Substrates and Inhibitors of Enzymes Involved in Exopolysaccharide Dependent Biofilms

Anthony Chibba

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Chemistry University of Toronto

Anthony Chibba

Doctor of Philosophy

Department of Chemistry University of Toronto

2014 Abstract

Many biofilm-forming bacteria produce a similar partially de-N-acetylated β-1,6-N-acetyl glucosamine homopolymer (dPNAG) to facilitate bacterial adhesion. In many medically important biofilm forming bacterial strains, including Escherichia coli, Staphylococcus epidermidis and Staphylococcus aureus, de-N-acetylation of the β-1,6-N-acetyl glucosamine homopolymer (PNAG) is catalyzed by a metal dependent de-N-acetylase. Sixty-five percent of all human persistent bacterial infections are considered to be biofilm related. In vivo studies have implicated the production and subsequent de-N-acetylation of PNAG is essential to bacterial virulence.

In this work, methods of monitoring and targeting PNAG-dependent biofilm processes are explored. A novel chromogenic glycosidase substrate based on a glycosyl carbamate was developed and used to effectively monitor the activity of Dispersin B (DspB), an enzyme capable of degrading PNAG/dPNAG. Additionally, an array of potential deacetylase inhibitors were synthesized with the goal of targeting the essential de-N-acetylase enzymes PgaB and IcaB, from

E. coli and S. epidermidis respectively, for biofilm formation. These inhibitors were based on a carbohydrate scaffold containing either a metal chelating moiety or a transition state mimic.

Finally, a novel coumarin based substrate was developed to monitor in vitro de-N-acetylase activity of PgaB and IcaB. Using this substrate, the potency of the synthesized inhibitors was evaluated through a competitive fluorogenic assay. The most effective inhibitor, a chemoenzymatically-synthesized pentasaccharide derivative showed a Ki value of 280 µM.

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Acknowledgments

Ahhhh – Acknowledgments! The only place where you won‘t find overwhelming amounts of science written in this wonderful document!

First and foremost, I would like to thank my supervisor, Prof. Mark Nitz. You encouraged me to take the plunge into this academic adventure and your years of guidance, mentorship, and especially utmost patience will never be forgotten. Furthermore, I would like to thank my Ph.D. committee members over the years: Prof. Andrew Woolley, Prof. Patrick Gunning, Prof. Ronald Kluger, Prof. Andrei Yudin, Prof. Zhao-an Zhang and Prof. Peter Adreanna.

Secondly, I would like to acknowledge the 1:20pm coffee group – Rodolfo, Colin, Andrew, Pengpeng for years of social therapy and humor and long publication records and the suggestion that worker‘s unions can be abusive to children. We also sometimes spoke about science, and rarely about our everyday lives, but it was always a part of the day to look forward to.

Thirdly, I would like to thank all the Nitz members, past and present, who shared both scientific and social and friendly endeavors with me! To list a few, in order of who I met first to last – Anna, Joanna, Yedi, Big Anthony, Dick Jacket, Rodolfo/Wolf, Carmen, Urja, Heather, Grace, Mike, Jason, Caroline, Kill Bill, Pengpeng, Landon, Melissa, Nesrin – Thank you for all the good times and wonderful memories!

Fourthly, I‘d like to acknowledge all the friends of mine outside of the lab, who have stuck around with me throughout the years, engaged in the utmost debauchery, played some awesome softball, and threw some great cottage weekends. In no particular order – Fu_Cam, Beubs, Dobbs, Grammo, Calvin, Xtina, Nat, Snosk, Alexandra, Ash, The Roxton Boys, José, Talon, Plona, Zsolt.

Finally, I‘d like to acknowledge my wonderful girlfriend Lisa Komar. I realize we have only began dating during the last year of my degree, but we‘ve always been great friends, and have shared many graduate school ordeals and adventures with each other over the years on our semi- annual sushi and coffee dates. I also appreciate you taking the time and effort to edit my scientific gibberish. I hope we can continue to have incredible adventures, perhaps even outside the graduate world!

Post-finally, MALDI-MS, you are the devil and the bain of my existence. I hope you burn down in an electrical fire.

Table of Contents

Acknowledgments ...... iv

Table of Contents ...... vi

List of Tables ...... ix

List of Schemes ...... x

List of Figures ...... xi

List of Abbreviations ...... xiv

1 General Introduction ...... 1

1.1 Biofilm Structure and Life Cycle ...... 1

1.1.1 Attachment ...... 1

1.1.2 Growth ...... 2

1.1.3 Release ...... 3

1.2 Biofilm Resilience and Antibiotic Resistance ...... 3

1.3 Extracellular Matrix ...... 5

1.3.1 Psl and Pel Matrices ...... 5

1.3.2 Alginate Matrix ...... 6

1.3.3 Cellulose Matrix ...... 7

1.3.4 Poly N-Acetyl Glucosamine Matrix ...... 7

1.4 Biofilm Treatment Strategies ...... 14

1.4.1 Prevention of Biofilm Formation ...... 15

1.4.2 Removal or Disruption of Existing Biofilms ...... 19

1.5 Purpose of the Current Study ...... 22

2 Substrate Development for Hexosaminidases ...... 24

2.1 Introduction ...... 24

2.2 Results and Discussion ...... 27

2.2.1 Synthesis of Acetal and Carbamate Substrates ...... 27

2.2.2 Substrate activity assay ...... 29

2.2.3 2nd Generation Carbamate Substrate ...... 34

2.3 Conclusions ...... 36

3 Synthesis and Development of Inhibitors towards PgaB and IcaB ...... 37

3.1 Introduction ...... 37

3.1.1 Inhibitor Design ...... 38

3.2 Results and Discussion ...... 48

3.2.1 Monosaccharide Inhibitors ...... 48

3.2.2 Chemoenzymatic Synthesis of a Polysaccharide Inhibitor ...... 58

3.3 Conclusions ...... 61

4 Assay Development and Inhibitor Evaluation with PgaB and IcaB ...... 63

4.1 Introduction ...... 63

4.2 Results and Discussion ...... 65

4.2.1 Enzyme Purification ...... 65

4.2.2 Assay Development ...... 67

4.2.3 Evaluation of Inhibitors towards IcaB and PgaB ...... 74

4.3 Conclusions ...... 87

5 Future Perspectives ...... 88

5.1 Substrate development for hexosaminidases ...... 88

5.2 Inhibitor development for PgaB and IcaB ...... 91

6 Experimental ...... 94

6.1 General Methods ...... 94

6.2 Procedures ...... 95

References ...... 116

Appendix A ...... 132 vii

Appendix B ...... 158

viii

List of Tables

Table 2.1 Kinetic characterization of substrate 2 and comparison with pNP-GlcNAc...... 32

Table 3.1 Relative inhibition activities of various SAHA analogs...... 40

Table 3.2 Inductive sigma values for various organic substituents...... 46

Table 4.1 Kinetic parameters for a selection of CE4 enzymes...... 65

Table 4.2 Absorbance / excitation and fluorescence emission wavelengths for the chromogenic substrates tested with PgaB...... 71

Table 4.3 Kinetic parameters for PgaB and IcaB with ACC...... 74

Table 4.4 Inhibition of PgaB and IcaB by N-acyl inhibitors...... 77

Table 4.5 Inhibition of PgaB and IcaB by N-acyl inhibitors...... 82

Table 4.6 Inhibition of PgaB and IcaB by N-alkyl inhibitors...... 83

Table 4.7 Inhibition of PgaB and IcaB by C-2 inhibitors...... 84

Table 4.8 Inhibition of PgaB and IcaB by polysaccharide inhibitor 64...... 86

List of Schemes

Scheme 2.1 Synthesis of acetal substrate 1...... 28

Scheme 2.2 Synthesis of carbamate substrate 2 ...... 29

Scheme 2.3 Synthesis of carbamate substrate 12...... 35

Scheme 3.1 Synthesis of divergent starting materials 15...... 49

Scheme 3.2 Synthesis of acyl inhibitors 19, 21, 23, 24, 25...... 50

Scheme 3.3 Synthesis of methylated starting material 29...... 51

Scheme 3.4 Synthesis of methylated acyl inhibitors 35 - 39...... 52

Scheme 3.5 Syntheis of inhibitors 42 and 45...... 53

Scheme 3.6 Syntheisis of alkyl inhibitor 47...... 54

Scheme 3.7 Synthesis of phosphonamidate inhibitor 51...... 55

Scheme 3.8 Synthesis of 2-C carbohydrate starting material 53a ...... 56

Scheme 3.9 Synthesis of 2-C inhibitors 54 and 56...... 56

Scheme 3.10 Synthesis of 2-C inhibitors 59 and 61 ...... 57

Scheme 3.11 Chemoenzymatic synthesis of pentasaccharide inhibitor 64 ...... 59

Scheme 4.1 Synthesis of chromogenic substrates ...... 69

List of Figures

Figure 1.1 Model of the biofilm life cycle ...... 1

Figure 1.2 3-D reconstruction of a bacterial biofilm by Vibrio cholerae ...... 3

Figure 1.3 Repeating unit of Psl polysaccharide produced by Pseudomonas aeruginosa ...... 6

Figure 1.4 Structure of the alginate polysaccharide...... 7

Figure1.5 Structure of cellulose polysaccharide...... 7

Figure 1.6 Structures of PNAG and dPNAG ...... 9

Figure 1.7 Model of dPNAG biosynthesis and export in Gram-negative E. coli...... 11

Figure 1.8 Model of dPNAG biosynthesis and export in Gram-positive S. epidermidis...... 12

Figure 1.9 Ribbon trace of the crystal structure of PgaB N-terminal deacetylase domain ...... 13

Figure 1.10 Outline of the stages in biofilm development and lists of strategies aimed at inhibiting/disrupting the biofilm at specific stages ...... 15

Figure 1.11 Mechanisms of anti-biofilm carbohydrates ...... 19

Figure 1.12 QS inhibitors against P. aeruginosa biofilms...... 21

Figure 1.13 Small molecule biofilm inhibitors ...... 22

Figure 2.1 Typical anchimeric participation mechanism for the hydrolysis of GlcNAc polymers by family 20 hexosaminidases...... 24

Figure 2.2 Common chromogenic aryl glycoside substrates used to measure hexosaminidase activity...... 25

Figure 2.3 Comparison of the β-1,4 and β-1,6 linkages in chitin and PNAG respectively...... 26

Figure 2.4 Fragmentation pathways for the acetal substrate 1 and carbamate substrate 2...... 27

Figure 2.5 1H NMR array of the acetal GlcNAc substrate and methyl-N-acetylglucosamine glycoside ...... 31

Figure 2.6 Double-reciprical plot used to obtain Michaelis-Menten kinetic parameters for the carbamate substrate 2...... 32

Figure 2.7 Graph of hydrolysis of substrate 2 and pNp-GlcNAc...... 34

Figure 3.1 Hydrolysis mechanism of CE4 family of enzymes...... 38

Figure 3.2 MMP-9 inhibitor that have undergone clinical trials for cancer treatment...... 39

Figure 3.3 X-ray crystal structure of SAHA bound to a zinc ion inside the HDAC active site.125 ...... 41

Figure 3.4 Biosynthesis of Lipid A, the role LpxC plays, and an LpxC inhibitor ...... 42

Figure 3.5 GPI de-N-acetylase catalyzed reaction and inhibitors synthesized and tested with GPI de-N-acetylase...... 42

Figure 3.6 Simplified reaction coordinate energy diagram of an enzyme-catalyzed reaction and an uncatalyzed reaction...... 43

Figure 3.7 Transition state proposed for HDAC1 and model for binding of a methyl phosphonamidate analog of SAHA...... 44

Figure 3.8 Reaction catalyzed by NagA and transition state analog designed to inhibit NagA ... 45

Figure 3.9 Equilibrium of ketones and geminal diols in aqueous solution...... 46

Figure 3.10 SAHA analog containing a trifluoromethyl moiety...... 47

Figure 3.11 Inhibitors of Angiotensin converting enzyme, a zinc protease...... 47

Figure 3.12 Proposed metal-chelating inhibitor structures for IcaB and PgaB...... 48

Figure 3.13 X-ray crystal structure of 19...... 54

Figure 3.14 MALDI mass spectra of compounds 63 and 64 ...... 61 xii

Figure 4.1 PgaB activity with varying substrate lengths and in the presence of various metals or small molecule metal chelaters...... 64

Figure 4.2 Reaction of fluorescamine with an amine to yield a highly fluorescent product ...... 68

Figure 4.3 Acetyl esters evaluated as substrates for PgaB under continuous assay conditions. ... 69

Figure 4.4 Rates of PgaB-catalyzed ACC deacetylation rates at varying ACC concentrations .. 73

Figure 4.5 Lineweaver-Burk inhibitor plots for compounds 23 and 35 with PgaB...... 78

Figure 4.6 Predicted Gauche projections for the acyl series of inhibitors ...... 80

Figure 4.7 Possible conformations of the amide bond in GlcNAc...... 81

19 Figure 4.8 F NMR of compound 61 in D2O...... 85

Figure 4.9 Hydration equilibrium of trifluoromethyl ketones in aqueous media...... 85

Figure 5.1 Potential novel acetal substrates...... 89

Figure 5.2 HexA enzymatic hydrolysis of GalNAc residue on GM2 ganglioside to produce GM3 ganglioside...... 90

Figure 5.3 Array of compounds to test using isothermal calorimetry ...... 92

Figure 5.4 Inhibitors developed against LpxC and pneumococcal peptidoglycan deacetylase PgdA...... 93

xiii

List of Abbreviations

9GlcNH2 β-1,6-Glucosamine Nonasaccharide

9GlcNH2-TT β-1,6-Glucosamine Nonasaccharide linked to Tetanus Toxoid

AcBr Acetyl Bromide

ACC 3-Carboxyumbelliferyl Acetate

AcOH Acetic Acid

ADEPT Antibody-Directed Enzyme Prodrug Therapy

AHL N-Acyl Homoserine Lactones

AMC 4-Methylumbelliferyl Acetate

5-AMQ 5-Acetoxymethylquinolinium

7-AMQ 7-Acetoxymethylquinolinium atm Atmosphere

BnBr Benzyl Bromide

BNCC Benzyl N-(Chlorosulfonyl)carbamate

CAZy Carbohydrate-Active enZYmes c-di-GMP Cyclic-di-guanosine Monophosphate

CE4 Carbohydrate Esterase Family 4 d Day(s)

DCC Dicyclohexylcarbodiimide

DCM Dichloromethane

xiv

DMAP 4-Dimethylamino Pyridine

DMF Dimethylformamide

DMSO Dimethylsulfoxide

DNA Deoxyribonucleic Acid dPNAG Partially de-N-acetylated β-1,6-N-acetyl Glucosamine Homopolymer

DTT threo-1,4-Dimercapto-2,3-butanediol

ELISA Enzyme-linked Immunosorbent Assay

ESI-MS Electrospray Ionization-Mass Spectrometry

ESI-MS/MS Tandem Electrospray Ionization-Mass Spectrometry

EtOAc Ethyl Acetate

EtOH Ethanol

GalNAc N-Acetylgalactosamine

GlcNAc N-Acetylglucosamine

GlcNH2 Glucosamine

GPI Glycosylphosphatidylinositol

GSL Glycosphingolipid h Hour(s)

