Design and Synthesis of Chemical Tools for Studies of Carbohydrate Active Enzymes

DESIGN AND SYNTHESIS OF CHEMICAL

TOOLS FOR STUDIES OF CARBOHYDRATE

ACTIVE ENZYMES

Marija Petricevic

Submitted in total fulfilment of the requirements of the degree of Doctor of Philosophy

October 2018

School of Chemistry

The University of Melbourne

ABSTRACT

Glycoside hydrolases (GH) are enzymes that catalyse the hydrolysis of glycosidic linkages. They are classified into over 150 families based on their primary amino acid sequence. Family GH99 endo-α-1,2-mannosidases/endo-α-1,2-mannases cleave α-

Glc/Man-1,3-α-Man-OR structures within mammalian N-linked glycans and fungal α- mannans. They are predicted to perform base-catalysed hydrolysis via an 1,2-anhydro sugar intermediate. The first part of this thesis reports the synthesis of a mechanism- inspired inhibitor, α-mannosyl-1,3-noeuromycin (ManNOE). ManNOE was found to be the most potent GH99 inhibitor to date. The success of ManNOE was attributed to a favourable interaction between the 2-OH on the NOE ring and active site residue E333, predicted to be important in mechanism. Also described is the synthesis of the inhibitor

α-mannosyl-1,3-(2-amino)deoxymannojirimycin (Man2NH2DMJ). Modest affinities were observed for Man2NH2DMJ. Structural studies revealed it binds in the same way as other iminosugar inhibitors, suggesting poor inhibition is not due to failure of the 2-NH2 to bind to reside E333, but rather a reduction in basicity of the endocyclic nitrogen in the presence 2-NH2 protonation.

The second part of this thesis examines a novel family GH134 β-1,4-mannanases.

A GH family 134 endo-β-1,4-mannanase from a Streptomyces sp. was found to possess a fold closely related to that of hen egg white lysozyme (HEWL) and to act with inversion of stereochemistry. X-ray crystallography and ab initio quantum mechanics

(QM)/molecular mechanics (MM) metadynamics reveal this enzyme utilizes a unique

1 3 ‡ 3 C4→ H4 → H1 conformational itinerary along the reaction co-ordinate, different to any known β-1,4-mannanases.

The third part of this thesis examines sulfoglycolysis. Sulfoquinovosyl glycerol

(SQGro) was synthesised as a mixture of diastereomers. A family GH31 sulfoquinovosidase (YihQ) isolated Escherichia coli (E. coli) was determined to have a

6-fold preference for the naturally occurring isomer (2’R-SQGro) but could cleave both isomers. SQGro supports growth of E. coli to find cell densities comparable to growth on glucose.

Preliminary studies revealed that the plant pathogen Agrobacterium tumefaciens performs sulfoglycolysis when grown on SQ as the sole carbon source. Growth of cultures and consumption of SQ were directly correlated to release sulfite into the media, which over time autooxidised to sulfate. Proteomics enabled the discovery of the sulfoglycolysis gene cluster in A. tumefaciens which enabled prediction of a novel sulfoglycolytic pathway.

DECLARATION

This is to certify that:

i. the thesis comprises only my original work towards the PhD except where indicated in the Preface,

ii. due acknowledgement has been made in the text to all other material used,

iii. the thesis is less than 100 000 words in length, exclusive of tables, bibliographies and appendices.

Marija Petricevic

October 2018

iii

PREFACE

All the work reported herein has been conducted by the candidate, except where indicated.

The inhibitors α-D-mannopyranosyl-1,3-(1,2-dideoxy)mannose (ManddMan), α-D- mannopyranosyl-1,3-glucal (ManGlucal) and α-D-mannopyranosyl-1,3-mannoimidazole

(ManManIm) in Chapters 2 and 3 were synthesised by Pearl Fernandes in the laboratory of Prof Spencer Williams (University of Melbourne). In Chapter 5 synthesis of (S)-2,3- dihydroxypropane-1-sulfonate (DHPS) was synthesised by Janice Mui under the guidance and supervision of the candidate. X-ray crystallographic studies and isothermal titration calorimetry with B. thetaiotaomicron, B. xylanisolvens GH99 enzymes as described in Chapters 2 and 3 were conducted by Lukasz Sobala in the laboratory of Prof. Gideon

Davies (University of York). Structural analysis of SsGH134 as described in Chapter 4 was performed by Dr Yi Jin in the laboratory of Prof. Gideon Davies (University of York).

Dissociation constants for the binding of ManddMan and ManGlucal to BtGH99 and

BxGH99 were determined by 2D NMR in the laboratory of Prof Jesus Jiminez-Barbaro

(Ikerbasque Basque Foundation for Science, Spain.) Classical molecular dynamics (MD) and QM/MM metadynamics described in Chapters 2 and 4 were conducted in the laboratory of Prof Carme Rovira with the assistance of Lluís Raich (Universitat de Bercelona, Spain).

Expression plasmids in Chapter 4 were assembled by Alan John in the laboratory of Dr

Ethan Goddard Borger (Walter and Eliza Hall Institute). Expression of proteins in Chapter

4 was conducted by the candidate with assistance of Alan John in the laboratory of Dr Ethan

Goddard Borger. YihQ and AtSQase proteins in Chapters 5 and 6 were prepared by James

Lingford in Dr Ethan Goddard-Borger's laboratory. Quantitative analysis of DHPS was performed with the assistance of Dr Eileen Ryan (University of Melbourne). A. tumefaciens C58 was a gift from Dr Monica Doblin (School of Botany, University of

Melbourne). Proteomics in Chapter 6 was performed in collaboration with Dr Nicholas

Scott (University of Melbourne).

The work discussed in this thesis has been published in part:

1) ACS Central Science, 2018, 4, 1266-1273.

Structural and Biochemical Insights into the Function and Evolution of

Sulfoquinovosidases.

2) Chemistry - A European Journal, 2018, 29, 7464–7473.

Exploration of strategies for mechanism-based inhibitor design for family GH99 endo-α-

1,2-mannanases.

3) Journal of the American Chemical Society, 2017, 139, 1089–1097.

Contribution of Shape and Charge to the Inhibition of a Family GH99 endo-α-1,2-

Mannanase.

4) ACS Central Science. 2016, 2, 896–903.

A β-Mannanase with a Lysozyme-like Fold and a Novel Molecular Catalytic Mechanism.

Additional work, not included here, performed by the candidate:

1) Chem. Comm. 2016, 52, 11096-11099.

Structural and mechanistic insights into a Bacteroides vulgatus retaining N-acetyl-β- galactosaminidase that uses neighbouring group participation.

ACKNOWLEDGEMENTS

First and foremost, I would like to thank my supervisor Professor Spencer J. Williams.

His enthusiasm for science is contagious and highly motivational. I cannot thank him enough for his invaluable support and guidance over the past few years.

Thank you to Dr Ethan Goddard-Borger for introducing me to the world of biochemistry and extending my love of science in new directions.

I gratefully acknowledge the funding sources that made this research possible. My PhD was funded by The Melbourne Research Scholarship and the Norma Hilda Schuster

Scholarship.

Thank you to all my colleagues for your friendship and help in the lab. You have made my PhD experience so much more enjoyable and memorable.

I would like to thank my parents and sister, although they have no idea what I’ve been studying, their love and support throughout my many years of university has been limitless.

A big thank you to my partner Charlie, who has been a constant source of moral support.

Thanks for listening to me vent about all those failed experiments and accompanying me to check bacterial cultures at 3 am. I really appreciate it!

TABLE OF CONTENTS

ABSTRACT ...... i

DECLARATION ...... iii

PREFACE ...... iv

ACKNOWLEDGEMENTS ...... vi

TABLE OF CONTENTS ...... vii

ABBREVIATIONS ...... x

CHAPTER ONE: Introduction ...... 1

1.1 Introduction to glycoside hydrolases ...... 2

1.2 Mechanisms of glycoside hydrolases ...... 3

1.3 Mechanism of hen egg white lysosome (HEWL) mechanism ...... 4

1.4 Mechanism of goose egg white lysosome (GEWL) ...... 5

1.5 Neighbouring group participation (NGP) mechanism...... 6

1.6 Anti/Syn lateral protonation ...... 8

1.7 Transition states of glycosidases ...... 9

1.8 Substrate distortion during enzymatic hydrolysis of glycosidases ...... 12

1.9 Reaction co-ordinate ...... 13

1.10 Scope of thesis ...... 20

CHAPTER TWO: Investigation of the effects of shape and charge on family GH99 endo-α-1,2-mannanases ...... 22

2.1 N-Linked glycan biosynthesis in the secretory pathway ...... 23

vii

2.2 Discovery of endo-α-1,2-mannosidase, a unique endo-acting glycoside hydrolase

...... 26

2.3 N-Linked glycan degradation by bacterial endo-α-1,2-mannanases ...... 29

2.4 Inhibitors of GH family 99 enzymes ...... 31

2.5 Aims ...... 33

2.6 Results and Discussion ...... 34

2.7 Conclusions ...... 54

2.8 Experimental ...... 56

CHAPTER THREE: Exploration of strategies for charge mimicry in inhibitor design for family GH99 endo-α-1,2-mannanases ...... 65

3.1 Importance of charge mimicry for binding to GH99 enzymes ...... 66

3.2 Aims ...... 66

3.3 Prior work with DMJ and NH2DMJ ...... 67

3.4 Synthetic strategy towards Man2NH2DMJ ...... 68

3.5 Results and Discussion ...... 69

3.6 Conclusions ...... 85

3.7 Experimental ...... 87

CHAPTER FOUR: GH134 a β-mannanase with a lysozyme-like fold and a novel molecular mechanism ...... 95

4.1 Use of β-mannanases in the biofuels industry ...... 96

4.2 β-mannanase and β-mannosidase GH families...... 97

4.3 Discovery of a novel β-1,4-mannanase and new GH family ...... 98

viii

4.4 Aims ...... 99

4.5 Results and Discussion ...... 100

4.6 Conclusions ...... 115

4.7 Experimental ...... 116

CHAPTER FIVE: Investigation of sulfoglycolysis in Escherichia coli ...... 119

5.1 Plant sulfolipid SQDG its role in the biosphere ...... 120

5.2 Occurrence and biosynthesis of SQDG ...... 121

5.3 Degradation of SQDG ...... 122

5.4 Catabolism of SQ in E. coli ...... 127

5.5 Aims ...... 129

5.6 Results and Discussion ...... 130

5.7 Conclusions ...... 141

5.8 Experimental ...... 143

CHAPTER SIX: Discovery of sulfoglycolysis in Agrobacterium tumefaciens ...... 146

6.1 Biomineralization of SQ ...... 147

6.2 Aims ...... 148

6.3 Results and Discussion ...... 148

6.4. Conclusion: The proposed sulfoglycolysis pathway of A. tumefaciens ...... 160

6.5 Experimental ...... 162

CHAPTER SEVEN: Summary and future work ...... 165

References ...... 174

ABBREVIATIONS

2D-PAGE two-dimensional polyacrylamide gel electrophoresis

Ab Alteromonadaceae

ABC ATP-binding cassette

ATP adenosine triphosphate

A. tumefaciens Agrobacterium Tumefaciens

B boat

BCD bromoconduritol

Bt Bacteroides thetaiotaomicron

Bx Bacteroides xylanisolvens

C chair

CAZy carbohydrate active enzymes

CNX calnexin

CRT calreticulin

CST castanospermine

CV collective variable

DAG diacylglycerol

DHPS (S)-2,3-dihydroxypropane-1-sulfonate

DMJ deoxymannojirimycin

DNJ deoxynojirimycin

DTT dithiothreitol

E envelope

E.coli Escherichia coli

ED Entner-Doudoroff

EDTA ethylenediaminetetraacetic acid

EM endo-α-1,2-mannosidase

EMP Embden-Meyerhof-Parnas

ER endoplasmic reticulum

ERAD endoplasmic reticulum associated degradation

ESI-MS electrospray ionisation mass spectrometry

FLA flavin mononucleotide

FOS oligosaccharide

GEWL goose egg white lysosome

GH glycoside hydrolase

Glc glucose

Glc1P glucose-1-phospate

GlcNAc N-acetylglucosamine

GlcNAcTI N-acetylglucosamine transferase

H half chair h hour(s)

HEWL hen egg white lysosome

Hs Herbaspirillium seropedicae

His histadine

IFG isofagomine

IPTG isopropyl ß-D-1-thiogalactopyranoside

kcat turnover number

KD dissociation constant

KIE kinetic isotope effect

KFN kifunensine

Km Michaelis constant

LC-MS liquid chromatography-mass spectrometry lyso-SQGro lyso-sulfoquinovosylglycerol min minute(s)

M2 mannobiose

M3 mannotriose

M4 mannotetraose

M5 mannopentaose

M6 mannohexaose

Mal maltose

Man/M1 mannose

xii

MANA mannanase A

MBP maltose-binding protein

MD molecular dynamics

NBD nucleotide-binding domain

NMR nuclear magnetic resonance

NGP neighbouring group participation

NOE noeuromycin

OD optical density

PA pyridylamino

PNPSQ 4-nitrophenyl-α-D-sulfoquinovoside ppm parts per million

P. putida Pseudomonas putida

QM/MM quantum mechanics/molecular mechanics

Rm Rhizopus microspores r.t. room temperature s second(s)

Sama Shewanella amazonensis

SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis

SL 3-sulfolactate

SLA 3-sulfolactaldehyde

xiii

SQ α-D-sulfoquinovose

SQase sulfoquinovosidase

SQD1 UDP-SQ synthases

SQD2 UDP-SQ:DAG sulfoquinovosyltransferase

SQDG α-D-sulfoquinovosyl diacylglycerol

SQGro sulfoquinovosylglycerol

SQMe methyl α-D-sulfoquinovose

Ss Streptomyces sp.

SW swainsonine

TCA trichloroacetimidate

Thr/T threonine

TLC thin layer chromatography

TM transmembrane

TS transition state

UDP uridine diphosphate

UDP3 UDP-glucose pyrophosphorylase3

UGT1 glycoprotein glucosyltransferase

UTP uridine triphosphate

xiv

CHAPTER ONE:

Introduction

1.1 Introduction to glycoside hydrolases

Carbohydrates are the most abundant class of biomolecule on earth. Carbohydrates exist in a diverse range of structures with the simplest building block termed a monosaccharide.1 A glycosidic linkage can be formed between one monosaccharide and the hydroxyl group of another, or with non-carbohydrate moieties. This leads to a variety of polysaccharides and glycoconjugates such as glycoproteins and glycolipids.

Collectively, this diverse group of glycoconjugates are involved in metabolic processes, cellular and protein recognition, the provision of structural frameworks, posttranslational modifications, immune responses and viral infection. Indeed, it is difficult to find a biological process in which a carbohydrate does not take part.2

A broad suite of carbohydrate-active enzymes catalyse reactions leading to the synthesis and breakdown of polysaccharides and glycoconjugates. They can be classified into groups including: glycoside hydrolases, glycosyltransferases, polysaccharide lyases, and lytic polysaccharide monooxygenases. The focus of this thesis is the glycoside hydrolases (glycosidases). Glycosidases catalyse the hydrolysis of acetal linkages of glycosides through the scission of the exocyclic C–O bond. Glycosidases are named with respect to the sugar unit and stereochemistry of the bond they cleave. For example, α- mannosidases cleave bond between α-linked D-mannose residues. Glycosidases are also usefully categorized into families based on their primary sequence similarity. As form defines function, the close sequence relationship of members of the same family generally means that they share similarities in their chemical mechanisms and stereochemical outcomes through the involvement of conserved enzymatic residues.3 There are over 150 families of glycosidases (www.cazy.org;www.cazypedia.org),4 which are denoted

GHXXX.

1.2 Mechanisms of glycoside hydrolases

Glycosidases can be classified as exo- or endo-acting depending on whether they cleave sugars from the end of the chain or internally (Figure 1.1a.).5 Structural studies have revealed that glycosidases possess binding subsites for one or more sugar residues. The subsites are numbered with respect to the site of hydrolysis. Negative integers are used to count away from the site of cleavage towards the non-reducing end and increasing positive integers are used toward the reducing end.6 Furthermore, glycosidases can be classified as either retaining or inverting depending on the stereochemical outcome of the product relative to the substrate (Figure 1.1b, c.).

Figure 1.1: Classification of glycosidases. a. exo- and endo-acting, showing subsite numbering; b. retaining glycosidase; c. inverting glycosidase.

Koshland retaining and inverting mechanisms

The classical retaining and inverting mechanisms for glycosidase hydrolases were first proposed by Koshland in the 1950s.7 Koshland speculated that retaining enzymes should proceed through a two-step double-displacement mechanism involving the formation of a glycosyl enzyme intermediate, or a cyclic intermediate formed by neighbouring group participation (NGP). On the other hand, he suggested that inverting enzymes would

proceed through a single-step substitution mechanism. Logically, the retaining mechanism involving a glycosyl enzyme intermediate, and the inverting mechanism should both require two participating catalytic residues: either a nucleophile and a general acid, or a general acid and general base, respectively. In the case of the neighbouring group participation mechanism a substrate-derived nucleophile positioned adjacent to the anomeric position is employed, along with a general acid residue. Koshland did not speculate on the identities of the enzymatic residues or nucleophilic group of the substrate.7

1.3 Mechanism of hen egg white lysosome (HEWL) mechanism

Hen egg white lysozyme (HEWL) is a retaining glycosidase belonging to GH family 22 and was the first enzyme to have its three-dimensional structure solved.8 HEWL is an example of a retaining glycosidase whose mechanism proceeds though a glycosyl enzyme intermediate (Figure 1.2). The catalytic residues were identified to be the amino acids aspartate (D) 52 and glutamate (E) 35, located 5.5 Å apart. E35 acts as a general acid/base residue while D52 acts as a nucleophile. In the first step nucleophilic attack by D52, with simultaneous protonation of the aglycon by E35, forms a glycosyl-enzyme intermediate with inversion of anomeric stereochemistry. In the second step, E35 acts as a base and deprotonates a water molecule, which substitutes the glycosyl enzyme intermediate, causing a second inversion and leading to an overall net retention.9 Most retaining glycosidases that use an enzymic nucleophile have a carboxylate residue fulfil this role; one exception includes the retaining sialidases of family GH33, which use a tyrosine residue.10

Figure 1.2: Mechanism of HEWL, a retaining glycoside hydrolase.

1.4 Mechanism of goose egg white lysosome (GEWL)

Inverting glycosidases proceed through a single displacement mechanism employing two carboxyl residues that are typically 9-10 Å apart. A notable example of a presumed inverting enzyme is the Emden goose egg white lysosome (GEWL) of family GH23

(Figure 1.3).11 E73 acts as a general base and deprotonates a water molecule, assisting nucleophilic attack on the anomeric position. D97 protonates the glycosidic oxygen to assist its departure, resulting in a hemiacetal with inverted anomeric stereochemistry.12

Figure 1.3: Mechanism of GEWL, an inverting glycoside hydrolase.

1.5 Neighbouring group participation (NGP) mechanism

Retaining glycosidases that act though a substrate-assisted neighbouring group participation mechanism (NGP) employ two catalytic acid/base residues borne on the enzyme, while the nucleophile is a group present on the substrate (Figure 1.4). The hexosaminidases of families GH18, 20, 25, 56, 84, 85 and 123 are retaining enzymes that act though NGP.13–18 They contain an N-acetyl (or in some cases an N-glycolyl) group at the C2-position, of which the carbonyl group acts as the nucleophile. In the first step, an enzymatic base orients and deprotonates the amido group at C2 promoting nucleophilic attack by the amide oxygen at the anomeric center, leading to the formation of an oxazoline (or oxazolinium ion) intermediate and departure of the leaving group, assisted by a general acid residue.19 In the second step the former acid residue acts as a base to deprotonate a water molecule, aiding substitution of the oxazoline intermediate, resulting in cleavage of the glycosidic linkage with net retention of stereochemistry. The former base meanwhile acts as an acid to reprotonate at nitrogen.

Figure 1.4: Mechanism of a retaining glycoside hydrolase acting through neighbouring group participation.

X-ray structures of retaining GHs in complex with the inhibitor isofagomine (IFG) in the active site typically have a nucleophile located <3 Å. Family GH99 mannosidases act with retention of stereochemistry, yet X-ray structures revealed that OE1 atom of

E333 lies 3.5 Å away and the OH of Y46 and Y252 4.0 Å away. This suggests that a classical retaining mechanism or a typical NGP mechanism cannot occur for this family.20

One unexpected feature of the X-ray structures was the observation of a conserved residue

(E333) that is located 2.6 Å away from the axial O2 group of the substrate, mutation of which resulted in a severely disabled catalyst. This led to the proposal of a second, unprecedented mechanism that invokes neighbouring group participation by the substrate

2-OH group leading to a 1,2-anhydro (epoxide) intermediate (Figure 1.5). In the first step

E333 deprotonates the 2-OH group to promote nucleophilic attack on C1 driving formation of an epoxide with simultaneous protonation and departure of the aglycon assisted by E336. In the second step E336 deprotonates a water molecule which drives nucleophilic attack at the anomeric centre and opening the epoxide; E333 serves as an acid to protonate O2.20

Figure 1.5: Proposed neighbouring group participation mechanism for GH99 enzymes.

1.6 Anti/Syn lateral protonation

X-ray crystallographic structures of glycosidases with ligands have revealed that the general acid is often located laterally to the pyranose ring. This has led to the classification of glycosidases based on the position of the acid residue. Anti-protonators have their acid residues positioned in front of the endocyclic oxygen – this has been argued to correspond to the position of the antibonding orbital on the glycosidic oxygen atom when the aglycon is in the orientation favoured by the exo-anomeric effect. Syn-protonators have their acid residues positioned on the same side as the endocyclic oxygen in a conformation not favoured by the exo-anomeric effect. Protonation of the glycosidic oxygen atom occurs as the oxocarbenium ion begins to form and results in a lengthening of the fissile C–O bond in a pseudoaxial direction (Figure 1.6). It has been argued that if the carboxyl group of the acid residue was located above or below the plane of the sugar ring, departure of

the aglycon would be hindered.21 A pseudoaxial orientation of the glycosidic bond has been observed in a large number of X-ray structures of glycosidases with ligands bound, including that of cellulases from Fusarium oxysporum22 and Bacillus agaradhaerans.23

Figure 1.6: Anti and syn ‘lateral’ protonation of a β-glycosidase.

1.7 Transition states of glycosidases

For hydrolysis of a glycosidic bond to occur at biological pH values on timescales relevant to life requires catalysis. In 1998 Wolfenden reported for the first time the rate of uncatalyzed hydrolysis of a glycoside.24 This work revealed that the uncatalyzed hydrolysis of methyl α-D-glucopyranoside occurs with a first order rate constant of 10-15 s-1. By comparison, the rate constant of sweet potato β-amalyse was reported to be 1360 s-1. Thus the uncatalyzed reaction is 1.4  10-17 times slower, leading to a calculated transition state affinity for the enzyme of 10-22 M.25

Koshland proposed that retaining glycosidases proceed through a glycosyl enzyme intermediate.7 Evidence for such an intermediate has been provided using kinetic isotope effect (KIEs) studies. A KIE is a ratio of the rates of reaction for a light substrate

(naturally abundant species), and a heavy substrate (wherein one atom in the reactant is replaced with its heavier isotope).26 Pioneered by Sinnott and Souchard, KIE studies have been used to study several glycosidases, notably by Kempton and Withers in their study of Agrobacterium β-glycosidase. Kinetic parameters were measured for the β-

glucosidase-catalysed hydrolysis of fifteen aryl-glucoside substrates. A Bronstead plot of log kcat versus leaving group pKa value showed a concave downward shape indicating a decrease in the rate-determining step with respect to leaving group ability (Figure 1.7).

Substrates with good leaving groups displayed no rate-change, whereas substrates with poor leaving groups (pKa >8) displayed reduced reactivity, identifying the glycosylation step to be the rate-limiting step. These results provide evidence for a two-step mechanism where one step requires glycosidic bond cleavage to form the glycosyl-enzyme intermediate.

Figure 1.7: Bronstead plot of Agrobacterium β-glucosidase catalysed hydrolysis of aryl-glucosidase substrates with decreasing leaving group ability.

Deuterium secondary KIEs were measured for 5 aryl-glucoside substrates.

Substrates poor leaving groups (pKa > 8) had small a isotope effect (kH/kD = 1.06) whereas those with good leaving groups (pKa <8) had a larger isotope effect (kH/kD = 1.11). These results are consistent with formation of a glycosyl-enzyme intermediate via oxocarbenium ion-like transition states.

The collective view arising from this study and many others are that the transition states of glycosidases contain partially formed or broken bonds between the anomeric center and leaving group, and the anomeric center and the nucleophile (indeed, such a

view was espoused in the seminal review of Koshland).7 Charge development at C1 of the transition state is stabilized by partial double bond character that develops between the anomeric carbon and the endocyclic oxygen. For efficient orbital overlap, this requires the C5–O5–C1–C2 atoms to be coplanar. With only the C3 and C4 atoms having the freedom to move, this results in a preference for boat (B) or half-chair (H) conformations at the transition state or the energetically less favourable envelope (E) conformations

(Figure 1.8).27

Figure 1.8: The major oxocarbenium ion-like transition state conformations.

1.8 Substrate distortion during enzymatic hydrolysis of glycosidases

The major pyranose conformations and their interrelations can be represented in a

Stoddard diagram (Figure 1.9; this plot excludes E conformations).1 As pyranose rings

4 4 assume a C1 conformation in the ground state, and since only the H3 conformation can be directly accessed from this conformation, access to the other transition state conformations would require substantial substrate distortion.

Figure 1.9: Pyranose ring conformational interconversions of the northern (left) and southern (right) hemispheres of the Stoddard plot. Oxocarbenium ion transition states depicted in box.

Substrate distortion was first proposed by Phillips in 1967 when he was exploring inhibitor/enzyme complexes of the HEWL enzyme. Molecular modelling predicted that binding of a sugar in the –1 subsite would result in hinderance between the enzyme and the C5 and C6 atoms on the pyranose ring. Phillips proposed that to achieve binding in

 1 The nomenclature of ring conformations (as described by IUPAC) is based on an italicized capital letter denoting the shape of the ring. Pyranoses can exhibit 26 conformations that are defined as: chair (C), half chair (H), boat (B), skewed (S) and envelope (E). Atoms that lie above the plane are detonated by superscript; those which lie below the plane are denoted with subscript.210 12

the –1 subsite the pyranose ring must distort to a conformation where the C5–C6 bond was axial, subsequently the C2–C1–O5–C5 atoms would lie in a plane such that the pyranose would adopt a half-chair conformation.28 In 1974 the X-ray crystal structure of tetra-N-acetylchitotetraose bound to HEWL was obtained, with the tetrasaccharide bound in the –4 to –1 subsites. The conformation of the sugar in the –1 subsite assigned a conformation close to an envelope or a boat.29

The first distorted enzyme/substrate (Michaelis) complexes were structurally characterized in 1996 in two independent studies of β-glycosidases. Upon binding,

4 substrates were distorted from their preferred C1 chair conformations to E and S conformations.22,30 In these conformations the leaving group was pseudo axial, which was proposed to allow for in-line nucleophilic attack at the σ* anti-bonding orbital of the anomeric carbon (Figure 1.10).31

Figure 1.10: First half of reaction mechanism of enzymatic hydrolysis of chitobiose. 4 4 The first step involves substrate distortion from C1 to E conformations.

1.9 Reaction co-ordinate

A key focus in the study of glycosidases is assigning the conformational changes that occur at the sugar in the –1 subsite as it progresses along the reaction co-ordinate (Figure

1.11). These conformational changes, also referred to as the ‘conformational itinerary’ may be determined using X-ray crystallography, kinetic and thermodynamic analysis, and

computational modelling. X-ray crystallography allows the viewing of several critical points along the reaction co-ordinate including: free enzyme, enzyme/substrate

(Michaelis), glycosyl enzyme, and product complexes. As well; transition state mimicking inhibitors can allow a view of features of the transition state.

Figure 1.11: Reaction co-ordinate of a retaining α-glycosidase.

Michaelis complex

The first step along the reaction co-ordinate of glycosidase catalysis is the binding of substrate by enzyme to form an enzyme/substrate [E·S] complex, also referred to as the

Michaelis complex. As glycosidases catalyse rate enhancements of up to 1017 fold, studying the Michaelis complex before hydrolysis occurs is challenging. Insight into the structure of the Michaelis complex can be achieved by perturbing the system to prevent catalysis in three main ways: by co-crystallizing non-hydrolysable substrate mimics with

wildtype enzymes; by co-crystallizing substrates with inactive mutant enzymes; or on rare occasions by employing pH conditions where the enzyme is inactive.32

Glycosyl-enzyme intermediate complex

The first chemical step is formation of a covalent glycosyl enzyme intermediate [E-S], which is formed via the first transition state [TS1] . There are two main ways in which the glycosyl-enzyme intermediate can be studied experimentally. The methods are conceptually similar: they both seek to speed up the glycosylation step whilst slowing the deglycosylation step, leading to accumulation of the glycosyl enzyme intermediate.

The first method achieves this through use of a substrate with a good leaving group and a mutant enzyme wherein the acid or base residue has been replaced with an inactive residue. Mutation of the catalytic acid/base residue in retaining enzymes reduces the rate of hydrolysis of glycosides; however, by incorporating a good leaving group this selectively speeds up the glycosylation step.33 The deglycosylation step, which requires base catalysis to promote attack of water on the glycosyl enzyme intermediate is slowed slow, leading to accumulation of the intermediate. The second method was pioneered by

Legler using glycals,34,35 and later refined by Withers and co-workers36,37 who developed

2-deoxy-2-fluoro, 2-deoxy-2,2-difluoro and 5-fluoro-glycosyl fluorides and glycosides, which act as time-dependent inhibitors. Fluorine substitution at the 2- or 5-positions destabilizes both the glycosylation and deglycosylation transition states of a retaining glycosidase and slows formation of the glycosyl enzyme intermediate and its hydrolysis.

Destabilization of the oxocarbenium ion-like transition states is attributed to the inductive effects of fluorine and loss of hydrogen bonding from the 2-OH. However, by installation of a good leaving group the glycosylation step is accelerated, leading to rapid formation and accumulation of the glycosyl-enzyme intermediate (Figure 1.12).38

Withers’ fluoro-sugar technology has been applied in the study of many retaining glycoside hydrolase families, most notably HEWL.9 Initial attempts by Vocadlo et al. focused on the first method: mutation of the catalytic acid/base residue of HEWL, and incubation of the E35Q mutant with β-chitobiosyl fluoride (NAG2F). While formation of the covalent-intermediate could be observed by ESI-MS, turnover to product was too fast to allow 3D-structural analysis. Consequently, efforts turned to the second method whereby the wild-type enzyme was incubated with a 2-deoxy-2-fluoro analogue of β- chitobiosyl fluoride, NAG-2FGlcF. However, while detectable by mass spectrometry, the intermediate was not stable enough to apply X-ray crystallography. Ultimately, a combination of these techniques employing the HEWL E35Q mutant and NAG-2FGlcF allowed the glycosyl-enzyme intermediate to be trapped for sufficient time to allow its characterization by X-ray crystallography. X-ray crystallography revealed the pyranose

4 ring in the –1 subsite adopts an undistorted C1 conformation. D52 was identified as the catalytic nucleophile that initiates the first step of the double-displacement retaining mechanism forming the glycosyl-enzyme intermediate.39

Figure 1.12: Reaction co-ordinate of a retaining α-glycosidase illustrating how a modified substrate can enable trapping of the glycosyl enzyme intermediate. 2-Deoxy-

2-fluoro-glycosyl fluorides accelerate the first step by lowering the energy of [TS1]

relative to [TS2] , leading to accumulation of the E-S complex. Mimicry of the enzyme transition state complex

Glycoside hydrolases have oxocarbenium ion-like transition states that are conformationally restricted and bear partial positive charge at O1 and C1 (Figure 1.13).

