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Insights into the catalytic mechanism of a retaining xylanase from Cellulomonas fimi Wicki, Jacqueline 2007

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INSIGHTS INTO THE CATALYTIC MECHANISM OF A RETAINING XYLANASE FROM CELLULOMONAS FIMI  by J A C Q U E L I N E WICKI B . S c , The University of British Columbia, 1998  A THESIS SUBMITTED IN PARTIAL FULFILLMENT O F THE R E Q U I R E M E N T S FOR T H E D E G R E E O F DOCTOR OF PHILOSOPHY  in  T H E F A C U L T Y O F G R A D U A T E STUDIES  (Chemistry)  THE UNIVERSITY O F BRITISH C O L U M B I A July 2007 © Jacqueline Wicki, 2007  ABSTRACT The family 10 xylanase from Cellulomonasfimi(Cex) is an important model enzyme on which numerous mechanistic studies have been performed. This enzyme catalyzes the hydrolysis of P-glycosidic linkages via a double-displacement mechanism involving the formation and subsequent breakdown of a covalent glycosyl-enzyme intermediate with net retention of stereochemistry at the centre undergoing substitution. The finer details of the mechanism of this enzyme were investigated in three studies in order to gain a better understanding of this family of enzymes. In the first study presented in Chapter 2, the roles of key active-site residues in the catalytic mechanism of Cex were investigated by utilizing site-directed mutagenesis in combination with steady state kinetic analyses and pH-rate dependencies. The ratedetermining step for the aryl substrates tested remains deglycosylation for many of the enzymes, while the altered pH profiles demonstrate a role for these highly conserved residues in the hydrogen-bond network responsible for maintaining the ionization state of the two catalytic residues. In Chapter 3, a second study addresses a fundamental enquiry of mechanistic enzymology; that is, how distal and proximal substrate interactions influence catalysis. By systematically removing hydrogen-bonding interactions through modification, individually, of substrate and enzyme, deep insight is gained into the effects of these modifications on each step of the hydrolysis reaction catalyzed by Cex and a family 11 xylanase (Bex). The data obtained provide significant insight into the contributions of hydrogen-bonding interactions at the distal and proximal sites.  The strongest bond  energies were measured in the proximal site, suggesting that these interactions are critical  ii  for substrate binding and bond hydrolysis. A particularly important finding of this study is that both 'uniform' and 'differential' binding interactions are recruited in the active site of a single enzyme. The third study, presented in Chapter  examines how well a series of five high  4,  *  2  3  affinity inhibitors mimic the transition state of Cex as a function of the sp - or sp hybridization state of the "anomeric carbon".  Kinetic parameters for o-nitrophenyl  P-xylobioside were determined, and very good correlations were observed in logarithmic plots relating the K\ value for the sp -hybridized class of inhibitor with 10 mutants and k JK a  m  for the hydrolysis of the substrate by the corresponding mutants. The dependence  was significantly less in the plot of log(& /AT ) versus log(lAfvi) for the sp -hybridized 3  cat  m  class of inhibitor, indicating that the sp -hybridized class of inhibitors more closely 2  mimics the geometry of the transition state than does the sp -hybridized class of inhibitors.  m  TABLE OF CONTENTS ABSTRACT  ii  TABLE OF CONTENTS  iv  LIST OF TABLES  xi  LIST OF FIGURES  xiii  LIST OF ABBREVIATIONS  xix  ACKNOWLEDGEMENTS  xxi  DEDICATION  xxii  1  GENERAL INTRODUCTION  1.1  1.1.1 1.1.2 1.1.3 1.2  1.2.1 1.2.2 1.2.3 1.2.4 1.2.5 1.2.6 1.3  1.3.1 1.3.2 1.4  2  GLYCOSIDASES  3  Traditional classification systems Subsite nomenclature Glycosidase families C A T A L Y T I C M E C H A N I S M OF RETAINING P-GLYCOSIDASES  Oxocarbenium ion-like transition states Nature of the covalent glycosyl-enzyme intermediate Nucleophilic participation Acid/base catalysis Non-covalent interactions Substrate distortion B I O D E G R A D A T I O N OF C E L L U L O S E A N D X Y L A N  4 6 7 7  11 14 18 22 28 30 34  Cellulomonasfimixylanase Bacillus circulans xylanase A I M S O F THIS T H E S I S  36 42 46  iv  TABLE OF CONTENTS ABSTRACT  ii  TABLE OF CONTENTS  iv  LIST OF TABLES  xi  LIST OF FIGURES  xiii  LIST OF ABBREVIATIONS  xix  ACKNOWLEDGEMENTS  xxi  DEDICATION  xxii  GENERAL INTRODUCTION  1 1.1  2  GLYCOSIDASES  1.1.1 1.1.2 1.1.3 1.2  Traditional classification systems Subsite nomenclature Glycosidase families C A T A L Y T I C M E C H A N I S M OF RETAINING (3-GLYCOSIDASES  1.2.1 1.2.2 1.2.3 1.2.4 1.2.5 1.2.6 1.3  Oxocarbenium ion-like transition states Nature of the covalent glycosyl-enzyme intermediate Nucleophilic participation Acid/base catalysis Non-covalent interactions Substrate distortion BIODEGRADATION OF C E L L U L O S E A N D X Y L A N  1.3.1 1.3.2 1.4  3  4 6 7 7  11 14 18 22 28 30 34  Cellulomonas fimi xylanase Bacillus circulans xylanase A I M S O F THIS T H E S I S  36 42 46  iv  2  CHARACTERIZATION OF HIGHLY CONSERVED RESIDUES IN THE ACTIVE SITE OF  CELLULOMONAS  FIMI  XYLANASE  49  2.1  INTRODUCTION  49  2.2  RESULTS  52  2.2.1 2.2.2 2.2.3 2.2.4  Generation of mutants of Cex Production, purification, and physical characterization Kinetic analysis: pH dependence of k tlK Kinetic analysis: steady state kinetics CSi  2.2.4.1 2.2.4.2 2.2.4.3 2.3  Probing the -2 subsite: steady state kinetic analyses of Glu43, Asn44, and Lys47 mutants Probing the -1 subsite: steady state kinetic analyses of His80 mutants Probing the -1 subsite: steady state kinetic analyses of Asnl26 and Asnl69 mutants  DISCUSSION  2.3.1 2.3.2 2.3.3  2.3.4 2.3.5 2.3.6  RoleofGlu43 RoleofAsn44 RoleofLys47 RoleofHis80  67 71  73 78 81 81 82 83 84  Amino acids that influence the catalytic residues  2.3.4.1 2.3.4.2  62  73  Effects of mutations on glycosylation and deglycosylation A second look at the rate-limiting step in the hydrolysis reaction by Cex and mutants: diffusion-controlled rate constants Importance of -2 subsite amino acids Glu43, Asn44, and Lys47  2.3.3.1 2.3.3.2 2.3.3.3 2.3.3.4  2.4  m  52 53 54 56  RoleofAsnl26 RoleofAsnl69  85 85 86  Analyses of the pH profiles for Cex variants Activity of family 10 enzymes CONCLUSIONS  87 90 92  v  3  AN ASSESSMENT OF THE BINDING ENERGY CONTRIBUTION OF HYDROGEN BONDS AT THE TRANSITION STATE USING DEOXYGENATED SUBSTRATES AND SITE-DIRECTED MUTAGENESIS  3.1  INTRODUCTION  3.1.1 3.1.2 3.1.3 3.1.4 3.2  Hydrogen-bonding energetics in water Hydrogen bonding to Electronic effects Objectives  fluorine  3.2.1.2 3.2.1.3  3.2.2  103  Kinetic analysis of family 10 C. fimi xylanase Kinetic analysis of family 11 B. circulans xylanase Comparison of results for two xylanases from distinct families  Binding energy contributions probed by using mutant enzymes  3.2.2.1  3.2.2.2 3.2.2.3  3.2.3  96 99 100 102  Binding energy contributions probed by using substrate analogues .... 103  3.2.1.1  3.3  94  RESULTS A N D DISCUSSION  3.2.1  94  Design of the mutations Subsite - 2 interactions Subsite - 1 interactions  104 112 118  123 124  125 129  Uniform and differential binding interactions  130 133  CONCLUSIONS  vi  4  INQUIRY INTO THE TRANSITION-STATE ANALOGY OF A SERIES OF IMINOSUGAR INHIBITORS FOR A XYLANASE  4.1  INTRODUCTION  136  4.1.1 4.1.2 4.1.3  Transition-state theory Enzyme catalysis Experimental approaches to enzymatic transition-state structure 4.1.3.1 K I E to determine TS structure 4.1.3.2 Computational approaches to the transition state 4.1.3.3 TS analogues 4.1.4 Criteria for transition-state analogy 4.1.4.1 Strong inhibition 4.1.4.2 Slow inhibition 4.1.4.3 Free energy relationships 4.1.4.4 Thermodynamic parameters 4.1.4.5 pH profiles 4.1.5 Inhibitors of glycoside hydrolysis 4.1.6 Inhibitors of Cex 4.1.7 Aims of this study 4.2  R E S U L T S A N D DISCUSSION  4.2.1 4.2.2 4.2.3 4.2.4  136  136 138 140 140 141 142  143 144 144 147 150 151  152 156 158 159  Inhibition of Cex mutants by iminosugar inhibitors 160 How good a transition-state analogue is the inhibitor? 163 The pK values of xylobio-derived inhibitors 168 The pH dependence of the binding of xylobio-derixed inhibitors 171 4.2.4.1 Xylobio-dehved lactam oxime, imidazole, and isofagomine lactam 172 4.2.4.2 Xylobio-derived deoxynojirimycin and isofagomine 174  4.3  a  C O N C L U S I O N : TRANSITION-STATE MIMICS OR FORTUITOUS BINDERS?  vii  175  5  MATERIALS AND METHODS  5.1  5.1.1 5.1.2 5.1.3 5.1.4 5.1.5 5.1.6 5.2  5.2.1 5.2.2 5.2.3 5.2.4 5.2.5 5.2.6 5.2.7  178  MOLECULAR BIOLOGY  178  Reagents, enzymes, and bacterial strains Oligonucleotides Site-directed mutagenesis by linear amplification Site-directed mutagenesis by PCR.... Overexpression and purification by active-site mutants of Cex Protein mass determination E N Z Y M E KINETICS  178 178 180 183 186 188 188  General materials and procedures Steady state kinetics pH stability studies pH dependence studies Kinetics of inhibition pK determinations for inhibitors Determination of extinction coefficients a  188 189 191 192 193 195 195  REFERENCES  198  APPENDICES  228  viii  A  GRAPHICAL REPRESENTATION OF DATA  A. 1  S T E A D Y S T A T E KINETICS FOR T H E HYDROLYSIS OF XYLOBIOSIDES OR CELLOBIOSIDES  A. 1.1 A.1.2 A.1.3 A.1.4 A.1.5 A.1.6 A.1.7 A.1.8 A.1.9 A.1.10 A.1.11 A.1.12 A.1.13 A.2  228  Native Cex Cex E43A CexN44A CexK47A CexH80A Cex IKON CexH80Q CexQ87M CexQ87Y CexN126A CexE127A Cex N169A CexH205N  228 231 235 239 242 246 250 254 257 260 263 264 267  S T E A D Y S T A T E KINETICS FOR INHIBITION B Y IMINO S U G A R INHIBITORS  A.2.1 A.2.2 A.2.3 A.2.4 A.2.5 A.2.6 A.2.7 A.2.8 A.2.9 A.2.10 A.2.11 A.3  228  Cex E43A CexN44A CexK47A CexH80A CexH80N Cex H80Q CexQ87M CexQ87Y CexN126A Cex E127A CexN169A  269  269 271 273 275 277 279 281 283 285 287 289  P H DEPENDENCIES OF k /K CAT  M  FOR T H E HYDROLYSIS OF XYLOBIOSIDES  OR CELLOBIOSIDES B Y N A T I V E A N D M U T A N T C E X  ix  292  B  COMPILATION OF CEX DATA  298  B.l  TABLES  298  B. 2  REFERENCES  310  C  FUNDAMENTALS OF ENZYME KINETICS  312  C l  B A S I C E N Z Y M E KINETICS  312  C. 2  T H E SIGNIFICANCE OF M I C H A E L I S - M E N T E N PARAMETERS  316  C.3  E N Z Y M E INHIBITION  322  C.3.1  Competitive inhibition  322  C.3.2  Noncompetitive, uncompetitive, and mixed inhibition  323  C.4  BINDING E N E R G Y A N D E N Z Y M E C A T A L Y S I S  325  C.5  T H E P H D E P E N D E N C E OF E N Z Y M E C A T A L Y S I S  327  C.5.1 C.5.2  The Michaelis-Menten mechanism Modifications and breakdown of the Michaelis-Menten mechanism  x  328 330  LIST OF TABLES Table 2.1: Total yields of Cex mutants produced  54  Table 2.2: Determination of molecular weight by mass spectrometry  54  Table 2.3: pH Dependence of k JK c  m  for C.fimixylanase and mutants  56  Table 2.4: Michaelis-Menten parameters for the hydrolysis of various aryl glycoside substrates by C.fimi xylanase  61  Table 2.5: Michaelis-Menten parameters for the hydrolysis of various aryl glycoside substrates by several family 10 xylanases  62  Table 2.6: Michaelis-Menten parameters for the hydrolysis of aryl glycosides by the C.fimixylanase (Cex) and mutants E43A, N44A, and K47A  63  Table 2.7: Michaelis-Menten parameters for the hydrolysis of aryl glycosides by the C.fimixylanase (Cex) and mutants H80A, H80N, and H80Q  68  Table 2.8: Michaelis-Menten parameters for the hydrolysis of aryl glycosides by the C.fimixylanase (Cex) and mutants N126A and N169A  72  Table 2.9: Kinetic parameters for the glycosylation and deglycosylation steps of the hydrolysis reaction by variants of Cex  74  Table 2.10: Free energy changes at the glycosylation and deglycosylation steps of the hydrolysis reaction by variants of Cex  76  Table 3.1: Kinetic parameters for the hydrolysis of deoxy/deoxyfluoro analogues by wild-type C.fimixylanase  105  Table 3.2: Summary of hydrogen-bonding interactions between the xylobiosyl intermediate and protein residues in the active site of the 2-deoxy2-fluoroxylobiosyl-enzyme complex of Cex 110 Table 3.3: Kinetic parameters for the hydrolysis of deoxy/deoxyfluoro analogues of PNPX2 by wild-type B. circulans xylanase  113  Table 3.4: Summary of hydrogen-bonding interactions between the xylobiosyl intermediate and protein residues in the active site of the 2-deoxy2-fluoroxylobiosyl-enzyme complex of Bex (81) 118 xi  Table 3.5: Binding energy contribution AAG* (kcalmol ') of individual hydroxyl groups in the hydrolysis of PNPX2 by Cex and Bex 120 Table 3.6: Kinetic parameters for the hydrolysis of mutant C.fimixylanase  PNPX2  by wild-type and 126  Table 3.7: Apparent binding energy contribution, AAG* (kcalmol ), of individual substrate hydroxyl groups in the hydrolysis of deoxy/deoxyfluoro analogues of PNPX by wild-type C.fimixylanase and selected mutants -1  2  128  Table 4.1: Inhibition constants (K\) for Cex and Bex with various jcy/06/0-derived iminosugars 157 Table 4.2: Inhibition constants (K() for C. fimi xylanase and mutants with various xylobio-derived iminosugars  161  Table 4.3: Xylobio-derived inhibitor pK values of the conjugate acid  170  Table 4.4: Parameters determined from the pH profile for the binding of xylobioderived inhibitors to wild-type Cex  172  Table 5.1: Primers used in the mutagenesis of C.fimixylanase  179  a  Table 5.2: pH Incubation mixture for enzyme stability studies  191  Table 5.3: Standard assay conditions for enzyme stability  192  Table 5.4: Standard assay conditions for pH profiles  193  Table 5.5: Extinction coefficients of substituted phenols at 37°C in sodium phosphate buffer, pH 7.00 196  xii  LIST OF FIGURES Figure 1.1: Glycone and aglycone moieties of a P-glucoside are shown with the arrow indicating the scissile glycosidic bond  3  Figure 1.2: Reactions catalyzed by retaining and inverting P-glucosidases  5  Figure 1.3: A schematic depiction of endo and exo sites of cleavage of an oligosaccharide by glycosidases  5  Figure 1.4: Schematic drawing of sugar-binding subsites in two glycosidases. By convention, the non-reducing end of the substrate is drawn on the left while the reducing end is drawn on the right. The arrow placed between the -1 and +1 subsites indicates the point of cleavage  6  Figure 1.5: The hydrolysis reactions catalyzed by retaining and inverting glycosidases  8  Figure 1.6: General double-displacement mechanism for a retaining glycosidase  10  Figure 1.7: Inactivation and reactivation by 2-deoxy-2-fluoro glycosides. (A) Inactivation by formation of the fluoroglycosyl-enzyme intermediate. (B) Reactivation by transglycosylation of the fluoroglycosyl-enzyme intermediate  16  Figure 1.8: Structures of several classes of the fluoro sugars that function as mechanism-based inactivators  17  Figure 1.9: Nucleophilic fluorination of 2,4-dinitrophenyl P-mannoside by the P-mannosidase nucleophile mutant (Glu519Ser) from Cellulomonas fimi  21  Figure 1.10. The anti orientation of the acid catalytic residue and protonation trajectory as the basis for rationalizing the relative potency of two triazole inhibitors. (A) ^nri-protonation of the glycosidic oxygen. (B) Absence of any possible hydrogen bonding to the 1,2,3-triazole inhibitor. (C) Possible aw/i-protonation of the 1,3,4-triazole inhibitor  23  Figure 1.11: Depiction of the pH dependence of k JK for Bex Asn35Asp mutant as the product of titration curves of catalytic residues derived from C-NMR. Although only 1 % of the mutant is in the correct ionization state for catalysis, it is 20% more active than the wild-type enzyme 27 c  m  13  xiii  Figure 1.12: The Asn35Asp mutant of Bex increases the inherent catalytic efficiency by at least 100-fold over the wild-type enzyme by forming a strong hydrogen bond between Asp35 and Glul72  28  Figure 1.13: Partial map of pyranoside interconversions adapted from Stoddart (78). Potential transition-state conformations that possess a coplanar arrangement of C5, 05, C2 and C l atoms are shown boxed  32  Figure 1.14: Orientation of the lone pairs on the endocyclic oxygen relative to the glycosidic oxygen of the leaving group. (A) The C\ ground state conformation of many common glycosides and covalent intermediate structures. This conformation features a lone pair on the ring oxygen that is antiperiplanar to the C l - O R bond in an a-configured glycoside. (B) The Hi conformation proposed as a possible transition-state conformation. (C) The 5 or E conformation and (D) the skew-boat ST, conformation that features a lone pair on the ring oxygen that is antiperiplanar to the C l - O R bond in a P-configured glycoside, as required for the optimal effect of the antiperiplanar lone pair hypothesis (ALPH)  34  Figure 1.15: Three-dimensional structure of Cex with bound 2 F - X 2 (76). The overall structure of Cex is depicted as a protein cartoon in divergent (walleyed) stereo, with a-helices coloured green, P-strands blue and loops red. The bound 2 F - X 2 is drawn as a stick model, with atoms coloured according to atom type, carbon in grey, oxygen in red, and fluorine in yellow. The water molecules in the active site have been removed for clarity. This figure and subsequent molecular representations were prepared with Chem3D Ultra 10.0 (http://www.cambridgesoft.com)  37  4  4 1  A  ]  Figure 1.16: Active site of the Cellulomonasfimixylanase trapped as a covalent 2-deoxy-2-fluorocellobiosyl-enzyme intermediate. Adapted from (32) 40 Figure 1.17: Hydrogen bonding at the transition state  42  Figure 1.18: Three-dimensional structure of Bex with bound 2 F - X 2 (81). The overall structure of Bex is depicted as a protein cartoon in divergent (walleyed) stereo, with a-helices coloured green, P-strands blue and loops red. The bound 2 F - X 2 is drawn as a stick model, with atoms coloured according to atom type, carbon in grey, oxygen in red, and fluorine in yellow. The water molecules in the active site have been removed for clarity. This figure was prepared with Chem3D Ultra 10.0 (http://www.cambridgesoft.com)  43  Figure 1.19: Active site of the Bacillus circulans xylanase trapped as a covalent 2-deoxy-2-fluoroxylobiosyl-enzyme intermediate  44  xiv  Figure 1.20: The 2-deoxy-2-fluoroxylobiosyl-enzyme intermediate formed on the Bacillus circulans xylanase. Green dashes represent hydrogen bonds, the blue dash depicts a non-hydrogen bond interaction, and the R group represents the saccharide unit bound in the -2 subsite of the active site. Adapted from Sidhu et al. (81)  46  Figure 2.1: Scheme of protein-carbohydrate interactions in the active site region of 2F-xylobiosyl-Cex-cd complex observed by three-dimensional structural analysis (76). Hydrogen-bonding interactions between heteroatoms are in Angstroms. The distance between 2F and the carbonyl oxygen of Glu233 is 2.6 A (not shown)  51  Figure 2.2: Overview of the QuikChange™ Site-Directed Mutagenesis Kit  53  Figure 2.3: Data for the hydrolysis of o-nitrophenyl xylobioside by wild-type Cex plotted in (A) and (B) Michaelis-Menten form; and (C) and (D) LineweaverBurk form. Kinetic parameters for the hydrolysis reaction were obtained by fitting data obtained at lower substrate concentrations (unbroken line), while those for the hydrolysis reaction were obtained by fitting data obtained at higher substrate concentrations (dashed line)  60  Figure 2.4: pH profiles of wild-type (O) and Glu43, Asn44, and Lys47 mutants of Cex. The pH dependence of wild-type Cex was measured using 2,4DNPC where pK \ = 3.9 and pK^ -1.1. (A) The E43A (•) mutant enzyme activity was measured using 2,4DNPC where pK \ =3.7 and pA^ = 6.9. (B) The N44A (•) mutant enzyme activity was measured using 2,4DNPC where pK \ = 3.9 and pK& = 8.0. (C) The K47A ( A ) mutant enzyme activity was measured using 2,5DNPX2 where pK \ =3.9 and pK^ = 6.0  66  Figure 2.5: pH profiles of wild-type (O) and His80 mutants of Cex. The pH dependence of wild-type Cex was measured using 2,4DNPC where pA^i =3.9 and pKa2 = 7.7. (A) The H80A (•) mutant enzyme activity was measured using PNPX where pK^ = 4.6 and pK& = 6.2. (B) The H80N (•) mutant enzyme activity was measured using PNPX2 where pA" i - 4.6 and pK& = 6.3. (C) The H80Q ( A ) mutant enzyme activity was measured using PNPX2 where pK \ = 4.6 and pK^ = 7.0  70  Figure 2.6: pH profiles of wild-type (O) and Asnl26 and Asnl69 mutants of Cex. The pH dependence of wild-type Cex was measured using 2,4DNPC where pK \ = 3.9 and pK& = 7.7. (A) The N126A (^mutant enzyme activity was measured using 2,4DNPC where p ^ i = 4.4 and pK^ = 8.5. (B) The N169A (•) mutant enzyme activity was measured using 2,4DNPC where pK = 4.0 and pK^ = 8.2  73  a  a  a  a  2  a  a  a  a]  xv  Figure 2.7: (A) Comparison of rate constants for glycosylation of PNPC and PNPX by variants of Cex. A slope of m = 1.0 and correlation coefficient of r = 0.87 was observed. (B) Comparison of rate constants for deglycosylation of cellobiosyl-enzyme intermediates and xylobiosyl-enzyme intermediates of variants of Cex. A slope of m = 0.94 and correlation coefficient of r = 0.95 was observed. (C) Comparison of rate constants for glycosylation and deglycosylation reactions for the hydrolysis of O N P X 2 by variants of Cex. Open circles correspond to mutations within the -2 subsite; and closed circles correspond to mutations within the -1 subsite of Cex. A slope of m = 0.97 and correlation coefficient ofr = 0.62 was observed  77  Figure 3.1: The hydroxyl can act as a hydrogen-bond donor or acceptor, whereas fluorine can only act as a hydrogen-bond acceptor  95  2  Figure 3.2: Chromogenic synthetic substrates. PNPX2, 2'-deoxy-PNPX2, 2'-deoxy2'-fluoro-PNPX , 3'-deoxy-PNPX , 3'-deoxy-3'-fluoro-PNPX , 4'-deoxyPNPX2, and 4'-deoxy-4'-fluoro-PNPX2 were prepared by Dr. Johann Schloegl. 3'-Deoxy-3'-fluoro-PNPX2 was prepared by Dr. Chris A. Tarling 104 2  2  2  Figure 3.3: Schematic representation of proposed intermolecular hydrogen-bond interactions between 4-nitrophenyl p-xylobioside and Cex based on substrate analogue data, X-ray crystallography and NMR studies 109 Figure 3.4: Scheme of protein-carbohydrate interactions in the active site region of the 2-fluoroxylobiosyl-Bcx complex observed by three-dimensional structural analysis (81). Hydrogen-bonding interactions between heteroatoms are in Angstroms 115 Figure 3.5: Binding energy contribution AAG* (kcalmoF ) plot for a series of deoxy- and deoxyfluoro-P-xylobioside analogues with B. circulans xylanase (Bex) and C.fimi xylanase (Cex) 121 1  Figure 3.6: Scheme of protein-carbohydrate interactions in the active site region of 2F-xylobiosyl-Cex-cd complex observed by three-dimensional structural analysis (76). Hydrogen-bonding interactions between heteroatoms are in Angstroms. The distance between 2F and the carbonyl oxygen of Glu233 is 2.6 A (not shown) 123 Figure 3.7: Reaction coordinate diagram showing the consequences of modifications to the (a) distal sugar and (b) proximal sugar 132 Figure 4.1: A reaction profile. The horizontal axis is the reaction coordinate and the vertical axis is the potential or Gibbs free energy. The transition state corresponds to the maximum of the Gibbs free energy 137 xvi  Figure 4.2: Reaction coordinate diagram to show how an enzyme can increase the rate of reaction of a bound substrate by stabilizing the transition state 139 Figure 4.3: Two potential mechanisms for slow-binding inhibition  145  Figure 4.4: Thermodynamic box illustrating the relationship between ground-state and transition-state binding. The designations E, S, and P refer to enzyme, substrate in the ground state form, and product, respectively, while S represents the transition state structure 147 x  +  Figure 4.5: Structures of some naturally occurring and synthetic glycosidase inhibitors in the g/wco-configuation 153 Figure 4.6: Structures of the 5 iminosugars studied  160  Figure 4.7: Inhibition of Cex N169A by xy/ofo'o-imidazole at varying concentrations of 2,4DNPC. Plot A is the Michaelis-Menten plot, plot B is the Lineweaver-Burk plot, and plot C is the Eadie-Hofstee plot. The concentrations of xylobio-imidazole were 0 (O), 0.123 (•), 0.370 (•), 0.925 (•), and 2.47 uM (A) 162 Figure 4.8: Transition state analogy plots for C. fimi xylanase Cex. Comparison between k JK for ONPX and inhibitor K\ values for xy/oZno-derived (A) lactam oxime 1; (B) imidazole 2; (C) isofagomine lactam 3; (D) deoxynojirimycin 4; and (E) isofagomine 5 167 c  m  2  Figure 4.9: Titration curves for the conjugate acid forms of the xy/o&z'o-derived lactam oxime (•), imidazole (•), deoxynojirimycin (T), and isofagomine (•). Note: amides are strong bases (conjugated acid pK between -6 and -10); therefore, the pK of the xy/ofoo-isofagomine lactam is likely well below -5 169 a  a  Figure 4.10: pH dependence of k JK for Cex (•), \IK\ for xylobio-derived lactam oxime pK 4.9 (•), imidazole pK 5.9 (•), isofagomine lactam pK < -5 ( A ) , deoxynojirimycin pK 6.9 (T), and isofagomine pK 8.8 (•). Fits to "bellshaped" profiles are shown for k JK (dashed line) and \IK\ (coloured solid lines) 171 c  a  m  a  a  a  a  c  m  xvii  Figure 5.1: Introducing mutations by the QuikChange™ Site-Directed Mutagenesis method. A supercoiled dsDNA vector with an insert of interest ( • ) and two synthetic oligonucleotide primers ( A ) containing the desired mutation are mixed. The oligonucleotide primers, each complementary to opposite strands of the vector, are extended during temperature cycling using Pfu DNA polymerase to generate mutated plasmid containing staggered nicks. Following temperature cycling, the methylated, nonmutated parental DNA template is digested with Dpn I. The circular, nicked dsDNA is transformed into X L 1-Blue supercompetent cells. After transformation, the nicks in the mutated plasmid are repaired by the XLl-Blue supercompetent cells 181 Figure 5.2: Restriction map of pUC 12-1.1 cejc(PTis)  183  Figure 5.3: Introducing mutations by in vitro overlap-extension PCR. Two PCRs are performed to produce two fragments that carry overlapping sequences. These two fragments are then mixed, denatured, and annealed to obtain mutant DNAs in a further PCR 184  xviii  LIST OF ABBREVIATIONS ALPH Bex 5cXynllA BSA CAZy Cex C/XynlOA DEGLY EC ESI-MS/MS GLY HEWL HPLC IPTG IUBMB kDa M MS/MS NMR PDB RP-HPLC TS WT  antiperiplanar lone pair hypothesis Bacillus circulans xylanase (family 11) Bacillus circulans xylanase (family 11) bovine serum albumin carbohydrate-active enzyme database Cellulomonas fimi xylanase (family 10) Cellulomonas fimi xylanase (family 10) deglycosylation Enzyme Commission (classification number) of the International Union of Biochemistry electrospray ionization tandem mass spectrometry glycosylation hen egg white lysozyme high performance liquid chromatography isopropyl P-D-thiogalactoside International Union of Biochemistry and Molecular Biology kiloDalton mutant enzyme tandem mass spectrometry nuclear magnetic resonance protein data bank reverse phase high performance liquid chromatography transition state wild-type enzyme  Substrates  3F-PNPX 2'H-PNPX 2'F-PNPX  />nitrophenyl 3-deoxy-3-fluoro-P-xylobioside  2  2  2  3'H-PNPX 3'F-PNPX 4'H-PNPX 4'F-PNPX 2F-DNPC DNPC PNPC PNPX ONPX 2,5DNPX  2  2  2  2  2  2  2  jp-nitrophenyl 2 '-deoxy-P-xylobioside />nitrophenyl 2'-deoxy-2'-fluoro-P-xylobioside />nitrophenyl 3'-deoxy-p-xylobioside />nitrophenyl 3 '-deoxy-3 '-fluoro-P-xylobioside />nitrophenyl 4'-deoxy-P-xylobioside /7-nitrophenyl 4'-deoxy-4'-fluoro-P-xylobioside 2,4-dinitrophenyl 2-deoxy-2-fluoro-P-cellobioside 2,4-dinitrophenyl P-cellobioside />nitrophenyl P-cellobioside />nitrophenyl P-xylobioside o-nitrophenyl P-xylobioside 2,5-dinitrophenyl P-xylobioside  xix  Amino  Ala Arg Asn Asp Cys Glu Gin Gly His He Leu Lys Met Phe Pro Ser Thr Tip Tyr Val  Acid  Abbreviations  A R N D C E Q G H I L K M F P S T W Y V  alanine arginine asparagine aspartic acid cysteine glutamic acid glutamine glycine histidine isoleucine leucine lysine methionine phenylalanine proline serine threonine tryptophan tyrosine valine  XX  ACKNOWLEDGEMENTS First and foremost, I would like to express my sincere gratitude to my supervisor, Professor Stephen G. Withers, for all of his guidance, encouragement, and tremendous patience. His insights and enthusiasm were a source of inspiration for me and I consider it a great privilege to have been part of his research group. I wish to thank all the members of the Withers group, past and present, for their friendship and all their assistance. The technical support of Shouming He, Ms. Karen Rupitz, and Ms. Emily Kwan is also greatly appreciated. I would like to extend my gratitude to the following people who contributed to this work: Professor Spencer Williams for a fruitful collaboration on the TSA project and for synthesizing the iminosugar inhibitors; Dr. Johann Schloegl for a fruitful collaboration on the deoxysugar project and for synthesizing all distal deoxy/deoxyfluoro xylobioside substrates; Dr. Chris Tarling for synthesizing the proximal xylobioside substrate; and Professor R. Anthony Warren for access to his "Cellulase" lab. And finally, to all my family and friends, thank you for your support and steadfast belief in me.  xxi  Mamie mojej Grazynie, ktorej nieustajqca cierpliwosc i oddanie sprawify, ze powstanie tej pracy stalo siq mozliwe.  xxii  CHAPTER 1  GENERAL INTRODUCTION  Chapter 1  1  General Introduction  2_  General Introduction Carbohydrates comprise a diverse array of simple and complex biological  molecules. They are the most abundant class of organic molecules found in living organisms. Unlike the other major classes of biopolymers such as proteins and nucleic acids, carbohydrate compounds are not synthesized according to a template, and this, in large part, accounts for their remarkable diversity. The vast assortment of biological functions expressed by carbohydrates is, at least in part, a consequence of the immense variety of distinctive structural motifs generated by synthetic enzymes. Carbohydrate polymers such as starch and glycogen are traditionally recognized as major sources of metabolic energy, while polymers such as cellulose and chitin are suitable for the maintenance of physical structures.  Various oligosaccharides are  ubiquitous as components of lipids, proteins, nucleic acids, and other complex biomolecules. The role of carbohydrates in molecular recognition and signal transduction is critical to the performance of biological systems.  As a result of their biological  importance and potential medical applications, carbohydrates, along with the enzymes responsible for their metabolism, are now the subjects of extensive research efforts. Glycosidases are one such group of enzymes, and are responsible for the transfer of glycosyl moieties from a glycoside to water, the result being hydrolysis. Due to the biologically widespread process of glycoside cleavage, glycosidase inhibitors have potential therapeutic value. An understanding of the mechanism by which these enzymes hydrolyse glycosidic bonds and of how to inhibit them is fundamental to their application in medicine and industry.  Chapter 1  1.1  General Introduction  3  Glycosidases Glycosidases, alternatively known as glycoside hydrolases, comprise a large  family of enzymes that catalyze the transfer of a glycosyl moiety from a donor sugar to an acceptor water molecule. These enzymes are typically involved in the biodegradation of complex polymeric carbohydrate molecules to their simple sugar constituents. glycosidic  bonds  that join polysaccharides  display  formidable  stability,  The as  is  demonstrated by the five million year half-life of a common glycosidic linkage under neutral aqueous conditions (7). Hydrolysis of the glycosidic scissile bond occurs between the glycpne, which is typically a carbohydrate moiety, and the aglycone, which commonly consists of another saccharide unit, but may also be an alcohol, phosphate, or thiol group (Figure 1.1).  Glycosidases catalyze the breakdown of such linkages by  factors on the order of 10 -fold, representing one of the largest rate accelerations attained 17  by any class of enzyme.  Furthermore, these enzymes are relatively promiscuous with  regards to aglycone specificity; consequently, a detailed kinetic analysis may be carried out using a wide array of synthetic substrates.  Glycone  Aglycone  Figure 1.1: Glycone and aglycone moieties of a B-glucoside are shown with the arrow indicating the scissile glycosidic bond;  Chapter 1  General Introduction  4  1.1.1 Traditional classification systems A number of methods have been devised to classify glycosidases.  A simple  system derived by the International Union of Biochemistry and Molecular Biology (IUBMB) classifies enzymes primarily according to the reaction they catalyze, and this, together with the name(s) of the substrate(s) provides a basis for classification of individual enzymes. Nevertheless, IUBMB rules are of limited value in that they do not provide any detailed information on the reaction mechanism. Several other classification systems exist, based on three criteria: glycone specificity, the anomeric configuration of the glycosidic linkage that is formed or broken, and the stereochemical outcome of the reaction. •  Regarding the glycone specificity, the classification is dependent on the sugar moiety for which the glycosidase is most reactive. For instance, glucosidases would be specific for a glucose moiety as the glycone while galactosidases would prefer a galactose unit as the glycone.  •  Glycosidases may also be classified as either "a" or "P" according to the absolute specificity for the anomeric configuration that is broken in the substrate. For instance, a P-glycosidase will only cleave P-glycosides and not a-glycosides.  •  Glycosidases are classified as "inverting" or "retaining" depending on whether the enzyme cleaves the glycosidic bond with net retention or inversion of anomeric configuration (Figure 1.2). For example, a retaining P-glycosidase cleaves a P-glycosidic linkage to yield a hemiacetal with the P-configuration,  Chapter 1  General Introduction  while an inverting P-glycosidase cleaves a P-glycosidic linkage to generate an a-hemiacetal.  Glycosidic linkage Inverting Glycosidase ~  OH  7  ~  HO HO  S  Retaining Glycosidase  OH Q  ROH H 0  X ' ^R  OH  2  f  \  H 0  ROH  2  HO  OH Q  OH  OH  Figure 1.2: Reactions catalyzed by retaining and inverting B-glucosidases.  •  Furthermore, the terms "exo" and "era/o" are used to categorize glycosidases that act on polymeric substrates such as cellulose and xylan. Enzymes that cleave terminal sugar residues from either end of the oligosaccharide chain, typically releasing mono- or disaccharides, are known as exro-glycosidases. Other enzymes that cleave at sites in the middle of the oligosaccharide chain are known as emfo-glycosidases. This distinction serves to indicate substrate specificity of a glycosidase and has been found to correlate with the tertiary structure of certain enzymes.  EXO  ENDO  Figure 1.3: A schematic depiction of oligosaccharide by glycosidases.  endo  EXO  and  exo  sites of cleavage of an  Chapter 1  General Introduction  6  1.1.2 Subsite nomenclature Glycosidases have evolved to bind the saccharide residues of oligo- and polysaccharides in distinct subsites.  The expansive  growth of available three-  dimensional structures has led to the identification of these binding subsites and consequently necessitated a more reliable system of nomenclature. Under a system put forth by Davies et al. (2), the position of the subsite relative to the point of cleavage is assigned by labelling subsites from -n to +n (where n is an integer). Subsites are labelled such that the —n represents the non-reducing end and +n the reducing end, with cleavage taking place between the -1 and +1 subsites.  Glucoamylase, for example, cleaves a  monosaccharide from the non-reducing end of the oligosaccharide chain (Figure 1.4A). Alternatively,  T. reesi  cellobiohydrolase I cleaves disaccharide units from the reducing  end of the substrate (Figure 1.4B).  A  B  NR  NR  R  R  Figure 1.4: Schematic drawing of sugar-binding subsites in two glycosidases. By convention, the non-reducing end of the substrate is drawn on the left while the reducing end is drawn on the right. The arrow placed between the -1 and +1 subsites indicates the point of cleavage.  Chapter  I  General Introduction  7  1.1.3 Glycosidase families  In addition to traditional criteria for classification, glycosidases have been grouped into distinct families by Henrissat based on amino acid sequence similarities and tertiary structure as evaluated through hydrophobic cluster analysis and other methods of sequence comparison (3-5). A recent compilation of carbohydrate-transforming enzymes is available from the continuously updated carbohydrate-active enzyme database (CAZy) at http://www.cazy.org/. To date, the CAZy classification lists ~100 sequence-derived glycoside hydrolase families and there are three-dimensional structural representatives for over 60 of these (6). The similarity of the tertiary structures and conserved chemical mechanisms amongst enzymes within the same family has long established the discriminatory power of this classification method.  Although more information is  required before one is able to infer structural information and reaction mechanism based solely on sequence, experimental results have confirmed the identity of key catalytic residues by extrapolation to all members of a particular family. Furthermore, enzymes belonging to different families have been grouped into different clans based on similar tertiary structure and location of catalytic residues, despite low sequence similarity. Therefore, useful insights into the evolution of these enzymes may be obtained by grouping into clans and families.  1.2 Catalytic Mechanism of Retaining |3-Glycosidases Glycosidases perform catalysis via two distinct mechanisms (7), originally proposed by Koshland (8), resulting in inversion or retention of anomeric configuration (Figure 1.5). In the case of inverting glycosidases, hydrolysis of the glycosidic bond  Chapter I  General Introduction  8  occurs in a single step with inversion of anomeric configuration through the general basecatalyzed attack of water at the sugar anomeric centre, with general acid-catalyzed departure of the aglycone.  OH  Figure 1.5: glycosidases.  The  hydrolysis reactions catalyzed by retaining and  inverting  In contrast, retaining glycosidases catalyze the hydrolysis of the glycosidic bond with net retention of anomeric configuration.  The reaction follows a double-  displacement mechanism involving the formation and hydrolysis of a covalent glycosylenzyme intermediate via oxocarbenium ion-like transition states, as illustrated in Figure 1.6. Binding and stabilization of the transition-state complexes is achieved through noncovalent binding interactions between the substrate and a number of amino acids within the active site of the enzyme. Two key amino acid residues play particularly important roles:  the catalytic nucleophile and the acid/base catalyst.  The acid/base catalyst  facilitates bond cleavage by donating a proton to the glycosidic oxygen during the formation of the glycosyl-enzyme intermediate.  The glycosylation step involves the  development of a covalent bond between the carboxyl group of the catalytic nucleophile  Chapter 1  General Introduction  and the anomeric carbon of the sugar substrate, while the deglycosylation step involves the hydrolysis of the glycosyl-enzyme intermediate. In this second step, the acid/base catalyst deprotonates the catalytic water which displaces the catalytic nucleophile resulting in a sugar product with retention of the original anomeric configuration. A crucial structural difference between retaining and inverting glycosidases is the difference in the distance separating their two key catalytic residues (9, 10). In inverting glycosidases, the general-base and the general-acid carboxyl groups are typically separated by an average distance of 7-11 A (O to O) in order to accommodate the nucleophilic water molecule, whereas in retaining glycosidases the nucleophile and acid/base catalyst are found approximately 5 A apart.  9  Chapter I  General Introduction  Figure 1.6: General double-displacement mechanism for a retaining glycosidase.  1_0  Chapter I  An  General Introduction  U_  alternative mechanism for (3-N-acetyl hexosaminidases that utilizes  acetamido-group participation during hydrolysis has been established wherein the 2-acetamido group of the substrate acts as an intramolecular nucleophile to displace the leaving group, forming a bicyclic oxazolinium-ion intermediate. Such a mechanism has been graphically demonstrated through determination of the structure of a family GH-20 hexosaminidase in complex with the inhibitor GlcNAc-thiazoline, which mimics the covalent oxazolinium intermediate of the substrate-assisted retentive hydrolysis (11). Another recent example of an enzyme utilizing substrate-assisted catalysis via acetamidogroup participation is that of family 56 hyaluronidase, a major allergen of bee venom (12).  The essential features of the general mechanism of action of glycosidases have been discussed in recent reviews  (6,  13-17).  A variety of experimental strategies  including kinetic isotope effects, crystal structure analyses of wild-type and mutated glycosidases and of their complexes with ligands, and structural and kinetic analyses of transition-state analogues have been employed to decipher the mechanistic details of catalysis. The key points that have been the focus of most of the mechanistic studies have been the nature of the intermediate and the transition state, the extent of nucleophilic participation, the role and nature of acid/base catalysis, and the role of substrate distortion in catalysis.  1.2.1  Oxocarbenium ion-like transition states  A key feature of the double-displacement mechanism for glycosidase-catalyzed cleavage is the structure of the oxocarbenium ion-like transition state through which  Chapter  1  General  Introduction  72  glycosylation and deglycosylation are postulated to proceed. The transition state occurs on the reaction coordinate somewhere between a distorted substrate and a species where the glycosidic bond is substantially elongated while the bond to the enzyme is hardly formed. A characteristic of bond cleavage in each step of the reaction includes the development of a positive charge on the C l and 05 atoms of the sugar ring in the -1 subsite. To distribute the positive charge across both C l and 05, the pyranose ring adopts a half-chair or boat conformation to accommodate an effective overlap of the lone pair of electrons on the endocyclic oxygen with the resulting vacant />orbital at the reaction centre. The majority (>75%) of the accumulating positive charge was shown to reside on the endocyclic oxygen of 2,4-dinitrophenyl (3-D-glycopyranosides during spontaneous hydrolysis (18).  The characteristics of the oxocarbenium ion-like transition  states for retaining glycosidases have been explored extensively through kinetic isotope effects and inhibition studies with transition-state analogues (19). Much of the evidence for positive charge development in the transition state has been derived from measurement of positive secondary a-deuterium kinetic isotope effects (k\\lkr)>  1). Through careful selection of the aglycone, thereby controlling rate-limiting  steps, these measurements have been reported for both the glycosylation and deglycosylation steps of the double-displacement mechanism. A normal kinetic isotope effect  (kwlkv  > 1) is associated with a change in the hybridization state of the anomeric  centre from sp to sp as the substrate goes from the ground state to the transition state, which is consistent with transition states for the formation and hydrolysis of the intermediate having significant oxocarbenium ion character. kinetic isotope effect  (knfko<\)  Conversely, an inverse  would be expected for a hypothetical ion-pair  Chapter I  General Introduction  I_3  intermediate in cases where the second step of the reaction, which would necessarily involve a change in hybridization from sp in the ion pair to sp in the product, is rate 2  3  determining. Nevertheless, in every known case, kinetic isotope effect measurements determined for the glycosidase-catalyzed hydrolysis of the substrates have been normal (kn/kD> 1) regardless of which step is rate determining, consistent with oxocarbenium ion character at both transition states. For example, kinetic isotope effects measured on the glycosylation transition state include values of h^lko =1.15-1.20 for Escherichia coli (lacZ) (3-galactosidase (20), k /k = 1.05-1.07 for Agrobacterium sp. (3-glucosidase (21), u  D  and ku/kr)= 1.06-1.12 for Cellulomonas fimi xylanase (22). Similarly, kinetic isotope effects for the deglycosylation transition state have been measured and include values of ku/ku = 1.20-1.25 for E. coli P-galactosidase (23) and  AT /A:D = H  1.10-1.12 for C. fimi  xylanase (22). Further evidence for the existence of the oxocarbenium ion-like transition states is obtained from studies with transition-state  analogue inhibitors.  Transition-state  analogues that closely resemble the proposed structure of the transition state will bind tighter to the active site of glycosidases than do the normal glycoside substrates since they mimic the transition state. The characteristic features proposed for an oxocarbenium ion-like transition state include a positive charge at the positions corresponding to either 05 or C l , an sp -hybridized centre at the position corresponding to C l and a conformation where the atoms corresponding to C5, 05, C l and C2 are coplanar, typically a half-chair. Evidence for the existence of oxocarbenium ion-like transition  Chapter 1  General Introduction  L4  states may be inferred through the tight binding by the enzyme to analogues that possess some or all of the characteristics discussed above. Physical organic studies have shown that although glycosyl oxocarbenium ions appear to have a finite lifetime in aqueous solution, they cannot exist in the presence of a closely located nucleophile, particularly when that nucleophile is anionic (24). The corresponding transition states of the glycosidase-catalyzed hydrolysis mechanism can therefore be expected to exhibit oxocarbenium ion  character,  with the nucleophile and  leaving group maintaining partial covalency to the anomeric carbon.  1.2.2  Nature of the covalent glycosyl-enzyme intermediate  A second feature of the nucleophilic substitution reaction characterizing glycoside cleavage is the presence of a covalent glycosyl-enzyme intermediate, as opposed to a stabilized oxocarbenium ion-pair intermediate. As alluded to earlier, strong evidence for the nature of the intermediate may be obtained from secondary a-deuterium kinetic isotope effect studies that provide insight regarding the changes in the geometry at the anomeric centre in the rate-limiting step of the reaction as it proceeds from the ground state to the transition state. A glycosyl-enzyme intermediate has more sp character than 3  the preceding and following transition states. A positive kinetic isotope effect on the deglycosylation step, whose value for k\\lkr> falls between 1.1 and 1.3, reflects significant 3  2  sp to sp rehybridization at the anomeric centre, consistent with the proposed geometry for the covalent glycosyl-enzyme intermediate. with a sp . 3  k /ko H  Conversely, an inverse isotope effect  of less than unity would be observed for a change in geometry from sp to 2  Chapter  I  75  General Introduction  Direct evidence for the existence of a covalent intermediate comes via the trapping of a relatively stable glycosyl-enzyme intermediate through a variety of different methods. One highly successful approach involves the use of mechanism-based inactivators such as the fluoro sugar inhibitors that function via the formation and accumulation of a relatively stable glycosyl-enzyme intermediate.  Several classes of  these mechanism-based inactivators have been developed to destabilize both transition states through the incorporation of a highly electron withdrawing fluorine substituent adjacent to the site of positive charge development, resulting in a decrease in the rate of both the formation and the hydrolysis of the intermediate.  Accumulation of the  intermediate is achieved by the incorporation of a good leaving group such as fluoride ion or 2,4-dinitrophenolate, which accelerates the glycosylation step of the reaction, but not deglycosylation (Figure 1.7A). The intermediate that forms breaks down very slowly, with typical half-lives for hydrolytic turnover measured in days. Reactivation of the enzyme occurs much faster by a transglycosylation reaction in the presence of a suitable glycosyl acceptor sugar according to Figure 1.7B.  Figure 1.7: Inactivation and reactivation by 2-deoxy-2-fluoro glycosides. Inactivation by formation of the fluoroglycosyl-enzyme intermediate. Reactivation by transglycosylation of the fluoroglycosyl-enzyme intermediate.  The first generation of such mechanism-based  (A) (B)  inhibitors, involving the  replacement of the 2-hydroxyl with a 2-fluorine substituent, were the 2-deoxy-2-fluoro(3-D-glycosyl fluorides (25) and the 2,4-dinitrophenyl 2-deoxy-2-fluoro-(5-D-glycosides (26). The removal of key hydrogen bonding interactions with the 2-position hydroxyl has the additional effect of destabilizing the transition states in the retaining P-glycosidase catalyzed hydrolysis of glycosides since interactions at this position have been shown to contribute up to 10 kcal mol to transition-state stabilization (27-29). -1  The fluorine group  therefore destabilizes the transition state both through its significant inductive effects and also by removal of important interactions at the 2-position. The development of alternative approaches was initiated due to the failure of the 2-fluoro sugar compounds to function as inactivators of a-retaining glycosidases, serving  Chapter I  General Introduction  17  rather as slow substrates for which the rate-determining step is formation of the glycosylenzyme intermediate. One approach to further slow the glycosylation step involved the installation of a second fluorine at C2 along with the simultaneous incorporation of a still better leaving group such as 2,4,6-trinitrophenol and resulted in the development of 2-deoxy-2,2-difluoro  glycosides  that  form  extremely  stable  glycosyl-enzyme  intermediates on a-retaining enzymes (30). A similar strategy that involves substitution of fluorine at C5 in place of the hydrogen was developed by McCarter and Withers (31). These 5-fluoro-glycosyl fluorides inactivate the glycosyl-enzyme intermediate of both aand P-retaining glycosidases by formation of a stabilized fluoroglycosyl-enzyme intermediate through a mechanism analogous to that of the 2-deoxy-2-fluoro glycosides. The three-dimensional structures of a growing number of these glycosyl-enzyme intermediates have been solved by single crystal X-ray diffraction. 2-fluorocellobiosyl-enzyme intermediate formed on the xylanase Cex from fimi  was the first such structure solved  Cellulomonas  (32).  N0  2-fluoro-p-D-glucosyl fluoride  The 2-deoxy-  2  TNP 2,2-difluoro glucoside  5-fluoro-a-D-glucosyl fluoride  Figure 1.8: Structures of several classes of the fluoro sugars that function as mechanism-based inactivators.  Chapter I  General Introduction  18  An alternate method of trapping the intermediate has been achieved by modification of key enzyme residues through mutation. covalent glycosyl-enzyme intermediate of a xylanase from  Using this approach, the C.  fimi  was the first to be  trapped without disrupting important interactions at the 2-position. The X-ray structure of this glycosylated enzyme once again confirmed that the intermediate is indeed covalent (33). Vocadlo et al. revealed that hen egg white lysozyme (HEWL) performs catalysis via the formation of a covalent intermediate in common with other retaining glycosidases (34).  The accumulation of a covalent 2-fluorochitobiosyl-enzyme  intermediate on the wild-type enzyme, as well as the nonfluorinated chitobiosyl-enzyme intermediate on the acid/base mutant, was demonstrated by mass spectrometry.  The  formation of a covalent intermediate was further confirmed by the X-ray crystallographic structural analysis of a trapped 2-fluorochitobiosyl-enzyme intermediate on an acid/base mutant of lysozyme. The mechanism of hen egg white lysozyme was unequivocally shown to proceed through a covalent glycosyl-enzyme intermediate and not a stabilized oxocarbenium ion pair as had long been held.  1.2.3 Nucleophilic participation  Retaining glycosidases perform nucleophilic catalysis with extraordinary ability. The identification of the essential nucleophilic residue involved in the doubledisplacement mechanism may be achieved through sequence alignments of enzymes within a family and identification of conserved amino acid residues as possible candidates to function in such a capacity. Site-directed mutagenesis studies followed by detailed kinetic analyses may lead to the correct assignment of catalytic residues.  Chapter I  General Introduction  [9  Essential residues in an enzyme may also be predicted using a high resolution, three-dimensional image obtained through X-ray crystallography.  