1H/13C NMR Proton/Carbon Nuclear Magnetic Resonance

HBTU N,N,N′,N′-Tetramethyl-O-(1H-benzotriazol-1-yl)uronium Hexafluorophosphate

HDAC Histone Deacetylase

HEPES 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic Acid

HPLC High Pressure Liquid Chromatography

ICP-AES Inductively Coupled Plasma-Atomic Emission Spectroscopy

LPS Lipopolysaccharide

MALDI Matrix Assisted Laser Desorption Ionization

MeOH Methanol min Minute(s)

MMP Matrix Metalloprotease

MsCl Mesyl Chloride

MSCRAMMs Microbial Surface Components Recognizing Adhesive Matrix Molecules

Mu-GlcNAc Methylumbellifery-β-N-acetylglucosamine

MWCO Molecular Weight Cutoff

NBS N-Bromosuccinimide

NOE Nuclear Overhauser Effect oC Degrees Celsius

OD600 Optical Density at 600 nm pip Piperidine pNA p-Nitroanilide

PNAG β-1,6-N-Acetylglucosamine Homopolymer

xvi

pNP p-Nitrophenolate pNPA p-Nitrophenyl Acetate pNP-GlcNAc p-Nitrophenyl-β-N-acetylglucosamine pNP-NCO p-Nitrophenyl Isocyanate py Pyridine

QS Quorum Sensing

RP-HPLC Reverse Phase-High Pressure Liquid Chromatography rt Room Temperature

SAHA Suberoylanilide Hydroxamic Acid

SAMA-OPFP Pentafluorophenyl S-acetylthioglycolate

SDS-PAGE Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis

TEA Triethylamine

TFA Trifluroracetic Acid

THF Tetrahydrofuran

TLC Thin Layer Chromatography

TMS-CF3 Trimethylsilyl Trifluoromethane

UDP-GlcNAc Uridine Diphosphate N-Acetylglucosamine

ZBG Zinc Binding Group

xvii 1

1 General Introduction 1.1 Biofilm Structure and Life Cycle

Biofilms are complex communities of bacteria that have adhered to a surface and are held together by an extracellular matrix.1 This matrix is principally composed of biological polymers including DNA, proteins, and polysaccharides, as well as other small molecules. These colonies often form mushroom-like structures that allow the bacteria to be sheltered from hostile environments and host defenses.2 Biofilm life cycles can be divided into three major stages: attachment, growth, and release (Figure 1.1).3

Figure 1.1 Model of the biofilm life cycle. 1-2) attachment of planktonic bacteria to a surface. 3-4) growth of the macroscopic biofilm structure. 5) release of planktonic to the environment. Image obtained from the Center of Biofilm Engineering, Montana State University.

1.1.1 Attachment

The development of a biofilm begins with planktonic bacteria adhering to a surface. The initial attachment is accomplished by weak van der Waals forces, electrostatic forces, and hydrophobic interactions between the bacteria and the surface.4 This initial attachment is

2 dynamic and reversible, during which hydrodynamic forces, repulsive forces, or nutrient availability may act as motives for bacteria to return to the planktonic population. Upon initial attachment to a surface, microbes may transition to a biofilm mode of existence. This involves the bacteria producing components necessary for permanent and irreversible attachment to the surface, known as MSCRAMMs (microbial surface components recognizing adhesive matrix molecules). Well-studied Gram-positive biofilm producers Staphylococcus aureus and Staphylococcus epidermidis can produce and export to the cell-surface up to 20 different MSCRAMMs.5 These components have a common structure that includes an exposed binding domain, a cell-wall spanning domain, and a domain responsible for covalent or non-covalent attachment to a surface.6 For example, Biofilm Associated Protein (BAP) produced by S. aureus and S. epidermidis facilitates adherence through non-covalent hydrophobic interactions on inert surfaces.7

1.1.2 Growth

Upon permanent attachment to a surface, bacteria in the newly-forming biofilm begin replication and matrix formation. Significant up-regulation of the genes responsible for cell envelope and extracellular matrix formation, along with a concurrent down-regulation of genes responsible for motility, is observed.8 Growth persists until the colony reaches a mature size, with mushroom-like or tower-like profiles often being observed in vitro, with its maximal size determined by the hostility of the environment. This stage of colony development and maturation is dependent on environmental conditions and can last for several days.4a, 9

Figure 1.2 3-D reconstruction of a bacterial biofilm by Vibrio cholerae. Bacterial cells (blue) attach themselves to the surface with glue-like proteins (green + grey). The bacterial clusters then cover themselves with a protective shell of proteins and polysaccharides (red).10

1.1.3 Release

The final stage of a biofilm life cycle involves a partial breakdown of the extracellular matrix and release of planktonic cells from the mature biofilm. This stage of development allows for the initiation of new biofilm colonies distant from the initial attachment site.9 In Actinobacillus actinomycetemcomitans, Dispersin B (DspB), a β-hexosaminidase enzyme, is secreted into the extracellular environment to partially degrade the polysaccharide matrix and subsequently allow for planktonic release.11 Likewise, some bacterial strains including Pseudomonas aeruginosa produce and release rhamnolipid surfactants which can also result in biofilm dispersal.12

1.2 Biofilm Resilience and Antibiotic Resistance

Biofilm-related infections typically occur from contact of a colonized surface with a host.13 Most commonly, these infectious surfaces include indwelling medical devices such as catheters, heart valves, and orthopedic devices,14 but biofilms can also be found on natural surfaces such as teeth,15 the lungs in the case of cystic fibrosis patients,16 and in the middle ear of patients with persistent otitis media infections.17 Common biofilm-forming bacteria that establish

4 colonies on indwelling medical devices include S. epidermidis,18 S. aureus,19 E. coli,20 Acinetobacter baumannii,21 Bordetella bronchiseptica,22 Bordetella pertussis,23 Yersinia pestis,24 Actinobacillus pleuropneumoniae,25 and P. aeruginosa.26 Once such a device has been compromised by biofilm bacteria, chronic infection of the host can follow.27 In most cases, removal or replacement of the device is the only effective means of treatment.2a

Bacterial infections are regularly treated with antibiotics; however, this form of treatment has shown limited success when attempting to treat a bacterial biofilm infection.2a, 27-28 It is generally the case that biofilm-residing bacteria demonstrate increased antibiotic resistance in comparison to their planktonic counterparts. Some reports suggest this increase of antibiotic resistance to be up to 1000-fold.29 This resistance is likely a result of the multicellular nature of biofilms, rather than through the mechanisms of efflux pumps or acquired mutation.30 There are several hypotheses to explain this observation, all relating to a central theme: the communal existence of biofilm bacteria leads to their increased antibiotic resistance and environmental resilience. In fact, it has been shown that dispersion of a biofilm structure, for example through mechanical means, negates this antibiotic resistance improvement.2a, 31

The extracellular matrix is thought to form a physical barrier, which sterically limits the penetration of some antibiotics including β-lactams32 aminoglycosides33 and host antibodies to the biofilm core, leaving only peripheral microbes susceptible. The steric barrier of the biofilm matrix also limits the activity of the host‘s phagocyte cells, which rely on an engulfing mechanism for bacterial removal, reserving their efficacy only towards planktonic bacteria.2a

Concurrently, the physiological heterogeneity of the bacteria in the biofilm can lead to antibiotic resistance. Planktonic bacteria exist in a uniform environment and thus live in similar physiological states. In a biofilm, gradients of nutrients, oxygen, waste, pH, etc exist which alter the physiological state of each bacterium in the biofilm depending on its specific local environment. It has been shown that biofilms consist of at least two subpopulations as a result of these chemical gradients: a growing aerobic subpopulation and a more dormant anaerobic subpopulation.34 Since many antibiotics such as aminoglycosides, β-lactams and fluoroquinolones target bacterial replication and metabolism pathways, the dormant subpopulation of bacteria is likely resistant to these antibiotic modes of action.35 In vitro

5 additions of compounds that stimulate anaerobic growth such as arginine and nitrates have shown to enhance the antimicrobial activity of these antibiotics.36

Finally, biofilm bacteria have also demonstrated a higher adaptive tolerance. The mutation frequency of biofilm-growing bacteria has been observed to be significantly increased compared to isogenic, planktonic-growing bacteria, likely through a stress-induced mechanism.37 Additionally, an increased rate of horizontal gene transfer among proximal bacteria is also observed. This allows for easy imprint of antibiotic resistance amid the communal biofilm bacteria.38

1.3 Extracellular Matrix

A commonality in all biofilm communities is the existence of an extracellular matrix.39 This matrix facilitates cohesion between the individual microbes in a biofilm, and it is composed of biomacromolecules including DNA, proteins, lipids, and long repeating polysaccharides, depending on the bacterial species/strain in question.40 Whereas DNA is thought to play an important role in the establishment of the biofilm structure, polysaccharides are considered to be the major structural component of the biofilm matrix.41 Bacterial biofilms that are devoid of polysaccharides are unable to form multilayer biofilms.42 The focus of this chapter will be on polysaccharide-based matrices.

1.3.1 Psl and Pel Matrices

In P. aeruginosa, an opportunistic pathogen, the exopolysaccharide matrix is composed of two main polysaccharides: the Pel exopolysaccharide, whose structure has not been fully characterized, and the Psl exopolysaccharide, a polysaccharide containing a pentasaccharide repeating unit consisting of D-mannose, L-rhamnose, and D-glucose (Figure 1.3). 43 In general, P. aeruginosa strains predominantly produce and secrete one of the two polysaccharides at any given time.44 The Psl exopolysaccharide is biosynthesized by the 12 gene operon pslABCDEFGHIJKL. It has been predicted that the Psl exopolysaccharide pathway is initiated on a lipid carrier on the inner membrane and subsequent oligosaccharide subunit polymerization is then achieved in the periplasm. The final polysaccharide is then translocated across the periplasm and exported through the outer membrane. This process is similar to E. coli capsule synthesis and export.45

Figure 1.3 Repeating unit of Psl polysaccharide produced by Pseudomonas aeruginosa

1.3.2 Alginate Matrix

Alginate is an acidic exopolysaccharide also found in P. aeruginosa, particularly strains involved in cystic fibrosis.16 Its importance in the pathology of cystic fibrosis has led to alginate being the first and best characterized biofilm matrix exopolysaccharide.46 The alginate polysaccharide consists of two monosaccharide uronic acid subunits: β-D-mannuronate residues and its C-5 epimer α-L-guluronate residues (Figure 1.4). The biosynthesis of alginate is accomplished by the gene operon algADEFGIJKL8(44).47 Biosynthesis involves 1,4- polymerization of β-D-mannuronic acid residues in the cytoplasm and concurrent transport into the resulting polymannuronate polymer into the periplasm. The polysaccharide is then modified by epimerases to produce α-L-guluronate residues within the mature polymer, and O-acetylated at various residues before final translocation across the periplasm and export into the environment.

The final mature alginate matrix contains homopolymeric regions of poly-β-D-mannuronate and 46 poly-α-L-guluronate, as well as heteropolymeric regions. Alginate can facilitate the formation of gel like structures in the presence of chelating cations including sodium and calcium, with functional properties strongly correlated to the mannurate/guluronate ratio and sequence.48

Figure 1.4 Structure of the alginate polysaccharide. It contains uronic acid residues β-D-mannuronatre (M) and its C-

5 epimer α-L-guluronate (G). R = H or Ac.

1.3.3 Cellulose Matrix

Cellulose is a polysaccharide comprised of β-1,4-glucose monomer subunits (Figure 1.5). It is the most abundant sugar polymer in the living world, existing as a structural component of cell walls of green plants.49 It has been identified as a biofilm matrix component of several bacterial species including Agrobacterium tumefaciens,50 E. coli,51 Pseudomonas fluorescens,52 and Gluconacetobacter xylinus.53 Cellulose consists of insoluble fibers that aggregate at the cell surface to form a gel-like substance. Hydrogen bonds between cellulose polymer strands lead to its fibrous structure. The gene operon responsible for cellulose polymer synthesis is the bcsABCD operon.50 It is polymerized by a multimeric protein complex located in the cytoplasmic membrane, followed by export into the extracellular environment, where multiple chains can begin to form higher order aggregates.

Figure1.5 Structure of cellulose polysaccharide.

1.3.4 Poly N-Acetyl Glucosamine Matrix

In the biofilms of Gram-negative E. coli and Gram-positive S. epidermidis, which are both medicinally relevant bacteria, the extracellular matrix is composed of a β-1,6-N-acetyl

8 glucosamine homopolymer (PNAG, poly-N-acetylglucosamine), which is de-N-acetylated at variable positions along the scaffold (dPNAG, de-N-acetylated poly-N-acetylglucosamine) (Figure 1.6).39, 54 De-N-acetylation widely varies among different bacterial species and strains, with isolated dPNAG exhibiting a range from 1% to 40% de-N-acetylation, with polymer strands on the scale of hundreds of monosaccharide units long.20b, 54-55 De-N-acetylation results in positive charges along the polymer at the de-N-acetylated sites, and is thus thought to lead to electrostatic interactions with negatively-charged functional groups including phosphates and carboxylates on a neighboring cell‘s cell wall, leading to robust adhesion among bacterial cells. Strains of bacteria displaying only PNAG polysaccharide showed limited ability to form biofilms and demonstrated compromised antibiotic resistance.55 Additionally, these strains also demonstrated diminished virulence in mice.56 For the remainder of this thesis, only this polysaccharide matrix will be discussed in detail.

Figure 1.6 Structures of PNAG (top) and dPNAG (bottom). Polymer length and degree of de-N-acetylation can vary among different bacterial strains.

1.3.4.1 Biosynthesis of PNAG and dPNAG

In both Gram-positive and Gram-negative strains of bacteria, PNAG and dPNAG matrices are synthesized similarly, with differences accommodating for export of the polysaccharide through the extra membrane of Gram-negative bacteria. In E. coli, synthesis is accomplished by the pgaABCD operon,54 whereas in S. epidermidis, this is achieved by the icaABCD operon.19 It is not well understood whether dPNAG remains covalently attached to the cellular membrane or remains loosely associated to the bacterial cells by electrostatic or other non-covalent interactions in either system.

1.3.4.1.1 PNAG Biosynthesis in Gram-negative Bacteria

In Gram-negative E. coli, synthesis and extracellular display of dPNAG is accomplished by four proteins. In a model proposed by Itoh et al., initially, UDP-GlcNAc is polymerized into the β-1,6-N-acetyl glucosamine homopolymer PNAG and consecutively transported to the periplasmic space by the transmembrane proteins PgaC and PgaD, with PgaC predicted to be a glycosyl transferase (Figure 1.7).55 PgaD has recently been found to act as an allosteric regulator of polysaccharide formation, and is dependent on the presence of cyclic di-GMP (c-di-GMP).57 Allosteric binding of c-di-GMP to PgaD allows for formation of a PgaCD complex, which in turn allows for the glycosyl transferase activity required for PNAG polymerization. Upon polymerization and transfer of the polysaccharide to the periplasmic space by PgaCD, PgaB is responsible for partial de-N-acetylation of the growing polysaccharide chain within the periplasm. PgaA is then thought to be responsible for the export of the developed polysaccharide into the extracellular space. Notably, ΔPgaB knockout strains of E. coli accumulate only PNAG into the periplasmic space; neither de-N-acetylation nor export is observed. The pgaABCD operon has also been implicated in dPNAG synthesis in other medically relevant Gram-negative bacterial strains including Aggregatibacter actinomycetemcomitans,58 A. baumanii,23 A. pleuropneumoniae,25 and Y. pestis59

Figure 1.7 Model of dPNAG biosynthesis and export in Gram-negative E. coli. PgaB is a periplasmic de-N-acetylase responsible for partial de-N-acetylation of PNAG. Image adapted and used with the permission of J. Poloczek.

