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Mechanistic studies on the cellulomonas fimi exoglycanase (Cex) Birsan, Camelia 1996

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MECHANISTIC STUDIES ON T H E CELLULOMONAS  FIMI  E X O G L Y C A N A S E (CEX) BY  C AMELIA B IRS AN B.Sc, University of Waterloo, 1994  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES DEPARTMENT OF BIOCHEMISTRY AND M O L E C U L A R BIOLOGY  We accept this thesis as conforming to the required standard:  THE UNIVERSITY OF BRITISH COLUMBIA July, 1996 © Camelia Birsan, 1996  In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholariy purposes may be granted by the head of my department or by his or her representatives.  It is understood that copying or  publication of this thesis for financial gain shall not be allowed without my written permission.  Department The University of British Columbia Vancouver, Canada  DE-6 (2/88)  ABSTRACT The objective of this study was to probe the roles of the conserved residues His205 and Gln87 in the active site of the retaining exoglycanase (Cex) from Cellulomonasfimi,a member of the family 10 glycosyl hydrolases. Cex has been previously shown to hydrolyze 6-1,4 glycosidic bonds in xylan, cellulose, and a range of soluble aryl-glycosides by a double displacement mechanism via formation (glycosylation) and hydrolysis (deglycosylation) of a covalent glycosyl-enzyme intermediate. X-ray crystallographic analysis of the enzyme reveals that His205 is positioned within hydrogen bonding distance of the catalytic nucleophile, Glu233, suggesting that it may play a role in modulating the ionization state of this key residue. Structural analysis also reveals that Gln87 is located within close proximity of the C-5 hydroxymethylene group of the distal glucose unit of a cellobioside substrate bound in the active site of Cex, thus suggesting a potential role of Gln87 in substrate binding and specificity. His205 has been systematically replaced with alanine and asparagine via sitedirected mutagenesis, and the resulting effect on overall structure, activity and pH profile has been determined. Kinetic analysis of mutants revealed that His205 is required for both the glycosylation and the deglycosylation steps; rate reductions of 300-5600 fold for His205Ala and 2000-9000 fold for His205Asn were observed on the glycosylation step, while the rate reductions on the deglycosylation step were found to be 4000-5000 fold for His205Ala and 8000-9000 fold for the His205Asn mutant. These effects are consistent with the proposed role of His205 in maintaining the proper ionization state of Glu233. Additionally, His205 may provide some acid catalytic assistance to the oxygen of the leaving group aglycone at the glycosylation transition state, thereby explaining the larger Br0nsted constants measured on the glycosylation step for the His205 mutants than the wild type enzyme. During the deglycosylation step, His205 may be important in stabilizing  Ul  the developing negative charge on the carboxylate group of the departing enzymic nucleophile and enhancing the nucleophilicity of the attacking water molecule, as suggested by the smaller secondary deuterium kinetic isotope effect on the deglycosylation step for the His205Asn mutant. The pH dependence of the glycosylation step showed that ionization states of key catalytic residues in the free enzyme were perturbed upon mutating His205 to either alanine or asparagine, indicating that His205 is important in maintaining the proper electronic environment in the active site of Cex. In order to compress the binding pocket of the C-5 hydroxymethylene groups of substrate to reduce the cellulase activity of Cex, Gln87 has been substituted with the bulkier residues methionine, tyrosine and histidine via site-directed mutagenesis. Kinetic investigation of the Gln87Met mutant revealed no significant changes in the substrate specificity relative to that of the native enzyme, and therefore the function of Gln87 in substrate recognition is not clear. It is hoped that future research activities will provide more insights into this question.  iv  TABLE OF CONTENTS  ABSTRACT TABLE OF CONTENTS LIST OF TABLES LIST OF FIGURES ABBREVIATIONS AND SYMBOLS ACKNOWLEDGEMENTS CHAPTER 1  1  INTRODUCTION  1  1.1 General introduction  2  1.2 Mechanism for hydrolysis of a B-l,4-glycosidic linkage by "retaining" glycosidases  4  1.2.1 Oxocarbonium ion-like transition states  7  1.2.2 Nature of the glycosyl-enzyme intermediate  9  1.2.3 Presence of the catalytic nucleophile  11  1.2.4 General acid assistance  14  1.2.5 Non-covalent enzyme-substrate interactions  16  1.3 Cellulomonas  fimi  B-l,4-exoglycanase  (Cex)  18  1.3.1 General background  18  1.3.2 Importance of the Asp235-His205-Glu233 triad  23  1.3.3 Xylanase vs. Cellulase Activity of Cex  24  1.4 Aims of project  25  CHAPTER 2  27  PROBING THE ROLE OF HIS205 IN THE ACTIVE SITE OF C. FIMI CEX  27  2.1 Introduction  28  2.2 Purification and physical characterization of His205 mutants  33  2.3 Kinetic analysis of Cex and His205 mutants  35  2.3.1 Probing the effect of His205 mutation on the individual steps of the reaction 43 2.3.2 pH dependence study 3.3 Concluding remarks CHAPTER 3  51 54 57  INVESTIGATION ON THE SUBSTRATE SPECIFICITY OF THE BIFUNCTIONAL  C. FIMI XYLANASE/CELLULASE (CEX)  57  3.1 Introduction  58  3.2 Generation of GIn87 mutants  63  3.3 Production, purification and physical characterization of the Gln87Met mutant  65  3.4 Catalytic properties of the Gln87Met mutant  66  3.5 Concluding remarks  69  CHAPTER 4  70  MATERIALS AND METHODS  70  4.1 Chemicals, media components and enzymes  71  4.2 Bacterial strains and plasmids  71  4.3 Media and growth conditions  71  vi 4.4 Recombinant DNA techniques  72  4.4.1 Site-directed mutation using PCR  72  4.4.2 Screening for mutants  76  4.5 Estimation of protein yields  76  4.6 Production of Cex and Cex mutants in E. coli  11  4.7 Protein mass determination  78  4.8 Confirmation of overall protein structure by circular dichroism spectroscopy  79  4.9 Enzymology  79  4.9.1 Determination of steady-state kinetic parameters  79  4.9.2 Determination of activity on CM-cellulose and xylan  80  4.9.3 Secondary deuterium kinetic isotope effect measurements  81  4.9.4 pH dependence studies  82  4.9.5 Burst experiment  82  4.9.6 Inactivation experiment  83  4.9.7 Determination of Extinction Coefficients  83  REFERENCES  85  APPENDIX A  91  BASIC CONCEPTS OF ENZYME CATALYSIS  91  A . l . Basic enzyme kinetics  92  A.2. The significance of the Michaelis-Menten parameters  95  A.3 Kinetics of inactivation of C. fimi Cex  99  APPENDIX B  101  BASIC INTRODUCTION TO LINEAR FREE ENERGY RELATIONSHIPS AND KINETIC ISOTOPE EFFECTS  101  vu B.l Linear free energy relationships  102  B. 1.1 General concept  102  B. 1.2 Linear free energy relationships in enzymology  102  B. 2 Kinetic isotope effects  105  B.2.1 General concept  105  B.2.2 Secondary deuterium kinetic isotope effects  106  APPENDIX C  109  GRAPHICAL REPRESENTATION OF KINETIC DATA  109  C. l Lineweaver-Burk plots for the hydrolysis of aryl (3-cellobiosides by Cex at 25° C, pH 7.0  110  C.2 Lineweaver-Burk plots for the hydrolysis of aryl (3-cellobiosides by Cex at 25°C, pH 5.5  111  C.3 Lineweaver-Burk plots for the hydrolysis of aryl (3-cellobiosides by the His205Ala mutant at 25° C, pH 7.0  112  C.4 Lineweaver-Burk plots for the hydrolysis of aryl (3-cellobiosides by the His205Ala mutant at 25° C, pH 5.5  113  C.5 Lineweaver-Burk plots for the hydrolysis of aryl [3-cellobiosides by the His205Asn mutant at 25°C, pH 7.0  114  C.6 Lineweaver-Burk plots for the hydrolysis of aryl (3-cellobiosides by the His205Asn mutant at 25°C, pH 5.5  115  C.7 Lineweaver-Burk plots for the hydrolysis of aryl (3-glycosides by the Gln87Met mutant at 37°C, pH 7.0  116  Vlll  LIST O F T A B L E S  Table 1.1 List of catalytic nucleophiles identified by 2-deoxy-2-fluoro-glycoside mechanism-based inactivators.  13  Table 2.1 Active site histidines hydrogen-bonded to catalytic nucleophiles.  30  Table 2.2 Active site tyrosines hydrogen-bonded to catalytic nucleophiles.  31  Table 2.3 Kinetic parameters for the hydrolysis of cellobioside substrates by Cex and His205 mutants at 25°C, pH 7.0.  38  Table 2.4 Kinetic parameters for the hydrolysis of cellobioside substrates by Cex and His205 mutants at 25°C, pH 5.5.  38  Table 2.5 Mass spectrometry of purified Cex and His205 mutants in the absence and presence of 2,4-DNPC.  41  Table 2.6 Br0nsted constants of logCk^/K,,,) vs pK for the hydrolysis of a  cellobiosides by Cex and His205 mutants. 45 Table 2.7 Secondary deuterium kinetic isotope effects on kcat for Cex and His205 mutants. Table 3.1 Properties of the individual amino acid residues.  50 63  Table 3.3 Kinetic parameters for the hydrolysis of 6-glycosides by Cex and the Gln87Met mutant at 37°C, pH 7.0. Table 4.1 Mutagenic primers used in site-directed mutation of Cex by PCR.  67 73  Table 4.2 Extinction coefficients of substituted phenols and phenyl cellobiosides at 25°C.  84  ix  LIST OF FIGURES  Figure 1.1 The structures of cellulose and xylan.  2  Figure 1.2 Synergistic model for cellulose hydrolysis.  3  Figure 1.3 Stereochemical classification of 6-glycosidases.  5  Figure 1.4 Double displacement mechanism proposed for retaining 6-glycosidases. 6 Figure 1.5 Structure of a glycosyl cation.  7  Figure 1.6 Resonance structures for an aldonolactone and an aldonolactam.  8  Figure 1.7 Nojirimycin, a transition state analogue for glycosidases.  8  Figure 1.8 Structure and reaction of activated 2-deoxy-2-fluoro-glycosides.  11  Figure 1.9 E. coli 6-galactosidase-catalyzed hydration of a heptenitol derivative.  15  Figure 1.10 Hydration of 2-acetamidoglycal by 6-N-acetyl hexosaminidases from Jack bean, bovine kidney and human placenta.  15  Figure 1.11 Structure of 6-D-galactosyl pyridinium cation.  16  Figure 1.12 Schematic representation of the intact exoglycanase (Cex).  18  Figure 1.13 Folding of the Cex catalytic domain.  20  Figure 1.14 Fluorocellobioside-binding region in Cex-cd.  21  Figure 1.15 Flurocellobioside interactions with Cex-cd.  22  Figure 1.16 Schematic diagram showing the Asp235-His205-Glu233 hydrogenbonding network in the active site of Cex.  23  Figure 1.17 Schematic diagram indicating the position of active site residues Gln87 and Trp281 relative to the C-5 hydroxymethylene groups of the cellobioside moiety.  25  Figure 2.1 Diagram showing the hydrogen bonding system around the catalytic nucleophile, Glu233, in wild type Cex (A), and the possible arrangements in His205Asn (B) and His205Ala mutant (C).  33  Figure 2.3 CD spectra of Cex and His205 mutants.  34  Figure 2.4 Thermal stability of Cex and His205 mutants.  35  Figure 2.5 Cellobioside substrates with different leaving group abilities.  37  Figure 2.6 The burst experiment for His205Ala (A) and His205Asn (B) using 2,4-DNPC. Figure 2.7 Inactivation of Cex (A) and His205Asn mutant (B) with 2F-DNPC.  40 42  Figure 2.8 Schematic diagram of the glycosylation step for the hydrolysis of cellobiosides by Cex.  44  X  Figure 2.9 Br0nsted plots relating log(k t/Km) versus pK of the aglycone phenols ca  a  at pH 7.0 (A) and pH 5.5 (B).  46  Figure 2.10 Schematic diagram showing the deglycosylation step for the hydrolysis of cellobiosides by Cex. 49 Figure 2.11 pH profiles of log(k at/K ) for the hydrolysis of PNPC by Cex and C  ra  His205 mutants.  52  Figure 2.12 Schematic diagram showing possible ionizations of Glu233 and His205.  53  Figure 3.1 Structure of glutamine, methionine, histidine and tyrosine side chains. 63 Figure 3.2 Generation of Gln87Met mutant.  64  Figure 3.2 CD spectra of Cex and Gln87Met mutant.  66  Figure 3.3 Structure of glutamine and serine side chains.  68  Figure 4.1 PCR site-directed mutagenesis scheme for mutating Gln87 in Cex gene. 74 Figure A. 1 Plot of velocity versus substrate concentration for a typical enzymatic reaction. Figure A.2 A typical Lineweaver-Burk plot for an enzymatic reaction.  92 94  Figure A.3 Hypothetical Gibbs free energy diagram for a retaining glycosidase showing rate-limiting deglycosylation.  97  Figure B.l Br0nsted plots relating rates of C.fimi Cex-catalyzed hydrolysis of aryl |3-glucosides with the leaving group ability of the phenols.  103  Figure B.2 Br0nsted plots relating rates of C. fimi Cex-catalyzed hydrolysis of aryl fi-cellobiosides with the leaving group ability of the phenols.  104  Figure B.3 Energy profiles illustrating the change in zero point energy difference between H- and D-substituted species on going to the transition state. 107  ABBREVIATIONS AND SYMBOLS  2,4-DNPC  2",4"-dinitrophenyl 6-D-cellobioside  2F-DNPC  2",4"-dinitrophenyl 2-deoxy-2-fluoro 6-D-cellobioside  3,4-DNPC  3" ,4" -dinitrophenyl 6-D-cellobioside  3,4-DNPG  3',4'-dinitrophenyl 6-D-glucoside  3,4-DNPX  3',4'-dinitrophenyl 6-D-xyloside  3,4-DNPX2  3",4"-dinitrophenyl 6-D-xylobioside  A  absorbance  Ala  alanine  Amp  ampicillin  Asn  asparagine  Asp  aspartate  Big  Br0nsted constant  bp  base pair  BSA  bovine serum albumin  CD  circular dichroism  Cex  Cellulomonas find 6-1,4-exoglycanase  CMC  carboxymethylcellulose  DMSO  dimethyl sulfoxide  EDTA  ethylenediamine tetra-acetic acid  ESI-MS  electrospray ionization mass spectrometry  Gin  glutamine  His  histidine  IPTG  isopropyl-6-D-thiogalactoside  K2HPO4  di-potassium hydrogen orthophosphate (monobasic)  kb  kilobase pairs  kcat  enzyme turnover number  kDa  kiloDaltons  Km  Michaelis constant  k bs  pseudo-first order rate constant  LB  Luria-Bertani medium  Met  methionine  min  minute  M  relative molecular mass  0  r  NaCl  sodium choride  NMR  nuclear magnetic resonance  PCR  polymerase chain reaction  PhC  phenyl B-D-cellobioside  Phe  phenylalanine  PhX2  phenyl 6-D-xylobioside  PNPC  4"-nitrophenyl 6-D-cellobioside  PNPG  4'-nitrophenyl 6-D-glucoside  PNPX  4'-nitrophenyl 6-D-xyloside  PNPX2  4"-nitrophenyl 6-D-xylobioside  s  second  SDS-PAGE  sodium dodecyl sulfate-polyacrylamide gel electroph  TE  tris-EDTA  TYP  tryptone, yeast extract, phosphate  Tyr  tyrosine  Vmax  maximum velocity of enzyme-catalyzed reaction  XIU  ACKNOWLEDGEMENTS  I would like to thank my supervisor, Dr. Steve Withers, for continual encouragement and advice throughout this study. I am also grateful to Dr. Tony Warren (Department of Microbiology, University of British Columbia) for including me as a part of his laboratory, and for his input into this project. My appreciation also goes to Dr. David Rose and Mr. Andre White (Department of Biophysics, University of Toronto) for collaborating on this research. Special thanks to my coworkers in the Withers' lab and the Cellulase group for their assistance, helpful discussions and especially their friendship, to Mr. David Chow and Dr. Shourning He for perforrning mass spectrometric analysis, and also to Mr. Mark Nitz and Ms. Margaret Li for their contribution to the project. Lastly, but not last, I would like to thank my friends and family for their love, support and encouragement.  CHAPTER 1 INTRODUCTION  2  1.1  General  introduction  Considerable effort has been directed towards the understanding of structurefunction relationships in enzymes which hydrolyze plant biomass. This derives from their importance in nature and their great potential for biotechnological applications in industry, such as in biomass degradation and fuel production, textiles, manufacture of pulp and paper, beverage and food processing (Coughlan & Hazlewood, 1993). Cellulose and xylan, the major components of plant biomass, are linear polymers of 6-1,4-linked D-glucose or D-xylose residues respectively (Figure 1.1).  Figure 1.1  The structures of cellulose and xylan (R=H, xylan; R=CH OH, 2  cellulose).  Biodegradation of these 6-1,4-glycans involves the action of 6-1,4-glycosidases, hydrolytic enzymes which act co-operatively to convert these polymeric, largely insoluble substrates into their constituent simple sugars. For example, cellulases are depolymerizing enzymes capable of hydrolyzing the B-l,4-glucosidic bonds of cellulose. The enzymes which cleave the internal 8-1,4 bonds are classed as endoglucanases (EC 3.2.1.4), while those that cleave off cellobiose units from the ends of cellulose chains are exoglucanases or cellobiohydrolases (EC 3.2.1.91). These enzymes act synergistically (Figure 1.2) as the action of endoglucanases on cellulose greatly increases the number of free non-reducing ends, which are sites for hydrolysis by exoglucanases. The final product of cellulose degradation, cellobiose, can then be cleaved to glucose by B-glucosidases (EC 3.2.1.21).  3 Similarly, xylan degradation involves xylanolytic systems that include B-l,4-xylanases (EC 3.2.1.8) and 6-xylosidases (EC 3.2.1.37); the former are generally considered to hydrolyze the xylan backbone, whereas the latter work to hydrolyze xylo-oligomers.  ii Crystalline region  i  Amorphous region A d s o r p t i o n of c e l l u l a s e s Endoglucanases  Figure 12  Synergistic model for cellulose hydrolysis (Adapted from Beguin & Aubert, 1994). Shaded glucopyranosyl residues indicate the reducing ends of the  cellulose chains.  4  8-Glycosidases are produced by a variety of organisms including microorganisms and plants. Many of these enzymes have a modular organization and comprise both catalytic and non-catalytic domains. Nearly all can be assigned to one of the eleven families in the recent classification of 6-glycosidases (Henrissat et al., 1989; Henrissat & Bairoch, 1993) on the basis of amino acid sequence alignment of their catalytic domain. Members of a single family group display similar catalytic mechanism and specificity. Families 5-10 contain endo- and exo-B-l,4-glucanases, although some members also have low activity towards related B-l,4-glycans. For example, significant B-l,4-xylanase activity is shown by several members of family 5 and 10. Family 11 contains only xylanases. Whereas mechanistic and specificity data are relatively abundant for families 5-11, there is as yet no full characterization of the catalytic activity of glycosidases from the most recently discovered families 12, 26, 44 and 45. Knowledge of the structure-function relationships of these enzymes is essential for their targeted bioengineering for use in the biotechnology industry. Much of the present understanding of the mode of action of such glycosidases and related enzymes has been delineated from biochemical and structural studies, discussed in some detail below.  1.2  Mechanism "retaining"  tor hydrolysis  of a 8-1,4-glycosidic  linkage  by  glycosidases  The most important mechanistic aspect of 8-glycosidases is the stereochemical outcome of the catalyzed reaction. Hydrolysis may occur with either inversion or retention at the anomeric centre of the reducing sugar product (Figure 1.3). Those glycosidases that hydrolyze the 8-1,4-glycosidic bond with net inversion of configuration at the anomeric carbon are termed inverting enzymes, and those that do so with net retention of anomeric configuration are termed retaining enzymes (Sinnott, 1990). This work will focus on retaining glycosidases.  5  "retaining"  .OH  .OR  OH  Figure 1.3  Stereochemical classification of 3-glycosidases.  It is generally accepted that hydrolysis of a B-l,4-glycosidic bond by a retaining glycosidase follows a double-displacement mechanism involving the formation and hydrolysis of a covalent cc-D-glycosyl-enzyme intermediate via oxocarbonium ion-like transition states (Koshland, 1953; Sinnott, 1990) as illustrated in Figure 1.4. A number of amino acids must be involved in binding and stabilizing the transition state complexes. Two active site residues, the catalytic nucleophile and the acid/base catalyst, play particularly important roles. In the formation of the glycosyl-enzyme intermediate (glycosylation step), the acid/base catalyst donates a proton to the glycosidic oxygen which facilitates bond cleavage by stabilizing the leaving group. The formation of the glycosyl-enzyme intermediate involves development of a covalent bond between the carboxyl group of the catalytic nucleophile and the anomeric carbon (C-l) of the sugar. The second step (deglycosylation step) involves the hydrolysis of the glycosyl-enzyme intermediate. In this step, the acid/base catalyst acts as a general base deprotonating the water, which then displaces the catalytic nucleophile resulting in a product which retains the 6-configuration at the anomeric centre.  