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Glycals as probes for mechanistic studies of glycosylases Lai, Ellen Chui-Kwan 1995

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Glycals as probes for mechanistic studies ofglycosylasesbyEllen Chui-Kwan LaiB.Sc., The University of Waterloo, 1985M.Sc., The University of Waterloo, 1987A THESIS SUBMITtED IN PARTIAL FULFILMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIESDepartment of ChemistryWe accept this thesis as conformingto the required standard:THE UNIVERSITY OF BRITISH COLUMBIANovember, 1994.© Ellen Chui-Kwan Lai, 1994.In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. it is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)Department of C_I4E-1 I 5 T&YThe University of British ColumbiaVancouver, CanadaDate Fight 91/.DE.6 (2/88)IIABSTRACTSeveral derivatives of D-gluco-heptenitol and D-glucal were synthesized and used to study the reactionmechanisms of three glycosylases (glycosyl mobilizing enzymes): glycogen phosphorylase, J3-glucosidase, andf3-N-acetythexosaminidase. l-Fluoro-D-gluco-heptenitol (Fjhept) and 1, 1-difluoro-D-gluco-heptenitol (F2hept)acted as competitive inhibitors of glycogen phosphorylase b, and collaborative X-ray crystallographic studiesrevealed that both F1hept andF2hept bind to the active site. Furthermore, F1hept and phosphate bindsimultaneously, allowing crystallographic investigation of a stable ternary complex.Various glycals were examined as potential substrates (catalytically hydrated by the enzyme), or aspotential inhibitors or inactivators (of the hydrolysis of a glucoside substrate), of Agrobacterium f3-glucosidase.Both heptenitol and methylglucal acted as substrates of this enzyme. Enzymatic protonation of the double bondof methylglucal occurred from below the ring. F1hept and Fhept acted as noncompetitive (or uncompetitive)inhibitors, as did heptenitol (at concentrations < 8 mM).a,f3-Unsaturated glucals, which might act as Michael acceptors for a nucleophilic residue in the activesite of a glycosylase, were investigated as a new class of potential inactivators of Agrobacterium [3-glucosidase.1-Nitroglucal functioned as a time-dependent inactivator, probably as a result of a Michael addition reactionbetween 1-nitroglucal and a nucleophilic residue in the active site. However, protein mass spectrometryrevealed that in most cases more than one equivalent of l-nitroglucal bound to the enzyme. Unfortunately, othercz,13-unsaturated glucals that were examined [1-cyano-, 1-(methyl carboxylate)-, sodium 1-(carboxylate)-, and2-cyano- derivatives of glucal] only acted as reversible inhibitors.Three f3-N-acetythexosaminidases (human placenta, jack bean, and bovine kidney) were also studied,and shown to hydrolyze an N-acetylglucosaminide substrate with net retention of anomeric configuration. Allthree enzymes hydrated 2-acetamido-D-glucal, yielding N-acetyl-D-glucosamine as the product in each case.This is the first time that proton transfer has been shown to occur from the top face during the hydration of aglycal by a ‘retaining’ [3-glycosidase. 2-Acetamido-D-glucal bound tightly to the human, bovine, and jack beanenzymes, with K1 values of 8.5, 25, and 29 M, respectively.111TABLE OF CONTENTSABSTRACTTABLE OF CONTENTS iiiLIST OF TABLESLIST OF FIGURES viiiLIST OF REACTION SCHEMES xABBREVIATIONS AND SYMBOLSACKNOWLEDGMENTSDEDICATION MVCHAPTER 1: GENERAL INTRODUCTION 11.1. Glycosylases and the reactions they catalyze 11.2. The classification of glycosylases 21.3. General features of the catalytic mechanism of retaining glycosylases 31.3.1. Overview 31.3.2. General acid catalysis 51.3.3. The carboxylate group in the active site 6a. X-ray structural studies 6b. Fluoro sugars 6c. Conduritol epoxides 7d. Cyclophellitol 8e. N-Acetylconduramine B trans-epoxide 9f. Other inhibitors of NAGases 101.3.4. The nature of the glycosyl-enzyme intennediate 12a. Evidence from lysozyme and Agrobacterium 3-glucosidase (pABG5) 12b. Evidence from 3-galactosidase 13c. Evidence from glycogen phosphorylase 131.3.5. The oxocarbonium ion-like transition states of glycosylase substrates 14a. cz-Secondary deuterium kinetic isotope effects 14b. Properties of transition-state analogues 15c. Examples of transition-state analogues 15d. The transition states of phosphorylase-catalyzed reactions 181.3.6. The binding energy attributable to noncovalent interactions 191.4. The reactions of glycals with glycosylases 191.4.1. The enzyme-catalyzed hydration of glycals 191.4.2. The enzyme-catalyzed hydration of heptenitols 231.4.3. The reactions of glycals with glycogen phosphorylase 251.5. The aims of this thesis 281.5.1. Research significance and objectives 281.5.2. Studies using fluorinated derivatives of D-gluco-heptenitol 281.5.3. Studies using derivatives of D-glucal 29CHAPTER 2: KINETIC STUDIES USING GLYCOGEN PHOSPHORYLASE 302.1. Introduction 302.1.1. Biochemical role in vivo 302.1.2. Regulation 312.1.3. Structural studies 31iv2.1.4. General features of the catalyticmhism .332.1.5. The cofactor pyridoxal phosphate (PLP) 35a. Structural and catalytic roles 35b. The proposed role of PLP as a BrØnsted acid catalyst. 37c. The proposed role of PLP as an electrophile 372.1.6. Glycogen phosphorylase catalysis as a rapid equilibrium, bireactant system 372.2. The aims of this study 412.3. Results and discussion 422.3.1. Syntheses of fluorinated heptemtols 422.3.2. The stereochemistry ofF1hept 46a. The trans and cis isomers ofF1hept 46b. 13CNMR experiments 46c. An NOE experiment on TBDMS-protectedF1hept 48d. 1H and 19F NMR experiments 482.3.3. Substrate and inactivation tests usingF2hept 492.3.4. Inhibition studies usingF2hept andF1hept 502.3.5. X-ray crystallographic studies 552.4 Conclusions 60CHAPTER 3: KINETIC STUDIES USING f3-GLUCOSIDASE 623.1. Introduction 623.1.1. Importance and general properties of 3-glucosidases 62a. Catalytic activity and role in the microbial cellulase complex 62b. Catalytic activity and role in lysosomal glycolipid metabolism in humans 623.1.2. Agrobacterium 13-glucosidase 633.2. The aims of this study 643.3. Results and discussion 653.3.1. The synthesis of heptenitol 653.3.2. Kinetic studies using heptenitol 66a. Heptenitol as a substrate of pABG5 66b. Heptemtol as an inhibitor of pABG5 69c. Inactivation tests using heptenitol 713.3.3. Kinetic studies using fluoroheptenitols 71a. Inactivation tests using fluoroheptenitols 71b. Substrate tests using fluoroheptenitols 72c. Fluoroheptenitols as inhibitors of pABG5 723.3.4. Kinetic studies using methylgiucal 76a. The synthesis of methyiglucal 76b. Methylglucal as a substrate of pABG5 76c. The stereochemistry of the catalytic hydration of methylglucal by pABG5 77d. Methylglucal as an inhibitor of pABG5 813.3.5. Kinetic studies using nitroglucal 82a. Nitroglucal as an inactivator of pABG5 82b. Determination of the site(s) of nitroglucal-mediated inactivation of pABG5 84c. Mass spectrometry of nitroglucal-inactivated pABG5 86d. Is niiroglucal-inactivated pABG5 capable of reactivation7 87e. Possible mechanisms for the inactivation of pABG5 by ct,3-unsaturated glucals. 873.3.6. Kinetic studies using other 3-unsaturated glucals 91a. Other x,f3-unsaturated glucals used in this study 91b. The synthesis of 1-cyanoglucal 91c. The synthesis of 2-cyanoglucal 92d. 1-Cyanoglucal as a reversible inhibitor of pABG5 94e. Other cc43-unsaturated glucals as reversible inhibitors of pABG5 96f. 2-Cyanoglucal as a reversible inhibitor of pABG5 97V3.3.7. Kinetic studies using 2-acetamidoglucal .99a. The selective deprotection of peracetylated 2-acetamidoglucal 99b. The inability of 2-acetamidoglucal to bind to pABG5 1003.4. Conclusions 100CHAPTER 4: KINETIC STUDIES USING -N-ACETYLHEXOSAMINIDASE (NAGase) 1044.1. Introduction 1044.1.1. Some general properties of 13-N-acetylhexosaminidase 1044.1.2. The clinical significance of 3-N-acetythexosaminidases 106a. Tay-Sachs disease and Sandhoff disease 106b. Abnormal levels of NAGase activity in cancer cells 1074.2. Aims of this study 1074.3. Results and discussion 1084.3.1. A “direct” colorimetric assay for NAGase-catalyzed hydrolysis of I3G1cNAcPNP 1084.3.2. The stereochemistry of J-NAGase-catalyzed hydrolysis of 3GlcNAcPNP 1114.3.3. NAGlucal as an inhibitor of NAGase 114a. Tests for irreversible inhibition (inactivation) 114b. Tests for reversible inhibition 1144.3.4. NAGlucal as a substrate of NAGase 116a. Preliminary TLC evidence 116b. HPLC analysis of the stereochemistry of the hydration reaction 116c. Determination of Km and Vm for the hydration reaction 1184.3.5. Kinetic studies with other glucal derivatives and glycosides 121a. Studies with D-glucal 121b. Studies with 2-cyanoglucal 123c. Studies with 293G1cDNP and fGlcDNP 1244.3.6. Substrate-enzyme interactions and the mechanism of NAGase-catalyzed reactions.... 1254.4. Conclusions 1284.5. Suggestions for future work 129CHAPTER 5: MATERIALS AND METHODS 1315.1. Organic synthesis 1315.1.1. Materials and routine experimental procedures 131a. Analytical methods 131b. Thin-layer chromatography and silica gel column chromatography 132c. Solvents and reagents 132d. Compounds synthesized and generously provided by colleagues 1325.1.2. Routine synthetic procedures 133a. Deacetylation with sodium methoxide in methanol 133b. Deacetylation with ammonia-saturated methanol 133c. Trimethylsilylation 133d. Triethylsilylation 134e. Tert-butyldimethylsilylation 1345.1.3. Syntheses 135a. The synthesis of heptenitol and its derivatives 135b. The synthesis of glucal and its derivatives 1415.2. Enzyme kinetics 1465.2.1. Miscellaneous procedures and definition of enzyme activity units 1465.2.2. Enzymes and enzyme assays used in this work 146a. Glycogen phosphorylase 1’16b. The assay for glycogen phosphoaylase activity 146c. Agrobacterium 3-glucosidase 147d. The assay for Agrobacterium f3-glucosidase activity 147e. j-N-Acetythexosaminidases 148f. Assays for f3-N-acetythexosaminidase activity 148vi5.2.3. The detennination of kinetic parameters for substes . 149a. Determinations of Km and Vm for various substrates 149b. Determinations of reaction rates for the catalytic hydration of heptenitol 149c. Determinations of reaction rates for the catalytic hydration of methylgiucal 1505.2.4. The determination of kinetic parameters for inhibitors 151a. Determinations ofK1 values (reversible inhibition) 151b. Determinations of K1 values using glycogen phosphorylase 151c. Irreversible inhibition (inactivation) tests: Experimental methods 152d. Irreversible inhibition (inactivation) tests: Theory and calculations 153e. The reactivation test for nitroglucal-inactivated pABG5 1545.2.5. The determination of kinetic parameters by HPLC 155a. Instrumentation 155b. Determination of the product of NAGase-catalyzed GlcNAcPNP hydrolysis. .. 155c. Determination of the rate of NAGase-catalyzed reactions 1565.3. Protein mass spectrometry of nitroglucal-inactivated pABG5 157APPENDIX I: SUPPLEMENTARY DATA 158APPENDIX II: SIMPLE THEORETICAL TREATMENT OF ENZYME CATALYSIS 161A-il. 1. Enzyme catalysis in the absence of inhibition 161A-il.2. The inhibition of enzyme catalysis 164A-il.2.1. Irreversible inhibition 164A-il.2.2. Reversible inhibition 165a. The three types of reversible inhibition 165b. Competitive inhibition 165c. Noncompetitive inhibition 166d. Uncompetitive inhibition 167e. Graphical methods for distinguishing different types of reversible inhibition 169REFERENCES 172VI’LIST OF TABLESTable 1.1. The reversible inhibition of glycosylases by some transition-state analogues 17Table 1.2. Comparison of the binding constants of some ligands of glycogen phosphorylase b 26Table 2.1. Dissociation constants of fluoroheptenitols and substrates with phosphorylase b 54Table 3.1. Summary of kinetic data obtained using a cloned [3-glucosidase, pABG5 103Table 4.1. Kinetic parameters for glycosylase-catalyzed reactions of glucals and related glucosides 121Table 4.2. Summary of kinetic data obtained using various [3-N-acetythexosaminidases 130Table A-lI. 1. Some kinetic parameters for different types of reversible inhibition 171vi”LIST OF FIGURESFigure 1.1. The structural similarities between the glycosyl cation and aldonolactones and aldonolactams 16Figure 1.2. The structural similarities between galactal and the transition state of a 3-galactoside 20Figure 2.1. The structure of the glycogen phosphorylase b monomer 32Figure 2.2. The Schiff base formed between Lys 680 and the cofactor pyridoxal phosphate (PLP) 35Figure 2.3. 13C NMR determination of the stereochemistry of the major isomer ofF1hept 47Figure 2.4. Kinetic parameters for the inhibition byF2hept of phosphorylase-catalyzed glycogen synthesis 51Figure 2.5. Kinetic parameters for the inhibition byF1hept of phosphorylase-catalyzed glycogen synthesis 52Figure 2.6. The inhibition by fluoroheptenitols of phosphorylase-catalyzed glycogen phosphorolysis 53Figure 2.7. The structure ofF1hept orF2hept bound in the active site of glycogen phosphorylase b 57Figure 2.8. Composite electron-density map of fluoroheptenitols bound in the active site of phosphorylase b. .. 58Figure 3.1. The structure of glucosylceramide 63Figure 3.2. 1H NMR determination of the stereochemistry of pABG5-catalyzed hydration of heptenitol 67Figure 3.3. Determination of Km and Vmax for pABG5-catalyzed hydration of heptenitol 69Figure 3.4. Determination of kinetic parameters for the inhibition of pABG5 by heptenitol 70Figure 3.5. Determination of kinetic parameters for the inhibition of pABG5 byF2hept 74Figure 3.6. Determination of kinetic parameters for the inhibition of pABG5 byF1hept 75Figure 3.7. 1H NMR determination of the stereochemistry of pABG5-catalyzed hydration of methylgiucal 78Figure 3.8. Interpretation of 1H NMR evidence for the stereochemistry of methylgiucal hydration 79Figure 3.9. Determination of Km and Vm for pABG5-catalyzed hydration of methylgiucal 80Figure 3.10. Estimation of the K for the inhibition of pABG5 by methylgiucal 82Figure 3.11. Determination of kinetic parameters for the inactivation of pABG5 by mtroglucal 83Figure 3.12. Competitive ligand-mediated protection against nitroglucal-mediated inactivation of pABG5 85Figure 3.13. Ring numbering systems used in this chapter for substituted glucals 88Figure 3.14. Estimation of the K1 for the inhibition of pABG5 by 1-cyanoglucal 94Figure 3.15. Estimation of the K1 values for methylcarboxylate glucal and sodium carboxylate glucal 95Figure 3.16. Minimization of electrostatic repulsion in the active site of the E17OG variant form of pABG5 96Figure 3.17. Estimation of the K for the inhibition of pABG5 by 2-cyanoglucal 98ixFigure 4.1. The catalytic removal of the GaINAc residue from ganglioside Gm2 by NAGase A 105Figure 4.2. Determination of Km and Vj for J-NAGase-catalyzed hydrolysis of 3GlcNAcPNP 110Figure 4.3. Determination of the stereochemistry of J-NAGase-catalyzed hydrolysis of f3G1cNAcPNP 113Figure 4.4. Determination of kinetic parameters for the inhibition of J-NAGase by NAGlucal 115Figure 4.5. HPLC determination of the stereochemistry of J-NAGase-caralyzed hydration of NAGlucal 117Figure 4.6. Determination of Km and Vmax for J-NAGase-catalyzed hydration of NAGlucal 120Figure 4.7. Estimation of the K1 for the inhibition of J-NAGase by D-glucal 122Figure 5.1. Structures ofCp2TiMe,heptenitol derivatives, and associated synthetic intermediates 135Figure 5.2. Structures of glucal derivatives and associated synthetic intermediates 140Figure A-I.1. Determination of Km and Vmax for f3G1cNAcPNP hydrolysis by K-NAGase and H-NAGase. ... 158Figure A-I.2. Estimation of K1 values for the inhibition of K-NAGase and H-NAGase by NAGlucal 159Figure A-I.3. Estimation of the K1 for the inhibition of J-NAGase by 2-cyanoglucal 160Figure A-lI. 1. Michaelis-Menten kinetics of an enzyme-catalyzed reaction 163Figure A-II.2. Double-reciprocal (or Lineweaver-Burk) plot of the Henri-Michaelis-Menten equation 164Figure A-II.3. Some graphical methods for distinguishing different types of reversible inhibition 169xLIST OF REACTION SCHEMESScheme 1.1. The hydrolytic and glycosyl transfer reactions catalyzed by glycosylases 1Scheme 1.2. The distinction between “retaining” and “inverting” glycosylases 2Scheme 1.3. The proposed mechanism for 13-glucosidase-catalyzed hydrolysis of 3-glucosides 4Scheme 1.4. Bacterial cell wail hydrolysis catalyzed by lysozyme 6Scheme 1.5. The proposed mechanism for the inactivation of 3-glucosidases by conduritol epoxides 8Scheme 1.6. The mechanism for the reaction of N-acetylconduramine B trans-epoxide with NAGase 10Scheme 1.7. The proposed mechanism for the slow inhibition of NAGase by N-acetylglucono-1,5-lactone 11Scheme 1.8. Isotope exchange between bridging and nonbridging oxygens through an enzyme intermediate. ... 14Scheme 1.9. The two forms of nojirimycin in aqueous solution at neutral pH 16Scheme 1.10. The proposed mechanism for the hydration of D-glucal inD20by f3-glucosidase 22Scheme 1.11. The hydration of D-galacto-octenitol by 3-galactosidase 24Scheme 1.12. The “proton transfer relay” for the deuteration of D-glucal by glycogen phosphorylase 27Scheme 2.1. The reaction catalyzed by glycogen phosphorylase 30Scheme 2.2. The proposed catalytic mechanism of glycogen phosphorylase 34Scheme 2.3. The proposed role of the phosphorylase cofactor PLP as a Brønsted acid catalyst 36Scheme 2.4. The proposed role of the phosphorylase cofactor PLP as an electrophule 38Scheme 2.5. The rapid equilibrium, random bi-bi mechanism for glycogen phosphorylase 39Scheme 2.6. Schematic representation of the ternary complexes of glycogen phosphorylase 40Scheme 2.7. The syntheses ofF2hept andF1hept 43Scheme 3.1. The synthesis of heptenitol from glucono-1,5-lactone using dimethyltitanocene 66Scheme 3.2. The f3-glucosidase-catalyzed hydration of heptenitol to form 1-deoxy-D-gluco-heptulose 68Scheme 3.3. The synthesis of methyiglucal 76Scheme 3.4. Possible mechanisms for the inactivation of pABG5 by nitroglucal 90Scheme 3.5. The synthesis of 1-cyanoglucal 92Scheme 3.6. The synthesis of 2-cyanoglucal 93Scheme 3.7. The selective deprotection of peracetylated 2-acetainidoglucal 100Scheme 4.1. The stereochemistry of I3G1cNAcPNP hydrolysis by 3-retaining and 13-inverting NAGases 111Scheme 4.2. The stereochemistry of the hydration of NAGlucal by NAGase 116Scheme 4.3. Comparison of the proposed mechanisms of 13-glucosidase and 13-N-acetythexosaminidase 126Scheme 5.1. A kinetic model for the inactivation of an enzyme (E) by an inactivator (I) 153xiABBREVIATIONS AND SYMBOLSAi) The absorbance (per unit pathlength) of a 0.1% (1 mg/mL) solution at 280 nm.A Absorbance at wavelength (where is given in nm).ADP Adenosme 5’-diphosphate.AMP Adenosine 5’-monophosphate.Asn L-Asparagine [(-i-)-3-aminobutanedioic acid 1-anude].Asp L-Aspartatic acid [(+)-2-aminobutanedioic acid].ATP Adenosine 5’-triphosphate.ax Axial.BES N, N-(Bis-2-hydroxyethyl)-2-aininoethanesulfonic acid.BSA Bovine serum albumin.C-terminal Carboxy terminal (end of a peptide or protein).conj. Conjugate.Cp Cyclopentadienyl.Cp2TiCl Titanocene dichloride.CpTiMe Dimethyl titanocene.DCI Desorption chemical ionization.1dfGlc l-Deoxy-f3-D-glucosylbenzene.cLDKJE c&Secondary deuterium kinetic isotope effect.DMF Dimethylformamide.D’IT Dithiotbreitol.E Enzyme.E.C. Enzyme Commission (classification number) of the International Union of Biochemistry.EDTA Ethylenediaminetetraacetic acid.(epsilon) Molar extinction coefficient at wavelength .eq Equatorial.ES Enzyme-substrate complex.F1hept 1-Fluoroheptenitol.F2hept 1, 1-Difluoroheptenitol.FAB Fast atom bombardment.2FI3G1cDNP 2’,4-Dinitrophenyl 2-deoxy-2-fluoro--D-glucopyranoside.G (or G+i) Glycogen.ctGlP a-D-Glucose-1-phosphate (oL-D-glucopyranosyl phosphate).I3G1cDNP 2’,4’-Dinitrophenyl f-D-glucopyranoside.GlcNAc N-Acetyl-D-glucosamine.I3GlcNAcMu 4’-Methylumbelliferyl 2-deoxy-2-acetamido-3-D-glucosaminide.GlcNAcPNP 4’-Nitrophenyl N-acetyl-f3-D-glucosaminide.fGlcPNP 4’-Niirophenyl f3-D-glucopyranoside.Glu L-Glutainic acid [(+)-2-aminopentanedioic acid].Gly Glycine [aminoethanoic acid].H-NAGase 3-N-Acetylhexosaminidase isolated from human placenta.HEPES N-[2-Hydroxyethyl]piperazine-N-[2-ethanesulfonic acid].His L-Histidine [(S)-2-amino-3-(4-imidazolyl)propionic acid].FIMPA Hexamethyl phosphoramide, (Me2N)P0.F[MPT Tris(dimethylamino)phosphine, (Me2N)3P.HPLC High-pressure liquid chromatography.I Inhibitor.IR Infrared (spectroscopy).xl’J Coupling constant.J-NAGase 13-N-Acetylhexosaminidase isolated from jack bean.K-NAGase 3-N-AcetyIhexosaminidase isolated from bovine kidney.kcat Catalytic rate constant (turnover number).Maximum first-order rate constant for inactivation.kobs Pseudo first-order rate constant for inactivation.K1 Dissociation constant for an enzyme-inhibitor complex.Km Michaelis constant of a substrate.Km,app Apparent Michaelis constant of a substrate (in the presence of an inhibitor).max Wavelength (nm) ofmaximum absorbance.Lys L-Lysine [(S)-2,6-diaminohexanoic acid].ManNAc N-Acetyl-D-mannosamine.mm Minutes.M Relative molecular mass.MS Mass spectrometry.N-terminal Amino terminal (end of a peptide or protein).3NADP 13-Nicotinamide adenine dinucleotide phosphoric acid.NAGase 3-N-Acetythexosaminidase.NAGlucal 2-Acetainido-D-glucal.NBS N-Bromosuccinimide.NMR Nuclear magnetic resonance.NOE Nuclear Overhauser effect.v (nu) Wavenumber.Nu Nucleophile.p Negative logarithm of (e.g., equilibrium constant, pK).p (or P) Phosphate.P Product.pABG5 Agrobacterium [3-glucosidase (cloned and expressed in E. coli).PL Pyndoxal.PLP Pyridoxal 5’-phosphate.Rf Retardation factor.Red Al® Sodium bis(2-methoxyethoxy) aluminum hydride.RF Range-finding (or approximate).Ser L-Serine [(S)-2-amino-3-hydroxypropionic acid].ThAF Tetra(n-butyl)ainmonium fluoride.TBDMS t-Butyldimethylsilyl.TEA Triethanolamine.THF Tetrahydrofuran.- (tilde) Approximately.TLC Thin-layer chromatography.U Units of enzyme activity.UV Ultraviolet (light).v Volume.v Reaction velocity.Vmax Maximal reaction velocity.VIS Visible (light).In most cases, the following were not listed:(1) SI units and prefixes.(2) Abbreviations and symbols used for systematic chemical nomenclature.xfflACKNOWLEDGMENTSI thank my supervisor, Prof. S. G. Withers, for his guidance and patience throughout the course of mydoctoral studies. I also thank Curtis Braun and Karen Rupitz for their technical assistance and adviceconcerning HPLC and enzyme kinetics, and Dr. Mark Namchuk and John McCarter for the syntheses ofI3G1cDNP and the peracetylated precursor of NAGlucal, tspectively. I am also very grateful for the advice andassistance of Dr. William Stirtan [who synthesized 1 -(methyl carboxylate glucal) and sodium 1 -(carboxylateglucal)], who was especially helpful with the work on glycogen phosphorylase, and Dr. Lothar Ziser, whogenerously shared his extensive knowledge of organic synthesis. I also thank Dr. S. Miao and Prof.R. Aebersold for their help with the protein mass spectrometry work. Dr. Qmgping Wang kindly informed meof her results prior to their publication, and I thank her for allowing me to compare some of her results withsome of my own. Thanks also to the other members of Prof. Withers’ laboratory for their valuable discussionsand helpful criticisms of various aspects of my work.Prof. A. Vasella of the University of Zurich generously provided 1-nitroglucal. Prof. L. N. Johnson andDr. E. Duke of Oxford University kindly agreed to collaborate with Prof. Withers and myself on the X-raystructural analysis of the binding of fluoroheptenitols in the active site of phosphorylase. I am most grateful tothese scientists for their thoughtfulness and assistance.Finally, I would like to thank my husband for his love and encouragement, and for typing thismanuscript. I also thank my parents for their patience and financial support, as well as their love andencouragement.xivDEDICATIONWith tlianfçfu(ness to qotifor my sear pareiits,Lai Wai 5(um atul Chung Sau Cñun.1CHAFfER 1: GENERAL INTRODUCTION1.1. GLYCOSYLASESAND THE REACTIONS THEY CATALYZE.Carbohydrates play central roles in energy metabolism, cell-cell recognition, and a variety of otherimportant biological processes, and glycosyl hydrolysis and glycosyl transfer are important biochemicalreactions. Glycosylases (or glycosyl mobilizing enzymes) are enzymes that utilize a glycoside (such asglycogen, a- or [3-glucosides, or 3-N-acetythexosaminides) as a substrate, and yield a product that contains aglycosyl residue. In the general reaction mechanism of these enzymes the aglycone moiety is replaced by eitherwater (for hydrolytic reactions) or some other glycosyl acceptor (i.e., in transfer reactions) to yield the product(see Scheme 1.1).Hydrolytic reactions OHOR1X = OH or NHAcV HOR2glycone aglyconeGlycosyl transfer reactions 0w.OR2xScheme 1.1. The hydrolytic and glycosyl transfer reactions catalyzed by glycosylases.For the enzymes studied in this thesis, the natural substrate substituents are:13-glucosidase: R1 = glucose (in cellobiose) or some other aglyconeglycogen phosphorylase: R1 = glycogen = phosphate3-N-acetythexosaminidase: R1 = a ganglioside X = NHAc (a 2-acetamido group)2The reaction mechanisms of three glycosylases were studied in this thesis: glycogen phosphorylase,3-glucosidase, and 3-N-acetylhexosaminidase (NAGase). Briefly, glycogen phosphorylase catalyzes theinterconversion of glucosyl-phosphate linkages (in a-glucose-i-phosphate, or aG1P) and glucosidic linkages (inglycogen, a highly branched homopolysaccharide of D-glucose units that is the major form of storagecarbohydrate in animal cells). f3-Glucosidase catalyzes the hydrolysis of (3-glucosides (e.g., cellobiose) to yieldglucose. NAGase catalyzes the hydrolytic removal of the N-acetylglucosamine (GIcNAc) residue fromgangliosides (oligosacchande-containing ceramide lipids). Some of the functions and specific features of eachenzyme will be discussed in subsequent chapters of this thesis.1.2. THE CLASSIFICATION OF GLYCOSYLASES.Glycosylases can be divided into several different classes based on several different characteristics.These enzymes can be classified as “retaining” or “inverting” according to the relative anomeric configurationsof the substrate cleaved and the product formed (see Scheme 1.2). Another classification is based on the sugar(glycone) moiety of the glycoside that the enzyme can accept as a substrate (e.g., glucosidases are most reactivetowards glucosides). Finally, these enzymes can be classified as “a” or “3” depending on the anomericconfiguration of the glycoside that the enzyme can accept as a substrate.__________io_______p._InversionOR2Scheme 1.2. The distinction between “retaining” and “inverting” glycosylases.3Of the enzymes studied in this thesis, glycogen phosphorylase is classified as an a-retainingglycosylase, whereas f3-glucosidase from Agrobacter sp. is 3 -retaining. In most cases the stereochemicaloutcome of the hydrolysis of 3-N-acetylglycosaminides by NAGases is not reported, and a determination of thestereochemistry of this reaction was in fact one of the objectives of this work.HO°[1.1]H([1.2]HThe anomeric specificity of glycosylases is typically absolute, e.g., a f3-glucosidase exhibits noenzymatic activity towards an ct-glucoside. However, certain compounds without a glycosidic bond—andtherefore lacking the appropriate a- or (3-aglycone group—can act as substrates for either a- or f3-glycosylases(or both). Examples of such nonglycosidic substrates are glycals (e.g., 1,5-anhydro-2-deoxy-D-hex-1-enitol, see1.1 above) and heptenitols (e.g., 2,6-anhydro-1-deoxy-D-gluco-hept-1-enitol, see 1.2 above) (Hehre et al., 1977).Examples of the corresponding reactions include the ability of glycogen phosphorylase to convert heptenitol(and phosphate) to 1-deoxy-D-gluco-heptulose-2-phosphate, and the ability of 3-glucosidase to hydrateheptenitol to form 1-deoxy-D-gluco-heptulose, and to hydrate D-glucal to form 2-deoxy-D-glucose.Glycosylases stereospecificafly catalyze these reactions with nonglycosidic substrates. The same stereochemicaloutcome (a or f3 product) is obtained from catalytic reactions with a glycal and with a natural substrate. Adetailed account of the glycosylase-catalyzed reactions of glycals is provided later in this chapter.1.3. GENERAL FEATURES OF THE CATALYTICMECHANISM OFRETAINING GLYCOSYLASES.1.3.1. Overview.Koshland (1953) was the first to propose that catalysis by retaining glycosylases involves a double-displacement reaction mechanism. This mechanism has subsequently received considerable experimentalsupport, and is now believed to include the following general features (see Scheme 1.3):HIH4AOH IHHO0HO ORHO-O 0I Enzyme IO 0\9/________________I Enzyme IOHHOO\HO-----.---OHHO-O 0I Enzyme I.OHHHO+HOI Enzyme Isyiation-ROH ifk1 A[Deglucosylation I k2+ H20LJAH HO 3+I;.’..QH+HOI Enzyme IScheme 1.3. The proposed mechanism for f-g1ucosidase-catalyzed hydrolysis of [3-glucosides.5(i) Acid catalysis promotes the departure of the aglycone group of certain substrates.(ii) In the active site of the enzyme, the carboxylate group of an acidic amino acid residue of the proteinis located next to the anomeric centre of the glycoside substrate. This carboxylate group is on theopposite side of the sugar ring relative to the aglycone.(lii) A covalent glycosyl-enzyme intermediate forms between the carboxylate of the enzyme and C-i ofthe sugar.(iv) Oxocarbonium ion-like transition states may be involved in both the formation and breakdown ofthe covalent glycosyl-enzyme intermediate.(v) Various noncovalent interactions accelerate the rate of the reaction.1.3.2. General acid catalysis.The departure of the aglycone leaving group is believed to be catalyzed by protonation of the sugar bythe side chain of an acidic amino acid residue in the enzyme’s active site. In hen egg white lysozyme, Glu 35has been identified as a catalytically important acidic residue by X-ray crystallography (Imoto et aL, 1972).However, the acidic residue may not need to be a carboxyl side chain.Additional evidence for the role of general acid catalysis was obtained from studies of theglycosylase-catalyzed hydration of glucal and heptenitol (Hehre et al., 1977; Schlesselmann et al., 1982). Thesestudies found that an acidic residue is essential for the protonation of the enolic double bond in such substrates,and that this protonation event is a prerequisite for subsequent nucleophilic attack by water (in hydrationreactions catalyzed by -glucosidase) or phosphate (in glycosyl transfer reactions catalyzed by glycogenphosphorylase) (Klein et aL, 1982).The importance of general acid catalysis in the reaction mechanism of glycosylases is variable, and insome cases it may not occur at all. For example, it is structurally impossible to protonate glycosyl pyridiniumsalts. Yet these compounds are hydrolyzed by glycosylases, and the observed rate increases for the hydrolysis ofthese compounds by 1 -galactosidase is 108 1013-fold compared with the rate of spontaneous hydrolysis(Sinnott, 1979). Clearly these dramatic rate increases are effected by a reaction mechanism that does not requiregeneral acid catalysis.61.3.3. The carboxylate group in the active site.a. X-ray structural studies.The first evidence for the presence of a strategically placed carboxylate group in the active site wasprovided by X-ray crystallographic studies of lysozyme, a glycosylase that catalyzes the hydrolysis ofpolysaccharides in the glycopeptide layer of bacterial cell walls (Scheme 1.4). X-ray diffraction methods havebeen used to determine the structures of the lysozymes of bacteriophage T4 and hen egg white (Anderson et al.,1981; Imoto et al., 1972), and strategically placed carboxylate groups were observed in the active sites of bothenzymes (the side chains of Asp 20 and Asp 52, respectively).4Scheme 1.4. Bacterial cell wall hydrolysis catalyzed by lysozyme.The glycosidic bond that is cleaved is indicated by the dotted lines.0•b. Fluoro sugars.Direct evidence for the role of a carboxylate group during catalysis was provided by Withers et al.(1990), who devised a novel class of inhibitors (exemplified by 2’,4’-dinitrophenyl 2-deoxy-2-fluoro--D-glucopyranoside, 2FI3G1cDNP, see 1.3 below) to trap and identify the amino acid residue involved. These“mechanism-based” inhibitors cause the enzyme to catalyze its own inactivation. The normal catalytic activitySSSSOSSNHpeptideSSSSSSNHpeptideSOHSSSS7of the target enzyme results in the formation of a covalent bond between the inhibitor molecule and acatalytically reactive residue in the enzyme’s active site.The substitution of an electron-withdrawing fluorine atom at C-2 in 2Ff3G1cDNP, immediately adjacentto the reaction centre at C-i, inductively destabilizes both transition states (see Scheme 1.3). This decreasesboth the rate of formation (k1) and the rate of hydrolysis (k2) of the glycosyl-enzyme intermediate. The presenceof a highly reactive leaving group (2,4-dinitrophenol) in the glycoside increases kj only (it does not affect k2).The combination of the inductive and leaving-group effects results in the accumulation of the glycosyl-enzymeintermediate. Thus when the radiolabeled, mechanism-based inhibitor [1-3H1-2Ff3G1cDNP was used withAgrobacterium [3-glucosidase, it identified Glu 358 as an active-site nucleophile of this enzyme (Withers et al.,1990).HOc. Conduritol epoxides.Similar evidence identifying the active-site nucleophile as a carboxylate moiety has been obtainedusing another class of inactivators of glycosylases. Conduritol epoxides (with the appropriate configuration,e.g., 1.4, conduritol B epoxide) are a class of synthetic mechanism-based inactivators that have been used in theaffinity labeling of active-site residues (an aspartate residue in each case) in three different retainingf3-glucosithses: enzymes isolated from the fungus Aspergillus wentii, bitter almonds, and lysosomes from humanplacenta (Bause & Legler, 1974; Dinur et al., 1986; Legler & Harder, 1978). D-Glucal has also been used tolabel the active-site nucleophile of Aspergillus wentii [3-glucosidase, and this study identified the same aspartateresidue as the active site nucleophile as was determined using conduritol B epoxide (Legler et al., 1979).FOH[1.3] [1.4] [1.5]OH8Conduritol C cis-epoxide (1.5) was used to identify Glu 461 as a residue that was thought to be theactive-site nucleophile of E. coli 13-galactosidase (Herrchen & Legler, 1984). However, a subsequent studyusing the tritiated, mechanism-based inhibitor [1-3H]-2FI3Ga1DNP identified Glu 537 as the active-sitenucleophile in this enzyme (Gebler et al., 1992). This latter study concluded that Gin 461 acts instead as ageneral acid-base catalyst during the reaction mechanism.Conduritol epoxides exploit the normal catalytic features of glycosylases to inactivate these enzymes(see Scheme 1.5). In the active site the side chain of an acidic amino acid (All) transfers a proton to the oxiranering of the inactivator. A carboxylate group of an amino acid in the enzyme’s active site then forms an esterbond with the activated oxirane, yielding a covalent enzyme-inactivator complex.OHHOHOO\,O 0\,0I Enzyme I I EnzymeScheme 1.5. The proposed mechanism for the inactivation of f3-glucosidases by conduritol epoxides.d. Cyclophellitol.Cyclophellitol (1.6) is a natural product that was initially isolated from culture filtrates of themushroom Phellinus sp. (Atsumi et al., 1990a), and subsequently prepared synthetically by Tatsuta et al. (1990).The structure of cyclophellitol differs from conduritol B epoxide (1.4) in that the former has a hydroxymethylgroup at C-5, and is therefore more similar to the structure of a [3-glucoside. Preliminary kinetic studies showedH09that cyclophellitol acts as an inhibitor of -glucosidases (Atsumi et al., 1990a, 1990b). More detailed kineticstudies with several cc- and 3-g1ucosidases showed that cyclophellitol is a highly stereospecific covalentmactivator of [3-glucosidases from almond and Agrobacter sp., with inactivation constants of [K1 = 0.34 mM,Ic1 = 2.38 min’] and [K = 0.055 mM, k1 = 1.26 mitf1], respectively (Withers & Umezawa, 1991). Bycomparison, conduritol B epoxide (1.4), which inactivates almond f3-glucosidase B with inactivation constantsof K1 = 1.7 mM, Ic1 = 0.13 min , is 92-fold less effective than cyclophellitol based on relative k1/K values(Legler & Hasnain, 1970). Due to the very high stereochemical specificity of cyclophellitol, no time-dependentinactivation of yeast a-glucosidase is observed, and only extremely slow inactivation (t112 > 5 hours) of E. coli3-galactosidase can be detected (Withers & Umezawa, 1991).OHHOHO[1.6]e. N-Acetylconduramine B trans-epoxide.The successful inactivation of various glycosylases by conduritol epoxides inspired the use ofN-acetylconduramine B trans-epoxide (1.7) in an attempt to inactivate NAGases from various sources (Legler &Bolthagen, 1992). Although this compound strongly inhibits NAGases isolated from bovine kidney, jack bean,and the gastropod Helix pomatsia (with K1 values of 0.50 to 1.6 IIM, i.e., 500-8,000-fold lower than the K1 forN-acetylglucosamine), no covalent inactivation is observed. An interesting phenomenon occurs when either ofthe first two of these NAGases is used at a reaction pH 5. Under these conditions the rate of substratehydrolysis in the presence of 1.7 increases with time. A proposed explanation for this result is given in Scheme1.6. At a reaction pH 5, these two NAGases catalyze the rapid conversion of the strong inhibitor 1.7 into anoxazoline (1.8; which has similar inhibitory potency to 1.7); however, 1.8 undergoes acid-catalyzed hydrolysisOH10to form the much less inhibitory compound N-acetylinosamine (1.9), and hence the rate of substrate hydrolysisin the presence of 1.7 increases with time (Legler & Bollhagen, 1992).I Enzyme I[1.8][1.9]OHHOScheme 1.6. The mechanism for the reaction ofN-acetylconduramine B trans-epoxide with NAGase.f Other inhibitors ofNAGases.Some indirect evidence for the participation of a catalytic carboxyl group in the reaction mechanism ofbovine kidney NAGase A was obtained from inhibition studies using N-acetylglucono-1 ,5-lactone (1.10) andN-acetylglucono-1,5-lactam (1.11) (Legler et al., 1991). Both the lactone and the lactam are good inhibitors ofthe enzyme, with K1 values of 0.036 and 0.67 j.iM, respectively. The lactam displays a normal approach to theinhibition equilibrium, whereas the lactone displays a slow approach. After the lactone is preincubated with thesubstrate, 4’-methylumbelliferyl 2-acetamido-2-deoxy-f3-D-glucopyranoside (3GlcNAcMu), the addition of theH0H,_HH_AHOI Enzyme I[1.7]FHBNH11enzyme catalyzes substrate hydrolysis at a rate that decreases slowly with time to a final value. The K1 for theinitial inhibition is about 50-fold higher than the final K1.OHCH3• 1+NCH3NHH3CCH3An explanation for (i) the much stronger inhibition displayed by the lactone (19 times stronger than thelactam), and (ii) the slow approach to the inhibition equilibrium of the lactone (1.10) compared with the lactam(1.11), has been proposed by Legler et al. (1991). They proposed that the lactone (1.10) is a better acceptor ofthe enzyme’s nucleophilic carboxylate group than the lactam (1.11). Thus the slow approach to the inhibitionequilibrium arises because of the formation of an orthocarboxylic acid derivative in the active site of the enzymeas a result of the attack on the lactone by a nucleophilic carboxylate group (see Scheme 1.7). However, no directproof of the existence of such a nucleophilic carboxylate group in the active site of NAGase has been obtained(e.g., by labeling and isolation), and the slow approach to the inhibition equilibrium may be due to reasons otherthan the formation of a covalent bond between the enzyme and the inhibitor (as discussed later in this chapter).HHO’-H3CC CH3CI Enzyme I I Enzyme IOHH0OHHO’H3C0[1.10] [1.111 [1.12]HO’OH OHNHScheme 1.7. The proposed mechanism for the slow inhibition of NAGase by N-acetylglucono-1 ,5-lactone.12Legler et al. (1991) have also studied the pH-dependence of the inhibition of bovine kidney NAGase Aby the cationic sugar 2-acetamido-1-deoxy-N,N-dimethylnojirimicin (1.12). The observed relationship betweenlog K1 and pH suggests that inhibition depends on the deprotonation of a functional group with a pKa near 5.0.This is probably a carboxylate group of a catalytic amino acid in the enzyme’s active site. However, no directproof of the existence of such a nucleophilic group has been obtained (e.g., by labeling and isolation).Compound 1.12 is useful for studying deprotonation of an active-site moiety because 1.12 possesses aquaternary amine. This removes any ambiguity as to whether the deprotonation event is attributable to acatalytically important amino acid residue of the enzyme, or a functional group on the inhibitor.1.3.4. The nature of the glycosyl.enzyme intermediate.a. Evidence from lysozyme andAgrobacterium /3-glucosidase (pABG5).A fundamental question concerning the nature of the glycosyl-enzyme intermediate is whether itinvolves an ion pair or a transient, covalent adduct. Based on evidence from X-ray structural studies of hen eggwhite lysozyme, Blake et al. (1967) suggested that a negatively charged Asp 52 carboxylate ion could stabilize apositively charged oxocarbonium ion intermediate, and that the lifetime of this ion pair could be long enough fora water molecule (or an alcohol) to attack the oxocarbonium ion. However, it is questionable whether the ionpair could last long enough for the departed leaving group to diffuse away and allow a glycosyl acceptor toapproach the active site and then react with the oxocarbonium ion.Convincing evidence that the reaction mechanism of glycosylases involves the formation of a covalentglycosyl-enzyme intermediate has been obtained from studies of the inactivation of Agrobacterium3-glucosidase (pABG5) with a novel class of mechanism-based inactivators, 2-deoxy-2-fluoro-3-D-glycosides,e.g., 2Ff3G1cDNP (Street, 1988). The reaction of pABG5 with 2Ff3G1cDNP permits the isolation of a2-fluoroglycosyl-enzyme intermediate whose half-life is over 500 hours. This covalent intermediate is stableenough to be characterized by various kinetic experiments and structural studies. 