1.3.4.1.2 PNAG Biosynthesis Gram-positive Bacteria

In Gram-positive Staphylococcus strains including S. epidermidis60 and S. aureus,61 the biosynthesis of dPNAG is similar to that of the Gram-negative strains, employing the icaABCD operon.62 In a model proposed by Götz et al, this system utilizes a complex of membrane- associated proteins IcaC, IcaA and IcaD to elongate and transport the growing PNAG into the extracellular space (Figure 1.8).63 IcaB, a secreted protein, is then subsequently responsible for the de-N-acetylation of the PNAG polysaccharide to form dPNAG. Akin to ΔPgaB E coli, ΔIcaB strains of S. epidermidis result in bacteria with compromised biofilm-forming ability, and isolation of the polysaccharide reveals only the fully acetylated PNAG polymer.

Figure 1.8 Model of dPNAG biosynthesis and export in Gram-positive S. epidermidis. IcaB is an extracellular de-N- acetylase responsible for partial de-N-acetylation of PNAG. Image adapted and used with the permission of J. Poloczek.

1.3.4.2 PgaB and IcaB

1.3.4.2.1 PgaB

PgaB, a lipoprotein localized in the periplasm, is a 672 amino acid enzyme with 2 domains: an N-terminal deacetylase domain and a C-terminal domain homologous to glycosyl hydrolases.64 Crystallographic studies of PgaB have revealed the presence of a metal ion in the predicted deacetylase active site of the N-terminal domain, which shares sequence homology with other enzymes in the Carbohydrate Esterase Family 4 (CE4), a family of metalloenzymes (Figure 1.9). A number of crystal structures have been produced, containing different metal ions including Fe2+, Ni2+, and Zn2+, all bound to the same Asp115-His184-His189 triad of residues.65 In vitro studies of PgaB expressed in the presence of different metals had varied rates of de-N- acetylase activity on synthetic β-1,6-N-acetyl glucosamine pentasaccharides, suggesting de-N- acetylation is metal-dependent, with Fe2+, Co2+,and Ni2+ loaded enzyme demonstrating the highest activity.64 Additionally, PgaB demonstrates size dependent de-N-acetylation activity when tested with oligomeric PNAG substrates, showing increased de-N-acetylase activity with increasing oligomer size ranging from tri- to pentasaccharide. Strong selectivity is also observed for de-N-acetylation of the third carbohydrate unit in a synthesized PNAG pentasaccharide,

13 suggesting a total of four flanking carbohydrate binding substites in the active site, two on each side.

Figure 1.9 Ribbon trace of the crystal structure of PgaB N-terminal deacetylase domain (residues 41-310). Trace is colored from blue at the N-terminus to red at the C-terminus. The active site nickel ion is shown in grey and is bound by Asp115-His184-His189 and an acetate molecule.64

1.3.4.2.2 IcaB

IcaB, a secreted protein found in Gram-positive S. epidermidis and S. aureus, is a 289 amino acid de-N-acetylase with sequence homology similar to that of the N-terminal domain of PgaB and other enzymes in the CE4 family.56 Similar to PgaB, it has been reported that deletion mutants of IcaB resulted in production of fully acetylated PNAG polysaccharide with limited cell surface retention, as well as a dramatically reduced pathogenicity in animal models of biofilm infections with S. epidermidis and S. aureus. Immunofluorescence spectroscopy has shown that IcaB does associate with the cellular membrane and is also present in the cellular medium.66 In vitro studies of IcaB in the presence of different metals also showed differential de-N-acetylase

14 activity with PNAG pentasaccharides, suggesting that de-N-acetylation is metal dependent.67 ICP-AES analysis on as-isolated IcaB showed zinc to be the predominant metal; however the addition of Co2+ to the as-isolated enzyme increased the de-N-acetylation activity four-fold. This sort of trend has been observed with many other Zn2+ metallohydrolases and other CE4 enzymes.68 In terms of substrate selectivity, IcaB displayed partial dependence on substrate length, showing negligible de-N-acetylation activity on monosaccharide and disaccharide PNAG substrates, and equivalent levels on trisaccharide to hexasaccharide PNAG substrates, suggesting a three unit binding site containing two occupancy sites and the active site. Tandem ESI-MS/MS analysis of the de-N-acetylated PNAG confirmed this assessment, and also showed a preference for de-N-acetylation to occur at the residue one sugar removed from the reducing terminus on pentasaccharides, signifying an extended binding motif for carbohydrate units on the non- reducing end of the PNAG polymer.

1.4 Biofilm Treatment Strategies

Sixty-five percent of all human persistent bacterial infections are considered to be biofilm related.69 Due to the antibiotic tolerance of biofilm-forming bacteria as previously discussed, novel strategies are required to combat growing biofilms. A number of anti-biofilm strategies have been proposed. These methods are based on several themes: prevention of initial formation, disruption or weakening of a maturing biofilm, or the targeted killing of the bacteria within a biofilm (Figure 1.10).34

Figure 1.10 Outline of the stages in biofilm development and lists of strategies aimed at inhibiting/disrupting the biofilm at specific stages

1.4.1 Prevention of Biofilm Formation

1.4.1.1 Antibodies

The use of antibodies against specific components of the biofilm has been investigated; particularly, the development of antibodies towards PNAG and dPNAG in the hopes of creating an anti-biofilm vaccine.61, 70 In one study, mice were immunized with goat antibodies towards chemically-modified native dPNAG (~85% de-N-acetylated) and subsequently infected with a lethal dose of S. epidermidis. Mice that were immunized against dPNAG showed significantly

16 decreased mortality rates. Interestingly, antibodies raised against native PNAG (~10% de-N- acetylation) showed low binding capability to dPNAG, whereas antibodies raised towards dPNAG showed equal binding capacity to both PNAG and dPNAG in ELISA.61 Attempts to conjugate dPNAG to a S. aureus-specific immunogenic protein carrier (clumping factor A) failed to improve the opsonic killing activity, and the protective activity of raised antibodies towards it.71

Future development in this field requires the availability of large amounts of well-defined dPNAG antigens. Work in our lab by Leung et al. developed pentasaccharide-sized PNAG and dPNAG glycoconjugates.72 These oligosaccharides are available for use as antigens for the development of anti-PNAG/anti-dPNAG antibodies. Conversely, work by Ninfantiev et al. has led to the synthesis of a PNAG nonasaccharide as well as several defined dPNAG nonasaccharide variants.73 One of these nonasaccharide variants, a β-1,6-glucosamine homopolymer (9GlcNH2), representing fully de-N-acetylated PNAG was glycoconjugated to 74 tetanus toxoid (9GlcNH2-TT) and was used as a potential antigen candidate. In vitro bactericidal killing of the following PNAG producing bacteria strains was accomplished using rabbit serum raised towards 9GlcNH2-TT: Streptococcus pneumonia (4/4 strains), Enterococcus faecalis (3/3 strains), Streptococcus pyogenes (3/3 strains), Neisseria gonorrheae (7/7 strains) and Neisseria meningitidis (5/5 strains).75 A control experiment using a P. aeruginosa alginate strain showed no bactericidal activity, verifying PNAG-dependent activity. Mice immunized with rabbit antibodies towards 9GlcNH2-TT demonstrated significantly reduced mortality when challenged with lethal doses of S. pyogenes, S. pneumoniae, Listeria monocytogenes, and N. meningtidis. Anti-malarial activity was also observed in mice.75 This antigen is currently undergoing clinical testing and has passed phase I clinical trials as a potential PNAG-based immunotherapy. Unfortunately none of the reported results targeted a pre-formed bacterial biofilm colony. Additionally, only limited utility of this antigen has been observed with S. epidermidis, S. aureus and E. coli, due to interference of its opsonic killing activity by antibodies toward capsular polysaccharides.76

1.4.1.2 Inhibition of Bacterial Attachment

One approach that has seen a significant undertaking is the modification of abiotic surfaces preceding biofilm exposure.77 A rough, irregular surface will increase the extent of

17 biofilm formation compared to a smooth surface, based on peak profile analysis of various polyurethane central venous catheters.78 The same can be said for a hydrophobic surface when compared to a charged surface, with the charged surface demonstrating a capability to inhibit S. epidermidis adherence in vitro and in vivo.79 Thus, material scientists have grafted surfaces with a variety of different agents in the hope of preventing initial microbial colonization, including antibiotics, disinfectants, and antiseptics.77

In one study, vancomycin-coated titanium has been shown to retain antibacterial activity for up to 11 months, even after multiple challenges in vitro.80. Surfaces designed to retain non- covalently bound antibiotic surfaces have also been explored, but no system has been developed that retains antibiotic release over the therapeutically relevant time scale of weeks to months.81 Unfortunately, the in vivo long-term efficacy of covalently bound antibiotics on the surface of implants has not been reported.82 This practice is most commonly employed with bone grafts, where grafts are impregnated with antibiotics prior to surgery.83 Long term efficacy is not an issue in this case since the graft is absorbed into the bone structure over the course of weeks.

Another strategy involves coating abiotic surfaces with silver nanoparticles. Historically, silver has been used to sterilize wound infections, and was used extensively during World War I.84 Though the potential toxicity to humans has diminished its use, the advent of nanotechnology has revived its popularity; the combination of small size and the large surface area-to-volume ratio of silver nanoparticles enables maximal proximate interactions with microbial membranes.85 Its bactericidal mode of action is owing to silver‘s interaction with enzyme thiol- groups which can inactivate essential enzymes involved in DNA replication and electron transport.84, 86 Silver nanoparticles coated implants in rabbits showed > 95% inhibition of P. aeruginosa and S. epidermidis biofilms. Moreover, silver accumulation in the host tissue was not observed 28 days after impregnation.87 Silver is now currently used as an exogenous wound dressing to combat biofilm formation.88

Finally, recent evidence has demonstrated that exogenous bacterial biofilm exopolysaccharides can inhibit or destabilize biofilm formation from other species. For example, it was found that Psl and Pel solutions from P. aeruginosa culture supernatants disrupted S. epidermidis and S. aureus biofilms without inhibiting bacterial growth.89 Furthermore, the presence of P. aeruginosa inhibited S. epidermidis biofilm formation in dual-species in vitro

18 experiments.90 Conversely, E. coli that produced the Ec300p anti-adhesion polysaccharide were able to quickly disperse S. aureus from E. coli/S. aureus mixed biofilms. Additionally, this E. coli strain was significantly protected from colonization by invading planktonic S. aureus.91 It has been proposed that the disruption caused by these anti-biofilm polysaccharides is due to several reasons: the foreign polysaccharide competitively inhibit multivalent carbohydrate- protein interactions essential for adhesion, it acts as regulators for biofilm formation, and it acts as a signaling molecule that can impact gene expression patterns involved in biofilm formation and maintenance (Figure 1.11).92 A range of polysaccharides have demonstrated a broad spectrum of biofilm inhibition and disruption capacity.93 As such, polysaccharides have been considered for use as surface coatings for indwelling medical devices; however, this utility has yet to be explored.

Figure 1.11 (A) Competition – Anti-biofilm polysaccharides can inhibit foreign biofilm formation (left) or enhance foreign biofilm dispersal (right). (B) Regulation – Anti-biofilm polysaccharides secreted to the extracellular medium can regulate adhesive behavior. (C) Signaling – Anti-biofilm polysaccharides secreted by competing bacteria can signal biofilm bacteria to alter their own gene expression.92

1.4.2 Removal or Disruption of Existing Biofilms

1.4.2.1 Enzymatic Disruption of Biofilms

The treatment of biofilms with enzymes has been explored; in particular, enzymes capable of breaking down the extracellular matrix.94 Enzymes that have been explored in this way include DNases for P. aeruginosa biofilms,55, 95 and glycosyl hydrolases for E. coli and S. epidermidis.40 Dispersin B, a 42 kDa exo-hexosaminidase, has been explored as an exogenous

20 treatment for PNAG biofilms. It has been shown to capably cleave β-1,6-N-acetyl glucosamine homopolymer into monosaccharide units.94a It has also demonstrated an ability to disperse mature biofilms, and in conjunction with the antimicrobial agent triclosan, nearly eliminate biofilm formation in vitro.29, 96 Recently, this synergistic combination has been marketed as topical wound treatment DispersinB®, and has been undergoing clinical trials since 2010. Unfortunately, as a protein, it is not hypothesized to be stable in blood, and therefore is not an ideal treatment for biofilms on implanted devices.

1.4.2.2 Chemical Disruption of Biofilms

Chemical treatment of biofilms has been explored in the context of quorum sensing (QS) inhibition.97 Quorum sensing is a system of stimulus and response outputs correlated to population density among bacteria; a method of coordinating gene expression among proximal microbes.97a, 98 In Gram-negative bacteria, such as E. coli and P. aeruginosa, crosstalk is accomplished by means of signal molecules such as N-acyl homoserine lactones (AHL).45 It has been proposed that this mechanism enables the arrest of virulence factors until enough bacteria have accumulated to conquer the host defense.99 One goal for researchers has been to develop compounds to inhibit QS, and thus prohibit bacteria from producing virulence factors. An array of inhibitors have been developed and seen moderate success with P. aeruginosa biofilms (Figure 1.12).100 Unfortunately, the inhibition of QS may not be a broad spectrum treatment, as it has been shown to increase biofilm formation in certain Staphylococcus strains.56, 101

Figure 1.12 QS inhibitors against P. aeruginosa biofilms.

Another investigation into chemical biofilm treatment by Losick et al. involved the utility 102 of D-amino acids to trigger biofilm disassembly. In particular, D-leucine, D-methionine, D- trypthophan, and D-tyrosine showed success against various bacterial strains, including PNAG dependent S. epidermidis and S. aureus. It was hypothesized that naturally-occurring biofilms incorporate D-amino acids other than D-alanine into their peptidoglycan as part of a signaling cascade to promote biofilm disassembly, and thus the accumulation of additional D-amino acid in the environment might enhance the rate of cascade.102-103 S. aureus cultures soaked in a medium containing either D-tyrosine (50 µM) or a D-amino acid cocktail containing the amino acids outlined above (15 nM each) inhibited biofilm formation in vitro over 24 hours. Surface impregnation of D-amino acids also demonstrated effectiveness in preventing S. aureus biofilm growth on poly-urethane surfaces.104 Unfortunately, this concept has yet to be tested in vivo, but does show great promise.102

Finally, there have been several reports of the aquatic natural product bromoageliferin, a 2-aminoimidazole compound disrupting mature biofilms of P aeruginosa and S. aureus (Figure 1.13a).105 Mechanistic analysis of bromoageliferin led to the discovery that its inhibitory mode of action is based on its Zn2+-chelating ability.106 Discovery of this natural product led to the synthesis and study of an array of 2-aminobenzimidazole derivatives based on the core structure of bromoageliferin (Figure 1.13b). One of the compounds, 2-ABI3, demonstrated IC50 values of 890 nM, 1.4 µM, and 570 nM towards methicillin-resistant S. aureus, vanomycin-resistant

Enterococcus faecium, and S. epidermidis respectively, where IC50 represents the concentration

22 of compound that inhibits biofilm development by 50% (Figure 1.13c). The addition of 200 µM of zinc suppressed the inhibitory ability of 2-ABI3, suggesting a zinc dependent mechanism. This chelative activity was also verified by 1H NMR analysis.106

Figure 1.13 Small molecule biofilm inhibitors

1.5 Purpose of the Current Study

As discussed previously, the dPNAG matrix is the defining component of biofilm formation and survival in medicinally relevant bacteria including E. coli and S. epidermidis. Studies with DspB, a glycosidase that specifically degrades the PNAG polysaccharide matrix, have demonstrated that depolymerization of the matrix in PNAG-based biofilms results in colony dispersal. Additional research on the pga and ica operons, specifically deletion studies, has shown that bacteria that is unable to de-N-acetylate the PNAG polymer is unable to effectively produce biofilm colonies.56, 101a This implies that PgaB and IcaB, the β-1,6-GlcNAc de-N- acetylase enzymes in Gram-negative and Gram-positive bacteria respectively are a crucial factor in the biofilm formation process, and thus represent attractive targets for potential antibiofilm compounds. To date, there are few reports of small molecules that specifically disrupt or target biofilms and none that target the extracellular matrix.