Figure 1.4  Double displacement mechanism proposed for retaining 8-glycosidases.  7  A variety of experimental strategies has served to support the above mechanism of action of retaining glycosidases, some of which are presented in the following sections.  1.2.1 Oxocarbonium ion-like transition states  Much of the evidence for the existence of oxocarbonium ion transition states is derived from studies of transition state analogues, competitive inhibitors which resemble the proposed structure of the transition state in that they have one or both of the following characteristic features of a glycosyl cation: C-l and 0-5 atoms share a full positive charge and the C-l, C-2, C-5 and 0-5 atoms are coplanar (Figure 1.5). As they mimic the transition state, such compounds bind to the active site of glycosidases significantly tighter than do the normal glycoside substrates.  OH  Figure 1.5  Structure of a glycosyl cation.  Classical transition state analogues include the aldonolactones and aldonolactams which resemble the glycosyl cation both in geometry and to some extent in charge (Figure 1.6).  These compounds have lfj2- to 10 -fold enhanced binding relative to the  corresponding hexoses (Legler, 1990).  4  8  Aldonolactam  Figure 1.6  Resonance structures for an aldonolactone and an aldonolactam.  The 5-amino-5-deoxy-aldose class of transition state analogues such as nojirimycin (Figure 1.7) are among the tightest binding glycosidase inhibitors. These compounds exist in several forms, and when protonated, can be both isosteric and isoelectronic with a glycosyl cation.  Figure 1.7  Nojirimycin, a transition state analogue for glycosidases.  Further evidence for the oxocarbonium ion nature of the transition states is the observation of a positive secondary deuterium kinetic isotope effect  (kfj/ko > 1)  for the  9  glycosylation and the deglycosylation steps. For example, kinetic isotope effects measured on the glycosylation transition state include values of kn/kD = 1.15 - 1.20 for E. coli (lac Z) 6-galactosidase (Sinnott & Withers, 1974), kj^/kD = 1-05 - 1.07 for Agrobacterium 8glucosidase (Kempton & Withers, 1992), and kn/kD = 1-06 - 1.12 for C. fimi exoglycanase (Tull & Withers, 1994). Similarly, kinetic isotope effects have also been reported on the deglycosylation transition state and these include values of kn/kD =1.21.25 for E. coli B-galactosidase (Sinnott, 1978), kjj/kD = 1-09 for Botrydiplodia theobromae B-glucosidase (Umezerike, 1988) and kjj/kD = 1.10 - 1.12 for C. fimi exoglycanase (Tull & Withers, 1994). The presence of a positive kinetic isotope effect is associated with a change in the hybridization state of the a carbon from sp^ to sp2 as the substrate goes from the ground state to the transition state. This is therefore consistent with the transition states for formation and hydrolysis of the intermediate having significant S N I character. If however, a value of kn/kD = 1 was measured, then that would indicate either an SN2 mechanism, as there would be no change in hybridization, or that another step was rate determining. Alternatively, inverse isotope effects (values of kpj/kD < 1) are indicative of a change in hybridization from sp2 at the ground state to sp3 at the transition state, suggesting the presence of an ion pair intermediate. Thus, the kinetic isotope effect measurements presented above, kpj/kD > 1, are entirely consistent with oxocarbonium ion character at both transition states.  1.2.2 Nature of the glycosyl-enzyme intermediate  Kinetic isotope effect measurements not only indicate that the transition states for the formation and hydrolysis of the glycosyl-enzyme intermediate have substantial oxocarbonium character (section 1.2.1), but they offer strong evidence for the covalent  10  nature of the glycosyl-enzyme intermediate. The positive kinetic isotope effects on the deglycosylation step reflect significant sp^ to sp2 rehybridization at the anomeric centre (C1) in going from the ground state glycosyl-enzyme intermediate to the deglycosylation transition state, and this can only occur if the intermediate has more sp3 character than the subsequent transition state, consistent with the proposed covalent glycosyl-enzyme intermediate. Furthermore, it is estimated that a glycosyl cation in water is relatively unstable, with estimated lifetime on the order of 10~10 seconds (Bennet & Sinnott, 1986), compared to lifetimes of 1-100 milliseconds for gly cosy 1-enzymes at ambient temperatures (Weber & Fink, 1980). It therefore seems unlikely that such an unstable species could exist, even within the stabilizing environment of an active site, without formation of a covalent bond (Sinnott & Souchard, 1973). Additional evidence for the covalency of the intermediate comes from studies with 2-deoxy-2-fluoro-B-D-glycosides, mechanism-based inactivators which function via the formation and accumulation of a relatively stable glycosyl-enzyme intermediate (Figure 1.8A) (Withers et al., 1987, 1988). These compounds inactivate B-glycosidases because the electronegative fluorine at C-2 destabilizes both oxocarbonium ion-like transition states, thereby slowing down both the formation and the hydrolysis of the intermediate, while the presence of the good leaving group (2,4-dinitrophenolate) speeds up the glycosylation step thereby leading to the accumulation of the intermediate. 19p_NMR studies of an inactivated Agrobacterium (3-glucosidase have demonstrated the formation of a covalent a-D-glycosylenzyme intermediate (Withers & Street, 1988), and furthermore, formation of a unique covalent linkage to a specific amino-acid and thus identification of the catalytic nucleophile have been demonstrated by identification and sequencing of the glycopeptide (Withers et al., 1990). This strategy has been used for other glycosidases to identify the catalytic nucleophile (Table 1.1, section 1.2.3.) and show that such glycosyl-enzyme species exist and are stable. In addition, the catalytic competence of the intermediate has been demonstrated by the fact that it turns over in the presence of a suitable glycosyl acceptor via  11  a transglycosylation reaction, thus resulting in reactivation (Figure 1.8B) (Withers et al., 1990). A. Inactivation  B. Reactivation via transglycosylation  Figure 1.8  Structure and reaction of activated 2-deoxy-2-fluoro-glycosides (R = benzene group).  1.2.3 Presence of the catalytic nucleophile  X-ray crystallographic analyses of enzymes or of enzyme/inhibitor complexes often reveals residues that are suitably positioned to act as catalytic residues. The hen's egg-white lysozyme (HEWL), a retaining 8-glycanase, was the first enzyme for which the threedimensional structure was determined by X-ray crystallography (Blake et al., 1965). An aspartate residue, Asp52, was observed to be correctly positioned in the active site of HEWL to function as the nucleophilic catalyst proposed by Koshland (1953). Later, X-ray  12  crystallographic studies on T4 lysozyme (Anderson et al., 1981) suggested that residue Asp20 plays a similar role to Asp52 in HEWL. Recent X-ray crystallographic studies on other retaining glycosidases including Trichoderma harzianum xylanase (Campbell et al., 1993), Cellulomonas fimi exoglycanase (White et al., 1994), E. coli 6-galactosidase (Jacobsen et al., 1994), and Streptomyces lividans Xylanase A (Derewenda et al., 1994) have also identified carboxylate residues appropriately located within the active site to function as nucleophiles. Even more evidence for the existence of a catalytic carboxylate is derived from glycosidases co-crystallized with either an inhibitor or a substrate in the active site. For example, the X-ray crystal structure of Bacillus circulans E172C xylanasexylotetraose complex revealed that Glu78 is correctly oriented and within close enough proximity of the substrate to act as the catalytic nucleophile (Campbell et al., 1993). Similarly, the structure of C. fimi exoglycanase with 2-deoxy-2-fluoro-B-D-cellobioside covalently bound at the active site confirms that Glu233 is the carboxylate nucleophile (White et al., 1996), while the X-ray structure of Trichoderma reesei cellobiohydrolase I complexed with 2-iodobenzyl-l-thio-G-D-cellobioside suggests Glu212 as the nucleophilic catalyst (Divne et al., 1994). In other cases, the catalytic carboxylate groups have been identified by labeling experiments using group-specific labels. For example, Glu87 has been proposed to be the catalytic nucleophile in the active site of Schizophyllum commune xylanase A using the carboxylate-modifying  reagent,  1 -(4-azonia-4,4-dimethyl-pentyl)-3-ethylcarbodiimide  iodide (EAC) (Bray & Clarke, 1994). Labeling experiments using the mechanism-based inactivators 2-deoxy-2-fluoroglycosides have successfully identified the catalytic nucleophiles in the active sites of several retaining glycosidases (Table 1.1). For more details see discussion in section 1.2.2. Identification of some of these nucleophilic catalysts has also been confirmed by the X-ray crystallographic structures of the enzymes, which indicate that these residues are correctly  13  positioned within the active site to perform such a role (White et al., 1994; Jacobsen et al., 1994).  Table 1.1  List of catalytic nucleophiles identified by 2-deoxy-2-fluoro-glycoside mechanism-based inactivators.  Glycosidase  Nucleophile  Inactivator  Reference  Agrobacterium P-Glucosidase  Glu358  2F-DNPG  (Withers et al., 1990)  C. fimi Exoglycanase  Glu233  2F-DNPG  (Tull etal., 1991)  C. thermocellum Endoglucanase C  Glu280  2F-DNPC  (Wang et al., 1993)  Glu537  2F-DNPGal  (Gebler et al., 1992)  2F-DNPG  (Miao et al., 1994)  E. coli P-Galactosidase  Human Glucocerebrosidase  B. subtilis Xylanase  Glu340  Glu78  2F-DNPX  2  (Miao et al., 1994)  14  1.2.4 General acid assistance  The mechanism proposed by Koshland for a retaining 6-glycosidase suggested that departure of the aglycone may proceed with general acid assistance. In support of this evidence, available 3D structures suggest suitably positioned carboxylic acid residues that could function as proton donor groups. From the crystal structure of FfEW lysozyme (Anderson et al., 1981), E. coli 6-galactosidase (Jacobsen et al., 1994), and S. lividans Xylanase A (Derewenda et al., 1994), residues Glu35, Glu461 and Glul28, respectively, were observed to be in appropriate positions to fulfill such a function. Even in the absence of structural information, highly conserved carboxylic acid residues, identified by means of sequence alignment, could be targeted for mutagenesis, and their role as acid catalysts probed by detailed kinetic analysis using substrates for which protonation of the leaving group is required compared with substrates requiring no acid assistance. By this means, Glu 127 was identified as the acid catalyst in the C. fimi exoglycanase (MacLeod et al., 1994) and this assignment has been confirmed by X-ray crystallographic analysis of the enzyme (White et al., 1994,  White et al., 1996). Similarly, on the basis of  mutagenesis/kinetic studies, Glu 170 is proposed to be the acid/base  catalyst in  Agrobacterium 8-glucosidase (Wang et al., 1995). Other evidence for strategically-placed acidic groups is derived from the E. coli Bgalactosidase hydration of C-l-unsaturated heptopyranosyl derivatives. Based on the stereochemistry of the hydrated product, protonation occured from the B-face of the galactose ring (Figure 1.9) (Lehmann et al., 1983), consistent with the presence of a residue in this enzyme that could function as a proton donor in the normal glycoside hydrolysis.  15  Figure 1.9  E. coli 3-galactosidase-catalyzed hydration of a heptenitol derivative.  Similar studies with B-N-acetyl hexosaminidases using 2-acetamidoglucal indicated that protonation was from the 8-face of the sugar ring, leading to formation of N-acetylglucosamine rather than N-acetyl-mannosamine (Figure 1.10) (Lai & Withers, 1994), and again this is consistent with the existence of an acid catalyst that could donate a proton in normal glycoside hydrolysis. However, in most cases, protonation of glycals occurs from the sugar face that is opposite to the acid catalyst (Sinnott, 1990). For example, stereochemical investigation of the hydration of cellobial by Aspergillus niger endo-B-(l,4)cellulase and Irpex lacteus exo-6-(l,4)-cellulase revealed that protonation occurred from the  Figure 1.10  Hydration of 2-acetamidoglycal by B-N-acetyl hexosaminidases from Jack bean, bovine kidney and human placenta.  16  a-face of the cellobiose ring (Hehre et al., 1977), the proton being delivered by the catalytic nucleophile rather than by the acid catalyst (Hehre et al., 1977; Kanda et al., 1986). Although general acid catalysis may assist aglycone departure, it is not essential for all substrates. The primary evidence for this is provided by the study of E. coli Bgalactosidase-catalyzed hydrolysis of 6-D-galactopyranosyl pyridinium salts (Figure 1.11). For such compounds, protonation of the leaving group is structurally impossible. Yet, rate enhancements of 10^ to lO^-fold for the enzymatic hydrolysis compared to spontaneous hydrolysis were observed (Jones et al., 1977), suggesting that acid catalysis may not be crucial in the hydrolysis of normal substrates.  OH  Figure 1.11  1.2.5  Structure of 6-D-galactosyl pyridinium cation.  Non-covalent enzyme-substrate interactions  Though nucleophilic and acid/base catalysis may play an important role in catalysis with glycosidases, non-covalent interactions between the enzyme and the substrate function to generate substrate specificity and to aid in catalysis, accounting for the majority of the catalytic power of these enzymes. The binding energy derived from these interactions is believed to stabilize the transition state and lead to a decrease in the activation energy of the reaction, resulting in rate acceleration.  17  Such interactions have been identified in the X-ray structures of several carbohydrate-binding proteins and glycosidases, most of which represent hydrogen bonds between the hydroxyl groups of the sugar and the enzyme. The importance of individual non-covalent interactions can be systematically explored using site-directed mutagenesis to remove the side chain on the protein which was reacting with the substrate, or through structure/activity correlation using modified substrates. The deoxy-glycoside substrates have been used to identify important non-covalent interactions in the catalytic mechanism of glycosidases. For example, with A. wentii 6glucosidase, the enzymatic rate of hydrolysis drops by at least a factor of 10 when the C-2 6  hydroxyl group of glucose is removed and replaced by a proton. The C-4 hydroxyl group is also important, with the 4-deoxy substrates being hydrolyzed 10^ to lO^-fold slower (Roeser & Legler, 1981). Similar studies with E. coli 8-galactosidase suggest the importance of the interactions at the 2 position, since hydrolysis of 2-deoxy-galactosides is lO^-fold slower than that of the normal substrates (Wentworth & Wolfenden, 1974). Interactions at the C-4 and C-6 hydroxyl groups must be less important, since rate reductions of only 500- and 1000-fold were observed with 4-deoxy and 6-deoxy analogues (McCarter et al., 1992). With Agrobacterium 8-glucosidase, hydrogen-bonding interactions at the 2 position also appear most important, with rate reductions of approximately 5x10^fold being observed for hydrolysis of 2-deoxy-glucosides. The interactions at the 3 and 6 positions fulfill a similar function, though they are much smaller in magnitude (Namchuk & Withers, 1995). Similarly, the C-2 hydroxyl group in C. fimi exoglycanase must also provide important non-covalent interactions at the transition state since the rate of hydrolysis of the 2-deoxy-cellobiosyl-enzyme intermediate is lO^-fold lower than that for the corresponding cellobiosyl-enzyme (Tull, 1995).  18  1.3  Cellulomonas  fimi R-1,4-exoglycanase  1.3.1  General background  (Cex)  The enzyme used in the present study is the exoglycanase from the soil bacterium, Cellulomonas fimi. The gene encoding the exoglycanase has been cloned, expressed into E. coli and subsequently sequenced (O'Neill et al., 1986a). When produced in C. fimi, the exoglycanase is glycosylated, but the recombinant form (Cex) is not. Glycosylation has no apparent effect on catalytic activity; its main function appears to be protection against proteolysis (Langsford et al., 1987). The exoglycanase (Cex) is a 47 kDa protein comprising an N-terminal catalytic domain (35 kDa) and a C-terminal cellulose binding domain (12 kDa) held together by a linker peptide rich in proline and threonine residues (Figure 1.12). The domains in Cex retain their respective catalytic and cellulose-binding properties when separated by limited proteolysis (Gilkes et al., 1988, 1989).  47 kDa  Figure 1.12  Schematic representation of the intact exoglycanase (Cex).  C. fimi  Cex has been well characterized. Previous investigations of the  exoglycanase revealed it to be active on cellulose, xylan, and a range of soluble aryl B-Dglycosides (Gilkes et al., 1984; Tull et al., 1991). From stereochemical studies using ^HN M R , the enzyme was found to be a "retaining" B-glycosidase (Withers et al., 1986).  19  Hence, hydrolysis of substrates occurs with retention of anomeric configuration, likely by a double-displacement mechanism involving formation and hydrolysis of a covalent glycosyl-enzyme intermediate via oxocarbonium ion-like transition states, as proposed by Koshland (Figure 1.4, section 1.2). This mechanism implies the existence of two catalytic residues in the active site of Cex. One, the nucleophile, that is present in the ionized form and stabilizes an oxocarbonium ion transition state, subsequently forming a glycosylenzyme intermediate; the other acts as the acid/base catalyst, protonating the glycosidic oxygen of the scissile bond. The nucleophilic residue has been identified as Glu233, by trapping a covalent 2-deoxy-2-fluoro-glucosyl-enzyme intermediate and sequencing of the purified glycopeptide (Tull et al., 1991), and the assignment was later confirmed by mutagenesis/kinetic analysis (MacLeod, 1994). Site-directed mutagenesis at Glu 127 yielded mutants with kinetic parameters fully consistent with the role of acid/base catalyst for this residue (MacLeod et al., 1994). In addition, a detailed kinetic study of Cex by Tull & Withers (1994) allowed delineation of the mechanism through identification of rate-limiting steps, as well as investigation of the transition-state structure for each step. Such mechanistic insights, derived from pre-steady and steady-state kinetics, Br0nsted relationships, kinetic isotope effect measurements, inactivation experiments, and pH studies, provide further supporting evidence for the double-displacement mechanism of Cex and other B-glycosidases. Moreover, the determination of the X-ray crystal structure of the Cex catalytic domain (White et al., 1994) provides a structural basis for explaining the wealth of biochemical information that is available for Cex and related glycosidases. The Cex-cd folds into an eight-parallel-stranded a/(3 barrel, with an open cleft at the carboxyl terminal end, proposed to be the active site (Figure 1.13). The two catalytic glutamates, Glu233 and Glu 127, are located on either side of the cleft, being separated by a distance of 5.5 A, consistent with a retaining mechanism. Such a separation is presumably optimal for the efficient formation of a glycosyl-enzyme intermediate on Glu233 of Cex-cd, while at the  20 same time allowing Glu 127 to protonate the aglycone in a concerted manner (White et al., 1994).  Figure 1.13 Folding of the Cex catalytic domain (the two catalytic residues, Glu233 and Glu127, are shown in yellow).  21 Recently, diffusion of the mechanism-based inactivator 2,4-DNP-2-deoxy-2fluorocellobioside into crystals of Cex-cd has led to the formation of a covalent 2-deoxy-2fluorocellobiosyl-enzyme intermediate. The structure of this glycosyl-enzyme complex has been determined by X-ray crystallography (Figure 1.14) (White et al., 1996), and together with the structural information on the catalytic domain alone, provides more insight into interactions at the active site that are crucial to the catalytic mechanism and substrate preference of Cex, as discussed below.  