Both modified and unmodified peptides could be identified from each reaction mixture and their apparent stoichiometry was consistent with the amount of benzoylation observed in the intact enzyme samples. Figure 21. ESI/MS/MS of a modified and an unmodified peptide isolated from BphDLB400 WT. For simplicity, the high m/z range is shown to highlight the differences in large b-series ions.  3.1.4 Stoichiometric incorporation of 18O into benzoic acid from reactions in H218O  Identification of an acyl-enzyme intermediate during the catalytic cycle of BphDLB400 conflicted with reports on the stoichiometry of 18O incorporation into the succinic acid product of the MhpC-catalyzed hydrolysis of 2-hydroxy-6-oxo-nona-2,4-diene-1,9-dioic acid. Importantly, these experiments were the impetus for the proposed general base mechanism in the MCP hydrolases[64]. To test whether BphDLB400 behaves similarly, 200 ?M HOPDA was hydrolyzed using 20 ?M BphDLB400 in KPi (I = 0.1 M), pH 7.5 at 92% H218O (v/v) and the resulting benzoic acid product was analyzed using EI/GC/MS. The observed distribution of ion fragments reflected both the extent of 18O incorporation and the isotopic abundance of silicon in the derivatization agent. Therefore, the average distribution   52 of ion fragments observed in control reactions performed using H2O was used to calculate the extent of 18O incorporation into benzoic acid observed from the reactions performed in H218O (Figure 22). In these calculations, two models were fit to the observed fragment ion distributions: (i) a two-species model, representing a single 18O incorporation event, and (ii) a three species model, representing single and double incorporation event (Table 22, Appendix II). In turnover experiments performed using H218O, the two-species model best fit the data. The result for the most abundant fragment, summarized in Table 4, indicates 85 ? 1% incorporation of a single 18O equivalent. Similar results were observed in all other oxygen-containing fragments of benzoic acid (Table 23, Appendix II). This experiment provided no evidence for the enzymatic incorporation of a second 18O equivalent into the benzoic product: in no case were ions at M+6 and M+7 observed, and modeling incorporation of a second 18O equivalent into even a small percentage of the benzoate increased the error associated with the overall relative abundance of the singly exchanged species. For example, using the data presented in Table 4, modeling the incorporation of a second 18O equivalent into just 3% of the product increased the error associated with the overall relative abundance of the single exchanged species by 5-fold. Moreover, the enzyme did not catalyze any detectable solvent exchange into benzoic acid, even over 300 min (Table 4). However, non-enzymatic 18O exchange from the solvent into HOPDA was observed at 18% over 20 min (Table 4). The pre-incubation of HOPDA in 18O-buffer for up to 20 min did not increase the stoichiometry of incorporation observed into the benzoic acid product. This result indicated that exchange occurred exclusively at the C2-substituent (Figure 22), suggesting that HOPDA exists predominantly as a 2-oxo-6-oxido-3,5-dienoate in solution at pH 7.5. Overall, the results are consistent with the occurrence of an acyl-enzyme intermediate and argue   53 against the non-productive release of a tautomerized substrate from BphDLB400. Figure 22. Results from representative EI/GC/MS analysis of (A) benzoate in H2O, (B) benzoate derived from BphDLB400-mediated hydrolysis of HOPDA in H218O, (C) HOPDA in H2O, and (D) HOPDA in H218O. Inset panels (C and D) highlight the lower intensity HOPDA ion fragments. (E) Parent ions and predicted fragments.     54 Table 4. Incorporation of 18O from H218O into benzoate and HOPDAa   Relative abundance (%) of benzoate base peak ions (m/z) Sample Time (min) 179 [M] 180 [M+1] 181 [M+2] 182 [M+3] 183 [M+4] 184 [M+5] 18O incorporation Benzoate in H2O  83.2 12.9 3.4 0.44    WT + benzoate in H218O 300 83 12.3 4.3 0.4   ND WT + HOPDA in H218O 20 12 2.7 71 10.7 3.6 0.3 85 ? 1% WT + HOPDA in H218O 20 PIb 16.7 3.3 65 10 4 0.5 79 ? 2%    Relative abundance (%) of HOPDA base peak ions (m/z)   245 [M] 246 [M+1] 247 [M+2] 248 [M+3] 249 [M+4] 250 [M+5]  HOPDA in H2O 20 78 16 5 0.5    HOPDA in H218O 20 65 12.7 17.9 3.2 1.0 0.12 18 ? 1% aThe errors reported for 18O incorporation represent the root mean square deviation from fitting the experimentally observed data to a single 18O incorporation event bPI: HOPDA was pre-incubated in 18O-buffer for 20 min prior to initiating the reaction.    55 3.1.5 Insight into BphDLB400 deacylation from nucleophile partitioning  The methanol-dependent steady-state kinetic turnover of HOPDA by BphDLB400 was initially reported as indirect evidence for acylation. Partitioning of an acyl-enzyme with water or methanol thus produced benzoate or methyl-benzoate, and a simple kinetic model of nucleophilic catalysis accounted for the observed methanol-dependent increase in kcat and Km and the independence of kcat/Km[101]. Accordingly, a more extensive number of alcohols were employed in order to identify the rate-limiting step during the hydrolysis of HOPDA by BphDLB400. The effect on initial reaction velocities (v0) and the partitioning between water and short primary alcohols were assessed. It was rationalized that the effect of distinct alcohols on the steady-state rate of C-C bond cleavage could allow for the dissection of the chemical and physical events that have been correlated to kcat[81]. More specifically, these experiments exploit differences in the pKa of the small alkoxides from water to provide insight into the deacylation event. The effects of several alcohols on the v0 of BphDLB400-catalyzed turnover of 10 ?M HOPDA were measured. In the absence of alcohol, v0 was 25 ? 1 ?M s-1 (Table 5). Alcohol concentration-dependent increases in v0 were apparent upon the addition of either methanol, ethanol, or 1-propanol. The observed effects ranged from an approximate 3.2-fold increase in the presence of 100 mM methanol to a more modest 1.2-fold increase at the same concentration of 1-propanol. Inspection of the series indicates that the increase in v0 reflect the lower pKa of the substituting alkoxide. Importantly, v0 was not significantly affected by the addition of 25 or 100 mM 2-propanol, suggesting that the tested alcohols compete to replace the nucleophilic water and do not contribute any allosteric effects that accelerate the reaction. The possible inhibitory effects of each alcohol were not tested in light of the   56 observed rate accelerations. In addition to the observed kinetic phenomena, HPLC analysis of the reaction products was used to correlate these observed accelerations in v0 directly to alcohol utilization by BphDLB400 (Table 5).  Table 5. The effect of alcohol on BphDLB400-mediated HOPDA hydrolysisa  [Alcohol] (mM) v0 (?M s-1) Product ratio acid:ester  0 25 ? 1   methanol 25 45.8 ? 0.7 1.06 ? 0.01  100 76 ? 1 0.24  ? 0.01 ethanol 25 29.4 ? 0.9 3.78  ? 0.05  100 45.9 ? 0.7 0.95 ? 0.02 1-propanol 25 26.0 ? 0.6 42.8 ? 0.8  100 29 ? 1 3.57 ? 0.07 2-propanol 25 25.8 ? 0.6 ND  100 25.5 ? 0.3 ND aSpectrophotometric determination of v0 and HPLC determination of product ratios were performed at 10 ?M and 40 ?M HOPDA, respectively.  Partitioning between the hydrolytic and alcoholytic reaction pathways could be linked to common v0 measurements. For instance, using either 25 mM methanol or 100 mM ethanol, the acid:ester ratio was ~1 and v0 ~46 ?M s-1. Similarly, the same v0 was observed using 25 mM ethanol and 100 mM 1-propanol (~29 ?M s-1), and the acid:ester product ratio was ~3.6. Taken together, the results suggest that hydrolysis of the acyl-enzyme is the rate-limiting step in catalysis by BphDLB400. 3.1.6 Proposal of a substrate-assisted mechanism of nucleophile activation  The His265-independent acylation of BphDL400 argues for a novel mechanism of nucleophile activation in this Ser-His-Asp containing enzyme. Instead of using the canonical His-Asp dependent mechanism of serinate formation described for the serine hydrolases, a substrate-assisted mechanism of nucleophile activation (Figure 23) could account for the observed kinetic and chemical quench data. In the proposed mechanism, the highly-  57 conserved MCP hydrolase active site acts allows the dianionic substrate to act as a general base for the activation of the nucleophilic Ser. In turn, a proton transfer from the serine hydroxyl to C5 of the substrate completes the requisite substrate ketonization. The following sections were aimed to investigate this mechanism. Figure 23. The proposed substrate-assisted mechanism of nucleophile activation. In the first panel, no formal charge is drawn on the dieneoate moiety of the HOPDA dianion and a grey arrow indicates rotation about the C4-C5 bond. The second panel shows a formal charge at C5 that is rationalized by the pre-requisite proton transfer reaction that occurs at this position. In the third panel, only the transferred proton is drawn on C5 to highlight this reaction.  3.2 Characterization of BphDLB400 esterase activity 3.2.1 Methanol-dependent esterase activity The in vitro MCP hydrolase esterase activity is well-documented[93,113-115]. A common kcat value of ~6.5 s-1 was determined for the steady-state hydrolysis of HOPDA and pNPB by BphDLB400. In the presence of 25 mM methanol, equal partitioning of the benzoyl-containing products was also observed for both C-C and C-O bond cleavage reactions[101]. The commonalities between the cleavage of pNPB and HOPDA represent indirect evidence for a shared benzoyl-enzyme intermediate, challenging the proposed general base mechanism for C-O bond cleavage[93]. To further investigate the mechanism of C-O bond cleavage by the MCP hydrolases, the methanol-dependent turnover of pNPB by BphDLB400 was measured by monitoring pNP formation at 400 nm (Table 6). The reactions were conducted at 25 ?C in   58 KPi (I = 0.1 M), pH 7.5 supplemented with 0.2% acetone (v/v) to facilitate the solubilization of pNPB. Unfortunately, the relatively low solubility of pNPB precluded reliable determination of steady-state kinetic parameters in the presence of methanol. Despite this limitation, the rate of BphD-catalyzed C-O bond cleavage at 2 and 4 ?M pNPB provided some mechanistic insight. As was observed for the transformation of HOPDA[101], the rate of pNPB turnover increased with methanol concentration. However, the kobs value also decreased at methanol concentrations greater than 250 mM. This apparent decrease in activity likely reflects an increase in the Km pNPB similar to the previously observed methanol-dependent increase in Km HOPDA. At lower methanol concentrations, the concentration of pNPB is assumed to be saturating, and the kobs values for turnover were within error of the kcat values determined for the cleavage of HOPDA. The methanol-dependent product formation for the BphDLB400-mediated of 4 ?M pNPB cleavage was measured by HPLC and correlated well with the observed acceleration to the rates of bond cleavage. In order to compare nucleophile partitioning for C-O and C-C bond cleavage, methanol-dependent product formation from steady-state HOPDA turnover was also measured by HPLC. As for the pNPB reactions, 4 nM BphDLB400 was used to cleave 40 ?M HOPDA at varying concentrations of methanol. The partitioning ratio, reported as the percent of benzoic acid produced, was the same for the two reactions (Table 6). Furthermore, these studies helped to definitively link the observed kinetic phenomenon to a simple nucleophilic model of catalysis, which accurately predicted the partitioning ratios[101].     59 Table 6. Observed rates of methanol-dependent pNPB and HOPDA cleavagea,b [MeOH] 2 ?M pNPB 4 ?M pNPB HOPDA (mM) kobs (s-1) kobs (s-1) % acidc kcat (s-1)a Km (?M)a % acidc 0 6.4 ? 0.3 6.5 ? 0 .1 100 ? 1 6.5 ? 0.5 0.2 ? 0.05 100 ? 2 25 9.9 ? 0.5 10.7 ? 0.5 49 ? 1 9.4 ? 0.5 0.4 ? 0.1 48 ? 2 62 12.8 ? 0.2 14.6 ? 0.3 29 ? 1 13 0.6 27 ? 2 124 16 ? 1 18.4 ? 0.1 20 ? 1 16 0.83 17 ? 1 247 17 ? 2 21 ? 1 14.4 ? 0.7 21 ? 4 0.8 ? 0.2 20 ? 1 618 14 ? 1 20.1 ? 0.8 5.7 ? 0.3 21 ? 5 0.99 ? 0.07 4.9 ? 0.3 1240 11 ? 1 17.0 ? 0.4 3.7 ? 0.2 28 ? 4 2.0 ? 0.4 3.0 ? 0.2 aData taken from [101] bkobs = v0/[E]T c% benzoic acid observed relative to a hydrolytic reaction. 3.2.2 Activation of the Serine nucleophile for C-O bond hydrolysis  While the methanol-dependent steady-state kinetic data and HPLC analysis of the corresponding nucleophile partitioning suggest that both the C-O and C-C hydrolytic reactions proceed through an acyl-enzyme intermediate, they do not establish the mechanism of nucleophile activation. The ester substrate lacks the requisite electron rich system proposed to facilitate serinate formation in the substrate-assisted mechanism for C-C bond cleavage. Accordingly, ester bond hydrolysis was further probed using stopped-flow spectrophotometry to monitor the hydrolysis of pNPB by BphDLB400 WT and three catalytic triad variants: S112A, H265A and H265Q. The turnover of 2 ?M pNPB by 1 or 2 equivalents of WT enzyme resulted in the biphasic production of pNP (Table 7), reminiscent of HPD formation during HOPDA turnover[81]. However, even at equal reactant concentrations, the second phase of pNP production, k2 ~41 s-1, was approximately 7-fold faster than kcat, suggesting that an additional process, downstream of pNP release, is rate-determining but cannot be directly observed at 400 nm. Again, these results are consistent with the rate-limiting deacylation during BphDLB400-catalyzed C-O bond cleavage.     60 Table 7. Single turnover kinetic data for pNPB turnover by BphDLB400 and catalytic variantsa [E]:[pNPB] k1 (s-1) [%Amp] k2 (s-1) [%Amp] 1 WT 210 ? 40 [18] 41 ? 5 [82] 2 WT 170 ? 10 [38] 74 ? 2 [62] 4 WT 206 ? 5     4 H265Ab 0.06     4 H265Q 0.014 ? 0.002     4 S112A 0.00016 ? 0.00002     aThe error associated with each amplitudes was < 10% bvalue from a single experiment  Turnover of pNPB by a 4-fold excess of WT enzyme resulted in monophasic formation of pNP at 210 s-1. Subsequently, experiments with BphDLB400 variants were performed using a 4-fold excess of enzyme (8 ?M) to pNPB (2 ?M). Substitution of Ser112 with alanine resulted in the near abrogation of C-O cleavage with pNP production occurring on an hour timescale, 0.58 h-1. The rate constant for ester bond hydrolysis by the His265 variants were reduced by at least 3 orders of magnitude (Figure 24). Specifically, the H265A and H265Q variants cleaved pNPB at 0.06 and 0.014 s-1, respectively. As compared to the modest reduction to the rate constant for acylation in the H265Q variant when reacting with HOPDA, the magnitude of the reduction in esterase activity argues for canonical Ser-His-Asp triad chemistry for C-O bond cleavage (Figure 25).   61 Figure 24. Results from representative stopped-flow experiments demonstrating the single turnover of 2 ?M pNPB by 8 ?M BphDLB400 WT (black) and H265Q (grey) at 400 nm. The timescales for the WT and H265Q reactions are indicated on the bottom and top of the plot, respectively.    Figure 25. Proposed roles for His265 during the hydrolysis of C-C and C-O bonds by BphDLB400. The rate-limiting step (rls) is indicated for the WT enzyme.  3.2.3 Biphasic pre-steady-state formation of pNP To further investigate the biphasic product release observed during single turnover experiments, we monitored the turnover of 4-, 8-, and 20-fold excesses of pNPB by BphDLB400. In contrast to previous reports[93], the kinetic traces from multiple pNPB turnovers were biphasic (Figure 26) and could be modeled by either burst or double-exponential equations (Table 8). Fitting a burst equation to the data revealed that the rate   62 constants for pNP formation during the burst phase (kburst ~42 s-1) and the steady-state phase (kss ~8 s-1) were independent of enzyme concentration. These values were also consistent with k2 observed at a 1E:1S ratio and the kcat, respectively. The amount of pNP formed during the burst phase corresponded to ~70% of the enzyme present in the assay. Similarly, analysis using a double-exponential equation implied that ~60% of the enzyme was active during the pre-steady-state phase (k1 ~50-1). Values of k2 from the fit of a double-exponential equation to the multiple-turnover data ranged from 0.8 to 2.8 s-1, demonstrating a dependence on enzyme concentration. These values were also significantly lower than the experimentally determined kcat ~6.3 s-1. The magnitude of the pre-steady-state burst of pNP was sub-stoichiometric but differed significantly from the value of 50% predicted from the two-conformation model proposed for C-C bond hydrolysis. While quantification of the pre-steady-state burst was helpful in defining the mechanism of HOPDA hydrolysis by BphDLB400, the observation of 60-70% activity in the first catalytic cycle for ester bond hydrolysis most likely reflects the unusual reaction of a non-cognate enzyme-substrate pair. Nevertheless, the ?burst-like? kinetic behaviour further supports the observation of rate-limiting deacylation in BphDLB400.   63 Figure 26. Results from representative stopped-flow experiments demonstrating the pre-steady-state formation of pNP by BphDLB400 in KPi (I = 0.1 M), 0.2% acetone (v/v), pH 7.5 at 25 ?C. The hydrolysis of 2 ?M pNPB by 0.1 (light grey), 0.25 (dark grey) and 0.5 ?M BphDLB400 (black) was monitored.  Table 8. Pre-steady-state kinetic parameters derived from monitoring pNB formationa  Fits to a burst equation [E] (?M) kburst (s-1) Ampburst (?A400nm) [pNP]burst (?M) vss (?A400nm s-1) kss (s-1) 0.10 42 ? 3 0.00077 ? 0.00004 0.060 ? 0.003 0.0101 ? 0.0005 7.9 ? 0.4 0.25 41 ? 4 0.0022 ? 0.0001 0.17 ? 0.01 0.024 ? 0.001 7.4 ? 0.4 0.50 44 ? 5 0.0044 ? 0.0003 0.35 ? 0.02 0.046 ? 0.001 7.1 ? 0.2  Fits to a double exponential equation  k1 (s-1) [% Amp] [pNP]k1 (?M) k2 (s-1) [% Amp] [pNP]total (?M)b 0.10 44 ? 2 [5] 0.059 ? 0.004 0.8 ? 0.1 [90] 1.2 ? 0.2 0.25 47 ? 6 [8] 0.152 ? 0.007 1.4 ? 0.1 [92] 1.8 ? 0.1 0.50 60 ? 10 [12] 0.27 ? 0.02 2.8 ? 0.4 [88] 2.2 ? 0.3 aall errors associated with amplitudes were ? 12% b[pNP]total was calculated from ?A400 nm 3.3 Mechanistic analysis of DxnB2 3.3.1 The oligomeric state of BphDLB400 and DxnB2 are different  BphDLB400 and DxnB2 share ~23% amino acid sequence identity and have been classified into phylogenetically distinct MCP hydrolase subfamilies. BphDLB400 belongs to the well-studied class I subfamily, which also includes MhpC and HsaD whereas DxnB2 groups with class III enzymes[39]. As the reported oligomeric state differs for class I and III   64 MCP hydrolases, the quaternary structure of DxnB2 was compared to that of BphDLB400 and HsaD in solution. Specifically, experiments were performed using enzyme samples at a concentration of 1.5 to 2.5 mg/ml in 20 mM HEPES, pH 7.5. SEC-MALS analysis indicated that DxnB2 exists as a dimer with a molecular weight of 58 ? 2 KDa (Figure 27). The tetrameric assemblies of BphDLB400 and HsaD, which measured 132 ? 2 KDa and 121 ? 1 KDa were also confirmed by SEC-MALS. Figure 27. Superposition of chromatograms from SEC-MALS analysis of BphDLB400 (grey), DxnB2 (black), and HsaD (dashed line). The data used for molecular mass determination has been overlaid on the chromatogram.  3.3.2 Evidence that DxnB2 also utilizes a nucleophilic mechanism of catalysis 3.3.2.1 A pre-steady-state kinetic burst of HPD formation  While previous steady-state kinetic analyses revealed that DxnB2 catalyzes the efficient (kcat/Km ~107 M-1s-1) hydrolysis of HOPDAs[39], no mechanistic information can be directly inferred from these studies. To further investigate the mechanism of catalysis by DxnB2, the pre-steady-state kinetic behaviour of the enzyme was probed using stopped-flow spectrophotometry. The hydrolysis of 20 ?M HOPDA by 1 or 2.5 ?M DxnB2 WT and 9-Cl HOPDA by 2.5 ?M enzyme was measured in KPi (I = 0.1 M), pH 7.5 at 25 ?C. These   65 substrates were selected on the basis of distinct steady-state kinetic behaviour: kcat values measured for the DxnB2-mediated hydrolysis of HOPDA and all other tested monochlorinated substrates ranged between 0.13 and 0.88 s-1 with the exception of 9-Cl HOPDA for which kcat was determined to be 5.1 s-1[39]. As in BphDLB400, reactions were monitored at 270 nm to follow the progress of HPD formation (Figure 28), and the values derived from fitting the stopped-flow measurements to a burst equation are summarized in Table 9. For HOPDA, a pre-steady-state burst occurred at ~62 s-1. Quantification of the amount of HPD produced during each burst phase indicated that hydrolysis was 1:1 with enzyme concentration, suggesting that half-site reactivity may not be a universal feature of the MCP hydrolases. The observed steady-state rate constant for the hydrolysis (kss) was ~0.4 s-1, similar to the previously reported value for kcat ~0.47 s-1[39]. Figure 28. Representative stopped-flow experiments demonstrating the pre-steady-state kinetic burst of HPD formation from HOPDA (black) and 9-Cl HOPDA (grey) by DxnB2. For clarity, 1/10 data points are shown and distinct y-axes are shown.    66 Table 9. Pre-steady-state kinetic parameters derived from monitoring HPD formation [E] (?M) Substrate kburst  (s-1) Ampburst (?A270nm) [P]burst (?M) vss  (?A270nm s-1) kss (s-1) 1 HOPDA 63 ? 