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An investigation of the mechanism of the Cellulomonas fimi exoglucanase Tull, Dedreia L. 1991

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A N INVESTIGATION OF THE MECHANISM OF THE CELLULOMONAS FIMI EXOGLUCANASE By DEDREIA L. TULL B.Sc, McGill University, 1989 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Department of Chemistry)  We accept this thesis as conforming to the required standard  THE UNIVERSITY OF BRITISH COLUMBIA September 1991 © Dedreia L. Tull  In  presenting  degree at the  this  thesis  in  University of  partial  fulfilment  of  of  department  this thesis for or  by  his  or  requirements  British Columbia, I agree that the  freely available for reference and study. I further copying  the  representatives.  an advanced  Library shall make it  agree that permission for extensive  scholarly purposes may be her  for  It  is  granted  by the  understood  that  head of copying  my or  publication of this thesis for financial gain shall not be allowed without my written permission.  Department  of  Qr *'SAMS~\ Q s ^ j  The University of British Columbia Vancouver, Canada  DE-6 (2/88)  ii  ABSTRACT The exoglucanase from Cellulomonasfimicatalyses the hydrolysis of cellobiose units from the non-reducing terminus of cello-oligosaccharides with overall retention of anomeric configuration. Its mechanism of action is therefore thought to involve a double displacement reaction, involving as the first step, formation of a glycosyl-enzyme intermediate (glycosylation) and as a second step, the hydrolysis of this intermediate (deglycosylation). This mechanism is investigated here through the study of the kinetics of hydrolysis of aryl (3-glucosides and aryl (3-cellobiosides and by employing the mechanismbased irreversible inactivators, 2', 4'-dinitrophenyl 2-deoxy-2-fluoro-p -D-glucoside (2F>  DNPG) and 2", 4"-dinitrophenyl 2-deoxy-2-fluoro-p-D-cellobioside (2F-DNPC). The study with the aryl P-glucosides revealed that this enzyme is indeed active on glucosides, a feature that had previously been undetected. A linear relationship was found to exist between the logarithm of V a x for hydrolysis and the phenol p K as well as m  between the logarithm of Vmax/Krn " ana  a  m  e  phenol pK , showing that glycosylation is both a  the rate determining step and the first irreversible step for all substrates. The reaction constant calculated, p = 2.21, indicates a considerable amount of charge build up at the transition state of glycosylation. The linear free energy relationship study of the aryl P-cellobiosides revealed no significant dependence of the logarithm of V a x on the pK of the phenol, indicating that m  a  deglycosylation is rate determining. However, the slight downward trend in this Hammett plot at higher p K values may suggest that the rate determining step is changing from a  deglycosylation to glycosylation. However, the logarithm of V a x / K m does correlate with m  the pK of the phenol, thus showing that thefirstirreversible step is glycosylation. The a  reaction constant (p = 0.60) which reflects the development of charge at the glycosylation transition state for the cellobiosides is less than that calculated for the glucosides, thus suggesting a glycosylation transition state with either a greater degree of acid catalysis or less C-O bond cleavage than that for the glucosides.  i ii  The inactivators, 2F-DNPC and 2F-DNPG, are believed to inactivate the exoglucanase by binding to the enzyme and forming covalent glycosyl-enzyme intermediates. The inactivated-enzyme was stable in buffer but reactivated in the presence of a suitable glycosyl-acceptor such as cellobiose, presumably via a transglycosylation reaction. These results indicate that covalent 2F-glycosyl-exoglucanase intermediates are stable and are catalytically competent to turn over to product, thus supplying further evidence for the Koshland mechanism. The exoglucanase is inactivated more rapidly by 2F-DNPC than by 2F-DNPG. However, both inactivated forms of the enzyme reactivated at comparable rates in the presence of cellobiose, showing that the second glucosyl unit present on the cellobiosides increases the rate of glycosylation relative to that found for the glucosides but not the rate of deglycosylation. The stable covalent nature of the 2F-glycosyl-enzyme intermediates provided an excellent opportunity to identify the enzymic nucleophile. This was accomplished by radiolabelling the exoglucanase with a tritiated analogue of 2F-DNPG cleaving the protein into peptides and purifying the radiolabelled peptides. Sequencing of this peptide resulted in the identification of the active site nucleophile as glutamic acid residue 274. This residue was found to be highly conserved in this family of (3-glycanases, further indicating its importance in catalysis.  iv  TABLE OF CONTENTS ABSTRACT TABLE OF CONTENTS LIST OF FIGURES LIST OF T A B L E S LIST OF ABBREVIATIONS AND DEFINITIONS ACKNOWLEDGEMENT  ii iV vi vii viii xi  CHAPTER 1 INTRODUCTION 1 GLYCOSIDASES  1 1 1  2  2 4 5 7 9 10 12 12 18 21 22 25 28  3 4 5 6  MECHANISM FOR RETAINING GLYCOSIDASES 2.1 Enzymic Carboxylate Group 2.2 Covalent Glycosyl-enzyme Intermediate 2.3 Oxocarbonium Ion-Like Transition States 2.4 Acid Catalysis 2.5 Noncovalent Interactions LINEAR FREE ENERGY RELATIONSHIPS 3.1 The Hammett Equation 3.2 Linear Free Energy Relationships in Enzymology IRREVERSIBLE INACTIVATORS 4.1 Irreversible Inhibition Studies in Enzymology THE CELLLASE COMPLEX THE AIM OF THIS STUDY  CHAPTER 2 RESULTS A N D DISCUSSION 1 2  3  4  HYDROLYSIS OF ARYL p-D-GLUCOSIDES AND ARYL p-DCELLOBIOSIDES BY C. FIMI EXOGLUCANASE INACTIVATION OF C.F1MI EXOGLUCANASE 2.1 Kinetics of inactivation 2.2 Covalent Inactivation 2.3 Reactivation of Inactivated C. fimi Exoglucanase IDENTIFICATION OF THE CATALYTIC NUCLEOPHTLE OF C. FIMI EXOGLUCANASE 3.1 Conservation of Glu-274 of C.fimiexoglucanase in other glycosidases CONCLUSION  29 29  29 39 40 41 42 45 46 48  CHAPTER 3 MATERIALS AND METHODS  50 50  1  50 50 51 52  SYNTHESIS 1.1 General Procedures and Materials 1.2 General compounds 1.3 Aryl 2,3,6,2*,3',4',6'-hepta-0-acetyl p-D-cellobiosides  V 1.4  2  Aryl B-D-cellobiosides 1.4.1 HCl/MeOH Method 1.4.2 NaOMe/MeOH Method ENZYMOLOGY 2.1 General procedures 2.2 Determination of the molar extinction coefficient of the C.fimi exoglucanase 2.3 Determination of K and V x for the hydrolysis of aryl 0-Dm  2.4  2.5 2.6  52 52 53 60 60 60  ma  glucosides & aryl P-D-cellobiosides by C.fimi exoglucanase Determination of Kj and lq for 2',4'-dinitrophenyl 2-deoxy-2fluoro-P-D-glucoside and 2",4"-dinitrophenyl 2-deoxy-2-fluoro-PD-cellobioside Reactivation of inactivated C.fimi exoglucanase Determination of the catalytic nucleophile of C.fimi exoglucanase with {1-3HJ-2F-DNPG  61  62 63 63  APPENDIX 1 BASIC CONCEPTS OF ENZYME CATALYSIS  66 66  1 BASIC E N Z Y M E KINETICS 2 INTERPRETATION OF kcat AND k c a t / K 3 BINDING ENERGY AND ENZYME CATALYSIS 4 INACTIVATION KINETICS OF C. FIMI EXOGLUCANASE  66 68 71 75  APPENDIX 2 GRAPHICAL REPRESENTATION OF KINETIC DATA  77 77  m  1 2 3  HYDROLYSIS OF ARYL p-GLUCOSIDES AND ARYL pCELLOBIOSIDES BY C. FIMI EXOGLUCANASE PLOTS OF THE INACTTVATTON OF C. FIMI EXOGLUCANASE WITH 2F-DNPC AND 2F-DNPG PLOTS OF THE REACTIVATION OF INACTIVATED C. FIMI EXOGLUCANASE  REFERENCES  77 83 85 86  LIST OF FIGURES Figure 1. Hydrolysis of glycosides by glycosidases Figure 2. Stereochemical classification of glycosidases. Figure 3. Koshland double displacement mechanism for retaining (J-glycosidases Figure 4. Figure 5. Figure 6. Figure 7. Figure 8.  such as a P-glucosidase The peptidoglycan substrate for lysozyme Labelling of the nucleophilic carboxylate of Agrobacterium |3glucosidase Comparison of aldonolactones with glycosyl cations Nojirimycins, transition state analogues for glycosidases Structure of P-D-galactopyranosyl pyridinium salt  Figure 9. FigurelO. Figure 11. Figure 12. Figure Figure Figure Figure Figure  Hydration of an octenitol derivative by E. coli P-galactosidase Substrate analogue for lysozyme The general compound used in Hammett correlations Schematic representation of a concave upward Hammett plot for a reaction with two possible paths (1 & 2) 13. Schematic representation of a concave downward Hammett plot for a multistep reaction 14. Schematic representation of papain-catalysed hydrolysis 15. Papain-catalysed hydrolysis of N-acetyl-L-phenylalanylglycine anilides 16. Hammett plot for papain-catalysed hydrolysis of N-acetyl-Lphenylalanylglycine anilides 17. Hammett plot for sweet almond P-glucosidase-catalysed hydrolysis of aryl P-glucosides  1 2 3 4 5 7 8 9 10 11 12 16 17 18 19 19 20  Figure 18. Structure of N-bromoacetyl-P-D-glycosylamines Figure 19. Inactivation of A. wentii P-glucosidase with conduritol B cis-epoxide Figure 20. Schematic representation of the degradation of cellulose to glucose by the cellulase complex Figure 21. Exoglucanase-catalysed hydrolysis of (a) aryl p-D-glucosides and (b) aryl P-D-cellobiosides Figure 22. The log V ax $ pK Hammett correlation for the hydrolysis of Pglucosides by exoglucanase Figure 23. The log V x / K vs pK Hammett correlation for the hydrolysis of Pglucosides by exoglucanase Figure 24. The log V ax vs pK Hammett correlation for the hydrolysis of Pcellobiosides by exoglucanase Figure 25. The log W /K vs pK Hammett correlation for the hydrolysis of Pcellobiosides by exoglucanase Figure 26. Varied degrees of acid catalysis at the glycosylation transition state for glucosides and cellobiosides Figure 27. Different degrees of C-0 bond cleavage at the glycosylation transition state for the glucosides and the cellobiosides Figure 28. Energy diagram showing the stabilization produced by the second  23 24 26 30  v  m  a  m a  m  m  32  a  33  a  max  m  34  a  35 36 37  vn glucosyl unit of the cellobiosides Figure 29. Proposed mechanism of inactivation of C./7miexoglucanase.by (a)2FDNPG and (b) 2F-DNPC Figure 30. The proposed mechanism for the reactivation of inactivatedexoglucanase by cellobiose Figure 31. Velocity vs substrate concentration for a typical enzyme reaction Figure 32. A typical Lineweaver-Burk plot Figure 33. Free energy diagram for the enzymatic reaction involving the interconversion of intermediates Figure 34. The free energy diagram for a typical reaction (solid line) and the corresponding uncatalysed reaction (dashed line) Figure 35. The free energy diagram when the enzyme is complementary to the ground state of die substrate Figure 36. The free energy diagram when the enzyme is complementary to the transition state of the substrate Figure 37. The Lineweaver-Burk plot for the hydrolysis of 2,4 DNPG Figure 38. The Lineweaver-Burk plot for the hydrolysis of 3,4 DNPG Figure 39. The Lineweaver-Burk plot for the hydrolysis of 2C14NPG Figure 40. The Lineweaver-Burk plot for the hydrolysis of PNPG Figure 41. The Lineweaver-Burk plot for the hydrolysis of MNPG Figure 42. The Lineweaver-Burk plot for the hydrolysis of 2,4 DNPC Figure 43. The Lineweaver-Burk plot for the hydrolysis of 3,4 DNPC Figure 44. The Lineweaver-Burk plot for the hydrolysis of PNPC Figure 45. The Lineweaver-Burk plot for the hydrolysis of 4C12NPC Figure 46. The Lineweaver-Burk plot for the hydrolysis of MNPC Figure 47. The Lineweaver-Burk plot for the hydrolysis of 3,5 DC1PC. Figure 48. The Lineweaver-Burk plot for the hydrolysis of 4 CNPC Figure 49. The inactivation of C.fimi exoglucanase with 2F-DNPC Figure50. Plot of 1/kobs vs. 1/TTJ for 2F DNPC Figure 51. The inactivation of C.fimi exoglucanase with 2F DNPG Figure52. Plot of 1/koos vs. 1/[I] for 2F DNPG Figure 53. Reactivation of 2F-DNPC-inactivated exoglucanase in the presence of a glycosyl-acceptor Figure 54. Reactivation of 2F-DNPG-inactivated exoglucanase in the presence of glycosyl-acceptors  39 42 44 67 68 69 73 74 74 77 77 78 78 79 79 80 80 81 81 82 ..82 83 83 84 84 85 .85  viii LIST OF TABLES Table 1. Kinetic parameters for the inactivation of E. coli P-galactosidase with Nbromoacetyl -P-D-glycosylamines  23  Table 2. Michaelis-Menten parameters for the hydrolysis of the p-glucosides and P-cellobiosides.by C.fimi exoglucanase Table 3. Kinetic parameters for the inactivation of C.fimi exoglucanase by 2FDNPG and 2F-DNPC Table 4. The rates of reactivation of inactivated exoglucanase in the presence of a glycosyl-acceptor  31 41 43  ; i  x  LIST OF ABBREVIATIONS AND DEFINITIONS  Abbreviations: BSA  Bovine serum albumin  SDS-PAGE  Sodium dodecyl sulfate polyacrylamide-gel electrophoresis  2,4 DNPG  2',4'-Dinitrophenyl P-D-glucoside  2C1 4NPG  2'-Chloro-4'-nitrophenyl P-D-glucoside  3,4 DNPG  3',4'-Dinitrophenyl P-D-glucoside  MNPG  3'-Nitrophenyl p-D-glucoside  PNPG  4'-Nitrophenyl P-D-glucoside  2,4 DNPC  2",4"-Dinitrophenyl P-D-cellobioside  3.4 DNPC  3",4"-Dinitrophenyl P-D-cellobioside  3.5 DC1PC  3",5"-Dichlorophenyl P-D-cellobioside  4C1 2NPC  4"-Chloro-2"-nitrophenyl P-D-cellobioside  4 CNPC  4"-Cyanophenyl P-D-cellobioside  MNPC  3"-Nitrophenyl P-D-cellobioside  PNPC  4"-Nitrophenyl P-D-cellobioside  2F-DNPC  2",4"-Dinitrophenyl 2-deoxy-2-fluoro P-D-cellobioside  2F-DNPG  2',4'-Dinitrophenyl 2-deoxy-2-fluoro P-D-glucoside  {1- H} 2F-DNPG  [l-3H]-2',4'-dinitrophenyl 2-deoxy-2-fluoro-p-D-glucoside  Arg  Arginine  Asn  Asparagine  Glu  Glutamic Acid  Gly  Glycine  lie  Isoleucine  Leu  Leucine  Thr  Threonine  3  X  Tyr  Tyrosine  Val  Valine  NMR  Nuclear magnetic resonance  m.p.  Melting point  Kinetic and Physical Constants  T  Temperature (K or °C)  k  Boltzmann constant  h  Planck's constant  R  Gas constant  Km  Michaelis-Menten constant (the apparent dissociation constant for all bound enzyme-substrate species)  Vmax  Maximal rate of an enzyme-catalyzed reaction  kcat  First-order rate constant for catalysis (turnover number)  Kj  Inhibition constant (the apparent dissociation constant for all bound enzyme inhibitor species)  ki  First-order rate constant of inactivation  X1  ACKNOWLEDGEMENT  I wish to express my appreciation to my supervisor, Dr. Stephen G. Withers, for his help and encouragement during the time of this research. Thanks also to my coworkers for their helpful discussion and insights. Special thanks to Karen Rupitz and Carola Ibe for their assistance. I would like to thank the University of British Columbia for financial support and also the nmr and elemental analysis staff of the Department of Chemistry for their help. Thanks to Sanny Chan for her help and support during the writing of this document. To my parents, I wish to say thanks for believing and having confidence in me. To them, I dedicate this thesis.  1  CHAPTER 1 INTRODUCTION  1.  GLYCOSIDASES Glycosidases are enzymes that catalyse the hydrolysis of the glycosidic linkage  between two sugar residues (Fig 1). The substrates are made up of two parts, a glycone portion and an aglycone portion (R). The natural substrates for most glycosidases contain a second sugar residue as the aglycone; however, since most of these enzymes are less specific for the aglycone than the glycone this sugar can frequently be replaced by an alkyl or aryl group.  Figure 1. Hydrolysis of glycosides by glycosidases.  