Figure 1.14  Fluorocellobioside-binding region in Cex-cd.  22  W84  . E233 - nucleophilic catalyst .  Figure 1.15  E127 - acid/base catalyst  Flurocellobioside interactions with Cex-cd.  23 1.3.2 Importance of the Asp235-His205-Glu233 triad  The details of the atomic architecture of the active site of Cex, including positions of Glu233, Glul27, and other residues involved in substrate binding and catalysis, are shown in Figure 1.15. The interactions involving the key catalytic residues are likely important in determining their ionization behavior during catalysis. Of particular interest is an Asp235His205-Glu233 hydrogen bonding network that may play a role in creating the proper electronic environment in the active site and modulating the ionization state of the nucleophilic catalyst, Glu233 (Figure 1.16).  Glu233  His205  Figure 1.16 Schematic diagram showing the Asp235-His205-Glu233 hydrogenbonding network in the active site of Cex.  Such an arrangement of hydrogen bonding interactions around the catalytic nucleophile involving the Asp-Ffis-Glu triad has been observed in the active sites of other retaining 8-glycosidases with known 3D structures. These residues are highly conserved within each family, suggesting an important role in the catalytic mechanism of these enzymes. The nucleophile probably exists in a deprotonated form in the free enzyme to facilitate attack at the anomeric centre of substrate. The dyad Asp-His is most likely assisting the hydrolysis of the glycosyl-enzyme complex by stabilizing the developing  24  negative charge on the carboxylate group of the nucleophile. For a more detailed discussion see Chapter 2, section 2.1.  1.3.3 Xylanase vs. Cellulase Activity of Cex  On the basis of amino acid sequence alignments, Cex has been assigned to family 10 of 8-glycanases (Henrissat & Bairoch, 1993). Family 10 members are primarily categorized as B-l,4-xylanases although activity against cellobiosides has been reported for several of them (Grepinet et al., 1988; Luthi et al., 1990; Lin et al., 1991; Shareck et al., 1991; Haas et al., 1992). In addition, low activity against carboxymethylcellulose (CMC) has been reported in some instances (Luthi et al., 1990). Clearly, glycosyl hydrolases of family 10 have a mixed specificity for both xylan and cellulose, in contrast to the low molecular weight family 11 enzymes which are only active against xylan. Cex has been reported to hydrolyze xylan 40-fold more efficiently than cellulose. In addition, kinetic studies revealed that kcat/Km values for aryl-xylobioside hydrolysis are in fact 30-100 times higher than those for equivalent aryl-cellobiosides. Since xylan is a 131,4-linked polymer of D-xylose, a saccharide unit similar to glucose but lacking the hydroxymethylene group on the carbon C-5 (Figure 1.1, section 1.1) , it seems likely that the presence of this group is somehow inhibitory to catalysis. Comparison of the complexed and uncomplexed Cex-cd structures reveals that upon binding the 2fluorocellobioside moiety, the side chains of Gln87 and Trp281 residues reorient to accomodate the C-5 hydroxymethyl groups of the distal and proximal glucosyl units, respectively (Figure 1.17).  25  Glu127  —n  H H  HO  %  HN 2  Gln87  Figure 1.17 Schematic diagram indicating the position of active site residues Gln87 and Trp281 relative to the C-5 hydroxymethylene groups of the cellobioside moiety.  Interestingly, these rearrangements do not seem necessary upon binding of the 2fluoroxylobioside moiety (preliminary electron density map, D. Rose). This suggests that the presence of the C-5 hydroxymethylene groups introduces repulsive interactions within the active site of Cex, causing movement of Gln87 and Trp281. This likely results in less binding energy available for catalysis, consistent with lower rates of hydrolysis observed for cellulose relative to xylan.  1.4  Aims  of project  C. fimi exoglycanase (Cex) has been classified as a retaining B-glycosidase with mixed xylanase/cellulase activity. The overall objective of this study is to contribute to a  26  better understanding of the substrate specificity and catalytic mechanism of the exoglycanase. Structural analysis of the Cex catalytic domain reveals that the positioning of the Gln87 side chain within the active site may be important in controlling substrate recognition and specificity of Cex. A mutagenesis/kinetic analysis approach will be undertaken to study the effect of mutation of Gln87 on substrate binding and catalysis. Such research studies directed at understanding substrate recognition by Cex and family 10 enzymes will ultimately allow engineering of such enzymes with altered specificity for use in various biotechnological processes. Some applications, such as enzymatic prebleaching of kraft pulps for paper manufacture, require xylanases that are free of cellulase activity, while others would benefit from enzymes with dual specificity. Secondly, this study aims to probe the role of His205 in the active site of Cex by detailed kinetic analysis. Studies have revealed that this histidine residue is conserved in the enzyme's sequence related family. It is believed that His205 influences the ionization state of the catalytic nucleophile by hydrogen bonding, thus mechanistic investigations of His205Ala and His205Asn mutants may help in clarifying the role of this residue. This will involve steady state kinetic analysis, linear free energy relationship studies, secondary kinetic isotope effect measurements and pH-dependence studies. Linear free energy relationship studies with His205 mutants should identify the rate determining step and provide insights into the degree of negative charge accumulation on the phenolate oxygen at the glycosylation transition state. Secondary kinetic isotope effects should provide a measure of the oxocarbonium ion character at the deglycosylation transition state. Furthermore, the pH-dependence studies of the His205 mutants' activity may provide insights into the ionization state of the residues at the active site of this enzyme.  27  CHAPTER 2 PROBING THE ROLE OF HIS205 IN THE ACTIVE SITE OF C. FIMI CEX  28  2.1  Introduction  In an effort to gain insight into the reaction mechanism of glycosidases, amino acids that are found to be conserved within a family, based on amino acid sequence alignments, have been a target for mutagenesis and kinetic analysis. Not only are the presumed nucleophile and acid/base catalyst residues the subject of study, but this has been extended to include many other residues which are thought to be important in substrate binding and catalysis. In particular, histidine residues have been found to be located in conserved positions within the active sites of several glycosidases and glycosyl hydrolases, such as amylolytic enzymes (Ishikawa et al., 1992; Svensson,  1994; Qian et al., 1994),  cyclodextrin glucanotransferases (Namakura et al., 1993), cellulases and xylanases (Ducros et al., 1995; Bortoli-Germani et ah, 1995, White et al., 1994). Histidine residues have also been implicated in the mechanism of other transferases such as ADP-ribosylating toxins (Locht & Antoine, 1995) and hexose-1-phosphate uridylyltransferases (Kim et al., 1990; Arabshahi et al., 1996). For example, the imidazole ring of His 166 in the active site of hexose-1-phosphate uridylyltransferase is proposed to function as a nucleophilic catalyst in the reaction of glucose-1-P with UDP-galactose to form galactose-1-P and UDP-glucose by a double displacement mechanism, involving the formation of a covalent uridylyl enzyme intermediate (E-Hisl^.TjjMP). A mutant enzyme in which His 166 is replaced by glycine exhibits no catalytic activity, but added imidazole could assume the function of the "deleted" imidazole ring of histidine in the wild-type enzyme, resulting in the synthesis of imidazolyl-UMP (Kim et al., 1990). Further evidence for a histidine catalytic residue came from studies with ADPribosylating toxins that are capable of catalyzing the cleavage of the N-glycosidic bond of N A D and the transfer of its ADP-ribose moiety to G proteins. These toxins include +  29  pertussis toxin (PTX), E. coli heat labile toxin (LTX), cholera toxin (CTX), and mosquitocidal toxin (MTX) of Bacillus sphaericus. Investigation of PTX by site-directed mutagenesis followed by kinetic analyses of mutant proteins revealed that His35 is a catalytic residue because alterations of this residue affect the turnover rate of NADglycohydrolysis by approximately two orders of magnitude without significantly affecting substrate binding (Antoine & Locht, 1994). It is speculated that this histidine residue together with the previously identified catalytic Glul29 (Antoine et al., 1993) constitute an enzymatic diad responsible for the N-glycosidic cleavage of the NAD+ molecule (Locht & Antoine, 1995). The two catalytic residues are conserved in CTX, LTX, and MTX, and are believed to play similar roles in catalysis. Most likely, the catalytic role of the histidine residue is to increase the nucleophilicity of the acceptor substrates of these toxins through hydrogen bonding interactions. The effects of mutation of the three histidine residues that are found in most aamylases and in all known cyclodextrin glucanotransferases (CGTases) have also been examined, but the precise roles of these residues in binding and catalysis are not clear. Based on kinetic analysis of two mutants of Taka amylase A (TAA), His 122 and His296 were suggested to participate in the stabilization of the transition state, rather than in substrate binding in the ground state as proposed from the X-ray crystallographic studies (Matsuura et al., 1984), since mutants generated at these positions show significant reduction in kc t but no effect on the K a  m  values relative to the wild-type enzyme. Similar  results were obtained for the corresponding histidines in porcine and human pancreatic aamylases (Ishikawa et al., 1992; Ishikawa et al., 1993; Qian et al., 1994), barley aamylase (Svensson, 1994), and Bacillus sp. 1011 CGTase (Nakamura et al., 1993), suggesting that their roles in catalysis are conserved among amylolytic enzymes. In several families of cellulases and xylanases, histidines have been found to be located within the vicinity of the catalytic machinery, and their role is being further explored. For example, from the crystal structure of the cellulase endoglucanase A from  30  Clostridium cellulolyticum (CelCCA), His254 (corresponding to Gln203 in C. fimi Cex) is involved in forming a hydrogen bond with the acid catalyst, Glu 170, and therefore seems to be implicated in the catalytic reaction (Ducros et al., 1995). Indeed, this position is found to be intolerant to amino acid substitutions, in the case of the catalytic activity of cellulase EGZ of Erwinia chrysanthemi (Bortoli-Germani et al., 1995). Another interesting feature is the presence of a conserved histidine within hydrogen bonding distance of the catalytic nucleophile in the active sites of the retaining glycosidases of families 7 and 10, as listed in Table 2.1. Interestingly, the histidine residue in these enzymes is also found within hydrogen bonding distance of an active site aspartate, which leads to the postulate that such a catalytic triad might be a common structural and mechanistic feature of these enzymes. Perhaps the most likely role of the histidine residue  Table 2.1  Active site histidines hydrogen-bonded to catalytic nucleophiles (shown underlined).  Enzyme  Family  Triad of residues  Reference  C. fimi exoglycanase  10  Asp235-His205-Glu233  White et al, 1994  (Cex)  White et al., 1996  C. thermocellum Xylanase  10  Asp756-His723-Glu754  Dominguez et al., 1995  S. lividans Xylanase A  10  Asp238-His207-Glu236  Derewenda et al.,1994  P.fluorescensXylanase A  10  Asp248-His215-Glu246  Harris et al., 1994  T. reesei  7  Asp214-His228-Glu212  Divne et al., 1994  XynZ  Cellobiohydrolase I  31  is to control the ionization state and orientation of the key catalytic nucleophile by hydrogen bonding. By such interaction in the free state, it may help in maintaining the nucleophile in the proper ionization state needed for nucleophilic attack. This hydrogen bond would be weakened upon formation of the glycosyl-enzyme intermediate, such that, during the deglycosylation step, it could serve to stabilize the departing carboxylate group of the nucleophile. It is noteworthy that in the case of several other retaining glycosidase families, a conserved active site tyrosine, rather than a histidine, is hydrogen-bonded to the nucleophilic catalyst, as illustrated in Table 2.2. Similarly, it is proposed that the equivalent  Table 2.2  Active site tyrosines hydrogen-bonded to catalytic nucleophiles (shown underlined). Enzyme  Family  Diad of residues  Reference  Agrobacterium faecalis B-glucosidase  1  Tyr298-Glu358  Gebleret al., 1995;  E. coli B-galactosidase  2  Tyr503-Glu537  Jacobsen et al., 1994  C. cellulolyticum endoglucanase A (CelCCA)  5  Tyr256-Glu307  Ducros et al., 1995  5 •  Tyr200-Glu280  Dominguez et al., 1995  C. thermocellum endoglucanase CelC  Wang et al, 1995  Bacillus circulans xylanase  11  Tyr69-Glu78  Wakarchuk et al., 1994  Barley (1-3, l-4)-glucanase  17  Tyrl70-Glu232  Varghese et al., 1994  32  tyrosine residues function in both orienting the nearby nucleophile and stabilizing its deprotonated state in the free enzymes (Wang et al. 1995; Gebler et al, 1995). Thus, it appears that the protonation state of the catalytic nucleophile in retaining glycosidases is conditioned by hydrogen-bond interactions with a conserved active site tyrosine (families 1,2,5, 11 and 17) or a histidine (families 7 and 10) side chain. An understanding of how this is effected is critical for a better understanding of the mechanism of retaining glycosidases. The present investigation of C. fimi xylanase/cellulase (Cex) may provide some insight into this question. In the active site of this enzyme, there is a His205 residue within hydrogen bonding distance of the catalytic nucleophile, Glu233, and also within hydrogen bonding distance of Asp235 (Figure 2.1 A). Such hydrogen bonding interactions are most likely involved in maintaining the proper ionization state of the nucleophilic catalyst, Glu233, by creating a favourable electronic environment within the active site, as speculated above. Moreover, His205 is in very close proximity to the substrate reaction centre, which suggests a possible role of this residue in the reaction mechanism, presumably in the stabilization of the transition states. Here, we probe the catalytic importance of His205 in the active site of Cex through kinetic analysis of the site-directed mutant proteins His205Asn and His205Ala. The imidazole ring of histidine can function either in an acid/base or in hydrogen binding capacity. The asparagine substitution retains some of the hydrogen binding ability of histidine, but has no useful acid/base properties (Figure 2. IB). Alanine substitution was chosen to minimize unfavourable steric contacts and to remove any possible charge interactions or hydrogen bonds at that position (Figure 2.1C).  33  A.  \  Asp235  Glu233  His205 O.  °<y < NH  Glu233  ©  6  Asp235  Asn205 0  C.  ©  0^0^,0  CH  Glu233  Figure 2.1  o  Asp235  3  Ala205  Diagram showing the hydrogen bonding system around the catalytic nucleophile, Glu233, in wild type Cex (A), and the possible arrangements in His205Asn (B) and His205Ala mutant (C).  2.2  Purification  and physical  characterization  of His205  mutants  The DNA codons replacing His205 with Ala and Asn were introduced into the C. fimi gene by Mark Nitz (summer student in Withers' lab, 1995) via PCR site-directed mutagenesis, as outlined in Figure 2.2. The protocol is described in Materials and Methods (section 4.4.1). The recombinant pUC12-l.lcex(PTIS) containing the desired mutations was then transformed into E. coli Topp5 for expression of the mutant proteins, which were produced and purified as described in section 4.6. Essentially no differences in behavior during purification were observed, the mutants eluting at the same position as wild-type from the cellulose-affinity column. The purified proteins ran as single bands on SDSPAGE, at the same position (47 kDa) as wild-type Cex. Final protein yields ranged from  34 0.5 to 1 mg per litre of liquid culture. Mass spectrometric analysis confirmed the molecular weight of mutant proteins (Table 2.5), and CD spectra C M N U T I H I Ihul ni< >in|'ir changes in the secondary structure of the protein were introduced by mutations (Figure 2.3). However, the mutant proteins show decreased temperature stability as compared to the native Cex, as reflected in the lower melting temperature (T ) values shown in m  Figure 2.4. For this reason kinetic investigations were performed at 25°C rather than at 37°C, a temperature suitable for both mutants and wild-type Cex.  8000  1  1  1  1  1 Cex wild-type  1  4000  _  HltfOSAsn His205Asn  o E E u ec. K  3  >•>  "-3  -\\  a  s  //  -4000  - %j i  -8000 200  Figure 2.3  i 220  i  i  i  1  240 260 Wavelength (nm)  CD spectra of Cex and His205 mutants  1  1 280  1  300  35  20  30  40  50  60  70  T(°C)  Figure 2.4  2.3  Thermal stability of Cex and His205 mutants monitored by CD at 220 nm.  Kinetic analysis of Cex and His205 mutants  The initial kinetic characterization of the mutant enzymes consists of comparison to wild-type with respect to the fundamental steady-state kinetic constants: kcat> the turnover number, K , the apparent dissociation constant, and kcat/Km, the ii|i|iiin'nt m  second-order rate constant, often referred to as the specificity constant. Based on knowledge of the mechanism of Cex, examination of these kinetic parameters will yield information on how particular mutations affect the rates of the individual glycosylation and deglycosylation steps. The generally-accepted double-displacement mechanism for Cex (Figure 1.4) can be expressed as follows:  36 glycosylation  E + G-OR  k.- l  deglycosylation  1  E.G-OR  r  EG  H 0  ROH  where,  E + GOH  2  E = unbound enzyme G-OR = unbound glycoside substrate E.G-OR = Michaelis enzyme-substrate complex E-G = glycosyl-enzyme intermediate ROH = leaving group phenol GOH = sugar product k = first order rate constant for glycosylation 2  k = first order rate constant for deglycosylation 3  As discussed in Appendix A, the kinetic parameters, k t, K , and kcat/K ^ can be ca  m  m  defined in terms of the individual rate constants as follows:  cat  k +k 2  3  k (k + k_ kj(k + k ) 3  1  2  3  As can be seen from these expressions, provided deglycosylation is the ratedetermining step, mutations which reduce the rate of deglycosylation (relative to glycosylation) will result in a greater accumulation of the glycosyl-enzyme intermediate [EG] and therefore result in a reduction in K . This arises since if k3 « k2, then m  37  Therefore, K  m  decreases as k.3/k_ decreases. Similarly, a reduction in the rate of  glycosylation would be expected to increase the K . The effect of a mutation on the rate of m  the glycosylation step can also be estimated by analysis of kc t/Km which reflects the first a  ;  irreversible step in the reaction, presumably the rate of formation of the glycosyl-enzyme intermediate from the free enzyme and substrate. This can be seen from the fact that no k3 term appears with expression for kcat/K . A more detailed interpretation of kcat, Km, and m  kcat/Km is presented in Appendix A. Nitrophenyl cellobiosides with aglycones having different p K values (Figure 2.5) a  were tested as substrates for Cex and its His205 mutants (collaboration with Ms. Margaret Li, Biochemistry 449 student) as described under Materials and Methods (section 4.9.1). Steady-state kinetic studies were initially performed at pH 7.0, the reaction pH used in previous studies with the native enzyme, and results are presented in Table 2.3. However, due to changes in the optimum pH range for the His205Ala mutant (see pH profile, section SUBSTRATE  P a » aglycone phenol K  f  3.96  5.96  7.18  Figure 2.5  Cellobioside substrates with different leaving group abilities. The site of bond cleavage by Cex is indicated by the arrow.  