While a stable, perfect mimic of the transition state is not possible, various aspects of this transition state can be mimicked. Transition state inhibitors of glycoside hydrolases are designed to either mimic the shape or charge of an oxocarbenium ion, and preferably both.

Figure 1.13: a. Oxocarbenium ion-like transition state. b. Oxocarbenium ion resonance forms

Shape based inhibitors mimic the flattened conformation at the transition state and typically have sp2-hybridized atoms at O1 or C1. The neutral glyconolactone, D-glucono-

1,5-lactone, is an effective β-glycosidase inhibitor. Its inhibition is predominately attributed to shape; however, it has been suggested that the polarized carbonyl group also partially mimics the positive charge of the oxocarbenium ion like transition state.

Glyconolactones are not very stable, and undergo hydrolytic ring opening under mildly basic conditions and convert to inactive 1,4-lactones under mildly acidic conditions, complicating measurement of inhibition constants.40,41 For these reasons researchers turned to the study of lactone analogues including gluconolactam, gluconohydroximolactam, and gluconoamidine, which are more stable under mild acid/base conditions.42–44 In 1992 Aoyagi and co-workers discovered nagstatin, a naturally occurring gluconoimidazole and discovered that it acts as a potent inhibitor of some glucosaminidases.45,46 The glyconoimidazoles are currently among the most potent glycosidase inhibitors that mimic the sp2 hybridization of the transition state.47

Importantly, the ‘glycosidic’ nitrogen can be protonated, providing some resemblance of the positive charge of an oxocarbenium ion. More recently, in 2018 Schröder and co- workers reported a new class of β-glucosidase inhibitors, the 1H-gluco-azoles (Figure

1.14).

Figure 1.14: Glycosidase inhibitors designed to mimic the sp2 hybridization of the oxocarbenium ion.

An oxocarbenium ion has two resonance forms, one where there is a positive charge on the endocyclic oxygen and another where there is a positive charge at the pseudo anomeric position. Nitrogen-containing heterocycles, imino- and azasugars, are frequently used to mimic these resonance forms. Iminosugars such as deoxynojirimycin

(DNJ) and castanospermine (CST) contain a nitrogen atom in place of the endocyclic oxygen that when protonated mimics the positive charge at O1 of an oxocarbenium ion.

Azasugars such as isofagomine (IFG) and noeuromycin (NOE) contain nitrogen in place of carbon at the pseudo-anomeric position that upon protonation mimics the partial positive charge found at C1 of the oxocarbenium ion. Imino/aza sugars typically bind to glycosidases with affinities that are orders of magnitude better than their corresponding natural substrates (Figure 1.15).48

Figure 1.15: Glycosidase inhibitors designed to mimic the charge of an oxocarbenium ion.

1.10 Scope of thesis

This thesis aims to explore structural and mechanistic features of representative enzymes from glycoside hydrolase families: GH99, GH134 and GH31. Chapters 2 and 3 will outline the design and synthesis of two GH99 mechanism-inspired inhibitors. Chapter 4 will examine the substrate preference, stereochemistry of hydrolysis, structural details and conformational itinerary of a newly discovered family of β-1,4-mannases (GH134).

Chapters 5 and 6 will investigate two GH31 enzymes isolated from two different sulfoglycolytic organisms and explore the related metabolic pathways these respective enzymes are part of.

CHAPTER TWO:

Investigation of the effects of shape and charge

on family GH99 endo-α-1,2-mannanases

2.1 N-Linked glycan biosynthesis in the secretory pathway

Protein glycosylation – a common post-translational modification

Post-translational modifications are chemical transformations of proteins that occur after their assembly (translation) on ribosomes. The majority of eukaryotic proteins undergo posttranslational modification, and protein glycosylation is an especially common and diverse post-translational modification that is critical for normal cellular function.49

Protein glycosylation involves the attachment of a carbohydrate to an amino acid residue resulting in nitrogen, oxygen, sulfur and carbon-linked glycoproteins. Linkage to the nitrogen of asparagine (N) results in N-linked glycoproteins. N-linked glycans play diverse roles in protein folding, stability, targeting and quality control, and influence the immunological properties of glycoproteins.50,51 N-Linked glycosylation occurs within the secretory pathway, commencing within the endoplasmic reticulum and continuing through the Golgi apparatus.52 There are two major pathways for N-linked glycan maturation, both of which involve the build-up, trimming and reconstruction of the N- glycan structure through the action of glycosidases and glycosyltransferases. The classical pathway utilizes only trimming exo-acting glycosidases whereas the 'endo- mannosidase pathway' involves the action of the endo-acting glycoside hydrolase endo-

α-1,2-mannosidase, among other trimming exo-glycosidases.

Biosynthesis of N-linked glycans – the classical pathway

N-Linked glycan biosynthesis commences in the ER with transfer of a highly a conserved

14 residue oligosaccharide, Glc3Man9GlcNAc2, which is preassembled on a lipidic carrier, dolichol phosphate. Transfer of the Glc3Man9GlcNAc2 14-mer occurs co- translationally within the lumen of the ER to the nitrogen of an asparagine residue on ribosome-bound peptides catalysed by oligosaccharyltransferase (Figure 2.1).53 The consensus sequence is Asn-Xxx-Ser/Thr, where Xxx is any residue except Pro.54

Figure 2.1: Transfer of the Glc3Man9GlcNAc2 14-mer to nascent protein catalysed by oligosaccharyltransferase.

Following the transfer of the preassembled 14-mer Glc3Man9GlcNAc2 to asparagine residues in peptides under assembly on the ribosome, assorted glycosidases sequentially trim specific residues from the oligosaccharide as it transits through the secretory pathway. In the classical processing pathway the first two glucose residues are sequentially trimmed by the exo-acting ER α-glycosidases I and II while the nascent glycopeptide is still attached to the ribosome.55 The resultant glycoproteins feature a terminal α-Glc-1,3-α-Man chain on the A branch and detach from the ribosome as a unfolded glycoproteins. These monoglucosylated glycoproteins bind to the lectins calnexin (CNX) and calreticulin (CRT), which act as molecular chaperones to facilitate proteins folding in concert with the disulfide oxidoreductase ERp57. The CNX/CRT proteins dissociate from the glycoprotein upon cleavage of the last glucose residue by the action of α-glucosidase II.56,57

Following release from the CNX/CRT chaperone, glycoproteins are assessed for their folding status by UDP-glucose-glycoprotein glucosyltransferase (UGT1) and

enzymes of the ER-associated degradation (ERAD) pathway. Newly synthesised but misfolded glycoproteins with intact B and C branches are substrates for UGT1, and are reglucosylated, which allows them to return to the CNX/CRT binding cycle for additional folding attempts.58,59 Concurrent with repeated folding attempts, exo-mannosidases slowly trim the B and C branches, allowing for identification of terminally-misfolded proteins, which are removed from the CNX/CRT binding cycle through the ERAD process, which involves further demannosylation and proteasomal degradation.

Correctly folded deglucosylated proteins are further modified with the removal of one or two mannose residues by ER α-1,2-mannosidase, and exit the ER to the Golgi apparatus for further processing.60,61 Within the Golgi, further trimming and reglycosylation steps ultimately lead to ‘hybrid-type’ and ‘complex-type’ structures

(Figure 2.2).62 Finally, mature glycoproteins are packaged into vesicles and transported to their various cellular destinations.

Figure 2.2: An overview of some major steps that occur in the classical N-linked glycan biosynthesis pathway. Trimming of Glc3Man9GlcNAc2 occurs in the endoplasmic reticulum though a series of exo-glycosidases. Correctly folded Man7GlcNAc2 N-linked glycans are transported to the Golgi apparatus for possessing into hybrid-type and complex-type glycans.

2.2 Discovery of endo-α-1,2-mannosidase, a unique endo-acting glycoside hydrolase

In 1987, Spiro and co-workers identified a novel enzymatic activity in rat liver acting on mammalian N-linked glycans that cleaved GlcNAc2Man9Glc1-2 to GlcNAc2Man8,

63 releasing intacts Glu1-2-1,3-α-Man units. Termed endo-α-1,2-mannosidase (EM), the enzyme was shown to be localized predominately within the Golgi apparatus. Subsequent work showed that EM can cleave the α-1,2 linkage between a glucose-substituted mannose and the rest of the glycan (Figure 2.3) in a variety of high mannose N-linked

64 glycans including GlcNAc2Man9Glc1-3 and GlcNAc2Man4-8Glc. Monoglucosylated

(GlcNAc2Man9Glc) substrates were hydrolysed faster than di- and triglucosylated

substrates (GlcNAc2Man9Glc2 and GlcNAc2Man9Glc3), and glycans with truncated mannose B and C chains (GlcNAc2Man4-8Glc) displayed increasing activity with a decreasing number of mannose residues. This suggests that EM is specific to a polymannose branch substituted with a glucose residue and that additional glucose residues or nonglucosylated mannose residues may sterically inhibit binding to the active site of EM.65 EM was purified from rat liver Golgi membrane and molecularly cloned.66,67

There was no sequence homology between EM and any other glycosidase, leading to its assignment to a novel glycoside hydrolase family, GH99.

Figure 2.3: Cleavage of GlcNAc2Man9Glc by endo-α-1,2-mannosidase.

Biosynthesis of N-linked glycans using the endo-α-1,2-mannosidase pathway

The cleavage of various glucosylated high mannose N-glycans involving EM is termed the endo-α-1,2-mannosidase pathway (Figure 2.4). It allows the bypass of the sequential action of several exo-glycosidases, specifically glucosidase II, and ER α-mannosidase.64

It has been shown that the EM pathway can support up to 80% of normal flux of N-glucan processing in the event of glucosidase II by CST and ER α-mannosidase I blockade by

DMJ.68–70 The major role of EM appears to be the Golgi processing of competently-

folded, ER-escaped glucosylated N-linked glycans, redirecting them to the N-glycan maturation processes within the secretory pathway. The pathway also plays a role in deglucosylating misfolded glycoproteins in the Golgi apparatus and intermediate compartments. Deglucosylated misfolded glycoproteins are recycled back to the ER where they can be eliminated from the secretory pathway through ERAD. This recycling process was demonstrated through FOS (free oligosaccharide) analysis.71 FOS are glycans that are released as deglucosylated glycoproteins undergo ERAD and can therefore be used as biomarkers for ERAD. Under glucosidase I/II inhibition, FOS glycans were deglucosylated by EM and did not accumulate in the cell. This demonstrated that EM action in the Golgi can deglucosylate glycans before they are transported back to the ER for recycling, preceding ERAD.

Figure 2.4: The endo-α-1,2-mannosidase pathway of N-linked glycan biosynthesis. ER escaped N-linked glycans are processed by the Golgi located endo--1,2-mannosidase, which bypasses the classical pathway.

2.3 N-Linked glycan degradation by bacterial endo-α-1,2-mannanases

Bacteria and archaea possess genes encoding proteins classified as GH99 family members.72 The first enzymatic characterization of a bacterial GH99 protein was reported in 2011. The gram negative, rod-shaped bacterium Shewanella amazonensis was isolated from shallow-water marine sediments of the Amazon River delta.73 This bacterium contained a gene which produced the GH99 protein, Sama99, sharing 45% homology with human EM. Sama99 was found to act on the pyridylamino (PA)-modified sugars,

Glc1Man9GlcNAc2-PA and Glc3Man9GlcNAc2-PA, yielding Man9GlcNAc2-PA in both cases. This is essentially identical to the mode of action of rat-liver EM, which acts on

Glc1-3Man9GlcNAc2, yielding Man8GlcNAc2. A subsequent study demonstrated that

GH99 enzymes from the gut bacteria Bacteroides thetaiotaomicron (Bt) and Bacteroides xylanisolvens (Bx) also possess the ability to act on glucosylated high mannose mammalian N-glycans sharing 42% and 41% sequence identity, respectively, to human

EM (Figure 2.5).20

Figure 2.5: Structures of mammalian high-mannose N-glycans and yeast high-mannose N-glycans. Box indicates identical chains with the exception of a single stereocenter (Glc versus Man).

However, as most bacteria do not produce N-linked glycans, it was unclear what role GH99 enzymes played in bacteria. Subsequent work demonstrated that the

Bacteroidete GH99 genes were located within one of three operons that were upregulated upon growth on fungal α-mannan.74 Yeast mannan is derived from mannoprotein, a component of the fungal cell wall (along with β-1,3-glucan and β-1,6-glucan). The yeast

N-linked glycan core is identical to mammalian N-glycans with the exception of a differing stereocenter: glucose in the mammal and mannose in the yeast. The yeast core is decorated with an outer chain of -linked mannose units, with long α-1,6-linked backbones and short α-1,2- and α-1,3-linked side chains (Figure 2.5). Both the core, and the short side chains, bear tetrasaccharide sequences that were shown to be preferred substrates for the bacterial GH99 proteins, leading to their assignment as endo-α-1,2- mannanases.75 It is argued that the Bacteroides, as common members of the human gut microbiota, regularly encounter yeast mannans in the human gastrointestinal tract.74 Such encounters are to be expected since the domestication of yeasts in food production occurred at least 7000 years ago, meaning that the fungal cell wall has comprised an enduring constituent of the human diet for millennia. Family GH99 is now known to contain enzymes that can process complex carbohydrate with two distinct enzymatic activities: mammalian GH99 enzymes are endo-α-1,2-mannosidases76 and bacterial

GH99 enzymes are endo-α-1,2-mannanases.75 Sequence alignment of several mammalian and bacterial GH99 enzymes revealed a difference in the amino acid at position 126. In the bacterial enzymes BtGH99 and BxGH99 it is a tryptophan whereas the mammalian counterpart is a tyrosine. Therefore, in mammalian GH99 enzymes, the phenolic amino acid side chain of tyrosine is predicted to hydrogen bond with the O2 position of glucoside, mammalian enzymes have a preference for glucose at the -2 subsite. It’s

speculated that the presence of tryptophan in bacterial enzymes results in a preference for mannose.75

Mechanistic analysis of GH99 enzymes

As discussed in Chapter 1, GH99 enzymes are proposed to perform substrate assisted catalysis though an unusual NGP mechanism. While there is no clear precedent for a 1,2- anhydro sugar intermediate in biological catalysis, it has been known for many years that basic solvolysis of 1,2-trans glycosyl fluorides and aryl glycosides occurs with a neighbouring group participation mechanism through a 1,2-anhydro sugar.77–79 Speciale and co-workers studied the transition state for the hydroxide-catalysed hydrolysis of 4- nitrophenyl α-D-mannopyranoside in aqueous media using kinetic isotope effect (KIE) measurements coupled with ab initio theoretical methods (Figure 2.6). 80 The results were consistent with a reaction mechanism involving rate-limiting formation of an intermediate epoxide that opens to give α-D-mannopyranose.

Figure 2.6: Alkaline hydrolysis of 4-nitrophenyl α-D-mannopyranoside through neighbouring group participation.

2.4 Inhibitors of GH family 99 enzymes

Since its discovery in 1987, significant effort has been applied to the discovery and development of inhibitors for EM. In 1988 known inhibitors of oligosaccharide processing exo-glycosidases including DNJ, CST, bromoconduritol (BCD), DMJ, SWN, and ethylenediaminetetraacetic acid (EDTA) were examined for their efficacy against the enzyme, but were ineffective.65 Some years later Spiro and co-workers performed an

extensive series of modifications to the characteristic product of the EM-catalysed reaction, Glc-α-1,3-Man, including deoxygenation, desaturation, fluorination, and the synthesis of several imino sugar variants. Compounds were assessed for their ability to inhibit EM processing of N-linked glycans. The most effective inhibitors identified were:

GlcddMan, GlcGlucal, GlcDMJ; their respective IC50 values were 3.8, 2.3, and 1.7 µM when incubated with EM from rat liver cells (Figure 2.7).81

Figure 2.7: The most effective inhibitors of eukaryotic EM as identified by Spiro and co-workers.81

The success of GlcDMJ prompted our laboratory to undertake the synthesis of the aza-sugar -glucopyranosyl-1,3-isofagomine (GlcIFG). Owing to the inability to obtain purified mammalian EM, the direct binding affinities to GH99 family members was assessed by studying the soluble bacterial orthologs BtGH99 and BxGH99. Isothermal titration calorimetry studies revealed that BtGH99 binds GlcDMJ and GlcIFG with KD values of 2400 nM and 635 nM, respectively (Figure 2.8).20,8275 Evidently changing the position of the positive charge improved enzyme-inhibitor affinity despite the loss of a hydroxyl group at C2. Based on the realization that BtGH99 enzymes are endo-α-1,2- mannanases that act on yeast mannan, α-D-mannosyl-α-1,3-isofagomine (ManIFG) was designed and synthesized. ManIFG binds with a four-fold higher affinity to BtGH99 with

75 a KD value of 171 nM.

Figure 2.8: Sequence of inhibitor development and associated KD values for BtGH99.

Isofagomine inhibitors lack a 2-OH, which has been proposed to be important for transition state stabilization of a range of glycosidases. Additionally, within GH99 enzymes it is proposed to be involved in the enzyme mechanism, as a nucleophile that undergoes neighbouring group participation at C1. In 2001 Lui et al. synthesised the IFG- like noeuromycin (NOE), which possesses a hydroxyl group reinstated at C2.83 NOE was

2-1000 times more potent than IFG against a range of glycosidases and mannosidases, which highlights the importance of the 2-OH for binding of aza-sugar inhibitors.

2.5 Aims

The aim of this chapter was to synthesize α-mannosyl-1,3-noeuromycin (ManNOE, 201), to investigate whether reintroduction of the ‘missing’ 2-hydroxyl group in ManIFG could deliver a more potent inhibitor. With this compound in hand, we would then investigate its affinity for endo-α-1,2-mannanases, pursue the three-dimensional structure of its complex, and seek to understand its binding using computational methods.

2.6 Results and Discussion

Previously approaches to noeuromycin (NOE) and related molecules

Several approaches to NOE have been developed. The key challenge in assembling this molecule is the synthesis of a branched carbon chain, particularly when starting from carbohydrate-based precursors that possess linear carbon backbones. In our opinion, the most practical approaches use cyanide to form a nitrile intermediate from D-arabinose or

L-xylose that possesses the branched carbon skeleton, which can be readily converted to

NOE in a few steps. For example, Meloncelli and Stick reported the synthesis of NOE via an isopropylidene nitrile intermediate obtained in 4 steps from L-xylose (Scheme 2.1).

The nitrile intermediate was converted to NOE using the procedure as described by

Andersch and Bols.84,85 The major drawbacks of this method are the high cost of L-xylose; the low yield (13%) obtained for preparation of benzyl α-L-xyloside; and an inefficient isopropylidenation step involving wasteful separation from a regioisomer.

Scheme 2.1: Meloncelli and Stick’s synthesis of noeuromycin from L-xylose.84

A synthesis of the related molecule isofagomine was reported by Goddard-Borger and Stick, beginning with inexpensive D-arabinose (Scheme 2.2).86 This approach provided a similar nitrile intermediate in a longer route of six steps but in good overall yield.

Scheme 2.2: Synthesis of a nitrile precursor according to the method of Goddard- Borger and Stick.86

A major consideration in the synthesis of glycosylated NOE is the timing of the formation of the disaccharide linkage relative to the formation of the NOE moiety. NOE contains a hemiaminal, which is somewhat unstable and prone to dehydration to an imine, and/or undergo an Amadori rearrangement. Meloncelli and Stick have reported the regioselective β-glucosylation of a nitrile-diol, at the 2-position, allowing formation of the β-glucosyl-1,3-NOE upon reduction and ring-closure (Scheme 2.3).87

Scheme 2.3: Meloncelli and Stick synthesis of GlcNOE.87 35

Synthetic strategy towards ManNOE

We chose to develop a route to α-mannosyl-1,3-noeuromycin (ManNOE, 201) involving a nitrile intermediate, which would then be elaborated to ManNOE. This would involve forming the glycosidic linkage prior to forging the noeuromycin moiety. Within this general framework two strategies were advanced (Figure 2.4). Strategy 1 involves an earlier mannosylation of the selectively protected alcohol, a disaccharide intermediate en route to the nitrile, elaboration to the disaccharide nitrile, and finally reduction and cyclization to the NOE heterocycle. Strategy 2 involves the regioselective mannosylation of the nitrile-diol, prepared in six steps as outlined by Stick, followed by elaboration to the nitrile disaccharide, reduction of the nitrile and formation of the NOE heterocycle

(ManNOE, 201).

Scheme 2.4: Outline of strategies for the synthesis of ManNOE. Strategy 1 involves an early glycosylation followed by nitrile formation and finally conversion of noeuromycin; Strategy 2 involves a late glycosylation of a nitrile followed by conversion to noeuromycin.

Strategy 1 was studied as part of my MSc studies (2013-2014) and will be summarized herein for the sake of presenting a complete report. Strategy 1 was initially pursued as it was unclear whether α-mannosylation of the nitrile-diol could be achieved

with the same regioselectively to give the 1,2-linked disaccharide as the -glucosylation precedent set by Meloncelli and Stick. Moreover, Strategy 1 appeared enticing because of its ready access to the protected arabinoside alcohol that could be glycosylated exclusively at the C2 position, avoiding issues of regioselectivity.

The alcohol (acceptor) was prepared in two steps from D-arabinose by Fischer glycosidation with benzyl alcohol and then isopropylidenation using dimethoxypropane/TsOH (Scheme 2.5). Condensation of the donor and acceptor proceeded uneventfully, affording the intermediate disaccharide in excellent yield

(Scheme 2.5). Acetonide cleavage, and regioselective alkylation of the diol (via a stannylene acetal) afforded the C4 alcohol. However, difficulty was encountered in the use of imidazolesulfonates to invert the stereocentre at C4 of the arabinosyl sugar through the Lattrell-Dax reaction; better results were obtained using a triflate. The resulting L- xylose-configured alcohol was converted to the triflate; however, cyanide substitution proceeded giving low yields of the D-arabino nitrile disaccharide; nevertheless, sufficient material was obtained to allow reduction of the nitrile and transformation to the Boc- protected amine. Disappointingly, the sole attempt at benzyl group and Boc carbamate deprotection were unsuccessful. Overall, while broadly successful, low yields throughout this sequence failed to deliver sufficient material to support the synthetic efforts.

Consequently, at the end of my MSc studies we abandoned this approach.

Scheme 2.5: MSc approach (213-2014), Strategy 1

With renewed optimism and a fresh outlook at the start of my PhD studies, we revisited the overall strategy towards ManNOE based on my MSc results. In retrospect, while no glaring shortcomings were evident in Strategy 1, difficulties were encountered in troubleshooting failed reactions at the disaccharide stage, and for reasons that were unclear it was not possible to obtain a high yield in the cyanide substitution on the disaccharide triflate. Consequently, we now chose to investigate Strategy 2, wherein the inversion and cyanide substitution steps follow literature protocols and are carried out on a monosaccharide, allowing for simpler, more easily interpreted spectra. It was hoped that conducting a glycosylation on a nitrile diol would allow more material to be brought through the sequence to support the synthetic endgame.

Synthesis of the arabinoside acceptor

The known arabinoside acceptor was synthesised from commercially-available D- arabinose in 7 steps (Scheme 2.6), as originally reported by Goddard-Borger and Stick

(and as was summarized in Scheme 2.2).86 In my hands, Fischer glycosylation of D- arabinose was performed with a suspension of D-arabinose in benzyl alcohol, with HCl generated in situ from acetyl chloride, to give benzyl β-D-arabinose (202) in 79% yield.88

Treatment of 202 with triethyl orthoacetate and catalytic p-toluenesulfonic acid gave the

3,4-orthoester. The acid was neutralised with triethylamine, and the orthoester regioselectively cleaved with TFA/H2O to give the acetate (203) in 85% yield.

Isopropylidenation of 203 with 2-methoxypropene and catalytic p-toluenesulfonic acid gave the 2,3-acetonide; saponification of the acetate using sodium hydroxide in methanol gave the alcohol (204) in 74% yield.

Scheme 2.6: Synthesis of acetonide (204). Reagents and conditions; a. CH3COCl, BnOH, 79%; b.i. MeC(OEt)3, TsOH, CH2Cl2; ii. Et3N; iii. TFA/H2O 9:1, 85%; c. i. CH2C(OMe)Me, TsOH, CH2Cl2; ii. NaOH, MeOH, 74% over two steps.

At this point we needed to achieve the stereochemical inversion of the alcohol 204 at C4. Epimerization of alcohols can be achieved though the installation of a good leaving group followed by the Lattrell-Dax reaction.89,90 Imidazole-1-sulfonates (imidazylates) and triflates are two examples of leaving groups commonly employed in the Lattrell-Dax reaction. Triflates are usually less stable than their imidazylate counterparts often requiring immediate use after preparation; for this reason imidazylates are preferred. They

can be synthesised through a one-pot procedure involving treatment of the alcohol with sulfuryl chloride in pyridine, leading to a chlorosulfate intermediate, followed by substitution with imidazole.91 Compound 204 was converted to an imidazylate (205), which when subjected to the Lattrell-Dax reaction gave the L-xylo-configured alcohol

(206) in only 10% yield over two steps (Scheme 2.7). Also isolated was the 4-chloro-4- deoxy-L-xyloside (207), arising from chloride substitution at C4. Presumably this compound arises from substitution of the intermediate chlorosulfate, or the imidazylate by chloride formed in the reaction mixture, and is unreactive under Lattrell-Dax conditions. Despite several attempts to improve the reaction we could not prevent chloride substitution. Ferro and co-workers have reported a similar problem in their attempts to form an imidazylate from a secondary alcohol.92

Scheme 2.7: Synthesis of L-xylo-configured alcohol (207) using imidazylate as the leaving group. Reagents and conditions: a. i. SO2Cl2, pyridine; ii. ImH; b. NaNO2, DMF, 90 °C, 10% over 2 steps.

In contrast to the difficulty in forming imidazylate 205, treatment of 204 with triflic anhydride in pyridine smoothly gave the triflate (208) (Scheme 2.8). Compound

208 was treated with sodium nitrite in DMF to afford the L-xylo-configured alcohol 206 in 45% yield over 2 steps.

Scheme 2.8: Synthesis of L-xylo-configured alcohol (206) using triflate as the leaving group. Reagents and conditions; a. Tf2O, pyridine, CH2Cl2; b. NaNO2, DMF 90 °C, 45% over 2 steps.

Whilst a triflate proved superior to an imidazylate, the yield for the substitution/inversion steps in the Lattrell-Dax reaction was still low. Consequently, we opted to apply a Mitsunobu reaction, inspired by Goddard-Borger and Stick.86 This involves substitution of an alcohol by a carboxylic acid, forming an ester with inversion of stereochemistry; subsequent ester cleavage affords the L-xylo-configured alcohol.93

Compound 204 was treated with diisopropyl azodicarboxylate, triphenylphosphine and

4-nitrobenzoic acid to give the nitrobenzoate (209). Transesterification with Et3N/MeOH provided 206 in 73% yield over 2 steps, a significant improvement compared to the triflate/Lattrell-Dax method (45%). Compound 206 was treated with triflic anhydride and pyridine to give the triflate (210). Substitution of 210 with KCN in DMF gave the nitrile

(211) in 47% over two steps. Compound 211 was treated with TFA/H2O 9:1 to cleave the acetonide affording diol 212.

Scheme 2.9: Synthesis of nitrile (212). Reagents and conditions: a. 4-O2NC6H4CO2H, DIAD, Ph3P; b. Et3N, MeOH, 73% over, 2 steps; c. Tf2O, pyridine, CH2Cl2, PhMe; d. KCN, DMF, 47% over 2 steps; e. TFA/H2O 9:1.

Synthesis of the mannosyl donor

The known mannosyl trichloroacetimidate (219)94 was synthesised from D-mannose in 8 steps (Scheme 2.10). D-Mannose was acetylated with acetic anhydride in the presence of sulfuric acid to give mannose pentaacetate (213) as a mixture of anomers in 80% yield.

Bromination with HBr in acetic acid gave the α-mannosyl bromide (214). Treatment of

214 with pyridine and methanol gave the triacetyl orthoester (215) in 52% yield over two steps. The formation of the orthester 215 can be rationalised by ionization of the bromide and capture of the resultant oxocarbenium ion by the acetate at C2 forming a dioxolenium ion; nucleophilic attack on the dioxolenium ion by methanol and deprotonation gives the orthoester. Deacetylation of the orthoester (215) under Zemplén conditions with catalytic sodium methoxide in methanol gave the triol (216). Benzylation of 216 with sodium hydride and benzyl bromide gave the orthoester (217), in 52% yield over 2 steps.

Scheme 2.10: Synthesis of 2-O-acetyl-3,4,6-tri-O-benzyl-α-mannoopyranosyl trichloroacetimide (219). Reagents and conditions: a. Ac2O, cat. H2SO4, 80%); b. HBr, AcOH; c. methanol, pyridine (52%); d. cat. NaOMe, MeOH; e. NaH, BnBr, DMF (62% over 2 steps); f. TFA/H2O 9:1; g. 1,8-diazabicyclo[5.4.0]undec-7-ene, CCl3CN, CH2Cl2, 80%.

Regioselective cleavage of the orthoester (217) in TFA/H2O, 9:1 gave the hemiacetal (218). The regioselectivity of this transformation merits a brief remark. cis-

Orthoesters on pyranose systems hydrolyse to orthoacids under acidic conditions, which cleave to form vicinal hydroxyl esters with the ester on the axial oxygen (Figure 2.10).

The regioselectivity of this reaction can be attributed to the oxocarbenium ion- intermediates of the orthoacid rearrangement, which is stabilised by two sets of lone pair electrons antiperiplanar to the scissile bond, one from each oxygen in the dioxolane ring system. This ring system has two low energy conformations that potentially can be adopted. In the first conformation the lone pair at O1 is anti-periplanar with the O2-C bond; however, this conformation is unfavourable as it results in a clash between substituents on the orthoacid carbon and the axial proton on C4. In the second conformation the lone pair at O2 is anti-periplanar with the O1-C bond, favouring scission

of this bond leading to formation of 218.95, Finally treatment of 218 with 1,8- diazabicylo[5.4.0]undec-7-ene and trichloroacetonitrile in CH2Cl2, gave 219 in 80% yield.

Figure 2.10: The two conformations for each epimeric orthoacid that could lead to cleavage of the dioxolane ring. The lone pair antiperiplanar to the scissile bond is depicted in black. The conformation on the left is preferred in both epimers.

Stereoselective trans-glycosylations can be readily achieved through the use of trichloroacetimidate as glycosyl donors. This class of compound was developed by the

Schmidt laboratory, and upon activation of the imidate by a proton or Lewis acid, trichloroacetimidate is lost giving rise to a resonance-stabilised oxocarbenium ion. 96

Participation by an ester at C2 results in a dioxolenium ion, which is opened in an SN2- like reaction by a nucleophile forming a trans linkage with excellent stereoselectivity. In our case the diol (212) may react at the 2- or the 3-OH positions. In 2007 Meloncelli et al. demonstrated the regioselective -glucosylation of diol 219 at the 2-position using a

D-gluco-configured trichloroacetimidate (Scheme 2.3). They attributed the regioselectivity to the electron-withdrawing effect of the nitrile, which electronically reduces the nucleophilicity of the 3-OH. However, it is also possible that this may have a significant contribution from stereochemical ‘matching’ of the chiral donor and chiral acceptor, and use of a different glycosyl donor may not give the same outcome. In the event, glycosylation of the diol 212 with the donor 219 catalysed by triflic acid, afforded a disaccharide alcohol as the major product that was isolated in 46% yield (Scheme 2.11).