Crystallographic  studies on retaining glycosidases have identified carboxyl residues appropriately located within the active site to function as nucleophiles. More substantial evidence for the correct assignment of a catalytic carboxyl group is derived from glycosidases cocrystallized with either an inhibitor or a substrate bound within the active site of the enzyme. Despite the fact that X-ray crystallography has provided many useful insights into the structures and functions of a diverse number of glycosidases, this technique does not unequivocally identify the important catalytic residues, and furthermore, is not always available for every enzyme. Although, group-specific labels have been used to identify catalytic groups in labelling experiments, a more popular and reliable approach to identifying and assigning the roles of the catalytic residues is the use of mechanism-based inactivators. Mechanism-based inactivators have been successfully applied in labelling experiments to identify the catalytic nucleophiles in the active sites of a number of retaining glycosidases.  A common approach involves proteolytic digestion of the  labelled enzyme to yield a mixture of short peptides, followed by separation of the resultant mixture by RP-HPLC and analysis by ESI-MS/MS (electrospray ionization tandem mass spectrometry).  The labelled peptide is characterized within the peptide  mixture by a tandem mass spectrometry method known as a neutral loss scan which detects those peptides that lose a characteristic reporter group by collision-induced fragmentation (35). Isolated, labelled peptides are subsequently sequenced by further MS/MS analysis, or by Edman degradation. This approach has been used successfully to  Chapter  I  General Introduction  :  20  identify the catalytic nucleophile in Bacillus subtilis xylanase as Glu78 (36) and human glucocerebrosidase as Glu340 (37). Leading references to this approach may be found in the following publications and reviews (26, 38-41). Mutagenesis studies involving replacement of specific side chain residues to function as alternative nucleophiles have led to insight into mechanistic details of catalysis. For instance, the replacement of specific side chain residues can result in a change of enzymatic mechanism.  An example of an inverting glycosidase being  converted into a retaining enzyme is that of the inverting T4 phage lysozyme (42). In the wild-type inverting enzyme, the hydroxyl side chain of Thr26 hydrogen bonds to the attacking water molecule, which is deprotonated by the neighbouring Asp20 during hydrolysis. The crystal structure shows that mutating Thr26 to a histidine residue results in a mutant that occupies the position normally assumed by the water molecule. It has been rationalized that the imidazole group acts as a nucleophile to form a covalent glycosyl-imidazolium-ion intermediate that is subsequently hydrolysed to form a product of retained configuration.  Conversely, converting a retaining fi-glycosidase to an  inverting enzyme by shortening a nucleophilic glutamate by substitution with aspartate has not been achieved (43, 44). Rather, these enzymes function as retaining enzymes, albeit at reduced rates relative to the wild-type enzymes. Replacement of the glutamic acid or aspartic acid by alanine, glycine or serine virtually eliminates all nucleophilic activity. Addition of small, external anions such as azide, which do not require general base catalytic assistance, can result in cleavage of the glycosidic bond of activated substrates such as 2,4-dinitrophenyl P-glycosides with inversion of configuration, yielding the a-glycosyl azide product. Rate accelerations upon addition of azide (2 M)  Chapter I  General Introduction  2l_  have been observed, with the catalytic rate constant (k ) rising from 1 x 10~ s to 1 s 5  H  _1  cat  for the Glu358Ala mutant of Agrobacterium sp. (3-glucosidase (45). In addition to formate and azide, halides have been shown to act as effective substitute nucleophiles. Indeed, the serine, glycine and alanine nucleophile mutants of Agrobacterium sp. P-glucosidase and Cellulomonas fimi P-mannosidase (46) were effective in catalyzing nucleophilic fluorination of 2,4-dinitrophenyl P-glycosides to form the corresponding oc-glycosyl fluorides (Figure 1.9, first step). The glycosyl fluoride, in turn, is used to glycosylate a second equivalent of substrate in a transglycosylation reaction, thereby forming glycosidic bonds (Figure 1.9, second step).  Glu519Ser  Glu519Ser  Glu519Ser  Figure 1.9: Nucleophilic fluorination of 2,4-dinitrophenyl B-mannoside by the P-mannosidase nucleophile mutant (Glu519Ser) from Cellulomonas fimi.  One application of mechanistic studies has led to the development of synthetically useful mutants termed glycosynthases (47). Mutation of the catalytic nucleophile of retaining glycosidases results in enzymes that catalyze the synthesis of oligosaccharides by using glycosyl fluorides as glycosyl donors (48-50).  Chapter  1  General Introduction  22  1.2.4 Acid/base catalysis  In Koshland's mechanism for inverting and retaining glycosidases, the hydrolysis of the glycosidic linkage proceeds with general acid/base assistance. As stated earlier, in the glycosylation step for retaining glycosidases, an enzymic carboxyl group acts as an acid catalyst to facilitate the departure of the aglycone. Indeed, X-ray crystal structures of several glycosidases reveal carboxyl group residues in one of two positions that could function as proton donor groups. Crystallographic analysis of glycosidases in complex with ligands has revealed two protonation trajectories, termed either anti or syn by Vasella and coworkers (19). The definition of the protonation trajectory is based on the orientation of the lone pair on the glycosidic oxygen with respect to the catalytic acid. The protonation trajectory is defined as anti if the catalytic acid is opposite to the endocyclic oxygen on the side of a plane formed from C l , 01, and HI. When the catalytic acid is on the same side of this plane as the endocyclic oxygen, then the protonation trajectory is defined as syn. The large discrepancy in the degree of inhibition by competitive triazole inhibitors with enzymes from different families of glycosidases has been explained to result from these two differences in the protonation trajectory (Figure 1.10). Inhibition kinetics and crystallographic analysis of the family 10 xylanase from Cellulomonasfimi(Cex) in complex with a xy/06/0-imidazole inhibitor and the xylobio-lactam oxime substantiates a«//-protonation.  The family 11 xylanase from  Bacillus circulans (Bex) interacted only weakly with these inhibitors.  The weak  inhibition was correlated with the proposed syw-protonation by family 11 glycosidases (51, 52). Predictably, because there is significant resemblance of the active site within glycosidase families, members within one family may thus be categorised as anti-  Chapter 1  General Introduction  23  protonators (GH-families 1, 2, 5, 10, 18 and 20) or .syra-protonators (GH-families 7, 11, 12, 16, 22, 23 and 45).  ?H^OH N HO  HO  HO  OH  A  ENZ  B  ENZ  ENZ  c  Figure 1.10. The anti orientation of the acid catalytic residue and protonation trajectory as the basis for rationalizing the relative potency of two triazole inhibitors. (A) /t«//'-protonation of the glycosidic oxygen. (B) Absence of any possible hydrogen bonding to the 1,2,3-triazole inhibitor. (C) Possible antiprotonation of the 1,3,4-triazole inhibitor.  The role of the acid/base catalyst has been probed extensively in several enzymes through mutagenesis studies. In the absence of a crystal structure, highly conserved carboxyl group residues can be identified using sequence alignment data, followed by site-directed mutagenesis and kinetic studies. Using this approach, Glul70 was identified as the acid/base catalyst in  Agrobacterium  sp. P-glucosidase  (53).  Detailed kinetic  analysis using substrates having poor leaving groups compared with substrates not requiring acid assistance may be performed on enzymes that have had the proposed residue mutated to confirm its role as the acid/base catalyst. For substrates having poor leaving groups that require efficient acid catalysis, deletion of the carboxyl group of the general acid/base catalytic residue results in a significant decrease in the rate of the glycosylation step of the hydrolysis reaction, which is reflected in a reduction in the second-order rate constant  (k /K ). cat  m  Conversely, for activated substrates that do not  require acid catalysis, the formation of the glycosyl-enzyme intermediate will occur at  Chapter 1  General Introduction  24  rates close to that of the wild-type enzyme. Accumulation of the glycosyl-enzyme intermediate is the net result observed when these acid/base mutants hydrolyse reactive substrates because the deglycosylation step of the double-displacement mechanism is also slowed upon deletion of the general aciaVbase catalytic residue. This accumulation of the intermediate is reflected in the kinetic analysis by an initial "burst" of the activated leaving group and an extremely low K value. m  Mutagenesis studies, where the position of the catalytic residue is changed in the active site, have demonstrated the flexibility of acid/base catalysis in glycosidases. This has been demonstrated in the xylanase from Bacillus circulans by changing the length of the acid/base catalyst (54), while incorporation of an appropriately positioned acid catalyst into the substrate was able to restore the activity of the acid/base mutant of Agrobacterium sp. P-glucosidase (Glul70Gly) (55). Additionally, oxidation of a mutant of glucoamylase in which the general base had been replaced by cysteine to convert it to cysteinesulfinic acid, and observation of activity demonstrates that acid/base catalysts are not limited to carboxyl groups (56, 57). The first step in the hydrolysis mechanism for retaining glycosidases, glycosylation, benefits from acid catalysis. Although the pK of a typical carboxylic acid a  is 4.5, the pK of the acid catalyst within the active site of many glycosidases is perturbed a  upward to a value between 6 and 8. This shift in pK is presumed to arise from an a  electrostatic field exerted by the nucleophilic carboxyl group that is held in close proximity to the acid/base residue.  A pK value above that of the medium pH is a  necessary to maintain a favourable protonation state of the general acid/base catalytic residue. The deglycosylation step of the hydrolysis reaction similarly benefits from base  Chapter I  General Introduction  25  catalysis. The deprotonated residue that formerly acted as a general acid catalyst is now poised to act as a general base catalyst in the deglycosylation step of the reaction. In fact, the general acid/base catalytic residue has been observed to be hydrogen bonded to a well-ordered water molecule positioned near the anomeric centre in the crystal structures of several glycosidases in which the glycosyl-enzyme intermediate has been trapped. The same residue that previously functioned as a general acid with a pK of 6-8 now a  fulfills that role of general base due to a drop in pK resulting from electrostatic field a  effects.  An electrostatic basis for pK cycling of the acid/base residue has been a  extensively studied in Bacillus circulans xylanase (Bex) through a series of elegant experiments that involved direct  C-NMR titrations of the enzyme that had been  C  labelled in the carboxyl group side chains of both the acid/base and the nucleophilic residues. The pK of the catalytic acid/base residue was found to be 6.7 in the free a  enzyme. Upon formation of a stable xylobiosyl-enzyme intermediate, thus neutralizing the charge on the nucleophile, the pK of the acid/base catalytic residue dropped by 2.5 a  units to 4.2 in order to provide the ionization state required to act as a general base. Furthermore, the effect of hydrogen bonding interactions on the acid/base catalyst of Bacillus circulans xylanase (Bex) has also been extensively studied through a series of elegant experiments. In the wild-type enzyme, Asn35 forms a hydrogen bond (3.1 A) to the acid/base catalyst Glul72.  This hydrogen bond is maintained at 3.2 A by the  mutation Asn35Asp, but the pH optimum is shifted from 5.7 to 4.6 while increasing the activity by 20% (see Figure 1.11). The pH dependence of k /K cat  m  for the Asn35Asp  mutant followed pK values of 3.5 and 5.8. These pK values were determined by a  13  a  C-NMR titrations on the enzyme to arise from Asp35 (pK = 3.7) and the nucleophile a  Chapter 1  General Introduction  26  Glu78 (pK = 5.7), but not the acid/base catalyst Glul72 (pK = 8.4). These pK values a  a  a  indicate that only 1% of the enzyme was in the correct ionization state. Thus, the active form of the Asn35Asp mutant was inherently 100 times more active than the wild-type enzyme. The increase in activity has been attributed to the Asn35Asp-Glul72 hydrogen bond.  Crystallographic studies on the Asn35Asp mutant trapped as the 2-fluoro-  xylobiosyl covalent intermediate revealed that this hydrogen bond had shortened substantially to 2.7 A in the intermediate. Likewise, C-NMR titrations on the trapped 13  intermediate determined a primary ionization pK \ of 2.6 for the Asn35-Aspl72 pair, a  indicating that the pK of Glul72 had effectively dropped 5.8 units upon formation of the a  intermediate, corresponding to an energy difference of ~8kcalmoF . 1  This is  dramatically greater than the pK drop found in the wild-type enzyme, in which the pK a  a  of Glul72 shifts only 2.5 units (3.5 kcal moF ) upon forming the covalent intermediate. 1  Therefore, the formation of a stronger hydrogen bond in the covalent intermediate drives the formation of that intermediate by making proton donation by Glul72 to the departing glycosidic oxygen increasingly more favourable.  Chapter 1  General  Introduction  27  Figure 1.11: Depiction of the pH dependence of k /K for Bex Asn35Asp mutant as the product of titration curves of catalytic residues derived from C-NMR. Although only 1% of the mutant is in the correct ionization state for catalysis, it is 2 0 % more active than the wild-type enzyme. cat  m  13  Chapter I  General Introduction  ;  28  Figure 1.12: The Asn35Asp mutant of Bex increases the inherent catalytic efficiency by at least 100-fold over the wild-type enzyme by forming a strong hydrogen bond between Asp35 and Glul72.  1.2.5  Non-covalent interactions  Non-covalent interactions between enzyme and substrate have been proposed to A-  account for the-majority of the catalytic power of enzymes by Pauling. The binding energy derived from these interactions leads to a decrease in the activation energy of the reaction by stabilizing the geometry and charge of the transition state preferentially over that of the ground state, thereby resulting in rate acceleration. X-ray crystal structures of  Chapter 1  General Introduction  29  carbohydrate-binding proteins and glycosidases have identified interactions between the hydroxyl groups of the sugar ligand and the enzyme, most of which represent hydrogen bonds. A systematic approach to explore the importance of individual non-covalent interactions has been achieved in which site-directed mutagenesis is carried out to remove the side chain residue on the protein that was interacting with the substrate. Another approach has been through the use of modified substrates in which each hydroxyl group has been replaced by a hydrogen or fluorine to correlate structure and activity. Kinetic parameters are then measured on the modified system in each case. Modified deoxy- and deoxyfluoro-glycoside substrates have been utilized to identify and quantify important non-covalent interactions in the catalytic mechanism of glycosidases.  For example, upon removal of the hydroxyl group of glucose at the  2-position and replacement by hydrogen, the rate of hydrolysis by Aspergillus wentii P-glucosidase drops by at least a factor of 10 , corresponding to >7 kcal mol 6  -1  of  stabilization (58). The hydroxyl group at position 4 is also important, with 4-deoxy substrates being hydrolysed 10 to 10 -fold slower relative to the unmodified substrates 4  5  (58). The importance of interactions at the 2-position is revealed through similar studies with E. coli p-galactosidase, in view of the fact that enzymatic hydrolysis of 2-deoxygalactosides is 10 -fold slower than is hydrolysis of the unmodified substrate (59). 5  Interactions at the 4- and 6-positions appear to be less important, with rate reductions of only 500 and 1000-fold being observed with 4-deoxy and 6-deoxy substrates (27). The interactions with the 3- and 6-hydroxyl groups during the chemical steps of the Agrobacterium sp. P-glucosidase catalyzed reaction each provide 2.2 kcal mol towards -1  stabilization of the transition state (28). Hydrogen-bonding interactions at the 2-position  Chapter 1  General Introduction  appear to be most important and provide 5 kcal mol  50  -1  worth of transition state  stabilization, with rate reductions of approximately 5000-fold being observed for the hydrolysis of 2-deoxy-glucosides (28).  Likewise, the rate of hydrolysis of the  2-deoxy-cellobiosyl-enzyme intermediate  is  10 -fold lower than that for 6  the  corresponding cellobiosyl-enzyme in C. fimi xylanase, revealing the importance of noncovalent interactions at position 2 for stabilization of the transition state (35). The formation of low-barrier hydrogen bonds, although controversial, has been proposed to account for significant rate accelerations (60). The formation of these specific, strong hydrogen bonds in enzyme transition states and intermediate complexes is effectively accomplished within the shielded environment of the enzyme active site.  1.2.6  Substrate distortion  Many aspects of the function and catalytic mechanism of glycosidases remain controversial despite the fact that the first X-ray structure of an enzyme was that of a glycosidase, hen egg white lysozyme (HEWL).  Hydrolysis by these enzymes is  performed via transition states that display considerable oxocarbenium ion-like character as the anomeric carbon becomes sp hybridized. Such hybridization requires a planar 2  arrangement of C5, 05, C l and C2 at or near the transition state (61). The resulting conformation for the transition state can only be accommodated by Hj, and # 4 half-chair 4  3  and ' B and Z?2,5 classical boat conformations (62). Support for the planar transition state has come from X-ray structural analysis of enzymes trapped in various states along the reaction coordinate that reveal distortion of the pyranosyl ring at the -1 subsite from its standard C\ conformation (63). This distortion was originally proposed in the late 1960s 4  Chapter 1  57  General Introduction  by Phillips and co-workers on the basis of the analysis of oligosaccharide complexes of HEWL (64), and later verified by the high-resolution structure determinations of a HEWL tetrasaccharide lactone derivative (65). Similar work on product and pseudo product complexes of HEWL (66, 67) and a mutant T4 lysozyme product complex (68) revealed distorted sugars.  The high-resolution crystal structures of "Michaelis"  complexes of two structurally unrelated e«Jo-glucanases from families GH-5 and GH-7 (69-72)  and a family GH-20 hexosaminidase  (73)  have been unambiguously observed.  For these P-retaining enzymes, the unhydrolysed substrates in the -1 subsite adopt a Sj l  skew-boat conformation (or the closely related E envelope conformation), while the 4  subsequent covalent intermediates have been observed as undistorted C\ chairs (33, 34, 4  74-77).  This implies a  '£3  Hs -> C\  4  4  pseudorotational itinerary for the glycosylation  step of the retaining mechanism (Figure 1.13).  Chapter 1  General Introduction  52  c  4  Figure 1.13: Partial map of pyranoside interconversions adapted from Stoddart (78). Potential transition-state conformations that possess a coplanar arrangement of C5, 05, C2 and C l atoms are shown boxed.  Other potential transition-state conformations have been observed. For instance, 25  evidence of a ' B boat conformation for the transition state has been observed for family GH-11 xylanases that display a covalent xylobiosyl-enzyme intermediate in a ' B 2 5  conformation  (79-81).  4  2  2  Ci -> HT,  Catalysis in this enzyme family presumably proceeds along a  25  So~^ ' B itinerary.  Furthermore, recent trapping of a family 26  P-mannanase in a S$ conformation and its subsequent covalent intermediate in a °S2 X  conformation suggest a different catalytic strategy via a #2,5 transition state (82) along a Ss  1  -> 7i 5 2i  enzyme.  -> °S2  pseudorotational itinerary for the glycosylation step for this class of  Similarly, the structures of three different covalent glycosyl-enzyme  intermediates for the family 38 Golgi a-mannosidase II from  Drosophila melanogaster  trapped by use of fluorinated sugar analogues revealed sugar intermediates bound in a  Chapter 1  General Introduction  33  distorted S$ skew boat conformation (83). Increasing evidence confirms that, despite l  sharing a common mechanism, glycosidases could utilize different transition state and intermediate structures due to substantial substrate distortion at the -1 subsite of the active site. Distortion of the -1 subsite sugar from the preferred ground state C\ chair A  conformation toward a boat or skew-boat form in which the glycosidic bond and leaving group become axial has been proposed to be driven by the extra interactions that develop within the +1 subsite. This distortion confers numerous catalytic benefits to the enzyme both in allowing a direct 'in-line' attack on the sugar anomeric centre by the nucleophile, concomitant with leaving group departure, and in removing potential repulsive interactions upon distortion to a boat or skew-boat form. As the substrate moves closer to the conformation of the oxocarbenium ion transition state, the interaction between a pseudoequatorial 2-hydroxyl substituent and the carbonyl group of the catalytic nucleophile, which in some cases is believed to contribute in excess of lOkcalmor  1  (28), is enhanced. Additionally, the pseudoaxial bond orientation is consistent with the dictates of stereoelectronic theory (84), which require an antiperiplanar arrangement of the sp lone pair electrons on the ring oxygen and the glycosidic bond in order to 3  facilitate electron donation to the transition state, and thus enhance the rate of hydrolysis (85). The antiperiplanar lone pair hypothesis (ALPH) requires a conformational change of the pyranosyl ring from a chair to a skew-boat or boat resulting in a pseudoaxial orientation of the aglycone (Figure 1.14).  Chapter I  General Introduction  A  B  C  34_  D  Figure 1.14: Orientation of the lone pairs on the endocyclic oxygen relative to the glycosidic oxygen of the leaving group. (A) The C\ ground state conformation of many common glycosides and covalent intermediate structures. This conformation features a lone pair on the ring oxygen that is antiperiplanar to the C l - O R bond in an a-configured glycoside. (B) The H conformation proposed as a possible transition-state conformation. (C) The B or E conformation and (D) the skewboat S conformation that features a lone pair on the ring oxygen that is antiperiplanar to the C l - O R bond in a (3-configured glycoside, as required for the optimal effect of the antiperiplanar lone pair hypothesis (ALPH). 4  4  3  4,i  4  l  3  Analysis of pyranoside ring conformations along the reaction pathway opens the capacity for the selective design of transition-state analogue inhibitors, which are highly valuable as mechanistic probes and potential therapeutic agents.  1.3 Biodegradation of Cellulose and Xylan Cellulose and xylan, which are linear polymers of (3-1,4-linked glucopyranosyl and xylopyranosyl residues respectively, are the most abundant polysaccharides on earth and are the major components of plant cell walls. Biodegradation of these p-l,4-glycans is achieved through the action of P-l,4-glycosidases or |3-l,4-xylosidases, hydrolytic enzymes which act cooperatively to convert these largely insoluble polymeric substrates into their simple constituent sugars. The depolymerization of cellulose occurs in a synergistic manner by the action of endo- and exo-glucanases. .EWo-glucanases cleave the internal P-1,4 bonds of cellulose, thereby increasing the number of free non-reducing  Chapter I  General Introduction  15  ends, which are sites for hydrolysis by exo-glucanases. The resulting product of cellulose degradation, cellobiose, is then hydrolysed to glucose by p-glucosidases. Xylan degradation involves similar xylanolytic enzymes for its complete hydrolysis that include e«Jo-l,4-xylanases (EC 3.2.1.8) and P-xylosidases (EC 3.2.1.37). Xylan is a heteropolysaccharide containing substituent groups of acetyl, 4-0-methylD-glucuronosyl and a-arabinofuranosyl  residues linked to the xylan backbone.  £W0-xylanases depolymerise xylan by the random hydrolysis of the xylan backbone and P-D-xylosidases hydrolyse small oligosaccharides. The side groups present in xylan are liberated by a-L-arabinofuranosidase, oc-D-glucuronidase, and acetyl xylan esterase, among others. A variety of microorganisms, mainly bacteria and fungi, along with plants are reported to produce xylanases (86-88). Many of these enzymes have a modular organization and comprise both catalytic and non-catalytic domains. On the basis of amino acid sequence alignment of their catalytic domains, they may be assigned to a specific family of P-glycosidases that displays similar catalytic mechanism and specificity. Most known xylanases belong to GH families 10 and 11 (over 300 gene sequences are known), and about 20 more xylanase genes are distributed between families 5, 8 and 43. Xylanases are fast becoming a major group of industrial enzymes (89), finding significant application in the biobleaching of wood pulp (86, 87, 90, 91). However, the use of xylanases is not restricted to the paper and pulp industry. Potential applications of xylanases in biotechnology include treating animal feed to increase digestibility (92, 93),  Chapter I  General Introduction  36_  clarification of juices and wines (94) and bioconversion of agricultural wastes to fermentation products (95).  The xylanolytic enzymes also have an application in rye  baking where they are used as dough strengtheners and result in an increased volume of baked bread (96-99). Processes used to produce ethanol as an alternative source of energy from biomass are an intensive area of research owing to the high cost and inevitable depletion of the world's petroleum supplies (100, 101).  There are reports  regarding the production of ethanol from hemicellulose wastes by incorporating xylanase treatment (95, 102, 103). Future work on the application of these enzymes in the biotechnology industry requires a complete understanding of the structure/function relationships obtained from biochemical and structural studies.  A n extensively studied family 10 enzyme is the  P-l,4-ewfo-glycanase from the soil bacterium Cellulomonas fimi. Family 10 members function primarily as 1,4-xylanases, although activity against cellobiosides has been reported. Members of GH-11 function exclusively as xylanases, of which the xylanase from Bacillus circulans is a representative enzyme.  1.3.1  Cellulomonas fimi xylanase The P-l,4-glycanase from the soil bacterium Cellulomonas fimi is an extensively  studied enzyme.  The gene encoding the glycanase has been cloned, expressed into E.  coli, and subsequently sequenced. The enzyme is glycosylated when produced in C. fimi, whereas the recombinant form expressed in E. coli is not.  The main function of  glycosylation appears to be protection against proteolysis, and has no apparent effect on catalytic activity. Also known as C/XynlOA, the glycanase enzyme is a 47 kDa protein  Chapter  I  General  Introduction  37  comprising an N-terminal catalytic domain (35 kDa) and a C-terminal cellulose-binding domain (12 kDa) connected by a proline and threonine-rich linker peptide (104). When separated by limited proteolysis, the domains of Cex retain their respective catalytic and cellulose binding properties. The catalytic domain (Cex-cd) is a member of family 10 of the glycoside hydrolases (105, 106).  Figure 1.15: Three-dimensional structure of Cex with bound 2F-X (76). The overall structure of Cex is depicted as a protein cartoon in divergent (wall-eyed) stereo, with a-helices coloured green, B-strands blue and loops red. The bound 2F-X is drawn as a stick model, with atoms coloured according to atom type, carbon in grey, oxygen in red, and fluorine in yellow. The water molecules in the active site have been removed for clarity. This figure and subsequent molecular representations were prepared with Chem3D Ultra 10.0 (http://www.cambridgesoft.com). 2  2  Cex is catalytically active on cellulose, xylan, and a range of soluble aryl P-D-glycosides (22, 106). Stereochemical studies using 'fi-NMR revealed a "retaining" enzyme that utilizes a double-displacement mechanism involving the formation and hydrolysis of a covalent glycosyl-enzyme intermediate via oxocarbenium ion-like  Chapter I  General Introduction  transition states during the hydrolysis of substrates  38  (107).  The catalytic nucleophile is  Glu233, identified by trapping of a covalent 2-deoxy-2-fluoro-glycosyl-enzyme intermediate and sequencing of the purified glycopeptide (35, 108). The assignment of the nucleophilic residue was later confirmed by mutagenesis followed by kinetic analysis (22). The acid/base catalyst was identified as Glul27 by detailed kinetic analysis of sitedirected mutants (105, 109). Insight into the kinetic mechanism was obtained though a detailed kinetic study which allowed the identification of rate-limiting steps, as well as investigation of the transition-state structure for each step. Support for the double-displacement mechanism of Cex was derived from pre-steady and steady state kinetics, Bronsted relationships, kinetic isotope effect measurements, inactivation experiments, and pH studies (22, 33, 35, 76, 105, 108-111).  The X-ray crystal structure of the catalytic domain of Cex (112) revealed an eightstranded parallel a/p-barrel, with an open cleft at the carboxyl-terminal end, proposed to be the active site (Figure 1.15). The two catalytic residues, Glu233 and Glul27, are separated by a distance of 5.5 A on either side of the cleft, consistent with a retaining mechanism as is presumably optimal for the efficient formation of a glycosyl-enzyme intermediate by Glu233 and the concomitant protonation of the aglycone by Glul27. Highly conserved residues within family 10 enzymes were identified that form the environments around the key catalytic carboxyl groups and likely, therefore, play important roles in catalysis. For example, His205 was hydrogen bonded to both the nucleophile, Glu233, and to Asp235.  Chapter I  General Introduction  59  Additional insights into the interactions at the active sight that are fundamental to the catalytic mechanism and substrate specificity may be obtained from crystallographic analysis of the structure of a glycosyl-enzyme intermediate complex. The formation of a covalent 2-deoxy-2-fluorocellobiosyl-enzyme intermediate was achieved by the diffusion of the mechanism-based inactivator 2,4-dinitrophenyl 2-deoxy-2-fluorocellobioside into the crystals of the catalytic domain of Cex (Cex-cd), followed by the determination of the structure of this glycosyl-enzyme complex by X-ray crystallography (32). No significant structural change was observed within the protein upon formation of the glycosylenzyme. The structure of the seemingly rigid cavity identified the amino acid residues involved in binding and catalysis.  Amino acid side chains interacting with the  6-hydroxyl, and thus likely responsible for determining the cellulose/xylan specificity, were revealed in the structure (Figure 1.16). Subsequent determination of the structure of the 2-deoxy-2-fluoroxylobiosyl-enzyme revealed that two of the residues that had moved position slightly in the fluorocellobiosyl-enzyme complex no longer moved, consistent with the higher specificity for xylan (76). However, when mutations at one of these positions were made there was no significant effect upon the cellulose/xylan specificity.  Figure 1.16: Active site of the Cellulomonasfimixylanase trapped as a covalent 2-deoxy-2-fluorocellobiosyl-enzyme intermediate. Adapted from (32).  Insights into the source of the very strong interactions at the 2-position were provided by the three-dimensional structure of stable intermediates formed from 2-deoxy2-fluoro glycosides (32, 76). The highly conserved Asnl25 residue and the carbonyl oxygen of the nucleophile itself were observed to be closest to the fluorine of the substrate moiety. Interactions at the 2-position are extremely important to transition-state stabilization of a number of glycosidases (28), contributing at least 10 kcal mol in the -1  case of Cex (32). Subsequent mutational studies have shown that interactions with the amide moiety of Asnl26 contribute only approximately 2.5 kcal mol to the stability of -1  Chapter I  General Introduction  £/  the transition state. Therefore, the most likely source of the remainder of this large interaction energy seems to be with the nucleophile. However, since the 2-position is substituted by fluorine in these trapped complexes, the interactions involved at this position have been impossible to unequivocally identify. To directly address the nature of the interactions between the enzyme and the 2-hydroxyl of the proximal cellobiosyl moiety, the catalytic domain of the Cex double mutant His205Asn/Glul27Ala was trapped with 2,4-dinitrophenyl cellobioside to form an unsubstituted cellobiosyl-enzyme complex and the crystal structure was subsequently solved to 1.8 A (33). The overall structure proved to be very similar to that of the wildtype enzyme in its 2-deoxy-2-fluorocellobiosyl-enzyme form, with only minor changes observed in the positioning of the nucleophile and in the hydrogen bonding patterns as a consequence of the mutations. The carbonyl oxygen of the nucleophile was involved in a short (2.37 A) hydrogen bond with the sugar 2-hydroxyl which was implicated to contribute >10 kcal moF to transition-state stabilization. Because the structure solved is 1  that of the reactive intermediate, presumably, this interaction is optimized at the transition state both geometrically, as the ring flattens, and electronically, as the positive charge develops at the anomeric centre, acidifying the 2-hydroxyl and thereby increasing its hydrogen bond donor capability, as shown in Figure 1.17.  Figure 1.17: Hydrogen bonding at the transition state.  1.3.2  Bacillus circulans xylanase  The xylanase from Bacillus circulans (Bex or BcXynMA)  is a 20.4 kDa protein  that is a member of the low molecular mass family 11 xylanases (113). Members of this family derive from both eukaryotic and bacterial species and share sequence identity varying from 40-90 %. This "alkaline" xylanase has been extensively characterized using a combination of structural, spectroscopic, and enzymatic techniques (36, 44, 54, 81, 114, 115). Previous studies have determined that Glu78 functions as the nucleophile while Glul72 functions as the general acid/base catalyst in a double-displacement mechanism (36, 81, 115). The pK values of all the carboxyl (775, 116) and imidazole (117) groups a  have been determined by NMR spectroscopy. Specifically, the pK values of Glu78 and a  Glul72 are 4.6 and 6.7, respectively, determined directly using C-NMR spectroscopy 13  and in close agreement with those determined from the bell-shaped pH-activity profile of this enzyme (115). The pH optima of family 11 xylanases range from 2 to 11 and are often correlated with the nature of the residue adjacent to the acid/base catalyst. An alkaline pH optimum is observed in xylanases where the amino acid hydrogen bonded to the catalytic acid/base  Chapter I  General Introduction  43_  is an asparagine residue, whereas an acid pH optimum is observed when this residue is aspartic acid. Substitution of Asn35 with Asp in Bex resulted in an increase in activity of approximately 20% accompanied by a reduction of the pH optimum from 5.7 to 4.6 (118). A  detailed study of this mutant demonstrated that this shift in the pH optimum,  together with the formation of a strong hydrogen bond between Asp35 and Glul72 in the glycosyl-enzyme intermediate, was the result of a reverse protonation mechanism, as discussed earlier.  Figure 1.18: Three-dimensional structure of Bex with bound 2F-X (81). The overall structure of Bex is depicted as a protein cartoon in divergent (wall-eyed) stereo, with a-helices coloured green, B-strands blue and loops red. The bound 2F-X is drawn as a stick model, with atoms coloured according to atom type, carbon in grey, oxygen in red, and fluorine in yellow. The water molecules in the active site have been removed for clarity. This figure was prepared with C It em 3D Ultra 10.0 (http://www.cambridgesoft.com). 2  2  Structures of the native enzyme, a non-covalent Michaelis complex, and the glycosyl-enzyme intermediate of Bex have been solved by X-ray crystallography (Figure 1.18) (81, 119). The structure of the active site of Bex revealed several highly conserved  Chapter 1  General Introduction  44  residues arranged in an intricate network of hydrogen bonds surrounding the two catalytically essential acidic residues, Glu78 and Glul72 {119). The structure of the 2-deoxy-2-fluoroxylobiosyl-enzyme intermediate formed on Bex revealed the sugar in the -1 subsite distorted into a ' B boat conformation (81), unlike the C i conformations 2 5  4  seen in other cases (Figure 1.19). This conformation places C5, 05, C l and C2 in a plane, as is required at the oxocarbenium ion-like transition state of the doubledisplacement mechanism. Analysis of the conformation of the intermediate also provides insight into specificity, as such a conformation is more readily accessible for xylan derivatives since sugars with bulky hydroxymethyl substituents at C5 would be forced axial resulting in steric clashes with the enzyme.  Figure 1.19: Active site of the Bacillus circulans xylanase trapped as a covalent 2-deoxy-2-fluoroxylobiosyl-enzyme intermediate.  Chapter 1  General Introduction  45_  A number of interesting observations are revealed by comparison of this structure of the glycosyl-enzyme intermediate and that of a non-covalent Michaelis complex (119).  Most notably, the hydroxyl group of conserved residue Tyr69 forms a bifurcated  hydrogen bond interaction with the endocyclic oxygen of the xylose residue in the -1 subsite and the oxygen of Glu78 in its ester linkage with the sugar (Figure 1.20). Indeed, mutation of Tyr69 to Phe leads to complete loss of enzyme activity (119), indicating a very important role for this hydroxyl group. It seems probable that a full hydrogen bonding interaction develops from Glu78 to Tyr69, while a stabilizing electrostatic or dipolar interaction develops between the endocyclic oxygen and the hydroxyl group of Tyr69 as the glycosidic bond in this intermediate cleaves, resulting in negative charge development on Glu78 and the generation of positive charge on the sugar ring oxygen at the transition state. Other interesting observations on this complex include the close approach of the fluorine at C2 to the carbonyl oxygen of the nucleophile. Additionally, the conserved residue Tyr80 is involved in a hydrogen bonding interaction with an ordered water molecule held in place for nucleophilic attack at the sugar anomeric centre through hydrogen bonding interaction with the acid/base catalyst Glul72 (Figure 1.20).  Chapter 1  General Introduction  46  Figure 1.20: The 2-deoxy-2-fluoroxylobiosyl-enzyme intermediate formed on the Bacillus circulans xylanase. Green dashes represent hydrogen bonds, the blue dash depicts a non-hydrogen bond interaction, and the R group represents the saccharide unit bound in the - 2 subsite of the active site. Adapted from Sidhu et al. (81).  1.4 Aims of this Thesis Numerous studies o f the glycanase Cex from Cellulomonas fimi have classified the enzyme as a dual xylanase/cellulase retaining (3-glycosidase. The overall objective o f this study is to contribute to a better understanding o f the catalytic mechanism o f the glycanase through a combination of studies.  Structural analysis of the catalytic domain of Cex reveals an intricate network o f amino acid residues that participate in the catalytic mechanism. Aside from the catalytic nucleophile and acid/base catalyst, the complex architecture o f the active site reveals amino acid residues that may be important in substrate recognition and specificity.  The  first part o f this thesis will consist o f a detailed kinetic analysis o f a number of active site mutants.  Mechanistic investigations o f these mutants may help clarify the roles o f the  Chapter 1  General Introduction  47_  corresponding residues. This will involve steady state kinetic analysis, linear free energy relationship studies, and pH dependence studies. Linear free energy relationship studies should give insight into the rate-determining step and provide information on the degree of negative charge accumulation on the phenolate oxygen at the glycosylation transition state. Furthermore, pH dependence studies may provide insights into the ionization state of the residues at the active site of this enzyme. Bex has been characterized extensively using a wide range of structural, spectroscopic, and enzymatic techniques. In this study, Bex will be further characterized using modified deoxy- and deoxyfluoro-substrates  to determine the individual  contributions of hydroxyl groups to enzyme catalysis. An evaluation of the transition-state mimicry of imino-sugar inhibitors may shed light on the structural details of the mechanism of action. Furthermore, rigorous criteria to establish the nature of enzyme inhibition through kinetic studies will aid in the development of therapeutic agents. In these combined biochemical studies, we hope to reveal the principles of catalysis and specificity for these enzymes.  CHAPTER 2  CHARACTERIZATION OF H I G H L Y C O N S E R V E D R E S I D U E S IN T H E A C T I V E S I T E O F CELLULOMONAS FIMI XYLANASE  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  49  2 Characterization of Highly Conserved Residues in the Active Site of Cellulomonas fimi Xylanase 2.1  Introduction The three-dimensional structure of the Cex catalytic domain provides a structural  basis for the clarification of the wealth of biochemical information that is currently available for the enzyme. Family 10 xylanases studied to date all consist of 8-fold parallel a/p barrels containing deep active site grooves, consistent with an endo-mode of hydrolysis. Members of this family cleave glycosidic bonds by a double-displacement mechanism involving formation and hydrolysis of a covalent glycosyl-enzyme intermediate via oxocarbenium ion-like transition states. During the glycosylation step of the reaction mechanism, a xylose residue at the -1 subsite becomes covalently linked to the enzyme's nucleophile, assisted by general acid-catalyzed protonation of the glycosidic oxygen as it departs. In the deglycosylation step, the covalent glycosylenzyme intermediate is hydrolyzed by an activated water molecule that is simultaneously deprotonated by the acid/base catalyst through a general base-catalyzed process. The location of the two key catalytic residues, the nucleophile and acid/base catalyst, is conserved in all family 10 enzymes  (120).  A series of studies have identified the  nucleophile and acid/base catalyst in the family 10 xylanase from Cex using a combination of mechanism-based inactivators, site-directed mutagenesis and detailed kinetic analysis (105, 108). Elucidation of the mechanism of action was achieved through identification of the rate-limiting steps, followed by an investigation of the transition-state structure for each step of the mechanism. This was derived from presteady and steady state kinetics, Bronsted relationships, kinetic isotope  effect  Chapter 2  Characterization of Hishlv Conserved Residues in C. fimi Xylanase  measurements, inactivation experiments, and pH studies. These data are compiled in Appendix B,  page 298.  The topology of the active sites of family 10 glycanases was investigated by Charnock  et  al. by testing the rate and pattern of xylooligosaccharide cleavage (121).  Data obtained showed that Cex contained three glycone binding sites and two aglycone binding sites.  The free energies of productive binding determined for a range of  oligosaccharides in the subsites of Cex showed that the -2 subsite plays an important role in substrate binding. Furthermore, productive complexes were generally only formed when xylotrioside and xylotetraoside substrates occupied the -2 and -1 subsites of the xylanase. Residues that have the potential to play an important role in ligand binding through hydrogen bonding with sugar hydroxyl groups or hydrophobic stacking interactions with the pyranose rings have been identified at the -1 and -2 subsites. The three-dimensional structures of several family 10 xylanases have revealed highly conserved residues on the surface of the active-site cleft  (112,  122-129).  In particular,  structural analysis of family 10 xylanases covalently linked to mechanism-based cellobiosyl and xylobiosyl inhibitors has facilitated the identification of active-site amino acids that play important roles in substrate binding and catalysis (Figure 2.1). Specifically, structural analysis of Cex shows that Glu43, Asn44, Lys47, His80, Gln87, Asnl26, and Asnl69 are part of an important hydrogen bond network involved in substrate recognition (32, 33, 76, 125). The high-resolution structures of the ligandenzyme complexes have also revealed amino acids that influence the positions and ionization states of the two key catalytic residues; for example, Asnl26 and Asnl69 are in the vicinity of the two catalytic residues.  These residues, based on amino acid  50  Chapter  2  Characterization  of Hishlv Conserved  Residues in C.  fimi  Xylanase  51  sequence alignments and structural analysis, have been a target for mutagenesis and kinetic analysis in order to gain insight into the reaction mechanism of glycosidases. In order to investigate the role of these active-site residues in the structure and function of Cex, mutant enzymes were produced at each position, namely, E43A, N44A, K47A, H80A, H80N, H80Q, N126A, and N169A.  -2  -1  +1  Figure 2.1: Scheme of protein-carbohydrate interactions in the active site region of 2F-xylobiosyl-Cex-cd complex observed by three-dimensional structural analysis (76). Hydrogen-bonding interactions between heteroatoms are in Angstroms. The distance between 2F and the carbonyl oxygen of Glu233 is 2.6 A (not shown).  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  The primary aim of this study is to investigate the roles of highly conserved residues in the -2 and -1 binding subsites of Cex. The biochemical properties of Cex and variants of Cex will be evaluated to achieve this objective through detailed steady state kinetic analyses and pH studies. The information derived from studies presented in this section will provide the basis for further work presented in the upcoming chapters of this thesis.  2.2 2.2.1  Results Generation of mutants of Cex  A linear amplification, site-directed mutagenesis method was used to produce E43A, N44A, K47A, H80A, H80N, and H80Q variants of Cex. The N126A and N169A mutants were produced by a similar procedure by Dr. Valerie Notenboom (University of Toronto), and generously provided for this study.  The procedure, based on the  QuikChange site-directed mutagenesis kit by Stratagene, utilized pUC12-l.lcex(PTlS) as the vector containing the 1.4 kb cex gene and two synthetic oligonucleotide primers (Figure 2.2). The complementary oligonucleotide primers were extended using Pwo DNA polymerase to generate a mutated plasmid containing staggered nicks. Following temperature cycling, the reaction mixture was treated with the Dpn I endonuclease which is specific for methylated and hemimethylated DNA, and was thereby used to select for mutated synthesized DNA by digesting the methylated, nonmutated parental DNA template.  The circular, nicked vector DNA was then transformed into X L 1-Blue  electrocompetent cells that repaired the nicks in the mutated plasmid. The plasmid DNA was isolated and subjected to restriction analysis to reveal positive clones, which were  52  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  then sequenced to verify the correct mutations. The recombinant pUC12-l.lcex(PTIS) was subsequently transformed into electrocompetent E. coli JM101 cells for expression of mutant protein.  Figure 2.2: Overview of the QuikChange  Site-Directed Mutagenesis Kit.  2.2.2 Production, purification, and physical characterization  The mutant proteins were produced and purified as described in Materials and Methods. No significant differences in behaviour were observed during purification. The mutant proteins eluted 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 at the same position (47 kDa) as the native enzyme with greater than 95% purity by inspection. Final protein yield ranged from 5 to 20mgL  _1  of liquid culture (Table 2.1). Mass  53  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  spectrometric analysis confirmed the molecular masses of the mutant proteins (Table 2.2). Table 2.1: Total yields of Cex mutants produced. "•i  1 n/\ me..  jy.^bVal'Yield .  : Cex E43A Cex N44A \ Cex K47A Cex H80A ! Cex H80N Cex H80Q  17 12 13 7.3 6.0 5.2  ; ! i  Table 2.2: Determination of molecular weight by mass spectrometry.  Enz\ me  ' *  Native Cex " 47124 ± 3 " Cex E43A 47064 ± 3 • Cex N44A 47082±3 Cex K47A 47070±3_ f Cex H80A " 47058 ± 3 Cex H80N """ 47100 ± 3 '. Cex H 8 0 Q 4 7 1 1 4 ± 3  2.2.3  0 ' 60 ± 5 42±5 54_±5 66 ± 5 ~" 24 ± 5 10 ± 5  0 58 43 57 66 23 9  Kinetic analysis: pH dependence of k IK cal  Michaelis-Menten  m  parameters for the hydrolysis of aryl cellobioside and  xylobioside substrates as a function of pH were measured to characterize the mutant  54  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  xylanases.  55  Substrates were chosen with high K values to enable k /K m  cat  to be  m  determined from a single experiment at a substrate concentration well below K . An m  initial concentration of about KJ5 was hydrolyzed by the enzymes to give a first-order increase in absorbance as the products were released with rate constant (k /K ) cat  m  x  [E] . 0  The pH dependence of k /K determined in this way agrees well with that determined cat  m  from Michaelis-Menten plots.  The method is experimentally easier, and more  reproducible and accurate (130) because it eliminates the need to correct for the variation of extinction coefficient of the substrate with pH, thereby reducing errors associated with the determination of substrate concentration. The parameter k /K was chosen to be cat  m  monitored, rather than k or K individually, in order to be able to interpret the pH cat  m  dependence in terms of ionization events related specifically to the unbound enzyme, since  k t/K ca  m  is the second-order rate constant for the reaction of free enzyme and  substrate. The first-order rate constant k reflects bound species including the enzymecat  substrate, -intermediate, and -product complexes. Since k /K reflects the events up to cat  m  and including the first irreversible step in the mechanism, its value will not be influenced by potential changes in the rate-determining step from glycosylation to deglycosylation or vice versa. Additionally, transglycosylation reactions may occur at elevated substrate concentrations making it experimentally difficult to extract K and k m  cat  values  individually. Conversely, a pseudo-first-order analysis of the reaction velocity under conditions of limited substrate accurately measures k /K . cat  m  The pH dependence plots for k /K were fit to a bell-shaped activity profile, caX  m  described by equation 2.1, to provide two pK values a  Table. 2.3.  (p.Kai  and pA^), as summarized in  Chapter 2  Characterization of Hishlv Conserved Residues in C. fimi Xylanase  56  (2.1) *.,+[H ])(^+[H ]) +  +  Table 2.3: pH Dependence of k JK for C.fimi xylanase and mutants. c  En/A me  Wild-Type E43A N44A K47A H80A H80N H80Q Q87M Q87Y N126A N169A  pA  3.9 3.7 3.9 3.9 4.6 4.6 4.6 4.0 3.9 4.4 4.0  pA,;  TJ  6.9 8.0 6.0 6.2 6.3 7,0 8.1 8.3 8.5 8.2  m  - Ooffai'tionsfr-.; • •  pll Optimum  5.8 5.3  S u b s t r a t c  2,4DNPC 2,4DNPC 2,4DNPC  4.9 5.4 5.5 5.8" 6.1 6.1 6.4 fir  2 5DNPX2 :  PNPX, PNPX PNPX 2,4DNPC 2,4DNPC 2.4DNPC 2,4DNPC 2  2  „  R a  „  3-5^8.5 3.5-8.5 3.5-8.5 3.5^.0 T5-73" 3.5-7.5 4.0-7.5 3.5-8.5 3.5-8.5 4.0-8.5 3.5-8.5  Graphical representation of this data is given in Appendix A, page 292. An error of ±0.1 pH units was estimated from a non-linear fit by GraFit 4.0 (131). a  2.2.4 Kinetic analysis: steady state kinetics  In order to investigate the role of highly conserved active-site residues in the mechanism of Cex, kinetic parameters for a range of substrates were determined for mutant enzymes of Cex; namely, E43A, N44A, K47A, H80A, H80N, H80Q, N126A, and N169A.  The kinetic parameters for these mutant enzymes may help reveal the  importance of these residues for the catalytic function of Cex.