38 2.3.2), kinetic parameters were redetermined at pH 5.5, and these are shown in Table 2.4. The average relative standard error in the data for individual experiments was 10-20%. For replicate experiments the average relative standard deviation was 5-10%.  Table 2.3  Kinetic parameters for the hydrolysis of cellobioside substrates by Cex and His205 mutants at 25°C, pH 7.0.  Enzyme  Substrate  k at (  s  C  Cex wild-type  His205Ala  His205Asn  Table 2.4 Enzyme Cex wild-type  His205Ala  His205Asn  )  K (mM)  k /K (s'mM")  m  1  cat  m  2,4-DNPC 3,4-DNPC  3.9 3.8  0.12  32.1  0.17  22.2  PNPC  3.5  0.56  6.2  2,4-DNPC  8.2 x 10"  0.007  1.2 x 10"  3,4-DNPC  8.6 x 10"  0.032  2.7 x 10"  PNPC  9.1 x 10"  0.82  1.1 x 10"  2,4-DNPC  4.9 x 10"  0.03  1.6 x 10'  3,4-DNPC  5.1 x 10"  0.083  6.1 x 10"  PNPC  4.6 x 10"  0.67  6.9 x 10"  4  4  4  4  4  4  1  2  3  2  3  4  Kinetic parameters for the hydrolysis of cellobioside substrates by Cex and His205 mutants at 25°C, pH 5.5. Substrate k at ( ) s  C  K (mM) m  WK  m  (s'W)  2,4-DNPC 3,4-DNPC  3.9  0.082  48  5.9  0.19  31  PNPC  4.1  0.70  5.9  2,4-DNPC  1.0 x 10"  0.0007  1.4 ± 0.6  3,4-DNPC  1.1 x 10"  0.023  4.8 x 10"  PNPC  1.2 x 10"  0.16  7.5 x 10~  2,4-DNPC  5.2 x 10"  0.008  7.4 x 10"  3,4-DNPC  6.6 x 10"  0.024  2.8 x 10"  PNPC  4.7 x 10"  0.19  2.5 x 10"  3  3  3  4  4  4  2  3  2  2  3  39  The data for the hydrolysis of these substrates by wild-type enzyme and His205 mutants were analyzed according to the scheme shown above. As can be seen, there is a marked decrease in kcat values relative to the wild-type. An approximately 4000 to 5000fold decrease is observed for His205Ala and 8000 to 9000-fold for the His205Asn mutant. Within each mutant, essentially the same kcat values were obtained for all substrates despite the different leaving group abilities of their aglycones, suggesting that the glycosylation step is unlikely to be rate-limiting. If it were, a dependence of kcat  o  n  the aglycone  structure should have been observed, since as the leaving group ability decreases kcat should also decrease, as the glycosylation step involves cleavage of the C-0 bond to the aglycone leaving group. As this was not observed, deglycosylation is most likely the rate determining step for the His205 mutants, as it is for the wild-type enzyme, since this step is necessarily independent of the aglycone leaving group. Further support that deglycosylation is the rate determining step is provided by the burst experiment for the His205 mutants with 2,4-DNPC, performed by following the release of dinitrophenol using the UV/VIS spectrophotometer, as described in Materials and Methods (section 4.9.5). As can be seen in Figure 2.6A, addition of 2.6 uM His205Ala to an excess (40 pM) of 2,4-DNPC resulted in a rapid initial release of dinitrophenol followed by a slower steady-state reaction. The linear portions of both phases were extrapolated back to time zero, and a burst magnitude of 2.4 pM dinitrophenolate was obtained. This corresponds well with the amount of enzyme present (2.6 pM), suggesting that this is indeed a true burst of dinitrophenolate arising from reaction of one equivalent of 2,4-DNPC with His205Ala mutant. The second, steady-state phase presumably arises from slow hydrolysis of the accumulated glycosyl-enzyme intermediate. The experiment was repeated with twice the amount of enzyme and the burst was also found to be increased two-fold (data not shown). Similar results were obtained with the His205Asn mutant, as illustrated in Figure 2.6B.  40  0  50  100  150  200  250  time (s)  0  1 0  I  1 100  I  I  I  200  300  •  I 400  i  I 500  time (s)  Figure 2.6  The burst experiment for His205Ala (A) and His205Asn (B) using 2,4DNPC.  Since the hydrolysis of the intermediate is rather slow, with a half-life [ti/2 = (ln2)/kcat] of approximately 14 minutes for His205Ala and 23 minutes for His205Asn, it was possible to observe the accumulation of the intermediate by mass spectrometric analysis. Reaction of the His205Ala(Asn) mutant with 2,4-DNPC prior to electrospray ionization mass spectrometric analysis resulted in an increase in molecular weight of the protein corresponding to the build-up of the glycosyl-enzyme intermediate (Table 2.5). Such an increase in mass could not be observed with the wild-type enzyme due to rapid hydrolysis of the intermediate.  41  Table 2.5  Mass spectrometry of purified Cex and His205 mutants in the absence and presence of 2,4-DNPC. Predicted  Determined M  M'  (a)  Native Cex  47 122.47  47 118 ± 5  nd*  nd*  His205Ala  47 056.41  47 058 ± 3  47 376 ± 4  320 ± 7  His205Asn  47 099.43  47 099 + 4  47 418 + 5  319 ± 9  Protein  r  r  Reacted with 2,4DNPC  1 , 2  Difference (b-a)  (b)  'predicted M is based on the amino acid sequence and is calculated as the average isotopic r  (MH+) mass The M of 2,4-DNPC is 508.4. The M of cellobiose (less OH) is 325.3 r  r  Substrate was added to the enzyme prior to mass spectrometric analysis. *n.d. = not determined  These results indicate that His205 plays a critical role in the reaction mechanism, particularly on the deglycosylation step, as reflected in the kcat values and the resulting accumulation of the glycosyl-enzyme intermediate. As mentioned previously, a measure of the effect on the glycosylation step can be determined by analysis of kcat/Km values, which reflect the first irreversible step of the reaction, most probably the formation of the glycosyl-enzyme  intermediate.  Thus,  kcat/Km values  were found  to decrease  approximately 300 to 5600-fold for the His205Ala mutant, larger decreases being observed for the His205Asn mutant (2000 to 9000-fold). Interestingly, in both cases the effects were greater with the substrates having poor leaving groups. It is known that as the leaving group ability decreases there is more requirement for acid catalysis to stabilize the departing leaving group. Thus, the results suggest that mutation of His205 likely removes some of the protonic assistance required. However, these effects could also be due to alterations in  42  the positioning and ionization of the catalytic nucleophile, Glu233, as this is situated within hydrogen bonding distance of His205. K  m  is the apparent dissociation constant and in this case, due to the dramatic  reduction in kcat, the decrease in K  m  likely represents a build-up of the glycosyl-enzyme  intermediate rather than a true increase in affinity for the substrates. As observed, the better the leaving group ability of the phenolate, the smaller the K , and thus the greater m  the accumulation of the intermediate. These lower K  m  values relative to wild-type  enzyme, in conjunction with the burst experiment and trapping of the glycosyl-enzyme intermediate by mass spectrometry, indicate that the activity observed is not due to contamination of completely inactive mutants by a small amount of wild-type; contaminating wild-type enzyme would rapidly hydrolyze the intermediate and release dinitrophenolate continuously, not in a burst. In addition, the values for k bs (pseudo0  first order rate constant of inactivation) with the His205Asn mutant and the wild-type enzyme, using the 2F-DNPC inactivator as described in Materials and Methods (section 4.9.5), were an order of magnitude different (Figure 2.7), indicating that the inactivation observed for His205Asn was not associated with the presence of wild-type activity.  time (min)  Figure 2.7  time (min)  Inactivation of Cex (A) and His205Asn mutant (B) with 2F-DNPC. Shown is residual enzymatic activity versus time.  43  The mutation of His205 therefore causes dramatic reductions in the rates of both the glycosylation step (formation of the glycosyl-enzyme intermediate) and of the deglycosylation step (hydrolysis of the glycosyl-enzyme intermediate). Similarly, changes in the rates of hydrolysis have been observed when mutating Tyr298, the Cex His205 equivalent in Agrobacterium 8-glucosidase, but to a lesser extent (Gebler et al., 1995). For example, with a Tyr298Phe mutant, there is a reduction of some 2000 to 3000-fold in kcat with substrates for which deglycosylation is rate-limiting, and some 400-fold slowing of the glycosylation step with the substrates for which glycosylation is rate-limiting. The rates of all substrates were reduced some 50 to 400-fold as measured through the kc t/K a  m  values. Similar to the results obtained for the His205 mutants, the relatively greater slowing of the deglycosylation step over the glycosylation step is also reflected in low K  m  values  seen for the more activated substrates.  2.3.1 Probing the effect of His205 mutation on the individual steps of the reaction  Although the above findings do not explain the exact role of His205 in catalysis, they clearly establish that this residue is of considerable importance. A more detailed analysis of the role of this histidine residue on the individual steps of the reaction mechanism was therefore performed.  (a) Glycosylation step: Linear free energy relationships provide deeper insight into understanding the effect of Ffis205 substitution on the glycosylation step. As described in Appendix B, this type of analysis correlates structural changes in the substrate with the activity of the enzyme, and can provide evidence regarding the degree of bond cleavage and charge distribution at the transition state.  44 Glul27  t  glycosyl-enzyme intermediate  Figure 2.8  Schematic diagram of the glycosylation step for the hydrolysis of cellobiosides by Cex (R = glucose residue).  A schematic view of the glycosylation step for the cleavage of the C-O bond by wild-type Cex is shown in Figure 2.8. Most likely, in going from the Michaelis enzymesubstrate complex (E.S) to the transition state complex (E.S+), the hydrogen bond between His205 and Glu233 is weakened, thus permitting nucleophilic attack of Glu233 on the  45  scissile bond. The acid catalyst, Glu 127, donates a proton to the glycosidic oxygen which facilitates bond cleavage by stabilizing the aglycone leaving group. It is anticipated that the protonated form of His205 may also play a similar role to Glu 127, assisting in the stabilization of the developing negative charge on the oxygen of the departing aglycone by hydrogen bonding interactions. Based on the modeling studies performed on the crystal structure of the 2-fluorocellobiosyl-Cex-cd complex, the His205 residue is located in the active site within approximately 3.3 A of the glycosidic oxygen, thus in a position potentially capable of forming a hydrogen bond. Presumably, disruption of such hydrogen bonding interactions by site-specific mutations will affect the extent of proton donation . In order to estimate the degree of negative charge accumulation on the oxygen of the aryl leaving group at the glycosylation transition state for the His205 mutants and the wildtype enzyme, values of log (kcat/Km) were plotted as a function of the phenol leaving group p K in the form of Br0nsted plots (Figure 2.9). The Br0nsted constants (Big) are a  given in Table 2.6. As discussed in Appendix B, plots of log(kcat/Km) vs p K having a a  large Br0nsted constant (Big near -1) reflect a large degree of negative charge build-up on the aglycone oxygen at the glycosylation transition state, which indicates either substantial C-0 bond cleavage or relatively little proton donation to the oxygen of the departing aglycone. Conversely, a smaller Br0nsted constant (Big « - 1 ) would indicate that there is either less C-0 bond breakage or more efficient proton donation.  Table 2.6  Bransted constants of log(k /K ) vs pKa for the hydrolysis of ca  m  cellobiosides by Cex and His205 mutants. Reaction conditions  Native Cex  Big His205Ala  pH 7.0  -0.23  -0.64  -0.43  pH 5.5  -0.29  -0.69  -0.46  His205Asn  46  Figure 2.9  Brensted plots relating log(kcat/K™) versus pK of the aglycone phenols at pH 7.0 (A) and pH 5.5 (B). a  As shown in Figure 2.9 and Table 2.6, larger reaction constants are observed for the His205 mutants relative to the wild-type, indicating a much greater degree of negative charge accumulation on the glycosidic oxygen at the glycosylation transition  47  state, suggesting that the mutations weaken the hydrogen-bonding interactions which serve to stabilize the leaving group. Thus, replacement of His205 by Ala or Asn results in less proton donation to the oxygen of the departing aglycone, which suggest that His205 could act directly as a proton donor or could indirectly affect the protonic assistance provided by the acid catalyst, Glu 127. It seems quite unlikely that the effects observed with the His205 mutants could be solely attributed to a reduction in the acid assistance from Glu 127, but this possibility cannot be ruled out. It is noteworthy that in a study by MacLeod et al., (1994), mutation of Glu 127 to Ala resulted in only modest changes (up to 20-fold reduction) in kc t/K for the a  m  hydrolysis of substrates which need relatively little protonic assistance (i.e., 2,4-DNPC, 3,4-DNPC, and PNPC), unlike the dramatic effects seen for the His205 mutants (300 to 9000-fold reduction) with the same substrates. It appears therefore that Glu 127 and His205 act independently. However, kinetic investigation of a His205Ala-Glul27Ala double mutant should determine whether the two mutations are independent or not. A completely additive effect on kcat/K would indicate that residues His205 and Glu 127 do not interact m  directly in the reaction mechanism that leads to the formation of the glycosyl-enzyme intermediate. Such studies are ongoing. Analogous studies with Agrobacterium B-glucosidase (Gebler et al., 1995; Wang et al., 1995) revealed essentially no changes in the degree of negative charge accumulation at the glycosylation transition state upon mutating the active site Tyr298 to Phe, which would make unlikely a direct role of Tyr298 in some form of acid catalysis during the formation of the glycosyl-enzyme intermediate, as proposed for Cex His205. Furthermore, the Tyr298 residue and the acid catalyst (Glu 170) in Agrobacterium 8-glucosidase appear to function independently, based on kinetic analysis of a double mutant in which both the Tyr298 and the acid catalyst, Glu 170, have been mutated. The changes observed with the Tyr298Phe  48  mutant on kcat/Km (50 to 400-fold reduction) may therefore be due to participation of Tyr298 in the orientation and activation of the catalytic nucleophile, and not in acid catalysis. In addition to its possible role in the acid catalysis at the glycosylation transition state in Cex, His205 may also be involved in controlling the ionization and/or positioning of the catalytic nucleophile, Glu233, which could account for the dramatic effects on the rates of glycosylation, reflected in the k^/Km values.  (b) Deglycosylation step: The mechanism for the deglycosylation step involves an oxocarbonium ion-like transition state, which requires a change in the hybridization state of the anomeric carbon (C-l) from sp3 to sp2. The extent of this rehybridization can be estimated by secondary deuterium kinetic isotope effect analysis, as discussed in Appendix B . A kinetic isotope effect of kn/kD > 1 (typically 1.1 to 1.3) is indicative of an S N I mechanism with an oxocarbonium ion-like transition state, whereas a kinetic isotope effect of kn/kD = 1 indicates an S N 2 mechanism, as there is no change in hybridization. This should provide valuable information pertaining to the structure of the deglycosylation transition state for the His205 mutants relative to the wild-type Cex. The hydrolysis of the glycosyl-enzyme intermediate by Cex is schematically shown in Figure 2.10. Glu 127 acts as a general base to deprotonate the water as it attacks at the anomeric centre of the glycosyl-enzyme intermediate to displace the catalytic nucleophile, Glu233. Some stabilization of the water hydroxyl group could be provided by hydrogen bonding to His205. This histidine residue may also be involved in stabilizing the developing negative charge on the carboxylate group of the Glu233 nucleophile. The dramatic effects on kc t observed upon mutating His205 to Ala or Asn would be consistent a  with such a role.  49  Figure 2.10  Schematic diagram showing the deglycosylation step for the hydrolysis of cellobiosides by Cex (R= glucose residue).  The degree of oxocarbonium ion character at the deglycosylation transition state is likely influenced by the degree of water preassociation and the leaving group ability of the glutamate nucleophile, Glu233. Since His205 is a likely candidate for assisting the  50  departure of Glu233, and may also be involved in assisting in deprotonation of the water molecule, substitution of this residue by Ala or Asn may affect the nature of the transition state. In order to determine the degree of oxocarbonium ion development at the deglycosylation transition state for His205 mutants, secondary deuterium kinetic isotope effects were measured on kcat using 2,4-DNPC as substrate, as described in Materials and Methods (section 4.9.3). Results are summarized in Table 2.7.  Table 2.7  Secondary deuterium kinetic isotope effects on k t for Cex and His205 ca  mutants. Enzyme Native Cex  kR/kD 1.10 ± 0.02*  His205Ala  1.12 ± 0.02  His205Asn  1.02 + 0.01  * Tull & Withers, 1994 The similar kinetic isotope effects measured for the native enzyme (kfj/kD = 1.10) and the His205Ala mutant (kjj/kD = 1.12) indicate that the extent of oxocarbonium ion development at the transition state for hydrolysis of the glycosyl-enzyme intermediate is similar in the two cases. These values of the kinetic isotope effects suggest that there is relatively little preassociation of the water nucleophile at the deglycosylation transition state for Cex and His205Ala mutant. By contrast, the smaller kinetic isotope effect (kn/kD = 1.02) measured for the His205Asn mutant indicates a transition state with lesser oxocarbonium ion character. This suggests a relatively early deglycosylation transition state for His205Asn with little bond cleavage and substantial preassociation of the water nucleophile. Together with the profound effects on kc t (8000 to 9000-fold decrease), such a  behavior would be consistent with a poorer enzymic "leaving group" Glu233 in the absence of stabilization from His205, thereby requiring greater nucleophilic assistance from the water, thus a more S]Sf2-like transition state. This value of the kinetic isotope effect is  51  comparable to that reported for this step with the Agrobacterium B-glucosidase Tyr298Phe mutant (kjj/kD = 1-03) (Gebler et al., 1995), thus supporting the similar role of Tyr298 in stabilizing the departure of the leaving group nucleophile Glu358. Interestingly, replacement of His205 with Ala apparently has no effect on kjj/kD* despite its significant effects on kcat (4000 to 5000-fold decrease). This mutation presumably generates a cavity in the active site (Figure 2.1C) which could be filled by a water molecule or the side chain of a nearby amino acid. Such an environment may somehow help to stabilize the released carboxylate group of the nucleophile in the His205Ala mutant. However, more definitive statements on this question await structural analysis of these mutants by X-ray crystallography. Remarkably, the extent of the C-O bond cleavage to the carboxylate in the deglycosylation step is not significantly affected by removing the acid/base catalyst or by truncation of the catalytic nucleophile; since the a-secondary deuterium kinetic isotope effects measured for hydrolysis of 2,4-DNPC by the acid/base catalyst mutant, Glu 127 Ala, and the nucleophile mutant, Glu233Asp, are comparable with that of the wild-type enzyme (Tull, 1995).  