In order to unequivocally assign its structure, this alcohol was acetylated to give a

diacetate; NMR analysis revealed a large downfield shift in H3 relative to the disaccharide alcohol, defining the structure of the disaccharide alcohol as 220, and the diacetate as 221.

A negligible amount of α-1,3-trans linked disaccharide was observed.

Scheme 2.11: Condensation of 212 and 219, and acetylation of the product. Reagents and conditions: a. TfOH, CH2Cl2, 46%; b. Ac2O, Pyridine.

Disaccharide alcohol 220 was treated with an excess of borane dimethyl sulfide complex, followed by addition of methanol, to give a crude amine (222) (Scheme 2.12).97

The amine was treated with di-tert-butyldicarbonate and aq. sodium carbonate to give the

Boc-protected disaccharide 223 in 68% yield over two steps.

Scheme 2.12: Synthesis of the Boc-protected disaccharide (223). Reagents and conditions: a. HCl, CH3OH, 80%; b. i. BH3.Me2S, THF; ii. MeOH; iii. 10% aq. Na2CO3, Boc2O, TFA/MeOH (1:2), 68% over 2 steps.

Careful planning of the final steps was crucial to successful attain our target,

ManNOE (201). There were two considerations: the order of the final deprotection steps and how the compound would be stored. The formation of the NOE moiety occurs though several intermediates (Scheme 2.13). Firstly, hydrogenolysis of the benzyl groups affords a polyhydroxylated lactol (step a); cleavage of the Boc group liberates an amine (step b);

finally, hemiacetal/hemiaminal interconversion (steps c, d). forms the NOE ring. Once formed, the NOE ring is prone to hemiacetal/hemiaminal interconversion and hence exists as a mixture of D-manno and D-gluco α-hydroxypiperidine and α-amino pyranose isomers.

Scheme 2.13: Proposed mechanism for the formation of ManNOE (201). a. Deprotection of benzyl groups; b. cleavage of the Boc group; c. ring opening; d. ring closure to give the NOE moiety.

The hydroxyl group of noeuromycin is prone to elimination to form the imine; this in turn could be reduced to give isofagomine if 201 is exposed to H2 in the presence of Pd-C (Scheme 2.14). Therefore, it was essential that hydrogenolysis of the benzyl groups was performed prior to liberation of the amine by cleavage of the Boc group.

Scheme 2.14: Subjecting NOE to standard palladium-catalysed hydrogenation gives the azasugar, IFG.

The sensitivity of the hemiaminal of NOE to elimination and Amadori rearrangement means that its stability as the free base is low. The Amadori rearrangement as applied to NOE involved dehydration to the imine, tautomerization to the enamine, and

hydration to give a gem-diol (Scheme 2.15). In our case this will lead to loss of the sugar at the 2-position. Fortunately, NOE is much more stable at low pH and has been isolated as a hydrochloride salt.85

Scheme 2.15: Amadori rearrangement of noeuromycin at neutral pH.

With these stability and reactivity considerations in mind, 223 was treated with

H2 over Pd(OH)2/C at 10 bar for 48 h to give a heptaol that upon mass spectrometric analysis revealed an m/z value consistent with that expected for the target (Scheme 2.16).

The crude product was treated with 1 M HCl to give the hydrochloride salt, ManNOE.HCl

(201) (as a mixture of α-hydroxypiperidine and pyranose isomers) in 80% yield over two steps.

Scheme 2.16: Synthesis of α-Man-1,3-noeuromycin.HCl (201). Reagents and conditions: a. i. H2, Pd(OH)2/C, THF/H2O; ii. 1 M HCl, 80% over 2 steps.

Binding affinity and 3D structural analysis of ManNOE

Isothermal titration calorimetry (ITC) and X-ray crystallography was performed by Mr

Lukasz Sobala from Prof. Gideon Davies laboratory (University of York). ITC revealed that ManNOE binds to BxGH99 with KD of 13 mM and to BtGH99 with KD of 30 mM

(Figure 2.11, Table 2.1). This constitutes 17- and 5-fold improvements in binding affinity

for the respective enzymes when compared to ManIFG, demonstrating that installation of the 2-OH provides improved inhibition.

Figure 2.11: Isothermal titration calorimetry of ManNOE binding to (left) BtGH99 and (right) BxGH99.

ManIFG ManNOE (201)

BxGH99 270 nM 13 nM

BtGH99 143 nM 30 nM

Table 2.1: Dissociation constants of ManIFG and ManNOE with BxGH99 and BtGH99

3D structures of a ternary complex of BxGH99-ManNOE-α-1,2-manobiose

(Figure 2.12) and a binary complex of BxGH99-ManNOE and were solved at 1.05 and

1.14 Å respectively. Figure 2.13 shows ManNOE bound in the -2/-1 subsites in both complexes. Notably, there is a short (2.58 Å) hydrogen bonding interaction between the

2OH and E333. Thus, reinstatement of the hydroxyl group missing in ManIFG, as part of

ManNOE, results in specific interactions with E333, the residue implicated in deprotonation of C2 in the neighbouring group participation mechanism and provides the most potent ligand yet reported for any GH99 enzyme.

Figure 2.12: Ternary complex of BxGH99-ManNOE-α-1,2-manobiose.

Figure 2.13 Zoom-in of the active site of BxGH99 in complex with ManNOE and α- 1,2-mannobiose. a. 3D-structure ManNOE binding in the -1/-2 subsites. b. Cartoon highlights key interactions with conserved active site residues: E336 (acid/base) and E333 (proposed base that deprotonates O2).

Synthesis of ManddMan and ManGlucal

The aforementioned work, conducted by myself, was part of a larger collaborative project that involved contemporaneous synthetic work by a co-worker, Pearl Fernandes. Owing to its bearing on the broad area of investigation and the conclusions to be drawn in this chapter, I will now take a detour to introduce that work in summery herein. Pearl

Fernandes synthesized two neutral compounds as putative inhibitors of endo--1,2- mannanase, ManddMan and ManGlucal, inspired by an earlier report by Spiro and co- workers (Figure 2.17).81 ManddMan and ManGlucal were synthesized from α-1,3- mannobiose. Acetylation, followed by bromination gave the mannobiosyl bromide, this was treated with Zn in MeOH to give the protected glucal. Zemplén transesterification afforded ManGlucal. Reduction using H2/Pd(OH)2-C followed by Zemplén transesterification afforded ManddMan.98,99

Scheme 2.17: Synthesis of ManGlucal and ManddMan by Pearl Z. Fernandes.

Binding affinity and 3D structural analysis of the neutral compounds ManddMan and ManGlucal

Dissociation constants for the binding of ManddMan and ManGlucal to BtGH99 and

BxGH99 were determined by 2D NMR in the laboratory of Prof Jesus Jiminez-Barbaro.

ManddMan was found to bind to BxGH99 with KD of 221 μM and to BtGH99 with KD of

53 μM. ManGlucal bound to BxGH99 with KD of 111 μM and to BtGH99 with KD of 15

μM (Table 2.2).

The nature of binding of ManGlucal to these GH99 enzymes was of interest.

Reaching back to seminal work by Legler, it is known that glucals are generally good inhibitors of retaining glycosidases that use a classical mechanism involving an enzymatic carboxylate as nucleophile.34,100 In these cases the enol ether of the glucal can be protonated by the conjugate acid of the carboxylate leading to addition to the glucal to give a 2-deoxy-glycosyl enzyme; this usually turns over to give the hydrated glycal.101

Retaining enzymes that do not utilize a carboxylate as nucleophile, such as N- acetylhexosaminidases that use neighbouring group participation, and the sialidases

(which use a tyrosine nucleophile), are competitively inhibited by glycals (the classic example being the drug Relenza); for hexosaminidases they are also slowly turned over to the hydrated glycal.102,103 In order to investigate whether inhibition by ManGlucal leads to a chemical reaction, the compound was incubated with the enzyme and the products analysed by MS. No hydrated product could be identified, and given that this family is

retaining, this observation provides further evidence for the neighbouring group participation mechanism proposed for GH99. The 3D structure of BxGH99 complexed

4 with ManGlucal revealed binding in the -2/-1 subsites with the Glucal ring in a H5 conformation in the -1 subsite. The 3D structure of ManddMan showed it to also bind in

4 the -2/-1 subsites with the ddMan moiety in a H1 conformation matching that expected for the ground state conformation of this compound.

ManGlucal ManddMan

BxGH99 111 μM 211 μM

BtGH99 15 μM 53 μM

Table 2.2: Dissociation constants (KD) and ManddMan with BxGH99 and BtGH99.

Conformational analysis of ManddMan ManGlucal and ManNOE

The conformational free energy landscapes (FELs) were calculated to give an insight into the intrinsic conformational preferences of the inhibitor headgroup in -1 subsite, and how these related to the conformations observed upon enzyme. FELs of the inhibitors were calculated in the laboratory of Prof Carme Rovira (Figure 2.13). FELs were computed by ab initio metadynamics, and the Cremer-Pople puckering coordinates θ and ϕ were used as collective variables, yielding a Mercator representation for each inhibitor FEL.

a. D-Glucal b. ddMan

c. NOE

Figure 2.13: Conformational free energy landscapes (FELs, Mercator projection); a. D- glucal, b. ddMan, and c. NOE, contoured at 1 kcal mol-1. Yellow star highlights conformation observed on-enzyme in complex with BxGH99.

4 The D-glucal ring of ManGlucal adopts a H5 conformation in the complex with

4 BxGH99. The FEL of D-glucal displays a main energy minimum centred at H5, and with only a minor energy penalty can also adopt the nearby 4E conformation predicted for the transition states leading to and from the proposed 1,2-anhydrosugar intermediate (Figure

2.13 a). Based on this data, D-glucal provides good shape mimicry of the intermediate of

4 the GH99 catalysed reaction. ManddMan adopts a C1 conformation in the complex with

BxGH99, which is also the lowest energy conformation in the FEL of ddMan (Figure 53

4 2.13b). The H5 conformation is much higher in energy, indicating that ManddMan cannot provide shape mimicry of the GH99 transition state; instead it appears to be a mimic of the conformation of the substrate in the Michaelis complex.

Given that NOE is a basic molecule it can be a neutral or a protonated species;

4 FELs for both forms were calculated. Neutral NOE has a minimum at C1 whereas that

1 1 for the protonated form is ring-flipped to C4; the S5 conformation is also a local minimum (Figure 2.13c). In the case of protonated NOE, the structure is dominated by

+ transannular hydrogen bonds between NH2 and O6 or O3, respectively. Such intramolecular hydrogen bonds are not expected to dominate the structure when bound to an enzyme as the protein supplies a large number of hydrogen bond donors and acceptors.

Thus, it was decided the FEL of the neutral species is a more accurate representation of

4 enzyme-bound conformations. Neutral NOE has a wide main minimum at C1, similar to

4 4 ddMan and neutral IFG. There is a 60° shift in ϕ in the transition state region H3/ E between the north pole and equator in NOE versus ddMan. This could be due to the

4 4 hydrogen bonding between 2-OH and 3-OH in NOE stabilizing the H3/ E conformations, which does not occur in ddMan or IFG as they do not have a 2-OH.

2.7 Conclusions

The FEL for D-glucal shows that this structure in the ground state provides mimicry of

4 4 the H5 intermediate conformation and also to some degree the proposed E transition state conformation. The 3D structures of BxGH99 with ManGlucal shows that it binds in

4 a H5 conformation, matching the proposed intermediate conformation. The dissociation constant for ManGlucal was modest suggesting that shape mimicry of the intermediate or transition state provides a weak affinity for the enzyme. The FEL for ddMan shows it

4 prefers a C1 conformation; the 3D structure of BxGH99 with ManddMan shows it to bind

4 in a C1 conformation. The dissociation constant for ManddMan is 8- and 18-fold worse for Bx- and BtGH99 enzymes, respectively, compared to ManGlucal. This data collectively represents the ability of shape-based inhibitors to provide transition state mimicry.

The best inhibitor reported prior to this work, ManIFG, mimics charge development at C1 of the transition state, yet lacks a 2-OH, a group proposed to be involved in the catalytic mechanism through interaction with E333. Accordingly,

ManNOE was synthesised and was shown to bind to Bx- and BtGH99 with KD values of

13 and 30 nM, a 17- and 5-fold enhancement of affinity for the respective enzymes compared to ManIFG. The 3D structure of ManNOE in complex with BxGH99 reveals a

4 C1 ground state conformation, mimicking the Michaelis complex. ManNOE is the most potent ligand of GH99 enzymes to date and we conclude that its high affinity derives in part from its ability to mimic the charge of the transition state, as well as achieve interactions similar to those involved in the catalytic mechanism.

2.8 Experimental Benzyl β-D-arabinopyranoside (202) Acetyl chloride (400 μL, 57.7 mmol) was added dropwise to a suspension of D-arabinose (2.03 g, 13.5 mol) and distilled benzyl alcohol (28 mL) at 35°C. The reaction was stirred vigorously at 40 ̊C for 18 h after which time product crystallized out of solution. The reaction was cooled to 25°C, diluted with ether (28 mL) and stirred for 0.5 h. The crude product was filtered and washed with ether. Recrystallization from 95% EtOH/H2O afforded benzyl β-D- arabinopyranoside (202)88 (2.51 g, 79 %), m.p. 172-173 ºC (lit (88) 172-173 ºC). 1H NMR (400 MHz, DMSO) δ 3.42-3.46, 3.59-3.70 (5 H, m, H2, H3, H4, 2 × H5), 4.43 (1 H, d, J

= 12.4 Hz, PhCH2), 4.50 (1 H, m, OH), 4.55-4.56 (1 H, m, OH), 4.60-4.62 (1 H, m, OH),

13 4.64 (1 H, m, PhCH2), 7.25-7.39 (5 H, m, Ph); C NMR (125 MHz, DMSO)  68.3, 68.4, 68.6, 69.1 (4 C, C2, 3, 4, 5), 98.9 (1 C, C1), 127.3, 127.4, 128.2, 138.2 (6 C, Ph).

Benzyl 2,3-O-isopropylidene-β-D-arabinopyranoside (204) p-Toluenesulfonic acid (328 mg, 1.9 mmol), was added to benzyl β-D- arabinopyranoside (202) (3.05 g, 12.7 mmol) and triethyl orthoacetate (3.65 mL, 25.4 mmol) in dichloromethane (50 mL) and the mixture stirred at 30 ˚C for 3 h. Triethylamine (0.15 mL, 1.9 mmol) was added and the mixture washed with water (3 × 20 mL), aq. NaHCO3, aq. brine, dried (MgSO4) and concentrated under reduced pressure. The residue was dissolved in AcOH/H2O (9:1, 20 mL) and the solution stirred at 40 ˚C for 0.5 h. The mixture was concentrated, azeotroped with toluene (3 × 20 mL) and was partially purified by flash chromatography (90:10, ethyl acetate/pet.spirits) to give a mixture of acetates colourless needles (3.08 g, 85%). p- Toluenesulfonic acid (27.6 mg, 0.16 mmol) was added to the diol (6.47 g, 22.9 mmol) and 2-methoxypropene (4.13 mL, 41.3 mmol) in CH2Cl2 (40 mL) and the mixture was stirred for 1 h. Et3N (22 µL) was added, the solution stirred 10 min. The mixture was washed with water (2 × 20 mL), aq. NaHCO3, aq. brine, dried (MgSO4) and concentrated under reduced pressure. The crude oil was purified by flash chromatography (EtOAc/pet. 86 1 sprits/Et3N 40:59:1) to afford the acetonide (204) (4.75 g, 74 %) as a yellow oil. H

NMR (500 MHz, CDCl3) δ 1.46 (3 H, s, CH3), 1.49 (3 H, s, CH3), 3.70 (1 H, dd, J = 1.5, 13 Hz, H5), 3.77 (1 H, dd, J = 1.5, 13, H5), 4.03-4.10 (2 H, m, H2, H3), 4.29 (1 H, m,

H4), 4.65 (1 H, d, J = 12.5 Hz, CH2Ph), 4.79 (1 H, d, J = 12 Hz, CH2Ph), 5.32 (1 H, d, J

13 = 2.5 Hz, H1), 7.29-7.37 (5 H, m, Ph); C NMR (125 MHz, CDCl3)  21.2, 21.3 (2 C,

CH3), 61.1, 68.8, 69.8, 70.0, 71.6 (5 C, C2, 3, 4, 5, CH2Ph), 98.5 (1 C, C1), 128.2, 128.2,

128.7, 137.0 (7 C, C(CH3)2 Ph).

Benzyl 2,3-O-Isopropylidene-4-O-(4-nitrobenzoyl)-α-L-xyloside (209)

Diisopropyl azodicarboxylate (260µL, 1.32 mmol), in PhMe (1 mL), was added dropwise to a solution of

alcohol (204) (132 mg, 0.47 mmol), Ph3P (346 mg, 1.32

mmol), and 4-O2NC6H4CO2H (220 mg, 1.32 mmol) in PhMe (2mL) at 0 ˚C. The resulting mixture was warmed (40 ˚C) and stirred 18h. Concentration of the mixture and purification by flash chromatography (EtOAc/pet. 86 spirits/Et3N 10:89:1) gave the 4-nitrobenzoate (209) (163 mg, 80%) as pale yellow 1 crystals. H NMR (500 MHz, CDCl3) δ 1.47 (3 H, s, CH3), 1.52 (3 H, s, CH3), 3.51 (1 H, apt. t, H5), 3.64 (1 H, dd, J = 3, 9.5 Hz, H2), 4.03 (1 H, dd, J = 5.5, 11 Hz, H5), 4.34 (1

H, atp. t, H3), 4.68 (1 H, d, J = 12 Hz, CH2Ph), 4.82 (1 H, d, J = 12 Hz, CH2Ph), 5.28 (1 H, d, J = 3.0, H1), 5.34 (1 H, m, H4), 7.30-7.41 (5 H, m, Ph), 8.21-8.29 (4 H, m, Ar); 13C

NMR (125 MHz, CDCl3)  26.7, 27.0 (2 C, CH3), 59.8, 70.2, 72.8, 73.7, 76.2 (C 2, 3, 4,

5, CH2Ph), 96.5 (1 C, C1), 111.3 (1 C, C(CH3)2), 123.7, 127.8, 128.1, 128.6, 131.2, 135.0, 137.2, 150.9 (12 C, Ph), 163.9 (1 C, C=O).

Benzyl 2,3-O-isopropylidene-β-D-zylopyranoside (206)

Triethylamine (200 µL) was added to the 4-nitrobenzoate (209) (156 mg, 0.36 mmol) in MeOH (2 mL) and stirred (40 ˚C, 0.5 h). The solution was concentrated and purified by flash chromatography

(EtOAc/pet. spirits/Et3N 25:74:1) to give the xylo-configured alcohol 86 1 (206) (92.7 mg, 91%) as a colourless oil. H NMR (500 MHz, CDCl3): δ 1.45 (6H, 2 s,

CH3), 2.58 (1 H, s, OH), 3.46 (1 H, dd, J = 5.4, 9 Hz, H5), 3.75 (1 H, dd, J = 6, 11 Hz,

H2), 3.97-4.04 (2 H, m, H3, H5), 4.63 (1 H, d, J = 12 Hz, CH2Ph), 4.79 (1 H, dd, J = 12 13 Hz, CH2Ph), 5.20 (1 H, d, J = 3 Hz, H1), 5.31 (1 H, m, H4), 7.29-7.38 (5 H, m, Ph); C

NMR (125MHz, CDCl3)  26.6, 27.0 (2 C, 2 × CH3), 62.9, 69.8, 70.3, 75.9, 77.3 (5 C,

C2, 3, 4, 5, CH2Ph), 96.4 (1 C, C1), 110.9 (1 C, C(CH3)2), 127.7, 127.9, 128.5, 137.5 (6 C, Ph).

Benzyl 2,3-O-isopropylidene, 4-cyano-4-deoxy-β-D-arabinopyranoside (211) Triflic anhydride (1.82 mL, 10.6 mmol) was added to a suspension of the

alcohol (206) (2.42 mg, 8.65 mmol) in CH2Cl2 (30 mL) at -60 ̊ C. The mixture was heated to 0 °C and stirred for 30 minutes. The mixture was concentrated under reduced pressure then dissolved in DMF (20 mL). KCN (1.86 g, 28.5 mmol) was added to the mixture and stirred for 18 h. The mixture was washed with water (3 × 20 mL), aq. NaHCO3 (3 × 20 mL), aq. brine, dried (MgSO4) and concentrated under reduced pressure. The crude was purified by flash chromatography (EtOAc/pet. spirits 20:70) to give the nitrile (211)86 (1.18 g, 47%) over two steps. 1H

NMR (500 MHz, CDCl3) δ 1.48 (6 H, s, 2 × CH3), 3.29 (1 H, s, H4), 3.76 (1 H, dd J = 2.0, 10.1 Hz, H5), 3.86-3.92 (2 H, m, H3, 5), 4.10 (1 H, dd, J = 3, 10 Hz, H2), 4.65 (1 H, d, J = 12.1 Hz, CH2Ph), 4.75 (1 H, d, J = 12.1 Hz, CH2Ph), 3.34 (1 H, d, J = 2.0 Hz, H1),

13 7.26-7.34 (5 H, m, Ph); C NMR (125 MHz, CDCl3)  26.5, 26.5 (2 C, 2 × CH3), 34.1 (1

C, C4), 46.2, 59.3, 60.2, 74.5 (4 C, C2, 3, 5, CH2Ph), 97.6 (1 C, C1), 110.8 (1 C, C(CH3)2) , 117.0 (1 C, CN), 127.6, 127.9, 128.4, 136.9 (6 C, Ph).

Benzyl 4-cyano-4-deoxy-β-D-arabinopyranoside (212)

The nitrile (211) (71.3 mg, 0.250 mmol) was treated with TFA/H2O (9:1, 200 µL) for 0.5 h. The solution was concentrated and azeotroped with toluene (3 × 5 mL) to give the diol (212) (52mg) without further 1 purification. H NMR (500 MHz, CDCl3) δ 3.11 (1 H, m, H4), 3.46-3.78 (2 H, m, H2, 5), 3.84 (1 H, dd, J = 2, 10 Hz, H5), 4.00 (1 H, dd, J = 4, 7.5 Hz, H3), 4.50 (1 H, d, J = 9.5

Hz, CH2Ph), 4.70 (1 H, d, J = 9.5 Hz, CH2Ph), 4.95 (1 H, d, J = 3 Hz, H1), 7.30-7.36 (5

13 H, m, Ph); C NMR (125 MHz, CDCl3)  35.7 (1 C, C4), 59.0, 67.1, 70.2, 70.9 (5 C, C2,

3, 5, CH2PH), 98.0 (1 C, C1), 118.5 (1 C, CN), 128.3, 128.3, 128.7, 128.8, 136.8 (6 C, Ph).

1,2,3,4,6-Penta-O-acetyl-α-D-mannopyranose (213)

Sulfuric acid 98% (10 μL) was added to a suspension of D-mannose (12.0 g, 66.7 mmoL) in acetic anhydride (50 mL) at 0 ̊C. The suspension was stirred for 30 m at room temperature then poured into ice water. The mixture was washed with aq. H2O (3 × 20.0 mL), aq. NaHCO3 (3 ×

20.0 mL), dried (MgSO4) and concentrated under reduced pressure to give the perbenzylated mannose (213) 94 (20.9 g, 80.3%) as the α-anomer. 1H NMR (400 MHz,

CDCl3) δ 2.00 (3 H, s, CH3), 2.08 (3 H, s, CH3), 2.09 (3 H, s, CH3), 2.16 (3 H, s, CH3),

2.17 (3 H, s, CH3), 4.03-4.06 (1 H, m, H5), 4.08-4.23 (1 H, m, H6a), 4.27 (1 H, dd, J 5,6a

= 3.9 Hz, J6a,6b, = 9.9 Hz, H6b), 5.29-5.35 (3 H, m, H2, 3, 4), 6.08 (1 H, d, J 1,2 = 1.4 Hz,

13 H1α); C NMR (100 MHz, CDCl3)  20.4, 20.5, 20.6, 20.7 (5 C, CH3), 61.9, 65.3, 68.1, 68.5, 70.4 (5 C, C2, 3 ,4 ,5, 6), 90.3 (1 C, C1), 167.9, 168.0, 169.5, 169.7, 169.9 (5 C, C=O).

3,4,6,-Tri-O-acetyl-1,2-O-(1-methoxyethylidene)-β-D-mannopyranose (215) HBr/AcOH 33% (1.6 mL) was added to perbenzylated mannose (213) (1.48 g, 3.79 mmol) at 0 ̊ C. The reaction was stirred at room temperature for 3 h then diluted with 20 mL

EtOAc. The mixture was poured into aq. NaCl (20 mL), washed with aq. NaHCO3 (3 ×

20 mL), dried (MgSO4) and concentrated under reduced pressure to give the bromide. Pyridine (3 mL) was added to a suspension of bromine (1.12 g, 2.2 mmol) in MeOH (1.5 mL) and stirred for 18h. The mixture was poured into ice water, washed extracted into dichloromethane (3 × 20 mL), dried (MgSO4) and concentrated under reduced pressure 94 1 to give the orthoester (215) (0.811 g, 52%). H NMR (400 MHz, CDCl3) δ 1.73 (3 H, s, Me), 2.05 (3 H, s, OMe), 2.07 (3 H, s, OMe), 2.11 (3 H, s, OMe), 3.27 (3 H, s, OMe),

3.65-3.70 (1 H, m, H5), 4.14 (1 H, dd, J5,6a = 2.8 Hz, J6a,6b = 12.2 Hz, H6a), 4.23 (1 H, dd, J5,6b = 4.8, J6a, 6b = 12.1Hz, H6b), 4.60-4.62 (1 H, m, H4), 5.14 (1 H, dd, J2,3= 4.0, J3,4 13 = 9.9 Hz, H3), 5.25-5.31 (1 H, m, H2), 5.47 (1 H, s, J1,2 = 2.6 Hz, H1); C NMR (100

MHz, CDCl3)  20.2, 20.3, 20.4, 24.4, 49.9 (5 C, 4 × MeCO, OMe, Me), 63.4, 65.5, 71.2, 71.5, 76.2 (5 C, C2, 3, 4, 5, 6), 96.5 (1 C, C1), 125.4 (1 C, C(OMe)Me), 170.1,170.5, 170.9 (3 C, 3 × C=O).

3,4,6,-Tri-O-benzyl-1,2-O-(1-methoxyethylidene)-β-D-mannopyranose (217) Sodium metal (catalytic) was added to a suspension of orthoester (215) (3.95 g, 10.9 mmol) in MeOH (40 mL) at 0 ̊C and stirred for 30 m. The crude was concentrated under reduced pressure. NaH (1.96 g, 49.1 mmol) was added to a solution of crude in DMF (30 mL) at 0⁰C and stirred for 30 m. BnBr (4.66 mL, 39.2 mmol) was added and stirred for 18 h. The mixture was poured into ice water, dried (MgSO4), extracted into EtOAc (3 × 20 mL) and concentrated under reduced pressure. The crude was purified by flash chromatography 94 (EtOAc/pet. spirits/Et3N 25:74:1) to give the benzyl protected orthoester (217) (3.44 g, 1 62% over two steps). H NMR (400 MHz, CDCl3) δ 1.75 (3 H, s, Me), 3.29 (3 H, s, MeO),

3.41-3.46 (1 H, m, H5), 3.70-3.78 (3 H, m, H4, 2 × H6), 3.93 (1 H, apt. t, H3), 4.40-4.41

(1 H, m, H2), 4.54-4.63 (3 H, m, 3 × CH2Ph), 4.79 (2 H, apt. s, 2 × CH2Ph), 4.91 (1 H, dd, J = 13.5 Hz, CH2Ph), 5.35 (1 H, dd, J1,2 = 2.6 Hz, H1), 7.23-7.42 (15 H, m, 3 × Ph);

13 C NMR (100 MHz, CDCl3)  25.2 (1 C, Me), 48.9 (1 C, OMe), 67.9, 68.2, 68.7, 72.4,

72.9, 73.1, 74.5, 76.3, 76.8 (9 C, C2, 3, 4, 5, 6, 3 × CH2Ph, C(OMe)Me), 98.2 (1 C, C1), 124.4, 126.9, 127.2, 127.3, 127.5, 128.2, 128.5, 128.6, 136.5, 136.7 (18 C, Ph).

2-O-Acetyl-3,4,6-tri-O-benzyl-α-D-mannopyranosyl trichloroacetimidate (219)

A solution of orthoester (217) (3.41 g, 6.73 mmol) in TFA/H20 9:1 (3.4 mL) was stirred for 30 mins. The crude was asioptroped with toluene (3 × 20 mL) and concentrated under reduced pressure without any further purification to give the diol (3.31

g, 6.73 mol). NCCCl3 (1.6 mL, 15.8 mmol) and DBU (24 L,

1.58 mmol) were added to a suspension of diol (1.56 g, 3.17 mmol) in CH2Cl2 (30 mL) at 0 ̊C. The mixture was stirred at room temperature for 2h. The crude was concentrated under reduced pressure and purified by flash chromatography (EtOAc/pet. spirits/Et3N 30:69:1) to give the TCA donor (219)94 (1.91 g, 95%) as a colourless oil. 1H NMR (500

MHz, CDCl3) δ 2.23 (3 H, s, Ac), 4.76 (1 H, dd, J = 1.5, 11 Hz, H6), 3.89 (1 H, dd, J =

4, 11.5 Hz, H6), 4.03-4.11 (3 H, m, H3, 4, 5), 4.54-4.80 (5 H, m, 5 × CH2Ph), 4.93 (1 H, d, J = 10.5 Hz, CH2Ph), 5.58 (1 H, m, H2), 6.37 (1 H, d, J = 2 Hz, H1), 7.40-7.22 (15 H,

13 m, 3 × Ph), 8.74 (1 H, s, NH); C NMR (125 MHz, CDCl3)  21.2 (1 C, MeCO), 66.9,

67.3, 71.5, 73.5, 73.8, 74.4, 75.1, 77.5 (8 C, C2, 3, 4, 5, 6, 3 × CH2Ph) 94.8, 107.9 (2 C,

C, CCl3 ), 127.5-140.3 (18 C, 3 × Ph,), 170.2 (1 C, C=O).