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  57  The initial kinetic characterization of the active-site mutants of Cex consisted of comparisons to the wild-type enzyme with respect to the fundamental steady state kinetic constants: & , the turnover number; cat  K, m  the apparent dissociation constant; and  k JK , c  m  the apparent second-order rate constant, often referred to as the specificity constant. Assessment of these kinetic parameters will yield insight into how particular mutations affect the rates of the individual glycosylation and deglycosylation steps. The generally accepted double-displacement mechanism for Cex can be expressed as follows: glycosylation deglycosylation E + G-OR , ,** E.G-OR — E - G — E %  ROH where,  + GOH  H 0 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 = second-order rate constant for association k_ =first-orderrate constant for dissociation k =first-orderrate constant for glycosylation x  x  2  k =first-orderrate constant for deglycosylation 3  As discussed in Appendix C, the kinetic parameters, & , cat  K, m  and  k JK , c  m  can be  defined in terms of the individual rate constants as follows:  k  - - t k k  ( 2  -  2 )  ( 2  '  3 )  + k)  3  2  Jt (* * ) 1  2+  3  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  K  m  58_  k_ k l+  2  Mutations that reduce the rate of deglycosylation (relative to glycosylation) will result in accumulation of the glycosyl-enzyme intermediate.  Therefore, provided  deglycosylation is the rate-determining step (k « k ), the K reduces to 3  K  2  m  j  Therefore, the K  m  decreases as the ratio k /k . Correspondingly, a relative 3  2  reduction in the rate of glycosylation would be expected to increase the K . The effect of m  a mutation on the rate of the glycosylation step can also be estimated by analysis of k JK , c  of k  m  caU  which reflects the first irreversible step in the reaction. Detailed interpretations K, m  and k JK c  m  are given in Appendix C.  Nitrophenyl cellobiosides and xylobiosides with aglycones having different pK  &  values were evaluated as substrates for Cex and its mutants. The enzymatic hydrolysis rates were measured by spectrophotometric detection of phenol release under the standard assay conditions described in Materials and Methods. The enzyme concentrations employed were those which gave a sufficiently large absorbance change to ensure accurate determination of rates, yet resulted in less than 10% conversion of the substrate to product to ensure linear kinetics. The maximum velocities (F  max  ) and the apparent  dissociation constants (K ) were determined when the highest substrate concentration m  assayed was greater than 2K by fitting the initial rates and substrate concentrations to m  the Michaelis-Menten equation using the program GraFit4.0 ( i i i ) . At low substrate concentration, when no saturation was reached ([S]  max  < 2K ), k JK m  c  m  was evaluated as  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  the slope of the linear region of  VQ  versus [S]. Alternatively,  k JK C  M  59_  was determined  using the substrate depletion method using very little substrate ([S] < 0.2AT ). Regardless m  of the method employed, all values are numerically equivalent, confirming the validity of this approach for these studies. In various cases, the Lineweaver-Burk plots for the xylobioside substrates were biphasic (Figure 2.3), revealing a rate enhancement over that expected at higher substrate concentrations. Such behaviour is likely due to transglycosylation of the xylobiosylenzyme intermediate to a second substrate molecule bound in the aglycone site at higher substrate concentrations.  This has been observed to occur with Agrobacterium sp.  (3-glucosidase (Abg) with PNPX (21). Values of k  cai  and K for both the hydrolytic m  reaction and the transglycosylation reaction were estimated by drawing a straight line through the two regions of the Lineweaver-Burk plot. Such an approach was used to estimate these two parameters for Abg (21). Thus, kinetic parameters determined at low substrate concentrations are those of the simple hydrolysis process, while those at high substrate concentration are for the transglycosylation process; the high K  M  reflecting the relatively poor binding at the aglycone site.  value  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  500  o 400 o c  E  j\  i  i  i >  60  /° •  / -  /  300 -  1  y  i  200  *  1 1 1  /  i y  -  -  100 0.5 1 1.5 [ONPJy (mM)  -20  r 0  20 40 60 80 100 l/IONPJy (mM" ) 1  150  oo <  100 h  E.  50 h  0.05  0.1 0.15 0.2 [ONPX,] (mM)  0.25  -2  0  2  4  6  8  10  1/[ONPX2] (mM" ) 1  Figure 2.3: Data for the hydrolysis of o-nitrophenyl xylobioside by wild-type Cex plotted in (A) and (B) Michaelis-Menten form; and (C) and (D) Lineweaver-Burk form. Kinetic parameters for the hydrolysis reaction were obtained by fitting data obtained at lower substrate concentrations (unbroken line), while those for the hydrolysis reaction were obtained by Fitting data obtained at higher substrate concentrations (dashed line).  Cex displays significant activity against soluble cellulose and xylan, and, as shown in previous studies and assembled in Table 2.4, is highly active against a range of aryl P-glucosides (22, 132). The enzyme displays considerably higher activity with aryl P-xylobiosides and p-xylosides, compared with the corresponding P-cellobiosides and P-glucosides, respectively.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  61  Table 2.4: Michaelis-Menten parameters for the hydrolysis of various aryl glycoside substrates by C.fimi xylanase. A in  Substrate  PNPG PNPX° PNPC" PNPX  0.033 2.6 15.8 40  a  a  2  2,4DNPG 2,4DNPC" a  a  (mM)  18 12.9  8.3 20 0.60 0.018 1.9 0.11  (s  1  m M  ')  0-P041 0.13 26 2200  9.5 117  Data taken from Tull and Withers (22).  The substrate specificities of family 10 enzymes vary considerably. Cex cleaves cellulosic substrates with significantly higher activity than do several other family 10 xylanases, as presented in Table 2.5. Although the xylanases from Streptomyces lividans and Cellvibrio japonicus cleave cellobiosides, the enzymes display far lower activity than Cex (121, 126). There is also some variability in terms of the rate-limiting steps in the double-displacement reaction mechanism. For example, for Cex the rate-limiting step for hydrolysis of either PNPC or 2,4DNPC is deglycosylation (22).  However, the rate-  limiting step in the hydrolysis of 2,4DNPC by XynlOA from Cellvibrio japonicus is glycosylation, as substantiated by the absence of a pre-steady state burst of DNP release (133).  Chapter 2  Characterization of Highly Conserved Residues in C, fimi Xylanase  62  Table 2.5: Michaelis-Menten parameters for the hydrolysis of various aryl glycoside substrates by several family 10 xylanases.  Cex, C. fimi"  PNPG  : XynlOA, S. lividans " h  0.033 0.011  XynlOA, C.japonicus'  PNPX  Cex, C.fimi" XynlOA, S. lividans Xynl OA, C.japqnicus' b  XynlOA, S. lividans XynlOA, C.japonicus 0  c  PNPX,  6.32 0.15 15.8  Cex,(^fimV  PNPC  2.6  Cex, C.fimi" I  3.6 2.6 40  XynlOA, C.japonicus  0  86  8.3 180 20 310 0.60 51 50 0.018 0.58 0.57  0.0041 0.00069 ND d 0.13 0.00081 0.00048 26 0.071 0.052 2200 51 150  Data taken from Tull and Withers (22). * Data taken from Ducros et al. (126). Data taken from Charnock et al. (121). ND, not detectable.  a  0  rf  2.2.4.1 Probing the -2 subsite: steady state kinetic analyses of Glu43, Asn44, and Lys47 mutants Residues in the active site are highly conserved among members of family 10 glycoside hydrolases.  For instance, the residues Glu43, Asn44, Lys47 are highly  conserved in the -2 subsite of the binding cleft of family 10 enzymes (Figure 2.1). Lys47 also interacts extensively with the -1 subsite. To evaluate the importance of conserved residues located at the -2 subsite of Cex, the effect on enzyme activity of creating E43A, N44A, and K47A mutations will be assessed. Cex E43A and N44A display very similar  Chapter 2  kcat  Characterization of Highly Conserved Residues in C. fimi Xylanase  63  values to those of native Cex, whereas K47A is significantly less active against the  aryl P-glucosides, particularly those with poorer leaving groups (Table 2.6). Table 2.6: Michaelis-Menten parameters for the hydrolysis of aryl glycosides by the C.fimi xylanase (Cex) and mutants E43A, N44A, and K47A. * Michaelis-Menten Parameters Enzyme  Substrate  2,4DNPC PNPC Native Cex  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex E43A  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex N44A  2,5DNPX PNPX  2  ONPX  Cex K47A  2  2  ^cat  Km  ^cat/A^m  (si)  (mM)  (s^mM )  0.068 ± 0.002 0.58 ± 0 . 0 1  124 ± 5 15.9 ± 0 . 5  DEGLY DEGLY  29.9 ± 0.6 32.0 ± 0 . 4 (i) 38.1 ± 0 . 8 ° (ii) 121 ± 8 "  0.0145 ±0.0009 0.0219 ±0.0007 (i) 0.068 ± 0.003 " (ii) 1.2 ± 0 . 1 °  2100 ± 2 0 0 1500 ± 7 0 (i) 560 ± 4 0 " (ii) 100 ± 2 0 "  DEGLY DEGLY  4.92 ± 0.02 1.29 ± 0 . 0 3  0.185 ±0.003 12.8 ± 0 . 5  26.6 ± 0 . 5 0.101 ±0.007  DEGLY DEGLY/GLY  (i) 19.5 ± 0 . 2 ° (ii) 35 ± 1" 21A ± 0.8 (i) 4 7 ± 3 ° (ii) 151 ± 7 °  (i) (ii) 5.4 (i) (ii)  (i) 3 1 0 ± 10° (ii) 100 ± 2 0 " 5.1 ± 0 . 4 (i) 3 0 ± 5 ° (ii) 19 ± 2 "  DEGLY  9.36 ± 0 . 0 6 10.1 ± 0 . 2  0.070 ± 0.002 1.74 ± 0 . 0 7  134 ± 4 5.8 ± 0 . 4  DEGLY DEGLY  (i) (ii) (i) (ii) (i) (ii)  (i) (ii) (i) (ii) (i) (ii)  (i) (ii) (i) (ii) (i) (ii)  43.1 ± 0 . 5 " 80±3" 89±7° 170 ± 10" 80±8° 430 ± 3 0 °  0.063 ± 0 . 0 0 1 " 0.35 ± 0 . 0 4 ° ±0.3 1.6 ± 0 . 2 ° 7.8 ± 0 . 6 °  0.038 ± 0 . 0 0 1 ° 0.27 ± 0 . 0 3 " 3.6 ± 0 . 4 " 8.9 ± 0 . 7 ° 0.64 ± 0 . 1 0 " 7.4 ± 0 . 9 "  4.20 ± 0.06  2.6 ± 0 . 1  PNPC  0.020 ± 0.002  29 ± 3  (i) 5 9 ± 2 " (ii) 120 ± 10" 0.36 ± 0 . 0 1 9.7 ± 0.2  (i) 0.20 ± 0 . 0 1 ° (ii) 0.76 ± 0 . 1 2 ° 9.6 ± 0 . 3 8.5 ± 0 . 3  PNPX ONPX  2  2  2  GM)  8.46 ± 0.07 9.24 ± 0.07  2,4DNPC  2,5DNPX  -1  RateLimiting Step (/Teat) (DEGLY or  1130 ± 5 0 ° 300 ± 5 0 ° 25 ± 4 ° 20 ± 2 " 130 ± 3 0 ° 6 0 ± 10"  1.6 ± 0.1 0.00071 ± 0.00015 (i) 300 ± 2 0 ° (ii) 150 ± 3 0 ° 0.038 ± 0.002 1.15 ± 0 . 0 6  DEGLY  DEGLY  DEGLY  DEGLY DEGLY DEGLY DEGLY GLY DEGLY GLY GLY  " Parameters are for the (i) hydrolysis and (ii) transglycosylation reactions. * Graphical representation of this data is given Appendix A, page 228. Errors were estimated from a non-linear fit by GraFit 4.0 (131).  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  64  For substrates that show no significant dependence of & on phenol leaving group cat  ability, the initial bond-cleavage step is unlikely to be rate limiting; thus, the deglycosylation step is the rate-limiting step for these substrates. Deglycosylation must be rate determining for substrates where transglycosylation is observed at elevated substrate concentrations. Conversely, for substrates that show a significant dependence of  &cat  on leaving group ability, the glycosylation step is likely to be the rate-limiting step  for these substrates. The  k  cat  values for hydrolysis of xylobioside substrates by the  E43A  variant are  essentially unchanged relative to those of the wild-type enzyme, indicating that the mutation does not affect the deglycosylation step, which remains rate determining. Hydrolysis of cellobioside substrates follows a somewhat different pattern, with 2,4DNPC  k  cai  for  and PNPC being 2- and 7-fold lower, respectively, than for wild-type enzyme,  implying that the rate-limiting step changes from deglycosylation to glycosylation, at least for PNPC. This is supported by the much higher K value observed. m  The kcm values for hydrolysis of cellobioside and xylobioside substrates by the N44A  variant are relatively unchanged, indicating that the mutation does not affect the  rate-limiting deglycosylation step.  Lys47  hydrolyzed aryl cellobioside and xylobioside  substrates with modest leaving groups with significantly lower than the wild-type enzyme. However, the values of k  t  ca  2,4DNPC  and  2,5DNPX2,  k t ca  and higher  K  m  values  for the more activated substrates  were similar to those of the wild-type enzyme. The mutation  changes the rate-limiting step for PNPC, as well as  PNPX2  and ONPX hydrolysis from 2  deglycosylation to glycosylation. Such a change in rate-limiting step is not seen for the more activated substrates since the glycosylation step is inherently faster in those cases.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  65_  As shown in Figure 2.4, the three variant enzymes have simple bell-shaped pH profiles, where the pAT of the acidic limb of the profile, corresponding to the ionization of a  the nucleophile, is largely unaffected.  The basic limb, corresponding to the acid/base  catalyst, is affected differently in each case. No significant shift is seen for the N44A enzyme, whereas the pK corresponding to ionization of the acid/base catalyst for Cex a  E43A and Cex K47A shifted to 6.9 and 6.0, respectively, compared to the wild-type value of 7.7 as shown in Table 2.3.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  2  4  6  8  10  2  4  6 PH  8  10  66  ~ 3000 |  2500  \  2000  * 1500 E  1000 ^  500 0  Figure 2.4: pH profiles of wild-type (O) and Glu43, Asn44, and Lys47 mutants of Cex. The pH dependence of wild-type Cex was measured using 2,4DNPC where p A a j = 3.9 and pK = 7.7. (A) The E43A (•) mutant enzyme activity was measured using 2,4DNPC where pJST = 3.7 and pK = 6.9. (B) The N44A (•) mutant enzyme activity was measured using 2,4DNPC where pK = 3.9 and pK = 8.0. (C) The K47A (•) mutant enzyme activity was measured using 2,5DNPX where pK = 3.9 and pK^ = 6.0. a2  al  a2  ai  a2  2  ai  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  57  2.2.4.2 Probing the -1 subsite: steady state kinetic analyses of His80 mutants In order to investigate the role of the His80 residue in the structure and function of Cex, three mutant enzymes were produced; namely, H80A, H80N, and H80Q. The properties of Cex H80A do not distinguish between its role as a hydrogen-bond (proton) donor or as an acceptor because Ala is capable of neither of these functions. Asn and Gin variants may distinguish between the models because these residues can participate in hydrogen bonding interactions that mimic the His nitrogens, but they cannot mimic the acid/base properties. The specific activity of these mutant enzymes is reduced by a significant amount, revealing the importance of this residue for the catalytic function of Cex. Fortunately, they retain sufficient activity to determine their kinetic parameters on the synthetic aryl glycoside substrates (Table 2.7). The properties of the H80A and H80N variants are nearly indistinguishable, while the activity of the H80Q mutant is further impaired by roughly 10-fold. These results indicate that the presence of an amide side chain at position 80 is not sufficient to restore Cex activity. Geometric limitations may hinder the amide-containing residues from mimicking the hydrogen-bonding contributions of His80, especially since it also interacts with Asp 123 in a manner reminiscent of a Ser—His—Asp triad in proteases. For the His80 mutations, deglycosylation remained the rate-limiting step for the hydrolysis of xylobioside substrates. For the cellobioside substrates, glycosylation may start to become rate limiting with PNPC.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  68  Table 2.7: Michaelis-Menten parameters for the hydrolysis of aryl glycosides by the C.fimi xylanase (Cex) and mutants H80A, H80N, and H80Q. * Michaelis-Menten Parameters Enzyme Substrate  Native Cex  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex H80A  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex H80N  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex H80Q  2,5DNPX  Km  or)  (mM)  8.46 ± 0.07 9.24 ± 0.07  0.068 ± 0.002 0.58 ± 0 . 0 1  124 ± 5 15.9 ± 0 . 5  DEGLY DEGLY  29.9 ± 0.6 32.0 ± 0 . 4 (i) 38.1 ± 0 . 8 ° (ii) 121 ± 8 °  0.0145 ±0.0009 0.0219 ±0.0007 (i) 0.068 ± 0 . 0 0 3 " (ii) 1.2 ± 0 . 1 "  2100 ± 2 0 0 1500 ± 7 0 (i) 560 ± 4 0 " (ii) 100 ± 2 0 "  DEGLY DEGLY  0.088 ± 0.004 0.048 ± 0.002  0.0071 ±0.0003 13± 1  12± 1 0.0037 ± 0.0004  (i) 0.25 ± 0 . 0 1 " (ii) 0.73 ± 0 . 0 3 " 0.42 ± 0.02 (i) 0.33 ± 0.02 " (ii) 2.9 ± 0 . 2 °  (i) 0.0021 ±0.0002" (ii) 0.32 ± 0 . 0 2 " 0.59 ± 0 . 0 3 (i) 0.039 ± 0 . 0 0 2 " (ii) 3.4 ± 0 . 2 "  (i) 120 ± 2 0 " (ii) 2.3 ± 0 . 3 " 0.72 ± 0.07 (i) 8.4 ± 0 . 8 " (ii) 0.84 ± 0 . 0 8 "  DEGLY  0.056 ± 0.003 0.040 ± 0.002  0.0051 ±0.0003 6.6 ± 0.4  11 ± 2 0.0060 ± 0.0006  DEGLY DEGLY  (i) (ii) (i) (ii) (i) (ii)  (i) (ii) (i) (ii) (i) (ii)  (i) (ii) (i) (ii) (i) (ii)  PNPX  2  ONPX  2  0.12 ± 0 . 0 1 " 0.22 ± 0 . 0 2 " 0.13 ± 0 . 0 1 ° 0.29 ± 0.03 " 0.14 ± 0 . 0 1 " 0.75 ± 0 . 0 7 °  0.012 ±0.002 0.0064 ± 0.0006 2  cat  Acat 1  2,4DNPC PNPC  RateLimiting Step (A ) (DEGLY or GLY)  (i) (ii) (i) (ii) (i) (ii)  0.034 ± o.o r 0.068 ± 0 . 0 1 ° 0.037 ± 0 . 0 1 " 0.10 ± 0 . 0 1 ° 0.039 ± 0 . 0 1 ° 0.26 ± 0 . 0 3 "  0.0012 ± 0 . 0 0 1 " 0.15 ± 0 . 0 3 " 0.099 ± 0 . 0 2 " 0.58 ± 0 . 0 6 " 0.016 ± 0 . 0 0 2 " 2.0 ± 0 . 2 "  ~ mM ) l  -1  99 ± 1 0 " 1.5 ± 0 . 3 " 1.3 ± 0 . 3 " 0.51 ± 0 . 1 " 8.6±1" 0.38 ± 0 . 1 "  0.0019 ±0.0002 3.1 ± 0 . 3  6.1 ± 1 0.0021 ±0.0004  (i) (ii) (i) (ii) (i) (ii)  (i) (ii) (i) (ii) (i) (ii)  0.00062±0.0001" 0.17 ± 0 . 0 2 " 0.062 ± 0 . 0 0 6 " 0.63 ± 0 . 0 7 " 0.010 ± 0 . 0 0 1 " 2.4 ± 0 . 3 "  54 ± 1 0 " 0.40 ± 0 . 0 8 " 0.61 ± 0 . 1 " 0.16 ± 0 . 0 3 " 3.8 ± 0 . 8 " 0.11 ± 0 . 0 2 "  DEGLY DEGLY DEGLY/GLY DEGLY  DEGLY  DEGLY DEGLY DEGLY DEGLY DEGLY DEGLY DEGLY DEGLY  Parameters are for the (i) hydrolysis and (ii) transglycosylation reactions. * Graphical representation of this data is given Appendix A, page 228. Errors were estimated from a non-linear fit by GraFit 4.0 (131).  a  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  69  The pH profiles of the modified enzymes were determined and compared to that of the wild-type enzyme (Figure 2.5). The shapes of the mutant Cex plots are simple bell-shaped profiles similar to that of the wild-type, but are much narrower because the pH profiles are affected on both the acidic and basic limbs. Therefore, the different mutations exhibit effects on the ionization states of both catalytic glutamic acid residues. For the three mutations studied, the calculated pK shifts are 0.7 pH units toward a more a  basic pH for the nucleophilic Glu233 and 0.7-1.5 pH units toward a more acidic pH for the acid/base catalyst Glul27. The profiles were fit using equation 2.1 to provide two pK values for each variant, as tabulated in Table 2.3. A role for His80 in the hydrogena  bond network responsible for maintaining the ionization state of the two catalytic residues is indicated by the significant shift in pK values of the two catalytic residues in a  all three mutations.  Chapter 2  Characterization of Highly Conserved Residues in C, fimi Xylanase  70  Figure 2.5: pH profiles of wild-type (O) and His80 mutants of Cex. The pH dependence of wild-type Cex was measured using 2,4DNPC where pK = 3.9 and pK = 7.7. (A) The H80A (•) mutant enzyme activity was measured using PNPX where pA" = 4.6 and pK = 6.2. (B) The H80N (•) mutant enzyme activity was measured using PNPX where pK = 4.6 and pA„ = 6.3. (C) The H80Q (A) mutant enzyme activity was measured using PNPX where pK = 4.6 and pK = 7.0 ai  a2  2  a2  al  2  al  2  2  ai  32  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  7/  2.2.4.3 Probing the -1 subsite: steady state kinetic analyses of Asnl26 and Asnl69 mutants N126A and N169A variants of Cex were previously constructed and purified to apparent homogeneity by Dr. Valerie Notenboom (University of Toronto) to investigate the role of Asnl26 and Asnl69 residues in the structure and function of Cex. The N169A mutant displays similar kinetic properties to that of native Cex with all substrates tested. The N126A mutant is considerably less active than wild-type Cex (Table 2.8). For both N126A and N169A, deglycosylation is still the rate-determining step in the hydrolysis of the xylobioside substrates.  For N126A, the k  cat  value for the hydrolysis of PNPC  decreases sharply, consistent with rate-limiting glycosylation for this substrate.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  72  Table 2.8: Michaelis-Menten parameters for the hydrolysis of aryl glycosides by the C.Jimi xylanase (Cex) and mutants N126A and N169A. 0  Michaelis-Menten Parameters Enzyme  Substrate  2,4DNPC PNPC Native Cex  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex N126A  2,5DNPX PNPX  2  2  ONPX  2  2,4DNPC PNPC Cex N169A  2,5DNPX PNPX  2  2  ONPX  2  RateLimiting Step (* ) (DEGLY or cat  ^cat  (mM)  (s  H  mM ) -1  8.46 ± 0.07 9.24 ± 0.07  0.068 ± 0.002 0.58 ±0.01  124 ± 5 15.9 ± 0 . 5  DEGLY DEGLY  29.9 ± 0 . 6 32.0 ± 0 . 4 (i) 38.1 ± 0 . 8 ° (ii) 121 ± 8 °  0.0145 ±0.0009 0.0219 ±0.0007 (i) 0.068 ± 0 . 0 0 3 " (ii) 1.2 ± 0 . 1 "  2100 ± 2 0 0 1500 ± 7 0 (i) 560 ± 4 0 " (ii) 100 ± 2 0 "  DEGLY DEGLY  0.49 ±0.01 0.052 ± 0.005  0.047 ± 0.002 12± 1  10.4 ± 0 . 5 0.0044 ±0.001  DEGLY GLY  (i) 1.30 ± 0 . 0 1 " (ii) 2.0 ± 0 . 1 " 3.5 ± 0 . 1 (i) 2.9 ± 0 . 1 ° (ii) 6.9 ± 0 . 4 °  (i) (ii) 7.7 (i) (ii)  2.89 ± 0 . 0 3 3.9 ± 0 . 2  0.036 ± 0.002 0.59 ± 0 . 0 3  81 ± 4 6.6 ± 0 . 5  (i) 19.4 ± 0 . 3 " (ii) 5 4 ± 4 " nd* (i) 29 ± 1" (ii) 160 ± 1 0 °  (i) 0.0076 ± 0 . 0 0 0 4 " (ii) 0.20 ± 0 . 0 3 " nd* (i) 0.050 ± 0 . 0 0 6 " (ii) 1.2 ± 0 . 1 "  (i) 2600±200" (ii) 270 ± 6 0 " nd* (i) 580 ± 9 0 " (ii) 130 ± 2 0 "  0.0025 ± 0.0001 0.056 ± 0 . 0 0 8 ° ±0.2 0.39 ± 0 . 0 2 " 1.6 ± 0 . 1 "  a  (i) 520 ± 2 0 " (ii) 3 6 ± 6 " 0.45 ± 0.05 (i) 7.3 ± 0 . 5 " (ii) 4.4 ± 0 . 6 "  DEGLY  DEGLY DEGLY DEGLY DEGLY DEGLY DEGLY DEGLY DEGLY  Parameters are for the (i) hydrolysis and (ii) transglycosylation reactions. * nd indicates that this kinetic parameter was not determined. Graphical representation of this data is given Appendix A, page 228. Errors were estimated from a non-linearfitby GraFit 4.0 (131).  a  c  The pH profiles of the N126A and N169A variants exhibit near wild-type pH profiles (Figure 2.6), although the pH optima are shifted toward a slightly more basic pH. Maximal activity is observed at pH values of 6.4 and 6.1 for the N126A and N169A variants, respectively. The data were fit using equation 2.1 to obtain the pK values a  shown in Table 2.3.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  73  Figure 2.6: pH profiles of wild-type (O) and Asnl26 and Asnl69 mutants of Cex. The pH dependence of wild-type Cex was measured using 2,4DNPC" where pAai = 3.9 and pK = 7.7. (A) The N126A (•)mutant enzyme activity was measured using 2,4DNPC where pK = 4.4 and pK = 8.5. (B) The N169A (•) mutant enzyme activity was measured using 2,4DNPC where pA^ = 4.0 and pK = 8.2. a2  al  a2  a2  2.3 Discussion 2.3.1 Effects of mutations on glycosylation and deglycosylation  Changes in the reaction parameters observed for Cex-catalyzed hydrolysis of the cellobiosides and xylobiosides upon mutation of the enzyme can be related to changes in the activation free energy. These values of AAG* for each mutation have been calculated  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  for the glycosylation step (from k JK c  m  74  values) and for the deglycosylation step k (from 3  k values). They are presented in Table 2.9 and summarized in Table 2.10. cat  As is evident from the values of the rate constants and the free energy changes, the mutations have a significant effect on each of the two steps of Cex catalysis. Further, the magnitude of the effect varies with both the mutation and the step in the reaction.  Table 2.9: Kinetic parameters for the glycosylation and deglycosylation steps of the hydrolysis reaction by variants of Cex.  Enzyme  2.4DNPC PNPC Native Cex  (A at/^m)  Substrate  2,5DNPX PNPX ONPX  2  2  (heat/Km)  ; 1  Cex E43A  2,5DNPX PNPX ONPX  2  2  2  2,4DNPC PNPC Cex N44A  2,5DNPX PNPX ONPX  2  2  2  2,4DNPC PNPC Cex K47A  2,5DNPX PNPX ONPX 2  2  2  ,v A A G *  . ,.  AAG*  DEGLY GLY 1 (kcal mor )^ (kcal mol ) -1  1 1  0 0  0 0  1 1 1  0 0 0  0 0 0  1.7 ± 0 . 0 2 na"  4.7 ± 0 . 2 160 ± 10  -0.3 ± 0 . 0 1 na"  -1.0 ± 0.1 -3.1 ± 0 . 1  1.5 ± 0 . 0 2 1.2 ± 0 . 0 4 0.8 ± 0 . 1  6.7 ± 0 . 7 290 ± 30 19 ± 4  -0.3 ± 0 . 0 1 -0.1 ± 0 . 0 2 0.1 ± 0 . 0 4  -1.2 ± 0.1 -3.5 ± 0 . 1 -1.8 ± 0.1  0.9 ±0.01 0.9 ± 0.02  0.9 ± 0 . 1 2.7 ± 0 . 3  0.1 ± 0 . 0 1 0.1 ± 0 . 0 1  0.0 ± 0 . 1 -0.6 ± 0 . 1  0.7 ±0.01 0.4 ± 0.03 0.5 ± 0.05  1.8 ± 0 . 3 5 9 ± 10 4.5 ± 1.1  0.2 ±0.01 0.6 ± 0.05 0.5 ± 0 . 1  -0.4 ± 0 . 1 -2.5 ± 0 . 1 -0.9 ± 0.2  2.0 ± 0.03 na°  77 ± 6 23000 ± 5000  -0.4 ±0.01 na"  -2.7 ± 0 . 1 -6.2 ± 0 . 1  0.5 ± 0.02 na" na°  7.0 ± 0 . 8 39000 ± 3000 490 ± 40  0.4 ± 0.02 na na°  -1.2 ± 0 . 1 -6.5 ± 0 . 1 -3.8 ± 0 . 1  2  2,4DNPC PNPC  W T  C  a  Chapter 2  ?;;Enzyme ;  Characterization of Highly Conserved Residues in C. fimi Xylanase  Substrate  AAG* DEGLY (kcal mol )  AAG* GLY (kcal mol )  -1  2,4DNPC PNPC Cex H80A  2,5DNPX PNPX ONPX  2  2  2  2,4DNPC PNPC Cex H80N  2,5DNPX PNPX ONPX  2  2  2  2,4DNPC PNPC Cex H80Q  2,5DNPX PNPX ONPX  2  2  2  2,4DNPC PNPC CexN126A  2,5DNPX PNPX ONPX  2  2  2  2,4DNPC PNPC CexN169A  2,5DNPX PNPX ONPX 2  2  2  75  -1  96 ± 4 na"  10± 1 4300 ± 500  -2.8 ± 0.03 na°  -1.4 ± 0.1 -5.2 ± 0 . 1  120 ± 5 76 ± 4 120 ± 7  17 ± 3 2100 ± 2 0 0 66 ± 8  -2.9 ± 0 . 0 3 -2.7 ± 0.03 -2.9 ± 0.04  -1.8 ± 0.1 -4.7 ± 0 . 1 -2.6 ± 0 . 1  150 ± 10 230 ± 10  11 ± 2 2600 ± 300  -3.1 ± 0 . 0 3 -3.4 ± 0 . 0 3  -1.5 ± 0 . 1 -4.9 ± 0 . 1  250 ± 20 250 ± 20 270 ± 20  21 ± 3 1100 ± 3 0 0 64 ± 9  -3.4 ± 0 . 0 5 -3.4 ± 0 . 0 5 -3.5 ± 0.04  -1.9 ± 0.1 -4.3 ± 0 . 1 -2.6 ± 0 . 1  700±100  20 ± 3 7 7 0 0 ± 1500  -4.0 ± 0 . 1 na"  -1.8 ± 0.1 -5.5 ± 0 . 1  880 ± 260 860 ± 230 980 ± 250  38 ± 8 2400 ± 400 140 ± 3 0  -4.2 ± 0.2 -4.2 ± 0.2 -4.2 ± 0.2  -2.2 ± 0 . 1 -4.8 ± 0 . 1 -3.1 ± 0 . 1  17 ± 0 . 4 na"  12± 1 3600 ± 800  -1.8 ± 0.01 na"  -1.5 ± 0 . 1 -5.0 ± 0 . 1  23 ± 0.3 9.2 ± 0 . 3 13± 1  4.0 ± 0 . 4 3200 ± 400 75 ± 7  -1.9 ± 0 . 0 1 -1.4 ± 0 . 0 2 -1.6 ± 0 . 0 2  -0.8 ± 0 . 1 -5.0 ± 0 . 1 -2.7 ± 0 . 1  2.9 ± 0 . 1 2.4 ± 0 . 1  1.5 ± 0 . 2 2.4 ± 0 . 3  -0.7 ± 0 . 0 1 -0.5 ± 0.03  -0.3 ± 0 . 1 -0.5 ± 0 . 1  1.5 ± 0 . 0 3 na" 1.3 ± 0 . 1  0.8 ± 0 . 1 na 1.0 ± 0 . 2  -0.3 ± 0 . 0 1  0.1 ± 0 . 1 na" 0.0 ± 0 . 1  " na indicates that this parameter is not available.  a  a  na -0.2 ± 0.02  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  76  Table 2.10: Free energy changes at the glycosylation and deglycosylation steps of the hydrolysis reaction by variants of Cex.  (kcal-mol*)  .. . • (kcalSmol'i) - •.  - t«Fello biosVI-fc', '*' JiSSXvlobiosvl-,'-?*' ;I3 *n /••*r;s* \nui.. ,  ,  • •.*."/«»!** "i'X*-." Cll/Mllf *  . ^f.V'-TfWft" • I'll/MIU'  -• ILK  ^"  i.  0 -1.0 0.0 -2.7 -1.4 -1.5 -1.8 -1.5  0 -3.1 -0.6 -6.2 -5.2 -4.9 -5.5 -5.0 -0.5  ill  HHMHHHHHHHBHII  Native Cex Cex E43A " Cex N44A Cex K47A Cex H80A Cex H80N CexH80Q CexN126A CexN169A  r  o "7 -o.3±o.r 0.1 ± 0 . l ' -0.4 ±6.1 -2.8 ± 0 . 1 -3.2 ± 6.1 _  4  Q  ±  Q  2  -1.8 ± 6.1 -0.6 ± 0 . 1  o " ~ -0.2 = 0.1 0.4-0.1 6.4 ± 0 . 1 -2.9 ± 0 . 1 -3.4 ± 0 . 1 -4.2 ± 0.2 -1.6 ±6.1 " -0.3 ± 0 . 1  " na indicates that this parameter is not available. * Error estimates for AAG* G L Y are given in Table 2.9.  -673  o " 0 -1.2 -3.5 -0.4 -2.5 -1.2 -6.5 -1.8 " -4.7 -1.9 -4.3 -2.2 -4.8 -0.8 -5.0 0.1 na"  6  1  -1.8 -0.9 -3.8 -2.6 -2.6 -3.1 -2.7 0.0 \ !  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  -6  -5 -4 -3  AAG*  G L Y  PNPXJ  -2  -1  77  0  (kcal mor ) 1  -5 - 4 - 3 - 2 - 1 0 1 1 AAG* 2 . 5 D N P X , (kcal mor ) D E G L Y  -5  - 4 - 3 - 2 - 1 0 1 1 AAG* O N P X , (kcal mol" ) G L Y  Figure 2.7: (A) Comparison of rate constants for glycosylation of PNPC and PNPX by variants of Cex. A slope of m = 1.0 and correlation coefficient of r = 0.87 was observed. (B) Comparison of rate constants for deglycosylation of cellobiosyl-enzyme intermediates and xylobiosyl-enzyme intermediates of variants of Cex. A slope of m = 0.94 and correlation coefficient of r = 0.95 was observed. (C) Comparison of rate constants for glycosylation and deglycosylation reactions for the hydrolysis of ONPX by variants of Cex. Open circles correspond to mutations within the —2 subsite; and closed circles correspond to mutations within the -1 subsite of Cex. A slope of m = 0.97 and correlation coefficient of r = 0.62 was observed. 2  2  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  78  The good linear free energy relationship (r = 0.87) observed between the rates of glycosylation of PNPC and PNPX indicates similarity of transition states for the 2  glycosylation of cellobiosides and xylobiosides by Cex and the series of mutants (Figure 2.7A). An even greater correspondence of transition states for the deglycosylation of cellobiosides and xylobiosides, as indicated by the linear free energy relationship (r = 0.95), was observed between the rates of deglycosylation of cellobiosides and xylobiosides by Cex and the series of mutants (Figure 2.7B). Comparison of the rate constants for the glycosylation and deglycosylation steps should provide insights into the relative amounts of positive charge (oxocarbenium ion character) generated at the two transition states. Such a free energy relationship is presented in Figure 2.7C. This plot has a weak correlation coefficient of r = 0.62; the modest correlation coefficient indicating that the mutations affect the two steps of the hydrolysis reaction differently.  2.3.2 A second look at the rate-limiting step in the hydrolysis reaction by Cex and mutants: diffusion-controlled rate constants  . Classically, the rate-determining steps in an enzyme-catalyzed reaction have been considered to be those involving chemical conversion of ES to EP complexes or dissociation of products (134). While this situation undoubtedly holds true for many enzymes, there is increasing evidence that for some enzymes the encounter of enzyme and substrate is the rate-determining step, and these enzymes are said to be at least partly diffusion-controlled. Albery and Knowles (135) have remarked that such enzymes have reached the end of their evolutionary kinetic refinement, since further improvements in catalytic efficiency would not be manifested in enhanced rates of reaction. The relevant  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  kinetic parameter is the second-order rate constant k JK , c  79-  which cannot exceed a  m  diffusion-controlled limiting value that has been estimated to be on the order of 1 0 8  10 s M l u  _ 1  for a number of enzyme-substrate complexes (135). The exact value  _ 1  depends in part on the possible contribution of electrostatic forces in the encounter of substrate and enzyme. There is, however, no clear lower limit for k JK c  m  below which an  enzyme reaction can no longer be considered to be diffusion controlled, for a number of reasons. For instance, steric constraints can arise from the location of active sites in deep grooves on an enzyme's surface. In addition, catalytically incompetent conformational states of an enzyme can coexist in equilibrium with the native state. The effect of such constraints is a reduction in the diffusion-controlled rate constant.  Frequently, the  observed association rate constants for enzyme-substrate interactions tend to fall in the range of 10 - 10 s" M " (136). 6  8  1  1  The rate constants for glycosidases acting on their natural substrates are typically several orders of magnitude below the diffusion-controlled values. Cex, however, is a broad specificity enzyme that hydrolyzes a number of synthetic substrates. For example, the rate constants for the formation of the xylobiosyl-enzyme of Cex, representing the first irreversible step in the reaction, can exceed 2 x 10 s M 6  presented in tables are in units of s m M ) . _1  _l  H  _ 1  (note that k JK c  m  values  This value approaches the diffusion-  controlled limiting value and consequently the encounter of enzyme and substrate may be the rate-determining step for this reaction when reactive leaving groups are present.  The observed rate constant for the formation of product is given by  k,  Chapter 2  Characterization of Highly Conserved Residues in C.fimiXylanase  where the rate constants k and x  dissociation.  80  represent the rate constants for association and  The well-established kinetic mechanism for Cex-catalyzed reactions is  given below. glycosylation deglycosylation E + G-OR ^=4=± E.G-OR -4— E-G -4— E + GOH ROH  H 0 2  The deglycosylation step, k , is overall rate determining for synthetic cellobioside 3  and xylobioside substrates with good to moderate leaving groups, while the glycosylation process, k , is limiting for synthetic substrates with poor leaving groups and the natural 2  substrates. The first irreversible step in both cases is glycosylation of the enzyme. Where Michaelis complex formation is at true equilibrium  » k ) , equation 2.6 2  reduces to  k=^  k k  k  k  (2.7)  =^ K  Conversely, covalent capture of the glycosyl moiety of the substrate by the enzyme with a rate constant greater than that for dissociation (k » &_,) leads to 2  (2.8)  K  Evidence for rate-limiting association was observed for  Agrobacterium  sp.  P-glucosidase (21). A biphasic relationship was obtained from the plot of the logarithm of  k /K cat  m  against the phenol  pK  a  of a series of aryl glucosides. It was reasoned that a  biphasic relationship in this case could not be due to a change to rate-limiting  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  8J_  deglycosylation at higher reactivities and was most likely due to association of enzyme and substrate becoming rate-limiting with the more reactive substrates. The second-order rate constants (k /K ) cat  m  for these substrates fell within the range of 2 x 10 6  3x lOV'M" . 1  It is possible that the rates of glycosylation of Cex and certain variants of Cex by highly reactive substrates approach the diffusion-controlled limits. For instance, the second-order rate constants (k /K ) for cat  m  2,5DNPX2  fall well within the range for which  such association has been shown to be rate limiting in other enzyme systems, including Agrobacterium  sp. (5-glucosidase (21) and the almond p-glucosidase (137).  2.3.3 Importance of -2 subsite amino acids Glu43, Asn44, and Lys47  2.3.3.1 Role of Glu43 Data obtained for the aryl P-cellobiosides suggest a change in the rate-limiting step from deglycosylation to glycosylation for the PNPC substrate given the 4-fold decrease in k obtained for the substrate relative to 2,4DNPC. Although the activity of cat  the E43A mutant is similar toward 2,4DNPC compared to wild-type Cex, the mutant is significantly less active against PNPC than the unmodified enzyme. The protonation of the glycosidic oxygen may be affected by the E43A mutation. The E43A modification may affect the way cellobiose sits in the active site such that the glycosidic oxygen is not close enough to Glul27 to be protonated. White et al. (32) showed that when 2-deoxy2-fluoro-p-cellobiose was covalently linked to the nucleophile of Cex, Glu43 forms a hydrogen bond with the C-2 hydroxyl of the distal saccharide unit. Consequently, a disruption of this bond in E43A could change the position of cellobioside substrates  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  82  within the active site, such that the glycosidic oxygen is no longer in close proximity to Glul27.  In contrast, in the glycosylated enzyme the covalent linkage between the  nucleophile Glu233 and the anomeric carbon of cellobiose positions the anomeric carbon in close proximity to the water molecule that is deprotonated by Glul27. Thus, the  k t ca  for E43 A against 2,4DNPC is similar to native Cex, as protonation of the leaving group is not essential for deglycosylation to occur. The mutation appears to have a slightly more detrimental effect on the binding of xylobiosides than cellobioside substrates. One possible explanation may be that the mutation generates more room for the accommodation of the C-5' hydroxymethyl moiety at the -2 subsite.  2.3.3.2 Role of Asn44 Crystal studies have shown that Asn44 in Cex forms a hydrogen bond with the hydroxyl at C-3' of the distal xylose of the covalently bound 2-deoxy-2-fluoroxylobiosylenzyme complex (76) and this suggests that it plays an important role in active-site binding (Figure 2.1). The N44A mutant of Cex exhibited similar properties to the native enzyme. The first-order rate for deglycosylation of the covalent cellobiosyl-enzyme intermediate was 9 s" for both the N44A mutant and native Cex. Therefore, the mutation presumably 1  affects the transition-state interactions at the transition state for the deglycosylation step to the same extent as the ground-state covalent intermediate complex. Furthermore, kcJKm values for both aryl P-cellobioside substrates remain comparable to those obtained for the wild-type enzyme. Similarly, the rate-determining step for the hydrolysis reaction for the aryl P-xylobioside substrates was deglycosylation. On average, however, the k  cai  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  value increased by 2-fold relative to the native enzyme.  83_  The mutation therefore  apparently destabilizes ground-state interactions to a greater extent than transition-state interactions for the second step of the hydrolysis reaction. The second-order rate constant k JK c  m  PNPX2  was affected to a greater degree, with at least a 70-fold decrease observed for the substrate, along with an increased K compared with native Cex. Overall, this m  asparagine residue plays a minimal role in sugar binding at the -2 subsite, and possibly the -3 subsite given that Asn44 is positioned in the distal region of the -2 site. A similar result was obtained against the aryl p-cellobiosides by the equivalent alanine mutant of Asn44 from C. japonicus XynlOA (133). 2.3.3.3 Role of Lys47  The data suggest that Lys47 plays an important role in positioning the substrate into the active site. The K47A mutant retains k values that are similar to native Cex cat  against 2,4DNPC and 2,5DNPX , although K values increase suggesting that the 2  m  mutation is influencing the glycosylation step. White et al. (32) showed that Lys47 in Cex formed hydrogen bonds with both the ring oxygen and C-3 hydroxyl of the distal and proximal glucose molecules of 2-deoxy-2-fluoro-P-cellobiose, respectively, in the glycosyl-enzyme complex. Removal of these hydrogen bonds could significantly alter the position of the substrate at the active site such that the nucleophile, Glu233, is not in close proximity with the anomeric carbon of the proximal sugar at the -1 subsite. Lys47 promotes proper substrate binding by hydrogen-bond donation, and may influence the orientation of the substrate to effect catalysis. However, it is also possible that K47A is having an indirect effect by altering the environment of aromatic residues at the active site of the enzyme. A potential candidate is Trp84, which is only ~4 A from Lys47, and  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  84  thus the removal of the charged nitrogen in the Lys47 side chain could enhance the hydrophobic environment of Trp84. This change could alter either the binding of the substrate at the active site or possibly the ionization state of the nucleophile.  2.3.3.4 Role ofHis80 The active site of the xylanase Cex from  Cellulomonas fimi  contains two histidine  residues (His80 and His205) that are almost completely conserved in family 10 glycanases. The structural analysis of the enzyme shows that His80 and His205 are part of an important hydrogen-bond network near the two catalytic residues (Glul27 and Glu233). The three-dimensional structure of Cex also suggests the involvement of His80 in substrate recognition, and shows that His80 is hydrogen bonded to 3-OH group of the proximal saccharide unit and the highly conserved residue Asp 123 (Figure 2.1). Co-crystal structures of xylanases from family 10 with xylopentaose (122) and with its 2-fluoro-2-deoxy-P-cellobiosyl intermediate have shown that the equivalent of His80 is hydrogen bonded with the C-3 hydroxyl of the saccharide moiety located in the -1 subsite in either case. The family 10 5.  lividans  XynlOA contains an equivalent residue,  His81 (Ns2) that is hydrogen bonded to Glu236 (Osl) through a water molecule and is hydrogen bonded through its N51 atom to Aspl24 (052)  (138).  The corresponding  histidine residues in other xylanases of family 10 have also been shown to be involved in maintaining the ionization state of the nucleophile (122, 124). To study the role of His80 in the structure and function of Cex, three mutations were made at this position. Since hydrogen bonding is involved, His80 was replaced with amino acids that have side chains capable of hydrogen bonding: asparagine (H80N) and glutamine (H80Q). The effects on the catalytic properties of the enzyme could be  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  85_  due to incorrect positioning of the nucleophile Glu233 in the catalytic site. Another explanation could be that an adequate stabilization of the catalytic intermediate is prevented by reduced interactions between His80 and the proximal sugar group. The three mutations at position His80 also significantly affect the pH profile of the enzyme (Figure 2.5). For the specific activity profiles, the effects of the three mutations studied affect both the acidic and basic sides, increasing the apparent pK by a  0.7 pH units toward a more basic pH for the nucleophilic glutamic acid residue and 0.71.5 pH units toward a more acid pH for the acid/base catalytic residue (Table 2.3). Therefore, in addition to its role in stabilizing the catalytic intermediate, His80 appears to be a very important residue in the hydrogen-bond network of the active site, and mutation of this residue modifies the interactions necessary to maintain the ionization state of the two catalytic glutamic acid residues of Cex. In conclusion, the results presented in this study show the importance of His80 for the structure and function of Cex. These residues are involved in a network of hydrogen bonds that are responsible for maintaining the ionization state of the two catalytic residues.  2.3.4 Amino acids that influence the catalytic residues  2.3.4.1 Role of Asnl26 An insight into the role of Asnl26 can be obtained from the study of White et al. (32), who suggested that this residue forms a hydrogen bond with the 2-OH group of the proximal glucose moiety of 2-deoxy-2-fluorocellobiose located at the -1 subsite. Modification of Asnl26 to alanine causes a significant decrease in the catalytic activity of  Chapter 2  Characterization of Highly Conserved Residues in C, fimi Xylanase  86  Cex against all the substrates tested. The rate-limiting step of hydrolysis of PNPC by N126A is glycosylation, whereas in the native enzyme deglycosylation is the limiting step in the cleavage of PNPC. These data suggest that Cex N126A is affecting both the efficient protonation of the substrate by Glul27 and the capacity of this residue to mediate subsequent general J?ase catalysis. It is possible that this interaction is important in positioning the glycosidic oxygen in close proximity to Glul27. The N126A mutation may cause an increase in the exposure of certain aromatic residues to a hydrophilic environment, and this subtle modification to the active site could alter the capacity of Glul27 to abstract protons from water.  Trp84 is only ~3 A from Asnl26, and the  removal of the amide component of the Asnl26 side chain could increase the hydrophilic environment of the tryptophan residue. Clearly, this is having a greater effect on the glycosylation step of PNPC cleavage, as PNP constitutes a moderate leaving group whereas protonation of 2,4DNP is not required for Cex to cleave 2,4DNPC; hence, the decrease in the rate at which this substrate is glycosylated results in a similar decrease in & to PNPC but also an associated increase in K . cat  m  2.3.4.2 Role of A s n l 6 9  Against the aryl (3-glycosides, N169A displayed similar k and k /K values as cat  cat  m  the native enzyme. This suggests that, although highly conserved, Asnl69 does not play a pivotal role in the binding of cellobiose or xylobiose in the active site of the enzyme. This is in contrast to the findings of White et al. (32) who suggested that the residue forms hydrogen bonds with Asnl26 and Glu233 in the structure of the covalently bound 2-deoxy-2-fluoro-cellobiosyl-enzyme intermediate.  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  87_  The pH profile for the N169A mutant enzyme exhibits a typical bell-shape, providing pK values of 4.0 and 8.2. The mutation does not alter the pH dependence of a  the nucleophilic residue greatly. However, this residue does influence the pK value of a  the acid/base catalytic residue.  2.3.5 Analyses of the pH profiles for Cex variants  The simple pH dependence of C. fimi xylanase arises from the critical roles played by two ionizable groups, the nucleophile (Glu233, pK -3.9) and the acid/base a  catalyst (Glul27, pK -7.7), with numerous other ionizable and non-ionizable groups a  modulating these pK s.  The effects of these other groups are brought out in the mutant  a  enzymes where a critical group is removed or perturbed in its properties. In general, residues that contribute positive charges and hydrogen bonds serve to lower the pK  a  values of Glu233 and Glul27.  In contrast, neighbouring carboxyl groups can either  lower or raise the pK values of the catalytic glutamic acids depending upon the a  electrostatic linkage of the ionization constants of the residues involved in the interaction. The pH optimum was shifted from -0.9 to +0.6 units by mutating residues within the active site. Some caution must be applied in interpreting the pH dependence of MichaelisMenten parameters and assignments of pK values (139). Nevertheless, comparison of a  the effects of pH on wild-type and mutant glycanases is mechanistically relevant and worthy of detailed discussion. The activity profile of Cex E43A follows apparent pK values of 3.7 and 6.9. The a  Cex E43A mutant is one where a potentially negatively charged residue was removed  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  88  from the active site. This resulted in a shift of the pH optimum of E43A from 5.8 to a more acidic value of 5.3. The most notable decrease in the apparent pK value was that a  of the basic limb, corresponding to the ionization of Glul27, by 0.8 pH units from 7.7 to 6.9. This is expected as a negative charge on Glu43 electrostatically destabilizes the conjugate base forms of the catalytic Glu residues, thereby serving to increase their pK  a  values relative to that of a neutral residue. This mutation has a greater effect upon the pK of Glul27 compared to Glu233, despite Glu43 being closer to Glu233 by 6.7 A in the a  crystalline wild-type enzyme. In the case of Cex N44A, replacement of an Ala for Asn resulted in minimal changes in the pK values controlling both limbs of its activity profile (p/C i =3.9 and a  a  PK& = 8.0). Removal of hydrogen-bonding residues generally serves to raise the pK  a  values of catalytic glutamic acids, as observed. Removal of a positive charge contributed by Lys47 resulted in a  decrease  in the  pH optimum of the enzyme from 5.8 to 4.9. The reduced pK observed for Glul27 is a  unexpected,  as removal of the positively charged Lys47 should lead to the opposite effect.  In general, a positive charge serves to stabilize the conjugate base forms of both catalytic Glu residues, thereby serving to lower their pK values relative to neutral residues. There a  is no obvious explanation for the precise nature by which the mutation leads to a decrease in the pK value of Glu 127 by a substantial 1.7 units, the largest observed decrease a  among the series of active-site residue mutants studied. It is possible that the K47A mutation results in localized structural perturbations.  The observed shift in the pH  optimum of the enzyme likely reflects delicate alterations in the complex hydrogenbonding network within the active site of Cex.  Chapter 2  Characterization of Highly Conserved Residues in C. firm Xylanase  89  His80 is positioned approximately 4 A from Glu233 and 7 A from Glul27 in the wild-type crystal structure and does not make direct contacts with either catalytic residue. In the structure of the 2F-xylobiosyl-Cex complex (76), however, the Ne2 atom of His80 is observed to make a direct contact with the 3-OH group of the proximal xylose saccharide and a highly conserved interaction is observed between His80 N81 with the 082 atom of Asp 123. Mutation of His80 influences the pK values of both catalytic a  residues. Mutation of His80 resulted in an increase in the pK of Glu233 from 3.9 to 4.6 a  for all variants of His80. This is as expected, as the hydrogen bonding properties of His80 stabilize the conjugate base form of Glu233, thereby serving to lower its pK value a  relative to that observed with a neutral residue at this position. Consequently, the increase in the pK value of the nucleophile should decrease the pK value of the a  a  acid/base catalyst due to electrostatic stabilization. The side chain of Asnl26 is positioned such that it can donate a strong hydrogen bond to Asnl69, which in turn is involved in a hydrogen-bonding interaction with the carboxyl group of the nucleophile Glu233 (Figure 2.1). When this interaction is removed through mutation of Asnl26 to Ala, the pK of Glu233 rises by 0.5 units to 4.4. The pH a  optimum of Cex shifts 0.6 units from 5.8 to 6.4, primarily due to elevation of the apparent pK value of Glul27 from 7.7 to 8.5. Asnl26 may influence the pK of Glul27 since the a  a  N82 atom of this asparagine is 4 A away from Glul27 Os2. The N82 atom of Asnl69 is involved in a hydrogen-bonding interaction with the Oe2 atom of the carboxyl group of the nucleophile Glu233 (Figure 2.1). When this interaction is removed through mutation of Asnl69 to Ala, the pK of Glu233 rises by 0.1 a  units to 4.0. Unexpectedly, the pK of Glul27 increases by 0.5 units to a value of 8.2. As a  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  90  seen for other Cex mutants, the pK value Glul27 is more sensitive to changes in the a  active site than is the pK of Glu233. It is clear that the hydrogen bond between Asnl69 a  and Glu233 can serve to lower the pK of this nucleophilic residue. However, the mode a  by which Asnl69 lowers the pK of Glul27 is less obvious since the N52 atom of this a  asparagine is >6 A away from either Glul27 Os2 or Osl. The position of the side chain of Asnl69 may change with pH such that it can form a hydrogen bond with Glul27. As mentioned previously, however, the  ionizations of Glu233 and Glul27  are  electrostatically coupled, and via this linkage the changes in the pK of one partner can a  influence the ionization of the other. In summary, the active site substitutions altered the pH-activity profile of Cex, in terms of both pH optimum and/or apparent pK values of the acidic and basic limbs. Any a  modifications to the local and global environments of Glu233 and Glul27 can result in changes in the pH optima of the enzyme.  2.3.6 Activity of family 10 enzymes  Data presented indicate that binding of substrates to the -2 subsite is important for efficient hydrolysis by Cex of xylobioside and cellobioside substrates. Data presented in various reports (133, 140, 141) show that there are significant differences in the capacity of cellobiose to bind to the active sites of family 10 xylanases. Although the k  cat  values for hydrolysis of both PNPC and 2,4DNPC by Cellvibrio japonicus XynlOA are similar to those of Cex, the K values of XynlOA were 2 to 3 orders of magnitude higher m  for the respective substrates, compared to those of Cellulomonas fimi xylanase. Furthermore, the rate-limiting steps for the hydrolysis of these two aryl P-cellobiosides  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  9J_  by QXynlOA and Cex are glycosylation and deglycosylation, respectively (705, 133). Surprisingly, although the k values for aryl P-cellobioside hydrolysis are similar for C3t  Cex and XynlOA, the Cellvibrio enzyme displays virtually no activity against polymeric cellulose substrates, in contrast to Cex. The difference in activity against cellulose is likely a reflection of the capacity of the + subsites of QXynlOA and Cex to accommodate glucose residues. The two family 10 xylanases, C. japonicus XynlOA and C. fimi xylanase Cex, hydrolyze aryl P-xylosides and xylobiosides at very different rates also, with  k JK c  m  values for Cex being 30-200 times higher than those of QXynlOA (22, 76, 133). Nonetheless, against the natural substrate xylan, the two enzymes have similar activities. In contrast, Cex is 20-fold more active against xylotetraose Xt, 10-fold more active against xylopentaose X 5 , and 3-fold more active against xylotriose X 3 than QXynlOA with respect to k JK c  m  (121).  The 30-fold difference between the activities of the two  enzymes against 4-nitrophenyl P-xylobioside, compared to the 3-fold difference for X 3 , has been suggested to result from the tighter binding of aryl moieties in the +1 subsite in the substrate binding cleft of Cex versus QXynlOA.  The efficient formation of a  productive complex to the C. fimi xylanase results from the tighter binding of the substituted aromatic ring. A fast glycosylation step, with deglycosylation being the ratedetermining step in the reaction, leads to a low K value for catalysis and the m  accumulation of the glycosyl-enzyme intermediate. In contrast, the glycosyl-enzyme intermediate does not accumulate in QXynlOA, hence the high K value obtained due to m  glycosylation being the rate-limiting step in the reaction presumably as a result of the poor binding of the aryl moiety in the substrate binding cleft (121).  