2.3.2 pH dependence study  The pH dependence of the Michaelis-Menten parameters for cellobioside hydrolysis upon mutation of His205 may provide some indication of the importance of the histidine in controlling the ionization states of active  site residues  involved in  substrate  binding/catalysis. The pH dependence of native Cex and the His205 mutants was investigated using PNPC, a substrate for which deglycosylation is rate limiting. Values for k t/K were determined by following the time course for the hydrolysis of PNPC at ca  m  concentrations much lower than K (0.2 x K ), since at [S] « Km, rates of hydrolysis are m  m  52 first order in substrate and the observed rate constants  (kob) correspond to kc /K S  at  m  (see  section 4.9.4). These results are presented as plots of log(kcat/Km) versus pH (Figure 2.11) and reveal ionizations of catalytic residues in the free enzyme. Values of kcat/Km for the native enzyme are seen to be dependent upon two ionizations, one of pK = 5.0 in a  the acidic limb of the pH profile and another of pK = 7.3 in the basic limb. These p K a  a  values are similar to those seen for the free enzyme when 2,4-DNPC was employed as substrate (pK = 4.1 and pK = 7.7) (Tull & Withers, 1994). Such pH activity curves do a  a  not provide much information on the actual identity, or number, of groups involved in binding or catalysis. However, the simplest explanation for the observed bell-shaped dependence on pH is that two catalytically-essential groups in the free enzyme can ionize, but the enzyme is active only when one is protonated and the other is not. The group in the free enzyme of pKa = 7.3 must be protonated to be catalytically active, then this ionization likely reflects that of the general acid catalyst, Glu 127, which is required to protonate the leaving group in the glycosylation step. The group in the free  <D  h  0  o  o  Cex wt  A  H205A  A  H205N  P o A  E  o  -2 A  J  I  I  I  I  A  I  A A  A  I  M  A  I  A  A  i  pH  Figure 2.11  pH profiles of log(k at/Km) for the hydrolysis of PNPC by Cex and His205 mutants C  53  enzyme of approximate p K = 5.0 must be deprotonated to be catalytically active, thus a  likely corresponding to the catalytic nucleophile, Glu233. No ionizations are revealed in the pH profile of His205Asn, likely due to rearrangements of hydrogen bonding interactions within the active site. However, a single ionization of p K = 6.2 in the basic limb, likely corresponding to the acid/base catalyst, is a  observed for the His205Ala mutant. The absence of an ionization at lower pH values upon removal of His205 may be due to a shift in the pK of the nucleophile outside the pH range a  studied. However, together with the drop in p K of the acid/base catalyst from 7.3 to 6.2 a  for the His205Ala mutant, this is not consistent with simple removal of positive charge from the active site; instead, this might indicate that mutation of His205 alters the electronic environment by disrupting important hydrogen bonding interactions. Perhaps the simplest explanation is that the p K of 5.0 observed in the native enzyme is that due to ionization of a  the proton shared between the nucleophile and the histidine, as shown in Figure 2.12. Such a deprotonation would render Glu233 more nucleophilic. Removal of His205 would then remove this ionization, as seen in the pH profiles of His205 mutants. The perturbation of  Asp235 Glu233  His205  Asp235 Glu233  Figure 2.12  His205  Schematic diagram showing the probable ionization states of Glu233 and His205.  54  the pK corresponding to the acid catalyst, Glu 127, is harder to interpret. Although His205 a  does not directly interact with Glu 127, it could likely be involved in hydrogen bonding/electrostatic interactions'which control the ionization of this residue. In light of the pH dependence of enzymatic reactions of other retaining glycosidases, especially in the case of mutant enzymes, it seems likely that changes in the structures of active sites are accompanied by changes in the protonation states of the catalytic groups. Evidence for this is provided by the study of Bacillus circulans Xylanase (Mcintosh et al., 1996), in which either mutation of the nucleophile, Glu78, to a neutral Gin or the glycosylation of this residue resulted in a decrease in the p K of the acid a  catalyst, Glu 127, by approximately 2.5 units. Similarly, mutation of Asp52 in hen egg white lysozyme to Asn, or ethylation of its carboxylate group, decreased the p K of the a  acid/base catalyst by approximately 1 unit (Parsons & Raftery, 1972; Inoue et al., 1992). Hence, the changes in the chemical environment of the active site of His205 mutants are not surprising, given the participation of His205 in an extended hydrogen-bonding network around the catalytic nucleophile. Although interpretation of the kinetic data is not trivial, the results suggest that Cex His205 is important in maintaining the proper ionization states of key catalytic residues in the free enzyme.  3.3  Concluding  remarks  His205 has been shown to play a critical role in the reaction mechanism of C. fimi exoglycanase (Cex). This histidine residue is found in a conserved position, within hydrogen bonding distance to the catalytic nucleophile, in the active sites of all glycosidase members of family 10 and 7 for which structural information is available to date, suggesting that it has a similar function in the two classes of enzymes. The data presented in this study indicate that His205 is required for both glycosylation and deglycosylation  55  steps; substitution by Ala or Asn significantly affects the rates of formation (kcat/Km) and hydrolysis (kcat) of the glycosyl-enzyme intermediate. Values of kcat/K are found to m  decrease some 300 to 5600-fold for His205Ala, and some 2000 to 9000-fold for the His205Asn mutant, depending on substrate. A greater effect is observed on kcat, with reductions of 4000 to 5000-fold for His205Ala and 8000 to 9000-fold for His205Asn. These effects are accompanied by an increased accumulation of the glycosyl-enzyme intermediate, as suggested by lower K  m  values coupled with mass spectrometric analysis.  During the glycosylation step, His205 is thought to enhance the nucleophilicity of Glu233, thus facilitating cleavage of the glycosidic bond. Additionally, His205 is likely involved in acid catalysis required to stabilize the oxygen of the leaving group aglycone at the glycosylation transition state, as suggested by the larger Br0nsted constants (Big values) measured on the glycosylation step for the His205 mutants. During the deglycosylation step, His205 may be important in increasing the nucleophilicity of the water molecule and stabilizing the released carboxylate group of the nucleophile; this is best reflected in the smaller kinetic isotope effects measured on the deglycosylation step for the His205Asn mutant. Thus, not only are the properties of His205 mutants consistent with the proposed role of His205 in modulating the ionization state of the catalytic nucleophile, but they also suggest a possible role of His205 in the stabilization of the glycosylation and deglycosylation transition states, via interaction with the glycosidic oxygen and the hydroxyl group of the water nucleophile, respectively. Whether His205 acts as a true acid/base catalyst in such interactions or it simply functions in a hydrogen bonding capacity is not as clear. The effects observed upon mutating His205 in Cex and the equivalent Tyr298 in Agrobacterium B-glucosidase are quite similar, but point out a slightly different function for the two residues, consistent with their characteristics. Although both histidine and tyrosine appear to function by modulating the pK of the catalytic nucleophile via hydrogen bonding a  56  interactions, the histidine residue might also provide some protonic assistance during the formation and hydrolysis of the glycosyl-enzyme intermediate in a role similar to that of the acid/base catalyst, thereby explaining the low Big values. Histidine, being a cationic acid, and having two possible protonation sites, is certainly a better candidate for such a role than is tyrosine.  57  CHAPTER 3 INVESTIGATION ON THE SUBSTRATE SPECIFICITY OF THE BIFUNCTIONAL C. FIMI XYLANASE/CELLULASE (CEX)  58  3.1  Introduction  Many P-l,4-glycanases hydrolyze a range of cellulosic substrates and artificial chromogenic substrates as well as xylans whereas others hydrolyze only the latter. Given the similarity in the chemical nature of the substrates, cross-specificity between cellulases and xylanases is not surprising. For example, although family 5 endoglucanases cleave internal 6-1,4-glucosidic linkages, producing rapid depolymerization of a model substrate, carboxymethylcellulose (CMC), several of these enzymes also display significant xylanase activity. The ratios of xylanase to CMCase activity observed are 1:10 for Clostridium thermocellum EGE (Hall et al., 1988) and 1:100 for C. thermocellum EGH (Yague et al., 1990), whereas Butyrivibrio fibrisolvens EG does not have any detectable xylanase activity (Berger et al., 1989). Conversely, family 10 enzymes predominantly hydrolyze xylan. For example, in the case of XynZ from Clostridium thermocellum (Grepinet et al., 1988), xylan is a 200-fold better substrate than is CMC. Family 11 enzymes, on the other hand, display only xylanase activity. The relaxed specificity of some xylanases versus the more restricted specificity of others must reflect differences between the active sites in the two groups of enzymes, those enzymes showing relaxed specificity accommodating the hydroxymethylene groups of cellulosic substrates while those of restricted specificity do not. A comparison of the binding environments within the active sites of cellulases and xylanases might therefore provide a rationale for their substrate preference. The structures of several such enzymes, in the presence and absence of inhibitors or slow substrates, have been determined over the last few years and these provide features of the active sites which accomodate both glucosidic and xylosidic residues. In all cases, a common distribution of aromatic residues and polar amino acids was found in the carbohydrate binding clefts, likely sites for the formation of hydrogen bonds with the hydroxyl groups of the sugar substrates (White et al., 1996; Ducros et al., 1995; Harris et  59  al, 1994; Wakarcuk et al., 1994; Spezio et al., 1993; Davies et al., 1993; Juy et al., 1992; Rouvinen et al., 1990; Philips, 1967; Strynadka & James, 1991). Initial studies on these systems have been an attempt to understand general enzyme-substrate binding interactions. Thus, residues which appear to be intimately involved in substrate binding have been a target for mutagenesis/kinetic analysis to further probe their roles. For example, kinetic analysis of a Tyr80Phe mutant of B. circulans xylanase (family 11) shows a 2000-fold decrease in the kcat value and an approximately 7-fold increase in K  m  for xylan, suggesting that Tyr80 may have a role in hydrogen bonding to  the substrate (Wakarchuk et al., 1994). As proposed from the X-ray crystallographic data of the xylanase-xylotetraose complex, this residue is within hydrogen bonding distance of the C-l of the second xylose unit of the xylotetraose substrate and also in close proximity of the enzymic acid/base catalyst, Glul72. Similarly, substitution of His 122 in the active site cleft of the cellulase endoglucanase A from Clostridium cellulolyticum (family 5) led to a drastic reduction in activity, which was less pronounced on substitution with phenylalanine (Belaich et al., 1992). The presence of a ring in both histidine and phenylalanine side chains, the rings being able to stack against sugar molecules, and the particular orientation of His 122 with respect to the active site (Ducros et al., 1995), support the proposal that Hisl22 is involved in substrate binding. This awaits the structure determination of an enzyme-substrate complex for further confirmation. It is interesting that, in the xylanase/cellulase Cex (family 10), a histidine (His80) is found in a similar position, as observed from the superposition of the two structures (Ducros et al, 1995), and this histidine is also known to be involved in recognition of the substrate molecule within the active site (Chapter 1, Figure 1.15) (White et al., 1996). Further evidence for oligosaccharide binding comes from the work on mutants of xylanase A from Streptomyces lividans (Moreau et al., 1994), a family 10 enzyme. On the basis of kinetic studies with a mutant in which the conserved residue Asn 173 was replaced  60  by Asp, it was suggested that this residue is involved in important hydrogen binding interactions with a xylose residue three units away from the cleavage site. However, stacking interactions of aromatic amino acids, tryptophans in particular, with the glycopyranosyl rings appear to be the most common binding interactions in these enzymes. Typically, the aromatic residues are located alternately on either side of the binding cleft, consistent with the arrangement of the hydrophobic faces of the pyranose rings in the substrate. For example, in the active site of the cellulase endoglucanase A from Clostridium cellulolyticum (CelCCA), a family 5 member, the side chains of Trp57 and Trp340 are situated on one side of the active site while Trp 180, Trpl81, Trp259, Trp287 and Tyr256 He on the opposite side (Ducros et al., 1995). Similarly, the B. circulans xylanase (family 11) contains six tyrosine and three tryptophan residues within the substrate binding cleft, likely involved in stacking interactions (Wakarchuk et al., 1994). In the active site of the exoglycanase (Cex) from C. fimi, which is a family 10 enzyme, the side chains of tryptophans 84, 281 and 273 are also involved in aromatic stacking interactions with the nonpolar faces of the cellobioside substrate (White et al., 1996). These are similar to the stacking interactions of Trp 135 and Trp367 with the glucose rings observed in the active site of T. reesei cellobiohydrolase II (CBHII) (Rouvinen et al., 1990). As more structural data become available, it is hoped that a larger number of residues will be identified and examined for their roles in substrate binding. Most likely, some amino acid positions involved in substrate recognition, invariant within each family but differing between cellulases and xylanases, may be associated with the distinct specificity for cellulose and xylan substrates. Although several studies on cellulases and xylanases have provided some information regarding positioning and binding stabilization of the cello- and xylo-oligosaccharide substrates within their active sites, they have not yet determined the specific residues involved in controlling their substrate preference. Understanding exactly how these enzymes accomplish this feat is still a major challenge.  61  The present study is a step towards localizing those particular residues which mediate the specificity of C. fimi exoglycanase (Cex). This enzyme is a family 10 bifunctional xylanase/cellulase, and thus shows preference for hydrolysis of xylan over cellulose. As discussed in Chapter 1, section 1.3.3, xylan and cellulose have highly similar chemical structures, the major difference being the presence of a hydroxymethylene group at the C-5 position on the sugar rings in cellulose (Chapter 1, Figure 1.1). Presumably, the observed lower specificity of Cex towards cellulose is due to destabilizing steric interactions induced within the active site by these C-5 hydroxymethylene groups. Thus, by targeting the residue(s) interacting with the hydroxymethylene groups by site-directed mutagenesis, it may be possible to block the binding sites for cellulose and impair the ability of Cex to catalyze cellulose breakdown. It is strongly desirable that such changes will not affect the xylanase activity of the enzyme. This approach has the potential to generate improved xylanase enzymes, free of cellulase activity, for the use in pulp and paper industry to reduce the amount of chemicals used in the traditional chemical bleaching stage of kraft pulps. It is known that parts of xylan initially dissolved in the kraft cooking liquor physically shield the residual lignin from bleaching chemicals. Thus, the disruption of xylan chains by xylanases would allow better access of bleaching chemicals to the residual lignin and easier extraction of lignin from pulp fibers. Cellulase activity in such xylanase preparations should be kept to a minimum or eliminated to avoid degradation of the cellulose fibrils during the bio-bleaching treatment. Thus, in order to undertake such "remodelling" of the binding site of Cex to confer an altered specificity to the protein, the binding interactions with the C-5 hydroxymethylene groups need to be considered. Inspection of the structure of the cellobiosyl-Cex-cd complex revealed that the conserved Gln87 and Trp281 residues lie within close proximity of the hydroxymethylene groups of the distal and the proximal glucosyl unit of the cellobioside substrate, respectively (Chapter 1, Figure 1.17). Interestingly, the comparison of the uncomplexed and complexed Cex-cd structures shows that the side chains of these  62  two residues must move away to "accomodate" the hydroxymethylene groups and to allow for binding of the respective glucosyl units. Moreover, this steric hindrance has not been observed upon binding a xylobioside substrate in the active site. Thus, a simplistic strategy to modify the binding site so that it would no longer accomodate a cello-oligosaccharide ligand is to replace the side chains of Gln87 and Trp281 with bulkier and potentially less flexible residues to compress the binding pocket and create a greater steric hindrance for the hydroxymethylene groups. This should not, however, be so extensive that it could not allow binding of xylo-oligosaccharides. On this basis, in an effort to reduce the cellulase activity of Cex while maintaining its xylanase activity, this investigation aims to replace Gln87 with bulkier residues via sitedirected mutagenesis and to determine the effect of mutations on substrate binding and specificity by kinetic analysis of generated mutant proteins. Evidently, replacement of the Gin side chain in the binding site could remove critical interactions, and in particular, disrupt the hydrogen bonds with the neighbouring residues, which could alter the structural integrity of the protein. In this regard, computer modelling was used to evaluate different amino acid replacements that would not only increase the bulkiness of the Gln87 side chain, but would also retain some of the hydrogen bonding capabilities of glutamine. The amino acids of choice were therefore methionine, tyrosine, and histidine. These residues have a larger molecular surface area, as shown in Figure 3.1 and also illustrated in Table 3.1. In addition, tyrosine and histidine have relatively rigid side chains; the side chain of methionine may still confer some flexibility, similar to that of glutamine. OH °VNH  Glutamine  Figure 3.1  C 2  N  3  Methionine  Histidine  Tyrosine  Structure of glutamine, methionine, histidine and tyrosine side chains.  63  Table 3.1  Properties of the individual amino acid residues*.  Residue  Residue mass (Da)  Van der Waals  pK  a  volume (A-*)  Side chain  Gin  128.14  114  Met  131.19  124  -  His  137.14  118  6.04 (imidazole)  Tyr  163.18  141  10.13 (phenol)  *Creigton, 1993  3.2  Generation  of Gln87  mutants  In vitro PCR site-directed mutagenesis was used to change Gln87 into Met, Tyr or His as described in Materials and Methods, section 4.4.1. Following mutagenesis, the recombinant pUC12-l.lcejc(PTIS) was transformed into E. coli JM101. Mutation anywhere within the codon for Gln87 resulted in destruction of a Pvu II restriction site, and restriction analysis of plasmid DNA isolated by small-scale alkaline lysis revealed that approximately 70% of the colonies had lost the Pvu II restriction site. The 500 base pair BamHl-Notl fragment isolated from one of the these clones was subcloned into pUC12l.lcex(PTIS), replacing the equivalent wild-type fragment of DNA as shown in Figure 3.2. DNA sequencing of the 500 bp mutant cassette indicated that it contained only the mutation of interest. pUC12-l.lcex(PTIS) was then transformed into various E. coli strains and tested for expression of the mutant protein. However, protein expression proved to be a very difficult task, especially for the Gln87Tyr and Gln87His mutants. To date, only the Gln87Met mutant has been successfully expressed (into E. coli Topp5 strain) and the following sections will focus on its characterization.  64  In vitro PCR site-directed mutagenesis  kb)  Digest each with BamHI/NotI  Figure 3.2  Generation of Gln87Met mutant. In vitro PCR site-directed mutagenesis was performed as described in section 4.4.1. A 500 bp BamH1-Not1 fragment was isolated from the PCR product and ligated to a 4.0 kb fragment from pUC12-1. ICex(PTIS). The mutant cassette replacing the wild-type DNA was sequenced to confirm that only the desired mutation was present. The position of mutation in the Cex gene is indicated by an asterisk.  