The glycosidases constitute a large class of enzymes which can be subdivided into smaller groups based on the following features. (1)  The type of glycone to which it is reactive such as pyranose or furanose and glucose or galactose (for example galactofuranosidase and galactopyranosidase). However, it should be noted that although a glycosidase may be active against a variety of substrates, that enzyme is classified based on the sugar against which it is most active.  (2)  The anomeric configuration (a or P) of the glycosidic bond that is to be cleaved. For example a P-glycosidase will only hydrolyse P-glycosides and not oc-glycosides.  2 (3)  The relative anomeric configuration of the substrate versus the initial product That is, if the anomeric configurations of the substrate and the product are the same then the enzyme is classified as a retaining glycosidase whereas if the anomeric configurations of the substrate and product are different then enzyme is classified as inverting (Fig 2).  Figure 2. Stereochemical classification of glycosidases.  The following discussion will be focused on studies carried out on retaining glycosidases.  2.  M E C H A N I S M FOR RETAINING GLYCOSIDASES A double displacement mechanism has been proposed by Koshland for retaining  glycosidases. This mechanism is outlined in Fig 3 and has the following mechanistic 1  features. (1)  There is a carboxylate group in the enzyme active site that is located on the opposite side of the sugar ring to the aglycone.  3  4  (2)  The carboxylate group forms a covalent axial glycosyl-enzyme intermediate at C-l of the glycone.  (3)  This covalent intermediate is formed and degraded via oxocarbonium ionlike transition states.  (4)  Acid catalysis may or may not assist bond cleavage.  (5)  Most of the observed rate acceleration is derived from the noncovalent interactions between enzyme and substrate.  There is much evidence in the literature for this mechanism . Some of this is presented below.  2.1  Enzymic Carboxylate Group Evidence for the presence of a carboxylate group in the active site of these  glycosidases comes from X-ray crystallographic and irreversible inhibition studies. The X-ray crystal structures of three lysozymes provided the most compelling evidence for the existence of a catalytic carboxylate group. These enzymes catalyse the  NH I CO CH  NH I CO 3  CH  NH I CO 3  CH  NH I CO 3  CH  3  R = CH CHCOOH 3  I  Figure 4. The peptidoglycan substrate for lysozyme: the arrow indicates the bond that is cleaved.  5 hydrolysis of the peptidoglycan component of the cell wall in bacteria (Fig 4). From the three dimensional structure of hen's egg white (HEW) lysozyme, aspartic acid (asp) 52 was observed to be correcdy positioned in the active site to be the carboxylate group proposed by Koshland as the nucleophile. Later X-ray crystallographic studies on T4 lysozyme and goose egg white (GEW) lysozyme identified asp 20 and asp 86 3  4  respectively, as the carboxylate groups corresponding to asp 52 of HEW lysozyme. In other cases the carboxylate groups were identified by labelling experiments with irreversible inactivators. Glutamic acid (glu) 358 was identified as the catalytic carboxylate in the Agrobacterium |}-glucosidase using the radiolabeled inactivator 2',4'-dinitrophenyl 2-deoxy-2-fluoro-p -D-glucoside (Fig 5) which operates by trapping the intermediate. >  5  Isolation and sequencing of the labelled peptide allowed identification of this residue.  Glu 358  Figure 5. Labelling of the nucleophilic carboxylate of Agrobacterium B-glucosidase.  2.2  Covalent Glycosyl-enzyme Intermediate Although Koshland's mechanism proposed the formation of a covalent glycosyl-  enzyme intermediate, Phillips and coworkers have suggested, based on the X-ray crystal 6  6 structure of the HEW lysozyme active site, that an ion-pair made up of asp 52 and the oxocarbonium ion would be long-lived enough for the leaving group to diffuse away from the active site and allow the glycosyl acceptor, water, to diffuse in and react. It is this concept which has been popularized in textbooks, however there has been no additional data to support the involvement of an ion-pair intermediate in the hydrolysis of glycosides. Studies of spontaneous (non-enzymatic) hydrolysis of glycosides have shown that glycosyl cations are quite unstable, with estimated lifetimes varying from 10" to 10" seconds in 10  12  aqueous solution. This can be compared to the lifetime of the glycosyl-enzyme which 7  o ranges from 1-100 ms at ambient temperatures.  It therefore seems more likely that the  glycosyl-enzyme intermediate is a covalent and not an ion-pair species. An example of an enzyme system in which such a covalent enzyme intermediate has been shown to exist, comes from the work done on the Agrobacterium (3-glucosidase using such substrates analogues as 2-deoxy-2-fluoro-|3-D-glucosides (Fig 5). These 29  deoxy-2-fluoro-P-D glucosides inactivate the enzyme because the presence of the electronegative fluorine at the C-2 position destabilizes the oxocarbonium ion-like transition states, thus slowing down both steps, glucosylation and deglucosylation (Fig 3). However, the presence of a good leaving group such as fluoride or 2,4 dinitrophenolate speeds up glucosylation and hence, allows for the accumulation of the intermediate. This covalent intermediate is stable with a half life of over 500 hours at 37 °C in buffer. The covalency and stereochemistry (a) of this intermediate have been demonstrated by F 1 9  NMR.  10  The trapping of this covalent glucosyl-enzyme intermediate shows that such  intermediates do exist and are stable. Furthermore, the catalytic competence of the intermediate is demonstrated by the fact that it turns over to product in the presence of a suitable glycosyl acceptor via a transglycosylation reaction, thus resulting in reactivation.  7 2.3  Oxocarbonium Ion-Like Transition States Although, there is evidence which indicates that a covalent glycosyl-enzyme is the  intermediate in enzymatic glycoside hydrolysis, there is also evidence which suggests that this intermediate is reached and hydrolysed via transition states with oxocarbonium ion character. Such information comes from studies with transition state analogues and from measurement of kinetic isotope effects. Transition state analogues are glycoside derivatives that sufficiently mimic the transition state in structure such that they bind to the enzyme significantlytighterthan do the substrates. Such transition state analogues are therefore expected to have one or both of the following characteristic features of a glycosyl cation: the C-l and 0-5 atoms share a full positive charge and the C-5, 0-5, C - l and C-2 atoms are coplanar.  11  Such features  distinguish the glycosyl cation from the parent glycoside. Aldonolactones resemble glycosyl cations both in geometry and to some extent in charge (due to the resonance form shown) and are generally considered transition state analogues (Fig 6). Indeed, these lactones bind to glycosidases with Kj values two to three orders of magnitude lower than the K values for normal substrates. s  Figure 6. Comparison of aldonolactones with glycosyl cations.  8 Nojirimycin (5-amino-5-deoxy-D-glucopyranose) and its other sugar analogues, for example mannonojirimycin, are powerful inhibitors of P-glycosidases. When these 5amino-5-deoxy aldoses are protonated, they are isoelectronic with the glycosyl cation and when they are also dehydrated, they are both isoelectronic and isosteric with respect to the glycosyl-cation (Fig 7). These compounds are transition state analogues and they have been found to inhibit a wide range of glycosidases with affinities ranging from weak (Kj = 4.6 mM for A. wentii (3-mannosidase) to strong (Kj = 2.7 u,M for A. wentii (3-glucosidase).  OH  Figure 7. Nojirimycins, transition state analogues for glycosidases.  The above transition state analogues have supplied evidence for an oxocarbonium ion-like transition state in the formation of the glycosyl enzyme intermediate. Deuterium kinetic isotope effect studies on glycosidases using specific substrates for which either glycosylation or deglycosylation is the rate determining step have also provided substantial evidence for oxocarbonium ion character at the transition state of these two steps. For example, an a-deuterium kinetic isotope effect of k n / k D = 1.21 was observed for the degalactosylation step of the lac z P-galactosidase of E. coli using 2',4'-dinitrophenyl p*-Dgalactoside as substrate.  An a-deuterium kinetic isotope effect of k n / k o = 1.15 - 1.20  was measured for the galactosylation step of the same enzyme system using (3-Dgalactopyranosyl pyridinium salts as substrates (Fig 8).  Figure 8. Structure offi-D-galactopyranosylpyridinium salt.  Such large a-deuterium kinetic isotope effects indicate substantial oxocarbonium ion character at the transition state of the rate determining step because the presence of an isotope effect reveals that the substituted carbon centre is converting from an sp hybridized ground state to a transition state with considerable sp character. If however, an adeuterium kinetic isotope effect of unity was observed then that would either indicate an SN2 mechanism, as there would be no change in hybridization, or that some other step was rate determining. An a-deuterium kinetic isotope effect of less than unity would indicate a change in hybridization from sp in the ground state to sp in the transition state. Indeed, in addition to these results indicating the intermediacy of oxocarbonium ion-like transition states, kinetic isotope effects also supply further evidence for the covalent nature of the intermediate. The kinetic isotope effect on degalactosylation (kn/kD = 1.21) indicates a change from sp to sp hybridization on approaching the transition state and this can only occur if the intermediate is covalent in nature.  2.4  Acid Catalysis Koshland proposed that the departure of the aglycone may or may not require acid  catalysis. Evidence in the literature which supports this proposal includes the identification of a suitably positioned carboxylic acid residue (glu 35) which could function as an acid catalyst in HEW lysozyme.  Also, from the crystal structures of T4 lysozyme and GEW  lysozyme, residues glu 73 and glu 11 respectively, were observed to be in appropriate  10  O  A  positions to fulfil such a function. ' Evidence which indicates that acid catalysis is not essential for the hydrolysis of glycosides comes from studies of the P-galactosidasecatalysed hydrolysis of the (3-D-galactopyranosyl pyridinium salts. compounds, Sinnott et a l  1 4  observed rate enhancements of 10 to 10 8  13  With these for enzymatic  hydrolysis when compared to the spontaneous hydrolysis. Since it is structurally impossible to protonate these compounds in a manner which will assist aglycone departure, then none of this rate acceleration can be due to acid catalysis. Clearly however, in some cases acid catalysis will assist the reaction and an example which illustrates this is the enzyme-catalysed hydration of the octenitol derivative shown in Fig 9.  15  This study also  showed that, in accordance with the stereochemistry of the hydrated product, the proton was delivered from the P-face of the galactose ring.  Figure 9. Hydration of an octenitol derivative by E. coli fi-galactosidase.  2.5  Noncovalent Interactions Noncovalent interactions are believed to be responsible for the majority of the rate  enhancement that is observed in catalysis by these enzymes. These interactions in glycosidases are likely to be predominantly hydrogen bonds formed between the sugar hydroxyl groups and the enzyme. The binding energy derived from such interactions, when realized at the transition state, stabilizes that transition state and leads to a decrease in the activation energy of the reaction, thus a rate acceleration occurs (Appendix 1).  11 For example, noncovalent interactions must contribute greatly to the rate enhancement observed for the enzymatic hydrolysis of P-D-galactopyranosyl pyridinium salts since protonation cannot assist with the departure of the leaving group. In addition, 14  the transition state appears to be oxocarbonium ion-like in nature, suggesting that there is only limited assistance from the catalytic nucleophile. Studies with deoxy-glycosides have given some indication of how much the interactions of each individual hydroxyl group can contribute to the rate acceleration. For example, with the P-glucosidase A3 from A. wentii, removal of the hydroxyl group at C-2 results in a 10 fold rate decrease and removal of the C-4 hydroxyl group results in a 10 6  4  10 fold rate decrease. The C-2 hydroxyl group is also important in the E. coli lac z p*5  16  galactosidase-catalysed hydrolysis since a 10 fold decrease in rate is observed when it is 4  removed. The C-5 hydroxymethyl group in lysozyme substrates must also be providing 17  an important noncovalent interaction at the transition state, as a rate acceleration of 1300 fold was observed with the C-5 hydroxymethyl compound compared to the 6-deoxy derivative (Fig 10).  Figure 10. Substrate analogue for lysozyme.  12 The above examples demonstrate the importance of noncovalent interactions of the substrate hydroxyl groups with the enzyme in stabilizing the transition state, thus leading to catalysis.  3.  LINEAR F R E E E N E R G Y RELATIONSHIPS A linear free energy relationship is mathematically expressed as  log ( k 2 / k2) = log ( k i / k i ) • constant x  x  where k represents either the rate (k) or equilibrium constant (K) of a reaction and k that x  of the reaction in which there has been a change in the reaction conditions (for example, addition of a substituent). In other words, there is a linear free energy relationship between a set of effects if the same change on two different reactions (1 and 2) produces those same effects on either their rate or equilibrium. The term "free energy" in this relationship is appropriate as the rate is a reflection of the free energy of activation and the equilibrium constant reflects the standard free energy change of the reaction.  3.1  The Hammett Equation One of the most important examples of a linear free energy relationship is that  proposed by Hammett to explain the electronic effects of substituents on aromatic systems such as that represented in Fig 11. In these systems, X is a meta or para substituent, Y is  Figure 11. The general compound used in Hammett correlations.  13  the reaction site and Z is a side chain. Ortho substituents were omitted, as substitutions at this position can perturb the outcome by introducing steric interference into the reaction centre. The relationship that Hammett proposed is the following :  log(k /k) = po x  where k can be either a rate constant or an equilibrium constant and k the corresponding x  value for the substituted reaction. The reaction constant, p, is characteristic of a given reaction under a particular set of conditions (for example, temperature). It is a measure of the sensitivity of the reaction to the electronic nature of that substituent and is independent of the substituents. The substituent constant, a, depends only on the substituent and its position on the benzene ring. This parameter measures the extent of the effect that the substituent has on the reaction. As a standard, Hammett arbitrarily assigned a value of 1 to the reaction constant for the dissociation of benzoic acid in aqueous solution at 25°C and defined the substituent constant of hydrogen as zero. The linear free energy relationship between benzoic acid dissociation (1) and another reaction, for example benzoate ester hydrolysis (2) can be seen in the following. The Hammett relationship for benzoic acid dissociation is represented by equation 1 whereas that for benzoate ester hydrolysis is represented by equation 2.  