Chapter 2  Characterization of Highly Conserved Residues in C. fimi Xylanase  The rate-limiting step for hydrolysis of X3 by both Q'XynlOA and Cex was found to be the glycosylation step (121). However, when the reducing end xylosyl residue of X  3  was substituted by a 4-nitrophenyl group (PNPX ), the k JK 2  10,000- and 1000-fold, respectively, for QXynlOA and Cex  c  values increased  m  These significant  (121).  increases in catalytic efficiency are probably a reflection of the capacity of the substituted phenolic rings to function as excellent leaving groups. Hence, there is little requirement for the glycosidic oxygen to be protonated during the glycosylation step.  2.4  Conclusions Site-directed mutagenesis studies provided insights into the roles of several  residues located in the - 2 and -1 subsites of the xylanase from C. fimi. The specific activity of these mutant enzymes is affected to various degrees, revealing the importance of these residues for the catalytic function of Cex. Furthermore, the pK values of the two a  catalytic residues are affected, demonstrating a role for these highly conserved residues in the hydrogen bond network responsible for maintaining the ionization state of the two catalytic residues.  92_  CHAPTER 3  AN ASSESSMENT OF T H E BINDING E N E R G Y C O N T R I B U T I O N O F H Y D R O G E N BONDS A T T H E TRANSITION STATE USING D E O X Y G E N A T E D SUBSTRATES AND SITE-DIRECTED MUTAGENESIS*  * A version of this chapter has been published. Wicki, J.. Schloegl, J., Tarling, C. A., and Withers, S. G . (2007) Recruitment of both uniform and differential binding energy in enzymatic catalysis: xylanases from families 10 and 11. Biochemistry, 46: 6996-7005.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  94  3 An Assessment of the Binding Energy Contribution of Hydrogen Bonds at the Transition State Using Deoxygenated Substrates and Site-Directed Mutagenesis 3.1  Introduction The extraordinary selectivities exhibited by most enzymes for their substrates are  necessarily derived from the interactions between the two partners, especially as they are expressed at the transition state. Of these, hydrophobic interactions are thought to contribute the major driving force for association, while hydrogen-bonding interactions and salt-bridge formation are thought to primarily provide the specificity  (142).  Interactions within a protein-ligand complex play an essential role in enzymatic catalysis and in cell-cell recognition (29,  143-148). Hydrogen-bonding  interactions are thought to  play a particularly critical role in the recognition of completely buried monosaccharides that contain a large number of hydroxyl groups. Many such protein-ligand interactions have been observed qualitatively from X-ray crystallographic studies of protein-ligand complexes  (143, 148-150),  while quantification of the contributions of specific hydrogen  bonds to a protein-ligand interaction has been obtained through a number of studies using two main approaches (58, 151, 152). One approach to providing an estimate of the contributions of various hydrogen bonds to binding is by using mutant enzymes in which single residues involved in specific interactions have been modified. In an elegant study by Fersht on a proteinligand complex of known three-dimensional structure, the side chains involved in hydrogen-bonding interactions with the substrate were systematically removed through site-specific mutagenesis and the affinity of the ligand for each mutant was then  Chapter 3  95  Binding Energy Contribution of Hydrogen Bonds  measured (151). More recently, this approach has been applied extensively to the study of glucoamylase (153, 154) and Bacillus 1,3-1,4-P-D-glucan 4-glucanohydrolases (155, 156). Alternatively, another approach to providing an estimate of the magnitude and polarity of a hydrogen bond involves the use of modified substrates in which the individual hydroxyl groups on the sugar moiety have been replaced by hydrogen or by fluorine. This approach has been applied to the study of P-glucosidase from Aspergillus wentii (58), glycogen phosphorylase (157, 158), phosphoglucomutase (159), Escherichia coli P-galactosidase  (27), Agrobacterium sp.  P-glucosidase  (28)  and trehalose  phosphorylase (160). The replacement of a sugar hydroxyl by hydrogen will remove all hydrogen-bonding interactions at that position, whereas substitution by fluorine will arguably allow hydrogen bonds in which fluorine acts as a hydrogen-bond (proton) acceptor, but not as a donor (Figure 3.1).  R  X.  donor  R  I  H  JO^  H  .X  ^  .X  acceptor  I  H  /X  acceptor  Figure 3.1: The hydroxyl can act as a hydrogen-bond donor or acceptor, whereas fluorine can only act as a hydrogen-bond acceptor.  Analysis of changes in binding interactions (K , K\, or in some cases K ) allows s  m  estimation of the contributions to ground-state interactions, whereas analysis of changes in rates (k /K csl  m  or & ) provides insights into transition-state interactions. For instance, cat  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  decreases in rate (k  t/K )  ca  m  96  observed for a deoxy sugar might be attributed to the loss of a  hydrogen-bonding interaction at that position and allow an estimation of the magnitude of the binding contribution at the transition state. Furthermore, the  type  of binding energy  change, whether "differential" or "uniform" in the parlance of Albery and Knowles (135), may be deduced from examination of kinetic parameters. An indication of the polarity of a hydrogen bond may be obtained when the rate of hydrolysis for a deoxyfluoro substrate is greater than that of the corresponding deoxy substrate.  Thus, if the enzyme acts as the hydrogen-bond donor, the deoxyfluoro  substrate may be able to retain some of the binding energy that is lost to the corresponding deoxy sugar substrate, resulting in a higher reaction rate. However, no simple deduction as to the polarity of the interaction at a particular position may be made when a lower rate of hydrolysis is observed for a deoxyfluoro compound versus the corresponding deoxy sugar.  3.1.1  Hydrogen-bonding energetics in water  Binding effects arise from the non-covalent interactions that develop between the enzyme and substrate at the transition state for the step in question, thereby stabilizing the transition state and providing catalysis. Hydrogen bonding between ligands and proteins in aqueous solution is a multifaceted process due to the competition of water for the hydrogen-bonding sites (161, 162). The very strong interaction of water with ions, dipoles, and hydrogen-bond acceptors or donors causes a reduction in, or disappearance, of most of the forces that would otherwise cause a strong interaction in a nonpolar solvent or vacuum. When two molecules A and B bind to one another, water has to be  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  97  displaced. The net change in energy is the observed binding energy, which measures the affinity of the two molecules for one another in competition with water. The difference between the energy of interaction of two molecules A and B with each other and the energy of interaction with water is small. A - B +H 0-H 0 ^ 2  2  A--H 0 + B - H 0 2  2  In fact, it is the strong energy of interaction of the water molecules with each other, rather than of a direct interaction of solute molecules with each other, that is responsible for the most important non-covalent intermolecular forces in water. It is difficult to pull water molecules apart in order to insert a weakly interacting molecule into the solution because of the large number of hydrogen bonds per unit volume of water. Relatively nonpolar molecules that have a small capacity to form bonds in a vacuum and cannot interact strongly with water are forced out of an aqueous solution and are therefore compelled to interact with each other. In this study, an overview of the hydrogen-bonding energetics in aqueous solution between enzyme and ligand, and of the ability of fluorine to hydrogen bond is required. A full hydrogen-bond inventory must be performed, as formulated previously by Jencks and Hine (161, 163), in order to understand the role of hydrogen bonding in the energetics of such interactions. A hydrogen-bonding interaction at a single site between a ligand and enzyme in an aqueous medium can be expressed as follows, E-H • • • OH + HOH • • • A-L ^ 2  [E-H • • • A-L] + HOH • • • OH  2  where a residue of enzyme E acts as a hydrogen-bond donor H which pairs with the acceptor A of the ligand L in the complex. The hydrogen-bonding roles of the enzyme  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  and the ligand could also be reversed. Although hydrogen-bond geometries may be quite different, both the numbers and types of hydrogen bonds are conserved in the process since there are two hydrogen bonds on each side. Binding energy therefore results partially from enthalpic differences due to different hydrogen-bond geometries but primarily from the increase in entropy associated with the release of water from the active site of the enzyme and the ligand into bulk water. A full loss of the energy of the hydrogen bond does not necessarily result from removal of a hydrogen-bonding group from the ligand L since in this case no hydrogen bond would exist between the substrate analogue and water before binding as shown below. E-H--OH +HOH + L ^ 2  [E-H L] + HOH • • • OH  2  Thus, the energetic difference between the two systems described above will not equal the full hydrogen-bond energy, but rather the difference in strengths of hydrogen bonds formed with the protein and the water. This difference may be rather significant when comparing a hydrogen bond in an evolved enzyme-ligand complex with that formed between a relatively mobile water molecule and a protein residue. In this study the two cases to be considered involve deoxygenation (OH to H) and deoxyfluorination (OH to F). The former is equivalent to the types of mutations performed by Fersht (151) and will result in complete removal of the hydrogen bond and a space in the complex that might accommodate a water molecule but is likely too small. The substitution of OH to F results in very little size difference, and thereby hampers coincident binding of water. The interaction is far more complex, since the fluorine  98  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  could be involved in a hydrogen bond but only as a hydrogen-bond acceptor (vide infra). Therefore, one may obtain two situations for the fluoro sugars as shown below. E - H - O H +HOH--F-L ^  [E-H • • • F-L] + HOH • • OH  E-A-H-OH + HOH-F-L ^  [E-A F-L] + HOH • • • OH  2  2  2  In the first case the enzyme acts as the hydrogen-bond donor at a particular position and fluorine acts as the hydrogen-bond acceptor, albeit a weak one. In the second case the enzyme accepts a hydrogen bond at a particular position and thus cannot interact favourably with the fluorine. A considerable decrease in affinity should result in this case since there is net loss of a hydrogen bond as the fluorine can still hydrogen bond as acceptor with the water. Thus, the deoxy sugar might have greater affinity than the fluoro sugar in such a situation. The ability of fluorine to act as a hydrogen-bond acceptor has been investigated in a number of studies discussed below.  3.1.2 Hydrogen bonding to fluorine  Fluorine has been documented to be involved in very strong hydrogen bonds. Fluorine is a stronger acceptor than the other halogens (164), but not as strong as oxygens and nitrogens (165). However, in their ionic and metal-bonded forms all the halogens act as considerable proton acceptors. In fact, the strongest known hydrogen bond is that in the (F—H—F)~ ion (166). In a large scale analysis by Howard and co-workers (167) of the X-ray data stored in the Cambridge Structural Database (CSD), short contacts of fluorine atoms to acidic hydrogens were discovered and reinforced by theoretical calculations from which half of the binding energy of a hydrogen bond to oxygen was  99  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  100  assigned to fluorine. Furthermore, a survey conducted by Shimoni and Glusker (168) in the CSD revealed that acidic hydrogens prefer to bind to stronger acceptors such as oxygens and nitrogens compared to fluorine atoms. Brammer and co-workers (169) examined the CSD and confirmed the weakness of fluorine hydrogen-bonding interactions, revealing shorter fluorine—donor contacts for C(sp )—F structures 3  compared to C(sp )—F, as previously observed by Howard and co-workers (167). 2  Increased acceptor ability of aliphatic over aryl-bonded fluorine atoms was observed; presumably because the fluorine lone-pairs are in conjugation with the 7i-orbital system and consequently less able to participate in hydrogen bonding. An exhaustive inspection of the Protein Data Bank (PDB) archive by Carosati and co-workers (170) was carried out to analyze those protein-ligand complexes with fluorines as acceptors and protein NH and OH groups as donors. Statistics were applied to all the PDB complexes whose protein structures had co-crystallized fluorine-containing ligands, and the effect of fluorine hydrogen bonds on the protein-ligand binding was estimated using the program GRID.  After protein-ligand complexes were identified as containing geometrically  possible hydrogen bonds, the contacts were investigated in detail by measuring the distance F—D and the angle at the donor.  Overall, fluorine atoms of ligands were  observed in hydrogen-bonding interactions in 18% of the complexes with a fluorinecontaining ligand (170).  3.1.3 Electronic effects  Electronic effects  have their origin in the considerable differences in  electronegativities of these substituents (H, OH, F), and the different effects these will  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  Wl_  therefore have on the stability of a positively charged, oxocarbenium ion-like transition state.  The magnitudes of these effects will be dependent upon the distance of the  substituent from the anomeric center, and will not be a major contributing factor beyond the first sugar ring.  The greatest electronic effects are those at the 2-position.  For  instance, replacement of a hydroxyl group at the 2-position with a hydrogen has been shown to favourably stabilize the oxocarbenium ion-like transition state for glycoside hydrolysis, resulting in substantial rate accelerations. The substitution of fluorine at the 2-position, on the  other  hand, destabilizes the  oxocarbenium transition  state.  Consequently, 2-deoxyfluoroglycosides, along with the 5-fluoroglycosides, have been successfully used as inactivators to characterize the covalent intermediate in the doubledisplacement mechanism as discussed previously (31, 41). Electronic effects were studied by Withers and co-workers (171) for a reaction that proceeds via an oxocarbenium ion-like transition state; that is, the acid-catalyzed hydrolysis of a series of deoxyfluoro-D-glucopyranosyl phosphates.  Significant rate  decreases were observed from fluorine substitution, with the greatest effect being upon the substitution of the 2-hydroxyl. Corresponding rate increases for a series of deoxyD-glucopyranosyl phosphates were observed as expected (172). More extensive studies were conducted on the rate constants for the spontaneous hydrolysis of substituted 2,4dinitrophenyl glycosides, thereby avoiding effects of the substituent on the concentration of the conjugate acid species and its pathway, and these gave similar results (18). The rate decreases observed for the fluoro sugars in the enzymic reaction are considerably greater than those observed for the corresponding non-enzymatic acidcatalyzed reaction, indicating that the loss of specific binding interactions that normally  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  W2_  contribute to stabilization of the transition state contributes to the rate reductions. The large rate reductions observed, as expressed in decreased k and k JK cat  0  m  values, likely  arise from some combination of intrinsic electronic effects and specific binding interactions. Although it is not possible to separate these two effects completely and determine their individual contributions, a reasonable estimate of the relative importance of the two effects can often be obtained, thereby providing useful insights into catalysis.  3.1.4 Objectives  In this study, the specificity constants, k JK , c  m  will be determined for the  hydrolysis of a series of deoxy and deoxyfluoro sugar derivatives by the family IOC. fimi xylanase (Cex) and the family 115. circulans xylanase (Bex), and these data will be interpreted in terms of active site hydrogen-bonding interactions. The binding energy contributions of individual sugar hydroxyl groups in the enzyme-substrate complexes at the transition state will be calculated using the relationship  AAG = -RT\n +  where x represents either a mutant enzyme or substrate analogue and y the wildtype enzyme or parent substrate. The series of calculated AAG* values or apparent binding energy contributions will then be compared to gain new insights into the role of hydrogen binding in catalysis by these two xylanases in particular as well as, more generally, into the molecular basis for remote binding effects upon catalysis. The carbohydrate-binding site of C. fimi xylanase will also be analyzed through a mutational analysis to probe the role of protein-carbohydrate interactions defining substrate  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  103,  specificity. Amino acid residues involved in substrate binding were mutated on the basis of the previously determined X-ray crystal structures of enzyme-substrate complexes. The effects of the mutations of various selected residues on catalysis and binding will be determined by steady state kinetics using a series of chromogenic substrates to assign the individual hydrogen-bond contributions. Comparison of the two methods will provide an evaluation of the validity of the use of such modified substrate analogues and appropriate active-site mutants to probe the role of hydrogen bonding in protein-ligand interactions, as well as an evaluation of the importance of hydrogen bonding in the generation of biological specificity.  3.2 Results and Discussion 3.2.1  Binding energy contributions probed by using substrate analogues  The  series  of  monosubstituted  deoxy  and  deoxyfluoro  4-nitrophenyl  P-xylobiosides shown in Figure 3.2 was synthesized by Dr. Johann Schloegl and Dr. Chris A. Tarling, and generously provided for this study. These represent a reasonably full set of substrate analogues for probing the role of hydrogen bonding at each hydroxyl. Unfortunately, despite many attempts, the substrate analogue 4-nitrophenyl 3-deoxyp-xylobioside (3-deoxy-PNPX2) could not be synthesized due to its lability (18). Likewise, synthesis of 4-nitrophenyl 2-deoxy-P-xylobioside (2-deoxy-PNPX2) was not attempted as it was expected to be yet more labile. However, fluorinated analogues were obtained for each position.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  HO HO, n u n v  PNPX  OH  NO,  2  104  -0  OH 3-deoxy-3-fluoro-PNPX  NO,  2  OH  o« nunv 2'-deoxy-PNPX  NUo  2  H O ^ ^ C . - ^ ^  H 8 ^ ^ S H - O \ ^ O _ OH  l^i  2 -deoxy-2'-fluoro-PNPX ,  N  2  °  2  C  OH  N0  3'-deoxy-PNPX  OH 3*-deoxy-3'-fluoro-PNPX  2  2  N0  2  2  OH  OH  4'-deoxy-PNPX  4'-deoxy-4 -fluoro-PNPX ,  2  NO  Z  2  Figure 3.2: Chromogenic synthetic substrates. PNPX , 2'-deoxy-PNPX , 2'-deoxy-2'-fluoroPNPX , 3'-deoxy-PNPX , 3'-deoxy-3'-fluoro-PNPX , 4'-deoxy-PNPX , and 4'-deoxy-4'-fluoroPNPX were prepared by Dr. Johann Schloegl. 3'-Deoxy-3'-fluoro-PNPX was prepared by Dr. Chris A. Tarling. 2  2  2  2  2  2  2  2  3.2.1.1 Kinetic analysis of family 10 C.fimi xylanase Kinetic parameters (k , K , and k /K ) for hydrolysis of the /7-nitrophenyl cat  m  cat  m  deoxy- and deoxyfluoro-p-xylobiosides by wild-type Cex are presented in Table 3.1. The calculated increase in activation free energy for the glycosylation step associated with each substitution (AAG*) was calculated from the ratios of the k /K values of the parent cat  m  substrate and the modified analogue according to the equation outlined in the introduction. Values of k /K characterize the transition-state interactions between the cat  m  substrate and the enzyme, since they reflect the activation barrier from free enzyme plus substrate to the transition state for the glycosylation step of the reaction.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  105  Table 3.1: Kinetic parameters for the hydrolysis of deoxy/deoxyfluoro analogues by wild-type C.Jimi xylanase. d  PNPX  40 ± 1  0.017 ±0.001  28 ± 4  1.4 ± 0 . 2  21 ± 5  2.9 ± 0 . 1  2  nd "~  nd  74 ± 7  2.1 ± 0 . 1  3'H-PNPX  47 ± 3  0.42 ± 0 . 0 3  110 ± 1 0  1.9 ± 0.1  48 ± 2  0.024 ± 0.004  2000 ± 400  0.1 ± 0 . 1  32 ± 3  0.034 ± 0 . 0 0 1  930 ± 90  0.6 ± 0 . 1  31 ± 3  0.015 ± 0 . 0 0 1  2100 ± 2 0 0  0.006 ± 0 . 0 0 1  0.96 ± 0 . 0 5  12.9  0.11  2  2'H-PNPX 2'F-PNPX  Distal  3'F-PNPX  2  c  2  2  4'H-PNPX 4T-PNPX  2  3F-PNPX  2  2  a  DNPC  c  2300 ± 1 0 0  0  0 ± 0.1  0.006 ± 0.001  7.9 ± 0 . 1  117  Proximal AT,  Inhibitor 2F-DNPC'  (mM) 0.0011  0.11  km (s m M ) _1  0.010  -1  5.8  "Enzyme kinetics courtesy of Dr. Chris A. Tarling using Mr. David K. Y. Poon's wildtype Cex protein preparation of the catalytic domain. Kinetic parameters taken from Tull and Withers (22). nd indicates that this kinetic parameter was not determined. Graphical representation of this data is given Appendix A, page 228. Errors were estimated from a non-linear fit by GraFit 4.0 (131). b  c  d  In previous kinetic studies on Cex the rate-limiting step in the reaction of Cex with />-nitrophenyl xylobioside was confirmed to be the hydrolysis of the xylobiosyl-  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  106  enzyme intermediate; that is, the deglycosylation step. Indeed the very low K value m  observed is consistent with this conclusion, as it indicates substantial accumulation of the glycosyl-enzyme intermediate. The parameter k JK c  m  reflects the first irreversible step;  in this case, formation of the glycosyl-enzyme intermediate. Therefore, by inspecting the effects of each substitution on these two kinetic parameters (k JK c  & at C  m  for glycosylation and  for deglycosylation) we can gain insights into the importance of the interactions at  subsite -2 and -1 on each step in catalysis. 3.2.1.1.1 Distal sugar (-2) interactions Interestingly, values of & remained essentially invariant upon removal of OH cat  groups at subsite -2, where the distal sugar unit binds (Table 3.1). This indicates that the deglycosylation step is insensitive to substitutions on the distal sugar ring. Values of K  m  varied up to 90-fold for the different deoxy xylobioside analogues compared to the parent value, while values of k JK v  c  m  correspondingly suffered up to a 100-fold reduction  compared to the parent value determined for  PNPX2.  By contrast, distal substitution has  very significant effects upon the glycosylation step as shown in the k JK c  m  values, with  binding energy contributions from each hydroxyl ranging up to 2.9 kcal mol . Since -1  kcai/K is the second-order rate constant corresponding to the first irreversible step in the m  reaction (glycosylation, in this case), it is evident that removal of hydroxyl groups on 4-nitrophenyl P-xylobioside decreases the rate of formation of the glycosyl-enzyme intermediate, but has little effect on rates of its breakdown. Given the distance of these substitutions from the reaction centre, it is a reasonable assumption that electronic effects on transition-state stabilization are minimized; thus, changes are assigned entirely to hydrogen-binding interactions.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  [07  Previous studies of the apparent strengths of hydrogen bonds between enzymes and ligands have suggested that hydrogen bonds between two neutral partners contribute up to about 1.5 kcal moF to overall binding free energy, while those between one neutral 1  and one charged species are worth 3 kcal mol or more (27, 151, 152, 157). On that -1  basis, and assuming that lost hydrogen bonding is responsible for the majority of the lost binding energy, this suggests that there are one or more hydrogen bonds at each hydroxyl, probably involving charged partners at the 2'-position, a neutral partner at the 3'-position, and a weak hydrogen bond at the 4'-position of the xylopyranose unit bound in the -2 subsite. For wild-type Cex, values of AAG* of 2.9 and 1.9 kcal mol (deoxy analogues) -1  thus identify 2'-OH and 3'-OH in subsite -2 of the second sugar unit as being involved in reasonably strong hydrogen bonds at the transition state for hydrolysis of xylobiosides. As expected, a fairly small AAG* value of 0.6 kcal mol  -1  contributed by the 4'-OH in  subsite -2 is in accord with the enzyme's specificity for xylan polymers which would include a sugar link at this position. This is consistent with the structure of the 2-deoxy2-fluoroxylobiosyl-Cex intermediate (76) where it is observed that this OH-group is involved in a hydrogen-bonding interaction with a water molecule. Interestingly, values of k JK c  m  for the deoxyfluoro xylobiosides were all greater in  comparison to the analogous deoxy substrate, confirming that the enzyme donates a hydrogen bond at the substituted position in each case. Amazingly, replacement by fluorine at C-3' had no significant overall energetic consequences as expressed in the apparent binding energy contribution. This suggests that the fluorine, thus the original 3'-OH, acts as the hydrogen-bond acceptor. Conversely, had the original hydrogen bond been of the opposite polarity, requiring the fluorine to act as a donor, a considerable loss  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  108  in transition-state affinity should have resulted. A more detailed interpretation of the data is encumbered by factors such as the slightly smaller overall size of the fluorine (2.74 A) relative to the oxygen (2.83 A), the unknown relative hydrogen-bond strengths of fluorine to water and its donor in the active site, and the possible role of the fluorine in promoting other hydrogen-bonding interactions (173). The deoxygenation at position C-2' results in a larger loss in affinity (2.9 kcal mol ) relative to that observed for position C-3'. Fluorination can partially -1  reinstate this binding energy, although the fluorosugar still binds 2.1 kcalmol more -1  weakly at the transition state than the parent substrate. A straightforward interpretation of the data implies that the 2'-OH acts as a hydrogen-bond acceptor since the 2'-deoxyfluoro sugar binds slightly better than the 2'-deoxy sugar. Further, an additional hydrogen bond of opposite polarity is suggested in which the sugar hydroxyl acts as the donor given the significant loss in binding energy at the 2'-position. Clearly, this analysis does not eliminate the possibility that sugar hydroxyl groups play dual roles as hydrogenbond donors and acceptors. Consequently, this method cannot detect the presence of multiple hydrogen-bonding interactions with a particular hydroxyl on the sugar ring. The observed order of hydrogen-bond strength and the role, as donor (D) or acceptor (A), of the sugar hydroxyl at each position in subsite -2 of the second sugar unit is summarized below. 2'(D +A)>3'(A)>4'(A) The role of the sugar hydroxyl as donor or acceptor at each position is summarized below in Figure 3.3.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  109  Charged  Neutral  N0  |  2  t  Charged Charged  Neutral  Figure 3.3: Schematic representation of proposed intermolecular hydrogen-bond interactions between 4-nitrophenyl (3-xylobioside and Cex based on substrate analogue data, X-ray crystallography and NMR studies.  The polarities of the hydrogen-bonding interactions predicted from the substrate analogue data are indeed consistent with the polarities of the hydrogen bonds observed in the active site of the xylobiosyl-enzyme complex observed by X-ray crystallography (Figure 3.6). A summary of the hydrogen-bonding interactions in the active site of the fluoroxylobiosyl-enzyme complex of Cex is given in Table 3.2.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  110  Table 3.2: Summary of hydrogen-bonding interactions between the xylobiosyl intermediate and protein residues in the active site of the 2-deoxy2-fluoroxylobiosyl-enzyme complex of Cex. Sugar Ring  Atom Involved  Role  Protein 5* Residue  Atom i Involved  Proximal  2-F ^ 3-OH 3-OH  Acceptor Donor Acceptor  Asnl26 His80 Lys47  N82 " Ns2 NC  Donor Acceptor Acceptor Acceptor Acceptor  Glu43 Trp273 Asn44 Gln87 Lys47  0E2  Distal,  2'-OH 2'-OH 3'-OH 5'-0 5'-0  Nel .. N82 : Ns2 NC  Role  Distance (± 0.2 A)  Donor Acceptor 'Donor  3.3 3.0 2.8  Acceptor Donor Donor Donor Donor  2.9 2.9 ,3.0 3.4 3.0  3.2.1.1.2 Proximal sugar (-1) interactions Unfortunately only a very limited data set is available for xylobioside substrates in which the proximal sugar has been modified, and, as noted earlier, these data are complicated by inductive effects upon transition-state stability. However comparison of the data for  3F-PNPX2  with that for  PNPX2  reveals that, indeed, both k and cat  k /K cat  m  are  severely affected, with the transition state for glycosylation being destabilized by a massive 7.9 kcal mol , while that for deglycosylation is destabilized by approximately -1  5.2 kcal mol . Part of the rate reduction arises from inductive effects, which can be -1  estimated from spontaneous hydrolysis data. In a study by Namchuk et al. (18) on the role of sugar substituents in spontaneous glycoside hydrolysis, the electronic effect of the ring substituent on the principal center of charge development at the transition state was the primary determinant of the rate of hydrolysis of a series of monosubstituted deoxy and deoxyfluoro 2,4-dinitrophenyl P-D-glycopyranosides.  The ratios of rates of  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  111  spontaneous hydrolysis obtained at the 3-position for F : OH : H substituted glucoses are 0.14 : 1 : 4 respectively; therefore, the inductive effect at the 3-position is relatively small.  A straightforward interpretation of the data therefore implies that the 3-OH  engages in multiple strong hydrogen-bonding interactions at the transition state, summing to  approximately  6.7 kcal mol  -1  and  4.0 kcal mol  -1  to the  glycosylation and  deglycosylation steps. Likewise, but difficult to evaluate, enormous effects upon k  cat  are seen for  substitution of the 2-hydroxyl in the proximal ring by fluorine. Indeed, the substantial decrease in reaction rate due to destabilization df the oxocarbenium ion-like transition state for the enzymatic hydrolysis of glycosides has been the basis for the use of such reagents as mechanism-based inactivators. In fact, 2,4-dinitrophenyl 2-deoxy-2-fluorocellobioside was shown to be^ an excellent time-dependent inactivator for Cex, functioning via the formation and accumulation of a relatively stable 2-deoxy-2-fluorocellobiosyl-enzyme intermediate (22). Unfortunately, data are not available to allow direct comparisons of aryl xylobiosides with their 2-deoxy or 2-deoxy-2-fluoro analogues. However, as shown in Table 3.1, inactivation parameters of k\ - 0.0011 s  _1  and K\ = Q).\ \ mM for 2,4-dinitrophenyl 2-deoxy-2-fluoro-[j-cellobioside, were obtained by Tull and Withers (22) according to the scheme below: E + Ir-^EI—^->E-I This scheme is analogous to that of the Michaelis-Menten mechanism. By comparing the inactivation parameter kJK\ for the modified "substrate" (2F-DNPC) to the k JK Q  m  value obtained for the parent 2,4-dinitrophenyl cellobioside (DNPC), a transition-  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  772  state stabilization energy of at least 5.8 kcal mol contributed by the 2-hydroxyl was -1  calculated for the glycosylation step. However, if the fluorine is involved in stabilizing interactions as a hydrogen-bond acceptor, the AAG* value obtained will be an underestimate. Effects on the deglycosylation step are much greater. Comparison of k  cat  for DNPC (reflecting the deglycosylation step) with the reactivation rate constant for the 2-fluorocellobiosyl-enzyme (22) results in a AAG* value of 11.3 kcalmol , of which -1  part is undoubtedly due to inductive effects.  For (non-enzymic) acid-catalyzed  hydrolysis, the ratios of rates obtained at the 2-position for F : OH : H substituted glycosides are 0.02 : 1 : 2000 respectively. The AAG* as a consequence of fluorination at the 2-position for the non-enzymic reaction is 2.4 kcal mol . -1  After applying this  correction, a AAG* value of approximately 9 kcal mol can be conservatively estimated -1  for the contribution of the interactions at the 2-hydroxyl.  3.2.1.2 Kinetic analysis of family 11 B. circulans xylanase Kinetic parameters for the hydrolysis of the deoxy and deoxyfluoro derivatives of 4-nitrophenyl p-xylobioside by wild-type Bex were determined, and the changes in activation free energy, AAG* for the glycosylation step associated with each substitution, are presented in Table 3.3. Values of k JK c  for the modified glycosides are one to three  m  orders of magnitude smaller than those for the parent substrate, PNPX2. Due to the high K values exhibited by Bex for aryl xylobioside substrates, and the scarcity and low m  solubility of the modified xylobiosides, full Michaelis-Menten kinetic parameters could not be determined. Instead, values of k  /K  cat  substrate depletion method.  m  for each substrate were determined by the  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  113  Table 3.3: Kinetic parameters for the hydrolysis of deoxy/deoxyfluoro analogues of P N P X 2 by wildtype B. circulans xylanase. Apparent Binding Energy Contribution  Substrate Depletion Kinetics  Substrate  AAG* (kcal mol )  (s mM ) 1  -1  -1  0.13 ±0.01  0  2  0.0012 ±0.0001 0.0035 ± 0.0004  2.9 ±0.1 2.2 ±0.1  3'H-PNPX 3T-PNPX  0.0046 ± 0.0005 0.050 ±0.005  2.1 ±0.1 0.6 ±0.1  PNPX  2  2'H-PNPX 2'F-PNPX  2  2  2  4'H-PNPX 4'F-PNPX  2  2  0.34 ±0.03 0.41 ±0.04  -0.6 ±0.1 -0.7 ±0.1  The values of AAG* for the deoxy sugars listed in Table 3.3 represent estimates of the loss of binding energy at each position at the transition state resulting from removal of the hydroxyl in question, since such substitution has resulted in the observed destabilization of the transition state. Very similar effects to those observed for Cex upon kcJKm are seen for Bex, with AAG+ values ranging from -0.7 up to 2.9 kcal moF (Table 1  3.3). Unfortunately, due to the relatively high K values exhibited by Bex and the m  scarcity of substrate it was not possible to obtain k values to probe effects upon the cat  deglycosylation step. The relative order of hydrogen-bond strengths measured for Bex in the -2 subsite is similar to that observed in the binding of the same substrates to wildtype Cex where position 2' > 3' > 4'. In the case of Bex, the strongest sum of hydrogenbond strengths measured in the -2 subsite was that at the 2'-position (2.9 kcal mol ), -1  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  114  with a slightly weaker interaction measured at the 3'-position (2.1 kcal mol ). -1  The  interaction at position-4' appears to play little role in substrate binding at the transition state, and even to be slightly destabilizing (-0.6 kcal mol ). This result is consistent -1  with this enzyme's specificity for an extended (3-1,4-linked polysaccharide substrate xylan  (174).  In studies of oligosaccharyl fluoride substrates for human pancreatic  a-amylase, similar results were obtained. Substrates in which the 4-hydroxyl group at the non-reducing end of the oligosaccharide were methylated were consistently better substrates, presumably because there were no solution-phase hydrogen-bonding partners that would be left unsatisfied at the transition state (175). Although the data obtained for Bex is less extensive, the same pattern of hydrogen-bond polarities and strengths once again emerges between the crystallographic and the kinetic data. The interactions between the protein side-chains and the 2-fluoroxylobioside moiety in the 2F-xylobiosyl-Bcx complex (Figure 3.4) were discussed extensively by Sidhu et al. (81).  Most notably, the distal xylose residue is maintained in the  conventional C\ (chair) conformation and forms hydrogen bonds to Tyrl66, Gln7, and 4  Tyr69.  The proximal xylose residue is distorted to a ' B (boat) conformation and 2 5  extensively hydrogen bonds to Tyr69, Glu78, Argll2, and the backbone carbonyl of Pro116.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  115  Figure 3.4: Scheme of protein-carbohydrate interactions in the active site region of the 2-fluoroxylobiosyl-Bcx complex observed by three-dimensional structural analysis (81). Hydrogen-bonding interactions between heteroatoms are in Angstroms.  Interactions with Tyr69 are of particular interest (Figure 3.4). In the structure of the covalent intermediate, the hydroxyl group of Tyr69 donates a hydrogen bond to the OE2 atom of Glu78 and accepts a hydrogen bond from the 2'-position O H of the distal (nonreducing) xylose ring. This latter hydrogen bond to 2 ' - O H helps to position the substrate, whereas the former may help orient the nucleophile for the reaction. Furthermore, the phenolic oxygen of Tyr69 also donates a hydrogen bond to the endocyclic oxygen (0-5) of the proximal xylose residue to form a bifurcated hydrogen bond. Deoxygenation at the 2'-position results in a larger loss in affinity (2.9 kcal mol ) -1  relative to that observed for the 3'-position (2.1 kcal mol ). The single hydrogen bond -1  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  U6  from Tyrl66 to the 3'-position OH of the second xylose moiety therefore appears to provide less transition-state stabilization than those at the 2'-OH. Although fluorination can partially reinstate binding energy at both positions, the 2'-fluorinated compound still binds 2.2 kcal mol  -1  more weakly than the parent substrate at the transition state. A  simple analysis of the data suggests that the 2'-OH acts as a hydrogen-bond acceptor in at least one interaction since the 2'-deoxyfluoro sugar binds slightly better than the 2'-deoxy sugar. Indeed, structural analysis shows that the 2'-OH plays dual roles as acceptor and donor with Tyrl66 and Tyr69, respectively. Furthermore, the hydroxyl at the 3'-position likely acts as a hydrogen-bond acceptor with Tyrl66 since the fluorine at C-3' is unable to act as a hydrogen-bond donor and a considerable loss in affinity should result. Accordingly, fluorination at the 3'-position had relatively minor overall energetic consequences as expressed in the apparent binding energy contribution (0.6 kcal mol ). -1  Although the data acquired are limited to interactions that occur in the -2 subsite of the Bex binding site, significant interactions in the -1 subsite contribute greatly to transition-state stabilization. An important interaction is that involving the Osl of Glu78 and the 2-substituent of the proximal sugar. Despite the fact that the interaction between the fluorine and the oxygen in the covalent complex must be destabilizing, the distance between these two atoms is shorter than that seen for the analogous yet stabilizing hydrogen-bonding interaction in the non-covalent complex of Bex with xylobiose. The shortening of this distance was suggested to be a consequence of the formation of the covalent bond at the anomeric center and the conformational change in the proximal saccharide. Similar interactions in other 2-fluorosugar glycosyl-enzyme complexes (32, 33, 76, 176) have suggested that short, strong hydrogen bonds are formed at this position  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  7/7  in the glycosyl-enzyme intermediate and that this hydrogen bond may be even shorter and stronger at the transition state. This is consistent with the very strong transition-state interactions (typically 5-10 kcalmol ) measured at the 2-position for a number of -1  glycosidases.  Another residue interacting with the 2-position fluorine in the  2F-xylobiosyl-Bcx complex is the highly conserved Argl 12. Although an important role has been proposed for the equivalent residue in Cex (Asnl26) in hydrogen bonding to the 2-hydroxyl of natural substrate (33), the mutation of Argl 12 in Bex causes a relatively small decrease in catalytic activity (119). Therefore, the most important interaction with the 2-substituent appears to be between the 2-position of the proximal sugar and Osl of Glu78. In summary, these data permit some quantification of the contribution of interactions at each individual hydroxyl group in the -2 subsite to catalysis by Bex. Atomic resolution crystallographic observation of such interactions would be useful in light of the many studies regarding the fluorosugar-protein interactions where such hydrogen bonding has been cited to explain the high affinity for deoxyfluoro derivatives as compared to deoxy derivatives. The similarities and differences between Cex and Bex, two xylanases from distinct families, will be discussed in greater detail in the following section.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  118  Table 3.4: Summary of hydrogen-bonding interactions between the xylobiosyl intermediate and protein residues in the active site of the 2-deoxy2-fluoroxylobiosyl-enzyme complex of Bex (81). Sugar Ring  Atom Involved  Proximal  Distal  Role"  Protein Residue  Atom Involved  2-F 2-F 3-OH 3-OH 5-0  Acceptor Acceptor Acceptor Donor Acceptor  Glu78 Argl12 Argl12 Proll6 Tyr69  Osl Ns NH 0 OH  2'-OH 2'-0H 2'-0H 3'-0H  Acceptor Donor Acceptor Acceptor  Gln7 Tyr69 Tyrl66 Tyrl66  Ns2 .; OH OH OH  2  Role  Distance (±0.2 A)  Acceptor* Donor Donor Acceptor Donor *  2.8 3.2  Donor Acceptor Donor Donor  3.1 2.8 3.0 2.9  2.7  Ligand and protein roles were derived with LPC software (177). The interaction between the fluorine and the oxygen in the covalent complex is destabilizing (81).  a  b  3.2.1.3 Comparison of results for two xylanases from distinct families Kinetic analyses from the previous two sections may be used to reveal information on active site similarities between two unrelated xylanases from two structurally distinct families. Linear free energy relationship (LFER) analysis has been used to show that two highly conserved a-glucan phosphorylases have essentially identical transition-state interactions  (178).  Similarly, LFER analysis has been used to  probe the active site similarities of three enzymes (179) acting on glucose or glucosidic substrates: glucose dehydrogenase (EC 1.1.1.47), glucose oxidase (EC 1.1.3.4), and glucoamylase (EC 3.2.1.3). Despite differences in specificity and structure, the high linear correlation coefficients obtained between AAG* values for a series of deoxygenated substrates with these enzymes indicate  significant  similarity in transition-state  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  119  interactions. Similarly, the transition-state binding interactions of Cex and Bex can be compared using LFER by plotting AAG* values for one enzyme versus the corresponding values for another (7 78, 179). An LFER with slope and correlation coefficient (p) close to 1 may suggest two enzymes with very similar active-site transition-state interactions, whereas unrelated enzymes would have very poor correlation coefficients. Therefore, this method reflects how well the binding energetics for different enzymes resemble each other. The specificity constants, k /K , determined for Cex and Bex with the different cat  m  substrate analogues are given in Table 3.1 and Table 3.3. Changes in activation energy (AAG*) due to substitution of individual hydroxyl groups are given in Table 3.5. Assuming that removal of the individual hydroxyl groups does not lead to a change in mechanism or rate-limiting step, the measured AAG* values reflect changes in hydrogenbonding interactions. The calculated AAG* values for the two enzymes (Table 3.5) show a very similar pattern in the involvement of the substrate hydroxyl groups for transitionstate stabilization. The activities of the two enzymes were found to depend heavily on the 2'-OH group, where a change in binding energy of 2.9 kcal mol  -1  suggested strong  hydrogen bonds between the enzymes and this substrate hydroxyl group. The 3'-OH was involved in weaker interactions, while the 4'-OH had a very small if any role in transition-state binding.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  120  Table 3.5: Binding energy contribution AAG* (kcal mol ) of individual hydroxyl groups in the hydrolysis of P N P X 2 by Cex and Bex. -1  Cex Substrate  Bex  AAG* (kcal mol )  AAG* (kcal mol )  -1  PNPX  -1  0  2  2'H-PNPX 2T-PNPX  2  2  3'H-PNPX 3T-PNPX  2  2  4'H-PNPX 4T-PNPX  2  2  2.9 ±0.1 ,^2.1*0.1 : 1.9 ± 0.1 0.1 ±0.1 0.6 ±0.1 0±0.1  0  v  2.9 ±0.1 2.2 ±0.1 2.1 ±0.1 0.6 ± f 1  * :  -0.6 ±0.1 -0.7 ±0:i  LFER plots of two evolutionarily related phosphorylases were previously reported to have a slope of 1.0 and p of 0.999 (178). Cex and Bex have no overall sequence homology, yet a LFER plot (Figure 3.5) yields a correlation coefficient of 0.92 and a slope of 1.1.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  -  1  1  0  2  121  4  3  Cex AAG*(kcal mol- ) 1  Figure 3.5: Binding energy contribution AAG* (kcalmol ') plot for a series of deoxy- and deoxyfluoro-P-xylobioside analogues with B. circulans xylanase (Bex) and C. fimi xylanase (Cex).  Such a strong correlation suggests the two enzymes possess similar transitionstate binding interactions, at least those interactions present in the -2 subsite. This result is surprising in light of the differences that exist between these two enzymes. A careful comparison of xylanases in 1988 by Wong et al. (94) indicated that these enzymes can be divided into two groups: one consisting of high molecular mass enzymes with low pi values, and the other consisting of low molecular mass enzymes with high pi values. Acidic high molecular mass xylanases (>30 kDa) were found to belong to one family, assigned as glycanase family 10, and basic low molecular mass xylanases, were found to belong to another family, designated as glycanase family 11 (4-6, 120, 180, 181). Family 10  xylanases,  such  as  Cex,  show  higher  affinity  for  shorter  linear  P-l,4-xylooligosaccharides than xylanases of family 11, such as Bex (182). Additionally, enzymes belonging to family 10 exhibit greater catalytic versatility or lower substrate specificity than enzymes of family 11.  Significant differences exist in the overall  architecture of the substrate binding sites: the substrate binding sites of the family 10  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  122  xylanases apparently consist of more shallow clefts versus the substrate binding sites of family 11 enzymes. Three-dimensional analysis of the binding sites of Cex and Bex in complex with a covalent xylobiopyranosyl residue may be utilized to investigate a basis for the transition-state interactions. In the structure of Cex, a glutamate and tryptophan residue interacts with the 2'-hydroxyl group of the substrate; in Bex, two tyrosine residues are positioned to hydrogen bond with the 2'-hydroxyl group. An asparagine residue in the -2 subsite of Cex hydrogen bonds with the 3'-hydroxyl group, while a tyrosine residue in Bex fulfills the same role. Although the properties of the amino acid residues of the two enzymes that interact with the hydroxyl groups are very different, very similar AAG* were obtained for both the deoxy and deoxyfluoro derivatives of the xylobioside substrate. The greatest divergence from the observed trend in Figure 3.5 was detected for the 4'-deoxy and 4'-deoxyfluoro derivatives, where it was observed that the 4'-hydroxyl group destabilizes the transition state to a greater extent in Bex than in Cex. This is consistent with the observation that family 10 xylanases show higher affinity for shorter linear (3-1,4-xylooligosaccharides than xylanases of family 11.  Further  examination and comparison of the "energetic fingerprint" of other glycosidases would be of interest. In summary, the energetics of transition-state interactions of the active sites of Cex and Bex have been probed using molecular recognition to indicate key roles for the 2'-OH and 3'-OH groups of the distal xylobiosyl residue. The LFER plot points out similar transition-state interactions despite diverse active-site architectures.  123 Chapter  3  Binding  Energy  Contribution  of Hydrogen  Bonds  3.2.2 Binding energy contributions probed by using mutant enzymes  Figure 3.6: Scheme of protein-carbohydrate interactions in the active site region of 2F-xylobiosyl-Cex-cd complex observed by three-dimensional structural analysis (76). Hydrogen-bonding interactions between heteroatoms are in Angstroms. The distance between 2F and the carbonyl oxygen of Glu233 is 2.6 A (not shown).  Michaelis-Menten kinetic parameters for the hydrolysis of p-nitrophenyl xylobioside by each of the mutants of Cex are presented in Table 3.6. All the mutants were found to be less active than the wild-type enzyme in terms of k /K cat  m  values. K  m  values are higher than for the wild-type enzyme, consistent with the involvement of the mutated residues in substrate binding. Again, the change in activation energy for the first (glycosylation) step associated with each mutation, calculated from relative  k JK c  m  values, is presented. Evaluation of a series of deoxygenated and deoxyfluoro xylobioside substrate analogues with both wild-type and mutant versions of Cex allows an assessment  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  \24_  of the contributions of transition-state stabilizing interactions involving hydrogen-bond donors/acceptors at the sites that are modified by mutagenesis. For mutant Cex, k JK c  m  values decreased by as much as 5 x 10 -fold relative to wild-type. 4  3.2.2.1 Design of the mutations  Figure 3.6 shows the putative protein-carbohydrate interactions that are proposed according to the X-ray structure of the glycosyl-enzyme intermediate (32, 33, 76). For subsite - 1 , the hydrogen-bonding interactions shown might not fully represent those formed at the transition state since ring distortion is expected for the covalently bound xylopyranose unit bearing the glycosidic bond to be hydrolyzed.  The protein-  carbohydrate contacts observed in the covalent fluoroxylobiosyl-Cex complex involve Lys47, His80 and Asnl26, but other interactions may hold if ring distortion occurs upon binding. Different mutants have been designed to evaluate hydrogen bonding.  For  hydrogen-bonding residues (Glu, Asn, Lys, His, and Gin), mutation to Ala will remove the interaction, and other mutations such as His —> Asn and His —• Gin will conserve elements of the functional side-chain group but modify or remove the interaction as a consequence of the larger or shorter side chain. Mutagenesis was performed using the QuikChange system as described in Materials and Methods and the mutants of Cex were characterized in Chapter 2. The Q87M and N126A mutants were described earlier (76, 183).  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  725  3.2.2.2 Subsite - 2 interactions  The kinetic parameters determined for hydrolysis of  PNPX2  by the mutants of  Cex are given in Table 3.6. When hydrogen bonds that are exclusively formed with the distal sugar  are destroyed (E43A; N 4 4 A ; Q87M) the k values for hydrolysis of PNPX  are essentially unaffected (Table  cat  3.6).  However, the effects upon k JK c  m  2  are substantial,  ranging from 1.0 to 3.7 kcal mol . The results indicate that hydrogen bonds formed with -1  the distal sugar are important for the glycosylation transition state, but not for the deglycosylation transition state.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  126  Table 3.6: Kinetic parameters for the hydrolysis of P N P X 2 by wild-type and mutant C. fimi xylanase."  • *  -2  -1  Native Cex  40 ± 1  Cex E43A  33 + 3  5± 1  Cex N44A  110 ± 10  4± 1  Cex Q87M  58 = 2  „  0.017 ±0.001 2 3 0 0 ± 1 3 0 ~  6.0 ± 1  0.12 ± 0 . 0 1  Cex K47A  0.43 ± 0.04  Cex H80A  0.50 ± 0.04  0.59 ± 0.07  Cex H80N  0.15 ± 0 . 0 1  0.099 ± 0.003  Cex H80Q  0.044 ± 0.002  0.061 ± 0 . 0 0 3  CexN126A  3.5 ± 0 . 3  10± 1  8±1  2100  29 ± 6  31  480 ± 40  nd"  0.045 ±0:005 0.86 ± 0 . 1 2  :  3.7 + 0.1  ]  2.7 ± 0 . 1 1.0 + 0 . 1 ;  0.054  6.7 ± 0 . 1  0.80  4.9 + 0.1  1.5 ± 0 . 1  nd*  0.72 ± 0.05  nd"  0.45 ± 0 . 0 7  0  0.49  4.5 ± 0 . 1  ;  5.0 + 0.1 ~ 5.3 = 0.1  Graphical representation of this data is given Appendix A, page 228. Errors were estimated from a non-linear fit by GraFit 4.0 (131). nd indicates that this kinetic parameter was not determined.  a  b  It is of interest to correlate the nature of the enzyme-substrate interactions, as observed crystallographically, with their strengths as, observed kinetically for the glycosylation transition state in this study.  