Benzyl 2-O-(2-O-acetyl-3,4,5-tri-O-benzyl-α-mannopyranosyl)-(1→2)-(4-cyano-4- deoxy-β-D-arabinopyranoside) (220) TfOH (4.4 µL, 0.050 mmol) was added to a mixture of acceptor (212) (125 mg, 0.50 mmol), 2-O-acetyl-3,4,5-tri-O- benzyl-α-mannopyranosyl trichloroacetimidate (219) (372 mg,

0.625 mmol) and 4 Å mol. sieves in CH2Cl2 at -40 ˚C. The mixture was stirred for 15 min, warmed to 0 ˚C and quenched with Et3N (7.0 µL, 5.07 mmol then concentrated under reduced pressure. Flash chromatography (EtOAc/pet. 23 spirits 40:60) gave the disaccharide (220) (168 mg, 46%) as a colourless oil; [α]D –44, 1 (c 0.9, CHCl3); H NMR (500 MHz, CDCl3) δ 1.21 (3 H, s, Ac), 3.17-3.18 (1 H, m, H4),

3.32 (1 H, s, OH), 3.5 (1 H, dd, J5',6a' = 6.5, J6a',6b' = 10.5 Hz, H6a'), 3.60 (1 H, dd, J5',6b' =

1.5, J6a',6b' = 10 Hz, H6b’), 3.67-3.96 (6 H, m, H2, 3', 4', 5a, 5b, 5'), 4.18-4.22 (1 H, m,

H3), 4.44-4.61 (6 H, m, 6 × CH2Ph), 4.69 (1 H, dd, J = 11 Hz, CH2Ph), 4.84 (1 H, dd, J

= 11 Hz, CH2Ph), 5.18 (1 H, dd, J1,2 = 3.5 Hz, H1), 5.23 (1 H, d, J1',2' = 1.5 Hz, H1'), 5.39- 13 5.40 (1 H, m, H2'), 7.14-7.32 ppm (20 H, m, Ph); C NMR (125 MHz, CDCl3) δ 21.1 (1 C, Ac), 36.5 (1 C, C4), 58.5 (1 C, C5), 64.3 (1 C, C3), 68.7, 69.3, 69.8, 72.1, 72.1, 73.7,

74.4, 74.9, 78.1, 78.9 (10 C, C2, 2', 3', 4, 5', 6', 4 × CH2Ph), 97.9 (1 C, C1), 99.1 (1 C, C1'), 127.6, 127.8, 127.9, 128.1, 128.2, 128.3, 128.4, 128.5, 128.7 ppm (24 C, 4 × Ph); + + HRMS (ESI) m/z 741.3401 [C42H45NO10 (M+NH4 ) requires 741.3382].

Benzyl 2-O-(2-O-acetyl-3,4,5-tri-O-benzyl-α-mannopyranosyl)-( 1→2)-(3-O-acetyl-

4-cyano-4-deoxy-β-D-arabinopyranoside) (221)

A mixture of acetic anhydride (250 µL) and disaccharide (220) (20 mg, 0.028 mmol) in pyridine (500 µL) was stirred at r.t. for 18 h. The solution was concentrated under reduced pressure and the residue was azeotroped with toluene (3 × 10 mL). Flash chromatography (EtOAc/pet. spirits 45:55) of the residue gave 24 1 the title compound (221) (10.8 g, 49%) as a colourless oil; [α]D –66 (c 0.29, CHCl3); H

NMR (500 MHz, CDCl3) δ 2.13 (3 H, s, Ac), 2.19 (3 H, s, Ac), 3.41-3.42 (1 H, m, H4),

3.50 (1 H, dd, J5',6a' = 1.5, J6a',6b' = 10.5, H6a'), 3.55-3.70 (2 H, m, H5', 6b'), 3.78-3.82 (2

H, m, H4', 5a), 3.90 (1 H, dd, J2',3' = 3.5, J3',4' = 9.4 Hz, H3'), 3.97-4.03 (2 H, m, H2, 5b),

4.45-4.67 (7 H, m, CH2Ph), 4.86 (1 H, d, J = 10.9 Hz, CH2Ph), 5.02 (1 H, d, J1',2' = 1.5

Hz, H1'), 5.08 (1 H, d, J1,2 = 3.6 Hz, H1), 5.21 (1 H, dd, J2,3 = 5.5, J3,4 = 10.4 Hz, H3), 13 5.24-5.25 (1, m, H2), 7.12-7.37 ppm (20 H, m, 4 × Ph); C NMR (125 MHz, CDCl3) δ 20.9 (1 C, Ac), 21.2 (1 C, Ac), 34.5 (1 C, C4), 58.2 (1 C, C5), 67.7 (1 C, C3), 68.9 (2 C,

C2', CH2Ph), 70.7 (1 C, CH2Ph), 72.1 (1 C, CH2Ph), 72.4 (1 C, C5'), 73.7 (1 C, CH2Ph),

74.3 (1 C, C4'), 74.9 (1 C, C2), 75.1 (1 C, CH2Ph), 77.9 (1 C, C3'), 97.2 (1 C, C1), 100.0 (1 C, C1'), 127.7, 127.9, 128.1, 128.2, 128.3, 128.4, 128.5, 128.6, 128.8 ppm (24 C, 4 × + + Ph); HRMS (ESI) m/z 788.3063 [C44H47NaO11 (M+Na) requires 788.3041]. The downfield shift of H3' relative to that in compound 220 provided evidence for the regiochemistry of the glycosylation reaction.

Benzyl 2-O-(3,4,5-tri-O-benzyl-α-mannopyranosyl)-(1→2)-(4-cyano-4-deoxy-β-D- arabinopyranoside) (222)

Acetyl chloride (860 µL) was added to disaccharide (220) (127 mg, 0.175 mmol) in MeOH (20 mL) and the mixture was

stirred for 18 h at rt. The reaction was quenched with Et3N (1.67 mL, 12 mmol) and the solvent evaporated under reduced pressure. Flash chromatography (EtOAc/pet. spirits 23 50:50) of the residue gave the diol (222) (84 mg, 70 %) as a colourless oil; [α]D –62, (c 1 2.26, CHCl3); H NMR (500 MHz, CDCl3) δ 2.59 (1 H, s, OH), 3.15 (1 H, m, H4), 3.30

(1 H, s, OH), 3.48, (1 H, dd, J5',6a' = 5.4, J6a',6b' = 8.6 Hz, H6a’), 3.57 (1 H, dd, J5',6b' = 1.6,

J6a',6b' = 8.5 Hz, H6b'), 3.69 (1 H, t, H4), 3.78 (1 H, dd, J4,5a = 1.5, J5a,5b = 10.0 Hz, H5a),

3.83-5.85 (3 H, m, H2, 3', 5b), 3.93 (1 H, m, H5'), 3.99 (1 H, m, H2'), 4.19 (1 H, dd, J3,4

= 4.6, J2,3 = 8.2 Hz, H3), 4.45-4.67 (7 H, m, CH2Ph), 4.80 (1 H, d, J = 9.3 Hz, CH2Ph),

5.20 (1 H, d, J1,2 = 3.0 Hz, H1), 5.27 (1 H, d, J1',2' = 1.5 Hz, H1'), 7.23-7.33 ppm (24 H, 13 m, 4 × Ph); C NMR (125 MHz, CDCl3) δ 36.4 (1 C, C4), 58.5 (1 C, C5), 64.9 (1 C, C3),

68.5, 69.3, 69.8, 71.8, 72.5, 73.6, 74.4, 74.8, 78.8, 79.8 (10 C, C2,2',3',4',5',6',4 × CH2Ph), 98.0 (1 C, C1), 100.6 (1 C, C1'), 127.7, 127.8, 1280, 128.1, 128.4, 128.5, 128.7 ppm (20 + + C, 4 × Ph); HRMS (ESI) m/z 699.3304 [C40H43NO9 (M+NH4) requires 399.3276].

Benzyl 2-O-(2-O-acetyl-2,4,5-tri-O-benzyl-α-mannopyranosyl)-(1→2)-(4-C-[(tert- butoxycarbonyl)amino]methyl-4-deoxy-β-D-arabinopyranoside) (223)

Borane dimethyl sulfide complex (35.7 µL, 0.379 mmol) was added dropwise to nitrile (220) (57.1 mg, 0.0838 mmol) in anhydrous THF (3 mL), and the mixture was stirred under reflux for 24 h. MeOH (3 mL) was added dropwise and reflux continued for an additional 18 h. The solvent was evaporated under reduced pressure and the residue was azeotroped with MeOH (3 × 20 mL) and evaporated to dryness under reduced pressure. The residue was dissolved in THF/MeOH (1:3, 12 mL) and treated with 10% aq. Na2CO3 (2 mL) and (Boc)2O (36.6 mg, 0.168 mmol). After 2 h the mixture was filtered and the filtrate concentrated under reduced pressure. Flash chromatography (EtOAc/pet. spirits 50:50) of the residue gave the carbamate (223) (44.7 23 1 mg, 68 %) as a colourless oil; [α]D –26 (c 1.2, CHCl3); H NMR (500 MHz, CDCl3) δ

1.45 (9 H, s, 3 × CH3), 2.11 (1 H, m, H4), 2.60 (1 H, m, OH), 3.24 (1 H, m, H6a'), 3.39-

3.48 (4 H, m, H5a, 6b', CH2NHBoc), 3.56 (1 H, dd, J4,5b = 4.5, J5a,5b = 11 Hz, H5b) , 3.76-

4.15 (6 H, H2, 2', 3, 3', 4', 5’), 4.41-4.80 (8H, m, 4 x CH2Ph), 5.01 (2 H, m, H1', NH), 13 5.15 (1 H, s, H1), 7.15-7.35 (20 H, m, 4 × Ph); C NMR (125 MHz, CDCl3): δ 28.5 (3

C, CH3), 38.1 (1 C, C6'), 40.7 (1 C, C4), 61.4 (C5), 66.9, 68.8, 69.0, 69.8, 71.3, 72.3,

73.6, 74.4, 74.7, 76.3, 79.9, 80.1 (10 C, C2, 2', 3, 3', 4', 5', CH2NHBoc, 4 × CH2Ph), 98.0 (1 C, C1), 99.8 (1 C, C1'), 127.6, 127.7, 127.8, 127.9, 128.0, 128.2, 128.3, 128.4, 128.5, + + 128.6 (24 C, 4 × Ph); HRMS (ESI) m/z 803.4132 [C45H55NO11 (M+NH4) requires 803.4113].

α-D-Mannopyranosyl-(1→3)-noeuromycin hydrochloride (ManNOE.HCl;201)

The carbamate (223) (21.7 mg, 00276 mmol) in THF/H2O (1:1,

5 mL) was treated with Pd(OH)2/C (10%, 10 mg) and H2 (15 atm, 18 h). The suspension was filtered, concentrated and

subject to flash chromatography (CHCl3/MeOH/H2O 70:27:1) to give a colourless oil. The oil was treated with 1 M aq. HCl (2 mL) and stirred for 30 min. The solution was concentrated under reduced pressure to give ManNOE.hydrochloride (201) as a colourless residue. NMR data of the noeuromycins (‘glc' has an equatorial 2-OH, ‘man' has an axial 2-OH, ‘p-glc' and ‘p-man' are the 25 1 pyranose tautomers); [α]D +38, 0.5 M; H NMR (500 MHz, D2O) δ 1.95-2.05 (m, H5- man, H5-glc), 2.96-3.94 (m, H2', 3, 3', 4, 4', 5', 6a, 6b, 6a', 6b'), 4.07 (dd, J1',2' = 1.5, J2',3'

= 3 Hz, H2'-glc), 4.09 (dd, J1',2' = 1.5, J2',3' = 3.5 Hz, H2'-man), 4.66 (m, H2-p-glc), 4.72

(m, H2-glc), 5.04 (d, J1',2' = 1.5 Hz, H1'-p-man), 5.12 (d, J1',2' = 1.5 Hz, H1'-man), 5.18

(d, J1',2' = 1.5 Hz, H1'-p-glc), 5.24 (d, J1',2' = 1.5 Hz, H1'-glc), 5.36 (d, J2,3 = 3.5 Hz, H2- 13 p-man), 5.43 ppm (d, J2,3 = 3.5 Hz, H2-man); C NMR (125 MHz, D2O) δ 37.3, 38.1, 40.7, 40.7, 40.9, 41.2, 55.4, 58.6, 58.8, 59.0, 60.6, 60.8, 60.9, 65.9, 66.3, 66.4, 66.5, 66.7, 69.7, 69.9, 70.0, 70.2, 70.3, 70.4, 72.3, 72.9, 73.2, 73.5, 77.4, 78.9, 80.0, 80.2, 91.8, 101.4. + + 101.6, 102.5 ppm; HRMS (ESI) m/z 326.1444 [C12H35NO9 (M+ NH4) requires 326.1446].

CHAPTER THREE:

Exploration of strategies for charge mimicry in

inhibitor design for family GH99 endo-α-

mannanases

3.1 Importance of charge mimicry for binding to GH99 enzymes

Chapter 2 explored the effect of charge versus shape mimicry to develop tight-binding ligands for GH99 enzymes. The most tightly binding ligands against bacterial BtGH99,

GlcDMJ (KD = 24 µM), GlcIFG (0.63 M), ManIFG (0.14 M) and ManNOE, 201 (30

4 nM) all bind in a ground state C1 conformation, mimicking the Michaelis complex. This data suggests potent inhibition can be achieved by mimicry of charge development in the transition state. We attribute the potency of ManNOE (201), the strongest binding ligand to date, to a favourable hydrogen bonding interaction between the 2-OH of the NOE ring and active site residue E333 (Figure 3.1).

Figure 3.1: Ligand binding to BxGH99 involving catalytic residues E333 and E336. Interatomic distances shown in red. a. ManIFG; b. ManNOE; c. GlcDMJ.

3.2 Aims

Based on the role of the 2-OH in the proposed 1,2-anhydrosugar mechanism of GH99 enzymes, and the interaction seen between the active site E333 residue and the 2-OH, we speculated that introduction of a 2-deoxy-2-amino group could promote a favourable ionic interaction between this nitrogen and E333 upon binding of the inhibitor. However, we deemed that installing a 2-NH2 group onto NOE would give rise to an unstable aminal that would hydrolyse to lose ammonia, and chose to work with DMJ (Figure 3.2).

Therefore, we proposed to synthesize α-mannosyl-1,3-(2-amino-1,2-

dideoxymannojirimycin) (Man2NH2DMJ, 301). With this compound in hand, we would investigate its affinity for endo-α-1,2-mannanase and obtain a three-dimensional structure of its complex to characterize its binding mode.

Figure 3.2: a. Proposed mechanism for family GH99 retaining endo-α-1,2- mannosidases/endo-α-1,2-mannanases. Only the first half of the catalytic cycle is shown; b. Proposed mode of 2NH2DMJ binding to Bt or BxGH99.

3.3 Prior work with DMJ and NH2DMJ

The iminosugars, DMJ and 2NH2DMJ, have been synthesized and their potency evaluated against several different glycosidases including: an α-glucosidase (from

Bacillus stearothermophilus), a β-glucosidase (from almonds), an α-mannosidase (from jack bean) and an α-L-fucosidase (from bovine kidney). For the α-glucosidase, DMJ (KI

105 = 18 µM) was 3-fold more potent than NH2DMJ (KI = 53 µM). However, in the case of the α-mannosidase DMJ (KI = 600 µM) was 30-fold less potent than 2NH2DMJ (KI =

20 µM). Based on these findings, we anticipated that Man2NH2DMJ (Figure 3.3b) could to be a more potent inhibitor of family GH99 α-mannanases compared to GlcDMJ (Figure

3.3a).

 ManDMJ would be a better comparison, however it had not been synthesized and studied. Additionally, given the preference of Bx and BtGH99 enzymes for a D-manno-configuration in the -2 subsite, ManDMJ would be expected to be around 4-fold better an inhibitor than GlcDMJ. 67

Figure 3.3: a. GlcDMJ; b. Man2NH2DMJ (301)

3.4 Synthetic strategy towards Man2NH2DMJ

Man2NH2DMJ was be synthesized by coupling, a mannosyl donor and DMJ-based acceptor, followed by, global deprotection will give the title compound (301). We planed to use the trichloroacetimidate (219) from Chapter 2; we therefore needed to identify a suitable precursor to the 2NH2DMJ fragment, set up with appropriate amino and hydroxyl group protection.

There are four syntheses of 2NH2DMJ reported, all of which use DNJ as a key intermediate.104–107 Broadly, the most practical convert DNJ to a 2-azido species, which is reduced to give the desired amine. The most relevant to our synthetic approach was disclosed by Khanna and co-workers (Scheme 3.1).104 According to their approach, DNJ was protected with a 4,6-O-benzylidene acetal. Stannylene acetal-mediated regioselective tosylation at C2 followed by treatment with base formed a 2,3-anhydro derivative, and opening of the epoxide gave a 1:1 mixture of gluco- and manno-configured azides; the latter of which would be ideal as an alcohol acceptor for glycosylation with donor 219, en route to Man2NH2DMJ. However, as we only require the D-manno-configured azide there is considerable wastage through formation of the unwanted D-gluco diastereomer.

104 Scheme 3.1: Synthesis of a protected 2NH2DMJ from DMJ by Khanna et al.

Inspired by Khanna and co-worker’s synthesis of 2NH2DMJ we planned to develop a stereoselective synthesis of the DMJ-derived alcohol and apply it in the synthesis of Man2NH2DMJ, as broadly outlined in Scheme 3.2.

Scheme 3.2: Synthetic strategy for synthesis of Man2NH2DMJ. Glycosylation of DMJ- derived alcohol by mannosyl TCA will afford a disaccharide. One pot azide reduction and global deprotection will give Man2NH2DMJ (301).

3.5 Results and Discussion

Synthesis of the NH2DMJ acceptor

DNJ was synthesized in 6 steps from commercially available 2,3,4,6-tetra-O-benzyl-D- glucopyranose using established approaches (Scheme 3.3). One pot oxidative amidation of 2,3,4,6-tetra-O-benzyl-D-glucopyranose was achieved by slow addition of I2 to a suspension in aqueous ammonia/THF, affording the amide 302.108 Albright-Goldman

oxidation of 303 with acetic anhydride in DMSO, followed by reductive lactamization with sodium cyanoborohydride and formic acid, gave the lactam 304 in an uninspiring yield of 28% over 2 steps.109

Scheme 3.3: Improved synthesis of lactam (304). Reagents and conditions: a. NH3 (30%), I2, THF, 89%; b. Ac2O, DMSO; c. NaCNBH3, HCO2H, CH3CN, 28% over 2 steps.

The yield of these oxidation/reductive amidation steps was increased by oxidation of 302 with pyridinium dichromate in CH2Cl2 to give 303 in quantitative yield (Scheme

3.4).110 Ketone 303 was treated with methanolic ammonia to give a mixture of D-gluco- and L-ido-hydroxylactams (305). ‘Ionic hydrogenation’ of the hydroxylactam mixture with sodium cyanoborohydride and formic acid gave the lactam 304 in 77% yield over 2 steps.109

Scheme 3.4: Synthesis of lactam (304). Reagents and conditions: a. PDC, CH2Cl2; b. NH3, MeOH; c. NaCNBH3, CH2O2, CH3CN, 77% over 3 steps.

The stereoselectivity of the reduction of the mixture of epimers 305 has been rationalized by Overkleeft and co-workers by consideration of molecular orbital overlap in the intermediate acyliminium ion (Scheme 3.5). Protonation and ionization of the hydroxylactams (305) leads to an intermediate acyliminium ion that is susceptible to

hydride attack. NaCNBH3 delivers hydride preferentially from the β-face of the ring resulting in a conformation where the nitrogen electron-pair maximizes overlap with the orbitals of the lactam carbonyl group, forming the D-gluco-lactam 304.

Scheme 3.5: Overkleeft’s rationale for preferential formation of the D-gluco-lactam (304).

An alternative explanation for the reaction stereoselectivity can be advanced by considering the Newman projection of the acyliminium ion (Figure 3.4). Attack of hydride from the β-face is hindered by a the 1,2-diaxial interaction with H4, additionally it would require a large movement of C6, eclipsing the C4 in its path. Attack of the hydride from the α-face is sterically more favourable and requires less movement of C6 generating the D-gluco-configured lactam.

Figure 3.4: Newman projection of acyliminium ion showing stereochemical outcomes of α- versus β-face hydride attack.

Lactam in hand, efforts then turned to the synthesis of the acceptor (Scheme 3.6).

The lactam (304) was reduced with borane dimethyl sulfide complex to give the fully protected DMJ (306).97,111 Hydrogenation of 306 with Pearlman’s catalyst and 10% HCl at 30 bar gave DNJ (307) in quantitative yield. Having obtained this key intermediate we were now able to consider the installation of appropriate protecting groups to allow for

C2-inversion and installation of the azide, and to provide a free 3-OH for subsequent glycosylation. Protection of the amine with benzyl chloroformate gave the Cbz-protected

DNJ (308) in 89% yield. The 4- and 6-positions were protected as a benzylidene acetal by treatment of 308 with benzaldehyde dimethyl acetal and catalytic TsOH, affording the benzylidene acetal (309) in 45% yield.104 A higher yield of this reaction (77%) was obtained by performing the reaction at 60 °C and 300 mbar, removing methanol as it was produced.

Scheme 3.6: Synthesis of protected DMJ (309). Reagents and conditions: a. (CH3)2SBH3, THF, 77%); b. H2, Pd(OH2)/C, 10% HCl, EtOAc:MeOH:H2O 2:2:1, >99%; c. CbzCl, NaHCO3, DMF, 89%; d. PhCH(OMe)2, TsOH, DMF, 77%.

With compound 309 in hand we focused our efforts to invert the stereocentre at

C2 and install the azide. While we considered using the stannylene acetal approach employed by Khanna and co-workers (Scheme 3.1);107 a simpler approach by Tanaka and co-workers attracted our attention.112 In this approach, regioselective mesylation was performed on a molecule similar to 309, by the slow, dropwise addition of mesyl chloride at low temperature. These authors attributed the selectivity for mesylation at the 2- position to the bulky nature of the benzylidene ring hindering reactivity at the 3-hydroxyl.

In our case the lack of a substituent at C1 should provide even greater selectivity for the

2-position. Accordingly, we treated a solution of 309 in pyridine at -40°C with mesyl chloride added dropwise, yet were disappointed to discover that upon purification the major products were an equimolar mixture of the 2-mesylate and the 3-mesylate and as well as trace amounts of dimesylate (Scheme 3.7). Our efforts to separate the mixture were unsuccessful, so nevertheless we progressed, hopeful that it would be separable at the next step. Treatment of the mixture of mesylates with sodium azide in DMF led to successful substitution, affording the azide-substituted products in a combined 46% yield, yet the mixture 2-N3 and 3-N3 configured azides remained inseparable. Thus, reluctantly, we returned to Khanna’s stannylene acetal approach.

Scheme 3.7: Synthesis of azide. Reagents and conditions: a. MsCl, pyridine, 48%; b. NaN3, DMF, 46%.

Treatment of a diol with dibutyl tin oxide in toluene under Dean-Stark conditions results in the formation of a stannylene acetal, which is reactive towards acylating agents.

The tin coordinates to the diol forming a 5-coordinate monomeric tin complex. However, these undergo multimerization, resulting in higher coordination at Sn. Certain stannylene acetals exist as dimers in which the monomeric units are joined through Sn-O bonds, in a parallelogram fashion, resulting in a trigonal bipyramidal geometry at Sn (Figure 3.5).

Around the trigonal bipyrimidal Sn atom within the dimer one alkoxy group coordinates to two Sn atoms, and is arranged to be apical to one Sn atom and equatorial to the other.

The other alkoxy group is coordinated to a single Sn atom and lies apical; it is this oxygen

that is regioselectively acylated. The nucleophilicity of this atom is possibly a result of electron ‘channelling’ from the Sn atom, preferentially activating this position over the equatorially orientated alkoxy group. 113

Figure 3.5: a. Proposed stannylene acetal dimer for regioselective alkylation with dibutyltin oxide; b. Intermediate proceeds through a trigonal bipyramidal tin complex where the 2-O is bound to the apical position on the tin and the 3-O is bound to equatorially to the tin center.

Compound 309 was refluxed with dibutyl tin oxide in methanol to form the stannylene acetal. The methanol was evaporated and mixture azeotroped with toluene then concentrated to dryness. The crude residue was re-suspended in toluene and treated with tosyl chloride to give the equatorial 2-tosylate (313) in 91% yield. With this leaving group installed, 309 was treated with sodium azide, inverting the stereocentre to give the

D-manno-configured azide (314) in 74% yield.

Scheme 3.8: Synthesis of azide (314). Reagents and conditions: a. i. Bu2SnO, MeOH, ii. TsCl, PhMe, 91%; b. NaN3, DMF, 74%.

Glycosylation, reduction and global deprotection to give Man2NH2DMJ

A mixture of 314 and 207 was treated with catalytic triflic acid to give the α-1,3-linked disaccharide (315) in 87% yield (Scheme 3.9). 1H NMR spectroscopy was used to confirm identity of compound 315; some key spectral features include: two doublets of the

mannosyl group at  5.28 and 5.59 ppm assigned to be H1 and H2 respectively; a doublet at  5.12 ppm assigned as the CH2 of the Cbz; a singlet at  5.64 ppm assigned as the CH of the benzylidene acetal.

Scheme 3.9: Glycosylation. Reagents and conditions: a. TfOH, CH2Cl2, -40 °C, 87%.

With disaccharide (315) in hand we required a strategy to reduce the azide and remove all the remaining protecting groups. We decided to first remove the acetate;

Zemplen deprotection of 315 afforded the alcohol (316) (Scheme 3.10). From here we explored a simultaneous global deprotection and reduction of the azide, both of which could potentially be achieved though hydrogenation over a Pd catalyst. Compound 316 was treated with Pearlman’s catalyst and H2 with 10% HCl at 35 bar for 96 h; however, no Man2NH2DMJ or identifiable species were observed by mass spectrometry. Owing to the number of transformations occurring in this final step and the complexity of the spectra, we were unable to unequivocally assign any intermediate species.

Scheme 3.10: Azide reduction and global deprotection. Reagents and conditions: a. NaOMe, MeOH, 87%; b. H2, Pd(OH)2, HCl, EtOAc:MeOH:H2O, 2:2:1.

To minimize the number of transformations in the final deprotection step we cleaved the benzylidene group in 315 with TFA/H2O 9:1 to give the triol (317) (Scheme

3.11). Compound 317 was treated with Pearlman’s catalyst and H2 with 10% HCl at 35 bar for 72 h. NMR analysis of the crude product revealed Man2NH2DMJ as the major species (95%) and an unexpected deaminated product, Man-1,2-dideoxynojirimycin

(MandDMJ, 5%). Mass spectrometry confirmed the exact mass of MandDMJ, 1H NMR showed two multiplets at  1.56 ppm and 2.15 ppm, which each integrate to two protons, and were assigned as H2a,b and H1a,b protons respectively. Our efforts to separate these compounds were unsuccessful. A similar observation was made in 2011 by Gomez and co-workers when they treated cyclohexane containing azide under hydrogenolysis conditions (H2, Pd/C) and observed complete elimination of the azide and reduction of the resultant double bond. 114 We speculate that formation of MandDMJ occurs by elimination of either N3 or NH2 forming an alkene that is hydrogenated under the reaction conditions (H2, Pd(OH)2, HCl).

Scheme 3.11: Deprotection and azide reduction. Reagents and conditions: a.i. NaOMe, MeOH, ii. TFA/H2O 9:1, 83% over 2 steps; b. H2, Pd(OH)2, HCl, EtOAc:MeOH:H2O, 2:2:1. In a bid to circumvent deamination we explored alternative methods of azide reduction that did not involve hydrogenation. The Staudinger reaction is a commonly employed method of azide reduction (Scheme 3.12). It is a multi-step process, the first step involves nucleophilic addition of a phosphine at the terminal nitrogen to form a triazaphosphane. This intermediate undergoes elimination of N2 via a four-membered

transition state, to give a phosphinimine intermediate. In the final step the phosphinimine is hydrolysed to the amine and triphenylphosphine oxide.115

Scheme 3.12: Staudinger reaction mechanism.

Compound 315 was treated under Staudinger conditions with triphenylphosphine, and TFA/H2O 9:1 (Scheme 3.13). Upon work up, NMR spectroscopy and mass spectrometry revealed the sole product to be the phosphinimine. However, this phosphinimine proved remarkably stable; efforts to effect its hydrolysis with both acid/base work up and reflux with strong acid or base were unsuccessful.

Scheme 3.13: Azide reduction. Reagents and conditions: a. PPh3, TFA/H2O 19:1.

We next explored azide reduction with Cleland’s reagent, dithiothreitol (DTT)

(Scheme 3.14). Compound 317 was treated with DTT and pyridine in a carbonate/bicarbonate buffer at pH 9.2 to give 318 in 80% yield. It is crucial that reactions with Cleland’s reagent are performed above pH 7 as the reaction proceeds through the thiolate (S-) the thiol (SH) is not reactive. The amine 318 was treated with Pearlman’s catalyst and H2 with 10% HCl at 35 bar for 48 h to give Man2NH2DMJ (301) in 70% yield.

Scheme 3.14: Azide reduction and global deprotection. Reagents and conditions: a. DTT, pyridine, NaHCO3/H2CO3 pH 9.2, 80%; b. H2, Pd(OH)2, HCl, EtOAc:MeOH:H2O, 2:2:1, 70%.

Binding constant and three-dimensional structure of ManNOE

Man2NH2DMJ was provided to our collaborators in the Davies group at the University of York for analysis. Isothermal titration calorimetry (ITC) performed by Lukaz Sobala revealed that Man2NH2DMJ binds to BtGH99 with KD = 97.7 ± 4.9 µM (Figure 3.6). This is 4-fold worse than GlcDMJ which binds to BtGH99 with KD = 24 µM; there is no equivalent data available for ManDMJ but as this enzyme has a preference for mannose, the difference is expected to be even greater.

Figure 3.6: Isothermal titration calorimetry thermogram showing binding of Man2NH2DMJ to BtGH99. DP = differential power. Binding parameters KD = 97.7±4.9 M, N = 1 (fixed) and H= –5.9±0.1 kcal mol–1.

Three-dimensional structure of Man2NH2DMJ

A three-dimensional structure of Man2NH2 DMJ in complex with BxGH99 was obtained at 1.1 Å resolution (Figure 3.7). The occupancy was complete and the inhibitor spanned

4 the -2/-1 subsites. The 2NH2DMJ ring sat in the -1 subsite in a C1 conformation, similar to complexes of the enzyme with GlcDMJ, Glc/ManIFG and ManNOE based inhibitors.

N2 was located 2.59 Å away from E333, within an appropriate distance for the pair to interact (Figure 3.7b, c).

a. b. c.

Figure 3.7: a. Three-dimensional structure of BxGH99 complex with Man2NH2DMJ. Distance between catalytic amino acid residue E333 and O2 or N2: b. GlcDMJ; c. Man2NH2DMJ (301)

Overlay of this crystal structure with that of BxGH99-GlcDMJ reveals the positioning of Man2NH2DMJ to be practically identical to that of GlcDMJ where residue

E333 is 2.54 Å away from O2; no amino acid residues appeared shifted (Figure 3.8).

Within the -2 subsite some small differences are noted arising from the difference between Glc/Man. These differences are not considered material. Given there was essentially no difference in binding position of the two inhibitor warheads, the poor binding affinity cannot be attributed to incorrect positioning of the inhibitor, hence there is a failure to form strong interactions.

Figure 3.8: Three-dimensional structure of BxGH99 overlay of GlcDMJ and Man2NH2DMJ.

Studies of pH dependence of binding show that iminosugars bind to glycosidases

116 in their protonated form. Man2NH2DMJ has two basic nitrogen residues that could potentially be protonated within the complex. Generally, pKa values of amine conjugate acids decrease as the number of electronegative groups (eg: hydroxyl, amine, acetamido) near the basic centre increases. Moreover for diamines protonation of one amino group can dramatically reduce the basicity the other. In the case of amino sugars, the magnitude of this effect depends heavily on the relative positions of the amine group within the structure and to a lesser extent on the stereochemistry. Sugars with amines (6-amino sugars) have pKa values of 8.8-8.9 whereas compounds with amines on a carbon adjacent

117,118 to the ring oxygen are much weaker bases with pKa values in the range of 5-6. In acyclic systems with vicinal amines the effect on pKa value of the second nitrogen has been estimated to be pKa = 3.6 units. In cyclic systems stereoelectronic and conformational effects come into play, some examples include (pKa1, pKa2): 4-2'- aminoethylmorpholine (9.5, 4.8);118 cis-1,3-diaminocyclohexane (10.3, 8.3);117 trans-1,3- diaminocyclohexane (10.4, 8.5); 4-hydroxy-3-hydroxymethylhexahydropyridazine (6.3,

5.3); 1-azafagomine (5.0, 4.7) (Figure 3.9).117,119

Figure 3.9: pKa values of: a. 4-2'-aminoethylmorpholine; b. cis-1,3- diaminocyclohexane c. trans-1,3-diaminocyclohexane; d. 4-hydroxy-3- hydroxymethylhexahydropyridazine e. 1-azafagomine. 117,119

Since, Man2NH2DMJ has 4-OH antiperiplanar to the endocyclic nitrogen, it is estimated this will reduce the basicity by 1.3 pKa units. Protonation at N2 may be favoured by protonation by E333, which would further decrease basicity of the endocyclic N by an additional 3.6 pKa units. This analysis suggest that formation of the di-cation is unlikely.