log(K i/Ki) = G  (1)  log (k /k2) = po"  (2)  x  x2  The values K i and K i are the equilibrium constants for the unsubstituted and substituted x  benzoic acid dissociation respectively whereas k2 and k 2 are rate constants for the x  14 unsubstituted and substituted ester hydrolysis. The difference in standard free energy between the substituted and unsubstituted benzoic acid dissociation is shown in equation 3 while the difference in the activation free energy between the substituted and the unsubstituted benzoate ester hydrolysis is shown in equation 4.  A G = -2.303 RTlogK 0  AG° i - AG°i = -2.303 RTc x  (3)  AG* = -2.303 RTlogk AG*x2 - A G * = -2.303 RT op  (4)  AG* 2-AG*2 = ( A G i - A G ° i ) p  (5)  2  0  X  x  Substitution of equation 3 into equation 4 shows that the difference in the activation free energies of the ester hydrolyses is related to the difference in the standard free energies of the corresponding benzoic acid dissociations (equation 5). That is, the extent to which the activation free energy of benzoate ester hydrolysis is changed by a substituent is linearly related to the extent to which that substituent changes the standard free energy of dissociation of benzoic acid. Although the a value of hydrogen has been assigned as zero, the a values of other substituents can be calculated from the Hammett equation using the acid dissociation constants (pKa) of the appropriately substituted benzoic acid and they can be either negative or positive. By convention, positive a values are indicative of electron-withdrawing groups whereas negative values indicate electron-donating groups. In certain reactions, some substituents deviate from the predictions of the Hammett equation. These deviations can be due to an enhanced effect of the substituents that are in direct conjugation with the reaction site. To accommodate these deviations, several new series of a values have been introduced. These new series include a c~ series (a* by  15 Jaffe ) which correlates substituents in conjugation with a negative charge at the reaction 19  site and a o series which correlates substituents in conjugation with a positive charge at +  the reaction site. Similarly, a o ° series has been determined for substituents which are not in conjugation with the reactive site. In some cases the Hammett relationship is violated no matter which series is used. These deviations can often be explained as due to solvent interactions (for example ionization) or due to a change in reaction mechanism with that particular substituent (for example intramolecular catalysis). Such cases are studied individually. The effects of more than one substituent on a compound are often additive such that the Hammett equation can be expressed as the following.  log (k /k) = p2xj x  However, deviations from this additive effect are sometimes observed and are explained as being due to neighbouring group steric interactions or cross conjugation between the reaction site and the substituent. As previously mentioned, ortho substituents were not originally correlated with meta and para groups by Hammett. However, attempts have been made by many '  to  construct an ortho sigma scale. This scale however cannot be applied as widely and reliably as those for the meta and para substituents since the effect depends on the individual reaction. The ortho effect can arise both from steric effects, including steric hindrance at the reactive centre, steric hindrance to solvation and steric inhibition to resonance, and from intramolecular secondary bonding forces such as hydrogen bonding and charge transfer reactions. The value of p for the ionization of benzoic acid was assigned a value of 1. The values of p for other reactions however, have been calculated from the slopes of Hammett plots where log (k) or log (K) have been plotted as functions of the o values of the  16 substituents. Since the absolute magnitude of p is a measure of the sensitivity of the reaction to the electronic nature of the substituents, then it is an indicator of the degree of charge build-up at the transition state. Values of p can be either negative or positive. A negative p value indicates that the transition state is electron-deficient and therefore, substituents that are electron-donors will accelerate the reaction. Alternatively, a positive p value indicates an electron rich transition state, this reaction is accelerated by electronaccepting substituents. Additional information about a reaction can be obtained from the shape of the Hammett plot For example, a "concave upward" plot such as the one schematically shown in Fig 12 indicates that the reaction can proceed by two different paths. Such a plot suggests therefore, that one path is favoured by one set of substituents (for example  /  \  A  B  log (k)  sigma  Figure 12. Schematic representation of a concave upward Hammett plot for a reaction with two possible paths (1 &2).  17 A B +  • c  D  log (k)  sigma  Figure 13. Schematic representation of a concave downward Hammett plot for a multistep reaction electron-donors) whereas the other path is favoured by another set of substituents (for example electron-acceptors) so that the rate observed (k( b ) = k(i) + k(2)) is a reflection of 0  S  the path favoured by that substituent. However, a "concave downward" plot (Fig 13) indicates that there is more than one step in the reaction and that the rate determining step changes at some point as the substituents are altered. Therefore, although linear free energy relationships are only empirical relationships, they have been extensively applied in the study of structure-reactivity patterns of many reactions.  From such applications many features of a reaction mechanism can be  determined. In summary, the following are some of the features of a Hammett correlation which make it possible to obtain useful mechanistic information. (1)  The size and sign of p  (2)  The shape of the plot  (3)  The a series used (o~, o , o°) in the correlation +  18 3.2  Linear Free Energy Relationships in Enzymology Two important aspects of enzymatic reactions that limit the application of linear free  energy relationships are binding and catalysis. Enzymes are rather specific for their substrates such that a modification of the substrate could affect its binding to the enzyme. Also, the electronic effects of these substituents may be obscured by the resultant binding effects. The orientation of the substrate reaction centre with respect to the catalytic residue could be affected by substrate modification and this in turn could interfere with the catalytic step. Nonetheless, in favourable circumstances linear free energy relationships can be applied. Linear free energy relationships have been applied to the study of enzymes such as glycosidases , acyl transferases  and proteases.  The following two examples have  been selected to illustrate the type of results obtained from such studies and how these are interpreted to provide mechanistic insights. Papain, a cysteine protease, catalyses the hydrolysis of the amide bond. The mechanism of this reaction is proposed to involve the formation of an acyl-enzyme intermediate which is formed and degraded via a tetrahedral intermediate (Fig 14). O E-SH  - NH R  O  2  +  +  H 0 2  R E acyl enzyme intermediate  O  O +  S  R  Figure 14. Schematic representation ofpapain-catalysed hydrolysis.  19 The rates of enzyme-catalysed hydrolysis for a series of aryl N-acetyl-Lphenylalanylglycine anilides  24  (Fig 15) have been determined and the results presented as  Hammett plots (Fig 16). Clearly, the reaction rate is dependent on thereactivityof the  "  »  »  / ^ k  ffCH CNHCHCNHCH ? ?COH  X  CH3 C NHCH C N H C H C N H — ( f-*\) ^ 2  x  /  v  3  2  Papain -•  Figure 15. Papain-catalysed hydrolysis ofN-acetyl-L-phenylalanylglycine anilides.  Figure 16.  Hammett plot for papain-catalysed hydrolysis of N-acetyl-LOA  phenylalanylglycine anilides.  20 leaving group (aniline), thus the rate determining step must be the formation of the acylenzyme intermediate. A p value of - 1.04 was calculated from the slope of the Hammett plot. Such a p value indicates an accumulation of positive charge at the transition state which suggests that the nitrogen is protonated. Lowe et a l  24  have interpreted this data to  imply a mechanism whereby the tetrahedral intermediate formation is general base catalysed and the degradation is general acid catalysed. Structure-reactivity studies have been also carried out on other classes of enzymes, including glycosidases. The results of such studies on sweet almond p-glucosidase with 99  substituted aryl glucosides are summarized in Fig 17.  The shape of the Hammett plot is  concave downward suggesting that there is a change in the rate determining step as the leaving group changes. That is, for substrates with good leaving groups (pK less than 8) a  the departure of the leaving group is not rate-limiting, thereby suggesting that deglucosylation is the rate determining step. However, for substrates with poor leaving groups (pK greater than 8), the rate is dependent on the leaving group, suggesting that a  f  «  J  1  »  •  1  1  i t  ;  1  1—  • ««  PK.  Figure 17. Hammett plot for sweet almond p-glucosidase-catalysed hydrolysis of ary glucosides.  21 glucosylation is the rate determining step. These results, in addition to providing evidence for a two step mechanism, also identify the rate determining step of the reaction. The above examples have shown that the application of linear free energy relationships in enzymology can provide useful insights such as, the identification of rate determining steps, the existence of intermediates, the presence of general acid and base catalysis and overall charge at the substrate reaction centre at the transition state.  4.  IRREVERSIBLE INACTIVATORS Irreversible inactivators have been used to study the mechanisms of many enzymes.  These inactivators are compounds that resemble the substrates such that they are targeted towards the enzyme active site, but contain a reactive moiety. Once in the active site they can form covalent complexes with the enzyme, thereby inactivating it. In order to be characterized as irreversible inactivators which can be used to identify amino acid residues, these compounds must satisfy the following criteria. 1)  12,25  The inactivation should be time-dependent and should follow first order kinetics.  (2)  Saturation kinetics should be observed, that is, the inactivation rate should be dependent upon the inactivator concentration at low concentrations but be independent at high concentrations.  (3)  The rate of inactivation should be lower in the presence of a substrate or competitive reversible inhibitor than in its absence, thus showing that the inactivator binds at the active site.  (4)  The inactivated enzyme should remain inactive after being subjected to techniques such as dialysis or gel filtration which remove excess and loosely bound inactivator. This indicates that the inactivation is irreversible.  (5)  The stoichiometry of the inactivation should be one mole of inactivator per mole of active site.  22 (6)  The inactivate* should be reasonably stable against spontaneous decomposition under the reaction conditions.  (7)  The inactivator must have a high noncovalent affinity for the active site so that the active site is specifically labelled at low inactivator concentrations.  (8)  The enzyme-inactivator bond must be stable against denaturation, proteolysis and conditions used for sequence analysis so that the labelled amino acid can be identified.  There are two types of irreversible inactivators that will be considered, affinity labels (also called active-site-directed inactivators) and mechanism-based inactivators (also called suicide inhibitors or kcat inhibitors). Affinity labels are compounds which contain a reactive functional group such as the bromoacetyl group. However, due to the high intrinsic reactivities of these compounds, they can react with several different residues both in and out of the active site. For example, the bromoacetyl group readily reacts with residues such as cysteine, methionine and histidine. Thus, the enzyme may be labelled nonspecifically. Mechanism-based inactivators are characterized as having low intrinsic reactivity; they do not contain highly reactive functional groups. These inactivators are designed such that they are specifically activated at the active site, usually by the catalytic residues on the enzyme, converting them into reactive species. In order for an inactivator to be classified as mechanism-based, a catalytic step must be shown to be involved in the inactivation. Therefore, due to their inherent lower reactivity and the catalytic activation required, mechanism-based inactivators are more specific for the enzyme active site and hence they label more specifically than affinity labels.  4.1  Irreversible Inhibition Studies in Enzymology Irreversible inactivators have been used to probe the active sites of many enzymes  including glycosidases. To illustrate the type of information obtained, work using the  affinity labels, N-bromoacetyl-B-D-glucosylarnine (N-BrAc-Glu) and N-bromoacetyl-p-Dgalactosylamine (N-BrAc-Gal)  26,27  (Fig 18) in the study of E.coli B-galactosidase will be  •JO •IQ discussed. The work on the mechanism-based inactivator, conduritol B epoxide  ' (Fig  19), used in the investigation of P-glucosidase A3 from A. wentii will be presented. In an attempt to probe the two binding subsites (presumably a galactose subsite for the glycone and a glucose subsite for the aglycone) of P-galactosidase, the inactivators Nbromoacetyl-p-D-glucosylamine and N-bromoacetyl-p-D-galactosylamine were used to inactivate the enzyme. The resulting kinetic constants for inactivation are shown in Table 1. OH  OH  Figure 18. Structure of N-bromoacetyl-fi-D-glycosylamines.  Inactivator  Kj (mM)  kjtrnin" )  kj/Ki (min* mM" )  N-BrAc-Gal  1.13  0.063  0.055  N-BrAc-Glc  110  7  0.032  1  1  1  Table 1. Kinetic parameters for the inactivation of E. coli P-galactosidase with Nbromoacetyl -(5-D-glycosylamines.  These inactivators were both found to label methionine 502, presumably through nucleophilic displacement of the bromine by sulfur. Replacement of this methionine in the  24 protein with norleucine resulted in no loss of enzymatic activity, indicating that it is not essential for catalysis. However, this methionine is believed to be located near the active centre since site-directed mutagenesis of its neighbour, tyrosine 503, to phenylalanine resulted in a 10 fold decrease in the kcat.  