1 0.019 ? 0.002  1.01 ? 0.09  0.0079 ? 0.0002 0.41 ? 0.01 2.5 HOPDA 61.7 ? 0.7 0.048 ? 0.002  2.5 ? 0.1  0.0173 ? 0.0009  0.36 ? 0.02  2.5 9CH 20.0 ? 0.5 0.032 ? 0.004 1.67 ? 0.02 0.208 ? 0.003 4.34 ? 0.06  In contrast to HOPDA, quantification of HPD formed during the burst from the first turnover of 9-Cl HOPDA suggested that only ~67% of the enzyme was active. The observation of significant substrate inhibition during the reaction of DxnB2 and 9-Cl HOPDA, Ksi ~6 ?M, may account for this observed reactivity. Alternatively, the 9-Cl substituent may provide an additional structural determinant required to invoke the half-site reactive kinetic behaviour observed in BphDLB400 or reflect the context of non-cognate substrate turnover by DxnB2. Finally, the quantification may also be less accurate for the turnover of 9-Cl HOPDA due to the comparable pre- and steady-state rates, kburst and kss, which differ by less than 5-fold. Despite the unexplained nature of the burst of HPD from the hydrolysis of 9-Cl HOPDA, the observation of pre-steady-state kinetic bursts for both HOPDA and 9-Cl HOPDA is consistent with a proposal that DxnB2 utilizes a covalent mechanism of catalysis in which kacylation > kdeacylation. 3.3.2.2 Identification of an acyl-enzyme intermediate in DxnB2 by LC ESI/MS  Similar to the manner in which an acyl-enzyme was initially identified in BphDLB400, chemical quench experiments were performed on whole protein samples recovered from the turnover of a 5-fold excess of HOPDA or 9-Cl HOPDA by DxnB2. Reactions of 4 ?M DxnB2 and 20 ?M HOPDA or 9-Cl HOPDA were allowed to age for 10 or 1 s before quenching with a final volume of 5% acetic acid (v/v). LC ESI/MS analyses of the reaction mixtures revealed acylation of DxnB2 in both cases (Figure 29). The observed mass of   67 DxnB2 was 30198 ? 7 Da (~220 ppm) was within error of that predicted based on the protein sequence, 30194 Da. Additional peaks with masses shifted by 104 or 138 Da were taken to correspond to benzoylation or 3-chlorobenzoylation of DxnB2. Beyond acylation, peaks were observed that were tentatively assigned to non-covalent adducts. Based on the shift in m/z, these additional peaks included (i) an acyl-enzyme in complex with an additional substrate molecule, M+324 and M+388, for the reaction with HOPDA and 9-Cl HOPDA, respectively, (ii) a non-covalent DxnB2:9-Cl HOPDA adduct or 3-benzoylated enzyme:HPD binary complex with M+250, and (iii) an M+640 peak in the 9-Cl HOPDA sample, which may consist of a 3-chlorobenzoylated enzyme with two additional substrate molecules adventitiously bound. The observation of additional non-covalent complexes is unusual, but may reflect preservation of active sites throughout the experimental treatment and the observed micromolar Ksi values for each substrate[39]. Moreover, the quenched reaction mixtures were concentrated to approximately 40 ?M DxnB2 prior to analysis by LC ESI/MS. Interestingly, ~85% of DxnB2 was acylated based on estimating the area of each of the three distinct species observed from the reaction with HOPDA. This observation is more consistent with the 1:1 reactivity determined from the pre-steady-state kinetic burst experiments than the expected half-site reactivity that has been reported or inferred for other MCP hydrolases.    68 Figure 29. Whole protein LC ESI/MS analysis of DxnB2 reacted with HOPDA for 10 s (dashed line) or 9-Cl HOPDA for 1 s (solid line).  3.3.3 Single turnover of HOPDA by DxnB2  To further evaluate the kinetic mechanism of DxnB2, single turnover experiments were performed using 16 ?M DxnB2 and 4 ?M HOPDA in KPi (I = 0.1 M), pH 7.5 at 25 ?C (Figure 30). A stopped-flow instrument outfitted with a PDA detector was utilized to identify any intermediates along the reaction pathway whereas a monochromator was employed to monitor reactions at a single wavelength. To distinguish intra- and intermolecular events, additional reactions were performed in the presence of a viscogen, either 30% sucrose (w/v) or 30% glycerol (v/v).  The hydrolysis of HOPDA by DxnB2 was biphasic. In this instance, ESred was characterized by a ?max at 487 nm and formed within the dead-time of the instrument (1/?1 > 500 s-1). Subsequently, the red-shifted intermediate decayed at ~69 s-1, which was mirrored by an increase in absorbance at 270 nm (Table 10). Following the hydrolysis of HOPDA, a relaxation attributed to HPD rebinding was also observed in the visible region, accounting for   69 ~1% of the total change in amplitude (Figure 55, Appendix III). Likewise, the ketonization of HPD was also detectable at 270 nm (see section 3.3.5), a post-catalytic event that was previously reported for BphDLB400[81]. Again, DxnB2 revealed no trace of half-site reactivity with respect to the hydrolysis of HOPDA, and the slowest rate constant determined from these experiments was ~150-fold faster than kcat, consistent with a mechanism in which deacylation is rate-limiting. Figure 30. Results from representative stopped-flow experiments showing the single turnover of 4 ?M HOPDA by 16 ?M DxnB2. (A) Reaction monitored using a PDA detector. (B) Reaction monitored at single wavelengths: ESred decay at 487 nm and HPD formation at 270 nm. For simplicity 1/10 data points are shown.   Interestingly, the addition of sucrose or glycerol to the reaction media did not affect 1/?2, measured as either the dissipation of ESred or increase in absorbance at 270 nm (Table 10). In BphDLB400, this transient kinetic phase was sensitive to both temperature and a viscogen, arguing for a dependence on product release, which in the two-conformation model is thought to precedes conformational changes that allow the second set of active sites to react[81]. The insensitivity of 1/?2 to viscogens in DxnB2 allows the provisional assignment of this transient kinetic phase to acylation, an on-enzyme event. While the data provide   70 accurate rate constants for acylation, an explicit assignment of the determinant(s) thereof cannot be inferred from a viscosity effect alone. Table 10. Solvent viscosity effects on a single turnover of HOPDA by DxnB2 Buffer composition ? (nm) 1/?2 (s-1) No viscogen 270 69 ? 2  487 68 ? 1 30% sucrose (w/v) 270 67 ? 5  487 69 ? 1 30% glycerol (v/v) 270 64 ? 5  487 72.7 ? 0.5  3.3.4 Transient state solvent kinetic isotope effect (SKIE) for HOPDA hydrolysis  In contrast to BphDLB400, the DxnB2-mediated cleavage of HOPDA was monophasic and impervious to solvent viscosity. Thus, it was rationalized that the measurement of a single turnover in deuterated reaction media could be employed to resolve the chemical step associated with ESred decay. The presence of an SKIE on 1/?2 would argue for a rate-determining proton transfer, ultimately affording the explicit assignment of this observable kinetic phase to substrate ketonization or acylation in DxnB2.  Single turnover experiments were performed using 8.2 ?M DxnB2 and 4 ?M HOPDA in KPi (I = 0.1 M), pH or pD 7.5 at 25 ?C (Figure 31). The isotopically enriched buffer contained 95% D2O (v/v), and pre-incubation of HOPDA resulted in solvent exchange into the dienoate (Figure 56, Appendix III). As in H2O, ESred decay was monophasic and matched by an increase in absorbance at 270 nm. Despite using approximately half of the concentration of enzyme, 1/?2 was within error to that observed at 4E:1S (Table 11). Upon measuring the reaction in deuterated buffer, an SKIE of ~2.5 was observed, indicating that a proton transfer limited this observable transient kinetic phase. Therefore, 1/?2 could be assigned to substrate ketonization in DxnB2. Thus, acylation of DxnB2 is determined by   71 substrate protonation, which is linked to nucleophile activation in the substrate-assisted mechanism proposed for the MCP hydrolases.  Figure 31. Representative stopped-flow experiments demonstrating a transient state SKIE on the rate of DxnB2-mediated HOPDA cleavage.  Table 11. Pre-steady-state solvent kinetic isotope effect on DxnB2 hydrolysis of HOPDA ? H2O 95% D2O (v/v)  ? (nm) 1/?2 (s-1) 1/?2 (s-1)? SKIE 270 71 ? 2 28 ? 1 2.5 ? 0.2  487 67.7 ? 0.7 27.0 ? 0.3 2.50 ? 0.05  3.3.5 An extended Br?nsted analysis of substrate ketonization Confident assignment of an observable transient kinetic phase to substrate ketonization in DxnB2 facilitated a more robust investigation of this enzymatic process. The DxnB2-mediated hydrolysis of a series of substituted HOPDAs with varying basicity was measured by stopped-flow using a similar experimental approach described for the unsubstituted substrate. Additional reactions between 16 ?M DxnB2 and 4 ?M 5-Cl , 5,8-diF, 8-Cl or 9-Cl HOPDA were performed in KPi (I = 0.1 M), pH 7.5 at 25 ?C (Table 12).   As previously described for HOPDA, the hydrolysis of 5-Cl HOPDA was biphasic.   72 Table 12. Kinetic data for the DxnB2-mediated hydrolysis of substituted HOPDAs Substrate ? (nm) 1/?1 (s-1) [% Amp] 1/?2 (s-1) [% Amp] 1/?3 (s-1) [% Amp] 1/?4 (s-1) [% Amp]a HOPDA 270     69 ? 2 [26 ? 0.2]     0.014 ? 0.001 [74 ? 5]  487 > 500 68 ? 1 [98.9 ? 0.1]     0.6 ? 0.3 [1.1 ? 0.1] 5-Cl  270     2.9 ? 0.2 [23.9 ? 0.2]     0.05 ? 0.02 [80 ? 30]  472 312 ? 5 [43 ? 1] 2.72 ? 0.01           5,8-diF  270         0.132 ? 0.002 [20.6 ? 0.3] 0.0018 ? 0.0001 [79 ? 2]  485 520 ? 20 [46 ? 4] 2.3 ? 0.3 [2.6 ? 0.6] 0.126 ? 0.001 [51.0 ? 0.5]     8-Cl  270     4.3 ? 0.1 [43.9 ? 0.6] 1.0 ? 0.2 [52 ? 1] 0.039 ? 0.001 [52 ? 1]  472 > 500 5.0 ? 0.1 [73 ? 1] 1.76 ? 0.08 [24 ? 1]     9-Cl  270     20 ? 1 [52 ? 7] 7 ? 1 [10 ? 5] 0.07 ? 0.02 [40 ? 10]  494 > 500 18.6 ? 0.5 [91 ? 5] 6 ? 2 [9 ? 4]     + 30% sucrose 270  17.4 ? 0.9 [43 ? 6] 8 ? 3 [6 ? 5] 0.060 ? 0.005 [52 ? 4] 494 > 500 17.8 ? 0.6 [91 ? 6] 6 ? 3 [8 ? 6]     a1/?4 is a measure of post-cleavage processes: HPD rebinding at 487 nm (see Appendix III) or enzymatic and non-enzymatic HPD ketonization at 270 nm. Note that there is a considerable amount of uncertainty in these measurements (reflected in the standard deviation between measurements) as the experiments were designed to investigate the pre-steady-state catalytic events    73 ESred was characterized by a ?max at 472 nm and a rate constant of ~312 s-1 was measured (Figure 32A and E). Subsequently, the ketonization reaction occurred at ~2.8 s-1, which was apparent from the kinetic traces at both 472 and 270 nm. For the remaining substrates, an additional relaxation event was observed after formation of ESred. The red-shifted intermediates, which all accumulated within or near the dead-time of the instrument (1/?1 ? 500 s-1) were defined by ?max values at 485, 472 and 494 nm for 5,8-diF, 8-Cl and 9-Cl HOPDA, respectively (Figure 32B-D). For 5,8-diF HOPDA, biphasic decay of ESred was not coupled to a biphasic increase in the absorbance at 270 nm (Figure 32F). Instead, a small decrease in the signal at 485 nm, accounting for only 2.3% of the total change in amplitude suggested that the two initial relaxation processes corresponded to substrate destabilization. The final observable phase, 1/?3 ~0.13 s-1 at 270 nm, matched the kcat value, indicating that ketonization rather than deacylation limited the hydrolysis of 5,8-diF HOPDA. The turnover of 8-Cl and 9-Cl HOPDA was also triphasic (Figure 32G and E), including the biphasic decay of ESred accompanied by HPD formation, reminiscent of the two distinct acylation events proposed for BphDLB400[81]. Thus, ketonization of 8-Cl HOPDA proceeded at 4.3 and 1.0 s-1 (1/?2 and 1/?3) whereas deacylation occurred at ~0.48 s-1 [39]. The corresponding rate constants associated with ketonization of 9-Cl HOPDA were faster, 20 s-1 and 7 s-1, and considering that 1/?3 was commensurate with kcat, the results suggested that this process may be rate-limiting even for more basic substrates. The assignment of the final observable phase to a chemical event was supported by repetition of the reaction of DxnB2 and 9-Cl HOPDA in the presence of 30% sucrose (w/v). The viscogen did not perturb the reaction under single turnover conditions. Finally, the biphasic nature of ESred decay and    74 Figure 32. Representative stopped-flow experiments monitoring the turnover of 4 ?M substrate by 16 ?M DxnB2: (A/E) 5-Cl , (B/F) 5,8-diF , (C/G) 8-Cl and (D/H) 9-Cl HOPDA.    75 the sub-stiochiometric burst of product formation from 9-Cl HOPDA prompted the hypothesis that the ring-substituents may be determinants that invoke half-site reactivity in DxnB2. Figure 33. Extended Br?nsted relationship of substrate basicity and 1/?2 or kcat/Km (inset; kcat/Km values were reported in [39], including 10-Cl HOPDA, which was not investigated in this work).  A plot of the experimentally observed pKa2 values of each substrate against the log(1/?2), demonstrated a positive linear correlation (R2 = 0.97). Thus, the extended Br?nsted analysis revealed that the ketonization reaction was sensitive to the nucleophilicity of the substrate, ?nuc = 1.17 ? 0.06 (Figure 33). Similarly, analysis of the steady-state specificity constants (kcat/Km) for the DxnB2 hydrolysis of substituted HOPDAs showed a positive correlation (R2 = 0.96) and ?nuc = 1.0 ? 0.1. 9-Cl HOPDA was excluded from the latter analysis since the C9-substitutent provided an additional substrate specificity determinant compared to all other tested substrates. The sensitivity of the reaction to the basicity of the substrate is consistent with the proposed substrate-assisted mechanism of nucleophile activation in the MCP hydrolases. Considering only the analysis on 1/?2, the more basic   76 substrates are more reactive towards an incipient proton and, therefore, are more readily ketonized. Along these lines, the more acidic substrates are better suited to stabilize the negative charge on the enzyme-bound dianionic dienoate moiety of ESred. For 5,8-diF HOPDA, the most extreme case reported herein, the rate of substrate ketonization is rate-determining, a consequence of the stability of ESred. 3.3.6 Probing the electronic nature of ESred by NMR  Despite the relevance of ESred to catalysis, the identity of this intermediate is unknown. In an attempt to characterize ESred, a uniformly labeled 13C-HOPDA was generated by enzymatic transformation of 13C-biphenyl by BphAEFG, BphB and BphC. The identities of individual carbons in the DxnB2 S105A:HOPDA complex were not assigned. Overall, the signals originating from HOPDA, whether free in solution or in complex to DxnB2 S105A, were observed over a similar chemical shift range (Figure 56, Appendix IV). Unfortunately, most of these broad signals occur within 1 ppm and 10 ppm in the 1H and 13C dimension, respectively. Instead, the ability of DxnB2 to hydrolyze fluorinated HOPDAs in a manner that resembled turnover of the unsubstituted substrate provided an opportunity to probe the electronic nature of this intermediate via the 19F nuclei. Moreover, the 19F signals could be definitively assigned through comparing the NMR spectra of 8-F and 5,8-diF HOPDA. Accordingly, 8-F and 5,8-diF HOPDAs were prepared and complexed to DxnB2 S105A. 19F NMR spectra of the solutions containing 400 ?M DxnB2 and 200 ?M 8-F or 5,8-diF HOPDA were compared to those of the free substrates (Figure 34) and model fluorine-containing compounds (Table 13). Interestingly, the 19F nuclei of 5,8-diF HOPDA were degenerate in solution whereas binding to DxnB2 S105A resulted in the appearance of a second signal at -157.71 ppm. The latter was assigned to C5-F based on the signal observed   77 from S105A:8-F HOPDA binary complex. For 8-F HOPDA, the enzyme effectively inverted the population of the major and minor species in solution. Relative to the model 2-fluorophenyl ketones, the observed -40.6 ppm shift of C5-F placed the observed species closer to the sp3- than the sp2-hybridized analog. In spite of this, a definitive assignment of the hybridization state of C5 in the S105A:5,8-diF HOPDA is not possible considering the broad and overlapping range of 19F chemical shifts associated with atoms connected to sp2 and sp3-hydbrized carbon centers and the sensitivity of the 19F to distinct environments. Figure 34. 19F NMR spectra of (A) 200 ?M 8-F HOPDA, (B) 200 ?M 8-F HOPDA in complex with 400 ?M DxnB2 S105A, (C) 200 ?M 5,8-diF HOPDA and (D) 200 ?M 5,8-diF HOPDA in complex with 400 ?M DxnB2 S105A in KPi (I = 0.1 M), pH 7.5 at 25 ?C.     78  Table 13. 19F chemical shifts observed in KPi (I = 0.1 M), pH 7.5 at 25 ?C. Sample 19F chemical shift (ppm) 2-fluorophenylpropanone  -184.32 (minor) and -185.13 (major) 2-fluorophenylprop-2-enone -111.73 (major), -116.46 (minor), -111.00 (minor) 8-F HOPDA -116.86 (minor), -111.00 (major) S105A:8-F HOPDA -116.79 (major), -112.04 (minor) 5,8-diF HOPDA -116.96 S105A:5,8-diF HOPDA -117.17 and -157.71   3.3.7 Probing the reactivity of ESred with molecular oxygen  Enzymes that mediate carbanion chemistry must protect their reactive species in order to prevent non-specific catalysis. In particular, the reaction media almost always contains undesired electrophiles such as molecular oxygen. Accordingly, the relative ability of enzymes to protect their carbanionic intermediates can be surveyed by monitoring O2-consuming side-reactions[116]. In light of the substrate-assisted proposal for nucleophile activation and the extended lifetime of ESred in the catalytic serine variants, the O2-sensitivity of a DxnB2 S105A:HOPDA binary complex was tested. The half-life (t1/2) of ESred was measured spectrophotometrically as a function of O2 concentration. Binary complexes were prepared at 650 ?M, 280 ?? or ~0.5 ?M O2 (? 5 ppm) in KPi (I = 0.1 M), pH 7.5 at 25 ?C. Interestingly, added O2 resulted in a 3.4-fold decrease in the t1/2 of ESred, which was measured to be 1.8 ? 0.2 and 6.2 ? 0.1 h at 650 and 280 ?M O2, respectively. In the absence of oxygen, the t1/2 of ESred could not be modeled by a single exponential decay function. Instead, the t1/2 was estimated to be 26 ? 2 h based on a single time point at which three quarters of starting absorbance remained.  To further assess the reactivity of O2 with ESred, the benzoyl-containing ESred decay products were analyzed by EI GC/MS. Benzoate was observed as the major decay product of   79 ESred after incubation overnight at ambient O2. Next, the products from the decay of 9-Cl HOPDA or a DxnB2 S105A:9-Cl HOPDA binary complex in a solution of KPi (I = 0.1 M), pH 7.5, saturated with O2 (~1 mM) were analyzed by EI GC/MS. It was rationalized that the natural abundance of the 35Cl and 37Cl isotopes could be used to guide the tentative assignment of any minor ESred or 9-Cl HOPDA decay products. Inspection of the chromatograms from the decay of ESred to that of a sample of 9-Cl HOPDA alone, revealed an additional DxnB2 S105A-associated product. This 3-chlorobenzoyl-containing product was defined by an Rt of 21.8 minutes and an m/z value of 241 (Figure 35). Strikingly, the 241 m/z product was produced at a 1:8 ratio to 3-chlorobenzoate (Rt 17.8 min; 228 m/z), which constituted the major decay product from the S105A:9-Cl HOPDA complex. The observed mass relative to 3-chlorobenzoate, M+13, is consistent with a compound possessing an additional C atom and H atom. This product has been provisionally assigned as 1-(3-chlorophenyl)-2-hydroxyethanone, which may result from an O2-consuming side reaction. Nevertheless, 18O2 and H218O isotope studies are required to identify the source of the oxygen atoms in the observed products and higher resolution experiments are required for the unambiguous assignment of the atomic mass for the newly identified 241 m/z product.   80 Figure 35. Mass spectrum of the O2-dependent ESred breakdown product. The chemical structure of a compound with a theoretical m/z value of 241 m/z is inset.  3.4 Characterization of DxnB2 P-subsite and His255 variants 3.4.1 Steady-state kinetic analysis of P-subsite and His255 variants   To help elucidate the role of the P-subsite and His255 in catalysis, a series of DxnB2 variants were generated that covered a relatively broad range of steric and electronic substitutions: Asn43 was substituted with each of Ala, Asp and His; Arg180 was replaced with each of Gln, Lys and Met; and His255 with each of Ala, Asp and Gln. P-subsite variants were characterized by a relatively wide range of steady-state kinetic parameters. In contrast, the His variants resembled those observed in other MCP hydrolases; the substitution of His255 in DxnB2 to Ala, Gln or Glu resulted in severely compromised enzymes. Steady-state turnover rate constants (v0/[E]T) estimated from v0 measurements of the turnover of 20 ?M HOPDA or 9-Cl HOPDA by each His255 variant ranged between 0.0001 and 0.002 s-1 (Table 14).      81 Table 14. The hydrolytic activity of DxnB2 His255 variants  20 ?M HOPDA 20 ?M 9-Cl HOPDA Enzyme v0/[E]t (s-1) v0/[E]t (s-1) H265A 0.00019 ? 0.00002 0.00032 ? 0.00007 H255E 0.00066 ? 0.00006 0.0015 ? 0.0004 H255Q 0.00011 ? 0.00003 0.00026 ? 