65 3.3  Production, purification Gln87Met mutant  and physical characterization  of the  The mutant protein was produced and purified as described in section 4.6. No significant changes in behavior during purification were observed, the mutant eluting at approximately the same position as wild-type enzyme from the cellulose-affinity column. The purified protein ran as a single band on SDS-PAGE (> 95% purity by inspection), at the same position (47 kDa) as native enzyme. Final protein yield ranged from 15 to 25 mg per litre of liquid culture. Mass spectrometric analysis confirmed the molecular weight of mutant protein as 47120 Da. Also, CD spectra (Figure 3.2) and thermal stability (data not shown) were found to be essentially identical to those of the wild-type enzyme, indicating that no major structural changes in protein folding were caused by the mutation.  4000 i  200  1  1  220  1  1  240  1  j  1  260  1  280  1  r  300  Wavelength (nm)  Figure 3.2  CD spectra of Cex and Gln87Met mutant  66  3.4  Catalytic  properties  of the Gln87Met  mutant  Previous kinetic studies with the wild-type xylanase/cellulase (Cex) have indicated that the specificity constant, kcat/Km, is 60 to 90-fold lower for aryl cellobiosides relative to aryl xylobiosides (Table 3.2). The decreased binding affinity (larger K  m  values) of the  cellobiosides is presumably due to the presence of the C-5 hydroxylmethylene groups. In order to determine the importance of interactions between the side chain of Gln87 and the C-5 hydroxymethylene group of the substrate, kinetic parameters for the Gln87Met-catalyzed hydrolysis of gluco-sugars and xylo-sugars were determined and compared to those for the wild-type Cex. The enzymic hydrolysis rates were measured by spectrophotometric detection of phenol release, under the standard assay conditions described in Materials and Methods, section 4.9.1, or by measuring the amount of reducing sugars liberated during hydrolysis of sugar polymers, as described in section 4.9.2. The enzyme concentrations employed were those which give a sufficiently large absorbance change to ensure accurate determination of the rates yet resulted in less than 10% conversion of the substrate to product during the observation period, thus ensuring linear kinetics. The maximum velocities (V ax) and the apparent dissociation constants (Km) were m  determined for the two series of substrates by fitting the initial rates and substrate concentrations to the Michaelis-Menten equation using the program GraFit (Leatherbarrow, 1990). These results are illustrated in Appendix C as Lineweaver-Burk plots for visual convenience although the Michaelis-Menten parameters were not calculated from these plots due to the nonlinear error span associated with such double reciprocal analysis. The kinetic constants are given in Table 3.3. The average relative standard error in the data for individual experiments was 10-15%. For replicate experiments the average relative standard deviation was 5-10%.  67  Table 3.3  Kinetic parameters for the hydrolysis of B-glycosides by Cex and the Gln87Met mutant at 37°C, pH 7.0.  Substrate  Enzyme  PNPG  K  (mM)  kcat/Km ( S ' V M - I )  Cex wt Gln87Met  kcat (s" ) 0.024 0.024  8.3 7.4  2.9 x 10" 3.2 x IO"  3,4-DNPG  Cex wt Gln87Met  2.9 1.2  6.5 6.0  0.45 0.20  PNPC  Cex wt Gln87Met  15.8 9.2  0.60 0.34  26.3 27.1  3,4-DNPC  Cex wt Gln87Met  9.7 8.3  0.14 0.065  69 128  PhC  Cex wt Gln87Met  -  -  1.0 x IO" 2.3 x IO"  PNPX  Cex wt Gln87Met  2.6 1.2  20 67  0.13 0.018  3,4-DNPX  Cex wt Gln87Met  22 9.0  7.9 7.2  2.8 1.2  PNPX2  Cex wt Gln87Met  39.8 57.9  0.018 0.12  2200 482  3,4-DNPX2  Cex wt Gln87Met  22 38.2  0.012 0.038  1840 1005  PhX  Cex wt Gln87Met  10.4 18.7  8.6 21.2  1.2 0.9  Cex wt Gln87Met  0.052* 0.025*  -  -  Cex wt Gln87Met  0.47* 0.40*  -  -  2  2% CMC 1% Xylan  1  m  3  3  4  4  *specific activity expressed as s" or nmoles/mL of reducing sugars liberated per sec per nmoles/mL of protein; '-' parameter not determined 1  As can be seen from these data, substitution of Gln87 by Met in the binding site of Cex does not significantly alter the kinetic properties of the enzyme. The K  m  and k_ t a  values for the two series of substrates remained essentially unchanged. Most likely, the small variations observed are due to perturbations in the local environment of the protein.  68  Thus, in contrast to what had been hoped, the Gln87Met mutant does not show a decreased affinity for gluco-oligosaccharide substrates. Presumably, the methionine residue is sufficiently flexible to undergo a similar rearrangement to that observed for Gln87 upon binding the gluco-substrates in the active site. A bulkier substitution with a more rigid side chain such as tyrosine or histidine might ehminate this possibility. However, attempts to create such mutant proteins have not succeeded. Interestingly, in a study by Dr. Alasdair MacLeod (postdoctoral fellow in the Cellulase lab, UBC, personal communication, 1995), a Cex mutant in which Gln87 was replaced with serine also retained the kinetic characteristics of the wild-type enzyme. Serine, having a smaller side chain than glutamine (Figure 3.3), would be expected to enlarge the binding  H O  Glutamine  Figure 3.3  Serine  Structure of glutamine and serine side chains.  pocket for the C-5 hydroxymethylene group of the distal glucose unit and thus potentially improve the binding of cello-substrates. Such an effect was not observed. Together with the above findings, this suggests that the binding interactions  with the C-5  hydroxymethylene groups in the active site are more complex than initially thought, and that a single point mutation in the catalytic domain of Cex may not be sufficient for introducing the desired changes in the specificity of the enzyme. It is also speculated that the cellulose binding domain (CBD) of the protein may play a role in recognition and  69  binding of the native polymeric substrates; however, an effect of the CBD on the binding of aryl oligosaccharides in the active site of the catalytic domain is very unlikely.  3.5  Concluding  remarks  The identification of enzymic residues involved in substrate recognition and binding is vital for engineering enzymes with altered specificities. However, changing the substrate specificity is by no means simple or routine at this point. It is a challenging biochemical task which requires a multi-disciplinary approach involving techniques such as X-ray crystallography, site-directed mutagenesis and kinetic analysis. It is hoped that this type of specific information will soon become available for the potentially valuable xylanase and cellulase enzymes. The present investigation aimed to contribute to a better understanding of the substrate recognition and specificity of C. fimi xylanase/cellulase (Cex), by probing the role of the conserved Gln87 residue in the active site binding cleft. The kinetic characterization of a Gln87Met mutant revealed no significant changes in substrate affinity relative to the wild-type enzyme, and thus the degree to which Gln87 contributes to the specificity of Cex is not clear. However, this study is only at the forefront of a protein engineering project to understand and modify the substrate preference of such cellulase/xylanase enzymes. Future studies will certainly bring more insight into this issue.  CHAPTER 4 MATERIALS AND METHODS  71  4.1  Chemicals,  media  components  and  enzymes  All chemicals used were of analytical or HPLC grade and were obtained from Sigma (St. Louis, MO) or BDH (Poole, England). All media components were from Difco (Detroit, MI). Restriction endonucleases, polymerases, ligase and nucleotides were obtained from Pharmacia (Baie d'Urfe, Quebec) or New England Biolabs (Beverly, MA). Sequenase Version 2.0 was from United States Biochemical (Oakville, ON). Avicel PH101, a microcrystalline cellulose, was from FMC International (Cork, Ireland) and CF1, a fibrous cellulose, was from Sigma. Buffers and solutions used in this study were prepared as described by Sambrook et al., (1989). CM-cellulose and birchwood xylan were from Sigma. All aryl glycoside substrates were synthesized by other members in this laboratory.  4.2  Bacterial  strains  and plasm ids  Escherichia coli strain JM101 (Yanisch-Perron et al., 1985) was used as the host strain for genetic manipulations, and Topp 5 (Stratagene) for protein production. DNA vector used was pUC12-l.lcex (PTIS) (O'Neill et al., 1986). Bacterial stocks were maintained at -70°C in LB medium containing 10% DMSO, while plasmid DNA was stored in TE buffer or water at -20°C.  4.3  Media  and growth  conditions  Luria-Bertani (LB) medium has been described previously (Sambrook et al., 1989). TYP medium contained the following (per litre): 16 g tryptone, 16 g yeast extract, 5 g  72  NaCl, 2.5 g K2HPO4. Small-scale cultures of E. coli containing pUC 12-1.Ice* (PTIS) vector were grown in shake flasks at 225 rpm at 30°C in LB or TYP supplemented with ampicillin (Amp) at 100 pg/ml. Growth of E. coli was monitored spectrophotometrically by measuring A600nm-  4.4  Recombinant  DNA  techniques  Most recombinant DNA techniques were carried out as described by Sambrook et al., (1989). Double-stranded plasmid DNA was isolated from E. coli by the small-scale alkaline lysis method (Sambrook et al., 1989). Sequencing grade double-stranded DNA was isolated using a plasmid purification kit (Plasmid midi kit, Qiagen, Chatsworth, CA). Restriction endonuclease digestions and ligations were carried out according to the directions of the manufacturers in the buffers provided. DNA fragments were isolated from agarose gels and purified using a gel extraction kit (Qiaex gel extraction kit, Qiagen). Electrocompetent E. coli  strains were transformed using a Gene pulser II apparatus  equipped with the Pulse Controller II unit (BioRad). Electroporation was carried out as described by the manufacturers (BioRad).  4.4.1 Site-directed mutation using PCR  Site-directed mutagenesis was performed by PCR using the overlap-extension procedure (Higuchi, 1990). The expression plasmid pUC12-l.lce;c(PTIS) containing the Cex gene was used as a template. Synthetic oligonucleotides used for mutagenesis were prepared by the UBC Nucleic acid and Protein synthesis unit (NAPS) with an Applied Biosystems 380A DNA synthesizer. The mutagenic primers are described in Table 4.1.  73  Table 4.1  Mutagenic primers used in site-directed mutation of Cex by PCR.  Name Oligonucleotide sequence (5'-3')  Mutation in Cex  QMf  GTA TGG CAC TCG ATG CTG CCC GAC TGG  Gln87Met  QMr  CCA GTC GGG CAG CAT CGA GTC CCA TAC  Gln87Met  QYf  GTA TGG CAC TCG TAC CTG CCC GAC TGG  Gln87Tyr  QYr  CCA GTC GGG CAG GTA CGA GTC CCA TAC  Gln87Tyr  QHf  GTA TGG CAC TCG CAC CTG CCC GAC TGG  Gln87His  QHr  CCA GTC GGG CAG GTG CGA GTC CCA TAC  Gln87His  HAf  TTC CAG TCG GCC CTC ATC GTC GGC CAG GTG CCT  His205Ala  GGC GAC TTC CGG HAr  CCG GAA GTC GCC AGG CAC CTG GCC GAC GAT GAG  His205Ala  GGC CGA CTG GAA HNf  TTC CAG TCG AAC CTC ATC GTC GGC CAG GTG CCT  His205Asn  GGC GAC TTC CGG HNr  CCG GAA GTC GCC AGG CAC CTG GCC GAC GAT GAG  His205Asn  GT1 CGA CTG GAA NOTE. Nucleotide changes are underlined. Silent mutations adding a restriction site are in italics.  74 PCR Tube # 1  PCR Tube # 2  QXr  CexPl  RUP  QXf  20 Cycles 95°C, 30 seconds 48°C, 30 seconds 72°C, 54 seconds  20 Cycles 95°C, 30 seconds 48°C, 30 seconds t 72°C, 45 seconds  1.1 kb fragment  =s=F  a: s a  0.7 kb fragment  1) Purify, dilute and combine fragments 2) Denature (95°C) and anneal (48°C)  03  1) Extend fragments (72° C, 1 minute) 2) Add flanking primers CexPl RUP  20 Cycles 95°C, 30 seconds 48°C, 30 seconds 72°C, 1 minute PCR product (1.8 kb)  Figure 4.1  PCR site-directed mutagenesis scheme for mutating Gln87 in Cex gene. "X" denotes the amino acid change (i.e., Met, Tyr or His).  75  The PCR protocol used for mutating Gln87 to either Met, Tyr or His is outlined in Figure 4.1. The reactions were designed so that the final PCR product contains the desired mutation and two unique restriction sites flanking it, to facilitate subcloning into the original vector. "X" denotes the amino acid change (i.e. M for Met, Y for Tyr, and H for His).The procedure is as follow. A PCR reaction mix was prepared by adding the following components to a 1.5 ml eppendorf tube: 15 uX of lOx Vent polymerase PCR buffer, 79.5 uL H2O, 30 uL of dNTP mix (2 mM each of dATP, dCTP, dGTP, dTTP) and 1.5 uL of Vent polymerase. In a separate 0.5 mL eppendorf tube, 1 uL each of PCR primers QXf and CexPl (20 pmol/uL) were combined with 1 uL of pUC12-l.lcejc (PTIS) DNA (10 ng/pL) and 5 pL of DMSO. A second 0.5 mL eppendorf tube contained 1 uL each of primers QXr and RUP (20pmol/pl), 1 uL of pUC12-l.lcex (PTIS) DNA (lOng/pX) and 5 p,L of DMSO. The 0.5 mL eppendorf tubes were heated to 96°C for 1 minute (hot start) in the PCR thermocycler (TwinBlock, Ericomp, San Diego, CA). At this time, 42 pL of the PCR reaction mix was added to each and 2 drops of mineral oil were placed on top of the mixtures. The mixtures were heated to 95°C for 30 seconds, 48°C for 30 seconds, and 72°C for 54 or 45 seconds. This cycle was repeated a total of 20 times. Afterwards, the mixtures were heated to 72°C for 1 minute, then cooled to room temperature. PCR products were analyzed by electrophoresis through a 1% agarose gel. The two resulting PCR products were diluted to about 50 ng/pL, combined, and 1 pL sample was then added to a 0.5 mL eppendorf tube along with 5 uL of DMSO. The tube was heated at 96°C for 1 minute in the PCR block as before. After the hot start, 42 uL of the reaction mix was added, then heated at 48°C for 30 seconds and 72°C for 1 minute. At this time, 1 pL of each flanking primer CexPl and RUP were added and 2 drops of mineral oil were placed on top of the PCR mixture. The mixture was heated to 95°C for 30 seconds, 48°C for 30 seconds and 72°C for 1 minute. This cycle was repeated a total of 20 times. Afterwards, PCR products were analyzed on a 1% agarose gel.  76  A similar PCR protocol was used for changing His205 to Ala or Asn (Mark Nitz, personal communication), with the following changes: the mutagenic primer QXf is replaced by HXf, and QXr is substituted with HXr, where X = A or N, respectively. Also, the flanking primer CexZAl is used instead of CexPl.  4.4.2 Screening for mutants  Following in vitro mutation, the PCR products of the correct size were purified by phenol/chloroform extraction and cut with BamHI and Notl/Pstl. The resulting DNA fragments were purified, cloned into pUC12-l.lCex(PTIS) and transformed into E. coli JM101. As a result of mutation, a PvuII site has been removed in the Gln87Met mutant DNA, while a silent mutation coding for a Bstxl restriction site has been introduced along with the mutations in the DNA of the His205 mutants. Thus, transformants were screened by restriction site analysis of purified double-stranded DNA. All mutations were confirmed by DNA sequencing, using the modified dideoxy chain termination method described previously (Tabor and Richardson, 1987) with the following modifications: in the primer extension reaction, T7 DNA polymerase (Sequenase) was used, the reaction temperature was increased to 43 °C and 7-deazaGTP was substituted for dGTP.  4.5  Estimation  of protein  yields  E. coli strains carrying the recombinant pUC12-l.lcex(PTIS) were grown for 1824 hours under the conditions described in section 2.3. The levels of gene expression were approximated from crude-cell extracts or culture supernatants as follow. Approximately 100 pL of cell extract (described in section 2.6) or 0.5 mL of culture supernatant was  77  added to 20 mg of Avicel (microcrystalline cellulose) in a final volume of 1 mL of 50 mM phosphate buffer, pH 7.0. The protein was adsorbed to the Avicel by mixing for 1 hr at 4°C. The Avicel was centrifuged for 30 seconds, washed with 1 M NaCl, then with 50 mM phosphate buffer, pH 7.0. The Avicel was collected by centrifugation, then boiled for 2 minutes in SDS loading buffer (Laemmli, 1970). The bound polypeptides were analyzed by SDS-PAGE (Laemmli, 1970) with a Mini-PROTEAN apparatus (BioRad). Protein bands were visualized by staining with Coomasie blue. An estimation of mutant protein yield could be made by comparison with a known quantity of wild-type Cex. From the variety of E. coli strains examined, Topp5 was selected for expression of the proteins.  4.6  Production  of Cex and Cex mutants  in E. coli  E. coli Topp5 containing the plasmid pUC12-l.lcex(PTIS) encoding wild-type Cex or mutants of Cex were grown in 2 L-cultures in shake flasks or in a 60 L Chemap FZ30000 fermenter. Cultures were grown overnight in TYP supplemented with ampicillin at 100 pg/mL. At an A600 of 6.0-7.0, IPTG was added to 0.1 mM and growth was allowed to continue for 3 hours. The cells were collected by centrifugation at 10000 x g for 10 minutes at 4°C and the cell pellet resuspended in approximately 1/100 of the original volume in 50 mM phosphate buffer, pH 7.0. Cells were ruptured by passage twice through a French pressure cell. PMSF was added to 0.5 mM and EDTA to 1 mM. Following clarification of the cell extract by centrifugation at 40000 x g for 30 minutes, streptomycin sulfate was added to 1.5% and the mixture stirred at 4°C overnight. The extract was centrifuged again at 40000 x g for 30 minutes. Cex was purified from the clarified cell extract by affinity chromatography on cellulose. To allow for binding, the cell extract was stirred with CF-1 cellulose (about 1 g  78  CF-l/mg Cex) in phosphate buffer for 3 hours at 4°C. The cellulose was packed into an XK-50 column (Pharmacia), attached to an FPLC system (Pharmacia), and washed with 2.5 column volumes of 1 M NaCl at a flow rate of 1.4 mL/min. Washing was continued by passing a column volume of phosphate buffer through the column at a flow rate of 1 mL/min. Adsorbed polypeptides were eluted with distilled water at a flow rate of 1 mL/min. The absorbance of the eluate was measured continuously at 280 nm and appropriate fractions were pooled and centrifuged at 40 000 x g for 30 minutes at 4°C to remove any cellofines. The supernatant was concentrated to greater than 1 mg protein/mL by diafiltration through an Amicon PM10 membrane and then analyzed by SDS-PAGE (Laemmli, 1970). To further purify, the protein sample was passed through a gel filtration column (Superose 12) in phosphate buffer. Purified protein solutions were stored at 4°C after being concentrated and passed through a 0.22 pm filter (Millipore). Final protein concentration was calculated from measured absorbance at 280 nm and the extinction coefficient of Cex (e  4.7  Protein  = 1.61 mL mg~l cm ). -1  280  mass  determination  The molecular weights of purified Cex and Cex mutants were determined by Ion Spray Mass Spectrometry performed on a PE SCIEX API 300 ion spray LC/MS system by David Chow and Shouming He. Enzyme samples (approximately 10 pg) were injected into the mass spectrometer to generate the mass spectra. The masses of generated mutants were confirmed by comparison between the expected molecular weights and those provided by the spectra.  