These results, therefore implied that tyrosine  503 was direcdy involved at the active centre. Conduritol epoxides are compounds which react with enzymes in which there is an acidic group appropriately aligned to protonate the epoxide oxygen, followed by attack of a well placed nucleophile (Fig 19). The acid catalyst and the nucleophile are expected to be equivalent residues involved in the regular enzymatic catalysis.  Figure 19. Inactivation of A. wentii B-glucosidase with conduritol B cis-epoxide  Conduritol B cis-epoxide inactivates A. wentii P-glucosidase by covalent attachment to an aspartate residue. This inactivated enzyme, when treated with hydroxylamine, released the label as inositol from the active site, suggesting the existence of an ester linkage between a carboxyl group at the active site and the inactivator. The  25 released inositol was the product of a trans-opening of the epoxide such that the two new hydroxyl groups are in a transdiaxial configuration. This suggests that the epoxide is bound as shown in Fig 19 and that the epoxide is protonated in much the same way as the normal glycoside, with the "normal" nucleophile presumably then attacking the epoxide. Therefore, in summary, irreversible inactivators such as the ones discussed have been employed in the labelling of residues in the active site of enzymes in attempts to determine their roles in catalysis.  5.  THE CELLULASE COMPLEX The degradation of cellulose to glucose is accomplished by the concerted action of  three types of enzymes that make up the cellulase complex. These three enzymes, endoglucanases  (endo-l,4-P-glucanases,  E C 3.2.1.4), exo-l,4-P~glucanases  (cellobiohydrolases, 3.2.1.91), and (3-glucosidases (cellobiases, E C 3.2.1.21), are produced by microorganisms, plants and animals (in this case they are actually produced by symbiotic microorganisms). These enzymes can be secreted, as is the case for most fungal cellulases, or they can be organized in supramolecular structures called cellulosomes as is found in some bacterial cellulases.  These three enzymes are known to hydrolyse  cellulose synergistically, however the detailed mechanism of this synergism is not well understood. The function of the individual enzymes in the complex is the following (Fig 20): the endoglucanase randomly cleaves the cellulose into oligosaccharides, the exoglucanase then cleaves cellobiose from the non-reducing end of these oligosaccharides and finally the glucosidase hydrolyses the cellobiose into glucose. The gene encoding the exoglucanase from the bacterium, Cellulomonas fimi, has been cloned, expressed in E. coli and subsequently sequenced . Based on sequence homology studies, this enzyme has been placed in the F-family of glycanases.  This  26 family also includes xylanases, endoglucanases and other exoglucanases from other organisms. The molecular weights (determined by SDS PAGE) of the secreted enzyme and the cloned enzyme are 49.3 and 47.3 kDa, respectively.  The difference in weight is  o-oooooo-o-o-oo-o ^Endoglucanase  o-o-o o-ooo o-o-oo-o o-o o o o o o o o o o o o oo o |Exoglucanase  JGlucosidase  Figure 20. Schematic representation of the degradation of cellulose to glucose by the cellulose complex.  presumably due to glycosyl groups on the secreted enzyme, however the glycosylated and nonglycosylated enzymes have similar enzymatic activity. All the work done in this study was carried out on the nonglycosylated cloned enzyme. The C. fimi exoglucanase comprises two domains, a catalytic domain, and a binding domain that binds cellulose.  These domains can be separated by proteolysis,  resulting in a 33 kDa fragment (SDS PAGE) and a 14 kDa fragment (SDS PAGE). The 33 kDa fragment retains full catalytic activity with small substrates but cannot bind cellulose. The 14 kDa fragment does not have catalytic activity but it can still bind cellulose.  27 The substrate specificity of this exoglucanase was investigated and the enzyme was found not only to be active on carboxymethyl cellulose but also on xylan and paranitrophenyl p"-D-cellobioside. It had been previously reported not to exhibit glucosidase 34  activity on para-nitrophenyl P-D-glucoside, however a reinvestigation has revealed that the C.fimi exoglucanase is indeed active on substituted phenyl P-D-glucosides (as discussed 35  in Section 2). From stereochemical studies performed by use of ^ - N M R , the enzyme was found to hydrolyse substrates with retention of anomeric configuration.  Hence, the mechanism  of this exoglucanase is assumed to be the same as that for other retaining glycosidases.  28  6.  T H E A I M OF THIS STUDY The objective of this study is to investigate the mechanism of the C. fimi  exoglucanase-catalysed hydrolysis of ^-glycosides by carrying out linear free energy relationship studies and irreversible inactivation studies. The presence of linear free energy correlations between the substrate structure (aryl |3-glucosides and aryl P-cellobiosides) and reactivity (determined from the kinetic parameters) may provide information about the mechanism of enzymatic hydrolysis such as evidence for a two step mechanism (as has been proposed for this retaining enzyme), the identification of the rate determining step and the degree of charge accumulation at the transition state of that rate determining step. In addition, a comparison of the results for the glucosides and the cellobiosides may reveal any mechanistic function of the second glucosyl unit of the cellobiosides. Irreversible inactivation studies with the mechanism-based inactivators, 2', 4'dinitrophenyl 2-deoxy-2-fluoro-p -D-glucoside and 2", 4"-dinitrophenyl 2-deoxy-2-fluoro>  (J-D-cellobioside, can provide substantial evidence for the covalent nature of the glycosylenzyme intermediate proposed for the mechanism of this enzyme. Hence, inactivation with the tritiated analogue of 2', 4'-dinitrophenyl 2-deoxy-2-fluoro-p-D-glucoside would then ,  be expected to radiolabel the catalytic nucleophile proposed to be involved in the formation of this covalent intermediate, thus leading to its identification.  29 CHAPTER 2 RESULTS AND DISCUSSION  1.  HYDROLYSIS OF ARYL-p-D-GLUCOSIDES AND ARYL-p-DCELLOBIOSIDES BY C. FIMI  EXOGLUCANASE  Previous studies on the C.fimi exoglucanase had indicated that PNPG was not a 34  substrate. Reinvestigation of this has revealed that C. fimi exoglucanase does in fact hydrolyse aryl P-D-glucosides to the corresponding substituted phenol and glucose, in addition to catalysing the hydrolysis of aryl P-D-cellobiosides to the corresponding substituted phenol and cellobiose (Fig 17). The hydrolysis of these substrates was monitored spectrophotometrically by measuring the absorbance of the phenol released at the appropriate wavelength in 50 mM sodium phosphate buffer (pH 7.0) and 1 mg/ml BSA. The enzyme concentrations employed were those which give a sufficiently large absorbance change to ensure accurate calculation of the rates yet resulted in less than 10% conversion of the substrate to product during the observation period, thus ensuring linear kinetics. The maximum velocities (Vmax) and the apparent binding constants (K ) were m  determined for the two substrate series by fitting the initial rates and substrate concentrations to the Michaelis-Menten equation (Appendix 1) using weighted least squares regression. These results are illustrated as Lineweaver-Burk plots for visual convenience although the Michaelis-Menten kinetic parameters were not calculated from these plots due to the nonlinear error span associated with such double reciprocal analysis. This study has revealed that the exoglucanase is active on several aryl P-Dglucosides with rates ranging from 96 pmol min mg- to 0.18 pmol min- mg- (Table 2). 1  1  1  1  30  (a)  R=H HO  -  O  H  'OH Figure 21. Exoglucanase-catalysed hydrolysis of (a) arylfi-D-glucosidesand (b) aryl fiD-cellobiosides.  It is particularly noteworthy that the fastest of these, 2,4 DNPG, has a comparable V a x m  value to that found for aryl P-D-cellobiosides such as PNPC ( V a x = 84 u.mol min  -1  m  mg ). However, PNPG itself does indeed hydrolyse slowly, thus explaining the previous 1  failure to observe any activity. In order to determine if there was a relationship between substrate structure and enzymatic activity, a linear free energy relationship study was undertaken. In this study, log Vmax and log V  m a x  /K  m  were plotted as functions of the p K of the phenol of the a  leaving group. The Hammett plots obtained (Fig 22-25) provide valuable support for the double displacement mechanism proposed by Koshland for a retaining glycosidase (Chapter 1 ) involving a covalent glycosyl-enzyme intermediate. The log V  m a x  vs pKa  Hammett plot (correlation coefficient = 0.91) for the hydrolysis of the B—glucosides (Fig 22) is linear with slope of -1 (corresponding to a p value of 2.21*), thus showing a strong  * For substituted phenols, pKax = p K  a H  - 2.21a; p K ^ = 9.99.  31 correlation between the enzymatic rate and the p K of the leaving group. This result a  indicates that glycosylation is the rate determining step as this is the step in which the C-O bond to the phenol is cleaved. A similar correlation is seen in the plot of log Vmax/Km  Glvcoside  Km  Ymax  (mM)  (umol min" mg")  (umol min" mg" mM")  1  V-max/Km 1  1  2,4 DNPG  3.96  1.87  96  51.3  3,4 DNPG  5.36  6.52  23  3.5  2 CI, 4 NPG  5.45  27.46  78  2.8  PNPG  7.18  8.33  0.18  0.022  MNPG  8.39  11.6  0.013  0.0011  2,4 DNPC  3.96  0.11  69  627  3,4 DNPC  5.36  0.19  83  437  4 CI, 2 NPC  6.45  0.12  75  625  PNPC  7.18  0.6  84  140  3,5 DC1PC  8.19  0.92  51  55  MNPC  8.39  1.29  68  53  4CNPC  8.49  1.11  48  43  1  1  *Phenol pK values were taken from the following references: 49,50,51,52. a  Table 2. Michaelis-Menten parameters for the hydrolysis of the B-glucosides and Bcellobiosides by C.fimi exoglucanase.  vs pK (Fig 23) which has a slope of -1 (correlation coefficient = 0.99), corresponding to a a  p value of 2.21. This reveals that the first irreversible step in the reaction is glycosylation. This value of p reflects the fact that electron-withdrawing groups on the phenolate increase the reaction rate which is consistent with a large degree of charge build up at the  glycosylation transition state. A large charge development suggests therefore, that at the transition state there is either little protonation of the phenolate by the acid catalyst or that the C-0 bond cleavage is rather advanced. Similar studies have been done on other B-glycosidases employing aryl Pglucosides. In the case of the sweet almond P-glucosidase-catalysed hydrolysis of such aryl p-glucosides, a p value of 1.16 was calculated again showing that electron22  withdrawing substituents increase the reaction rate. This p value is less than the value calculated for the exoglucanase-catalysed hydrolysis of B—glucosides (p = 2.21) thus indicating that there is less charge separation at the transition state for the  33  E X CO  E >  o  -4.0 • J.u  •  •  3  •  1  4  •  I  l  5  6  7  "  •  8  "  9  pKa  Figure 23. The log V IK max  m  vs pK Hammett correlation for the hydrolysis of pa  glucosides by exoglucanase.  P-glucosidase-catalysed reaction. The slope of -1 indicates that there is full charge development at the glycosylation transition state for the exoglucanase-catalysed hydrolysis of these substrates, implying a greater amount of C-0 bond cleavage. The log V  m a  x / K vs pK Hammett plot for the (J-cellobiosides (Fig 25), like that m  a  for the substituted phenyl glucosides, shows that the first irreversible step for these substrates is also glycosylation. In contrast to the plot of log V m a x / K vs pK , the log m  V  m a  a  x vs pK plot (Fig 24) reveals no significant dependence of the rate on the pK^ of the a  leaving group and indicates that deglycosylation is likely the rate determining step. However, at the higher p K values there is a downward trend suggesting that the rate a  determining step is changing from deglycosylation to glycosylation. This change in rate determining steps has been reported for other glycosidases, for example the sweet almond  34 p-glucosidase-catalysed hydrolysis of aryl P-glucosides (Fig 17). These results provide 22  further support for the double displacement mechanism proposed by Koshland.  1.51.41.3-  3,4 DNPC •  1.2-  log(Vm  a  • 1.1 • 2,4 DNPC 1.00.90.80.73.5  Figure 24.  •  i  4.5  5.5  The log V  max  PNPC " MNPC 4C12NPC • 3,5 DC1PC • 4 CNPC m  6.5  7.5  pKa  vs pK  a  8.5  9.5  Hammett correlation for the hydrolysis of (3-  cellobiosides by C.fimi exoglucanase.  The slope of the plot of log V  m a x  /K  m  vs pK (Fig 25) for the P-cellobiosides is a  -0.28 (correlation coefficient = 0.82), corresponding to a p value of 0.60. This reaction constant, like those for the P-glucosides, reflects the fact that electron-withdrawing substituents on the leaving phenolate increase the reaction rate. The reaction constant for the hydrolysis of the glucosides is much greater than that for the cellobiosides, indicating a greater charge separation at the transition state for glycosylation with the glucosides than with the cellobiosides. This larger reaction constant for the glucosides may be an indication of a reaction mechanism in which there is less acid catalysis at the transition state for glycoside hydrolysis than for cellobioside hydrolysis since the degree of protonation will  35 affect  the extent of charge build up (Fig 26).  If however, both the  2.8-  2.3-  "E  4C1 2NPC 2,4 DNPC  X  jf  1.8-  3,4 DNPC  >  PNPC  o  ' 1.3-  0.8 3  4  •  I  5  6  T  7  3,5 DCIPC^ MNP™ 4 CNPC 8  9  pKa  Figure 25. The log Vmaxl^m Hammett correlation for the hydrolysis of B-cellobiosides by exoglucanase.  glucosides and the cellobiosides experience the same degree of acid catalysis at the glycosylation transition state, then the larger reaction constant for the glucosides would indicate greater C-O bond cleavage of the glucosides (i.e. a later transition state) than the cellobiosides (Fig 27). This study does not provide sufficient information to define the degree of acid catalysis and the degree of bond cleavage present at the transition states. However, simple a-deuterium kinetic isotope effect measurements on several of these glucosides and cellobiosides could provide supporting evidence for either one or the other of the above explanations. If similar a-deuterium kinetic isotope effects were measured for both the glucosides and the cellobiosides, this would indicate that there was approximately the same  36  X  _ J  Figure 26. Varied degrees of acid catalysis at the glycosylation transition state for glucosides and cellobiosides.  amount of sp /sp hybridization at the transition states and thus the same degree of 2  3  oxocarbonium ion character. This suggests that the same amount of bond cleavage occurs at the transition states for the glucosides and the cellobiosides which would be more consistent with the larger reaction constant of the glucosides resulting from less acid catalysis than the cellobiosides. However, if a larger a-deuterium kinetic isotope effect is measured for the glucosides than for the cellobiosides, it would suggest that there was greater bond cleavage at the transition state for the glucosides than for the cellobiosides. In  37 this case, the kinetic isotope effects would be more consistent with the larger reaction constant of the glucosides resulting from a later transition state than that of the cellobiosides.  