As a result, we propose that Man2NH2DMJ binds with only a modest KD value as it fails to be protonated at the endocyclic N and thereby mimic an oxocarbenium-ion like transition state (Figure 3.10).

Figure 3.10: a. First transition state in proposed mechanism for family GH99 retaining endo-α-1,2-mannosidases/endo-α-1,2-mannanases; b. protonation state of ManDMJ; c. protonation state of Man2NH2DMJ.

Exploration of strategies for shape mimicry in inhibitor design for family GH99 endo-α-1,2-mannanases

The work discussed so far was conducted by myself as part of a larger collaborative project that involved concurrent synthetic work by a co-worker, Pearl Fernandes. I will now provide a summary this work.

Pearl Fernandes considered approaches to develop inhibitors of GH99 that could

4 achieve transition state mimicry. Although charge-based inhibitors which bind in a C1 conformation mimicking the Michaelis complex have experienced much success, neutral compounds have also been shown to be relatively effective inhibitors of mammalian EM.

In 1993, Spiro and co-workers reported that the neutral compound GlcGlucal (IC50 2.3

M) was an effective inhibitor of mammalian EM, only slightly less effective than charge based inhibitor GlcDMJ (IC50 1.7 M), which was the most potent inhibitor at that time.

In Chapter 2, it was shown that the equivalent molecule, ManGlucal, binds with modest affinity to BtGH99 (KD 15 M). Computational analysis of the conformational free energy landscape for D-glucal suggests the inhibition of these neutral compounds arise

4 from mimicking the H3 conformation proposed for of the GH99-catalyzed reaction intermediate, not the conformation proposed for the TS.

A basic class of flattened inhibitors, the glycoimidazoles, has not yet been explored in the study of GH99 enzymes. This class of inhibitors was developed following the discovery of the natural product nagstatin and its potent inhibition of N-acetyl-β- glucosaminidase (NAGase).120 The potency of the glycoimidazoles is attributed to their ability to mimic the shape of the oxocarbenium-ion-like transition state and ability of the imidazole glycosidic nitrogen form a hydrogen bond with an appropriately situated active site carboxylate residue, often the acid/base.21 As part of this collaborative body of work,

Pearl Fernandes chose to evaluate the glycoimidazole ManManIm, for its potency against bacterial GH99.

Synthesis of ManManIm

Pearl Fernandes synthesise ManManIm is shown in Scheme 3.15. Starting from the known alcohol (Scheme 3.15), treatment with napthylmethyl bromide (NapBr)/NaH gave the thioglycoside. Hydrolysis with NIS in H2O/acetone gave the hemiacetal. The hemiacetal was converted to the aminals over 4 steps, involving (1) oxidation to a lactone under Albright-Goldman conditions (2) aminolysis of the lactone to give an acyclic amide

(3) Albright-Goldman oxidation and ring-closure with methanolic ammonia to afford a mixture of aminals and (4) reduction with NaCNBH3 gave a 2:1 mixture of the D-manno and L-gulo lactams, from which the D-manno lactam was isolated in 38% yield. The D- manno-lactam was converted to the thionolactam with Lawesson’s reagent and the imidazole ring annulated with aminoacetaldehyde dimethyl acetal via the amidine, which when treated with tosic acid afforded a mixture of the D-gluco and D-manno imidazoles in a 2:1 ratio. The D-manno imidazole was isolated in 32% yield and Nap group removed under oxidative conditions (DDQ, CH2Cl2/H2O) to give the alcohol. Triflic acid catalyzed glycosylation of the alcohol with a mannose derived trichloroacetimidate donor afforded the disaccharide in 47%. Deprotection of the acetate with K2CO3/MeOH, followed by global deprotection (Pd(OH)2, H2) gave ManManIm.

Scheme 3.15: Pearl Fernandes’ synthesis of ManManIm.

Binding data and three-dimensional structure of ManManIm

ManManIm was provided to our collaborators in the Davies group at the University of

York for analysis. Isothermal titration calorimetry (ITC) was performed by Lukasz

Sobala. However, no heat flow was observed and so a KD value could not be obtained. A three-dimensional structure of ManManIm in complex with BxGH99 was obtained at a

1.3 Å resolution. The compound spanned the -2/-1 subsites, however even after prolonged soaking the enzyme occupancy was only 80%, indicating poor affinity of ManManIm to

2 the enzyme. The structure shows the piperidine ring to be in an unusual H3/E3

conformation. Overlay with GlcDMJ complex reveals the -2 sugar to be in a similar position (Figure 3.11). However, the mannoimidazole headgroup projects downward into the active site with a 2.65 Å distance between residue E336 and N1 of the imidazole. This is a typical distance for related glycoimidazole complexes; as demonstrated by Varrot and co-workers in the case of a cellobiose-derived imidazole bound to Cel5A from B. agaradhaerens, the distance between N of imidazole and E139 (acid/base residue) is 2.58

Å.121 Furthermore, if the mannoimidazole headgroup were to be shifted up to the same position as the piperidine of GlcDMJ, it would give rise to a steric interaction with conserved residue Y252 that forms the top of -1/+1 pockets.

a. b.

Figure 3.11: Three-dimensional structure of BxGH99. a. Complex with ManManIm; b. overlay of GlcDMJ and ManManIm.

3.6 Conclusions

In this chapter, we report the synthesis of novel molecules Man2NH2DMJ and

ManManIm, mechanism-inspired inhibitors of bacterial EM enzymes. ITC revealed

Man2NH2DMJ binds to BtGH99 with KD = 97.7 ± 4.9 µM, 4-fold worse than its GlcDMJ counterpart (KD = 24 µM). The three-dimensional structure revealed the NH2DMJ ring to bind virtually identically to the DMJ ring and the catalytic residue E333 to be located

2.59 Å away from N2 of NH2DMJ, an appropriate bond forming distance. The poor

binding of Man2NH2DMJ is likely a result of the protonated 2NH2 substituent reducing basicity of N1 and preventing it from undergoing protonation in the active site failing to mimic charge on the oxocarbenium-ion like transition state. No binding was observed for

ManManIm by ITC. Yet a moderate quality complex with BxGH99 was obtained. This revealed the ManIm headgroup failed to bind in the expected pose likely as consequence of the inability of the enzyme to accommodate the ManIm ring, a result of the unfavourable interaction with conserved residue Y252, as well as the location of the acid/base E366, below the mean plane of the substrate pyranose ring.

3.7 Experimental

2,3,4,6-Tetra-O-benzyl-D-gluconamide (302) 2,3,4,6-Tetra-O-benzyl-D-glucopyranose (5.00 g, 9.25 mmol) was dissolved in THF (20 mL). A 30% aqueous ammonia solution (100 mL) was added, precipitation of the sugar was observed. Iodine (3.29 g, 12.8 mmol) was added to afford a black mixture. After 18 h the reaction mixture was colourless and iodine (0.47 g, 1.85 mmol) was added. After a further 24 h of stirring,

5% Na2SO4 (10 mL) was added. The resulting mixture was extracted into Et2O (3 × 100 mL). The combined organic layers were washed with brine (100 mL), dried MgSO4, filtered and concentrated under reduced pressure. Flash chromatography (EtOAc/pet. 108 1 spirits 60:40) gave the amine (302) (4.57 g, 89%) as a white solid. H NMR (CDCl3, 500 MHz) δ 2.82 (1 H, d, J = 4 Hz, OH), 3.59 (1 H, dd, J = 9.5, 5 Hz, H6a), 3.65 (1 H, dd, J = 10, 3 Hz, H6b), 3.85- 3.93 (2 H, m, H4, 5), 4.07 (1 H, dd, J = 5.5, 3.5 Hz, H3),

4.25 (1H d, J = 3 Hz, H2), 4.49-4.73 (8 H, m, 4 × CH2Ph), 5.46 (1 H, s, NH), 6.59 (1 H, 13 s, NH), 7.21-7.36 (20 H, m, 4 × Ph); C NMR (CDCl3, 125 MHz) δ 71.2 (1 C, C6), 71.5

(1 C, C5), 73.6, 73.9, 74.3, 75.4 (4 C, 4 × CH2Ph), 77.8 (1 C, C4), 79.8 (1 C, C2), 80.7 (1 C, C3), 127.8, 127.9, 128.0, 128.1, 128.4, 128.5, 128.6, 128.8, 136.9, 137.9, 138.2, 138.3 + + (24 C, 4 x Ph), 174.1 (1 C, C1); HRMS (ESI) m/z 556.2692 [C34H37NO6 (M+H) requires 556.2694].

2,3,4,6-Tetra-O-benzyl-5-dehydro-5-oxo-D-gluconamide (303) PDC (1.35g, 3.60 mmol) was added to a solution of amine (302)

(1.00 g, 1.80 mmol), in CH2Cl2 (10 mL) over 4 Å sieves. The

solution was stirred for 3 h, filtered through a silica plug (Et2O), dried over anhydrous MgSO4 and concentrated under reduced pressure. The resulting crude amide (303) (0.940 g, 0.170 mmol) was used in the next step without purification.110 1 H NMR (CDCl3, 500 MHz) δ 4.16- 4.67 (13 H, m, H2-6b, CH2Ph), 5.89 (1 H, s, NH), 6.60 (1 H, s, NH), 7.18-7.33 (20 H, m, Ph).

2,3,4,6-Tetra-O-benzyl-5-dehydro-5-hydroxy-D-glucono-and L-idonolactam (305) Crude amide (303) (0.940 mg, 0.170 mmol), was stirred in sat.

methonic NH3 (10 mL) for 2 h. The solution was evaporated under

reduced pressure and purified by flash chromatography (Et2O/pet. spirits 50:50) to give a mixture of hydroxy lactams (D-gluco and D-iodo’, 305)109 (0.940

1 mg, 0.170 mmol). H NMR (CDCl3, 500 MHz) δ 2.25 (1 H, d, J = 9.5 Hz, H6a), 3.27 (1H, s, OH), 3.34 (1 H, d, J = 9.6 Hz, H6b), 3.52 (1 H, d, J =9.2 Hz, H6a’), 3.62 (1 H, d, J =9.2 Hz, H6b’), 3.73-3.75 (2 H, m, H4, 4’), 3.90 (1 H, dd, J = 6.6, 4.2 Hz, H3’), 4.02 (1 H, d, J = 8.5 Hz, C2), 4.20-4.23 (1 H, m, H3), 4.34 (1 H, d, J = 6.6 Hz, H2’), 4.38-5.19

(16 H, m, 4 × CH2Ph, 4 × CH2Ph’), 6.17 (1 H, s, NH), 6.27 (1 H, s, NH), 7.15-7.13 (40 H, m, 4 × Ph, 4 × Ph’). Based on the NMR-spectra the major product is assigned to be the D-gluconolactam and the minor product to be the L-idonolactam.

2,3,4,6-Tetra-O-benzyl-D-gluconolactam (304) The hydroxyl lactams (305) (0.940 mg, 0.170 mmol) were dissolved in acetonitrile (17 mL) and formic acid (4.5 µL). Sodium cyanoborohydride (256 mg, 4.08 mmol) was added and the solution stirred at reflux for 2 h. The mixture was cooled to 0˚C and quenched with 10% HCl (4.0 mL) and left to stir for 15 min. The mixture was poured into EtOAc:sat.NaHCO3 (1:1, 100 mL). The aqueous layer was extracted with EtOAc (2 × 50 mL), washed with brine

(2 × 50 mL), dried over anhydrous MgSO4, filtered and concentrated under reduced pressure. The resulting oil was purified with flash chromatography (EtOAc/pet. spirits 109 1 30:70) to afford the lactam (304) (745 mg, 77%) as a white solid. H NMR (CDCl3, 500 MHz) δ 3.25 (1 H, m, H6a), 3.51-3.61 (3 H, m, H4, 5, 6b), 3.91 (1 H, t, J = 8.2 Hz,

H3), 4.00 (1 H, d, J = 8.0, H2), 4.44-4.49 (2 H, m, 2 × CH2Ph), 4.72 (1 H, d, J = 11.1 Hz,

CH2Ph), 4.77 (1 H, d, J = 11.2 Hz, CH2Ph), 4.83-4.86 (2 H, m, CH2Ph), 5.17 (1 H, d, J = 13 11.2 Hz, CH2Ph), 5.87 (1 H, s, NH), 7.18-7.42 (20 H, m, CH2Ph); C NMR (CDCl3, 125

MHz) δ 53.91, 70.13, 73.45, 74.73, 74.84, 77.24, 78.90, 82.45 (9 C, C2-C6, 4 × PhCH2), 127.90, 127.99, 128.11, 128.24, 128.33, 128.47, 128.52, 128.66, 137.39, 138.69, 137.95, + + 138.14 (24 C, Ar), 170.58 (1C, C=O); HRMS (ESI) m/z 538.2588 [C34H35NO5 (M+H) requires 538.2588].

2,3,4,6-Tetra-O-benzyl-1-deoxynojirimycin (306) Borane dimethyl sulfide (927 µL, 9.77 mmol) was added dropwise to the lactam (304) (1.5 g, 2.79 mmol) in THF (15 mL). The solution was stirred at refluxed for 24 h. The solution was cooled to 0 ˚C and methanol (30 mL) was added dropwise, then the solution refluxed for 24 h. The crude was asiotroped with methanol (3 x 20 mL), reduced under concentrated pressure. Flash chromatography (EtOAc/pet. spirts 30:70) gave tetra benzyl DNJ (306)97, 111 (1.46g, 1 >99%) as white crystals. H NMR (CDCl3, 500 MHz) δ 2.52 (1 H, dd, J = 12.2, 10 Hz,

H1), 2.73 (1 H, ddd, J = 9.4, 5.9, 2.6 Hz, H5), 3.26 (1 H, dd, J = 12.3, 4.9 Hz, H1), 3.37 (1 H, m, H4), 3.48-3.59 (3 H, m, H2, 3, 6a), 3.68 (1 H, dd, 9.0, 2.6, H6b), 4.43-5.00 (8 13 H, m, 4 × CH2Ph), 7.20-7.37 (20 H, m, 4 × Ph); . C NMR (CDCl3, 125 MHz) δ 48.25,

59.86, 70.38, 72.86, 73.48, 75.26, 75.74, 80.21, 80.77, 87.44 (10 , C1-C6, 4 × CH2Ph), 127.73, 127.86, 127.93, 128.01, 128.09, 128.42, 128.44, 128.47 (24 C, Ph); HRMS (ESI)+ + m/z 524.2793 [C34H37NO4 (M+H) requires 524.2795].

1-Deoxynojirimycin (DNJ, 307)

The benzylated DNJ (306) (400 mg, 0.744 mmol) was dissolved in

EtOAc/MeOH/H2O 2:2:1 (12 mL). To this Pd/C (80 mg) and 10% HCl in

MeOH (1.2 mL) was added. The suspension was treated with H2 (20 bar) and stirred for 24 h. The crude was filtered through celite, washed with MeOH/H2O 2:1 (100 mL), and concentrated under reduced pressure. The crude was purified through anion exchange chromatography to give DNJ (307)122 (124 mg, >99%) as a colourless 1 oil. H NMR (D2O, 500 MHz) δ 2.94 (1 H, t, J = 12.0 Hz, H1a), 3.16 (1 H, J = 3.1, 5.1, 10.1, H5), 3.47-3.60 (3 H, m, H1b, 3, 4), 3.78 (1 H, ddd, J = 5.1, 9.3, 11.4 Hz, H2), 3.87 13 (1 H, dd, J = 5.3, 12.6 Hz, H6a), 3.95 (1 H, dd, J = 3.0, 12.6 Hz, H6b); C NMR (D2O, 100 MHz) δ 48.5 (1 C, C1), 60.2 (1 C, C5), 62.9 (1 C, C6), 69.8 (1 C, C2), 70.7 (1 C, C4), + + 78.8 (1 C, C3); HRMS (ESI) m/z 164.0916 [C6H13NO4 (M+H) requires 164.0917].

N-Benzyloxycarbonyl-1-deoxynojirimycin (308) DNJ (307) (160 mg, 0.981 mmol) was dissolved in a mixture of 1,4-

dioxane:water (1:1, 64 mL), then NaHCO3 (165 mg) and benzylchloroformate (211 µL, 1.47 mmol), were added and the reaction stirred overnight. The crude was concentrate under reduced pressure and purified by flash chromatography (AcOEt:MeOH:H2O, 85:10:5) to give the 123 1 chlorobenzoate (308) (255 mg, 89 %) as a colourless oil. H NMR (CD3OD, 500 MHz) δ 3.48 (1 H, d, J = 13.8 Hz, H1a), 3.71-3.91 (5 H, m, H2, 3, 4, 6a, 6b), 4.04 (1 H, d, J = 13 14Hz, H1b), 4.24-4.27 (1 H, m, H5), 5.17 (2 H, s, CH2), 7.26-7.44 (5 H, m, Ph); C NMR

(CD3OD, 125 MHz) δ 61.15, 61.41, 61.53, 68.32, 70.29, 71.14 (C 7, C1-6, CH2), 128.68, + 128.93, 129.45 (6 C, Ph), 172.97 (C=O); HRMS (ESI) m/z 298.1284 [C14H19NO6 (M+H)+ requires 298.1285].

4,6-O-(R-benzylidene)-N-benzyloxycarbonyl-1-deoxynojirimycin (309) The chlorobenzoate (308) (250 mg, 0.84 mmol) was dissolved in dimethylformamide (1 mL). To this benzaldehyde dimethyl acetal (140 µL, 1.09 mmol) and p-toulenesulfonic acid (2.5 mg) was added and the mixture heated to 60 ˚C for 2 h. The reaction mixture was concentrated under reduced pressure. Flash chromatography (AcOEt/pet. spirits 60:40) gave the diol (309)104 (250 mg, 77 %) 1 as a white solid. H NMR (600 MHz, CDCl3) δ 3.06 (1 H, dd, J = 13.7, 9.6 Hz, H1a), 3.27 (1 H, td, J = 10.2, 4.6 Hz, H5), 3.55-3.72 (2 H, m, H3, 4), 4.28 – 4.40 (3 H, m, H1b,

H2, H6a), 4. (1 H, dd, J = 11.3, 4.0 Hz, H6b), 5.10 (2 H, d, J = 13.4 Hz, CH2), 5.50 (1 H, 13 s, CHPh), 7.26- 7.78 (10 H, m, Ph); C NMR (100 MHz, CDCl3) δ 49.30, 55.23, 67.78,

69.56, 69.59, 77.26, 80.28 (7 C, C1-6, CH2), 101.75 (1 C, CHPh), 126.37, 128.29, 128.52, 128.80, 129.48, 136.02, 137.33 (12 C, Ph), 154.88 (1 C, C=O); HRMS (ESI)+ m/z + 386.1597 [C21H23NO6 (M+H) requires 386.1598].

4,6-O-(R-benzylidene)-N-benzyloxycarbonyl-1,5-dideoxy-1,5-imino-2-O-(p- toluenesulfony1)-D-glucitol (313)

The diol (309) (100 mg, 0.25 mmol) was refluxed with dibutyltin oxide (78 mg, 0.31 mmol) in methanol (5mL) for 2h. The mixture was concentrated to dryness and azeotroped with toluene (2 x 5mL). The residue was dissolved in dichloromethane (2 mL and) triethylamine (43 µL, 0.31 mmol). The solution was cooled to 0˚C and p-toluenesulfonyl chloride (54 mg, 0.29 mmol) was added. The solution was stirred at 0˚C for 30 min then warmed to room temperature and stirred for 2 h. The reaction mixture was poured into sat. aq. sodium carbonate, washed with 0.5M KHSO4 (3 × 10 mL), water (3 × 10mL), dried MgSO4 and concentrated under reduced pressure. Column chromatography (AcOEt/pet. spirits 30:70) 104 1 gave the tosylate (313) (127 mg, 91%) as a white solid. H NMR (500 MHz, CDCl3) δ

2.41 (3 H, s, CH3), 2.74 (1 H, d, J = 2.2, OH), 3.05 (1 H, dd, J = 9.5, 13.8 Hz, H1a), 3.26 (1 H, dt, J = 1, 4.6, 10.2 Hz, H5), 3.59 (1 H, dd, J = 5.0, 10 Hz, H4), 3.78 (1 H, m, H3), 4.31 (2 H, m, H1b, 6b), 4.39 (1 H, ddd, J= 4.8, 7.6, 9.4, H2), 4.77 (1 H, d, J = 4.6, 11.5,

H6b), 5.12 (2 H, s, CH2), 5.50 (1 H, s, CH), 7.26 (2 H, d, J = 8Hz, Ph), 7.33 (10 H, m, + + Ph), 7.78 (2 H, d, J = 8.4 Hz, Ph) ; HRMS (ESI) m/z 428.1582 [C28H29NO8S (M+H) requires 428.1506].

2-Azido-4,6-O-benzylidene-N-benzyloxycarbonyl-1,2,5-trideoxy-1,5-imino-D- mannitol (314) Sodium azide (57.8 mg, 0.890 mmol) was added to a solution of tosylate (313) (120 mg, 0.222 mmol) in DMF (1 mL). The suspension was refluxed for 18 h, poured into ice, extracted into EtOAc (3 × 20 mL), washed with brine (2 × 20mL), dried over anhydrous MgSO4 and evaporated to dryness. Column chromatography (AcOEt:pet. spirits 20:80) gave the azide (314) (67.7 mg, 74%) 24 1 as a white solid; [α]D –21.9 (c 1.12, CHCl3); H NMR (CDCl3, 500 MHz) δ 2.74 (1H, s, NH), 2.82 (1 H, d, J = 1.6, 14.5 Hz, H1a), 3.06 (1 H, td, J = 4.6, 10.2 Hz, H5), 3.74 (1 H, dd, J = 3.8, 9.2 Hz, H3), 3.79-3.93 (2 H, m, H2, 4), 4.31 (1 H, dd, J = 3.0, 14.5 Hz, H1b) 4.46 (t, J = 1, 11 Hz, H6a), 4.66 (1 H, dd, J = 4.6, 11.6 Hz, H6b), 5.01 (2 H, d, J = 13 3.1 Hz, CH2), 5.48 (1 H, s, CH). C NMR (CDCl3, 125 MHz) δ 48.1, 55.8, 60.1, 67.8,

69.2, 73.6, 78.2 (7 C, C1-6, CH2, 101.8 (1 C, CH), 126.3, 128.3, 128.4, 128.5, 128.7, 129.4, 136.0, 137.3 (12 C Ph), 155.0 (1 C, C=O); HRMS (ESI)+ m/z 411.1664 + [C21H22N4O5 (M+H) requires 411.1663].

2-O-Acetyl-3,4,6-tri-O-benzyl-α-D-mannopyranosyl-(1→3)-2-azido-4,6-O- benzylidene-N-benzyloxycarbonyl-1,2,5-trideoxy-1,5-imino-D-mannitol (315) TfOH (0.043 µL, 0.0049 mmol) was added to a mixture of acceptor (314) (20 mg, 0.049 mmol) and 2-O-acetyl-3,4,6-tri- O-benzyl-α-D-mannopyranosyl trichloroacetimidate (219)

(37 mg, 0.058) in CH2Cl2 over 4 Å sieves at -30 ̊ C, The mixture was stirred for 30 min, warmed to 0 °C and quenched with Et3N (7 µL, 0.05 mmol) then concentrated under reduced pressure. Flash chromatography (EtOAc/pet. 24 spirits 25:75) gave the disaccharide (315) (37.4 mg, 87%) as a colourless oil. [α]D -4.2 1 (c. 0.89, CHCl3); H NMR (CDCl3, 500 MHz) δ 2.80 (1 H, J1,1 =14.4, J1,2 = 0.9, H1a), 3.15 (1 H, dt, J = 10.1, 4.6, 1 Hz, H5), 3.70-4.00 (6 H, m, H3, 4, 4', 5', 6a', 6b'), 4.03 (1 H, dd, J = 9.3, 3.4, H3'), 4.17-4.20 (1 H, m, H2), 4.28 (1 H, dd, J = 14.5, 2.2, H1b), 4.47-

4.52 (3 H, m, 3 × CH2Ph), 4.60-4.64 (2 H, m, H6a, CH2), 4.69 (1 H, d, J = 11 Hz, CH2Ph),

4.76 (1 H, dd, J = 11.6, 4.5 Hz, H6b), 4.86 (1 H, d, J = 11 Hz, CH2Ph), 5.12 (2 H, J = 3.6,

CH2), 5.28 (1 H, d, J = 1.6 Hz, H1'), 5.59 (1 H, J = 3.3, 1.8 Hz, H2'), 5.64 (1 H, s, CH), 13 7.17-7.46 (25 H, m, Ph); C NMR (CDCl3, 125 MHz) δ 48.3 (1 C, C1), 56.3 (1 C, C5),

60.0, 72.7, 74.4, 77.8 (4 C, C3,4,4',5), 67.7 (1 C, CH2), 68.5 (1 C, C2'), 69.1 (1 C, C6),

69.3 (1 C, C6'), 72.2, 73.6, 75.1 (3 C, CH2Ph), 78.1 (1 C, C2), 78.2 (C1, H3'), 99.5 (1 C,

C1'), 100.90 (1 C, CH), 100.92, 126.0, 127.77, 127.79, 127.83, 127.9, 128.0, 128.2, 128.28, 128.29, 128.41, 128.44, 128.5, 128.7, 128.9 (C30, Ph); HRMS (ESI)+ m/z + 907.3544 [C50H52N4O11 (M+Na) requires 907.3525].

3,4,6-Tri-O-benzyl-α-D-mannopyranosyl-(1→3)-2-azido-N-benzyloxycarbonyl- 1,2,5-trideoxy-1,5-imino-D-mannitol (317) A solution of sodium methoxide in methanol (0.1 M, 10 µL, 1 µmol) was added to (315) (60 mg, 0.068 mmol) in methanol (0.5 mL) and stirred for 1 h. The mixture was concentrated under reduced pressure to give an alcohol, which was used without purification. TFA/H2O 9:1 (100 µL) was added to the crude alcohol, the mixture was stirred for 30 min, concentrated and azeotroped with toluene (3 × 10 mL). Flash 25 chromatography (EtOAc/pet. spirits 90:10) gave the triol (317) (42.5 mg, 83%). [α]D 1 44.6 (c. 1.03, MeOH); H NMR (500 MHz, CD3OD) δ 3.67-4.20 (13 H, H1a-6b, H2'-

H6'b), 4.43-4.46 (2 H, m, CH2), 4.58 (1 H, d, J = 12.0 Hz, CH2Ph), 4.67 (2 H, s, J = 12.4

Hz, CH2Ph), 4.78 (1 H, d, J = 11.0 Hz, CH2Ph), 5.12 (2 H, s, CH2), 5.15 (1 H, apt. s, 13 H1'), 7.03-7.42 (20 H, m, 4 × Ph); C NMR (CDCl3, 125 MHz) δ 59.5, 68.0, 68.9, 69.0,

71.9, 72.5, 73.5, 74.2, 74.9, 79.5 (13 C C1, 2, 3, 4, 5, 6, 1', 2', 3', 4', 5', 6', CH2) 127.8, 127.9, 128.0, 128.1, 128.16, 128.19, 128.4, 128.5, 128.6, 128.7, 137.9, 138.0, 138.3 (24 + + C Ph), 156.5 (1 C, C=O); HRMS (ESI) m/z 755.3300 [C41H46N4O10 (M+H) requires 755.3287].

3,4,6-Tri-O-benzyl-α-D-mannopyranosyl-(1→3)-2-amino-N-benzyloxycarbonyl- 1,2,5-trideoxy-1,5-imino-D-mannitol (318) DTT (51 mg, 0.331 mmol) was added to a solution of azide (317) (25 mg, 0.0331 mmol) in pyridine (1 mL) and

NaHCO3/H2CO3 buffer (0.625 mL, pH 9.16). The mixture was stirred at room temperature for 4 h, concentrated and azeotroped toluene (5 × 10 mL). Flash chromatography (EtOAc/MeOH/H2O 94:4:2) to 1 give the amine (318) (80%, 19.2 mg). H NMR (500 MHz, CD3OD) δ 2.89 (1 H, t, J =

12.4 Hz, H2), 3.21-4.13 (13 C m, H1a,1b, 3, 5, 6a, 6b, 1'-6b'), 4.36 (1 H, t, J = 7.8 Hz,

H4), 4.46-4.54 (2 H, m, CH2Ph), 4.58 (1 H, d, J = 12.0 Hz, CH2Ph), 4.66 (d, J =11.8 Hz,

CH2Ph), 4.77-4.81 (2 H, m, CH2Ph), 4.98 (1 H, d, J = 2.5 Hz, H1'), 5.15 (2 H, s, CH2), 13 7.16-7.47 (20 H, m, Ph); C NMR (CDCl3, 125 MHz) δ 46.8, 59.9, 65.6, 68.5, 69.4, 70.4,

72.6, 73.7, 74.4, 75.4, 75.7, 78.1, 80.1, 100.8 (16 C C1-6, C1'-6', 4 × CH2), 128.81,

128.84, 129.2, 129.28, 128.30, 129.3, 129.4, 129.5, 138.0, 139.3, 139.5, 139.6 (24 C Ph); + + HRMS (ESI) m/z 729.3398 [C41H48N2O10 (M+H) requires 729.3385].

α-D-Mannopyranosyl-(1→3)-2-amino-N-benzyloxycarbonyl-1,2,5-trideoxy-1,5- imino-D-mannitol (301)

The triol (318) (19.2 mg, 0.0264 mmol) in MeOH/H2O (2:1, 3 mL) and 10% HCl in methanol (0.3 mL) was treated with

PdOH/C (50 mg) and H2 (20 atm, 18h). The suspension was filtered, concentrated and purified with cation and anion resin

(eluted with aqueous NH3) to give Man2NH2DMJ (301) (70%, 6.02mg) as a colourless 25 1 oil. [α]D 17.2 (c. 0.08, H2O); H NMR (500 MHz, D2O) δ 2.78-2.84 (1 H, m, H5), 3.09

(1 H, dd, J1a,1b = 14.0, J1a,2 = 2.1, H1a), 3.25 (1 H, dd, J1a,1b = 14.0, J1a,2 = 3.2 Hz, H1b),

3.62-3.95 (9 H, m, H2-4, 6a, 6b, 4’-6b'), 3.98 (1 H, dd, J3',4' = 9.2, J2',3' = 4.3 Hz, H3'), 13 4.09 (1 H, dd, J2',3' = 3.3, J1',2' = 1.8 Hz, H2'), 5.24 (1 H, d, J1',2' = 1.6 Hz, H1'); C NMR

(125 MHz, D2O) δ 44.5, 50.4, 60.0, 60.8, 61.0, 66.6, 67.3, 69.7, 70.1, 73.7, 77.3, 101.6; + + HRMS (ESI) m/z 325.1606 [C12H24N2O8 (M+H) requires 325.1605].