79  4.8  Confirmation  of overall  protein  structure  by circular  dichroism  spectroscopy  CD spectra were recorded with a spectropolarimeter (model J-720, JASCO, Tokyo, Japan) controlled with J-700 software. The spectra were obtained at a protein concentration of 0.3 mg/mL in 5 mM phosphate buffer, pH 7.0, 25°C using a 100 pL silicon quartz cell with a 1 mm path length. They were recorded 4 times from 190 to 300 nm at a scan rate of 50 nm/min, using a 2.0 s response and a sensitivity of 10 mdeg. The thermal stabilities of the proteins were assessed by monitoring changes in circular dichroism at 220 nm.  4.9  Enzymology  4.9.1 Determination of steady-state kinetic parameters  Michaelis-Menten parameters for the enzymatic hydrolysis of all aryl glycoside substrates, with the exception of PhC and PhX2, were determined using a continuous assay, by recording changes in absorbance using a UV/Vis Pye-Unicam 8700 spectrophotometer equipped with a temperature-controlled circulating water bath. Solutions of the appropriate substrate concentrations in 50 mM citrate or phosphate buffer, pH 5.5 or 7.0 respectively, 1 mg/mL BSA at 25°C or 37°C were preincubated within the spectrophotometer until thermally equilibrated, and reactions were initiated by the addition of enzyme. Hydrolysis was continuously monitored at a wavelength at which there was a convenient absorbance difference between the initial glycoside and the phenol product as previously reported (Kempton and Withers, 1992). In order to ensure linear kinetics and to obtain a sufficient absorbance change for accurate initial rate calculations, the concentration  80  of the enzyme added and the reaction time were selected such that less than 10% of the total substrate was converted to product. Hydrolysis rates for the substrates PhC and PhX_ were determined at 37°C using a stopped assay. Different concentrations of substrate in 50 mM phosphate buffer, 1 mg/mL B S A (pH 7.0) (190 pL) were prewarmed at 37°C and the reaction initiated by the addition of a 10 p L enzyme. After an appropriate time, 0.6 mL of 2.0 M Na3P04/H20 (pH 12.15) was added to stop the reaction. The absorbance of the released phenolate at 288 nm was determined immediately and corrected for the spontaneous hydrolysis of substrate and the background absorbance of the enzyme (Ae = 2.17 mM~l cm~l for phenol at pH 12.15). For each substrate, approximate values of K  m  and V a x were determined by m  measuring initial rates of hydrolysis at three widely ranging substrate concentrations. Accurate values were then measured using various substrate concentrations, which generally ranged from 0.2 to 5 times the K  m  value. The program Grafit (Leatherbarrow,  1990) was used to perform nonlinear regression analysis in the determination of K  m  and  Vmax-  4.9.2 Determination of activity on CM-cellulose and xylan  CM-cellulase and xylanase activity were measured by the p-hydroxybenzoic acid hydrazide (HBAH) method (Miller et al, 1960) which quantitates the production of reducing sugar. Initial rates of CM-cellulose hydrolysis were determined by incubating 100 p L of an enzyme solution with 900 p L of 2% CM-cellulose in 50 m M citrate buffer, pH 6.8 at 30°C. Aliquots of 100 p L were removed at 0, 2, 4, 6, 8 and 10 minute intervals and added to 1 mL of 5 m M NaOH in glass test tubes. After all points were taken 1 mL of H B A H reagent was added to each tube and the samples were heated in a steam bath for 12 minutes.  81  When tubes had cooled to room temperature, A420 m  w  n  a  s  measured and the amount of  reducing sugar released was determined using a glucose standard curve generated under the same conditions. The activity is expressed as min" or nmoles/mL reducing sugars released 1  per minute per nmoles/mL of enzyme. To ensure the validity of the assays for enzyme activity, concentrations of enzyme were chosen such that the production of reducing sugar in the assay fell within the linear response range of the standard curve. Appropriate controls for background reducing sugar and/or for the presence of chemical reagents that may interfere with the assay were always included with the assay procedures. Substrate for the xylanase assays was prepared by dissolving 6-10 g of birchwood xylan in 100-200 mL distilled water at room temperature for 3-4 hours with continuous stirring. Insoluble material was removed by centrifugation at 16000 x g in a GSA rotor for 20 minutes. The clear supernatant was lyophilized, and the dry, water-soluble product was used at a concentration of 0.2% for determination of reducing sugars as described above.  4.9.3 Secondary deuterium kinetic isotope effect measurements  Isotope effects were determined by comparison of the initial rates of hydrolysis of high (3-10 times the  value) concentrations of protio and deuterio substrate (2,4-DNPC)  determined spectrophotometrically. Quartz cells were filled with the appropriate concentration of substrate in 50 mM phosphate buffer, 1 mg/ml BSA (pH 7.0) and incubated at 25°C, reaction being initiated by the addition of a small volume (50 pL) of thermally equilibrated enzyme. Rates of protio and deuterio substrate hydrolysis were determined in alternation until a total of 7 or 8 rates for each (protio and deuterio) substrate had been measured. Average rates for the protio and deuterio substrates were then calculated and the rate taken to give the isotope effect. Errors are the standard deviation of the average kinetic isotope effect.  82  4.9.4 p H dependence studies  The dependence of kc t/K on pH for Cex wild-type and H205 mutants was a  m  determined as follows. A final PNPC concentration of 0.2 x K  m  in the appropriate buffer  containing 1 mg/mL BSA and 145 mM NaCl was incubated at 25°C until thermally equilibrated. Following the addition of enzyme, the release of p-nitrophenolate was monitored by following the absorbance at 400 nm until substrate depletion was observed. The change in absorbance with respect to time was fitted to a first order rate equation using the program Grafit (Leatherbarrow, 1990) which yielded pseudo-first order rate constants at each pH value. At low substrate concentrations, the reaction rates are given by the equation v =- ^  L  - [E ][S] 0  Thus, pseudo first order rate constants measured correspond to Eo(kcat/K ), from which m  kcat/Km is easily obtained. The pH of each reaction mixture was measured after completion of reaction to ensure that the pH had not fluctuated during the reaction. Also, assays were performed at extreme pH values to check for any loss in enzymatic activity due to enzyme degradation/instability.  4.9.5 B u r s t e x p e r i m e n t  The burst experiment for the His205 mutants with the 2,4-DNPC substrate was performed  by  following  the  release  of  2,4-dinitrophenol  using  a  UV/Vis  spectrophotometer. To 2,4-DNPC (190 pi, 40 uM) in 50 mM phosphate buffer, 0.1%  83  BSA, pH 7.0 at 25°C was added 10 pi enzyme (2.6 pM His205Ala or 2.3 pM His205Asn). Extrapolation of the linear portion of the burst phase and steady state phase back to time zero determined the size of the burst. The extinction coefficient used for 2,4DNP was determined as outlined in section 4.9.7.  4.9.6 Inactivation experiment  The pseudo-first order rate constant (k bs) f ° inactivation was calculated for Cex r  0  wild-type and the Cex H205N mutant using the inactivator 2F-DNPC at a concentration of 1.5 mM. In each case, the enzyme (10 pi, 2 mg/mL) was added to the inactivation mixture containing 2F-DNPC (90 pi, 1.5 mM) in phosphate buffer, 0.1% BSA, pH 7.0 at 25°C. Aliquots (10 pi) were removed at different time intervals and diluted into reaction cells containing a large volume of substrate (2,4-DNPC) at saturating concentration (1.2 mM). This stopped the inactivation both by dilution of the inactivator and by substrate competition for the enzyme, as the substrate is present in large excess. The residual enzymatic activity was then determined from the rate of hydrolysis of substrate, which is directly proportional to the amount of active enzyme. The inactivation was monitored until 80-90% of the enzymatic activity was lost. For each enzyme, the pseudo-first order rate constants (kobs) f ° inactivation were calculated by direct fit of the residual activity versus r  time curve to a first-order function using the program GraFit (Leatherbarrow, 1990).  4.9.7 Determination of Extinction Coefficients  Extinction coefficients (e) were measured at 25°C at pH 5.5 and pH 7.0 in 50 mM citrate buffer and 50 mM phosphate buffer, respectively, using matched quartz cells. Phenols and cellobiosides were dried in vacuo, weighed, and dissolved in a known volume  84  of buffer. Absorbance values were taken at a wavelength of 400 nm for six different concentrations of substrate. Extinction coefficients were determined from Beer's Law:  where A is the absorbance at 400 nm, c is the concentration of solution (M) and 1 is the cell pathlength (1 cm). To convert the observed rate of change of absorbance (AA/min) to rate of phenol release (AmM/min), the extinction coefficient difference (Ae) between phenol and cellobioside is used:  v  __ AA/min Ae  It should be noted that the same Ecellobioside values were used for both pH 5.5 and pH 7.0 since the substrate does not ionize and change in extinction coefficient.  Table 4.2  Extinction coefficients of substituted phenols and phenyl cellobiosides at 25°C. Ephenol (M" cm )  Phenol substituent  Wavelength monitored (nm)  pH 5.5  pH 7.0  Ecellobioside (M^cm" )  pNP  400  0.65  8.21  0  2,4-DNP  400  12.30  12.46  0  3,4-DNP  400  8.18  12.87  0.041  1  85  REFERENCES  86 Anderson, W. F., Grutter, M. G., Remington, S. J., Weaver, L. FL, & Matthews, B. W. (1981) J. Mol. Biol. 147, 523. Antoine, R., & Locht, C. (1994) J. Biol. Chem. 269, 6450. Antoine, R., Tallett, A., Heyningen, S., & Locht. C. (1993) J. Biol. Chem. 268, 24149. Arabshahi, A., Ruzicka, F. J., Geeganage, S., & Frey, P. A. (1996) Biochemistry 35, 3424. Beguin, P. & Aubert, J.-P. (1994) FEMS Microbiol. Rev. 13, 25. Belaich, A., & Belaich, J. P. (1992) J. Bacteriol. 174, 4677. Bennet, A., & Sinnott, M. L. (1986) J. Am. Chem. Soc. 108, 7287. Berger, E., Jones, W. A., & Woods, D. T. (1989) Mol. Gen. Genet. 219, 193. Blake, C. C. F., Mair, G. A., North, A. C. T., Philips, D. C., & Sharma, V. R. (1967) Proc. R. Society London Ser. B 167, 365. Bortoli-Germani, I., Haiech, J., Chippaux, M., & Barras, F. (1995) J. Mol. Biol. 246, 82. Bray, M. R., & Clarke, A. J. (1994) Eur. J. Biochem 219, 821. Campbell, R., Rose, D., Wakarchuk, W., To, R., Sung, W., & Yaguchi, M.(1993) A comparison of the structures of the 20 kD xylanases from Trichoderma harzianum and Bacillus circulans, in Proceedings of the 2nd TRICEL symposium on Trichoderma reesei cellulases and other hydrolases, (Suominen, P., & Reinikainen, T., Ed.) p 63. Foundation for Biotechnical and Industrial Fermentation Research, Helsinki, Finland. Coughlan, MP, & Hazlewood, G. (1993) Biotechnol. Appl. Biochem. 17, 259. Creighton, T. E. (1993) in Proteins, 2nd ed. W. H. Freeman & Company, NY. Davies, G. J., Dodson, G. G., Hubbard, R. E., Tolley, S. P., Dauter, Z., Wilson, K. S., Hjort, C , Mikkelsen, J. M., Rasmussen, G., & Schulein, M. (1993) Nature 365, 362. Derewenda, U., Swenson, L., Green, R., Wei, Y., Morosoli, R., Shareck, F., Kluepfel, D., & Derewenda, Z. S. (1994) / . Biol. Chem. 269, 20811. Divne, C , Stahlberg, J., Reinikainen, T., Ruohonen, L., Petterson, G., Knowles, J. K. C , Teerei, T., & Jones, T. A. (1994) Science 265, 524. Dominguez, R., Souchon, H., Spinelli, S., Zbigniew, D., Wilson, K. S., Chauvaux, S., Beguin, P., & Alzari, P. M. (1995) Nature Stuct. Biol. 2, 569. Ducros, V., Czjzek, M., Belaich, A., Gaudin, C , Fierobe, H.-P., Belaich, J.-P., Davies, G. J., & Haser, R. (1995) Structure 3, 939.  87 Gebler, J. C , Aebersold, R., & Withers, S. G. (1992) J. Biol. Chem. 267, 11126. Gebler, J. C., Trimbur, D. E., Warren, R. A. J., Aebersold, R., Namchuk, M., & Withers, S. G. (1995) Biochemistry 34, 14547. Gilkes, N. R., Langford, M., Kilburn, D. G., Miller, R. C. Jr., & Warren, R. A. J. (1984) J. Biol. Chem. 259, 10455. Gilkes, N. R., Warren, R. A. J., Miller, R. C. Jr., & Kilburn, D. G. (1988) J. Biol. Chem. 263, 10401. Gilkes, N. R., Kilburn, d. G., Miller, R. C. Jr., & Warren, R. A. J. (1989) J. Biol. Chem. 264, 17802. Grepinet, O., Chebrou, M.-C., & Beguin, P. (1988) /. Bacteriol. 170, 4582. Haas, H., Herfurth, E., Stoffler, G., & Redl, B. (1992) Biochim. Biophys. Acta 1117, 279. Hall, J., Hazlewood, G. P., Barker, P. J., & Gilbert, H. J. (1988) Gene 69, 29. Harris, G. W., Jenkins, J. A., Connerton, I., Cummings, N., Leggio, L. L., Scott, M., Hazlewood, G. P., Laurie, J. I., Gilbert, H. I., & Pickersgill, R. W. (1994) Structure 2, 1107. Hehre, E. J., Genghof, D. S., Sternlicht, H., & Brewer, C. F. (1977) Biochemistry 16, 1780. Henrissat, B., & Bairoch, A. (1993) Biochem. J. 293, 781. Henrissat, B., Claeyssens, M., Tomme, P., Lemesle, L., & Mornon, J.-P. (1989) Gene 81,83. Higuchi, R. (1990) In: Innis MA, Gelford, D.H,. Sninsky, J. J., White, T. J., eds. PCR protocols: A guide to methods and applications. San Diego, California: Academic Press, p 117. Inoue, M., Yamada, H., Yasukochi, T., Kuroki, R., Miki, T., Horiuchi, T., & Imoto, T. (1992) Biochemistry 31, 5545. Ishikawa, K., Matsui, I., Honda, K., & Nakatani, H. (1992) Biochem. Biophys. Res. Commun. 183, 286. Ishikawa, K., Matsui, I., Kobayashi, S., Nakatani, H., & Honda, K. (1993) Biochemistry 32, 6259. Jacobsen, R. H., Zhang, X-J., DuBose, R. F., & Matthews, B. W. (1994) Nature 369, 761. Jones, C. C , Sinnott, M. L., & Souchard, I. J. L. (1977) J. Chem. Soc. Perkin Trans. II, 1191.  88 Juy, M., Amit, A. G., Alazri, P. M., Poljak, R. J., Claeyssen, M., Beguin, P., & Aubert, J. P. (1992) Nature 357, 89. Kanda, T., Brewer, C. F., Okada, G., & Hehre, E. J. (1986) Biochemistry 25, 1159. Kempton , J. B., & Withers, S. G. (1992) Biochemistry 31, 9961. Kim, J., Ruzicka, F., & Frey, P. A. (1990) Biochemistry 29, 10590. Koshland, D. E. (1953) Biol. Rev. 28, 416. Laemmli, U. K. (1970) Nature 227, 680. Lai, E. C. K., & Withers, S. G. (1994) Biochemistry 33, 14743. Langsford, M. L., Gilkes, N. R., Singh, B., Moser, B., Miller, R. C. Jr., Warren, R. A. J., & Kilburn, D. G. (1987) FEBS Lett. 225, 163. Leatherbarrow, R. J. (1990) GraFit Version 2.0, Erithacus Software Ltd., Staines, U. K. Legler, G. (1990) Adv. Carb. Chem. Biochem. 48, 319. Lehmann, J., & Schlesselmann, P. (1983) Carbohydrate Res. 113, 93. Lin, L.-L., & Thompson, J. A. (1991) Mol. Gen. Genet. 228, 55. Locht, C , & Antoine, R. (1995) Biochimie 77, 333. Luthi, E., Love D. R., McAnulty, J., Wallace, C., Caughey, P.A., Saul, D., & Berguist, P. L. (1990) Appl. Environ. Microbiol. 57, 694. MacLeod, A. M. (1994) Ph.D. Thesis, University of British Columbia, Vancouver, Canada MacLeod, A. M., Lindhorst, T., Withers, S. G., & Warren, R. A. J. (1994) Biochemistry 33, 6571. Matsuura, Y., Kusunoki, M., Harada, W., & Kakudo, M. (1984) J. Biochem. 95, 695. McCarter, J. D., Adam, M., & Withers, S. G. (1992) Biochem. J. 286, 721. Mcintosh, L. P., Hand, G., Johnson, P. E., Joshi, M. D., Korner, M., Plesniak, L. A., Ziser, L., Wakarchuk, W. W., & Withers, S. G. (1996) Biochemistry Miao, S., McCarter, J. D., Grace, M., Grabowski, G., Aebersold, R., & Withers, S. G. (1994) J. Biol. Chem. 269, 10975. Miao, S., Ziser, L., Aebersold, R., & Withers, S. G. (1994) Biochemistry 33, 7027. Miller, G. L., Blum, R., Glennon, W. E., & Burton, A. L. (1960) Anal. Biochem. 1, 127. Moreau, A., Sharek, F., Kluepfel, D., & Morosoli, R. (1994) Eur. J. Biochem. 219, 261.  89 Nakamura, A., Haga, K., & Yamane, K. (1993) Biochemistry 32, 6624. Namchuk, M. N., & Withers, S. G. (1995) Biochemistry O'Neill, G., Goh, S. H., Warren, R. A. J., Kilburn, D. G., & Miller, R. C. Jr. (1986a) Gene 44, 325. O'Neill, G. P., Kilburn, D. G., Warren, R. A. J., & Miller, R. C. Jr. (1986b) Appl. Environ. Microbiol. 52, 737. Parsons, S. M., & Raftery, M. A. (1972) Biochemistry 11, 1623. Philips, D. C. (1967) Proc. Natl. Acad. Sci USA 57, 484. Qian, M., Haser, R., Buisson, G., Duee, E., & Payan, F. (1994) Biochemistry 33, 6284. Roeser, K.-R., & Legler, G. (1981) Biochim. Biophys. Acta 657, 321. Rouvinen, J., Bergfors, T., Teerei, T., Knowles, J. K. C., & Jones, T. A. (1990) Science 249, 380. Sambrook, J., Fritsch, E. F., & Maniatis, T. (1989). in Molecular Cloning: A laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Shareck, F., Roy, C , Yaguchi, M., Morosoli, R., & Kluepfel, D. (1991) Gene 107, 75. Sinnott, M. L. (1978) FEBS Lett. 94, 1. Sinnott, M. L. (1990) Chem. Rev. 90, 1171. Sinnott, M. L., & Souchard, I. J., L. (1973) Biochem. J. 133, 89. Sinnott, M. L., & Withers, S. G. (1974) Biochem. J. 143, 751. Spezio, M., Wilson, D. B., & Karplus, P. A (1993) Biochemistry 32, 9906. Strynadka, N. C. J., & James, M. N. G. (1991) / . Mol. Biol. 220, 401. Svensson, B. (1994) Plant Mol. Biol. 25, 141. Tabor, S., & Richardson, C. C. (1987) Proc. Natl. Acad. Sci. USA 84, 4767. Tull, D., & Withers, S. G. (1994) Biochemistry 33, 6363. Tull, D., Withers, S. G., Gilkes, N. R., Kilburn, D. G., Warren, R. A. J., & Aebersold, R. (1991) J. Biol. Chem. 266, 15621. Tull, D. (1995) Ph.D Thesis, University of British Columbia, Vancouver, Canada. Umezerike, G. M. (1988) Biochem. J. 254, 73.  90 Varghese, J. N., Garrett, T. P. J., Colman, P. M., Chen, L., Hoj, P. B., & Fincher, G. B. (1994) Proc. Natl. Acad. Sci. USA 91, 2785. Wakarchuk, W. W., Campbell, R. L., Sung, W. L., Davoodi, J., & Yaguchi, M. (1994) Protein Sci. 3, 467. Wang, Q., Tull, D., Meinke, A., Gilkes, N. R., Warren, R. A. J., Aebersold, R., & Withers, S. G. (1993) / . Biol. Chem. 268, 14096. Wang, Q., Trimbur, D., Graham, R., Warren, R. A. J., & Withers, S. G. (1995) Biochemistry 34, 14554 Weber, J. P., & Fink, A. L. (1980) J. Biol. Chem. 255, 9030. Wentworth, D. F., & Wolfenden, R. (1974) Biochemistry 13, 4715. White, A., Withers, S. G., Gilkes, N. R., & Rose, D. R. (1994) Biochemistry 33, 122546. White, A., Tull, D., Johns, K., Withers, S. G., Rose, D. R. (1996) Nature Struct. Biol. 3, 149. Withers, S.G., & Aebersold, R. (1994) Protein Sci 4, 361. Withers, S. G., & Street, I. P. (1988) J. Am. Chem. Soc. 110, 8551. Withers, S. G., MacLennan, D. J., & Street, I. P. (1986) Carbohydr. Res. 154, 127. Withers, S. G., Street, I. P., Bird, P., & Dolphin, D. H. (1987) J. Am. Chem. Soc. 109, 7530. Withers, S. G., Dombroski, D., Berven, L. A., Kilburn, D. G., Miller, R. C. Jr., Warren, R. A. J., & Gilkes, N. R. (1986) Biochem. Biophys. Res. Commun. 139, 487. Withers, S. G. , Street, I. P., & Percival, M. D. (1988) in Fluorinated carbohydrates: Chemical and Biochemical Aspects (Taylor, N. F., Ed.) p 59, American Chemical Society, Washington, D. C. Withers, S. G., Warren, R. A. J., Street, I. P., Rupitz, K., Kempton, J. B., & Aebersold, R. (1990). J. Am. Chem. Soc. 112, 5887. Yague, E., Beguin, P., & Aubert, J.-P. (1990) Gene, in press. Yanish-Perron, C , Vieira, J., & Messing, J. (1985) Gene 33, 103.  APPENDIX A BASIC CONCEPTS OF ENZYME CATALYSIS  92  A.1.  Basic  enzyme  kinetics  Consider a typical enzyme-catalyzed reaction, in which the substrate forms a complex with the enzyme that subsequently decomposes to products and enzyme:  E +S  -  " ES  *- E + P  The general expression for the initial velocity (rate) of this reaction is given by the Michaelis-Menten equation: v  =  k [E ][S] K + [S] cat  0  m  where [Eo] is the total enzyme concentration; [S] is the substrate concentration; kc t is the a  catalytic constant; and K is the Michaelis constant. This expression is the basic equation m  of enzyme kinetics, and describes a rectangular hyperbola such as is plotted in Figure A . l .  Figure A. 1  Plot of velocity versus substrate concentration for a typical enzymatic reaction (Fersht, 1985).  93  In the Michaelis-Menten equation two assumptions are made; the enzyme concentration is negligible compared to that of the substrate and, the velocity measured is the initial rate of product formation, thus there is no significant accumulation of product (or depletion of substrates) hence, the reverse reaction can be ignored. Therefore, the change in substrate concentration is generally linear with time. The K is the substrate concentration at which the reaction velocity is half-maximal m  ( =V v  m  /2).  It may be treated as an apparent dissociation constant of all the bound enzyme  species and as such is expressed as:  K  = m  K  m  JE][S]_ Z [ES]  is therefore used as an indicator of the stability of the bound enzyme-substrate  complex. A substrate with a low K is said to bind tightly to the enzyme. m  At low substrate concentration ([S] « K ) , the Michaelis-Menten equation reduces m  to v  _  [E ][S]k n  rat  whereas at saturating concentrations ([S] » K ) the equation becomes m  v=V  m  = kcat[Eo].  At low [S] most of the enzyme is unbound such that the total enzyme concentration, which is a sum of the concentration of the free and bound enzyme, can be approximated to the concentration of the free enzyme, [E]. The Michaelis-Menten equation under these conditions is expressed as  v  =  DB]_[S]  94  The kcat/Km from the above equation is an apparent second-order rate constant which relates the reaction rate to the concentration of the free enzyme and free substrate. This kinetic parameter is also referred to as a specificity constant which is a measure of the catalytic efficiency for the substrate. The Michaelis-Menten equation is often changed to a linear form which is useful for analyzing data graphically and detecting deviations from the ideal behavior. An example of the Michaelis-Menten equation transformed is where both sides of the Michaelis-Menten equation have been inverted.  