The purpose of this study was thus to develop and synthesize novel inhibitors to target PgaB and IcaB, in order to develop an agent that would ultimately inhibit biofilm formation and/or promote biofilm colony degradation. Additionally, we endeavored to develop assays to

23 study the effectiveness of current biofilm treatment methods as well as to determine the effectiveness of synthesized inhibitors.

Methods were also developed to evaluate the efficacy of DspB to treat biofilm-related infections. This was accomplished by developing a novel carbamate substrate to measure DspB activity in vitro via a continuous absorbance spectroscopy assay. This method improved on the existing method of detecting DspB activity in several ways, including utilizing a continuous assay as opposed to a discontinuous assay, and achieving a 30-fold greater analytical sensitivity resulting from improved enzyme recognition and improved catalytic consumption of the novel substrate. The scope of this study is presented in chapter 2.

Subsequently, an array of potential inhibitors was developed towards PgaB and IcaB. Previous work by Poloczek and Pokrovskaya in our lab has demonstrated both enzymes to be metal dependent with respect to their catalytic ability.64, 67 Thus the design of the inhibitors was based on a carbohydrate GlcNAc scaffold, mimicking the natural enzyme substrate and containing a metal binding moiety expected to bind to the enzymatic metal ion resulting in inhibition of its catalytic ability. Transition state mimic inhibitors based on trifluoromethyl ketone moieties were also explored. The scope of this study is presented in chapter 3.

Finally, a new method was developed to measure PgaB and IcaB activity. Earlier methods included a cumbersome discontinuous and indirect assay for measuring de-N- acetylation activity utilizing fluorescamine to detect the resulting free amines at specific time- points over a lengthy 24 hour period using PNAG oligomers (tri- through hexasaccharides) as substrates, which were both tedious to synthesize and purify. The new method involved the development of a new assay using a substrate that could be readily synthesized, and its activity monitored continuously over a short period (20 minutes). With this new approach, it was then possible to measure inhibitor effectiveness quickly and accurately. The scope of this work is presented in detail in chapter 4.

2 Substrate Development for Hexosaminidases

The synthesis of the acetal substrate was done by Dr. Somnath Dasgupta. This chapter has been reproduced in part with full permissions from:

Chibba, A.; Dasgupta, S.; Yakandawala, N.; Madhyastha, S.; and Nitz, M. (2011) Chromogenic carbamate and acetal substrates for glycosamidases. Journal of Carbohydrate Chemistry, 30, 549.

2.1 Introduction

β-Hexosaminidases are a class of glycoside hydrolases that are responsible for the hydrolysis of hexosamine subunits from a carbohydrate polymer. Many hexosaminidases belong to the family 20 glycoside hydrolases class of enzymes. These exo-hexosaminidases use the 2- acetamido group as an anchimeric participating group to achieve glycoside hydrolysis and act on the non-reducing hexosamine residues (Figure 2.1).107 The mechanism of hydrolysis involves intramolecular nucleophillic attack of the glycosidic bond by the 2-acetamido carbonyl to form an oxazolinium ion intermediate, which is stabilized by enzymatic carboxylate residues. Upon expulsion of the polysaccharide chain, the oxazolinium intermediate is subsequently attacked by water to release the hexosamine monosaccharide.

Figure 2.1 Typical anchimeric participation mechanism for the hydrolysis of GlcNAc polymers by family 20 hexosaminidases.

β-Hexosaminidase activity has been implicated in a number of human genetic lysosomal disorders. As such, a considerable research effort has been committed to developing different assays for monitoring β-hexosaminidase activity.108 Assays using fluorogenic and chromogenic

25 substrates, such as p-nitrophenyl-β-N-acetylglucosamine (pNP-GlcNAc) or 4-methylumbellifery- β-N-acetylglucosamine (Mu-GlcNAc), are commonly used (Figure 2.2).109 Alternatively, using a synthesized radiolabeled ‗natural polysaccharide‘, incorporating 3H onto a GlcNAc or N- acetylgalactosamine (GalNAc) residue, is also a viable method.110 Whereas chromogenic and fluorogenic substrates are ideal for patient diagnoses in hospital laboratories, radiolabeled substrates would be used for more specialized assays and diagnoses. The remainder of this chapter will focus on chromogenic substrates.

Figure 2.2 Common chromogenic aryl glycoside substrates used to measure hexosaminidase activity.

Dispersin B, an exo-acting hexosaminidase from the oral pathogen A. actinomycetemcomitans, is capable of cleaving PNAG-like β-1,6-linked N-acetylglucosamine polymers from the non-reducing end.111 In vivo, it is a secreted protein that mediates A. actinomycetemcomitans biofilm attachment and detachment properties.58a It has since been developed as a topical treatment for exogenous biofilm infections, and marketed as DispersinB®.112

Previous work on DspB has shown that it is capable of cleaving PNAG isolated from biofilm-forming E. coli, and results in reaction products that include GlcNAc monomers and various uncharacterized GlcNAc oligomers.94b Interestingly, no hydrolytic products were generated when tested with chitin, a β-1,4-linked N-acetylglucosamine homopolymer. Analysis of DspB activity with the chromogenic carbohydrate substrates pNP-GlcNAc or Mu-GlcNAc

26 showed a heavy dependence on the identity of the aglycone (Figure 2.2).94b DspB‘s preference for the β-1,6-linkage has also been verified through molecular modeling.113 The kinetic parameters for DspB with pNP-GlcNAc and Mu-GlcNAc were poor in comparison to other family 20 hexosaminidases.94b We proposed that due to the selectivity of DspB, a substrate that more closely mimics the natural β-1,6-linkage may show improved catalytic activity, in contrast to pNP-GlcNAc or Mu-GlcNAc, whose linkages directly to aryl rings may be sterically disfavored and better mimic linkages to secondary hydroxyls (Figure 2.3).

Figure 2.3 Comparison of the β-1,4 and β-1,6 linkages in chitin and PNAG respectively.

The work described in this chapter will discuss an ongoing effort to develop more suitable substrates for DspB that better mimic the β-1,6-linkage, by introducing less steric bulk around the glycosidic linkage.114 This was accomplished through the synthesis of glycosyl acetal and glycosyl carbamate substrates that release chromophores upon enzymatic hydrolysis.

Traditionally, β-glucuronic carbamates were synthesized to attach to organic drug molecules to form a prodrug to improve solubility and biological uptake and aid in site selective delivery to a cancer site via the ADEPT (Antibody Directed Enzyme Prodrug Therapy) principle.115 In vivo hydrolysis of the pro-moiety carbamate by native β-glucuronidase enzymes then reveals the active drug molecule. Alternatively, p-nitroaniline carbamates have been previously used to monitor protease activity through a cascade degradadion mechanism involving the production of azaquinone methides, carbon dioxide, and p-nitroaniline. Prior to our work, neither glycosyl carbamates nor glycosyl acetals had been explored as chromogenic glycosidase substrates.116

2.2 Results and Discussion

2.2.1 Synthesis of Acetal and Carbamate Substrates

The synthesized substrates are derived from N-acetyl glucosamine with either an acetal or carbamate linkage at the reducing end. These linkages were chosen for their flexibility proximal to the glycosidic oxygen in comparison to the traditionally-used aryl substrates and for their potential fragmentation upon enzymatic hydrolysis, which would lead to a signal from either p- nitrophenolate (pNP) or p-nitroanilide (pNA) moiety (Figure 2.4). A substrate with greater activity towards DspB would overcome the challenges inherent to the system, including dilute conditions or assays conducted in biological media.

Figure 2.4 Fragmentation pathways for the proposed acetal substrate 1 and carbamate substrate 2.

2.2.1.1 Synthesis of Acetal Substrate 1

The synthesis of the acetal substrate 1 began from the alkylation of the known compound

3,4,6-triacetyl-2deoxy-2-phthalimido-D-glucose with 1-chloromethoxy-4-nitrobenzene, synthesized in two steps from pNP and chloromethylthioether, in basic conditions, yielding the β-glycoside product in 40% yield (Scheme 2.1).117 Removal of the phthalimido group with ethylenediamine in butanol, followed by acetylation of the resulting free amine in acetic

28 anhydride/pyridine yielded the per-acetylated substrate in 78% yield. Final global deprotection in Zemplén conditions afforded the final compound 1 in 90% yield.

Scheme 2.1 (a) ClCH2SMe, NaI, NaH, DMF (b) SOCl2, DCM. (c) NaI, K2CO3 (d) i. NH2CH2CH2NH2, Butanol, 120 o C. ii. Py, Ac2O (e) MeOH, NaOMe.

2.2.1.2 Synthesis of Carbamate Substrate 2

The carbamate substrate 2 was synthesized in four steps from commercially available N- acetylglucosamine (Scheme 2.2). Following peracetylation, selective deacetylation of the anomeric hydroxyl with benzylamine gave the free hemiacetal. This was then treated with p- nitrophenyl isocyanate (pNP-NCO) in toluene with a catalytic base to afford the protected glycosyl carbamate 8, which precipitated out of the reaction mixture exclusively as the β-anomer in 91% yield. This result is consistent with the results found in the literature, where glucose derivatives were reacted with various aryl and alkyl isocyanates to yield only β-carbamates when conducted in the presence of weak bases.115b Final deprotection of the remaining acetyl groups proved challenging: Zemplén conditions resulted in the return of N-acetyl glucosamine and p- nitroisocyanate, whereas acidic HCl in methanol resulted in hydrolysis of the carbamate, yielding only N-acetyl glucosamine and pNA. The final substrate 2 was obtained by careful addition of 3.3 equivalents of hydrazine monohydrate in methanol, while monitoring the reaction by TLC

29 over six hours. Crude NMR analysis of the reaction mixture suggested a 62% yield of the desired product by integration of the anomeric signals; however, 48% yield was obtained after flash column chromatography.

Scheme 2.2 (a) pNP-NCO, cat TEA, Tol, rt, 12 h, 91%. (b) NH2NH2·H2O, MeOH, rt, 6 h, 48%.

2.2.2 Substrate activity assay

Substrates 1 and 2 were submitted to enzymatic assays with either DspB, a β-1,6 hexosaminidase, or jack bean β-hexosaminidase, an enzyme with a preference for hydrolyzing β-1,4-linkages, while monitoring reaction progress by continuous absorbance spectroscopy analysis. Substrate 1 hydrolysis was monitored at A405 over a range of concentrations (0.1- 5 mM 1, 0.1 µM DspB or 25 nM jack bean β-hexosaminidase, 50 mM phosphate, 100 mM NaCl, pH

6.0) resulting in the production of pNP, whereas substrate 2 was monitored by A410 (same conditions as above), resulting in the subsequent production of pNA. These results were then compared to the enzymatic activity against the benchmark pNP-GlcNAc substrate under the same conditions, monitored at A405. The major point of difference for these assays is that they were run continuously, whereas pNP-GlcNAc hydrolysis assays are typically measured discontinuously at specific time-points after alkaline spiking to improve the phenolate analytical signal.

2.2.2.1 Acetal Substrate 1

Upon introduction of substrate 1 to the assay conditions outlined above, no increase in

A405 was detected over a 30 minute time period in the presence of either DspB or jack bean β- hexosaminidase. Further experiments were performed to determine whether this was a result of the glycosidic linkage not being cleaved or the resulting hydroxyl-pNP-acetal not fragmenting to formaldehyde and pNP. 1H NMR analysis of the product from the enzyme incubation showed no glycosidic bond cleavage, suggesting that 1 was not a suitable substrate for either hexosaminidase. Further competition assays with pNP-GlcNAc (0.5 mM) in the presence and

30 absence of 1 (5 mM) showed no difference in the pNP-GlcNAc hydrolysis rate, signifying that 1 did not inhibit either enzyme, and the lack of activity could be a result of inadequate enzyme recognition towards 1.

Due to the similarity of 1 to pNP-GlcNAc, it was confounding to discover that no affinity for DspB or jack bean β-hexosaminidase was observed. From the 1H NMR analysis of 1, one notable feature is the resonance of the acetyl protons at 1.46 ppm, which is significantly upfield from the typical 1.9 to 2.1 ppm region common to N-acetylglucosamine derivatives (Figure 2.5). We hypothesized that this was due to an anisotropic effect from the aryl ring, which may be in close proximity to the acetyl protons, leading to an enzymatically inactive conformation. Analysis of crystallized 2-phenylethyl glycosides have shown the aryl rings folded back towards the pyranoside ring, with similar 1H NMR resonance shifts being observed in similar large, flexible aromatic glycosides; napththylmethyl-N-acetylglucosamine glycoside was reported to have acetyl proton resonance of 1.68 ppm.118 Unfortunately, 1H NMR NOE experiments showed no through space contacts between the acetyl protons and aryl protons in 1, and thus our hypothesis could not be verified.

Figure 2.5 1H NMR array of the acetal GlcNAc substrate in MeOD (bottom) and methyl-N-acetylglucosamine glycoside in D2O (top). Note the difference in chemical shift for the acetyl protons (1.49 ppm vs 2.05 ppm).

2.2.2.2 Carbamate Substrate 2

Upon introduction of substrate 2 to the previously mentioned assay conditions, promising results were obtained. With DspB, a 20-fold improvement of Km was observed, combined with a

200-fold kcat/Km when compared to the benchmark pNP-GlcNAc (Table 2.1). When 2 was assayed in the presence of jack bean β-hexosaminidase, there was only a small improvement in

Km compared to pNP GlcNAc and a three-fold improvement in kcat. Experimentally-obtained Km and kcat values for pNP-GlcNAc were consistent with literature values for both DspB and jack bean β-hexosaminidase. Due to the low solubility of the substrates, saturation could not be achieved. Michaelis-Menten kinetic parameters were obtained from double-reciprocal (1/v vs 1/[2]) plots using first order kinetics (Figure 2.6).

Enzyme/ Km kcat kcat/Km Literature kcat/Km Substrate (mM) (s-1) (mM/s-1) (mM/s-1) DspB/2 2.3 ± 0.2 14.0 ± 0.1 6.1 - DspB/pNP-GlcNAc 46 ± 2 1.0 ± 0.1 0.021 0.0294b Jack bean/2 0.43 ± 0.02 22 ± 2 51 - Jack bean/pNP GlcNAc 0.58 ± 0.02 7.9 ± 0.3 13.6 35.6119

Table 2.1 Kinetic characterization of substrate 2 and comparison with pNP-GlcNAc.

Figure 2.6 Double-reciprical plot used to obtain Michaelis-Menten kinetic parameters for the carbamate substrate 2. This method was used to acquire the kinetic parameters in every case.