CHAPTER FOUR:

GH134 a β-mannanase with a lysozyme-like fold

and a novel molecular mechanism

4.1 Use of β-mannanases in the biofuels industry

The major carbohydrates of the plant cell wall are cellulose, hemicellulose and pectin.124

The cell wall also contains significant quantities of lignin, a polyphenolic polymer that is covalently linked to hemicellulose and which strengthens the cell wall. Second generation biofuels are obtained by conversion of lignocellulosic material from the plant cell wall to ethanol, by firstly depolymerisation to component monosaccharides, and then fermentation of the monosaccharides.125 Unlike first generation biofuels, which are derived from starch-based polysaccharides, second generation biofuels aim to uncouple the competitive link between food and fuel production that has adversely affected the cost of food in many poorer countries.

A challenge in obtaining beneficial economics in second generation biofuel production is ensuring the complete saccharification of lignocellulose, a particularly recalcitrant substrate.126 Consequently much effort is being expended upon the discovery and industrial optimization of diverse plant cell wall degrading enzymes. Enzymes that degrade hemicelluloses are seen as one important part of the solution. Hemicelluloses include a range of polysaccharides such as xylan, xyloglucan and -mannans, which along with pectin bind cellulose fibers to provide a complex cross-linked matrix. The - mannans are comprised of a range of polysaccharides, including pure -1,4-linked mannan, glucomannan, galactomannan and galactoglucomannan (Figure 4.1).

Glucomannan is comprised of β-1,4-linked glucose and mannose units,127 and galactomannan and galactoglucomannan are made up of a -1,4-linked mannan backbone modified by galactose side chains linked through α-1,6-bonds. Reflecting the wide variety of -mannans, a range of mannanolytic enzymes with different substrate specificities are necessary for complete saccharification of the cell wall.128

Figure 4.1: Examples of some common β-mannans which comprise hemicelluloses

4.2 β-Mannanase and β-mannosidase GH families.

Up until 2014, β-mannanase activity had been reported for three families: GH 5, 26 and

113. These three families act with retention of stereochemistry though a classical

Koshland double-displacement mechanism, share an (α/β)8 barrel fold and have similar catalytic machinery.129–133 β-1,4-Mannosidase activity has been reported for two families,

GH 2 and 130. Family GH2 contains a retaining enzymes that use the classical Koshland

134,135 mechanism and consist of five domains. The first domain is a ‘jelly-roll’ (β)8 barrel and the second and fourth have a fibronectin-type III fold.136 The third domain forms a distorted TIM barrel, in contrast to a typical TIM which consist of eight β/α repeats, GH22 has an unusual (α/β)8 barrel with a distortion in the sixth parallel barrel strand and missing fifth helix. The fifth domain is an 18-stranded, antiparallel sandwich.135 In contrast, family GH130 contains inverting enzymes acting though a single-displacement mechanism, and which have a five-bladed β-propeller fold with long α-helices attached to the N- and C-termini.137 Enzymes from families GH 2, 5, 26, 113 and 130 all operate

1 ‡ O though conserved S5→B2,5 → S2 conformational itineraries (for the glycosylation half reaction or the complete reaction in the case of GH130) (Scheme 4.1).47,138,139

Scheme 4.1: Conformational itinerary for the glycosylation half reaction of retaining, β- 1,4-mannanases and β-1,4-mannosidases of families GH 2, 5, 26 and 113. OR is the leaving group and X is a hydrogen or another sugar moiety.

4.3 Discovery of a novel β-1,4-mannanase and new GH family

In 2015 a new β-1,4-mannanase, Man134A, was identified from Aspergillus nidulans.140

It had no amino acid sequence similarity with any β-1,4-mannanases or any other known glycosidases, and consequently became the foundation member of a new family, GH134.

Man134A acted on -1,4-mannooligosaccharides, with optimal activity at neutral pH.

Man134A cleaved β-mannohexaose (M6) with a high catalytic efficiency, as measured by kcat/KM, into mannobiose (M2), mannotriose (M3) and mannotetraose (M4), with M3 the predominant product (Scheme 4.2). Mannopentaose (M5) was cleaved to M2 and M3 with much lower efficiency. No transglycosylation products were detected in reactions catalyzed by Man134A. By contrast, β-mannanases from families GH5 and GH113 are known to generate transglycosylation products.141

Scheme 4.2: Man134A cleavage of β-mannohexaose (M6).

GH26 and GH113 enzymes are of high molecular weight (typically 40-46 kDa) and thus their expression imposes a considerable metabolic expense onto producer strains.142,143 By contrast, GH134 proteins are much shorter, comprising of approximately

200 amino acids in length (hen egg white lysozyme is even shorter – 129 amino acids), corresponding to a much lower molecular weight (approx. 18 kDa), which could be attractive from an industry perspective.140

4.4 Aims

This chapter seeks to study substrate preference, mode of cleavage, and determine the stereochemistry of hydrolysis of a representative from the newly created family GH134, identified by Dr Ethan Goddard-Borger. In collaboration with Prof Gideon Davies’ laboratory, we determine the first tertiary structure of a representative of this family, including in complex with substrate. In collaboration with Prof Carme Rovira’s laboratory, and using this data, quantum mechanical calculations are performed to define

the free energy landscape for the reaction and assign a conformational itinerary along the reaction coordinate.

4.5 Results and Discussion

Identification of a GH134 representative

Four representative GH134 members were selected by Dr Ethan Goddard-Borger at the

Walter and Eliza Hall Institute and their genes synthesized in codon-optimized form for expression in E. coli. Mr Alan John assembled seven expression plasmids by sub-cloning as various His6- and maltose binding protein (MBP)-tagged constructs. These included:

Aspergillus niger GH134 (XP_660314.1), Alteromonadaceae bacterium (AIF91558.1),

His6/MBP-tagged Alteromonadaceae bacterium (AIF91558.1) (pHisMALP-AbGH134),

Streptomyces sp. (WP_030268297.1), His6/MBP-tagged Streptomyces sp. GH134

(WP_030268297.1), and Rhizopus microsporus (CEG81953.1), His6/MBP-tagged

Rhizopus microsporus (CEG81953.1), With the assistance of Mr Alan John in the

Goddard-Borger laboratory, I transformed these plasmids into E. coli, log-phase cultures were induced by isopropyl β-D-1-thiogalactopyranoside (IPTG). After lysis and clarification by centrifugation, the supernatant was filtered and subjected to immobilised metal-ion affinity chromatography. Fractions containing product were identified by SDS- page, combined and purified by size exclusion gel filtration.

With the seven GH134 proteins in hand, digestion assays were used to determine if the enzymes displayed any activity towards the β-mannooligosaccharide M5. Assays were performed by incubating each enzyme (5µL) with commercially available M5 (0.5 mM) in 50 mM NaPi buffer (1 mL) for 1 h at 30 ˚C. TLC analysis of the reaction mixture allowed identification of Streptomyces sp. GH134 with an N-terminal hexahistidine-

100

tagged MBP fusion partner, termed SsGH134, as the most active; M5 was hydrolyzed into two products.

Thin layer chromatography (TLC) analysis of M1-M6 cleavage by SsGH134

Based on its superior activity, we sought to assess the ability of SsGH134 to process β-

1,4-oligomannosides. SsGH134 was incubated with mannose (M1) and one of five potential β-1,4-oligomannoside substrates (M2-M6), for 30 min. Aliquots of each reaction mixture were loaded onto a TLC plate and eluted with butanol/ethanol/water

(10:8:7, v/v) then stained with an orcinol/sulfuric acid reagent. Figure 4.2 shows the TLC of the reaction mixtures, with (+) or without (-) enzyme, and with a ladder of the six sugars (M1-M6) as a reference. No activity was observed with M2, M3 or M4, suggesting that SsGH134 requires occupancy of at least 5 subsites for catalysis. M5 was cleaved into two components, which were assigned as M2 and M3. M6 displayed two modes of hydrolysis, one yielding two molecules of M3, the other yielding M2 and M4. Based on the intensity of staining there appears to be more M3 present, suggesting this is the preferred mode of hydrolysis.

101

Figure 4.2: TLC analysis of M2-M6 hydrolysis by SsGH134. M1, mannose; M2, mannobiose; M3, mannotriose; M4, mannotetraose; M5, mannopentaose; M6, mannohexaose; (-) without enzyme; (+) with enzyme. TLC was eluted with butanol/ethanol/water (10:8:7, v/v) and visualized by staining with an orcinol/sulfuric acid reagent.

18 Mass spectrometric analysis of SsGH134 hydrolysis of M5 and M6 in H2O and O- labelled H2O

To confirm the digestion products of M5 and M6, the experiment was conducted in H2O and monitored by mass spectroscopy. Using this technique, we confirmed that M5 hydrolysis gave rise to M2 and M3; and that M6 hydrolysis gave rise to M2, M3 and M4.

No transglycosylation products could be detected by mass spectroscopy.

To define the site of cleavage within M5 and M6 oligosaccharides enzymatic hydrolysis was performed in 18O-labelled water. Upon hydrolysis 18O-labelled water is

142,144,145 18 incorporated into the non-reducing fragment. M5 cleavage in O-labelled water produces M2 and M3+2 quasi-molecular ions, with 18O incorporation into the latter fragment indicating hydrolysis when substrate bound across the -3 → +2 subsites (Figure

4.3a).146 Analysis of the products of M6 hydrolysis revealed M3 and M3+2, and M4 and

M2+2 quasi-molecular ions, showing that hydrolysis occurs preferentially for substrate

102

bound across the -3 → +3 subsites, with lower amounts of product formed for substrate binding across the -4 → +2 subsites (Figure 4.3b,c).

[M3[M3+NH + 2 + 4+2 [M2+NH4] + NH4] [M2 +

+ NH4]

[M3 + [M3+N [M3[M3+NH + 2 + 4+

+ NH4] + NH4]

[M2 + [M4 + 2 + [M2+NH4] + [M4+NH4+2 ] + + NH4] NH4]

18 Figure 4.3: ESI-mass spectra of the products of hydrolysis by SsGH134 in H2 O. Isotopic distribution reveals: a. M5 hydrolysis to M3 and M2+2; b. M6 hydrolysis yielding M3 and M3+2; c. M6 hydrolysis yielding M4 and M2+2.

103

Determination of stereochemistry of hydrolysis

With cleavage sites defined, we next assessed the stereochemistry of substrate cleavage to determine if SsGH134 acts with retention or inversion of anomeric configuration. A similar experiment has been performed Bolam and co-workers in 1996. They used 1H

NMR spectroscopy to demonstrate the stereochemical outcome of hydrolysis of M4 to

M2 by a family GH26 enzyme, mannanase A (MANA), isolated from Pseudomonas

147 fluorescens. MANA was incubated with M4 in D2O and the course of the reaction was monitored over the course of several hours (Figure 4.4a). For both M4 and M2, the H1α and H1β peaks are located at  5.17 ppm and  4.90 ppm, respectively. Given that the substrate M4 and the product M2 have the same H1 shifts, it was not possible to monitor the formation of a new signal. Instead, the progress of the reaction was monitored by observing the changing integration ratios of the peaks at t = 2, 30 and 960 min (Figure

4.4b, c, d, e). They found that addition of MANA changed the starting δ

5.17(H1a):4.90(H1b) ppm ratio from 65:35 (representing anomeric equilibrium) to a maximum of 165:35 at t = 2 min, and with time the peaks reverted to the equilibrium

65:35 ratio. This indicated that the first formed product was α-configured and hence the enzyme acted with retention of anomeric configuration.

104

b. c. d. e.

B A B A

B A

Figure 4.4: a. Mannotetrose (M4) cleavage by MANA. 1H NMR time-course of the reaction; the signals are H1-α (A), H1-β (B) of mannobiose and mannotetraose and H-1′ (C) of mannobiose; b. spectrum at 0; c. 2; d. 30; e. 960 min.147

In our case SsGH134 was not active on M4; however, given that the H1 chemical shifts of M5 and M6 are identical to that of M2, M3 and M4 this approach was viable and we chose to work with commercially available M5. Upon hydrolysis of M5 we expected to see an initial increase in either the H1α or H1β peaks, then a return to the equilibrium

65:35 ratio upon mutarotation. Accordingly, SsGH134 was incubated with M5 in 10 mM sodium phosphate buffer in D2O and the hydrolysis reaction was monitored over 2 h. At this time, TLC indicated complete consumption of substrate and formation two products, which were purified by silica gel chromatography and identified as M2 and M3. However, there was no detectable change in the integration of the H1 peaks over the course of the reaction. The analysis was complicated by a significant overlap of the H1β peak and the

HOD peak, which hindered accurate integration of the H1β peak. Water suppression

105

techniques were not useful in suppressing the water peak as they also suppressed the H1β peak. HOD in D2O has a steep dependence of chemical shift on temperature: every 5 °C increase shifts the HDO resonance approximately 0.05 ppm upfield.148 The experiment was therefore repeated at a higher temperature (35 ˚C) in a bid to separate the H1β and

HOD peak. While the HDO peak moved upfield and reduced overlap with the H-1β signal, at the elevated temperature no changes in the integration of the anomeric protons could be distinguished over the course of the reaction, possibly because the rate of mutarotation was enhanced. In their original study of A. nidulans Man134A, Shimizu and

-1 -1 co-workers reported a much higher catalytic activity for M6 (kcat/Km = 4,600 s mM )

-1 -1 140 compared to M5 (kcat/Km = 48 s mM ). We speculated that a similar situation may apply for SsGH134, and therefore we repeated the NMR experiment with SsGH134 and

M6 at 35 ˚C, hoping this substrate would have a higher catalytic efficiency than M5 and be hydrolysed faster than the rate of mutarotation. This approach was not fruitful and therefore it was clear we had to address the issue of spectral overlap in a different way.

In considering the problem we faced, it was clear that one of the challenges was that the anomeric protons of the substrate overlapped with the product(s), so that a large change in intensity was needed to unequivocally determine the stereochemical outcome of hydrolysis. Our attention therefore turned to establishing a means to chemically convert the substrate into a derivative that had a different chemical shift for the anomers.

It is known that acyl hydrazines react readily with sugar hemiacetals in the presence of acetic acid in water to form β-glycosyl acylhydrazides We therefore treated M6 with benzoyl hydrazine in refluxing water and acetic acid to afford the mannohexaosyl benzoylhydrazide (401), resulting in a shift of the reducing end anomeric proton to δ 4.42 ppm and providing a clear spectral window in the range δ 4.85-5.25 ppm (Scheme 4.3).

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Scheme 4.3: Synthesis of β-mannohexaosyl benzoylhydrazide (401). Reagents and conditions: a. benzoyl hydrazide, AcOH, H2O, 68%.

Before conducting an NMR time-course, we first monitored the hydrolysis of 401 by mass spectrometry to determine if this compound is a substrate for SsGH134 (Figure

4.5). Like M6, compound 401 displayed two modes of hydrolysis, one giving a molecule of M3 and M3 benzoylhydrazine, the other providing M4 and M2 benzoylhydrazine, consistent with the results of SsGH134 catalysed hydrolysis of M6 in 18O-labelled water.

+ [M3N2H4COPh+H]

+ + [M3+Na] [M2N2H4COPh+H]

[M4+Na]+

Figure 4.5: Mass spectrum of the products of hydrolysis of M6-benzoylhydrazine by SsGH13. M6 hydrazine (401) to M3 and M3 hydrazine, and to M4 and M2 hydrazine.

An aliquot of SsGH134 was added to a solution of M6 benzoylhydrazine in 10 nM sodium phosphate buffer in D2O and the hydrolysis reaction was monitored over 40

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min (Figure 4.6). At t = 0 there are no anomeric protons corresponding to the hemiacetal.

At t = 2 min two new peaks appear at δ 5.20 and 4.92 ppm assigned as H1α and H1β of

M2, M3 and M4; the ratio of peaks was 0.85:0.15 (H1α:H1β). By 40 min the reaction was complete and the anomeric mixture had mutarotated to a final ratio of 0.69:0.31

(H1α:H1β). These results demonstrated initial formation of the α-anomer of M2, M3 and

M4, thereby classifying SsGH134, and by inference family GH134 enzymes, as inverting.

Figure 4.6: M6-benzoylhydrazide was incubated with SsGH134 and the stereochemistry of the reaction was monitored by 1H NMR spectroscopy. At t = 0 there are no anomeric protons; at t = 2 min, two new peaks are visible at δ 5.20 and 4.92 ppm, assigned as H1α and H1β, respectively, of M2, M3 and M3. The enzyme-catalyzed reaction and mutarotation is at equilibrium by t = 40 min.

The rate of mutarotation of the mixture of α-M2, -M3 and -M4 to the corresponding β-anomers was determined to be 0.0307 min-1 at 25.1˚C (Figure 4.7). This

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is similar to the rate of mutarotation of α-D-mannose to β-D-mannose, reported as 0.0297 min-1 at 24.9 ˚C.149

0.5

0.4

(ppm)  0.3

0.2

0.1

Integration of H1- Integration 0.0 0 5 10 15 20 25 30 35 40 45 Time (min)

Figure 4.7: Plot of the rate of mutarotation of the mixture of α-M2, -M3 and -M4 to the corresponding β-anomers. Integration ratio of H1β = (integral H1β)/(sum of integrals of H1α and H1β).

Structural analysis of SsGH134

Structural analysis of SsGH134 was performed by Dr Yi Jin in Prof. Gideon Davies laboratory at the University of York. The key details are summarized below. The three- dimensional structure of SsGH134, the first GH134 representative, was solved at 1 Å resolution, revealing a mixed α-helix/β-sheet fold. This type of structure bears a superficial resemblance to family GH19 chitinases,150 GH22 C-type lysozymes,8 GH23

G-type lysozymes,11 and GH124 cellulases.151 Structural similarity searches using

PDBeFold152 determined GH134 is most similar to GH22 C-type lysozymes. The most notable member of GH22 is HEWL, a retaining enzyme that acts though a classical double displacement mechanism catalysed by amino acid residues D52 and E35.8 In contrast

GH134 enzymes act with inversion of anomeric configuration most likely through a classical single displacement mechanism requiring two residues acting as an acid/base.

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The structure of SsGH134 was overlaid with that of family GH22 members in complex with substrate. This led to the identification of a likely active site cleft, and possible catalytic residues. The SsGH134 E45Q variant was constructed by site-directed mutagenesis and a complex obtained with M5 (Figure 4.8).

a. b.

Figure 4.8: Three-dimensional structure of SsGH134 enzyme in complex with β-1,4- mannopentose; a. Overview of the C- to N-terminus; b. Surface representation of complex coloured by conservation according to sequence alignment (cyan = variable, burgundy = conserved).

Comparison of the SsGH134-M5 complex with that of HEWL in complex with chitohexoase shows the acid/base of HEWL, E35, overlays well with E45 (Q in the mutated variant) of SsGH134 (Figure 4.9). The nucleophile D52 of HEWL overlays well with N65 of SsGH134. Given asparagine is a poor nucleophile, it is unlikely that this residue is involved in mechanism, which is consistent with the observation that GH134 enzymes act with inversion of stereochemistry. Similar overlap was seen for active site residues from representatives of families GH19, 23, and 124.

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Q45(E) N65 Q35(E) D52

D57

N44

Figure 4.9: Close up overlay of the catalytic residues of SsGH134 in green and HEWL (GH22) in yellow.

A three-dimensional structure of wild type SsGH134 in complex with M3 was solved at 1.2 Å, in which M3 binds to the −3 → −1 subsites, and in which the reducing

1 end sugar was bound as the -anomer in a C4 conformation (Figure 4.10a). This is the stereochemistry of the product formed in the inverting mechanism, and thus this crystal structure represents a product complex. This crystal structure also implicated E45 as positioned to act as the acid; D57 was located in an ideal position to act as base to deprotonate a water molecule in a single displacement inverting mechanism. Mutagenesis studies confirmed that D57N and E45Q variants were inactive against M5 or M6.

a. b.

Figure 4.10: Complex of a. wildtype SsGH134 and M3; b. SsGH134-E45Q and M5.

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The electron density for mannopentose in the pseudo Michaelis complex with the catalytically inactive mutant SsGH134-E45Q is shown in Figure 4.10b. This revealed that, with the exception of the −1 subsite, all the other sugar rings are in a ground state

4 1 C1 conformation. The sugar in the −1 subsite adopts an unusual ring-flipped C4 conformation where the C1-O bond is orientated axially, and the nucleophilic water is ideally positioned for an in-line attack at C1. This is suggestive of a Southern hemisphere

1 3 ‡ 3 C4→ H4 → H1 conformational itinerary along the reaction co-ordinate, with the product

1 relaxing to a Michaelis-mimicking C4 conformation upon completion of the reaction.

This is a novel conformational itinerary that has never been reported for a β-mannosidase

3 3 ‡ 1 or β-mannanase. However a reversed itinerary H1→ H4 → C4 itinerary has been reported for the α-mannanases of family GH47.153

Computational studies

Classical molecular dynamics (MD) and QM/MM metadynamics were performed by Mr

Lluis Raich in the laboratory of Prof Carme Rovira at the University of Barcelona. The structure of SsGH134 in complex with mannopentaose was used as the starting point. To simulate the wild type enzyme, the mutation of the acid residue (E45Q) was manually reverted (changing atom N by O without modifying its orientation). The free energy landscape (FEL) of the reaction was explored using the metadynamics approach with three collective variables (CVs): (CV1) nucleophilic attack of water, (CV2) proton transfer between D57 and the water molecule, and (CV3) transfer of the E45 proton to the glycosidic oxygen (Figure 4.11a). The reactant initially adopt an conformation with

1 3 the -1 sugar between C4 and E. Upon commencement of the reaction, the glycosidic bond elongates and a proton is transferred from the E45 to the glycosidic oxygen (Figure

1 3 3 3 4.11b). The mannopyranose ring in the -1 subsite distorts from C4/ E to a H4/ E conformation where upon proton transfer from E45 is complete, the glycosidic bond is

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broken, and a new bond between the water molecule and anomeric carbon is partially

3 3 3,O 3 formed. The mannopyranose ring in the -1 subsite changes from H4/ E to B/ S1 conformation as proton transfer from the water to D57 occurs. The sugar in the -1 subsite

1 then relaxes to a C4 conformation, losing the interaction between the anomeric OH and

D57. This conformation matches that observed in the product complex of SsGH134 and

4 154 M3 and likely relaxes to the C1 conformation as the product exits the active site.

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Figure 4.11: QM/MM simulations of SsGH134 mechanism. a. FEL of SsGH134 catalyzed reaction. b. Reaction co-ordinate itinerary for SsGH134 inverting endo- β-1,4- mannanases, R represents the reactants (Michaelis complex), TS the transition state, P the products and P’ the product complex after a water molecule enters and replaces OH of mannose.

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4.6 Conclusions

β-1,4-Mannanases from families GH5, 26 and 113 are retaining enzymes that operate

1 ‡ O through S5→B2,5 → S2 conformational itineraries. In this work we revealed that β-1,4- mannanases of family GH134 differ from other β-mannanase families in both their mechanism and tertiary structure. We showed that a representative GH134 from

Streptomyces sp. has a fold closely related to that of the retaining GH22 hen egg white lysozyme (HEWL) but acts with inversion of stereochemistry. GH134 utilizes a unique

1 3 ‡ 3 C4→ H4 → H1 conformational itinerary along the reaction co-ordinate, different to any known β-1,4-mannanase.

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4.7 Experimental

Protein Expression Escherichia coli BL21 (DE3) transformed with pHisMALP-SsGH134 were grown in 1 L LB media with shaking (200 rpm) at 37 C (100 μg ml–1 ampicillin) until the culture reached an OD600 of 0.8. The culture was cooled to room temperature, IPTG added to a final concentration of 200 μM, and then it was incubated with shaking (200 rpm) at 18 °C for 16 h. Cells were harvested by centrifugation (17,000 g, 20 min, 4 C) and resuspended in 40 ml of binding buffer (50 mM NaPi, 500 mM NaCl, 5 mM imidazole, pH 7.5) containing EDTA-free protease inhibitor cocktail and lysozyme (0.1 mg/ml) by nutating at 4°C for 30 min. Benzonase (1 ul) was added to the mixture and lysis was effected by sonication. The lysate was centrifuged (17,000 g, 20 min, 4 C) and the supernatant collected. The supernatant was filtered (0.45 μm) and then subjected to immobilized metal-ion affinity chromatography. Fractions containing product (as determined by SDS-PAGE) were combined and further purified by size exclusion chromatography (GE HiLoad 16/600 Superdex 200) using 50 mM sodium phosphate, 150 mM NaCl, pH 7.5 buffer. The protein obtained was estimated to be >95% pure by Coomassie-stained SDS-PAGE. Protein concentration was determined by Bradford assay. The yield of MBP-GH134 fusion protein was >50 mg L–1.

TLC analysis of M1-M6 hydrolysis by SsGH134. TLC plates (Silica gel 60, 20 × 20, Merck) were cut to 10 cm in height. 2 μL of samples were spotted on the plate, separated by 10 mm. Solvent (50 ml) comprising freshly made 1-butanol/ethanol/water (10:8:7, v/v) was poured into a glass chromatography tank (23 × 23 × 7.5) and covered tightly. Vapors were allowed to equilibrate for at least 2 h before use. SsGH134 (25 µM ) was incubated with substrate (0.1 mg) in H2O (25 µL) at r.t. for 30 min; M1, mannose; M2, mannobiose; M3, mannotriose; M4, mannotetraose; M5, mannopentaose; M6, mannohexaose; (-) without enzyme; (+) with enzyme. The TLC plate was loaded and placed into the tank and samples allowed to migrate until the running buffer reached approximately 1 cm from the top of the plate. The plate was dried gently using a hairdryer and returned to the tank and eluted a second time. The spots were visualized with orcinol/sulfuric acid reagent (H2SO4/EtOH/H2O 3:70:20 v/v, 1% orcinol), dried carefully and heated until sugars were developed, at 120 °C (5-10 min). Standards consisting of known monosaccharides and oligosaccharides were spotted on the TLC plate. 116

18O isotope-mapping of SsGH134 catalysed cleavage of β-1,4-mannopentaose Solutions of 0.5 mg of mannotetraose (M4), mannopentose (M5), and mannohexose 18 (M6), respectively, were each dissolved into 50 μL of H2 O. 1 μL of SsGH134 (25 mg ml-1in 50 mM NaPi and 150 mM NaCl at pH 7.5) was added to each solution. After 30 min the solution was analyzed using an Agilent ESI-TOF mass spectrometer using electrospray ionisation in positive ion mode.

β-1,4-mannohexaosyl benzoylhydrazine (M6-benzoylhydrazine) (401)

Mannohexaose (4 mg, 0.004 mmol) was refluxed in H2O (1 ml) with benzoyl hydrazide (0.8 mg, 0.006 mmol) and acetic acid (25 μl of a 0.5% solution in water) for 2.5 h. The solution was evaporated to dryness. Reverse phase chromatography (Alltech Prevail C18- silica), eluted with water gave M6-benzoylhydrazide (3 mg, 68%); 1H NMR (500 MHz,

CDCl3): δ 3.46-3.42 (44H, m, 5 × H1, 6 × H2, 6 × H3, 6 × H4, 6 × H5, 12 × H6, 4 × NH), 4.44 (1H, s, H1β), 4.75 (1H, s, H1α), 7.54-7.89 (12H, m, 2 × Ph); HRMS (ESI)+ m/z + 1109.3818 [C43H68N2O31(M+H) requires 1109.3879].

Determination of stereochemistry of catalysis by 1H NMR analysis SsGH123 catalysed hydrolysis of M6-benzoylhydrazide was monitored by 1H NMR spectroscopy using a 500 MHz instrument. A solution of SsGH123 in buffered D2O (0.15 ml, 0.095 mM in 10 mM sodium phosphate, pD 7.4) was added to a solution of M6- hydrazine (3.0 mg, 0.0027 mmol) in buffered D2O (0.6 ml, 10 mM sodium phosphate, pD 7.4 ) at 25 °C. 1H NMR spectra were acquired at time points (t = 0, 2, 3, 10 and 40 min).

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CHAPTER FIVE:

Investigation of sulfoglycolysis in Escherichia coli

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5.1 Plant sulfolipid SQDG its role in the biosphere

Discovery of plant sulfolipid: SQDG

Sulfur is an important element that is required by organisms for the biosynthesis of essential amino acids (cysteine and methionine), cofactors (biotin, glutathione, coenzyme

A lipoic acid) and the osmolyte dimethylsulfoniopropionate.77 Sulfur exists in a wide

2- 2- range of redox states from +6 (SO4 ) to -2 (S ) and its fate is heavily dependent on microbial activity.155 Although mostly overlooked, the plant sulfolipid sulfoquinovosyl diacylglycerol (SQDG) is now recognized as a major component of the global sulfur cycle, produced in amount of 10 billion tonnes per annum, commensurate with estimates of cysteine and methionine biosynthesis (Figure 5.1a.).

Despite its ubiquity, the discovery of SQDG was only reported relatively recently, by Benson and co-workers in 1959.156 Several photosynthetic microorganisms and higher

35 2- plants were grown in media containing radiosulfate ( SO4 ), and the cells were centrifuged and extracted into ethanol or ethanol/chloroform (75:25). Thin layer chromatography of the soluble lipids revealed a sulfolipid with high (90%) 35S activity.

This sulfolipid could be cleaved by a β-galactosidase isolated from Escherichia coli to give glycerol and a 6-sulfo-sugar initially proposed to be 6-deoxy-6-sulfo-D-galactose, suggesting that the sulfolipid was a 6-deoxy-6-sulfo-β-D-galactoside. Subsequent work revealed that cleavage of the sulfoglycosyl glyceride by the β-galactosidase was anomalous and the proposed D-galacto stereochemistry was called into question.157 A sample of the sulfolipid was deacylated and yielded a sulfoglycoside that exhibited an optical rotation characteristic of an alkyl α-D-glucopyranoside, and the low-field NMR spectrum revealed a chemical shift consistent with an equatorial anomeric proton of D- glucose. Furthermore, the glyceride was converted to a methyl sulfoglycoside whose structure was verified as methyl 6-deoxy-6-sulfo-α-D-glucopyranoside (sulfoquinovose,

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SQ, Figure 5.1 b.).158 Based on this analysis, a revised structure for the sulfolipid was proposed: 6-deoxy-6-sulfo-α-D-glucoside glyceride (SQGro, Figure 5.1c.). The glycerol configuration was determined using a radiochemical tracer method. SQGro isolated from the hydrolysis of 14C-labelled Chlorella pyrenoidosa sulfolipid was oxidized to the glyceric acid glycoside, and the glycoside then cleaved and its configuration determined as L-(+)-glyceric acid by radiotracer analysis by co-precipitation with unlabelled authentic enantiomers.159 Unambiguous confirmation of its structure was subsequently achieved by X-ray crystallographic studies of the rubidium salt of SQGro, one of the earliest uses of X-ray diffraction in the determination of a natural product structure.160

Figure 5.1: Plant sulfolipid SQDG and its metabolites.

5.2 Occurrence and biosynthesis of SQDG

SQDG is produced within the thylakoid membranes of chloroplasts of photosynthetic organisms (mosses, ferns, higher plants and algae), photosynthetic apicomplexa, dinoflagellates, diatoms and most photosynthetic bacteria.161 It typically comprises 4-7% of total plant lipids and 2-50% of total polar lipids in marine plants and algal tissues.162

Isolated chloroplasts contain all the necessary enzymes and substrates required for synthesis of SQDG. In 1989, Heinz and co-workers loaded chloroplasts with radiolabelled diacylglycerol (DAG), which they permeabilized and then treated with several sulfosugar nucleotides: ADP-SQ, CDP-SQ, GDP-SQ and UDP-SQ.