Figure 27. Different degrees ofC-0 bond cleavage at the glycosylation transition state for the glucosides and the cellobiosides.  As previously discussed, the log V  m a  x vs p K Hammett correlation for the a  cellobiosides shows deglycosylation to be the rate limiting step with a rate of about 84 pmol min" mg" . Interestingly, the rate of hydrolysis of the fastest glucoside, 2,4 DNPG, 1  1  (96 pmol min" mg" ) is comparable to the rate of deglycosylation of the cellobiosides. A 1  1  minimum estimate of the rate of deglycosylation for the glucosides can therefore be  38 obtainedfromthe rate of hydrolysis of 2,4 DNPG. Thus, the rates of deglycosylation for the glucosides and the cellobiosides are either approximately the same or the deglycosylation rate for the glucosides is greater than that for the cellobiosides. This assumption is supported by the reactivation data (to be discussed later) which shows that samples of the exoglucanase inactivated with 2F-DNPG and 2F-DNPC reactivate at comparable rates. However, in contrast to these similar rates of deglycosylation for the glucosides and the cellobiosides, the rates of glycosylation for all the cellobiosides are greater than those for the corresponding glucosides. Thus, the presence of the second sugar unit in the cellobiosides increases the glycosylation rate, but not the deglycosylation rate. A possible free energy diagram which illustrates this situation is shown in Fig 28, the energy levels being arbitrarily chosen. In this diagram it is assumed that the transition states for glycosylation and deglycosylation are similar in energy and that the energy difference between the glycosylation transition states and the deglycosylation transition states for the cellobiosides and the glucosides is approximately the same. According to the Hammond postulate, which states that the structure of an unstable intermediate in a reaction path will more resemble the structure of the transition state than the ground state, then as a first approximation it can be assumed that the energy difference between the glucosylenzyme intermediate and the cellobiosyl-enzyme intermediate will be approximately the same as the energy difference between the two glycosylation transition states. The result of this would be that the rates of deglycosylation for the cellobiosides and the glucosides would be approximately the same. The energy difference between the cellobioside-enzyme complex and the glucoside-enzyme complex is less than the energy difference between the cellobiosyl-enzyme intermediate and the glucosyl-enzyme intermediate, thus the rates of glycosylation for the cellobiosides will be greater than those for the glucosides. This would seem to be quite reasonable. Furthermore, this diagram is consistent with the much larger p value for the glucosides than that for the cellobiosides resulting from a later transition state.  39  E + S "  w  ES  •  ES*  • EP  • EP*  ES*  P  EP*  AG*c(d)  A G 1(g)"  AG  *-E +  n  EP E+ S I A G  I  t  ES  ES  A G  ES  Reaction coordinate  Figure 28. Energy diagram showing the stabilization produced by the second glucosyl un of the cellobiosides (dashed line - glucosides, solid line = cellobiosides).  40 2.  I N A C T I V A T I O N O F C.FIMI  EXOGLUCANASE  As previously discussed, the mechanism-based inactivator, 2\ 4'-dinitrophenyl 2deoxy-2-fluoro-p-D-glucoside, has been observed to inactivate the Agrobacterium |5glucosidase by formation of a covalent 2-deoxy-2-fluoro-glucosyl-enzyme intermediate. 5  Similar inactivation studies have been undertaken with the C.fimi exoglucanase and the results will be presented and discussed below.  2.1  Kinetics of inactivation The inactivators used in this study were 2', 4'-dinitrophenyl 2-deoxy-2-fluoro-|J-D-  glucoside (2F-DNPG) as the enzyme is now known to have glucosidase activity and 2", 4"-dinitrophenyl 2-deoxy-2-fluoro-P-D-cellobioside  (2F-DNPC) which is closer in  structure to the regular substrates of the enzyme. The rates of inactivation by 2F-DNPG were assayed with both PNPC and 2,4 DNPG and found to be identical, thereby suggesting that both substrates bind to a single active site. The inactivation in both cases wastime-dependentand exhibited saturation kinetics. Control experiments performed by incubating the enzyme in the absence of either inactivator showed that it retained full activity over the inactivation period, thus all inactivation observed was associated with the presence of the inactivator. The inactivation kinetic parameters listed in Table 3, Ki (the inactivation binding constant) and kj (the rate constant of inactivation) were determined by first calculating the kobs (pseudo-first order rate constant of inactivation) from the slopes of plots of ln(residual activity) vs time (Appendix 2) and then fitting the kobs ° d inactivator concentrations to a a  variation of the Michaelis-Menten equation using the Curvefitter weighted nonlinear regression program. However, for convenient visual inspection, the kobs inactivator vs  concentration is presented plotted according to Lineweaver and Burk (Appendix 2). The value of Kj for 2F-DNPG, 4.5 mM, was in the range observed for aryl |3-glucosides and the K, value for 2F-DNPC, 0.11 mM, was similarly found to be in the range for aryl 3 |—  41 cellobiosides. Since these compounds inactivate the enzyme through stabilization of their glycosyl-enzyme intermediates (to be discussed), then the inactivation rate constants measured, 8.0 x 10 min- (t(i/2) = 87 min) and 2.5 x 10" min* (t/\p) = 2772 min) for 3  1  4  1  2F-DNPC and 2F-DNPG respectively, reflect the rates of glycosylation of the enzyme. In addition, the greater rate measured for 2F-DNPC inactivation of the exoglucanase compared to that for 2F-DNPG is consistent with the fact that the rate of glycosylation for the parent cellobioside (2,4 DNPC) is greater than that for the corresponding glucoside (2,4 DNPG).  This indicates that the presence of the fluorine at C-2 alters the rate of  glycosylation for the cellobioside and the glucoside in a similar manner.  Inactivator  Ki (mM)  lq (min )  2F-DNPG  4.5  2.5 x lO"  2F-DNPC  0.11  8.0x10-3  -1  4  Table 3. Kinetic parameters for the inactivation of C.fimi exoglucanase by 2FDNPG and 2F-DNPC  2.2  Covalent Inactivation The Agrobacterium (3-glucosidase has previously been inactivated with 2F-DNPG  by formation of a covalent bond between the inactivator and the enzyme. The covalency and the stereochemistry of the trapped covalent glycosyl-a-glucosidase have been characterized by NMR experiments and the carboxylate to which the inactivator is bound has been identified as glutamate 358. 5  Similar inactivation studies had been done on the C.fimi exoglucanase where samples of the enzyme were separately inactivated with 2F-DNPG and 2F-DNPC and then the excess inactivators were extensively dialysed away from the enzyme. In both cases the enzyme remained inactive after dialysis providing evidence for the covalent nature of this  5  42 inactivation. The proposed mechanism of inactivation for both inactivators (Fig 29) is the same in which the enzyme is trapped as the covalent glycosyl-a-exoglucanase intermediate.  Figure 29. Proposed mechanism of inactivation of C.fimi exoglucanase Jjy (a)2F-DNPG and(b)2F-DNPC.  2.3 Reactivation of Inactivated C. fimi Exoglucanase Samples of the exoglucanase which have been inactivated with 2F-DNPG and 2FDNPC were assayed (see Materials and Methods) for return of enzymatic activity and the reactivation rates were calculated (Table 4). The reactivation rate for the 2F-DNPGinactivated exoglucanase in buffer was 1.3 x 10' min (t(i/2) = 880 hours) and that for the 5  -1  2F-DNPC-inactivated exoglucanase was 8.5 x 10 min" (t(i/2) = 1340 hours), showing -6  1  that these inactivated-enzyme species are relatively stable in buffer. However, in the presence of an appropriate glycosyl acceptor, the exoglucanase slowly reactivated. For example, the 2F-DNPG-inactivated exoglucanase reactivated in the presence of glucose, (}D-glucopyranosyl benzene and cellobiose (Appendix 2), the fastest reactivation being in the presence of cellobiose (Table 4), as expected since the enzyme is known to have a greater affinity for oligosaccharides. The reactivation of the 2F-DNPC-inactivated enzyme was also accelerated in the presence of cellobiose.  Reactivator  Rates of reactivation for 2F- Rates of reactivation for 2Fglucosyl-enzyme  cellobiosyl-enzyme  Buffer  1.3 x lO' min-  1  8.5 x lO^min-  Glucose (55 mM)  1.6 x lO^min  1  ND*  P-D-Glucopyranosyl  1.8 x lO-Smhr  1  5  1  ND*  benzene (55 mM) 4.4 x 10" min-  Cellobiose (55 mM)  5  1  1.9 x 10" min5  1  * Not determined Table 4. The rates of reactivation of inactivated exoglucanase in the presence of a gtycosyl-acceptor.  The reactivation rate of the 2F-DNPG-inactivated exoglucanase in the presence of cellobiose is 4.4 x 10" min* and that for the 2F-DNPC-inactivated exoglucanase is 1.9 x 5  1  44 10 rnirr . These rates are quite comparable, thus indicating that the second glucosyl unit -5  1  of the 2F-cellobiosyl moiety does not contribute to the rate of reactivation. This suggests therefore, that this glucosyl unit is not important for deglycosylation which is consistent with the results for the hydrolysis of the aryl |3-glucosides and the aryl ^-cellobiosides. This reactivation is assumed to occur by transglycosylation, as had previously been observed for the reactivation of the Agrobacterium 2F-glucosyl-|3-glucosidase by |$-Dglucopyranosyl benzene (previously discussed). In this case, transglycosylation is assumed to involve the nucleophilic attack by the C-4 hydroxyl of the glycosyl-acceptor onto the anomeric centre of the 2F-glycosyl-enzyme (Fig 30). The reactivation of the exoglucanase in this manner is therefore consistent with the proposed formation of a glycosyl-enzyme  RO HO OH —O OH  REACTIVATION .OH  AH  R=H  Figure 30. The proposed mechanism for the reactivation of inactivated-exoglucanase by cellobiose.  intermediate and that intermediate's ability to turn over to product in the presence of an appropriate glycosyl-acceptor as outlined in Fig 30.  3.  IDENTIFICATION O F T H E C A T A L Y T I C N U C L E O P H I L E O F C. FIMI E X O G L U C A N A S E The compounds 2F-DNPG and 2F-DNPC are mechanism-based inactivators and  therefore, by definition should inactivate the exoglucanase through the involvement of the catalytic residues. Also, it is observed that the inactivated enzyme, presumably the covalent 2F-glycosyl-enzyme intermediate formed, is very stable. This intermediate is assumed to be formed between the catalytic nucleophile and the inactivator. These features suggest therefore that the catalytic nucleophile of C.fimi exoglucanase could be identified with such inactivators. The following then, is a discussion of the labelling and identification of the active site nucleophile of the exoglucanase with the tritiated analogue of 2F-DNPG ({13H} 2F-DNPG) The exoglucanase was incubated in the presence of {1- H) 2F-DNPG until 80% of 3  the enzymatic activity had been lost. Activity checks of the enzyme in the absence of inactivator showed that the enzyme retained full activity over this inactivation period. The inactivation process was also monitored by SDS-PAGE which showed that the exoglucanase (47 kDa) had been cleaved to a 33 kDa peptide fragment in both the absence and presence of inactivator. This 33 kDa fragment has been identified previously as the catalytic domain of the exoglucanase which retains activity on small substrates.  33  The  other fragment from the cleavage (14 kDa) corresponds to the cellulose binding domain and the linker region between the two domains (PT box). This proteolytic cleavage of the intact enzyme was the result of a contaminating endogenous protease of C.fimi  30  and has been  observed previously in prolonged incubation. This proteolysis proved, however, to simplify the subsequent steps of digestion and peptide purification.  46 Following the inactivation of the enzyme, it was extensively dialysed against buffer and concentrated resulting in the removal of excess inactivator and of the 14 kDa fragment. The proteolysis of the 33 kDa fragment was accomplished by 1:100 pepsin digestion and the peptide fragments were separated by reverse phase HPLC as described in Materials and Methods. The HPLC purification yielded two radioactive peptide fragments (A+B) of sufficient purity for sequencing. The peptides, when sequenced by Edman degradation, both yielded the amino acid sequence, val-arg-ile-thr-glu-leu, showing that both peptides clearly arise from the same region of the protein. However, these two peptides are in fact different as they were observed to have different retention times on the HPLC columns. This difference must presumably be on the carboxyl terminal side of leucine and therefore, must be due to incomplete digestion by pepsin. The glutamate residue was identified as the labelled residue in the sequence because the derivatized glutamic acid (PTH-glu) intensity was drastically decreased compared to the intensity of the other derivatized residues in the sequence. This decrease in intensity of the PTH-glu is expected if the glutamate is modified to the 2-fluoro-glucosyl ester, a compound known to be stable under Edman degradation conditions. These sequenced 5  peptides overlapped with the known amino acid sequence of the enzyme such that the labelled glutamate corresponds to Glu-274 of the proenzyme.  31  Therefore, Glu-274 is  proposed to be the nucleophilic carboxylate of C.fimi exoglucanase.  3.1  Conservation of Glu-274 of C. fimi exoglucanase in other  glycosidases The C.fimi exoglucanase belongs to the F family of glycanases (based on sequence homology) which comprise eleven enzymes including other exoglucanases, endoglucanases and xylanases. In this family, this ile-thr-glu-leu (asp) motif is conserved in all these enzymes with the exception of one xylanase. It is interesting to note that  although this motif does not occur at the same position in the actual amino acid sequence, it does occur in the catalytic domain at about the same position in all cases. The conservation of residues in a family of proteins usually indicates that those residues are important in maintaining enzymatic activity. Therefore, this motif is expected to have an important function and it is probable that the conserved glutamate is the catalytic nucleophile for all the enzymes of this family. It is therefore reasonable to assume that the enzymes of this family (with the possible exception of the xylanase) all hydrolyse their substrates with retention of anomeric configuration, thus are retaining enzymes. This conserved motif of the F family of P-glycanases has considerable similarity to a conserved region of another family of P-glycosidases comprising glucosidases and galactosidases. The conserved sequence in this other family is tyr-ile-thr-glu-asn-gly. The glutamate in this sequence has also been identified as the catalytic residue in one enzyme of this family, the Agrobacterium P-glucosidase, which is a retaining enzyme. Therefore, it would not be unexpected if the glutamate is determined to be the catalytic nucleophile in the rest of the enzymes and if these enzymes all perform hydrolysis with retention of configuration. However, it is not suggested that all retaining glycosidases must contain similar amino acid motifs in their active sites since the Trichoderma reesei cellobiohydrolase I which is a retaining enzyme of the C family does not have any such amino acid motif in 32  its active site.  48  4.  