0.00009  For the P-subsite variants, v0 data was measured by varying the concentration of HOPDA or 9-Cl HOPDA from 0.5 to 50 ?M or from 10 to 200 ?M, depending on the enzyme under investigation. Steady-state kinetic parameters were obtained from plots of v0 versus substrate concentration fit with either the Michaelis-Menten or substrate inhibition equation (Appendix V). As summarized in Table 15, all of the variant possessed greater specificity for 9-Cl HOPDA, a characteristic shared with the WT enzyme. However, the variants showed one of two kinetic behaviours. The first group, N43A, N43H and R180K, was similar to the WT in that each was subject to substrate inhibition. Nevertheless, the observed inhibition was lesser in variants and for substituted HOPDAs. For example, the N43H variant was inhibited by HOPDA but not 9-Cl HOPDA, and the effects of the N43A, N43H and R180K substitutions on Ksi varied between a 1.7 to 50-fold relative to the WT. Along with sharing kinetic behaviour with the WT enzyme, these variants were the most active towards the substrates with kcat/Km values determined between 104 and 105 M-1s-1. The second group of variants, N43D, R180M and R180Q, were defined by the lack of substrate inhibition and significantly greater Km values (> 100-fold). Remarkably, for both HOPDA and 9-Cl HOPDA the Km increased by up to three orders of magnitude whereas the kcat values were typically within an order of magnitude the WT-catalyzed reaction. Taken together, these steady-state kinetic results emphasize the importance of the P-subsite for catalysis and implicate the P-subsite as a determinant for substrate inhibition in DxnB2.  82 Table 15. Steady-state kinetic parameters of DxnB2 P-subsite variants for HOPDA and 9-Cl HOPDAa,b  HOPDA 9-Cl HOPDA  kcat (s-1) Km (?M) kcat/Km (105 M-1 s-1) Ksi (?M) kcat (s-1) Km (?M) kcat/Km (105 M-1 s-1) Ksi (?M) WTc 0.47 ? 0.02 0.039 ? 0.006 120 ? 40 37 ? 8 5.1 ? 0.6 0.11 ? 0.04 500 ? 100 6 ? 2 N43A 0.12 ? 0.01 0.25 ? 0.03 4.9 ? 0.5 63 ? 7 0.85 ? 0.02 0.19 ? 0.03 46 ? 6 190 ? 40 N43D 0.045 ? 0.002 57 ? 4 0.0079 ? 0.0003 NA 0.45 ? 0.01 39 ? 4 0.115 ? 0.007 NA N43H 0.023 ? 0.001 1.00 ? 0.07 0.23 ? 0.01 90 ? 10 0.077 ? 0.001 0.28 ? 0.03 2.8 ? 0.3 NA R180K 0.31 ? 0.01 1.26 ? 0.09 2.5 ? 0.1 130 ? 20 2.36 ? 0.05 0.37 ? 0.03 63 ? 5 280 ? 8 R180M 0.022 ? 0.001 76 ? 9 0.0029 ? 0.0002 NA 0.98 ? 0.01 190 ? 30 0.053 ? 0.003 NA R180Q 0.035 ? 0.001 64 ? 4 0.0054 ? 0.0002 NA 1.17 ? 0.04 94 ? 7 0.124 ? 0.006 NA aApparent Km and kcat/Km values for enzymes in which Km < 1 ?M bThe complete set of steady-state kinetic curves for the P-subsite variants are available in Appedix V. cValues for the WT enzyme were taken from [39].    83 3.4.2 Investigating the ability of P-subsite and His255 variants to accumulate ESred  While sequence conservation and steady-state kinetic data have implicated the P-subsite in catalysis, the specific roles of these residues remain unknown. To study the role of Asn43, Arg180 and His255 in substrate binding and destabilization (ESred formation), each residue was mutated in a DxnB2 S105A background. The electronic absorption spectra of ES complexes were measured in KPi (I = 0.1 M), pH 7.5 at 25 ?C. Both HOPDA and 9-Cl HOPDA were examined (Figure 36). In nearly all the complexes, at least a small population of ESred was observed. The replacement of Arg180 with a neutral amino acid appeared to have the most deleterious effect on substrate destabilization. For HOPDA, no red-shifted species was observed at 4E:1S, and only a very small population was observed for 9-Cl HOPDA in solution with the S105A/R180M and S105A/R180Q variants. Moreover, these variants appeared to stabilize the enol form of the substrates, possessing a maximal absorbance near 340 and 350 nm for HOPDA and 9-Cl HOPDA, respectively. Finally, an unidentified species characterized by a ?max between 400 to 420 nm was observed for 9-Cl HOPDA in complex with the S105A/R180M, S105A/R180Q and S105A/H255E variants. Although the electronic nature of this species is unknown, absorption at wavelengths greater than 390 nm is typically associated with dianionic HOPDA isomers (Appendix I). The ability of each P-subsite variant to at least partially stabilize some red-shifted species suggests that each residue contributes to substrate destabilization.     84  Figure 36. Perturbations to the electronic absorption spectra observed upon mixing 40 ?M enzyme with 10 ?M HOPDA (top) or 10 ?M 9-Cl HOPDA (bottom)  85 3.4.3 Kd Determinations  The observed spectral perturbations observed at 4E:1S was further exploited to determine dissociation constants for HOPDA and 9-Cl HOPDA to key DxnB2 variants. The enzymes with high Km values, N43D/S105A, S105A/R180M and S105A/R180Q were of particular interest in order to delineate the deleterious effects associated with altered substrate binding or chemistry. Thus, enzymes were titrated into a solution of either 4 ?M HOPDA or 3 mM 9-Cl HOPDA in KPi (I= 0.1 M), pH 7.5 at 25 ?C. Electronic absorption spectra were recorded upon each successive addition of enzyme, and difference spectra were used to determine the dissociation constants from fitting either hyperbolic or quadratic binding equations to plots of ?Abs versus enzyme concentration (Appendix V). The assay was not sensitive enough to determine Kd HOPDA or Kd 9-Cl HOPDA from the N43D/S105A variant. An upper bound for the Kd in both cases was set at 20 nM (Figure 59, Appendix V), suggesting that the high Km value determined for the N43D variant reflected a deficiency in chemistry rather than affinity for the substrates. Alternatively, the dissociation constants for S105A/R180Q and especially S105A/R180M indicated that poor substrate binding contributed significantly to the reduced hydrolytic activity of the R180M and R180Q variants (Table 16 and Figure 37). At least two distinct species were observed from complexes of HOPDA and S105A/R180M or S105A/R180Q (Figure 19D). Up to four electronic transitions were observed from complexes of 9-Cl HOPDA and S105A/R180M or S105A/R180Q (Figure 19C). For HOPDA, the variants perturbed the enol:enolate equilibrium with a prominent species at ~340 nm appearing with the addition of enzymes. For the 9-Cl HOPDA complexes, a blue-shifted species possessing the highest absorptivity was centered between 400 and 410 nm, and an enol form of the substrate was also apparent   86 (?max ~350 nm). Interestingly, despite the replacement of Arg180 with uncharged residues, a small population of ESred was still observed. Table 16. Kd for DxnB2 S105A/R180M and S105A/R180Q:HOPDA complexes Enzyme ?? (nm) Kd HOPDA (?M) ?? (nm) Kd 9-Cl HOPDA (?M) S105A/R180M 337 140 ? 20 350 180 ? 30 S105A/R180Q 337 52 ? 2 512 14 ? 2  Figure 37. Titration of 4 ?M HOPDA or 3 ?M 9-Cl HOPDA with (A) DxnB2 S105A/R180M and (B) S105A/R180Q. (C) The spectra of 3 ?M 9-Cl HOPDA (dashed lines) before and after titration with DxnB2 S105A/R180M (black) or S105A/R180Q (grey). (D) The spectra of 4 ?M HOPDA before and after titration with DxnB2 S105A/R180M or S105A/R180Q.     87 3.4.4 Single turnover of HOPDA and 9-Cl HOPDA by P-subsite variants  Each of the P-subsite variants perturbed the substrate in a distinct manner. In order to determine the catalytic relevance of these perturbations, the single turnover of HOPDA and 9-Cl HOPDA by each variant was examined. DxnB2 R180M and R180Q were excluded from this analysis since the corresponding variants in the S105A background possessed significantly reduced substrate affinity. Reactions of 16 ?M enzyme with 4 ?M substrate in KPi (I = 0.1 M), pH 7.5 at 25 ?C, were first monitored using a PDA detector in order to identify relevant intermediates (Figure 38 and 39A-D). In contrast to the observations in the S105A/P-subsite double variants, a red-shifted intermediate was only detected during turnover by DxnB2 N43H. Following the detection of catalytic intermediates, reactions were monitored at 270 nm and the ?max of the enolate or, in the case of N43H, a red-shifted wavelength, 453 and 455 nm for the turnover of HOPDA and 9-Cl HOPDA, respectively (Figures 38 and 39 E-H; Table 17). For some reactions, a small and rapid spectral perturbation, 1/?1 ranging from 6 to 150 s-1, preceded enolate decay assigned here to substrate ketonization based on the ability to  monitored changes at both stated wavelengths. A third, post-reaction relaxation, associated with HPD ketonization could also be observed at 270 nm. However, the accuracy in defining this phase was almost always limited by the timescale of the experiments (designed to capture early catalytic events). Notably, the rate constants for substrate ketonization, 1/?2, matched the kcat values determined for the N43H variant as well as that for the R180K turnover of 9-Cl HOPDA. Under single turnover conditions, the N43D enzymes were consistently 4-fold slower than under steady-state conditions. The rate constant for ESred decay rather than formation limited substrate ketonization in the N43H variant. Indeed, the   88 most common effect of perturbing the P-subsite was the generation of an enzyme in which ESred formation was rate-limiting, characterized by its absence from kinetic traces, as previously described for the BphDLB400-mediated turnover of recalcitrant PCB metabolites[86] or the activity associated with the His265 variants[66].     89 Figure 38. Results from representative stopped-flow experiments demonstrating the turnover of 4 ?M HOPDA by 16 ?M DxnB2 P-subsite variant: (A/E) N43A, (B/F) N34D, (C/G) N43H and (D/H) R180K. (A/E) N43A, (B/F) N34D, (C/G) N43H and (D/H) R180K. For clarity, selected traces showing the progress of the reaction between 300 and 600 nm are shown, and 1/10 data points are shown in the plots of relative ?Abs versus time.     90 Figure 39. Results from representative stopped-flow experiments demonstrating the turnover of 4 ?M 9-Cl HOPDA by 16 ?M DxnB2 P-subsite variant: (A/E) N43A, (B/F) N34D, (C/G) N43H and (D/H) R180K. (A/E) N43A, (B/F) N34D, (C/G) N43H and (D/H) R180K. For clarity, selected traces showing the progress of the reaction between 300 and 600 nm are shown, and 1/10 data points are shown in the plots of relative ?Abs versus time     91 Table 17. Kinetic data for HOPDA and 9-Cl HOPDA turnover by P-subsite variants  ? HOPDA DxnB2 (nm) 1/?1 (s-1) [%Amp] 1/?2 (s-1) [%Amp] 1/?3 (s-1) [%Amp] N43A 270     1.3 ? 0.1 [79 ? 5] 0.3 ? 0.2 [29 ? 9]  434 120 ? 20 [4.6 ? 0.1] 1.33 ? 0.01 [95 ? 1]     N43D 270     0.019 ? 0.001 [45 ? 3] 0.08 ? 0.01 [55 ? 1]  434 6 ? 2 [1.2 ? 5] 0.017 ? 0.001 [99 ? 1]     N43H 270     0.026 ? 0.001 [50 ? 10] 0.013 ? 0.002 [50 ? 10]  453 29 ? 4 [4.9 ? 0.2] 0.035 ? 0.001 [95 ? 1]     R180K 270     1.09 ? 0.04        434 110 ? 40 [5.7 ? 0.3] 1.01 ? 0.02         9-Cl HOPDA N43A 270     2.40 ? 0.07        436     2.38 ? 0.02       N43D 270     0.149 ? 0.004 [49 ? 1] 0.006 ? 0.004 [50 ? 20]  436 9 ? 1 [2.2 ? 0.4] 0.152 ? 0.002 [97 ? 1]     N43H 270     0.062 ? 0.004 [56 ? 5] 0.011 ? 0.004 [44 ? 1]  455 150 ? 50 [6.1 ? 0.2] 0.069 ? 0.001 [94 ? 1]     R180K 270     2.53 ? 0.08        436     2.42 ? 0.02        3.4.5 Single turnover of HOPDA and 9-Cl HOPDA by the His255 variants  The accumulation of red-shifted intermediates in DxnB2 S105A/H255 variants contrasts with what was reported for BphDLB400 S112A/H265 variants, which stabilized the HOPDA enolate dianion in an apparently planar conformation[66]. Stopped-flow experiments were used to measure the rate of a single turnover of HOPDA or 9-Cl HOPDA by the DxnB2 His255 variants in KPi (I = 0.1 M), pH 7.5 at 25 ?C. Detection of reaction intermediates using a PDA indicated that only the H255A and H255Q variants accumulated detectable quantities of a red-shifted species (Figure 40 and 41A-C). Although the rate constants for the turnover of the substrates by each enzyme were similar (Table 18), the spectroscopic changes that were observed during the turnover of 9-Cl HOPDA were more pronounced, again reflecting the additional determinant of specificity carried by the   92 substitution at C9.  The spectra of the HOPDA species observed during the H255A-mediated turnover were bathochromically-shifted by up to 40 nm. For both substrates, the electronic absorption spectrum was complex, containing at least three spectroscopically distinct species throughout the single turnover. For HOPDA, the ?max of ESred was near 450 nm. However, a series of overlapping bands gave rise to a relatively broad signal between 375 to 525 nm. For 9-Cl HOPDA, the ESred ?max was better defined at 457 nm. Upon formation of ESred, the signal decayed at 0.5 and 0.2 s-1 for HOPDA and 9-Cl HOPDA, respectively, with a concomitant increase in the absorbance at 270 nm. In either case, this represented an ~100-fold reduction compared to the WT enzymes but was ~1000-fold faster than the steady-state rate constant for H255A turnover, ~0.0002 or ~0.0003 s-1 of HOPDA or 9-Cl HOPDA, respectively.  The absence of ESred from the kinetic traces of H255E-catalyzed reactions indicated that the formation of this intermediate limits the rate of ketonization and acylation in this enzyme. Moreover, the rate of ESred formation likely contributes to the steady-state rate of hydrolysis by this enzyme, which differed by only 16- or 6-fold from the measured 1/?4 values for HOPDA or 9-Cl HOPDA. Along with the absence of ESred, the H255E variant apparently shifted the enol:enolate equilibrium from that observed free in solution. A detectable quantity of the enol, absorbing at ~340 nm, accumulated during the turnover of both substrates.  Among the three His255 variants, the single turnover kinetic behaviour of H255Q was the most complex, and as in the WT the two distinct substrates were hydrolyzed in a different number of observable kinetic phases. The enzyme appeared to bind a protonated form of the substrate, indicated by the first recorded spectrum, which possessed bands at 340   93 and 370 nm. Next, a small population of ESred accumulated at 100 or 50 s-1 (1/?1), which was coupled to a decrease in the absorbance attributed to the bound enol species. The latter was measured at 340 nm for HOPDA and at both 345 and 370 nm for 9-Cl HOPDA. The ESred of HOPDA and 9-Cl HOPDA decayed in a mono and biphasic fashion, respectively. Nevertheless, the ketonization reaction could be modeled by single exponential process yielding rate constants of 0.036 (1/?3) or 0.0017 s-1 (1/?4) for HOPDA or 9-Cl HOPDA. Figure 40. Results from representative stopped-flow experiments demonstrating the turnover of 4 ?M HOPDA by 16 ?M DxnB2 His255 variants: (A/D) H255A, (B/E) H255E, (C/F) H255Q. For clarity, selected traces showing the progress of the reaction between 300 and 600 nm are shown and 1/10 data points are shown in the plots of relative ?Abs versus time.    94 Figure 41. Results from representative stopped-flow experiments demonstrating the turnover of 4 ?M 9-Cl HOPDA by 16 ?M DxnB2 His255 variants: (A/D) H255A, (B/E) H255E, (C/F) H255Q. For clarity, selected traces showing the progress of the reaction between 300 and 600 nm are shown and 1/10 data points are shown in the plots of relative ?Abs versus time.   .    95 Table 18. Kinetic data for HOPDA and 9-Cl HOPDA turnover by DxnB2 His255 variantsa  ? HOPDA DxnB2 (nm) 1/?1 (s-1) [%Amp] 1/?2 (s-1) [%Amp] 1/?3 (s-1) [%Amp] 1/?4 (s-1) [%Amp] H255A 270         0.49 ? 0.01 [79 ? 2]      340 120 ? 40 [7 ? 1] 1.8 ? 0.4 [12 ? 4] 0.49 ? 0.02 [81 ? 5]      474 70 ? 20 [6.2 ? 0.1] 1.7 ? 0.2 [13 ? 2] 0.50 ? 0.01 [81 ? 2]     H255E 270             0.0128 ? 0.0006 [54 ? 5]  434 70 ? 20 [19 ? 9] 14 ? 4 [30 ? 9] 1.19 ? 0.05 [37 ? 7] 0.0108 ? 0.0005 [17 ? 2] H255Q 270         0.036 ? 0.002 [61 ? 1]      340 102 ? 7 [10.3 ? 0.1] 1.9 ? 0.2 [3.9 ? 0.1] 0.063 ? 0.04 [48 ? 5]  0.019 ? 0.003 [35 ? 5]  464 90 ? 6 [35.4 ? 0.8]     0.053 ? 0.001 [59 ? 6] 0.006 ? 0.001 [25 ? 3]   9-Cl HOPDA H255A 270         0.204 ? 0.005 [79 ? 2]      436 19 ? 2 [5.1 ? 0.7]     0.202 ? 0.002 [95 ? 1]      457 18 ? 3 [3.5 ? 0.3]     0.203 ? 0.003 [96.5 ? 0.3]      474 17 ? 4 [2.8 ? 0.6]     0.203 ? 0.001 [97 ? 1]     H255E 270             0.0088 ? 0.006 [51 ? 1]  436 140 ? 30 [32 ? 4] 21 ? 2 [42 ? 6] 2.2 ? 0.3 [13 ? 1] 0.096 ? 0.003 [13.1 ? 0.3] H255Q 270 40 ? 10 [4.4 ? 0.8]     0.33 ? 0.02 [1.4 ? 0.2] 0.016 ? 0.001 [50.1 ? 0.7]  345 49 ? 2 [29 ? 2] 15 ? 2 [9 ? 1] 0.259 ? 0.003 [23.7 ? 0.5] 0.0174 ? 0.003 [39 ? 1]  370 48.3 ? 0.8 [30 ? 2] 18 ? 3 [9 ? 2] 0.264 ? 0.005 [23 ?1] 0.0176 ? 0.001 [37 ? 1]  476 49 ? 3 [45 ? 3] 10 ? 5 [7 ? 2] 0.29 ? 0.02 [19 ? 1] 0.018 ? 0.002 [28.3 ? 0.4] aFor the H255E and H255Q variants, the combined enzymatic and non-enzymatic rates of HPD ketonization (based on a decrease in A270 nm) were 0.004 and 0.008 s-1, respectively. For H255A, the rate was not accurately determined due to the decreased timescale of the experiment.     96  Interestingly, despite the accumulation of ESred in the H255A and H255Q variants, the rate constants associated with ketonization were reduced from ~100 to > 1000-fold for HOPDA and 9-Cl HOPDA. For H255E, in which ESred formation is rate-determining, the timescale for the ketonization reaction was > 2000-fold slower than in the WT enzyme. The disparity between the ability to generate ESred and the measured rate of HPD formation observed at 270 nm in the DxnB2 His255 variants has thus implicated this residue in the ketonization reaction. The specific role of His255 remains unclear since the effect of His255 substitution in DxnB2 was one to two orders of magnitude greater than the effects observed in BphDLB400, in which acylation (HPD formation) occurred at rates that were 5 to 40-fold slower than the WT. 3.5 DxnB2-mediated cleavage of 3-Cl HOPDAs 3.5.1 DxnB2:3-Cl HOPDAs complexes do not reveal substrate destabilization Although this work has emphasized DxnB2 as a model system for the study of substrate ketonization by the MCP hydrolases, the enzyme was initially characterized due to its remarkable ability to catalyze the hydrolysis of 3-Cl HOPDAs[39]. To better understand the manner in which DxnB2 hydrolyzes these recalcitrant PCB metabolites, the S105A variant was employed to investigate substrate destabilization of 3-Cl and 3,9,11 tri-Cl HOPDA, a model and biologically relevant PCB metabolite, respectively. The electronic absorption spectra of 3-Cl and 3,9,11 tri-Cl HOPDA were measured in the presence and absence of DxnB2 S105A (Figure 42). Binding of 3-Cl HOPDA to DxnB2 S105A resulted in a hyperchromically-shifted spectrum whereas the DxnB2 S105A:3,9,11 tri-Cl HOPDA complex revealed a 21 nm hypsochromic shift from 438 to 417 nm. Neither ES complex accumulated a detectable population of a red-shifted species, suggesting that the HOPDA   97 molecules are bound in a planar, non-productive conformation. Indeed, a planar binding mode was directly observed in an S105A:3-Cl HOPDA binary complex[117]. Figure 42. Electronic absorption spectra of 10 ?M (A) 3-Cl HOPDA or (B) 3,9,11 tri-Cl HOPDA in solution or complexed to 40 ?M DxnB2 S105A in KPi (I = 0.1 M), pH 7.5 at 25 ?C.  3.5.2 DxnB2 is able to access ESred faster than BphDLB400. In order to gain kinetic insight into the ability of DxnB2 to overcome the apparent inhibitory or stabilizing effect attributed to the 3-chloro substituent, hydrolysis of 3-Cl and 3,9,11 tri-Cl HOPDA was monitored under single turnover conditions, as described for other substrates. A PDA detector was utilized for the identification of intermediates (Figure 43, inset). For more accurate rate constant determination, reactions were monitored at single wavelengths. While decay of the HOPDA enolates (?max 432 or 438 nm) were readily monitored, the formation of 3-Cl HPD (?max ~244 nm) could not be distinguished from other underlying processes and the resulting multiphasic kinetic traces could not be reliably interpreted. To distinguish intra- and intermolecular events, additional reactions were performed in the presence of a viscogen, 30% sucrose (w/v).   98 The hydrolysis of both substrates by DxnB2 was biphasic (Figure 43 and Table 19). The first observable kinetic phase was attributed to ES complex formation, which occurred at 220 and 26 s-1 for the mono- and trichlorinated substrate, respectively. In both cases, the presence of sucrose slowed this bimolecular event by 1.3 to 2-fold. For 3-Cl HOPDA, ES  Figure 43. Results from representative stopped-flow experiments demonstrating the turnover of 4 ?M 3-Cl HOPDAs by 16 ?M DxnB2 (A) Hydrolysis of 3-Cl HOPDA monitored at 432 nm. (B) Hydrolysis of 3,9,11 tri-Cl HOPDA monitored at 417 and 438 nm. For each substrate, the time-dependent changes to the visible spectrum from 300 to 600 nm are inset. For clarity, 1/10 data points are shown.     99 Table 19. Kinetic values for the DxnB2-mediated hydrolysis of 3-Cl HOPDAs Substrate ? (nm) k1 (s-1) [Amp (%)] k2 (s-1) [Amp (%)] kcata 3-Cl 432 220 ? 20 [11.5 ? 0.3] 0.152 ? 0.001 [88 ? 2] 0.25 ? 0.1 + 30% sucrose 432 100 ? 10 [7.4 ? 0.3] 0.152 ? 0.001 [92.6 ? 0.7]  3,9,11-triCl 417 26 ? 1 [12.2 ? 0.6] 0.056 ? 0.001 [88 ? 2]   438 22 ? 2 [32 ? 1] 0.056 ? 0.001 [68 ? 1] 0.13 ? 0.1 + 30% sucrose 417 18 ? 2 [12 ? 1] 0.054 ? 0.001 [88 ? 2]   438 16.9 ? 0.2 [32.9 ? 0.8] 0.054 ? 0.001 [67 ? 2]  BphDLB400 + 3-Clb 432 15 [19] 0.0077 [81] 0.0089 adata from [39]. bdata from[86].  complex formation was accompanied by a small hypochromic perturbation whereas the spectrum of 3,9,11-tri-Cl HOPDA was hypsochromically-shifted by 21 nm as in the stable S105A complex. Following accumulation of the spectrophotometrically distinct ES complexes, the respective signals decayed and rate constants of 0.152 and 0.056 s-1 were modeled using exponential equations The observed insensitivity of 1/?2 to sucrose and the relatively large loss in amplitude (~90%), corresponding to the dissipation of the HOPDA enolate chromophore, support the assignment of this kinetic step to substrate ketonization as described for other substrates above. The absence of ESred from the kinetic traces suggests that formation of this intermediate is rate-limiting to the hydrolysis of 3-Cl HOPDAs. The observed 1/?2 values differed by only 2-fold from the reported kcat values[39], which may be explained by differences in the enzyme quantification methods employed. Nevertheless, substrate destabilization and ketonization still occur 20-fold faster than in BphDLB400, accounting for the increased specificity of the DxnB2 for 3-Cl HOPDAs. 