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Radiolabelled SQDG synthesis increased dramatically only upon addition of UDP-SQ, identifying it as the donor used in the final step of SQDG biosynthesis.163,164

Our current understanding of SQDG biosynthesis can be illustrated through the process that has been characterized at the molecular level in the model plant, Arabidopsis thaliana (Arabidopsis) (Figure 5.2). Within the stroma of Arabidopsis chloroplasts, UDP- glucose pyrophosphorylase3 (UDP3) catalyses the formation of UDP-Glc from glucose-

1-phospate (Glc1P) and uridine triphosphate (UTP).165 In the second step of SQDG biosynthesis, UDP-SQ synthase (SQD1) substitutes the 6-OH of Glc-UDP with sulfite, this occurs in a three step process via an enone in a NADPD-dependent manner. Lastly

UDP-SQ:DAG sulfoquinovosyltransferase (SQD2) catalyses the transfer of SQ from

UDP-SQ to DAG. A related pathway has been discovered in the cyanobacterium

Rhodobacter sphaeroides.166,167

Figure 5.2: Biosynthesis of SQDG in Arabidopsis thaliana.

5.3 Degradation of SQDG

The first step of SQDG catabolism in both plants and mammals appears to be delipidation.

In 1962 a sulfoglycolipase was discovered in the plant Scenedesmus obliquus, with has

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the ability to cleave the acyl groups of SQDG to afford lyso-SQGro and then SQGro

(Figure 5.3).168 In mammalian organisms, lipases secreted by pancreatic cells can deacylate SQDG to SQGro; furthermore intestinal gut microflora can metabolize the

2- 169 resultant SQGro, releasing free sulfate (SO4 ).

Figure 5.3: Overview of SQDG degradation.

Once SQDG has been deacylated to SQGro, it must be cleaved to SQ in order to enter the sulfoglycolytic pathway. This is accomplished by the action of a dedicated glycoside hydrolase, a sulfoquinovosidase (SQase). SQases are retaining glycosidases of family GH31 and operate though a classical double displacement mechanism. SQ- specific glycoside hydrolyses also have the ability to cleave SQDG to SQ; however, it is likely that they more commonly encounter the delipidated form, SQGro due to their periplasmic location within the cell.170

Two sulfoglycolytic pathways have been reported for the catabolism of SQ. In both pathways SQ is processed into a C3 intermediate, 3-sulfolactaldehyde (SLA, Figure 5.3), and then to different C3 products. The first sulfoglycolytic pathway reported produces

(S)-2,3-dihydroxypropane-1-sulfonate (DHPS) and mimics the Embden-Meyerhof-

Parnas (EMP) glycolytic pathway and hence has been named the sulfo-EMP pathway.171

The second sulfoglycolytic pathway to be reported produces 3-sulfolactate (SL) and mimics the Entner-Doudoroff (ED) glycolytic pathway and has been named the sulfo-ED pathway accordingly.172

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Figure 5.3: SQ metabolites.

Sulfo-EMP pathway

In 2014 Denger and co-workers reported that E. coli K-12 performs sulfoglycolysis in addition to normal glycolysis.171 Minimal media supplemented with 4 mM SQ as sole carbon source was shown to support growth of E. coli yielding half the amount of total cellular protein compared to media containing 4 mM glucose.173 Denger and co-workers showed that sulfoglycolysis by E. coli led to production of DHPS, which is excreted into the media. Spent media from E. coli K-12 cultures grown on SQ were filter sterilized and inoculated with Cupriavidus pinatubonensis JMP134, an organism that can grow on

DHPS as sole carbon source. This organism consumed the DHPS and released an equimolar amount of sulfate. This demonstrated that collectively, this two member community can achieve the complete biomineralization of SQ to sulfate.171,174

In order to identify the pathway used to perform sulfoglycolysis, proteins from E. coli grown on either glucose or SQ were subject to two-dimensional polyacrylamide gel electrophoresis (2D-PAGE), allowing visualization of proteins that were upregulated in the presence of SQ. The upregulated proteins were excised from the gel and identified by peptide fingerprinting–mass spectrometry. The SQ inducible proteins were localized to a

10-gene cluster, annotated as ompL and yihO-yihW (Figure 5.4). Transcriptional analysis showed that genes within this cluster were strongly transcribed when cells were grown on SQ. Confirmation of the role of this cluster in encoding the capacity to grow on SQ

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was obtained by the observation that single gene knockouts of yihO, yihS, yihT and yihV did not grow on SQ.171

Figure 5.4: Gene loci encoding sulfo-EMP in E. coli K12 (experimentally determined genes outlined).

Denger and co-workers biochemically characterized four core enzymes, YihS,

YihT, YihU and YihV, to understand their roles in metabolism of SQ, which allowed them to propose a pathway for sulfoglycolysis in E. coli. They proposed that the gene cluster (Figure 5.4) encodes for a predicted transporter (YihO) for importing SQ and SQ glycosides; a predicted SQase (YihQ) to hydrolyze SQ glycosides to SQ; a predicted mutarotase (YihR) to catalyze conversion of α-SQ to β-SQ; an isomerase (YihS), to isomerase SQ to sulfofructose (SF); a kinase (YihV) to phosphorylate SF to SF-1- phosphate (SFP); an aldolase (YihT) to convert SFP into dihydroxyacetone phosphate and (S)-SLA; a reductase (YihU) to convert SLA into (S)-2,3-dihydroxypropane-1- sulfonate (DHPS) and a second predicted transporter (YihP) to export DHPS from the cell.171,161 The proposed pathway and location of the sulfoglycolytic proteins is shown in

Figure 5.5, along with a comparison of equivalent steps in the EMD glycolysis.

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Figure 5.5: a. Overview of sulfo-EMP sulfoglycolytic pathway in E. coli as described by Denger and co-workers;171 b. Overview of EMP glycolytic pathway.

Sulfo-ED pathway

The ED sulfogycolytic pathway is used by some prokaryotes and was first characterised in the organism Pseudomonas putida SQ1 (Figure 5.6a.) by Felux and co-workers.173 P. putida can utilize SQ as a sole carbon source, releasing equimolar SL. P. putida; expressed NAD+ dependent dehydrogenase activity that oxidised SQ to 6-deoxy-6-sulfo-

D-gluconate lactone (SLA), analogous to the glycolytic ED pathway (Figure 5.6b). Cells were grown in glucose or SQ and cell lysates compared by 2D-PAGE, peptide fingerprint

MS and total-proteome analysis.172 This data allowed the identification of a gene cluster

(Figure 5.7) containing; two predicted transported proteins (PpSQ_00095, PpSQ_00098); a predicted SQase to hydrolyse SQ glycosides to SQ (PpSQ_00094); a predicted SQ mutarotase (PpSQ_00092); an SQ dehydrogenase (PpSQ_00090) to oxidize SQ to SGL; a sulfogluconate lactonase (PpSQ_00091) to hydrolyze SGL to 6-deoxy-6-sulfo-D- gluconate (SG); an SG dehydratase (PpSQ_00089) to convert SG into 2-keto-3,6- dideoxy-6-sulfo-D-gluconate (KDSG); an aldolase (PpSQ_00100) to convert KDSG into pyruvate (PYR) and SLA; an SLA dehydrogenase (PpSQ_00088) to convert SLA into

SL and the predicted transporter (PpSQ_00097), which may export SL from the cell.172,173

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Figure 5.6: a. Overview of sulfo-ED sulfoglycolytic pathway in P. putida SQ1 as described by Felux and co-workers172; b. Overview of ED glycolytic pathway.

Figure 5.7: Gene loci encoding sulfo-ED in P. putida SQ1 (experimentally determined

genes outlined).

5.4 Catabolism of SQ in E. coli

SQ itself is rarely found as the free sugar and exists almost exclusively as a glycoconjugate (SQDG, lyso-SQDG or SQGro). For organisms to catabolise SQ though either the EMP or ED sulfoglycolysis pathways, it needs to be freed from its glycoconjugates. Within the sulfo-EMP E. coli K-12 gene cluster there is a gene (YihQ) that encodes a protein assigned to family GH31 within the CAZy sequence-based classification. This family contains enzymes with α-glucosidase, α-glucan lyase and α- xylanase activities. Early attempts to define the function of this protein failed to provide convincing evidence for any catalytic activity for YihQ or its function in E. coli.115,175

Levels of YihQ are upregulated upon growth of E. coli on SQ as sole carbon source.171 In 2016 Speciale and co-workers identified E. coli YihQ as the first dedicated

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sulfoquinovosidase and gateway enzyme to sulfoglycolytic pathways.170 YihQ was cloned and expressed then incubated with SQDG or SQGro. LCMS analysis of the reaction mixture revealed complete conversion to SQ. The hydrolysis of the SQ glycoside

4-nitrophenyl α-D-sulfoquinovoside (PNPSQ) by YihQ was monitored with 1H NMR spectroscopy, revealing that like other family GH31 members, YihQ acts with retention of stereochemistry. YihQ was shown to act through a classical Koshland retaining mechanism employing two catalytic residues, a catalytic nucleophilic carboxylate (D405) and an acid/base residue (D472), through kinetic studies with mutants and by trapping a glycosyl enzyme intermediate using 5-fluoro-β-L-idopyranosyl fluoride. Kinetic studies of YihQ did not detect any α-glucosidase activity, defining this enzyme as a dedicated sulfoquinovosidase. The three-dimensional structure of YihQ reveals an (αβ)8 barrel, similar to other members of family GH31, however the active site contains residues that bind the sulfonate moiety and provide specificity for SQ over Glu (Figure 5.8).

Figure 5.8: Cartoon depicting active site showing major hydrogen-bonding interactions of PNPSQ with YihQ D472N mutant.

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In work published recently, a homolog of E. coli YihR from Herbaspirillium seropedicae was shown to be an SQ mutarotase, with a 17,000-fold preference for SQ versus glucose-6-phosphate.176 SQ mutarotase activity was demonstrated using NMR exchange spectroscopy (EXSY), a method that allows chemical exchange of magnetization between the α- and β-SQ anomers at equilibrium. HsSQM was found to catalyse the mutarotation of several other aldohexoses containing an equatorial 2-OH including: D-galactose, D-glucose and D-glucuronic acid, but not D-mannose.

Figure 5.9: Processing of SQ-glycosides (SQOR) by E. coli YihQ and YihR in preparation for sulfoglycolysis.

Collectively, this data suggests that YihQ is required for the growth of E. coli on

SQ glycosides. However, the ability of E. coli to grow on SQ glycosides has never been studied. In fact, while several other bacteria have been reported to grow on SQ, the only bacterium known to grow on an SQ glycoside is Flavobacterium sp., which as described earlier operates through a sulfo-ED pathway. It was found to be able to grow on the methyl a-glycoside of SQ (SQMe) as a sole carbon source.177

5.5 Aims

This chapter aims to synthesize a diastereomeric mixture of the SQ glycoconjugate,

SQGro, and determine if both stereoisomers are substrates for the SQase YihQ from E. coli. With this compound in hand we will investigate if it and SQMe can solely support growth of E. coli.

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5.6 Results and Discussion

Previous syntheses of SQGro

The synthesis of SQGro was first described in 1962 by Miyano and Benson beginning with 1,2:5,6-di-O-isopropylidene-α-D-glucofuranose (Scheme 5.1).178 Selective hydrolysis of the 5,6-isopropylidene group followed by tosylation of the primary position gave 1,2-O-isopropylidene-6-O-tosyl-α-D-glucofuranose. Sulfite substitution of the tosylate, followed by cleavage of the remaining isopropylidene ring afforded SQ, which was isolated as the barium salt. The SQ barium salt was converted to the free acid then treated with allyl alcohol to perform a Fischer glycosidation. The mixture of anomers of the allyl sulfoquinovosides were converted to the cyclohexylammonium salt and recrystallized from ethanol/ethyl acetate. The cyclohexylamine salt was converted to the free acid and the double bond dihydroxylated with potassium hydroxide and potassium permanganate. Manganese dioxide was removed by filtration and the filtrate was passed through three different ion-exchange columns, providing the sulfonic acid. Treatment of the acid with cyclohexylamine and selective recrystallization afforded a pure sample of cyclohexylammonium SQGro as the 2’R isomer.

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Scheme 5.1: Miyano and Benson’s 1962 synthesis of SQGro.178,179

In 1998 Roy and Hewlins reported an improved preparation of SQGro (Scheme

5.2).180 This procedure started by synthesis of SQ as the potassium salt (following the method of Miyano and Benson178) which was refluxed with Dowex 50 resin (H+ form) in allyl alcohol to give allyl SQ in 62% yield, marginally higher than the 55% yield obtained by Miyano and Benson. Dihydroxylation was again achieved using NaOH and KMnO4, however the work up was simplified by isolating the SQGro as a brucinium salt thus eliminating the need for an additional tedious ion exchange column. The brucinium salt was converted to the cyclohexylammonium salt from which the pure 2’R isomer was isolated by recrystallisation in 12.5%.

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Scheme 5.2: Roy and Hewlins’ 1998 synthesis of SQGro.180

Synthesis of SQGro

In considering a possible approach to SQGro, we considered the approaches of both

Miyano and Benson, and Roy and Hewlins. Both require the initial synthesis of SQ, and then use a cumbersome approach to obtain the pure allyl glycoside. The dihydroxylation of the alkene to afford SQGro is especially low yielding, and again uses cumbersome approaches to isolate the product. We therefore considered an alternative approach that would more directly provide the target. We proposed to start from commercially available allyl α-D-glucopyranoside (Scheme 5.3). Installation of a thiobenzoate at the 6 position could be achieved with high regioselectivity with the Mitsunobu reaction. Oxidation of the thiobenzoate should lead to formation of the sulfonate, and subsequently, the double bond could be dihydroxylated to yield SQGro as a mixture of diastereomers. For the purposes of our experiments we were planning to use a mixture of the 2’R SQGro and

2’S SQGro isomers.

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Scheme 5.3: Proposed synthesis of SQGro (501)

Accordingly, we treated allyl α-D-glucopyranoside with triphenylphosphine,

DIAD and thiobenzoic acid in a Mitsunobu reaction, which gave the thiobenzoate (502) in 79% yield (Scheme 5.4). The thiobenzoate was oxidized with Oxone and potassium acetate in acetic acid to the sulfate (503), but in a disappointingly low yield of just 13%.

Careful analysis of the products of this reaction allowed identification of several by- products: the 4-O-benzoyl sulfonate (504, 8%), the epoxides (505, 34%) and the 4-O- benzoyl sulfonate epoxides (506, 42%). Compounds 505 and 506 were entirely unexpected; epoxidation of diols with Oxone typically requires a catalytic ketone and proceeds via a dioxirane intermediate.181

Scheme 5.4: Synthesis of the sulfonate (506): a. PPh3, DIAD, BzSH, THF, 79%; b. Oxone, AcOK, AcOH, 13%.

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Despite this discouraging result, we recognized that the epoxides potentially could be converted to SQGro (Scheme 5.5). Treatment of 506 with sodium carbonate in methanol led to transesterification of the benzoate ester to give 505 in 90% yield.

Treatment of 505 with potassium acetate in refluxing aqueous acetic acid, followed by potassium carbonate gave SQGro (501) as a 11:9 mixture of 2’R and 2’S diastereoisomers in 60% yield.

Scheme 5.5: Synthesis of SQGro (501). a. Na2CO3, MeOH, 90%; b. i. AcOK, H2O; ii K2CO3, MeOH, 60%.

We were thrilled that the epoxide could be cleaved to afford SQGro, and our attention turned to improve the epoxidation step so that epoxides 505 and 506 were formed exclusively. As remarked upon earlier, epoxidations achieved under the agency of Oxone usually require a catalytic ketone and proceed via a dioxirane. Nucleophilic

- - attack of the carbonyl carbon of the ketone by KHSO5 with subsequent loss of KHSO4 generates a dioxirane in situ. Transfer of oxygen from the dioxirane to the substrate forms the epoxide and regenerates the ketone allowing its recycling (Scheme 5.6).181 This process is less efficient below pH 7 and above pH 8.5 due to a competing Baeyer-Villiger reaction consumes the ketone.182

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Scheme 5.6: Overview of ketone-mediated epoxidation of a double bond.

Based on these considerations, we attempted epoxidation of 502 with Oxone and

KHCO3 in acetone/water (4:1), which gave the mixture of epoxides 505. Compound 505 was refluxed with potassium acetate for 1 h to afford SQGro as a 11:9 mixture of 2’R and

2’S diastereomers in 48% yield over 2 steps. The identity and ratio of these diastereomers was assigned by careful analysis of the 1H NMR spectrum with reference to Roy and

Hewlins’ assignment of 2’R-SQGro.180 We assigned the characteristic H1 protons of the diastereomers as the overlapping doublets δ 4.90-4.92 ppm (J = 3.80 Hz), these were the most downfield shifted signals in our spectrum. Our assignment agrees well with Roy and

Hewlins’ who observed H1 at δ 4.72 ppm (J = 3.80 Hz). There was only one signal in our spectrum that corresponded to a well-resolved single proton. This key signal was a doublet of doublets at δ 3.47 ppm (J = 7.3, 10.3 Hz). We assigned this proton as H1a’R, consistent with Roy and Hewlins’ (δ 3.27 ppm, J = 7, 10 Hz).180

Scheme 5.7: Synthesis of SQGro (501): a. Oxone, KHCO3, acetone/H2O 9:1; b.i. AcOK, H2O; ii. K2CO3, MeOH, 48% over 2 steps.

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Evaluation of SQGro as a substrate for YihQ

With SQGro in hand we sought to determine if both isomers were substrates for E. coli

YihQ (Scheme 5.7). This enzyme was recombinantly expressed, purified and kindly provided to us by James Lingford from Dr Ethan Goddard-Borger's lab at the Walter and

Eliza Hall Medical Institute. We monitored the hydrolysis of SQGro by YihQ by 1H NMR spectroscopy (Figure 5.10). Prior to addition of enzyme (t = 0), there were two overlapping doublets at  4.9 ppm corresponding to the H1 protons of the 2’R and 2’S diastereoisomers. A spectrum acquired 14 min after enzyme addition revealed the formation of a new doublet at  5.2 ppm (J = 3.9 Hz) which was assigned as α-H1 of SQ.

A spectrum acquired at 18 h revealed a new doublet at  4.7 ppm (J = 8 Hz), assigned as

β-H1 of SQ and indicating that mutarotation had occurred, and that both diastereoisomers of SQGro had been completely consumed.

Scheme 5.8: Hydrolysis of SQGro (501) by YihQ.

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Figure 5.10: Hydrolysis of SQGro (501) by YihQ monitored by 1H NMR spectroscopy. t = 0 prior to enzyme addition, H1 of 2’R and 2’S SQGro stereoisomers at 4.9 ppm; t = 14 min after enzyme addition shows appearance of new peak at 5.2 ppm assigned as α- H1 of SQ; t = 1 h after enzyme shows almost complete consumption of SQGro, appearance of β-H1 SQ at 4.7 ppm; t = 18 h complete consumption of SQGro and mutarotation of SQ α and β anomers

The disappearance of the signals for each diastereoisomer were plotted as a function of time and the data was fit to an exponential decay equation, allowing calculation of a first order rate (Figure 5.11). This revealed that the 2’R stereoisomer, which corresponds to the natural stereochemistry of SQDG/SQGro, is hydrolysed 6-fold faster than the 2’S stereoisomer. Incidentally, this data also confirms that hydrolysis of

SQGro catalyzed by YihQ occurs with retention of anomeric configuration, as previously reported using PNPSQ.170

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Figure 5.11: Rates of consumption of individual SQGro diastereoisomers by YihQ, fitted to an exponential decay.

With good evidence that YihQ SQase can hydrolyse both epimers of SQGro, we next sought to examine whether E. coli strain BW25113 can grow on SQGro as sole carbon source. We based our growth experiments on those reported by Denger et al. who studied growth of this E. coli strain on phosphate-based minimal media containing SQ as sole carbon source.171 E. coli strain BW25113 was initially cultured on glucose- containing M9 minimal media, then was sub-cultured into SQ-containing minimal media.

Growth in this media suffered an extended induction period, but after around 2 weeks, cells began to grow, as indicated by an increase in absorbance measured at 580 nm by a

UV/Vis spectrophotometer (OD580). Cells adapted to growth on SQ M9 media were used to inoculate 3 mL cultures containing 4 mM concentrations of the SQ glycosides methyl

α-D-sulfoquinovose (SQMe) and SQGro in M9 media. Upon sub-culturing SQ-adapted cells into fresh SQ-containing M9 media, subsequent recovery was much faster, with an induction period of 1-3 days. Cultures grown on SQMe took approximately 2 weeks to adapt to growth and subsequently 1-3 days to regrow. Cells adapted to growth on SQGro in only 3-4 days and subcultures just 1-2 days to re-grow, significantly faster than both

SQ and SQMe, suggesting SQGro is a preferred carbon source.

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With E. coli adapted for growth on several carbon sources, we chose to examine growth on Glu, Gro, SQGro and SQ as a sole carbon sources, by determining and comparing the growth curves. Growth was determined spectroscopically by taking samples at various times and monitoring optical density (OD580) (Figure 5.12). Cultures grown on SQ and Gro grew to a similar optical density, approximately half that of cultures grown in Glc and SQGro. The similar cell densities obtained for Gro and SQ, and Glc and SQGro, suggest that these pairs provide similar amounts of carbon to E. coli. SQ and

Gro both provide similar amounts of carbon to support growth of E. coli, namely 3 carbons each, while Glc and SQGro provide similar and approximately double amounts of carbon to E. coli suggesting that SQGro provides 6 carbons. The E. coli growth rates on Glc, Gro, SQGro and SQ were determined to be 0.11, 0.045, 0.086 and 0.0034 h-1 respectively, revealing SQGro is preferred to both SQ and Gro.

Figure 5.12: Growth of E. coli BW25113 in M9 minimal media containing 4 mM Glc, Gro, SQGro or SQ as sole carbon source at 30 °C. This data is representative of two independent experiments.

In order to confirm that growth on SQGro results in sulfoglycolysis, we sought to determine the concentration of DHPS in a sample of the spent culture supernatant. In order to quantify DHPS levels we undertook the synthesis of authentic DHPS, as the cyclohexylammonium salt (Scheme 5.8). This was performed by an undergraduate

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student, Ms Janice Mui, under my supervision. The approach taken was that developed by Friese in 1938, and more recently adapted by Mayer in 2010.174,183 Thus, allyl alcohol was treated with H2SO4 and Ac2O at 80° C. The solvent was evaporated and excess acetic acid azeotroped with water. Excess H2SO4 and DHPS were neutralised with BaCO3 to give Ba(DHPS)2. Ion exchange chromatography gave free DHPS, and this was treated with cyclohexylamine to give DHPS as the crystalline cyclohexylammonium salt.

Scheme 5.8: Janice Mui’s synthesis of DHPS as the cyclohexylammonium salt.

Quantitative analysis of DHPS was performed with the assistance of Dr Eileen

Ryan using LC-MS. Using the authentic cyclohexylammonium DHPS, a standard curve was established by LC-MS on a 5 cm HILIC column (Figure 5.13). Samples of the culture supernatant from spent M9 minimal media of E. coli grown to stationary phase on SQGro were studied by LC-MS revealing the complete consumption of SQGro and equimolar release of DHPS.

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6.0E+8 y = -2.095 x 106 x2 + 78.65 x 106 x 5.0E+8

4.0E+8

3.0E+8

Peak area Peak 2.0E+8

1.0E+8

0.0E+0 0.0 2.0 4.0 6.0 8.0 10.0 Concentration (mM)

Figure 5.13: a. ZIC-HILIC LC/MS chromatogram (extracted ion m/z 155) of spent M9 culture media of E. coli strain BW25113 grown on SQGro as sole carbon source. The peak at tR 10.45 was assigned as the anion derived from orotic acid. MS spectrum shows fragmentation pattern in negative ion mode for DHPS. b. Calibration curve for detection of DHPS in spent M9 minimal media. Final concentration of DHPS for E. coli strain BW25113 grown on 4 mM SQGro as sole carbon source.

5.7 Conclusions

In this chapter we have reported a novel approach to synthesizing SQGro as a mixture of

2’R and 2’S isomers in three steps from allyl α-D-glucopyranoside. We have shown that both SQGro isomers are substrates for YihQ, cleaving SQGro into SQ + Gro (Scheme

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5.9). The natural stereoisomer, 2’R-SQGro a 6-fold better substrate for YihQ than 2’S-

SQGro. SQGro can act as a carbon source for E. coli strain BW25113, and results in growth at a rate higher than that of either SQ or Gro. SQGro supports growth on par with glucose (which is known to provide 6-carbons to central metabolism) and releases equimolar DHPS through sulfoglycolysis. Collectively, this data supports the contention that SQGro is the naturally encountered form of SQ in the environment.

Scheme 5.9: Overview of SQGro metabolism. SQGro is imported into the cell by putative importer YihO. The SQase YihQ hydrolyses SQGro to SQ and Gro. SQ undergoes mutorotation cataluzed by YihR and enters the sulfoglycolysis pathway and is converted to DHPS which is ultimately exported from the cell by the putative transporter YihP. Gro is metabolized in the normal glycolysis pathway.

E. coli is largely found in the gastrointestinal tract of humans and other warm bodied animals.184 It is the primary facultative anaerobe of the human gastrointestinal tract, meaning it can grow both aerobically and anaerobically, whereas most intestinal microbes are obligate anaerobes. E. coli is found in the thin layer of mucus that lines the gut where it consumes oxygen and maintains the anaerobic nature within the core of the gut lumen.185 Consumption of SQDG leads to rapid deacylation and breakdown into

SQGro169 which is available for E. coli within the gut to metabolise. The SQGro may therefore promote growth of E. coli within the gut, maintaining the anerobic gut microenvironment and contributing to beneficial effects of E. coli colonization.

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5.8 Experimental

Allyl 6-5-benzoyl-6-deoxy-6-thio-α-thio-α-D-glucopyranoside (502) A solution of allyl α-D-glucopyranoside (1.00 g, 4.50 mmol) and thiobenzoic acid (0.65 mL, 5.45 mmol) in dry THF (15 mL) at 0

°C was added to a mixture of PPh3 (1.43 g, 5.45 mmol) and DIAD (1.07 mL, 5.45 mmol) in THF at 0 °C. The mixture was warmed to r.t. and stirred for 18 h. The mixture was concentrated and the crude residue purified by flash chromatography (EtOAc/pet. spirits 80:20) to give the thiobenzoate (5002) (79%, 1 1.23g) as a colourless oil. H NMR (500 MHz, CDCl3) δ 2.88 (1H, d, J = 8.6 Hz, H6a), 3.34 (t, J = 7.7 Hz, H4), 3.52-3.61 (d, J = 3.3 Hz, H2, H6b), 3.78 (t, J = 9.3 Hz, H3), 3.87 (1H, 3.3, 5.8, 9.3 Hz, H5), 4.03 (1H, m, H1a’), 4.21 (1H, J = 12.7, 5.2, H1b’), 4.90 (1H, J = 3.7 Hz, H1), 5.18 (1H, d, J = 10.2 Hz, H3a’), 5.28 (1H, dd, J = 17.2, Hz, H3b’), 5.90 (1H, ddd, J = 22.4, 11.0, 5.8 Hz, H2’) 7.45 (2H, t, J = 7.8 Hz, Ar), 7.60 (1H, t, J = 7.5 - + Hz, Ar), 7.97-7.99 (2H, m, Ar); HRMS (ESI) m/z 341.1056 [C16H20O6S (M+H ) requires 341.1053].

Sulfoquinovosyl glycerol (501) Oxone (1.36 g, 2.20 mmol)

and KHCO3 (449 mg, 4.48 mmol) were added portion- wise to a solution of 502 (95 mg, 0.28 mmol) in acetone/H2O 80:20 (2.5 mL). The mixture was stirred for 18 h, filtered with MeOH/H2O 2:1 (20 mL) and concentrated to dryness to give an enantiomeric mixture of epoxides (57 mg) as a white solid. Potassium acetate (138 mg, 0.696 mmol) was added to the crude epoxides in water (1 mL) and the mixture was refluxed for 1 h then the mixture was concentrated. K2CO3 (98.0 mg, 0.696 mmol) was added to the crude residue in MeOH (1mL) for 18 h, the mixture was concentrated and crude purified by flash chromatography (EtOAc/MeOH/water, 60:27:13) to give a mixture of (2’R)-D-glycer-1’-yl 6-deoxy-6-C-sulfonato-α-D-glucopyranoside and (2’S)- D-glycer-1’-yl 6-deoxy-6-C-sulfonato-α-D-glucopyranoside (501)178 (42.2 mg, 48% over 1 2 steps) (44:56) as a colourless oil. H NMR (500 MHz, D2O) 3.07-3.12 (2H, m, H6aS,

H6aR), 3.29 (2H, t, J= 9.5 Hz, H4S, H4R), 3.39-3.43 (2H, m, H6bS, H6bR), 3.47 (1H, J =

7.2, 10.3 Hz, H1a’R) 3.57-3.78 (9H, m, H1a’S, H2S, H2R, H3S, H3R, H3a’S, H3a’R, H3b’S,

H3b’R), 3.92-4.10 (6H, m, H1b’ S, H1b’R, H2’ S, H2’R, H5S, H5R), 4.90-4.92 (2H, m, H1S,

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13 H1R); C NMR (125 MHz, D2O) 51.9, 62.5, 62.7, 68.0, 68.5, 68.8, 70.6, 70.7, 71.3, 72.4, - - 72.9, 97.8, 98.2; HRMS (ESI) 317.0547 m/z [C9H17O10S (M) requires 317.0548].

YihQ hydrolysis of SQGro by 1H NMR analysis YihQ catalysed hydrolysis of SQGro was monitored by 1H NMR spectroscopy using a 500 MHz instrument. A solution of YihQ (20 µL, 0.084 mM, 50 mM sodium phosphate and 150 mM NaCl, pH 7.5) was added to a solution of SQGro (4 mg, 12.6 mmol) in D2O (0.6 ml) at 25 °C. 1H NMR spectra were acquired at time points (t = 0, 14 and 60 min, 20 h).

Bacterial culture of E. coli E. coli K-12 substrain BW25113 was grown in a sterile phosphate-buffered mineral salts medium (pH 7.2)186 with glucose, glycerol, SQGro or SQ as the sole carbon source. Cultures were grown at 30 ˚C with glucose, glycerol displaying growth within 1-2 days, SQGro within 3-4 days and SQ within 1-2 weeks. Subcultures were inoculated (1%) with pre-cultures grown with the same substrate and grown aerobically at 30˚C. Culture volumes were 2 mL and 4mM substrate concentrations were used. Growth was measured using a Varian Cary50 UV/visible spectrophotometer to measure the OD580. All growth experiments were replicated (n=2).

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CHAPTER SIX:

Discovery of sulfoglycolysis in Agrobacterium

tumefaciens

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6.1 Biomineralization of SQ

The two well-described sulfoglycolysis pathways convert sulfoquinovose to the C3- sulfonates DHPS and SL, which are excreted from the organism.171172 Metabolism of

DHPS and SL to inorganic sulfur to complete mineralization of SQDG most commonly occurs in separate organisms to that which perform sulfoglycolysis. A range of pathways have been reported that enable biomineralization of DHPS and SL. 173,187,188

In 2000 Roy and co-workers reported the isolation of a soil bacterium identified as Agrobacterium sp. Strain ABR2 that could grow on SQ and achieve its mineralization to inorganic sulfate (Figure 6.1). Agrobacterium sp. strain ABR2 grew smoothly on SQ with formation of sulfate, yet neither DHPS or SL could be detected at any point in its growth.187 Interestingly there was a delay in the consumption of SQ and the formation of sulfate, consistent with the formation of an intermediate. The authors report a growth curve over 24 h; over this period consumption of SQ was incomplete, suggesting that the culture had yet to completely reach stationary phase. 13C NMR spectroscopy of the culture medium grown on 13C labelled SQ led to detection of bicarbonate as the sole metabolic end-product. Cell extracts from SQ conditioned cells of Agrobacterium sp. displayed SQ kinase activity, yet did not display SQ dehydrogenase activity. These authors suggested that these results were consistent with the accumulation of an organosulfonate intermediate and it was proposed that Agrobacterium sp. Strain ABR2 utilized a sulfo-

EMP pathway; nonetheless these authors highlighted the inability to detect any sulfonated intermediate including DHPS.