The improved Km for 2 with DspB supports the hypothesis that reduced steric congestion around the glycosidic linkage would improve recognition of the substrate as compared to pNP- GlcNAc. Consistent with previous work on 1,4-cleaving hexosaminidases, jack bean β- hexosaminidase showed no improved affinity for either aglycone, demonstrating a similar Km for

33 both 2 and pNP-GlcNAc. Fortuitously, both enzymes showed improved turnover of the glycosyl carbamate in comparison to pNP-GlcNAc, shown by the improved kcat values, demonstrating that glycosyl carbamates are ideal substrate candidates for other family 20 hexosaminidases. Overall, a 35-fold improvement in analytical sensitivity was observed with DspB and substrate 2 over pNP-GlcNAc (Figure 2.7). This is a result of improved kinetics and improved extinction coefficients of pNA over pNP in the reaction conditions at pH 6.0 (5450 M-1 cm-1 for pNA vs 7280 M-1 cm-1 for pNP).

Figure 2.7 Hydrolysis of substrate 2 (♦) monitored at 410 nm and hydrolysis of pNp-GlcNAc (█) monitored at 405 o nm over time in the presence of 50 nM DspB, 50 mM NaPO4 (pH 5.8), and 50 mM NaCl at 23 C.

2.2.3 2nd Generation Carbamate Substrate

Upon the successful application of carbamate substrate 2 with DspB, a new substrate was designed. The limitations of substrate 2 included limited solubility in aqueous media (5 mM) and the relatively low extinction coefficient of the pNA chromophore (5450 M-1 cm-1 at pH 6). To this end, the next generation carbamate contained a charged N-methyl-quinolinium moiety as the chromogenic species, which would be expected to expel 7-amino-1-N-methylquinolinium upon hexosaminidase cleavage.

2.2.3.1 Synthesis

The carbamate substrate 12 was synthesized in four steps from the commercially available N-acetylglucosamine (Scheme 2.3). Following peracetylation and selective deacetylation of the anomeric hydroxyl with benzylamine, the known free hemiacetal 7 was then treated with p-nitrobenzoic anhydride to afford the mixed anhydride 9. Nucleophillic attack of the anhydride center by 7-aminoquinoline yielded the carbohydrate carbamate 10, which could then be globally deprotected in the presence of hydrazine monohydrate in methanol. Methylation of the ring nitrogen with methyl iodide in DMF followed by precipitation upon the addition of ethanol led to the final product 12.

Scheme 2.3 (a) 3 eq (pNP)2CO, TEA, DCM, 12 h, rt, 80%. (b) 2 eq 5-quinamine, TEA, DCM, rt, 6 h, %. (c)

NH2NH2·H2O, MeOH, rt, 6 h 78%. (d) 1 eq MeI, DMF, rt, 3 h, 47%.

2.2.3.2 Substrate Activity

Compound 12 was found to be soluble in aqueous solution to at least 20 mM, compared to the maximal solubility of 5 mM for the carbamate substrate 2, which was expected due to its charged state. Unfortunately compound 12 also showed a limited stability in aqueous solution,

36 exhibiting a moderate background hydrolysis rate in the absence of enzyme (data not shown). As a result it was not tested in the presence of DspB.

2.3 Conclusions

In conclusion, novel acetal and carbamate substrates were synthesized and tested against two hexosaminodases. This is the first reported instance for both a glycosyl acetal and a glycosyl carbamate to be tested as an analytical substrate for glycosyl hydrolases. Traditionally, glycosyl carbamates have been used for development of prodrugs to improve solubility and site-specific delivery. Using the glycosyl carbamate structure 2, it was possible to monitor DspB activity in a continuous assay over a short time period with 35-fold greater sensitivity than the traditional pNP-GlcNAc under the same conditions.

3 Synthesis and Development of Inhibitors towards PgaB and IcaB

The synthesis of the phosphonamidate inhibitor was done by Dr. Vavara Pokrovskaya. This chapter has been reproduced in part with full permissions from:

Chibba, A.; Poloczek, J.; Little, D.J.; Howell, P.L.; Nitz, M. (2012) Synthesis and evaluation of inhibitors of E. Coli PgaB, a polysaccharide de-N-acetylase involved in biofilm activity. Organic and Bimolecular Chemistry, 10, 7103.

3.1 Introduction

Based on sequence homology, IcaB and PgaB, proteins involved in the biofilm pathway for S. epidermidis and E. coli respectively, are predicted to have carbohydrate deacetylase activity. These proteins are homologous to other enzymes classified in the Carbohydrate Esterase Family 4 in the CAZy database (www.cazy.org). Well studied enzymes within this family have de-N- or de-O-acetylation activity on the carbohydrate polymers xylan or chitin, or peptidoglycan residues, and commonly exhibit metal-dependent activity.120 Based on crystallographic data acquired for PgaB, and comparison to other known CE4 enzymes, a catalytic mechanism for PgaB has been hypothesized utilizing a metal center to achieve de-N- acetylation of PNAG to produce dPNAG (Figure 3.1).64-65

Figure 3.1 A) Hypothesized mechanism of PgaB based on homology to SpPgdA, a mechanistically-studied CE4 enzyme. B) Reaction performed on PNAG polymers by PgaB. C) X-ray diffracted active site of PgaB containing a nickel metal center and catalytically-important residues.

Due to the necessity of PNAG de-N-acetylation for efficient bacterial biofilm formation in E. coli and S. epidermidis, these enzymes make attractive drug targets. To date, no inhibitor has been identified for these enzymes. In this chapter, the synthesis of potential inhibitors against these proteins is presented. These inhibitors are based on a metal-chelating scaffold, and on transition state analogs. Finally, the development of a pentasaccharide inhibitor synthesized using a chemoenzymatic approach will be described.

3.1.1 Inhibitor Design

3.1.1.1 Metal Chelating Inhibitors

Metalloenzymes catalyze a variety of hydrolytic reactions on many different substrates in important metabolic pathways. In particular, deacetylation is an example of a reaction commonly catalyzed by zinc-dependent enzymes.121 The biological importance of this metal center has made these enzymes an attractive pharmaceutical target for the development of inhibitors. Generally, these enzymes adopt a mechanism where metal-bound water acts as a nucleophile and stabilization of the resulting tetrahedral intermediate is also provided by the active site metal.

Increased understanding of these enzymes and their mechanistic pathways has led to the design of different types of inhibitors.

One class of metallohydrolases inhibitor includes compounds that specifically bind the metal center, with the most common subclass being coined ―zinc binding groups‖ (ZBG). Notable examples include inhibitors targeting matrix metalloproteases (MMPs) a family of zinc dependent endopeptidases,122 inhibitors targeting histone deacetylases (HDACs),123 and glycosyl deacetylase inhibitors, specifically, LpxC inhibitors.124 A general principle for metalloenzyme drug design is that the metal binding group is responsible for binding energy, whereas the other functional groups on the small molecule provide binding selectivity.125

3.1.1.1.1 Matrix Metalloprotease Inhibitors

Great efforts have gone into developing MMP inhibitors with anti-cancer activity.122a A common feature found in the most effective inhibitors was a metal chelating moiety (Figure 3.2). These metal chelating moieties can be mono- or bi-dentate in character. Due to the sheer number of homologous MMP enzymes (>20), inhibitor discovery has been performed via broad spectrum library screening on all MMP enzymes to ensure target selectivity.122c In one example, it was computationally determined that in a small molecule selective for MMP-9 binding the hydroxamic acid moiety accounted for 96% of the binding energy. 126

Figure 3.2 MMP-9 inhibitor examples that have undergone clinical trials for cancer treatment. Metal chelating hydroxamic acid group is denoted in red.

3.1.1.1.2 Histone Deacetylase Inhibitors

Histone deacetylases (HDACs) have been a popular target for scientists for many years as anti-cancer drug targets. Breslow et al. were the first to discover the potent HDAC inhibitor suberoylanilide hydroxamic acid (SAHA), marketed in the United States and Canada as

Vorinostat for the treatment of T cell lymphoma (Figure 3.3).123a The mode of action of this small molecule entails binding of the HDAC Zn2+ metal center in a bidentate fashion through the hydroxamic acid and the aliphatic chain interacts with a hydrophobic pocket. Work by Miyata et al. explored the binding of SAHA analogs to HDAC8, where each analog contained a substituted metal binding domain (Table 3.1).123b This study provided great insight into the relative binding capacities of various common metal binding moieties for Zn2+.

Functional Functional IC50 (µM) IC50 (µM) Group (R) Group (R)

-CONHOH 0.28 -NHCOCH NH >100 (SAHA) 2 2

-NHCONHOH 80 -NHCOCH2SH 0.39

-NHCONHNH2 150 -NHSO2Me 7500

-NHCOCH OH >100 -SO Me 230 2 2

Table 3.1 Relative inhibition activities of various SAHA analogs.

Figure 3.3 X-ray crystallography structure of SAHA bound to a zinc ion inside the HDAC active site.127

3.1.1.1.3 Glycosyl Deacetylase Inhibitors

LpxC, a zinc-dependent lipopolysaccharide (LPS) de-N-acetylase, provides an attractive target for drug design as a potential antibiotic against Gram-negative bacteria.124a LPS is a potent endotoxin found in the outer membrane of Gram-negative bacteria consisting of 3 parts: lipid A, the core oligosaccharide, and the O antigen, with the first two parts considered to be highly conserved among bacteria (Figure 3.4).126, 128 Early in the LPS biosynthesis, the core N-acetyl glucosamine residue is de-N-acetylated by LpxC, which in turn is re-acylated with a fatty chain before subsequent functionalization and polysaccharide elongation. Inhibition of LpxC leads to the formation of attenuated LPS, which subsequently lead to bacteria with compromised viability.129 Work by Hindsgaul et al. demonstrated the synthesis of an inhibitor with a carbohydrate scaffold and a metal-chelating hydroxamic acid moiety.124a This compound demonstrated 50% inhibition of LpxC at 1 ug/mL.124b Although LpxC belongs to a different carbohydrate esterase family than PgaB and IcaB (CE11 vs CE4), LpxC shares some similarity with PgaB and IcaB in that it acts on the same GlcNAc scaffold and is metal dependant.

Figure 3.4 Left – Biosynthesis of Lipid A and the role LpxC plays. Right – LpxC inhibitor developed by Hindsgaul et al.124a, 125

More recently, Urbanaik et al. have developed a library of carbohydrate-based metal binding inhibitors targeting glycosylphosphatidylinositol (GPI) de-N-acetylases, a causative agent in African sleeping sickness, based on the previously mentioned LpxC inhibitor (Figure 130 130c 3.5). These compounds showed IC50 values ranging from micromolar to molar.

Figure 3.5 Left – GPI de-N-acetylase catalyzed reaction. Right – inhibitors synthesized and tested with GPI de-N- 130a acetylase. Metal chelating regions in red. Values below inhibitors represent IC50 values.

3.1.1.2 Transition State Mimics

Transition state analogs are commonly employed to target enzymes. These analogs mimic a substrate‘s transition state during the enzymatic reaction. Since the transition state is the highest energy state during the reaction course, enzymes stabilize this state in order to lower the energy barrier of the chemical reaction. As a result, the transition state is the tightest bound enzyme complex over the reaction course (Figure 3.6).131 Therefore, molecules that mimic the transition state of an enzyme-catalyzed reaction are ideal inhibitor candidates. A number of transition state analog inhibitors have also been developed for various metallodeacetylase enzymes and will be discussed in detail.

Figure 3.6 Simplified reaction coordinate energy diagram of an enzyme-catalyzed reaction and an uncatalyzed reaction.

3.1.1.2.1 Phosphonamidate Inhibitors

Phosphonamidates have long been employed in medicinal chemistry as transition state inhibitors for de-N-acetylase enzymes. Phosphorous-containing compounds are found to be amongst the most potent inhibitors of metalloproteases. The phosphorous center is tetrahedral with heteroatoms approximately positioned to mimic the tetrahedral transition state of amide

44 bond hydrolysis (Figure 3.7). Additionally, the phosphorous-heteroatom bonds are longer than ground state carbon-heteroatom bonds and are thus capable of mimicking the longer bonds in the transition state of the reaction with the native substrate.

Figure 3.7 Left – Transition state proposed for HDAC1. Right – Model for binding of a methyl phosphonamidate analog of SAHA.

NagA is a metal dependent enzyme that catalyzes the deacetylation of N-acetyl-D- glucosamine-6-phosphate to form D-glucosamine-6-phosphate (Figure 3.8), similar to the action of IcaB and PgaB. This enzyme is found in E. coli and homologs have also been identified in 300 different bacterial strains.132 Its overall function is involved in the process of catabolizing N- acetyl-D-glucosamine from chitobiose and recycling the peptidoglycan, an essential component of the bacterial cell wall, thus making it an attractive drug target.133 In E. coli, this enzyme contains a zinc ion in its active site coordinated by a glutamic acid residue and two histidine residues in a tetrahedral conformation.132

Figure 3.8 Top left - Reaction catalyzed by NagA. Top right - Transition state analog designed to inhibit NagA. Bottom – X-ray diffracted active site of NagA containing a zinc metal center and bound phosphonamidate transition state analog inhibitor.132

Recently, work by Raushel et al involved development of a carbohydrate-based phosphonamidate to act as an inhibitor towards the metallo de-N-acetylase NagA (Figure 3.8).134 This inhibitor mimicked the substrate exactly with the exception of a methyl phosphonamidate moiety in place of the N-acetyl group. In vitro kinetic analysis of recombiant NagA showed it to be a moderately fast-acting enzyme when introduced to its natural N-acetyl-D-glucosamine-6- 5 -1 -1 phosphate substrate (kcat/Km of 5.7 x 10 M s ). Introduction of the glycosyl phosphonamidate showed the molecule to be a remarkably strong competitive inhibitor, with a Ki value of 34 nM.

The authors also pointed out that this Ki value is among the lowest ever measured for the inhibition of a deacetylase.134

3.1.1.2.2 Fluoro Ketone Inhibitors

The incorporation of fluorine can result in profound changes in the physical and chemical properties of organic molecules. These changes can also affect the biological activity of the molecules.135 Trifluoromethyl groups in particular are exceptionally inductively electron withdrawing, boasting an inductive sigma constant (σI) of 0.43 (Table 3.2). When placed alpha to a ketone, it has the implication of drastically altering the equilibrium between the ketone and the corresponding geminal diol. In the case of acetone versus hexafluoroacetone, a 109-fold shift in the equilibrium constant towards the geminal diol in aqueous media was observed (Figure 3.9).136

Inductive Sigma Inductive Sigma Substituent Substituent Value (σI) Value (σI)

H 0 CF3 0.43

CH3 -0.04 Cl 0.47

C6H5 0.10 NO2 0.76

+ OH 0.29 N(CH3)3 0.93

Table 3.2 Inductive sigma values for various organic substituents.136 Positive values represent electron-withdrawing power relative to hydrogen.

Figure 3.9 (A) Equilibrium of ketones and geminal diols in aqueous solution. (B) Comparison of equilibrium constants between acetone and hexafluoroacetone to their corresponding geminal diols. In this case, a 109-fold difference was observed.

The ability of trifluoromethyl-ketone moieties to adopt the geminal diol conformation is an important measure of their role as potential metallohydrolase inhibitors.137 This conformation best resembles the tetrahedral intermediate of an acyl hydrolysis reaction. A crystal structure of carboxypeptidase A and a trimethylfluoro-ketone inhibitor shows the inhibitor bound to the active site in its geminal diol form. A SAHA mimic replacing the hydroxamadate group with a trifluoromethyl ketone moiety has also demonstrated comparable inhibition efficacies, exhibiting 138 an IC50 value of 6.7 µM (Figure 3.10).