3.5.3 Comparison of the tertiary and quaternary structure of MCP hydrolases  To investigate the basis of the difference in oligomeric state and the possible link to substrate specificity, the seven structurally characterized MCP hydrolases were compared using a suite of bioinformatic tools. SEC-MALS analysis indicates DxnB2 is a dimer   100 whereas BphDLB400 and HsaD are tetramers (Figure 27). CumD, CarC, and MhpC have also been described as dimers. However, the crystal structure of MhpC displays a tetrameric assembly of the same form as BphDLB400 and HsaD. BphDRHA1 purified as an octamer. To investigate the basis of the differences in oligomeric state, the seven structurally characterized MCP hydrolases were superposed and a structure-based sequence alignment was established (Figure 44). As noted previously for BphDLB400[81], all of the crystal structures present a two-fold symmetric dimer mediated in part by anti-parallel ?-interactions between edge strands (?8) of the monomer's core domains; the associated protein surfaces are defined as elements as the dimeric interface, the A-B interface identified for BphDLB400. Of interest here are the distinctions in sequence and structure that mediate assembly of the BphDLB400, HsaD, and MphC tetramers through additional interfaces that have been labeled as such: the tetrameric A-B' and A-A' interfaces.  The protein segment that connects the ?/?-hydrolase core and the N-terminus of the lid domain is referred to as the NC-loop in other enzyme families[46,118]. Prior to the NC-loop, a conserved LMG sequence (DxnB2 127-9) completes the ?6 strand in all known structures. The glycine residue, at position i, has a left-handed ? conformation, which creates a sharp turn as the backbone departs from the core. The next highly conserved sequence positions are a pair of branched non-polar residues at positions j and j+3 (DxnB2 residues 146 and 149); the length of the protein segment between i and j varies between 12 and 17 residues in the known structures. In all structures except that of CarC (wherein these residues are disordered and not defined by the electron density maps), the side chains of residues j and j+3 project from a helical segment of the lid domain into the NP-substrate binding subsite. Importantly, the residues between positions i and j are prominent at the tetrameric interfaces,    101 Figure 44. Sequence alignment resulting from the structural superposition of seven MCP hydrolases. The NC-loop is indicated by a dashed line. Secondary structural elements are numbered. The catalytic triad is coloured blue whereas the P-subsite and ?oxyanion hole? residues appear in green. The residues defined as i and j are coloured red. The helix labeled 11 might also be considered a pair of shorter helices joined by a turn.   102 and the sequences and structures of BphDLB400, HsaD, and MhpC present features distinguishing them from the dimeric MCP hydrolases (Figure 45). Figure 45. (A) Ribbon drawings of the BphDLB400 protomer (PDB ID: 2OG1, chain A). The regions that are involved either the A-B? and/or A-A? tetrameric interfaces, which include the NC-loop and lid domain helices, are colored green. (B) Superposition of the DxnB2 S105A:3-Cl HOPDA binary complex and BphDLB400 demonstrating differences in the NC-loop. (C) The apolar phenylalanine cluster conserved among MCP hydrolase tetramers. The dimer pairs (A-B pairs) are color coded blue and yellow.  In particular, the tetrameric MCP hydrolases of known crystal structure all share an F-X-P-X-P motif that begins at position i+9 and contributes to the tetrameric interface that associates a pair of lid domains, the A-B? interface. In fact, Fi+9 makes a visually striking contribution to both tetrameric interfaces as illustrated in Figure 45. Thus Fi+9 lies at the apex of a loop that protrudes away from the hydrolase and lid domains, and Fi+9 residues from all four monomers cluster to form a buried apolar core at the junction of the A-B' and A-A' interfaces. Residues immediately preceding Fi+9 as well as residues in the linker and the helix (?9) that follows the triad Asp also contribute to tetramer formation, but they do not belong to sequence motifs that distinguish dimers and tetramers. BphDLB400, HsaD, and MhpC belong to the same class, class I, on the basis of amino acid sequences(10). The sequences of other homologs in this class such as PcbD from Pseudomonas strain sp. DJ12 and Ro09014 F145a/b-hydrolase coreMCP hydrolase lidF145BAb6a5a5a6C  103 from R. jostii RHA1 show variations at the first position of the F-X-P-X-P motif. However, the oligomeric states are not known. In contrast to the protruding structure observed for the tetramers, the first few residues of the NC-loop in DxnB2 associate with helix ?10 of the hydrolase core domain as well as residues just prior to helix ?9 via several hydrogen bonds. The NC-loop of CumD is five residues shorter than that of DxnB2 and, while it follows a similar path, is not involved in the same suite of H-bonding interactions. However, CumD and several other class II enzymes share an F-P-XP-Q-X4-D motif that disrupts helix ?8 in CumD and places the Gln side chain from this motif in position to hydrogen bond with the backbone carbonyl and amide of NC-loop i+6 and i+8 residues, respectively. Indeed, class-specific lid domain architecture was apparent upon superposition (Figure 46). Figure 46. Superposition of MCP hydrolase lid domains from BphDLB400 (class I; green), CumD (class II; blue) and DxnB2 (class III; white). Some of the relevant helix-helix interactions are shown. Y152F204P44E193R195N140Y160S184F145NC-loopa5a8a7a6  104 In general, the beginning and end of the lid show greatest variability among the known MCP hydrolase crystal structures (Figure 44). Thus in many pair-wise or multi-protein comparisons it can be difficult to structurally align residues from the NC-loop and ?5 and/or residues within ?8 and the segment that links ?9 to ?10. In addition, the NC-loop and ?5 are prone to disorder and are sensitive to the occupancy of ligands in the active site. These uncertainties should be considered in evaluating sequence alignments, including those based on structures. By contrast, the segment of the lid spanning helices ?6 and ?7, residues 155-186 of DxnB2 and residues 164-196 of BphDLB400, is more readily superposed and aligned. The position of the helix ?5 is particular interesting when considering the observed differences in the rates of 3-Cl HOPDA hydrolysis by DxnB2 and BphDLB400. Accordingly, the lid domain architecture of all available DxnB2 S105A:MCP binary complexes was also compared by structural superposition. Despite the conserved interaction between helix ?5 and helix ?6 in all DxnB2 structures, mediated in part by the H-bonding interaction between Tyr152 and Ser184, the orientation of the N-terminus of the former helix is variable (Figure 47). More specifically, the lid domain in the S105A:3-Cl HOPDA binary complex appeared to adopt a more ?open? conformation than those observed in the S105A:HOPDA or S105A:5,8-diF HOPDA complexes[117,119]. Indeed, the different lid conformations were also reflected by two distinct substrate binding modes: a planar, and apparently non-productive binding mode was observed for 3-Cl HOPDA, whereas non-planar dienoate binding modes, which could be correlated to ESred by single crystal spectroscopy, were observed for HOPDA and 5,8-diF HOPDA[112,117]. Together, the analysis suggests that lid dynamics help to mitigate the hydrolysis of recalcitrant PCB metabolites.   105 Figure 47. Differences in the position of DxnB2 helix ?5. (A) Structural superposition of DxnB2 S105A in complex with 3-Cl HOPDA (white), HOPDA (yellow) and 5,8-diF HOPDA (orange). The difference in the positioning of helix ?5 is highlighted at the top. (B) Root mean square distance (RMSD) in ? between the C? atoms of DxnB2 S105A:3-Cl HOPDA and S105A:HOPDA binary complexes. The 5,8-diF HOPDA complex is nearly isomorphous with the latter. (C) RMSD in ? between chain A and B of a crystalline dimeric assembly of BphDLB400 (PDB ID: 2OG1). The largest observed differences are in helix ?9, which can be partially explained by uncertainties in C? position.   0 50 100 200 25015064200 50 100 200 2501506420residue numberresidue numberRMSD (A)RMSD (A)BCA  106 Chapter  4: DISCUSSION 4.1 The chemical mechanism of the MCP hydrolases  This thesis describes the chemical mechanism of the MCP hydrolases using two phylogenetically distinct homologues that share less than 30% identity and have distinct oligomeric states. First, stopped-flow analysis and chemical quench experiments on BphDLB400 provide evidence for an acyl-enzyme rather than a gem-diolate intermediate. Acylation was reproduced in DxnB2 using both HOPDA and 9-Cl HOPDA as substrates. These findings were complemented with a crystallographic study of BphDLB400 in which electron density extending from the catalytic serine in the H265Q variant could be unambiguously modeled with a benzoyl moiety[119]. Stoichiometric incorporation of 18O into benzoate from the BphDLB400-mediated turnover of HOPDA also provided indirect support for a covalent mechanism of catalysis. Finally, comparison of the hydro- and methanolysis of HOPDA and pNPB by BphDLB400 suggests that both C-C and C-O bond cleavage occurs via a covalent mechanism. Nevertheless, acylation during the former is histidine-independent, and a substrate-assisted mechanism of nucleophilic catalysis has been forwarded. Overall, the body of work overturned the paradigm for these enzymes in the literature.  The study of DxnB2 was particularly informative with respect to the mechanism of substrate ketonization by the MCP hydrolases. Solvent viscosity and isotope effects allowed for the identification of substrate ketonization as the rate-limiting step in the acylation reaction. Furthermore, the influence of the P-subsite on catalysis, specifically ESred formation and substrate ketonization, was clarified using stopped-flow and other spectrophotometric assays of systems at equilibrium.    107 4.1.1 An updated substrate-assisted nucleophilic mechanism of catalysis On the basis of the evidence presented herein, emphasizing the histidine-independent acylation, an updated mechanism of catalysis using BphDLB400 as the prototypical MCP hydrolase is proposed (Figure 48). To begin, the active site architecture, observed binding phenomena and changes in the visible spectra suggest preferential binding of a dianionic HOPDA isomer. The pKa of the catalytic histidine is unknown, although a possible interaction with Asn111 suggests that it may be protonated in the resting state. In this case, His265 may serve as a general acid to protonate the oxido group of a dianionic HPD produced during bond cleavage. Alternatively, a lone pair of electrons on the His265 N?2 may help to favor localization of negative charge in the bound dianion near the ?oxyanion hole?. Importantly, a gauche+ conformation around the C4-C5 bond is required to maintain all interactions between the substrate and the NP- and P-subsites. Ultimately, the substrate destabilization, empirically observed as ESred, is thought to be manifested as the increased reactivity at C5. Thus, the reactivity of ESred towards protonation at C5 replaces the His-Asp dyad in the activation of Ser112, thereby completing ketonization via a proton transfer. After ketonization, which generates a requisite electron sink, the resulting serinate can attack the C6-carbonyl, forming the first tetrahedral oxyanion intermediate that collapses to a Ser112 benzoyl-ester and HPD. If His265 N?2 were uncharged the lone pair may act to destabilize product binding, accelerating its release. Alternatively, it may act as general acid as described above. Nevertheless, upon binding of H2O to the acyl-enzyme, the His-Asp dyad acts as a general base to activate it. Nucleophilic attack on the acyl-enzyme generates the second tetrahedral oxyanion intermediate, which ultimately leads to benzoate production. At the end of the catalytic cycle, protonation of Ser112 regenerates the resting state enzyme.   108 Figure 48. The proposed MCP hydrolase mechanism. Although the pKa of His265 is unknown, it is drawn as neutral. P-subsite residues (not shown) play a critical role in the ketonization reaction. The top line of the mechanism can be considered as MCP hydrolase-specific chemistry whereas the bottom line represents chemistry shared by other ?/?-hydrolases. The second equilibrium arrow bridging the second and third panels on the top line represents an enzyme-substrate complex equilibrium (the two HOPDA dianions drawn are resonance forms). This equilibrium (for example, a subtle change in the active site microenvironment to induce substrate destabilization or conformational dynamic of the MCP hydrolase lid domain, vide infra) is thought to result in the formation of ESred and modulate it?s reactivity. A grey arrow is drawn in the second panel to indicate rotation about the C4-C5 bond resulting in a gauche+ conformation at the active site.  109 For the MCP hydrolases studied herein, the turnover of HOPDA is typically limited by the deacylation reaction. Alcoholysis experiments suggest that this step is limited by the rate at which the His-Asp dyad activates water. Interestingly, the mechanistic investigations of DxnB2 and BphDLB400 have also revealed that substrate destabilization and ketonization may be rate-limiting for the hydrolysis of environmentally relevant PCBs. More specific to BphDLB400 and other tetrameric homologues, which appear to be half-site reactive, a two-conformation model of catalysis can be inferred. The possibility that product release may limit acylation at the second set of active sites in these half-site reactive homologues remains unknown. Interestingly, two (Fo-Fc) difference electron density features approximately the size and shape of a phenylalanine side chain were observed in a nonpolar surface pocket bounded largely by the side chains of Ile153, Phe157, Phe204, and Leu205 near the periphery of the NP subsite in BphDLB400[119]. These features approach within 5 and 7 ? of the active site HOPDA? s phenyl group and may represent the phenyl groups of additional HOPDA molecules with disordered dienoate groups. Furthermore, an additional HOPDA density was observed at the opposite side of the active site in the BphDLB400 H265Q:HOPDA structure. Thus, it appears that BphDLB400 may have two routes for substrate/product entry/egress, which may also influence reactivity. 4.1.2 Evidence for a nucleophilic mechanism  This thesis presents three independent lines of evidence that support a nucleophilic mechanism of catalysis. First, an acyl-enzyme was observed by mass spectrometry in DxnB2, BphDLB400 WT and H265Q. Complementary X-ray crystallography experiments also revealed the nature of the reported Ser112-benzoyl intermediate[119]. Second, a single equivalent of 18O was incorporated into the benzoic acid product of HOPDA when   110 hydrolyzed by BphDLB400 in H218O. Finally, common kcat values and equal nucleophilic partitioning ratios were observed for the methanol-dependent C-C and C-O bond cleavage of HOPDA and pNPB by BphDLB400. 4.1.2.1 Direct evidence for an acyl-enzyme intermediate  The chemical quench data provide evidence for the occurrence of a covalent catalytic intermediate in a wild type MCP hydrolase. LC ESI/MS on intact protein samples of DxnB2 and BphDLB400 revealed modifications that were consistent with acylation. Furthermore, peptide matching from LC ESI/MS/MS of pepsin-digested reaction mixtures of BphDLB400 WT or H265Q and HOPDA substantiated the observations that were made on the intact protein samples. In both cases, benzoylation was uniquely identified as a modification to Ser112. LC ESI/MS on intact BphDLB400 reactions mixtures of varied age indicated that the modification occurs on a timescale commensurate with turnover. These data represent the first unambiguous identification of a catalytic intermediate on the MCP hydrolase pathway. All other reports on possible reaction intermediates have relied on indirect methods and variant enzymes[64,89]. In line with the rapid acylation of BphDLB400 H265Q, observed by MS, a Ser112-benzoyl species was trapped in a crystals of this variant upon exposure to HOPDA[119]. A 1.6 ? resolution crystal structure of H265Q incubated with HOPDA revealed clear electron density extending from the catalytic serine, which was unambiguously fit with a benzoyl moiety at full occupancy (Figure 48). The benzoyl-oxo formed H-bonds with the amides of each of the ?oxyanion hole? residues, Met113 and Gly42, highlighting a capacity to stabilize the formation of tetrahedral oxyanion intermediates during catalysis. The side chain of Gln265 was also oriented by polar contacts, including an H-bond between the Gln265 N?2   111 and O?2 of Asp237 of the catalytic triad. The Gln-Asp effectively constitute a dyad, with no apparent interactions with the Ser-benzoyl moiety, which is located more than 4.5 ? from the N?2 of Gln265. Additional H-bonding linking Gln265 O?1 and Asn111 N?2 was also observed. The stability of the crystalline acyl-enzyme can be attributed to the inability of the H265Q variant to activate water for deacylation. This was observed in solution as the rate constant for acylation of the BphDLB400 H265Q was ~10 s-1 although steady-state turnover or deacylation proceeded at only ~10-5 s-1. Figure 49. Ball-and-stick representation of the BphDLB400 H265Q:HOPDA complex active site, including the unbiased Fo ? Fc density (green) for the benzoyl adduct, contoured at 3 ?. H-bonding is indicated between the ?oxyanion hole? residues and the benzoyl moiety.  The identification of an acyl-enzyme in BphDLB400 and DxnB2 conflicts with previous reports on the mechanism of MhpC and BphDLB400. Two major arguments against covalent catalysis were based on the inability to observe an acyl-enzyme in MhpC[64] and the assignment of a 13C NMR signal to a gem-diolate species in both MhpC and BphDLB400 variants[89]. First, the failure to trap an acyl-enzyme intermediate in MhpC may be explained by the reaction and quenching conditions that were used. In some cases, the quenching reagents appear to be insufficient to halt turnover. In other cases, the reaction time appears to be too long, leading to complete turnover prior to the addition of a proper quenching reagent.  Asp237Gln265Ser112Met113Gly42  112 The assignment of the 13C NMR signal to a gem-diolate in samples of catalytically impaired hydrolase variants turning over HOPDA is weak for at least three reasons. In these studies, performed using [13C6]-HOPDA and each of BphDLB400 S112A, H265A or MhpC H114A[89], assignment of the signal at 128 ppm was based primarily on a predicted chemical shift value for a gem-diolate H-bonding to serine hydroxyl. However, the quality of the signal is compromised by the poor signal-to-noise ratio in the spectra. Moreover, a signal with the same chemical shift but having a sharper linewidth was present in the enzyme-free substrate sample (Figure 17) and was attributed to a hydrated ?-lactone, a HOPDA decay product. The authors did not rule out the presence of this decay product in the enzyme samples. Indeed, an enzyme:lactone complex would account for the observed increase in linewidths. Finally, the observation of a shared 13C NMR signal in samples of BphDLB400 S112A and H265A variants is inconsistent with the spectrophotometric evidence that each accumulates a distinct intermediate[66]. 4.1.2.2 Evidence for an acyl-enzyme intermediate from isotope studies  The stoichiometric incorporation of 18O from solvent into benzoate by BphDLB400 contrasts to a study of MhpC, in which 3 to 6% of its succinic acid product contained two 18O equivalents[64]. Unfortunately, the MhpC study did not report on the possibility of an enzyme-catalyzed 18O incorporation into succinate. Significantly, both aspartic and serine proteases have been reported to exchange 18O into their acid products despite using different catalytic mechanisms for hydrolysis have been reported[88,120,121]. While BphDLB400 did not catalyze solvent exchange into benzoate, differences between the systems may account for the differential incorporation of 18O into the respective reaction products. First, several distinct binding modes have been observed for succinate and malonate in complex with   113 MhpC H263A[65] and BphDLB400[86], respectively. Along these lines, small diacids interact with the hydrolase active site differently than monoacids such as benzoate. Furthermore, MhpC contains an additional active site histidine, His114, which contributes to catalysis and coordinates succinate in crystallo[65,113]. This residue, which corresponds to Ala116 in BphDLB400, may influence the reactivity of small diacids with MhpC. Interestingly, MhpC can activate hydroxylamine[114], and the rate of 5-carboxyvalero-hydroamic acid and ethanol formation from the reaction of ethanolamine and monoethyl adipate was ~50% slower in the H114A variant than in the WT. Finally, the resolution of the product detection method (GC/MS) cannot be used to define the position of the second 18O-incorporation event. In particular, either carboxylate group of the succinic acid product may be susceptible to exchange. However, the proposal that an 18O-incorporated gem-diolate could be misprocessed, released to the bulk solvent and dehydrated to form an 18O-substrate implies that a single carboxylate is doubly incorporated[64]. 