CONCLUSION This investigation of the C.fimi exoglucanase-catalysed hydrolysis of {^-glycosides  has revealed many significant features about the substrate specificity and the mechanism of hydrolysis. The results show that the enzyme is capable of hydrolysing ^-glucosides (previously undetected) with rates ranging from 96 u.mol min mg* to 0.013 umol rrjur -1  1  1  mg , where the fastest rate measured is comparable to rates measured for the fastest $ |— -1  cellobiosides (for example PNPC). The Hammett correlations of this glucosidase activity with pK of the leaving group revealed glycosylation as the rate determining step and the a  first irreversible step in the reaction. The reaction constant (p = 2.21) reflects a significant amount of charge build-up at the transition state for glycosylation. The results of a similar investigation on the cellobiohydrolase activity of this enzyme showed glycosylation to be the first irreversible step for the cellobiosides and the rate determining step to be deglycosylation which appears to be changing to glycosylation at the higher p K values. a  The reaction constant calculated from the plot of log V  m a x  vs pK for these cellobiosides a  (p = 0.60), like that for the glucosides (p = 2.21), reflects charge build up at the transition state of glycosylation, although to a lesser extent. This smaller reaction constant may be due to either a greater extent of acid catalysis at the transition state or to a much earlier transition state for the cellobiosides. These two situations cannot be distinguished with the data from this study, but this may be accomplished by measuring a-deuterium kinetic isotope effects for both cellobiosides and glucosides. The inactivation studies showed the enzyme to be covalendy inactivated in the presence of the mechanism-based inactivators, 2F-DNPG and 2F-DNPC, with dissociation constants (K\) consistent with the enzyme's greater affinity for oligosaccharides. The rate constants measured showed 2F-DNPC to be a faster inactivator of the enzyme than 2FDNPG. The inactivated enzyme is stable in buffer, but is reactivated in the presence of  glycosyl-acceptors such as glucose, P-glucosyl-benzene and cellobiose. This reactivation occurs presumably via a transglycosylation reaction, thereby indicating the catalytic competence of covalent glycosyl-exoglucanase intermediates. The rates of reactivation of the 2F-DNPG and the 2F-DNPC-inactivated enzyme in the presence of cellobiose were comparable. The similar rates of reactivation of the 2F-DNPG- and 2F-DNPC-inactivated enzyme and the greater rate of inactivation by 2F-DNPC demonstrate that the second glucosyl unit of the cellobioside selectively increases the rate of glycosylation relative to deglycosylation. This is consistent with the results of the Hammett plots with aryl Pglycosides where the rates of glycosylation of the cellobiosides are greater than those for the corresponding glucosides and where the minimum estimated rate of deglycosylation of the glucosides is similar to the rate of deglycosylation of the cellobiosides. The exoglucanase was radiolabelled with the tritiated analogue of 2F-DNPG, allowing the identification of the catalytic nucleophile as glutamate 274. This is consistent with Koshland's mechanism which proposes the involvement of a carboxylate in the hydrolysis of glycosides. This glutamate is part of a highly conserved sequence in the Pglycanases of family F, thus suggesting that this glutamate in these enzymes is likely the catalytic nucleophile.  50 CHAPTER 3 MATERIALS AND METHODS 1.  SYNTHESIS  1.1  General Procedures and Materials Melting points (m.p.) were determined using a Laboratory Devices Mel-temp II  melting-point apparatus, and are uncorrected. Proton nuclear magnetic resonance ("HNMR) spectra were recorded either on a 400 MHz Bruker instrument or on a 300 MHz Varian XL-300 instrument. Chemical shifts are listed in the delta (8) scale. Compounds dissolved in CDCI3 or C D 3 O D are referenced against the internal standard tetramethylsilane (TMS, 8 = 0.00 ppm). F-NMR spectra were recorded either on a 200 MHz Bruker AC19  200 spectrophotometer or on a 300 MHz Varian XL-300 instrument. Chemical shifts values are reported relative to CFCI3 (8 = 0.00 ppm) and were referenced either against external trifluoroacetic acid (8 = 76.53 ppm) or hexafluorobenzene (8 = 162.9 ppm). Micro-analyses were performed by Mr. P. Borda, Microanalytical laboratory, University of British Columbia, Vancouver. Solvents and reagents used were either reagent grade, certified or spectral grade. Solvents were dried as follows: methanol was distilled from magnesium methoxide prepared in situ by reaction of methanol with magnesium turnings in the presence of iodine; dimethylformamide was stirred over MgS04 for several hours and distilled under reduced pressure.  Acetyl chloride was dried by refluxing over PCI5. followed by  distillation. Thin layer-chromatography (tic) was carried out on Merck Kieselgel 60 F-254 plates. Acetylated compounds were run either in ethyl acetate/petroleum ether (1:1) or diethyl ether/chloroform (2:1) solvent mixtures whereas deprotected compounds were run in mixtures of methanol/ethyl acetate (3:2). Compounds were detected visually by employing U.V. light and/or by charring with 5% sulfuric acid in methanol. The method on  of Still et al  was used to carry out column chromatography using a silica gel column of  Kiesegel 60 (180-230 mesh).  51 The following compounds were prepared by other members of this laboratory: 2\4'-dinitrophenyl p-D-glucopyranoside by Adam Becalski; 3',4'-dinitrophenyl p*-Dglucopyranoside, 3'-nitrophenyl P-D-glucopyranoside and 2',4'-dinitrophenyl 2-deoxy-2fluoro-{l-^H}-P-D-glucopyranoside fluoro-P-D-glucopyranoside  by Julie Kempton; T^'-dinitrophenyl 2-deoxy-2-  by Mark Namchuk; 2'-chloro-4'-nitrophenyl P-D-  glucopyranoside by Marc Claeyssens.  The following compounds were prepared in  collaboration with Carola Ibe: 4"-chloro-2"-nitrophenyl P-D-cellobioside, 3"-nitrophenyl P-D-cellobioside and 3",5"-dichlorophenyl P-D-cellobioside.  1.2  General compounds  1,2,3,6,2 ',3',4',6'-Octa-0-acetyl-a-D-cellobiose (1) This compound was prepared by the method of Wolfrom and Thompson.  40  Cellobiose (5.3 g, 14.5 mmol) was added to a stirring mixture of acetic anhydride (40 mL) and pyridine (70 mL) at 0° C. After three days the reaction mixture was poured over icewater (200 mL) which precipitated the product. The product was collected by vacuum filtration and recrystallized from ethanol to give white needle-like crystals (5.02 g, 7.40 mmol; 50%). M.p. 197-198 °C (lit  40  m.p. 202-202.5 °C).  2 J,62 ',3\4',6'-Hepta-0-acetyl- a-D-cellobiosyl bromide (acetobromocellobiose) (2) This compound was synthesized by the Fischer and Zemplen method with the 41  following modifications. The peracetate, (1) (10 g, 14.75 mmol) was dissolved in glacial acetic acid (385 mL) and 45% HBr/glacial acetic acid (15 mL). The reaction proceeded to completion at 4 °C overnight. The reaction mixture was poured over ice-water and the product was extracted with chloroform. The chloroform layer was washed (3 x 20 mL) with saturated aqueous sodium bicarbonate and dried over MgS04. The MgS04 was removed by gravity filtration and the chloroform was evaporated in vacuo to leave an orange oil. The product, (2), was recrystallized from chloroform and diethyl ether to give  52 small white needle-like crystals (8.10 g, 11.59 mmol, 79%). M.p. 169-170 °C ( l i t  41  m.p. 180 °C).  1.3  Aryl  2,3,6,2\3\4\6'-hepta-0-acetyl  P-D-cellobiosides  The cellobiosides, with the exception of 2",4" dinitrophenyl P-D-cellobioside, were prepared according to the method of Koenigs-Knorr  4  2  To acetobromocellobiose (2)  dissolved in acetone (~ 0.4 mmol acetobromocellobiose/mL), the appropriate substituted phenol dissolved in 1M NaOH (1 mmol phenol/mL base) was added. The reaction mixture was stirred at room temperature for 24-48 hours. The solvent was then evaporated in vacuo leaving behind a syrup which was diluted with water and extracted with chloroform. The organic phase was washed (3 x 50 mL) with saturated sodium bicarbonate, dried over MgS04, filtered and evaporated in vacuo. The products were then crystallized with the appropriate solvent mixtures.  1.4  Aryl p-D-cellobiosides The aryl P-D-cellobiosides were prepared by one of the following deacetylation  procedures; (1) HCl/MeOH or (2) NaOMe/MeOH. 43  44  The HCl/MeOH method was used  to deacetylate cellobiosides with activated leaving groups (pKa < 6) whereas the NaOMe/MeOH method was used to deacetylate cellobiosides with leaving groups with pKa > 6.  1.4.1  HCl/MeOH Method The acetylated cellobioside was suspended in methanol (16 mg/mL), cooled to 0 °C  and acetyl chloride added to generate a final HC1 concentration of about 4%. The reaction was allowed to proceed at 4 °C for 16-24 hours until completion. The solvent was removed in vacuo, and the product washed (5x10 mL) with anhydrous diethyl ether to remove excess acid. The product was recrystallized from appropriate solvent mixtures.  53  1.4.2  NaOMe/MeOH Method The protected cellobioside was dissolved in methanol (~ 3 mg/mL) and sodium  methoxide in methanol (0.22 g sodium metal in 10 mL methanol) was added to make a final sodium methoxide concentration of 0.1 M. The reaction was stirred at room temperature until completion (30-60 minutes) and then the excess base neutralized with Amberlite 1R120 (H) cation exchange resin. The resin was removed by gravity filtration and washed several times with methanol, the solvent was evaporated in vacuo and the product crystallized from the appropriate solvents.  3",4"-Dinitrophenyl 2,3,6,2',3',4',6'-hepta-0-acetyl B-D-cellobioside (3)  To acetobromocellobiose (500 mg, 0.72 mmol) dissolved in acetone, 3,4 dinitrophenol.(250 mg, 1.36 mmol) was added. The reaction proceeded for 48 hours to completion at room temperature over K2CO3. The product was recrystallized from ethyl acetate and ethanol as a pale yellow solid (85 mg, 0.11 mmol, 15 %). M.p. 239-240 °C (lit m.p. 240-242 ° C ) , *H NMR (CDCI3): 5 8.00 (d, J », " 9 Hz, H(5")), 7.40 (d, 1 \6" 4 45  5  Hz, H(2")), 7.25 (m, H(6")), 3.65-5.35  6  2  (m, H(l-6) and H(r-6')), 2.00-2.15 (7 s,  7(OAc)). Elemental analysis for C32H38N2O22 ; calculated: C, 47.90%; H, 4.74%; N, 3.49%. Found: C, 47.92%; H, 4.87%; N, 3.53%.  3",4"-Dinitrophenyl R-D-cellobioside (4)  This compound was prepared from the acetylated compound (3) (80 mg, 0.10 mmol) via the HCl/MeOH method. The product was recrystallized from methanol and diethyl ether as a pale yellow solid (20 mg, 0.04 mmol, 40%). M.p. 198-199 °C (lit m.p. 187-192 ° C ) , *H NMR (CD3OD): 8 8.15 (d, J «, » 11 Hz, H(5")), 7.65 (d, J ", -11 Hz, 45  5  6  5  6  H(5")), 7.65 (d, J ",6 2 Hz,H(2")), 7.47 (dd, J «, " 2 Hz, J " " H Hz, H(6")), 5.20 (d, B  2  Jl 9 > 2  2  6  6  5  Hz, H(l)), 3.20-5.20 (m, H(2-6) and H(l'-6')). Elemental analysis for  54 C32H38N2O22 ; calculated: C, 42.50%; H , 4.72%; N, 5.51%. Found: C, 42.27%; H , 4.91%; N, 5.42%.  4"-Chloro-2"-nitrophenyl 2,3,6,2'j',4',6'-hepta-0-acetyl p-D-cellobioside (5) The acetylated compound (5) was prepared from acetobromocellobiose (1.5 g, 2.15 mmol) and 4-chloro-2-nitrophenol (798 mg, 4.6 mmol) according to the general method (Section 1.3). The product was recrystallized from ethyl acetate and ethanol as a white solid (350 mg, 0.44 mmol, 20%). M.p. 213-214 °C. "H NMR (CDCI3): 8 7.80 (d, J3"^" 1 Hz, H(3")), 7.50 (dd, J ", " J5\6" 1 Hz, H(5")), 7.25 (d, J «, « 3 Hz, H(6")), 3.65-5.03 5  3  6  5  (m, H(l-6) and H(l'-6')), 2.00-2.15 (7 s, 7 OAc). Elemental analysis for C32H38O20CIN; calculated: C, 48.51%; H, 4.80%; N, 1.76%. Found: C, 48.67%; H, 4.88%; N, 1.70%.  4"-Chloro-2"-nitrophenyl B-D-cellobioside (6) The protected compound (5) (330 mg, 0.42 mmol) was deacetylated by the NaOMe/MeOH method. The product (6) was recrystallized from methanol and diethyl ether to yield a white solid (41 mg, 0.084 mmol, 21%). M.p. 155-158 °C. *H NMR ( C D 3 O D ) : 8 8.00 (d, J -,5"2 Hz, H(3")), 7.65 (dd, J ",5"2 Hz, J «, » 11 Hz, H(5")), 3  3  5  6  7.40 (d, J ",5" 11 Hz, H(6")), 5.25 (d, J r , " 9 Hz, H(l)), 3.25-4.90 (m, H(2-6) and 6  2  H(l'-6')). Elemental analysis for C18H24O13NCI; calculated: C, 43.41%; H , 4.82%; N, 2.81%. Found: C, 43.26%; H, 4.98%; N, 2.74%.  3"y-Dichlorophenyl2,3,6X,3',4',6'-hepta-0-acetyl-p-D-cellobioside (7) The protected glycoside (7) was synthesized from acetobromocellobiose (1.4 g, 2.06 mmol) and 3,5-dichlorophenol (650 mg, 4.0 mmol) according to the general method (Section 1.3) and was recrystallized from ethanol to yield white crystals (175 mg, 0.22 mmol, 11%). M.p. 219 - 221 °C. "H NMR (CDCI3): 8 7.01 (m, H(4")), 6.90 (d, J 2 Hz, H(2",6")), 3.53-5.03 (m, H(l-6) and H(l'-6')), 2.00-2.15 (7 s, 7(OAc)). Elemental  55 analysis for C32H37O18CI2; calculated: C, 49.20%; H, 4.74%; Found: C, 49.31%; H , 4.88%.  3"j"-Dichlorophenylfi-D-cellobioside(8) The acetylated compound (7) (160 mg, 0.21 mmol) was deprotected using NaOMe/MeOH method to give the desired product (8) which recrystallized from methanol, diethyl ether and hexane as a white solid (33 mg, 0.067 mmol, 33%). M.p. 250-253 °C. -H NMR (CD3OD): 8 7.20 (m, H(4")), 7.10 (d, J 2 Hz, H(2",6")), 5.11, (d, Ju HQ)),  10 Hz,  3.27-4.97 (m, H(2-6) and H(l'-6')). Elemental analysis for C18H24O11CI2,  calculated: C, 44.35%; H, 4.92%. Found: C, 44.18%; H, 5.08%.  3"-Nitrophenyl 2,3,6,2\3',4',6'-hepta-0-acetyl-P-D-cellobioside (9) The protected cellobioside (9) was prepared according to the method of KoenigsKnorr from acetobromocellobiose (1.5 g, 2.21 mmol) and 3-nitrophenol (625 mg, 4.50 mmol) and recrystallized from ethanol to give a white solid (581 mg, 0.77 mmol, 35%). M.p. 160-163 °C, "H NMR (CDCI3): 8 7.90 (m, H(2",4")), 7.45 (m, H(5",6")), 5.70 (d, J  1 > 2  , 8 Hz, H(l)), 3.65-5.30 (m, H(l-6) and H(l'-6')), 2.00-2.15 (7 s, (7 OAc)).  Elemental analysis for C32H39O20N; calculated: C, 50.73%; H, 5.15%, N; 1.85%; Found: C, 50.65%; H , 5.34%; N, 1.50%.  3 "-Nitrophenylfi-D-cellobioside(10) The protected cellobioside (9), (250 mg, 0.33 mmol) was deacetylated using the NaOMe/MeOH method to yield the product (10) which recrystallized from methanol, diethyl ether and hexane to give a white solid (17 mg, 0.037 mmol, 10%). M.p. 220-222 °C. -H NMR (D 0): 8 8.00 (m, H(2",4")), 7.55 (m, H(5",6")), 5.27 (d, J^9 2  Hz, H(l)),  3.30-5.00 (m, H(2-6) and H(l'-6')). Elemental analysis for C18H25O13N, calculated: C, 46.60%; H, 5.39%; N, 3.02%. Found: C, 46.19%; H, 5.47%; N, 2.89%.  