Figure 3.10 Left – Ketone binds to the hydrolase metal center in its geminal diol form. Right – SAHA analog containing a trifluoromethyl moiety, exhibiting a IC50 value of 6.7 µM.

The effectiveness of fluoro ketone inhibitors is best observed when compared to similar non-fluorinated derivatives. A study using fluoro ketones as potential enzyme inhibitors was performed by Abeles et al.135 Working with angiotensin converting enzyme, a zinc metalloprotease, they demonstrated a 3000-fold improvement in inhibition capability of a trifluoro methyl ketone when compared to a simple methyl ketone (Figure 3.11). A similar 1000- fold improvement was observed when making similar modifications to a ketone inhibitor for pepsin, an aspartate protease demonstrating that fluoro ketone inhibitors are not metal dependent.

Figure 3.11 Inhibitors of angiotensin converting enzyme, a zinc protease.

3.2 Results and Discussion

3.2.1 Monosaccharide Inhibitors

3.2.1.1 Acyl and Sulfonyl Inhibitors

Many metal-chelating inhibitors have been designed for metal-dependent N- acetylamidases, such as histone de-N-acetylases and other carbohydrate processing de-N- acetylases such as Lipopolysaccharide de-N-acetylase LpxC. A library of inhibitors with ranging

IC50 values (100 – 500 uM) has also been identified for a Streptococcal peptidoglycan de-N- acetylase spPgdA from the CE4 family via a large in silico library screen. A common feature of these de-N-acetylamidase inhibitors has been a metal-coordinating functional group, capable of binding the active site metal. Common metal coordinating functional groups include free thiols and hydroxamic acid moieties, the latter being used as a successful inhibitor for LpxC, under the name TU-514 (Figure 3.12).124a, b, 125 Due to these past results, part of the library of potential PgaB/IcaB inhibitors was based on a glucosamine scaffold with an installed metal coordinating group in the acetamide position.139 Additionally, methylation of the amido nitrogen was investigated, as this would potentially allow for different orientations of the metal chelating groups, and has been shown to improve inhibitor efficacy for Zn metalloproteases, which have been proposed to act in a similar mechanism to PgaB.

Figure 3.12 A) TU-514, a proposed metal-chelating lipopolysaccharide de-N-acetylase inhibitor for LpxC. B)

Proposed metal-chelating inhibitor structures for IcaB and PgaB, where R = H or Me, Y = CH2NH2, CH2OH,

CH2SH, NHOH.

The acyl and sulfonyl inhibitor array was initiated from the starting scaffold molecule methyl β-D-2-deoxy-2-aminoglucopyranoside, which could be obtained in three steps from the

49 commercially available N-acetylglucosamine (Scheme 3.1). Using protection-group-free chemistry, condensation of N-acetylglucosamine with a small excess of tosyl hydrazide in DMF, followed by precipitation with diethyl ether resulted in a multigram scale production of 13.140 Oxidation of the glycosyl hydrazide donor with NBS in the presence of methanol yielded the methyl glycoside 14, which could then be subsequently de-N-acetylated in the presence of hydrazine monohydrate to quantitatively yield the divergent starting material 15 upon acidic workup.141 Conversely, 15 could also be obtained in 3 steps from glucosamine hydrochloride; initial acetylation and bromination of the carbohydrate by acetyl bromide yielded the 3,4,6- triacetyl-2-deoxy-2-aminoglucosyl bromide 16.142 Direct methanolysis of 16 in the presence of pyridine, followed by subsequent de-O-acetylation in acidic conditions also yielded the divergent starting material 15, in large scale.

o Scheme 3.1 A) (a) 1.1 eq Tosyl Hydrazide, cat AcOH. 5:1 DMF/H2O, 37 C, 24 h, 92%. (b) 2.2 eq NBS, 20 eq o MeOH, DMF, rt, 30 min, 72%. (c) NH2NH2·H2O (neat), 110 C, 48 h, 97% yield. B) (a) AcBr (neat), rt, 48 h, (b) 10 eq MeOH, py, rt, 3 h. (c) HCl · MeOH, 2 h, rt, 68% yield over 3 steps.

Upon synthesis of the divergent starting material, the free amine of 15 could then be acetylated to yield a variety of potential inhibitors (Scheme 3.2). HBTU-assisted condensation of 15 with benzyloxyacetic acid or Cbz-Gly-OH followed by hydrogenation over Pd/C yielded the hydroxyl and amino inhibitors 19 and 21 respectively. Conversely, condensation with pentafluorophenyl S-acetylthioglycolate (SAMA-OPFP) followed by alkaline deprotection under Zemplén conditions yielded the thiol inhibitor 23. The hydroxylurea inhibitor 24 and the sulfonylate inhibitor 25 could also be accessed by acetylation of 15 with 1-(4-nitrophenol)-N- hyxroxycarbamate or with mesyl chloride respectively.143

Scheme 3.2 (a) 1 eq HBTU, 1eq benzyloxyacetic acid , 1.2 eq TEA, DMF, rt, 12 h, leading to 18, 84%. (b) 1 eq HBTU, 1eq CBZ-Gly-OH , 1.2 eq TEA, DMF, rt, 12 h, leading to 20, 76%. (c ) 1 eq SAMA-OPFP, 1.2 eq TEA, DMF, rt, 12 h, leading to 22, 78%. (d) 1 eq 1-(4-Nitrophenol)-N-hydroxycarbamate, 1.2 eq TEA, DMF, rt, 12 h, leading to 24, 54%. (e) 1 eq MsCl, 1.2 eq TEA, DMF, rt, 12 h, leading to 25, 58%. (f) 10% Pd/C, 1 atm H2, MeOH, rt, 12 h, 89%. (g) 10% Pd/C, 1 atm H2, MeOH, rt, 12 h, 81%. (h) 0.1 eq NaOMe, MeOH, rt, 10 min, 92%.

Synthesis of inhibitors with a methylated amide required the production of a divergent monosaccharide containing a secondary methylated amine. To this end, benzyl protected methyl 2-aminomethyl-2-deoxy-β-D-glucoside was synthesized in four steps from 14 (Scheme 3.3).144 Initial per-O-benzylation of 15 with benzyl bromide followed by N-carbamylation of the amide with Boc2O in the presence of catalytic DMAP yielded the mixed imide 27. De-N-acylation of the mixed imide with NaOMe resulted in the expulsion of acetate, yielding the carbamate monosaccharide 28. Reduction with LiAlH4 yielded the secondary methylamine 29 in good yield, which could then be divergently functionalized to produce the secondary methyl amide inhibitors.

Scheme 3.3 (a) 3.5 eq BnBr, 3.5 eq NaH, DMF, 0 °C, 16 h, 76%. (b) 2 eq Boc2O, 0.25 eq DMAP, THF, 60 °C, 12 h. (c) 0.2 eq NaOMe, MeOH, rt, 1 h, 72% over 2 steps. (d) 3 eq LiAlH4, THF, 60 °C, 6 h, 68%.

Synthesis of the methylated inhibitors proceeded from compound 29 in a similar fashion to the previously described non-methylated inhibitors (Scheme 3.4). DCC-assisted condensation of 29 with benzyloxyacetic acid or Cbz-Gly-OH followed by global deprotection employing hydrogenation over Pd/C yielded the hydroxyl and amino inhibitors 35 and 36, respectively. Similarly, the hydroxylurea and sulfamate inhibitors could be accessed by condensation of the amine with 1-(4-nitrophenol)-N-hyxroxycarbamate or mesyl chloride followed by global deprotection employing hydrogenation over Pd/C to yield compounds 37 and 38, respectively. Finally, condensation of the secondary amine with the sulfamylating agent benzyl N- (chlorosulfonyl)carbamate (BNCC) yielded the sulfamate inhibitor 39 after global deprotection of the benzyl groups via hydrogenation over Pd/C.145

Scheme 3.4 (a) 1eq DCC, 1 eq benzyloxyacetic acid, TEA, EtOAc:DCM (1:1), rt, 12 h, leading to 30, 89%. (b) 1eq DCC, 1 eq Cbz-Gly-OH, TEA, EtOAc:DCM (1:1), rt, 12 h, leading to 31, 92%. (c) 1 eq 1-(4-Nitrophenol)-N- hydroxycarbamate, TEA, DMF, rt, 12 h, leading to 32, 72%. (d) 1 eq MsCl, TEA, EtOAc, rt, 12 h, leading to 33,

88%. (e) 1.2 eq BNCC, 1.2 eq TEA, DCM, rt, 3 h, leading to 34, 88%. (f) 10% Pd/C, 1 atm H2, MeOH, rt, 12 h, 75– 82%. Lastly, modified chemical schemes were required for the synthesis of the sulfamate derivative and the methylthioglycolyl amide derivative, due to their inherent incompatibility with the previously described schemes (Scheme 3.5). Owing to the reactivity of the sulfamylating agent BNCC, hydroxyl protecting groups were required. To this end, 28 was treated with TFA to afford the benzyl-protected glucosamine derivative 40, which could then be condensed with BNCC followed by subsequent global deprotection by hydrogenation over Pd/C to yield the sulfamate derivative 42. Conversely, due to the incompatibility of sulfur with hydrogenation conditions, an alternative synthesis for the methylthioglycolyl amide derivative was as follows; the methyl glucosamine derivative 29 was initially debenzylated by hydrogenation over Pd/C to yield the unprotected glucosamine 43, which could then be acylated using SAMA-OPFP after which subsequent base-catalyzed hydrolysis of the thioester would yield the methylthioglycolyl amide inhibitor 45 as a symmetric disulfide.

Scheme 3.5 A) (a) 1:1 TFA:DCM, rt, 30 min, 81%. (b) 1.5 eq BNCC, 1.2 eq TEA, DCM, rt, 3 h, 88%. (c) 10%

Pd/C, 1 atm H2, MeOH, 12 h, 75%. B) (a) 10% Pd/C, 1 atm H2, MeOH, 12 h, 78%. (b) 1 eq SAMA-OPFP, 1.2 eq TEA, DMF, rt, 12 h, 69%. (c) 0.1 eq NaOMe, MeOH, rt, 10 min, 64%.

3.2.1.1.1 Crystal Structure of 35

Compound 19 was dissolved in a minimum of methanol, whereupon ethyl acetate was slowly added to that solution, resulting in a transition from clear and colourless solution to slightly cloudy. Sonication of the mixture for 10 minutes yielded a separation of the sugar from the solution as a slightly yellow oil. The reaction mixture was then begrudgingly left open to air for 60 hours, after which colourless, needle-shaped crystals (2 – 4 cm in length) grew in place of the oil. The crystals were then filtered, dried, and submitted to X-ray crystallography analysis (Figure 3.13).

Figure 3.13 X-ray crystal structure of 19. Hydrogen bond denoted with dashed line.

3.2.1.2 N-Alkyl Inhibitor Synthesis

Starting from the glucosamine scaffold 15, reductive amination with glyoxylic acid in the presence of refluxing formic acid through a transfer hydrogenation mechanism led to the formation of the N-alkyl-N-formyl carbohydrate 46 (Scheme 3.6).146 Several methods were attempted for the deformylation of compound 46, including basic Zemplén conditions (MeONa/MeOH) and methanolic HCl, however these methods proved unsuccessful. The reported route of aqueous 2M HCl was unavailable to us due to the potential hydrolysis of the glycosidic linkage. It was finally determined that hydrazine hydrate at 50 ºC afforded the final inhibitor 47 in modest yield after recrystallization.

o Scheme 3.6 a) 4 eq glycoxylic acid, formic acid, reflux, 4 h, 62%. B) NH2NH2·xH2O, 50 C, 12 h, 52 %.

3.2.1.3 Phosphonamidate Inhibitor Synthesis

Synthesis of the glycosyl phosphonamidate inhibitor was adapted from a procedure reported by Xu et al.134 Starting from glucosamine hydrochloride, initial per-O-acetylation of the carbohydrate by acetyl bromide yielded the 2-deoxy-2-amino-1,3,4,6-tetra-O-acetyl-β-D- glucopyranosyl hydrochloride 49 (Scheme 3.7).147 Phosphonylation of the amine was then accomplished by using methyl methylchlorophosphonate in chloroform in the presence of TEA under reflux conditions to obtain 50 in moderate yield. Introduction of the compound to Lewis base-activated glycosidation conditions resulted in the thiobenzyl glycoside 51. Finally, global hydrolysis of the acetyl groups and the methyl phosphonyl ester with aqueous ammonia in methanol resulted in the final phosphonamidate inhibitor 52.

Scheme 3.7 (a) Methyl methylchlorophosphonate, CHCl3,TEA, reflux, 5 h, 48%. (b) BnSH, BF3·OEt2, DCM, rt,

63%. (c) NH3(aq), MeOH/H2O, rt, 4 h, 98%.

3.2.1.4 2-C Carbohydrate Inhibitors

Synthesis of the 2-C inhibitor array was accomplished from the scaffold molecule 52a, which could be synthesized in two steps from the commercially available D-Glucal (Scheme 3.8). 148 Initially, per-O-benzylation of D-Glucal with benzyl bromide and sodium hydride resulted in the benzylated glucal in good yield. This was then followed by the radical addition of dimethyl malonate to the double bond using cerium ammonium nitrate as a radical initiator in the presence of methanol and sodium bicarbonate to yield a diastereomeric mixture of the gluco product 52a and the manno product 52b in 65% and 10% yield respectively, according to crude 1H NMR. Fortunately, the gluco diastereomer was easily isolable by flash column chromatography.

Scheme 3.8 (a) 4.5 eq BnBr, 3.3 eq NaH, DMF, 0oC – rt, 12 h, 74%. (b) 2 eq cerium ammonium nitrate, 10 eq dimethyl malonate, NaHCO3, MeOH, rt, 3 h, 65% for the gluco-product, 10% for the manno-product (relative yields obtained by crude 1H NMR).

With compound 52a in hand, synthesis of 2-C inhibitors containing a bis-binding moiety could be synthesized (Scheme 3.9). Initial base catalyzed hydrolysis of the methyl esters with 2 M sodium hydroxide in dioxane yielded the bis-carboxylic acid glycoside 53 in excellent yield. This was followed by a global debenzylation by hydrogenation over Pd/C to yield the final inhibitor 54 in moderate yield.

Scheme 3.9 (a) 1:1 2M NaOH/Dioxane, 12 h, rt, 86%. (b) 10% Pd/C, 1 atm H2, MeOH, 12 h, rt, 77% for top, 72% for bottom. (c) 5 eq TMS-CF3, cat CsF, DME, rt, 12 h, 88%.

Transformation of the methyl esters to a trifluoromethyl ketone was accomplished using a method developed by Shreeve et al.149 Briefly, the glycosyl ester was dissolved in ethylene glycol dimethyl ether along with the trifluroromethylating agent TMS-CF3 and the catalytic initiator CsF, resulting in excellent yield of the bis-trifluoromethyl ketone product 55. Following

57 this step, global debenzylation of the carbohydrate by hydrogenation over Pd/C yielded the final inhibitor 56 in moderate yield.