4.1.2.3 Evidence for an acyl-enzyme from steady-state kinetic approaches  The detection of reaction intermediates by steady-state kinetics is indirect and relies on inference: relating results to direct observations and kinetic models of catalysis[95]. At a minimum, the detection of an intermediate from steady-state methods depends on: (i) the ability of an accumulated species to react with an acceptor whose concentration may be varied, and (ii) the generation of a common intermediate from substrates that share chemical substructure. Herein, water and methanol represent acceptors for a common Ser112-benzoyl intermediate produced during the turnover of HOPDA and pNPB. The similar methanol-dependent kcat or kobs values for HOPDA and pNPB hydrolysis, respectively, together with the common partitioning of these reactions between the hydro- and alcoholytic pathways   114 mirror what has been reported for other hydrolases that utilize a covalent mechanism of catalysis. Thus, constant partition ratios have been reported in serine proteases, such as chymotrypsin[122], and non-serine hydrolases, such as prostatic acid phosphatase[123], which are proposed to proceed via acyl-enzyme intermediates. For the BphDLB400-mediated methanol-dependent turnover of HOPDA, a simple kinetic model for a nucleophilic mechanism correctly predicted the observed increase in kcat and Km as well as the independence of kcat/Km[101]. Furthermore, the experimentally determined product ratios from the turnover of both HOPDA and pNPB, respectively, also matched the values predicted by the nucleophilic model of catalysis. Collectively, the data suggest that C-C and C-O bond hydrolysis proceed through a shared Ser112-benzoyl intermediate. This conclusion contradicts an earlier proposal that Ser112 acts as a hydrogen bond donor during C-O bond hydrolysis. This previously assigned role for Ser112 was deduced from a Hammet analysis using para-substituted pNPBs[93] and further supported using substrate docking experiments[115]. In particular, the Hammet plot was nonlinear, a feature that was attributed to a change in the rate-limiting step from ester bond cleavage to product release for substrates for which ? ? 0. An alternative explanation for the nonlinearity is that the activation of water for nucleophilic attack on the Ser112-benzoyl ester may become rate-limiting for substrates for which ? ? 0. This alternate explanation is consistent not only with the dependence of kcat on alcohol but also with the presence of a SKIE during turnover of esters[93]. Moreover, the strikingly similar kcat values observed for the DxnB2-mediated hydrolysis of several HOPDAs[39], supports a rate-limiting step that is independent of or shows little dependence on the MCP?s identity.     115 4.1.3 Evidence for rate-limiting hydrolysis The observed accelerations in the initial reaction velocities and the ability of BphDLB400 to utilize longer alcohols, such as 2-propanol, is consistent with a covalent mechanism of catalysis. The crystal structure of BphDLB400 S112A in complex with HOPDA reveals a complex suite of interactions[66,86], which is thought to reflect the high degree of active site residue conservation. Little residual volume is present within the substrate-occupied active site, suggesting that acylation and release of HPD precede the binding of additional nucleophiles such as water or alcohols. Indeed, product inhibition studies support a mechanism in which HPD is released before benzoate during turnover of HOPDA[81]. In contrast, significant movement of the MCP hydrolase lid domain would be required to bind additional molecules to a HOPDA-occupied active site. Considering the full suite of interactions within the enzyme-substrate complex, it is difficult to conceive how such lid movement would not disrupt productive substrate binding. The alcohol-dependent acceleration of v0 provides novel insight into the MCP hydrolase catalytic cycle by implicating deacylation as the rate-limiting catalytic step. To date, experiments in MhpC and BphDLB400 have assigned kcat to either release of the non-covalently bound product[83,93] or a conformational change associated with chemistry[81]. The observed increase in the rate of turnover in the presence of alternate acceptors suggests breakdown of the acyl-enzyme intermediate rather than product release limits catalysis. Indeed, the methanol-induced acceleration to kcat, 28 s-1 versus 6.5 s-1, is similar in magnitude to that reported for the turnover of an ester substrate by ?-chymotrypsin, 482 s-1 versus 144 s-1[122]. While the data do not unambiguously refute the possibility that product release is rate-determining, the observation of similar accelerations to v0 by different alcohols suggests   116 that this scenario is unlikely. Specifically, the limiting of turnover by product release would imply that methyl, ethyl, and propyl benzoates are released from the active site at an equal rate that is faster than that of benzoic acid. By contrast, assigning a chemical dependence to kcat is consistent with the presence of an SKIE during BphDLB400-mediated ester bond hydrolysis[93]. Interestingly, the lack of a viscosity effect reported for the single turnover of HOPDA by BphDLB400[81] implies that this rate-limiting deacylation at one set of active sites must precede acylation at the second set of active sites in the two-conformation model of catalysis.  4.1.4 The role of the catalytic histidine The catalytic histidine is the only universally conserved catalytic residue in the ?/?-hydrolase superfamily. As such, it has been implicated in both general base and nucleophilic mechanisms of catalysis. In the general base mechanisms proposed for the cofactor-less dioxygenases, the His-Asp dyad mediates a proton abstraction from a hydroxyl group on the substrate[63]. Cyanohydrin cleavage by the hydroxynitrile lyases is also initiated by a substrate deprotonation catalyzed by the triad residues[62]. In contrast, the generally accepted nucleophilic mechanism of catalysis posits a multifunctional role for the catalytic histidine, acting as (i) a general base for the activation of the serine nucleophile, (ii) a general acid for the protonation of the leaving heteroatom and (iii) a general base for the activation of water during the deacylation reaction. The role of the catalytic histidine in the MCP hydrolases is more obscure. Histidine-independent acylation was directly observed in BphDLB400. However, this histidine was required for acylation during pNPB turnover. Thus, while the catalytic triad of the MCP hydrolases behaves in a ?textbook? fashion for C-O bond cleavage, C-C bond cleavage   117 appears to proceed through a substrate-assisted mechanism of nucleophile activation. During this reaction, the catalytic histidine can be considered as a contributor to the P-subsite, which influences substrate binding, destabilization and ketonization. 4.1.4.1 The role of the catalytic histidine in the acylation reaction Kinetic characterization of histidine variants of BphDLB400 suggested that acylation was histidine-independent, reduced by only 5- to 40-fold compared to the WT. In the case of BphDLB400 H265Q, this was confirmed by mass spectrometry of a chemically quenched reaction mixture. Moreover, a BphDLB400 D237N variant behaved similarly to His265 variants during single turnover experiments[89], exhibiting a hyperchromic-shift at 434 nm and a slower rate constant for acylation. These initial studies were the impetus for the hypothesis of a substrate-assisted mechanism of nucleophile activation in the MCP hydrolases. Further inspection of the rates provisionally associated with acylation/HPD formation (limited by ketonization) in DxnB2 His255 variants revealed a more complex role for this catalytic residue. Specifically, acylation of the DxnB2 variants was more drastically perturbed, and the measured rates of HPD formation were reduced by up to 6000-fold. The moderate decrease in the rate of acylation observed in BphDLB400 His265 variants likely reflects cognate enzyme-substrate interactions. Thus, the catalytic histidine is integral to a properly organized P-subsite, which is required for efficient ketonization (see section 4.4). Differences in enzyme-substrate interactions, originating from both enzyme dynamics and active site microenvironment may also contribute to the large differences in the effect of histidine substitution in DxnB2 compared to BphDLB400. Along these lines, BphDLB400 is a tetramer whereas DxnB2 is a dimer, and the active site of BphDLB400 contains an additional polar interaction between its Tyr52 hydroxyl and the backbone carbonyl of Asn111 in the P-  118 subsite (Figure 50A). This additional interaction may account for stabilization of the P-subsite in BphDLB400 compared to DxnB2, in which a phenylalanine is present in the same position (Figure 50B). In the observed DxnB2 S105A:MCP binary complexes but not the ligand-free structures, the position of Asn104 (Asn111 in BphDLB400) and the organization of the P-subsite is the same as that observed in the BphDLB400 WT active site. Overall, the results indicate that the catalytic histidine contributes to but is not required for serine activation and acylation in MCP hydrolases, showing a remarkable divergence from the canonical mechanism typified by the serine proteases[73,121]. Figure 50. Ball-and-stick representations of BphDLB400 and DxnB2 active sites. (A) The BphDLB400 WT active site (PDB ID: 2OG1, chain A) in the absence of an MCP. (B) The DxnB2 S105A:5,8-diF HOPDA binary complex (coordinates provided by Bhowmik & Bolin, [117]). Possible polar interactions are indicated by dashed lines and the potential role of Tyr52 in the pre-organization of the P-subsite is highlighted by the comparison.  In the histidine-independent mechanism of nucleophile activation proposed for the MCP hydrolases, the substrate rather than the His-Asp pair acts as a general base to deprotonate Ser112, which also completes substrate ketonization. Interestingly, concerted movement of the catalytic His and the adjacent, conserved Trp residue away from the A BH265R190N51Y52L39G114S112H255R180N43F44V31G107S105AN111N104  119 nucleophile has also been observed in unpublished structures of DxnB2 upon substrate binding[117,119]. Thus, the kinetic and crystallographic data highlight the divergent catalytic mechanism of the Ser-His-Asp containing MCP hydrolases, which have evolved to utilize the chemical potential of the MCP substrates. 4.1.4.2 The role of the catalytic histidine in ESred formation The catalytic histidine is not absolutely required for the formation of ESred in the MCP hydrolases. Contrary to the rate constants reported for ketonization, which were greater for BphDLB400 than for DxnB2 catalytic histidine variants, the DxnB2 His255 variants accumulated ESred while the BphDLB400 His265 variants did not. Interestingly, the fact that a red-shifted species is observed in DxnB2 but is less reactive indicates that the catalytic His also has a role in the ketonization reaction. As discussed above, the histidine is fundamental for the maintenance of the P-subsite. The crystal structure of the BphDLB400 S112A/H265Q variant reinforces the importance of the catalytic histidine in forming ESred and maintaining the P-subsite. The comparison of the BphDLB400 S112A and S112A/H265Q variants provides unique insight into the nature of cognate enzyme-substrate interactions and disruption thereof. Thus, geometrically distinct HOPDA conformations are correlated to ESred or a species that possesses a hyperchromically-shifted absorption spectrum. As previously outlined, the conformation of ESred is non-planar and includes a gauche+ conformation about the C4-C5 bond. Ultimately, the observed conformation is thought to be poised for ketonization but is trapped in these structures due to the lack of the serine hydroxyl. In contrast, the X-ray structure of a BphDLB400 S112A/H265Q:HOPDA complex revealed a presumably non-productive binding mode[119]. Restrained refinements of HOPDA at the active site   120 suggested that a (3E,5Z)-2-oxo-6-oxido-dienoate was compatible with the X-ray data although the binding of a (2E,4E)-2-hydroxy-6-oxo- or a (3E)-2,6-dioxo isomer could not be eliminated based on the electron density observed for the substrate. The latter isomer is disfavored in light of the visible absorption spectra observed in solution: the enol has an expected ?max at 340 nm whereas an enolate spiecies, ?max 434 nm, is observed in BphD S112A/H265Q. In the absence of enzyme, the existence of a (3E,5Z)-2-oxo-6-oxido-dienoate was validated by observations of solvent 18O exchange into the substrate. The observed binding mode, in which the dienoate carbons, C1 to C6, are coplanar, is stabilized in part by a number of polar contacts (Figure 51A). In particular, the 6-oxido group is H-bonded to both the ?oxyanion hole? amides. At the P-subsite, the HOPDA carboxylate hydrogen bonds with Gly43, Arg190, Trp266 and a water molecule that also forms H-bond with Gly41, Asn51, and Asn111. In the absence of HOPDA, a second water molecule (not shown) replaces the interactions of a carboxylate oxo-group in the P-subsite. The orientation of the planar (3E,5Z)-2-oxo-6-oxido- isomer and its interaction with the active site differ significantly from the observed binding mode in the S112A:HOPDA complex, which has been fit to a non-planar (3E)-2,6-dioxo isomer (Figure 51B). The contribution of the catalytic His to the maintenance of the P-subsite is highlighted in structures of the BphDLB400 S112A/H265Q variant with and without an MCP. The substitution of His265 to glutamine results in significant perturbations to the P-subsite. First, the orientation of the two asparagine side chains, Asn51 and Asn111, are flipped compared to the conformation observed in the WT and S112A variant structures (Figure 52A). Typically, an H-bonding network is observed between His265, Asn111 and Asn51: the His N?2 acts as an H-bond donor to the Asn111 O?1, and the Asn111 N?1 interacts with the   121 O?1 of Asn51 (Figure 52B). Second, the electron density for three P-subsite residues (Asn51, Asn111 and Trp266) resulted in modeling two distinct conformers for each.   Figure 51. Ball-and-stick representations of the stable HOPDA binding modes observed in crystalline ES complexes. Polar contacts equal to or less than 3.4 ? are drawn as dashed lines. (A) The planar dienoate bound to BphD S112A/H265Q showing an additional water molecule in the P-subsite, below the dienoate plane (PDB ID: 3V1M). (B) The non-planar, electronically excited species, ESred, bound by BphD S112A (PDB ID: 2PUH).  These two distinct P-subsite conformations were observed at a 3:2 ratio and the first or major set of sites is thought to be occupied by the substrate (colored green in Figure 52A). The hydrogen bonding network in the crystal structure of the S112A/H265Q:HOPDA binary complex differs significantly from that reported for the S112A variant. The Gln265 O?1 is AArg190Gln265Asn111Asn51Gly42Gly43Met113Ala112Trp266BArg190His265Ala112Asn51Gly42Gly43Met113Trp266Asn111watO2  122 within H-bonding distance of the Asn111 N?1, and the Asn111 O?1 can interact with the N?1 of Asn51. This inversion of H-bond donors and acceptors, centered at Asn111, also results in the orientation of the Asn51 O?1 towards the Trp256 N?1, which is located at ~3.6 ?. Thus, in addition to the differences between the geometric isomers of HOPDA observed at the active site, the hydrogen bonding network at the P-subsite is significantly distinct from that correlated to ESred or observed in WT structures. The results from the crystallographic analysis further underline the importance of His265 in maintenance of the P-subsite.   Figure 52. BphDLB400 P-subsite variability upon substitution of His265. (A) The active site of a BphDLB400 S112A/H265Q:HOPDA complex (PDB ID: 3V1M). The active site conformations were modeled at a 3:2 ratio and are coloured green and purple, respectively. (B) The active site of a BphDLB400 S112A:HOPDA complex (PDB ID: 2PUH).  The distinct sets of P-subsite interactions that associated with either the planar and non-planar substrate binding modes in BphDLB400 suggest that changes to the substrate conformation are accompanied by changes at the enzyme active site. In line with this proposal and the defined role of the MCP hydrolase lid in substrate recognition and specificity, a role for MCP hydrolase lid movement in catalysis is probable (see section 4.5). Further structural studies on the DxnB2 S105A/H255X variants may resolve the nature of binding in the histidine-independent ESred species that are not observed in BphDLB400. Asn111Asn51Trp266Gln265Asn111Asn51Trp266Gln265A B  123 4.1.4.3 The role of the catalytic histidine in the deacylation reaction Overall, the data indicate that the MCP hydrolases utilize the His-Asp dyad to activate water for enzyme deacylation, a conserved role among serine hydrolases. Thus, while histidine-independent acylation was directly observed in BphDLB400, and inferred in DxnB2, steady-state hydrolysis was essentially abrogated in catalytic histidine variants of both enzymes. In fact, the turnover of HOPDA and/or 9-Cl HOPDA occurred on the timescale of hours rather than milliseconds or seconds, as observed for the WT enzymes.  4.1.5 Pre-steady-state kinetic behaviour of wild type enzymes and half-site reactivity  The existence of a pre-steady-state kinetic burst is consistent with a covalent mechanism of catalysis in which a process following acylation is rate-limiting. Remarkably, the pre-steady-state kinetic behaviour of BphDLB400 and DxnB2 differed with respect to the ratio of active enzyme during HOPDA cleavage. For the reaction of DxnB2 and HOPDA, 1:1 reactivity was observed during the first catalytic cycle. Interestingly, for 9-Cl HOPDA turnover, quantification of the amount of HPD produced during the burst phase indicated that only 67% of the enzyme was active. This peculiar reactivity may reflect either (i) an additional specificity determinant at C9 that invokes half-site reactivity or (ii) the relatively strong substrate inhibition by 9-Cl HOPDA, Ksi = 6 ? 2 ?M[39]. The accuracy of the values reported for kburst, kss and their associated amplitudes may be limited by the fact that the burst and steady-state rate constants differ by only a factor of five. Finally, full occupancy of the HsaD S114A active site was observed when HOPDA, a poor substrate, was incubated with crystals of this enzyme[41], suggesting that non-cognate substrates may also perturb reactivity.  The magnitude of the pre-steady-state burst in BphDLB400 is consistent with the   124 proposed two-conformation model of catalysis; quantification of HPD produced during the pre-steady-state burst corresponded to only half of the available hydrolase active sites. Moreover, the relative ratio of modified to unmodified enzyme and peptides as measured by MS from chemically quenched reactions was also more consistent with half-site reactivity. This kinetic feature of the MCP hydrolases was first proposed based on the half-site occupancy of MhpC complexed with an inhibitor[69] and has since been observed in two HsaD:MCP complexes[41]. Unfortunately, crystallographic symmetry of all BphDLB400:MCP complexes requires equivalence across all active sites, which limits information regarding differential active site binding affinities[66,81,86]. Nevertheless, in some enzymes, substrate binding is stoichiometric but reactivity is not; recently LuxG was shown to be a full-site binding, half-site reactive flavin reductase[124]. Here, analysis of substrate binding to DxnB2 P-subsite variants, specifically N43D/S105A, was indicative of 1:1 substrate binding. For BphDLB400, the two-conformation model provides a more satisfactory explanation for the biphasic formation of HPD observed under single turnover conditions. In the two-conformation model, deacylation at one set of sites allows the second set to become active. In contrast, the kinetic behaviour of DxnB2 is less defined, changing with the substrate. However, the relevance of this aberrant behaviour is limited to in vitro studies of this enzyme since the physiologically relevant substrate is unknown.  