56  4"-Cyanophenyl 2,3,6,2'j',4',6'-hepta-0-acetyl-f$-D-cellobioside (11) The protected cellobioside 11 was synthesized from acetobromocellobiose (1.5 g, 2.14 mmol) and 4-cyanophenol (510 mg, 4.28 mmol) according to the general method (Section 1.3) and recrystallized as a white powdery solid (500 mg, 0.68 mmol, 32%) from ethyl acetate and ethanol. M.p. 217-219 °C. H NMR (CDCI3): 6 7.60 (d, J 9 Hz, !  H(3",5")), 7.05 (d, J 9 Hz, H (2",6")), 5.70 (d, J i , 8 Hz, H(l)), 3.65-5.55 (m, H(2-6) 2  and H(l'-6')), 2.00-2.15 (7 s, 7(OAc)). Elemental analysis for C33H39NO18; calculated: C, 53.73%; H, 5.29%; N, 1.90%. Found: C, 53.77%; H, 5.40%; N, 1.86%.  4"-Cyanophenyl B-D-cellobioside (12) Compound 12 was prepared from the acetylated cellobioside (11) (200 mg, 0.27 mmol) by NaOMe/MeOH deacetylation and crystallized and recrystallized as a white powder from methanol and diethyl ether (70 mg, 0.16 mmol, 59%). M.p. 241-242 °C. H NMR ( C D 3 O D ) : 5 7.70 (d, J 9 Hz, H(3",5")), 7.23 (d, J 9 Hz, H(2",6")), 5.12 (d,  l  J l , 9 Hz, H(l)), 3.20-5.05 (m, H(2-6)and H(l'-6')). 2  C19H25O11N,  Elemental analysis for  calculated: C, 51.47%; H, 5.64%; N, 3.16%. Found: C, 50.60%; H ,  5.96%; N, 2.97%.  2,5,6,2 '3',4',6'-Hepta-0-acetyl-D-cellobiose  (13)  The peracetate (1) (2.0 g, 2.80 mmol) and hydrazine acetate (330 mg, 3.60 mmol) were dissolved in dimethylformamide (6 mL).  The mixture was stirred at 50 °C until  complete dissolution and then allowed to proceed for three hours at room temperature to completion. The reaction mixture was diluted with approximately 15 mL of ethyl acetate and washed (2 x 20 mL) with saturated sodium chloride. The ethyl acetate was evaporated in vacuo leaving a pale yellow oil which was then redissolved in toluene and the solvent  57 evaporated to remove any residual DMF. The product (13) was recrystallized from ethanol as a white solid (1.133 g, 1.78 mmol, 63%). M.p. 206-208 °C. (lit. m.p. 208 °C). 46  2;'4"-Dinitrophenyl2,3,6,2\3\4\6'-hepta-0-acetyl-^  (14)  The cellobioside (13) (1.08 g, 1.70 mmol) and l,4-diazabicyclo(2.2.2)octane (660 mg, 5.90 mmol) were stirred over molecular sieves (4 A) in DMF (20 mL) for three hours. Fluorodinitrobenzene (409 mg, 2.20 mmol) was added and the reaction was allowed to proceed at room temperature for 24 hours. The sieves were removed by gravity filtration, washed with chloroform and the combined extracts evaporated in vacuo. The remaining yellow solid was dissolved in chloroform (40 mL) and washed (3 x 50 mL) with saturated sodium bicarbonate. The organic layer was dried over anhydrous MgS(X filtered and the solvent evaporated in vacuo leaving a yellow solid which recrystallized from ethyl acetate and low boiling petroleum ether to yield compound 14 (505 mg, 0.63 mmol, 37%) as a white solid. M.p. 213-214 °C.  ]  H NMR (CDC1 ): 8 8.70 (d, J », " 4 Hz, H(3")), 8.45 3  3  5  (m, H(5")), 7.40 (d, J \5"9 Hz, H(6")), 3.65-5.53 (m, H(l-6) and H(l'-6')), 2.00-2.15 6  (7 s, 7 OAc). Elemental analysis for C32H38N2O22. calculated: C, 47.90%; H, 4.74%; N, 3.49%. Found: C, 48.12%; H, 4.74%; N, 3.39%.  2",4"-Dinitrophenyl R-D-cellobioside (15) The protected cellobioside (14) (300 mg, 0.59 mmol) was deacetylated using the HCl/MeOH method and the product (15) was recrystallized from methanol and diethyl ether as a white powdery solid (128 mg, 0.16 mmol, 27 %). M.p. (dec.) 180 °C. *H NMR (CD3OD): 8 8.72 (s, H(3")), 8.45 (d, J ", " 10 Hz, H(5")), 7.62 (d, J \5" 10 Hz, 5  6  H(6")), 5.30 (d, J i , 6 Hz, H(l)), 3.22-5.00 (m, H(2-6) and H(l*-6')). 2  6  Elemental  analysis for Ci8H 4N20i5. calculated: C, 42.50%; H , 4.72%; N, 5.51%. Found: C, 2  42.20%; H , 4.87%; N, 5.33%.  58 3,62'j',4',6'-Hexa-0-acetyl cellobial  (16)  To acetobromocellobiose (2.0 g, 2.86 mmol) was dissolved in glacial acetic acid (60-70 mL) was added Zn/AgOAc (zinc metal (4.5 g) activated with 1 0 % HC1 was added to silver acetate (150 mg) boiling in acetic acid). The reaction proceeded to completion (16 hours) at 4 °C. The Zn(Ag) residue was removed by gravity filtration and the resultant clear solution poured over ice-water. The product was extracted with chloroform (2 x 5 0 mL) and the combined organic layers washed (3 x 50 mL) with saturated sodium bicarbonate, then dried over anhydrous MgSCM, filtered and evaporated in vacuo to leave a pale orange oil. The product was purified by flash column chromatography using chloroform/diethyl ether (2:1) solvent mixture and recrystallized from chloroform and low boiling petroleum ether to give 16 as a white powder (400 mg, 0.71 mmol, 25%). M.p. 134-135 °C (lit. m.p. 134-135 °C). 47  l,3,6,2\3',4',6'-Hepta-0-acetyl-2-deoxy-2-fluoro-p-D-cellobiose (17) The cellobial (16) (80 mg, 0.14 mmol) was dissolved in freon-11 (10 mL) and acetonitrile (1 mL). Acetyl hypofluorite (generated by bubbling diluted fluorine gas (in neon gas) through a column of sodium acetate in freon-11/acetic acid ) was bubbled into 48  the reaction mixture. After 15-20 minutes at room temperature, the solvent was evaporated in vacuo leaving a white solid. This procedure was repeated 12 moretimesas the column was only designed to generate hypofluorite to react with a maximum of 0.14 mmol of reactant. The products from the individual reactions were pooled and compound 17 was recrystallized from diethyl ether and chloroform as white crystals (206 mg, 0.32 mmol, 17 %). The 2F-manno-analogue of compound 17, a side product of this reaction was left behind in the filtrate. M.p.193-194 °C. *H NMR (CDC1 ): 8 6.35 (d, J-,2.4 Hz,H(l)), 3  3.45-5.60 (m, H(2-6) and H(l'-6')), 2.00-2.15 (7 s, 7 OAc). CDCI3):  5 -201.8 (dd, JF2H2 50 Hz, JF2H3 12 Hz, F(2)).  1 9  F NMR ( H coupled, !  59 2-Deoxy-2-fluoro-3,6,2'j',4',6'-hexa-0-acetyl-D-cellobiose (18) The selective deacetylation of the fluorinated peracetate (17) (200 mg, 0.31 mmol) was accomplished with hydrazine acetate (35 mg, 0.38 mmol) as previously described for compound 13. The product (18) was recrystallized as the anomeric mixture from chloroform/low boiling petroleum ether as a white solid (130 mg, 0.22 mmol, 71%). M.p. 213-215 °C. (OAc)).  1 9  !  H NMR (CDCI3): 8 3.40-5.60 (m, HQ-6) and H(l'-6')), 1.95-2.05 (6 s, 6  F NMR ( H decoupled.CDCls): -200.10 (s, F(2)), -199.48 (s, F(2)). !  2",4"-Dinitrophenyl 2-deoxy-2-fluoro-3,6,2',3',4',6'-hexa-0-acetyl R-D-cellobioside (19) Compound 19 was synthesized using the procedure previously described for the synthesis of acetylated 2", 4"-dinitrophenyl cellobioside (14). Fluorodinitrobenzene was added (48 mg, 0.26 mmol) to the partially protected disaccharide (18) (130 mg, 0.22 mmol) and l,4-diazabicyclo(2.2.2)octane (72 mg, 0.64 mmol) stirring in DMF (3 mL) over molecular sieves. The reaction was allowed to proceed to completion at room temperature. The product (19) was isolated by flash column chromatography using a ethyl acetate/low boiling petroleum ether (1:1) solvent mixture. Evaporation of the solvent in vacuo left a yellow solid which was recrystallized from low boiling petroleum ether and ethyl acetate as a pale yellow solid (55 mg, 0.072 mmol, 33%). M.p. 182-183 °C. *H NMR (CDCI3): 8 8.75 (d, J », " 4 Hz, H(3")), 8.45 (dd, J \3" 4 Hz, J " " 9 Hz, H(5")), 3  5  5  5  t6  7.40 (d, J6",5"9 Hz, H(6")), 3.65-5.45 (m, H(l-6) and H(l'-6')), 2.00-2.15 (6 s, 6 (OAc)).  1 9  F NMR (*H decoupled, CDCI3): 8-197.125 (s, F(2)). Elemental analysis for  C30H35O20F. calculated: C, 47.24%; H , 4.59%; N, 3.67%. Found: C, 46.95%; H , 4.67%; N, 3.62%.  60  2",4"-Dinitrophenyl2-deoxy-2-fluoro R-D-cellobioside (20) The protected glycoside  1 9 ( 4 8  mg,  mmol) was deacetylated using the  0 . 0 6 3  HCl/MeOH method to give product ( 2 0 ) which was recrystallized from ethanol to give a white solid ( 1 1 mg, 0 . 0 2 2 mmol, 3  J" 3  >5  H ( 6 " ) ) ,  2  Hz, H ( 3 " ) ) ,  (d, Ji,2  5 . 6 5  8 . 5 0  9  5 % ) .  (dd, J " 5  Hz, H(l)),  coupled, C D 3 O D ) : 8 - 1 9 8 . 4 1 (dd, for C18H23N2O14F, calculated: C, 4 . 6 7 % ;  N,  3  2  M.p.  Hz, J ", " 5  3 . 2 0 - 5 . 1 0  JF2,H3  1 6  4 2 . 3 5 % ;  °C *H NMR (CD OD): 8 8 . 7 5 (<L  1 7 8 - 1 7 9  6  9  3  Hz, H ( 5 " ) ) ,  7 . 6 5  (m, H ( 2 - 6 ) and H(l'-6')). Hz,  H,  JF2,H2  4 . 5 1 % ;  5  0  Hz,  F ( 2 ) ) .  (d, 1 9  J  6  \ 5 "  9  Hz,  F NMR ( H !  Elemental analysis  N, 5 . 4 9 % . Found: C,  4 1 . 7 3 % ;  H,  5 . 1 9 % .  2.  ENZYMOLOGY  2.1  General procedures The C.fimi exoglucanase stock ( 2 4 mg/mL as determined by the BIORAD protein  assay using bovine serum albumin as the standard) was provided by Dr. Neil Gilkes of the Department of Microbiology, University of British Columbia. measurements were recorded on a Pye Unicam 8  8 0 0  A l l absorbance  UV/Vis spectrophotometer equipped  with a circulating water bath. The bovine serum albumin was purchased from Sigma and the buffers from BDH.  2.2  Determination of the molar extinction coefficient of the C.fimi exoglucanase The molar extinction coefficient of the exoglucanase was measured as follows.  Aliquots of the stock exoglucanase were diluted in sodium phosphate buffer ( 5 0 mM, pH 7 . 0 ) and incubated at 3 7 °C in the spectrophotometer. The absorbance at 2 8 0 nm was recorded and the approximate molar extinction coefficient of the exoglucanase determined to be 2 . 3 cm" mg" mL" from Beer's law: 1  1  1  61 A where A is the absorbance, b is the cell path length (1 cm) and c is the concentration (mg/mL).  2.3  Determination of K  and V  m  m a  x  for the hydrolysis of aryl  P-D-  glucosides & aryl p-D-cellobiosides by C. find exoglucanase Solutions of substrate (-0.7 mL) in sodium phosphate buffer (50 mM, pH 7.0) and bovine serum albumin (1 mg/mL) were incubated at 37 °C in spectrophotometer cells within the spectrophotometer until thermally equilibrated (-20 rnin). The reaction was initiated by the addition of enzyme (-10 uL) and the reaction progress was followed by the change in absorbance due to the release of the phenolate. The rates of hydrolysis of the glycosides were followed at wavelengths where there was a maximal absorbance difference between the initial glycoside and the phenol product. In order to ensure linear kinetics and to obtain a sufficient absorbance change for accurate calculation of the rates, the concentration of the enzyme added and the length oftimethat thereactionwas monitored were chosen such that less than 10% of the total substrate was converted to product. Initially, approximate K  m  and V x values were calculated from three poini m a  Lineweaver-Burk plots where the initial rates of hydrolysis for three widely varied substrate concentrations were measured. More accurate values were then determined by measuring the initial rates of hydrolysis for 6-10 different substrate concentrations. Generally rates were measured over a wide range of substrate concentrations (0.2 to 5 times K ) . However, due to the insolubility of some substrates, much narrower substrate m  ranges were studied in several cases. The results are illustrated as Lineweaver-Burk double reciprocal plots for visual convenience in Appendix 2 where the enzyme concentrations, the wavelengths and the molar extinction coefficients used are indicated in the legends. However, due to the inaccuracy introduced by the nonlinear error span of the double  62 reciprocal analysis, the K and V m  m a x  values reported were actually determined from a  weighted nonlinear regression analysis, using a Curvefitter program written for an Apple lie computer by Ian P. Street  2.4  Determination of K, and kj for 2',4'-dinitrophenyl 2-deoxy-2-fluoroP-D-glucoside and 2",4"-dinitrophenyI 2-deoxy-2-fluoro-P-Dcellobioside The equilibrium binding constant (KO and the inactivation rate constant (ki) were  measured as follows. C.fimi exoglucanase was added to inactivation mixtures containing BSA (1 mg/mL) and varying concentrations of the inactivators in sodium phosphate buffer (50 mM, pH 7.0) incubated at 37 °C. Aliquots were removed at differenttimeintervals and diluted into reaction cells containing a large volume of substrate (2,4 DNPG or 2,4 DNPC) at saturating concentrations. This stopped the inactivation both by dilution and by substrate competition for the enzyme as the substrate is present large excess. The residual enzymatic activity was then determined from the rate of hydrolysis of the substrate, which is directly proportional to the amount of active enzyme. The inactivation was monitored until 80-90% of enzymatic activity was lost From the slope of the plot of natural logarithm of the residual activity versus time, pseudo-first order rate constants (kobs)  w e r e  calculated for each inactivator concentration.  These plots are illustrated in Appendix 2 and the inactivator concentrations are indicated in the legends. Plotting the reciprocal of kobs  as a  function of the reciprocal of the inactivator  concentration, using the Curvefitter weighted nonlinear regression program, afforded values for Ki and ki. These results are illustrated as Lineweaver-Burk plots for visual convenience in Appendix 2.  63 2.5  Reactivation of inactivated C.fimi exoglucanase Samples of exoglucanase inactivated by either 2',4'-dinitrophenyl 2-deoxy-2-  fluoro-P-D-glucoside or 2",4"-dinitrophenyl 2-deoxy-2-fluoro-P-D-cellobioside  were  extensively dialysed at 4 °C against several changes of phosphate buffer (50 mM, pH 7.0) in order to remove the excess inactivator. Aliquots of the 2F-DNPG-inactivated enzyme were removed and added to buffer solutions containing BSA (1 mg/mL) and either (1) sodium phosphate buffer (50 mM, pH 7.0) only, (2) 55 mM P-D-glucopyranosyl benzene, (3) 55 mM glucose or (4) 55 mM cellobiose. Aliquots of the 2F-DNPC-inactivated enzyme were removed and added to buffered solutions containing BSA (1 mg/mL) and either (1) sodium phosphate buffer (50 mM, pH 7.0) only or (2) 55 mM cellobiose. These solutions were incubated at 37 °C and monitored for return of activity by periodic removal of samples and addition to spectrophotometer cells containing sodium phosphate buffer, BSA (1 mg/mL) and substrate (2,4 DNPG or 2,4 DNPC). The enzyme activity was determined from the rate of release of 2,4-dinitrophenolate which is directly proportional to the enzymatic activity. The reactivation rate constants were calculated from the slopes of plots of ln(full rue - observed rate) vs time as illustrated in Appendix 2. The concentrations of the reactivators used are given in the legends. The linearity of these plots indicates that the reactivation process was first order.  