Finally, in order to attain a mono-functionalized inhibitor, an additional step was necessary, involving microwave-assisted decarboxylation of one of the methyl esters (Scheme 3.10).148 This was accomplished by dissolving the bis-methyl ester 52a in DMSO in the presence of an excess of lithium iodide, and submitted to microwave irradiation (200 W, 10 bar, 100 oC, 20 min) to yield the mono ester product 57 in decent yield. This step was also accomplished by using the same reagents, but refluxing at 180 ºC for 5 hours. The reflux method, however, resulted in a lower yield. This product could then be subjected to similar reaction conditions as described above to obtain the monocarboxylic acid inhibitor 59 and the mono- trifluoromethyl inhibitor 61.

Scheme 3.10 (a) 3 eq LiI, microwave irradiation (200 watts, 10 bar, 100 oC, 20 min), DMSO, 62% (b) 1:1 2M

NaOH/Dioxane, 12 h, rt, 82%. (c) 5 eq TMS-CF3, cat CsF, DME, rt, 12 h, 84%. (d) 10% Pd/C, 1 atm H2, MeOH, 12 h, rt,, 72% for top, 81% for bottom

3.2.2 Chemoenzymatic Synthesis of a Polysaccharide Inhibitor

Previous work has shown PgaB to have no measurable de-N-acetylaiton activity on GlcNAc or any monosaccharide derivative. However, increased catalytic activity has been observed against larger PNAG substrates up to pentasaccharide in length.64 Thus, the synthesis of a larger oligosaccharide inhibitor was envisioned. An added advantage of a chemo-enzymatic approach is that PgaB de-N-acetylation of the pentasaccharide provides optimal regiochemistry of the metal binding group (Scheme 3.11).

Scheme 3.11 (a) HF·Pyridine (neat), rt, 5 days. 40 mg per gram of GlcNAc used after HPLC size exclusion purification. (b) PgaB, 50 mM HEPES, pH 7.5, 37 oC, 24 h. (c)10 eq N-hydroxysuccinyl 2-(octanoylthio)acetate,

1:1 H2O:MeOH, rt, 2 h. (d) 0.1 eq NaOMe, MeOH, rt, 1 h.

PNAG pentasaccharide was obtained through a one-step procedure involving the dissolution of GlcNAc in a HF·pyridine solution (a procedure modified from work done by Defaye et al) to produce a mixture of oligomers from length two to eight.72, 150 Size exclusion

60 chromatography was then used to isolate substrates of uniform lengths. Purified PNAG pentasaccharide was subjected to PgaB-catalyzed de-N-acetylation, resulting in an inseparable mixture of PNAG and mono-de-N-acetylated dPNAG (PgaB expression and methodology is presented in Chapter 4). The mixture was then carried forward to an amine acylation step with activated S-octanoylthioglycolate-N-hydroxysuccinamide ester to provide a product that could be separated from the starting material PNAG pentasaccharide using RP-HPLC. Base-catalyzed hydrolysis of the octanoyl thioester resulted in the pentasaccharide inhibitor 64 with a thioglycolamide metal binding moiety (Figure 3.14).

Figure 3.14 MALDI Mass spectra of compounds 63 (top) and 64 (bottom)

3.3 Conclusions

In this chapter, the synthesis of an array of inhibitors was presented. These inhibitors include compounds designed to be metal-chelating in their mode of inhibition, as well as inhibitors representing transition state analogs. This array represents a diverse library of functional groups and inhibitor styles, improving the chances of a successful hit. The development of a pentasaccharide inhibitor through a chemoenzymatic approach was also

62 described. Since both IcaB and PgaB have shown size dependence for PNAG substrate length in terms of de-N-acetylation activity, it is hypothesized that a pentasaccharide inhibitor would demonstrate an increased binding affinity.

4 Assay Development and Inhibitor Evaluation with PgaB and IcaB

Development of the PgaB expression methodology was done by Dr. Joanna Poloczek. Expression and testing of inhibitors with IcaB was performed by Dr. Varvara Pokrovskaya. This chapter has been reproduced in part with full permissions from:

Chibba, A.; Poloczek, J.; Little, D.J.; Howell, P.L.; Nitz, M. (2012) Synthesis and evaluation of inhibitors of E. coli PgaB, a polysaccharide de-N-acetylase involved in biofilm activity. Organic and Bimolecular Chemistry, 10, 7103.

4.1 Introduction

Based on sequence homology, the N-terminal domain of PgaB and IcaB have predicted carbohydrate deacetylase functionality similar to enzymes classified in the Carbohydrate Esterase Family 4 (CE4) family of enzymes in the CAZy database. This class of enzymes consists of metal dependant hydrolases containing a His-His-Asp metal binding triad, along with other conserved catalytic residues. The recently solved structure of the N-terminal domain of PgaB has confirmed that the enzyme is structurally homologous to other well-characterized CE4 family enzymes like the chitin de-N-acetylase ClCDA from Colletotrichim lindemuthianum and the peptidoglycan de-N-acetylase SpPgdA from Streptococcus pneumonia.

Previous work in our lab was done to characterize these enzymes and their substrate activity in vitro. Several factors have been shown to play a role in de-N-acetylase activity for PgaB and IcaB (Figure 4.1). The activity of both of these enzymes was shown to be dependent on the length of the PNAG substrate length, with PgaB activity peaking with pentasaccharide substrate, whereas IcaB showed maximal activity with trisaccharide. Carbohydrate polymer linkage also affected activity, with both enzymes showing no de-N-acetylase activity on short polymers of chitin, which is comprised of β-1,4-linked N-acetylglucosamine units, despite their similarity to the chitin de-N-acetylase ClCDA. Finally, both enzymes demonstrated varied metal dependence. PgaB showed optimal activity with, Co2+, Fe2+and Ni2+, in contrast to Zn2+ dependency observed in other CE4 deacetylases. IcaB, on the other hand demonstrated optimal activity with Co2+ and Zn2+, as has been observed with other CE4 enzymes.

Figure 4.1 A) PgaB activity with varying substrate lengths. B) PgaB activity with PNAG tetrasaccharide and a chitin tetrasaccharide. C) PgaB activity in the presence of various metals or small molecule metal chelaters.64

A significant challenge in monitoring IcaB and PgaB activity is the low activity of the enzymes necessitating long incubations. Using PNAG pentasaccharides, activity could be measured using a discontinuous fluorescamine-based assay to probe for the production of free amines. Despite the optimization of reaction conditions for both enzymes, they have still demonstrated extremely slow kinetic parameters compared to other well-studied CE4 enzymes (Table 4.1). As a result, meaningful assay results required large amounts of substrate, and had to be monitored discontinuously over a timescale of hours to days. However, an assay capable of

65 evaluating deacetylase activity quickly and continuously is required for efficient inhibitor evaluation.

-1 -1 Enzyme Substrate kcat/Km (M s )

IcaB PNAG pentasaccharide 0.0367

PgaB PNAG pentasaccharide 0.2664

spPgdA Chitin trisaccharide 150124c

ClCDA Chitin pentasaccharide 89000151

Table 4.1 Kinetic parameters for a selection of CE4 enzymes.

To this end, new methodology was explored for monitoring PgaB and IcaB activity using synthetic substrates. Once a suitable method was established, the efficacy of potential PgaB and IcaB inhibitors was explored.

4.2 Results and Discussion

4.2.1 Enzyme Purification

4.2.1.1 PgaB

An MBP-PgaB(22-672) construct in a pIADL plasmid (prepared by Joanna Poloczek) was expressed in E. coli BL21(DE3) cells. A 1 L culture was grown at 37 ºC in the presence of kanamycin (100 mg/L) to an OD600 of 0.3. Nickel (II) sulfate (0.26 g, 100 mmol) was then added to the culture and the temperature was lowered to 10 ºC. At this point, the cells were induced with isopropyl 1-thio-β-D-galactopyranoside (0.123 g, 0.5 mmol) for 16 hours. After centrifugal pelleting (3750 x g, 70 min) of the culture, the cells were then resuspended in 10 mL of buffer containing HEPES (50 mM, pH 7.5), NaCl (300 mM), and a Complete Mini protease inhibitor cocktail tablet (Roche) at a final concentration of 2 ml per gram of pellet weight. Lysing of the cells was achieved by sonication of the suspension on ice, followed by centrifugal pelleting of the cellular debris (16 000 x g, 1 hour). The resulting soluble fraction was then passed through an amylose resin (Sigma-Aldrich) column. MBP-PgaB was then eluted with buffer containing

HEPES (50 mM, pH 7.5). The fractions containing MBP-PgaB were concentrated using centrifugal filter units (Millepore, 30K MWCO) and stored at 4oC. The enzyme was stable and active for approximately 1 month under these conditions. Protein concentrations were measured using a predicted molar extinction coefficient of 218 070 M-1. Purity of the fractions could also be assessed by sodium dodecyl sulfate−polyacrylamide gel electrophoresis (SDS-PAGE).64

4.2.1.2 IcaB

An IcaBHis10 construct in a UT032 plasmid was expressed in E.coli BL21(DE3) cells. A 1

L culture was grown at 37 °C in LB media with 100 mg/L ampicillin to an OD600 of 0.5. The cells were then cooled to 10 °C, and allowed to grow until an OD600 of 0.7−0.9 was reached. At this point, the cells were induced with isopropyl β-D-thiogalactoside (0.185 g, 0.75 mmol). After 24 h at 10 °C, the cells were centrifuged (3430 x g, 40 min) and the cell pellet was suspended in 20 mL of buffer containing sodium phosphate (20 mM, pH 7) NaCl (1 M) and glycerol (5% (v/v)) (buffer A). Lysing of the cells was accomplished by sonication of the resulting suspension on ice. The resulting crude extract was centrifuged to pellet and remove cell debris (16260 x g, 40 min). The soluble fraction was then gently mixed with 0.5 mL of Ni(II)-NTA resin (QIAGEN) for 1 hr before being loaded into a column (1 cm diameter). Unbound proteins were washed from the resin with 10 mL of buffer A containing 20 mM imidazole. IcaBHis10 was then eluted with buffer A containing 150 mM imidazole. All purification steps were performed at 4 °C. The purified protein was dialyzed overnight against buffer A and subsequently washed with buffer containing sodium phosphate (50 mM, pH 8) and NaCl (0.5 M) (buffer B) by centrifugation in an Amicon Ultra-15 10 000 Da MWCO centrifugal filter (Millipore), and concentrated to 3 mg/mL and stored at 4 ºC in buffer B. The purity of the protein as analyzed by 15% SDS-PAGE was >95%. Protein mass was confirmed by electrospray ionization mass spectrometry (ESI-MS). The protein concentrations were determined using a molar extinction coefficient of 39 400 M−1 cm−1 calculated from a BCA assay on purified protein. The purified

IcaBHis10 was digested with Factor Xa protease (New England Biolabs, Ipswich, MA) according to the recommended protocol to remove the His10 tag. Briefly, 1 mg of IcaBHis10 was incubated with 9 μg of Factor Xa at room temperature for 4 h in Factor Xa reaction buffer (75 mM HEPES, pH 6.5, 75 mM NaCl, 100 mM Na2SO4, 2 mM CaCl2). After protease digestion, Factor Xa protease was removed from the reaction mixture by affinity chromatography using Xarrest Resin. Xarrest resin slurry (100 μL, pre-equilibrated with Factor Xa reaction buffer) was added to the

67 cleavage reaction. The resin was resuspended by gentle mixing and incubated for 10 min at room temperature. The mixture was then centrifuged (1000 x g, 5 min) and the soluble fraction collected. In order to remove cleaved His-tag peptide, the sample was washed 4 times on a 10 000 Da MWCO centrifugal filter with buffer B, resulting in purified IcaB enzyme. The effectiveness of the digestion could be analysed by SDS-PAGE.67

4.2.2 Assay Development

Previous work on measuring PgaB and IcaB activity involved using PNAG oligomers as substrates and indirectly measuring free amine production with the aid of fluorescamine (Figure 4.2).64, 67, 152. Several unresolved challenges with this method included the low catalytic -1 -1 efficiency of de-N-acetylation on PNAG oligomers (kcat/Km = 0.25 M s for PNAG pentasaccharide with PgaB), the inefficient methods of obtaining purified PNAG pentasaccharide, which featured a 5 day reaction time and 4 size exclusion chromatography steps for purification, and the low solubility of PNAG pentasaccharides (1 mM in aqueous solution), which, in turn, prevented determination of isolated Km or kcat values. In addition, monitoring the de-N-acetylation reaction of the pentasaccharide substrate required a cumbersome discontinuous fluorescamine assay.

Figure 4.2 A) Reaction of PgaB with PNAG oligomers. B) Reaction of fluorescamine with an amine to yield a highly fluorescent product. R in this case represents dPNAG. C) Competing hydrolysis reaction of fluorescamine yielding a non-fluorescent by-product.

4.2.2.1 Synthesis of Chromogenic Enzyme Substrates

A series of chromogenic acetyl esters were synthesized or purchased, and evaluated as PgaB substrates (Figure 4.3). Such esters have been used to study other de-O- and de-N-acetylase enzymes.153

Figure 4.3 Acetyl esters evaluated as substrates for PgaB under continuous assay conditions.

5-Acetoxymethylquinolinium (5-AMQ) and 7-acetoxymethylquinolinium (7-AMQ) were easily and efficiently synthesized in 2 steps from 5-quinolinol and 7 quinolinol respectively (Scheme 4.1). 3-Carboxyumbelliferyl acetate (ACC) was synthesized in two steps: initial condensation of Meldrum‘s acid and 2,4-dihydroxybenzaldehyde in ethanol to give 3- carboxylumbelliferyl, followed by acetylation of the phenol group with acetic anhydride in DMF.154 p-Nitrophenyl acetate (pNPAc) and 4-methylumbelliferyl acetate (AMC) were commercially available.

Scheme 4.1 A) (a) Ac2O, Py, 3 h, 97% for 5-quinolinol, 96% for 7-quinolinol. (b) 3eq MeI, THF, rt 2 h. 82% for 5- o quinolinol, 78% for 7-quinolinol 82%. B) (a) EtOH, cat AcOH, cat pip, 6 h, 74%. (b) DMF, 60 C, 20 eq Ac2O, cat TEA, 3 h, 83%.

4.2.2.2 Evaluation of Proposed Chromogenic Enzyme Substrates

Enzyme reactions were carried out in 50 µL solution volumes containing HEPES (100 mM, pH 7.5), PgaB (10 µM), and varying concentrations of pNPAc, AMC, 5-AMQ, 7-AMQ, and ACC. (0.05–1 mM for pNPAc, AMC, 0.05-10m M for 5-AMQ, 7-AMQ and ACC) at 25 °C in a 96 well plate. Substrates were all predissolved in DMSO and added to the reaction mixtures directly as such. The final DMSO concentration was 10% (v/v) in all cases. Due to substrate solubility constraints and background absorbance levels, 1-10 mM was used as a maximum concentration of substrate. Reactions were monitored in real time using the increase of either absorbance signal or fluorescence signal at the wavelengths found in Table 4.2, resulting from the hydrolysis and release of acetate over 10 min. Background hydrolysis rate were also monitored and subtracted from the values measured for the enzyme-catalyzed reactions.

Substrate Absorbance/Excitation (nm) Fluorescence Emission (nm)

405 -

354 445

460 -

454 -

386 446

Table 4.2 Absorbance / excitation and fluorescence emission wavelengths for the chromogenic substrates tested with PgaB.

When tested under the conditions described above, all of the tested compounds were found to be substrates for PgaB; however, the pNPAc, 5-AMQ, and 7-AMQ substrates suffered from high rates of background hydrolysis, making them unsuitable for further studies. AMC and ACC were more stable to hydrolysis, so experiments were continued with these two substrates only. The limiting solubility of AMC (~1 mM) prevented saturation of enzymatic activity, thus this substrate was not evaluated further. Fortunately, ACC was soluble up to 10 mM, which allowed for the determination of the kinetic parameters of PgaB and subsequently IcaB.