19F NMR studies of the2-fluoromannosyl-enzyme intermediate formed by 2FI3ManDNP and pABG5 showed that the sugar wascovalently bonded to the enzyme through an axial (CL-) glycosidic linkage (Withers & Street, 1988).13b. Evidence from f3-galactosidase.Additional evidence that the reaction mechanism of glycosylases involves the formation of a covalentglycosyl-enzyme intermediate has been obtained from cc-secondary deuterium kinetic isotope effect (aDKIE)studies using E. coli 3-galactosidase and a series of aryl galactosides (Sinnott, 1978). Reactions where theglycosylation step is rate-limiting yield ccDKIEs with ki-i/kD = 1.05, and reactions where the deglycosylation stepis rate-limiting yield aDKIEs with kH/kj 1.2-1.25. A normal aDKIE indicates that there is an increase in sp2character at the cc-carbon as the substrate changes from its sp3 ground state to the transition state, whereas aninverse aDKIE indicates that there is an increase in sp3 character at the a-carbon as the substrate changes fromits sp2 ground state to the transition state. Therefore the ccDKJE results of both types of studies (where theglycosylation or the deglycosylation step is rate-limiting) indicate that a covalent intermediate is formed, andthen hydrolyzed, via oxocarbonium ion-like transition states.c. Evidence from glycogen phosphorylase.A glucosyl-enzyme covalent intermediate has yet to be isolated for the glucosyl transfer reactioncatalyzed by glycogen phosphorylase. The complexity of the reaction mechanism—which requires the enzymeto form a ternary complex with both substrates (glycogen as well as phosphate or ccGlP)—and the rapidturnover of the enzyme make it difficult to accumulate sufficient amounts of the putative glucosyl-enzymeintermediate. To overcome this problem, Kokesh & Kakuda (1977) used cc-cyclodextrin (cyclohexaamylose, acyclic oligosaccharide composed of six glucose residues) as a substrate for potato phosphorylase. Althougha-cyclodextrin activates the enzyme ternary complex, it cannot act as a glucosyl acceptor due to the absence of afree hydroxyl group at C-4. This reduces the rate of turnover of any reaction intermediate that may be formed.The use of cc-cyclodextrin as a substrate analogue for potato phosphorylase allowed Kokesh & Kakuda (1977) todetect enzyme-catalyzed exchange between the bridging and nonbridging phosphoryl oxygen atoms of180-labeled ccGlP. This result is evidence for the involvement of a covalent glucosyl-enzyme intermediate inthe proposed catalytic mechanism (see Scheme 1.8).I Enzyme I<011.0 uHOHO HO180)O=P—OH0-I Enzyme IHO\/%JHOHOScheme 1.8. Isotope exchange between bridging and nonbridging oxygens through an enzyme intermediate.Withers & Rupitz (1990) have demonstrated the mechanistic similarity between potato phosphorylaseand rabbit muscle phosphorylase. This study used a series of deoxy- and deoxyfluoro-aG1P substrates in alinear free-energy study of the nature of the transition state of each enzyme. The reaction catalyzed by rabbitmuscle phosphorylase therefore probably involves the formation of a covalent glucosyl-enzyme intermediate.1.3.5. The oxocarbonium ion-like transition states of glycosylase substrates.a. a-Secondary deuterium kinetic isotope effects.The results of a-secondary deuterium kinetic isotope effect (xDKIE) studies indicate that bothtransition states in the proposed reaction mechanism of glycosylases have oxocarbonium ion-like character (seeH(14I Enzyme IOH INuO=P—OHEnzyme IOH INuH(0P—OH180-0P—OH180-15Scheme 1.3). As mentioned previously, these studies provide insights into the changes in the hybridization atthe reactive centre (the a-carbon) as it proceeds from the ground state to the transition state of the reaction. Theresults of Sinnott (1978) and Kempton & Withers (1992)—obtained using a series of aryl galactosides andE. coli 3-galactosidase, or a series of aryl glucosides and Agrobacterium 3-glucosidase, respectively—showedthat the (normal) aDKIEs observed when the deglycosylation step is rate-limiting are greater than thoseobtained when the glycosylation step is rate-limiting. Thus the second transition state has more oxocarboniumion-like character (i.e., involves a process that is less SN2-like) than the first transition state.b. Properties of transition-state analogues.A transition-state analogue is a tight-binding, reversible inhibitor whose structure resembles thetransition state of the enzyme-catalyzed reaction under study. The ability of transition-state analogues to bindmore tightly to the enzyme’s active site when compared with the binding of the normal substrate is aphenomenon that agrees well with the hypothesis that the structure of the enzyme’s active site is morecomplementary to the transition state than to the ground state of the substrate (Pauling, 1948).Studies performed using transition-state analogues have also provided evidence that the transition statesof glycosylase-catalyzed reactions have oxocarbonium ion-like character. The oxocarbonium ion form of theparent glycoside substrate can be distinguished from the latter in two ways. In the oxocarbonium ion form of theglycoside (i) the 0-5 and C-i atoms share a full positive charge, and (ii) the C-5, 0-5, C-i, and C-2 atoms arecoplanar (Sinnott, 1987). Compounds whose structures possess these properties have been shown to act asreversible inhibitors of glycosylases, and are classified as transition-state analogues for these enzymes (seeTable 1.1).c. Examples of transition-state analogues.An aldonolactone (or aldonolactam) is structurally similar to a glycosyl cation due to its half-chairconformation and the partial positive charge carried by the ring oxygen as a result of the conthbution of theresonance structure shown in Fig. 1.1. Glucono- 1,5-lactone and glucono- 1,5-lactam are transition-state16analogues that bind to A. wentii f3—glucosidase 100- to 300-fold more strongly than the corresponding aldoses(see Table 1.1).4 5[ ::b.I + 1 /\L.,0 NH NH ÷1Eoj +Aldonolactone glycosyl cation AldonolactamFigure 1.1. The structural similarities between the glycosyl cation and aldonolactones and aldonolactams.Nojirimycin and related compounds are transition-state analogues that are among the tightest-bindingglycosylase inhibitors known (Niwa et aL, 1970). These compounds contain a positively charged ring nitrogen.Nojirimycin may exist in different forms (see Scheme 1.9), each of which is able to act as a glycosylase inhibitorby forming an ion pair with a negatively charged residue in the active site (presumably an amino acid side chaincarboxylate) (Legler, 1990).OH+ OH-H20HO7\. +HOHO OH ÷H20OHScheme 1.9. The two forms of nojirimycin in aqueous solution at neutral pH.Several transition-state analogues have been examined as inhibitors of bovine kidney NAGase A(Legler et aL, 1991). When compared with G1cNAc (K1 = 1.97 mM), NAGase A was inhibited 2,600-fold morestrongly by N-acetyldeoxynojirimycin (1.14); 2,900-fold more strongly by N-acetylglucono-1,5-lactam (1.11);1755,000-fold more strongly by N-acetylglucono-1,5-lactone (1.10); and about 106-fold more strongly byN-acetylnojirimycin (1.13).OHHO’4NHHO.-w.OHNHo=zç[1.13]HO[1.14]HO°[1.15] [1.16]The transition-state analogues D-glucono-1,5-lactone (1.15) and nojirimycin tetrazole (1.16), with K1values of 25 and 14 j.tM, respectively, bind approximately 100-fold more strongly than glucose (K = 2 mM) tothe complex formed by glycogen phosphorylase, glycogen, and orthophosphate (Gold et a!., 1971; Withers &Rupitz, unpublished results). The strong binding of these compounds suggests that the reaction catalyzed byglycogen phosphorylase involves oxocarbonium ion-like transition states.Table 1.1. The reversible inhibition of glycosylases by some transition-state analogues.Enzyme Inhibitor K1 (suM) Reference13-glucosidase glucose 2800 Legler et al., 1980(A. wentii) 3-glucosylamine 1.6 ibid.glucono-1,5-lactone 9.6 ibid.glucono-1,5-lactam 36.0 ibid.nojirimycin 0.36 * Legler & Julich, 19843-N-acetythexosaminidase A N-acetylglucosamine 1970 Legler et al., 1991(Bovine kidney) 2-acetamido-2-deoxyglucosylamine 4.3 ibid.N-acetylglucono-1,5-lactone (1.10) 0.036 * ibid.N-acetylglucono-1 ,5-lactam (1.11) 0.67 ibid.N-acetylnojirimycm (1.13) 0.002 * ibid.N-acetyldeoxynojirimycin (1.14) 0.76 ibid.glycogen phosphorylase glucose 2000 Gold et al., 1971(Rabbit muscle) glucono-1,5-lactone (1.15) 25 ibid.nojiriinycin tetrazole (1.16) 14 Withers & Rupitz,unpublished resultsAn asterisk (*) denotes a slow approach to the inhibition equilibrium.18As was briefly mentioned earlier, the reaction of some glycosylases in the presence of certain inhibitorsis characterized by a slow approach to the inhibition equilibrium (see Table 1.1). This phenomenon is morefrequently observed with inhibitors that have a nitrogen atom in the sugar ring. When a slow approach to theinhibition equilibrium is observed, the K1 for the initial inhibition is about 50- to 100-fold greater than thefmal K1. One possible explanation for this phenomenon is that a loose complex is formed initially, which thenundergoes a slow conformational change to form a tight enzyme-inhibitor complex (Legler, 1990). Anotherpossible explanation is that the enzyme is capable of assuming two different conformations, characterized byeither a high or low affinity for the inhibitor, and that these conformations are in equilibrium. When a slowapproach to the inhibition equilibrium is observed, the inhibitor binds to the low-concentration, high-affinityconformer, which then shifts the confonnational equilibrium towards the high-affinity state (Legler, 1990).d. The transition states ofphosphoiylase-catalyzed reactions.The transition states in the glycogen phosphorylase reaction have been studied using deoxy anddeoxyfluoro analogues of the substrate cxGlP (Street et al., 1989). These derivatives are turned over by theenzyme, but at greatly reduced rates (1,000- to 10,000-fold slower than xG1P). These rate reductions have beenattributed to two effects: destabilization of transition states by the disruption of normal enzyme-ligandinteractions, and inductive destabilization of electron-deficient transition states by the introduction of anelectronegative fluorine atom adjacent to the reaction centre (as seen in the inactivation of Agrobacterium3-glucosidase by 2-deoxy-2-fluoro-J3-D-glucosides).A linear free-energy relationship, with a correlation coefficient p = 0.90, was obtained from alogarithmic plot of the rate constants for the enzyme-catalyzed reaction versus the nonenzymatic, acid-catalyzedhydrolysis of deoxy and cleoxyfluoro analogues of ctGlP (Street et al., 1989). This indicates that the electronicstructures of the transition states of the two reactions are similar. The mechanism of the nonenzymatic,acid-catalyzed hydrolysis of xG1P has been shown to involve transition states with substantial oxocarboniumion-like character (Bunton & Humeres, 1969). Thus the enzymatic reaction should also involve transition stateswith substantial oxocarbonium ion-like character, given the good correlation in the linear free-energyrelationship for the enzyme-catalyzed and nonenzymatic reactions.191.3.6. The binding energy attributable to noncovalent interactions.Pauling (1946) was the first to propose that most of the catalytic power of an enzyme comes fromnoncovalent interactions between the transition state of the reaction and the enzyme’s active site. Attempts toevaluate the importance of noncovalent interactions in the reaction mechanism of glycosylases have been carriedout using various substrate analogues in which an individual hydroxyl group has been replaced by a hydrogen ora fluorine atom.Noncovalent interactions between the C-2 hydroxyl group of the substrate and the active site ofA. wentii [3-glucosidase account for a 106-fold increase in the rate of glucoside hydrolysis, whereas the sameinteractions involving the C-4 hydroxyl group account for a104-105fold rate increase (Roeser & Legler, 1981).Similar studies by Namchuk (1993) using Agrobacterium 13 -glucosidase showed that the binding energyattributable to noncovalent interactions between the enzyme’s active site and different hydroxyl groups of thesubstrate decreased in the order C-2> C-3 > C-6 C-4.Deoxy derivatives of ctGlP have been used as substrate analogues in a study of noncovalentinteractions in the glucopyranose binding site of glycogen phosphorylase b (Street, 1988). This study showedthat hydrogen-bonding interactions between C-3 and C-6 hydroxyl groups and the enzyme’s active site areimportant in the stabilization of the transition state. Stabilization of the transition state lowers the activationenergy of the reaction and increases the reaction rate. The structure of the transition state of the reaction (not theground state) has the highest complementarity to the enzyme’s active site, and noncovalent interactions areclearly an important aspect of this complementarity.1.4. THE REACTIONS OF GLYCALS WITH GLYCOSYLASES.1.4.1. The enzyme-catalyzed hydration of glycals.Glycals, with their half-chair conformations, bear a structural resemblance to the transition states for theenzymatic hydrolysis of glycosides (see Fig. 1.2). These compounds were once considered to be transition-stateanalogues, and they have been investigated as inhibitors of glycosylases. Indeed, D-galactal was found to be avery strong inhibitor (K = 18-90 jIM) of several [3-D-galactosidases (Lee, 1969). However, a close inspection of20the structures of glycals and the transition states of glycosylase substrates shows that their similarities are limited(Fig. 1.2). In addition to the absence of a hydroxyl group at C-2, glycals have their double bond located betweenC-i and C-2, whereas the double bond of the transition state of a glycosylase substrate is located between C-iand 0-5. The stereochemistry of some of the ring substituents on each structure also differs (see Fig. 1.2).These considerations prompted more detailed kinetic studies of the mechanism of the inhibition ofE. coli (3-galactosidase by D-galactal (Wentworth & Wolfenden, 1974). The extent of the steady-state inhibition(K = 14 jiM) was interpreted as the result of the slow formation of a covalent, 2-deoxy-galactosyl-enzymeintermediate, and the subsequent hydrolysis of this adduct to form 2-deoxy-(3-galactose (keat 0.004 sd).However, under pre-steady-state conditions unmodified (i.e., unreacted) D-galactal is only a weak inhibitor(K1> 1 mM). It is noteworthy that the 2-hydroxyl group of normal substrates seems to play an important role inincreasing the rate of enzymatic hydrolysis (Namchuk, 1993).HOOHHOHO° C C,_(OH—ROH HOFigure 1.2. The structural similarities between galactal and the transition state of a (3-galactoside.Subsequent research has shown that glycals generally act as substrates, and that the hydration of theenolic double bond in glycals is catalyzed by both c’- and f3-glycosylases. Examples of this catalytic activityinclude the hydration of D-glucal by both Candida tropicalis a-glucosidase and sweet almond (3-glucosidase21(Hehre et a!., 1977), the hydration of maltal by sweet potato 3-amylase (Hehre et a!., 1986), and the hydration ofcellobial by exo- and endo- type ceiltilases (Kanda et aL, 1986).The hydration of glycals (I) by glycosylases (E) can be represented by the following reaction scheme(Legler, 1990):K1’ icon H20E + I ... E.I .. E—I E + 2-deoxyhexosek0ff khydrowhere K1’ is the dissociation constant of a loose, rapidly formed, pre-steady-state complex (E.I). The inhibitionconstant for the steady state, K, is:Kj’K =______________11 ic01 (k0ff+ hy)and the rate constant for the approach to the steady state, kpr, is:• [IIkappr =____ __+ k0ff + khydroK1’ + [IJIn the presence of substrate, (S), with a Michaelis constant, Km, as defined in the case of inhibition studies, then:I [SI 1 1k011’[IIkar 1 +— .= + k0ff + khydroKm JFrom the above scheme, depending on the magnitude of the kinetic parameters k0,k0’, and khyo, differentcombinations of glucal—glycosylase interactions are possible.For A. wentii f3-glucosidase the sum of andk0ff is smaller than k0, and this together with the useof [14C1-labeled D-glucal made it possible for Legler et a!. (1979) to trap the covalent glucal-enzymeintermediate before glucal was turned over to 2-deoxyglucose. Before the enzyme had time to turn over the[14C1-glucal the protein was rapidly denatured, and a catalytic aspartate residue was identified by subsequentproteolytic digestion and amino-acid sequencing of the radiolabeled peptide. The same nucleophilic aspartateresidue that is involved in the catalytic hydration of glucal by 3-glucosidase was also labeled by the irreversible22inhibitor conduritol B epoxide (Bause & Legler, 1974) and the slow substrate p-nitrophenyl-2-deoxy-f3-D-glucopyranoside (Roeser & Legler, 1981).The stereochemistry of the enzyme-catalyzed hydration and protonation at C-2 of glycals has beeninvestigated with cx- and f3-glucosidases (Hehre et al., 1977) and with three “inverting” exo-a-glucanases (Chibaet al., 1988). These reactions result in the production of 2-deoxy-cc-glucose by cx-glucosidases, whereas2-deoxy-fi-glucose is produced by fi-glucosidases and inverting exo-cc-glucanases. These are the samestereochemical outcomes as observed with the glycosylase-catalyzed hydrolysis reactions of glycosides.I NMR studies of the catalytic hydration of glycals inD20 (where exchangeable protons are replacedby deuterium) showed that retaining a-glucosidases protonate D-glucal from above the double bond, whereas13—glucosidases and inverting exo-cc-glucanases protonate D-glucal from below the double bond. Thus thedirection of protonation of the double bond in D-glucal is opposite from the direction that would be expectedbased on the known stereochemistry of the glycosylase-catalyzed hydrolysis reactions of glycosides.OHHOR OD- HO—------.7D7/,,deglycosylationHOHYglycosylationScheme 1.10. The proposed mechanism for the hydration of D-glucal inD20by f3-glucosidase.23The available kinetic data on the glycosylase-catalyzed hydration reactions of glycals shows thatalthough the initial protonation step appears to differ significantly from that seen in the glycosylase-catalyzedhydrolysis reactions of normal glycosides, the remainder of the reaction mechanism is the same, involving thesame active site residues and yielding products with the same configuration. Thus in the proposed mechanismfor the catalytic hydration of glycals (Legler et al., 1979), carboxylate groups act as both a general acid catalystand a nucleophile (Scheme 1.10).1.4.2. The enzyme-catalyzed hydration of heptenitols.D-gluco-heptenitol (1.2) is a glycal with an exocydic enolic double bond. In general, heptenitols donot inhibit glycosylases as effectively as hexenitol glycals (glycals with an endocyclic enolic double bond).Glycosylases are able to hydrate D-gluco-heptenitol to form 1-deoxy-D-gluco-heptulose (1.17). [Note that inboth (1.2) and (1.17) the exocyclic carbon atom is designated as C-i.] However, if the reaction is performed inthe presence of an alcohol, a glycoside may be formed. Thus -galactosidase is able to synthesize glyceryl2,6-anhydro-i-deoxy--D-galacto-heptuloside (1.18) from D-galacto-hept-1-enitol and glycerol. The initialanomeric configuration (i.e., prior to subsequent ct/13 equilibration) of the product of the catalytic hydration (orglycoside synthesis reaction) of a heptenitol is determined by the anomeric specificity of the glycosylase.HO”CH3[1.2] [1.17] [1.18]The greater catalytic activity of glycosylases towards heptenitols compared with hexenitol glycals hasbeen attributed to the greater structural resemblance of heptenitols to the normal glycoside substrates of theseHO24enzymes. In the three-dimensional structure of heptenitol there are four hydroxyl groups that can assumepositions that are equivalent to the hydroxyls of x- or 3-D-glucoside substrates, and this increased structuralsimilarity may allow for better binding of heptenitols in the active site. In addition, the exocyclic double bond ofa heptenitol occupies a position in the active site that is extremely close to the position normally occupied by theglycosidic bond of regular glycoside substrates, and hence the double bond of the heptenitol is susceptible to thesame catalytically important protonation event (i.e., general acid catalysis).The catalytic hydration of D-galacto-heptenitol by E. coli f3-galactosidase has a Michaelis constant,Km, of 50-70 mM, and a turnover number, kcat, of 41 to 64 s (Brockhaus & Lehmann, 1977). Similar kineticdata have been obtained for the catalytic hydration of D-gluco-heptenitol by c-glucosidase from C. tropicalis(rice), sweet almond f3-glucosidase, and an inverting exo-a-glucanase from A. globiformis (Hehre et al., 1980).The stereochemistry of the hydration of heptenitols by retaining cx-glucosidases has been studied using2-[H]-D-gluco-octenitol. Enzymes from A. niger and C. tropicalis were used in this study, and both protonatedthe substrate from the top (Si, re) face, the same direction of protonation that several a-glucosidases use toprotonate D-glucal and D-galactal (Weiser et al., 1988), but the opposite direction of protonation used by(x-glucosidases with normal glycoside substrates.HO OHH- HO_0H [1.20]HO.cD H3C’DScheme 1.11. The hydration of D-galacto-octenitol by f3-galactosidase.In contrast, when2-[H}-D-galacto-octenitol (1.19) is hydrated by E. coli f3 -galactosidase, thestereochemistry for the protonation of the double bond is the same as that observed with normal glycosidesubstrates (Lehmann & Schlesselman, 1983). During the catalytic hydration of 1.19, protonation is exclusivelyfrom above the ring, and the hydration reaction yields 1.20, 1,2-dideoxy-2-deutero-D-galacto-octulopyranose[1.19]25(see Scheme 1.11). The two different directions that can be taken for the protonation step implies that theprotonatmg amino acid residue may be located on either face of the ring of a bound substrate in the active site ofa particular glycosylase enzyme.The results of stereochemical investigations of the glycosylase-catalyzed hydration of hexenitol andheptenitol glycals suggests that there are two mechanistically distinct and separately controlled aspects of thesereactions (Chiba et aL, 1988). In the so-called “plastic” aspect of the reaction, various glycosylases showconsiderable heterogeneity in the direction of protonation, a phenomenon that is substrate-dependent. In theso-called “conserved” aspect of the reaction, the creation of a product with a specific configuration is aconsequence of the acceptor’s direction of attack against the reaction centre. The latter phenomenon is mainlydetermined by the particular enzyme, with little influence by the substrate.1.43. The reactions of glycals with glycogen phosphorylase.Glycogen phosphorylase catalyzes the addition of D-glucal (as a 2-deoxyglucosyl residue) to thenonreducing end of a polysaccharide composed of glucose subunits:glucal + (glucose) ‘ (2—deoxyglucose)—(glucose)One of the products of the reaction is also 2-deoxy-xG1P, which is probably formed by the enzymaticphosphorolysis of the modified polysaccharide (Klein et al., 1982). The maximum rate of utilization ofD-glucalis approximately 20-30% of the rate of glycogen synthesis using aG1P.In the presence of arsenate (i.e., in place of orthophosphate), phosphorylase catalyzes the arsenolysis ofD-gluco-heptenitol to form 1-deoxy-D-gluco-heptulose-2-arsenate, which undergoes rapid spontaneoushydrolysis with retention of configuration to form 1-deoxy-cz-D-gluco-heptulose. The rate of the phosphorylasecatalyzed formation of 1-deoxy-c&D-gluco-heptulose (in the presence of arsenate) with 50 mM heptenitol is17-32% of the rate of arsenolysis of polysaccharides, depending on the source of the enzyme (Klein et al., 1986).In the presence of orthophosphate, phosphorylase converts D-gluco-heptenitol to form 1-deoxy-a-D-gluco-heptulose-2-phosphate (heptulose-2-phosphate, 1.21), which binds tightly to the enzyme and acts as aninhibitor (with K1 = 14 IIM) (Klein et al., 1984). To date, heptenitol is the only known substrate of26phosphorylase that can be utilized in the absence of a polysaccharide primer. The rate of this reaction cannot bemeasured accurately due to the formation of the “dead end” inhibitor, heptulose-2-phosphate.In the presence of glycogen and arsenate (i.e., with arsenate acting as a phosphate analogue), heptenitolacts as a competitive inhibitor (with K = 4.3 mM) of rabbit skeletal muscle phosphorylase b (Klein et al., 1986).However, transfer of the heptulosyl residue to a polysaccharide acceptor is not detectable. Therefore heptenitolis used only as a substrate for the degradative pathway of the phosphorylase reaction.Table 1.2. Comparison of the binding constants of some ligands of glycogen phosphorylase b.Ligand K (or I() (pM) Referenceglucose-i-phosphate 2,200 Hu & Gold, 1978glucose-i,2-(cyclic phosphate) [1.22] 500 ibid.i-deoxy-heptulose-2-phosphate [1.21] 14 Klein et al., 1984(also referred to as heptulose-2-phosphate)OH OHHO—P o0—[1.211 [1.22]Heptulose-2-phosphate binds to phosphorylase b with an affinity that is 100-fold greater than thesubstrate xG1P (see Table 1.2). X-ray structural analysis of the (heptulose-2-phosphate)—[phosphorylase bjcomplex showed that the phosphate group of the inhibitor was located underneath the sugar ring, orientedtowards the C-3 hydroxyl group, probably to avoid steric hindrance from the C-i methyl group (McLaughlin etal., 1984).27Heptulose-2-phosphate (1.21) and glucose-1,2-(cyclic phosphate) (1.22) are considered to betransition-state analogues for the degradative reaction of glycogen phosphorylase because of their chemicalstructures and low values (Hu & Gold, 1978). The preferred orientation of the phosphate group of bothcompounds is under C-2 of the sugar ring, in contrast with aG1P with its phosphate group away from the ring(trans to C-2) (O’Connor et al., 1979).The stereochemistry of the protonation of D-glucal by glycogen phosphorylase b is from below thesugar ring (Klein et al., 1982). Thus if the reaction is carried out in a deuterated buffer, deuterium isincorporated into the equatorial position at C-2—the opposite stereocheniical outcome to that observed whenhexenitol glucals are protonated by other ce-retaining glycosylases. Klein et al. (1982) also found that inaddition to an absolute requirement for phosphate (or arsenate), the catalytic reaction of D-glucal is alsodependent on the dianionic form of the enzyme-bound cofactor, pyridoxal phosphate.CD L%.r%._J CD- B tran.er to a. saccharideacceptorDC\/0 0-j] OD OzPL—O—P—OzPL—O—P—0yl II y IIml mjY = P or As B = nucleophile (or base) PL = pyridoxalDOH+D HEScheme 1.12. The “proton transfer relay” for the deuteration of D-glucal by glycogen phosphorylase.28On the basis of these observations, Klein et al. (1982) have proposed that a “proton transfer relay” is afeature of the enzyme’s reaction mechanism (Scheme 1.12). Orthophosphate (or arsenate) is located below theD-glucal ring, in a region of the active site that is adjacent to the phosphate group of the enzyme’s cofactor. Theenzyme-bound cofactor pyridoxal phosphate is believed to mediate a proton transfer relay between the enzymeand D-glucal as shown in Scheme 1.12.1.5. THE AIMS OF THIS THESIS.1.5.1. Research significance and objectives.Both reversible and irreversible inhibitors (or inactivators) are useful in studies of enzyme reactionmechanisms and the properties of transition states. These compounds may also find applications as therapeuticdrugs. Identification of catalytic residues in the enzyme’s active site, and gaining an understanding of theirspecific roles in catalysis, are also possible using irreversible inhibitors. The aims of this thesis were tosynthesize several derivatives of D-gluco-heptenitol and D-glucal, and examine the potential utility of thesecompounds as mechanism-based inhibitors of three glycosylases: glycogen phosphorylase, f3-glucosidase, and3-N-acetythexosaininidase.1.5.2. Studies using fluorinated derivatives of D-gluco-heptenitol.Withers et al. (1990) have shown that 2-fluoro-glucosides rapidly inactivate I3-glucosidase, and thatsuch compounds are useful probes for studying the reaction mechanism of this enzyme. Theelectron-withdrawing fluorine atom inductively destabilizes the oxocarbonium ion-like transition states, and theoverall effect of these inhibitors is the covalent inactivation of the enzyme by the accumulation of theglucosyl-enzyme intermediate. Fluorinated heptenitols, with one or two electron-withdrawing fluorine atomssituated adjacent to the reaction centre, were therefore considered to be potential inactivators of glycosylases.One advantage of these compounds over 2-fluoro-sugars is that the former can be used to study the effect ofvarying the degree of withdrawal of electron density from the reaction centre. Hence difluoroheptenitol29(F2hept) and monofluoroheptenitol (Fihept) were synthesized, and kinetic studies of these compounds wereearned out using glycogen phosphorylase and f3-glucosidase.Fluoroheptenitols also proved to be useful for structural studies of glycogen phosphorylase. It hadpreviously been established that glycogen phosphorylase is able to convert heptenitol in the presence ofphosphate to 1-deoxy-heptulose-2-phosphate (Klein et al., 1984). The synthesis of F2hept and Fihept—compounds that are not turned over by the enzyme as rapidly as heptenitol—assisted collaborators in the U.K.,who performed X-ray structural studies using crystals of the enzyme containing either of the fluoroheptenitols.These studies yielded insights on the binding of phosphate in the enzyme’s active site, studies that could not beperformed with heptenitol.1.5.3. Studies using derivatives of D-glucal.An initial aim of this study was to determine whether methylglucal might act as a substrate, or possiblyeven an inhibitor or inactivator, of -glucosidase. If promising results were obtained with the parent compound,the synthesis of fluorinated derivatives of methyiglucal would be attempted, and their effects on the activity ofthe enzyme would also be studied to provide more insight into the role of inductive effects on glycal hydration,or possibly the utility of fluorinated glycals as inhibitors or inactivators.Another aim of this study was to examine whether c,f3-unsaturated glucals, which can act as Michaelacceptors for the nucleophiic residue present in the active site of f3-glucosidase, might constitute a new class ofinactivators of this enzyme. 1-Nitroglucal and other x,f3-unsaturated glucals [1-cyano-, 1-(methylcarboxylate)-,sodium 1-(carboxylate)-, and 2-cyano- glucal] were therefore examined as potential inhibitors of f3-glucosidase.Finally, 2-acetamidoglucal (NAGlucal) was used to study the reaction mechanism of the clinicallyimportant enzyme f3-N-acetythexosaminidase (NAGase). Stereochemical studies and other kinetic experimentswere performed on NAGases isolated from jack bean, bovine kidney, and human placenta. The stereochemistryof the hydrolysis of the commonly used substrate GlcNAcPNP (4’-nitrophenyl N-acety1-3-D-glucosaminide)was also investigated.30CHAPTER 2: KINETIC STUDIES USING GLYCOGEN PHOSPHORYLASE2.1. INTRODUCTION.2.1.1. Biochemical role in vivo.Glycogen phosphorylase (cx-1,4-glucan-orthophosphate glucosyl iransferase, E.C. 2.4.1.1) plays a keyrole in carbohydrate metabolism. It is found in all cell types, from microbial species to the various complextissues of higher plants (starch phosphorylase) and mammals. Glycogen phosphorylase catalyzes the reactionshown below, where the terminal a(1-4) glucosidic linkage at the nonreducing end of a glycogen side chainundergoes phosphorolysis. The cleavage of the terminal glucosidic bond results in the removal of the terminalglucose residue as x-D-glucose-1-phosphate (cxGlP), leaving behind a glycogen chain with one less glucoseunit. Although glycogen phosphorylase catalyzes the formation and breakdown of glycogen, its primaryfunction in vivo is to catalyze the breakdown of glycogen and provide a regulated supply of xG1P.I HOo — Phosphorolysis— —Synthesis IHOOHH0HO glycogen HO glycogenOHOHScheme 2.1. The reaction catalyzed by glycogen phosphorylase.312.1.2. Regulation.Rabbit muscle glycogen phosphorylase was first isolated and characterized by Con & Con (1936).This enzyme has been studied extensively, and has been shown to be subject to complex regulatory mechanisms.The activity of this enzyme is carefully controlled by covalent modification (phosphorylationdephosphorylation), as well as an allosteric regulatory mechanism (ligand-mduced conformational changes).Glycogen phosphorylase exists in two forms; the less active phosphorylase b and the more activephosphorylase a. Phosphorylase b is converted to the “a” form by phosphorylase kinase, which phosphorylatesthe enzyme at Ser 14 (Fischer & Krebs, 1955). A phosphoprotein phosphatase dephosphorylates the phosphateester at Ser 14 in phosphorylase a, thereby converting the enzyme back to phosphorylase b (Wosilait &Sutherland, 1956).Phosphorylase b assumes at least two conformations, the catalytically active R-conformation, and theinactive T-conformation. Phosphorylase b is activated by adenosine-5’-monophosphate (AMP) and is subject toallosteric inhibition by adenosine-5-triphosphate (ATP), glucose, adenosine-5’-diphosphate (ADP) andglucose-6-phosphate (Madsen & Shechosky, 1967). Phosphorylase a exists mainly as the active R-conformer,and is not subject to aflosteric regulation (Madsen & Withers, 1986).2.1.3. Structural studies.Glycogen phosphorylase can exist as a tetramer or a dimer in vivo, but the catalytically active R-statesof phosphorylase a orb exist as dimers (Metzger et aL, 1967). Each monomer has an Mr of 97,444, and consistsof 842 amino acids (Johnson, 1992). The amino acid sequence of rabbit muscle phosphorylase was determinedby Titani et al. (1977). The tertiary structure of the monomer can be divided into N-terminal (residues 1-489)and C-terminal (residues 490-842) domains. In each domain, approximately 45% and 25% of the amino acidresidues are in a-helical or 13-sheet secondary structural elements, respectively.The X-ray crystal structures of several forms of glycogen phosphorylase have been determined, andthese studies have identified the primary ligand binding sites on the enzyme (Acharya et al., 1991; Sprang &Hetterick, 1979; McLaughlin et al., 1984). The catalytic site (site C) is located at the centre of the enzyme,32Glycogenstoragsite IGIFigure 2.1. The structure of the glycogen phosphorylase b monomer.Towercontact to .Ilost.rICof ofher subunitphosphat.The structure shown is from McLaughlin et al. (1984). a-Helices and f3-strands are represented by cylinders andarrows, respectively. Please see the text for descriptions of the primary ligand binding sites.33near the boundary between the N-terminal and C-terminal domains (Madsen & Withers, 1986). The active siteconsist of the following elements:1. A highly specific pocket for the glucopyranosyl moiety of the substrate, cxGlP. The inhibitorglucose also binds here.2. A phosphate subsite for the binding of the phosphate moiety of ctGlP.3. A cofactor subsite for the binding of pyridoxal-5’-phosphate (PLP). PLP is covalently bound to theenzyme via a Schiff base with Lys 680. The phosphate group of PLP is adjacent to the phosphatesubsite of the substrate (Madsen & Withers, 1986).At the entrance to the catalytic site there is an inhibitor binding site (site I) where aromatic inhibitorssuch as caffeine, and various nucleosides and nucleotides, bind to the enzyme. About 32 A from site C, nearSer 14 and close to the subunit-subunit interface, there is an allosteric effector binding site (Site N) where AMPbinds to the enzyme. The glycogen storage site (site G) is located on the surface of the N-terminal domain of theenzyme. Site G is located about 30 A and 39 A from the catalytic and allosteric effector sites, respectively. Theenzyme attaches to the glycogen particle via site G, and smaller oligosaccharides such as maltoheptaose alsobind strongly to this site (McLaughlin et al., 1984).2.1.4. General features of the catalytic mechanism.Phosphorylase is an a-retaining glycosylase, and hence the reaction it catalyzes is thought to involve adouble-displacement mechanism (see Scheme 2.2). The main features of the proposed catalytic mechanism (inthe direction of glycogen synthesis) include: general acid catalysis (to facilitate cleavage of theglucosyl-phosphate linkage); formation of a glucosyl-enzyme intermediate (either covalently orelectrostatically); and general base catalysis to facilitate nucleophilic attack (by the 4-hydroxyl group ofglycogen’s terminal glucose residue) directed against the anomeric centre of the glucosyl-enzyme intermediate.These features of the reaction, and the experimental support for their occurrence, were reviewed in Chapter 1 ofthis thesis. Other important features of the catalytic mechanism of glycogen phosphorylase include the role ofthe cofactor, pyridoxal phosphate (PLP), and the fact that the enzyme catalyzes a two-substrate reaction.34HO#’..