4.2 Distinct kinetic features of DxnB2 wild type Although the half-site reactive behaviour of BphDLB400 is an interesting kinetic phenomenon, the associated kinetic complexity had contributed to a minimal assignment of observed kinetic steps during C-C bond hydrolysis. Moreover, the observation of biphasic turnover also influenced arguments in support of a general base mechanism, assigning   125 ketonization and HPD formation to distinct observable steps under single turnover conditions[65,83]. In the context of the covalent mechanism, the biphasic decay of ESred can be attributed to HPD formation/acylation at distinct catalytic sites. Nevertheless, the apparent two-conformation behaviour of BphDLB400 limits information on the ketonization reaction and the determinants thereof. The timescale of acylation observed by mass spectrometric studies on BphDLB400 allowed for the provisional assignment of 1/?2 and 1/?3 to this process. The later phase also represents the rate-determining deacylation of the first set of active sites, 1/?3 ~kcat ~6.5 s-1. However, the determinants of 1/?2 are unclear, and are complicated by its sensitivity to solvent viscosity and temperature in BphDLB400[81]. Thus, the mechanisms of substrate destabilization, ketonization, and ultimately acylation in the MCP hydrolases were gleaned from studies of DxnB2 as discussed below. 4.2.1 Defining observable kinetic steps in DxnB2 Pre-steady-state kinetic experiments indicated that the DxnB2-mediated turnover of HOPDA could be used to better define the mechanism of acylation. The reaction was apparently 1:1 with respect to protomer concentration, and therefore devoid of the kinetic complexity of the two-conformation model (Figure 53). The DxnB2-catalyzed hydrolysis of HOPDA is defined by the spectrophotometric observation of three kinetic steps. The first two, ESred formation (1/?1 > 500 s-1) and decay or enzyme-acylation (1/?2 ~69 s-1), could be monitored under single turnover conditions at 487 and/or 270 nm. The third step, deacylation, was reported from steady-state characterization of the enzymatic reaction, kcat ~0.4 s-1[39]. In particular, the rate constant for the second step was insensitive to solvent viscosity and had an SKIE of ~2.5. The involvement of a rate-limiting proton transfer in this   126 step further suggests that it represents substrate ketonization and that the generation of an electron sink limits acylation in the MCP hydrolases. Figure 53. Kinetic scheme for DxnB2 hydrolysis of HOPDA showing the underlying chemical process that has been assigned to observable transient state kinetic phase that was previously assigned to acylation/HPD formation in BphDLB400.   While the SKIE on 1/?2 indicates that substrate ketonization limits acylation in the MCP hydrolases, the specific source of the reactive proton remains unknown. In the context of the proposed substrate-assisted mechanism, the proton is abstracted from the Ser105 hydroxyl, which is expected to be deuterated under the reaction conditions[125]. In spite of the consistency between the proposed mechanism and observed effect, several other proton sources exist within the reaction media, including other exchangeable protein sites, the solvent itself and the substrate?s own protons, which can exchange with the solvent.  4.2.2 A correlation between substrate basicity and ketonization by DxnB2  Kinetic characterization of the substrate ketonization reaction using a series of HOPDA analogues supported the notion of substrate-assisted nucleophile activation in the MCP hydrolases. An extended Br?nsted analysis revealed a positive linear correlation between substrate basicity and the rate constant for substrate ketonatization. Here, substrate basicity is defined by the enol:enolate equilibrium, which is empirically measured as   127 pKa2[37], and substrate ketonization is measured as 1/?2 during a single turnover of the tested substrates (HOPDA, 5-Cl, 8-Cl, 9-Cl and 5,8-diF ). More explicitly, the results suggest that the reactivity of the substrates towards protonation at C5 mirrors the enol:enolate equilibrium (protonation of a hydroxyl). The positive linear correlation is in agreement with the assignment of 1/?2 to a proton transfer step. The observed slope, ?nuc ~1, is particularly interesting since White and Jencks interpreted a similar analysis of succinyl-CoA:3-ketoacid CoA transferase to be representative of a change in charge from -1 to 0 when the substrate reached the transition state[126]. This change in charge is expected to occur upon substrate ketonization in the MCP hydrolases. The observable ?nuc also argues against a concerted mechanism for acylation in the MCP hydrolases. Instead, the results indicate that ketonization and acylation occur in a stepwise mechanism whereby the substrate ketonization precedes nucleophilic attack at the carbonyl. Interestingly, the substrate?s reactivity or inherent ability to stabilize negative charge may ultimately limit catalysis. For example, this protonation reaction was determined to be rate-limiting for the turnover of 5,8-diF HOPDA, which is characterized by a low pKa2. 4.2.2.1 Substrate-derived determinants of biphasic kinetics  Curiously, substitution of the phenyl ring altered the kinetic behaviour of DxnB2. More particularly, the DxnB2-mediated cleavage of HOPDA or dienoate-substituted analogs was biphasic under single turnover conditions, while the hydrolysis of ring-substituted analogs occurred in three distinct transient kinetic phases. In the case of 8 or 9-Cl HOPDA, the large amplitudes associated with 1/?2 and 1/?3 suggest that both phases can be assigned to acylation. In this way, the ring-substituents appear to invoke sub-stoichiometric turnover that is somewhat reminiscent of the half-site reactivity or, more so, pNPB turnover observed in   128 BphDLB400. The kinetic behaviour under single turnover conditions is also consistent with the pre-steady-state burst of HPD formation observed for 9-Cl HOPDA. In contrast for 5,8-diF HOPDA, the small change in absorbance associated with 1/?2, accounting for only 2.6 % of the total amplitude, and the absence of a change in absorbance at 270 nm, indicate substrate destabilization was biphasic rather than product formation. The rate constant associated with substrate ketonization was reduced > 500-fold for 5,8-diF HOPDA, and the relatively slow turnover may have also influenced the number of observable transients preceding chemistry. Thus, ESred decay and acylation was monophasic for 5,8-diF HOPDA instead of the biphasic behaviour for 8-Cl and 9-Cl HOPDA. Again, it is unclear whether the observation of multiphasic ESred decay can be correlated to a two-conformation model of catalysis or if the kinetic behaviour reflects the fact that HOPDAs are not the cognate substrate of DxnB2. 4.3 Insight into the identity of ESred  Prior to this study, two proposals on the identity of ESred had been forwarded. In one, the spectral perturbation was ascribed to bond strain in an enzyme-bound (2E, 4E)-2-oxido-6-oxodienoate[65,81]. This strain hypothesis was rationalized by analogy to the rhodopsin family, which covalently bind retinal through a Schiff base at a lysine residue, forming a cationic species. The mechanism of the red-shift in rhodopsins is still debated, and includes arguments for conformational and electrostatic manipulation of the protein-bound chromophore[127]. Recently, extreme bathochromic shifting, > 200 nm, was observed from rationally designed variants of human cellular retinal binding protein II (hCRBPII), and complementary crystallographic studies revealed apparently little dependence on conformational strain[128]. Instead, sequestration of the chromophore from bulk solvent and balanced distribution of the cationic charge across the polyene, mediated by evenly   129 distributed electrostatic potential in proteins that lacked a cationic-counter ion interaction, were largely responsible for the spectral perturbations in hCRBPII variants. The strain hypothesis has also been challenged by crystallographic analysis of the BphDLB400 S112A:HOPDA or DxnB2 S105A:HOPDA binary complexes. These data, which have been correlated to that observed in solution by single crystal spectroscopy, suggest that ESred has a geometric conformation that is most consistent with a (3E)-2,6-dioxo isomer[81,119]. Unfortunately, studies on the electrostatic potential of the MCP hydrolase active site are lacking. A second hypothesis for the identity of ESred which is more consistent with the results of restrained refinements of the dienoate, is polarization of the ? electrons of an ?,?-unsaturated ketone. In fact, binding of an inhibitor to crotonase resulted in a 90 nm bathochromic shift[85]. Interestingly, the introduction of a negatively charged residue into the P-subsite of DxnB2 (N43D) did not dramatically alter the identity of ESred: while the relative population of species observed at ~475 and 500 nm in ESred for the N43D/S105A and S105A variants were inverted, the ?max values were not significantly shifted. While these results argue against a uniform electrostatic environment and polarization, this interpretation is complicated by the timescale of ESred formation in the P-subsite variants, which is drastically reduced. In contrast to the contributions of the P-subsite, the identity of ESred was very similar between DxnB2 and BphDLB400, which possess distinct NP-subsites, arguing for a minimal contribution from this site in ESred formation. Here, a third proposal is presented that posits ESred may represent an enzyme-stabilized carbanionic species with charge localized at C5. An enzyme-stabilized C5 carbanion is compatible with the results of the restrained refinements of MCPs to (3E)-2,6-  130 dioxo isomer. Mechanistically, such a carbanion is also consistent with the negative charge that must accumulate on C5 for substrate ketonization. Although enzymatic carbanion formation is typically energetically unfavorable, it is only ~9 kcal/mol for an enolate[129], and delocalization into a carbonyl stabilized by coordination to a Lewis acid, such as an ?oxyanion hole?, is a common catalytic motif[77]. Additionally, the requisite energy could be easily supported by the large number of polar and nonpolar contacts provided by the NP- and P-subsites (Table 24, Appendix V). In BphDLB400 and DxnB2, the theoretical change in energy associated with the observed electronic transitions characteristic of ESred versus the free enolate in solution range between ~7 to 12 kcal/mol. Indeed, these values are comparable to the energy required to stabilize an enolate carbanion. Finally, the spectrophotometric contribution of a carbanion is also consistent with a red-shifted intermediate, explained by an n to ?* transition. A similar, 50 nm bathochromic shift was observed between different species of triphenylmethyl lithium: the contact ion pair exhibited a ?max at 446 nm whereas the absorption spectra of the solvent separated ion pair was centered around 496 nm[130]. In nature, the quinoid intermediates of PLP-dependent enzymes provide prevalent examples of carbanionic intermediates with red-shifted absorption profiles. For example, the quinonoid intermediates of the C-C bond hydrolase kynureninase are characterized by a ?max between 490 and 500 nm, which are shifted ~80 nm from their respective external aldimine intermediates[131,132]. Despite consistency with the observed data, the absence of any direct evidence makes any assignment of ESred provisional. 4.3.1 Preliminary experimental investigation of ESred by NMR  Two indirect approaches were used to examine the electronic nature of ESred in DxnB2. First, 19F NMR experiments hinted at the development of sp3 character at C5 in a   131 DxnB2:5,8-diF HOPDA binary complex by comparing the chemical shifts of F5 to purchased standards. Nevertheless, these results provide minimal insight into the electronic nature of ESred as the assignment of the hybridization state of C5 via the 19F probe is ambiguous. Conversely, the major signals observed in the 1H13C HSQC spectrum of uniformly 13C-labeled HOPDA complexed to DxnB2 S105A were at chemical shifts values between 100 to 130 ppm, suggesting that the system of ? electrons is intact in ESred. Future experiments should include the direct investigation of a site-specifically labeled [13C5]-HOPDA to clarify the electronic nature of C5 in ESred. 4.3.2 EI GC/MS analysis of ESred decay products The reactivity of ESred, specifically at C5, was particularly intriguing. The reaction of a carbanionic intermediate with molecular oxygen, an electrophile, is an expected result. However, in the absence of heavy isotopes, the observations do not uniquely identify O2 as the reactive species. Second, and most importantly, the resolution of the experiment is not sufficient for the identification of the product as a 1-(3-chlorophenyl)-2-hydroxyethanone. Isolation of the ESred decay product for chemical characterization by NMR in conjunction with higher resolution MS analyses of products formed in the presence of 18O2 and H218O should facilitate confident identification in the future. Additional control experiments that include the decay of an ES complex that is not red-shifted should also be performed. Finally, in the MCP hydrolases, the apparent sensitivity to O2 may be further complicated by the possibility of an enzymatic affinity for and/or ability to utilize O2. Specifically, while the cofactor-less dioxygenases of the ?/?-hydrolase superfamily share only ~10% amino acid identity with DxnB2, comparison of their three-dimensional structure reveals a high degree of conservation. Structural superposition of DxnB2 and 1-H-3-hydroxy-4-oxoquinaldine 2,4-  132 dioxygenase (HOD) from Arthobacter nitroguajacolicus R?61a and 1-H-3-hydroxy-4-oxoquinoline 2,4-dioxygenase from Pseduomonas putida 33/1 revealed root mean square deviations between 2.9 to 3.1 ? across 217-224 structurally conserved positions (~75% of the DxnB2 structure). Thus, the affinity and ability of DxnB2 and other MCP hydrolases to bind and utilize O2 should be carefully characterized before concluding on the nature of ESred. This is especially pertinent since substrate deprotonation to form an anionic intermediate is thought to precede O2 activation and O2 has been proposed to occupy the ?oxyanion hole? in these cofactor-less dioxygenases[63]. 4.4 The role of P-subsite residues in catalysis  The current studies greatly extend previous investigations of the P-subsite residues. More particularly, previous studies of MhpC reported on the respective roles of three P-subsite residues: (i) Trp264 in substrate binding, (ii) Arg188 in MCP ketonization, and (iii) Asn109 in positioning the active site loop containing Ser110[65]. Substitution of each of these residues resulted in enzymes with greater C-S and/or C-O bond cleavage activity than the WT[114]. Nevertheless, the aforementioned roles for the P-subsite residues were forwarded based on steady-state kinetic or specific activity measurements, and the interpretation was guided by limited structural data. The experiments presented here better resolve the individual roles of Arg180 and Asn43 in DxnB2, although structural data of the P-subsite variants in complex with HOPDA and/or 9-Cl HOPDA would greatly augment these kinetic data. Finally, the contribution of the catalytic histidine to the P-subsite was identified and has been discussed above.     133 4.4.1 Kinetic behaviour of P-subsite variants The role of Asn43 and Arg180 in substrate destabilization and ketonization is consistent with structural data showing that each directly contacts the dienoate moiety and/or other P-subsite residues. In fact, structural data for several MCP hydrolases suggests that Asn43 may also be involved indirectly in catalysis by properly orienting His255 during turnover through a conserved interaction with Asn104. In the crystal structures of DxnB2 S105A:HOPDA or 5,8-diF HOPDA complexes[117,119], which display the productive non-planar substrate binding mode characteristic of ESred, the Asn43 O?1 is within H-bonding distance of the Asn104 N?2, and the Asn104 O?1 may or may not interact with the His255 N?1, depending on the residue?s protonation state, which is unknown. In addition, Asn43 N?2 is within H-bonding distance of the HOPDA carboxylate in crystalline complexes that are characteristic of ESred[41,65,66,117,119] but not in complexes that are characterized by planar HOPDA dienoate binding[86,119]. For Arg180, the gunidinium group also H-bonds with the substrate carboxylate in ESred or with both the carboxylate and C2-oxo in planar conformations, excluding the the DxnB2 S105A:3-Cl HOPDA complex[117].  Disruption of polar contacts between the P-subsite residues and the substrate perturbed the rate of HOPDA hydrolysis. The removal of a positively charged residue, R180M and R180Q, or the addition of a negatively charged residue, N43D, significantly reduced hydrolytic activity. In the case of Arg180 variants, the >100-fold increases in Km values was due to reduced substrate affinity, reflecting the role of the guanidinium group in carboxylate binding. In contrast, the N43D variant bound and destabilized the substrate, shown by ESred formation upon binding to the N43D/S105A variant. The measured kcat/Km values for N43D were reduced by 15000- and 4300-fold for HOPDA and 9-Cl HOPDA,   134 respectively, compared to the WT enzyme. Interestingly, while the Asn43 is typically located ~4 ? from Arg180 in MCP hydrolase structures, the closest distances between the two residues have been observed in class III homologues, and is as little as 3.6 ? in the structure of CarC (PDB ID: 1J1I). Accordingly, replacement of Asn43 with an aspartic acid may result in the introduction of a new interaction between Asp43 and Arg180. While this variant interaction does not appear to disrupt substrate destabilization in the N43D/S105A variant, the timescale of ESred formation was in fact reduced (see section 4.4.2).  The greater activity of the second group of P-subsite variants reflects the more conservative nature of the amino acid substitutions: N43A, N43H and R180K. In each case, the substitution reduced the specificity of the enzyme for cleavage and the degree of substrate inhibition. For example, substrate inhibition was ~50-fold lower for R180K than in the WT for 9-Cl HOPDA. In light of the strong substrate inhibition observed for the DxnB2-mediated hydrolysis of ring-substituted HOPDAs (typically Ksi < 10 ?M) and the relatively modest 10-fold decrease in substrate specificity for R180K, this variant may be an interesting target for directed evolution experiments aimed at expanding the substrate range of the MCP hydrolases. 4.4.2 The role of the P-subsite in substrate destabilization and ketonization The full suite of P-subsite interactions is required for efficient C-C bond cleavage. In particular, the ability of the enzyme to destabilize the substrate is critical for ketonization. The ability of S105A/P-subsite variants to accumulate ESred further highlights the importance of this intermediate on the reaction pathway. Moreover, the absence of ESred from kinetic traces under single turnover conditions argue that substrate destabilization is at least partially rate-limiting for ketonization, and thus, acylation in the N43A, N43D and R180K variants.   135 Overall, these results highlight the evolution of a finely tuned active site with extensive enzyme-substrate interactions. 4.5 Structural determinants in MCP hydrolases  The structural and catalytic role of the P-subsite, including the catalytic histidine has been discussed within the context of differences observed in HOPDA turnover by BphDLB400 and DxnB2. Upon the analysis of the available structures, several other distinguishing features of phylogenetically diverse MCP hydrolase subfamilies became evident and may be linked to improved specificity of DxnB2 towards recalcitrant PCB metabolites. 4.5.1 Differences in MCP hydrolase structures and activity. The structural analyses of the MCP hydrolases further support a significant contribution of the NC-loop to catalysis. This loop, which joins the core and lid domains, has been proposed to contribute to catalysis in several other ?/?-hydrolase families including the lipases[46], haloalkane dehalogenases[133] and epoxide hydrolases[118]. Furthermore, substitution of the NC-loop was required to convert an esterase into an epoxide hydrolase[47]. In the MCP hydrolases, domain-swapping experiments between BphD (BphDLA4) and MfphA from Dyella ginsengisoli LA-4 further established that the identity of the lid domain is a strong determinant of substrate specificity[134], and a single amino acid substitution located in the NC-loop of BphDLA4 reduced kcat/Km for HOPDA by nearly 300-fold[135]. Molecular dynamics simulations have also been used to argue the relevance of NC-loop flexibility in the haloalkane dehalogenases[133] and significant ?5 helix motions were observed in the Candida antarctica lipase B (CALB)[136]. Interestingly, while the CALB and MCP hydrolase lid domains are distinct, the first helix of each, which follows the NC-loop, occupy very similar three-dimensional space in relation to the ?/?-hydrolase core.   136 Indeed, the sum of the available data for the ?/?-hydrolases sets a precedent for the involvement of the NC-loop in substrate binding and catalysis by the MCP hydrolases. The identification of the NC-loop as a determinant of MCP hydrolase oligomeric state provides a further example of the ability of the ?/?