The distal sugar at subsite -2 contributes  through hydrogen bonding with Glu43, Asn44 arid Lys47. Significantly, the strongest interactions with the distal sugar are those at the 2'-position, consistent with a hydrogen bond involving a potentially charged residue (Glu43). Since the 2'-hydroxyl must donate the hydrogen bond to the carboxylate of E43, the loss of almost all the interaction energy  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  727  in the 2'-fluorosugar case (2.1 kcal mol" ) is also understandable, since fluorine cannot 1  donate a hydrogen bond. It could however, still accept a hydrogen bond, if more weakly, from W272, hence the lesser loss of interaction energy than was the case for the 2'deoxy sugar (167-170). The mutation of Glu43 to Ala decreases k only minimally, but results cat  in a 300-fold increase in K . m  For the analogous 2'-deoxygenated and 2'-fluorinated  substrates, losses of 5.1 kcal mol and 3.4 kcalmol (Table 3.7), respectively, relative to -1  -1  the wild-type enzyme were measured. The additional destabilization due to removal of the 2'-OH group on the distal xylobiose sugar unit may be due to the loss of the interaction between the 2'-hydroxyl and the Trp273 residue worth 1.4 kcalmol . -1  Removal of the acidic group may also influence other charged binding side chains contributing to destabilization of the complex.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  128  Table 3.7: Apparent binding energy contribution, AAG* (kcal mol ), of individual substrate hydroxyl groups in the hydrolysis of deoxy/deoxyfluoro analogues of P N P X 2 by wild-type C.fimi xylanase and selected mutants -1  Apparent Binding Energy Contribution AAG* (kcal mol") 1  Substrate E43A PNPX  3.7 ± 0 . 1  2  2'H-PNPX 2T-PNPX  3T-PNPX  2.7 ± 0 . 1  0 2.9 ± 0 . 1  3.4 ± 0 . 1  nd  2.1 ± 0 . 1  a  nd nd  2  • ,WT4>;,  nd*  nd* 2  N44A  5.1 ± 0 . 1  nd  2  2  4'H-PNPX 4T-PNPX  2  2  3'H-PNPX  a  •  a  3.1 ± 0 . 1  1.9 ± 0.1  2.7 ± 0 . 1  0.1 ± 0 . 1  nd  fl  nd  a  0.6 ± 0 . 1  fl  0±0.1  a  nd indicates that this kinetic parameter was not determined.  Asn44 forms a bidentate hydrogen-bond pattern with the xylosyl 3'-OH and the NC, atom of the Lys47 side chain at subsite -2 in the crystal structure of Cex (76) (Figure 3.6) with a contribution of 2.7 kcal m o l to transition-state stabilization (Table 3.7). For -1  the analogous 3'-deoxygenated and 3'-fluorinated substrates, losses of 3.1 kcalmol and -1  2.7 kcalmol , respectively, relative to the wild-type enzyme were measured (Table 3.7). -1  An apparent binding energy contribution of 1.9 kcalmol measured for the 3'-OH group -1  (Table 3.1) indicates that the additional loss of interaction energy caused by mutation at Asn44 may be due to the disruption of the interaction of the amide carbonyl with the charged K47. No significant hydrogen-bonding interactions are seen at the 4'-position of  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  ;  12R  the distal sugar since further sugar residues are normally appended at this site in the intact xylan substrate. Again, consistently, the deoxy and' deoxyfluorosugars are turned over at essentially the same rates as the parent species. 3.2.2.3 Subsite -1 interactions  A different model emerges when considering hydrogen bonds formed with the proximal sugar. In all mutants in which side chains interacting with the proximal sugar are modified (K47A; H80A; H80N; H80Q; N126A), values of both £ , and k JK ca  (Table 3.6).  The largest effect is upon k JK , c  m  c  m  drop  with AAG* values ranging from  4.5-6.7 kcal mol" . 1  The particularly important transition-state hydrogen-bonding interactions at the 3-position of the proximal sugar arise from a pair of charged hydrogen-bonding interactions.  The 3-OH engages in strong hydrogen-bonding interactions with a  positively charged Lys47 residue, recently unequivocally shown to be charged by 15  N-NMR analysis of the trapped intermediate (184), and His80, also recently shown by  15  N-NMR to be present in its charged, conjugate acid form in the free enzyme, but to lose  its proton upon formation of the glycosyl-enzyme intermediate (185). Mutation of His80 to Ala introduces 4.9 kcal mol  -1  (Table 3.6) overall destabilization with  PNPX2,  which  includes the elimination of the hydrogen bond between the carboxylate function of Asp 123 and the 3-OH of the proximal xylose residue. Mutating the side chain to Asn or Gin has approximately the same effect. This effect, too large for a single hydrogenbonding interaction, probably indicates an additional role of His80 in transition-state stabilization. Mutation of Lys47 to Ala introduces 6.7 kcal mol  -1  (Table 3.7) overall  destabilization with PNPX2, which includes the elimination of a hydrogen bond between  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  130  the endocyclic oxygen and the 081 atom of Asn44 at subsite -2, as well as the 3-hydroxyl group at subsite -1 (worth only 1.9 kcal mol ). This substantial effect results -1  from removal of a positively charged side chain that will likely influence other side chains interacting through a hydrogen-bond network and/or influence the pK of a  neighbouring charged groups. Again, mutation of either of these residues is seen to have drastic effects on rates, as would be expected when a charged hydrogen-bonding network such as this is destroyed. Finally, removal of interactions at the 2-position of the proximal sugar either by mutation  (N126A:  AAG* = 5.3 kcal mol ) -1  or  via  fluorine  substitution  (AAG* = 5.8 kcal mol ) slows the glycosylation step enormously. Interactions at this -1  position of the saccharide are important for transition-state stabilization in a number of glycosidases, providing up to 10 kcal mol  -1  stabilization (27, 28), but the identity of the  amino acid residues involved is not clearly established.  However, the principal  transition-state interaction at OH-2 is thought to be that with the carbonyl oxygen of the nucleophile (E233). The loss of transition-state stabilization due to the mutation of Asnl26 to Ala may also result from the loss of a hydrogen bond from Asnl26 to Asnl69, which in turn aids in positioning Glu233.  3.2.3 Uniform and differential binding interactions  Interactions with the distal (-2) sugar contribute significantly to k /K cat  m  (up to  3.7 kcal mol ), thus to the formation of the glycosyl-enzyme intermediate, but not to k -1  caU  thus to the hydrolysis of the glycosyl-enzyme. Interactions with the proximal (-1) sugar are much more substantial, contributing up to 6.7 kcal mol" to both k /K and k . The 1  cat  m  cat  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  1_3J_  implication of these results, as is illustrated in Figure 3.7, is that interactions with the distal sugar maintain similar magnitudes in the transition states for glycosylation and deglycosylation, as well as in the intervening glycosyl-enzyme intermediate.  These  interactions can thus be referred to as "uniform binding interactions" in the parlance of Albery and Knowles (135, 136).  They form in the first Michaelis complex, and are  maintained with equal strength in all the intervening states along the reaction coordinate. Such interactions serve to improve k JK , c  A: t. ca  m  but have no positive effect whatsoever upon  By contrast, interactions with the proximal sugar are considerably stronger at the  two transition states than in the intermediate, and thus fall into the category of "differential binding interactions" of Albery and Knowles (135, 136). Interactions of this class are harder to recruit as they must necessarily exploit the bonding arrangements and charge distributions present at the transition state that differ from those in the ground state in order to achieve this selectivity.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  [32.  Distal sugar interactions: uniform binding  G  Proximal sugar interactions: differential binding  G  Figure 3.7: Reaction coordinate diagram showing modifications to the (a) distal sugar and (b) proximal sugar.  the  consequences  These observations may be explained by the substantial  of  conformational  rearrangement that occurs in the proximal sugar to accommodate oxocarbenium ion centre formed at the transition state, most likely via a  the planar half-chair  conformation in the case of Cex (186). Interactions will be optimal at this transition-state conformation. By contrast, no conformational changes are expected at the distal sugar. It therefore appears that Cex recruits both uniform binding interactions (at the distal sugar) and differential binding interactions (at the proximal sugar) in effecting catalysis on oligosaccharides.  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  133  Interestingly, some clues of this behaviour had been suggested earlier, in a study of the mechanisms of hydrolysis of aryl cellobiosides and aryl glucosides by Cex (22). The rate-limiting step for aryl glucosides was the glycosylation step, while that for aryl cellobiosides was deglycosylation, as revealed by Bransted plot analyses. Therefore, the presence of an extra glucose moiety appears to selectively accelerate the glycosylation step, consistent with the reaction coordinate diagrams in Figure 3.7A. Uniform lowering of the energy levels of the two transition states and the intermediate will change the ratelimiting step from glycosylation to deglycosylation. Furthermore, examination of the slopes of the two Brensted plots derived from  k JK c  m  values, which reveal the charge  development on the aglycone oxygen at the glycosylation transition state in each case, provide further insight. A slope of -1 for the  glucosides  development, while a slope of -0.3 for the  indicated full negative charge  cellobiosides  revealed less charge  development when both subsites were occupied. This likely results from a greater degree of nucleophilic pre-association by the nucleophile (Glu233) and/or a greater extent of proton donation from the acid catalyst (Glu 127).  The finding of slightly lower  a-deuterium kinetic isotope effects measured for the cellobiosides is consistent with greater pre-association as at least one of the contributing factors.  3.3 Conclusions Molecular recognition using substrate analogues has successfully been used to identify and quantify interactions between various functional groups and modified sugar substrates in Cex and Bex. While hydrogen bonds at the 2'- and 3'-positions of the distal saccharide aid in stabilizing the oxocarbenium ion-like transition state for glycosylation,  Chapter 3  Binding Energy Contribution of Hydrogen Bonds  [34  the strongest stabilization energies were those measured for hydroxyl groups of the proximal saccharide in subsite -1 of Cex. This is in accord with the mechanism for Cex and Bex, where no distortion of the distal sugar ring occurs, whereas considerable distortion is expected for the proximal sugar unit. Furthermore, a remarkably good agreement on the contributions of specific hydrogen bonding interactions to catalysis by two different xylanases was demonstrated using two different approaches.  Kinetic  analyses with a series of 8 mutants of Cex, in which active-site residues interacting with the substrate were mutated, yielded complementary insights on the role of amino acid residues in the binding cleft and the magnitude of protein-substrate interactions. Interaction strengths measured were consistent with the nature of the hydrogen bonds involved: charged or neutral. In those enzyme variants where the ionization properties at the active site were potentially altered (i.e., the mutations of Glu43 and Lys47), evidence of disturbances of hydrogen-bonding interactions with hydroxyl groups at subsite -2 and -1 were observed. The change in pK values resulting from the change in local charge a  may account for these disturbances. The strongest bond energies were measured in subsite - 1 , suggesting that these interactions are critical for substrate binding and bond hydrolysis.  Particularly important was the revelation that both 'uniform' and  'differential' binding interactions are recruited in the active site of a single enzyme.  CHAPTER 4  INQUIRY INTO T H E TRANSITION-STATE A N A L O G Y O F A SERIES OF IMINOSUGAR INHIBITORS F O R A XYLANASE*  * A version of this chapter has been published. Wicki, J., Williams, S. J., and Withers, S. G . (2007) Transition-state mimicry by glycosidase inhibitors: a critical kinetic analysis. Journal of the American Chemical Society, 129: 4530-4531.  Chapter 4  4  Transition-State Analogy of a Series of Iminosugar Inhibitors  136.  Inquiry Into the Transition-State Analogy of a Series of Iminosugar Inhibitors for a Xylanase  4.1 Introduction Transition-state analogues have proved to be an extremely important class of enzyme inhibitors, not only as therapeutic agents, but also as probes of the enzyme mechanism itself.  Before applying this investigative tool to discern pertinent  fundamental features of the transition state, an enzyme inhibitor must be determined to be a transition-state mimic rather than a fortuitously binding inhibitor. Therefore, the criteria used for designating enzyme inhibitors as transition-state analogues will be reviewed in this section.  4.1.1 Transition-state theory  Transition states are the balance point of catalysis where bonds are partially made and/or broken. The potential energy surface in a typical chemical reaction consists of two distinct regions corresponding to the reactant(s) and product(s) that are separated by an energy barrier at a saddle point on the surface. A common representation of the reaction kinetics passing through the activation barrier is given by the reaction coordinate diagram (Figure 4.1).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  137  Transition state  e c  <D  « e B  o a.  Reactants / Products  Reaction coordinate  Figure 4.1: A reaction profile. The horizontal axis is the reaction coordinate and the vertical axis is the potential or Gibbs free energy. The transition state corresponds to the maximum of the Gibbs free energy.  The structure represented by this saddle point is the activated complex although the term transition state is often applied. Transition-state theory originated in the 1930s with the work of Eyring (187). The theory makes two basic assumptions: (1) that there is a thermodynamic equilibrium between the transition state and the state of reactants, and (2) that the rate of a chemical reaction is proportional to the concentration of the particle(s) in the high-energy transition state. These assumptions lead to the Eyring equation for the rate constant A: of a chemical reaction (Equation 4.1).  k = K^eh  AGt/RT  (4.1)  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  [38  In this equation K is the transmission coefficient which defines the fraction of the molecules that go to the product state when they have reached the transition state, k is D  Boltzmann's constant, h is Plank's constant, T is the absolute temperature, and AG** is the Gibbs free energy of activation. This is the difference between the Gibbs energy of the transition state and the Gibbs energy of the reactants. In fact, the Eyring equation provides theoretical validation of the empirically derived relationship by Arrhenius, which states that: (4.2)  k = Ae-^  jRr  where A is the pre-exponential factor, E is the activation energy of the reaction, R a  is the gas constant, and T is the absolute temperature.  4.1.2 E n z y m e catalysis  Enzymes accelerate chemical reactions by stabilizing the transition-state structure of the substrate relative to that of the ground state, and often involve alternative pathways from that of the non-catalyzed reaction (29, 188-194). Four common mechanisms by which an enzyme might be expected to bring about rate acceleration include approximation of the reactants, covalent catalysis, general acid/base catalysis, and the induction of distortion or strain in the substrate, the enzyme, or both. Utilizing these mechanisms enzymes lower the energy of the transition state, thereby allowing a greater population of the starting material to attain the energy needed to overcome the activation energy and proceed to product.  Chapter 4  Transition-State Analogy of a Series of lminosuzar Inhibitors  139  Ci  c  o 15 0)  o E+S E+P  Reaction coordinate  Figure 4.2: Reaction coordinate diagram to show how an enzyme can increase the rate of reaction of a bound substrate by stabilizing the transition state.  Transition-state theory as applied to enzymatic catalysis provides an approach to understanding enzymatic catalysis in terms of the factors that determine the strength of binding of ligands to proteins. The prediction that the transition state should bind to the enzyme much more tightly than does the substrate is supported by experimental results with stable analogues of transition states (29, 188-194). Transition-state analogues have great potential for use in understanding enzymatic catalysis and are discussed in greater detail in the following section.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  [40  4.1.3 Experimental approaches to enzymatic transition-state structure 4.1.3.1 K I E to determine T S structure  Kinetic isotope effects (KIE) permit experimental access to the TS structure and are perhaps the only way to obtain detailed TS information. The defining features of enzymatic transition states (bond-lengths, bond-angles and molecular electrostatic potential surfaces) can be approached experimentally by the measurement of intrinsic kinetic isotope effects (195, 196). The theory and application of isotope effects to establish TS structures for enzymatic reactions is well documented elsewhere (192, 197199).  KIE analysis of transition-state structures has not been widely investigated  compared to structural techniques that measure stable states (NMR and crystallography) because its application requires thorough knowledge of each enzymatic catalytic mechanism. Nevertheless, transition-state structures derived from kinetic isotope effect analysis have provided an atomic outline for the design of transition-state analogues (188, 196, 198, 199).  Protein structural similarity has not been a strong predictor of transition-state structure.  For example, the catalytic sites of human and bovine purine nucleoside  phosphorylases are identical within the error of X-ray diffraction experiments, yet the transition states differ considerably (200).  Thymidine phosphorylase, another  ribosyltransferase, was predicted from structural and computational analysis to have a dissociative transition state, but kinetic isotope effect analysis established a near-classical SN2 transition state (201, 202).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  ;  141  4.1.3.2 Computational approaches to the transition state Ab initio computations of enzymatic transition states starting from crystal structure models have become increasingly accessible as computational power has increased together with better atomic models from crystallographic starting points (203205).  Use of quantum-mechanical and molecular-mechanical (QM/MM) allows the  calculation of reaction trajectories, thereby providing transition-state structures and energies of reaction. Quantum-mechanical  and  molecular-mechanical  (QM/MM)  separation  of  enzymes into catalytic site and the non-catalytic regions permits inclusion of both dynamic and electronic/electrostatic components to transition state formation. Transition states located by ab initio searches of this type have found agreement with experimental intrinsic KIE measurements in some cases, but not in others (206). A major barrier for computational approaches to enzymatic transition states is the structural model for the starting point. Computational chemistry can simulate dynamic motion for periods of only a few nanoseconds (10  s), while catalysis occurs on the scale of milliseconds (10  s).  On the nanoseconds scale, covalent bond equilibration occurs and local motion can be optimized (e.g., amino acid side chain torsion angles) but there can be no significant loop, domain or group movements of the type that are well known to convert Michaelis complexes to the transition state. Thus computational approaches to the transition state from these enzymatic ground state complexes are only approximations of actual transition states. They are unlikely to return robust predictions of the transition state unless the close approach of enzymatic groups to the substrate that constitute the transition state interactions can be reproduced. This requirement is best met when the starting point for  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  142  computational chemistry is a high-resolution crystal structure of an enzyme with a transition-state analogue bound at the catalytic site. The significance of experimental KIEs as the basis for computational approaches to enzymatic transition states cannot be overemphasized. 4.1.3.3 T S analogues  Transition-state analogue inhibitors are designed from the hypothesis that chemically stable analogues that mimic geometric and molecular electrostatic features of the transition state will be bound to the enzyme tighter than the substrate by a factor equal to the ratio of enzymatic and non-catalytic processes (188, 207).  Transition-state  analogues with nearly irreversible binding can hypothetically be designed for enzymes with typical catalytic rate enhancements (10  10  to 10 ). 15  However, the dissociation  constants (Xi) for transition-state analogues designed to date fall short of attaining the affinity calculated for the transition states, since the actual enzymatic transition state has non-equilibrium bond lengths and charges that cannot be accurately copied in chemically stable molecules. Nevertheless, dissociation constants in the picomolar range are not uncommon (208, 209). For instance, zebularine binds to cytidine deaminase with an estimated K[ value of 1.2  x  10~ M (210). Analogues in the femtomolar range (10~ M) have also 12  been designed.  15  Recently, Singh et al  tested inhibitors of Escherichia coli  5'-methylthioadenosine/S-adenosylhomocysteine nucleosidase (MTAN) (211). The most powerful inhibitor was 5'-p-Cl-phenylthio-DADMe-immucillin-A with a K, value of 47 fM (47 x 10" M), representing one of the most powerful non-covalent inhibitors 15  reported for any enzyme.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  143  The specific arrangements of many transition-state analogue inhibitors within the active sites of enzymes have been determined by X-ray crystallography (212, 213). These structures have provided insight into the interactions that might be responsible for transition-state stabilization. For instance, a number of transition-state analogue designs have incorporated geometric differences between ground and transition states (194, 214218). The orientation of hydrogen bonds between enzyme and substrate in the transition state may be investigated using transition-state analogue inhibitors with a defined conformation and orientation of the functional groups. The electronic characteristics of charged intermediates have also been exploited. Nevertheless, many questions remain concerning the details of TSA binding and the origin of the high binding affinities observed.  4.1.4 Criteria for transition-state analogy  Many potent inhibitors are known to bind in a very different manner than the transition state; therefore, the observation of binding affinity that is greater than that of the ground state is not sufficient proof of transition-state analogy. While the structural analogy between inhibitor and transition state has been investigated in a limited number of studies (194, 219), a number of more rigorous criteria have been suggested to characterize transition-state analogy. These criteria, discussed below, include (a) strong inhibition, (b) slow inhibition, (c) free energy relationships (d) thermodynamic parameters, and (e) pH profiles.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  144  4.1.4.1 Strong inhibition Although a good transition-state analogue must necessarily be a strong inhibitor, a strong inhibitor does not have to be a transition-state analogue. In fact, a strong inhibitor may not even bind to the active site. Therefore, one cannot classify an inhibitor as a transition-state analogue based solely on strong inhibition.  4.1.4.2 Slow inhibition Two basic kinetic mechanisms (Figure 4.3) have been described to account for the slow-binding inhibition of enzyme-catalyzed reactions (220, 221). One mechanism involves the slow interaction of an inhibitor with enzyme (mechanism A), while the other involves the rapid formation of an enzyme-inhibitor complex that undergoes a slow conformational change (mechanism B). Mechanism A assumes that the formation of an EI complex is a single slow step and that the magnitude of k [l] is quite small relative to 3  the rate constants, &,[S] and k for the conversion of substrate to product (136). 2  Mechanism B, which is more general, assumes the rapid formation of an EI complex that then undergoes a slow conformational change to an EI* complex. Again, all other steps, and in particular & [I] and k_ are assumed to be fast relative to & and k_ . 3  v  4  A  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  *i[S] v  v  ES  E +P  ES  E +P  145  -i  > slow  -3  EI  B *i[S]  k [l] 3  EI  slow EI*  Figure 4.3: Two potential mechanisms for slow-binding inhibition.  Slow-binding inhibition has been suggested as a criterion for transition-state analogy based on the following hypothesis. Some enzymes are in a conformation that binds the substrate in a ground-state conformation, but then undergo a conformational change in order to bind the transition state better. TSA inhibitors can only bind to the conformationally changed enzyme, and the slow-binding behaviour is associated with the time spent for the enzyme to change conformation (222).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  1_46  However, in most cases, slow binding is an inevitable consequence of the low concentrations of tight-binding inhibitor used to determine its K\ (193, 223). When working with very low inhibitor concentrations that are comparable with the concentration of enzyme, the enzyme-inhibitor complex is present at concentrations comparable with that of the unbound enzyme; changes in experimental conditions change appreciably as a result when working with slow-binding inhibitors. Also, when working with low concentrations of inhibitor and enzyme, the rates for the bimolecular process are necessarily low. Association is a slower event if the concentrations of both inhibitor and enzyme are low and the two ligands have to find each other in solution (the probability of collision is low). For the condition that [I] » [E] , the rate equations for mechanisms A and B are 0  readily derived and it has been possible to distinguish between these mechanisms by analysis of the time courses of the reactions as a function of initial substrate and inhibitor concentrations. It has also been possible to analyze data corresponding to mechanisms A and B when the inhibition constant of a tight-binding inhibitor is of similar magnitude to the enzyme concentration by performing experiments at elevated enzyme concentrations (221).  Also, methods that take substrate depletion into account have been described for  analysing experiments in which the initial substrate concentration is below K  m  (224).  The slow-binding component may be eliminated if the enzyme and inhibitor are pre-incubated and the reaction started by addition of substrate (225). Nevertheless, slow onset inhibition is too poorly understood a process to be a safe criterion. Alternatively, it may be more appropriate to apply "fast-binding" behaviour as a criterion for "substrate analogy" rather than the reverse.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  1_47_  4.1.4.3 Free energy relationships Rigorous criteria have been proposed by Wolfenden (226) and by Thompson (227) to address the question of whether a particular glycosidase inhibitor is a transitionstate analogue  or simply a fortuitously  binding inhibitor.  Alterations in  inhibitor/substrate structure or enzyme active site should produce parallel effects on inhibitor binding (reflected in K\) to those seen on transition-state stabilization, as inferred from the inverse second order rate constant K /k . m  Therefore, as a consequence  cat  of the thermodynamic cycle of Figure 4.4, a correlation between the inhibitor K\ value and the substrate K /k m  cat  value is indicative of transition state analogy, as derived below.  Similarly, a correlation of inhibitor K\ versus substrate K values provides evidence of m  ground state analogy. This approach has been reviewed by Mader and Bartlett (194). Such alterations can be achieved through modification of the ligands or of the protein.  t  uncat  E +S ^=  il ES  E + S*  E+P  ES*  E +P  K,cat  Figure 4.4: Thermodynamic box illustrating the relationship between ground-state and transition-state binding. The designations E, S, and P refer to enzyme, substrate in the ground state form, and product, respectively, while S represents the transition state structure. }  Equation 4.3 relates the dissociation constant Ks and the hypothetical dissociation constant K to the pseudo equilibrium constants ^„ ts  K**  at  for the enzyme-catalyzed reaction.  ncat  for the uncatalyzed reaction and  Using a variation of the Eyring equation  Chapter 4  (k —  Transition-State Analogy of a Series of Iminosugar Inhibitors  with the assumption that the transmission coefficient  KVK*)  K  148  and the frequency of  the normal mode oscillation v are equivalent for the uncatalyzed and catalyzed reactions, the hypothetical dissociation constant for the transition-state structure, K , may be related ts  to the substrate dissociation constant, Ks, by the ratio of the uncatalyzed and catalyzed rate constants,  & ncat U  and k x, as shown in equation 4.4. The transition-state structure is ca  bound to an enzyme with an increase in affinity over the ground state that is proportional to the increase in the rate constant for the catalyzed over the uncatalyzed reaction. This transition-state analogy was first recognized by Pauling (147) in 1946 and later implemented by Lienhard (228) and Wolfenden (190) to provide a basis for the design of tight binding enzyme inhibitors. K.  Kc -  t s  K*  uncat  (4.3)  s  K  l  cat  uncat  K  (4.4)  s  V ^cat J  As no inhibitor will be a perfect mimic of the transition state, the relationship between K\ and K may be obtained by introducing a proportionality constant (d). ts  k  r  K dK =dK l=  ts  s  (4.5) uncat  V ^cat J  Equation 4.6 may be derived by rewriting equation 4.5 in logarithmic form and separating the proportionality constant d and the uncatalyzed rate constant & £s/& at C  w  i  t  n  u n c a  t  (equating  ^m/#cat)-  log ^ = log  + l0g^unca, V ^cat J  (4-6)  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  Through the relationship  AG = -RT \nK, 0  the plot of log K  149  t  against  log(AT /£ ) is directly proportional to the free energies of binding for the inhibitor and m  cat  transition state since the proportionality constant d along with the unmeasurable become part of the constant log dk . ancat  A  &  u n c a  t  slope of 1 in the correlation of equation 4.6  means that structural changes produce the same effects on binding of the inhibitor and the binding of the substrate transition state. Likewise, ground-state analogy can be sought through a correlation between log K and log K , assuming K « Ks. {  m  m  One approach to obtain a correlation involves the synthesis of a series of modified substrates and identically modified inhibitors, followed by a kinetic evaluation with the wild-type enzyme, along with measurement of K\ values for the inhibitors. Previous correlations of this type have shown that a series of phosphorus-containing peptide inhibitors are transition-state analogues for thermolysin (229, 230), carboxypeptidase A (231),  pepsin (232), and a collagenase (233). Similarly, nojiritetrazoles were shown by  this method to be true transition-state analogues for a range of glycosidases (234). Most recently, this approach has been applied to investigate the binding of two known potent inhibitors to human 0-GlcNAcase; a family 84 glycosidase (235). Another approach involves generating a series of mutant enzymes, and then measurement of k JK c  m  and K\ values with each mutant for a single substrate and  transition state analogue inhibitor, respectively. The latter approach has been applied to rat carboxypeptidase A (236), glucoamylase (237), and cyclodextrin glycosyltransferase (238).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  150  A linear relationship will only gauge the degree to which the varied part of the inhibitor or enzyme structure mimics the corresponding region of the transition state, and therefore depends on the substrates and inhibitors binding in the same fashion across the series  (194).  The tightness of the correlation may in fact provide an indication of the  degree to which the binding geometries and electronic nature of the inhibitor and transition state remain constant. However, parts of the inhibitor that are not varied, may not resemble the transition state. For example, a study by Vasella's and Withers' groups found a linear relationship between ln(A; /^ ) for hydrolysis of 4-nitrophenyl glycoside cat  m  by a series of different mannosidases, glucosidases, and galactosidases and the \n(K\) value for  gluco-  and ma««o-tetrazole  (234).  Surprisingly, despite the use of different  substrates and different enzymes, a linear correlation was obtained. However, the only varied part of the inhibitor was the 2-hydroxyl group and no variation in the charge of the inhibitor was made. The only conclusion that can be drawn from this correlation is that the 2-hydroxyl group interacts with the protein active site in the same manner that it does in the transition state. These results do not reveal anything about the transition-state mimicry of any other part of the molecule. Similar free energy relationship studies have been made for other glycosidase inhibitors (238, 239).  4.1.4.4 Thermodynamic parameters Wolfenden suggested that the binding of true transition-state mimics should be driven by enthalpy (207, 240).  A comparison of the temperature dependences of  uncatalyzed single-substrate and hydrolytic reactions with those of the corresponding enzyme reactions showed that these enzymes invariably lower the reaction's enthalpy of activation substantially, by an amount that tends to determine the effectiveness of the  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  151  enzyme as a catalyst, while the entropy of activation of a reaction changes only to a modest extent. In laboratory experiments (240), spontaneous uncatalyzed reactions were shown to be much more sensitive to temperature than generally assumed. For enzyme reactions, k x is less temperature-sensitive. ca  corresponding rate enhancements (k /k ) ceLt  non  As a result of this tendency, the  increase with decreasing temperature, and  enzyme affinities for transition-state analogue inhibitors are expected to increase with decreasing temperature, more sharply than do enzyme affinities for the corresponding substrates in the ground state. This sharp temperature dependence of K, provides a new criterion for testing potential transition state analogue inhibitors. 4.1.4.5 p H profiles  It has been proposed that a transition-state mimic should be a true analogue of the transition state if the pH dependence of inhibition correlates with the pH dependence of kcJK  m  of the enzyme (194). However, an incredibly complex expression for the pH  dependence of inhibition occurs if the inhibitor is ionisable since the pH dependence of K  {  depends upon the pH dependence of the free enzyme, free inhibitor and the enzyme/inhibitor complex (139). In this type of analysis, the pH dependence of catalysis will include the pH dependence of free enzyme and something mimicking the enzyme/inhibitor complex, but, by definition, will not include the pH dependence of the free  inhibitor.  Therefore, the pH dependence of catalysis should mimic the pH  dependence of inhibition for a transition-state analogue if the inhibitor is non-ionisable. Nevertheless, this criterion has been used to disqualify inhibitors whose binding profile as a function of pH did not correlate with that of catalysis (241-243), which occurred when the inhibitor changed charge over the observed pH range. Nevertheless, the inhibitor  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  752  could still be a transition-state analogue, albeit an imperfect one, at one pH and not another.  4.1.5 Inhibitors of glycoside hydrolysis The best small molecule inhibitors currently available for glycosidases (219, 244) have K, values of just 10" to 10" M . 9  10  Transition states for glycoside hydrolysis feature  extensive sp hybridization and partial positive charge (predominantly along the bond between the anomeric carbon and endocyclic oxygen) and likely involve pyranoside distortion to half-chair ( H^ or / Y or their closely related envelope form) or boat ( ' B or 4  3  2 5  4  B ' ) conformations (63). 2 5  In the design of transition-state analogues of glycoside  hydrolysis, much of the focus has been on mimicking these features.  Most designs  focusing on geometry have tried to create inhibitors that are in a half-chair conformation. It should be noted, however, that the most substantial evidence for this conformation comes from measurement of kinetic isotope effects that show various degrees of sp 2  character at the anomeric carbon of the transition state (245).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  1-Deoxynojirimycin  Amidine  Amidrazone  Isofagomine  Nojirimycin tetrazole  X>H  ISSS^ O N OH N Y O  A/-Phenylcarbamate  OH Nojirimycin imidazole  Figure 4.5: Structures of some naturally occurring and synthetic glycosidase inhibitors in the £/Hco-configuation.  153  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  1_54_  Some of the first tight-binding glycosidase inhibitors investigated were the glyconolactones, depicted in Figure 4.5 in the g/wco-configuration  (246).  These were  argued to function as transition-state analogue inhibitors by virtue of the resemblance to the oxocarbenium ion.  Unfortunately, their usefulness has been limited by their  instability. Another class of naturally occurring glycosidase inhibitors are that of the glycosylamines and glycosamines, as shown in Figure 4.5. The good inhibition in these cases is attributed to the presence of a protonated (or protonatable) amine moiety, which was proposed to interact electrostatically with the carboxylate groups at the active site. Nojirimycin was discovered as an isolate from  Streptomyces  by Ishida  et al. (247),  and  resembles glucose very closely, differing only by the presence of an endocyclic nitrogen rather than oxygen. Subsequently, 1-deoxynojirimycin was obtained by Inouye et al. (248) by reduction of nojirimycin.  Over three decades a large number of naturally  occurring or synthesized hydroxylated piperidines, pyrrolidines and other related structures have been reported and tested. Particularly noteworthy are the isofagomines, which incorporate a nitrogen atom in place of C l  (249).  Isofagomine was the first azasugar prepared with nitrogen in the  pseudo-anomeric position (250, 251), and because of the close resemblance between this azasugar and the natural product fagomine, the compound was named isofagomine. The 2-hydroxy group had been omitted for stability reasons. Isofagomine is a potent inhibitor of P-glucosidases, while a-glucosidase inhibition is moderate. The onset of inhibition of isofagomine with sweet almond P-glucosidase was found to be slow, and the binding was found to be driven by a large increase in entropy in contrast to the binding of deoxynojirimycin, which was found to bind due to a decrease in enthalpy (252).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  155  Attempts to develop stable analogues of glyconolactones have resulted in compounds  such  as  the  hydroximolactones,  hydroximolactams  and the  N-phenylcarbamates (Figure 4.5). These compounds have a flattened chair conformation and an sp -hybridized anomeric carbon. The gluco-, manno-, and ga/acto-derivatives of hydroximolactams have been made, and have been characterized as broad spectra inhibitors of glycosidases (253-256).  The most stable tautomer of the gluco-  hydroximolactam has an exocyclic double bond. This has been shown, both in the solid state and in solution, by X-ray, *H NMR, and by  1 5  N NMR of the N-labeled 15  isotopomers (257, 258). Thus, the double bond was not endocyclic as assumed by Tong et al. but exocyclic (257). The location of the double bond is important because it has a  large effect on the conformation of the compound and also on the interpretation of the inhibitory properties. An approach to mimic both the flattened chair conformation and the positive charge of the assumed transition state was begun in 1991 by Vasella's group (259). They tried to mimic the conformation and charge of the transition state by synthesizing compounds that consisted of an azasugar fused with heterocycles such as tetrazoles, triazoles, or imidazoles (19). In this way it was possible to make a series of compounds which mimicked the transition state in the desired manner and which had a wide range of p/C values (260). Furthermore, another benefit was that the azoles were expected to be a  more stable than the amidines and amidine derivatives that were already known. It should also be pointed out that imidazole itself and derivatives of L-histidine also have turned out to be moderate to strong inhibitors of different glycosidases (261).  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  Subsequently, Tatsuta's group also synthesized compounds of this class (262, 263) . Some of the most powerful glycosidase inhibitors contain imidazole moieties (260, 264) , and are more selective than the amidines and their derivatives. The azoles are not only interesting because of their strong inhibitory properties, but also have provided a better understanding of the enzymatic mechanism of the hydrolysis of glycosides (19).  In 1996 Heightman et al. noticed that the moderate to  strong P-glycosidase inhibitors all had a heteroatom (N or O) at the glycosidic position and that similar compounds which did not have a heteroatom in this position were weak inhibitors; therefore, they suggested that there is an important interaction with the enzyme at this position (265). Assuming that the inhibitors are bound in the same way in the enzyme as the substrate, it is reasonable to propose that the enzyme-inhibitor interaction is an interaction between the catalytic acid and the heteroatom, thus mimicking the protonation of the glycosidic oxygen at the substrate.  It should also be possible to  determine whether a glycosidase is a syn- or a«r/-protonator by investigating the inhibitory properties of a series of carefully selected compounds.  For example, the  imidazoles, which contain a heteroatom at the glycosidic position, would be expected to be selective cwrz'-protonators because the orientation of the lone pair at the glycosidic nitrogen should allow it to interact with the catalytic acid only in this type of enzymes.  4.1.6  Inhibitors of Cex .A/y/oo/o-derived inhibitors have been made and investigated by the Withers group  as inhibitors (Table 4.1) of two different xylanases, namely, the retaining enzymes Cex and Bex.  156  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  157  Table 4 . 1 : Inhibition constants (K\) for Cex and Bex with various xylobioderived iminosugars. Cex;;.  Compound"'' ' " . V  (uM)  ... J H M ) , „  4.9 ±0.1  0.37"  1400"  5.9 ±0.1  0.15  520"  <-5  0.34*  9000*  6.9 ±0.1  5.8*  1500*  5  8.8 ±0.2  0.13°  1100"  6  nd  1  .  OH  2  a  OH  3 0  4  OH  c  110"  3100"  " Data taken from Williams et al. (52). Data taken from Williams et al. (183). nd indicates that this parameter was not determined. b  0  The inhibitor 5 was an inhibitor of Cex in the nanomolar region (K\ = 0.13 uM) and as such the strongest inhibitor of this enzyme yet described. A l l the inhibitors studied were poor inhibitors of Bex with inhibition constants ranging from 520 to 9000 u.M. This low activity against the family 11 xylanase was explained by the enzyme being a svfl-protonator that favors boat conformers in the enzyme-catalyzed transition state, while Cex was believed to prefer a half-chair conformation.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  Structures of all the above-mentioned xy/00/0-derived azasugars, except 6, complexed in the Cex active site were determined by X-ray crystallography (57). It was shown that, as expected, all inhibitors were bound in the -2 and -1 subsites. It was further revealed that for deoxynojirimycin analogue 4 together with the isofagomine 5, the azasugar moiety adopted a C\ conformation. Even though ry/06/0-isofagomine 5 is a A  very potent inhibitor of Cex, it demonstrated only a few contacts with the enzyme. The most important interaction was proposed to be between the protonated anomeric nitrogen and the oxygen of the catalytic nucleophile.  Because of the small number of  enzyme/inhibitor interactions and the conformation of the isofagomine, its strong binding was proposed to be fortuitous (51). Deoxynojirimycin analogue 4 binds 830 times better than xylobiose, although no direct interactions were observed to the azasugar's nitrogen atom. Two water molecules, however, were found to form hydrogen bonds to the basic center, one of which forms a hydrogen bond back to the enzyme (57).  4.1.7 Aims of this study  The primary aim of the work to be presented in this chapter is to discern the important features that mimic the transition state of a model family 10 xylanase (Cex). Since very little is known about the structure of the transition state, insights from our study of Cex could provide a starting point towards a greater understanding of the nature of interactions that can potentially occur between the enzyme and the substrate during the course of the catalytic reaction. To investigate this aspect of the enzyme, the kinetic parameters for inhibition of four classes of potential transition-state  analogue inhibitors will be examined.  158  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  Discernment between the crucial features of a transition-state analogue will be achieved through a series of inhibition studies with five inhibitors and a series of mutants of Cex. Linear free energy relationship plots will be constructed to show whether the inhibitors are indeed transition-state analogues. Additionally, the pH dependence of inhibition will be investigated. The information gained from these studies will provide insight about the enzyme-substrate interactions at the transition state of the Cex-catalyzed hydrolysis reaction. An understanding of the transition state is a valuable asset, for it will greatly aid in the design of compounds in the quest for highly potent inhibitors and specific therapeutic drugs.  4.2  Results and Discussion The binding of several different active site mutants of Cellulomonasfimixylanase  to a series of iminosugar inhibitors (1-5) has been investigated through measurement of K\ values. The mutations represent several key amino acid positions, most of which are believed to play important roles in ground-state and transition-state binding interactions of Cex.  Michaelis-Menten parameters  (k , K , cat  m  and  were determined for each  k JK ) c  m  mutant enzyme with o-nitrophenyl xylobioside (ONPX2) and are reported in Chapter 2. The structural changes within these mutant enzymes give rise to k JK c  about three orders of magnitude. Such a wide variation in k JK c  m  m  values varying by  values is valuable for  establishing a meaningful correlation with K values for the inhibitors in order to gain x  insight into the nature of the strong inhibition afforded by these nitrogen-containing compounds.  159  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  [60  4.2.1 Inhibition of Cex mutants by iminosugar inhibitors  Inhibition constants for a series of iminosugar inhibitors were measured with a series of Cex mutants, and these are presented in Table 4.2. For the wild-type errzyme and the N169A mutant, the K\ values were determined from a full analysis using a range of substrate and inhibitor concentrations as is shown for the inhibition of Cex N169A by xylobio-imidazole 2 in Figure 4.7. Inhibition was strictly competitive in all cases. In all other cases, K\ values were determined from data obtained at a series of inhibitor concentrations using a fixed concentration of substrate as described under Materials and Methods.  Figure 4.6: Structures of the 5 iminosugars studied.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  I6J_  Table 4.2: Inhibition constants (Ki) for C. fimi xylanase and mutants with various xylobio-derived iminosugars. c  Wild-Type Glu43Ala : Asn44Ala Lys47Ala His80Ala His80Asn His80Gln Gln87Met Gln87Tyr Asnl26Ala Glul27Ala Asn 169 Ala  0.37" 0.15" 0.34* 110 22 ~~ 96 23 8.9 24 2300 "J . 1000 2300 140 43 " 1000 350 1 87 340 1300 "~ 350 " ~ 340 4.9 1.1 2.1 16 '_ " " 6.4 Ill T"'1" 1300* 200* 2000* " 13 ~ " 99 28 2.1 0.41 1.5 -  5.8" 2100 180 6600 800 370 590 18 ^ 6 2 ~ 4800* 660 17  :  0.13" 45 7.1 870 5.9 13 " 6.7~" 0.39 " 1 . 2 1.6* 35 0.12  Data taken from Williams et al. (52). * Data taken from Williams et al. (183). Graphical representation of this data is given Appendix A, page 269. A relative error of 10% is estimated for the inhibition constants (K().  a  0  0  40 80 120 1/[2,4DNPC] (mM" ) 1  160  0 0.4 0.8 \Z /[2,4DNPC] ( A A Q  1.2 1.6 mirT mM- ) 1  400  1  Figure 4.7: Inhibition of Cex N169A by xylobio-imidazole at varying concentrations of 2,4DNPC. Plot A is the Michaelis-Menten plot, plot B is the Lineweaver-Burk plot, and plot C is the Eadie-Hofstee plot. The concentrations of jcy/otoo-imidazole were 0 (O), 0.123 (•), 0.370 (•), 0.925 (•), and 2.47 uM (A).  The mutant enzymes bind to all inhibitors, but less tightly than does wild-type Cex, with widely varying K values. x  For example, for xy/o6/o-imidazole, affinities  ranging from a K\ of 160 nM for wild-type Cex to 1.0 mM for Cex K47A were measured. This large loss of affinity upon removal of the Lys47 side chain is again consistent with the key role played by this residue.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  [63_  4.2.2 How good a transition-state analogue is the inhibitor?  A rigorous approach to establish transition-state analogy of the series of iminosugar inhibitors for Cex-catalyzed xylanase hydrolysis has been to use free energy correlations between inhibitor binding and transition-state stabilization (194, 229). A strong correlation with a slope of 1 reveals the inhibitor is a transition-state analogue. Any slope greater or smaller than this may indicate a lesser degree of mimicry. It is not, however, clear that a slope of 1.0 is required to indicate any degree of mimicry as has been suggested (194), a conclusion that was developed on the assumption of direct proportionality of rate constants and equilibrium constants themselves (188, 228). Conversely, scattered plots that have weak correlations suggest the inhibitor is a poor transition-state analogue or a fortuitously binding inhibitor (266). Furthermore, strong correlations having a slope of 1 between \og(K\) of the inhibitor and log(X ) values for s  binding of the substrate reveal ground-state analogy.  For enzymes with a rapid-  equilibrium substrate binding step, log(AT ) may be used, or alternatively \og(K\) for a m  known ground-state analogue may also be used. It is useful to review previous LFER studies on glycosidases with similar classes of inhibitor. The first published study of a correlation in glycosidases between substrate and inhibitor specificity, as a probe of transition-state mimicry, was that on the glucoand /wa««0-nojirimycin tetrazoles with a series of glucosidases and mannosidases (234). In this study a pair of substrates and inhibitors was studied with the corresponding pair of glycosidases from a variety of sources. A plot of \og(K /k ) for each of these substrates m  cat  versus the \og(K\) value for the tetrazole inhibitor of the corresponding configuration was linear with a slope of 0.96 and a correlation coefficient of 0.9. The plot of log(A^ ) for the m  Chapter 4  Transition-State Analogy of a Series of Iminosuear Inhibitors  substrate versus \og(K\) for the corresponding tetrazole was a scatter plot with a correlation coefficient of 0.2. The tetrazoles were therefore classified as transition-state analogues. It is noteworthy that in two studies to check the transition-state analogy of acarbose, correlations of log(K\) versus log(AT /A: t) have yielded slopes other than 1 (237, m  ca  238). The approach followed in both cases involved the generation of a series of mutants of the enzyme in question, then measurement of kinetic parameters for the hydrolysis of a defined substrate by each of these mutants. The inhibition of each of these mutants by acarbose was then investigated, and K\ values determined. The first such study was performed with the family 15 Aspergillus niger glucoamylase using maltose as substrate. The plot was linear with a slope of 0.38 and a correlation coefficient of 0.88, indicating some degree of transition-state mimicry. A second study of acarbose as a transition-state analogue was performed on the retaining family 13 cyclodextrin glycosyltransferase from Bacillus circulans.  