(NUOHHO HO?HO”\ç — P=0 HO—AH —O glycogenHONuHO HOA-HO\glycogenHO—P=O—O—0+ HO—P0—OI glycogenScheme 2.2. The proposed catalytic mechanism of glycogen phosphorylase.The flow of electrons is shown for the glycogen synthesis reaction only, but it should be noted that the reactionis reversible.OH NuHOOHAH352.1.5. The cofactor pyridoxal phosphate (PLP).a. Structural and catalytic roles.One molecule of PLP is present in each phosphorylase monomer. PLP is attached to the enzyme by aScuff base involving Lys 680, and this adduct plays an important role in maintaining the quaternary stnicture ofthe enzyme through subunit interactions (Kastenschmidt et aL, 1968). PLP also plays an important role duringcatalysis. Removal of PLP yields the apo-enzyme, which has no catalytic activity, but catalytic activity can berestored by reconstitution (Iuingworth et al., 1958).aminoacid68l I0—\O0—Figure 2.2. The Schiff base formed between Lys 680 and the cofactor pyridoxal phosphate (PLP).In the proposed reaction mechanism of phosphorylase, the 5’ phosphate group of PLP is believed to actas either a BrØnsted acid catalyst (Klein et al., 1982, 1984; Palm et al., 1990) or as an electrophile, with the5’ phosphate of PLP remaining in the di-anionic form throughout the course of the reaction (Withers et al., 1982)(see Schemes 2.3 and 2.4 for the mechanistic details of how PLP might act as an acid catalyst or as anelectrophile, respectively). Although researchers agree that PLP plays a crucial role during phosphorylasecatalysis, there is a lack of agreement on the precise details of the role this cofactor plays during the reactionmechanism.36c-OOHHO\c-glycogenn glycogenOHQ% OHP_4O 0— glycogenThe cofactor pyridoxal phosphate(attached to Lys 680)}Scheme 2.3. The proposed role of the phosphorylase cofactor PLP as a Brønsted acid catalyst.Adapted from Palm et al. (1990).37b. The proposed role ofPLP as a BrØnsted acid catalyst.In this proposal, the phosphate group of PLP acts as a proton donor and acceptor, as shown in Scheme2.3 (Helmreich, 1992; Palm et al., 1990). When the reaction proceeds in the direction of glycogen synthesis, thephosphate group of PLP first acts as an acid catalyst and protonates ctGlP. This protonation event facilitatescleavage of the glucosidic bond, which in turn facilitates the general base-catalyzed attack against theglucosyl-enzyme intermediate by the 4-hydroxyl group of the terminal glucosyl residue of the glycogen particle.When the reaction proceeds in the direction of glycogen phosphorolysis, the phosphate group of PLPprotonates the bound phosphate substrate, which in turn protonates the glucosidic linkage at the terminalglucosyl residue of the glycogen particle. The phosphate group of PLP subsequently acts as a base catalyst thatfacilitates nucleophilic attack against the glucosyl-enzyme intermediate by phosphate. This mechanismprobably involves oxocarbonium ion-like transition states (Helrnreich, 1992).c. The proposed role ofPLP as an electropizile.An alternative proposal for the role of the phosphorylase cofactor PLP is that this moiety acts as anelectrophilic catalyst (reviewed in Madsen & Withers, 1984, 1986). This proposal states that in the activeternary complex the geometry of the 5 phosphate of PLP is distorted (by basic amino acid side chains) into atrigonal pyramidal configuration, with an empty apical position oriented towards the substrate phosphate (seeScheme 2.4). The interaction of the substrate phosphate with this PLP electrophile labiizes the glucosidic bondvia the transient formation of a partial pyrophosphate bond. In the proposed mechanism the transition stateinvolves both oxocarbonium ion-like sugar and trigonal bipyrantidal phosphate species.2.1.6. Glycogen phosphorylase catalysis as a rapid equilibrium, bireactant system.Initial-rate, binding, and inhibition studies, as well as studies of isotopic exchange at equilibrium, haveall demonstrated that glycogen phosphorylase is a two-substrate enzyme with a rapid equilibrium, random bi-bireaction mechanism (Engers et al., 1969, 1970; Gold et al., 1970; Maddaiah & Madsen, 1966) (see Scheme 2.5).HO(NU26-8—16+HO—P—0O68 Q—V6iNHO CH338—O—P=OHONuOHHO HOOHglycogenglycogenHO CH3OH0II+ HO—P—0glycogen 0The cofactor pyridoxal phosphate(attached to Lys 680)}HO CH3Scheme 2.4. The proposed role of the phosphorylase cofactor PLP as an electrophile.39E—G E—G(fl+1)/_/\E E—G—GP E—G(fl+l)—P EE—GP E—PScheme 2.5. The rapid equilibrium, random bi-bi mechanism for glycogen phosphorylase.Adapted from Engers et al. (1969). The symbols are defmed as follows:E = Enzyme E—G—GP = ternary complexE—G = Enzyme—glycogen complex E—G(n+1)--P = ternary complexE—GP = Enzyme—c.G1P complex E—P = enzyme—phosphate complexIn the glycogen synthesis direction of the proposed reaction mechanism, glycogen phosphorylase firstbinds to either one of its two substrates (G or GP in Scheme 2.5). Either of these two binary complexes maythen bind to the remaining substrate. The result is a ternary complex (E—Gn—GP in Scheme 2.5) that may beformed randomly in either of the two pathways shown (on the left-hand side of Scheme 2.5).In the opposite direction—glycogen phosphorolysis—the enzyme first binds either of its two substrates(G(n+1) or P in Scheme 2.5), and then either of these two binary complexes may bind to the remaining substrate.The result is a ternary complex (E—G(n÷1)—P in Scheme 2.5) that may be formed randomly in either of the twopathways shown (on the right-hand side of Scheme 2.5).Both of the ternary complexes (i.e., formed from either direction) are interconvertible (see Scheme 2.6),and this interconversion is the rate-limiting step of the reaction. The fact that the isomerization of the ternarycomplex is rate-limiting indicates that this step is slow compared with the formation of the product. However,40no product may be formed until after the ternary complex has formed. To simplify matters, in most kineticstudies (including those reported herein) one of the substrates (e.g., glycogen) is kept at saturatingconcentrations, and under these conditions the model reduces to a single-substrate system.Scheme 2.6. Schematic representation of the ternary complexes of glycogen phosphorylase.Synthesis modeiiPhosphorolysis modeThe two modes of glycogen binding are mutually exclusive. Circles represent glucosyl residues. Other symbolsare defined in Scheme 2.5. Adapted from Segel (1975).2.2. THE AIMS OF THIS STUDY.Glycogen phosphorylase is known to be extremely specific for its substrate, and very few substrateanalogues have been found that are turned over by this enzyme. One such substrate analogue is heptenitol (seeChapter 1 of this thesis). Heptenitol acts as a substrate analogue for glycogen phosphorylase exclusively in thedegradation reaction (McLaughlin et al., 1984).As was mentioned in the first Chapter, 2-deoxy-2-fluoroglycosides with the appropriate anomericconfiguration rapidly inactivate p-retaining glycosidases, and these compounds also partially inactivate somea-glycosidases (Withers et al., 1988). Fluorinated heptenitols, with one or two electron-withdrawing fluorineatoms situated adjacent to the reaction centre, were therefore considered to be potential inactivators (irreversibleinhibitors) of glycosylases. One advantage of these compounds over 2-deoxy-2-fluoro sugars is that the formercan be used to study the effect of varying the degree of withdrawal of electron density from the reaction centre,while maintaining interactions between the 2-hydroxyl of the sugar and the enzyme. Hence difluoroheptenitol(F2hept), (2.1), and monofluoroheptenitol (F1hept), (2.2), were synthesized, and kinetic studies of thesecompounds were carried out using two glycosylases, glycogen phosphorylase and 3-glucosidase. The results ofthe kinetic studies performed with (3-glucosidase will be discussed in Chapter 3.The fluorinated heptemtols studied herein have the same basic structure as heptenitol, except that one orboth of the vinylic hydrogens is replaced by fluorine. These fluorinated heptenitols were therefore expected tobind to the active site of glycogen phosphorylase. One or both of these fluoroheptenitols might act as asubstrate of phosphorylase (which would be expected if both glycosylation and deglycosylation of the enzyme41H0OH0OHF[2.1] [2.2]H42were rapid). If either of the fluoroheptenitols acted as a substrate, the influence of inductive effects on thereaction mechanism could be studied.Alternatively, one or both of these fluoroheptenitols might act as an irreversible inhibitor (inactivator)by forming a covalent glucosyl-enzyme intermediate (which would be expected if the rate of glycosylationexceeded the rate of deglycosylation), or act as a reversible inhibitor. Enzyme inhibitors are useful for studies ofreaction mechanisms, and often yield insights into the nature of the binding interactions between the inhibitorand the active site. Such information may be obtained from kinetic or structural studies (e.g., X-raycrystallography). Fluorinated inhibitors are especially useful in enzyme kinetic studies because fluorinesubstitution provides an active-site probe for the spectroscopic analysis of trapped glycosyl-enzymeintermediates (e.g., by 19F NMR). If either of the fluoroheptenitols acted as an inhibitor, then a stable analogueof heptenitol (one that is not turned over) would be particularly useful for X-ray structural studies of thephosphorylase-inhibitor complex.2.3. RESULTSAND DISCUSSION.2.3.1. Syntheses of fluorinated heptenitols.F2hept was synthesized using a procedure that was briefly described by Motherwell et al. (1989). Thefirst step in this procedure is a Wittig-type reaction using persilylated glucono-1,5-lactone (2.3) (see Scheme2.7). F1hept was obtained by selective reduction of the appropriately persilylated F2hept derivative (2.5) usingRed Al® [sodium bis(2-methoxyethoxy) aluminum hydride] according to the procedure of Hayashi et al. (1979).The original, briefly described procedure of Motherwell et al. (1989) for the difluoromethylenation ofpersilylated glucono-1,5-lactone (2.3) involves refluxmg the starting material (in anhydrous T1{F) together with5 equiv. of each of the following reagents: (i) tris(dimethylamino)phosphine (HMPT, [Me2N]3P);(ii) dibromodifluoromethane (CF2Br);and (iii) zinc dust. Unfortunately, there are serious problems associatedwith this procedure. It requires the use of the toxic reagent FIMPT, and a byproduct of the reaction is the oxide(Me2N)3P0 (HMPA, hexamethyl phosphoramide), one of the most potent carcinogens known. The reaction isalso plagued by variable and generally poor yields (Houlton et al., 1993).43Me3SiOOSiMe3Me3SiOMjOo[2.3](Me2N)3P, CF2BrZn, THEMe3SiOOSiMe3eiOFMe3SiO[2.4] FMe4WFmay be used here(to remove TMS groups),butBu4WFis requiredto remove TBDMS groups.(t-Bu)Me2SiOOSiMe2(t-Bu)(t-Bu)MeiOF(t-Bu)Me2SIOF[2.5]toluene(t-Bu)Me2SiCIimidazole, DMFNa(CH3OC2CH)A1(Red AIHOHO 0HO F[2.1]F[orohenitoI(t-Bu)Me2SiOOSiMe2(t-Bu)(t-Bu)MeSi ,,,,H(t-Bu)Me2SiOF[2.6]Bu4NFBu4NFOHH’H[2.2]nofIuoroheptenitoI IScheme 2.7. The syntheses ofF2hept andF1hept.44Professor Motherwell’s research group has recently published a complete, detailed experimentalprotocol for the difluoromethylenation of persilylated carbohydrates such as 2.3 (Houlton et aL, 1993). Theauthors attributed the poor reproducibility and low yields of the original version of this procedure (Motherwell etal., 1989) to the use of a heterogeneous system employing zinc dust, which may lead to the decomposition ofreaction intermediates.The deprotection of 2.4 to yield the desired product, F2hept (2.1), presented several problems.Regardless of whether Me4NF or Bu4NF was used to effect the deprotection, it was found that F2heptco-migrated with the tetraalkylammonium salt during TLC analysis. The tetraalkylammonium salt could not beremoved even after repeated column chromatography (27:2:1 ethyl acetate:methanol:water) of the deprotectionreaction mixture.Several cation-exchange resins were then examined to see if they could remove the tetra(n-butyl)ammonium ion and allow for the purification ofF2hept. These resins (and their counterions) were:1. Dowex® 50WX12 (Na) 5. Dowex® MR-3X8 (Li, — OH)2. Biorex® 70 (Naj 6. Dowex® 50WX12 (Lii3. Dowex® 50 WX2 (Na) 7. Dowex® 50 WX2 (Lij4. Amberlite® JR 120 (LijOf the resins exaniined, only #6 and #7, i.e., those containing the lithium salt form of Dowex® 5OWX,were able to remove the tetra(n-butyl)ammonium ion and allow for the purification ofF2hept. Resin #7,Dowex® 50 WX2 (Lii, gave the best results of the resins examined, and after cation-exchange, the LiF saltcould be easily removed from the crude deprotection reaction mixture by subsequent column chromatographyusing silica gel.Difluoroalkenes can be selectively reduced to monofluoroalkenes using the procedure of Hayashi et al.(1979). They found that monofluorinated alkenes (such as 1-fluoro-l-octene and 2-fluoro-1-phenylethene)could be successfully prepared from the corresponding difluorinated derivatives using Red Al® [sodiumbis(2-methoxyethoxy) aluminum hydride] as a reducing agent. Hayashi et al. (1979) found that the strong45reducing agent LiA1H4was nevertheless sluggish and impractical for use in these reactions. Unfortunately, itwas found that the difluoroheptenitol derivative 2.4 could not be used as a starting material for the selectivereduction reaction because the trimethylsilyl protecting groups were unstable under the reaction conditions. Itwas therefore necessary to explore the use of alternative protecting groups. Triethylsilyl and t-butyldimethylsilylethers are about 100 and 10,000 times more stable than trimethylsilyl ethers, respectively (Greene & Wuts,1991), thus these groups were suitable candidates. Unfortunately, all attempts to persilylate glucono-1,5-lactonewith TBDMS chloride were unsuccessful, and if glucono-1,5-lactone was first persilylated withEt3SiC1, thedifluoromethylenation reaction (i.e., analogous to 2.3 —* 2.4) was unsuccessful. It was therefore necessary toproceed as shown in Scheme 2.7, first synthesizing the TMS-protected difluoroheptenitol, then replacing theprotecting groups.The purified persilylatedF1hept (2.6) yielded only one spot after TLC. However, NMR spectroscopy(1H and 19 showed that two isomers of persilylatedF1hept were formed in a ratio of 28:1 (from the ratio ofintegrated H-i resonances in the 1H NMR spectra). The spectral (1H, 13C, and 19FNMR as well as MS) datafor the major isomer (transF1hept and its persilylated precursor) are reported in the Materials and MethodsSection.The selective reduction of difluoroalkenes to monofluoroallcenes using the procedure of Hayashi et al.(1979) generally yields a mixture of the trans (major) and cis (minor) isomers. Hayashi et al. (1979) found thatalthough the ratio of trans and cis isomers varied, they obtained substantial amounts ( 20% of the product) ofthe cis isomer in each of the reactions studied, except for the synthesis of 2-fluoro-1 -phenylethene, where only7% of the product was the cis isomer. Hence it was noteworthy that the synthesis of Fhept yielded only 3.5%of the product as the cis isomer.In most cases the stereochemistry of monofluoroalkenes can be easily identified by the value of theor 1H NMR coupling constant,3JH,F. The3JH,F values for fluorine and vicinal vinylic hydrogens are 40and 20 MHz for trans and cis coupling, respectively (Gordon & Ford, 1972). However, the fluorine atom inF1hept lacks a vicinal hydrogen, and hence it was impossible to use 3JH,F values to determine thestereochemistry ofF1hept.462.3.2. The stereochemistry ofF1hept.a. The trans and cis isomers ofFjhept.The trans and cis isomers ofF1hept are shown below:7HQ”LJOH F[ cisVarious NMR experiments were performed in order to confirm the stereochemistry of the major isomerofF1hept.b. 13CNMR experiments.In the1H-decoupled, 13C NMR spectrum of TBDMS-protectectF2hept (2.5, see Scheme 2.7), c-i and-2 each resonated as a doublet of doublets, and the multiplicity of these resonances arose from coupling withthe two fluorine atoms. There was one other doublet in the spectrum (at 6 = 66.9 ppm, J 2 Hz), and thisresonance was assigned to c-3, the multiplicity presumably arising from trans aflylic coupling with one of thefluorine atoms (see Fig. 2.3a). The other resonances in the1H..decoupled, ‘3c NMR spectrum ofTBDMS-protectedF2hept were singlets (see the Materials and Methods Section for detailed analytical data).In the1H-decoupled, 13c NMR spectrum of TBDMS-protected F1hept (2.6, see Scheme 2.7) C-3resonated at 6 = 65.2 ppm as a singlet (see Fig. 2.3b and Materials and Methods). It is known that the couplingconstant arising from trans allylic coupling between a fluorine atom and a carbon atom in a given molecule isgreater than the coupling constant that would arise from cis allylic coupling between two such atoms in the sameallylic moiety of the molecule in question (or a structurally related molecule) (Gaudemer, 1977). The absence ofHIF47(a) (b)11 1 I I ii 11 I ) I i t ‘ ‘ ‘ ‘ ‘ I68 66 66 646 (ppm)Figure 2.3. 13CNMR determination of the stereochemistry of the major isomer of Fjhept.75 M1-Iz, H-decoupled, 13C NMR spectra were obtained using the samples (in CDC13)indicated below. Onlythe relevant regions of the spectra are shown, which contain the signal atthbutable to C-3 as indicated below.(a) TBDMS-protected Fhept (6= 66.9 ppm, d, 1 2 Hz, C-3).(b ThDMS-protectedF1hept (8=652 ppm, s, C-3).48coupling in the C-3 resonance (6= 65.2 ppm) in the 13C NMR spectrum of TBDMS-protectedF1hept ruled outa trans allylic relationship between the fluorine atom and C-3, for if this was the case, a doublet would havebeen observed. Hence the geometric relationship between the fluorine atom and C-3 inF1hept is cis, andtherefore 0-5 is trans with respect to the fluorine atom.c. An NOE experiment on TBDMS-protected Fjhept.The nuclear Overhauser effect (NOE; reviewed in Derome, 1988) frequently yields information on theinternuclear distance between atoms. When an NOE experiment on TBDMS-protected F1hept was performed,no enhancement of the vinylic hydrogen (H-i) signal (at 6 = 6.8 ppm) was observed when the H-3 signal (at6=4.57 ppm) was irradiated, and vice versa (note that only one peak of the doublet attributable to the vmylichydrogen could be irradiated at a time due to the magnitude of the coupling constant, J13 = 80 Hz). This resultindicated that the vinylic hydrogen and H-3 are not very close to each other. These two hydrogens are mostlikely trans with respect to each other, and hence 0-5 is most likely trans with respect to F-i. This result agreedwith the 13C NMR data that showed that the predominant isomer ofF1hept was the trans isomer. However, theabsence of an NOE does not provide defmitive proof for the stereochemistry of a molecule.d.1Ho,zd19FNMR experiments.The 1H NMR spectrum ofF1hept provided compelling evidence for the stereochemistry and abundanceof the major (trans) and minor (cis) isomers of this compound. Two doublets attributable to H-i were observedin this spectrum; one each at 6 = 7.02 and 6.58 ppm, with integration ratios of 28:1, respectively. The moreabundant H-i signal was downfield (6= 7.01 ppm) relative to the less abundant H-i signal (6 = 6.58 ppm). Anexhaustive compilation of experimental data obtained from olefinic compounds (see Table 5 of Gaudemer,1977) has shown that an olefinic proton that is cis to an O-alkyl group resonates downfield from an olefinicproton that is trans to the same O-alkyl group. Hence the H-i atom of the major isomer must have been cis tothe oxygen atom of the sugar ring, which is exactly what one would expect if the major and minor isomers weretrans and cis, respectively.49The 19F NMR spectrum ofF1hept also provided evidence for the stereochemistry of the major (trans)and minor (cis) isomers of this compound. Two doublets attributable to F-i were observed in this spectrum; aminor doublet at 8 = -163.78 ppm and a major doublet at 6 = -169.71 ppm. Here the less abundant F-i signalwas downfield (6 = -i63.78 ppm) relative to the more abundant F-i signal (6 = 469.71 ppm). Hence the F-iatom of the minor isomer must have been cis to the oxygen atom of the sugar ring, which again is exactly whatone would expect if the major and minor isomers were trans and cis, respectively.The above results are consistent with the assignment of 1 and 19F NMR signals arising fromstereoisomeric H-i and F-i atoms in the cis (2.7) and trans (2.8) monofluoroalkenes shown below (Houlton etal., 1993). In the cis isomer (2.7), a compound whose stereochemistry has been determined by X-raycrystallography, the vinylic H-i and F-i atoms give rise to NMR signals of 6 = 6.37 and -160 ppm in the 1 and19F spectra, respectively (Houlton et aL, 1993). However, in the trans isomer (2.8), the vinylic H-i atom givesrise to a signal of 6 = 7.00 (downfield from 6 = 6.37 ppm) in the 1H NMR spectrum, and the vinylic F-i atomgives rise to a signal of 6 = -i77.4 (upfield from 6 = -i60 ppm) in the 19FNMR spectrum (Houlton et al., 1993).These data are what one would expect based on the geometric relationship of the stereoisomenc H-i and F-iatoms to the ring oxygen.H F[2.7] (cis) 00 [2.8] (trans)2.3.3. Substrate and inactivation tests usingF2hept.F2hept was tested as a substrate of phosphorylase b in the glycogen phosphorolysis reaction. In thisexperiment, i7 mMF2hept was incubated with iOO mM KC1, i mM EDTA, 1 mM DTr, 50mM thethanolaminehydrochloride (pH 6.8), 1% glycogen, 5 mM orthophosphate, i .5 mM AMP, and 0.6 mg of enzyme. After fivedays at room temperature there was no detectable turnover of F2hept as measured by TLC and 19fNMR.50The substrate test was run in parallel with a control reaction without F2hept but otherwise containingthe same reaction components. During the course of the above test, small aliquots (10 pL) were removed fromthe substrate test and the control reaction, and then added to fresh tubes, each of which contained 0.500 mL of areaction mixture containing a saturating concentration (16 mill) of ctGlP, 1 mM AMP, 1% glycogen, 100 mMKC1, 1 mM EDTA, 1 mMDT, and 50 mM triethanolamine hydrochloride (pH 6.8). Initial reaction rates werethen assayed in the direction of glycogen synthesis as described (Engers et al., 1970). No significant decrease inphosphorylase activity was observed over 5 days when the results obtained using aliquots from the substrate testwere compared with those from the parallel control reaction.2.3.4. Inhibition studies usingF2hept andF1hept.Inhibition studies were performed using phosphorylase acting in either direction of the reaction. In theglycogen synthesis direction, the experiments were performed using a constant concentration (1%) of glycogen,5 different concentrations of aG1P, and 4-5 different concentrations of the inhibitor (as well as in the absence ofthe inhibitor). About 2.5 jig of phosphorylase b (from rabbit muscle) was used in each reaction. The exactreaction conditions are given in the legends to Figs. 2.4 and 2.5. A standard assay (Engers et al., 1970) was usedto determine the amount of inorganic phosphate released during the reaction.When the data for Figs. 2.4a and 2.5a were plotted as double-reciprocal or Dixon plots, curved lineswere obtained, indicating that the fluoroheptenitols bind to and stabilize the T-state conformation ofphosphorylase. Straight lines were obtained from Hill plots (ln[vI(Vmax — v)] vs. ln[SJ) with both inhibitors(Figs. 2.4a and 2.5a). The values of the inhibition constants were determined from replots of the apparent Kmvalues (Km,app) versus the inhibitor concentrations (Figs. 2.4b and 2.5b). F2hept andF1hept both acted ascompetitive inhibitors of phosphorylase. These conclusions were based on the fact that for both compounds—over the range of inhibitor concentrations studied—the Vmax was constant, while the Km value showed anapparent increase of a factor of (1+ [I]IK) (Segel, 1975). The K1 values ofF2hept andF1hept for the glycogensynthesis reaction were 4.9 ± 0.4 and 4.0 ± 0.4 mM, respectively. The values of the kinetic parameters for thesecompounds and other substrates and inhibitors of phosphorylase are summarized in Table 2.1.5120-20in [zGlP]414121086420-5 0 15 20(b)Figure 2.4. Kinetic parameters for the inhibition byF2hept of phosphorylase-catalyzed glycogen synthesis.(a) Hill plots used to determine Km,app for the inhibition by F2hept of phosphorylase b-catalyzed utilization ofcLG1P. Reactions (0.500 mL) were performed at 30 °C and contained 100 mM KC1, 50 mM TEA-HC1 (pH6.8), 1 mM EDTA, 1 mM DTT, 1 mM AMP, 1% glycogen, and 2.5 .Lg of rabbit muscle phosphorylase b.Initial rates for the 5-mm reactions were determined using a standard phosphate assay (Engers et al., 1970).The concentrations ofF2hept in the reactions were 0 (÷}, 3.0 (o), 6.0 { .}, 10 {), and 20 (I) mM.(b) A replot of data obtained from panel (a) in order to estimate the value of the K1 by visual inspection. Thefollowing values of Km,app were determined from panel (a): 2.6,4.1,5.5, 7.1, and 13 mM.(a)1 2 35 10[F2hept] (mM)520—420Figure 2.5. Kinetic parameters for the inhibition by Fjhept of phosphorylase-catalyzed glycogen synthesis.(a) Hill plots used to determine Km,app for the inhibition byF1hept of phosphorylase b-catalyzed utilization ofctGlP. Reactions were performed as described in Fig. 2.4 except that the concentrations of Fhept in thereactions were 0 (÷}, 0.60 (01, 1.2 (• 1 4.0 (DI, 8.0 { }, and 12 { L} mM.(b) A replot of data obtained from panel (a) in order to estimate the value of the K1 by visual inspection. Thefollowing values of Km,app were determined from panel (a): 3.3, 3.7, 4.5, 6.0, 8.7, and 13 m!vf.32—1-21412‘1o80 0.5 1 1.5 2 2.5 3 3.5 4in [xG1PJ(a)(b)-4 -2 0 2 4 6 8 10 12[F1hept] (mM)53Figure 2.6. The inhibition by fluoroheptenitols of phosphorylase-catalyzed glycogen phosphorolysis.Initial rates were measured by quantitating the reduction of NADP spectrophotometrically (Engers et al., 1969).Reactions (0.205 mL, 30 °C, and pH 6.8) contained 0.15 .tg of rabbit muscle phosphorylase b, 35 mMimidazole, 20 mM sodium glycerophosphate, 10 mMMg(OAc)2,5 mM DTT, 1 mM EDTA, 1 mM AMP, 0.5%glycogen, 1 mM (3NADP, 15 U/mL phosphoglucomutase, and 3.4 UImL of glucose-6-phosphate dehydrogenase.(a) Inhibition byF2hept. The concentrations ofF2hept in the reactions were 0, 0.83, 3.3, 8.3, 11.2, and 16.5 mM.(b) Inhibition byF1hept. The concentrations of Fjhept in the reactions were 0, 2.4, 4.9, 9.8, 17.1, and 24.4 mM.0.10.080.060.040.0200.180.160.140.120.10.080.060.040.020E10 5 10 15[F2hept] (mM)(a)••— [= 1(b)•—0 5 10 15 20[F1hept] (mM)2554Table 2.1. Dissociation constants of fluoroheptenitols and substrates with phosphorylase b.K1 or Km (mM)Compound Referenceglycogen synthesis glycogen phosphomlysisF2hept 4.9 ± 0.4 -.2.8 this workF1hept 4.0±0.4 -.0.9 ibid.aG1P 1.5 — Engersetal., 1969D-glucose 2.0 — Street et al., 1986heptenitol — 43(a) Klein et al., 1986orthophosphate — 11(b) Engers et at, 1969Except as noted in (b) below, data were obtained from reactions with 1 mM AMP and a saturating concentrationof glycogen (1%).(a) Obtained using arsenate in lieu of phosphate.(b) A value of 15 mM was obtained with 1 mM AMP and a nonsaturating concentration of glycogen.“Range-finding” (or approximate) K values (RF K1 values) forF2hept andF1hept were determined forthe glycogen phosphorolysis reaction (Fig. 2.6). In these experiments, initial rates were determined using asingle concentration of the substrate orthophosphate (21 mM), and 5 different inhibitor concentrations (as wellas with no inhibitor). As before, glycogen was present in all phosphorolysis reactions at a saturatingconcentration (1%). Initial rates were determined using a phosphoglucomutase/glucose-6-phosphatedehydrogenase coupled assay (Engers et al., 1969). The approximate K1 values forF2hept andF1hept weredetermined from the appropriate Dixon plot (1/v vs. [I]) (Fig. 2.6).The linear plots obtained in this case (Fig. 2.6) suggest that in the presence of orthophosphate,F2heptandF1hept were able to bind to the R-state conformation of the enzyme. Similar behaviour has been observedpreviously with glucono-1,5-lactone and nojirimycin tetrazole (Gold et at, 1971; Withers & Rupitz, unpublishedresults). The approximate K1 values forF2hept andF1hept were 2.8 and 0.9 mM, respectively, for the glycogen55phosphorolysis reaction. The dissociation constants (K or Km) of the fluoroheptenitols, their parent compound(heptenitol), and other substrates in either direction of the reaction catalyzed by phosphorylase b are given inTable 2.1.The monosaccharides listed in Table 2.1 all exhibited similar affinities for the enzyme’s active site (i.e.,K1 or Km values in the 5 mM range). These monosaccharides have closely related glycone moieties (glucose orgluco-heptenitol structures), but quite different aglycone moieties. However, the monosaccharides listed inTable 2.1 all have aglycone moieties that are either negatively charged (e.g., the phosphate moiety in aGIP), oruncharged but electron-rich (e.g., C=C moiety in the heptenitols). This suggests that there must be one or morepositively charged (or electron-accepting) amino acid side chains adjacent to the anomenc carbon atom of themonosaccharide substrate (or inhibitor) in the enzyme’s active site.2.3.5. X-ray crystallographic studies.X-ray crystallography has been used to study the binding of heptenitol (and heptulose-2-phosphate) byglycogen phosphorylase b (Hajdu et al., 1987; Johnson et a!., 1990), and these studies have yielded insights intothe catalytic mechanism of the enzyme. As an example, the high-resolution crystal structure of theheptulose-2-phosphate-enzyme complex provided considerable experimental support for the “proton transfer”role of the cofactor PLP (McLaughlin et al.,1984; see also Johnson et al., 1990).Another example of the utility of X-ray crystallographic studies is the work of Hajdu et a!. (1987), whoused a synchrotron radiation source and fast data-collection techniques to ‘observe” the progress of the catalyticconversion of heptenitol to heptulose-2-phosphate in crystals of phosphorylase b. Their objective was to detectthe transient accumulation of a ternary complex (involving the enzyme, heptenitol, and phosphate) prior to theformation of the product (heptulose-2-phosphate). They did indeed obtain data indicating occupancy of thephosphate-binding site, as well as occupancy of the sugar-binding site, at early stages of the reaction.Occupancy of the phosphate-binding site increased with time, and its electron density eventually becamecontiguous with that of the reaction centre on the sugar, consistent with the formation of the product,heptulose-2-phosphate. However, the enzyme has a much higher affinity for the product, heptulose-2-phosphate(K = 14 tiM; Klein et a!.,1984), than for phosphate (K = 15 mM for AMP-activated phosphorylase; Engers56et aL, 1969). It is therefore debatable whether the crystal structure observed at the early stages of the reaction inquestion represents a ternary complex (involving the enzyme, heptenitol, and phosphate) or a mixture of twobinary complexes: (i) the enzyme and heptenitol, as well as (ii) the enzyme and heptulose-2-phosphate (Hajduetal., 1987).Hence even with the fastest available X-ray crystallographic data-collection methods, it is stillnecessary to reduce the phosphorylase reaction rate as much as possible in order to detect phosphate bound inthe enzyme’s active site (i.e., prior to the formation of any product). Part of the reason for this is that therate-limiting step in data accumulation is the diffusion of the ligands into the crystal. Thus it is necessary toreduce the reaction rate to below the rate of diffusion of small ligands into the crystal. The use of fluorinatedheptenitols proved to be especially useful for this purpose. The results of the kinetic studies reported hereinshowed that fluoroheptenitols are not turned over by the enzyme. Furthermore, it is known that in the absence ofheptenitol, phosphate alone cannot bind in the active site of phosphorylase b (E. Duke, pers. comm.). However,from the data discussed below, it appears that the binding ofF1hept promotes the binding of phosphate, andallows for the detection of phosphate bound in the active site, since in this case there is no detectable formationof a pmduct.X-ray crystallographic studies of each of the fluoroheptenitols bound at the active site of glycogenphosphorylase b were carried Out in collaboration with Dr. E. Duke and Prof. L. N. Johnson of the Laboratory ofMolecular Biophysics at Oxford University. Crystals of T-state phosphorylase b were soaked in a solution of50 mM fluoroheptenitol, 50 mM phosphate, 10 mM Mg(OAc)2,2.5 mM AMP, in 10 mM BES-NaOH, pH 6.7.Each crystal was soaked for a minimum of 30 mm to allow for diffusion of the ligands into the crystal.The three-dimensional electron-density map of the phosphorylase—F1eptco-crystal showed a weak butreadily detectable region of electron density attributable to the binding of phosphate (Fig. 2.7a). Phosphate wasbound in a region of the active site directly below (i.e., on the a-face of) the sugar ring ofF1hept, close to thefluorine atom ofF1hept and also adjacent to the 5’ phosphate of the cofactor PLP (Fig. 2.7a). The structural datafrom the phosphorylase—Fihept co-crystal agree with the results of mechanistic studies which postulated that thebound substrate phosphate and the 5’ phosphate of PLP are very close together in the active site of57Figure 2.7. The structureof Fhept orF2hept boundin the active site ofglycogen phosphorylase b.(a)Co-crystals of T-stateglycogen phosphorylase band either F1hept (a) orF2hept (b) were preparedas described in the text.Data were collected to2.3 A resolution by Dr. E.Duke in Prof. L. N.Johnsons laboratory atOxford. Contours ofelectron density areindicated.(a) F1hept—phosphorylase.Electron density forphosphate was observedbelow the sugar ring. Thisperspective of thestructure does not permit aclear view of the oxygenatom of the C-4 hydroxylgroup.(b)F2hept—phosphoiylase.No electron density forphosphate was observed.For the purpose ofcomparison only, thiselectron density mapshows the location of thephosphate bound in thephosphorylase—Fheptco-crystaLllX17 /17 -N284/../ “‘.• A’Figure 2.8. Composite electron-density map of fluoroheptenitols bound in the active site of phosphorylase b.This electron density map is a composite of two separate maps of either F1hept (yellow) orF2hept (blue) boundin the active site of T-state glycogen phosphorylase b (see Fig. 2.7). The electron density attributable tophosphate is from theF1hept-phosphorylase co-crystal only.58Asp 2?3Asn 2H4Difluoroheptcnj(oI : cyani\loiioiluoroheptcnitol yeflowii’I\;ip4fIR 377_, •‘/vi•.../ ..