2.6  Determination of the catalytic nucleophile of C. fimi exoglucanase with {1- H}-2F-DNPG 3  The exoglucanase (2 mg, 0.54 mL) was radiolabelled by incubation in the presence of tritiated 2',4'-dinitrophenyl 2-deoxy-2-fluoro-P-D-glucoside  (1.07 mM) in sodium  phosphate buffer (50 mM, pH 7.0) at 37 °C over a 34 day period. The residual enzymatic activity was monitored at regular intervals. Enzyme incubated in the absence of inactivator over this 34 day period showed no significant loss of activity. Aliquots of these mixture  64 were also removed over the inactivation period and added to a "stop" mixture (5 pX, 20 mM Tris, 2 mM EDTA, 0.05% bromophenol blue, 5% SDS, 10% p-mercaptoethanol) to monitor protein stability. These samples were heated at 95 °C for 5 minutes and then electrophoresed on a Pharmacia Phast 20% acrylamide gel. After 80% of the activity had been lost, the excess inactivator was removed by extensive dialysis against phosphate buffer (50 mM, pH 7.0) at 4 °C and the resultant enzyme solution was then concentrated using a Millipore UFC-10 polysulfone membrane concentrator. The radiolabelled enzyme was proteolysed by pepsin digestion(l:100, pepsimexoglucanase, w/w) in sodium phosphate buffer (50 mM, pH 2.1) at room temperature which was monitored by SDSPAGE. Aliquots (5 uL) were removed at differenttimesover the six hour digestion period and added to a "stop" mixture (5 pL 20 mM Tris, 2 mM EDTA, 0.05% bromophenol blue, 5% SDS, 10% p-mercaptoethanol). These samples were heated at 95 °C for 5 minutes and then electrophoresed on a Pharmacia Phast 20% acrylamide gel which allowed for visualisation of the resultant peptide fragments. Separation of the peptide fragments was accomplished by high performance liquid chromatography (HPLC) and monitored by UV absorption at 215 nm using a Waters peptide analyser Model 990 and 600 E at 50 °C. The peptide mixture was loaded onto a Vydac C4 column (2.1 x 150 mm), eluted with a gradient of 0.1% trifluoroacetic acid (TFA) in water to 0.1% TFA in 70% acetonitrile/30% water and the samples collected manually. The radioactivity of the fractions collected was determined on a Packard TriCarb Liquid Scintillation analyser 2200CA. Two of the fractions collected were radioactive (A and B). These were concentrated on a Savant Speedvac and rechromatographed on a Vydac CI8 column using the same eluting solvent mixtures. This additional purification of the radioactive peptide B on the CI8 column resulted in a sufficiently pure peptide for sequencing. Peptide A required further purification and was reconcentrated and reloaded onto the CI8 column and eluted with a gradient of 150 mM sodium chloride/water to 150  mM sodium chloride in 70% acetonitrile/30% water. The resultant radiolabelled peptide was sufficiently pure to be sequenced. Peptides A and B were sequenced by solid phase Edman degradation on a Milligen/Biosearch model 6600 protein sequencer by Ruedi Aebersold at the Biomedical Research Center, UBC, Vancouver. This involved the sequential derivatisation of the Nterminal amino acids to phenylthiohydantoins (PTH) followed by the separation and identification of the PTH-residues by HPLC.  66  APPENDIX 1 BASIC CONCEPTS OF ENZYME CATALYSIS  1.  BASIC ENZYME KINETICS The basic equation of enzyme kinetics is the Michaelis-Menten equation where v is  the velocity of the reaction measured either as the initial rate of formation of the products or depletion of the substrates; [E] is the total concentration of the enzyme; [S] is the substrate concentration; k ^ is the catalytic constant; and KJJ, is the Michaelis constant [E ][S]k  V  V  0  "  K  M  c a t  V*  + [S]  In the Michaelis-Menten equation two assumptions are made; the enzyme concentration is negligible compared to that of the substrate which is generally true since enzymes catalyse reactions with a high efficiency and, the velocity measured is the initial rate of product formation, thus there is no significant accumulation of product (or depletion of substrates) hence, the reverse reaction can be ignored. Therefore, the change in substrate concentration is generally linear with time. The KJJJ is the substrate concentration when v = V  m a x  / 2 . It may be treated as an  apparent dissociation constant of all the bound enzyme species and as such is expressed as:  K  M  =1*^1  L [ES]  The value of  (2)  can be a measure of the enzyme affinity for the substrate, for example, a  low KJJJ means that the enzyme has a high affinity for the substrate. At low substrate concentration ( [ S ] « K ) the Michaelis-Menten equation becomes: M  v  =  kcat[E ][S3 9  whereas at saturating concentrations ( [ S ] » K ) the equation becomes: m  max = catI ol  v = V  k  E  At low [S] most of the enzyme is unbound such that the total enzyme concentration, which is a sum of the concentration of the free and bound enzyme, can be approximated to the concentration of the free enzyme, [E]. The Michaelis-Menten equation under these condition is expressed as  v  The k  c a t  /K  m  .k«JHJSi  from the above equation is an apparent second-order rate constant  which relates the reaction rate to the concentration of the free enzyme and free substrate. This kinetic parameter is also referred to as a specificity constant which is a measure of the catalytic efficiency for the substrate. The Michaelis-Menten equation is often changed to a linear form which is useful for graphical analysis of data and detection of deviations from the expected values. An example of the Michaelis-Menten equation transformed is where both sides of the Michaelis  68 -Menten equation have been inverted. Plotting 1/v as a function of 1/[S] gives the Lineweaver-Burk plot (Fig 32) where the y-intercept is 1/V  max  , the x-intercept is -1/Kg,  and the slope is KJJ/VJ^.  ./Is!  o  Figure 32. A typical Lineweaver-Burk plot.  56  2.  INTERPRETATION OF k  The rate constant k  Mt  c a t  AND k  which equals V  max  c a t  /K 58 m  / [ E ] is a reflection of the rate of the rate 0  determining step and the rate constant, k^j/K,,, reflects the rate of the first irreversible step in the reaction. In order to show this, first consider the following general mechanism shown below, where the formation of ES is referred to as the association step, the interconversion of the ES and EP as the chemical step and the final step as product-release.  E  •  S  ki  ES  *  f  EP  ki  —  — ^ — • E  • P  The corresponding free energy diagram for this mechanism is shown in Fig 33 where the energy levels are arbitrarily chosen. It can be shown that the kinetic parameters for the reaction are given by equations 1,2 and 3.  E - P  Reaction Ccordinaic Figure 33. Free energy diagram for the enzymatic reaction involving the interconversion of intermediates  _  cat  K  -  m  kk ic.2 + k ->- k 2  3  2  (1)  3  _ k (k.i + k ) + k.ik. ki (k. +k +k ) 3  2  2  kcat  2  2  3  k]k2k  3  KnT k (k.i + k ) + k.ik.2 =  3  (2)  (3)  2  Now, consider the above reaction where there is a rapid, reversible association step followed by a rate determining chemical step. The kinetic relationships that describe this situation are k-i » k , k » k. and k » k . When these conditions are applied to 2  3  2  3  2  equations 1 and 3, the kinetic parameters are reduced to:  70 k  cat= 2 K  Kai  =  k]k  2  which expressed in Eyring form become:  _kT  -  h  l^at _ kT K  h  _ m  -AG_*/RT  e c r.  e  (G  ± - G )VRT E+S  -AGpt/RT  _kT  where k is the Boltzmann constant and h is Planck's constant. Thus, under these conditions, k  cat  and k^/K^ both give information pertaining to the transition state of the  chemical step. However, the initial reference point for k  cat  is the ES complex and for  k ^K , it is free enzyme (E) and free substrate (S). ca  m  If however, the restrictions on the reaction were rapid, reversible association followed by an irreversible chemical step and then rate-determining product release: the kinetic relationships would be k.j »  k , k » k and k » k- . The kinetic constants 2  then can be reduced to:  k^ ~ 3 k  kcat kik K " k.i m  and the Eyring equations are:  2  2  3  3  2  71 _kT  -  h  e  ^cat-El  Kin"  h  -AG t/RT D  P •AG */RT T  e  In this case, kcat refers to the transition state of product release with the enzyme-product complex as the initial reference point and k  c a t  /K  m  refers to the transition state of the  chemical step with E and S as the initial reference points. Thus, these examples, show that k  c a l  refers to the rate-determining step while  ^cat/Km corresponds to the first irreversible step in the reaction with E and S as reference states. These general concepts can be extended to the hydrolysis of (3-glucosides and Pcellobiosides by C. fimi exoglucanase where the chemical step corresponds to glycosylation, and product-release to deglycosylation. Thus, the first example of rapid, reversible association followed by a rate determining chemical step is equivalent to the situation when glycosylation is the rate determining step and the second example of rapid, reversible association, an irreversible chemical step and then rate-determining product release corresponds to the situation when deglycosylation is rate determining.  3.  BINDING E N E R G Y AND E N Z Y M E CATALYSIS The function of any catalyst, including an enzyme, is to lower the activation energy  of a reaction, thus leading to rate acceleration. Enzymes are known to bind specifically to their substrates and the binding energies involved may be quite large. However, since the structure of the substrate changes as it is converted to product via a transition state, the  72 enzyme can only be fully complementary to one 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 of the substrate. Consider a typical enzymatic reaction consisting of a binding step and a catalytic step such as that shown below.  E  +  S "  Km  ES  kcat  •  E +  P  The energy diagram for this reaction is illustrated in Fig 34 where k-i » k2, so that K  M  =  K and the kinetic parameters may be expressed as the following. S  v  _  -AG /RT  .  _kT  CC  -AG^t/RT  Kcat - jj e kcat _ kT Kin" h  c  -AG_* /RT e  Now, consider the above situation when an extra amount of binding energy, A G R , has become available, for example a hydrogen bond between the enzyme and the substrate. If that extra binding energy is realized at the ground state (Fig 35) rather than the transition state, then the ground state is stabilized and A G E S increases by A G R , thus K  M  is decreased  73  G  E+S  E+P Reaction Coordinate Figure 34. Thefreeenergy diagram for atypicalreaction (solid line) and the corresponding uncatalysed reaction (dashed line).  The value of kc is also reduced since AGc* (Michaelis complex (ES) proceeding to the al  transition state (ES*)) is increased by the ground state stabilization, A G R . However, kcat/K is unaffected as this is the rate constant for free enzyme (E) and free substrate (S) m  proceeding to the transition state (ES*). This shows that if the enzyme is complementary to the ground state, then there is tighter substrate binding but a slower reaction rate, kc . at  Alternatively, if the extra binding energy, A G R , is realized at the transition state (Fig 36), then the transition state is stabilized and AGc* and AGj will be lowered by A G R , thus, kcat and kc t/K will be increased. The value of A G E S is unchanged, and hence, K a  m  m  remains the same. This shows that when the enzyme is complementary to the transition state rather than the ground state, then the rate of the reaction and the enzyme efficiency (kcat/K ) is increased m  74  ES  Figure 35. The free energy diagram when the enzyme is complementary to the ground st of the substrate  Figure 36. The free energy diagram when the enzyme is complementary to the transition state of the substrate.  4.  INACTIVATION KINETICS OF C. FIMI EXOGLUCANASE A schematic representation of the mechanism of the exoglucanase-catalysed  hydrolysis of substrates is the following where k2 corresponds to glycosylation and k3 to deglycosylation.  E + GX + ±  k. j  EGX - J * E-G "HX +H 0  E GOH +  2  The inactivators 2F-DNPG and 2F-DNPC have been designed so that k 3 « k 2 while  k - » k 2  thus leading to the accumulation of the covalent glycosyl-enzyme  intermediate. The kinetic equation for this inactivation, is the following variation of the Michaelis-Menten equation,  v v  kj[E ][GX] ~ K j + [GX] 0  u  ;  where kj is the inactivation rate constant and Kj is the apparent dissociation constant for all bound enzyme species. If [GX]»[Eo], then [GX] appears constant throughout the inactivation process and the kinetics are pseudo-first-order with respect to the enzyme concentration. Thus, the Michaelis-Menten equation can be expressed as  v = kobs[E ] 0  (8)  kj [GX] ~ Ki + [GX]  w  The value of kobs can be calculated from the slope of the natural logarithm of the residual enzymatic activity plotted as a function of time. That is, since the rate of formation of HX is equal to the rate of inactivation of the enzyme then  76  v  dJHX]_ dTEcJ. " dt ~ dt  a K  f  J  )  '  The rate can then be expressed as  L  ^  ^  e  l  = k [Eo] obs  (11)  = kobsdt  (12)  Therefore, ln[Eo] = -kob t S  (13)  The values of kj and Ki can then be calculated by substituting kobs into equation (9), or graphically by the reciprocal analysis.  77 APPENDIX 2 GRAPHICAL REPRESENTATION OF KINETIC DATA 1. HYDROLYSIS OF ARYL P-GLUCOSIDES AND ARYL BCELLOBIOSIDES BY C. FIMI EXOGLUCANASE. 100  o H — • — i — i — i — i — | — i — i — | — i  0  2  4  1/[2,4  i  6  8  (1/mM)  DNPG]  Figure 37. The Lineweaver-Burk plot for the hydrolysis of 2,4 DNPG. [Enzyme] = 7.98 x 10- mglmL; X = 400 nm; Ae = 10.91 M^cm' 4  0 -| 0  1  1  1  1  1  1 1/(3,4  2  •  1  >  3  4  DNPG] (1/mM)  Figure 38. The Lineweaver-Burk plot for the hydrolysis of 3,4 DNPG. [Enzyme] 8.0 x= 10- mglmL; A = 400 nm; Ae = 11.01 M-hmr 4  1  78  800  1 oo H — i — i — i  0  2  •—i—i—i—i—i—i—i—«—  •  4 6 1/J2CI4NPG] (1/mM)  8  10  Figure 39. The Lineweaver-Burk plot for the hydrolysis of2Cl 4NPG. [Enzyme] = 5.95 x lQr mg/mL; X = 294 nm; Ac = M-kmr 4  10  H 0.0  1  1  1 0.1  1  —i  1  —  •  0.2  —  0.3  1/[PNPG] (1/mM) Figure 40. The Lineweaver-Burk plot for the hydrolysis ofPNPG. [Enzyme] - 4.83 x JO' mg/mL; A = 400 nm; Ae = 7.28 Nf-kmr 2  1  79  Figure 41. The Lineweaver-Burk plot for the hydrolysis ofMNPG. [Enzyme] = 2.94 x 10- mglmL; X = 380 nm; Ae = 0.385 Af-lcmr . ]  1  40  1/[2,4 DNPC]  (1 / mM)  Figure 42. The Lineweaver-Burk plot for the hydrolysis of 2,4 DNPC. [Enzyme] = 1.79 x 10 mglmL; A = 400 nm; Ae = 10.91 h^kmr 4  1  80  5  1 - J — i — i — |  0  2  i  i  |  i  i — | — i — — i — |  4  6  1/[3,4 DNPC]  i  8  •—|  10  • •  12  (1/mM)  Figure 43. The Lineweaver-Burk plot for the hydrolysis of 3,4 DNPC [Enzyme] = 3.75 x 10-3 mg/mL; X = 400 nm; Ae = 11.01 M-kmr 1  140 -j E c o o  min  < <  120 100 80 60 40 20 0  1  2 1/[PNPC]  3  4  5  6  (1/mM)  Figure 44. The Lineweaver-Burk plot for the hydrolysis ofPNPC. [Enzyme] = 352 x 1(H mg/mL; X = 400 nm; Ae = 7.28 M-kmr 1  81  = 750 x Figure 46. The Lineweaver-Burk plot for the hydrolysis ofMNPC. [Enzyme] lO' mg/mL; X = 380 nm; Ae = 0.385 f^km' 3  1  82  10 H 0  •  1  «  1 2 1/[3,5 DCIPC]  1  1  3 (1/mM)  Figure 47. The Lineweaver-Burk plot for the hydrolysis of 3,5 DCIPC. [Enzyme] = 1.13 x 10' mglmL; X = 280 nm; Ae = 0.732 M-kmr 2  1  83  2. PLOTS OF THE INACTIVATION OF C. FIMI EXOGLUCANASE WITH 2F-DNPC AND 2F-DNPG  Figure 49. The inactivation of C.fimi exoglucanase with 2F-DNPC.  84  Figure 51. The inactivation of C.fimi exoglucanase with 2F DNPG.  70 T 0.15  1  1  0.25  i  |  i  0.35  |  0.45  1/[l]  Figure 52.Plot of 1/k  obs  |  1  0.55  (1/mM)  vs. llll] for 2F DNPG.  •  |  0.65  i  0.75  85 3.  PLOTS  O F T H E REACTIVATION O F INACTIVATED EXOGLUCANASE  C.FIMI  •  Buffer  •  57 mM Cellobiose  Time (h)  Figure 53. 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