-hydrolase core to accommodate family-specific insertions to determine substrate and chemical specificity[42,44]. Thus, comparison of the seven structurally characterized MCP hydrolases representing three different oligomeric states revealed high structural variation for the NC-loop. The NC-loop residues appear to be most highly conserved among the class I enzymes, and to at least partially mediate tetramer formation. In particular, the NC-loop contributes to two distinct tetrameric interfaces and the phenylalanine residue of its F-X-P-X-P motif lies at the core of the biological assembly. Moreover, differences in the NC-loop are propagated to the lid domain architecture: the position of and interactions involving the first helix of the MCP hydrolase lid, ?5, are not conserved between the phylogenetically distinct classes. Each class possesses a distinguishing set of intra-lid interactions, which are likely to mitigate substrate specificity. 4.5.2 The DxnB2 lid domain and specificity for 3-Cl HOPDAs The single turnover data together with the proposed catalytic mechanism of MCP hydrolases indicate that the higher specificity of DxnB2 for 3-Cl HOPDAs (kcat/Km) versus BphD is due to higher rate of ESred formation in the former. More specifically, the hydrolysis of HOPDAs by DxnB2[137] and BphDLB400[66,81,86] is defined by three observable steps for which the rate constants have been determined: (i) ESred formation (> 500 s-1); (ii) ESred decay (50 to 70 s-1), which is coupled to acylation/HPD formation and limited by the ketonization reaction; and (iii) deacylation (kcat ~0.4 to 6.5 s-1). In contrast, the hydrolysis of 3-Cl HOPDAs by these enzymes is characterized by just two observable processes:   137 (i) formation of an alternative, presumably non-productive, binding mode named ESe in BphDLB400[86] and characterized by a hypo- and/or hypsochromically-shifted absorption spectrum in DxnB2; and (ii) ESe decay, which is coupled to HPD formation and corresponds to kcat. The failure to detect ESred during 3-Cl HOPDA cleavage indicates that it is consumed at a rate that precludes detection. Thus, the similarity between the observed rate constants for HPD formation and steady-state turnover indicate that ESred formation is rate-limiting to catalysis[39]. For 3-Cl HOPDA, this rate constant (1/?2) is at least 3000-fold slower than for HOPDA in DxnB2[138]. However, formation of ESred from 3-Cl HOPDA in DxnB2 (0.152 s-1) is 20-fold faster than in BphDLB400 (0.0077 s-1), accounting for the increased reactivity of the former towards recalcitrant PCB metabolites. Interestingly, ESred formation during turnover of 3,9,11-triCl HOPDA was only ~3-fold slower than 3-Cl HOPDA. This small reduction is in line with the general observation that ring-substituents are minor contributors to the substrate specificity of BphDLB400 and DxnB2 with the exception of 9-Cl HOPDA for the latter enzyme[37,39]. The variable disorder in helix ?5 observed in crystal structures of DxnB2 suggests that this region has increased mobility relative to other MCP hydrolases with the exception of CarC. Such movement may be restricted in the tetrameric enzymes based on the location the tetrameric interfaces. Interestingly, superposition of three DxnB2 S105A:HOPDA binary complexes reveals a difference in the position of helix ?5 of up to ~6 ? based on placement of the C? atoms. The position of helix ?5 drastically affects the form of the NP-subsite. Specifically, it carries Leu146 and Val149, which are further removed from the substrate in crystal structures of DxnB2 than in BphDLB400[86,117]. Accordingly, the comparatively limited restraint placed by the DxnB2 NP-site on the substrate is also likely to contribute to   138 the relative increase in hydrolytic activity towards recalcitrant PCB metabolites. Thus, the possible mobility of this helix may account for the increased accessibility to ESred in DxnB2 versus BphDLB400. The high structural variance of the NC-loop contributes to the ability of DxnB2 to overcome the stabilization of 3-Cl HOPDAs in non-productive, planar dienoate binding modes. Moreover, the NC-loop is prominent at MCP hydrolase tetrameric interfaces suggesting that it at least partially stabilizes the oligomeric state of class I family members. The attributes of this loop have implications for both environmental bioremediation and the design of therapeutics. Replacement of BphDLB400 (or other bona fide BphD enzymes) by DxnB2 in the Bph pathway may alleviate the metabolic bottleneck caused by 3-Cl HOPDAs. The dimeric MCP hydrolases could also represent better starting points for future enzyme engineering efforts. The plasticity of the NC-loop should also be considered in designing inhibitors of HsaD, which appears to be required for survival of Mycobacterium tuberculosis in macrophages[139]. Finally, while the results further establish the catalytic adaptability of the ?/?-hydrolase superfamily and highlight the influence of lid architecture within a specific enzyme family, the correlation of loop and lid movements to catalysis is relevant to a broader understanding of enzyme dynamics[140]. 4.6 Concluding Remarks  This thesis described the chemical mechanism of the MCP hydrolases. Thus, considering the conservation of the P-subsite throughout the family, it provides an outline for the development of mechanism-based inhibitors for the M. tuberculosis enzyme. In addition, through the kinetic characterization of two homologues of differing oligomeric state, DxnB2 and BphDLB400, substrate ketonization was found to be rate-limiting for the acylation   139 reaction. A more definitive role for the P-subsite, including the catalytic histidine, in the ketonization reaction has been established. Finally, while the identity of ESred remains unknown, progress towards its identification has been reported, and future experiments have been proposed.     140 REFERENCES 1. Boerjan W, Ralph J, Baucher M (2003) Lignin biosynthesis. Annu Rev Plant Biol 54: 519-546. 2. Carmona M, Zamarro MT, Blazquez B, Durante-Rodriguez G, Juarez JF, et al. (2009) Anaerobic catabolism of aromatic compounds: a genetic and genomic view. Microbiol Mol Biol Rev 73: 71-133. 3. Vaillancourt FH, Bolin JT, Eltis LD (2006) The ins and outs of ring-cleaving dioxygenases. Crit Rev Biochem Mol Biol 41: 241-267. 4. Dagley S (1978) Microbial catabolism, the carbon cycle and environmental pollution. Naturwissenschaften 65: 85-95. 5. Mason JR, Cammack R (1992) The electron-transport proteins of hydroxylating bacterial dioxygenases. Annu Rev Microbiol 46: 277-305. 6. Palfey BA, McDonald CA (2010) Control of catalysis in flavin-dependent monooxygenases. Arch Biochem Biophys 493: 26-36. 7. Harayama S, Rekik M (1989) Bacterial aromatic ring-cleavage enzymes are classified into two different gene families. J Biol Chem 264: 15328-15333. 8. Gibson J, C SH (2002) Metabolic diversity in aromatic compound utilization by anaerobic microbes. Annu Rev Microbiol 56: 345-369. 9. Olivera ER, Minambres B, Garcia B, Muniz C, Moreno MA, et al. (1998) Molecular characterization of the phenylacetic acid catabolic pathway in Pseudomonas putida U: the phenylacetyl-CoA catabolon. Proc Natl Acad Sci U S A 95: 6419-6424. 10. Rather LJ, Knapp B, Haehnel W, Fuchs G (2010) Coenzyme A-dependent aerobic metabolism of benzoate via epoxide formation. J Biol Chem 285: 20615-20624. 11. Lunt D, Evans WC (1970) The microbial metabolism of biphenyl. Biochem J 118: 54P-55P. 12. Denef VJ, Park J, Tsoi TV, Rouillard JM, Zhang H, et al. (2004) Biphenyl and benzoate metabolism in a genomic context: outlining genome-wide metabolic networks in Burkholderia xenovorans LB400. Appl Environ Microbiol 70: 4961-4970. 13. Furukawa K, Miyazaki T (1986) Cloning of a gene cluster encoding biphenyl and chlorobiphenyl degradation in Pseudomonas pseudoalcaligenes. J Bacteriol 166: 392-398. 14. Furukawa K, Tomizuka N, Kamibayashi A (1979) Effect of chlorine substitution on the bacterial metabolism of various polychlorinated biphenyls. Appl Environ Microbiol 38: 301-310. 15. Baker P, Seah SY (2012) Rational design of stereoselectivity in the class II pyruvate aldolase BphI. J Am Chem Soc 134: 507-513. 16. Hurtubise Y, Barriault D, Powlowski J, Sylvestre M (1995) Purification and characterization of the Comamonas testosteroni B-356 biphenyl dioxygenase components. J Bacteriol 177: 6610-6618. 17. Faroon OM, Keith LS, Smith-Simon C, De Rose CT (2003) Polychlorinated Biphenyls: Human Health Aspects. World Health Organization, Geneva. 18. AgencyforToxicSubstancesandDiseaseRegistry (2000) Toxicological Profile for Polychlorinated Biphenyls (PCBs). Atlanta, GA: U.S. Department of Human Health and Human Services.   141 19. Hopf NB, Ruder AM, Succop P (2009) Background levels of polychlorinated biphenyls in the U.S. population. Sci Total Environ 407: 6109-6119. 20. Haddock JD, Gibson DT (1995) Purification and characterization of the oxygenase component of biphenyl 2,3-dioxygenase from Pseudomonas sp. strain LB400. J Bacteriol 177: 5834-5839. 21. Fortin PD, Horsman GP, Yang HM, Eltis LD (2006) A glutathione S-transferase catalyzes the dehalogenation of inhibitory metabolites of polychlorinated biphenyls. J Bacteriol 188: 4424-4430. 22. Gomez-Gil L, Kumar P, Barriault D, Bolin JT, Sylvestre M, et al. (2007) Characterization of biphenyl dioxygenase of Pandoraea pnomenusa B-356 as a potent polychlorinated biphenyl-degrading enzyme. J Bacteriol 189: 5705-5715. 23. Iwasaki T, Takeda H, Miyauchi K, Yamada T, Masai E, et al. (2007) Characterization of two biphenyl dioxygenases for biphenyl/PCB degradation in A PCB degrader, Rhodococcus sp. strain RHA1. Biosci Biotechnol Biochem 71: 993-1002. 24. Pieper DH, Seeger M (2008) Bacterial metabolism of polychlorinated biphenyls. J Mol Microbiol Biotechnol 15: 121-138. 25. Iwai S, Chai B, Sul WJ, Cole JR, Hashsham SA, et al. (2010) Gene-targeted-metagenomics reveals extensive diversity of aromatic dioxygenase genes in the environment. ISME J 4: 279-285. 26. Standfuss-Gabisch C, Al-Halbouni D, Hofer B (2012) Characterization of biphenyl dioxygenase sequences and activities encoded by the metagenomes of highly polychlorobiphenyl-contaminated soils. Appl Environ Microbiol 78: 2706-2715. 27. Kim D, Chae JC, Zylstra GJ, Kim YS, Kim SK, et al. (2004) Identification of a novel dioxygenase involved in metabolism of o-xylene, toluene, and ethylbenzene by Rhodococcus sp. strain DK17. Appl Environ Microbiol 70: 7086-7092. 28. Yoo M, Kim D, Zylstra GJ, Kang BS, Kim E (2011) Biphenyl hydroxylation enhanced by an engineered o-xylene dioxygenase from Rhodococcus sp. strain DK17. Res Microbiol 162: 724-728. 29. Cho HJ, Kim K, Sohn SY, Cho HY, Kim KJ, et al. (2010) Substrate binding mechanism of a type I extradiol dioxygenase. J Biol Chem 285: 34643-34652. 30. Kumamaru T, Suenaga H, Mitsuoka M, Watanabe T, Furukawa K (1998) Enhanced degradation of polychlorinated biphenyls by directed evolution of biphenyl dioxygenase. Nat Biotechnol 16: 663-666. 31. Barriault D, Sylvestre M (2004) Evolution of the biphenyl dioxygenase BphA from Burkholderia xenovorans LB400 by random mutagenesis of multiple sites in region III. J Biol Chem 279: 47480-47488. 32. Mohammadi M, Sylvestre M (2005) Resolving the profile of metabolites generated during oxidation of dibenzofuran and chlorodibenzofurans by the biphenyl catabolic pathway enzymes. Chem Biol 12: 835-846. 33. Viger JF, Mohammadi M, Barriault D, Sylvestre M (2012) Metabolism of chlorobiphenyls by a variant biphenyl dioxygenase exhibiting enhanced activity toward dibenzofuran. Biochem Biophys Res Commun 419: 362-367. 34. Furukawa K, Suenaga H, Goto M (2004) Biphenyl dioxygenases: functional versatilities and directed evolution. J Bacteriol 186: 5189-5196.   142 35. Fortin PD, MacPherson I, Neau DB, Bolin JT, Eltis LD (2005) Directed evolution of a ring-cleaving dioxygenase for polychlorinated biphenyl degradation. J Biol Chem 280: 42307-42314. 36. Bedard DL, Haberl ML (1990) Influence of Chlorine Substitution Pattern on the Degradation of Polychlorinated Biphenyls by Eight Bacterial Strains. Microbial Ecology 20: 87-102. 37. Seah SY, Labbe G, Nerdinger S, Johnson MR, Snieckus V, et al. (2000) Identification of a serine hydrolase as a key determinant in the microbial degradation of polychlorinated biphenyls. J Biol Chem 275: 15701-15708. 38. Seah SY, Labbe G, Kaschabek SR, Reifenrath F, Reineke W, et al. (2001) Comparative specificities of two evolutionarily divergent hydrolases involved in microbial degradation of polychlorinated biphenyls. J Bacteriol 183: 1511-1516. 39. Seah SY, Ke J, Denis G, Horsman GP, Fortin PD, et al. (2007) Characterization of a C-C bond hydrolase from Sphingomonas wittichii RW1 with novel specificities towards polychlorinated biphenyl metabolites. J Bacteriol 189: 4038-4045. 40. Van der Geize R, Yam K, Heuser T, Wilbrink MH, Hara H, et al. (2007) A gene cluster encoding cholesterol catabolism in a soil actinomycete provides insight into Mycobacterium tuberculosis survival in macrophages. Proc Natl Acad Sci U S A 104: 1947-1952. 41. Lack NA, Yam KC, Lowe ED, Horsman GP, Owen RL, et al. (2010) Characterization of a carbon-carbon hydrolase from Mycobacterium tuberculosis involved in cholesterol metabolism. J Biol Chem 285: 434-443. 42. Ollis DL, Cheah E, Cygler M, Dijkstra B, Frolow F, et al. (1992) The alpha/beta hydrolase fold. Protein Eng 5: 197-211. 43. Nardini M, Dijkstra BW (1999) Alpha/beta hydrolase fold enzymes: the family keeps growing. Curr Opin Struct Biol 9: 732-737. 44. Holmquist M (2000) Alpha/Beta-hydrolase fold enzymes: structures, functions and mechanisms. Curr Protein Pept Sci 1: 209-235. 45. Larsen NA, Lin H, Wei R, Fischbach MA, Walsh CT (2006) Structural characterization of enterobactin hydrolase IroE. Biochemistry 45: 10184-10190. 46. Li B, Yang G, Wu L, Feng Y (2012) Role of the NC-loop in catalytic activity and stability in lipase from Fervidobacterium changbaicum. PLoS One 7: e46881. 47. Jochens H, Stiba K, Savile C, Fujii R, Yu JG, et al. (2009) Converting an esterase into an epoxide hydrolase. Angew Chem Int Ed Engl 48: 3532-3535. 48. Semeriva M, Chapus C, Bouvier-Lapierre C, Desnuelle P (1974) On the transient formation of an acetyl enzyme intermediate during the hydrolysis of p-nitrophenyl acetate by pancreatic lipase. Biochem Biophys Res Commun 58: 808-813. 49. Chapus C, Semeriva M, Bovier-Lapierre C, Desnuelle P (1976) Mechanism of pancreatic lipase action. 1. Interfacial activation of pancreatic lipase. Biochemistry 15: 4980-4987. 50. Koller W, Kolattukudy PE (1982) Mechanism of action of cutinase: chemical modification of the catalytic triad characteristic for serine hydrolases. Biochemistry 21: 3083-3090.   143 51. Froede HC, Wilson IB (1984) Direct determination of acetyl-enzyme intermediate in the acetylcholinesterase-catalyzed hydrolysis of acetylcholine and acetylthiocholine. J Biol Chem 259: 11010-11013. 52. Li J, Szittner R, Derewenda ZS, Meighen EA (1996) Conversion of serine-114 to cysteine-114 and the role of the active site nucleophile in acyl transfer by myristoyl-ACP thioesterase from Vibrio harveyi. Biochemistry 35: 9967-9973. 53. Nazi I, Wright GD (2005) Catalytic mechanism of fungal homoserine transacetylase. Biochemistry 44: 13560-13566. 54. De Pascale G, Nazi I, Harrison PH, Wright GD (2011) beta-Lactone natural products and derivatives inactivate homoserine transacetylase, a target for antimicrobial agents. J Antibiot (Tokyo) 64: 483-487. 55. Lejon S, Ellis J, Valegard K (2008) The last step in cephalosporin C formation revealed: crystal structures of deacetylcephalosporin C acetyltransferase from Acremonium chrysogenum in complexes with reaction intermediates. J Mol Biol 377: 935-944. 56. Verschueren KH, Seljee F, Rozeboom HJ, Kalk KH, Dijkstra BW (1993) Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 363: 693-698. 57. Stennicke HR, Mortensen UH, Breddam K (1996) Studies on the hydrolytic properties of (serine) carboxypeptidase Y. Biochemistry 35: 7131-7141. 58. Medrano FJ, Alonso J, Garcia JL, Romero A, Bode W, et al. (1998) Structure of proline iminopeptidase from Xanthomonas campestris pv. citri: a prototype for the prolyl oligopeptidase family. EMBO J 17: 1-9. 59. Pathak D, Ashley G, Ollis D (1991) Thiol protease-like active site found in the enzyme dienelactone hydrolase: localization using biochemical, genetic, and structural tools. Proteins 9: 267-279. 60. Chakravarty B, Gu Z, Chirala SS, Wakil SJ, Quiocho FA (2004) Human fatty acid synthase: structure and substrate selectivity of the thioesterase domain. Proc Natl Acad Sci U S A 101: 15567-15572. 61. Yin DT, Kazlauskas RJ (2012) Revised molecular basis of the promiscuous carboxylic acid perhydrolase activity in serine hydrolases. Chemistry 18: 8130-8139. 62. Andexer JN, Staunig N, Eggert T, Kratky C, Pohl M, et al. (2012) Hydroxynitrile lyases with alpha/beta-hydrolase fold: two enzymes with almost identical 3D structures but opposite enantioselectivities and different reaction mechanisms. Chembiochem 13: 1932-1939. 63. Steiner RA, Janssen HJ, Roversi P, Oakley AJ, Fetzner S (2010) Structural basis for cofactor-independent dioxygenation of N-heteroaromatic compounds at the alpha/beta-hydrolase fold. Proc Natl Acad Sci U S A 107: 657-662. 64. Fleming SM, Robertson TA, Langley GJ, Bugg TD (2000) Catalytic mechanism of a C-C hydrolase enzyme: evidence for a gem-diol intermediate, not an acyl enzyme. Biochemistry 39: 1522-1531. 65. Li C, Li JJ, Montgomery MG, Wood SP, Bugg TD (2006) Catalytic role for arginine 188 in the C-C hydrolase catalytic mechanism for Escherichia coli MhpC and Burkholderia xenovorans LB400 BphD. Biochemistry 45: 12470-12479. 66. Horsman GP, Bhowmik S, Seah SY, Kumar P, Bolin JT, et al. (2007) The tautomeric half-reaction of BphD, a C-C bond hydrolase. Kinetic and structural evidence   144 supporting a key role for histidine 265 of the catalytic triad. J Biol Chem 282: 19894-19904. 67. Seah SY, Terracina G, Bolin JT, Riebel P, Snieckus V, et al. (1998) Purification and preliminary characterization of a serine hydrolase involved in the microbial degradation of polychlorinated biphenyls. J Biol Chem 273: 22943-22949. 68. Lack N, Lowe ED, Liu J, Eltis LD, Noble ME, et al. (2008) Structure of HsaD, a steroid-degrading hydrolase, from Mycobacterium tuberculosis. Acta Crystallogr Sect F Struct Biol Cryst Commun 64: 2-7. 69. Dunn G, Montgomery MG, Mohammed F, Coker A, Cooper JB, et al. (2005) The structure of the C-C bond hydrolase MhpC provides insights into its catalytic mechanism. J Mol Biol 346: 253-265. 70. Fushinobu S, Saku T, Hidaka M, Jun SY, Nojiri H, et al. (2002) Crystal structures of a meta-cleavage product hydrolase from Pseudomonas fluorescens IP01 (CumD) complexed with cleavage products. Protein Sci 11: 2184-2195. 71. Riddle RR, Gibbs PR, Willson RC, Benedik MJ (2003) Purification and properties of 2-hydroxy-6-oxo-6-(2'-aminophenyl)hexa-2,4-dienoic acid hydrolase involved in microbial degradation of carbazole. Protein Expr Purif 28: 182-189. 72. Diaz E, Timmis KN (1995) Identification of functional residues in a 2-hydroxymuconic semialdehyde hydrolase. A new member of the alpha/beta hydrolase-fold family of enzymes which cleaves carbon-carbon bonds. J Biol Chem 270: 6403-6411. 73. Hedstrom L (2002) Serine protease mechanism and specificity. Chem Rev 102: 4501-4524. 74. Grogan G (2005) Emergent mechanistic diversity of enzyme-catalysed beta-diketone cleavage. Biochem J 388: 721-730. 75. Phillips RS (2011) Structure, mechanism, and substrate specificity of kynureninase. Biochim Biophys Acta 1814: 1481-1488. 76. Lam WW, Bugg TD (1997) Purification, characterization, and stereochemical analysis of a C-C hydrolase: 2-hydroxy-6-keto-nona-2,4-diene-1,9-dioic acid 5,6-hydrolase. Biochemistry 36: 12242-12251. 77. Begley TP, Ealick SE (2004) Enzymatic reactions involving novel mechanisms of carbanion stabilization. Curr Opin Chem Biol 8: 508-515. 78. Van Vleet JL, Reinhardt LA, Miller BG, Sievers A, Cleland WW (2008) Carbon isotope effect study on orotidine 5'-monophosphate decarboxylase: support for an anionic intermediate. Biochemistry 47: 798-803. 79. Bornscheuer UT, Kazlauskas RJ (2004) Catalytic promiscuity in biocatalysis: using old enzymes to form new bonds and follow new pathways. Angew Chem Int Ed Engl 43: 6032-6040. 80. Li JJ, Bugg TD (2005) Stereochemistry of the reaction catalysed by 2-hydroxy-6-keto-6-phenyl-hexa-2,4-dienoic acid 5,6-hydrolase (BphD). Chem Commun (Camb): 130-132. 81. Horsman GP, Ke J, Dai S, Seah SY, Bolin JT, et al. (2006) Kinetic and structural insight into the mechanism of BphD, a C-C bond hydrolase from the biphenyl degradation pathway. Biochemistry 45: 11071-11086. 82. Catelani D, Colombi A (1974) Metabolism of biphenyl. Structure and physicochemical properties of 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoic acid, the meta-cleavage   145 product from 2,3-dihydroxybiphenyl by Pseudomonas putida. Biochem J 143: 431-434. 83. Henderson IM, Bugg TD (1997) Pre-steady-state kinetic analysis of 2-hydroxy-6-keto-nona-2,4-diene-1,9-dioic acid 5,6-hydrolase: kinetic evidence for enol/keto tautomerization. Biochemistry 36: 12252-12258. 84. Dahl KH, Dunn MF (1984) Carboxymethylated liver alcohol dehydrogenase: kinetic and thermodynamic characterization of reactions with substrates and inhibitors. Biochemistry 23: 6829-6839. 