\ \59phosphorylase, and that their interaction with each other is the major source of the catalytic power of the enzyme(Helmreich, 1992; Madsen & Withers, 1984, 1986; Palm et al., 1990).The three-dimensional electron-density map of thephosphorylase—F2eptco-crystal did not show anyregion of electron density attributable to the binding of phosphate in the active site (Fig. 2.7b). For the purposeof comparison, this electron density map shows the location of the phosphate bound in the phosphorylase—FJzept co-crystal, but it is clear that in the same regionof the electron density map of the phosphorylase—F2eptco-crystal there is no corresponding electron density (compare the electron-density contours of Figs. 2.7a and2.7b in this region). It may be that the presenceof two electron-rich fluorine atoms at the C-i position of aheptenitol derivative bound at the active site destabilizes the enzyme-substrate interactions necessary for thebinding of a negatively charged phosphate immediately adjacent to these fluorine atoms.BothF1hept and Fhept bound to the same location in the active site of phosphorylase b as heptenitol,and all three heptenitols appeared to exploit the same binding interactions with the same amino acid residuesofthe enzyme (Fig. 2.8; see also McLaughlin etal., 1984 for the binding of heptenitol in the enzyme’s activesite).Extra lobe(s) of electron density representing thefluorine atoms were evident (Figs. 2.7-2.8). When the electrondensity maps of the fluoroheptenitol-enzyme co-crystals were compared with that of the enzyme alone,itappeared that the position of the Asn 284 side chain had moved slightly in order to accommodate the fluorineatom(s) in each case (F. Duke, pers. comm.).Interestingly, closer examination of the structuresshown in Fig. 2.8 suggested that the two boundsugars adopted different conformations. F1hept appeared to bind in a full-chair conformation, as had beenfoundpreviously for heptenitol (Hajdu et al., 1987), whereasF2hept appeared to adopt a structure resembling ahalfchair (F. Duke, pers. comm.). However, the resolutionof the data (2.3 A) is insufficient to place muchconfidence in these interpretations.Only one isomer (trans) of Fihept was observed in theX-ray structure of the phosphorylase—Fiheptco-crystal, as evidenced by the single lobe of electrondensity due to fluorine at C-i (see Fig. 2.7a). Prior to theX-ray crystallographic studies, analytical data forF1hept already had indicated the formation of one major60(>96%) isomer. The results of various NMR experiments (reported earlier in this chapter) indicated that themajor isomer was transF1hept, and hence the X-ray crystallographic and NMR results were in good agreement.2.4 CONCLUSIONS.The 1H NMR spectrum ofF1hept showed that the major (>96%) isomer formed during the selectivereduction ofF2hept was the trans isomer, and that a minor (<4%) but detectable amount of the cis isomer wasalso formed. The assigned stereochemistry of the major isomer ofF1hept was confirmed by 1H, 1H NOE, 13C,and 19F NMR experiments.BothF2hept and Fihept were found to be competitive inhibitors of phosphorylase, binding to the activesite in the absence of phosphate and stabilizing the (inactive) T-state of the enzyme, as seen with glucoseanalogues (Martinet al., 1991; Street et al., 1986).The binding constants ofF2hept,F1hept and czGlP in the glycogen synthesis direction are all similar(1-5 mM; see Table 2.1). The inhibition constants of F2hept, F1hept and heptenitol in the glycogenphosphorolysis direction are also similar (1-4 mM; see Table 2.1). These results indicate that fluorinesubstitution did not significantly affect the affinity of the enzyme for the heptenitol derivatives under study.As was discussed in Chapter 1, heptenitol acts as a “suicide substrate” for phosphorylase in theglycogen degradation reaction. Heptenitol is rapidly turned over by the enzyme to form 1-deoxy-D-gluco-heptulose-2-phosphate, which acts as a tight-binding inhibitor (Klein et al., 1984). Given that heptenitol andF2hept possess similar affinities for glycogen phosphorylase b, the inability ofF2hept to act as either a substrateor an inactivator must be due to a very slow rate of hydration (k1). This slow rate of hydration of F2hept isprobably atthbutable to the inductive effect of the electron-withdrawing fluorine atoms at the C-i position,which would destabilize any development of positive charge at the reaction centre. Hence the kinetic results arein agreement with the proposal that the reaction mechanism of phosphorylase involves oxocarbonium ion-liketransition states.The synthesis of fluoroheptenitols also proved to be a valuable conthbution to X-ray crystallographicstudies of glycogen phosphorylase carried out by collaborators in Great Britain. These studies yielded important61insights into the binding of phosphate in the enzymes active site. In particular, they provided the first proof thata heptenitol derivative and phosphate can bind simultaneously, something which was not clear from earlier workperformed with heptenitol (due to conversion to the product). Phosphate was bound in a region of the active sitedirectly below (i.e., on the cL-face of) the sugar ring of transF1hept, close to the fluorine atom and also adjacentto the 5’ phosphate of the cofactor PLP (see Fig. 2.7a). Both transF1hept andF2hept bound to the same locationin the active site of phosphorylase b as heptenitol, and all three heptenitols appeared to exploit the same bindinginteractions with the same amino acid residues of the enzyme. Finally, the assigned stereochemistry of themajor isomer of Fhept (trans) was confirmed by the results of the X-ray crystallographic studies of theF1hept-phosphorylase co-crystal.Since fluoroheptenitols possess no anomeric configuration, they may also prove useful for kinetic andstructural studies of other a- or -glucosyl mobilizing enzymes. These fluoroheptenitols might act as reversibleor possibly even irreversible inhibitors of these enzymes. Indeed, the next chapter of this thesis shows that thesecompounds also reversibly inhibit Agrobacterium 3-glucosidase.62CHAPTER 3: KINETIC STUDIES USING [3-GLUCOSIDASE3.1. INTRODUCTION.3.1.1. Importance and general properties of f3-glucosidases.a. Catalytic activity and role in the microbial cellulase complex.13-Glucosidase ([3-D-glucoside glucohydrolase or cellobiase, E.C. 3.2.1.21) is an enzyme that catalyzesthe hydrolysis of various compounds with J3-D-glucosidic linkages. Cleavage of the bond between the anomericcarbon and the glucosidic oxygen yields glucose as the product, as well as the free aglycone moiety.13-Glucosidases also catalyze reactions with nonglycosidic substrates, such as hydrolytic cleavage of the C—Fbond in glucosyl fluorides, and hydration of the double bond of glucals.j3-Glucosidases are widely distributed in nature, and have been isolated from bacteria (Han &Srinivasan, 1969), yeast (Duerksen & Halvorson, 1958) and other fungi (Berghem & Pettersson, 1974), andplants (Hultson, 1964). As a component of the cellulase complex, [3-glucosidase catalyzes the hydrolysis ofcellobiose and cellodextrins (formed by the action of endo- and exo-glucanases) to form glucose. Sincecellobiose acts as an inhibitor of both endo- and exo-glucanases, J3-glucosidase plays an important role in thedegradation of cellulose (Berghem et al., 1975).b. Catalytic activity and role in lysosomal glycolipid metabolism in humans.Glucosylceramide is an important glycolipid in animal cells. It is the substrate of lysosomal (or acid)13-glucosidase, which hydrolyzes the f3-D-glucosidic linkage of this glycolipid (Fig. 3.1). In humans, thehereditary metabolic disorder known as Gaucher disease is a result of a deficiency of lysosomal [3 -glucosidase(glucosylceramidase, E. C. 3.2.1.45). Defective activity of this enzyme may lead to one of several variants ofGaucher disease, which include the non-neuropathic (Type 1), neuropathic infantile (Type2), and neuropathicjuvenile (Type 3) forms (Barranger & Ginns, 1989; Grabowski, 1993; Grabowski et al., 1990). Prominentsymptoms of Gaucher disease include the accumulation of glucosylceramide in tissues, and enlargement of thespleen and liver.63Figure 3.1. The structure of glucosylceramide.Gaucher disease was the first lysosomal storage disease discovered (Gaudier, 1882), and it remains themost common type of lysosomal storage disease. It is also the first lysosomal storage disease to have beensuccessfully treated by enzyme therapy (Barton et al., 1991; Beutler et al., 1991; Fallet et a]., 1992). There arecurrently many studies in clinical and basic research directed at improved treatment and a possible cure for thisimportant disorder of 3-glucosidase function (Barton et al., 1991; Beutler et a!., 1991; Grabowski, 1993).3.1.2. Agrobacterium f-g1ucosidase.The [3-glucosidase used for this work is a protein from Agrobacreriumfaecalis (previously typed asAlcaligenesfaecalis) (Han & Srinivasan, 1969). The purification and characterization of Agrobacrerium[3-glucosidase have been reported by Day & Withers (1986).The Agrobacterium [3-glucosidase monomer has a relative molecular mass of -.50,000. Thecatalytically active form of this enzyme is a dimer, with one active site per monomer (Day & Withers, 1986).The enzyme is 3-retaining and has a ‘flexible” specificity for both the glycone and aglycone moieties of thesubstrate. Agrobacterium f3-glucosidase hydrolyses a wide variety of different glycoside substrates, including[3-glucosides, 13-mannosides, J3-galactosides, and cellobiose (Day & Withers, 1986). The substrate flexibility ofthe enzyme even includes the ability to hydrolytically cleave C—S, C—N, and C—F bonds.The gene encoding Agrobacterium f3-glucosidase has been cloned and expressed in E. coli(Wakarchuck et a!., 1986). The expression product of the cloned gene is basically identical with the naturallyoccurring enzyme. Both proteins have the same amino acid sequence, and display the same kinetic behaviour(Wakarchuck et a!., 1986). The j3-glucosidase enzyme (commonly referred to as pABG5) used for this workwas the expression product of the cloned gene.643.2. THE AIMS OF THIS STUDY.Various glycals were examined as potential substrates (catalytically hydrated by the enzyme), or aspotential inhibitors or inactivators (of the hydrolysis of a glucoside substrate), of Agrobacterium 3-glucosidase.One or more of these compounds might inactivate f3-glucosidase by the formation of a covalent glycosyl-enzymeintermediate, which would be expected if the rate of glycosylation significantly exceeded the rate ofdeglycosylation, and this latter rate was extremely slow. Alternatively, one or more of these compounds mightact as a substrate, which would be expected if both glycosylation and deglycosylation were rapid. Under thesecircumstances it might be possible to use a series of related substrates to study inductive effects on the reactionmechanism, or to study the stereochemistry of hydration of the substrate. Finally, these compounds should atleast act as inhibitors of the catalytic hydrolysis of glucosides. The presence of the double bond in a glycalmight result in increased affinity of these compounds for f3-glucosidase, due to the flattening of the sugar ringtowards the half-chair conformation of the transition state. Tight-binding enzyme inhibitors have frequentlyproven useful in basic and applied research.D-Glucal is a hexenitol glycal with an endocydic enolic double bond. This compound has been shownto act as a slow substrate of Agrobacterium 13-glucosidase (Street, 1988). D-Gluco-heptenitol is a glycal with anexocyclic enolic double bond, and the effect of this compound on pABG5 activity has yet to be investigated.Such an investigation was part of this study, and hence heptemtol was synthesized for this work.Both Fhept andF1hept were found to act as reversible inhibitors of glycogen phosphorylase b, an(x-glycosylase (see Chapter 2). Since heptenitols possess no anomeric configuration, the effects ofF2hept andF1hept on Agrobacterium 3-glucosidase activity were investigated. Methylgiucal was also synthesized, and itseffects on f3-glucosidase activity were examined. If promising results were obtained with the parent compound,the synthesis of fluorinated derivatives of methylgiucal would be attempted, and their effects on the activity ofthe enzyme would also be studied to provide more insight into the role of inductive effects on glycal hydration.a,3-Unsaturated glucals, which might act as Michael acceptors for a nucleophilic residue in the activesite of a glycosylase, were investigated as a new class of potential inactivators of Agrobacterium 3-g1ucosidase.The first a,[,3-unsaturated derivative of D-glucal that was examined was 1-nitro-D-glucal, which was generously65provided by Prof. A. Vasella of the University of Zurich. Other cx,[3-unsaturated derivatives of D-glucal weresynthesized (or in some cases generously provided by colleagues) and their effects on Agrobacterium(3-glucosidase activity were also studied.3.3. RESULTS AND DISCUSSION.3.3.1. The synthesis of heptemtol.In 1977, Brockhaus & Lehmann published a multistep procedure for the synthesis ofD-galacto-heptenitol. Later, Prof. Lehmann’s group and their collaborators used a similar, multistep syntheticprocedure to prepare D-gluco-heptenitol as part of a study of the catalytic hydration of this compound by a- and3-glucosidases and exo-a-glucanases (Hehre et al., 1980).H3C%N,CH3AH2C*\ •CI CI ••‘C H3C*\ CH3[3.1] [3.2] [3.3]RajanBabu & Reddy (1986) have also reported the synthesis of D-gluco-heptenitol. They usedpersilylated D-glucono-1,5-lactone as a starting material, and Tebbe’s reagent (3.1) (Tebbe et al., 1977) to effectthe methylenation. However, there are several difficulties associated with the use of Tebbe’s reagent. Inaddition to being very expensive, Tebbe’s reagent has a short shelf-life, and its use requires special techniquesand equipment due to its extreme sensitivity to air and water.Dimethyltitanocene (3.3) has been used for the methylenation of several carbonyl compounds,including esters and lactones (Petasis & Bzowej, 1990). In this procedure titanocene dichlonde (Cp2TiC1,where Cp = cyclopentadienyl) (3.2) is used to prepare dimethyltitanocene (Cp2TiMe)(Claus & Bestian, 1962),which is used immediately to effect the methylenation of carbonyl compounds via a Wittig-type reaction. In this66work the procedure of Petasis & Bzowej (1990) was used for the one-step conversion of persilylatedglucono- 1,5-lactone to persilylated D-gluco-heptenitol (Scheme 3.1). The reaction was relatively simple(compared with the reaction using Tebbe’s reagent) and efficient (with an 80% yield for the reaction shown inScheme 3.1). Triethylsilyl ethers were used to protect hydroxyl groups during the synthesis ofD-gluco-heptenitol. These ethers were stable enough for the reaction conditions, yet the silyl groups could beeasily removed later using tetra(n-butyl)ammonium fluoride (TBAF).OSiEt3 .OSiEt3El3Cp2TiMeEt3SiO’E,°EtSiQ>’QTHF, 65 °C, 26 hEtSiO>___ H(80%)HScheme 3.1. The synthesis of heptemtol from glucono-1,5-lactone using dimethyltitanocene.3.3.2. Kinetic studies using heptenitol.a. Heptenitol as a substrate ofpABG5.Thin-layer chromatography provided the first evidence that heptenitol was catalytically hydrated by thecloned 3-glucosidase, pABG5. The enzymatic hydration product appeared to be identical to a sample of1-deoxy-D-gluco-heptulose that had been prepared by the quantitative (nonenzymatic) hydration of heptenitol in0.025 N H2504 after 10 mm at 100 °C. 1H NMR was then used to determine which anomer of1-deoxy-D-gluco-heptulose was formed initially during the enzymatic reaction. This determination was basedon the assignment of axial and equatorial methyl protons of 1-deoxy-D-gluco-heptulose reported by Hehre et al.(1980).67‘ ‘..-,-.I.-..-.,.I I. — I(a)(b)(c)2.0 1.8 1.6 1.4(ppm)Figure 3.2. 1H NMR determination of the stereochemistry of pABG5-catalyzed hydration of heptenitol.400 MHz NMR spectra were obtained using the samples indicated below. Only the relevant regions of thespectra are shown, which contain the signals attributable to equatorial (3 = 1.52 ppm) or axial (6 = 1.50 ppm)methyl protons of the product, 1-deoxy-D-gluco-heptulose (Hebre et aL, 1980). In all cases 18 mM heptenitolwas incubated in 50mM deuterated sodium phosphate buffer (pD 6.8) and 0.1% BSA at room temperature.(a) Nonenzymatic hydration at 8.5 hours. No enzyme was added.(b) Catalytic hydration at 13-27 minutes. pABG5 (0.32 mg/mL) was present.(b) Catalytic hydration at 51-66 minutes. pABG5 (0.32 mg/mL) was present.[H1ax144 --I ‘ I I68HOOHOH<OH 1H0 3-glucosidase HO CH3HO 2H HOOHCatalytic hydration Spontaneous anomerizationScheme 3.2. The 3-glucosidase-catalyzed hydration of heptenitol to form i-deoxy-D-gluco-heptulose.In a control reaction without enzyme, heptenitol was incubated for 8.5 hours at room temperature.During this time some spontaneous hydration occurred. In the 1H NMR spectrum (Fig. 3.2a) of this reactionmixture the only signal attributable to methyl protons was the signal at 6= 1.52 ppm (assigned to protons of anequatorial C-i methyl group) (Hehre et al., 1980). Hence only the more stable a-anomer of 1-deoxy-D-gluco-heptulose was formed as a result of the nonenzymatic hydration of heptenitol.When heptenitol was briefly incubated with pABG5 (for 13-27 mm at room temperature), the 1H NMRspectrum (Fig. 3 .2b) of this reaction mixture showed two distinct methyl 1 resonances, one at 6= 1.50 ppm(assigned to protons of an axial C-i methyl group), and another at 6 = 1.52 ppm (assigned to protons of anequatorial C-i methyl group). The signal at 6= 1.50 ppm increased as the duration of the enzymatic reactionincreased to 51-66 mm at room temperature (Fig. 3.2c; cf. Fig. 3.2b). Hence the f3-anomer of 1-deoxy-D-gluco-heptulose was formed as the initial product of the pABG5-catalyzed hydration of heptenitol (Scheme 3.2).Presumably this product subsequently anomerized spontaneously to form the more stable (X-anomer.For kinetic studies the cuprimetric method described by Hehre et al. (1980) was used to measure therate of formation of the product, kcat, and the Michaelis COflStaIIt, Km. The enzyme hydrated heptenitol at areasonable rate, with a keat value of 640 ± 60 mm4. This is about 7% of the rate of pABG5-catalyzedhydrolysis of I3G1cPNP, but about 300 times faster than the rate of pABG5-catalyzed hydration of D-glucal(kcat = 2.28 miir1,Km = 0.85 mlvi) (Street, 1988). The affmity between heptenitol and the enzyme’s active siteappeared to be very poor, as the Km measured by the cuprimetric method was 270 ± 40 mM (Fig. 3.3).1 I691.81.61.41.210.80.60.40.20 0.02 0.04 0.06[heptenitol] —1Figure 3.3. Determination of Km and Vm for pABG5-catalyzed hydration of heptenitol.Reactions (0.500 mL) were performed at 37 °C in 50 mM sodium phosphate buffer (pH 6.8) with 30 jig ofpABG5. Heptenitol concentrations were 10, 30, 60, 80, 121, 161, 201, and 251 mM.b. Heptenitol as an inhibitor ofpABG5.As a further probe of this apparently poor binding, a reversible inhibition test was performed usingheptenitol, the enzyme, and the substrate I3G1cPNP: The resulting double-reciprocal plot indicated a verycomplicated pattern of binding (Fig. 3.4a). The results suggested that heptenitol acts as a mixed-type inhibitorof pABG5-catalyzed hydrolysis of I3G1cPNP. The change (from uncompetitive or noncompetitive tocompetitive) in the type of inhibition observed as heptenitol concentration increased can be seen clearly in aDixon plot (Fig. 3.4b).However, the substrate specificity for pABG5-catalyzed hydration of heptenitol, kcatlKm = 2.4mintM,wasabout the same as that for pABG5-catalyzed hydration of D-glucal, kcat/Km = 2.7min1M4(Street, 1988).I00.08 0.1(rnM)’6040120I0-201001806040• —01008070I I I IAI I I(a)-5O-4O-3OOO 0 10 20 30 40[I3GIcPNPI—1 (mM) —1‘I’ ‘I’I I’ I’l’(b)IIIIIlIIIIJII-40 -20 0 20 40 60 80 100 120 140[heptenitol] (mM)Figure 3.4. Determination of kinetic parameters for the inhibition of pABG5 by heptenitol.(a) Double-reciprocal plot of kinetic data. Reactions (0.500 mL) were performed at 37 °C in 50 mM sodiumphosphate buffer (pH 6.8) with 0.06 .tg of pABG5 and 0.1% BSA. The concentrations of heptenitol in thereactionswere0{o},1.6(V},4.1{I},8.2(.},20.4(L},40.8([]},81.6{A},and122(x}mM.(b) Dixon plot of data obtained from panel (a). The concentrations of 3GlcPNP in the reactions were 0.33 { a),0.17 {.}, 0.10 {[]}, 0.067 (I), and 0.033 (La) mM.71These results suggest that at low inhibitor concentrations, heptenitol occupies a binding site other thanthe active site, and acts as a noncompetitive inhibitor (K = 13 ± 1 mM). This K1 value was obtained by fittingthe data from low inhibitor concentrations (0, 1.6, 4.1, and 8.2 mM) to equations describing different types ofenzyme inhibition, using the computer program GraFit (Leatherbarrow, 1990). The best fit of the data wasobtained for noncompetitive inhibition. The next best fit was obtained for uncompetitive inhibition, whichwould have yielded a value ofK1 = 8.5 ± 0.7 mM.As the concentration of heptenitol increased, the K increased also. It appeared that at higher inhibitorconcentrations, heptenitol may also act as a competitive inhibitor, binding to the enzyme’s active site. Thisexplanation might account for the large difference in the values of the dissociation constants obtained from thesubstrate test (Km = 270 mM) and the reversible inhibition test (K = 13 mM).c. Inactivation tests using heptenitol.Heptenitol was examined as a potential inactivator of pABG5 by incubating the enzyme with 10 mMheptenitol and the appropriate reaction buffer for 20 hours at 37 °C (see Materials and Methods). At varioustime intervals small aliquots were removed and residual enzyme activity was assayed using the substrateI3G1cPNP. No time-dependent inactivation of pABG5 was detected during these tests when compared withparallel control reactions.3.3.3. Kinetic studies using fluoroheptenitols.a. Inactivation tests using fluoroheptenitols.F2hept andF1hept were examined as potential inactivators of pABG5 by incubating the enzyme with10 mM of either fluoroheptenitol and the appropriate reaction buffer for 19 hours at 37 °C (see Materials andMethods). At various time intervals small aliquots were removed and residual enzyme activity was assayedusing the substrate I3G1cPNP. No time-dependent inactivation of pABG5 was detected during these tests whencompared with parallel control reactions.72b. Substrate tests using fluoroheptenitols.TLC and 19F NMR were used to determine if F2hept orF1hept acted as a substrate of pABG5. Eitherpotential hydration product (1, 1-difluoro- or 1-fluoro- derivative of 1-deoxy-D-gluco-heptulose) was expected toyield a TLC spot with an Rf lower than that of the precursor fluoroheptenitol. The 19F NMR spectrum of eitherpotential hydration product was also expected to differ from that of the precursor fluoroheptenitol.There were no detectable differences in the thin-layer chromatograms and 19F NMR spectra obtainedusing aliquots taken over a period of 3 days from a 0.500-mL reaction mixture containing 17 mM F2hept, 0.6 mgof pABG5, 0.1% BSA, and 50 mM deuterated phosphate buffer (pH 6.8). comparable results were obtained inanother experiment using the same conditions, but whereF2hept was replaced by 10 mMF1hept. These resultsindicated that neither F2hept norF1hept acted as either substrates or inactivators of pABG5.c. Fluoroheptenitols as inhibitors ofpABG5.Reversible inhibition tests were performed using either F2hept or Fihept. In each case, an approximatevalue of the inhibition constant (a range-finding K1 or RF K) was determined by measuring initial hydrolysisrates using a single substrate concentration (85 JiM I3G1cPNP) and 5 or 6 inhibitor concentrations. Dixon plotsof the respective data yielded RF K1 values. Inhibition constants for eitherF2hept or F1hept were thendetermined accurately. This involved measuring initial rates using 5 substrate concentrations bracketing the Kmvalue (determined in the absence of inhibitor), and carrying out a set of reactions at each substrate concentrationusing 5-7 inhibitor concentrations bracketing the respective RFK value determined earlier.The initial-rate data obtained at each substrate and inhibitor concentration were then fitted to equationsdescribing different types of enzyme inhibition, using the computer program GraFit (Leatherbarrow, 1990).The data fit very well to equations describing noncompetitive and uncompetitive inhibition. The fit to theequation describing uncompetitive inhibition yielded K1 values of 0.37 ± 0.01 mM and 2.1 ± 0.1 mM forF2heptand Fihept, respectively, while the fit to the equation describing noncompetitive inhibition yielded K1 values of0.57 ± 0.03 mM and 3.2 ± 0.2 mM forF2hept andF1hept, respectively.Double-reciprocal plots were used to illustrate the nature of the observed inhibition (see Figs. 3.5a and3.6a for theF2hept andF1hept plots, respectively). These plots showed that neitherF2hept norF1hept acted as a73competitive inhibitor of pABG5. In each case it appeared that uncompetitive (or noncompetitive) behaviour wasobserved. Thus bothF2hept andF1hept displayed similar inhibitory behaviour to that observed earlier with lowconcentrations of heptenitol. This behaviour was clearly evident in Dixon plots (see Figs. 3.5b and 3.6b for theF2hept andF1hept plots, respectively), and by a comparison of these plots with that of heptenitol (Fig. 3.4b).The theory, mathematical expressions, and graphical methods for the determination of different types ofinhibition have been reviewed by Segel (1975) (see also Appendix II). By definition a competitive inhibitor isone that competes with the substrate for binding to the active site of the free enzyme. Although other modelshave been proposed to describe this phenomenon, the classical model postulates that competitive inhibitioninvolves a single site that can bind either a substrate or an inhibitor molecule, but not both.By definition, both noncompetitive and uncompetitive inhibition involve two sites on the enzyme, onethat is specific for the substrate (the active site), and a second, distinct site that is specific for the inhibitor(Segel, 1975). In noncompetitive inhibition, the inhibitor (I) can bind to either the free enzyme or the enzyme-substrate (ES) complex; in either case, however, the result is either the formation of (i) an inactive ternarycomplex, i.e., ES + I—*(ESI), or (ii) a binary complex (El) which (in the forward direction of the reaction) canonly lead to the formation of such an inactive complex, i.e., El + S—*(ESI). An uncompetitive inhibitor cannotbind to the free enzyme, but can only bind to and inhibit the enzyme-substrate complex, i.e., ES + I—>(ESI).Although heptenitol acted as a mixed-type inhibitor, and F2hept and F1hept acted as uncompetitive (ornoncompetitive) inhibitors, it was quite surprising that none of these heptenitols acted as competitive inhibitorsof pABG5. These heptenitols all possess a pyranosyl ring that is structurally similar to that found in normalsubstrates. The pyranosyl ring of these heptenitols contains all of the hydroxyl groups found in normalsubstrates, thus these heptenitols are presumably able to exploit all of the noncovalent interactions involving theglycone that contribute to the stability of binding of normal substrates in the enzymes active site.Noncompetitive and uncompetitive inhibition are most commonly observed in steady-statemultisubstrate systems, but are only rarely observed in unisubstrate enzymes like pABG5. However,noncompetitive inhibition of a -glucosidase has been reported, since D-glucal is a mixedcompetitive/noncompetitive inhibitor of Aspergillus wentii f3-glucosidase (Legler et al., 1979).:: ‘ ‘(a){f3G1cPNPJ —‘ (mM) —‘40(b)2:/.[F2hept] (mM)Figure 3.5. Determination of kinetic parameters for the inhibition of pABG5 byF2hept.(a) Double-reciprocal plot of kinetic data. Reactions (0.500 mL) were performed at 37 °C in 50 mM sodiumphosphate buffer (pH 6.8) with 0.051 p.g of pABG5 and 0.1% BSA. The concentrations ofF2hept in thereactions were0 (o},O.33 (.},0.49 {D},0.82(L), 1.2 (A), and 1.6 {7) mM.(b) Dixon plot of data obtained from panel (a). The concentrations of J3G1cPNP in the reactions were 0.34 (o },0.17 (.),0.10 {C), 0.069 (•}, and 0.034 {} mM.75Figure 3.6. Determination of kinetic parameters for the inhibition of pABG5 byF1hept.(a) Double-reciprocal plot of kinetic data. Reactions (0.500 mL) were performed at 37 °C in 50 mM sodiumphosphate buffer (pH 6.8) with 0,053 ig of pABG5 and 0.1% BSA. The concentrations ofF1hept in thereactions were 0 {o}, 0.16 1+1,0.49 {.} 1.6 (}, and 3.3 (•) mM.(b) Dixon plot of data obtained from panel (a). The concentrations of I3G1cPNP in the reactions were 0.34 { o),0.161.1,0.10 ([]}, 0.068 (.}, and 0.034 {} mM.4020040200—I(a)(b)-20 0 20 40{3G1cPNP] -1 (mM) -‘‘ I I I-6 -4 -2 0 2 4[F1hept] (mM)763.3.4. Kinetic studies using methylglucal.a. The synthesis ofmethyiglucal.Methyiglucal was synthesized from persilylated glucal according to the procedure of Lesimple et al.(1986). This reaction involved deprotonation at C-i of protected D-glucal, and then reaction of the lithio anionwith methyl iodide to afford the protected methylgiucal (Scheme 3.3). Tetra(n-butyl)ainmonium fluoride(TBAF) was used for the deprotection of silylated hydroxyl groups.RO70tBuLTHF[R$o IR— = (t-B:)Me2Si—Mel, —78 °CHO°TBAF, THFRO°CH3 CH3IScheme 3.3. The synthesis of methylgiucal.The low yield of the reaction was probably attributable to the use of t-butyldimethylsilyl ethers toprotect the hydroxyl groups, which may have led to the fonnation of (x-silyl carbanions (Friesen et al., 1991).The use of t-butyldiphenylsilyl or tri-isopropylsilyl ethers as hydroxyl protecting groups is now thought to be amore appropriate synthetic approach for reactions such as the one shown in Scheme 3.3 (Friesen et aL, 1991).b. Methyiglucal as a substrate ofpABG5.TLC provided initial evidence (the appearance of a spot with an Rf lower than that of the startingmaterial) that methyiglucal was hydrated by pABG5. However, methyiglucal was unstable in the 50 mMphosphate buffer (pH 6.8) that is normally used for kinetic studies with pABG5. The phosphate buffer in thereaction mixture was replaced with 10 mM F[EPES-NaOH (pH 7.0), as both the enzyme and methylgiucal were77relatively stable in this buffer. As a control, the values of kinetic parameters for the hydrolysis of the substrateI3G1cPNP were obtained using the 10 mM HEPES-NaOH buffer (Km = 75 jiM, heat = 8400 niin1), and thesevalues agreed well with those obtained using the 50 mM phosphate buffer (Km = 78 .LM, kcat = 9500mm1).The product was clearly identified by 1H NMR spectroscopy as 1,3-dideoxy-D-gluco-heptulose, thesignal atthbutable to the methyl hydrogens of the product (8= 1.43 ppm) being clearly distinguishable from thesignal attributable to the methyl hydrogens of the substrate methylglucal (5 = 1.74 ppm) (see Figs. 3.7 and 3.8).The initial rate of the hydration reaction was determined after a very lengthy incubation period (110 h).The activity of the enzyme (in control reactions without methylglucal) was checked periodically throughout thisperiod, and after 120 hours the enzyme retained over 91% of its initial activity. Reactions were performed using5 different concentrations of methylglucal. Each 0.500-mL reaction also contained 0.65 mg of pABG5, 0.1%BSA, and 10 mM HEPES-NaOH (pH 7.0). At the end of the incubation period, the 1H NMR spectrum of eachD20-exchanged reaction solution was determined and then the rate of formation of the product was calculatedfrom the 1H NMR data (see Materials and Methods Section).The values of Km and heat were determined by fitting the data to the Michae]is-Menten equation usingthe computer program GraFit (Leatherbarrow, 1990). The Lineweaver-Burk plot for this data is shown inFig. 3.9. Methyiglucal was found to be a very poor substrate for pABG5, with a Km of 57± 8 mM and akcat of0.056 ± 0.004 mm4 Given these results, the original intention of proceeding with the synthesis and kineticanalysis of fluorinated derivatives of methylglucal was abandoned.c. The stereochemistry of the catalytic hydration of inethyiglucal by pABGS.The stereochemistry of the enzyme-catalyzed addition of a proton (or deuteron) to C-3 of methylglucalwas determined by 1H NMR analysis of the hydration product derived from methyiglucal after reaction withpABG5 in aD20-containing solution. This reaction (0.500-mL) contained 14 mM methylgiucal and 0.62 mg ofpABG5 in HEPES buffer (pH 7.0). All of these solutes were dissolved inD20; substrates and buffers havingbeen lyophiized from (or in the case of the enzyme, dialysed using)D20. The reaction mixture was incubatedfor 10 days (241 h) at 37 °C, lyophilized, resuspended in D20, and then the stereochemistry of deuteriumincorporation was determined by 1HNMR spectroscopy (Figs. 3.7c and 3.8c).78UFigure 3.7. 1HNMR detennination of the stereochemistry of pABG5-catalyzed hydration of methylgiucal.400 MHz 1H NMR spectra were obtained using the samples indicated below. Spectra were recorded using asweepwidth of 5 kHz and signals were averaged over 1,000 transients. Only the relevant regions of the spectraare shown, which contain the signals attributable to equatorial (6 = 2.10 ppm, dd) or axial (6 = 1.56 ppm, t) H-3protons of the hydration reaction product, l,3-dideoxy-D-gluco-heptulose. Samples (b) and (c) were obtainedafter incubation of 14 mM methyiglucal with or without 0.31 mg/mL pABG5 for 10 days at 37° C in fullydeuterated reaction mixtures containing 10 mM HEPES buffer (pH 7.0).(a) A sample of 1,3-dideoxy-D-gluco-heptulose obtained after incubation of methylglucal for 8 days at 37° C in50 mM sodium phosphate buffer (pH 6.8). The spectrum was recorded with solvent suppression.(b) Sample without enzyme (nonenzymat.ic hydration only).(c) Sample with pABG5 (enzymatic and nonenzymatic hydration).(a)(b)(c)2.5 2.2 2.1 2. 1.9 191’ 1.6 1.5 . S 279(a,b)Without ABG5,J = HnonenzymatichydrationOR leads to either ORstereoisomerCH3R(c) DOO_______________With pABG5,t:nonenzymatichydrationleads to eitherCH3Figure 3.8. Interpretation of 1HNMR evidence for the stereochemistiy of methylglucal hydration.HOR ORenzyme-catalyzedstereospecifichydrationAxial H-3:5=1.56 ppm(t, J3a,3e = J3a,4 = 12 Hz)Equatorial H-3:8= 2.10 ppm(dd,J3a,e= 12 Hz,J3e,4 = 5 Hz)CDPanels (a,b) and (c) represent interpretations of the relevant 1F1 NMR results obtained from the correspondingreactions described in the legend of Fig. 3.7.80600040001 200:-0.02 0 0.02 0.04 0.06 0.08[methyiglucal] (mM)’Figure 3.9. Determination of Km afld Vm for pABG5-catalyzed hydration of methylgiucaLReactions (0.500 mL) were performed at 37 °C in 10 mM HEPES buffer (pH 7.0) with 0.65 mg of pABG5 and0.1% BSA. Methyiglucal concentrations were 10, 20, 40, 60, and 90 mM.Two control reactions were also carried out. In one control reaction, methylgiucal was incubated for8 days at 37 °C in anH20-containing, phosphate buffer (Figs. 3.7a and 3.8a). These conditions resulted in thenonenzymatic hydration of methyiglucal to form 1,3-dideoxy-D-gluco-heptulose. At the end of this controlreaction, water was removed by lyophilization, and after several rounds of resuspension in D20 andlyophilization, the 1H NMR spectrum was taken (Fig. 3.7a). Chemical shifts and1Hcoupling constantsobtained from this spectrum were used to assign resonances that were specific for the axial (H-3) andequatorial (H3eq) protons at C-3 of the product (see Figs. 3.7 and 3.8).0.181Another control reaction contained no enzyme (Figs. 3.7b and 3.8b), but was otherwise identical to (andwas run in parallel with) the enzymatic reaction described previously. This control reaction (no enzyme,10 days, inD20-containing buffer) yielded a 1H NMR spectrum containing resonances attributable to bothH-3 and H-3l protons. In this spectrum the integration ratio of these two signals was 1:1, indicating that thenonenzymatic hydration ofmethylglucal was not stereospecific (Figs. 3.7b and 3.8b).The parallel, enzymatic reaction (10 days, in D20-containing buffer) yielded a 1H NMR spectrum thatalso contained resonances atthbutable to both H-3 and H-3l protons. In this spectrum the integmtion ratio ofthese two signals (H-3 : H3eq) was 3:2 (Figs. 3.7c and 3.8c). During the (very slow) enzymatic hydrationreaction, spontaneous nonenzymatic hydration was unavoidable, and hence it was not surprising that the1H NMR spectrum contained resonances attributable to both H-3m’ and H-3l protons. However, the extrapresent in the enzymatic reaction must have resulted from the catalytichydration of methylglucal by pABG5 in theD20-containing buffer, which led to the increase in the [lfl..3 ax]:[lH3eq] ratio (from 1:1 to 3:2). Hence during the enzymatic hydration of methylglucal, the deuteron must haveadded from below the CL-face of the sugar ring, the same stereochemistry that has been observed during thehydration of D-glucal by pABG5 (Street, 1988).d. Methylgiucal as an inhibitor ofpABGS.A reversible inhibition test was perfonned using methylgiucal. Due to a shortage of methylglucal, onlyan approximate value of the inhibition constant (a range-finding K1 or RF K) was determined. A Dixon plot ofthe data (Fig. 3.10) yielded an RF K1 value of 40 mM, which differed from the Km value (57 ± 8 mM)determined earlier for methylglucal. However, the Km (determined using an NMR-based assay instead of amore accurate spectrophotometric assay) and the RF K1 (an approximation of the actual K1 value) were bothdetermined using methods of limited accuracy, and hence the agreement between the two values was satisfactorygiven the error limits involved in their determination.821.0 111111111 11111111130 -20 — -100iiii ii iii I I iiii I It-40 -20 0 20 40[methyiglucal] (mA4)Figure 3.10. Estimation of the K1 for the inhibition of pABG5 by methyiglucal.Reactions (0.500 mL) were performed at 37 °C in 10 mM HEPES buffer (pH 7.0) with 0.10 .tg of pABG5 and0.1% BSA. The concentration of I3G1cPNP in the reactions was 100 tiM. The concentrations of methylgiucal inthe reactions were 0, 1.0, 4.0, 10, 20, and 40 mM.3.3.5. Kinetic studies using nitroglucal.a. Nitroglucal as an inactivator ofpABG5.1-Nitroglucal (1 ,5-anhydro-2-deoxy-1 -nitro-D-arabino-hex-1-enitol; also named as 1 ,2-dideoxy-1-nitro-D-arabino-hex-1 -enopyranose by Beer et al., 1986) was the first cz,13-unsaturated derivative of glucalexamined. This compound (see below) was kindly provided by Prof. A. Vasella of the University of Zurich (seeBeer et al., 1986; Baumberger et aL, 1986).HO°831z[nitroglucal]—1 (mM)Figure 3.11. Determination of kinetic parameters for the inactivation of pABG5 by nitroglucal.