Values of & , K cat  m  and k JK c  m  were determined for the reaction of  both a-glucosyl fluoride and a-maltotriosyl fluoride with each of 8 mutants and with the wild-type enzyme. In addition, K\ values were measured for the inhibition of each mutant by acarbose.  A plot of \og(K lk A for a-glucosyl fluoride versus the \og(K\) value m  ca  acarbose was linear with a slope of 2.2 and a correlation coefficient of 0.98. Interesting, acarbose functions as a transition-state analogue for both an inverting a-glucosidase and retaining a-glucosyl transferase. Although castanospermine and deoxynojirimycin are good competitive inhibitors of Agrobacterium sp. p-glucosidase with K\ values of 2.8 and 12 uM respectively (21, 28),  plots of \og(KJk A versus \og(K\) for these inhibitors failed to show any significant ca  164  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  \_65_  correlation. Despite their high affinity binding, neither of these classes of compound was found to function as a transition-state analogue, but rather appeared to act as a fortuitous inhibitor. In the present study, plots of log (k  /K )  cat  ONPX2  m  for the Cex-catalyzed hydrolysis of  versus log (l/K[) for the inhibitors in Figure 4.8 were constructed. Excellent  correlations (r = 0.96-0.97) were observed with slopes close to 1 for the sp -hybridzed 2  inhibitors (compounds 1-3), clearly indicating a substantial degree of mimicry (Figure 4.8, plots A, B, and C). Changes in binding interactions with the inhibitors (compounds 1-3) therefore correlate quite well with changes in transition-state binding interactions with substrate. Somewhat weaker correlations were seen for 4 and 5, with correlation coefficients of 0.89 and 0.77, respectively (Figure 4.8, plots D and E). The poorer correlations observed for the sp -hybridized inhibitors (compounds 4 and 5) clearly indicate that both 3  inhibitors have some properties of a transition-state analogue, presumably their charge, but are not as good analogues as the sp -hybridized inhibitors (compounds 1-3).  The  poorer correlation observed for the isofagomine 5 over the deoxynojirimycin 4 might be attributed to the absence of a 2-hydroxyl, which interacts with the nucleophile and Asnl26, and has been shown to contribute greatly to transition-state stabilization in P-glycosidases (17). The inferior correlation relative to the deoxynojirimycin may also reflect the poorer mimicry of transition-state charge distribution. Indeed, this conclusion of poorer transition-state mimicry by 5 is consistent with the findings of Davies et al. (218),  who detected a large entropic contribution in a calorimetric study of binding of  (+)-isofagomine to  Thermotoga maritima  family 1 P-glycosidase (7mGHl). The binding  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  166  of deoxynojirimycin, on the other hand, was largely driven by a large, favourable, enthalpic contribution, consistent with Wolfenden's suggestion that the binding of true transition-state analogues should be driven by enthalpy (240). Correlations with K values were not possible, as K is not the true dissociation m  m  constant of the enzyme-substrate complex, but rather represents the overall dissociation constant of all enzyme-bound species. In the case of Cex and the series of mutants, deglycosylation, resulting in accumulation of the glycosyl-enzyme intermediate, was shown to be the rate-limiting step in the hydrolysis of aryl xylobiosides  (Chapter 2).  Correlations probing for ground-state analogy were therefore not possible. Nevertheless, interesting patterns emerge when comparing the results of this study to those mentioned above. It seems apparent that the sp -hybridized class of inhibitor, 2  including tetrazoles, acarbose, and inhibitors 1-3, function as transition-state analogues. On the other hand, weak correlations observed for the sp -hybridized class of inhibitor, in 3  particular deoxynojirimycin and isofagomine, seem to suggest that these two inhibitors are ground-state analogues or adventitious binders to the enzymes studied (21, 28, 237, 266).  Surprisingly, recent studies have shown that the sp -hybridized 'PUGNAc' inhibitor is a poor mimic for the family 84 glycosidase O-GlcNAcase, which utilizes substrate-assisted catalysis.  NAG-thiazoline, which possesses an sp -hybridized  anomeric centre, is a better transition-state mimic in this case  3  (235).  The future design of sp - and sp -hybridized classes of compounds as glycosidase inhibitors should be undertaken with these findings under consideration.  Chapter 4  Transition-State  I  " -1 1  i  l  i  l  0  1  i  l  2  i  Analogy of a Series of Iminosugar Inhibitors  l  3  i  [67  I  4  log(1/Ki)  Figure 4.8: Transition state analogy plots for C. fimi xylanase Cex. Comparison between for ONPX and inhibitor K-, values for xy/06/0-derived (A) lactam oxime 1; (B) imidazole 2; (C) isofagomine lactam 3; (D) deoxynojirimycin 4; and (E) isofagomine 5.  *cat/*m  2  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  168  4.2.3 The pK values of xy/oo/o-derived inhibitors a  The pK values of the conjugate acid forms of xylobio-derived iminosugar a  inhibitors were determined by potentiometric titration (see Figure 4.9) and the results are summarized in Table 4.3. Determination of the pK values of the azasugar inhibitors is a  important in order to gain insight into the inhibition of these compounds. Potentiometric titration of the acidic form of g/MCO-lactam oxime revealed a pK  a  value of 5.6 (254), although a lower value of 4.8 for the lactam oxime has been reported as well (257). The xy/oZj/o-lactam oxime used in this study yielded a pK value of 4.9. a  Furthermore, the pK of the xylobio analogue of imidazole was measured to be 5.9, close a  to that reported for the gluco and cellobio analogue ($K 6.1) (219). a  The pK of deoxynojirimycin and its marino and galacto analogues were reported a  (267) to be 6.3, 7.2, and 7.1, respectively. The xylobio-dexxvQd analogue yielded a pK  a  value of 6.9. The pK of the protonated forms of isofagomine and its galacto analogue a  were measured to be 8.4 (250) and 8.8 (268), respectively (269). The xy/06/o-derived analogue used for this study yielded a pK value of 8.8. a  The pK of the xy/ofo'o-isofagomine lactam was not measured, but is likely well a  below -5, as amides are very strong bases (conjugated acid pK between -6 and -10). a  Chapter 4  Transition-State  Analogy of a Series of Iminosugar  Inhibitors  169  Figure 4.9: Titration curves for the conjugate acid forms of the xylobio-derived lactam oxime (•), imidazole (•), deoxynojirimycin ( T ) , and isofagomine (•). Note: amides are strong bases (conjugated acid pK between -6 and -10); therefore, the pK of the Jtj/oA/o-isofagomine lactam is likely well below -5. a  a  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  Table 4.3: acid.  170  Xylobio-derived inhibitor pK values of the conjugate a  Xylobio-derived Inhibitor  4.9 ±0.1 OH  5.9 ±0.1 OH  <-5 0  6.9 ±0.1 OH  8.8 ±0.2  At the pH at which inhibition constants (K\) were obtained (pH 7.0), the sp 2  hybridized xylobio analogues of the lactam oxime 3  1,  imidazole 2 and isofagomine lactam  are likely to be in their neutral, unprotonated form.  The sp -hybridized 3  xylobio  analogue of the deoxynojirimycin 4 is half-protonated in solution, while the isofagomine 5 is in its charged, protonated form.  Chapter  4  Transition-State  Analogy  of a Series  of Iminosugar  171  Inhibitors  4.2.4 The pH dependence of the binding of xy/ofa/o-derived inhibitors  I  |  I  |  I  |  I  |  4  2  i  1  1  i  i  180  i .  \  1.5  /  f  1  n  [ J  i  i  0.5 =2  2  \  7  J  v i  L-ir'* i 1 i 1 L  3  \  4  5  6  i  7  i  i  8  i  120 100 80 60 40  E •„ ~ ^  J  _  9  10  PH  i  *  i /  1  3  Jr 4  \  JT  l_k-T  5  i  i  y  1  i  |  •4  1  f -  i  \\  1  -  |  -  \  J  1 l 1  6  i  7  i  8  \ l  ~-  10  180 160 140 120 100 80 60 40  I  I  I  I  8 E 3  it (8  / * i  6 4 2  -  -  I  I  I  I  r  1  1  *; i I  ' 3  5  6  7  180 160 140 120 100 80 : 60 40 20 0 10  : \.\ :  J 4  1 1  J \l \\ ,  2  10  1  ^  i , N ^ _  9  I  8  i  ,  9  pH  PH  Figure 4.10: pH dependence of k /K for Cex (•), l/K, for xy/oo/o-derived lactam oxime pK 4.9 (•), imidazole pK 5.9 (•), isofagomine lactam pK < - 5 ( A ) , deoxynojirimycin pK 6.9 ( T ) , and isofagomine pK„ 8.8 (•). Fits to "bell-shaped" profiles are shown for k JK (dashed line) and l/K, (coloured solid lines). caLt  a  a  m  a  a  c  m  E E  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  172  Table 4.4: Parameters determined from the pH profile for the binding of xylobioderived inhibitors to wild-type Cex.  ^optimum 'inhibition -luieiivirv •  Xylobip^der.lyed inhlbitovt  ;-;(»iMjr • 'iv'  OH OH  0.59 ±0.06  \ 5.3 ±0.1  7.5 ±0.1  6.4 ± 0.2  0.16 ±0.04  6.2 ±0.1  7.3 ±0.1  6.7 ±0.2  0.36 ± 0.03  4.7 ±0.1  7.6 ±0.1  6.2 ± 0.2  3.2 ±0.1  6.5 ±0.1  8.0 ±0.1  7.3 ± 0.2  0.16 ±0.03  7.2 ± 0.2  0  HB^^SH-O^S  OH  WTCex  •  ~  3.9 ±0.04  7.5 ± 0.2  7.6 ± 0.04  5.8 ±0.1  4.2.4.1 Xylobio-derWed lactam oxime, imidazole, and isofagomine lactam The pH profile of k JK c  m  for Cex is bell-shaped with an optimum at 5.8, and  acidic and basic limbs of pA^ 3.9 and 7.6 (Figure 4.10). As with other related systems a  (109, 115, 270, 271), this pH profile most likely represents titration of the nucleophile and acid/base, respectively.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  I_73_  The pH dependence of \/K[ for 1-3 are similarly bell-shaped, with slight alkaline shifts relative to catalysis (Figure 4.10). The pH optimum for inhibition of 2 is around 6.7, with acidic and basic limb pK values of 6.2 and 7.3, respectively. The simplest a  explanation of the pH profiles is that the acidic limb is likely to reflect protonation of the inhibitor (which has a solution pK of 5.9) and the basic limb protonation of the acid/base a  residue.  This suggests that at the pH optimum for Cex activity, 2 is bound in a  predominantly protonated form, as experimentally observed with the cellobio-derived form of 2 in complex with a family 5 endoglucanase using atomic resolution X-ray data (272).  The pH optimum for inhibition by 1 is around pH 6.4, with acidic and basic limb pK values of 5.3 and 7.5, respectively. The basic limb is likely to reflect the protonation a  of the acid/base residue, and the acidic limb is likely to reflect protonation of the inhibitor (1 has a solution pK of 4.9). This suggests that at the pH optimum for Cex activity, 1 is a  bound in a predominantly unprotonated form. The pH optimum for inhibition by 3 is around pH 6.2, with acidic and basic limb pK values of 4.7 and 7.6, respectively. a  The titration profile for compound 3 is  interesting to interpret in that the protonation state of the compound is unlikely to change (since 3 has a solution pK of < -5, and will remain unprotonated throughout the pH a  range of enzyme activity). Therefore, the acidic limb is likely to reflect protonation of the nucleophile whose pK has increased approximately 0.8 units from a pK value of 3.9. a  a  The basic limb is likely to reflect the protonation of the acid/base residue, whose pK  a  remains unchanged.  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  174  As has been pointed out by others, the interpretation of pH profiles of inhibition is notoriously difficult since they are a combination of the events surrounding the free enzyme, the inhibitor and the enzyme-inhibitor complex (139). 4.2.4.2 Xylobio-derWed deoxynojirimycin and isofagomine  Cex is inhibited by 4 and 5 with K, values of 3.2 uM and 160 nM at pH 7.0„ respectively. The pH optimum for inhibition of 4 is around 7.3, with acidic and basic limb pK values of 6.5 and 8.0, respectively. The pH dependence of \/K{ of 5 gives an a  acidic limb of pK 7.2, although the basic limb could not be determined with accuracy a  because the alkaline shift in pH dependence did not allow for sufficient data points to be collected. Consistent with other studies of inhibition by 5, maximal binding occurs at high pH where the enzyme has lost significant activity (218, 271, 273). This has been interpreted in other studies as the protonated inhibitor optimally binding to an enzyme with a deprotonated acid/base and deprotonated nucleophilic residues (218, 271). In light of X-ray crystallographic data of cellobio-dchved 5, the alkaline shift of the acidic limb may simply indicate that a protonated inhibitor cannot bind tightly to an enzyme in which the acid/base catalytic residue is also protonated (271). In previous studies, deoxynojirimycin and its derivatives have yielded acid and basic limbs for l/K[ that are difficult to interpret. In a study by Gloster et al (273), a pH dependence of \/K\ for 4 with S. lividans xylanase gave acidic and basic limbs of pK 6.6 a  and 7.8. The atomic resolution structure of 4 in complex with the xylanase from S. lividans  XynlOA suggested unusual pK changes of the enzyme's active-site residues a  upon binding (273). Given that the pK value for 4 is 6.9 (Table 4.3) and the pH of a  crystallization was 5.8, it was expected that the inhibitor would be present in the active  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  775  site of the enzyme in protonated form. At pH 5.8, the nucleophile was protonated when xylobzodeoxynojirimycin was bound to STXynlOA (273). One interpretation of the pH profile that was suggested was that the drop of \IK\ with an acidic limb of 6.6 reflected the titration of the nucleophile whose pK had increased approximately 2 units from a pK a  a  value of 4.1 (273). A similar interpretation was suggested for the pH profile of inhibition for 5. Indeed, NMR spectrometry has been used to demonstrate significant liganddependent shifts in pK values for catalytic residues within the active sites of glycosidases a  (115). As with the inhibition of other glycosidases (241, 243, 271) by iminosugars, the inhibition of Cex as a function of pH does not correlate with the pH dependence of k JK Q  m  of the enzyme. Optimal inhibition occurs at pH values where the acid/base of the  enzyme becomes partially or fully ionized and catalysis is reduced or absent. Indeed, optimal inhibition occurs at pH values where the enzyme displays reduced activity.  4.3 Conclusion: Transition-state mimics or fortuitous binders? By definition, a "transition-state mimic" is never perfect and will not capture all of the binding energy available in an enzyme active site for the reaction transition state. In assessing a transition-state analogue, one must consider what enzyme-inhibitor interactions are relevant to transition-state binding, as well as what interactions are absent in the analysis.  Atomic structural analyses of inhibitor complexes with Cex have  identified important interactions with active-site residues in the enzyme (51, 183). In this study, the correlation of K\ with k /K in linear free energy relationships (LFERs) (194) cat  m  Chapter 4  Transition-State Analogy of a Series of Iminosugar Inhibitors  [76  has provided a mathematically rigorous means of investigating transition-state mimicry (207, 240).  Certainly, the sp -hybridized inhibitors, deoxynojirimycin and isofagomine, achieve some resemblance to the enzyme-catalyzed transition state.  In fact, the  sp -hybridized isofagomine was the most potent inhibitor despite it being the poorest 3  transition-state mimic studied, demonstrating that powerful inhibition can be obtained with partial resemblance to the transition state.  Nevertheless, the sp -hybridized 2  inhibitors (lactam oxime, imidazole, and isofagomine lactam) exhibited a higher degree of transition-state mimicry as assessed by the stronger correlations obtained in the transition-state analogy plots. The finding that sp -hybridized inhibitors are true transition state analogues, while 3  * •  2  sp -hybridized inhibitors are not, indicates that true transition-state mimicry demands sp hybridization at the anomeric centre, at least for the enzyme studied. Attempts to take advantage of the 10 -fold affinity increases available to transition-state analogues should 16  therefore focus upon maintaining an sp geometry at the anomeric centre.  CHAPTER 5  MATERIALS AND METHODS  Chapter 5  5  Materials and Methods  178  Materials and Methods  5.1 Molecular Biology 5.1.1  Reagents, enzymes, and bacterial strains  Growth media components were obtained from Sigma. Plasmid-containing strains were grown in Luria Broth containing 100u,g/mL ampicillin or 50 ug/mL kanamycin. Pwo DNA polymerase and deoxynucleoside triphosphates were obtained from Roche. Pfu DNA polymerase was from Promega. Restriction endonucleases and T4 DNA ligase were from New England BioLabs unless otherwise indicated. Escherichia coli  One Shot™ TOP 10 competent cells (F~ mcrA A(mrr-hsdRMS-mcrBC)  (|>80/acZAM15 AlacXIA deoR recA\ araD139 A(ara-leu)1691 galU galK rpsL endA\ nupG) and  the Zero Blunt™ PCR Cloning Kit were from Invitrogen. The QuikChange™  Site-Directed Mutagenesis Kit and Epicurian Coli® XLl-Blue electroporation-competent cells (recAl endAX gyrA96 thi-l hsdWM supEAA relAX lac [V pro AB lacfZAMl5 TnlO (Tet )]) were from Stratagene. PCR DNA fragment purification and plasmid purification r  kits were from Qiagen or Promega.  5.1.2 Oligonucleotides  Synthesis of oligonucleotide primers (Table 5.1) and DNA sequencing was performed by the Nucleic Acids and Peptide Services (NAPS) Unit at the University of British Columbia (Vancouver, Canada).  Chapter 5  Materials and Methods  179  Table 5.1: Primers used in the mutagenesis of C.Jimi xylanase. Name „  v  F f  .  QEr  Oligonucleotide Sequence  QAr jy^f  Forward Q203E mutagenic primer with engineered Kpn I restriction site.  5'-GTC GGG T T C GAG TCG CAC CTC A T C GTC GGC CAG GTA CCG GGC GAC-3'  Reverse Q203E mutagenic primer with engineered Kpn I restriction site.  5 ' - G T C GCC CGG TAC CTG GCC GAC GAT GAG GTG CGA CTC GAA  QAf  Important Characteristics  5'-GTC  ccc GAC-3'  5'-GTC  Forward Q203A mutagenic primer i t h engineered Kpn I restriction site.  GGG T T C GCG TCG CAC CTC A T C  GTC GGC CAG GTA CCG GGC GAC-3'  w  Reverse Q203A mutagenic primer with engineered Kpn I restriction site, j  GCC CGG TAC CTG GCC GAC GAT  GAG GTG CGA CGC GAA CCC GAC- 3 '  5 ' - G T G CGC ATC A C C GAG CTA GCC A T C CGC A T G CGG A C G - 3 '  I i  5 ' - C G T CCG CAT GCG GAT GGC TAG CTC  j  GGT GAT GCG C A C - 3 ' 5'-GCC  GAC A C C GGT AAG GAG CTG TAC  GGC GCC ACG CTC G - 3 ' 5' -  Forward D235A mutagenic primer with engineered Nhe I restriction site. Reverse D235A mutagenic primer with engineered Nhe I restriction site. Forward H80A mutagenic primer with engineered Age I restriction site.  c GAG CGT GGC GCC GTA CAG CTC Reverse H80A mutagenic primer with engineered Age I restriction site.  CTT ACC GGT GTC G G C - 3 '  p^j.  5 ' - G C C GAC A C C GGT AAG GAG CTG TAC  GGC AAC ACG CTC G - 3 ' 5'CTT  HQf  HQr j^^.  c GAG CGT GTT GCC GTA CAG CTC Reverse H80N mutagenic primer ACC GGT GTC G G C - 3 ' with engineered Age I restriction site.  5 ' - G C C GAC ACC GGT AAG GAG CTG TAC  GGC CAG ACG CTC G -3' 5' -  Forward H80N mutagenic primer with engineered Age I restriction site.  Forward H80Q mutagenic primer with engineered Age I restriction site.  c GAG CGT CTG GCC GTA CAG CTC Reverse H80Q mutagenic primer with engineered Age I restriction site.  CTT ACC GGT GTC G G C - 3 '  5'-GTC  GTC GCC GAG AAT GCG ATG GCG  TGG GAC GCC A C C - 3 '  Forward K47A mutagenic primer i t h engineered Bsm I restriction site.  w  Chapter 5  Materials and Methods  Name  Oligonucleotide Sequence  \  GGC GTC C C A C G C CAT C G C ATT  5'-GGT  CTC GGC GAC G A C - 3 ' -vT JL r-  NAi  5'-GAC G  C  C  G  A  180  Important Characteristics  Reverse K47A mutagenic primer with engineered Bsm I restriction site. Forward N44A mutagenic primer „ \_ . . RI restriction site.  AGC GAA TTC AAC CTC GTC GTC , . , „ ^(3.3, with engineered Eco G C C GC"G A T G U  Q  T  Reverse N44A mutagenic primer , . T  NAr  5 ' - C T T CAT CGC G G C CTC GGC GAC GAC GAG GTT GAA TTC GCT G T C - 3 '  ' J EAf  5  ' ~  C  C  T  C  G  T  C  ^_ G  T  C  ATG AAG TGG G A C - 3 JJ^.  5'-GTC  G  C  C  '  °-  G  G  C  G  ^  5'-TCG F P 1  ccc  ™  0  5'-CGA  F  2  P  ccc  , „  &  _  T  \ .  .  ^ E43A mutagenic primer with engineered Bsm I restriction site. Reverse E43A mutagenic primer with j engineered Bsm I restriction site. ;  CCC GGG GAT C C T AGG A C C A C G „  GCA  .  o r w a r <  C C A C T T CAT C G C ATT CGC GGC  GAC GAC GAG G - 3 '  ..  wim engmeered £ c o RI restriction site.^ _  ,  A  ..  Forward flanking primer.  GGC C A C - 3 '  C T C GCC CGA CAG GGG GAT GGG „  CGT GAA ccc GAT C G C - 3 '  „  ,•  Reverse flanking primer.  SP1  5'-GTG  TGG AAT TGT GAG C G G - 3 '  Sequencing primer.  SP2  5'-GTC  GCC GAC CAC T T C G A G - 3 '  Sequencing primer.  Note: Mutated residues are underlined. Mutated amino acid codons are in bold. Restriction sites are in italics.  5.1.3  Site-directed mutagenesis by linear amplification  The genes encoding for cex D235A, H80A, H80N, H80Q, K47A, N44A, and E43A  were linearly amplified  following  the  protocol  QuikChange™ Site-Directed Mutagenesis Kit (Figure 5.1).  outlined  in Stratagene's  Chapter 5  Materials and Methods  \8]_  Figure 5.1: Introducing mutations by the QuikChange™ Site-Directed Mutagenesis method. A supercoiled dsDNA vector with an insert of interest ( • ) and two synthetic oligonucleotide primers (—A—) containing the desired mutation are mixed. The oligonucleotide primers, each complementary to opposite strands of the vector, are extended during temperature cycling using Pfu DNA polymerase to generate mutated plasmid containing staggered nicks. Following temperature cycling, the methylated, nonmutated parental DNA template is digested with Dpn I. The circular, nicked dsDNA is transformed into XLl-Blue supercompetent cells. After transformation, the nicks in the mutated plasmid are repaired by the XLl-Blue supercompetent cells.  The reaction mixture contained 150 ng oligonucleotide primers, 250 uM each of the four deoxynucleoside triphosphates, and 50 ng of plasmid pUC12-l.lcex(PTIS) in 50 u.L of Pfu DNA polymerase reaction buffer containing 8% DMSO.  Plasmid  pUC12-l.lcex(PTIS) contains the 1.4 kb cex gene (274). The reaction was initiated by  Chapter 5  Materials and Methods  l_82_  adding 4 U of Pfu DNA polymerase (Promega). After heating the reaction mixture to 95°C for 30 s, sixteen cycles (30 s at 95°C, 1 min at 55°C, and 9 min at 68°C) were performed in a thermal cycler (GeneAmp® PCR System 2400, Perkin Elmer). Agarose gel electrophoresis confirmed the presence of sufficient amplification product. The parental supercoiled dsDNA was digested with 20 U Dpn I restriction enzyme at 37°C for 1 hour. Epicurian Coli® X L 1-Blue electroporation-competent cells (Stratagene) were subsequently transformed with 1 uL of the Dpn I treated DNA from the amplification reaction using the BioRad GenePulser® II and 0.1 cm cuvettes under the following conditions: 1.8 kV, 25 p.F, 200 Q. The cells were plated on LB agar plates containing 100 ug/mL ampicillin and incubated at 37°C overnight. Single colonies were selected and grown overnight in LB media containing 100 ug/mL ampicillin. Plasmid DNA was isolated using the QIAprep® Spin Miniprep Kit from Qiagen or the Wizard®  Plus  Minipreps DNA Purification System from Promega. Plasmid DNA was stored in water at -20°C.  Restriction endonuclease mapping revealed positive clones, which were  subsequently sequenced to verify the correct mutations. subsequently transformed into electrocompetent  E. coli  The cloned products were  JM101 cells using the BioRad  GenePulser® II and 0.2 cm cuvettes under the following conditions: 2.5 kV, 25 uF, 400 Q. Cells were selected by the ampicillin resistance conferred by pUC12-l.l on LBamp  agar plates. Single colonies were selected and grown overnight in LBamp for long-  term storage in 10% DMSO at -70°C.  Chapter 5  Materials and Methods  4517 BamHI 4499 E c o R I 4 A v r l l 4499 A p o l 4 S t y I 4143  217  183  Sgfl  Pcil  524 NotI 681 E c o N I 700 B s s H I I  979 B b r 7 I  979 Bbsl  3165 B p m l  2448  Sspl  1485 A p a l 1485 PspOHI 1487 S r f l 1651 F s e l 1748 H l u l 1832 B s i ¥ I 1851 S b f l 1864 H i n d l l l 2080 H d e l  Figure 5.2: Restriction map of pUC12-l.lcex(PTlS).  5.1.4 Site-directed mutagenesis by PCR A connecting PCR mutagenic strategy outlined in Figure 5.3 was used to introduce the desired Q203A and Q203E mutations and a silent mutation that would introduce a unique Kpn I restriction site into plasmid pUC12-l.lcex(PTIS).  PCR  Mix, denature and anneal  Figure 5.3: Introducing mutations by in vitro overlap-extension P C R . Two P C R s are performed to produce two fragments that carry overlapping sequences. These two fragments are then mixed, denatured, and annealed to obtain mutant DNAs in a further P C R .  Two separate PCRs were performed using pUC12-l.lcejc(PTlS) as the template and either an outside-forward flanking primer and inside-reverse mutagenic primer, or inside-forward mutagenic primer and outside-reverse flanking primer, to produce two DNA fragments that carry overlapping sequences and intended mutations.  The PCR  reaction mixtures contained 30 ng pUC12-l.lcex(PTlS), 150 ng oligonucleotide primers and 250 uM each of the four deoxynucleoside triphosphates in 50 uL Pwo polymerase buffer containing 8% DMSO. After heating to 95°C for two minutes, the PCR reaction was initiated by adding 5 U of Pwo DNA polymerase (Roche). Fifteen PCR cycles (30 s at 95°C, 30 s at 65°C less 1°C per cycle, and 1.5 min at 72°C) followed by 20 PCR cycles  Chapter 5  Materials and Methods  1_85_  (30 s at 95°C, 30 s at 55°C, and 1.5 min plus 5 s per cycle at 72°C) and a final extension for 5 minutes at 72°C were performed in a thermal cycler (GeneAmp® PCR System 2400, Perkin Elmer). Agarose gel electrophoresis of the PCR reaction mixture revealed two DNA fragments of 780 bp and 1100 bp that were isolated using the QIAEXII DNA Extraction Kit according to the manufacturer's protocol (Qiagen). These two fragments were mixed and heated to 95°C for two minutes.  After adding 5 U of Pwo DNA  polymerase (Roche), 20 PCR cycles were performed to extend the fragments. Fifteen PCR cycles (30 s at 95°C, 30 s at 60°C less 1°C per cycle, and 1.5 min at 72°C) followed by 20 PCR cycles (30 s at 95°C, 30 s at 48°C, and 1.5 min plus 5 s per cycle at 72°C) and a final extension for 5 minutes at 72°C were performed in a thermal cycler (GeneAmp® PCR System 2400, Perkin Elmer). Agarose gel electrophoresis of the PCR reaction mixture failed to reveal the expected 1845 bp fragment. An alternative strategy described below was attempted. The resulting two fragments (1.2 p,g) from the first PCR were digested with 30 U Kpn I restriction endonuclease.  Following purification of the digested PCR products  using the QIAquick™ PCR Purification Kit, a ligation was performed using 4 U T4 DNA ligase (Invitrogen) for 1 hour at 16°C. Subsequently, a blunt-end ligation into vector pCR®-Blunt was performed using the Zero Blunt™ PCR Cloning Kit from Invitrogen. E. coli  One Shot™ TOP 10 competent cells (Invitrogen) were then transformed with the  ligation mixture.  The cells were plated on LB agar plates containing 50 ug/mL  kanamycin and incubated at 37°C overnight. Single colonies were selected and grown overnight at 37°C in LB media containing 50 p.g/mL kanamycin. Plasmid DNA was isolated using the QIAprep® Spin Miniprep Kit from Qiagen or the Wizard®  Plus  Chapter 5  Materials and Methods  186  Minipreps DNA Purification System from Promega. Restriction endonuclease mapping revealed positive clones, which were subsequently sequenced to verify the correct mutations.  5.1.5 Overexpression and purification by active-site mutants of Cex  The E. coli JM101 transformants from the previously prepared DMSO stocks were selected on LBamp (100 ug/mL) agar plates. A single colony was picked and grown for 6 hours in 3 mL of LBamp, and this culture was subsequently used to inoculate 1 L of TYPamp.  After the culture grew to an O D  600  of 2-3 at 30-37°C, 0.2 mM isopropyl  P-D-thiogalactoside (IPTG) was added to induce protein expression and grown for an additional 4 hours at 25 °C. Overexpression of the enzyme was monitored by sampling of both induced and non-induced cells using SDS-polyacrylamide gel electrophoresis. Induced cells were then harvested by centrifugation at 5000 rpm for 20 minutes at 4°C. A cell extract was prepared using BugBuster™ Protein Extraction Reagent. The cell pellet from a 1 L culture was resuspended in 50 mL of BugBuster reagent at room temperature and 50 uL of Benzonase.  EDTA was added to 0.5 mM and PMSF to  1.0 mM. The mixture was incubated with shaking for 10-20 minutes. Following removal of the insoluble cell debris by centrifugation at 16,000 x g for 20 minutes at 4°C, the soluble cell extract was added to 450 mL of 500 mM NaCl in 50 mM sodium phosphate buffer (pH 7.0). EDTA and PMSF were added to 0.5 mM. Cex was purified from the clarified cell extract by affinity chromatography on cellulose. Approximately 25 g of CF1 cellulose (Sigma) was gently stirred with distilled water, allowed to settle, and the "cellofines" were decanted. The cellulose was packed  Chapter 5  Materials and Methods  187_  into an XK50/20 column (Pharmacia), yielding a column volume of approximately 150 mL. The column was attached to an FPLC system (Pharmacia) and equilibrated with 5 column volumes of 500 mM NaCl in 50 mM sodium phosphate buffer (pH 7.0) at a flow rate of 5 mL/min. The soluble cell extract from the 1 L cell culture was applied to the column at a flow rate of 5 mL/min. The column was washed with 2 column volumes of 500 mM NaCl in 50 mM sodium phosphate buffer (pH 7.0) followed by 1 column volume of 50 mM sodium phosphate buffer (pH 7.0) at a flow rate of 5 mL/min. Cex protein was eluted with 2.5 column volumes of distilled H2O at a flow rate of 5 mL/min. The absorbance of the eluate was measured continuously at 280 nm, and appropriate fractions were pooled and passed through a 0.22 pm filter (Millipore). The protein sample was further purified on a 5 mL HiTrap Mono Q HP column attached to an FPLC system (Pharmacia).  The column was washed with 5 column  volumes of 20 mM Tris-HCl buffer (pH 7.5), 5 column volumes of 20 mM Tris-HCl buffer containing 1 M NaCl (pH 7.5), and then 10 column volumes of 20 mM Tris-HCl buffer (pH 7.5) at a flow rate of 5 mL/min. The protein sample was adjusted to pH 8.0 with 20 mM Tris-HCl and applied onto the HiTrap Mono Q HP column at a flow rate of 5 mL/min. The column was washed with 4 column volumes of 20 mM Tris-HCl buffer (pH 7.5). Cex protein was eluted from the anion-exchange column with a linear NaCl gradient from 0 to 500 mM in a buffer containing 20 mM Tris-HCl (pH 7.5). Fractions (8 mL) were collected at a flow rate of 5.0 mL/min and analyzed using SDSpolyacrylamide gel electrophoresis on a Mini-PROTEAN II apparatus (BioRad). Protein bands were visualized by staining with Coomassie Blue.  Fractions containing pure  enzyme were pooled and stored at 4°C. If necessary, the enzyme was concentrated to at  Chapter 5  Materials and Methods  [88  least 2.0 mg/mL using a Centriprep concentrator (30 kDa cutoff) from Amicon. Final protein concentration was calculated from measured absorbance 280 nm using the previously determined extinction coefficient for Cex ( S 2 8 0 =1.61 mL mg~' cm ). -1  5.1.6  Protein mass determination  The molecular weights of purified Cex and Cex mutants were determined by ion spray mass spectrometry.  Mass spectra were recorded on a PE-Sciex API 300 triple-  quadrupole mass spectrometer (Sciex, Thornhill, Ontario, Canada) equipped with an ion spray ion source by Mr. Shouming He. Enzyme samples (10 u.g) were injected into the mass spectrometer to generate the spectra. The masses of the generated mutants were confirmed by comparison between the expected molecular weights and those provided by the spectra.  5.2 Enzyme Kinetics 5.2.1  General materials and procedures  All buffer chemicals and other reagents were obtained from Sigma or Aldrich Chemical Companies unless otherwise noted. The following compounds were prepared and generously provided by co-workers in this laboratory. Xylobiosyl lactam oxime, xylobiosyl deoxynojirimycin, xylobiosyl isofagomine, and xylobiosyl isofagomine lactam were synthesized by Dr. Spencer J. Williams while xylobiosyl imidazole was prepared by Dr. Roland Hoos. 4-Nitrophenyl P-xylobioside, 4-nitrophenyl 2'-deoxyp-xylobioside, 4-nitrophenyl 2'-deoxy-2'-fluoro-P-xylobioside, 4-nitrophenyl 3'-deoxy-  Chapter 5  Materials and Methods  [89  P-xylobioside, 4-nitrophenyl 3'-deoxy-3'-fluoro-P-xylobioside, 4-nitrophenyl 4'-deoxyP-xylobioside, and 4-nitrophenyl 4'-deoxy-4'-fluoro-P-xylobioside were prepared by Dr. Johann Schloegl. 2,4-Dinitrophenyl P-D-cellobioside and 4-nitrophenyl P-D-cellobioside were synthesized by Lloyd Mackenzie while 2,5-dinitrophenyl P-xylobioside and 2-nitrophenyl P-xylobioside were prepared by Dr. Lothar Ziser. All absorbance measurements were recorded on a Unicam UV4 UV/Vis spectrophotometer equipped with a circulating water bath.  5.2.2 Steady state kinetics  Michaelis-Menten parameters for aryl glycosides were determined by continuous measurement of the release of the substituted phenol product using a Unicam UV4 spectrophotometer as described previously (21).  Reactions were monitored at  appropriate wavelengths using the extinction coefficients given in Table 5.5. Phenol pK  a  values used were those reported in Kempton and Withers (21). The buffer used for Cex was 50 mM sodium phosphate buffer, pH 7.00, containing 0.1% w/v bovine serum albumin (BSA) at 37°C. The kinetic studies for Bex were performed at 40°C in 20 mM MES, pH6.00, containing 50 mM NaCl and 0.1% w/v BSA. Concentrations of aryl glycosides were confirmed by total hydrolysis of the glycoside and determination of the final concentration of phenol released. Initial rates of enzyme-catalyzed hydrolysis of aryl glycosides were determined by incubating solutions of the appropriate substrate concentrations  in 1 cm cuvettes within the  spectrophotometer until thermally  equilibrated. Reactions were initiated by the addition of enzyme, and the release of phenol product was monitored at the appropriate wavelength. In order to ensure linear  Chapter 5  Materials and Methods  190  kinetics and to obtain a sufficient absorbance change for accurate calculation of the rates, the concentration of enzyme added and the length of time that the reaction was monitored was selected such that less than 10% of the total substrate was converted to product. Enzyme-catalyzed hydrolysis for each substrate was measured at 8 to 10 different substrate concentrations ranging from about 0.1 K to 7 K , where practical. Values for m  m  K and /c were determined from the initial rates of hydrolysis (V ) versus substrate m  cat  0  concentration by non-linear regression analysis using the computer program GraFit 4.0 (131). In cases where significant transglycosylation was observed a non-linear regression  was performed on data from 0.1 to approximately 2 times K . The values of k and K m  cat  m  so obtained were then compared to those determined from linear regression of the reciprocal data as plotted according to Lineweaver-Burke. In almost all cases the kinetic constants K , V , and k were calculated from a fit to the Michaelis-Menten equation. m  max  cat  For those mutants with very high K values, thus for which saturation behavior could not m  be observed, an accurate value of k /K was obtained from the slope of the Lineweavercat  m  Burk plot, and very approximate estimates of the individual parameters were obtained from the intercepts. In addition, the second-order rate constants (k /K ) were determined from cat  m  progress curves at low substrate concentrations as described in section 5.2.4 on page 192. The individual contribution of each substrate hydroxyl group to transition-state stabilization  with  the wild-type enzyme  was determined  from the relation  AAG =-i?rin^(/c / AT )^/'(/c /AT )^ j , where x represents a substrate analogue or {  cat  m  cat  m  mutant enzyme and y the parent substrate or wild-type enzyme.  Chapter 5  5.2.3  Materials and Methods  191  pH stability studies Enzyme stabilities over the assay time of the pH dependence studies were  examined by incubating enzyme in 50 m M sodium phosphate buffer containing 0.1% w/v B S A at extreme pH values of 3.5, 4.0 and 10.0. The components of the pH incubation mixture are given in Table 5.2.  Table 5.2: pH Incubation mixture for enzyme stability studies.  HB3H  loKll  1  "ff. Volume  <&iy WT E43A N44A K47A H80A I H80N H80Q Q87M Q87Y N169A N126A  \ ol'ume  1 n/\me . ,.\ olimic-  (ijil.);'•_« •'  300 200 " " " " 500 200 " 100 100 100 300 300 3(Xf 100  Bui l e i '  \ olunK  . f  •(ul  30 20 50 20 10 10 10 30 30 30 10  10  io 10 20 20 20 20 10 10 20 20  260 170 440 160 70 70 70 260 260 250 " " " 70  f  e  nil 2.00  •  j  2.48 3.00 1.91 1.46 1.49 1.19 2.13 2.18 2.58 2.90  : !  1 "!  At various time intervals, aliquots of the mixture were removed and injected into another preincubated solution of 2,4DNPC in 50 m M sodium phosphate buffer (pH 7.00) containing 0.1% w/v B S A (Table 5.3).  The half-lives were determined by nonlinear  analysis of the residual activity as a function of time of incubation. In all cases, no more than 10% enzyme death had occurred over a period of 1 hour. determinations of k  cat  Subsequent  and K were then only made at pH values at which the enzyme was m  stable (>95% activity retained) over a 5 minute period.  Chapter 5  Materials and Methods  192  Table 5.3: Standard assay conditions for enzyme stability.  1 n/\ me• • *>* . .*  Iotal Assay \ olume •  WT | E43A N44A K47A H80A H80N H80Q Q87M Q87Y N169A N126A  1 n/MiK  Vlolume'ci'  800 800 800 800 200 200 200 800 800 800 800  0.076 0.22 0.076 ~ 0 . 5 5 "" "' 0.022 0.022 0.022 r  _  o  Q  5  5  0.11 0.055 0.055  -  10 10 10 20 20 " 20 20 10 10 10 20  imii ml > 8.33 x 10" 1.55 x 10~ 7.50 x 10" 4.78 x i o 2.92 x 10" 2.98 x 10" 2.38 x 10"  04 03  04  -03 02  02 02  g  8  g  9.08 2.15 1.45  1 Q  x  x x x  -04  10" 10" 10"  04 03 02  5.2.4 pH dependence studies  The dependence of k JK c  m  on pH for wild-type Cex and mutants was determined  as follows. Appropriate substrate at a concentration of less than less than 1/5 K . in the m  appropriate buffer containing 0.1% w/v BSA and 150 mM NaCl was incubated at 37°C until thermally equilibrated (Table 5.4). Following the addition of 10 uL enzyme, the release of 2,4-dinitrophenolate 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 that yielded pseudo firstorder rate constants at each pH value. At low substrate concentrations, the reaction rates are given by the equation:  V =  ^cat  [E] [S] 0  Chapter 5  Materials and Methods  193  Thus, pseudo first-order rate constants measured correspond to [E]  , from  0  which k JK c  m  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.  Table 5.4: Standard assay conditions for pH profiles.  •.  P  Engine  WT E43A N44A K47A H80A H80N H80Q  I  *  *  Substrate  2.'  2,4DNPC 2,4DNPC 2,4DNPC 2,5DNPX _ PNPX PNPX PNPX  2  2  2  2  Assa\ [SI • 1  w  i'  "  \olume  1  0.0109 0.0109 0.0109 0.0422 0.0586 0.0293 0.0293  "_  800 800 800 _ 800 _ 200 200 200  "  Assav IF. 1 , L  5.24 6.50 7.86 5.01 1.48 1.51 1.20  J  x 10~ x 10~ * 10" x KT x 10" x 10" x 10"° c  c  c  C  c  C  5.2.5 Kinetics of inhibition  K\ values for inhibitors were determined by estimating an approximate K\ value by using a "range-finding" assay at a fixed substrate concentration with a range of inhibitor concentrations (6-10), which encompassed the K value ultimately determined, generally x  from 0.3 to 3 K\. The observed rates were plotted in the form of a Dixon plot and the K\ value was determined (assuming competitive inhibition) by an intersection of this line with a horizontal line drawn through l/V . All reactions were performed at 37°C in max  50 mM sodium phosphate buffer (pH7.00) containing 0.1% w/v BSA. A full K\ determination was then carried out at a series of six different substrate concentrations bracketing the K\ value with each of five inhibitor concentrations bracketing the  Chapter 5  Materials and Methods  194  approximate K\ value. The nonlinear regression analysis program GraFit 4.0 (131) was used for fitting of all such data. The pH dependence for inhibition was determined via the substrate depletion method, with substrate present at less than 1/5 K .  The inhibition constant of the  m  enzyme-inhibitor complex, K\, was calculated from the effect of the inhibitor on the ratio of the two steady-state parameters: 1 +[I] V *cat J app  V ^cat  J  At each pH, appropriate substrate at a concentration of less than 1/5 K in the m  appropriate buffer containing 0.1% w/v BSA and 150 mM NaCl and a range of inhibitor concentrations was incubated at 37°C.  The reaction was initiated upon addition of  enzyme, and the substrate was depleted to obtain  /c b 0  S  The parameter K lk m  determined conveniently under first-order conditions ([S]«. K ). m  constants  *cat  cat  was  The first-order rate  were evaluated graphically from a plot of absorbance versus time. app  The absorbance was monitored at a wavelength that gave a convenient absorbance  K2  r  change. The K\ value was obtained from plot of  V  cat J pp  dependence for inhibition, l/K versus pH was plotted. {  versus [I]. To obtain a pH  k a  Chapter 5  5.2.6  Materials and Methods  1_95_  pKa determinations for inhibitors A l l pH determinations were carried out using a Radiometer P H M 82 pH meter  equipped with an Orion 8103 Ross pH electrode. Dilute solutions of the inhibitors were prepared in deionized water at room temperature.  The amine solution (100 uL) was  acidified by addition of 2-3 uL of HC1. The solution was then back-titrated by addition of 1 uL aliquots of NaOH using a micrometer burette (Gilmont® Instruments). The pH after each addition was recorded and fit to a pH titration curve using GraFit 4.0 (131).  5.2.7  Determination of extinction coefficients Extinction coefficients (s) were measured at 37°C at pH 7.00 in 50 m M sodium  phosphate buffer. Phenols and glycosides were dried in vacuo, weighed, and dissolved in a known volume of buffer. Absorbance values were taken at an appropriate wavelength for five different concentrations of substrate. Extinction coefficients were determined from Beer's Law:  e=  A cxl  where A is the absorbance at a particular wavelength, c is the concentration of solution (mM) and / is the cell pathlength (1 cm). To convert the observed rate of change of absorbance (A^/min) to rate of phenol release (mM/min), the extinction coefficient difference (As) between the phenol and the glycoside was used:  Chapter 5  Materials and Methods  196  A^/min  It should be noted that E iy g  CO  side  values were negligible.  Table 5.5: Extinction coefficients of substituted phenols at 37°C in sodium phosphate buffer, pH 7.00.  IMVehol 1  - •  SufiNiitiici r&t •  2,4-Dinitro ! 2,5-Dinitro  *\li»iiiii-»rc*.l-i nni"i *•* j i nAl^cnivty-- mftrV-iivV-i ..." •'• *400 11.40 0.00  ciiiXI cnV) 11.40  440  4.10  0.00  4.10  4-Nitro  400  9.42  0.00  9.42  2-Nitro  400  2.36  0.00  2.36  ;  REFERENCES  References  198  References 1.  Wolfenden, R., Lu, X., and Young, G. (1998) Spontaneous hydrolysis of glycosides. J. Am. Chem. Soc. 120, 6814-6815.  2.  Davies, G. J., Wilson, K. S., and Henrissat, B. (1997) Nomenclature for sugar-binding subsites in glycosyl hydrolases. Biochem. J. 321, 557-559.  3.  Henrissat, B., Callebaut, I., Fabrega, S., Lehn, P., Mornon, J.-P., and Davies, G. (1995) Conserved catalytic machinery and the prediction of a common fold for several families of glycosyl hydrolases. Proc. Natl. Acad. Sci. USA. 92, 7090-7094.  4.  Henrissat, B., and Bairoch, A. (1996) Updating the sequence-based classification of glycosyl hydrolases. Biochem. J. 316, 695-696.  5.  Henrissat, B., and Davies, G. (1997) Structural and sequence-based classification of glycoside hydrolases. Curr. Opin. Struct. Biol. 7, 637-644.  6.  Davies, G. J., Gloster, T. M., and Henrissat, B. (2005) Recent structural insights into the expanding world of carbohydrate-active enzymes. Curr. ' Opin. Struct. Biol. 15, 637-645.  7.  Sinnott, M . L. (1990) Catalytic mechanism of enzymic glycosyl transfer. Chem. Rev. 90,  8.  1171-1202.  Koshland, D. E. (1953) Stereochemistry and mechanism of enzymatic reactions. Biol. Rev. 28, 416-436.  9.  McCarter, J. D., and Withers, S. G. (1994) Mechanisms of enzymatic glycoside hydrolysis. Curr. Opin. Struct. Biol. 4, 885-892.  10.  Zechel, D. L., and Withers, S. G. (1999) Glycosyl transferase mechanisms, in Comprehensive Natural Products Chemistry  (Poulter, C. D., Ed.) ed., pp 279-  314, Pergamon, Amsterdam. 11.  Mark, B. L., Vocadlo, D. J., Knapp, S., Triggs-Raine, B. L., Withers, S. G., and James, M . N. G. (2001) Crystallographic evidence for substrate-assisted catalysis in a bacterial ^-hexosaminidase. J. Biol. Chem. 276, 10330-10337.  Markovic-Housley, Z., Miglierini, G., Soldatova, L., Rizkallah, P. J., Muller, U., and Schirmer, T. (2000) Crystal structure of hyaluronidase, a major allergen of bee venom. Structure. 8, 1025-1035. Rye, C. S., and Withers, S. G. (2000) Glycosidase mechanisms. Curr. Opin. Chem. Biol. 4,  573-589.  Zechel, D. L., and Withers, S. G. (2001) Dissection of nucleophilic and acidbase catalysis in glycosidases. Curr. Opin. Chem. Biol. 5, 643-649. Vasella, A., Davies, G. J., and Bohm, M . (2002) Glycosidase mechanisms. Curr. Opin. Chem. Biol. 6,  619-629.  Withers, S. G. (2001) Mechanisms of glycosyl transferases and hydrolases. Carbohydr. Polym. 44, 325-337.  Zechel, D. L., and Withers, S. G. (2000) Glycosidase mechanisms: anatomy of a finely tuned catalyst. Acc. Chem. Res. 33, 11-18. Namchuk, M . N., McCarter, J. D., Becalski, A., Andrews, T., and Withers, S. G. (2000) The role of sugar substituents in glycoside hydrolysis. J. Am. Chem. Soc. 122,  1270-1277.  Heightman, T. D., and Vasella, A. T. (1999) Recent insights into inhibition, structure, and mechanism of configuration-retaining glycosidases. Angew. Chem. Int. Ed. 38,  750-770.  Sinnott, M . L., and Withers, S. G. (1974) The (3-galactosidase-catalysed hydrolyses of P-D-galactopyranosyl pyridium salts. Rate-limiting generation of an enzyme-bound galactopyranosyl cation in a process dependent only on aglycone acidity. Biochem. J. 143, 751-762. Kempton, J. B., and Withers, S. G. (1992) Mechanism of Agrobacterium pglucosidase: kinetic studies. Biochemistry. 31, 9961-9969. Tull, D., and Withers, S. G. (1994) Mechanisms of cellulases and xylanases: a detailed kinetic study of the exo-P-l,4-glycanase from Cellulomonas fimi. Biochemistry. 33,  6363-6370.  Sinnott, M . L. (1978) Ions, ion-pairs and catalysis by the lacZ P-galactosidase of Escherichia coli. FEBS Lett. 94, 1-9.  Huang, X., Surry, C , Hiebert, T., and Bennet, A. J. (1995) Hydrolysis of (2deoxy-P-D-glucopyranosyl)pyridinium salts. J. Am. Chem. Soc. 117, 1061410621. Withers, S., Rupitz, K., and Street, I. (1988) 2-Deoxy-2-fluoro-D-glycosyl fluorides. A new class of specific mechanism-based glycosidase inhibitors. J. Biol. Chem. 263, 7929-7932.  Withers, S. G., and Street, I. P. (1988) Identification of a covalent a-Dglucopyranosyl enzyme intermediate formed on a (3-glucosidase. J. Am. Chem. Soc. 110, 8551-8553.  McCarter, J. D., Adam, M . J., and Withers, S. G. (1992) Binding energy and catalysis. Fluorinated and deoxygenated glycosides as mechanistic probes of Escherichia coli (lacZ) p-galactosidase. Biochem. J. 286, 721-727.  Namchuk, M . N., and Withers, S. G. (1995) Mechanism of Agrobacterium pglucosidase: kinetic analysis of the role of noncovalent enzyme/substrate interactions. Biochemistry. 34, 16194-16202. Wolfenden, R., and Kati, W. M . (1991) Testing the limits of protein-ligand binding discrimination with transition-state analogue inhibitors. Acc. Chem. Res. 24,209-215. Braun, C., Brayer, G. D., and Withers, S. G. (1995) Mechanism-based inhibition of yeast a-glucosidase and human pancreatic a-amylase by a new class of inhibitors: 2-deoxy-2,2-difluoro-a-glycosides. J. Biol. Chem. 270, 26778-26781. McCarter, J. D., and Withers, S. G. (1996) 5-Fluoro glycosidases: a new class of mechanism-based inhibitors of both a- and P-glucosidases. J. Am. Chem. Soc. 118, 241-242. White, A., Tull, D., Johns, K., Withers, S. G., and Rose, D. R. (1996) Crystallographic observation of a covalent catalytic intermediate in a Pglycosidase. Nat. Struct. Biol. 3, 149-154.  References  33.  201  Notenboom, V., Birsan, C , Nitz, M., Rose, D. R., Warren, R. A. J., and Withers, S. G. (1998) Insights into transition state stabilization of the P-1,4glycosidase Cex by covalent intermediate accumulation in active site mutants. Nat. Struct. Biol. 5, 812-818.  34.  Vocadlo, D. J., Davies, G. J., Laine, R., and Withers, S. G. (2001) Catalysis by hen egg-white lysozyme proceeds via a covalent intermediate. Nature. 412, 835-838.  35.  Tull, D., Miao, S., Withers, S. G., and Aebersold, R. (1995) Identification of derivatized peptides without radiolabels: tandem mass spectrometric localization of the tagged active-site nucleophiles of two cellulases and a Pglucosidase. Anal. Biochem. 224, 509-514.  36.  Miao, S., Ziser, L., Aebersold, R., and Withers, S. G. (1994) Identification of glutamic acid 78 as the active site nucleophile in Bacillus subtilis xylanase using electrospray tandem mass spectrometry. Biochemistry. 33, 7027-7032.  37.  Miao, S., McCarter, J. D., Grace, M . E., Grabowski, G. A., Aebersold, R., and Withers, S. G. (1994) Identification of Glu340 as the active-site nucleophile in human glucocerebrosidase by use of electrospray tandem mass spectrometry. J. Biol. Chem. 269, 10975-10978.  38.  Wicki, J., Rose, D. R., and Withers, S. G. (2002) Trapping covalent intermediates on P-glycosidases. Methods Enzymol. 354, 84-105.  39.  Mosi, R. M., and Withers, S. G. (2002) Trapping of a-glycosidase intermediates. Methods Enzymol. 354, 64-84.  40.  Withers, S. G., and Aebersold, R. (1995) Approaches to labeling and identification of active site residues in glycosidases. Protein Sci. 4, 361-372.  41.  Street, I. P., Kempton, J. B., and Withers, S. G. (1992) Inactivation of a pglucosidase through the accumulation of a stable 2-deoxy-2-fluoro-a-Dglucopyranosyl-enzyme intermediate: a detailed investigation. Biochemistry. 31, 9970-9978.  42.  Kuroki, R., Weaver, L. H., and Matthews, B. W. (1999) Structural basis of the conversion of the T4 lysozyme into a transglycosidase by reengineering the active site. Proc. Natl. Acad. Sci. USA. 96, 8949-8954.  References  43.  202  Gebler, J. C , Trimbur, D. E., Warren, R. A. J., Aebersold, R., Namchuk, M., and Withers, S. G. (1995) Substrate-induced inactivation of a crippled Pglucosidase mutant: identification of the labeled amino acid and mutagenic analysis of its role. Biochemistry. 34, 14547-14553.  44.  Lawson, S. L., Wakarchuk, W. W., and Withers, S. G. (1996) Effects of both shortening and lengthening the active site nucleophile of Bacillus circulans xylanase on catalytic activity. Biochemistry. 35, 10110-10118.  45.  Wang, Q., Graham, R. W., Trimbur, D., Warren, R. A. J., and Withers, S. G. (1994) Changing enzymatic reaction mechanisms by mutagenesis: conversion of a retaining glucosidase to an inverting enzyme. J. Am. Chem. Soc. 116, 11594-11595.  46.  Zechel, D. L., Reid, S. P., Nashiru, O., Mayer, C , Stall, D., Jakeman, D. L., Warren, R. A. J., and Withers, S. G. (2001) Enzymatic synthesis of carbonfluorine bonds. J. Am. Chem. Soc. 123, 4350-4351.  47.  Mackenzie, L. F., Wang, Q., Warren, R. A. J., and Withers, S. G. (1998) Glycosynthases: mutant glycosidases for oligosaccharide synthesis. J. Am. Chem. Soc. 120,  48.  5583-5584.  Faijes, M., Fairweather, J. K., Driguez, H., and Planas, A. (2001) Oligosaccharide synthesis by coupled e«do-glycosynthases of different specificity: a straightforward preparation of two mixed-linkage hexasaccharide substrates of 1,3/1,4-P-glucanases. Chemistry. 7, 4651-4655.  49.  Fort, S., Boyer, V., Greffe, L., Davies, G. J., Moroz, O., Christiansen, L., Schulein, M., Cottaz, S., and Driguez, H. (2000) Highly efficient synthesis of P(l—>4)-oligo- and -polysaccharides using mutant cellulases. J. Am. Chem. Soc. 122, 5429-5437.  50.  Trincone, A., Perugino, G., Rossi, M., and Moracci, M . (2000) A novel thermophilic glycosynthase that affects branching glycosylation. Bioorg. Med. Chem. Lett. 10,  365-368.  Notenboom, V., Williams, S. J., Hoos, R., Withers, S. G., and Rose, D. R. (2000) Detailed structural analysis of glycosidase/inhibitor interactions: complexes of Cex from Cellulomonasfimiwith xylobiose-derived aza-sugars. 11553-11563.  Biochemistry. 39,  Williams, S. J., Hoos, R., and Withers, S. G. (2000) Nanomolar versus millimolar inhibition by xylobiose-derived azasugars: significant differences between two structurally distinct xylanases. J. Am. Chem. Soc. 122, 22232235. Wang, Q., Trimbur, D., Graham, R., Warren, R. A. J., and Withers, S. G. (1995) Identification of the acid/base catalyst in Agrobacterium faecalis pglucosidase by kinetic analysis of mutants. Biochemistry. 