v  v  VJS]  m  Plotting 1/v as a function of 1/[S] gives the Lineweaver-Burk plot (Figure A.2) where the y-intercept is 1/V , the x-intercept is -1/K and the slope is K / V . m  m  m  (  ^ —  o  Figure A.2  m  /^Slope= A^/l/max  l/^max  i/[s]  A typical Lineweaver-Burk plot for an enzymatic reaction (Fersht, 1985).  95  A.2.  The significance  of the Michaelis-Menten  parameters  The Michaelis-Menten scheme may be extended to cover a variety of cases in which additional intermediates occur on the reaction pathway. It is found in all examples that the Michaelis-Menten equation still applies, although K  m  and kcat are now a combination of  various rate and equilibrium constants. Here, we will discuss the general mechanism proposed by Koshland for a retaining glycosidase (see Chapter 1, section 1.2). Assuming only the chemical steps are kinetically significant, the mechanism may be depicted as follows.  E +S  ki  *  ES  -l  k  X T 2  »  kc,  EP  ^  -  E+P  H 0 2  The formation of E S is referred to as the association step, the interconversion of the E S and EP as the glycosylation step and the product release as the deglycosylation step. Assuming a steady state concentration for E S and E P is reached during the reaction, then k [ES] 2  =  k [EP] 3  and =  k [E][S]-k [ES]-k [ES 1  1  2  dt =0 The total concentration of enzyme, Eo is the sum of the concentration of free enzyme and all enzyme-bound species: [ E o ] = [ E ] + [ E S ] + [ E P ] Therefore, by substituting for [EP] and rearranging, we have  k i [EQ][S] = (k j + k )[ES] + kj[ES][S] + 2  K L K  2  [ E S ] [ S  k = (k.i  + k )[ES] 2  + (kptykg  +  J  3  k k /k )[ES][S] 1  2  3  96  Solving the equation for [ES], gives k E ][S] 1L  0  [ES] = k  1  k  +  2  - ^  +  2 [S] 3  k  Since at steady state, the rate of product formation, vp is given by ;  ^I=k [EP] 3  dt  3  = k [ES] 2  then kk 2  3  [E ][S] 0  k}  Vp=  + ko  k? — k +k 2  ki + k — kj  3  2  +[S]  This follows the standard form of the Michaelis-Menten equation „ _ k [E ][S] K + [S] cat  0  m  Therefore, the kinetic parameters for this mechanism are k cat K  -  kk 2  3  k +k 2  3  k (k! + k ) 3  2  k,(k + k ) 2  k  v at-mC  k  l  3  K  2 ,  -1 * 2 T  It can be shown that kc t is the rate constant for the rate-determining step of the a  reaction and will always be associated with the highest free energy step in the pathway. kcat/Km will always be the pseudo second-order rate constant for the free enzyme and free substrate proceeding to the transition state of the first irreversible step (Fersht, 1985).  97  Now, consider the above reaction where the rate of glycosylation was increased sufficiently relative to deglycosylation, thus deglycosylation becomes rate determining, i.e. k2 » k . Further, assuming a rapid, reversible association of enzyme and substrate, then 3  k-i » k2. When these conditions are applied to the above equations, the kinetic parameters are reduced to kcat = k  3  cat' m  which expressed in Eyring form become: kcat = (kT/h)exp{-(G p*-G p)/RT} = (kT/h)exp{-AG */RT} E  E  P  kcat/Km = (kT/h)exp{-(G *-GE + s)/RT} = (kT/h)exp{-AG */RT} ES  T  where k is the Boltzmann constant and h is Planck's constant. The Eyring equations relates these rate constants to differences in energy levels among the various species on the reaction pathway. The free energy diagram for this situation is depicted schematically in Figure A.3. EP  t  G  E +P  Reaction coordinate  Figure A3  Hypothetical Gibbs free energy diagram for a retaining glycosidase showing rate-limiting deglycosylation, i.e. k3 « k2-  98  Thus, under these conditions, k c refers to the transition state of deglycosylation with the a t  enzyme-product complex as the initial reference point and kcat/K refers to the transition m  state of the glycosylation step with E and S as the initial reference points. If however, the rate of deglycosylation was increased sufficiently relative to glycosylation, then the kinetic relationships would be k3 » k 2 , and the kinetic constants can be reduced to kcat = k  k  cat  /K = m  2  k l k z  k  1 +  k  2  and the Eyring equations are  kcat = (kT/h)exp{-(G s*-G s)/RT} = (kT/h)exp{-AG */RT} E  E  C  kcat/Km = (kT/h)exp{-(G_s*-G_ + s)/RT} = (kT/h)exp{-AG */RT} T  In this case, kcat and kcat/Km both give information pertaining to the transition state of the glycosylation step. However, the initial reference point for kcat is the ES complex and for kcat/Km, it is free enzyme (E) and free substrate (S). kcat refers to the transition state of deglycosylation with the enzyme-product complex as the initial reference point and k c / K a t  m  refers to the transition state of the glycosylation step with E and S as the initial reference points. Thus, these examples, show that kcat refers to the rate determining step while kcat/Km corresponds to the first irreversible step in the reaction with E and S as the reference states.  99  A.3  Kinetics  of inactivation  of C. fimi Cex  The inactivation mechanism of a glycosidase such as C. fimi Cex can be expresed as follows:  E +I *  -  E.I  — • E-I  The first step involves a reversible binding (K ) of the inactivator (I) and the free enzyme (  (E). The second step which is rate limiting, involves an irreversible, bond forming step (kj) that forms an inactivated-enzyme intermediate (E-I). If the concentration of inactivator is much greater than the enzyme concentration (I »  E), then the amount of inactivator  remains essentially unchanged during the reaction, and the kinetics are pseudo-first order with respect to the enzyme concentration. The Michaelis-Menten equation for this inactivation can be written as:  v = k [E ][I]/(K + [I]) i  0  i  Where v is the rate of inactivation, kj is the rate constant of inactivation, and K; is the apparent dissociation constant for all species of enzyme bound inactivator, given by  K = [E][TJ/Z[EI] i  Since [I] is assumed constant over the course of the reaction, the Michaelis-Menten equation becomes:  v = k [E ] obs  t  where  k^kjPMKj + P])  100 The value of k  obs  (the pseudo first-order rate constant of inactivation) at one inactivator  concentration can be determined from the time-dependent first-order decay in residual activity by plotting the initial rates to the equation shown below, using the computer program GraFit (Leatherbarrow, 1990).  [E]  =  [EQ]  where [ E 0 ] = the initial enzyme concentration [ E ] = the active enzyme concentration  e W  APPENDIX B BASIC INTRODUCTION TO LINEAR FREE ENERGY RELATIONSHIPS AND KINETIC ISOTOPE EFFECTS  102  B. 1  Linear  free energy  relationships  B. 1.1 General concept The classic analysis of transition state structure is based on the premise that the rate constant (k) for a given reaction should be a simple function of the equilibrium constant (K) for the reaction when the structures of the reactants are systematically varied such that the following linear free energy relationship holds: k = ARG  or  logk = p(logK) + constant  where A and B are constants such that a plot of logk against logK should be a straight line with slope B. For the usual case, 8 falls in the range 0 to -1. Accordingly, Br0nsted plots have been used to examine the dependence of a rate of an enzymatic reaction with the equilibrium constant for the phenol ionization as the phenol substituents are varied, and thereby to infer the nature of the transition state relative to the structural variation explored. The term "free energy" in these relationships is appropriate as the rate is a reflection of the free energy of activation an d the equilibrium constant reflects the standard free energy change of the reaction.  B. 1.2 Linear free energy relationships in enzymology Linear free energy relationships are valuable tools in elucidating the mechanism of enzymatic reactions since they can provide valuable insights into the relationship between substrate structure and enzymatic activity. However, many enzymes are rather specific for their substrates, and therefore cannot accomodate the structural changes necessary for linear free energy analysis. Furthermore, substituent groups interact with the binding site of the enzyme in very specific ways, thus substrate modifications may affect its binding to the active site and obscure the electronic effects of the substituents on the substrate. Substrate  103 modification could also affect the orientation of the substrate reacting centre relative to the enzymatic catalytic residues and this in turn could interfere with catalysis. Despite these limitations, however, linear free energy relationships can provide strong mechanistic evidences. These studied have been extensively applied to the study of glycosidases (Dale et al., 1986; Kempton & Withers, 1992; Sinnott, 1990), including Cellulomonas fimi exoglycanase (Cex) (Tull & Withers, 1994). The results from linear free energy studies on the Cex enzyme are summarized below. The C.fimiexoglycanase (Cex) is quite reactive towards a large range of aryl 6D-glycosides. Both logkcat and log(kc t/Km) were strongly correlated with the phenol a  leaving group pK for the glucoside substrates (Figure B.l), indicating that both the ratea  determining step and the first irrevesible step in catalysis are the formation of the glycosyl-enzyme intermediate, since this is the step in which the C-0 bond to the phenolate is cleaved. The large value of the reaction constant Big = -1, which is similar to those found for other 8-glucosidases using aryl glucopyranosides [for example, Big = 0.7 for both 8-glucosidases from Agrobacterium (Kempton & Withers, 1992) and sweet  Figure B. 1  Bronsted plots relating rates of C. fimi Cex-catalyzed hydrolysis of aryl figlucosides with the leaving group ability of the phenols (Reproduced from Tull& Withers, 1994).  104 almond (Sinnott, 1990)], reflects a large degree of negative charge accumulation on the phenolate oxygen at the glycosylation transition state. This indicates that there is almost complete C-0 bond cleavage and relatively little proton donation. However, Br0nsted correlations with the cellohiosides were found to be quite different. The absence of any significant dependence of logkcat upon the leaving group p K across most of the range a  (Figure B.2A) indicates that glycosylation is not rate-limiting with these substrates, but rather the hydrolysis of the cellobiosyl-enzyme for these more active cellobiosides. The downward break observed at higher p K values (Big = -0.3) suggests that the ratea  determining step changes from deglycosylation to glycosylation with the poorer leaving groups. By contrast, the Br0nsted plot for log(kcat/Km) (Figure B.2B) shows a modest (Big = -0.3) linear dependence across the entire pKa range, as expected if the initial C-0 bond cleavage is the first irreversible step. This slope is considerable less than that determined for the glucosides ((Big = -1), reflecting some degree of charge acuumulation on the phenolate oxygen, but not as much as seen for the glucosides. Presumably, there is either less C-0 bond cleavage at the glycosylation transition state for cellobiosides, or more proton donation from the general acid catalyst in cellobioside hydrolysis.  Figure B.2  Bmnsted plots relating rates of C. fimi Cex-catalyzed hydrolysis of aryl P-cellobiosides with the leaving group ability of the phenols (Reproduced from Tull & Withers, 1994).  105  The above example demonstrates that application of linear free energy relationships in enzymology can provide useful mechanistic information regarding identification of rate determining steps, general acid-catalysis, existence of intermediates along the reaction path, as well as bond cleavage and charge distribution at the transition state.  B.2  Kinetic  isotope  effects  B . 2.1 General concept A kinetic isotope effect is defined as a change in reaction rate resulting from an isotopic substitution in a reactant. It is expressed by the ratio of the rate constants for the isotopically unsubstituted and substituted compounds (kfj/kD for deuterium isotope effects). The rate difference is dependent on the nature of the transition state. A primary isotope effect occurs in a reaction in which a bond to the isotopically substituted atom is formed or broken in the rate-determining step. Alternatively, a secondary kinetic isotope effect is a difference in reaction rate brought about by the isotopic substitution of atoms to which no bonds are formed or broken during the course of the reaction. These secondary isotope effects are the result of rehybridization of the reacting center in the rate-determining step. a-Substitutions are those where the isotope is directly attached to the atom undergoing covalency change. Isotopic substitutions do not alter the electronic nature of the isotopically-labeled compounds and have no affect on the binding of the molecule at the active site of an enzyme. This is especially important in exploring enzyme mechanisms, where altered substrates could cause a change in the reaction pathway. Isotopic substitution has most often involved replacing protium by deuterium, but the principle is applicable to nuclei other than hydrogen. Although such isotopic substitution has no effect on the qualitative chemical reactivity of the substrate, it has an  106  easily measured effect on the rate at which reaction occurs. The mass difference between the isotopes will be reflected in the frequency of the vibrating atoms according to  *-W(F) where K is the force constant of the bond and p is the reduced mass of the vibrating system, given by LL =  m,m, — — mj + m  2  with mi and m being the masses of the two atoms forming the bond. K does not change 2  with isotopic substitution The energy associated with these vibrations is called the zero-point energy, and it is defined as: E  0  hv  Thus the zero-point energy is related to the mass of the vibrating atoms by  Because of the greater mass of deuterium, the vibrations associated with a C-D bond contribute less to the zero-point energy of a molecule than does the corresponding C-H bond. For this reason, substitution of protium by deuterium lowers the zero-point energy of a molecule.  B . 2.2 Secondary deuterium kinetic isotope effects These effects have been studied especially thoroughly in the case of nucleophilic substitution reactions. They arise from changes in the bending frequencies of the isotopic bonds in reaching the transition state. If the reacting centre undergoes sp3 to sp2  107  rehybridization at the transition state of the rate determining step, then the frequency of the C-H(D) bending mode vibration is lowered, thus decreasing the force constant, K, of the vibration. Since the difference in zero-point energies of the C-H and C-D bonds is proportional to (KJ\x)^^, the energy levels will be closer at the transition state than at the ground state. This will result in a lower activation energy, and thus a faster rate, for the hydrogen-substituted reaction than that for the deuterium-substituted reaction. This is a positive kinetic isotope effect, with kn/kD > 1.0 (Figure B.3A). Alternatively, if the reacting centre undergoes sp2 to sp3 rehybridization at the transition state of the rate determining step, then the force constant for bond vibration increases, thus resulting in a greater difference in zero-point energies at the transition state than that at the ground state. The activation energy for the hydrogen-substituted reaction is now greater than that for the deuterium-substituted reaction (Figure B.3B). Inverse kinetic isotope effects of kn/kD < 1.0 are measured for this reaction type.  E  Figure B.3  Energy profiles illustrating the change in zero point energy difference between H- and D-substituted species on going to the transition state. (A) positive kinetic isotope effect, (B) inverse kinetic isotope effect.  108  A positive a-secondary deuterium kinetic isotope effect can have a maximum value of kjj/kD = 1.40, but generally values of kj-r/kD = 1.10 - 1.25 are considered indicative of a significant degree of sp3 to sp2 rehydridization at the transition state for the rate determining step. An a-secondary deuterium kinetic isotope effect of kfj/kD = 1.0 suggests little or no change in hybridization at the transition state, consistent with an S N 2 mechanism. However, S N 2 reactions can have kj-r/kD ^ 1 . 0 since temperature dependence studies as well as theoretical calculations indicate that the relative degree of bond breaking/formation and temperature can affect the magnitude of the observed isotope effect. a-Secondary deuterium kinetic isotope effects have been particularly useful as transition state probes in mechanistic studies of glycosidases. The magnitude of these effects may indicate the hybridization change on approaching the transition state, and provide insight into the bonding changes which occur during the course of the reaction.  APPENDIX C GRAPHICAL REPRESENTATION OF KINETIC DATA  110  C. 1  Lineweaver-Burk plots for the hydrolysis Cex at 25°C, pH 7.0  0  of aryl ^-cellobiosides  20  40  60  1/[3,4-DNPC] (mM ) 1  [Enzyme] = 0.0052J mg/ml, A = 400 nm, Ae = 8.18 mAf'cm'.  0  2  4  6  8  1/[PNPC] (mM ) 1  [Enzyme] =0.027 mg/ml, X = 400 nm, Ae = 0.65 mM'cm . 1  by  Ill  C.2  Lineweaver-Burk plots for the hydrolysis Cex at 25°C, pH 5.5  -10  0  of aryl ^-cellobiosides  10  20  30  1/[3,4-DNPC] (mM ) 1  [Enzyme] = 0.0103mg/ml, X = 400 nm, Ae = 12.83 mAf'cm' . 1  by  112  C.3  Lineweaver-Burk plots for the hydrolysis the His205Ala mutant at 25°C, pH 7.0  of aryl (5-cellobiosides  5000 4000  c  1  3000  >  2000 1000  1 -50  0  50  100 150 200 250 300  1/[2,4-DNPC] (mM ) 1  [Enzyme] = 0.094 mg/ml, A = 400 nm, Ae = 12.46 mAf'cni . 1  i -50  0  I i 50  I i  I i 1 i I i  I i  100 150 200 250 300  1/[3,4-DNPC] (mM") 1  [Enzyme] = 0.0908 mg/ml, X = 400 nm, Ae = 12.83 mAf'cm'  0  2  4  6  8  10  1/[PNPC] (mM") 1  [Enzyme] = 0.08mg/ml, X = 400nm, Ae = 8.21 mAf'cm'.  by  113  C.4  Lineweaver-Burk plots for the hydrolysis the His205Ala mutant at 25°C, pH 5.5  of aryl (3-cellobiosides  1600. c  ^^1200  1  800 400 1/ \  1  -1600  ,  1  -1200  1  1  -800  l „  -400  ' 0  1 400  1/[2,4-DNPC] (mM ) 1  [Enzyme] = 0.0567 mg/ml, X = 400 nm, Ae = 12.30 mAf'cm' . 1  6000 c E  4000  > 2000 . I . I . I . I . I . I  1 «t -40  -20  0  20  40  60  80  100 120  1/[3,4-DNPC] (mM ) 1  [Enzyme] = 0.0625 mg/ml, X = 400 nm, Ae = 8.18 mAf'cm' . 1  20000 15000  c 1  10000  >  1  -6  '  -3  1  .  n  0  0  i  3  i  i  i  6  i  9  i  i  12  i  i  15  1/[PNPC] (mM ) 1  [Enzyme] =0.15 mg/ml, X = 400 nm, Ae = 0.65 mAf'cm' . 1  by  114  C.5  Lineweaver-Burk plots for the hydrolysis the His205Asn mutant at 25°C, pH 7.0  of aryl ^-cellobiosides  4000 3000  c  2000  >  1000^, i  JC  in  -50  I  I  0  50  I  I 100  I  I  I  150  I  I  200  1/[2,4-DNPC] (mM ) 1  [Enzyme] = 0.158 mg/ml, A = 400 nm, Ae = 12.46 mAf'cm' . 1  12000 *  9000 6000 3000 0 -10  '  0  I  10  20  30  40  l  l  50  1/[3,4-DNPC] (mM ) 1  [Enzyme] = 0.05mg/ml, A = 400nm, Ae = 12.83 mAf'cm'.  by  Lineweaver-Burk plots for the hydrolysis the His205Asn mutant at 25°C, pH 5.5  of aryl  fi-cellobiosides  by  116  C. 7  Lineweaver-Burk plots for the hydrolysis the Gln87Met mutant at 37°C, pH7.0  -0.2  0  0.2  0.4  of aryl ^-glycosides  0.6  by  0.8  1/IPNPG] (mM' ) 1  [Enzyme] = 0.028 mg/ml, X = 400 nm, Ae = 7.28 mAf'cm . 1  250 200 150 100 50 i  ^  I  0  0.4  0.8  1.2  i  i  1.6  1/[3,4-DNPG] (mM ) 1  [Enzyme] = 2.5 x Iff mg/ml, X = 400 nm, Ae = 11.03 mAf'cm' . 3  1  117  0  0.2  0.4  0.6  1/IPNPX] (mM ) 1  [Enzyme] =0.01562 mg/ml, X = 400 nm, Ae = 7.28 mAf'cm . 1  1/13,4-DNPX] (mM ) 1  [Enzyme] = 1.27 x 10' mg/ml, X = 400 nm, Ae = 7.28 mAf'cm' . 3  1  0  100  200  300  400  1/[3,4-DNPXJ (mM ) 1  [Enzyme] = 2.55 x 10' mg/ml, X = 400 nm, Ae = 11.03 mAf'cm' . 4  1  1/[PNPXJ (mM ) 1  [Enzyme] = 2.55 x 10' mg/ml, X = 400 nm, Ae = 7.28 mAf'cm' . 4  1  1/[PhXJ (mM ) 1  [Enzyme] = 0.0156 mg/ml, X = 400 nm, Ae =2.172 mAf'cm' . 1  

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