(a) Time-dependent decrease in pABG5 activity with increasing concentrations of nitroglucal. Inactivationreactions were performed at 37 °C in 50 mM sodium phosphate buffer (pH 6.8) with a fixed concentration ofpABG5 and 0.1% BSA. Residual activity assays were performed as described in the text. Theconcentrations of mtroglucal in the inactivation reactions were 1.2 (+), 3.9 {x}, 5.9 (o), 7.8 {.}, and 11.7(} mM, and the values of kobs were 0.0019, 0.0044, 0.0056, 0.0069, and 0.0073 mm4,respectively.(b) A double-reciprocal plot of the kobs values from panel (a), and the nitroglucal concentrations at which theywere obtained.j—1-2-3-4-56004002000-0.20 200 400 600time800 1000 1200 1400(minutes)(a)(b)0 0.2 0.4 0.6 0.8 184When pABG5 was incubated with nitroglucal there was a time-dependent decrease in the activity of theenzyme (relative to a control reaction without nitroglucal) when aliquots were withdrawn and assayed withI3GIcPNP (Fig. 3.1 la). This time-dependent decrease in activity indicated that nitroglucal was covalentlyinactivating the enzyme, and hence experiments were perfonned as described below to determine theinactivation constants k and 1(1.Inactivation constants were determined using reactions containing one of 5 different concentrations ofnitroglucal (ranging from 1.2 to 12 mM) in 50 mM phosphate buffer and 0.1% BSA. At different time intervalssmall aliquots of each reaction mixture were removed and then assayed for residual enzyme activity (seeMaterials and Methods Section) to determine if nitroglucal-mediated inactivation of pABG5 involved theformation of a covalent bond at the active site. In these assays (which contained 1.00 mL of 1 mM f3G1cPNP)each small aliquot of the inactivation mixture (10 iL) was diluted 100-fold into a solution containing asaturating concentration of the substrate. Under these conditions, any nitroglucal which was not covalentlybound to the enzyme’s active site would be competed out of the active site by the large excess of substrate.Hence after dilution, this fraction of the enzyme (previously containing noncovalently bound nitroglucal) wouldnow be available for the catalysis of [3G1cPNP hydrolysis.The pseudo first-order rate constant (kobs) obtained at each concentration of nitroglucal was fitted to thenonlinear form of the equation kobs = k.[I]I{K1+ [I]) using the computer program GraFit (Leatherbarrow,1990). This equation was used to calculate the binding constant, K1 = 5.5 ± 0.9 mM, and the inactivation rateconstant, k1 = { 1.1 ± 0.1) x 10-2 mint. A double-reciprocal plot of the data is shown in Fig. 3.1 lb.b. Determination of the site(s) ofnitroglucal-mediated inactivation ofpABG5.Another test for the active site-directed nature of an inactivator is whether the presence of a competitiveligand decreases the rate of inactivation at a fixed concentration of the inactivator. A competitive ligand bindsto the active site of the free enzyme, thus it transiently protects the active site from inactivation, but since thebinding is reversible, any active site-directed inactivation of the enzyme should eventually go to completion.In the above-mentioned “protection” experiment that was used in this work, the competitive ligand wasl-deoxy--D-glucosylbenzene (ldGlcØ), which binds to the active site of pABG5 with a K1 = 3.4 mM (Street,85CIC1988). The enzyme pABG5 was incubated at 37 °C in solutions containing a phosphate buffer (pH 6.8), 7.8 mMnitroglucal, and either 0 or 9.9 mM ldJ3Glc(p. Residual activity assays were carried out at different time intervalsexactly as described earlier for the inactivation test.At the concentration of 1df3Glc that was used (—3 xK1), the rate of nitroglucal-mediated inactivationof pABG5 was reduced approximately 2-fold compared to the rate in the absence of ldI3GlcØ (from 0.0069 to0.0036 mind), as expected if the two ligands bind at the same site (Fig. 3.12).__-2-2.5-3-3.5-4-4.5—CC0 200 400 600time (minutes)Figure 3.12. Competitive ligand-mediated protection against nitroglucal-mediated inactivation of pABG5.Inactivation reactions were performed at 37 °C in 50 mM sodium phosphate buffer (pH 6.8) with a fixedconcentration of pABG5, 0.1% BSA, and 7.8 mM nitroglucal. Residual activity assays were performed asdescribed in the text. The concentrations of 1dGlcØ in the inactivation reactions were 0 { • } or 9.9 to) mM,and the values of kobs obtained were 0.0069 or 0.0036 mm4,respectively.86c. Mass spectrometry ofnitroglucal-inactivatedpABG5.Recent developments in protein mass spectrometry have made it possible to use this technique toidentify a catalytically important amino acid residue in an enzyme’s active site. Typically an active site-specificmodifying reagent is used to covalently modify and label the amino acid in question, and enzyme mactivatorsare frequently ideal for such applications. Mass spectrometry is then used to determine the amino acid sequenceof the modified tryptic peptide, and identify the modified amino acid. An example of this approach is the workof Miao et al. (1994), who used the inactivator 2’,4’-dinitrophenyl-2-deoxy-2-fluoro-f3-xylobioside to label andidentify Glu 78 as an active-site nucleophile in the enzyme xylanase from Bacillus subtilis.In this work, protein mass spectrometry data were collected using native pABG5 and thenitroglucal-inactivated enzyme, in collaboration with Dr. S. C. Miao and Prof. R. Aebersold of the University ofBritish Columbia. The mass spectrum of the native enzyme indicated an Mr of 51,205 ± 8. In contrast, the massspectrum of the nitroglucal-inactivated enzyme showed that this sample consisted of many species. Eachenzyme monomer was probably labeled with several molecules of nitroglucal, but the sample was soheterogeneous that the mass spectrometry data could not yield a meaningful estimate of the average number ofequivalents of nitroglucal covalently bound per enzyme monomer.Although disappointing, the above results were not surprising. Nitroglucal is a highly reactivea,f3-unsaturated glucal, and was probably able to react with several nucleophilic amino acid residues present inpABG5, including one located in the active site of the enzyme. Sample heterogeneity has also been observed inthe mass spectrum of nitroglucal-inactivated glycogen phosphorylase b (Stirtan, 1993). Multiple-sitemodification of glycogen phosphorylase b by nitroglucal has also been detected by X-ray crystallographicstudies. In these studies a covalently bound nitroglucal molecule was detected at the active site, and also at asecond site near the surface of the enzyme (Stirtan, 1993). At the second site, a covalent bond had been formedbetween the C-2 position of the 2-deoxy-3-nitroglucosyl moiety and an imidazole nitrogen of the side chain ofHis 73.87d. Is nitroglucal-inactivatedpABG5 capable of reactivation?Street (1988) showed that if pABG5 was inactivated with 2FfGlcDNP, there was a slow return ofenzymatic activity when the inactivated enzyme was removed from excess inactivator. Under these conditionsthe covalent inactivator-enzyme intermediate slowly completed the deglycosylation step of the catalytic reactionmechanism, and liberated the free enzyme. Transglycosylation acceptors such as 1d(GlcØ and cellobiose werefound to increase the rate of reactivation of 2FfGlcDNP-inactivated pABG5 (Street, 1988). Transglycosylationacceptors bind at the “second binding site” immediately adjacent to the occupied active site of f-glucosidase. Ifa compound such as idGlcØ or cellobiose is present, a covalent inactivator-enzyme intermediate undergoestransglycosylation (with a sugar as the acceptor) more rapidly than deglycosylation (with water as the acceptor),but the net result is still liberation of the free enzyme (Street, 1988).In this work, reactivation of nitroglucal-inactivated pABG5 was attempted after the removal of excessinactivator by dialysis using 50 mM phosphate buffer (pH 6.8). The “nitroglucal-free” retentate containingnitroglucal-inactivated pABG5 was divided into three equal-size aliquots. Each of these aliquots was in turnadjusted to the same volume using 50 mM phosphate buffer (pH 6.8) and either (i) nothing else; or sufficienttransglycosylation acceptor to give a final concentration of 21 mM (ii) 1d3Glc4; or (iii) cellobiose.In all three cases there was no detectable reactivation of nitroglucal-inactivated pABG5 after anincubation period of 48 hours at 37 °C.e. Possible mechanismsfor the inactivation ofpABGS by a;13-unsaturated glucals.Nitroglucal was one of several a ,f3-unsaturated glucals that were studied as potential inactivators ofpABG5. Unfortunately, the conventional ring numbering systems used for substituted glycals makes it difficultto compare the results of these studies, and the results of work carried out using glycoside inactivators. This isbecause the C-i position in an unsubstituted glycal does not maintain the designation “1” in a derivative thatbears a substituent (at C-i of the unsubstituted glycal) with one or more carbon atoms (e.g., in heptenitol, theexocyclic carbon atom is C-i; see Houlton et al., 1993). Hence for the purposes of the discussion in this chapter,it was deemed necessary to modify slightly the conventional ring numbering systems used for substituted glucalsto allow for a clearer comparison of the results obtained in this work and in previous studies (Fig. 3.13).88HO° HO° HO°1-nitroglucal 1-cyanoglucal I -(methyl carboxylate)glucalHO° HO,°NHN0 Nasodium 1-(carboxylate)glucal 2-cyanoglucal 2-acetamidoglucalFigure 3.13. Ring numbering systems used in this chapter for substituted glucals.Conventional ring numbering systems for substituted glycals that were part of this study are given in Figs. 5.1and 5.2.The structure and reactivity of c3-unsaturated glucals suggest three possible ways for thesecompounds to inactivate pABG5 (see Scheme 3.4). One possible mechanism for inactivation would involve aMichael addition reaction between the a,f3-unsaturated glucal and a nearby nucleophilic residue in the enzyme’sactive site. The reaction of 2-substituted czj3-unsaturated glucals (e.g., 2-cyanoglucal) in the active site may befavoured because with these compounds a Michael (or 1,4-) addition reaction involves C-i, and the formation ofa covalent bond between C-i of the glucal and the “normal” active-site nucleophile in pABG5 (Glu 358; seeWithers et al., 1990) may be particularly facile. Furthermore, the reaction of 1-substituted a 4-unsaturatedglucals (e.g., 1-nitroglucal) in the active site may also be possible. However, with these compounds a Michael89addition reaction involves C-2, thus although formation of a covalent bond between C-2 of the glucal and Glu358 may be less likely, another active-site nucleophile in pABG5 may be close enough to react with C-2 in thesecompounds.A second possible inactivation mechanism would involve a nucleophilic attack at the C-i position ofthe glucal to give a glycosyl-enzyme intermediate. The electron-withdrawing nature of the C-i or C-2substituent in an a,-unsaturated glucal should slow down the deglycosylation step (due to inductivedestabilization of the transition state), resulting in accumulation of the intermediate. This mechanism isanalogous to that proposed to explain the inactivation of 3-glucosithse by 2-deoxy-2-fluoro-glucosides (Street,1988).There is a third possible mechanism for the inactivation of 3-glucosidase by ct43-unsaturated glucalsthat possess a functional group with an electrophilic centre (e.g., a carbonyl group). This mechanism wouldinvolve a 1,2-addition reaction between an active-site nucleophile and the electrophilic centre of the substituent.In contrast with the results obtained with 2Ff3G1cDNP-inactivated pABG5 (Street, 1988), it was notpossible to reactivate nitroglucal-inactivated pABG5 (i.e., with 1d(GlcØ or cellobiose, see above). This suggeststhat the second of the above-mentioned mechanisms does not explain how nitroglucal mactivates pABG5. Onthe other hand, the bulky nitro group at the C-i position may have prevented water or a transglycosylationacceptor from approaching close enough to attack the C-i position of the inhibitor in the enzyme-nitroglucalintermediate. This steric hindrance would have left the nitroglucal bound to the enzyme’s active site, and hencereactivation of pABG5 would not have been possible.Another more likely explanation for the absence of any reactivation may have been that the mechanismof nitroglucal-mediated inactivation of pABG5 involved a Michael addition reaction at the C-2 position of theglucal, and that the resulting covalent bond in the glucosyl-enzyme intermediate was resistant to hydrolysis (ortransglycosylation) due to its sheltered position in the enzyme’s active site (Scheme 3.4). The results of theprotein mass spectrometry of nitroglucal-inactivated pABG5 provided support for the hypothesis that themechanism of inactivation involved a Michael addition reaction. These results were very similar to thoseobtained by Stirtan (1993) with nitroglucal-mactivated glycogen phosphorylase b.0NO2OxO/r/F// /f /7Glu358Normal catalyticglucosylationNO20 0Glu358NO2/7/_F_F /7/77Glu35890Nu7”-,,OH AHH0[Michael addition\N.OHr srOScheme 3.4. Possible mechanisms for the inactivation of pABG5 by nitroglucal.913.3.6. Kinetic studies using other a,f3-unsaturated glucals.a. Other a13-unsaturated glucals used in this study.The high reactivity of nitroglucal is attributable to the nitro substituent, which is probably the mostactivating group for Michael addition reactions (Shenhav et aL, 1970). Other less reactive cc4-unsaturatedderivatives of glucal (see Fig. 3.13) were therefore examined in an attempt to find an inactivator of pABG5 thatwould selectively label an active-site nucleophile. Two glucal derivatives (1-cyanoglucal and 2-cyanoglucal)were synthesized for this work. Two other compounds (methyl carboxylateglucal and sodiumcarboxylateglucal) were synthesized and kindly provided by Dr. William Stirtan of Prof. Wither& laboratory.b. The synthesis of 1-cyanoglucal.The synthesis of 1-cyanoglucal was accomplished using the mukistep procedure shown in Scheme 3.5.The peracetylated glucosylcyanide (3.5) was obtained by heating a neat mixture of glucosyl bromide (3.4) andmercuric cyanide (Fuchs & Lehmann, 1975). This step gave a much higher yield (60%) than other routes thatrequire the use of a solvent (e.g., 12%, Coxon & Fletcher Jr., 1963; 20%, Myers & Lee, 1984).Photobromination of 3.5 was accomplished using N-bromosuccinimide (Lichtenthaler & Jarglis, 1982), and thisstep was performed by Mr. K. Mok in Prof. Withers’ laboratory. The product, 2,3,4,6-telra-O-acetyl-a-bromo-(3-D-glucosylcyanide (3.6), was formed stereospecificafly.Peracetylated 1-cyano-D-glucal (3.7) was prepared from 3.6 under aprotic conditions using zinc andone equivalent of pyridine. Somsák et al. (1990) published this procedure, and noted its superiority over themethod involving zinc dust and acetic acid (Roth & Pigman, 1963). Use of the latter method to prepare 3.7leads to a mixture of a- and f3-glucosylcyanides as side products, and chromatographic separation of theseanomers from the desired glucal is difficult (Somsâk et al., 1990). Deacetylation of the base-sensitive compound3.7 was accomplished using ammonia-saturated methanol under mild conditions (see Fritz et al., 1983).92AcO”Hg(CN)2Br NBS,hv[3.4] [3.5]OAc0AcO CNAcO AcOBr[3.6]/Zn pyr.benz , AOH OAcHO NH3, MeOH AcOHO _0°CAcO C’CN CN[3.8] [3.7]Scheme 3.5. The synthesis of 1-cyanoglucal.c. The synthesis of2-cyanoglucal.Peracetylated 2-cyanoglucal was synthesized from peracetylated D-glucal (see Scheme 3.6) usingchiorosulfonyl isocyanate and subsequent trealment with triethylamine (Hall & Jordaan, 1973). Deacetylationwas accomplished as before using ammonia-saturated methanol under mild conditions (Fritz et al., 1983).In the first step of the reaction sequence shown in Scheme 3.6, an N-chlorosulfonyl carboxamidederivative of glucal was probably formed initially, prior to the elimination of chlorosulfuric acid bytriethylainine to yield the unsaturated nitrile. The low overall yield (27%) of the desired product was probably aconsequence of the competing side-reaction shown. In this side-reaction the 3-acetoxy group of the startingmaterial (peracetylated D-glucal) was probably cleaved with the assistance of the trans 4-acetoxy group, which93led to the formation of the dimer 3.9 (Hall et al., 1973). Hall & Jordaan (1973) found that a similar side-reactionwas much less of a problem when peracetylated galactal was used to synthesize 2-cyanogalactal (which theyobtained with a 60% overall yield), as in this case the C-3 and C-4 acetoxy groups were cis to one another,precluding trans-assisted dimer fonnation (i.e., analogous to 3.9).OAcAcOAcO _OAcCIEt3N, 0°C — RTN”NH3, MeOH0°COAcAcOAcO _N”OAcAcO0II0= C= N—S— CI, ether, 0°CII0[3.9]OAcH(OHScheme 3.6. The synthesis of 2-cyanoglucal.941812____60-12 -6 0 6 12 18 24 30[1 -cyanoglucall (mM)Figure 3.14. Estimation of the K1 for the inhibition of pABG5 by 1-cyanoglucal.Reactions (0.500 mL) were performed at 37 °C in 50 mM sodium phosphate buffer (pH 6.8) with 0.05 jig ofpABG5 and 0.1% BSA. The concentration of [3G1cPNP in the reactions was 83 jiM. The concentration of1-cyanoglucal in the reactions was 0, 1.7, 3.4, 4.3, 8.4, 16.9, and 25.3 mM.d. 1-Cyanoglucal as a reversible inhibitor ofpABG5.1-Cyanoglucal was examined as a potential inactivator of pABG5 by incubating the enzyme with33 mM 1-cyanoglucal and the appropriate reaction buffer for 8 hours at 37 °C (see Materials and Methods). Atvarious time intervals small aliquots were removed and residual enzyme activity was assayed using the substrate3GlcPNP. No time-dependent inactivation of pABG5 was detected during these tests when compared withparallel control reactions. The 1 -cyanoglucal remained unchanged (as determined by TLC) throughout thisincubation period, which indicated that the compound was also not a substrate of the enzyme. Further kineticstudies showed that 1-cyanoglucal acted as a reversible inhibitor of pABG5-catalyzed hydrolysis of 3GlcPNP.An approximate value for the inhibition constant, K1 9.0 mM, was obtained from a Dixon plot of the kineticdata (Fig. 3.14).954020-100 -80 -60 -40 -20 0 20 40 60(b)— = v[sodium 1 -(carboxylate)glucal] (mM)Figure 3.15. Estimation of the K1 values for methylcarboxylate glucal and sodium carboxylate glucal.In both cases reactions were performed as described in Fig. 3.14 and as specified further below:(a) RF K1 determination for the inhibition of pABG5 by methylcarboxylate glucal. The concentration ofI3G1cPNP in the reactions was 67 (•) or 101 (a) jiM. The concentration of methylcarboxylate glucal in thereactions was 0, 1.7, 2.3, 3.3, 6.6, and 9.9 mM.(a) RF K determination for the inhibition of pABG5 by sodium carboxylate glucal. The concentration ofG1cPNP in the reactions was 101 jiM. The concentration of sodium carboxylate glucal in the reactions was0, 30, and 60 mM.10(a)-10 -5 0 5 10[1-(methyl carboxylate)glucall1614(mM)12108696e. Other a,/3-unsaturated glucals as reversible inhibitors ofpABGS.1-(Methyl carboxylate)glucal and sodium 1-(carboxylate)glucal are a,3-unsaturated derivatives ofglucal that bear a carboxyl substituent at the C-i position of the ring (see Fig. 3.13). Each compound wasexamined as a potential inactivator of pABG5 by incubating the enzyme with 10 or 11 mM methylcarboxylateglucal or sodium carboxylateglucal, respectively, and the appropriate reaction buffer for 24 hours at37 °C (see Materials and Methods). At various time intervals small aliquots were removed and residual enzymeactivity was assayed using the substrate 3GlcPNP. No time-dependent inactivation of pABG5 was detectedduring these tests when compared with parallel control reactions. Both of these a4-unsaturated derivatives ofglucal remained unchanged (as determined by TLC) throughout their respective incubation periods, whichindicated that neither compound acted as a substrate of the enzyme.(a) I GIu170 (b) [GIyl7oICOO HOH OH 00CHO0,0_QHO0,o_b[3.10] OOC [3.10]COO— COO—I IGlu 358 Glu 358Figure 3.16. Minimization of electrostatic repulsion in the active site of the E17OG variant form of pABG5.Shown above are possible orientations of the aglycone moiety of 3.10, 2’-carboxyphenyl f3-D-glucoside, in theactive site of (a) wild-type, and (b) the site-directed E17OG variant, of pABG5 3-glucosidase (Q.P. Wang, pers.comm.). Carboxylate groups are shown in their fully ionized forms.97Reversible inhibition tests were then performed using either methyl carboxylateglucal or sodiumcarboxylateglucal (Fig. 3.15). Methyl carboxylateglucal bound to the enzyme with an inhibition constant, K1, of—3.2mM (Fig. 3.15a). Sodium carboxylateglucal bound very poorly to the enzyme, with an inhibition constant,K, of —96 mM (Fig. 3.15b).The finding that sodium carboxylateglucal bound very poorly to pABG5 agreed well with unpublishedresults recently obtained by Dr. Qingping Wang of Prof. Withers’ laboratory. Glycosides with an aglyconemoiety bearing a negatively charged carboxylate group (such as 3.10) bound very poorly to pABG5 (Q.P. Wang,pers. comm.). Kinetic studies have shown that although 3.10 binds very poorly to the active site of wild-typepABG5 (K1 = 60 mM), this glycoside binds about 65 times more strongly (K1 = 0.92 mM) to a variant form ofthe enzyme obtained by site-directed mutagenesis (Q.P. Wang, pers. comm.). In the site-directed mutant protein(E17OG) an active-site glutamate residue (Glu 170) had been replaced by a glycine residue, which removed onenegative charge from the active site of this variant of pABG5 (see Fig. 3.16). Hence the very poor binding ofnegatively charged compounds such as sodium carboxylateglucal and 3.10 to wild-type pABG5 appears to be aconsequence of electrostatic repulsion between the carboxylate group on the sugar and catalytically importantcarboxylate residues in the active site of the enzyme.f 2-Cyanoglucal as a reversible inhibitor ofpABG5.When the C-i derivatized a43-unsaturated glucals (1-nitro-, 1-cyano-, methyl carboxylate-, and sodiumcarboxylate- glucal) under study were tested as inactivators of pABG5 they were not reactive enough, or in onecase (1-nitroglucal) too reactive, to act as specific labeling reagents for an active-site nucleophile in the enzyme.In most cases C-i derivatized x,f-unsaturated glucals might not function as inactivators because the mechanisminvolves a Michael addition reaction, one that would require the formation of a covalent bond between theenzyme and the C-2 position of the sugar (see Scheme 3.4). Perhaps C-i derivatized a43-unsaturated glucalshave difficulty binding to the enzyme in such a manner as to bring the C-2 position of the sugar close enough toa potentially reactive nucleophile in the active site, and hence only exceptionally reactive C-i derivatizedcz,J3-unsaturated glucals (such as 1-nitroglucal) are able to react.98 Hence for steric reasons C-2 derivatized a,p-unsaturated glucals might be better able to act as Michael acceptors for an active-site nucleophile in pABG5. In this case a Michael addition reaction would involve attack by an active-site nucleophile of pABG5 at the "usual", C~l position of the glucal. However, this would require that a bulky substituent at C-2 be accommodated in the active site. Nevertheless it was worth investigating. Several attempts were made to synthesize 2-nitroglucaI using the procedure of Holzapfel et al. (1988), but only the peracetylated form of this compound was stable. Unfortunately, even under the mildest deprotection conditions, 2-nitroglucal was simply too reactive, and decomposed spontaneously into several different compounds. I I I ' 1 I ' I • I I I I I I I I I I I I I I I I y = -20 -10 0 10 20 30 40 50 60 70 80 [2-cyanoglucal] (mM) Figure 3.17. Estimation of the K{ for the inhibition of pABG5 by 2-cyanoglucal. Reactions were performed as described in Fig. 3.14 and as specified further below. The concentrations of (3GlcPNP in the reactions were 67 {D}, 101 {A} or 170 {O} \i,M. The concentrations of 2-cyanoglucal in the reactions were 0, 8.3,13, 25,33, and 67 mM. 992-Cyanoglucal was therefore synthesized, and then tested as a potential inactivator of pABG5 byincubating the enzyme with 33 mM 2-cyanoglucal and the appropriate reaction buffer for 20 hours at 37 °C (seeMaterials and Methods). At various time intervals small aLiquots were removed and residual enzyme activitywas assayed using the substrate 3GlcPNP. No time-dependent inactivation of pABG5 was detected during thesetests when compared with parallel control reactions. The 2-cyanoglucal remained unchanged (as determined byTLC) throughout this incubation period, which indicated that the compound was also not a substrate of theenzyme.Reversible inhibition tests were then performed using 4-5 different concentrations of 2-cyanoglucal and3 different concentrations of the substrate f3G1cPNP. The initial-rate data obtained at each substrate andinhibitor concentration were fitted to equations describing different types of enzyme inhibition, using thecomputer program GraFit (Leatherbarrow, 1990). The best fit of the data was to the equation describingcompetitive inhibition, which yielded a K value of 26± 3 mM. The Dixon plot obtained from the data is shownin Fig. 3.17, and this graphical approximation method yielded a similar value for the K1. Although2-cyanoglucal acted as a competitive inhibitor of pABG5, it bound poorly to the enzyme’s active, and itsK1 =26± 3 mM represented even poorer binding to the enzyme than was observed with 1-cyanoglucal (RF K =9.0 mM). The poorer affinity of 2-cyanoglucal for the active site of pABG5 must have been a consequence ofeither the size, geometry, or properties of the nitrile group at the C-2 position of this sugar. In many ways this isnot surprising given the known specificity of this enzyme for interactions at the C-2 position (Namchuk, 1993).3.3.7. Kinetic studies using 2-acetamidoglucal.a. The selective deprotection ofperacetylated 2-acetamidoglucal.The peracetylated precursor (3.11) of 2-acetamidoglucal was synthesized from G1cNAc by JohnMcCarter of Prof. Withers’ laboratory, using the procedure of Pravdi et al. (1975). In this study theperacetylated precursor was first purified by column chromatography. Deacetylation of 3.11 using sodiummethoxide in methanol (Pravdi & Fletcher Jr., 1967) selectively removed one of the two N-acetyl groups andall three O-acetyl groups to yield the desired product (3.12).100AcO,° . HO°(Ac)2N HN,CH3[3.11] [3.12]Scheme 3.7. The selective deprotection of peracetylated 2-acetamidoglucal.b. The inability of2-acetamidoglucal to bind to pABG5.Inactivation tests and reversible inhibition tests were performed using 2-acetainidoglucal and pABG5.This compound did not inactivate or even bind to the enzyme, even at 2-acetamidoglucal concentrations as highas 33 mM. The reversible inhibition tests also showed that there was no effect on enzyme activity, even at2-acetamidoglucal concentrations as high as 33 mM.The inability of 2-acetamidoglucal to bind to pABG5 was probably due to the presence of the N-acetylgroup at the C-2 position. Comparable results have been reported for the interaction between 2-deoxy-2-acetamidoglucose and sweet almond f3-glucosidase (Dale et al., 1985). An inhibition constant, K, of> 900 mMwas obtained, and this extremely poor affinity was attributed to the polar nature of the acetyl group (Dale et al.,1985). Sweet almond f3 -glucosidase displays much greater affinity for compounds such as 2-deoxy-2-[(p-chlorobenzyl)amino]glucose (K = 3.0 mM) and 2-deoxy-2-[(p-methoxybenzyl)amino]glucose (K = 5.9mill), where the acetyl group has been replaced by a nonpolar benzyl group (Dale et al., 1985).3.4. CONCLUSIONS.Heptenitol and two fluorinated derivatives (F2hept andF1hept) were synthesized and kinetic studieswere performed using a cloned Agrobacterium f3-glucosidase, pABG5. Heptenitol showed a mixed inhibitionpattern. F2hept andF1hept both acted as uncompetitive (or noncompetitive) inhibitors, and there was nodetectable binding of eitherF2hept orF1hept in the active site of pABG5. There were no changes in the101thin-layer chromatograms or 19F NMR spectra of the reaction mixtures after prolonged incubation of pABG5with either of the fluoroheptenitols. The fluoroheptenitols did not act as substrates or inactivators of pABG5.A comparison of K values showed that heptenitol, with two olefinic hydrogen atoms at C-i, did notbind to pABG5 as strongly as the more polar F1hept andF2hept (Table 3.1), whose only structural difference(s)with the parent compound are the presence of one or two olefmic fluorine atoms at C-i, respectively. Fluorine isnot only the more polar of the olefinic C-i substituents in these compounds (and thus more likely to strengtheninteractions between the compound and a polar environment in the enzyme); fluorine is also, unlike hydrogen,capable of acting as a hydrogen-bond acceptor. Hydrogen bonding involving such an acceptor and ahydrogen-bond donor on the enzyme may in part account for the increased affinity of pABG5 forfluoroheptenitols compared with heptenitol.Heptenitol was catalytically hydrated by pABG5 to yield 1-deoxy--D-gluco-heptulose as the initialproduct. Although this hydration reaction was reasonably fast (keat = 640 min* or —7% of the rate of I3GIcPNPhydrolysis), the affinity of heptenitol for the active site of pABG5 was extremely low (Km = 270mM).Heptenitols have been tested as substrates with other 3-glycosidases. As an example, E. coli13-D-galactosidase hydrates D-galacto-heptenitol with a Km = —50-70 mM and a kcat = 2460-3840 mind, or—1-2% of the rate of f3-galactoside hydrolysis by the same enzyme (Brockhaus & Lehmann, 1977). However,this author is unaware of any studies that have tested heptenitols as inhibitors of -glycosidases. Hence it wasnot possible to compare the type of inhibition of pABG5 displayed by heptenitol with published results obtainedusing other f -glycosidases. It is therefore unclear if heptenitol (and the other heptenitol derivatives studiedherein) would have the same effects on 13-glucosidases from other sources as these compounds had on pABG5.Further kinetic studies using 13-glucosidases and cc-glucosidases (given the lack of an anomeric carbon at the C-iposition of heptenitols) from other sources would help to answer these questions.Methylgiucal acted as a very poor substrate of pABG5, yielding 1,3-dideoxy-D-gluco-heptulose as theproduct of the hydration reaction. A Km of 57 mM and a keat of 0.056 min1 were obtained, thus methyiglucalwas —2400- or [1.2 x 108j fold less effective as a substrate compared with heptenitol or I3G1cPNP, respectively(based on the ratio of their kcatlKm values). Methylgiucal itself was unstable, and the rate of its spontaneous102hydration was quite significant, especially in phosphate-containing buffers. Presumably the poor turnover ofmethylgiucal is a consequence of several factors. Firstly, methyiglucal lacks a C-3 hydroxyl group, and enzyme-substrate interactions involving the comparable position in a glycoside (in this case the C-2 hydroxyl) have beenshown to contribute 8 kcallmol or more to transition-state stabilization in the reactions catalyzed by pABG5(Namchuk, 1993) and other enzymes (McCarter et al., 1992). Secondly, the C-i methyl group may hinderapproach of the nucleophile at C-2 (in methylgiucal; the comparable position in a glycoside being C-i), or itmay interact unfavourably with other active-site groups upon conversion to the glycosyl-enzyme intermediate.Thirdly, the apolar C-i methyl group may not fit well into the generally polar “aglycone pocket” (adjacent to theactive site) expected in a glycosidase.1-Nitroglucal, an czj3-unsaturated glucal, acted as an inactivator of pABG5. Kinetic studies showedthat inactivation occurred at the active site, most likely as a result of a Michael addition reaction between anactive-site nucleophile and C-2 of the inactivator. The mass spectrum of nitroglucal-inactivated pABG5 showedthat the sample was quite heterogeneous, indicating that on average several equivalents of nitroglucal werecovalently bound to the enzyme. The synthesis of radiolabelled nitroglucal may prove useful in future studiesdirected at identifying active-site amino-acid residues of glycosylases, and thereby confirm that a Michaeladdition reaction provides the mechanistic basis for the inactivation behaviour of reactive cs,3-unsaturatedglucals.Unfortunately, the other cz,13-unsaturated glucals that were tested (1-cyano-, methyl carboxylate-,sodium carboxylate-, and 2-cyano- glucal) did not act as inactivators or substrates of pABG5. These compoundsonly acted as relatively poor-binding, reversible inhibitors of the enzyme.For a summary of the kinetic data obtained using pABG5, please see Table 3.1.Table 3.1. Summary of kinetic data obtained using a cloned f3-glucosidase, pABG5.Compound Structure Inhibition Inactivation SubstrateI3G1cPNP A Km=78JJMkcat= 9,500 mindheptenitol B K1 = 13 ± 1 mM no Km =270±40mMRi = R2 = H (for noncompetitive) keat 640±60 mm4F2hept B K1 = 0.67 ± 0.04 mM no noRi = R2 = F (for noncompetitive)Fihept B K1=3.2±0.2mM no noRi = H (for noncompetitive)R2 = Fmethylgiucal C K1 -40 mM no Km 57±8 mMR3 = H kcat = 0.056 miir1R4 = CH3 ± o.oo4 miir11-nitroglucal C K1 - 4.8 mM K1 = 5.5 ± 0.9 mM noR3H k=0.011minR4 = N02 ± 0.001 mind1-cyanoglucal C I( — 9.0 mM no noR3 = HR4 = CN1-(methyl C K — 3.2mM no nocarboxylate) R3 = Hglucal R4 = COOCH3sodium C K —96 mM no no1-(carboxylate) R3 = Hglucal R4 = C00 Na2-cyanoglucal C K1 =26±3 mM no noR3 = CN (competitive)R4=H2-acetamidoglucal C no no noR3 = NHCOCH3R4=HOHHO’NO2H0*<RiOH R4HO R3A B C103104CHAPTER 4: KINETIC STUDiES USING [3-N-ACETYLHEXOSAMINIDASE (NAGase)4.1. INTRODUCTION.4.1.1. Some general properties of —N—acetylhexosaminidase.j3—N-Acetylhexosaminidase (2-acetamido-2-deoxy--D-hexoside acetamidodeoxyhexohydrolase, E.C.3.2.1.52)—abbreviated herein as NAGase—is an enzyme that is widely disthbuted in nature, and can be isolatedfrom microbial, plant, and animal sources (Stirling, 1983). The enzyme possesses both f—N-acetylglucos.aminidase and [3—N--acetylgalactosaminidase activities, and the former activity was first discovered by Helferich& Iloff (1933) in an emulsion prepared from almonds. A f3—N—acetylglucosaminidase activity was first detectedin mammalian tissue by Watanabe (1936).Mammalian lysosomal NAGases are dimeric enzymes, consisting of a- or u-subunits held together bynoncovalent forces. The relative molecular mass of each a and f3 subunit is in the [5-71 x i0 range(Conzelmann & Sandhoff, 1987). There are three isozymes of human lysosomal NAGase; NAGase A is anheterodimer, whereas the two homodimeric isozymes, NAGase B and NAGase S, are composed of 43 and aasubunits, respectively. NAGase S is unstable and appears to have only negligible enzymatic activity. NAGaseA and NAGase B have isoelecthc points near pH 5 and between pH 7.0-7.5, respectively, and hence are referredto as the acid and neutral forms of the enzyme (Conzelmann & Sandhoff, 1987). This difference in isoelectricpoints has been exploited in biochemical procedures (ion-exchange chromatography and isoelectric focusing) forthe purification of these two clinically important isozymes of NAGase (Dance et aL, 1970; Sandhoff, 1968).NAGase A and NAGase B have different substrate specificities, and they also differ in their ability towithstand elevated temperatures (Okada & O’Brien, 1969). NAGase B is a heat-stable enzyme that is mostactive towards neutral substrates such as globoside, glycoproteins, glycosaminoglycans, and oligosaccharides.NAGase A is less stable at elevated temperatures, and is most active towards Gm2 gangliosides (see Fig. 4.1), aswell as acidic substrates such as glucuronic acid-containing oligosaccharides, and even 6-sulfo-glucosaminides(Bearpark & Stirling, 1978; Ludolph et al., 1981). However, in vitro studies by Legler et al. (1991) have shown105that NAGase A and NAGase B display similar kinetic properties and pH-activity profiles, with the exceptionthat NAGase B is inactive towards acidic substrates, e.g., the 6-sulfate ester of N-acetyl-f3-glucosaminide.HO’eHO”1CeramideHOGaIOHOHH&OH€0AcNHNAGase A+COOHH•HOHOHH2COHFigure 4.1. The catalytic removal of the GaINAc residue from ganglioside Gm2 by NAGase A.106The ability of NAGase A to remove the N-acetylgalactosamine residue from Gm2 gangliosides isparticularly important (see Fig. 4.1). Gangliosides are sugar-containing lipids found in high concentrations inthe nervous system (especially in gray matter), and they are important constituents of neuronal membranes.Inside lysosomes (a type of subcellular organelle) gangliosides are enzymatically degraded by the sequentialremoval of their sugars, and disorders of ganglioside breakdown can have extremely serious clinicalconsequences.In vivo, the interaction between the water-soluble enzyme NAGase A and its membrane-boundglycolipid substrate is mediated by the noncatalytic activity of the Gm2-activator protein (Conzelmann et al.,1982). This activator protein binds to the a-subunit of the enzyme, but not to the 3-subunit. The amphipathicGm2-activator protein is sufficiently hydrophilic to bind to NAGase A, yet it is also sufficiently lipophilic tobind membrane-bound Gm2 gangliosides and solubiize them as [Gm2-activator protein].