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Figure 6.1: Growth curves of Agrobacterium sp. strain ABR2 as reported by Roy and 187 co-workers. Symbols: full circles, optical density at OD600; empty circles, SQ (mM); triangles, sulfate.

6.2 Aims

We planned to perform growth studies on a genetically defined strain of Agrobacterium tumefaciens to establish if it can grow on SQ. We aimed to gather evidence for a putative sulfoglycolysis pathway in this organism using proteomics. We then aimed to study the mode of sulfoglycolysis to identify possible intermediates between SQ and the metabolic end-product sulfate. In parallel, we sought to determine if SQGro is a substrate for an

SQase cloned and recombinantly expressed from A. tumefaciens (AtSQase) and which lies in a putative sulfoglycolytic operon.

6.3 Results and Discussion

A. tumefaciens growth studies

In the growth studies of Agrobacterium sp. strain ABR2 by Roy and co-workers, SQ consumption was incomplete and the amount of sulfate released did not correlate to amount of SQ consumed.187 No sulfur containing metabolic intermediates were identified. These results give rise to many questions about what is biochemically occurring in sulfoglycolysis performed by this organism.

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We chose to repeat these experiments using a genetically well-defined strain of A. tumefaciens. A sample of A. tumefaciens C58 was obtained as a gift from Dr Monica

Doblin (School of Botany, University of Melbourne). We assessed growth of A. tumefaciens C58 in M9 minimal media on 5 different carbon sources: Glc, Gro, SQGro,

SQMe, and SQ. Growth was observed on all 5 sources, confirming that this organism can conduct sulfoglycolysis.

In order to probe the details of substrate consumption and to explore the formation of metabolic intermediates and end-products, we chose to work with SQ as it is commercially available, using Glc as a positive control (Figure 6.2). Cultures of A. tumefaciens were grown in M9 minimal media containing SQ (10 mM) or Glc (10 mM), and samples were taken at various time-points, and were analysed for residual substrate using a reducing sugar assay (the PAHBAH assay), and for the formation of sulfate (as barium sulfate) by turbidimetry.

The PAHBAH assay was performed as reported by Blakeney and Mutton.189 A freshly prepared PAHBAH working solution consisting of p-hydroxybenzoic acid hydrazide dissolved in an alkaline diluent (0.5 M trisodium citrate, 0.01 M calcium chloride, 0.5 M sodium hydroxide) was added to a sample of culture supernatant. This mixture was incubated at 98 °C for 4 mins, diluted with water and the absorbance measured using a UV/Vis spectrophotometer at 415 nm. The concentration was determined by comparison with a standard curve constructed using SQ of Glu.

For the sulfate assay, we used a turbidimetric method as reported by Sörbo which

precipitates sulfate as a BaSO4 precipitate, allowing for its detection by monitoring light

190 scattering. A freshly prepared Ba-PEG reagent was made by dissolving BaCl2 (barium source) and propylene glycol 6000 (a stabilizing agent for the BaSO4 precipitate) in

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distilled water followed by dropwise addition of Na2SO4. Addition of Na2SO4 to the Ba-

PEG reagent forms BaSO4 seed crystals that promote uniform precipitate formation in the assay, which is crucial for obtaining consistent results.191 HCl was added to each sample to reduce interference from anions such as phosphate,192 then treated with freshly prepared Ba-PEG reagent. The samples were vortex mixed and their absorbance measured at 600 nm. However, we found the sensitivity to be low at this wavelength. As absorbance of the sample is wavelength dependent (because it relies upon light scattering) and decreases with increasing wavelength.192 We investigated the sensitivity at 400, 500 and 600 nm. Measurements made at 400 nm gave the best results and hence we used this wavelength for all studies.

In cultures grown on Glc, cell density increased over a period of approx. 100 h to a maximal final OD600 of 1.2, and growth correlated closely with Glc consumption. No change in sulfate concentration was detected, suggesting that under these conditions consumption of sulfate was low and undetectable. Cultures grown on SQ also grew over a period of approx. 100 h, but achieved a much lower final OD600 of approx. 0.4.

However, consumption of SQ was not correlated with growth; rather, while around 60% of the SQ was consumed during the exponential growth phase, the remainder required an additional 150 h to be completely consumed, during which time there was little change in OD600. During exponential growth, sulfate was released, but the amount formed lagged behind consumption of SQ, with only around 30% sulfate released at the end of exponential growth. However, over the next 150 h of incubation, levels of sulfate continued to rise and eventually ended up approximately equivalent in concentration to the starting concentration of SQ. The apparent excess formed may arise because of evaporation of water during the 11 day experiment. Broadly the results of our experiment

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in its early phases is similar to the observation of Roy and co-workers,187 however, they only grew their culture for 24 h and did not observe complete consumption of SQ.

Figure 6.2: Growth of A. tumefaciens in M9 minimal media at 30 °C containing: a. 10 mM Glc as the sole carbon source; b 10 mM SQ as sole carbon source.

To explore whether SQ could be used as a source of sulfur, we grew cultures of

A. tumefaciens on Glc (10 mM) and separately on SQ (10 mM) in sulfate-free M9 minimal media (Figure 6.3). We did not expect to see significant growth in the sulfate-free Glc culture, however, we expected the SQ culture to be unaffected as SQ consumption releases sulfate which can be incorporated into the biosynthesis of amino acids and sulfur- containing cofactors such as biotin and lipoic acid.193,194 Additionally, we grew cultures in sulfate free media with no carbon source as a negative control. In the sulfate-free Glc culture, the OD600 reached 0.4, significantly lower than the 1.2 it reached in the sulfate containing media. When the culture reached its maximum OD600, around 80% of the Glc remained unconsumed. Interestingly, despite no increase in OD600, Glc consumption

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continued slowly, such that there was just 20% of Glc present after 340 h. We attribute the limited growth to carry-over of sulphur species from the inoculating sample which was grown in sulfate-containing media (1 µM), and carryover of sulfur containing amino acids within the bacteria that were likely broken down to recycle sulfate. We attribute the steady consumption of Glc in stationary phase to metabolic processes that consume energy but without cellular division and consumption of sulfur. Cultures grown on SQ- containing sulfate-free M9 media grew to an OD600 of 0.55, similar to the level of OD600

= 0.4 recorded in sulfate-containing M9 media. SQ consumption was coincidental and inversely correlated with growth, however, sulfate release was slow and not proportional to SQ consumption. No growth was detected in sulfate free media with no carbon source.

Figure 6.3: Growth of A. tumefaciens in sulfate-free M9 minimal media at 30 °C containing: a. 10 mM Glc as the sole carbon source; b 10 mM SQ as sole carbon source.

On the basis of these observations, we speculated that sulfate was not formed directly from SQ, and that there was an intermediate that is subsequently converted to sulfate. Our hypothesis was that this intermediate was sulfite. We established a sulfite

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quantification assay as reported by Brychkova and co-workers.195,196 The colour reagent was freshly prepared from basic fuchsin, sulfuric acid and formaldehyde. The colour reagent was added to a mixture of sample and 0.5 mM Na2SO3, incubated for 10 minutes at r.t. and measured spectroscopically at 570 nm. Analysis of the samples obtained from the growth experiment using the basic fuchsin assay revealed that sulfite was produced upon metabolism of SQ and was released into the media (Figure 6.4). Production of sulfite was released in a manner coincidental and inversely correlated with growth. Subsequent to cells entering stationary phase at around 80 h, the concentration of sulfite slowly declines while the concentration of sulfate increases. This data is consistent with a model in which SQ metabolism releases sulfite as the first formed mineral form of sulfur, and which is exported from the cell. Subsequently, sulfite autooxidizes to sulfate, the final mineral form of sulfur.

Figure 6.4: Growth of A. tumefaciens in sulfate free M9 minimal media containing 10 mM SQ as sole carbon source at 30 °C

Proteomic studies

In order to identify the proteins whose expression increases upon growth of A. tumefaciens on SQ, we conducted proteomics. Five samples of A. tumefaciens were grown in M9 minimal media on 4 mM Glc and separately another five samples were grown on 4 mM SQ. The cells were collected in mid-log phase by centrifugation and

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frozen, then were supplied to our collaborator Dr Nicholas Scott. The samples were subjected to a proteomics workflow that allowed determination of the identity of the studied proteins by multistage mass spectrometry and querying a database of predicted proteins derived from the genome sequence of the organism. In addition, the relative abundance of peptides was used to determine the relative abundance of the proteins present in the samples. Statistical analysis of the results allowed presentation of the results as a ‘volcano’ plot, in which the logarithm of fold-change in abundance (y-axis) is plotted versus significance on the (x-axis) (Figure 6.5). Shown in red are proteins that exhibited a change with high significance as measured by a -Log(p-value) > 3. In this data is it striking that a set of proteins that lie largely contiguous in the genome are highly upregulated (5 to 10-fold) during growth on SQ versus growth on Glc, namely Atu3277,

3278, 3279, 3280, 3281, 3282 and 3285.

Figure 6.5: Volcano plot showing relative abundance of proteins in A. tumefaciens grown on 4 mM Glu (left) or 4 mM SQ (right). Proteins with high significance are highlighted in red.

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These proteins were identified to belong to an operon that additionally contains

Atu3283 and 3284. The existing bioinformatics annotation of this operon is shown in

Figure 6.6, and includes putative flavin reductase, alkanesulfonate monooxygenase, ABC transporters, and sugar transporters and binding proteins with similarity to the maltose- binding protein and importation apparatus. Additionally, a putative glycosidase

(Atu3285) is present that is a member of family GH31. Based on sequence-alignment with the known sulfoquinovosidase YihQ from E. coli Atu3285 was tentatively assigned as a sulfoquinovosidase. The bioinformatic annotation of Atu3283 and 3284 is as membrane associated proteins; possibly, their association with membranes led to them being undetected in the proteomics experiment.

Figure 6.6: Genes within the operon identified by proteomics of SQ-inducible proteins.

Several genes in this operon display similarity to the ssuEADCB gene cluster in

E. coli (Figure 6.7). In situations of sulfate starvation, E. coli expresses several proteins from this operon which provide the capacity to grow on assorted alkylsulfonates including: ethanesulfonate, propanesulfonate, butanesulfonate, pentanesulfonate, hexanesulfonate, ethanedisulfonate, octanesulfonate, decanesulfonate, isethionate, sulfoacetate, MOPS, HEPES, MES, and PIPES.197

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Figure 6.7: Genes within the E. coli ssuEADCB operon.197

SsuABC proteins display amino acid sequence similarity to the ATP-binding cassette (ABC)-type transporter systems and function as an uptake system for alkanesulfonates. SsuD and SsuE are specifically involved in desulfonation of alkanesulfonates, working together in a two-component system as FMNH2-dependent monooxygenases. SsuD is catalyzes the conversion of alkanesulfonates to aldehyde and sulfite. This catalysis is dependent on availability of reduced flavin mononucleotide

(FMNH2) and oxygen. SsuE is a NAD(P)H:flavin oxidoreductase that reduces flavin

198 mononucleotide (FMN) to FMNH2 for SsuD to utilize (Scheme 6.1).

Scheme 6.1: Overview of desulfonation system of alkanesulfonates in E. coli.198

Other genes in the A. tumefaciens operon display similarity to proteins within the maltose-importation system of E. coli. Maltose transport across the plasma membrane in

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E. coli is mediated by ABC transporters that harness the energy of ATP to transport substrates.199 This requires the action of four proteins; maltose-binding protein (MBP) expressed by the malE gene and proteins expressed from the malF, malG and malK genes

(Scheme 6.2).200 MBP is a soluble protein located in the periplasm, MalF201 and MalG202 are hydrophobic proteins that span the membrane and the MalK203 protein is hydrophilic, located in the inner face of the membrane. The maltose transporter (MalFGK2) is located in the membrane and comprised of two transmembrane subunits formed by one MalF, one MalG and two subunits of MalK, which act as nucleotide-binding domains (NBDs)

204 that bind and hydrolyse ATP. The MalFGK2 transporter is inactive in the absence of

MBP and rests in an inward-facing conformation (Scheme 6.2a) where the two NBDs are separated and inactive; and the transporter is open to the intracellular side. In the periplasm, MBP binds ligand and undergoes a conformational change from an open (no substrate) to closed form (with substrate).205 The maltose bound MBP binds to the

MalFGK2 transporter, causing a conformational change within the transporter bringing the NBDs closer together, allowing ATP to interact with the protein. ATP binding opens the transport pathway to the periplasmic side, with simultaneous release of the ligand from MBP, transporting it through the channel into the cytoplasm (Scheme 6.2b). ATP is hydrolysed into ADP and inorganic sulfate and the transporter returns into its original inward-facing inactive conformation.206,207

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Scheme 6.2: Summary of maltose transporter (MalFGK2). a. MalFGK2 inactive complex with inward facing conformation; b. MPB- maltose complex binds to MalG and MalF causing a conformational shift in the MalFGK2 complex; c. NBDs bind ATP and conformational change opens transporter to periplasm, maltose enters channel; d. 3- 206,207 ATP is hydrolysed to ADP and PO4 .

Atu3285 is a sulfoquinovosidase that can process SQGro

The gene Atu3285 (accession code WP_035199431) was synthesized and the corresponding protein (hereafter AtSQase) was recombinantly expressed in E. coli by Mr

James Lingford in Dr Ethan Goddard-Borger’s laboratory at the WEHI. In work performed by a colleague, Palika Abayakoon, AtSQase was shown to exhibit high specificity for 4-nitrophenyl α-D-sulfoquinovoside (PNPSQ) relative to PNPGlc, defining it as a sulfoquinovosidase.208

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I sought to establish whether this AtSQase could utilize SQGro as a substrate

(Scheme 6.3). The hydrolysis of SQGro by AtSQase was monitored using 1H NMR spectroscopy (Figure 6.7). Prior to addition of enzyme (t = 0), there were two overlapping doublets at  4.9 ppm corresponding to the H1 of the 2’R and 2’S diastereoisomers. 18 min after addition of AtSQase a new peak appeared at  5.2 ppm consistent with the H1 of α-SQ. At 18 h both diastereoisomers of SQGro were fully consumed and SQ had undergone mutarotation to equilibrium, with H1 protons of α- and β-SQ evident.

Scheme 6.3: Hydrolysis of SQGro by AtSQase.

Figure 6.7: Hydrolysis of SQGro by AtSQase monitored by 1H NMR spectroscopy. t = 0 min prior to enzyme addition, H1 of 2’R and 2’S SQGro stereoisomers at 4.9 ppm; t = 18 min enzyme addition shows appearance of new peak at 5.2 ppm assigned as H1 of α- SQ; t = 1 h after enzyme shows appearance of H1 of β-SQ at 4.7 ppm; t = 18 h complete consumption of SQGro and mutarotation of SQ α and β anomers.

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A time-course for the disappearance of individual peaks corresponding to each diasteroisomer of SQGro over the first 25 minutes revealed that the 2’R stereoisomer, which corresponds to the natural stereochemistry of SQDG, hydrolysed 3-fold faster than the 2’S stereoisomer (Figure 6.8).

Figure 6.8: Rates of consumption of individual SQGro diastereoisomers by AtSQase.

6.4. Conclusion: The proposed sulfoglycolysis pathway of A. tumefaciens

Collectively, our data provide compelling evidence that A. tumefaciens can grow on SQ and utilizes a novel sulfoglycolysis pathway and allows us to propose a model for this pathway. The three key pieces of data are that: (1) the release of sulfite is coincident with growth, (2) a 9-gene operon is upregulated upon growth on SQ, and (3) the gene product of Atu3285 is a sulfoquinovosidase. The similarity of the 9-gene operon to the alkylsulfonate utilization ssuEADCB gene cluster and maltose importation system malEFGK of E. coli allow us to propose a putative sulfoglycolysis pathway in A. tumefaciens (Scheme 6.4). We propose that an SQ binding protein (Atu3282) binds SQ and its glycosides in the periplasm and delivers them to a predicted ABC transporter system (Atu3283, 3284), importing them into the cytosol. SQ glycosides are then hydrolysed to SQ by the AtSQase (Atu3285). SQ is likely converted to an aldehyde by

Atu3277 and 3279, acting like the SsuD/SusE alkenesulfate monooxygenase/flavin

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reductase two-component system. This releases sulfite and a possible exporter transports the sulfite into the periplasm where it autoxidizes to sulfate. Finally, Atu3278 is an

NAD(P)H-dependent oxidoreductase that acts on the glucose-6-aldehyde to either oxidize it to GlcA or reduce it to Glc, where it can enter central carbon metabolism.

Scheme 6.4: Proposed sulfoglycolysis pathway for SQGro metabolism in A. tumefaciens.

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6.5 Experimental

Growth studies Cultures of A. tumefaciens were grown in a phosphate-buffered M9 mineral salts medium (pH 7.2), with or without sulfate, with glucose or SQ as the sole carbon source. Cultures were grown at 30 ˚C and displayed growth within 2-3 days. Subcultures were inoculated with 1% volumes of pre-cultures growth with the same substrate and grown aerobically at 30˚C. Culture volumes were 10 mL and 10 mM substrate concentrations were used. Growth was assessed using a Varian Cary50 UV/visible spectrophotometer to measure the OD600. All growth experiments were replicated (n=2).

Reducing sugar assay. Alkaline diluent: Sodium hydroxide (20 g, 0.5 mol) was added to a solution of 0.10 M trisodium citrate (0.05 mol, 0.5 L) and 0.02 M calcium chloride (0.013 mol, 0.5 L). PAHBAH working solution: 4-Hydroxybenzhydrazide (PAHBAH) (0.25 g, 1.64 mmol) was dissolved in alkaline diluent (50 mL). The PAHBAH working solution should be freshly made when needed.

Procedure: 0.90 mL of PAHBAH working solution was added to 0.10 mL of culture supernatant. The mixture was heated at 98 °C for 4 min. 0.5 mL of the mixture was diluted into 1 mL of deionized water and the absorbance read at 415 nm. The amount of sugar present in the sample was then obtained from a standard curve prepared with known amounts of Glucose or SQ (0.0-0.25 mM).189

Standard curve:

Sulfate Assay

Ba-PEG reagent: BaCl2 (41.7 mg, 0.20 mmol) and polyethylene glycol 6000 (750 mg) were dissolved in deionized water (5 mL). Na2SO4 (10 µL, 50 mM) was added to the solution with efficient magnetic stirring.

2- Procedure: A sample of culture supernatant (containing up to 2.5 µmol of SO4 ), was diluted to 0.1 mL with deionized water. 0.5 M HCl (0.1 mL) was added followed by Ba- PEG reagent (0.1 mL). The mixture was vortex-mixed and the absorbance of the sample was measured at 400 nm. The amount of sulfate present in the sample was then obtained 190 from a standard curve prepared with known amounts of sulfate (0.0-0.5 mM Na2SO4).

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Sulfite Assay Reagent A: Basic fuchsin (4.0 mg, 0.012 mmol) was added to deionized water (8.25 mL) at 0°C. 98% H2SO4 (1.25 mL) was added.

Reagent B: Formaldehyde 36% in H2O (0.32 mL) was added to deionized water (9.68 mL) at 0°C.

Reagent C: Reagent A (1 mL) was added to deionized water (7 mL). Reagent B (1 mL) was then added.

Procedure: Reagent C (516 µL) was added to a mixture of culture supernatant (72 µL) and 0.5 mM Na2SO3 (12 µL). The sample was incubated at r.t. for 10 min and the absorbance of the sample was measured at 570 nm. The amount of sulfite present in the sample was then obtained from a standard curve prepared with known amounts of NaSO3 (0.0-0.5 mM).195,196

AtSQase hydrolysis of SQGro by 1H NMR analysis AtSQase catalysed hydrolysis of SQGro was monitored by 1H NMR spectroscopy using a 500 MHz instrument. A solution of AtSQase (10 µL, 0.062 mM, 50 mM sodium phosphate and 150 mM NaCl, pH 7.5) was added to a solution of SQGro (4 mg, 12.6 1 mmol) in D2O (0.6 ml) at 25 °C. H NMR spectra were acquired at time points (t = 0, 18 min and 1, 20 h).

Growth conditions for proteomics sample Cultures of A. tumefaciens were grown in a phosphate-buffered M9 mineral salts medium (pH 7.2), with glucose or SQ as the sole carbon source. Cultures were grown at 30 ˚C and displayed growth within 2-3 days. Subcultures were inoculated with 1% volumes of pre- cultures growth with the same substrate and grown aerobically at 30˚C. Culture volumes were 10 mL and 4 mM substrate concentrations were used. When cultures reached exponential growth (OD600 = 0.2) cells were spun down and supernatant was removed. Cells were resuspended and washed with KCM buffer (3 × 1.0 mL) and submitted subject to proteomics. All growth experiments were replicated (n=5).

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164

CHAPTER SEVEN:

Summary and future work

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This thesis focused on the mechanistic and structural analysis of members of three glycoside hydrolase families: GH99, GH134 and GH31. Additionally, this thesis examined aspects of sulfoglycolysis in E. coli and A. tumefaciens, and has led to the proposal of a novel sulfoglycolytic pathway. This overview chapter presents a summary of the major findings and future prospects for this work.

Chapters 2 and 3: GH99 endo-α-1,2-mannanase and endo-α-1,2-mannosidases

Summary of findings

• The mechanism inspired inhibitor α-D-mannopyranosyl-1,3-noeuromycin

(ManNOE, 201) was successfully synthesized to explore its interaction with

bacterial GH99 endo-α-1,2-mannanases.

• ITC (performed by Lukasz Sobala from Prof Gideon Davies’ lab) with two

bacterial GH99 endo-α-1,2-mannanases revealed 201 to be the most tightly

binding ligand identified to date.

• X-ray crystallography (performed by Lukasz Sobala) and quantum mechanical

calculations (performed by Lluís Raich from Prof Carme Rovira’s lab) of the free

energy landscapes revealed that 201 mimics the conformation of the enzyme-

bound substrate. The tighter binding relative to ManNOE was attributed to a

hydrogen bonding interaction between the 2-OH of the noeuromycin ring and

active site residue E333.

• Comparison of 201 with the less tightly binding neutral inhibitors α-D-

mannopyranosyl-1,3-glucal (ManGlucal) and α-D-mannopyranosyl-1,3-(1,2-

dideoxy)mannose (ManddMan) (synthesized by Williams group member Dr Pearl

Fernandes) demonstrated the importance of charge over shape mimicry for

effective inhibitor design for GH99 enzymes.

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• The mechanism inspired inhibitor α-D-mannopyranosyl-1,3-(2-

amino)dideoxynoijirimycin (Man2NH2DMJ, 301) was synthesized to assess

whether interactions with the DMJ ring and the active site residue E333 could be

enhanced by incorporation of a basic group.

• ITC and X-ray crystallography (performed by Lukasz Sobala) with two bacterial

GH99 endo-α-1,2-mannanases revealed weak binding of 301, four times lower

than the α-D-glucosylpyranosyl-1,3-dideoxynonijimycin (GlcDMJ) counterpart.

• It was proposed that the low affinity occurred because protonation of the 2-NH2

group reduced the basicity of N5, preventing it from protonating within the active

site.

Figure 7.1: Ligands of endo-α-1,2-mannanase: ManNOE (201) and Man2NH2DMJ (301). Current and future work

Although family GH99 has been extensively studied, it has not been definitively shown that this family of enzymes do indeed act though the proposed neighbouring group participation mechanism. Such studies are in train, through kinetic isotope studies initiated by another member of the Williams group, Dr Gaetano Speciale, who synthesized stable isotope labelled substrate analogues and in collaboration with Prof

Andrew Bennet (Simon Fraser University) measured KIEs at various sites. At the time of writing the results of these studies are not known.

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Most efforts to develop inhibitors has been targeted towards bacterial endo-α-1,2- mannanases, as our efforts have relied heavily upon ligand/enzyme co-complexes, crystallization of mammalian endo-α-1,2-mannosidase had not been achieved, owing to difficulties in expression of the mammalian protein. Exploiting the recently reported method for recombinant expression of human endo-α-1,2-mannosidase developed by

Kajihara and co-workers,209 the Davies group has recently crystallized and solved the structure of the human enzyme. In future work, using the knowledge developed leading to the identification of ManNOE as the most potent inhibitor known for bacterial endo-α-

1,2-mannanase, the stereochemically matched compound GlcNOE could be synthesized and investigated as an inhibitor of the human enzyme (Figure 7.2).

Figure 7.2: GlcNOE, a putative inhibitor of human endo-α-1,2-mannosidase, and GlcMan3OMe, a putative substrate. Glucose units depicted in blue

As part of our efforts to characterize human endo-α-1,2-mannosidase, Dr Pearl

Fernandes synthesised methyl α-D-glucopyranosyl-(1,3)-α-D-mannopyranosyl-(1,2)-α-D- mannopyranosyl-(1,2)-α-D-mannopyranoside (GlcMan3OMe), a substrate for the mammalian enzyme. Efforts are afoot to soak this molecule into an inactive form of the enzyme to obtain a Michaelis complex.

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The low affinity of α-D-mannopyranosyl-1,3-(2-amino)dideoxynojirimycin

(Man2NH2DMJ, 301) for endo-α-1,2-mannanases was attributed to protonation of the N2 reducing the basicity of the endocyclic N and preventing it from undergoing protonation.

To circumvent the perturbing effects of the two nitrogen atoms, one might be tempted to consider replacing the endocyclic N in DMJ with O to give Man2NH2Man (Figure 7.3).

This compound would still be protonated at N2 allowing for an ionic interaction with

E333, but like Man2NH2DMJ, would not develop positive charge at the endocyclic position, so may not be improved relative to Man2NH2DMJ. An alternative suggestion would be synthesis of a mannose-derived episulfide (Man1,2SMan), which would mimics the 1,2-anhydrosugar intermediate, yet would be more stable, and could provide a means to obtain a complex with the protein to characterize interactions and conformation of the intermediate.

Figure 7.3: Man2NH2Man and Man1,2SMan as putative inhibitors of endo-α-1,2- mannanases.

Chapter 4: GH134 endo-β-1,4-mannanase

Summary of findings • SsGH134, a representative of a newly discovered family of β-1,4-mannanases,

was structurally and mechanistically characterized.

• NMR spectroscopic and enzyme digestion studies revealed substrate preference,

cleavage site, and stereochemistry of cleavage catalyzed by SsGH134.

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• QM/MM studies (performed by Lluís Raich in Prof Carme Rovira’s lab) revealed

1 3 ‡ 3 SsGH134 possesses a different mechanism and utilizes a unique C4→ H4 → H1

conformational itinerary along the reaction co-ordinate, distinct from other known

β-1,4-mannanases.

Current and future work

Currently, no research aimed at GH134 enzymes is ongoing in our laboratory. A possible direction for furthering research into GH134 enzymes, certainly in theme with this thesis, could be development of inhibitors. A kifunensine (KFN) based inhibitor may be useful in the study of GH134 enzymes. KFN is known to be a potent mannosidase inhibitor, as exemplified by its ability to inhibit mannosidase I of the N-glycan trimming pathway.

KFN is well-known as an inhibitor of family GH47 α-mannosidases, which utilise a

3 3 ‡ 1 H1→ H4 → C4 conformational itinerary, which is reversed compared to that we propose for GH134 enzymes.153 Consequently, one possible avenue to developing inhibitors of

SsGH134 would be to elongate this inhibitor warhead with a -1,4-mannobiosyl unit to generate a pseudo-trisaccharide that is able to occupy the –3 to –1 subsites (Figure 7.4).

Figure 7.4: Man2-KFN, a putative inhibitor of GH134 β-1,4-mannanases.

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Chapters 5 and 6 – new insights into sulfoquinovosidases and sulfoglycolysis

Summary of findings

• The substrate sulfoquinovosyl glycerol (SQGro, 501) was synthesized as a

mixture of 2’R and 2’S stereoisomers.

• SQGro (501) was determined to be a substrate for the family GH31 E. coli

sulfoquinovosidase YihQ. This enzyme exhibited a preference for the naturally

occurring 2’R isomer.

• Based on the approximate doubling of final culture density relative to SQ, SQGro

(501) was proposed to act as a 6-carbon nutrient source for E. coli compared to

the 3-carbon nutrient, SQ.

• The soil bacterium A. tumefaciens strain C58 was shown to grow on SQ as sole

carbon source.

• Time-course analysis of the growth media revealed consumption of SQ was

proportional to release of sulfite, which, over time, autooxidised to sulfate.

• Proteomics revealed a 9-gene operon was upregulated upon growth on SQ.

• This operon bore some similarity to the alkylsulfonate utilization ssuEDCB gene

cluster in E. coli, as well as the maltose/maltodextrin system malEFGK.

• Collectively these results led to the proposal of a novel sulfoglycolytic pathway

in A. tumefaciens, involving binding and import of SQ/SQGro (501), flavin-

dependent monooxygenation to release sulfite and 6-oxo-glucose, reduction of 6-

oxo-glucose to glucose and entry into central carbon metabolism, and export of

sulfite followed by autooxidation to sulfate.

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Current and future work

In parallel to this work, fellow group member Palika Abayakoon set out to biochemically characterize the putative SQase in A. tumefaciens. This work, recently published with the

E. coli studies outlined above, was conducted in collaboration with Dr Ethan Goddard-

Borger and Prof Gideon Davies, and identified a key active site amino acid difference between E. coli YihQ and AtSQase.208 Structural and mutagenic studies revealed an extended QQRWY motif in E. coli and a ‘charge neutral’ KERWY motif in A. tumefaciens. This work enhanced our knowledge of the sequence determinants of SQase enzymes. Very recently, in collaboration with Dr Ethan Goddard-Borger we have shown that the putative SQ/SQGro binding protein (Atu 3282) binds 2’R-SQGro with sub- micromolar affinity. As part of this work a new synthesis leading to pure 2’R-SQGro has been developed by Dr Ruwan Epa.

In other work aimed at experimentally verifying the proposed pathway for A.

13 tumefaciens, Ms Janice Mui (Hons 2018) has synthesised C6-labelled SQ in an effort to characterise metabolites of the sulfoglycolysis. Preliminary studies using NMR reveal

13 disappearance of C6-labelled SQ from the growth medium, and the transient appearance

13 – 187 of H CO3 , a result consistent with the early observations of Roy. No other carbon metabolites were observed in the culture media. Current work is focussed on the cloning and expression of the putative carbohydrate binding proteins and enzymes within the operon, and their biochemical characterization to support the proposed sulfoglycolysis pathway.

Our group continues its efforts to characterize other proteins within the sulfo-EMP operon. SQ-SF isomerase YihS, SF kinase YihV and SLA reductase YihU. Meanwhile, structural studies in Prof Gideon Davies’ laboratory has resulted in 3-D structures for all

172

four core sulfoglycolysis enzymes (YihS, YihV, YihT and YihU), determined by X-ray crystallography. Kinetic characterization is being conducted in parallel, and ligands for complex formation have been synthesized. Collectively this work promises to provide deep molecular insight into the sulfo-EMP pathway.

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Author/s: Petricevic, Marija

Title: Design and synthesis of chemical tools for studies of carbohydrate active enzymes

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