85. D'Ordine RL, Tonge PJ, Carey PR, Anderson VE (1994) Electronic rearrangement induced by substrate analog binding to the enoyl-CoA hydratase active site: evidence for substrate activation. Biochemistry 33: 12635-12643. 86. Bhowmik S, Horsman GP, Bolin JT, Eltis LD (2007) The molecular basis for inhibition of BphD, a C-C bond hydrolase involved in polychlorinated biphenyls degradation: large 3-substituents prevent tautomerization. J Biol Chem 282: 36377-36385. 87. Bugg TD, Fleming SM, Robertson TA, Langley GJ (2002) 2-hydroxy-6-keto-nona-2,4-diene 1,9-dioic acid 5,6-hydrolase: evidence from 18O isotope exchange for gem-diol intermediate. Methods Enzymol 354: 106-118. 88. Hyland LJ, Tomaszek TA, Jr., Roberts GD, Carr SA, Magaard VW, et al. (1991) Human immunodeficiency virus-1 protease. 1. Initial velocity studies and kinetic characterization of reaction intermediates by 18O isotope exchange. Biochemistry 30: 8441-8453. 89. Li JJ, Li C, Blindauer CA, Bugg TD (2006) Evidence for a gem-diol reaction intermediate in bacterial C-C hydrolase enzymes BphD and MhpC from 13C NMR spectroscopy. Biochemistry 45: 12461-12469. 90. Speare DM, Olf P, Bugg TD (2002) Hammett analysis of a C-C hydrolase-catalysed reaction using synthetic 6-aryl-2-hydroxy-6-ketohexa-2,4-dienoic acid substrates. Chem Commun (Camb): 2304-2305. 91. Speare DM, Fleming SM, Beckett MN, Li JJ, Bugg TD (2004) Synthetic 6-aryl-2-hydroxy-6-ketohexa-2,4-dienoic acid substrates for C-C hydrolase BphD: investigation of a general base catalytic mechanism. Org Biomol Chem 2: 2942-2950. 92. Bender ML, Nakamura K (1962) The effect of strructure on the rates of some ?-chymotrypsin-catalyzed reactions. J Am Chem Soc 84: 2577-2582. 93. Li JJ, Bugg TD (2007) Investigation of a general base mechanism for ester hydrolysis in C-C hydrolase enzymes of the alpha/beta-hydrolase superfamily: a novel mechanism for the serine catalytic triad. Org Biomol Chem 5: 507-513. 94. Seydoux F, Malhotra OP, Bernhard SA (1974) Half-site reactivity. CRC Crit Rev Biochem 2: 227-257. 95. Fersht A (1999) Structure and mechanism in protein science: a guide to enzyme catalysis and protein folding: W.H. Freeman and Company. 96. Silverman RB (2000) The organic chemistry of enzyme-catalyzed reactions: Academic Press. 97. Couture MM, Colbert CL, Babini E, Rosell FI, Mauk AG, et al. (2001) Characterization of BphF, a Rieske-type ferredoxin with a low reduction potential. Biochemistry 40: 84-92.   146 98. Patil DN, Tomar S, Sylvestre M, Kumar P (2010) Expression, purification, crystallization and preliminary crystallographic studies of cis-biphenyl-2,3-dihydrodiol-2,3-dehydrogenase from Pandoraea pnomenusa B-356. Acta Crystallogr Sect F Struct Biol Cryst Commun 66: 1517-1520. 99. Maniatis T, Fritsch EF, Sambrook J (1982) Molecular cloning: a laboratory manual. New York: Cold Spring Harbor Laboratory. 100. Bauchop T, Elsden SR (1960) The growth of micro-organisms in relation to their energy supply. J Gen Microbiol 23: 457-469. 101. Horsman GP (2008) Characterization of BphD, a C-C boond hydrolase involved in the degradation of polychlorinated biphenyls. Vancouver: The University of British Columbia. 102. Cornish-Bowden A (1995) Analysis of enzyme kinetic data. New York: Oxford University Press. 103. Team RDC (2010) R: A language and environment for statistical computing. Vienna, Austria: R Foundation for Statistical Computing. 104. Chan QW, Howes CG, Foster LJ (2006) Quantitative comparison of caste differences in honeybee hemolymph. Mol Cell Proteomics 5: 2252-2262. 105. Russell RB, Barton GJ (1992) Multiple protein sequence alignment from tertiary structure comparison: assignment of global and residue confidence levels. Proteins 14: 309-323. 106. Roberts E, Eargle J, Wright D, Luthey-Schulten Z (2006) MultiSeq: unifying sequence and structure data for evolutionary analysis. BMC Bioinformatics 7: 382. 107. Humphrey W, Dalke A, Schulten K (1996) VMD: visual molecular dynamics. J Mol Graph 14: 33-38, 27-38. 108. Emsley P, Cowtan K (2004) Coot: model-building tools for molecular graphics. Acta Crystallogr D Biol Crystallogr 60: 2126-2132. 109. Nandhagopal N, Senda T, Hatta T, Yamada A, Masai E, et al. (1997) Three-dimensional structure of microbial 2-hydroxyl-6-oxo-phenylhexa-2,4-dienoic acid (HPDA) hydrolase (BphD enzyme) from Rhodococcus sp. strain RHA1, in the PCB degradation pathway,. Proceedings of the Japan Academy, Ser B 73: 154. 110. Habe H, Morii K, Fushinobu S, Nam JW, Ayabe Y, et al. (2003) Crystal structure of a histidine-tagged serine hydrolase involved in the carbazole degradation (CarC enzyme). Biochem Biophys Res Commun 303: 631-639. 111. Pollard JR, Henderson IM, Bugg TD (1997) Chemical and biochemical properties of 2-hydroxypentadienoic acid, a homologue of enolpyruvic acid. Chem Commun: 1885-1886. 112. Ghosh S (2012): Purdue University. 113. Li C, Montgomery MG, Mohammed F, Li JJ, Wood SP, et al. (2005) Catalytic mechanism of C-C hydrolase MhpC from Escherichia coli: kinetic analysis of His263 and Ser110 site-directed mutants. J Mol Biol 346: 241-251. 114. Li C, Hassler M, Bugg TD (2008) Catalytic promiscuity in the alpha/beta-hydrolase superfamily: hydroxamic acid formation, C--C bond formation, ester and thioester hydrolysis in the C--C hydrolase family. Chembiochem 9: 71-76. 115. Zhou H, Qu Y, Kong C, Wu Y, Zhu K, et al. (2012) Promiscuous esterase activities of the C-C hydrolases from Dyella ginsengisoli. Biotechnol Lett 34: 1107-1113.   147 116. Abell LM, Schloss JV (1991) Oxygenase side reactions of acetolactate synthase and other carbanion-forming enzymes. Biochemistry 30: 7883-7887. 117. Bhowmik S (2008) Structural investigation into the mechanism of catalysis and inhibition of two carbon-carbon bond hydrolases. Ann Arbor: Purdue University. 118. Barth S, Fischer M, Schmid RD, Pleiss J (2004) Sequence and structure of epoxide hydrolases: a systematic analysis. Proteins 55: 846-855. 119. Ghosh S (2012) Structural Studies of BphD LB400 Variants : Insights into the reaction mechanism of MCP hydrolases.: Purdue University. 120. Sprinson DB, Rittenberg D (1951) Nature of the activation process in enzymatic reactions. Nature 167: 484. 121. Blow DM, Birktoft JJ, Hartley BS (1969) Role of a buried acid group in the mechanism of action of chymotrypsin. Nature 221: 337-340. 122. Fastrez J, Fersht AR (1973) Demonstration of the acyl-enzyme mechanism for the hydrolysis of peptides and anilides by chymotrypsin. Biochemistry 12: 2025-2034. 123. Ostrowski W, Barnard EA (1973) Evidence for a phosphoryl-enzyme intermediate in the catalytic reaction of prostatic acid phosphatase. Biochemistry 12: 3893-3898. 124. Nijvipakul S, Ballou DP, Chaiyen P (2010) Reduction kinetics of a flavin oxidoreductase LuxG from Photobacterium leiognathi (TH1): half-sites reactivity. Biochemistry 49: 9241-9248. 125. Liepinsh E, Otting G, Wuthrich K (1992) NMR spectroscopy of hydroxyl protons in aqueous solutions of peptides and proteins. J Biomol NMR 2: 447-465. 126. White H, Jencks WP (1976) Mechanism and specificity of succinyl-CoA:3-ketoacid coenzyme A transferase. J Biol Chem 251: 1688-1699. 127. Nielsen MB (2009) Model systems for understanding absorption tuning by opsin proteins. Chem Soc Rev 38: 913-924. 128. Wang W, Nossoni Z, Berbasova T, Watson CT, Yapici I, et al. (2012) Tuning the electronic absorption of protein-embedded all-trans-retinal. Science 338: 1340-1343. 129. Schloss JV, Hixon MS (1997) Enol chemistry and enzymology, in Comprehensive Biological Catalysis; Sinnod M, editor. London: Academic Press. 130. Buncel E, Menon B (1979) Carbanion Mechanisms. 9. Spectrophotometric Study of Triphenylmethyl Alkali Metal Salts. Contact and Solvent-Seperated Ion Pairs in Ethereal Solvents. J Org Chem 44: 317-320. 131. Phillips RS, Holtermann G (2005) Differential effects of temperature and hydrostatic pressure on the formation of quinonoid intermediates from L-Trp and L-Met by H463F mutant Escherichia coli tryptophan indole-lyase. Biochemistry 44: 14289-14297. 132. Kumar S, Gawandi VB, Capito N, Phillips RS (2010) Substituent effects on the reaction of beta-benzoylalanines with Pseudomonas fluorescens kynureninase. Biochemistry 49: 7913-7919. 133. Otyepka M, Damborsky J (2002) Functionally relevant motions of haloalkane dehalogenases occur in the specificity-modulating cap domains. Protein Sci 11: 1206-1217. 134. Zhou H, Qu Y, Shen E, Kong C, Zhang X, et al. (2012) Tuning the substrate selectivity of meta-cleavage product hydrolase by domain swapping. Appl Microbiol Biotechnol.   148 135. Zhou H, Qu Y, Kong C, Shen E, Wang J, et al. (2013) The key role of a non-active-site residue Met148 on the catalytic efficiency of meta-cleavage product hydrolase BphD. Appl Microbiol Biotechnol. 136. Skjot M, De Maria L, Chatterjee R, Svendsen A, Patkar SA, et al. (2009) Understanding the plasticity of the alpha/beta hydrolase fold: lid swapping on the Candida antarctica lipase B results in chimeras with interesting biocatalytic properties. Chembiochem 10: 520-527. 137. Ruzzini AC, Bhowmik S, Yam KC, Ghosh S, Bolin JT, et al. (2013) unpublished. 138. Ruzzini AC, Horsman GP, Eltis LD (2012) The catalytic serine of meta-cleavage product hydrolases is activated differently for C-O bond cleavage than for C-C bond cleavage. Biochemistry 51: 5831-5840. 139. Rengarajan J, Bloom BR, Rubin EJ (2005) Genome-wide requirements for Mycobacterium tuberculosis adaptation and survival in macrophages. Proc Natl Acad Sci U S A 102: 8327-8332. 140. Nashine VC, Hammes-Schiffer S, Benkovic SJ (2010) Coupled motions in enzyme catalysis. Curr Opin Chem Biol 14: 644-651. 141. Wilkinson AJ, Fersht AR, Blow DM, Winter G (1983) Site-directed mutagenesis as a probe of enzyme structure and catalysis: tyrosyl-tRNA synthetase cysteine-35 to glycine-35 mutation. Biochemistry 22: 3581-3586.       149 APPENDIX I: CHEMICAL PROPERTIES OF SUBSTRATES    Figure 54. The HOPDA enol:enolate equilibrium, defined by pKa2. An 18O-incorporation event into the C2-oxo was observed in KPi (I = 0.1 M), pH 7.5, suggesting that the 2-oxo-6-oxido- isomer exists under those conditions.  Table 20. HOPDA pKa2 values and extinction coefficients in KPi (I = 0.1 M), pH 7.5 Substrate/Product ?max (nm) ? (mM-1cm-1) pKa2 HOPDA 434 25.7 7.3 3-Cl HOPDA 432 40.6 6.1 5-Cl HOPDA 402 40.1 6.1 8-Cl HOPDA 393 40.3 6.5 9-Cl HOPDA 436 28.2 6.8 10-Cl HOPDA 438 26.3 6.9 3,9,11 tri-Cl HOPDA 438 35.6 ND 8-F HOPDA 398 ND ND 5,8-diF HOPDA 405 31.2 5.0 ? 0.2 HPD 270 19.7 ND      150 APPENDIX II: MASS SPECTROMETRY RESULTS Table 21. b-series ion fragment matches to a BphDLB400 peptide analyzed by ESI/MS/MSa,b,c b-series ion  Peptide  WT (unmodified) WT S112-benzoyl H265Q S112-benzoyl b9+ 977.52 / 977.5163 977.54 / 977.5163 977.51 / 977.5163 b10+ 1091.58 / 1091.5592 1091.58 / 1091.5592 1091.57 / 1091.5592 b11+ 1178.61 / 1178.5913  1282.64 / 1282.6175 b12+ 1309.66 / 1309.6317 1413.69 / 1413.6580 1413.73 / 1413.6580 b13+ 1366.65 / 1366.6532 1470.69 / 1470.6794  b14+ 1423.67 / 1423.6747 1527.73 / 1527.7009  b15+ 1494.74 / 1494.7118 1598.74 / 1598.7380  b17+ 1666.82 / 1666.7966   b11++ 641.83 / 641.8124 589.81 / 589.7993  b13++ 735.85 / 735.8434  735.83 / 735.8434 b14++ 764.36 / 764.3541 712.34 / 712.3410 764.33 / 764.3541 b15++ 764.85 / 799.8726   b17++ 885.97 / 885.9150   b19++ 999.95 / 999.4785   b11*++ 581.30 / 581.2860   b16*++ 789.89 / 789.8701 841.91 / 841.8832  b17*++  877.43 / 877.4018  b11o+ 1160.61 / 1160.5807   b16o+ 1577.72 / 1577.7489   b11o++ 580.80 / 580.7940   b16o++ 789.39 / 789.3781 841.41 / 841.3912 841.37 / 841.3912 b17o++  876.91 / 876.9098 876.91 / 876.9098   Search Result Statistics  Ions score 85 63 50 Expect Value 3.8 E-6 0.00057 0.022 Matches (out of 212) 25 34 23 RMS error (ppm) 207 220 276 a ? values of the observed fragments based on manual inspection of raw data and fits to Gaussian peaks followed by MASCOT match results b ? MASCOT search of B. xenovorans LB400 proteins resulted in a match to the BphD peptide, DIDRAHLVGNSMGGATALNF c ? Symbols * and o indicate ions with an additional loss of NH3 and H2O, respectively.     151 Table 22. Example of modeling EI GC/MS data to 18O incorporation events  Observed relative abundance of ions from EI/GC/MS (%) m/z 179 [M] 180 [M+1] 181 [M+2] 182 [M+3] 183 [M+4] 184 [M+5]  Controla 83.5 12.4 3.9 0.35    WT + HOPDAb 12 2.7 71 10.7 3.6 0.3  Modeling the relative abundance (R.A.) of benzoate species in the sample (%) Model 1: Two species accounting for a single 18O incorporation event  R.A. [M] R.A. [M+1] R.A. [M+2] R.A. [M+3] R.A. [M+4] R.A. [M+5] Average  R.A.c,d 16O/16O 14 22 16 14   15 ? 1 16O/18O   84 86 92 86 85 ? 1 Model 2: Three species including 3% of doubly 18O-incorporated benzoatee 16O/16O 14 22 16 14   15 ? 1 16O/18O   84 86 28 -21 82 ? 3 a ? Control values represent the average of all benzoate ion fragments observed in H2O b ? Benzoate derived from HOPDA incubated with WT BphD for 20 minutes without any pre-incubation c ? The relative abundance of each species (16O/18O and 18O/18O) was calculated from the observed intensity of each ion (Obs) from the WT reaction and the control (Co) as follows: R.A. [M]16O/16O = [M]Obs/[M]Co R.A. [M+1]16O/16O = [M+1]Obs/[M+1]Co, R.A. [M+2]16O/16O = ([M]Co - [M+2]Obs)/([M]Co-[M+2]Co) R.A. [M+3]16O/16O = ([M+1]Co - [M+3]Obs)/([M+1]Co-[M+3]Co) R.A. [M+4]16O/16O = ([M+2]Co - [M+4]Obs)/([M+2]Co-[M+4]Co) R.A. [M+5]16O/16O = ([M+3]Co - [M+5]Obs)/([M+3]Co-[M+5]Co) R.A. of [M+N] 16O/18O = 1 - [M+N]16O/16O d ? The averages and errors were weighted based on fragment ion intensity e ? The calculation in Table S4 used a model in which 3% of the benzoate contains two equivalents of 18O. Ion fragments derived from this species only contribute to the observed signals at M+4 and M+5: no M+6 or M+7 ions were observed.       152 Table 23. Relative abundance of ion fragments analyzed by EI GC/MS    Relative abundance (%)a of benzoate ion fragments (m/z)  Sample and Incubation time 105 [M] 106 [M+1] 107 [M+2] 108 [M+3] 109 [M+4]   18O incorporationb benzoate 300 min H2O 92 ? 1 7 ? 1 1.0 ? 0.1     H218O 92.2 ? 0.2  6.7 ? 0.1 1.1 ? 0.1    ND WT + benzoate 300 min H2O 92.3 ? 0.1  6.6 ? 0.2 1.1 ? 0.1     H218O 91.5 ? 0.7 7.2 ? 0.5 1.3 ? 0.2    ND WT + HOPDA 20 min H2O 92.2 ? 0.4 6.8 ? 0.8 1.0 ? 0.4     H218O 53 ? 2 4.9 ? 0.4 39 ? 3 2.7 ? 0.2 0.3 ? 0.2  42 ? 1% WT + HOPDA 5 min PI H2O 92 ? 2 7 ? 1 1.0 ? 0.3     20 min rxn H218O 52.9 ? 0.3 5.5 ? 0.9 38 ? 1 3.1 ? 0.2 0.17 ? 0.02  42 ? 2% WT + HOPDA 20 min PI H2O 91 ? 2 8 ? 1 0.9 ? 0.4     20 min rxn H218O 52.5 ? 0.6 4.8 ? 0.3 38.5 ? 0.4 3.8 ? 0.1 0.29 ? 0.03  43 ? 2%    179 [M] 180 [M+1] 181 [M+2] 182 [M+3] 183 [M+4] 184 [M+5]  benzoate 300 min H2O 83.2 ? 0.6 12.9 ? 0.5 3.4 ? 0.1 0.44 ? 0.08    H218O 83.1 ? 0.5 12.4 ? 0.3 4.1 ? 0.1 0.4 ? 0.2   ND WT + benzoate 300 min H2O 84.1 ? 0.4 11.9 ? 0.4 3.6 ? 0.1 0.31 ? 0.01    H218O 83 ? 0.2 12.3 ? 0.1 4.3 ? 0.1 0.4 ? 0.04   ND WT + HOPDA 20 min H2O 84 ? 1  12 ? 1 3.9 ? 0.1 0.3 ? 0.1    H218O 12 ? 1 2.7 ? 0.1 71 ? 2 10.7 ? 0.5 3.6 ? 0.1 0.3 ? 0.1 85 ? 1% WT + HOPDA 5 min PI H2O 82.7 ? 0.9  12 ? 1 4.4 ? 0.5 0.4 ? 0.1 0.23 ? 0.01   20 min rxn H218O 14 ? 3 3.9 ? 0.3 68 ? 2 11 ? 1 3.1 ? 0.1 0.3 ? 0.2 80 ? 2% WT + HOPDA 20 min PI H2O 82.7 ? 0.1 12.8 ? 0.4 4.0 ? 0.4 0.33 ? 0.07 0.12 ? 0.04   20 min rxn H218O 16.7 ? 0.4 3.3 ? 0.2 65 ? 1 10.0 ? 0.1 4 ? 1 0.5 ? 0.2 79 ? 2%    193 [M-1] 194 [M] 195 [M+1] 196 [M+2] 197 [M+3] 198 [M+4]  benzoate 300 min H2O 15.7 ? 0.1 70.2 ? 0.7 11.2 ? 0.3 3 ? 1    H218O 18 ? 2  65.6 ? 0.4 12.6 ? 0.1 3 ? 1   ND WT + benzoate 300 min H2O 15.5 ? 0.3 69 ? 1 12 ? 2 4.0 ? 0.7    H218O 14.3 ? 0.8 69.5 ? 0.3 11.8 ? 0.3 3.9 ? 0.4 0.39 ? 0.06  ND WT + HOPDA 20 min H2O 17 ? 5 66 ? 1 11 ? 2 5 ? 1    H218O 3 ? 1 11 ? 4 15 ? 2 57 ? 8 11 ? 2 2.7 ? 0.8 85 ? 4 % WT + HOPDA 5 min PI H2O 16 ? 2 69 ? 2 11 ? 4 3 ? 5    20 min rxn H218O 3.0 ? 0.4 15 ? 1 12 ? 2 55 ? 1 11.5 ? 0.8 3.0 ? 0.9 72 ? 9 % WT + HOPDA 20 min PI H2O 14 ? 3 71 ? 1 10.7 ? 0.1 5 ? 1    20 min rxn H218O 4 ? 1 10 ? 1 12.5 ? 0.4 59 ? 2 10 ? 1 4.4 ? 0.1 80 ? 10 %    Relative abundance (%) of HOPDA ion fragments (m/z)     245 [M] 246 [M+1] 247 [M+2] 248 [M+3] 249 [M+4] 250 [M+5]  HOPDA 20 min H2O 78 ? 2 16.0 ? 0.2 5 ? 2 0.5 ? 0.2    H218O 65.0 ? 0.1 12.7 ? 0.2 17.9 ? 0.1 3.2 ? 0.1 1.0 ? 0.2 0.12 ? 0.01 18 ? 1%    257 [M] 258 [M+1] 259 [M+2] 260 [M+3] 261 [M+4]   HOPDA 20 min H2O 71 ? 5 22 ? 5 6.7 ? 0.1     H218O 62.3 ? 0.3 15 ? 3 18 ? 8 3.1 ? 0.1 1.1 ? 0.1  17.? 1 %    347 [M] 348 [M+1] 349 [M+2] 350 [M+3] 351 [M+4]   HOPDA 20 min H2O 73 ? 6 18 ? 2 8 ? 2 1 ? 1    H218O 58 ? 3 17 ? 2 16 ? 2 6.3 ? 0.8 2 ? 1  17 ? 5% a ? errors for relative abundance measurements are a standard deviation from two replicates b ? errors for 18O incorporation represent the root mean square error from fitting the experimentally observed data to a single 18O incorporation or to a model that accounts for 2% incorporation of a second 18O equivalent in parentheses c ? overall rms error based on weighted residual plot analysis, where residual = predicted % intensity from model ? observed value    153 APPENDIX III: PRE- OR POST-CATALYTIC EVENTS Figure 55. Visible spectra of post-reaction mixtures of DxnB2 and 20 ?M HOPDA under single or multiple turnover conditions. The signal at 405 nm has tentatively been assigned to a DxnB2:HPD complex based on the enzyme-dependent increase in absorptivity.    Figure 56. EI GC/MS demonstrating deuterium exchange into HOPDA over 2 hours. (A) The HOPDA base peak (245 m/z) at t = 0 h (top panel) and t = 2 h (bottom panel). The mass spectrum of HOPDA at t = 0 h is inset. (B) The time-dependent exchange of deuterium into the HOPDA base peak and calculation of the exponential rate of exchange using the M and M+2 species. Two equivalents of deuterium were incorporated into approximately 63% of the sample. The incorporation was previously determined to occur at C3 and C5 of the dienoate[76].     154 APPENDIX IV: 1H13C HSQC SPECTRUM OF DxnB2 S105A:HOPDA COMPLEX  Figure 57. (A) 1H 13C HSQC spectra of HOPDA in acetone (blue) and phosphate buffer (red). (B) The 1H 13C HSQC spectrum of a DxnB2 S105A:[13C]-HOPDA binary complex. Definitive assignment of the carbons in the binary complex is not possible from the HSQC experiment. Background signal origination from the enzyme is also clearly visible in this region, and was especially significant for nuclei with 1H shifts < 5 ppm (not shown).  155 APPENDIX V: STEADY-STATE KINETICS AND EQUILIBRIUM BINDING  Equations used for the determination of steady-state kinetic parameters  (1) Michaelis-Menten equation    (2) Substrate Inhibition    Equations used for the determination of dissociation constants  (3) Hyperbolic binding equation   where A is the difference in absorbance (?Abs) of the sample upon the addition of enzyme, [E], and Amax is the absorbance once all the substrate is bound.  (4) Quadratic binding equation    where S is the concentration of substrate and E is the concentration of enzymeVmax[S] Km + [S] v0 = Vmax[S] Km + [S] 1 +  [S] Ksi v0 = Amax[E] Kd + [E] A = S + E + Kd ?   (S + E + Kd)2 ? 4SE 2E A = Amax   156 Figure 58. Steady-state kinetic curves (v0 versus [S]) for the DxnB2 P-subsite variant catalyzed hydrolysis of HOPDA. The fit to each dataset is shown as a line.     157 Figure 59. Steady-state kinetic curves (v0 versus [S]) for the DxnB2 P-subsite variant catalyzed hydrolysis of 9-Cl HOPDA. The fit to each dataset is shown as a line.   158 Figure 60. Titration of 4 ?M HOPDA (grey) or 3 ?M 9-Cl HOPDA (black) with DxnB2 N43D/S105A in KPi (I = 0.1 M), pH 7.5 at 25 ?C. The data were fit with a quadratic binding equation. In each case, the Kd was determined to be ~0.02 ?M, indicating the limit of the assay. Overall, the results from fitting the quadratic equation to each dataset also poorly modeled the amount of substrate known to be present in each assay: estimates of the known amount 4 or 3 ?M of HOPDA or 9-Cl HOPDA were only 75 and 82%, respectively. Thus, a Kd < 20 nM was used to describe the binding of both substrates to the N43D/S105A.     159 Table 24. Transition state stabilization by MCP hydrolase side chains to HOPDA. Enzyme kcat/Km (M-1s-1) ??GR| (kcal mol-1) BphD wild type, pH 7.5a 3.2 x 106  S112C 9.5 x 104 2.1 BphD wild-type, pH 8.0b 3.3 x 106  R190Q 5.7 x 102 5.1 R190K 2.8 x 103 4.2 MhpC, pH 8.0b 4.1 x 106  N109A 3.3 x 104 2.9 N109H 1.9 x 104 3.2 F173G 1.3 x 104 2.0 F173D 2.6 x 104 3.0 R188Q 1.4 x 103 4.7 R188K 1.9 x 104 3.2 C261A 2.9 x 106 0.2 W264G 2.1 x 104 3.1 DxnB2, pH 7.5 1.2 x 107  N43A 4.9 x 105 1.9 N43D 7.9 x 102 5.7 N43H 2.3 x 104 3.7 R180K 2.5 x 105 2.3 R180M 2.9 x 102 6.3 R180Q 5.4 x 102 5.9 a,bkcat/Km values were taken from references [81] and [65], respectively.  Analysis of steady-state kinetic data. Contributions of individual residues to transition state stabilization were estimated from previously measured steady-state kinetic parameters for BphDLB400, MhpC and those presented in this work for DxnB2. The analysis was performed as previously described for a C35G variant of tyrosyl-tRNA synthetase[141]. Briefly, the specificity constants were compared using the following equation: ?GR = RTln[(kcat/Km)mut / (kcat/Km)WT], which assumes that the transition state energy is unaffected by substitution, and that the mutated side-chains do not affect productive binding of a second substrate, in this case H2O.    

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