—lipid complexes.4.1.2. The clinical significance of 13.N-acetylhexosaminidases.a. Tay-Sachs disease and Sandizoffdisease.In the hereditary disorder known as Tay-Sachs disease there is a deficiency in NAGase A activity dueto a defect in the production of the a-subunit (Okada & O’Brien, 1969). The symptoms of Tay-Sachs disease areusually evident before an affected infant is one year old. Weakness and retarded psychomotor development aretypical early symptoms. By the age of two, the child is demented and blind, and death before the age of four isthe inevitable clinical outcome. The ganglioside content of the brain of an infant with Tay-Sachs disease isgreatly elevated due to the deficiency of NAGase A activity.Sandhoff disease is a hereditary disorder that is similar to but much rarer than Tay-Sachs disease. InSandhoff disease there are deficiencies in both NAGase A and NAGase B activities due to a defect in theproduction of the 13-subunit (Sandhoff et al., 1968). Both of these neuronal lipid storage diseases have similarsymptoms and clinical outcomes, but Sandhoff disease is also characterized by the abnormal accumulation ofglobosides, glycopeptides, and oligosaccharides in extraneural tissues.107b. Abnormal levels ofNAGase activity in cancer cells.Some serum glycosylases, and especially NAGases, are produced at elevated levels and secreted intothe extracellular medium by many different tumor cell types in Vitro (Bernacki et al., 1985). These observationshave led to a great deal of interest in the use of glycosylase inhibitors as pharmaceuticals. 2-Acetamidoglucal(NAGlucal) is an example of a NAGase inhibitor that has been evaluated as a therapeutic drug for Lewis lungcarcinoma in mice (Bemacki et aL, 1985). The administration of 100 mg of NAGlucallkg body weight/dayintrapeneoneally for five consecutive days (days 3-8 following the subcutaneous implant of a tumor in C57B l/6J female mice) resulted in a 63% increase in the life span of drug-treated mice compared to untreatedcontrols.4.2. AIMS OF THIS STUDY.Several kinetic studies have been performed using various glycosylases and glycal derivatives (seeChapter 1). However, very few studies have been performed using NAGases and glycal derivatives such asNAGlucal. As mentioned above, Bernacki et al. (1985) noted the potential efficacy of NAGlucal as achemotherapeutic agent. Another report briefly mentioned that 2-acetamidoglucal and 2-acetamidogalactal arereversible inhibitors of NAGase from boar epididymis, with inhibition constants, K, of 58 and 40 IIM,respectively (Pokorny et al., 1975).NAGase-catalyzed reactions probably follow the same mechanism as those catalyzed by other(3-retaining glycosylases. However, no X-ray structural data is available for NAGases, and thus no informationis available about catalytically important amino acids in the active sites of these enzymes. Furthermore,although some metal ions (such as Ag and Hg2j are known to inactivate NAGases (Li & Li, 1970), nomechanism-based inactivators appear to have been discovered for these clinically important enzymes. Legler &Bollhagen (1992) did find that N-acetylconduramine B trans epoxide (see Scheme 1.6) acts as a reversibleinhibitor of NAGases from various sources, with values of 0.5 to 1.6 j.iM. However, in contrast to theinteraction of other glycosidases with cyclohexane polyol epoxides with the appropriate configuration (seeChapter 1), no covalent, irreversible inhibition is observed.108In this thesis, three different NAGases—isolated from jack bean, bovine kidney, or human placenta(J-, K-, and H-NAGase, respectively)—were studied. It was felt that NAGlucal might inactivate one of theseNAGases, in the same way that D-glucal mactivates A. wentii 13-glucosidase (Legler et al. 1979). This wouldoccur if the rate of deglycosylation is much slower than the rate of glycosylation (see Scheme 1.3). If this werethe case, it might be possible to trap an N-acetylglucosaminyl-NAGase intermediate, and identify a nucleophiicresidue in the enzymes active site.NAGlucal might instead act as a substrate of one of the NAGases under study. This would occur if therates of both deglycosylation and glycosylation are reasonably fast. If this were the case, the stereochemistry ofthe hydration reaction could be studied. In addition, the effects of the 2-acetainido substituent on the kinetics ofthe hydration reaction could also be studied, and these results could be compared with those obtained using otherglycal derivatives and glycosides.NAGlucal should at least act as a reversible inhibitor of one of the NAGases under study (and reducethe rate of hydrolysis of an N-acetylglucosaminide substrate, 3GlcNAcPNP). Kinetic and structural studiescarried out using enzyme inhibitors often yield insights into the nature of the binding interactions between theinhibitor and the active site.This study also determined the stereochemistry of the NAGase-catalyzed hydrolysis of 3GlcNAcPNP(4’-nitrophenyl 2-acetainido-2-deoxy-f3-D-glucosaminide), a commonly used substrate for the assay of NAGaseactivity.4.3. RESULTSAND DISCUSSION.4.3.1. A “direct” colorimetric assay for NAGase-catalyzed hydrolysis of I3GIcNAcPNP.In previously published studies NAGase activity has been assayed using fluorometric or colorimetricmethods. In the fluorometric assay, 4-methylumbelliferyl 2-acetamido-2-deoxy43-D-glucosaminide(I3G1cNAcMu) is used as a substrate, and fluorometry is used to determine the concentration of the aglyconeproduct, 4-methylumbelliferone, liberated as a result of enzyme-catalyzed hydrolysis.109In the colorirnetric assay of NAGase activity, a chromogenic substrate such as 4-nitrophenyl2-acetamido-2-deoxy-3-D-glucosaminide ([IG1cNAcPNP) is used. The concentration of the aglycone product(p-nitrophenol) liberated as a result of the enzyme-catalyzed hydrolysis of I3G1cNAcPNP is measured at awavelength of 400 urn, where there is very little absorbance by the substrate. p-Nitrophenol has a pKa of 7.15,and it is only the p-nitrophenolate anion that absorbs 400-urn light. However, the pH optimum for NAGaseactivity is usually pH 5.0; thus the percentage of the liberated aglycone in the anionic, light-absorbing form isless than 1% of the total product at this pH, resulting in small absorbance changes. The colorimetric assaytherefore requires the addition of an alkaline “stop” buffer to raise the pH well above the PKa ofp-nitrophenol inorder to determine the concentration of the product.There are several disadvantages associated with a “stopped” colorirnetric assay of enzyme activity. A“stopped” assay is not as convenient as a “direct” assay of the product. A “stopped” assay also requires that thereaction rate is linear over the time period in question, and any variation in the reaction rate during this periodwill seriously affect the results. For these reasons a “direct”, continuous colorimetric assay of NAGase activitywas developed for this work, based on the procedure of Kempton & Withers (1992).This direct assay involved monitoring the reaction (carried out in a cuvette thermostatted at 25 °C) at awavelength of 360 nm. This wavelength was chosen after recording the absorption spectrum of a standardsolution of f3G1cNAcPNP before and after hydrolysis at pH 5.0 (performed by adding a small volume of enzymesolution). At pH 5.0 the difference in the molar extinction coefficients of the product and the substrate of thereaction (, and respectively) was greatest at 360 um. The values of £1, and , and Ar = (e — r) at 360 urnwere then calculated at pH 5.0 (for the assay of J-NAGase) and pH 4.25 (for the assay of K-NAGase andH-NAGase) using carefully prepared standard solutions of the product and the substrate. Because of thestoichiometry of the hydrolysis reaction, the concentration of the substrate consumed (—AC) is equal to theconcentration of the product formed (AC) during the reaction, and hence (where 1= path length of 1.00 cm):= change inA360 during the reaction= AAp + AA = the change inA360 due to the formation of the product plusthe change inA360 due to the consumption of the substrate.= e•(AC)’l + r•(—AC)•l110= [Ep —=and hence the initial rate of product formation, v, can be measured as follows:v = (C)Imin= [AA/min] ÷ {i.l}The kinetic parameters Km and Vmax describing 3GIcNAcPNP hydrolysis were determined by fittingthe initial-rate data to the nonlinear form of the Michaelis-Menten equation using the computer programGraFit (Leatherbarrow, 1990). The initial-rate data for the hydrolysis of f3G1cNAcPNP by J-NAGase isshown in Lineweaver-Burk form in Fig. 4.2. The values of Km and Vmax for the hydrolysis of 3GlcNAcPNP byeach of the NAGases under study are given in Table 4.2. The values of kinetic parameters determined for othercompounds using NAGase are also listed.0.080.06‘ 0.04.‘ 0.0200[3G1cNAcPNP] -lFigure 4.2. Determination of Km and Vaç for J-NAGase-catalyzed hydrolysis of I3G1cNAcPNP-2 2 4(mM)’Reactions were performed at pH 5.0 and 25 °C in 50 ruM citrate buffer containing 0.1% BSA, 100 mM NaCI,and 1.0 p.g/mL of f -N-acetythexosaminidase from jack beans. The concentration of 3G1cNAcPNP in thereactions was 0.24, 0.34, 0.49, 0.78, 1.36, and 1.95 mM.111Ac13-retainingH’/’+ 1f1f3GIcNAcPNPOH 4% NO2HO°,..C__Scheme 4.1. The stereochemistry of I3G1cNAcPNP hydrolysis by 13-retaining and 13-inverting NAGases.The first 3-N-acetylhexosaminidase studied was J-NAGase. The isolation, crystallization, and generalproperties of J-NAGase have been reported by Li & Li (1970). The Km value obtained for J-NAGase-catalyzedhydrolysis of [3G1cNAcPNP using the “direct” colorimethc assay (0.62 ± 0.04 mM) agrees well with the valueobtained by Li & Li (1970) for the same reaction using a “stopped” assay (0.64 mM). The Vmax value obtainedusing the “direct” assay, 50± 1 U/mg, is also very close to the vendor’s (Sigma Chemical Co.) specification forthis enzyme-catalyzed reaction, 53 U/mg. It is therefore reasonable to conclude that the “direct” colorimetricassay yields reliable values for enzyme kinetic parameters.4.3.2. The stereochemistry of J-NAGase-catalyzed hydrolysis of 13G1cNAcPNP.Koshland (1953) was the first to propose that there are important differences between the reactionmechanisms of retaining and inverting glycosidases. The reactions catalyzed by retaining glycosidases probablyinvolve a double-displacement mechanism (see Scheme 1.3). In this reaction mechanism an amino acid residueacting as a nucleophile is involved in the formation of a glycosyl-enzyme intermediate, and this intermediate isOHHO_0NO2NHo=çOH13-invertingOH+OH NO2NH[aGIcNAcI112subsequently attacked by a water molecule. In contrast, the reactions catalyzed by inverting glycosidasesprobably involve a single-displacement mechanism, where a water molecule acts as the nucleophile that directlydisplaces the aglycone leaving group (Koshland, 1953).Determination of the stereochemistry of catalysis by NAGase is a prerequisite to understanding thereaction mechanism. It appears that the stereochemistry of the hydrolysis of f3G1cNAcPNP by jack beanNAGase has not been reported. Indeed only one NAGase, isolated from boar epididymis, has been subjected tosuch an analysis: Ya Khorlin et al. (1972) used 1H NMR to show that this NAGase is a 13-retaining enzyme.In this thesis, HPLC was used as an analytical tool to determine the stereochemistry of the hydrolysis off3G1cNAcPNP by J-NAGase. A DextroPak® column (operated at room temperature using 100% H20 as theeluant) was able to resolve the two anomers of 2-acetamido-2-deoxy-D-glucopyranose (GlcNAc). This columnwas used to determine which anomer was the initial product of the reaction (i.e., before a/f3 equilibration).When an equilibrated standard solution of GIcNAc was analyzed by HPLC, two peaks (with retention times of3.38 and 3.55 minutes) eluted from the DextroPak® column (Fig. 4.3c). When a freshly prepared anomericallypure standard solution of aGlcNAc was analyzed by HPLC, a single peak (with a retention time of 3.55minutes) eluted from the same column (not shown), identifying the first and second peaks as being attributableto the elution of the 13— and a— anomers of GlcNAc, respectively. The anomeric configuration of the freshlyprepared aGlcNAc standard (prepared inD20) was confirmed by 1H NMR (8 of H-i = 5.2 ppm, J1,2 = 3 Hz)and by determination of the melting point of the solid (203-205 °C; lit. mp 211 °C, Aldrich Chemical Co. Cat.).If the standard solution of aGlcNAc was left standing prior to HPLC analysis, the sample formed an equilibrium(a/13) mixture, and the f3-anomer appeared as a shoulder of the ct-anomer peak (as in Fig.4.3c). 1H NMRanalysis of the equilibrium mixture indicated that the a-anomer was the major component.When aliquots of an incubation mixture of f3G1cNAcPNP and J-NAGase were analyzed by HPLC overtime, the initial product observed was the f3 -anomer of GlcNAc, demonstrating that J-NAGase is a 13-retainingenzyme (see Fig. 4.3a). This product subsequently anomerizes to form an equilibrium (aI[3) mixture (Fig. 4.3b).The enzyme-catalyzed hydrolysis of 13GIcNAcPNP by K-NAGase or H-NAGase also yielded the f3-anomer ofG1cNAc (not shown), thus all three of the NAGases examined are 13—retaining enzymes.1130.140.120.10.080.060.040.0200.140.120.10.080.060.14G1cNAc Standard (c)0.120.10.08 70.060.040.020 II I I I0 2 4 6time [mm]Figure 4.3. Determination of the stereochemistry of J-NAGase-catalyzed hydrolysis of 3GlcNAcPNP.Shown are the HPLC chromatograins obtained using a DextroPak® column (Waters®) and the indicatedsamples. The eluant was 100%H20. The initial (off-scale) peak in panels (a) and (b) represents the elution ofthe enzyme reaction buffer. The GIcNAc standard (panel c) was an equilibrium ci43 mixture: the -anomereluted as a shoulder peak (with a retention time of 3.38 mm) immediately before the x-anomer peak (with aretention time of 3.55 mm).G1cNAcPNP + NAGase (a)(7mm)G1cNAcPNP + NAGase (b)(3days)1144.3.3. NAGlucal as an inhibitor of NAGase.a. Testsfor irreversible inhibition (inactivation).NAGlucal was examined as a potential inactivator of NAGase by incubating each of the enzymes understudy with 10 mM NAGlucal and the appropriate reaction buffer for 16 hours at 25 °C (see Materials andMethods). At various time intervals small aliquots were removed and residual enzyme activity was assayedusing the substrate I3GIcNAcPNP. No time-dependent inactivation of any of the NAGases under study wasdetected during these tests when compared with parallel control reactions.b. Testsfor reversible inhibition.NAGlucal was then examined as a reversible inhibitor of J-NAGase-catalyzed hydrolysis ofGlcNAcPNP. The initial rates obtained at each concentration of substrate and inhibitor were fitted to equationsdescribing different types of inhibition, using the computer program GraFit (Leatherbarrow, 1990). The bestfit of the data was to the equation describing competitive inhibition, which yielded a value of K1 = 29±2 jiM. Adouble-reciprocal plot of this data illustrates the competitive nature of the inhibition (Fig. 4.4a), and a replot ofthe Km,app values obtained at each inhibitor concentration (Fig. 4.4b) yielded an estimate of K that was clearlyin the same range as that obtained using numerical methods.NAGlucal was then examined as a reversible inhibitor of each of the other two NAGases under study.In these cases, approximate values of K1 (RFK) were derived from Dixon plots of data obtained using differentconcentrations of NAGlucal and a fixed concentration of the substrate. The RF K values obtained forK-NAGase and H-NAGase were —25 and —8.5 jiM, respectively. Thus in each case, the binding of NAGlucal tothe NAGases under study was of high affmity, about 170-fold better than that of G1cNAc itself (K = 5 mM forJ-NAGase; Li & Li, 1970).115• —C12010080604020021.510.50-2 0 2 4[I3G1cNAcPNPI (mM)(a)(b)-0.03 0 0.03 0.06 0.09[NAGlucal] (mM)Figure 4.4. Determination of kinetic parameters for the inhibition of J-NAGase by NAGlucal.(a) Determination of Km, app for the inhibition by NAGlucal of J-NAGase-catalyzed hydrolysis off3G1cNAcPNP. Reactions were performed as described in Fig. 4.2 except as noted. The concentrations ofNAGlucalinthereactionswereO {o}, 8.2 {+}, 16.4(x), 32.8 (A), 57.5 {L},and82.1 {1} ILM.(b) A graphical method—involving replotting data obtained from panel (a)—which allows the reader to estimatethe value of the K1 by visual inspection.116OHHO HNAc HO Hcl.0 0HO’J,OH >< H HO OH[3ManNAc 2-acetamidoglucal I3GIcNACprotonation from below the ring protonation from above the ringScheme 4.2. The stereochemistry of the hydration of NAGlucal by NAGase.4.3.4. NAGlucal as a substrate of NAGase.a. Preliminary TLC evidence.NAGlucal was not only found to be a good inhibitor of NAGase; preliminary TLC analysis usingAgNO3/NaOH for detection (Hough & Jones, 1962) indicated that NAGlucal was also hydrated by the enzyme.Control incubations of NAGlucal in buffer solution alone (i.e., without NAGase) showed no decomposition,even after extended incubation (3 days). Analysis of aliquots taken from the enzyme-catalyzed reaction mixturerevealed that the NAGlucal spot decreased as the reaction progressed, while a product spot increased. Thisproduct spot co-migrated with GIcNAc and ManNAc standards.b. HPLC analysis of the stereochemistry of the hydration reaction.The hydration of NAGlucal should yield one of two possible products, depending on thestereochemistry of protonation of the double bond. If protonation of the double bond is from below the sugarring, N-acetylmannosainine (ManNAc) should be formed, whereas if protonation of the double bond is fromabove the sugar ring, N-acetylglucosamine (G1cNAc) should be formed (see Scheme 4.2). Identification of theproduct was therefore important. The isomers G1cNAc and ManNAc could not be resolved by HPLC using a2C0)Cuw0CCu-o0Cl)0Cu>D0 2 4 6 8 10 12 14time [miniFigure 4.5. HPLC determination of the stereochemistry of J-NAGase-catalyzed hydration of NAGlucal.Shown are the HPLC chromatograms obtained using an Aminex® HPX-87H column (BioRad®). The eluantwas 13 mMH2S04.Peaks with retention times < 10 mm represent the elution of components other than theproduct of the enzyme reaction (e.g., buffer, BSA, or NAGase). The ManNAc and GIcNAc standards elutedwith retention times of 11.0 and 11.5 mm, respectively, when 13 mMH2S04was used as the eluant.Reaction mixture(NAGlucal, NAGase + buffer)117as in (a) + ManNAc standard21.81.61.41.210.80.60.40.21.41.2I0.80.60.40.2I0.80.60.40.2(a)(b)(c)ManNAc + G1cNAcstandards only.(in protein-free buffer)118DextroPak® column, as both compounds eluted as a doublet (i.e., an CL/[3 equilibrium mixture) with the sameretention time. However, an Aminex® HPX-87H column (BioRad®) was able to separate G1cNAc andManNAc, although their elution peaks were very close together (Fig. 4.5c). The ManNAc standard eluted fromthis column before the GlcNAc standard (with retention times of 11.0 and 11.5 minutes, respectively) when13 mMH2S04was used as the eluant.When NAGlucal was incubated with J-NAGase (and the appropriate buffer) for several days, andaliquots of this reaction were analyzed by HPLC, the sole product observed was G1cNAc, which was identifiedby its elution at the same retention time (11.5 mm) as that of an authentic sample (compare Fig. 4.5a and 4.5c).Furthermore, if the reaction product sample was spiked with an authentic ManNAc sample, HPLC analysisshowed that the previously observed product peak acquired a prominent, leading shoulder (Fig. 4.5b), and thusManNAc was not the product of the hydration reaction. GlcNAc was also the sole product of the hydration ofNAGlucal by K-NAGase or H-NAGase (not shown). These results indicate that NAGase—a [3-retamingglycosidase—protonates the double bond of NAGlucal from above the ring. In contrast (as described in Chapter1), all other [3-retaining glycosidases studied to date protonate the double bond of their glycal substrates frombelow the ring (Hehre et al., 1977; Legler, 1990).c. Determination ofKm and Vm for the hydration reaction.The initial-rate data necessary to calculate Km and Vmax for the hydration of NAGlucal by J-NAGasecould be obtained in two possible ways. One way was measurement of the rate of consumption of the substrate(NAGlucal), and the other would involve measurement of the rate of formation of the product (GlcNAc).However, the product of the hydration reaction does not absorb light very strongly; hence attempts were firstmade to measure the rate of consumption of the substrate by measuring the decrease in absorbance of 220-nmUV light. This wavelength was chosen after the absorption spectra were taken and the extinction coefficientscalculated for NAGlucal and GlcNAc (max = 220 nm; 8220 = 5.36 and 0.17 mMcm,respectively).However, at a wavelength of 220 nrn there was too much interfering absorption from other components in thereaction (the buffer, BSA, and NAGase), and this approach was abandoned.119Several colorimetric methods are available for measuring the concentration of G1cNAc (e.g., Reissiget a]., 1955). However, the results of the inhibition studies showed that NAGlucal binds very tightly to NAGase(K = 29 ± 2 pM for J-NAGase). Thus in order to determine the kinetic parameters Km and Vmax for thereaction, initial rates (i.e., with < 10% consumption of the substrate) of enzyme-catalyzed hydration had to bemeasured with substrate concentrations in the range of 10 to 100 aiM. This meant that the concentration ofGlcNAc formed would be in the 1 to 10 pM range, concentrations well below the detection limits of mostcolorimetric methods. Another difficulty with existing colorimetric methods is that NAGlucal was found toreact with the reagents used to detect GlcNAc.As a result of these difficulties neither a cuprimetric method (Hehre et a]., 1980), nor the colorimetricmethods developed by Park & Johnson (1949) and Reissig et a]. (1955) were able to accurately measure GlcNAcconcentration with the required sensitivity. The same sensitivity problem was encountered in attempts to silylatethe components of the stopped kinetic reactions, and then analyze them by gas chromatography using theprocedure of Sweeley et al. (1963).HPLC (with an Aminex® HPX-87H column) was eventually used to measure the concentration ofG1cNAc. The instrumentation used was unable to accurately quantitate G1cNAc in the 1 to 10 pM range. Toovercome this difficulty the assay volume was scaled-up 10-fold, from 0.100 to 1.00 mL. These 1.00-mi.reactions contained 20 to 200 jiM NAGlucal, citrate buffer (pH 5.0), and J-NAGase, and were incubated at25 °C. Reactions were stopped by boiling for exactly 30 seconds, a procedure which was found to irreversiblydenature the enzyme, but which did not result in significant decomposition of other components of the reaction.Water was removed by lyophiization, the residue remaining in each reaction tube was resuspended with 0.100mL of double-deionizedH20, and then 0.080 mL of each reaction was analyzed by HPLC. The lyophilizationstep effectively concentrated the GlcNAc 10-fold, and this permitted the quantitation of the reaction product byHPLC. The area of the G1cNAc peak was determined, and then corrected by subtracting the area of the G1cNAcpeak in appropriate controls (reactions with the same concentrations of NAGlucal but without enzyme).120654, 3—=0-0.04 0.06[NAGlucal] —1Figure 4.6. Determination of Km and Vmax for J-NAGase-catalyzed hydration of NAGlucal.Reactions were performed at pH 5.0 and 25 °C in 5 mM citrate buffer containing 0.01% BSA and 2.0 p.g/mL of3-N-acetythexosaminidase from jack beans. The concentration of NAGlucal in the reactions was 21, 50, 80,150, and200).IM.Fig. 4.6 shows the Lineweaver-Burk plot for the enzyme-catalyzed hydration of NAGlucal byJ-NAGase. A Km of 27 ± 3 pM and a Vmax of 0.48 ± 0.01 U/mg were obtained by fitting the data to thenonlinear form of the Michaelis-Menten equation using the computer program GraFit (Leatherbarrow, 1990)(see Table 4.1). The Km for the J-NAGase-catalyzed hydration of NAGlucal (27 pM) was in good agreementwith the K1 of 29 pM obtained for the inhibition of the enzyme by this compound.The Km for the J-NAGase-catalyzed hydrolysis of 3GlcNAcPNP (620 pM: see Table 4.1) indicates thatthe affmity of NAGlucal for J-NAGase is about 20-fold greater than the affinity of 3GlcNAcPNP for theenzyme. However, J-NAGase hydrated NAGlucal about tOO-fold more slowly than the enzyme hydrolyzed3GlcNAcPNP (Table 4.1). Hence the overall substrate efficiency, Vm!Km, for NAGlucal is about 4.5-fold-0.02 0 0.02 0.04121lower than that of I3G1cNAcPNP. The kinetic parameters for the substrate activities of NAGlucal andI3G1cNAcPNP with J-NAGase are summarized in Table 4.1. For comparison, the corresponding values forD-glucal and GlcPNP with Agrobacterium f-glucosidase (pABG5) are also listed.In contrast, Agrobacterium 1 -glucosidase binds D-glucal 10-fold less strongly and hydrates thiscompound about 4,400-fold more slowly than it binds to and hydrolyzes I3G1cPNP. This means that D-glucal isabout 48,000-fold less efficient as a substrate for pABG5 than I3G1cPNP (based on their VmaxlKm values).Clearly D-glucal, which lacks the C-2 hydroxyl group found on glycosides such as 3GlcPNP, is a much poorersubstrate for pABG5. Thus, even though NAGlucal was not as efficient a substrate for NAGase asf3G1cNAcPNP, the difference between the efficiencies of these two substrates of NAGase was much less than thedifference in the efficiencies of D-glucal and I3G1cPNP as substrates of 3-glucosidase. These results stress theimportance of the 2-substituent for interactions in the active sites of glycosylases that are required for efficientcatalysis.Table 4.1. Kinetic parameters for glycosylase-catalyzed reactions of glucals and related glucosides.Enzyme SubstrateVmax Km Vmax/Km Reference(U/mg) (pM) (U/[mg.jsM])J-NAGase NAGlucal 0.48 ± 0.01 27 ± 3 0.018 ± 0.002 This workfGlcNAcPNP 50 ± 1 620±40 0.081 ± 0.005 ibid.Agrobacterium D-glucal 0.046 850 5.4 x i0- Street (1988)f3-glucosidase I3G1cPNP 200 78 2.6 Kempton &(pABG5) Withers (1992)4.3.5. Kinetic studies with other glucal derivatives and glycosides.a. Studies with D-glucal.A substrate test showed that J-NAGase did not hydrate D-glucal. A reversible inhibition test showedthat D-glucal binds to J-NAGase very poorly, with an RF K1 of 39 mM (see Fig. 4.7). The affinity of J-NAGasefor D-glucal is therefore 1,300-fold less than the affinity of the enzyme for NAGlucal (K = 29 .tM).12211111111 I I I 1111111111 I 11111I___max0 Iiiiliii IIIIIIIIII[jIII)III-40 -20 0 20 40 60 80 100[D-glucal] (mM)Figure 4.7. Estimation of the K1 for the inhibition of J-NAGase by D-glucal.Reactions were performed at pH 5.0 and 25 °C in 50 mM citrate buffer containing 0.1% BSA and 1.0 j.Lg/mL off3-N-acetythexosaminidase from jack beans. The concentration of GIcNAcPNP in the reactions was 0.58 mM.The concentrations of D-glucal in the reactions were 0, 22, 36, 42, 63, and 90 mM.The 1,300-fold difference in the inhibition constants corresponds to a difference in binding energyt of17.8 kJ/mol (or 4.2 kcal/mol) at 25 °C. This result, together with the results of the inhibition and substrate testsof NAGlucal, confirms the importance of the noncovalent interactions between the 2-acetamido group and theenzyme’s active site. Indeed these interactions are likely much greater at the transition state, and would accountfor the relatively high rate of hydration of NAGlucal, and the lack of reaction with D-glucal.t The free-energy difference, AAG°, is calculated from the expression SAG° = RT( ln(K,2I1i)) whereK,2 andK1, are the inhibition constants for the two inhibitors being compared, R is the gas constant, and T is theabsolute temperature.123b. Studies with 2-cyanoglucal.The results of the inhibition studies of J-NAGase using NAGlucal and D-glucal showed that removingthe 2-acetamido group drastically reduced the affinity of the enzyme for the compound. It was therefore ofinterest to study the effect of replacing the 2-acetamido group with another C-2 substituent—in this case, thecyano group—to see if this resulted in improved binding in the enzyme’s active site compared with D-glucal.2-Cyanoglucal is also an ct,13-unsaturated glucal, and thus it might act as a Michael acceptor for an active-sitenucleophile and covalently inactivate NAGase.2-Cyanoglucal was examined as a potential inactivator of NAGase by incubating each of the enzymesunder study with 33 mM 2-cyanoglucal and the appropriate reaction buffer for 17 hours at 25 °C (see Materialsand Methods). At various time intervals small aliquots were removed and residual enzyme activity was assayedusing the substrate 3GlcNAcPNP. No time-dependent inactivation of any of the NAGases under study wasdetected during these tests when compared with parallel control reactions.A reversible inhibition test showed that 2-cyanoglucal binds to J-NAGase very poorly, with a K1 of-57 mM, i.e., even worse binding than D-glucal (K1 — 39 mM). Despite the fact that NAGlucal and2-cyanoglucal both have a substituent at C-2, there is clearly a profound difference in the ability of the enzymeto accept these substituents. Either there is some steric repulsion (due to the linear geometry of the nitrile), ormore likely, the mtrile is incapable of the hydrogen bonding that the N-acetyl group of NAGlucal exploits whenit binds in the active site.The configuration of the 2-acetamido substituent is also very important. Li & Li (1970) reported thatManNAc at concentrations up to 10 mM does not inhibit J-NAGase. Furthermore, althoughN-acetyldeoxynojirimycin (4.1) acts as a potent competitive inhibitor of J-NAGase (with a K1 = 0.23 tiM), themannosamine analogue (4.2) of N-acetyldeoxynojirimycin does not act as a NAGase inhibitor (Fleet et al.,1986). These results show that NAGase binds very poorly to sugars if the 2-acetamido group is in the axialposition, presumably because of repulsive steric interactions and/or the absence of hydrogen bonding.OH OHHNAc[4.1] HO._j’1 [4.2]HNAc124c. Studies with 2F/3G1cDNP and /3G1cDNP.2f93G1cDNP inactivates f3-retaining glucosidases very rapidly, and it does so by forming a 2-deoxy-2-fluoroglucosyi—enzyme intermediate whose rate of deglycosylation is extremely low (Withers et al., 1988). Thestereochemical analysis of the hydrolysis of f3GIcNAcPNP showed that J-NAGase is a f3-retaining enzyme witha catalytic mechanism that is similar to that of a f3-retaining glucosidase. Hence it was possible that 2Ff3G1cDNPmight also act as an inactivator of NAGase, even though NAGase has very little affmity for sugars without a2-acetamido group. The fluorine atom at C-2 in 2FI3G1cDNP could possibly act as a hydrogen bond acceptor,thereby increasing the affinity of the enzyme for this compound, and the excellent leaving group could drive theglycosylation step.2F(3G1cDNP was examined as a potential inactivator of J-NAGase or K-NAGase by incubating eitherof these enzymes with 3 mM 2FI3G1cDNP and the appropriate reaction buffer for 48 hours at 25 °C (seeMaterials and Methods). At various time intervals small aliquots were removed and residual enzyme activitywas assayed using the substrate f3G1cNAcPNP. No time-dependent inactivation of any of the NAGases understudy was detected during these tests when compared with parallel control reactions.A substrate test was also performed with f3G1cDNP and J-NAGase. Rates of I3G1cDNP hydrolysis weredetermined using an assay that was similar to the one used for f3GIcPNP In both of these assays the reactionwas monitored at 400 nm (whereas for I3G1cNAcPNP the reaction was monitored at 360 urn), and the productformed during the reaction was quantitated spectrophotometricafly (see Materials and Methods).The results of the substrate test with 3GlcDNP and J-NAGase showed that there was a very smallamount of an unknown contaminant in the f3G1cDNP preparation that appeared to act as a substrate ofJ-NAGase, but that f3G1cDNP itself did not. Thus when f3G1cDNP (6.5 and 1.5 mM were tested) was incubatedat 25 °C in a 0.200-mL reaction containing 50 mM citrate (pH 5.0) and 6 ig of J-NAGase, there was a burst ofproduct (DNP) formation, but the reaction rate decreased over time and eventually equaled the spontaneous(nonenzymatic) rate after 90 minutes. The amount of this contaminant “substrate” was estimated (from theinitial burst of product formation) to be about 0.4% of the f3G1cDNP used in the assay. After this burst ofproduct formation had ended (i.e., about 90 mm after the start of the assay), the addition of more J-NAGase125(6 jig) to the assay did not result in any additional product formation, despite the fact that over 99% of the initialamount of I3GIcDNP in the assay was still present. Only spontaneous (nonenzymatic) hydrolysis was observedwhen more enzyme was added after the contaminant “substrate” had been consumed during the initial burst ofDNP formation.f3G1cDNP itself, then, did not act as a substrate for J-NAGase. The absence of hydrolytic activity byJ-NAGase towards f3G1cDNP suggests that the failure of 2Ff3G1cDNP to inactivate NAGase was probablyattributable to an extremely low rate of cleavage of the glycosidic bond.4.3.6. Substrate-enzyme interactions and the mechanism of NAGase-catalyzed reactions.Each of the NAGases under study catalyzed the hydrolysis of f3G1cNAcPNP with net retention ofanomeric configuration. This suggests that all three enzymes operate through a double-displacement mechanismin which an N-acetylglucosaminyl-enzyme intermediate is formed and then hydrolysed. This mechanism wasoriginally proposed by Koshland (1953), and has subsequently received considerable experimental support(reviewed in Sinnott, 1990).The binding of NAGlucal to each NAGase was shown to be of high affinity, about 170-fold better thanthat of G1cNAc itself (K1 = 5 mM for jack bean NAGase; Li & Li, 1970), which is particularly noteworthy giventhe absence of an anomeric hydroxyl group on NAGlucal. There are two possible explanations for this tightbinding. It could genuinely be a case of tight binding, i.e., due to a resemblance of NAGlucal, with its planargeometry around C-i, to the structure of an oxocarbonium ion-like transition state. Alternatively, the tightbinding could be a consequence of the accumulation of an N-acetylglucosaminyl-enzyme intermediate duringthe hydration reaction, comparable to that which occurs during the inhibition of E. coli 13-galactosidase byD-galactal (Wentworth & Wolfenden, 1974), or the inhibition of Aspergillus wentii 3-glucosidase by D-glucal(Legler et al., 1979).126(a)HO,°Yglycosylation___HO°\,ODYdeglycosylationD0 OD777/JHO,OHAcNHScheme 4.3. Comparison of the proposed mechanisms of -g1ucosidase and 13-N-acetythexosaminidase.(a) (3-Glucosidase-catalyzed hydration of D-glucal in D20.(b) 3-N-Acetythexosaminidase-cata1yzed hydration ofNAGlucal.0 CD(b)HH\\\\0 OHYglycosylation/77/YdeglycosylationJ127The latter hypothesis is unlikely given that the deglycosylation step for the reaction of NAGlucal isidentical to that for the hydrolysis of 3GlcNAcPNP (as both steps involve the hydrolysis of anN-acetylglucosaminyl-enzyme intermediate). The Vmax value for 3GlcNAcPNP hydrolysis catalyzed by jackbean NAGase (50 p.mollminlmg) therefore represents a minimum estimate of the rate of the deglycosylation step.Since the Vmax value for NAGlucal hydration catalyzed by jack bean NAGase was only 0.48 jmiollminlmg, thesecond step cannot be rate-limiting for NAGlucal, and the value observed likely reflects the rate of formation ofthe intermediate.The tight binding observed is therefore a genuine case of a high-affinity interaction, and is most likely aconsequence of some critical structural features of the inhibitor resembling those of the transition state, and theconsequent exploitation of transition-state binding interactions. Such tight, reversible binding of a glycal to aglycosidase has never been observed previously, presumably because in all cases examined the key 2-hydroxylsubstituent was necessarily missing. In fact, glycals typically bind 10 to 100-fold worse to glycosidases than dothe corresponding glycosides (Legler, 1990). Comparable behaviour was observed with D-glucal (which lacksthe 2-acetamido substituent of NAGlucal) and jack bean NAGase. The binding of D-glucal to jack beanNAGase was about 8-fold worse than that of G1cNAc (Li & Li, 1970), and 1300-fold worse than that ofNAGlucal.The 1300-fold greater binding affinity of NAGlucal compared with D-glucal establishes thecontribution of the 2-acetamido functionality to the binding affinity of NAGlucal as 4.2 kcallmol, and in turnestablishes, to at least some extent, the contribution of this substituent to transition-state stabilization in thenormal reaction. NAGlucal is, at best, an imperfect transition-state analogue, given the absence of an anomencoxygen atom and the imposed planarity around (0-5, C-i, C-2, and C-3) rather than (C-5, 0-5, C-i, and C-2).The value of 4.2 kcallmol therefore represents a minimum estimate of the contribution of the 2-acetamido groupto transition-state stabilization, as indeed is reflected in the complete absence of reaction of the enzyme withD-glucal or f3G1cDNP, despite the superiority of the leaving group in the latter substrate.The stereochemistry of NAGase-catalyzed proton donation to C-2 of NAGlucal is without precedent inprevious studies of ‘retaining’ f-glycosidases. Previously these enzymes have only been shown to effect proton128donation to glycals from the bottom face, most likely through the concerted process shown in Scheme 4.3a(Legler, 1990). In the absence of other constraints, this more commonly observed stereochemistry ispresumably a consequence of the known preference for the reaction of acetals via oxocarbonium ion-liketransition states to occur in a pre-associative process (Banait & Jencks, 1991). Such pre-association is mostreadily achieved in the concerted process shown in Scheme 4.3a. Furthermore, with unsubstituted glycals theintermediate formed involves a 2-deoxy sugar, thus the correct stereochemical orientation of a bulky substituentat C-2 is not a consideration. However, if proton donation came from the bottom face during the catalytichydration of NAGlucal, it would result in the formation of an N-acetylmannosantinyl-enzyme intermediate inwhich the bulky C-2 substituent assumes the enzymatically unfavourable axial position. By contrast,protonation of NAGlucal at C-2 from the top face generates an N-acetylglucosaminyl-enzyme intermediate inwhich normal transition-state binding interactions with the equatorial 2-acetamido group can be exploited toassist the formation and subsequent hydrolysis of this intermediate (Scheme 4.3b). NAGase-catalyzedprotonation of NAGlucal from the top face, as observed, is therefore quite reasonable and reflects both theconstraints and benefits of having the correct substituent at C-2.4.4. CONCLUSIONS.The direct colorimetric assay measuring the absorbance of 360-nm light proved to be a reliable,convenient method for assaying the hydrolysis of 3GlcNAcPNP by NAGase. HPLC was used to determine thestereochemical outcome of this reaction for all three NAGases studied, all of which were found to be 3-rerainingenzymes that yielded 3GlcNAc as the initial product of the hydrolysis reaction.NAGlucal was found to be a reasonably good substrate for J-NAGase, with a Km = 27 ± 3 j.IM and aVmax = 0.48 ± 0.01 U/mg. Indeed, all three NAGases studied catalyzed the hydration of NAGlucal to formG1cNAc—not ManNAc—which indicated that the double bond was protonated from above the plane of thesugar ring. This is the first time that a f3-retaining glycosidase has been shown to protonate an endocyclicdouble bond from the “correct” face. These results demonstrate firstly that the “incorrect” protonation observed129in other cases is the consequence of an alternative pathway adopted in the absence of the C-2 substituent, andsecondly that protonation of glycals from the “correct” face is indeed possible.Kinetic parameters for the NAGase-catalyzed hydration of NAGlucal, D-glucal, and various otherglucal derivatives and glycosides were determined (see Table 4.2), and a comparison of the values of theseparameters showed that the 2-acetamido group is extremely important for binding of NAGlucal by the enzyme.The relatively tight, reversible binding of NAGlucal suggests that glycals may indeed be regarded astransition-state analogues for glycosidases, but that the relatively low affinities generally observed are aconsequence of their lack of the critical 2-substituent.4.5. SUGGESTIONS FOR FUTURE WORK.Bernacki et al. (1985) carried out studies using mice that showed that NAGlucal has promise as anantitumor and antimetastatic agent, and that these properties of the compound are attributable to its ability to actas an inhibitor of NAGase. A particularly noteworthy property of NAGlucal is its lack of cytotoxicity towardsnormal cells, a property that is very desirable for a chemotherapeutic agent (Bernacki et al., 1985).The research described herein showed that NAGlucaL has a low K1 for human placental NAGase, andhence that it might be worthwhile to investigate the effect of this compound on the growth and spread of varioustypes of human tumors in vitro. In addition, it might well be worth investigating analogues of NAGlucal that aresubstituted with a sugar at C-i; such compounds could have considerably higher affinity and specificity forNAGase.At a more fundamental level, no mechanism-based inactivators of NAGase have been discovered. Thesynthesis and kinetic analysis of potential mechanism-based inactivators of 3-N-acetythexosaminidase would bevery desirable, not only for possible practical applications, but also to provide more insights into the catalyticmechanism of this clinically important enzyme.Table 4.2. Summary of kinetic data obtained using various f3-N-acetythexosaminidases.130Temperature (°C) 25 25 25pH 5.0 4.25 4.253GIcNAcPNP A Km = 620 ± 40 pMVmax =50 ± 1 U/mgKm 1100±lOOj.iMVm =24±2 U/mg580± 60 pMVmax = 8.0 ± 0.4 U/mgStereochemistry f3-retaining 3-retainmg n-retainingNAGlucal B= NHAcR4 = HProduct fromNAGlucalK1 =29±2 pMKm =27±3 .LMV=0.48±0.01U/mgG1cNAcK-25iM K-8.5jiMG1cNAc G1cNAcD-glucal BR3 = H= H2-cyanoglucal BR3 = CNR4 = Hnot an inactivator not an inactivator2Ff3GIcDNP CR1 =Fnot an inactivator not an inactivator3GIcDNP CR1 =OHnot a substrateNHOR4Parameter or Structure J-NAGase K-NAGase H-NAGasecompound (jack bean) (bovine kidney) (human placenta)K1 -. 