34, 14554-14562. Lawson, S. L., Wakarchuk, W. W., and Withers, S. G. (1997) Positioning the acid/base catalyst in a glycosidase: studies with Bacillus circulans xylanase. Biochemistry. 36,  2257-2265.  Wang, Q., and Withers, S. G. (1995) Substrate-assisted catalysis in glycosidases. J. Am. Chem. Soc. 117, 10137-10138. Fierobe, H.-P., Mirgorodskaya, E., McGuire, K. A., Roepstorff, P., Svensson, B., and Clarke, A. J. (1998) Restoration of catalytic activity beyond wild-type level in glucoamylase from Aspergillus awamori by oxidation of the Glu400-»Cys catalytic-base mutant to cysteinesulfinic acid. Biochemistry. 37, 3743-3752. Fierobe, H.-P., Clarke, A. J., Tull, D., and Svensson, B. (1998) Enzymatic properties of the cysteinesulfinic acid derivative of the catalytic-base mutant Glu400—»Cys of glucoamylase from Aspergillus awamori. Biochemistry. 37, 3753-3759. Roeser, K. R., and Legler, G. (1981) Role of sugar hydroxyl groups in glycoside hydrolysis. Cleavage mechanism of deoxyglucosides and related substrates by P-glucosidase A3 from Aspergillus wentii. Biochim. Biophys. Acta. 657,  321-333.  References  59.  204  Wentworth, D. F., and Wolfenden, R. (1974) Slow binding of D-galactal, a "reversible" inhibitor of bacterial P-galactosidase. Biochemistry. 13, 47154720.  60.  Cleland, W. W., Frey, P. A., and Gerlt, J. A. (1998) The low barrier hydrogen bond in enzymatic catalysis. J. Biol. Chem. 273, 25529-25532.  61.  Davies, G. J., Sinnott, M . L., and Withers, S. G. (1997) Reactions of electrophilic carbon, phosphorus and sulfur, in Comprehensive Biological  Catalysis (Sinnott, M. L., Ed.) Vol. 1, Academic Press, London. 62.  IUPAC-IUB (1980) Eur. J. Biochem. Ill, 295-298.  63.  Davies, G. J., Ducros, V. M., Varrot, A., and Zechel, D. L. (2003) Mapping the conformational itinerary of p-glycosidases by X-ray crystallography. Biochem. Soc. Trans. 31, 523-527.  64.  Blake, C. C. F., Johnson, L. N., Mair, G. A., North, A. C. T., Phillips, D. C , and Sarma, V. R. (1967) Crystallographic studies of the activity of hen eggwhite lysozyme. Proc. R. Soc. Lond. Ser. B Biol. Sci. 167, 378-388.  65.  Ford, L. O., Johnson, L. N., Machin, P. A., Phillips, D. C , and Tjian, R. (1974) Crystal structure of a lysozyme-tetrasaccharide lactone complex. J. Mol. Biol. 88, 349-371.  66.  Hadfield, A. T., Harvey, D. J., Archer, D. B., MacKenzie, D. A., Jeenes, D. J., Radford, S. E., Lowe, G., Dobson, C. M., and Johnson, L. N. (1994) Crystal structure of the mutant D52S hen egg white lysozyme with an oligosaccharide product. J. Mol. Biol. 243, 856-872.  67.  Strynadka, N. C , and James, M. N. (1991) Lysozyme revisited: crystallographic evidence for distortion of an A^-acetylmuramic acid residue bound in site D. J. Mol. Biol. 220, 401-424.  68.  Kuroki, R., Weaver, L. H., and Matthews, B. W. (1993) A covalent enzymesubstrate intermediate with saccharide distortion in a mutant T4 lysozyme. Science. 262, 2030-2033.  69.  Varrot, A., Schiilein, M., and Davies, G. J. (2000) Insights into ligandinduced conformational change in Cel5A from Bacillus agaradhaerens revealed by a catalytically active crystal form. J. Mol. Biol. 297, 819-828.  Davies, G. J., Mackenzie, L., Varrot, A., Dauter, M., Brzozowski, A. M., Schulein, M., and Withers, S. G. (1998) Snapshots along an enzymatic reaction coordinate: analysis of a retaining P-glycoside hydrolase. Biochemistry.  37, 11707-11713.  Sulzenbacher, G., Driguez, H., Henrissat, B., Schulein, M., and Davies, G. J. (1996) Structure of the Fusariwn oxysporum endoglucanase I with a nonhydrolyzable substrate analogue: substrate distortion gives rise to the preferred axial orientation for the leaving group. Biochemistry. 35, 1528015287. Sulzenbacher, G., Schulein, M., and Davies, G. J. (1997) Structure of the endoglucanase I from Fusarium oxysporum: native, cellobiose, and 3,4epoxybutyl p-D-cellobioside-inhibited forms, at 2.3 A resolution. Biochemistry. 36,  5902-5911.  Tews, I., Perrakis, A., Oppenheim, A., Dauter, Z., Wilson, K. S., and Vorgias, C. E. (1996) Bacterial chitobiase structure provides insight into catalytic mechanism and the basis of Tay-Sachs disease. Nat. Struct. Biol. 3, 638-648. Juers, D. H., Heightman, T. D., Vasella, A., McCarter, J. D., Mackenzie, L., Withers, S. G., and Matthews, B. W. (2001) A structural view of the action of Escherichia coli (lacT) P-galactosidase. Biochemistry. 40,  14781-14794.  Sulzenbacher, G., Mackenzie, L. F., Wilson, K. S., Withers, S. G., Dupont, C., and Davies, G. J. (1999) The crystal structure of a 2-fluorocellotriosyl complex of the Streptomyces lividans endoglucanase CelB2 at 1.2 A resolution. Biochemistry. 38, 4826-4833. Notenboom, V., Birsan, C , Warren, R. A. J., Withers, S. G., and Rose, D. R. (1998) Exploring the cellulose/xylan specificity of the P-l,4-glycanase Cex from Cellulomonasfimithrough crystallography and mutation. Biochemistry. 57,4751-4758.  Mackenzie, L. F., Sulzenbacher, G., Divne, C , Jones, T. A., Woldike, H. F., Schulein, M., Withers, S. G., and Davies, G. J. (1998) Crystal structure of the family 7 endoglucanase I (Cel7B) from Humicola insolens at 2.2 A resolution and identification of the catalytic nucleophile by trapping of the covalent glycosyl-enzyme intermediate. Biochem. J. 335, 409-416. Stoddart, J. F. (1971) Stereochemistry of Carbohydrates, in Stereochemistry of Carbohydrates John Wiley and Sons, New York. Sabini, E., Sulzenbacher, G., Dauter, M., Dauter, Z., Jorgensen, P. L., Schulein, M., Dupont, C , Davies, G. J., and Wilson, K. S. (1999) Catalysis and specificity in enzymatic glycoside hydrolysis: a 2>5# conformation for the glycosyl-enzyme intermediate revealed by the structure of the Bacillus agaradhaerens  family 11 xylanase. Chem. Biol. 6, 483-492.  Sabini, E., Wilson, K. S., Danielsen, S., Schulein, M., and Davies, G. J. (2001) Oligosaccharide binding to family 11 xylanases: both covalent intermediate and mutant product complexes display 2,5_g conformations at the active center. Acta Crystallogr. Sect. D Biol. Cryst. 57, 1344-1347. Sidhu, G., Withers, S. G., Nguyen, N. T., Mcintosh, L. P., Ziser, L., and Brayer, G. D. (1999) Sugar ring distortion in the glycosyl-enzyme intermediate of a family G/l 1 xylanase. Biochemistry. 38, 5346-5354. Ducros, V. M., Zechel, D. L., Murshudov, G. N., Gilbert, H. J., Szabo, L., Stoll, D., Withers, S. G., and Davies, G. J. (2002) Substrate distortion by a pmannanase: snapshots of the Michaelis and covalent-intermediate complexes suggest a 7i2 5 conformation for the transition state. Angew. Chem. Int. Ed. ;  Engl. 41,  2824-2827.  Numao, S., Kuntz, D. A., Withers, S. G., and Rose^, D. R. (2003) Insights into the mechanism of Drosophila melanogaster Golgi a-mannosidase II through the structural analysis of covalent reaction intermediates. J. Biol. Chem. 278,  48074-48083.  Deslongchamps, P. (1983) Stereoelectric Effects in Organic Chemistry, in Organic Chemistry Pergamon Press, Oxford.  Kirby, A. J. (1984) Stereoelectronic effects on acetal hydrolysis. Acc. Chem. Res. 77,305-311. Kulkarni, N., Shendye, A., and Rao, M . (1999) Molecular and biotechnological aspects of xylanases. FEMS Microbiol. Rev. 23, 411-456. Subramaniyan, S., and Prema, P. (2002) Biotechnology of microbial xylanases: enzymology, molecular biology, and application. Crit. Rev. Biotechnol. 22, 33-64.  Cleemput, G., Hessing, M., Oort, M . v., Deconynck, M., and Delcour, J. A. (1997) Purification and characterization of a P-D-xylosidase and an endoxylanase from wheat flour. Plant Physiol. 113, 377-386. Shallom, D., and Shoham, Y. (2003) Microbial hemicellulases. Curr. Opin. Microbiol. 6, 219-228.  Beg, Q. K., Kapoor, M., Mahajan, L., and Hoondal, G. S. (2001) Microbial xylanases and their industrial applications: a review. Appl. Microbiol. Biotechnol. 56, 326-338.  Bissoon, S., Christov, L., and Singh, S. (2002) Bleach boosting effects of purified xylanase from Thermomyces lanuginosus SSBP on bagasse pulp. Process Biochem. 37, 567-572.  Malathi, V., and Devegowda, G. (2001) In vitro evaluation of nonstarch polysaccharide digestibility of feed ingredients by enzymes. Poult. Sci. 80, 302-305. Mathlouthi, N., Saulnier, L., Quemener, B., and Larbier, M. (2002) Xylanase, (3-glucanase, and other side enzymatic activities have greater effects on the viscosity of several feedstuffs than xylanase and P-glucanase used alone or in combination. J. Agric. Food Chem. 50, 5121-5127. Wong, K. K., Tan, L. U., and Saddler, J. N. (1988) Multiplicity of p-1,4xylanase in microorganisms: functions and applications. Microbiol. Rev. 52, 305-317. Lynd, L. R., Cushman, J. FL, Nichols, R. J., and Wyman, C. E. (1991) Fuel ethanol from cellulosic biomass. Science. 251, 1318-1323.  Harbak, L., and Thygesen, H. V. (2002) Safety evaluation of a xylanase expressed in Bacillus subtilis. Food Chem. Toxicol. 40, 1-8. Courtin, C. M , Roelants, A., and Delcour, J. A. (1999) Fractionationreconstitution experiments provide insight into the role of endoxylanases in bread-making. J. Agric. Food Chem. 47, 1870-1877. Ingelbrecht, J. A., Verwimp, T., and Delcour, J. A. (2000) Endoxylanases in durum wheat semolina processing: solubilization of arabinoxylans, action of endogenous inhibitors, and effects on rheological properties. J. Agric. Food Chem. 48, 2017-2022.  Figueroa-Espinoza, M . C , Poulsen, C , Soe, J. B., Zargahi, M . R., and Rouau, X. (2002) Enzymatic solubilization of arabinoxylans from isolated rye pentosans and rye flour by different enafo-xylanases and other hydrolyzing enzymes. Effect of a fungal laccase on the flour extracts oxidative gelation. J. Agric. Food Chem. 50, 6473-6484.  Mielenz, J. R. (2001) Ethanol production from biomass: technology and commercialization status. Curr. Opin. Microbiol. 4, 324-329. Galbe, M., and Zacchi, G. (2002) A review of the production of ethanol from softwood. Appl. Microbiol. Biotechnol. 59, 618-628. Chandrakant, P., and Bisaria, V. S. (1998) Simultaneous bioconversion of cellulose and hemicellulose to ethanol. Crit. Rev. Biotechnol. 18, 295-331. Sommer, P., Georgieva, T., and Ahring, B. K. (2004) Potential for using thermophilic anaerobic bacteria for bioethanol production from hemicellulose. Biochem. Soc. Trans. 32, 283-289.  O'Neill, G. P., Goh, S. EC., Warren, R. A. J., Kilburn, D. G., and R. C. Miller, J. (1986) Structure of the gene encoding the exoglucanase of Cellulomonas fimi. Gene. 44, 325-330.  MacLeod, A. M., Lindhorst, T., Withers, S. G., and Warren, R. A. J. (1994) The acid/base catalyst in the exoglucanase/xylanase from Cellulomonas fimi is glutamic acid 127: evidence from detailed kinetic studies of mutants. Biochemistry. 33, 6371-6376.  References  106.  209  Gilkes, N. R., Henrissat, B., Kilburn, D. G., R. C. Miller, J., and Warren, R. A. J. (1991) Domains in microbial P-l,4-glycanases: sequence conservation, function, and enzyme families. Microbiol. Rev. 55, 303-315.  107.  Withers, S. G., Dombroski, D., Berven, L. A., Kilburn, D. G., Jr, R. C. M., Warren, R. A., and Gilkes, N. R. (1986) Direct K n.m.r. determination of the l  stereochemical course of hydrolyses catalysed by glucanase components of the cellulase complex. Biochem. Biophys. Res. Commun. 139, 487-494. 108.  Tull, D., Withers, S. G., Gilkes, N. R., Kilburn, D. G., Warren, A. J., and Aebersold, R. (1991) Glutamic acid 274 is the nucleophile in the active site of a "retaining" exoglucanase from Cellulomonas fimi. J. Biol. Chem. 266, 15621-15625.  109.  MacLeod, A. M., Tull, D., Rupitz, K , Warren, R. A. J., and Withers, S. G. (1996) Mechanistic consequences of mutation of active site carboxylates in a retaining P-l,4-glycanase from Cellulomonas fimi. Biochemistry. 35, 1316513172.  110.  Tull, D., Burgoyne, D. L., Chow, D. T., Withers, S. G., and Aebersold, R. (1996) A mass spectrometry-based approach for probing enzyme active sites: identification of Glu 127 in Cellulomonas fimi exoglycanase as the residue  modified by Af-bromoacetyl cellobiosylamine. Anal. Biochem. 234, 119-125. 111.  Ziser, L., Setyawati, I., and Withers, S. G. (1995) Syntheses and testing of substrates and mechanism-based inactivators for xylanases. Carbohydr. Res. 274, 137-153.  112.  White, A., Withers, S. G., Gilkes, N. R., and Rose, D. R. (1994) Crystal structure of the catalytic domain of the P-l,4-glycanase Cex from Cellulomonasfimi.Biochemistry. 33,  113.  12546-12552.  Sung, W. L., Luk, C. K , Zahab, D. M., and Wakarchuk, W. (1993) Overexpression of the Bacillus subtilis and circulans xylanases in Escherichia coli. Protein Expr. Purif. 4, 200-206.  Birsan, C , Johnson, P., Joshi, M., MacLeod, A., Mcintosh, L., Monem, V., Nitz, M., Rose, D. R., Tull, D., Wakarchuck, W. W., Wang, Q., Warren, R. A., White, A., and Withers, S. G. (1998) Mechanisms of cellulases and xylanases. Biochem. Soc. Trans. 26, 156-160. Mcintosh, L. P., Hand, G., Johnson, P. E., Joshi, M . D., Korner, M., Plesniak, L. A., Ziser, L., Wakarchuk, W. W., and Withers, S. G. (1996) The pK of the a  general acid/base carboxyl group of a glycosidase cycles during catalysis: a l^C-NMR study of Bacillus circulans xylanase. Biochemistry. 35, 99589966. Joshi, M . D., Hedberg, A., and Mcintosh, L. P. (1997) Complete measurement of the p/C values of the carboxyl and imidazole groups in a  Bacillus circulans  xylanase. Protein Sci. 6, 2667-2670.  Plesniak, L. A., Connelly, G. P., Wakarchuk, W. W., and Mcintosh, L. P. (1996) Characterization of a buried neutral histidine residue in Bacillus circulans  xylanase: NMR assignments, pH titration, and hydrogen exchange.  Protein Sci. 5,  2319-2328.  Joshi, M . D., Sidhu, G., Pot, I., Brayer, G. D., Withers, S. G., and Mcintosh, L. P. (2000) Hydrogen bonding and catalysis: a novel explanation for how a single amino acid substitution can change the pH optimum of a glycosidase. J. Mol. Biol. 299,  255-279.  Wakarchuk, W. W., Campbell, R. L., Sung, W. L., Davoodi, J., and Yaguchi, M . (1994) Mutational and crystallographic analyses of the active site residues of the Bacillus circulans xylanase. Protein Sci. 3, 467-475. Davies, G., and Henrissat, B. (1995) Structures and mechanisms of glycosyl hydrolases. Structure. 3, 853-859. Charnock, S. J., Spurway, T. D., Xie, H., Beylot, M.-H., Virden, R., Warren, R. A. J., Hazlewood, G. P., and Gilbert, H. J. (1998) The topology of the substrate binding clefts of glycosyl hydrolase family 10 xylanases are not conserved. J. Biol. Chem. 273, 32187-32199.  References  122.  211  Harris, G. W., Jenkins, J. A., Connerton, I., Cummings, N., Leggio, L. L., Scott, M., Hazlewood, G. P., Laurie, J. I., Gilbert, H. J., and Pickersgill, R. W. (1994) Structure of the catalytic core of the family F xylanase from Pseudomonasfluorescensand identification of the xylopentaose-binding  sites. Structure. 2, 1107-1116. 123.  Derewenda, U., Swenson, L., Green, R., Wei, Y., Morosoli, R., Shareck, F., Kluepfel, D., and Derewenda, Z. S. (1994) Crystal structure, at 2.6-A resolution, of the Streptomyces lividans xylanase A, a member of the F family of p-l,4-D-glycanases. J. Biol. Chem. 269, 20811-20814.  124.  Dominguez, R , Souchon, H., Spinelli, S., Dauter, Z., Wilson, K. S., Chauvaux, S., Beguin, P., and Alzari, P. M . (1995) A common protein fold and similar active site in two distinct families of p-glycanases. Nat. Struct. Biol. 2, 569-576.  125.  Pell, G., Szabo, L., Charnock, S. J., Xie, H., Gloster, T. M., Davies, G. J., and Gilbert, H. J. (2004) Structural and biochemical analysis of Cellvibrio japonicus xylanase 10C: how variation in substrate-binding cleft influences  the catalytic profile of family GH-10 xylanases. J. Biol. Chem. 279, 1177711788. 126.  Ducros, V., Charnock, S. J., Derewenda, U., Derewenda, Z. S., Dauter, Z., Dupont, C , Shareck, F., Morosoli, R., Kluepfel, D., and Davies, G. J. (2000) Substrate specificity in glycoside hydrolase family 10. Structural and kinetic analysis of the Streptomyces lividans xylanase 10A. J. Biol. Chem. 275, 23020-23026.  127.  Xie, H., Flint, J., Vardakou, M., Lakey, J. H., Lewis, R. J., Gilbert, H. J., and Dumon, C. (2006) Probing the structural basis for the difference in thermostability displayed by family 10 xylanases. J. Mol. Biol. 360, 157-167.  128.  Vardakou, M., Flint, J., Christakopoulos, P., Lewis, R. J., Gilbert, H. J., and Murray, J. W. (2005) A family 10 Thermoascus aurantiacus xylanase utilizes arabinose decorations of xylan as significant substrate specificity determinants. J. Mol. Biol. 352, 1060-1067.  References  129.  212  Payan, F., Leone, P., Porciero, S., Furniss, C , Tahir, T., Williamson, G., Durand, A., Manzanares, P., Gilbert, H. J., Juge, N., and Roussel, A. (2004) The dual nature of the wheat xylanase protein inhibitor XIP-I: structural basis for the inhibition of family 10 and family 11 xylanases. J. Biol. Chem. 279, 36029-36037.  130.  Renard, M., and Fersht, A. R. (1973) Anomalous pH dependence of &cat^M in enzyme reactions. Rate constants for the association of chymotrypsin with substrates. Biochemistry. 12, 4713-4718.  131.  Leatherbarrow, R. J. (1989) Erithacus Software Limited,  132.  Gilkes, N. R., Langsford, M. L., Kilburn, D. G., Robert C. Miller, J., and Warren, R. A. J. (1984) Mode of action and substrate specificities of cellulases from cloned bacterial genes. J. Biol. Chem. 259, 10455-10459.  133.  Charnock, S. J., Lakey, J. H., Virden, R., Hughes, N., Sinnott, M . L., Hazlewood, G. P., Pickersgill, R., and Gilbert, H. J. (1997) Key residues in subsite F play a critical role in the activity of Pseudomonas fluorescens subspecies cellulosa xylanase A against xylooligosaccharides but not against highly polymeric substrates such as xylan. J. Biol. Chem. 272, 2942-2951.  134.  Cleland, W. W. (1975) What limits the rate of an enzyme-catalyzed reaction? Acc. Chem. Res.  135.  §,145-151.  Albery, W. J., and Knowles, J. R. (1976) Evolution of enzyme function and the development of catalytic efficiency. Biochemistry. 15, 5631-5640.  136.  Fersht, A. (1999) Structure and mechanism in protein science: a guide to enzyme catalysis and protein folding,  in W. H. Freeman and Company, New  York. 137.  Dale, M . P., Kopfler, W. P., Chait, I., and Byers, L. D. (1986) P-Glucosidase: substrate, solvent, and viscosity variation as probes of the rate-limiting steps. Biochemistry. 25,  138.  2522-2529.  Moreau, A., Roberge, M., Manin, C , Shareck, F., Kluepfel, D., and Morosoli, R. (1994) Identification of two acidic residues involved in the catalysis of xylanase A from Streptomyces lividans. Biochem. J. 302, 291-295.  Knowles, J. R. (1976) The intrinsic pK -values of functional groups in a  enzymes: improper deductions from the pH-dependence of steady-state parameters. Crit. Rev. Biochem. 4, 165-173. Biely, P., Kratky, Z., and Vrsanska, M . (1981) Substrate-binding site of endo1,4-P-xylanase of the yeast Cryptococcus albidus. Eur. J. Biochem. 119, 559564. Moreau, A., Shareck, F., Kluepfel, D., and Morosoli, R. (1994) Alteration of the cleavage mode and of the transglycosylation reactions of the xylanase A of Streptomyces lividans 1326 by site-directed mutagenesis of the Asnl73 residue. Eur. J. Biochem. 219, 261-266. Fersht, A. R. (1984) Basis of biological specificity. Trends Biochem. Sci. 9, 145-147. Bourne, Y., Tilbeurgh, H. v., and Cambillau, C. (1993) Protein-carbohydrate interactions. Curr. Opin. Struct. Biol. 3, 681-686. Bundle, D. R., and Young, N. M. (1992) Carbohydrate-protein interactions in antibodies and lectins. Curr. Opin. Struct. Biol. 2, 666-673. Quiocho, F. A. (1993) Probing the atomic interactions between proteins and carbohydrates. Biochem. Soc. Trans. 21, 442-448.  Vyas, N. K. (1991) Atomic features of protein-carbohydrate interactions. Curr. Opin. Struct. Biol. 1, 732-740.  Pauling, L. (1946) Chem. Eng. News. 24, 1375. Williams, S. J., and Davies, G. J. (2001) Protein-carbohydrate interactions: learning lessons from nature. Trends Biotechnol. 19, 356-362. Gloster, T. M., Roberts, S., Ducros, V. M.-A., Perugino, G., Rossi, M., Hoos, R., Moracci, M., Vasella, A., and Davies, G. J. (2004) Structural studies of the P-glycosidase from Sulfolobus solfataricus in complex with covalently and noncovalently bound inhibitors. Biochemistry. 43, 6101-6109. Varrot, A., and Davies, G. J. (2003) Direct experimental observation of the hydrogen-bonding network of a glycosidase along its reaction coordinate revealed by atomic resolution analyses of endoglucanase Cel5A. Acta Crystallogr. D Biol. Crystallogr. 59,  447-452.  References  151.  214  Fersht, A. R., Shi, J. P., Knill-Jones, J., Lowe, D. M., Wilkinson, A. J., Blow, D. M., Brick, P., Carter, P., Waye, M. M . Y., and Winter, G. (1985) Hydrogen bonding and biological specificity analysed by protein engineering. Nature. 314,  152.  235-238.  Fersht, A. R. (1987) Dissection of the structure and activity of the tyrosyltRNA synthetase by site-directed mutagenesis. Biochemistry. 26, 8031-8037.  153.  Sierks, M . R., and Svensson, B. (2000) Energetic and mechanistic studies of glucoamylase using molecular recognition of maltose OH groups coupled with site-directed mutagenesis. Biochemistry. 39, 8585-8592.  154.  Frandsen, T. P., Staffer, B. B., Palcic, M . M., Hof, S., and Svensson, B. (1996) Structure and energetics of the glucoamylase-isomaltose transitionstate complex probed by using modeling and deoxygenated substrates coupled with site-directed mutagenesis. J. Mol. Biol. 263, 79-89.  155.  Piotukh, K., Serra, V., Borriss, R , and Planas, A. (1999) Proteincarbohydrate interactions defining substrate specificity in Bacillus 1,3-1,4-pD-glucan 4-glucanohydrolases as dissected by mutational analysis. Biochemistry. 38,  156.  16092-16104.  Gaiser, O. J., Piotukh, K., Ponnuswamy, M. N., Planas, A., Borriss, R., and Heinemann, U. (2006) Structural basis for the substrate specificity of a Bacillus  157.  1,3-1,4-P-glucanase. J. Mol. Biol. 357, 1211-1225.  Street, I. P., Armstrong, C. R , and Withers, S. G. (1986) Hydrogen bonding and specificity. Fluorodeoxy sugars as probes of hydrogen bonding in the glycogen phosphorylase-glucose complex. Biochemistry. 25, 6021-6027.  158.  Street, I. P., Rupitz, K., and Withers, S. G. (1989) Fluorinated and deoxygenated substrates as probes of transition-state structure in glycogen phosphorylase. Biochemistry. 28, 1581-1587.  159.  Percival, M . D., and Withers, S. G. (1992) Binding energy and catalysis: deoxyfluoro sugars as probes of hydrogen bonding in phosphoglucomutase. Biochemistry. 31, 498-505.  Eis, C , and Nidetzky, B. (2002) Substrate-binding recognition and specificity of trehalose phosphorylase from Schizophyllum commune examined in steady-state kinetic studies with deoxy and deoxyfluoro substrate analogues and inhibitors. Biochem. J. 363, 335-340. Jencks, W. P. (1969) Catalysis in Chemistry and Enzymology, in Catalysis in Chemistry and Enzymology McGraw-Hill, New York. Klotz, I. M., and Franzen, J. S. (1962) Hydrogen bonds between model peptide groups in solution. J. Am. Chem. Soc. 84, 3461-3466. Hine, J. (1972) Structural effects on rates and equilibriums. Hydrogenbonded intermediates and stepwise mechanisms for proton-exchange reactions between oxygen atoms in hydroxylic solvents. J. Am. Chem. Soc. 94,5766-5771. West, R., Powell, D. L., Whatley, L. S., Lee, M . K. T., and Schleyer, P. v. R. (1962) The relative strengths of alkyl halides as proton acceptor groups in hydrogen bonding. J. Am. Chem. Soc. 84, 3221-3222. Murray-Rust, P., Stallings, W. C , Monti, C. T., Preston, R. K., and Glusker, J. P. (1983) Intermolecular interactions of the carbon-fluorine bond: the crystallographic environment of fluorinated carboxylic acids and related structures. J. Am. Chem. Soc. 105, 3206-3214. Harrell, S. A., and McDaniel, D. H. (1964) Strong hydrogen bonds. II. The hydrogen difluoride ion. J. Am. Chem. Soc. 86, 4497-4497. Howard, J. A. K., Hoy, V. J., O'Hagan, D., and Smith, G. T. (1996) How good is fluorine as a hydrogen bond acceptor? Tetrahedron. 52, 12613-12622. Shimoni, L., and Glusker, J. P. (1994) The geometry of intermolecular interactions in some crystalline fluorine-containing organic compounds. Struct. Chem. 5,  383-397.  Brammer, L., Bruton, E. A., and Sherwood, P. (2001) Understanding the behavior of halogens as hydrogen bond acceptors. Cryst. Growth Des. 1, 277290.  References  170.  216  Carosati, E., Sciabola, S., and Cruciani, G. (2004) Hydrogen bonding interactions of covalently bonded fluorine atoms: from crystallographic data to a new angular function in the GRID force field. J. Med. Chem. 47, 51145125.  171.  Withers, S. G., MacLennan, D. J., and Street, I. P. (1986) The synthesis and hydrolysis of a series of deoxyfluoro-D-glucopyranosyl phosphates. Carbohydr. Res. 154, 127-144.  172.  Withers, S. G., Percival, M . D., and Street, I. P. (1989) The synthesis and hydrolysis of a series of deoxy- and deoxyfluoro-ct-D- "glucopyranosyl" phosphates. Carbohydr. Res. 173, 43-66.  173.  Withers, S. G., Street, I. P., and Rettig, S. J. (1986) The preferred conformation of 2-fluoro-2-deoxy (3-D-mannopyranosyl fluoride. An X-ray crystallographic and 2-dimensional proton nuclear magnetic resonance study. Can. J. Chem. 64, 232-236.  174.  Torronen, A., and Rouvinen, J. (1997) Structural and functional properties of low molecular weight endo-l,4-P-xylanases.J Biotechnol. 57, 137-149.  175.  Damager, I., Numao, S., Chen, H., Brayer, G. D., and Withers, S. G. (2004) Synthesis and characterisation of novel chromogenic substrates for human pancreatic a-amylase. Carbohydr. Res. 339, 1727-1737.  176.  Burmeister, W. P., Cottaz, S., Driguez, H., Iori, R., Palmieri, S., and Henrissat, B. (1997) The crystal structures of Sinapis alba myrosinase and a covalent glycosyl-enzyme intermediate provide insights into the substrate recognition and active-site machinery of an ^-glycosidase. Structure. 5, 663675.  177.  Sobolev, V., Sorokine, A., Prilusky, J., Abola, E. E., and Edelman, M . (1999) Automated analysis of interatomic contacts in proteins. Bioinformatics. 15, 327-332.  178.  Withers, S. G., and Rupitz, K. (1990) Measurement of active-site homology between potato and rabbit muscle a-glucan phosphorylases through use of a linear free energy relationship. Biochemistry. 29, 6405-6409.  References  179.  217  Sierks, M . R., Bock, K., Refn, S., and Svensson, B. (1992) Active site similarities of glucose dehydrogenase, glucose oxidase, and glucoamylase probed by deoxygenated substrates. Biochemistry. 31, 8972-8977.  180.  Henrissat, B. (1991) A classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem. J. 280, 309-316.  181.  Henrissat, B., and Bairoch, A. (1993) New families in the classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem. J. 293, 781-788.  182.  Biely, P., Vrsanska, M., Tenkanen, M., and Kluepfel, D. (1997) Endo-P-1,4xylanase families: differences in catalytic properties. J. Biotech. 57, 151-166.  183.  Williams, S. J., Notenboom, V., Wicki, J., Rose, D. R., and Withers, S. G. (2000) A new, simple, high-affinity glycosidase inhibitor: analysis of binding through X-ray crystallography, mutagenesis, and kinetic analysis. J. Am. Chem. Soc. 122,  184.  4229-4230.  Poon, D. K. Y., Schubert, M., Au, J., Okon, M., Withers, S. G., and Mcintosh, L. P. (2006) Unambiguous determination of the ionization state of a glycoside hydrolase active site lysine by ! H - N heteronuclear correlation 15  spectroscopy. J. Am. Chem. Soc. 128, 15388-15389. 185.  Schubert, M., Poon, D. K. Y., Wicki, J., Tarling, C. A., Kwan, E. M., Nielsen, J. E., Withers, S. G., and Mcintosh, L. P. (2007) Probing electrostatic interactions along the reaction pathway of a glycoside hydrolase: histidine characterization by NMR spectroscopy. Biochemistry, submitted,  186.  Money, V. A., Smith, N . L., Scaffidi, A., Stick, R. V., Gilbert, H. J., and Davies, G. J. (2006) Substrate distortion by a lichenase highlights the different conformational itineraries harnessed by related glycoside hydrolases. Angew. Chem. Int. Ed. 45, 5136-5140.  187.  Eyring, H. (1935) The activated complex and the absolute rate of chemical reactions. Chem. Rev. 17, 65-77.  188.  Wolfenden, R. (1969) Transition state analogues for enzyme catalysis. Nature. 223,  704-705.  References  189.  218  Wolfenden, R. (1972) Analog approaches to the structure of the transition state in enzyme reactions. Acc. Chem. Res. 5, 10-18.  190.  Wolfenden, R. (1976) Transition-state analog inhibitors and enzyme catalysis. Annu. Rev. Biophys. Bioeng. 5,  191.  271-306.  Radzicka, A., and Wolfenden, R. (1995) Transition-state and multisubstrate analog inhibitors. Methods Enzymol. 249, 284-312.  192.  Schramm, V. L. (1998) Enzymatic transition states and transition state analog design. Annu. Rev. Biochem. 67, 693-720.  193.  Morrison, J. F., and Walsh, C. T. (1988) The behavior and significance of slow-binding enzyme inhibitors. Adv. Enzymol. Relat. Area Mol. Biol. 61, 201-301.  194.  Mader, M . M., and Bartlett, P. A. (1997) Binding energy and catalysis: the implications for transition-state analogs and catalytic antibodies. Chem. Rev. 97, 1281-1302.  195.  Schramm, V. L. (2003) Enzymatic transition state poise and transition state analogues. Acc. Chem. Res. 36, 588-596.  196.  Schramm, V. L. (2005) Enzymatic transition states: thermodynamics, dynamics and analogue design. Arch. Biochem. Biophys. 433, 13-26.  197.  Cleland, W. W. (1995) Isotope effects: determination of enzyme transition state structure. Meth. Enzymol. 249, 341-373.  198.  Schramm, V. L. (1999) Enzymatic transition-state analysis and transitionstate analogs. Methods Enzymol. 308, 301-355.  199.  Berti, P. J. (1999) Determining transition states from kinetic isotope effects. Methods Enzymol. 308,  200.  355-397.  Lewandowicz, A., and Schramm, V. L. (2004) Transition state analysis for human and Plasmodium falciparum purine nucleoside phosphorylases. Biochemistry. 43,  1458-1468.  Mendieta, J., Martin-Santamaria, S., Priego, E.-M., Balzarini, J., Camarasa, M.-J., Perez-Perez, M.-J., and Gago, F. (2004) Role of histidine-85 in the catalytic mechanism of thymidine phosphorylase as assessed by targeted molecular dynamics simulations and quantum mechanical calculations. Biochemistry.  43,405-414.  Birck, M . R., and Schramm, V. L. (2004) Nucleophilic participation in the transition state for human thymidine phosphorylase. J. Am. Chem. Soc. 126, 2447-2453. Garcia-Viloca, M., Gao, J., Karplus, M., and Truhlar, D. G. (2004) How enzymes work: analysis by modern rate theory and computer simulations. Science. 303, 186-195.  Gao, J., Ma, S., Major, D. T., Nam, K., Pu, J., and Truhlar, D. G. (2006) Mechanisms and free energies of enzymatic reactions. Chem. Rev. 106, 31883209. Bruice, T. C. (2006) Computational approaches: reaction trajectories, structures, and atomic motions. Enzyme reactions and proficiency. Chem. Rev. 706,3119-3139. Garcia-Viloca, M., Truhlar, D. G., and Gao, J. (2003) Reaction-path energetics and kinetics of the hydride transfer reaction catalyzed by dihydrofolate reductase. Biochemistry. 42, 13558-13575. Wolfenden, R., and Snider, M . J. (2001) The depth of chemical time and the power of enzymes as catalysts. Acc. Chem. Res. 34, 938-945. Lewandowicz, A., Ringia, E. A. T., Ting, L.-M., Kim, K., Tyler, P. C , Evans, G. B., Zubkova, O. V., Mee, S., Painter, G. F., Lenz, D. H., Furneaux, R. H., and Schramm, V. L. (2005) Energetic mapping of transition state analogue interactions with human and Plasmodium falciparum purine nucleoside phosphorylases. J. Biol. Chem. 280, 30320-30328. Singh, V., Shi, W., Evans, G. B., Tyler, P. C , Furneaux, R. H., Almo, S. C , and Schramm, V. L. (2004) Picomolar transition state analogue inhibitors of human 5'-methylthioadenosine phosphorylase and X-ray structure with MTimmucillin-A. Biochemistry. 43, 9-18.  References  210.  220  Guo, H., Rao, N., Xu, Q., and Guo, H. (2005) Origin of tight binding of a near-perfect transition-state analogue by cytidine deaminase: implications for enzyme catalysis. J. Am. Chem. Soc. 127, 3191-3197.  211.  Singh, V., Evans, G. B., Lenz, D. H., Mason, J. M., Clinch, K., Mee, S., Painter, G. F., Tyler, P. C , Furneaux, R. H., Lee, J. E., Howell, P. L., and Schramm, V. L. (2005) Femtomolar transition state analogue inhibitors of 5'methylthioadenosine/5-adenosylhomocysteine nucleosidase from Escherichia coli. J. Biol. Chem. 280,  212.  18265-18273.  Lolis, E., and Petsko, G. A. (1990) Transition-state analogs in protein crystallography: probes of the structural source of enzyme catalysis. Annu. Rev. Biochem. 59, 597-630.  213.  Kirby, A. J. (1994) Crystallographic approaches to transition state structures, in Advances in Physical Organic Chemistry, Vol 29 ed., pp 87-183, Academic Press Ltd, London.  214.  Wolfenden, R. (1999) Conformational aspects of inhibitor design: enzymesubstrate interactions in the transition state. Bioorg. Med. Chem. 7, 647-652.  215.  Gloster, T. M., Macdonald, J. M., Tarling, C. A., Stick, R. V., Withers, S. G., and Davies, G. J. (2004) Structural, thermodynamic, and kinetic analyses of tetrahydrooxazine-derived inhibitors bound to (3-glucosidases. J. Biol. Chem. 279, 49236-49242.  216.  Gloster, T. M., Madsen, R , and Davies, G. J. (2006) Dissection of conformationally restricted inhibitors binding to a p-glucosidase. Chembiochem. 7, 738-742.  217.  Gloster, T. M., Roberts, S., Perugino, G., Rossi, M., Moracci, M., Panday, N., Terinek, M., Vasella, A., and Davies, G. J. (2006) Structural, kinetic, and thermodynamic analysis of glucoimidazole-derived glycosidase inhibitors. Biochemistry. 45,  11879-11884.  Zechel, D. L., Boraston, A. B., Gloster, T., Boraston, C. M., Macdonald, J. M., Tilbrook, D. M. G., Stick, R. V., and Davies, G. J. (2003) Iminosugar glycosidase inhibitors: structural and thermodynamic dissection of the binding of isofagomine and 1-deoxynojirimycin to P-glucosidases. J. Am. Chem. Soc. 125,  14313-14323.  Lillelund, V. H., Jensen, H. H., Liang, X. F., and Bols, M . (2002) Recent developments of transition-state analogue glycosidase inhibitors of nonnatural product origin. Chem. Rev. 102, 515-553. Cha, S. (1975) Tight-binding inhibitors: kinetic behavior. Biochem. Pharmacol. 24,  2177-2185.  Perdicakis, B., Montgomery, H. J., Guillemette, J. G., and Jervis, E. (2005) Analysis of slow-binding enzyme inhibitors at elevated enzyme concentrations. Anal. Biochem. 337, 211-223. Legler, G. (1999) in Iminosugars as glycosidase inhibitors (Stiitz, A., Ed.) ed., pp 31-67, Wiley-VCH, Weinheim. Bartlett, P. A., and Marlowe, C. K. (1987) Possible role for water dissociation in the slow binding of phosphorus-containing transition-state-analogue inhibitors of thermolysin. Biochemistry. 26, 8553-8561. Waley, S. G. (1993) The kinetics of slow-binding and slow, tight-binding inhibition: the effects of substrate depletion. Biochem. J. 294, 195-200. Frieden, C , Kurz, L. C , and Gilbert, H. R. (1980) Adenosine deaminase and adenylate deaminase: comparative kinetic studies with transition state and ground state analog inhibitors. Biochemistry. 19, 5303-5309. Westerik, J. O. C , and Wolfenden, R. (1972) Aldehydes as inhibitors of papain../. Biol. Chem. 247, 8195-8197. Thompson, R..C. (1973) Use of peptide aldehydes to generate transition-state analogs of elastase. Biochemistry. 12, 47-51. Lienhard, G. E. (1973) Science. 180, 149-154. Bartlett, P. A., and Marlowe, C. K. (1983) Phosphonamidates as transitionstate analogue inhibitors of thermolysin. Biochemistry. 22, 4618-4624.  References  230.  222  Morgan, B. P., Scholtz, J. M., Ballinger, M. D., Zipkin, I. D., and Bartlett, P. A. (1991) Differential binding energy: a detailed evaluation of the influence of hydrogen-bonding and hydrophobic groups on the inhibition of thermolysin by phosphorus-containing inhibitors. J. Am. Chem. Soc. 113, 297-307.  231.  Hanson, J. E., Kaplan, A. P., and Bartlett, P. A. (1989) Phosphonate analogues of carboxypeptidase A substrates are potent transition-state analogue inhibitors. Biochemistry. 28, 6294-6305.  232.  Bartlett, P. A., and Giangiordano, M . A. (1996) Transition state analogy of phosphonic acid peptide inhibitors of pepsin. J. Org. Chem. 61, 3433-3438.  233.  Mookhtiar, K. A., Grobelny, D., Galardy, R. E., and Wart, H. E. V. (1988) Ketone-substrate analogues of Clostridium histolyticum collagenases: tightbinding transition-state analogue inhibitors. Biochemistry. 27, 4299-4304.  234.  Ermert, P., Vasella, A., Weber, M., Rupitz, K , and Withers, S. G. (1993) Configurationally selective transition state analogue inhibitors of glycosidases. A study with nojiritetrazoles, a new class of glycosidase inhibitors. Carbohydr. Res. 250, 113-128.  235.  Whitworth, G. E., Macauley, M. S., Stubbs, K. A., Dennis, R. J., Taylor, E. J., Davies, G. J., Greig, I. R., and Vocadlo, D. J. (2007) Analysis of PUGNAc and NAG-thiazoline as transition state analogues for human O-GlcNAcase: mechanistic and structural insights into inhibitor selectivity and transition state poise. J. Am. Chem. Soc. 129, 635-644.  236.  Phillips, M. A., Kaplan, A. P., Rutter, W. J., and Bartlett, P. A. (1992) Transition-state characterization: a new approach combining inhibitor analogues and variation in enzyme structure. Biochemistry. 31, 959-963.  237.  Berland, C. R., Sigurskjold, B. W., Stoffer, B., Frandsen, T. P., and Svensson, B. (1995) Thermodynamics of inhibitor binding to mutant forms of glucoamylase from Aspergillus niger determined by isothermal titration calorimetry. Biochemistry. 34, 10153-10161.  References  238.  223  Mosi, R., Sham, H., Uitdehaag, J. C. M., Ruiterkamp, R., Dijkstra, B. W., and Withers, S. G. (1998) Reassessment of acarbose as a transition state analogue inhibitor of cyclodextrin glycosyltransferase. Biochemistry. 37, 17192-17198.  239.  Withers, S. G., Namchuk, M., and Mosi, R. (1999) Iminosugars as glycosidase inhibitors: nojirimycin and beyond, in Iminosugars as glycosidase inhibitors:  nojirimycin and beyond (Stiitz, A. E., Ed.) ed., pp  188-206, Wiley-VCH, Weinheim. 240.  Wolfenden;R, Snider, M., Ridgway, C., and Miller, B. (1999) The temperature dependence of enzyme rate enhancements. J. Am. Chem. Soc. 727,7419-7420.  241.  Dale, M . P., Ensley, H. E., Kern, K., Sastry, K. A. R , and Byers, L. D. (1985) Reversible inhibitors of p-glucosidase. Biochemistry. 24, 3530-3539.  242.  Hartman, F. C., LaMuraglia, G. M., Tomozawa, Y., and Wolfenden, R. (1975) The influence of pH on the interaction of inhibitors with triosephosphate isomerase and determination of the pK of the active-site a  carboxyl group. Biochemistry. 14, 5274-5279. 243.  Axamawaty, M . T., Fleet, G. W., Hannah, K. A., Namgoong, S. K., and Sinnott, M. L. (1990) Inhibition of the a-L-arabinofuranosidase III of Moniliniafructigena  by l,4-dideoxy-l,4-imino-L-threitol and 1,4-dideoxy-  1,4-imino-L-arabinitol. Biochem. J. 266, 245-249. 244.  Asano, N. (2003) Glycosidase inhibitors: update and perspectives on practical use. Glycobiology. 13, 93R-104R.  245.  Lee, J. K., Bain, A. D., and Berti, P. J. (2004) Probing the transition states of four glucoside hydrolyses with C kinetic isotope effects measured at natural 1 3  abundance by NMR spectroscopy. J. Am. Chem. Soc. 126, 3769-3776. 246.  Conchie, J., and Lewy, G. A. (1957) Inhibition of glycosidases by aldonolactones of corresponding configuration. Biochem. J. 57, 389-395.  247.  Ishida, N., Kumagai, K., Niida, T., Tsuruaka, T., and Yumato, H. (1967) J. Antibiot. 20, 66-71.  248.  Inouye, S., Tsuruoka, T., Ito, T., and Niida, T. (1968) Structure and synthesis of nojirimycin. Tetrahedron. 24, 2125-2144.  Kajimoto, T., Liu, K. K. C , Pederson; R. L., Zhong, Z., Ichikawa, Y., Porco, J. A., and Wong, C. H. (1991) Enzyme-catalyzed aldol condensation for asymmetric synthesis of azasugars: synthesis, evaluation, and modeling of glycosidase inhibitors. J. Amer. Chem. Soc. 113, 6187-6196. Jespersen, T. M., Dong, W., Sierks, M . R., Skrydstrup, T., Lundt, I., and Bols, M . (1994) Isofagomine, a potent, new glycosidase inhibitor. Angew. Chem., Int. Ed. Engl. 33, 1778-1779.  Jespersen, T. M., Bols, M., Sierks, M. R., and Skrydstrup, T. (1994) Synthesis of isofagomine, a novel glycosidase inhibitor. Tetrahedron. 50, 13449-13460. Biilow, A., Plesner, I. W., and Bols, M . (2000) A large difference in the thermodynamics of binding of isofagomine and 1-deoxynojirimycin to Pglucosidase. J. Am. Chem. Soc. 122, 8567-8568. Ganem, B., and Papandreou, G. (1991) Mimicking the glucosidase transition state: shape/charge considerations. J. Am. Chem. Soc. 113, 8984-8985. Papandreou, G., Tong, M . K., and Ganem, B. (1993) Amidine, amidrazone, and amidoxime derivatives of monosaccharide aldonolactams: synthesis and evaluation as glycosidase inhibitors. J. Am. Chem. Soc. 115, 11682-11690. Vonhoff, S., Heightman, T. D., and Vasella, A. (1998) Inhibition of glycosidases by lactam oximes: influence of the aglycon in disaccharide analogues. Helv. Chim. Acta. 81, 1710-1725. Hoos, R., Vasella, A., Rupitz, K., and Withers, S. G. (1997) DGlyconhydroximolactams strongly inhibit a-glycosidases. Carbohydr. Res. 298, 291-298. Hoos, R., Naughton, A. B., Thiel, W., Vasella, A., Weber, W., Rupitz, K., and Withers, S. G. (1993) D-Gluconhydroximo-1,5-lactam and related N arylcarbamates - theoretical calculations, structure, synthesis, and inhibitory effect on P-glucosidases. Helv. Chim. Acta. 76, 2666-2686. Tong, M . K., Papandreou, G., and Ganem, B. (1990) Potent, broad-spectrum inhibition of glycosidases by an amidine derivative of D-glucose. J. Am. Chem. Soc. 772,6137-6139.  References  259.  225  Ermert, P., and Vasella, A. (1991) Synthesis of a glucose-derived tetrazole as a new P-glucosidase inhibitor - a new synthesis of 1-deoxynojirimycin. Helv. Chim. Acta. 74,  260.  2043-2053.  Panday, N., and Vasella, A. (1999) Synthesis of glucose- and mannosederived N-acetylamino imidazopyridines and their evaluation as inhibitors of glycosidases. Synthesis. SP, 1459-1468.  261.  Field, R. A., Haines, A. H., Chrystal, E. J., and Luszniak, M . C. (1991) Histidines, histamines and imidazoles as glycosidase inhibitors. Biochem. J. 274, 885-889.  262.  Tatsuta, K., Miura, S., Ohta, S., and Gunji, H. (1995) Syntheses and glycosidase inhibiting activities of nagstatin analogs. J. Antibiot. (Tokyo). 48, 286-288.  263.  Tatsuta, K., Ikeda, Y., and Miura, S. (1996) Synthesis and glycosidase inhibitory activities of nagstatin triazole analogs. J. Antibiot. (Tokyo). 49, 836-838.  264.  Panday, N., Canac, Y., and Vasella, A. (2000) Very strong inhibition of glucosidases by C(2j-substituted tetrahydroimidazopyridines. Helv. Chim. Acta. 83,  265.  58-79.  Heightman, T. D., Locatelli, M., and Vasella, A. (1996) Synthesis of fused triazoles as probes for the active site of retaining P-glycosidases: From which direction is the glycoside protonated? Helv. Chim. Acta. 79, 2190-2200.  266.  Stiitz, A. E. (1999) Iminosugars as glycosidase inhibitors: nojirimycin and beyond,  in Iminosugars as glycosidase inhibitors: nojirimycin and beyond  (Stiitz, A. E., Ed.) Vol. Wiley-VCH, Weinheim. 267.  Legler, G., and Pohl, S. (1986) Synthesis of 5-amino-5-deoxy-Dgalactopyranose and l,5-dideoxy-l,5-imino-D-galactitol, and their inhibition of a- and P-D-galactosidases. Carbohydr. Res. 155, 119-129.  268.  Ichikawa, Y., and Igarashi, Y. (1995) An extremely potent inhibitor for Pgalactosidase. Tetrahedron Lett. 36,4585-4586.  Jensen, H. H., Lyngbye, L., and Bols, M . (2001) A free-energy relationship between the rate of acidic hydrolysis of glycosides and the pK of a  isofagomines. Angew. Chem. Int. Ed. 40, 3447-3449. Vocadlo, D. J., Wicki, J., Rupitz, K., and Withers, S. G. (2002) A case for reverse protonation: identification of Glul60 as an acid/base catalyst in Thermoanaerobacterium saccharolyticum  P-xylosidase and detailed kinetic  analysis of a site-directed mutant. Biochemistry. 41, 9736-9746. Varrot, A., Tarling, C. A., Macdonald, J. M., Stick, R. V., Zechel, D. L., Withers, S. G., and Davies, G. J. (2003) Direct observation of the protonation state of an imino sugar glycosidase inhibitor upon binding. J. Am. Chem. Soc. 125, 7496-7497. Varrot, A., Schiilein, M., Pipelier, M., Vasella, A., and Davies, G. J. (1999) Lateral protonation of a glycosidase inhibitor - structure of the Bacillus agaradhaerens  Cel5A in complex with a cellobiose-derived imidazole at 0.97  A resolution. J. Am.  Chem. Soc. 121,  2621-2622.  Gloster, T. M., Williams, S. J., Roberts, S., Tarling, C. A., Wicki, J., Withers, S. G., and Davies, G. J. (2004) Atomic resolution analyses of the binding of xylobiose-derived deoxynojirimycin and isofagomine to xylanase XynlOA. Chem. Commun. 16,  1794-1795.  O'Neill, G. P., Kilburn, D. G., Warren, R. A. J., and Robert C. Miller, J. (1986) Overproduction from a cellulase gene with a high guanosine-pluscytosine content in Escherichia coli. Appl. Environ. Microbiol. 52, 737-743.  APPENDIX A  GRAPHICAL REPRESENTATION OF KINETIC DATA  Appendix A  A  228  Graphical Representation of Data  GRAPHICAL REPRESENTATION OF DATA  A. 1 Steady State Kinetics for the Hydrolysis of Xylobiosides or Cellobiosides A. 1.1 Native Cex 0.025  250 200 % 150 £• 100 0.05  0.1  0.15  0.2  [2.5DNPX2] (mM)  50 Ij l i I i \ i_ 0 -100 -50 0 50 100 150 200 250 1/I2.5DNPX2] (mM" ) 1  Figure A . l : Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by wild-type Cex.  Figure A.2: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by wild-type Cex.  Figure A.3: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by wild-type Cex.  Figure A.4: Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by wild-type Cex.  2  2  2  2  Appendix A  Graphical Representation of Data  Figure A.5. Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by wild-type Cex. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the MichaelisMenten plot for the hydrolysis reaction. 2  229  Figure A.6. Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by wild-type Cex. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  Figure A.7: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by wild-type Cex.  Figure A.8: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by wild-type Cex.  Figure A.9: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by wild-type Cex.  Figure A.10: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by wild-type Cex.  230  Appendix A  231  Graphical Representation of Data  A.1.2 CexE43A  150  0  0.05 0.1 0.15 [2.5DNPX,] (mM)  Figure A . l l : Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex E43A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  0.2  -5  0 5 10 15 1/P.5DNPXJ (mM" ) 1  Figure A.12: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex E43A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  20  Appendix A  Graphical Representation of Data  Figure A.13: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by Cex E43A.  2  232  Figure A . H : Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by Cex E43A. 2  Appendix A  Graphical Representation of Data  250  1  1  1  233  1  1  1  o o •<*  _  E  /  -  5  1 1  Jh  1 1  / / / / / / f  . I I I !  2  4 6 [ONPXj] (mM)  -2  -1  0 1 2 3 4 5 6 7 1/[0NPX,] (mM" )  8  0 0.5 1 1.5 l/fONPXg] (mM" )  2  1  0.02  100 o  f  60 -  E,  40 -  ^° 0  0.2  0.4 0.6 0.8 [ONPXg] (mM)  Figure A. 15: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex E43A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  1  80 _  -  20 0 L -0.5  1  Figure A.16: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex E43A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  0.25  0  0.5  1  [2.4DNPC]  1.5  2  (mM)  Figure A.17: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex E43A.  Figure A.18: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex E43A.  0 . 0 3 |—i—|—i—|—i—|—i—|—i—|—i—|—r  800  [PNPC]  I—i—I—i—|—i—|—i—i—i—|—r  (mM)  Figure A. 19: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex E43A.  Figure A.20: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex E43A.  234  Appendix A  Graphical Representation of Data  235  A.1.3 C e x N 4 4 A 0.07  I  I  A  0.06  1  I  1  1  ^  175  1  o- ~  0.05  -  0.04  1  0.03  -  o" -  0.02  -cT / p/  0.01  ?/  -  -  150 o  < <  125 100  E  75  *°  50 25  0  0 0.2  0.4  0.6  0.8  -40  0 40 80 120 1/[2,5DNP)C,] (mM" )  [2,5DNP>y (mM)  160  1  0.04  < E,  5 0  0.05  0.1  0.15  0.2  0  [2.5DNPX,] (mM) Figure A.21: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex N44A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  5  10  15  1/[2,5DNPX2] ( m M ) -1  Figure A.22: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex N44A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  20  Appendix A  Graphical Representation of Data  0.4  o o  0.3  i ^  o o o  0.2  h  E  0.1  35  r  30  -  25  -  20  -  15  -  10  -  5  -  236  0 L 2  4  6  -0.5  0  0.5  o  1.5  12  o o  0.15 h  10  <  8  0.1 h  <  6  0.05 h  5  4  //-  IB AS  -  -  y? i  0 1  1.5  2.5 3  i ' i i /  1  2 0.5  2 1  0.2 "c c E  1  1/[PNPX,] (mM" )  [PNPXj] (mM)  2  2.5  [PNPXg] (mM)  -0.2  0  0.2 0.4  i  i  .  0.6 0.8  l / f P N P ^ ] (mM" ) 1  Figure A.23: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by Cex N44A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.24: Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by Cex N44A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  0  0.2 0.4 0.6 [ONP>y (mM)  0.8  -0.5  0 0.5 1 1.5 l/IONP)^] (mM" ) 1  Figure A.25: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex N44A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.26: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex N44A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  2  Appendix A  Graphical Representation of Data  Figure A.27: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex N44A.  Figure A.28: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex N44A.  Figure A.29: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex N44A.  Figure A.30: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex N44A.  238  Appendix A  Graphical Representation of Data  239  A. 1.4 C e x K 4 7 A  0.06  30  o E  5  25 20  Figure A.31: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex K47A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  0.4  _  ,  1  B  /  10 0  J * '  —  15 5  0.1 0.2 0.3 [2.5DNPX2] (mM)  •  I-  / /  7 _ >  -2  •  .  i  .  0 2 4 1/I2.5DNPX2] (mM' ) 1  Figure A.32: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex K47A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  240  Figure A.33: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by Cex K47A.  Figure A.34: Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by Cex K47A.  Figure A.35: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex K47A.  Figure A.36: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex K47A.  2  2  2  2  Appendix A  Graphical Representation of Data  241  40  o o  E  %  o  <  30 h 20 h  <  10  2  4 6 8 [2.4DNPC] (mM)  -0.5  0 0.5 1 1.5 1/[2,4DNPC] (mM' ) 1  Figure A.37: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex K47A.  Figure A.38: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex K47A.  Figure A.39: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex K47A.  Figure A.40: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex K47A.  Appendix A  Graphical Representation of Data  242  A.1.5 C e x H 8 0 A  0.025  60  o  i  40 20  0.01 0.02 0.03 0.04 0.05 [2.5DNPX,] (mM) Figure A.41: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex H80A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  0 -20  0 20 40 60 80 100 1/[2,5DNPX2] (mM" ) 1  Figure A.42: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex H80A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  Figure A.43: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by Cex H80A. 2  243  Figure A.44: Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by Cex H80A. 2  Appendix  A  Graphical  Representation  0.08  of  244  Data  250 ^o o  200 h 150 h  E -  1  2  3  100 h 50 h  4  -40 -20  50 40  o <* <  -  0.15  30  10 0  [ONP)^] (mM) Figure A.45: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex H80A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  i  1  —  i  '  B  /  i  /  -  20 0.1  80  1  [ONPXg] (mM)  0.05  0 20 40 60 1/[0NP)y (mM" )  p  /  /  ?  /  :  0 1 2 1/[0NP)C,] (mM" )  3  1  Figure A.46: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex H80A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  0.025  0  200  I  0  1  1  1——I  1  0.05 0.1 [2.4DNPC] (mM)  i—i—\—i—I—i—\—i—|—i—|—r  1 0.15  Figure A.47: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex H80A.  Figure A.48: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex H80A.  Figure A.49: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex H80A.  Figure A.50: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex H80A.  245  Appendix A  Graphical Representation of Data  246  A.1.6 CexH80N 0.02  I  I  I  A -  i  i  i  >  i  . —o  P  ; _  _  ?1 / 0.005  >  I y '  i  i  i  0.2 0.4 0.6 0.8 [2,5DNP)C,] (mM)  1  -1000  0 1000 2000 1/[2,5DNP>C,] (mM" )  120  o  <  < E  100 -  /  B  /  80 60 40 20 -  0.01 0.02 [2,5DNP^] (mM) Figure A.51: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex H80N. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  3000  1  0.03  /  0 L -10  /  /  0 5 10 1/[2,5DNP>y (mM )  -5  -1  Figure A.52: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex H80N. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  15  Appendix A  Graphical Representation of Data  247  0.06 I  o o  0.04  5  0.02  0.5 1 1.5 [PNPX^ (mM)  2  -20  0 20 40 l/tPNPXg] (mM" )  60  1  0.02  o o  <  <  0  0.05 0.1 0.15 [PNPX^ (mM)  Figure A.53: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by Cex H80N. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  0.2  0  5 10 15 l/IPNPXj] (mM" ) 1  Figure A.54: Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by Cex H80N. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  248  0.03 o o  <  < E  5 1  2  3  4  -100 -50  [ONPX,] (mM)  0  50  100  150  1/[0NP)y (mM" ) 1  0.008  200  0.05  0.1  0.15  [0NP)y (mM)  1/[0NP)y (mM" ) 1  Figure A.55: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex H80N. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.56: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex H80N. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  200  Appendix A  Graphical Representation of Data  0.02  249  300  o o  200  E  5 0.02 0.04 [2.4DNPC] (mM)  0.06  -i  I  1  I  1  I  1  I  4 6 8 [PNPC] (mM)  i  -250  0 250 500 1/[2,4DNPC] (mM" )  750  1  Figure A.57: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex H80N.  0.008  100  I  Figure A.58: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex H80N.  r  10 12  Figure A.59: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex H80N.  0  0.5 1 1.5 1/[PNPC] (mM" ) 1  Figure A.60: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex H80N.  2  Appendix A  Graphical Representation of Data  250  A.1.7 CexH80Q  Figure A.61: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex H80Q. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.62: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex H80Q. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  0.015  1  I  1  I  A |  -  cr  • /  < ~  i  ^ cr ~~~ -  0.01  o  i  i  o o  -  -  0.005  'f f,  251  ,  , ,  0.5  5  i  1  1.5  2  0  [PNP>y (mM)  20  40  60  l / I P N P J y (mM" ) 1  0.006  400  o o  300  I  -  i  i  B  200  /  i  i  i  z.  /  /  -  E.  i  r r ^ -  _  5  p /  0.05  0.1  0.15  0  [PNPX,] (mM)  5  10  15  1/[PNPX>] (mM" ) 1  Figure A.63: Michaelis-Menten plot for the hydrolysis of PNPX catalyzed by Cex H80Q. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.64: Lineweaver-Burk plot for the hydrolysis of PNPX catalyzed by Cex H80Q. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  20  Appendix A  Graphical Representation of Data  0.008  1  1  1  1  1  1  1500  1  A  o o  :  o o  0  /  x  -  V  ,  E  \  1  1000  c  -.0/  V  252  2 3 [ONPX,] (mM)  5 ?  4  500  -100  -50 0 50 100 150 l/IONPJy (mM' ) 1  0.002  1000  o E  5  800 -  / /  600 -  /  400 200 0  0.05 0.1 0.15 [ONPXj] (mM)  B  o  r^-  /  O  \  i  7  /  cP —i  0  \  \  I  \  \  i _  2 4 6 8 1/[0NPX2] (mM" ) 1  Figure A.65: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex H80Q. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.66: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex H80Q. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  10  Appendix A  Graphical Representation of Data  0.003 I  o o  <  253  2000  o o 1500  0.002 \-  <  < = 1000 E  0.001  500 h  0.01 0.02 0.03 [2.4DNPC] (mM)  0.04  -1000  0 1000 1/[2,4DNPC] (mM" )  2000  1  Figure A.67: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex H80Q.  Figure A.68: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex H80Q.  0.001  8000  _c 0.0008 •% o 0.0006 % 0.0004 0.0002 0  0  2  4 6 [PNPC] (mM)  8  Figure A.69: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex H80Q.  10  -0.5  0  0.5 1 1.5 1/[PNPC] (mM" ) 1  Figure A.70: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex H80Q.  2  Appendix A  Graphical Representation of Data  254  A.1.8 C e x Q 8 7 M 0.04 -i  1  0  r  0.1 0.2 0.3 [2.5DNPX,] (mM)  Figure A.71: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex Q87M. 2  0.4  -50  0 50 100 1/[2,5DNPX,] (mM" ) 1  Figure A.72: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex Q87M. 2  150  Figure A.73: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex Q87M. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.74: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex Q87M. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  Figure A.75: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex Q87M.  Figure A.76: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex Q87M.  Figure A.77: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex Q87M.  Figure A.78: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex Q87M.  256  Appendix A  Graphical Representation of Data  257  A.1.9 C e x Q 8 7 Y  Figure A.79: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex Q87Y. 2  Figure A.80: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex Q87Y. 2  Appendix A  c  E  o  <  <  5  Graphical Representation of Data  0.14 - 1 — i — i — r 0.12 A 0.1 h 0.08 0.06 h 0.04 0.02 g -i  i  2  i —  1  —  1  —  1  —  1  258  200  — J  JO  o < c  150 h 100 h  E 50 h i  i  3  i  i  4  i  i  i_  5  -10  0 10 20 30 40 1/[0NP>y (mM" ) 1  [ O N P X J (mM)  0.025  o o  5 0.1 0.2 [ONPXj] (mM)  0  2 4 6 1/[0NP)y (mM ) -1  Figure A.81: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex Q87Y. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.82: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex Q87Y. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  Figure A.83: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex Q87Y.  0  2  4 6 8 [PNPC] (mM)  Figure A.85: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex Q87Y.  259  Figure A.84: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex Q87Y.  10  -1  0 1 2 1/[PNPC] (mM' ) 1  Figure A.86: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex Q87Y.  3  Appendix A  Graphical Representation of Data  260  A . 1 . 1 0 C e x N126A 0.008  I  1  I  I  A  1  1  1  1  600  1  _o  o  f,  5  V I  1  I  ,  |  0.004  ~  0.002  II  i  i  /  a  / ~  1 -  200 h s  0.006  i  /  400  1  0.1 0.2 0.3 0.4 0.5 [2.5DNPX2] (mM)  1  .A  E  )i  1  i  -500  1  1  -250 0 250 1/[2,5DNP)y (mM" )  500  0 50 100 1/[2,5DNPX2] (mM" )  150  1  300  0.02 0.04 0.06 [2.5DNPX,] (mM) Figure A.87: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex N126A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  0.08  1  Figure A.88: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex N126A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  0.2 c  1  0.15 h  E  i  o  ^  A  1  1  i  •  I  120  i  o o  100  <  261  1  •  :A _  E  /  0.05 .  I  .  i  *  / y -  /  /  /  /  ir  I  /  //  -  i/ i  i  -  _  0.1 h  0  i  -  i /'  i  0.5 1 1.5 [ONPX,] (mM)  2  -10  0 10 20 30 l/tONPXJ (mM" )  40  1  30 g  o  E  1  i  B  i  .  y  /  20  o  i  /  -  £  10  ' O  y •  0.1 0.2 [ONPX^ (mM)  0.3  •  i  _  i  i  i  i  0 2 4 l/fONPX^ (mM" ) 1  Figure A.89: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex N126A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.90: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex N126A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  0.12  100  262  i  /  80 -  / . -  /  60 40  o  /  20 0  0.5  1  [2.4DNPC] (mM) Figure A.91: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex N126A.  1.5  0 -50  0 50 100 150 200 1/[2,4DNPC] (mM ) -1  Figure A.92: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex N126A.  Appendix A  Graphical Representation of Data  263  A.1.11Cex E127A 0.008  -i  1  1  1  1  1  0.1 0.2 0.3 [PNPC] (mM) Figure A.93: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex E127A.  1200  r  0.4  0  100 200 300 400 1/[PNPC] (mM" ) 1  Figure A.94: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex E127A.  /  Appendix A  Graphical Representation of Data  264  A . 1 . 1 2 C e x N169A  0.008  200  o  <  •< E  • j  —  B  150  Figure A.95: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex N169A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  0.08  i  i  .  i  i  8  —  i  •  0.02 0.04 0.06 [2.5DNPX,] (mM)  i  1  •  0  i  "—/  100 50  i  /  -20  0 20 40 1/I2.5DNPX2] (mM" ) 1  Figure A.96: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex N169A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  60  Graphical Representation of Data  Appendix A  265  600  0  0.2 0.4 0.6 0.8 [ONPXg] (mM)  1 1.2  -25 0 25 50 75 100 125 150 l/fONPXg] (mM" ) 1  0.012  120  o o  5 0.05 0.1 [ONPX^ (mM)  0.15 1/[0NP)y (mM ) -1  Figure A.97: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex N169A. Plot A shows all data points with a solid line drawn through the points that represent the hydrolysis reaction and a dashed line through those that represent the transglycosylation reaction. Plot B shows the Michaelis-Menten plot for the hydrolysis reaction. 2  Figure A.98: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex N169A. The solid line represents the hydrolysis reaction and the dashed line represents the transglycosylation reactions. Plot A shows all data points. Plot B magnifies data points obtained for the transglycosylation reaction. 2  Appendix A  Graphical Representation of Data  0.08  150  0  0.5 1 [2.4DNPC] (mM)  Figure A.99: Michaelis-Menten plot for the hydrolysis of 2,4DNPC catalyzed by Cex N169A.  1.5  Figure A.100: Lineweaver-Burk plot for the hydrolysis of 2,4DNPC catalyzed by Cex N169A.  266  Appendix A  Graphical Representation of Data  267  A.1.13Cex H205N  Figure A.101: Michaelis-Menten plot for the hydrolysis of 2,5DNPX catalyzed by Cex H205N.  Figure A.102: Lineweaver-Burk plot for the hydrolysis of 2,5DNPX catalyzed by Cex H205N.  2  2  0.0006  12000  0.0005I  10000 o o  0.0004 P 1 0.00031-  %  8000 -  <  6000 -  E  5  ~ 0.0002  4000 2000 -  0.0001  a  <  2  4  6  0L  8  10  [ONPX2] (mM)  0  2  4  1/[ONP>y (mM" ) 1  Figure A.103: Michaelis-Menten plot for the hydrolysis of ONPX catalyzed by Cex H205N.  2  Figure A.104: Lineweaver-Burk plot for the hydrolysis of ONPX catalyzed by Cex H205N. 2  Appendix A  Graphical Representation of Data  Figure A.105: Michaelis-Menten plot for the hydrolysis of PNPC catalyzed by Cex H205N.  Figure A.106: Lineweaver-Burk plot for the hydrolysis of PNPC catalyzed by Cex H205N.  268  Appendix A  Graphical Representation of Data  A.2 Steady State Kinetics for Inhibition by Imino Sugar Inhibitors A.2.1 C e x E 4 3 A  Figure A.108: Inhibition of Cex E43A by Xylobiosyl Imidazole.  269  Appendix A  Graphical Representation of Data  270  40  /  2000 4000 [Xylobiosyl DNJ] ( M) U  6000  1  1  \  I 20 5 10  1  %  I  o ou  1  DO  1  -  0 -5000  w.  / ,  i  ,  0 5000 10000 [Xylobiosyl DNJ] ( M)  Figure A.109: Inhibition of Cex E43A by Xylobiosyl Deoxynojirimycin.  Figure A . l 11: Inhibition of Cex E43A by Xylobiosyl Isofagomine Lactam.  U  max  Appendix A  Graphical Representation of Data  A.2.2 C e x N 4 4 A  [Xylobiosyl Imidazole] (uM)  [Xylobiosyl Imidazole] (uM)  Figure A.113: Inhibition of Cex N44A by Xylobiosyl Imidazole.  Figure A.114: Inhibition of Cex N44A by Xylobiosyl Deoxynojirimycin.  271  Appendix A  Graphical Representation of Data  E-272  40  ^ ^  o o 30 -  < E  5 0  10  20  30  40  [Xylobiosyl Isofagomine] (^M)  50  20 10 0 -10  0  10  20  30  40  [Xylobiosyl Isofagomine] (|Jv1)  Figure A.115: Inhibition of Cex N44A by Xylobiosyl Isofagomine.  Figure A. 116: Inhibition of Cex N44A by Xylobiosyl Isofagomine Lactam.  50  Appendix A  Graphical Representation of Data  A.2.3 C e x K 4 7 A  Figure A . l 19: Inhibition of Cex K47A by Xylobiosyl Deoxynojirimycin.  273  Appendix A  Graphical Representation of Data  8  DO .  1  1  1  1  6 4  V  E,  1  s  1 I  o  I  1  E  1  I  10 o o  c  1I 1  12  E-274  2  w,  max  1 , 1 0 -2000 0 2000 4000 6000 [Xylobiosyl Isofagomine] (^M) I  0  2000 4000 6000 [Xylobiosyl Isofagomine] (fiM)  Figure A . 120: Inhibition of Cex K47A by Xylobiosyl Isofagomine.  I  I  i  I  i  i  B o o <  o o  <  I  5  4  2  1000 2000 [Xylobiosyl IL] ( M) U  3000  1  IW.  , i • i i 0 -3000 -1500 0 1500 3000 [Xylobiosyl IL] ( u M )  Figure A.121: Inhibition of Cex K 4 7 A by Xylobiosyl Isofagomine Lactam.  Appendix A  Graphical Representation of Data  A.2.4 C e x H 8 0 A  Figure A.124: Inhibition of Cex H80A by Xylobiosyl Deoxynojirimycin.  275  Appendix A  Graphical Representation of Data  [Xylobiosyl Isofagomine] (p.M)  E-276  [Xylobiosyl Isofagomine] ( M)  Figure A.125: Inhibition of Cex H80A by Xylobiosyl Isofagomine.  Figure A.126: Inhibition of Cex H80A by Xylobiosyl Isofagomine Lactam.  U  Appendix A  Graphical Representation of Data  277  _ 100  1  1  1  1  1  1  1  I  1  I  1  o  1  1  120  <° 80  1  <  1  0.04  11  A.2.5 CexH80N  1  1  .1 60 o 40 h  0  100 200 300 400 [Xylobiosyl Imidazole] (uM)  q  1  20  0 -100  w.  max  0 100 200 300 [Xylobiosyl Imidazole] (uM)  Figure A.128: Inhibition of Cex H80N by Xylobiosyl Imidazole.  0.03  800  5000 10000 [Xylobiosyl DNJ] ( M) U  15000  -5000  0 5000 10000 15000 [Xylobiosyl DNJ] ( M)  Figure A.129: Inhibition of Cex H80N by Xylobiosyl Deoxynojirimycin.  U  max  Appendix A  Graphical Representation of Data  0.03 c E o  £  E-278  500  0.02  0.01 h  0  100  200  300  -100  [Xylobiosyl Isofagomine] (JIM)  0  100  200  300  [Xylobiosyl Isofagomine] (JIM)  Figure A.130: Inhibition of Cex H80N by Xylobiosyl Isofagomine.  0.04  800  MV. 2500  5000  7500  [Xylobiosyl IL] (^M)  10000  max  -5000  0  5000  10000  [Xylobiosyl IL] ( M ) U  Figure A.131: Inhibition of Cex H80N by Xylobiosyl Isofagomine Lactam.  15000  Appendix A  A.2.6  Graphical Representation of Data  CexH80Q  0.01  250  % 0.004 h  1  \  50  0.002 h  ,  00  1  :  1  0.006 h o  I  ^200 o <? < 150 c E 100  1  1  0.008Jb-  1  1  1 I  ^  279  1000  2000  3000  [Xylobiosyl Lactam Oxime] (jxM)  1  0 -2000-1000  ,  0  1000 2000 3000  [Xylobiosyl Lactam Oxime] ( ^ M )  Figure A.132: Inhibition of Cex H80Q by Xylobiosyl Lactam Oxime.  Figure A.134: Inhibition of Cex H80Q by Xylobiosyl Deoxynojirimycin.  Appendix  A  Graphical  Representation  0.006  E-280  Data  1200 _1000  |  of  i  i  i  i  i  i  i  i  •  i  i  i  i  \  B  o  0.004  <° 800  o  <  <:  0.002 h  0  20 40 60 80 100 120 [Xylobiosyl lsofagomine| (p.M)  |  600  J  400  MV.max J  i_  ^ 200 -20 0 20 40 60 80 100 120 0 [Xylobiosyl Isofagomine] (JIM)  Figure A.135: Inhibition of Cex H80Q by Xylobiosyl Isofagomine.  Figure A . 136: Inhibition of Cex H80Q by Xylobiosyl Isofagomine Lactam.  Appendix A  Graphical Representation of Data  281  A.2.7 CexQ87M  Figure A. 138: Inhibition of Cex Q87M by Xylobiosyl Imidazole.  [Xylobiosyl DNJ] ( M ) U  [Xylobiosyl DNJ] ( M)  Figure A.139: Inhibition of Cex Q87M by Xylobiosyl Deoxynojirimycin.  U  Appendix A  Graphical Representation of Data  0.08  E-282  100 o o  80 60 _  E  40 -  20 H w .m a x  0^ 5  10  15  20  [Xylobiosyl IL] ( M) U  25  0  5  10  15  20  [Xylobiosyl IL] ( M) M  Figure A.141: Inhibition of Cex Q87M by Xylobiosyl Isofagomine Lactam.  25  Appendix A  Graphical Representation of Data  A.2.8 C e x Q 8 7 Y  Figure A.144: Inhibition of Cex Q87Y by Xylobiosyl Deoxynojirimycin.  283  Appendix A  Graphical Representation of Data  [Xylobiosyl Isofagomine] ( M) U  E-284  [Xylobiosyl Isofagomine] (jJVI)  Figure A.145: Inhibition of Cex Q87Y by Xylobiosyl Isofagomine.  [Xylobiosyl IL] ( M) U  [Xylobiosyl IL] ( M) U  Figure A.146: Inhibition of Cex Q87Y by Xylobiosyl Isofagomine Lactam.  Appendix A  Graphical Representation of Data  285  A.2.9 Cex N126A  0.08  40  i  5  0.02  i  i  B  § 30  I  i  1  20 10k^  1  1W.  max  -  0  2500 .5000 7500 10000 [Xylobiosyl DNJ] ( M ) U  0 -5000  i  0 5000 10000 [Xylobiosyl DNJ] ( M)  Figure A.149: Inhibition of Cex N126A by Xylobiosyl Deoxynojirimycin.  U  Appendix A  Graphical Representation of Data  0.08  E-286  200  MV.  max  0  10 20 30 40 [Xylobiosyl Isofagomine] ^M)  -10  0 10 20 30 40 [Xylobiosyl Isofagomine] (|iM)  Figure A.150: Inhibition of Cex N126A by Xylobiosyl Isofagomine.  Figure A.151: Inhibition of Cex N126A by Xylobiosyl Isofagomine Lactam.  Appendix A  Graphical Representation of Data  287  A.2.1 OCex E127A  0.005  500  I  I  I  I  I  i  i  i  i  i  -2,400  o  <\ 300  5  0.001 h 0 0  J  I  I  1_  J  l_  50 100 150 200 [Xylobiosyl Imidazole] (uM)  200  w „  100 0 -150 -100 -50 0 50 100 150 [Xylobiosyl Imidazole] (uM)  Figure A.153: Inhibition of Cex E127A by Xylobiosyl Imidazole.  Figure A.154: Inhibition of Cex E127A by Xylobiosyl Deoxynojirimycin.  Appendix A  Graphical Representation of Data  0.005  E-288  1200  0.004 J-o o 0.003 h S 0.002 h 0.001 h  MV. max  0 50 100 150 200 250 300 [Xylobiosyl Isofagomine] (^M)  -50 0 50 100 150 200 250 300 [Xylobiosyl Isofagomine] {^M)  Figure A.155: Inhibition of Cex E127A by Xylobiosyl Isofagomine.  0.005  2000  0.004<  MV„  100 200 300 [Xylobiosyl IL] ( M) U  400  -100  0 100 200 300 [Xylobiosyl IL] (uM)  Figure A.156: Inhibition of Cex E127A by Xylobiosyl Isofagomine Lactam.  400  Appendix A  Graphical Representation of Data  Figure A.157: Inhibition of Cex N169A by Xylobiosyl Lactam Oxime.  289  Appendix A  Graphical Representation of Data  Figure A.158: Inhibition of Cex N169A by Xylobiosyl Imidazole at varying concentrations of 2,4DNPC. Plot A is the Michaelis-Menten plot, plot B is the Lineweaver-Burk plot, and plot C is the Eadie-Hofstee plot. The concentrations of xylobiosyl imidazole were 0 (O), 0.123 (•), 0.370 (TJ), 0.925 P), and 2.47 uM (A).  290  Figure A.161: Inhibition of Cex N169A by Xylobiosyl Isofagomine Lactam.  Appendix A  Graphical Representation of Data  292  A.3 pH Dependencies of k /K for the Hydrolysis of Xylobiosides or Cellobiosides by Native and Mutant Cex ca{  m  0.06 c E  o  <i  5 8  Figure A.162: pH dependence of for the hydrolysis of 2,4DNPC catalyzed by wild-type Cex. knJK  m  100  10  20  40 60 Time (min)  80  Figure A.163: pH stability of wild-type Cex at pH 3.0 (O), 3.5 (•), and 10.0 (TJ).  0.08  CO  o  0  Figure A.164: pH dependence of for the hydrolysis of 2,4DNPC catalyzed by Cex E43A.  k JK c  m  20  40 60 Time (min)  Figure A.165: pH stability of Cex E43A at pH 3.0 (O), 3.5 (•), and 10.0 (TJ).  80  Appendix A  Graphical Representation of Data  293  0.08  (0  20  Figure A.166: pH dependence of k mlK for the hydrolysis of 2,4DNPC catalyzed by Cex N44A. K  40 60 Time (min)  Figure A.167: pH stability of Cex N44A at pH 3.0 (O), 3.5 (•), and 10.0 (TJ).  m  3000 c E  2000 h  o  %  ^ 1000 V  6 PH Figure A. 168: pH dependence of kcat'Km for the hydrolysis of 2,5DNPX catalyzed by Cex K47A. 2  10  20  40 60 Time (min)  Figure A. 169: pH stability of Cex k47A at pH 3.0 (O), 3.5 (•), and 10.0 (•).  80  Appendix A  Graphical Representation of Data  294  Figure A.170: pH dependence of k /K for the hydrolysis of PNPX catalyzed by Cex H80A. cat  2  m  4  2  6  8  10  PH Figure A.171: pH dependence of k JK Cex H80N. c  m  Figure A.172: pH dependence of k JK Cex H80Q. c  m  for the hydrolysis of PNPX catalyzed by 2  for the hydrolysis of PNPX catalyzed by 2  Appendix A  Graphical Representation of Data  295  0.06  o o  £  20  Figure A.173: pH dependence of kiJK for the hydrolysis of 2,4DNPC catalyzed by Cex Q87M.  Figure A.174: pH stability of Cex Q87M at pH 3.0 (O), 3.5 (•), and 10.0 (•).  80  0.08  m  60 •S  40 60 Time (min)  c E  r-  o  40 \-  %  20 h  0.04 h 0.02 h  6 PH Figure A.175: pH dependence of for the hydrolysis of 2,4DNPC catalyzed by Cex Q87Y. *cat/^m  0.06 h  10  20  40 60 Time (min)  Figure A.176: pH stability of Cex Q87Y at pH 3.0 (O), 3.5 (•), and 10.0 (•).  Appendix A  Graphical Representation of Data  296  0.08  2  4  6 PH  8  10  0  cM  m  for  2,4DNPC N126A.  the  hydrolysis  catalyzed  by  40 60 Time (min)  80  Figure A. 178: pH stability of Cex N126A at pH 3.0 (O), 3.5 (•), and 10.0 (•).  Figure A.177: pH dependence of k IK  20  of  Cex  0.08  80 60 h  c  0.06 h  E  o  40 h  £  20 h  0.04 h 0.02 h  10 PH Figure A. 179: pH dependence of kcat/Km for the hydrolysis of 2,4DNPC catalyzed by Cex N169A.  0  20  40 60 Time (min)  Figure A.180: pH stability of Cex N169A at pH 3.0 (O), 3.5 (•), and 10.0 (TJ).  80  APPENDIX B  COMPILATION OF C E X DATA  Appendix B  B  Compilation of Cex Data  298  Compilation of Cex Data  B.1 Tables Table B.1: Dependence upon pH of kinetic parameters by C.fimixylanase and mutants. Conditions  Kinetic Parameter  Enzyme  al  3.9 teat  Wild-Type  k JK k JK t-cat' ^-m kca\l Km kcatlK kcat/Km kcat/K kcat/K kcat/Km kcat/Km kcat/Km kcat/K kcat/Km kcat/Km kcatlK kcat kcatlKm kcat _ kcdi[K a  b  Glu43Ala Asn44Ala Lys47Ala His80Ala HisSOAsn His80Gln Gln87Met Gln87Tyr Asnl26Ala Asnl69Ala Mis205Ala* His205Asn* Glul27Ala  c  Glu233Asp  c  m  m  m  m  4.1 5.0 3.7 3.9 3.9 J5 4.6 4.6 4.0 3.9 4.4 4.0  4  m  5.9  Data taken from Tull and Withers (i). * Data taken from Notenboom et al. (2). Data taken from MacLeod et al. (J).  c  Substrate  7.7 '  2,4DNPC 2,4DNPC 2,4DNPC PNPC 2,4DNPC  ~  7.7 7.3 6.9 8.0 6.0 6.2 " 6.3 7.0 " 8.1 8.3 8.5 8.2 6.2 5.0 "  m  a  PAa2  ~  .  2  4  D  N  p  "  c  2,5DNPX PNPX PNPX PNPX 2,4DNPC 2,4DNPC 2,4DNPC " 2,4DNPC PNPC PNPC 2,4DNPC PNPC 2,4DNPC 2,4DNPC 2  2 2 2  pH Range 3.5-8.5 4.5-9.5 4.5-9.5 4.0-8.5 3.5-8.5 3.5-8.5 3.5-8.0 3.5-7.5 3.5-7.5 4.0-7.5 3.5-8.5 3.5-8.5  4.6-8.5 3.5-8.5 4.0-8.5 4.0-8.5 4.5-9.0 5.5-9.5 4.5-8.0 4.5-8.0  Appendix B  Compilation of Cex Data  299  Table B.2: Brensted relationships for the hydrolysis of various glycosides by C./imi xylanase and mutants.  Enzyme  Substrate  Aryl P-D-Glucosides  Parameter  Pie  -l  ^cat  -\  K  m  a  a  (f  ^cat ^cat  -0.3" -0.3 -0.3 -0.3° -0.23* -0.29* -0.8 -0.9 0 a  Aryl p-Cellobiosides  K  fl  k /K 2  Wild-Type  s  W^m(pH 7.0). W^m(pH5.5) Aryl p-D-Xylosides  C  ^cat  C  kcat/K  m  Glu233Asp  Aryl P-Xylobiosides  kcat  Aryl p-Cellobiosides  kcat  C  kcallKyn  ~ f> - "•  kcaJK  -0.3  kcat  o '""  d  m  j  dr  -l.0 -0.8* d  kcatlK  m  K \ Glul27Ala  Aryl p-Cellobiosides  k /K  -0J  *ca.(pH 7.0)  0* -0.64* 0* -0.69* 0* -0.43* 0* -0.46*  2  d  s  k /K (pH caX  7.0)  m  £ (pH5.5) cat  His205Ala  Aryl p-Cellobiosides  k J K  m  (pH  5.5)  * (pH 7.0) cat  ! His205Asn >;r;,-;,Aryl P-Cellobiosides i  •%  hatlK  m  (pH  £ (pH5.5) W^m(pH5.5) cat  Data taken from Tull and Withers (1). * Data taken from Notenboom et al. (2). Data taken from Tull, D. (1995) Ph.D. Thesis, UBC. ^Data taken from MacLeod et al. (J). a  c  7.0)  P^a Range  4.0-8.4 4.0-8.4 4.0-8.0 8.0-9.3 4.0-9.3 4.0-7.2 4.0-7.2 4.0-7.2 4.0-7.2 4.9-8.5 4.9-8.5 5.2-7.2 5.2-7.2 4.0-9.3 4.0-9.3 4.0-7.5 4.0-9.3 4.0-7.2 4.0-7.2 4.0-7.2' 4.0-7.2 4.0-7.2 4.0-7.2 4.0-7.2 4.0-7.2 4.0-7.2 4.0-7.2  Appendix B  Compilation of Cex Data  300  Table B.3: Inactivation parameters for C.fimi xylanase and mutants.  pBHHHHB  A,  1 n/\mv'v  :•  Inacthator  V 2F-DNPG" 2F-DNPC* 2F-DNPX  Wild-Type  2  Glul27Ala His205Asn  2F-DNPC 2F-DNPC* rf  "IIMIHIIWIV  •'(mil) ) . \ ^ m M a ^ m i r , I'sjam  1  2.5 x 10~ 6.7 x 10" 5.7 x 10  4.5 0.11 0.0035  5.56 x 10" 5 6.12 x 10" 1 16  1.3 x 10~ 8.5 x 10" nd^  nd 1.3 x 10"  nd nd  1.6 nd  nd nd  4  2  C  A./A,  A,  -2  7  3  7 7  7  5  6  7 7  Data taken from Tull et al. (4). Data taken from Tull and Withers (7). Data taken from Ziser et al. (5). Data taken from MacLeod et al. (6). Data taken from Notenboom et al. (2). ^nd indicates that this kinetic parameter was not determined. a  b  c  rf e  Table B.4: Secondary deuterium kinetic isotope effects measured with C. fimi xylanase and mutants. •Enzyme"' ;•,\-  i  j Wild-Type  6  Glu233Asp 1 Gliil27Ala His205Ala c  rf  i His205Asn  rf  c  - v s - - ki)si_-  Substrate-- 2,4DNPG PNPG 2,4DNPC PNPC 4BrPC 2,4DNPC 2,4DNPC 2,4DNPC 2,4DNPC  "  :  degly gly degly degly gly degly degly degly degly  ° RDS, rate-determining step. Data taken from Tull and Withers (7). Data taken from MacLeod et al. (J). Data taken from Birsan, C. (1996) M.Sc. Thesis, UBC. 6  c  rf  • -.->-?'  1.12 1.12 1.10 1.10 1.06 1.09 1.08 1.12 1.02 - — "'" -  i i "  ""71  J  Appendix B  Compilation of Cex Data  301_  Table B.5: Michaelis-Menten parameters for the hydrolysis of 2,4-dinitrophenyl P-cellobioside by C.fimi xylanase and mutants.  --7-  K (mM)  ^cat  Enzyme -  ;  Wild-Type •  Glu43Ala Asn44 Ala . Lys47Ala His80Ala His80Asn His80Gln Gln87Met Gln87Tyr Asp 123 Ala Asnl26Ala Glul27Ala Glul27Asp Glul27Gly Asn 169 Ala His205Ala" His205Asn Glu233Asp  m  8.5 12.9" 7.0* 3.9 4.9 9.4 4.2 0.088 0.056 0.012 8.8 7.0 5.4 0.49 0.040* 0.027* 0.035* 2.9 0.00082 0.00049 0.003\ C  C  c  c  d  " Data taken from Tull and Withers (1). * Data taken from MacLeod et al. (6). Data taken from Notenboom et al. (2) Data taken from MacLeod et al. (3). 0  rf  0.068 0.11* 0.06* 0.12 Q  1  0  Q  7  m  1  c  9  0.070 2.6 0  ta  120 117° 116* 32 27  c  ....  k \IK (s^mM" )  1  0.0051 0.0019 0.056 0.11 0.53 0.047 0.0003* 0.0017* " 0.0004* 0.036 ~ 0.007 0.03 0.086 c  c  c  d  1.6 12 11 6.1 160 66 10 10 130* !6* 92* 81 0.12 0.016 0.036 c  C  c  rf  Appendix B  Compilation of Cex Data  302  Table B.6: Michaelis-Menten parameters for the hydrolysis of 4-nitrophenyl P-cellobioside by C.fimi xylanase and mutants. Km  Enzyme  Wild-Type  Gln87Met Gln87Tyr Asp 123 Ala Asnl26Ala Glul27Ala Glul27Asp Glul27Gly " Asnl69Ala His205Ala His205Asn Glu233Asp  " " i o ' ~ ~ ~ 0.020 0.048 0.040 0.0064 9.6 " " 9.2 7.4 0.007< 0.052 0.033 0.038* 0.033^ 0 . 0 5 5 * " 0.033* 3.9 0.0009 l 0.00074 0.00046 e  /  c  c  6.003V  " Data taken from Tull and Withers (i). * Data taken from MacLeod et al. (6). Data taken from Notenboom et al. (2) ^Data taken from MacLeod et al. (3). Data taken from Notenboom et al. (7) 0  e  0.58 0.60* 0.53* 0.56 13 1.7 ~ 29 13 6.6 3.1 0.35 0.34 2.6 lS 11.8 0.020 0.025* 0.020 ~ 7 1.7* 0.029* 0.59 0.82 1.05 0.67 0.66 c  C  Glu43Ala Asn44Ala Lys47Ala His80Ala His80Asn His80Gln  ca  (mM) 9.2 15.8° 11.3* 3.5 1.3  A t//r (s^mM )  e  d  rf  c  c  d  m  -1  16 26" 21* 6.2 0.10 5.8 0.00071 0.0037 0.0060 C  0  0  0  2  ]  27 27 ~ ~ 2 . 8 0.0004"' 0.0044 1.6 1.5* U e  d  0.033* 1.2* 6.59 0.001 l 0.00070 0.00069 0.0058^ c  c  Appendix B  Compilation of Cex Data  303  Table B.7: Michaelis-Menten parameters for the hydrolysis of 2,5-dinitrophenyl P-xylobioside by C.fimi xylanase and mutants.  Glu43Ala Lys47Ala His80Asn •BOGlnGln87Met  S87T>r  Asp 123 Ala  [f4Hl»Ala Glul27Ala Glul27Gly  IW§Ala ' His205Ala  H205Asn  Glu233Asp  30" 20  -v-  i?pi^Riiiiiiissiis ~ 59 • 0.25 • 0.12 0.034 47  isisiisiiisipiiiiiiii nd"  1:3- • 0.11  IllllBillllllilllliiiiiiHI nd  a  liS^ittlfflill|ii| nd°  ()'.0()059  :  o;0i>5, • 0.063 0.0 0.20 0*0021 ' •: 0.0012 IM # 6 2 . . 0.047 0.052 :  nd"  nd"  " " .520 >210  <0.0005 nd"  nd" nd*  nd"  • 010078 nd"  . ,0.013 nd"  * nd indicates that this kinetic parameter was not determined.  2100 310" 1 lOd 300 '120 99 54 1000 750  •  2600 nd"  0.044  Appendix B  Compilation of Cex Data  304  Table B.8: Michaelis-Menten parameters for the hydrolysis of 4-nitrophenyl P-xylobioside by C.Jimi xylanase and mutants.  \Mild-1 \ pe Glu43Ala Asn4;4&la Lys47Ala His80Asn •  •Hjs80Glh'" V Gln87Met Gln87Tyr Asp 123 Ala Glul27Ala Glul27Asp. Glul27Gly Asnl^^l'a. His205Ala ' Mis2p5'A5sn Glu233Asp  :  27 89 0.36 • 0:42 0.13 * -0i37 58* rid nd c  c  nd nd nd nd' 0.01\  c  ,  c  c  .'  c  c  c  d  rf  Data taken from Tull and Withers (7). * Data taken from Notenboom et al. (7). nd indicates that this kinetic parameter was not determined. Data taken from MacLeod et al. (5).  a  0  rf  480* nd nd 0.45 nd "" nd nd "nd nd nd 0.61 c  c  c  c  .  6.61  6  c  -  1500 22001"*": ~ 5.1 " 25 . 0.038 " 0.72 1.3  c  c  nd  0.022 0.018'5.4 3.6 9.6 0.59 0.099 .0.061 ' 0.12* nd nd ' 7.7 nd nd "' ~ nd "nd nd nd 0.018  c  '  c  c  c  c  c  rf  Appendix B  Compilation of Cex Data  305  Table B.9: Michaelis-Menten parameters for the hydrolysis of 2-nitrophenyl P-xylobioside by C.fimi xylanase and mutants.  : .,, 38 0.068 560 ; Wild-Type ^ o.06 ^ 720" Glu43Ala 47 _ JL6 _ 30^ A s n 4 4 A l a " " 8 0 ~ " ~ " " " " 0.64\_'~~IZ~130" Lys47Ala 9.7 ~ " J 8.5 ~_~ _ " 1.2 J His80Ala 0.33 0.039 8.4 His80Asn 0.14 _ ~0.016 ~ ~ ~ 8 . 6 His80Gln ""_ 0.039 " T£^L~~3.8 Gln87Met """55 '_' " 0.13 ~ ~ 4 2 0 Gln87Tyr'" 46 "™ 0.22 210 """ Asp 123 Ala _ jid* _ _ nd* _"_ nd* Asnl26Ala ~ ~~~~ 2.9 ZZ 6-39"ZLTZZ?-" Glul27Ala 0.080 ~ ~" <0.001 >80 Glul27Asp " nd* M" " I nd* ~~ " " ~ nd* ZZ Glul27Gly ncf_ _ ' _ nd "'nd*"~ _~ Asn 169 Ala 29 0 . 0 5 0 580 His205Ala nd* nd* d* His2()5Asn J 0.0029 1.1 ~' J Tl 0.0026 Glu233Asp nd* " nd* nd* w  x  a  :  1  Z~1_Z~~ 6  _  n  Data taken from Tull and Withers (7). * nd indicates that this kinetic parameter was not determined.  a  Appendix B  Compilation of Cex Data  306  Table B.10: Inhibition constants (K ) for C.Jimi xylanase and mutants with various xylobiose-derived iminosugars. x  Compound . *TFSi.»S..il,«V>*t'? . -  Wild-Type Glu43Ala Asn44Ala Lys47Ala His80Ala His80Asn His80Gln Gln87Met Gln87Tyr Asnl26Ala Glu 127 Ala Asnl69Ala  '  0.37° 110 " 23 2300 140 350 1300 4.9 16 1300* 13 2.1  0.15 22 8.9 1000 " 4 3 " " " 87 350 1.1 6.4 200* 99 0.41  Data taken from Williams et al. (8). * Data taken from Williams et al. (9).  a  . . . . . .  5  g  a  fl  2100 180 6600 800 370 590 18 62 4800* 660 17  "  45 7.1 870 5.9 13 6.7 0.39 1.2 1.6* 35 0.12  "  0.34* 96 24 2300 1000 340 340 2.1 11 2000* 28 1.5  Appendix B  Compilation of Cex Data  307  Table B . l l : Pre-steady state kinetic parameters for the hydrolysis of various aryl glycosides by C.fimi xylanase and mutants. A'cat/A', l'.n/\ me  Substrate (mM)  2,4DNPC PNPC"  Wild-Type „. Glul27Ala  b  1660 244  fl  2,4DNPC* p  N  p  c  10.0 12.4  190  C  Q  3  (s  0.68 0.17  Q  1  (s • m M " )  m M ')  1  166 20  117 26  280 1.8  130 1.5  "Data taken from Tull and Withers (7). * Data taken from MacLeod et al. (J). Data taken from MacLeod et al. (6). c  Table B.12: Michaelis-Menten parameters for the hydrolysis of various aryl glycoside substrates by C.fimi xylanase.  Substrate  (s  P N P G " " " " " PNPX" PNPC" ~ PNPX "  K ')  0,033 2.6 15.8 40  2  ...*•;.> "(mM)  8 . 3 ~" " 2 0 " 0 . 6 0 ~™ 0.018 _ _  (s  ~ "  2,4DNPG" 18 ; 2,4DNPX "" nd* _ 2,4DNPC" ~' ~~1 2 . 9 ~ , 2 , 4 D N P X ~ n d * r  —  2  •  —  '  ,.T.I^:^M-\< m M ')  0.0041 0.13 " 2 6 . " 2200  1.9 9.5 nd* ~_IT nd* ~ H 0.11 ~ " " 117 " n d * 7~ " " " n d *  " Data taken from Tull and Withers (1). * nd indicates that this kinetic parameter was not determined.  Appendix B  Compilation of Cex Data  308  Table B . 1 3 : Michaelis-Menten parameters for the hydrolysis of various aryl glycoside substrates by several family 1 0 xylanases.  Cex, C. fimf  PNPG  b  ....  —  XyllOA, £lividans XyJIOA, P. cellulose?  -  2  Cex, C. fimf : Xyl 1 OA, S. lividans  PNPX  h  . - --  XyllOA, P. celluloscf  PNPC  Cex, C. fimf XyllOA, S. lfvidans  —  XyllOA, P. celluloscf  —  -  PNPX  b  —--  Cex, C. fimf  , XyllOA, S. lividans ' b  2  XyllOA, P. cellulosa  0  Data taken from Tull and Withers (i). * Data taken from Ducros et al. (10). ° Data taken from Charnock et al. (11). *ND, not detectable. a  0.033 0.011 6  0.32 0.15 15.8 3.6 2.6 40 29 86  8.3 180 ND* -  0.0041 0.00069 ND*  20 380 310  0.13 0.00081 0.00048  0.60 51 50 0.018 0.58 Q  5  7  " "  26 0.071 0.052 2200 51 150  Appendix B  Compilation of Cex Data  Figure B . l : Scheme of protein-carbohydrate interactions in the active site region of 2F-xylobiosyl-Cex-cd complex observed by three-dimensional structural analysis (7). Hydrogen-bonding interactions between heteroatoms are in Angstroms. The distance between 2F and the carbonyl oxygen of Glu233 is 2.6 A (not shown).  309  Appendix B  Compilation of Cex Data  310  B.2  References  1.  Tull, D., and Withers, S. G. (1994) Mechanisms of cellulases and xylanases: a detailed kinetic study of the exo-P-l,4-glycanase from Cellulomonas fimi. Biochemistry. 33, 6363-6370. Notenboom, V., Birsan, C., Nitz, M., Rose, D. R., Warren, R. A. J., and Withers, S. G. (1998) Insights into transition state stabilization of the P-l,4-glycosidase Cex by covalent intermediate accumulation in active site mutants. Nat. Struct. Biol. 5, 812-818. MacLeod, A. M., Tull, D., Rupitz, K., Warren, R. A. J., and Withers, S. G. (1996) Mechanistic consequences of mutation of active site carboxylates in a retaining P1,4-glycanase from Cellulomonas fimi. Biochemistry. 35, 13165-13172. Tull, D., Withers, S. G., Gilkes, N. R., Kilburn, D. G., Warren, A. J., and Aebersold, R. (1991) Glutamic acid 274 is the nucleophile in the active site of a "retaining" exoglucanase from Cellulomonasfimi.J. Biol. Chem. 266, 1562115625. Ziser, L., Setyawati, I., and Withers, S. G. (1995) Syntheses and testing of substrates and mechanism-based inactivators for xylanases. Carbohydr. Res. 274, 137-153. MacLeod, A. M., Lindhorst, T., Withers, S. G., and Warren, R. A. J. (1994) The acid/base catalyst in the exoglucanase/xylanase from Cellulomonas fimi is glutamic acid 127: evidence from detailed kinetic studies of mutants. Biochemistry. 33, 6371-6376. Notenboom, V., Birsan, C , Warren, R. A. J., Withers, S. G., and Rose, D. R. (1998) Exploring the cellulose/xylan specificity of the P-l,4-glycanase Cex from Cellulomonasfimithrough crystallography and mutation. Biochemistry. 37, 47514758. Williams, S. J., Hoos, R., and Withers, S. G. (2000) Nanomolar versus millimolar inhibition by xylobiose-derived azasugars: significant differences between two structurally distinct xylanases. J. Am. Chem. Soc. 122, 2223-2235. Williams, S. J., Notenboom, V., Wicki, J., Rose, D. R., and Withers, S. G. (2000) A new, simple, high-affinity glycosidase inhibitor: analysis of binding through X ray crystallography, mutagenesis, and kinetic analysis. J. Am. Chem. Soc. 122, 4229-4230. Ducros, V., Charnock, S. J., Derewenda, U., Derewenda, Z. S., Dauter, Z., Dupont, C , Shareck, F., Morosoli, R., Kluepfel, D., and Davies, G. J. (2000) Substrate specificity in glycoside hydrolase family 10. Structural and kinetic analysis of the Streptomyces lividans xylanase 10A. J. Biol. Chem. 275, 2302023026. Charnock, S. J., Spurway, T. D., Xie, H., Beylot, M.-H., Virden, R., Warren, R. A. J., Hazlewood, G. P., and Gilbert, H. J. (1998) The topology of the substrate binding clefts of glycosyl hydrolase family 10 xylanases are not conserved. J. Biol. Chem. 273, 32187-32199.  2.  3. 4.  5. 6.  7.  8. 9.  10.  11.  APPENDIX C  FUNDAMENTALS OF E N Z Y M E KINETICS  Appendix C  C  Fundamentals of Enzyme Kinetics  312  Fundamentals of Enzyme Kinetics  C.1 Basic Enzyme Kinetics In 1913, L. Michaelis and M. L. Menten* proposed the following scheme: E +S ^ - ^ E S  *  cat  >E + P  (Cl)  The typical enzyme-catalyzed reaction occurs in two steps.  The enzyme  reversibly forms an initial complex with the substrate to give an enzyme-substrate complex, ES. Also known as the Michaelis complex, ES subsequently decomposes to products and enzyme in a second step with the first-order rate constant k . cat  The general expression for enzyme kinetics is known as the Michaelis-Menten equation where v is the initial velocity of the reaction; [E] is the total enzyme 0  concentration; [S] is the substrate concentration; k is the catalytic constant; and K is cat  m  the Michaelis constant.  V  =  0  cat  * +[S]  (C.2)  m  The Michaelis-Menten equation makes two assumptions.  First, the enzyme  concentration is assumed to be negligible compared to that of the substrate. This is generally true since enzymes catalyze reactions with high efficiency. Second, the reverse reaction can be ignored since the velocity measured is the initial rate of product release. Thus, there is no significant accumulation of products or depletion of substrates, and the change in substrate concentration is generally linear with time.  * Michaelis, L.; Menten, M . L . Biochem. Z49, 333 (1913).  Appendix C  Fundamentals of Enzyme Kinetics  313  The Michaelis-Menten mechanism assumes that the enzyme-substrate complex is in thermodynamic equilibrium with free enzyme and substrate, which is true only if k « 2  k_ in the following scheme: x  E + S,  ^ES—^->E + P  * In Briggs-Haldane kinetics , K  (C.3)  m  k +k =— . Nonetheless, the K  m  is the substrate  K concentration when the reaction velocity is half the maximum velocity (v = V l2). It msK  may be treated as an apparent dissociation constant of all enzyme-bound species and as such is expressed as:  (CO  * . = ™ -  £[ES]  Therefore, in the simple Michaelis mechanism, the K  m  k simplifies to K = — and s  K  as such is a measure of the enzyme affinity for the substrate. A low K indicates that the m  enzyme has a high affinity for the substrate. At low substrate concentration, where [S] « K , the Michaelis-Menten equation m  simplifies to  v  =  »  -  (C. ) 5  m  whereas at saturating concentrations, where [S] »  * Briggs, G. E.; Haldane, J. B. S. Biochem. J. 19, 338 (1925).  K , the equation becomes m  Appendix C  Fundamentals of Enzyme Kinetics  314  The Michaelis-Menten expression above describes a rectangular hyperbola such as is plotted in Figure C l .  Figure C l : Plot of reaction rate v versus substrate concentration [S] for a typical enzymatic reaction obeying Michaelis-Menten kinetics.  At low [S] the enzyme is largely unbound such that the total 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 [E][S]A-  cat  cal  v =  (C.7)  K  m  The k JK c  m  from the above equation is an apparent second-order rate constant that  relates the reaction rate to the concentration of free enzyme and free substrate. This kinetic parameter is also referred to as a specificity constant that is a measure of the catalytic efficiency for the substrate. It is not a true microscopic rate constant except in  Appendix C  Fundamentals of Enzyme Kinetics  315  the case in which the rate-determining step in the reaction is the encounter of enzyme and substrate.  The rate constant is then said to be diffusion-controlled and has been found  experimentally to be in the range of 10 to 10 s M . * 6  9  _1  -1  The Michaelis-Menten equation is often transformed into a linear form that is useful for the graphical analysis of the data and detection of deviations from ideal behaviour.  The double-reciprocal or Lineweaver-Burk plot* is a common method of  graphical analysis that is obtained by inverting both sides of the Michaelis-Menten equation.  - = v  +  v  —  v max  (C.8)  rsi  max L ^ J  Plotting 1/v as a function of 1/[S] (Figure C.2) gives an intercept of l/V  max  y axis and -l/K  m  on the x axis. The slope of the line is  * Eigen, M . ; Hammes, G. G. Adv. Enzymol. 25, 1 (1963). Lineweaver, H.; Burk, D. J. Am. Chem. Soc. 56, 658 (1934).  f  K /V . m  max  on the  Appendix C  Fundamentals of Enzyme Kinetics  316  Figure C.2: A Lineweaver-Burk plot for an enzymatic reaction.  C.2 The Significance of Michaelis-Menten Parameters The Michaelis-Menten equation remains applicable to a variety of cases in which additional intermediates occur on the reaction pathway, although £  cat  and K are now a m  combination of various rate and equilibrium constants. The general double-displacement mechanism for glycoside hydrolysis by a retaining glycosidase, originally proposed by Koshland,* proceeds through equation C.9 below. E + S^JL^ES—^->E-S—^->E + P  (C.9)  The formation of the Michaelis-Menten complex ES is referred to as the association step. The conversion of ES to the covalent intermediate E-S is known as the glycosylation step, while the release of product is referred to as the deglycosylation step. * Koshland, D. E . Biol. Rev. 28, 416 (1953).  Appendix C  Fundamentals of Enzyme Kinetics  317  Assuming a steady state concentration of ES and E-S is reached during the reaction, the Michaelis-Menten equation is given by ^[E] [S] 0  + [S] ^1 (^2  ^3 )  The equation above follows the standard form of the Michaelis-Menten equation with the following kinetic parameters for the mechanism:  ^cat =  (Cll)  k k 2+  K  =  m  3  * 3 < * - '  +  ^ \ (^2  k K  cM  m  =  * » >  (  k^k-y  1  2  )  (C.13)  k_ +k x  C  ^3 )  2  It may be shown that k  t  ca  is the rate constant for the rate-determining step of the  reaction and will always be associated with the highest free energy step in the pathway. The pseudo second-order rate constant, k JK , refers to the properties and the reactions c  m  of the free enzyme and free substrate proceeding to the transition state of the first irreversible step. Glycosylation becomes rate determining when the rate of deglycosylation is much greater than glycosylation (A: » k ). Furthermore, if association of enzyme and substrate 3  2  are rapid and reversible (k_ » k ), the kinetic parameters reduce to x  k  cat  =k  2  2  (C.14)  Appendix C  Fundamentals of Enzyme Kinetics  318  (C.15)  ^cat  _ k  _  2  (C.16)  and the Eyring equations are  Kat  kT  exp  -  'kT^ \  n  J  RT  (  exp  ( ES~ E s)^ G  G  +  RT  AG-  exp  kT\  (C.17)  RT  f  — exp  ES  (C.18)  }  E+P Reaction Coordinate  Figure C.3: Hypothetical Gibbs free energy diagram for a retaining glycosidase showing rate-limiting glycosylation (k « k ). 2  3  In this case, the kinetic parameters, k  cat  and k JK , both provide information c  m  pertaining to the transition state of the glycosylation step. However, the initial reference  Appendix C  Fundamentals of Enzyme Kinetics  319  point for k is the ES complex and for k JK , it is free enzyme (E) and free substrate csl  c  m  (S). If, however, the rate of deglycosylation becomes rate determining when the rate of glycosylation is much greater than deglycosylation (k » k ), and association of 2  3  enzyme and substrate are rapid and reversible (&_,» k ), the kinetic parameters are 2  reduced to ^cat  £  (C.19)  K  _  _  _ £  k  x  Kat  k  2  2  _ Jh_  _  K  k_  m  (C.20)  3  kk  (C.21)  K  x  s  which in Eyring form become (GI_ -G _ ) S  ( ) { h kT  Kat  -  V «  J  J  exp V  exp  E  kT  S  RT  exp  h  (^ES-^E+s)  =  RT J  (  (kT)  I  —  h  RT  exp J  —  AG*) RT ,  (C.22)  (C.23)  Appendix C  Fundamentals of Enzyme Kinetics  320  AG*. E-S E-S* AG\  A G  E +S E+P Reaction Coordinate  Figure C.4: Hypothetical Gibbs free energy diagram for a retaining glycosidase showing rate-limiting deglycosylation (A: « k ). 3  In this case,  k t ca  2  refers to the transition state of deglycosylation with the covalent  intermediate complex as the initial reference point and k JK refers to the transition state a  m  of the glycosylation step with free enzyme [E] and free substrate [S] as the initial reference points. Thus, in these examples, k  refers to the rate-determining step while k JK c  cat  m  corresponds to the first irreversible step in the reaction with free enzyme and free k  k  substrate as the reference states. In general, K Ki= — , except for the Briggs-Haldane rn S m  mechanism in which the enzyme-substrate complex is not in thermodynamic equilibrium with the free enzyme and substrate. If the first chemical step of the reaction accelerates to such an extent that the encounter of the enzyme and the substrate become rate determining apparent second-order rate constant k JK c  m  « k ) , the 2  approaches the diffusion-controlled limit of  Appendix C  Fundamentals of Enzyme Kinetics  10 s M . 9  _1  321  The k . is unaffected and is dependent on whether glycosylation or  _1  C3  t  deglycosylation is the rate-determining step of the reaction. Assuming product release is rate determining, the kinetic parameters reduce to ^cat  -  (C.24)  ^3  (C.25)  (C.26)  which in Eyring form become  ^cat  _  V  '"cat  nJ  exp  'kT^ v h j  exp  (^E-S-^E-s)  RT  (GES-GE+S)  RT  h)  =  fkT^ —  exp  RT J  f  exp  AG*") —  1  RT ,  (C.27)  (C.28)  Appendix C  Fundamentals of Enzyme Kinetics  322  Figure C.5: Hypothetical Gibbs free energy diagram for a retaining glycosidase showing rate-limiting association (k_ « k ). }  2  C.3 Enzyme Inhibition Enzymes may be  reversibly  inhibited by the non-covalent binding of inhibitors  according to four binding modes.  C.3.1 Competitive inhibition  An inhibitor is said to be a  competitive  inhibitor when it binds reversibly to the  active site of an enzyme and prevents the substrate S from binding and C.6).  vice versa  (Figure  Appendix C  Fundamentals of Enzyme Kinetics  E + S + I  ^  ES  323  E + S  EI  Figure C.6: The simple Michaelis-Menten mechanism showing an addition equilibrium where K, = |E][I]/[EI|.  The Michaelis constant K appears to increase by a factor of (1 + [I]/AT) as in the m  ;  equation below.  v=  [  E  ]  °  [  S  ]  ^'  (C.29)  [sj+^a+ti]/^) Infinitely high concentrations of S displace I from the enzyme, therefore competitive inhibition affects only K and not Vmaxm  m  C.3.2 Noncompetitive, uncompetitive, and mixed inhibition  Simultaneous binding of I and S to the enzyme instead of competition for the same binding site results in different inhibition patterns (Figure C.7).  Appendix C  Fundamentals of Enzyme Kinetics  E +S  324  ES + I  +  I K  K  EI + S  EIS  Figure C.7: Thermodynamic box where the inhibition constant of the free enzyme is K. ; that of the enzyme-substrate complex is K.; and the dissociation constants of EIS and ES are K and K , respectively. s  s  Noncompetitive inhibition  occurs when the dissociation constant of S from ES is  the same as that from EIS (that is, K = K ), and EIS does not react so that 5  „ , BUS]*.  s  /Q+[!]«,)  In the above case of noncompetitive inhibition, (1  + [IJ/TCj) and K is unaffected. m  Mixed inhibition  k  cat  is lowered by a factor of  occurs when the dissociation constant  of S from ES is different than that from EIS (K * K ), and results in altered values of s  K  m  and  k .  Uncompetitive inhibition  cat  s  occurs when I binds to ES but not to E. In this  case, both K and A" are altered by the same factor, therefore m  cat  k /K cat  m  remains unchanged.  Appendix C  Fundamentals of Enzyme Kinetics  325  C.4 Binding Energy and Enzyme Catalysis In 1978 Schowen* proposed "...that the entire and sole source of catalytic power is the stabilization of the transition state; that reactant-state interactions are by nature inhibitory and only waste catalytic power." Enzymes, like other catalysts, increase reaction rates by lowering activation energies. A unique feature of enzyme catalysis is that the enzyme binds specifically to the substrate and the binding energies involved are used to lower activation energies. However, since the structure of the substrate changes during the reaction, the enzyme can be complementary to only the ground-state or transition-state form of the substrate. It will be shown that it is catalytically advantageous for the enzyme to be complementary to the transition-state structure rather than to the ground-state form of the substrate. Consider a typical enzymatic reaction consisting of a binding step followed by a catalytic step such as that shown below. E +S^SL^ES  *  >E + P  cal  (C.31)  The energy diagram for this reaction is illustrated in Figure C.8 where k_ » k , x  so that K = Ks and the kinetic parameters may be expressed as the following. m  \ h j  exp  AG  RT J  V  r  () exp { h ) kT  S  AG*  \  (C.33)  —  RT  (C.32)  /  * Schowen, R. L. Transition States of Biochemical Processes, Plenum, New York (1978).  2  Appendix C  Fundamentals of Enzyme Kinetics  fkT^ V  exp  " J  326  (C.34)  " RT  Figure C.8: The reaction coordinate diagram for a typical reaction (solid line) and the corresponding uncatalyzed reaction (dashed line).  Consider the above situation when an additional amount of binding energy, A G R , has become available in the form of a hydrogen bond between the enzyme and the substrate. If that extra binding energy is realized in the enzyme-substrate (ES) complex (Figure C.9, left) rather than the transition-state (ES*) complex, then the ground state is stabilized and AGs increases by A G R resulting in a decrease of K . m  The value of k is cat  also reduced since AG£ (enzyme-substrate complex, ES, proceeding to the transition state, ES*) is increased by the ground-state stabilization,  AGR.  However k /K is cat  m  unaffected as this is the rate constant for free enzyme (E) and free substrate (S) proceeding to the transition state (ES*). Therefore, if the enzyme is complementary to the ground state, then there is tighter substrate binding but a slower reaction rate, k . cat  Appendix C  Fundamentals of Enzyme Kinetics  327  Alternatively, if the extra binding energy, A G R , is realized only in the transition-state complex (Figure C.9, right), then the transition state is stabilized and AG£ and A G | will be lowered by  AGR,  thus,  k  cai  and k  /K  cat  m  will be increased.  The value of AGs is  unchanged, and hence, K remains the same. Transition-state stabilization can therefore m  lower the activation energies of k  cat  consequence.  and k  /K ,  cai  m  increasing reaction rates as a  Ground-state stabilization has no effect on k /K , although the k is cat  m  cat  decreased.  Figure C.9: The reaction coordinate diagram illustrating complementarity of the enzyme to (left) the ground state, and (right) the transition state of the substrate.  C.5 The pH Dependence of Enzyme Catalysis Enzymes contain a significant number of ionizing groups, some of which are directly involved in catalysis at the active site, while others may be responsible for maintaining the active conformation of the enzyme. Enzyme activity can vary with pH since the active sites contain important acidic or basic groups, where only one protonic  Appendix C  Fundamentals of Enzyme Kinetics  328  form of the acid or base is catalytically active. Plots of rate against pH take the form of simple single or double ionization curves because, although enzymes contain a multitude of ionizing groups, the only ionizations that are of importance are those of groups that are directly involved in catalysis at the active site. Four simplifying assumptions include: 1.  The groups act as perfectly titrating acids or bases.  2. Only one ionic form of the enzyme is active. 3.  Proton transfers are faster than chemical steps.  4. The rate-determining step does not change with pH.  C.5.1 The Michaelis-Menten mechanism In the simple Michaelis-Menten mechanism, the following equilibria may be set up:  E + S  + H  ES + H  slow  ES  H E + S "==F  HES  Figure C.10: Thermodynamic box where the ionization constant of the free enzyme is K ; that of the enzyme-substrate complex is A' ; and the dissociation constants of ES  E  HES and ES are K and K , respectively. s  s  Appendix C  Fundamentals of Enzyme Kinetics  329  Due to the cyclic nature of the equilibria, all four equilibrium constants are related by the equation: (C.35)  -^E^S - ^ E S ^ S  Assuming K = K , the pH dependence of v is given by the following expression: m  s  *ca,[E] [S] 0  V  H  = '  (C.36)  [H ] +  K +[S] S  ^ES J  V  When [S] »  Ks,  (^ax)H=[E]o(*  the pH dependence of k is derived from equation C.36: cat  c a t  )  H  =  [E]p -^ES  ^cal  (C.37)  ^ES+[H ] +  The pH dependence of V  max  or k follows the ionization constant of the enzymecat  substrate complex, KESThe apparent value of K (where K = K ) at each pH is derived by rearranging M  M  S  equation C.36: [FTf 1+  (  (C.38)  1+^ K  V  c  ES J  K also follows the ionizations of the enzyme-substrate complex. m  When [S] « k  f  the pH dependence of k  K, m  /K  cat  ^  m  is derived from equation C.36:  (C.39) \  E J  K  Appendix C  Fundamentals of Enzyme Kinetics  The p H dependence o f k /K cat  m  330  follows the ionizations in the free enzyme and free  substrate.  C.5.2 Modifications and breakdown of the Michaelis-Menten mechanism  If a mechanism more complicated than the simple Michaelis-Menten occurs, additional considerations must be taken into account.  E +S^Jl—1ES—^-»E-S—^->E +P  (C.40)  The presence o f additional intermediates on the reaction pathway does not affect the p H dependence o f k /K . cat  m  The pAT 's obtained from the p H profile will correspond to a  the pAT 's o f the free enzyme and free substrate.* However, the p H dependence of Ar and a  cat  K will give the pAV s of the major enzyme-bound species. The p H profiles will give the m  p A V s that are dependent on the concentration o f the intermediates and the enzymesubstrate complex.*  When a substrate binds in a non-productive as well as in the productive mode, the p H dependence of k  cai  or K may give an apparent pA: that significantly deviates from the m  a  real value. This occurs i f the ratio o f productive to non-productive binding is dependent on ionization o f a group in the active site of an enzyme. The p H dependence of k /K cat  not affected by non-productive binding modes.*  * Bender, M . L.; Clement, G. E.; Kezdy, F. J.; Heck, H. d'A. J. Am. Chem. Soc. 86, 3680 (1964). Fersht, A . R.; Requena, Y . J. Am. Chem. Soc. 93, 7079 (1971). Fersht, A . R.; Fastrez, J. Biochemistry 12, 1067 (1973).  f :  m  is  Appendix C  Fundamentals of Enzyme Kinetics  331  The simple rules for pH dependence breakdown in the case of Briggs-Haldane kinetics:* E + S,  ^ES—^->E + P  (C.41)  A change of rate-determining step with pH may give rise to anomalous pAVs. When the rate constant for the chemical step is larger than that for the dissociation of the enzyme-substrate complex,  k JK c  m  is equal to k , the association rate constant of the x  enzyme and the substrate. Suppose that the catalytic activity of an enzyme depends on the ionization of a base so that k will be faster than k_ at high pH, but will decrease as 2  x  the pH is lowered and the base becomes protonated until k is slower than k_ . The 2  apparent  pA'a  obtained from a pH profile of k JK c  m  results in a  x  kinetic pK  a  that is lower  than the actual pK of the group. If the ionization constant of the group on the enzyme is a  K , the apparent ionization constant is given by a  K = 3PP  K(k'+k ) ' I k_ x  x  * Schmidt, D. E., Jr.; Westheimer, F. H . Biochemistry 10, 1249 (1971).  (C42)  Appendix C  Fundamentals of Enzyme Kinetics  332  pH  Figure C . l l : Illustration of how a change in rate-determining step from a chemical event at low pH to the encounter of enzyme and substrate at high pH may lead to an anomalous pK in the k JK pH profile. a  c  m  In general, kinetic pA^ 's occur whenever there is a change of rate-determining a  step with pH.  * Renard, M . ; Fersht, A. R.; Biochemistry 12, 4713 (1973).  

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