39 mMK-57mMNO2A B C131CHAPTERS: MATERIALS AND METHODS5.1. ORGANIC SYNTHESIS.5.1.1. Materials and routine experimental procedures.a. Analytical methods.Melting points (mp) were determined on a Laboratory Devices Mel-Temp II melting-point apparatus.Melting points were uncorrected.1H Nuclear magnetic resonance (NMR) spectra were recorded on the following instruments at theindicated field strengths: a Bruker AC-200 at 200 MHz, a Varian XL-300 at 300 MHz, or a Bruker WH-400 at400 MHz. When possible, tetramethylsilane (TMS) was used as an external reference (6 = 0.00 ppm). Spectraobtained in D20 were referenced externally to 2,2-dimethyl-2-silapentane-5-sulfonate (6 = 0.00 ppm). Allchemical shifts were reported using the 6 scale.19F NMR spectra were recorded with proton coupling on a BrUker AC-200 instrument at a fieldstrength of 188 MHz. Chemical shifts were reported using the 6 scale referenced to CFC13 (6 = 0.00 ppm),although the external reference used wasCF3OOH (6= -76.53 ppm). Signals upfield of CFCI3 were assignednegative values.13C NMR spectra were recorded with proton decoupling on the following instruments at the indicatedfield strengths: a BrUker AC-200 at 50 MHz, a Varian XL-300 at 75 MHz, or a Bruker WH-400 at 125.76 MHz.The solvent peak (either CDC13 orD20) was used as a reference.Low resolution electron-ionization mass spectra were recorded on a Kratos MS 50 mass spectrometeroperating at 70 eV. Desorption chemical ionization (DCI) mass spectra were recorded on a Delsi NermagR10-1OC mass spectrometer using NH3 as the chemical ionization gas. Fast atom bombardment (FAB) massspectra were recorded on a AEI-MS 9 mass spectrometer with xenon as the FAB gun, operating at 7-8 kV and1 mA current.132Microanalyses were performed by Mr. Peter Borda in the Microanalytical Laboratory, Department ofChemistry, at The University of British Columbia, Vancouver, B. C.b. Thin-layer chromatography and silica gel column chromatography.Thin-layer chromatography (TLC) was performed using analytical plates (silica gel 60F254, Merck).Compounds were visualized under UV light or after charring with 10%H2S04inmethanol or iodine vapour.Column chromatography was performed using Kieselgel 60 (230-400 mesh) silica gel.c. Solvents and reagents.Solvents and reagents were either reagent, certified, or spectral grade. Dry solvents and reagents wereprepared as described below. Dichioromethane was first washed with concentrated sulfuric acid followed bywater and sodium bicarbonate, predriecl over calcium chloride, and finally distilled over calcium hydride.Dimethyl formamide was stirred overnight over magnesium sulfate, then distilled under reduced pressure onto4-A molecular sieves. Diethyl ether and tetrahydrofuran were distilled over sodium metal and benzophenone.Methanol was distilled from magnesium methoxide (formed in situ by reaction of methanol with magnesiumturnings in the presence of a catalytic amount of iodine). Pyridine was predried for several days by standingover pellets of NaOH, and then distilled over calcium hydride. Toluene was distilled over calcium hydride.d. Compounds synthesized and generously provided by colleagues.Some of the compounds used in this work were synthesized and generously provided by colleagues.With the exception of Prof. A. Vasella, all of the people mentioned were coworkers in Professor Withers’laboratory. Prof. A. Vasella of the University of Zurich provided 1-nitroglucaL Dr. Mark Namchuk provided2FGlcDNP {2’,4’-dinitrophenyl 2-deoxy-2-fluoro-3-D-glucopyranoside} and J3G1cDNP {2’,4’-dmitrophenyl 3-D-glucopyranoside). Dr. William Stirtan provided 1-(methyl carboxylate)-D-glucal (methyl 2,6-anhydro-3-deoxy-D-arabino-hept-2-enolate) as well as sodium 1-(carboxylate)-D-glucal (sodium 2,6-anhydro-3-deoxy-D-arabino-hept-2-enolate}. John McCarter provided the peracetylated precursor of 2-acetamidoglucal{ 1,5-anhydro-2-deoxy-2-(N-acetylacetamido)-3,4,6-tri- O-acetyl-D-arabino-hex-1-enitol). Ken Mok performed133 the photobromination of 2,3,4,6-tetra-O-acetyl-P-D-glucopyranosyl cyanide to give 2,3,4,6-tetra-0-acetyl-l-a-bromo-P-D-g/Mco-pyranosyl cyanide. 5.1.2. Routine synthetic procedures. a. Deacetylation with sodium methoxide in methanol. The following procedure was adapted from the work of Zemplen & Pacsu (1929). The acetylated sugar derivative (~1 g) was dissolved in -50 mL of dry methanol. A catalytic amount of solid sodium methoxide (-1-2 mg per 50 mL) was added, and then the reaction mixture was stirred at room temperature until the reaction was complete. The sodium methoxide was removed by passing the reaction solution through a small column of silica gel. The column eluate was then evaporated to dryness in vacuo. The resulting oil was crystallized and then recrystallized from the solvents specified for the individual compounds. b. Deacetylation with ammonia-saturated methanol. The following procedure was adapted from the work of Fritz et al. (1983). The acetylated glycal (~1 g) was dissolved in -30 mL of dry methanol. About 10 mL of cold (0 °C), dry MeOH was saturated with NH3 by bubbling, and then the solution was poured into the reaction flask. The mixture was left at room temperature until the reaction was complete, and then the solvent was evaporated in vacuo. The resulting residue was crystallized and then recrystallized from the solvents specified for the individual compounds. c. Trimethylsilylation. The following procedure was adapted from the work of Horton & Priebe (1981). The starting material (-1 g) was dissolved in -10 mL of dry pyridine and the resulting solution was cooled in an ice bath. Syringes were used to add one equiv. of hexamethyldisilazane (Me3 SiNHSiMe3) and one-half equiv. of trimethylsilyl chloride (Me3SiCl) for every hydroxyl group of the starting material. A white precipitate formed immediately. The ice bath was removed, and then the reaction was stirred at room temperature for -25 min until the reaction was complete. The contents of the reaction flask were vacuum filtered through Celite in a fine porosity sintered glass funnel. The solvent in the filtrate was evaporated in vacuo, and then the resulting oil was purified further 134 by vacuum distillation. The trimethylsilyl group was not stable enough for the product to be purified by silica gel column chromatography. d. Triethylsilylation. The following procedure was adapted from the work of RajanBabu & Reddy (1986). The starting material (-0.5 g) was dissolved in -5 mL of a 1:1 v:v mixture of dry pyridine and dry methylene chloride, and then a small amount of triethylamine (-0.5 mL) was added. Triethylsilyl chloride (Et3SiCl, 1.1 equiv. for every hydroxyl group of the starting material) was added, followed by stirring at room temperature until the reaction was complete, and then pentane, saturated KH2PO4, and H2O were added. The product was isolated by repeated extraction into pentane, followed by washing of the combined pentane layer with saturated sodium bicarbonate and then H2O. The pentane layer was dried over MgS04, and the solvent was evaporated in vacuo. The triethylsilyl group was stable enough for the product to be purified by silica gel column chromatography. e. Tert-butyldimethylsilylation. The following procedure was adapted from the work of Corey & Venkateswarlu (1972). The starting material (-0.4 g) was dissolved in -10 mL of dry DMF. For every hydroxyl group of the starting material, 1.5-2.0 equiv. of (f-butyl)dimethylsilyl chloride (TBDMS chloride) and 2.5 equiv. of imidazole were added. The reaction was stirred at room temperature for 2 hours, a condenser was attached to the flask, and then the mixture was stirred at 45 °C for 10 hours until the reaction was complete. The product was extracted four times into ether, and then the combined ether layer was washed with saturated sodium bicarbonate and then H2O. The ether layer was then dried over MgS04, and the solvent was evaporated in vacuo. The product was purified by silica gel column chromatography. 135 <UOR [3.3] OR -R [2.3] [5.1] SiMe3 SiEt3 RO-RO-[1.2] [2.1] [2.2] [2.4] [2.5] [2.6] [5.2] RO -R H H H SiMe3 SiMe2(NBu) SiMe20-Bu) SiEt3 Ra -Ra H F F F F F H RH -Rb H F H F F H H Figure 5.1. Structures of Cp2TiMe2, heptenitol derivatives, and associated synthetic intermediates. The numbering systems used for D-glucono-l,5-lactone and D-g/uco-heptenitol derivatives are indicated. 5.1.3. Syntheses. a. The synthesis of heptenitol and its derivatives. Dimethyltitanocene (Cp2TiMe2, 3.3) (see Claus & Bestian, 1962). Titanocene dichloride (3.2, Cp2TiCl2, obtained from Aldrich Chemical Co.) [1.4 g, 5.6 mmol] was dissolved in -30 mL of dry ether. A dropping funnel was used to add 8.4 mL of a 1.4 M solution of methyl lithium [2.1 equiv., 12 mmol] dropwise at -20 °C under N2. The reaction mixture was stirred in the dark at -20 °C under N2 for one hour. Ice was then added to quench the reaction, and the resulting solution was extracted three times with ether; during this procedure, the solution was kept < 15 °C and shielded from light. The combined ether layer was dried over MgS04, and the solvent was evaporated in vacuo at low temperature. The resulting bright orange solid [0.95 g, 81%] was dried over P2O5 in a dessicator in vacuo for 2 hours at room temperature to give 3.3 (mp 110-117 °C, 136dec), which was stored shielded from light at -20 °C as 9.1 mL of a 0.5 M solution in THF. 1H NMR data (300MHz, THF): 6 5.75 (s, 10 H, Cpa); —0.08 (s, 6 H, Me2).2,3,4,6-Tetra-O-(triethylsilyl)-D-glucono-1,5-lactone (5.1) (see Raj anB abu & Recldy, 1986).Glucono-i,5-lactone (1.15, Sigma Chemical Co.) [0.50 g, 2.81 mmol] was persilylated as described above usingthethyl silyl chloride (Et3SiC1) [2.1 mL, 12 mmol, 4.4 equiv.], 5.6 mL of a 1:1 v:v mixture of dry pyridine anddry methylene chloride, and 0.5 mL of triethylainine. The product was stable enough to be purified by silica gelcolumn chromatography (4:1 hex:CHC13)to give a colorless oil of 5.1 [1.63 g, 91%]. 1H NMR data (300MHz, CDC13): 34.56 (ddd, 1 H, J45 = 7.6 Hz, J,6 = 4.2 Hz, i’5,6 = 2.7 Hz, H-5); 4.12 (dd, 1 H, J23 = 3.6 Hz,= 0.9 Hz, H-2); 4.00 (dt, 1 H,J34 =f4,5 = 7.6 Hz, J2,4 = 0.9 Hz, H-4); 3.92-3.86 (m, 2 H, H-6, H-3); 3.80(dd, 1 H, J = 10.8, 4.2 Hz, H-6’); 1.02-0.95 (m, 36 H, [{CJICH2)Si-014;0.70-0.58 (m, 24 H,[{CH3CkI2}Si-O]4).l3NMR data (75 MHz, CDC13): 6 169.9 (C=O); 81.4, 77.2, 73.7, 71.0 (C-2 to C-5);61.7 (C-6); 6.76, 6.72, 6.67, 6.59 (12 C, [{kj3CH2}3Si-O]4); 4.93, 4.58, 4.45 (12 C, [{CHH)Si-O].2,6-Anhydro-1-deoxy-3,4,5,7-tetra-O-(triethylsilyl)-D-gluco-hept-1-enitol (5.2) (see Petasis &Bzowej, 1990). The lactone 5.1 [0.62 g, 0.98 mmol] was dried overnight in vacuo. Dry THF [9 mU was addedand then the flask was fitted with a condenser. Cp2TiMe (3.3, prepared as described above) [2.1 equiv., 2.1mmol] was added in the dark underN2, and the mixture was refluxed at about 70 °C for 2 days. Heating wasstopped, the solution was filtered through a bed of silica (stirred with 1% Et3N in EtOAc), the product waseluted with 5% ether in hexane, and the solvent was evaporated in vacuo. The resulting residue was purifiedtwice by column chromatography using 2% ether in hexane as the eluant. The solvent was evaporated in vacuoto give an oil [0.52 g, 84%} of 5.2. 1ff NMR data (300 MHz, CDC13): 64.38 (s br, 1 H, H-i); 4.11 (ddd, 1 H,= 9.0 Hz, f6,’ = 4.8 Hz, J6,-j = 1.9 Hz, H-6); 4.02 (s br, 1 H, H-i’); 3.98 (d, 1 H, J3,4 = 3.0 Hz, H-3); 3.85(dd, 1 H, f7, = 11.3, J6,7 = 2.0 Hz, H-7); 3.76-3.65 (m, 3 H, H-7’, H-5, H-4); 1.04-0.93 (m, 36 H,[{ki3CH2}3Si-O]4); 0.70-0.60 (m, 24 H, [{CHjj}Si-O]).2,6-Anhydro-1-deoxy-D-gluco-hept-1-enitol (1.2). After the heptenitol 5.2 [0.52 g, 0.82 mmol] hadbeen purified by column chromatography it was dried overnight in vacuo and then dissolved in 6.6 mL [8 equiv.,6.6 mmol] of a 1 M solution of TBAF in THF and then stirred for -.2 hours at room temperature. The solvent137was evaporated in vacuo, and then an excess of the Li form of Dowex 50W X2 resin [-5-10 mL of solidsuspended in MeOH] was added to remove the ainmonium salt (Bu4Nj. The mixture was stirred overnight atroom temperature, and then the resin was filtered and washed with MeOH. The solvent was evaporated invacuo, and then the residue was purified by column chromatography (27:2:1 EtOAc:MeOH:H0) andcrystallized and recrystallized from MeOH, ether, and EtOAc to give white crystals [0.12 g, 85%] of 1.2 (mp131-133 °C; mp 95-97 °C when recrystallized from 5:1 EtOAc:MeOH). 1HNMR data (400 MHz,D20): 64.78 (t, 1 H, Ji,1 = J2,i < 1 Hz, H-i); 4.74 (t, 1 H, Ji,i =J2,i’ < 1 Hz, H-i’); 3.95-3.89 (m, 2 H, H-3, H-7); 3.76(dd, 1 H, f7 = 12.5 Hz, J67 = 5.3 Hz, 11-7’); 3.55 (t, 1 H, J3,4=J4,5= 9.1 Hz, H-4); 3.45 (ddd, 1 H,J56 = 9.9Hz, J67’ = 5.3 Hz, J6,-j = 2.1 Hz, H-6); 3.41 (dd, 1 H, J56 = 9.4 Hz, J4,5 = 9.1 Hz, H-5). 13C NMR data (50MHz,D20): 3 159.45 (C-2); 94.95 (C-i); 81.91, 77.45, 71.29, 69.98, 61.45 (C-3 to C-7). DCI MS data: mlz177 (MHj; 194 (M+NH,j. Anal. Calc. forC7H1205(176.17): C, 47.72; H, 6.87. Found: C, 47.76; H, 6.84.2,3,4,6.Tetra-O-(trimethylsilyl)-D-glucono-1,5-Iactone (2.3) (see Horton & Priebe, 1981). Gluconoi,5-lactone (1.15, Sigma Chemical Co.) [5.0 g, 28 mmol] was dissolved in 47 mL of dry pyridine and thenpersilylated as described above using hexamethyldisilazane [23 mL, 110 mmol, 4 equiv.j and trimethyl silylchloride (Me3SiC1) [58 mmol, 2 equiv.]. Vacuum distillation (bp = 128 °C, 0.4 Torr) yielded a colorless oil of2.3 [10.7 g, 82%]. 1HNMR data (300 MHz, CDC13): 34.18 (dt, 1 H, J5,6 =J,6 = 2.3 Hz, J45 = 7.5 Hz, H-5);4.00 (d, 1 H, J23 = 8 Hz, H-2); 3.91 (t, 1 H, J3,4=J4,5 = 7.5 Hz, 11-4); 3.82 (dd, 1 H, J56 = 2.3 Hz, J6, = 11Hz, 11-6); 3.78 (dd, 1 H, J5,6 = 2.3 Hz, J6,o’ = ii Hz, H-6); 3.75 (t, 1 H, J2,3 =J3,4= 7.5 Hz, H-3); 0.22, 0.20,0.19, 0.12 (4 s, 36 H, [{CH3)Si-0j4. 13CNMR data (75 MHz, CDC13): 6 171.02 (C-i); 81.28, 75.77, 73.00,70.71 (C-2 to C-5); 61.30 (C-6); 0.75, 0.55, 0.44, 0.26 (12 C, [{CH3)Si-O]4.DCIMS data: mJz 467 (MHj.2,6-Anhydro-1-deoxy-1,1-difluoro-3,4,5,7-tetra-O-(trimethylsilyl)-D-gluco-hept-1-enitol (2.4) (seeMotherwell et al., 1989). Freshly distilledCF2Br (obtained from Aldrich Chemical Co.) [4.5 g, 22 mmol, 5equiv.1 was weighed in 15 mL of dry THF and then added to a dry 3-necked flask. The solution was cooled inan ice bath, and then (Me2N)3P(obtained from Fluka Chemical Co.) [3.9 mL, 22 mmol, 5 equiv., dissolved in 5mL of dry THFI was added dropwise from a dropping funnel under N2. A white precipitate formedimmediately. The mixture was stirred at room temperature for 30 mm. Freshly activated zinc [1.41 g, 21.5138mmol, 5 equiv.] (Zn was activated with 10% HC1 for -1 miii, rinsed 3 times with acetone, 3 times with ether,and then dried in vacuo for -.2 hours) was added, the dried lactone 2.3 [2.0 g, 4.3 mmol, dissolved in —10 mL ofdry THF] was added by syringe, and then the reaction was stirred for 90 mm at 70 °C. Heating was stopped, thepmduct was extracted three times into ether, and then the combined ether layer was washed with NaC1 andH20.The ether layer was dried over MgSO4,the solvent was evaporated in vacuo, and then the residue was purifiedby column chromatography (1:1 CHC13:hex) to give an oil of 2.4 [0.64 g, 30%]. 1H NMR data (200 MHz,CDC13): 64.15 (t, 1 H, J3,4 =J3,F = 3.6 Hz, H-3); 3.86 (m, 1 H, H-6); 3.82 (dd, 1 H, J6,-j = 2Hz, J7,7 = 11.6Hz, H-7); 3.74 (dd, 1 H, J6,7’ = 4 Hz, J7, = 11.6 Hz, H-7’); 3.70-3.65 (m, 2 H, H-4, H-5); 0.14 (s, 27 H,[(CH3}3Si-O]); 0.12 (s, 9 H, {CH3}3Si-O). 19F NMR data (188.3 MHz, CDC13): 6 -103.03 (d, J = 78.7 Hz,Fb); -117.75 (d, J = 78.9 Hz, Fa); DCI MS data: nilz 500 (Mj; 501 (MHj.2,6-Anhydro-1-deoxy-1,1-difluoro-D-gluco-hept-1-enitol. (2.1). The heptenitol 2.4 [3.4 g, 6.8 mmol]was dissolved in —20 mL of MeOH,Me4NF.3H20[8.8 g, 5 equiv.J was added, and then the mixture was stirredfor 60 miii at room temperature. The Li+ form of Dowex 50W X2 resin was added to remove the ammoniumsalt (Me4Nj. The mixture was stirred overnight at room temperature, and then the resin was filtered andwashed with MeOH. The solvent was evaporated in vacuo, and then the product was purified further by columnchromatography (27:2:1 EtOAc:MeOH:H0). Lyophilization of the eluate gave a white powder [0.90 g, 62 %]of 2.1 (mp 67-70 °C). 1H NMR data (400 MHz,D20): 6 4.18 (m, 1 H, H-3); 3.89 (dd, 1 H, J77’ = 12.6 Hz,= 2 Hz, H-7); 3.75 (dd, 1 H, .17,7 = 12.6 Hz, J6,7 = 5.6 Hz, H-7’); 3.62-3.50 (m, 3 H, H-6, H-5, H-4).l3 NMR data (125.8 MHz,D20): 6 154.2 (t, JC,Fa = 1C,Fb = 283 Hz, C-i); 115.25 (dd, JC,Fa = 25 Hz, Jc,a=12.6 Hz, C-2); 82.19, 76.30, 69.12 (s, C-4, C-5, C-6); 68.13 (d, JC,F = 2.5 Hz, C-3); 60.74 (s, C-7). l9J’ NMRdata (188.3 MHz,D20): 6 -97.36 (d, JF,F = 74.3 Hz, Fb); -112.55 (dd, JF,F = 74.0 Hz, Fa). DCI MS data: mlz230 (M+N114j. Anal. Calc. forC7H10F205(212.15): C, 39.63; H, 4.75. Found: C, 39.72 H, 4.90.2,6-Anhydro-1-deoxy-1,1-difluoro-3,4,5,7-tetra-O-(t-butyldimethylsilyl)-D-gluco-hept-1-enitol(2.5) (see Corey & Venkateswarlu, 1972). The heptenitol 2.1 [0.374 g, 1.76 mmol, dissolved in -.10 mL of dryDMF] was persilylated as described above using TBDMS chloride [2.25 g, 8.4 equiv.] and imidazole [1.9 g, 10equiv.]. The reaction was stirred at room temperature for 2 hours under N2, a condenser was attached to the139flask, and then the mixture was stirred at 100 °C for 2 hours until the reaction was complete. Work up of theproduct was as described above. The product was purified by column chromatography (5:1 hex:CHC13)to yield2.5 [0.9 g, 76%]. 1HNMR data (400 MHz, CDC13): 6 4.27 (m, 1 H, H-3); 4.02 (ddd, 1 H, J56 = 7.6 Hz, J67’= 4.1 Hz, J6,7 = 2.9 Hz, H-6); 6 3.89 (d, 1 H, J5,6 = 7.9 Hz, H-5); 3.82 (dd, 1 H, .J-j, = 11.8 Hz, J6,7 = 2.8 Hz,H-7); 3.79-3.73 (m, 2 H, H-4, H-7’); 0.87 (x3), 0.84 (2 s, 36 H, [(M3C)MeSi-0j4);0.09, 0.08, 0.07 (x2), 0.05(x2), 0.04, 0.03 (6 s, 24 H, [(Me3C)MSi-0]). 13CNMR data (75 MHz, CDC13): 6 152.23 (dd, JC,Fb =268.9 Hz, JC,Fa = 287.2 Hz, C-i); 114.86 (dd, JC,Fa = 42.38 Hz, Jc,Fb = 12.2 Hz, C-2); 78.29, 75.64, 71.68 (s,C-4, C-5, C-6); 66.90 (d, J = 2 Hz, C-3); 61.87 (s, C-7); 25.84, 25.82, 25.65, 25.55 (s, 12 C, [(M3C)Me2Si-014); 18.30, 17.93, 17.88, 17.77 (s, 4 C, [(MeC)Me2Si-0j),-5.47, -5.11{x2}, -5.02{x2}, -4.73, -4.36, -4.12 (s,8 C, [(MeC)M2Si-0]4). 19k? NMR data (188.3 MHz, CDC13): 6 -104.90 (d, J = 85.0 Hz, Fb); -121.24 (dd, J= 85.0 Hz, Fa). FAD MS data: m/z 669 (MHj.(E)-2,6-Anhydro-1-deoxy-i-fluoro-3,4,5,7-tetra-O-(t-butyldimethylsilyl)-D-gluco-hept-1-enitol(2.6) (see Hayashi et al., 1979). The dried heptenitol 2.5 [0.250 g, 0.37 mmol] was combined with 1.8 mL ofRed Al® (obtained from Aldrich Chemical Co.) [0.89 M in toluene; diluted from a 3.4 M stock in toluene; anexcess of Red Al® was needed for complete reaction] in a reaction flask at 0 °C. The mixture was stirred at 0°C, allowed to warm slowly to room temperature, and stirring was continued until the reaction was complete.The reaction was quenched with ice, acidified with conc. HC1, the product was extracted four times into ether,and then the combined ether layer was washed with NaC1 and then H20. The ether layer was dried overMgSO4, the solvent was evaporated in vacuo, and then the residue was purified by column chromatography(20:1 hex:CH2C1)to yield an oil of 2.6 [0.11 g, 47%]. 1H NMR data (400 MHz, CDC13): 6 6.82 (d, 1 H, JH,F= 81.3 Hz, H-i); 4.57 (dd, 1 H, J34 = 3.1 Hz, J3,F = 2.2 Hz, H-3); 4.01 (ddd, 1 H, .156 = 8 Hz, f6,7 = 4.4 Hz,= 2.5 Hz, H-6); 3.83 (d, 1 H, J56 = 8 Hz, H-5); 3.80 (dd, 1 H, J67 = 2.5 Hz,J7 = 11.7 Hz, H-7); 3.79 (m,1 H, H-4); 3.72 (dd, 1 H, f6,7 = 4.3 Hz, J7,’= 11.7 Hz, H-7’); 0.87 (x3), 0.83 (2 s, 36 H, [(M3C)Me2Si-0]4);0.10 (x2), 0.09, 0.08, 0.07 (x3), 0.03 (5 s, 24 H, [(Me3C)M2Si-0]). 13CNMR data (75 MHz, CDC13): 6143.27 (d, JC2,F = 29.9 Hz, C-2); 138.42 (d, Jc1,F = 232.9 Hz, C-i); 77.90, 75.83, 72.27 (s, C-4, C-5, C-6);65.24 (s, C-3); 62.33 (s, C-7); 25.89, 25.84, 25.70, 25.63 (s, 12 C, [Uk3C)Me2Si-0]4); 18.33, 17.95{x2}, 17.85140(s, 4 C, [(Me3C)Me2Si-O]4), -5.28, -5.15, -5.03, -4.99, -4.94, -4.70, -4.25, -4.02 (s, 8 C, [(Me3C)M2Si-O]4).19FNMR data (188.3 MHz, CDC13): 6 -181.5 (d, JH-1,F = 81 Hz).(E)-2,6-Anhydro-1-deoxy-1-fluoro-D-gluco-hept-1-enitol (2.2). The heptenitol 2.6 [0.230 g, 0.35mmol] was dissolved in 1.75 mL [5 equiv.J of a 1 M solution of TBAF in THF and then stirred for 60 mm atroom temperature. The Li form of Dowex 50W X2 resin was added to remove the ammonium salt (Bu4Nj.The mixture was stirred for several hours at room temperature, and then the resin was filtered and washed withMeOH. After removal of the solvent the residue was purified by column chromatography (27:2:1EtOAc:MeOH:H20)to give an oil of 2.2 [0.060 g, 87%]. 1HNMR data (400 MHz,D20): 6 7.01 (dd, 1 H, J1,F= 78.6 Hz, J1,3 = 1.8 Hz, H-i); 4.18 (ddd, 1 H, J5,6 = 7.5 Hz, J6,-j’ = 4.3 Hz, J6,7 = 1.5 Hz, H-6); 3.81 (dd, 1 H,= 2.0 Hz,J7, = 12 Hz, H-7); 3.68 (dd, 1 H, J6,7 = 5.4 Hz, J7, = 12 Hz, H-7’); 3.54 (t, 1 H, J45 = J5,6 =7.7 Hz, H-5); 3.42 (m, 1 H, H-4). NOTE: No resonance could be assigned to H-3, presumably because thissignal was hidden by the resonance attributable to water (6 = 4.88-4.65 ppm). 19F NMR data (188.3 MHz,D20): 6 -169.71 (d, JH-l,F = 78 Hz). Anal. Caic. forC7H11F05(194.16): C, 43.30; H, 5.67. Found: C, 43.05;H, 5.70.6 OR OAc OR_______ ____AO_::R1cN_______ ____[1.1] H H [3.5] H [3.7] Ac CN[3.12] H NHAc [3.6] Br [3.8] H CN[5.3] SiMe2(t-Bu) H [5.4] SiMe2(t-Bu) CH3[5.6] Ac CN [5.5] H CH3[5.7] H CNFigure 5.2. Structures of glucal derivatives and associated synthetic intermediates.The numbering systems used in this chapter for the various compounds are indicated.141b. The synthesis ofglucal and its derivatives.1,5-Anhydro-2-deoxy-D-arabino-hex-1-enitol (1.1). l,5-Anhydro-2-deoxy-3,4,6-tri-O -acetyl-Darabino-hex-1-enitol (Sigma Chemical Co.) was deacetylated using NaOMe/MeOH as described above (seeZemplen & Pacsu, 1929). The product was recrystallized from hot EtOAc to yield D-glucal 1.1 [99%] (mp57-58 °C; lit. 57-59 °C, Roth & Pigman, 1963). 1H NMR data (400 MHz,D20): 8 6.42 (dd, 1 H, Ji,2 = 7.0Hz, J1,3 = 1.7 Hz, H-i); 4.82 (dd, 1 H, f1,2 = 7.0 Hz, J2,3 = 2.8 Hz, H-2); 4.23 (di; 1 H, J45 = 7.0 Hz, J56 =J5,6 = 1.7 Hz, H-5); 3.94-3.84 (m, 3 H, H-3, H-6, H-6’); 3.69 (dd, 1 H, J45 = 7.0 Hz, .134 = 9.4 Hz, H-4).1,5-Anhydro-2-deoxy-3,4,6-tri-O-(t-butyldimethylsilyl)-D-arabino-hex-1-enitol (5.3) (see Corey &Venkateswarlu, 1972). A dry 3-necked flask was charged with the glucal 1.1 [0.5 g, 3.4 mmol], which waspersilylated as described above using TBDMS chloride [2.58 g, 17 mmol, 5 equiv.] and imidazole [2.33 g, 34mmol, 10 equiv., dissolved in -8 mL of dry DMF]. The reaction was stirred at room temperature for 2 hoursunderN2, a condenser was attached to the flask, and then the mixture was stirred at 45 °C for 10 hours until thereaction was complete as determined by TLC. Work up of the product was as described above. The productwas purified by column chromatography (3:1 hex:CHC1)to yield an oil of 5.3 [1.35 g, 81%]. 1H NMR data(300 MHz, CDC13): 66.33 (d, 1 H, J12 = 6.4 Hz, H-i); 4.65 (ddd, 1 H, .11,2 = 6.4 Hz, J2,3 = 4.5 Hz,J24= 1.0Hz, H-2); 3.99 (m, 1 H, H-5); 3.95 (dd, I H, J5,6 = 3.7 Hz, J6,6 = 11 Hz, H-6); 3.89 (m, 1 H, H-3); 3.80-3.73 (m,2 H, H-4, H-6’); 0.90, 0.89 (x2) (2 s, 27 H, [(MC) eSi-O]3); 0.10 (x2), 0.08 (x2), 0.06, 0.05 (4 s, 18 H,[(Me3C)M2Si-O]3).2,6-Anhydro-1,3-dideoxy-4,5,7-tri-O-(t-butyldimethylsilyl)-D-arabino- hept-2-enitol (5.4) (seeLesimple et al., 1986). The glucal 5.3 [1.30 g, 2.66 mmol] was combined with 1.5 mL of dry THF, and then thesolution was cooled to -78 °C. Immediately after the addition of 4.4 mL of t-BuLi [1.84 M in pentane, 8 mmol,3 equiv.] the colorless solution turned yellow (due to the formation of the t-BuLi-THF complex). The mixturewas stirred at -78 °C for 15 mm, then at 0 °C for 45 mm. About 5 mL of THF was added at 0 °C to react withthe excess t-BuLi, and then the solution was cooled to -78 °C. Methyl iodide [1.0 mL, 2.3 g, 16 mmol, 6 equiv.,dried by passage through a pipette filled with flame-driedAl203]was added at -78 °C, and then the solution wasstirred and slowly allowed to warm up to room temperature. The reaction was stopped by the addition ofN1{4C1142thenH20, and then the product was extracted 4 times with ether. The combined organic layer was washed withH20, filtered, and then the solvent was evaporated in vacuo. 1 NMR analysis showed that the reaction yieldeda mixture of 5.3 and 5.4, and it was not possible to purify the product by column chromatography, as the Rfvalues of 5.3 and 5.4 were too similar.2,6-Anhydro-1,3-dideoxy-D-arabino-hept-2-enitol (5.5). The unpurified mixture from the precedingsynthesis [0.90 g, —1.8 mmol of a mixture of 5.3 and 5.4] was cooled to 0 °C underN2, 11 mL of t-Bu4NF [1.0M in THF, 11 mmol, 6 equiv.] was added, and the mixture was stirred under N2 at 0 °C for 15 mm and then atroom temperature for 2 hours. The solvent was evaporated in vacuo, the residue was purified by columnchromatography (4:1:2 CHC13:MeOH:hex) 3 times (to remove t-Bu4Nj, and then the final eluate wascrystallized and recrystallized from ether, MeOH, and pet. ether to yield crystals of 5.5 [0.110 g, 27% from 5.3](mp 100-102 °C). 1HNMR data (300 MHz,D20): 34.58 (dd, 1 H, J34 = 2.9 Hz, J13 = 1.0 Hz, H-3); 4.14(ddd, 1 H, Jj4 = 1.7 Hz, J34 = 2.9 Hz, J4,5 = 6.7 Hz, H-4); 3.88 (ddd, 1 H, J67 = 3.3 Hz, J6,’ = 4.8 Hz, J5,6 =8.4 Hz, H-6); 3.82 (m, 2 H, H-7, H-7’); 3.60 (dd, 1 H, J4,5 = 6.7 Hz, J5,6 = 8.4 Hz, H-5); 1.73 (dd, 3 H, J1,4=1.7 Hz, J1,3 = 1.0 Hz, H-i). DCI MS data: m/z 178 (M+NH4j. Anal. Caic. forC7H1204(160.2): C, 52.49;H, 7.55 Found: C, 52.29 H, 7.76.1,5-Anhydro-2-deoxy-2-acetamido-D-arabino-hex-1-enitol (3.12). The peracetylated precursor1,5 -anhydro-2-deoxy-2(N-acetylacetamido)-3,4,6-tri-O-acetyl-D-arabino-hex- 1-enitol (3.11) was first purifiedby column chromatography (1:1 hex:EtOAc). Deacetylation of 0.44 g of this starting material using sodiummethoxide in methanol (Pravdi’c & Fletcher Jr., 1967) removed one of the two N-acetyl groups and all threeO-acetyl groups. The product was purified by column chromatography (25:10:4 EtOAc:EtOH:H20)[0.4 g,91%] and then recrystallized from 2-PrOH, CH3N, and hexane to yield 3.12 [0.25 g, 57%] (mp 120- 122 °C; lit.124-125 °C, Pravdiô & Hetcher Jr., 1967). 1H NMR data (300 MHz,D20): 6 6.71 (d, 1 H, Jj3 = 1 Hz, H-i);4.29 (dd, 1 H, J13 = 1 Hz, J3,4 = 6.4 Hz, H-3); 4.02 (dt, 1 H, .15,6 = .15,6 = 4.2 Hz, J45 = 8.5 Hz, H-5); 3.89 (d,2 H, J5,6 = J5,6’ = 4.2 Hz, H-6, H-6’); 3.79 (dd, 1 H, J3,4 = 6.4 Hz, J4,5 = 9.0 Hz, H-4); 2.10 (s, 3 H,]j3CCONH-). 13C NMR data (50 MHz,D20): 6 175.29 (H3CCONH-); 142.46 (C-i); 113.97 (C-2); 79.32,14369.56, 69.15, 60.63 (C-3, C-4, C-5, C-6); 22.64 (H3CCONH-). Anal. Calc. forC8H13N05(203.20): C, 47.29;H, 6.45; N, 6.89. Found: C, 47.29; H, 6.39; N, 6.75.1,5-Anhydro-2-deoxy-2-cyano-3,4,6-tri-O-acetyl-D-arabino-hex-1-enitol (5.6) (see Hall & Jordaan,1973). 1 ,5-Anhydro-2-deoxy-3,4,6-tri-O-acetyl-D-arabino-hex-1-enitol (obtained from Sigma Chemical Co.)[0.50 g, 1.8 mmolJ was dried in vacuo for a few hours, then dissolved in 5 mL of dry ether. A solutioncontaining 5 mL of dry ether and 0.16 mL of chiorosulfonyl isocyanate was prepared, and then a droppingfunnel was used to add this solution to the reaction at 0 °C over a period of 30 mm. The reaction was stirredovernight at 0 °C, and then a solution containing 1.3 mL of dry CH21 and 0.24 mL of dry Et3N [0.19 g,1 equiv., dried over KOHl was added at 0 °C. The reaction was slowly allowed to warm to room temperature,poured into 25 mL ofH20, and then the organic solvent was evaporated with a stream of N2. The aqueousresidue was extracted 4 times withCH21,the combined organic layer was washed withH20, the solvent wasevaporated in vacuo, and then the residue was purified by column chromatography (2:1 hex:EtOAc) to give 5.6[0.15 g, 27%]. 1H NMR data (200 MHz, acetone-D6): 8 7.58 (s, 1 H, H-i); 5.53 (d, 1 H,J34 = 5 Hz, H-3);5.23 (dd, 1 H, .13,4 = 5 Hz, J45 = 6 Hz, H-4); 4.70 (td, 1 H, J4,5 = J5,6 = 6 Hz, J5,6’ = 3 Hz, H-5); 4.49 (dd, 1 H,= 6 Hz, Jo,o’ = 13 Hz, H-6); 4.28 (dd, 1 H,J56= 3 Hz, J6 = 13 Hz, H-6’); 2.09, 2.05, 2.02 (3 s, 9 H,[AcO-]3). 13C NMR data (50 MHz, CDC13): 6 170.21, 169.50, 169.11 (s, [H3CCOO-]); 157.84 (s, C-i);115.54 (s, conj. CN); 88.19 (s, C-2); 75.46, 65.20, 64.25 (3x s, C-3, C-4, C-5); 60.44 (s, C-6); 20.61 {x2}, 20.57(s, {H3COO-]). IR data: Vmax (cm4): 2223 (conj. CN); 1757 (C=O; 1631 (conj. C=C). DCIMS data: in/z315 (M+NH4j.1,5-Anhydro-2-deoxy-2-cyano-D-arabino-hex-1-enitol (5.7). The glucal 5.6 was dried in vacuoovernight prior to deacetylation usingNH3-saturated MeOH as described above. The reaction was left at 4 °Cfor a few hours until the reaction was complete as determined by TLC (27:2:1 EtOAc:MeOH:H0). The solventwas evaporated in vacuo to give a solid that was recrystallized from acetone and hexane to give colorlessneedles of 5.7 [0.17 g, 57%] (mp 123-125 °C). 1ff NMR data (200 MHz,D20): 8 7.30 (d, 1 H, J13 = 1 Hz,H-i); 4.29 (dd, 1 H, Ji,3 = 1 Hz, J3,4= 7 Hz, H-3); 4.12 (ddd, 1 H, J56 = 3 Hz J5,6 = 4 Hz, J4,5 = 9 Hz, H-5);3.89 (d, 1 H, J56 = 3 Hz, H-6); 3.86 (d, 1 H, J5,6’ = 4 Hz, H-6’); 3.73 (dd, 1 H, J34 = 7 Hz, J4,5 = 9 Hz, H-4).144l3 NMR data (50 MHz,D20): 6 159.11 (s, C-i); 118.70 (s, CN); 91.10 (s, C-2); 80.89, 67.55, 67.39 (s, C-3,C-4, C-5), 60.25 (C-6). IR data: Vmax (cm1): 2220 (conj. CN); 1627 (conj. C=C). DCI MS data: m/z 189(M+NH4j. Anal. CaIc. forC7H9N04(171.15): C, 49.12; H, 5.30; N, 8.18. Found: C, 49.08; H, 5.38; N, 8.12.2,3,4,6-Tetra-O-acetyl-J3-D-glucopyranosyl cyanide (3.5) (see Fuchs & Lehmann, 1975). 2,3,4,6-Tetra-O-acetyl-c-D-glucopyranosyl bromide (3.4, see Lemieux, 1963) [10 g, 24 mmol] and Hg(CN)2 [13 g,51 mmol, 2 equiv.] were dried in a dessicator in vacuo overP205 and KOH for 2 days. The starting materialswere ground in a mortar and mixed together, and then disthbuted evenly on the bottom of a 125-mL Erlenmeyerflask. The mixture was heated at 80-85 °C under N2 for 30 mm, and then the melted solid was dissolved in-.100 mL of wann CHCI3. The solution was filtered through a bed of Celite, the filtrate was washed 3 timeswith 10% KBr (aq), the organic solvent was evaporated in vacuo, leaving a solid that was crystallized andrecrystallized from EtOH to give white crystals of 3.5 [5.2 g, 60%] (mp 113 °C; lit. 114-115 °C, Myers & Lee,1984). 1H NMR data (200 MHz, CDC13): 6 5.30 (t, 1 H, J12 =J2,3 = 9 Hz, H-2); 5.24-5.03 (m, 2 H, H-3,H-4); 4.30 (d, 1 H, J1,2 = 9Hz, H-i); 4.23 (dd, 1 H,J56 = 4.8 Hz, .16,6 = 12Hz, H-6); 4.10 (dd, 1 H, Js,6’=2 Hz, J6,6’ = 12 Hz, H-6’); 3.69 (ddd, 1 H, J4,5 = 10 Hz, J5,6 = 4.8 HzJ5,6 = 2 Hz, H-5); 2.09 (x2), 2.01, 2.00(3 s, 12 H, [AcO-]4). 13C NMR data (50 MHz, CDCJ3): 6 170.46, 170.02, 169.10, 168.69 (s, [H3COO-]4);114.10 (conj. CN); 76.81, 72.80, 68.94, 67.25, 66.46, 61.41 (s, C-i, C-2, C-3, C-4, C-5, C-6); 20.64, 20.48 (x2),20.36 (s, [H3COO-]4).2,3,4,6-Tetra-O-acetyl-1-a-bromo-13-D-glucopyranosyl cyanide (3.6) (see Lichtenthaler & Jarglis,1982). The glucosyl cyanide 3.5 [2.8 g, 7.8 mmol] was dried in vacuo overnight and then dissolved in 100 mLof CC!4. N-Bromosuccinimide [5.5 g, 31 mmol, 4 equiv., recrystallized from hotH20and then dried overP205in vacuo] and a catalytic amount of BaCO3 [0.6 g, 3 mmol] were added, and then a condenser and a drying tubewere connected to the reaction flask. The reaction was refluxed for ...90 mm while it was irradiated with a250 W tungsten lamp. The mixture was cooled on ice to give the product as a solid, which was collected byfiltration prior to further purification by column chromatography (3:1 hex:EtOAc) and recrystallization fromanhydrous ether and pet. ether to yield white needles of 3.6 [2.1 g, 60%] (mp 100-101 °C; lit. 92 °C,Lichtenthaler & Jarglis, 1982). 1H NMR data (200 MHz, CDC13): 6 5.39 (t, 1 H, J23 =J3,4 = 9.6 Hz, H-3);1455.25 (d, 1 H, J2,3 = 9.6 Hz, H-2); 5.16 (t, 1 H, J3,4 =J4,5 = 9.6 Hz, H-4); 4.31 (dd, 1 H, J5,6= 3 Hz, J66’ = 12Hz, H-6); 4.25 (ddd, 1 H, J4,5 = 9.6 Hz,J5 6’ = 1.6 Hz, J5,6 = 3 Hz, 11-5); 4.08 (dd, 1 H, J5,6’ = 1.6 Hz,J6’ = 12Hz, H-6’); 2.16, 2.12, 2.04, 2.01 (4 s, 12 H, [AcO-]4). DCI MS data: m/z 455, 453 (M+NTj.2,6-Anhydro-3-deoxy-4,5,7-tri-O-(acetyl)-D-arabino-hept-2-enononitrile (3.7) (see Somsák et al.,1990). The brommated glucosyl cyanide 3.6 [2.82 g, 6.4 rnmol] was dissolved in benzene, Zn [1.7 g, 29 minol,4.5 equiv.] was added, and then the mixture was stirred under N2 and heated to reflux. Pyridine [0.50 mL,6.4 mmol, 1 equiv.] was added to the refluxing suspension and then the reaction was continued for 30 mm. Thesolid was evaporated by filtration and washed with ether. The filtrate was washed with KHSO4 then 1120, andthen the organic layer was dried overMgSO4. Evaporation of the solvent in vacuo gave a white solid which waspurified further by column chromatography (2:1 hex:EtOAc) to give crystals of 3.7 [1.72 g, 90%]. 1H NMRdata (400 MHz, CDC13): 6 5.68 (d, 1 H, J34 = 4 Hz, H-3); 5.33 (dd, 1 H, J34 = 4 Hz, .145 = 5.5 Hz, 11-4); 5.19(dd, 1 H, .145 = 5.5 Hz, J5,6= 6.4 Hz, H-5); 4.41-4.36 (m, 2 H, H-6, H-7); 4.15 (dd, 1 H, J-j,-j’ = 15 Hz, J67’ = 5Hz, H-7’); 2.08, 2.04, 2.02 (3 s, 9 H, [AcO-]3).2,6-Anhydro-3-deoxy-D-arabino-hept-2-enononitrile (3.8) (see Fritz et al., 1983). The cyanoglucal3.7 [2.82 g, 6.4 mmol] was dried in vacuo overnight prior to deacetylation usingNH3-saturatecl MeOH asdescribed above. The reaction was left at 4 °C for 4.5 hours until the reaction was complete. The solvent wasevaporated in vacuo to give a solid that was purified further by column chromatography (27:2:1 EtOAc:MeOH:H20) and recrystallized from acetone, ether and hexane to give 3.8 [0.20 g, 70%] (mp 95-97 °C).1H NMR data (200 MHz,D20): 6 5.80 (d, 1 H,J34 = 3 Hz, H-3); 4.30 (dd, 1 H, J3,4 = 3 Hz, .145 = 7 Hz, H-4);4.05 (ddd, 1 H, J5,6 = 9 Hz, J6,7 = 2.8 Hz, J6,7’= 4 Hz, H-6); 3.88 (d, 1 H, .167 = 2.8 Hz, H-7); 3.86 (d, 1 H,J6,7’=4 Hz, H-7’); 3.71 (dd, 1 H, J4,5 = 7 Hz, J5,6 =9 Hz, H-5). 13C NMR data (50 MHz,D20): 8 129.04 (s,C-2); 118.98 (s, C-3); 114.72 (s, CN); 81.03, 68.68, 68.02, 60.34 (s, C-4-7). DCI MS data: m/z 189(M-i-NH4j. Anal. Calc. forC7H9N04(171.15): C, 49.12; H, 5.30; N, 8.18. Found: C, 49.42; H, 5.29; N, 8.26.1465.2. ENZYME KINETICS.5.2.1. Miscellaneous procedures and defmition of enzyme activity units.Absorbance measurements were made using either a Perkin-Elmer Lambda 2 model, or a Pye-UnicamPU-8800 or PU-8700 model UV-VIS spectrophotometer, each of which was equipped with a circulating waterbath and thermostatted cuvette holders. In all cases absorbance measurements were made using cuvettes with a1.00-cm pathlength.All pH measurements were performed using a Radiometer PHM 82 pH meter equipped with an Orioncombination electrode (Model no. 8103). Prior to use this instrument was standardized using commerciallyobtained standard pH buffers.In all cases, one unit (U) of enzyme activity represents an amount capable of catalyzing the formationof one micromole of product per minute under the conditions specified for each assay.5.2.2. Enzymes and enzyme assays used in this work.a. Glycogen phosphoiylase.Glycogen phosphorylase b (E.C. 2.4.1.1) was prepared from rabbit muscle by (now Dr.) WilliamStirtan of Prof. Withers laboratory (see Stirtan, 1993). The concentration of this protein in stock solutions wasdetermined from A2s0 measurements using an A9°b0 = 1.32 cm-1 (Buc & Buc, 1968).b. The assayfor glycogen phosphorylase activity.In most cases phosphorylase activity was assayed in the direction of glycogen synthesis, and in thesecases initial rates were determined by measuring the amount of inorganic phosphate released in a 5-minute timeperiod (Engers et al., 1970).Except as noted, the buffer for all kinetic experiments using phosphorylase in the glycogen synthesisreaction contained 100 mM KC1, 1 mM EDTA, 1 mM DTT, and 50 mM triethanolamine hydrochloride pH 6.8.Reaction mixtures (0.500 mL) also contained 1 mM AMP and 0.5-1% glycogen. Reactions were carried out at30 °C for 5 mm. In most cases the enzyme was preincubated with AMP and glycogen and then added to a147solution containing a substrate with or without an inhibitor. The enzyme concentration in the reaction mixturedepended on the kinetic parameter under study, and is given in the figure legends.When phosphorylase activity was assayed in the direction of glycogen phosphorolysis, initial rates weredetermined using a phosphoglucomutase/glucose-6-phosphate dehydrogenase coupled assay (Engers et at,1969). The reaction buffer for these assays was adjusted to pH 6.8 and contained 35 mM imidazole, 20 mMsodium glycerophosphate, 10 mM Mg(OAc), 5 mM DTT, 1 mM EDTA, and also contained 1 mM AMP, 0.5%glycogen, 1 mM (3NADP, 15 U/mL phosphoglucomutase, and 3.4 U/mL of glucose-6-phosphate dehydrogenase.In these coupled assays the reduction of NADP is quantitated by following the change in absorbance at 340 urn.These assays were performed by Ms. Karen Rupitz.Technical assistance with the kinetics experiments using glycogen phosphorylase was also provided by(now Dr.) William Stirtan.c. Agrobacterium /3-glucosidase.The cloned Agrobacterium 13-glucosidase pABG5 was prepared as described (Kempton & Withers,1992). The concentration of this protein in stock solutions was determined fromA280 measurements using anA9% = 2.184 cm (Street, 1988).d. The assayfor Agrobacteriuni /3-glucosidase activity.Except as noted, initial rates of 3-glucosidase activity were measured by adding aliquots of the enzymeto a cuvette containing 1.00 mL of the substrate f3G1cPNP in 50 mM sodium phosphate buffer (pH 6.8)containing 0.1% BSA. The cuvette and its contents were equilibrated to 37 °C in the spectrophotometer justprior to addition of the enzyme. Absorbance readings at 400 urn (A400)were measured to give initial rates inA400 units per minute. Enzyme concentration and reaction times were chosen to ensure that <10% of thesubstrate was consumed, thus ensuring linear kinetics. The £41J of 4-nitrophenol at pH 6.8 and 37 °C is7280M4cnr1(Street, 1988).148e. /3-N-Acetylhexosaminidases.Sigma Chemical Company supplied 3-N-acetythexosaminidases (NAGases) isolated from jack bean(J-NAGase), bovine kidney (K-NAGase), and human placenta (H-NAGase). The catalogue numbers of theseproducts were A-2264, A-2415, and A-6152, respectively.f Assays for J3-N-acerylhexosaminidase activity.Kinetic studies of hydrolysis reactions were performed by following changes in UV-VIS absorbanceusing a spectrophotometer equipped with a circulating water bath that maintained the 1.00-cm cuvettes at 25 °C.Except as noted, reaction buffers contained BSA (0.1%), NaCl (100 mM) and citrate buffer (50 mM), and wereadjusted to pH 5.0 (used with jack bean NAGase) or pH 4.25 (used with human placenta or bovine kidneyNAGase). Molar extinction coefficients for 4-nitrophenol and GlcNAcPNP were determined at 25 °C bymeasuring the absorbance at 360 rim of carefully prepared stock solutions of each compound in the appropriateenzyme reaction buffer (pH 5.0 or 4.25). The molar extinction differences at 360 rim (Ae360) determined for3GlcNAcPNP and 4-nitrophenol at pH 5.0 and pH 4.25 were 2280 and 2150Mcm, respectively.The initial rates of 3GlcNAcPNP hydrolysis were determined by incubating the reaction buffer at 25 °Cin the thermostatted cuvette-holder of the spectrophotometer. An appropriate volume of stock substrate solutionwas added to the cuvette —4 mm before the reaction (to ensure negligible spontaneous hydrolysis of the substrateduring the pre-incubation and reaction periods). Reactions were initiated by the addition of enzyme (inBSA-containing buffer) by syringe, and the reactions were monitored at 360 um. Initial rates were determinedusing 6-7 different substrate concentrations, which ranged from about one-third to (in most cases) three times thevalue of the Km ultimately determined. However, due to the high background absorbance of 3GlcNAcPNP at360 rim, its concentration was kept below 2 mM. The rates of I3G1cDNP hydrolysis were determined using acomparable assay, except the reaction was monitored at 400 rim. In these (0.200-mL) reactions, 1.5 or 6.5 mM3GlcDNP was incubated in the appropriate buffer for several hours at 25 °C with (or without) 6 jig ofJ-NAGase. The &400 determined for 3GlcDNP and 2,4-dinitrophenol at pH 5.0 was 9750M4cm.1495.2.3. The determination of kinetic parameters for substrates.a. Determinations ofKm and Vm for various substrates.Approximate values of Km and Vmax were determined by measuring initial reaction rates using3 different (and wide-ranging) concentrations of the substrate. Accurate values of these parameters were thendetermined by measuring initial reaction rates using 5-8 different concentrations of the substrate, concentrationsthat typically ranged between 0.3-5 times the approximately determined Km value (Lineweaver & Burk, 1934).Values of Km and Vmax, together with the errors associated with the scatter of the data, were calculatedby fitting the data to a weighted nonlinear regression of the Michaelis-Menten equation using the computerprogram GraFit (Leatherbarrow, 1990).Initial-rate data were also plotted according to the method of Lineweaver and Burk (1934), but thismethod was not used for calculating kinetic parameters due to inaccuracies associated with such plots.However, these plots are included in this thesis because they are a useful tool for recognizing deviations fromlinear behaviour caused by allostery or various types of inhibition.b. Determinations of reaction rates for the catalytic hydration of heptenitol.Eight 0.500-mL reaction buffer mixes containing sodium phosphate buffer (50 mM, pH 6.8), no BSA,and one of eight different concentrations of heptenitol were prepared. Appropriate control reactions (with thesame concentrations of heptenitol but without enzyme) were also carried out. Reactions were pre-incubated andcarried out at 37 °C, and were started by the injection of 30 .tg of pABG5 (in 10 iL) into each solution.Reactions were carried out for 10 mm.The reaction product, 1-deoxy-D-gluco-heptulose, as well as standard solutions of the same, wasassayed by a cuprimetric method (Hehre et al., 1980) with slight modifications. The basis of this method is thestoichiometric reduction of Cu(II) (by the reducing sugar) to Cu(I) oxide, which in turn reducesarsenomolybdate to molybdenum blue. The absorption at 540 nm of the latter is a measure of the sugarconcentration. Each reaction was stopped by transferring 0.200 mL of the 0.500-niL reaction mixture into afresh tube containing 2.00 mL of Somogyi’s reagent (Somogyi, 1952). Each stopped reaction mixture was150placed in a boiling water bath for 10 mm and then cooled. Nelson’s reagent [2.00 mU (Nelson, 1944) and 1.5mL of double-deionized water were added per tube, each reaction was mixed, and then the absorbance at 540 nmwas measured on a Pye-Unicam PU-8700 spectrophotometer. Nine dilutions of stock l-deoxy-D-glucoheptulose were used to construct the standard curve for the assay. There was a linear relationship between theA540 of the cuprimetric assay and the amount of 1-deoxy-D-gluco-heptulose over the range of 0.56 to 7.9 mM(in the original 0.500-mL reactions).Initial rates were calculated after correction for the rate of nonenzymatic hydration in appropriatecontrol reactions. Determinations of Km and Vmax values were as described above using the computer programGraFit (Leatherbarrow, 1990). The standard curve was used to quantitate the reaction product.c. Determinations of reaction rates for the catalytic hydration ofmethylgiucal.This assay utilized 1H NMR to quantitate the reaction product. Five 0.500-mL reaction buffer mixescontaining HEPES-NaOH (10 mM, pH 7.0), BSA (0.1%), and one of five different concentrations ofmethylglucal were prepared. Reactions were pre-incubated and carried out at 37 °C, and were started by theinjection of 0.65 mg of pABG5 into each solution. Reactions were carried out for 110 hours and then stoppedby cooling to 0 °C. Water was removed by lyophiization and then several cycles of resuspension withD20andlyophilization were carried out. The 1H NMR spectrum was obtained for eachD20-exchanged reaction using a400 MHz Bruker WH-400 instrument. The concentration of the reaction product was determined by integrationof the product methyl hydrogen peak (at 6 = 1.43 ppm), which was corrected by subtracting the integral of thesame peak in the appropriate control (reactions with the same concentration of methylglucal but withoutenzyme). The number of micromoles of product formed was calculated from the ‘H NMR data using theequation shown below. Determinations of Km and Vmax values were as described above.I (Area of the product 1 o1 oftmol of j methyl 1H peak ) I substrateproduct = X originallyformed 1 (Area of the product’\ + (Area of the substrate \ j present in\ methyl H peak ) methyl H peak ) J the assay1515.2.4. The determination of kinetic parameters for inhibitors.a. Determinations ofK values (reversible inhibition).Approximate values for inhibition constants (range-finding or RF K1 values) for reversible inhibitorswere determined by measuring initial reaction rates at a single substrate concentration (usually equal to the Km)and 5 or 6 different concentrations of the inhibitor. An accurate Vmax value was then determined in a parallelexperiment using several different concentrations of the substrate, but the same amount of enzyme as was usedin the first experiment (i.e., with the inhibitor present). The data from the first experiment was then plotted asthe inverse of the initial reaction rate vs. the inhibitor concentration (Dixon, 1972). The RF K1 value is easilyobtained from the intercept (—K) of the line through the plotted data and a horizontal line drawn at l/Vmax(Dixon, 1972).In some but not all cases, inhibition constants were determined accurately by measuring initial reactionrates at 4 or 5 substrate concentrations bracketing the Km value, at each of 4 or more inhibitor concentrationsbracketing the ‘range-fmding’ K1 value, and also in the absence of inhibitor. The initial-rate data obtained at eachsubstrate and inhibitor concentration were then fitted to equations describing different types of enzymeinhibition (and which yield a K1 value for each case), using the nonlinear regression analysis computer programGraFiV’ (Leatherbarrow, 1990). The K1 value for the best fit of the data was reported in all cases, but in somecases more than one K value was reported because the data fit well to equations describing different types ofinhibition. Double-reciprocal (or Dixon) plots were also used as a convenient graphical method to show thepattern of inhibition.b. Determinations ofK values using glycogen phosphorylase.In most cases the inhibition of glycogen phosphorylase activity was assayed in the direction ofglycogen synthesis (Engers et al., 1970). Initial rates were determined by measuring the amount of inorganicphosphate released in a 5-minute time period (Engers et al., 1970; described above). Inhibition constants weredetermined accurately by measuring initial reaction rates at 5 substrate concentrations, at each of 5 or moreinhibitor concentrations. The data were then analyzed using Hill plots (i.e., plots of hi[V/{Vmax — v}] V5. ln[S]).152The Km,app values are easily obtained from the intercept (in [Km,appl) of the line through the plotted data and theabscissa (ln[SJ axis). Replots of the Km,app values vs. the inhibitor concentrations yielded the inhibitionconstant, K, from the intercept (—K1)of the line through the plotted data and the abscissa ([I] axis).The inhibition of glycogen phosphorylase activity was also assayed in the direction of glycogenphosphorolysis for the fluoroheptenitol inhibitors. Initial rates were determined using a phosphoglucomutase/glucose-6-phosphate dehydrogenase coupled assay (Engers et at, 1969; described above). In these cases RF K1values were determined using Dixon plots as described above.c. Irreversible inhibition (inactivation) tests: Experimental methods.F2hept was tested as an inactivator of glycogen phosphorylase b by incubating 17 mMF2hept, 100 mMKC1, 1 mM EDTA, 1 mM DTT, 50 mM triethanolamine hydrochloride (pH 6.8), 1% glycogen, 5 mlviorthophosphate, 1.5 mM AMP, and 0.6 mg of enzyme, for 5 days at room temperature. Over the course of thisincubation period, small aliquots (10 giL) of the above reaction were removed, and then added to fresh tubes,each of which contained 0.500 mL of a reaction mixture containing a saturating concentration (16 mM) ofaG1P, 1 mM AMP, 1% glycogen, 100 mM KC1, 1 mM EDTA, 1 mM DTT, and 50 mM triethanolaminehydrochloride (pH 6.8). Initial reaction rates were then assayed in the direction of glycogen synthesis asdescribed (Engers et al., 1970).The following inactivation tests were performed with either [3-glucosidase or NAGase, and in thesecases the substrates employed were the chromogenic glycosides [3GIcPNP and [3G1cNAcPNP, respectively. Inboth cases the residual activity was detennined by measuring the release of 4-nitrophenol spectrophotometricallyas described earlier. In these inactivation tests, reaction mixtures were set up containing one of several differentconcentrations of the putative inactivator in the appropriate buffer for each enzyme (see above; in all cases thesebuffers contained 0.1% BSA), and then the reactions were incubated at the enzyme’s optimal temperature. Equalaliquots of enzyme were added to each reaction, the reactions were incubated at the indicated temperature, andthen the residual enzyme activity was measured at various time intervals. This was done by removing a 10-IlLaliquot of the inactivation reaction mixture and then adding this aliquot to a fresh tube containing a much largervolume (typically 0.600 or 1.00 mL) of a saturating concentration of the substrate (as well as the appropriate153buffer for each enzyme; see above). For assays of residual j3-glucosidase activity, -4.0 mM f3G1cPNP was used(Km = 78 pM, Kempton & Withers, 1992). For assays of residual NAGase activity, -.1.4 mM I3G1cNAcPNP wasused (Km 0.62 mM, Li & Li, 1970, and this work) due to the high background absorbance of the substrate at360 urn.d. Irreversible inhibition (inactivation) tests: Theory and calculations.The inactivation of a glycosylase can be expressed by the following Scheme (5.1):K1I+E - El E—IScheme 5.1. A kinetic model for the inactivation of an enzyme (E) by an inactivator (I).Here the inactivator (I) and the free enzyme (E) are involved in a reversible, initial binding step (K1),followed by an irreversible, rate-limiting, bond-forming step (k) that results in the formation of a covalent,glucosyl-enzyme complex (E—I). If[II >> [El; i.e., {[I] — [E]} [I],then [I] is essentially constant over the course of the reaction, and the kinetics are pseudo first-order with respectto the enzyme concentration. The inactivation equation can then be written in Michaelis-Menten form as:= lq [E]0 [I] Equation (1)K1 + [1]where: = rate constant of inactivationK1 = the apparent dissociation constant for all species of enzyme-bound inactivatorK1 = [E] [I] Equation (2)E [El]154If [I] is constant, Equation (1) becomes:v = kobs Wit Equation (3)where:k — k1 [1] Equation (4)obs —K1 + [1]In those cases where a time-dependent, first-order decay in activity (after correction for the controlassay without the putative inactivator) was observed, the residual activity was plotted against time, andk0b5 Wascalculated at each inactivator concentration by fitting the initial rates (which measure WI) to Equation (5), usingthe computer program GraFit (Leatherbarrow, 1990).[El = WIoe_{(obst} Equation (5)The pseudo first-order rate constant (kobs) obtained at each concentration of the mactivator was fitted toEquation (4) above using the computer program GraFit (Leatherbarrow, 1990). This equation was used tocalculate the binding constant, K, and the inactivation rate constant, k. These data were also presented in theform of a double-reciprocal plot for convenient visual inspection.e. The reactivation testfor nitroglucal-inactivated pABG5.A sample (0.13 mg) of pABG5 was inactivated after incubation in a solution containing 16 mMnitroglucal. This solution was placed in dialysis tubing, and then excess inactivator was removed by dialysisusing three changes of a large volume of 50 mM phosphate buffer (pH 6.8).As a control, another sample (0.13 mg) of pABG5 went through the same procedure except that noinactivator was added. After the control sample was dialyzed, the retentate was divided into two equal-sizealiquots. To one control aliquot was added BSA (to a final concentration of 0.1%) in 50 mM phosphate buffer.To the second control aliquot was added B SA (to a final concentration of 0.1%) and cellobiose (to a final155concentration of 21 mM) in 50 mM phosphate buffer. The final volume of each of these control “reactivation”reactions was 0.110 mL.The “nitroglucal-free” retentate containing nitroglucal-inactivated pABG5 was divided into threeequal-size aliquots. To one aliquot was added BSA (to a final concentration of 0.1%) in 50 mM phosphatebuffer. To the second aliquot was added BSA (to a final concentration of 0.1%) and cellobiose (to a finalconcentration of 21 mM) in 50 mM phosphate buffer. To the third aliquot was added BSA (to a finalconcentration of 0.1%) and ldI3GlcØ (to a final concentration of 21 mM) in 50 mM phosphate buffer. Thevolume of each of these reactivation reactions was 0.110 mL.The five 0.1 10-mL reactions mentioned above were incubated at pH 6.8 at 37 °C. At various timeintervals, aliquots were removed from each reaction, transferred to cuvettes containing 1.00 mL of I3G1cPNP,and then assayed for enzymatic activity.5.2.5. The determination of kinetic parameters by IIPLC.a. Instrumentation.HPLC analyses were carried out using instrumentation from Waters®, including the HPLC apparatus,Model 712 WISP® auto-sampler (injector), Model 410 differential refractometer detector, Model 486 tunableabsorbance detector, and an analytical DextroPak® column (100 x 8 mm; operated using water as the eluent;used to separate anomers of GIcNAc). A GuardPak® pre-column was used to remove protein before thesample entered the Dextropak® column. A BioRad® Aminex® HPX-87H column was also used(300 x 7.8 mm; operated using 13 mM H2S04 as the eluant; used to separate G1cNAc and ManNAc). Data wascollected using the Baseline® 810 chromatography workstation, and analytes were identified by their retentiontime in comparison with authentic standards. Chromatographs from the workstation were exported as ASCIIdata files to the computer program GraFit (Leatherbarrow, 1990) for printing.b. Determination of the product ofNAGase-catalyzed /3G1cNAcPNP hydrolysis.The stereochemical course of the enzymatic hydrolysis of fGlcNAcPNP was determined by HPLCanalysis using a DextroPak® column. Assignment of peaks was achieved by loading a freshly prepared sample156of aGlcNAc onto the column and measuring its retention time. ‘H NMR analysis of a similar, freshly preparedsample of ccGlcNAc confirmed the identity of the sample, while similar analysis of an equilibrated mixture wasused to determine which anomer was the major component. HPLC analysis was carried out using samples of3GlcNAcPNP (in 3 mM citrate buffer without BSA) incubated for 7 mm at 25 °C with one of each of the threetypes of NAGase under study. Control reactions (without GlcNAcPNP or the enzyme) were also analyzed.The analyses of the enzymatic reactions were repeated using samples obtained after 1-3 days of incubation at25 °C.c. Determination of the rate ofNAGase-catalyzed reactions.The following procedures were used to determine kinetic parameters for NAGlucal hydration catalyzedby jack bean NAGase. Seven l.00-mL reaction buffer mixes containing NaCl (100 mM), citrate (5 mM, pH5.0), BSA (0.01%), and one of 7 different NAGlucal concentrations were prepared. Reactions (at 25 °C) werestarted by injecting 2 j.ig of jack bean NAGase into each solution. Reactions were incubated at 25 °C for 8 mm,then stopped by boiling for 30 sec (the latter step irreversibly denatured the enzyme, but did not result insignificant decomposition of other components of the reaction). Water was removed by lyophilization, and thenthe residue remaining in each reaction tube was resuspended with 0.100 mL of double-deionized water.Aliquots (0.080 mL) of each reaction were analyzed by HPLC using a BioRad® Aminex® HPX-87H column(300 x 7.8 mm) with 13 mMH2S04as the eluant, which separated GlcNAc and ManNAc. A G1cNAc standardcurve was prepared by following the above procedure for the enzymatic reactions (but without addition of theenzyme) using 8 different concentrations of G1cNAc standard. The concentration of the enzyme reactionproduct was determined by measuring the area of the GlcNAc peak, which was corrected by subtracting the areaof the G1cNAc peak in appropriate controls (reactions with the same concentration of NAGlucal but withoutenzyme), and then comparing the corrected peak area with the standard curve. Determinations ofK and Vmaxvalues were as described above.1575.3. PROTEINMASS SPECTROMETRY OFNITROGLUCAL-INACTIVATED pABG5.Ion-spray protein mass spectrometry was carried Out Ofl a PE-Sciex API III triple quacinipole instrument(Sciex, Thornhill, Ontario) in the laboratory of Prof. R. Aebersold at the Biomedical Research Centre of theUniversity of British Columbia. Spectra were collected in the LC-MS mode (single MS) by Dr. S. C. Miao.Prior to mass spectrometry the protein samples were centrifuged to remove insoluble matter. Low molecularmass compounds were removed by HPLC using a 1.00-mm microbore PLRP-S column (Michrom BioresourcesInc.). The following gradient was applied to elute the protein from the column: 20% solvent B in solvent A to100% solvent B over 10 mm, followed by 100% solvent B over 2 mm. The composition of solvent A was0.05% TFA, 2% acetonitrile in water. The composition of solvent B was 0.045% TFA, 80% acetonithle inwater.The experimental conditions for the preparation of nitroglucal-inactivated pABG5 (0.100 mg,2.5 mg/mL) were the same as those used in previous inactivation experiments (described above).APPENDIX I: SUPPLEMENTARY DATAFigure A-I.1. Determination of Km and Vmax for G1cNAcPNP hydrolysis by K-NAGase and H-NAGase.158Reactions were performed at pH 4.25 and 25 °C in 50 mM citrate buffer containing 0.1% BSA, 100 mM NaC1,and f3-N-acetythexosaminidase and I3G1cNAcPNP as indicated below.(a) K-NAGase (2.0 j.tg/mL). Reactions contained 0.25, 0.34, 0.49, 0.79, 1.38, or 1.96 mM GlcNAcPNP.(b) H-NAGase (4.2 .tgImL). Reactions contained 0.20, 0.25, 0.34, 0.49,0.79, 1.38, or 1.96 mM I3G1cNAcPNP.I I I I I0.250.20.150.1noEI I I I i I I i(a)(b)0.0500.60.40.200 2 4[I3G1cNAcPNPJ-1 (mM)1LZV/>i:Z-2 0 2 4 6[I3G1cNAcPNPJ-l (mM)’-10 0 10 20 30 40 50[NAGlucal] (jiM)Figure A-I.2. Estimation of K values for the inhibition of K-NAGase and H-NAGase by NAGlucal.Reactions were perfonned at pH 4.25 and 25 °C in 50mM citrate buffer containing 0.1% BSA, 100 mM NaCI,and 3-N-acetythexosaminidase, fGlcNAcPNP, and NAGlucal as indicated below.(a) RFK determination for the inhibition of K-NAGase (2.0 p.g/mL). The concentration of j3G1cNAcPNP in thereactions was 0.98 mM. Reactions contained 0, 8.2, 16, 58, or 82 I.iM NAGlucal.(a) RFK1 determination for the inhibition of H-NAGase (4.2 .tgJmL). The concentration of I3G1cNAcPNP in thereactions was 0.79 mM. Reactions contained 0, 3.3, 8.2, 16, 25, 33, or 41 pM NAGlucal.II III I 1111111111111111iii ii I 11111 ilti itii i IiECC,.,EI6040200806040200159(a)Imax(b)=Vmax-20 0 20 40 60 80[NAG1ucall (jiM)I ‘ I I I I I II I I I I I i I I I iI[2-cyanoglucal] (mM)Figure A-L3. Estimation of the K1 for the inhibition of J-NAGase by 2-cyanoglucal.160-, Vma2520151050I-60 -40 -20 0 20 40Reactions were performed at pH 5.0 and 25 °C in 50 mM citrate buffer containing 0.1% BSA and 1.0 p.gImL of3-N-acetythexosaminidase from jack beans. The concentration of f3G1cNAcPNP in the reactions was 0.58 mM.The concentrations of 2-cyanoglucal in the reactions were 0, 4.1, 8.2, 33, and 41 mM.161APPENDIX U: SIMPLE THEORETICAL TREATMENT OF ENZYME CATALYSIS.A-lI. 1. ENZYME CATALYSIS IN THEABSENCE OF INHIBITION.A general theory of enzyme action and kinetics was developed by Michaelis & Menten (1913), whichconfirmed the earlier work of Henri (1902) (see Segel, 1975). Later, a more general treatment was given byBriggs & Haldane (1925), who introduced the idea of the steady-state. In the absence of inhibition, a simplereaction with one substrate and one enzyme (in this case a protein that acts as a catalyst) can be expressed asshown below, where the free enzyme (E) first combines with the substrate (S) to form one distinctenzyme-substrate complex (ES), which may then be converted to the free enzyme and one product (P).E+S ES E+Pk_1The other assumptions (in addition to those described above) underlying the Henri-Michaelis-Mententreatment of enzyme catalysis are listed below (Segel, 1975):(i) The concentration of the substrate is much greater than that of the enzyme, thus the formation of theenzyme-substrate complex does not significantly alter the concentration of the substrate.(ii) The enzyme and the substrate react rapidly to form the enzyme-substrate complex.(lii) The enzyme, substrate, and enzyme-substrate complex establish a rapid equilibrium, thus the rateat which the enzyme-substrate complex dissociates to form the free enzyme and the substrate (k i)greatly exceeds the rate of conversion of the enzyme-substrate complex to form the free enzymeand the product (k2).(iv) The rate-limiting step in the reaction is therefore the conversion of the enzyme-substrate complexto form the free enzyme and the product.(v) Only the initial velocity of the reaction is considered, thus the reverse reaction (from the freeenzyme and the product) to form the enzyme-substrate complex is ignored.Under steady-state conditions (Briggs & Haldane, 1925):d [ES]dt= k1[E][S] — k_1[ES] — k2[ES] 0 (1)k1[EJ[S] = k_1[ES] +k2[ES] (2)162The total concentration of the enzyme, [El0, is the sum of the concentrations of free andsubstrate-bound enzyme species.{E]0 = [El + [ESJ (3)Solving for [ES] using Equations 2 and 3, one obtains:k1[E]0[5][ES] = (4)k_j +k2 + k1[S]The initial velocity (v) of the reaction is equal to the initial rate of formation of the product:d [P]= dt= k2[ES] (5)Substituting Equation 4 into 5, one obtains:k21[E]0Sk_1 +k2 + k1[SJwhich can be expressed as the Michaelis-Menten (or Henri-Michaelis-Menten) equation:[E]0SJK+[S](6)Some kinetic parameters derived from Equation 6 are given below:Catalytic constant: = The first-order rate constant for the conversion ofthe enzyme-substrate complex to form the product.k_1 + l2 When k....1 >> k2, K K = k_1/k, theMichaelis constant: Km =k1 dissociation constant of the ES.k12 A second-order constant that relates the reactionSpecificity constant: = rate to the concentration of the free enzyme, [E],Km k_1 + k2 rather than that of the total enzyme, [E]0.{E]0[SNVVmax[S1Km163IVmax = [E]0VmaxVm20 KmSubstrate concentration [SIFigure A-II.1. Michaelis-Menten kinetics of an enzyme-catalyzed reaction.The effect of substrate concentration on the rate of the enzyme-catalyzed reaction is shown inFig. A-II.1. At low concentrations of substrate, where [SI <<Km, the initial rate of the reaction is proportionalto the substrate concentration:Km(7)In contrast, at saturating concentrations of the substrate, where [SI >> Km, the initial rate of the reactionbecomes independent of the substrate concentration, and approaches a constant maximum rate, V:V = kcat [E]0 = Vmax (8)It also follows from the above that when the initial rate of the reaction is equal to one-half of themaximal velocity (v = VmI2), the substrate concentration is equal to the Michaelis Constailt, Km. The value ofKm gives a measure of the stability of the enzyme-substrate complex. An enzyme has a high affinity for asubstrate with a low Km.164The Henri-Michaelis-Menten equation is often transformed into a linear form for analyzing datagraphically and detecting deviations from ideal behaviour. One of the most commonly used transformations isthe double-reciprocal form introduced by Lineweaver & Burk (1934). This type of plot is shown in Fig. A-II.2,and the equation used to plot the data is obtained by simply taking the reciprocal of both sides of Equation 6:—1X-intercept = —KmL__+ Km (iV Vmax [SIFigure A-U.2. Double-reciprocal (or Lineweaver-Burk) plot of the Henri-Michaeis-Menten equation.A-II.2. THE INHIBITION OF ENZYME CATALYSIS.A-ll.2.1. Irreversible inhibition.Irreversible inhibition (or inactivation) has already been dealt with in Section 5.2.4 (d) of Chapter 5.I KmSlope = Vmax1Vmax0(Substrate concentration) —l [SI165A-J1.2.2. Reversible inhibition.a. The three types of reversible inhibition.There are three major types of reversible inhibition: competitive, noncompetitive, and uncompetitive.In each case the effects of these types of inhibition can be analyzed using the simple theoretical treatmentdiscussed above.b. Competitive inhibition.A competitive inhibitor competes with the normal substrate for binding in the active site of the freeenzyme. The binding of either the competitive inhibitor or the substrate in the active site is mutually exclusive.The reactions describing competitive inhibition are shown below:E÷S ES+IThe substrate constant (Ks) is the dissociation constant for the enzyme-substrate complex, and theinhibition constant (K1) is the dissociation constant for the enzyme-inhibitor complex. The catalytic constant(keat) is the rate constant for the conversion of the enzyme-substrate complex to form the free enzyme and theproduct.[El0 = [E] + [ES] + [El] (9)Using Equation 9, the steady-state assumption (d [ESJId t = 0), and the expression v = k2[ES], onecan derive the rate equation for competitive inhibition:. E+PKi![El[Sl [E1[I1’K K[ElI[ES]166[Sj/ [I]\ (10)Kmtl + — ) + [SJK1)A competitive inhibitor increases the apparent Km by a factor of (1 + [I]IKj). With a competitiveinhibitor the factor (1 + [I]/K) is a function of [I] that reflects the statistical distribution of the enzyme betweenthe E and El forms. The value of Km,app increases as [I] increases because the formation of the El removessome of the free enzyme, which drives the first step of the reaction to the left. The value of Vmax does notchange, but the substrate concentration necessary to achieve any fraction of Vmax increases as a result of thepresence of the competitive inhibitor.Four parameters affect the degree of competitive inhibition observed: [SI, [I], Km, andK1. The degreeof inhibition decreases as [SI increases and [I] remains constant, whereas the degree of inhibition increases as [IIincreases and [SI remains constant. The degree of inhibition at any given [SI and [II is larger with smallervalues of K1. When [I] = K1, the slope of the double-reciprocal plot (1/v vs. 1I[SI) is twice that of the uninhibitedreaction, and the value ofKm,p is2Km.C. Noncompetitive inhibition.A noncompetitive inhibitor can bind to the free enzyme or the ES. In both cases the inhibitor binds at asite other than the active site, thus the binding of a noncompetitive inhibitor and the substrate are independentevents. The reactions describing noncompetitive inhibition are shown below:KE+S ES E+P+ +I I____________IEI[SI [EI][SI=[ES] [ESI]K1 K[E][I] [ESI[IIK K1 =___________[El] [ESI]EI+S -.. ESI___ ___167[E]0E]+[ S]+[EI]+[ESI] (11)Using Equation 11, the steady-state assumption (d [ES]/d t = 0), and the expression v = k2[ES], onecan derive the rate equation for noncompetitive inhibition:[S](12)A noncompetitive inhibitor does not affect the value of Km because at any [I] both of the substrate-binding forms of the enzyme (E and El) have equal affmities for the substrate. The inhibitor decreases the valueof the apparent Vmax by a factor of (1 + [Il/K1). With a noncompetitive inhibitor the factor (1 + [I]IKj) is afunction of [I] that reflects the statistical distribution of the enzyme-substrate complexes between the ES and ESIforms. In the presence of a noncompetitive inhibitor, the steady-state concentration of the ES is decreased at all[Si, and can never approach [E]0, no matter how high [5] may be. However, the value of keat is unchanged. Theoverall effect of a noncompetitive inhibitor is to reduce the effective concentration of the enzyme.With a noncompetitive inhibitor the degree of inhibition only depends on [I] and K. The degree ofinhibition increases as [I] increases. For a given [I], the degree of inhibition is constant, regardless of the valuesof [5] or Km. The degree of inhibition at any given [I] is larger with smaller values ofK1. The [I] that causes50% inhibition (at all [Si) equals K1.d Uncompetitive inhibition.An uncompetitive inhibitor can only bind to the enzyme-substrate complex. The inhibitor is unable tobind to the free enzyme. This type of inhibition is rarely observed in one-substrate reactions, but is common insteady-state multireactant systems. The reactions describing uncompetitive inhibition are shown below:Km + [Si168K kcatE+S - ES - E+P+I_________________________[E][S]KK[ES][ES] [I]K1[ESI]ESI[E]0 = [E] + [ES] + [ESI] (13)Using Equation 13, the steady-state assumption (d {ES]Id t = 0), and the expression v = k2[ES], onecan derive the rate equation for uncompetitive inhibition:[5](14)+ [SIAn uncompetitive inhibitor decreases the values of both the apparent Vmax and the apparent Km by afactor of (1 + [I]IK). With an uncompetitive inhibitor the factor (1 + [I]IK) is a function of [I] that reflects thestatistical distribution of the enzyme-substrate complexes between the ES and ESI forms. In the presence of anuncompetitive inhibitor the value of keat is unchanged, but the steady-state concentration of the ES is decreasedat all [S], and can never approach [El0, no matter how high [5] may be. The value of Km app decreases as [I]increases because the formation of the ESI removes some of the ES, which drives the first step of the reaction tothe right. Thus an uncompetitive inhibitor can be viewed as an activator with respect to Km (as Km,app < Km),and at low [SI (i.e., first-order kinetics), the effects of the inhibitor on Km and Vmax essentially cancel, and littleif any inhibition is observed.169Four parameters affect the degree of uncompetitive inhibition observed: [SI, [I], Km, and K1. Thedegree of inhibition increases as [I] increases and [SI remains constant. However, in contrast to the case ofcompetitive inhibition, with uncompetitive inhibition the degree of inhibition increases as [SI increases and [I]remains constant. This is because the [ES] increases as the [5] increases, and an uncompetitive inhibitor canonly bind to the ES. For the same reason the degree of uncompetitive inhibition is inversely related to the valueof Km, again in contrast to the case of competitive inhibition. The degree of inhibition at any given [SI and [I] islarger with smaller values ofK.e. Graphical methodsfor distinguishing different types of reversible inhibition.The three types of reversible inhibition discussed above can be identified and distinguished bygraphical methods. The two most commonly used graphical methods are double-reciprocal plots and Dixonplots. These are shown in Fig. A-II.3, along with the appropriate forms of the rate equations used to plot thedata.In this very brief Appendix, only the three “pure” types of reversible inhibition have been discussed.Table A-II.1 provides a summary of some of the kinetic parameters that may be obtained for these three cases.Intermediate (or mixed) inhibition behaviour may also occur. For a thorough treatment of these cases the workof Segel (1975) is highly recommended.Figure A-ll.3. Some graphical methods for distinguishing different types of reversible inhibition.This figure appears on the following page. Double-reciprocal plots (1) appear on the left, and Dixon plots (2)appear on the right. The appropriate form of the rate equation appears at the bottom of each panel.(C) Competitive inhibition; (N) Noncompetitive inhibition; (U) Uncompetitive inhibition.170c.1I C.2jy-intercept =/ Km\ 1—1 0Decreasing[Si1= x-intercept [SIKm (1+1 Kmf [‘I’1 I+N.1m1Vx-intercept = 0_K(1+--)1 / Km ‘\ / Km\1VVXK[Sl)rn +Increasing[II/N.2y-intercept/ Enil+ —/vx /1Vy-intercept =/ Km\ 10—1 —— x-interceptDecreasing/::Z)cI[Sjx-intercept = —K1 Km / [II\i / [II\ 1+0U.1EnIV= [(‘+)]i + (i+)__VmKiIncreasing[IIU.2y-intercept/ [Ii=____1Vy-intercept =/ Km\ 1Decreasing[SI/ [I1\0—11+—i IS\ Kj j = x-interceptKmI /K\i / [‘I\ I+0/ Km\—K,çl+ = x-lntercept[1]1 /1 \ / Km\1;-= +Table A-IT.1. 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