4.2.2.3 Determination of Enzyme Kinetics

4.2.2.3.1 PgaB

Enzyme reactions were carried out in 50 µL solutions volumes containing HEPES (100 mM, pH 7.5), PgaB (10 µM), and varying concentrations of ACC (0.05 –10 mM) at 25 °C in a Corning 3686 half-area 96 well black plate. A TECAN Safire2 plate reader was used to quantify fluorescence. ACC was predissolved in DMSO and added directly to the reaction mixtures as such. The final DMSO concentration was 10% (v/v) in all cases. Reactions were monitored in real time in a continuous assay measuring the increase of a fluorescence signal at 446 nm, resulting from the hydrolysis of the acetyl moiety with excitation at 386 nm over a 20 min period to determine steady-state initial velocity kinetics. Michaelis-Menten kinetic parameters were obtained using non-linear regression analysis of the data (Figure 4.4). A slow but measurable background hydrolysis rate was also monitored and subtracted from the enzyme-catalyzed reaction before data analysis. A control experiment replacing PgaB with bovine serum albumin (BSA, 10 μM) showed no increase in reaction rate above background. All assays were performed in triplicate. A calibration curve for ACC was obtained under the reaction conditions using 7- hydroxycoumarin-4-carboxylic acid as a standard to calculate reaction rate. A kcat value of -1 0.013s and a Km value of 1.2 mM were determined for the activity of PgaB on ACC (Table 4.3).

0.00012

0.00010

0.00008

0.00006

0.00004 Rate (mM/s) Rate

0.00002

0.00000

0 2 4 6 8 10 [ACC] (mM)

Figure 4.4 Rates of PgaB-catalyzed ACC deacetylation rates at varying ACC concentrations. Conditions were as follows: HEPES buffer (100 mM, pH 7.5) containing 10% (v/v) DMSO, 10 µM PgaB, ACC (0.5 mM – 10 mM). Assays were done in triplicate. Rates were obtained by measuring the increase in fluorescence emission at 446 nm resulting from the acetylosis of ACC. Vmax, Km and kcat were determined from non-linear regression analysis of the data to obtain a curve.

4.2.2.3.2 IcaB

For the continuous fluorogenic assay, purified IcaBHis10 (30 uM) in buffer B and CoCl2 (30uM) were incubated at room temperature for 30 min. ACC was predissolved in DMSO and added directly to the reaction mixtures as such. The final DMSO concentration was 10% (v/v) in all cases. To determine the steady-state initial velocity kinetics the substrate concentrations were varied from 0.01 to 5 mM. The sample was excited at 386 nm and continuously monitored for the increase in fluorescence emission at 446 nm. The production of phenol was quantified using 7-hydroxycoumarin-4-carboxylic acid as a standard. The fluorescence intensity data were

74 analyzed as above, using nonlinear regression analysis. All assays were performed in triplicate. -1 A kcat value of 0.0013s and a Km value of 0.5 mM were determined for the activity on IcaB on ACC (Table 4.3).

-1 -1 -1 Enzyme kcat (s ) Km (mM) kcat/Km (s mM )

PgaB 0.013 ± 0.002 1.2 ± 0.2 0.01

IcaB 0.0013 ± 0.0001 0.5 ± 0.1 0.0025

Table 4.3 Kinetic parameters for PgaB and IcaB with ACC.

4.2.3 Evaluation of Inhibitors towards IcaB and PgaB

The potential synthesized de-N-acetylase inhibitors presented in chapter 3 were tested with Ni2+-loaded PgaB and IcaB in a competitive assay in the presence of the fluorogenic substrate ACC. Previous work in our lab has shown that PgaB loaded with a selection of divalent ions was active in vitro, particularly with Fe2+; however this variant was unstable in ambient atmospheric conditions and therefore the Ni2+ loaded enzyme was used. Inhibitors stored as disulphides were pre-incubated with DTT for 10 minutes prior to assay onset to reduce the disulphide and bring the inhibitor into its active form. Overnight incubation of the inhibitors at 10 mM with the enzymes under assay conditions followed by 1H NMR analysis showed no observable hydrolysis of the amide bonds, indicating that none of the inhibitors acted as substrates for the enzymes. Inhibitors demonstrating greater than 10% inhibition of enzymatic ACC deacetylation at 10 mM were carried forward to full kinetic analysis. Methyl 2-amino-2- deoxy glucopyranoside 4 showed no inhibition of the enzyme at 1 mM.

4.2.3.1 Obtaining Inhibitor Activity

Enzyme reactions were carried out in 50 µL solution volumes in a Corning 3686 half-area 96-well black plate. The solution contained HEPES (100 mM, pH 7.5), PgaB (10 µM), and ACC

(5 mM) for PgaB, or IcaBHis10 (30 uM), sodium phosphate (50mM, pH 8), and NaCl (0.5M). In the IcaB case only, CoCl2 (30uM) was also added to the reaction mixture. ACC was predissolved in DMSO in both cases and added directly to the reaction mixtures as such. The final DMSO

75 concentration was 10% (v/v) in all assays. Reactions were carried out in the presence of each inhibitor (all at 10 mM, except for 1 mM in the cased of 21, 36, and 64). Reactions containing a free thiol (23, 45, 64) were pre-incubated with 1.1 equivalents of DTT for 5 min before assay onset. Reactions containing an inhibitor candidate with a primary amine were run at a maximal concentration of 1 mM due to its high background signal. Reactions were monitored in real time using the increase of fluorescence signal at 446 nm resulting from the hydrolysis of the acetyl moiety, over a 20 min period. Percent activity was measured as the relative difference of ACC acetolysis rate in the presence and absence of each inhibitor. Samples were corrected for background hydrolysis by measuring fluorescence signals of reactions lacking only the PgaB or IcaB enzyme. All assays were performed in triplicate.

4.2.3.2 Determining Kinetic Parameters of Inhibitors

Enzyme reactions were carried out in 50 µL solution volumes in a Corning 3686 half-area 96-well black plate. The solution contained HEPES (100 mM, pH 7.5), PgaB (10 µM), and varying concentrations of ACC (0.05 – 10 mM) for PgaB, or IcaBHis10 (30 µM) sodium phosphate (50 mM, pH 8) and NaCl (0.5 M), CoCl2 (30 µM), and varying concentrations of ACC (0.05 – 10 mM) for IcaB. Reactions were carried out in the presence of 3 different inhibitor concentrations (1 mM, 5 mM, 10 mM or 0.1 mM, 0.5 mM 1 mM for 64 only). Final DMSO concentration was 10% in every assay reaction mixture. Reactions containing a free thiol (23, 45) were pre-incubated with 1.1 equivalents of DTT for 5 min before assay onset. Reactions were monitored in real time over a 20 min period using the increase of fluorescence signal at 446 nm resulting from the hydrolysis of the acetyl moiety. A slow but measurable background hydrolysis rate was also monitored and subtracted from the enzyme-catalyzed reaction. All assays were performed in duplicate.

Ki values were obtained for each inhibitor by line-fitting the double reciprocal (1/rate vs 1/[ACC]) of each curve to obtain an α coefficient at each concentration, representing the change in slope (slope = αKm/Vmax) . From the derived α values, Ki values could be obtained from the following formula (α = 1 + [I]/Ki). The average Ki value (n = 3) is reported. Obtained Lineweaver-Burk plots revealed that all the inhibitors were competitive in nature, as they had similar y-intercepts. Obtained Lineweaver-Burk plots revealed that all the inhibitors were competitive in nature, as they had similar y-intercepts.

4.2.3.3 N-Acyl Inhibitors

Of the primary amide inhibitors, the thioglycolyl amide derivative 23 showed significant inhibitory activity towards PgaB (Ki = 480 µM), whereas the hydroxyglycolyl and aminoglycolyl amide derivatives 21 and 23 showed only minimal inhibitory capability (Table 4.4). This is consistent with previously reported inhibitors designed for other de-N-acetylases, as thioglycolyl amides have shown greater inhibitory capacity than their hydroxy and amino counterparts. The secondary amide inhibitors as a whole displayed much better inhibitory capacity than their primary amide counterparts. In particular, the hydroxyglycolyl inhibitor 35 was found to be the best inhibitor for PgaB activity (Ki = 320 µM). The amino counterparts 21 and 36 demonstrated no measurable inhibitory capacity (Table 4.4). Interestingly, the amino derivatives were the best inhibitors towards IcaB, in contrast to the PgaB results.

Structure

Residual PgaB 92 ± 3 98 ± 5b 24 ± 2c 106 ± 4 Activity (%)a IcaB 100 74 99 95

PgaB - - 480 ± 10 -

Ki (µM) IcaB - - - -

Structure

Residual PgaB 17 ± 2 96 ± 4b 52 ± 3c 103 ± 3 Activity (%)a IcaB 100 65 - 101

PgaB 320 ± 10 - 920 ± 20 -

Ki (µM) IcaB - - - -

Table 4.4 Inhibition of PgaB and IcaB by N-acyl inhibitors. a Remaining PgaB or IcaB enzyme activity in the presence of 10 mM inhibitor. b Remaining PgaB or IcaB enzyme activity in the presence of 1 mM inhibitor. c Inhibitor preincubated with 1.1 eq DTT prior to assay onset.

Figure 4.5 Lineweaver-Burk inhibitor plots for compounds 23 and 35 with PgaB.

One factor to take into consideration is the conformation of the amide bond. Crystal structures of inhibitors bound to other metallodeacetylases show the metal-binding heteroatom and the carbonyl oxygen in an eclipsed gauche conformation (Figure 4.6a).155 A crystal structure of compound 19 shows the hydroxyl moiety hydrogen bonded with the amido nitrogen, locking it in an anti-conformation relative to the carbonyl oxygen, and thus unsuitable for bivalent metal binding (Figure 4.6b,c).156 This factor could partly account for the limited inhibitory activity from the primary amide series of inhibitors, particularly the glycolyl inhibitors 19, 21, and 157 reduced 23, where this hydrogen bond network would limit rotation around the C(O)-Cα bond. Methylation of the amido nitrogen, as in the secondary amide series of inhibitors would effectively prevent this hydrogen bonding from occurring. As a result, adopting the anti- conformation relative to the carbonyl oxygen would be less energetically favourable, in turn promoting the eclipsed gauche conformation (Figure 4.6d).

Figure 4.6 A) Crystal structure of SAHA bound to zinc active site of human HDAC2155 B) Crystal structure of inhibitor 19 in the anti-conformation. C) Gauche projection of the primary amide series of inhibitors. D) Gauche projection of the secondary amide series of inhibitors. X in all cases refers to a heteroatom binding moiety (OH,

NH2, SH)

Another conformational factor to take into account is the rotation around the C2-N bond and the N-C(O) bond. There are 4 plausible states in which the acetamido functionality could 158 exist (Figure 4.7). NMR-based conformational analysis of methyl 2-deoxy-2-acetamido-β-D- glucopyranoside has shown that a significant population exists in the (E)-anti conformation, whereas methyl 2-deoxy-2-N-methyl-N-acetamido-β-D-glucopyranoside adopts primarily the (Z)- anti conformation. The improved inhibitory activity of the N-methyl inhibitors suggests that the preferred bound conformation of the acyl metal chelates may be the (E)-anti conformation. Oddly, the best inhibitor of the N-methyl series, 35, contains a metal chelating group that has not been previously reported as an effective inhibitor of amidases.

Figure 4.7 Possible conformations of the amide bond in GlcNAc derivatives. (Z)-Anti was found to be the principle conformation when R = H. (E)-Anti was found to be the principle conformation when R = Methyl.

4.2.3.4 N-Sulfonyl and N-Phosphonyl Inhibitors

Of the sulfonyl inhibitors, the N-methylated inhibitors 38 and 39 exhibited greater inhibitory potential than their non-sulfonyl counterparts towards PgaB in line with the acyl results; the sulfonamide inhibitor 39 was shown to have a Ki value of 680 µM (Table 4.5). The increased inhibitory capability of the sulfamide inhibitors 42 and 39 over the sulfonamide derivatives 25 and 38 could be due to additional hydrogen bonding. Unfortunately a conclusive study exploring the comparative inhibitory effects of sulfamides versus sulphonamides has not 159 been explored. The phosphonamidate inhibitor 51 showed a similar Ki value of 620 µM similar to the Ki value calculated for the sulfamide inhibitor 39. This similarity is not surprising due to the conformational and electronic similarity of phosphonamidates and sulfonamides/sulfamides. Unfortunately, these compounds only showed limited inhibition of IcaB.

Structure

Residual PgaB 93 ± 3 91 ± 2 32 ± 2 Activity (%)a IcaB - 80 -

PgaB - - 620 ± 10

Ki (µM) IcaB - - -

Structure

Residual PgaB 82 ± 3 33 ± 4 Activity (%)a IcaB 90 93

PgaB 5800 ± 100 680 ± 20

Ki (µM) IcaB - -

Table 4.5 Inhibition of PgaB and IcaB by N-acyl inhibitors. a Remaining PgaB or IcaB enzyme activity in the presence of 10 mM inhibitor.

4.2.3.5 N-Alkyl and C-2 Inhibitors

The N-alkyl inhibitor 47 was tested against PgaB and IcaB activity and was found to show no inhibition towards both enzymes (Table 4.6). It was hoped that the carboxylate group would mimic the substrate tetrahedral intermediate chemically and proximally. It is possible that the poor inhibitory capacity of 47 could be a result of suboptimal placement of the carboxylate to effectively mimic the tetrahedral intermediate.

Structure

PgaB 98 ± 5 Residual Activity (%)a IcaB 104

PgaB -

Ki (µM) IcaB -

Table 4.6 Inhibition of PgaB and IcaB by N-alkyl inhibitors. a Remaining PgaB or IcaB enzyme activity in the presence of 10 mM inhibitor.

Carbon-branched carbohydrates have long been used as important subunits in many antibiotics. C-2 carbohydrates have demonstrated activity against a number of bacterial carbohydrate deacetylase enzymes including LpxC and other GPI de-N-acetylase inhibitors.124a, 130a These C-2 derivatives provide a number of advantages. A methylene connection provides a greater degree of rotational freedom over an amide, allowing for a greater diversity of potential binding modes to an enzyme. Additionally, these derivatives are non-hydrolyzable, and as such, there is no risk of potential inhibitors acting as a substrate for the targeted enzyme.

Of the C-2 inhibitors tested, none of them showed significant inhibitory capacity towards PgaB relative to the other acyl and sulfonyl inhibitors tested (Table 4.7). Fortunately, compound

59 was shown to be an adequate inhibitor towards IcaB, with a Ki value of 460 µM.

Structure

Residual PgaB 98 ± 5 77 ± 5 84 ± 3 86 ± 5 Activity (%)a IcaB 104 102 22 82

PgaB - - -

Ki (µM) IcaB - - 460 ± 20 -

Table 4.7 Inhibition of PgaB and IcaB by C-2 inhibitors. a Remaining PgaB or IcaB enzyme activity in the presence of 10 mM inhibitor.

This result was unexpected, given the success of trifluoromethyl ketones with other metalloenzymes. Their lack of efficacy could be due to the molecules not hydrating to the 19 geminal diol. F NMR studies of these compounds in D2O showed primarily the presence of the ketone, and minuscule amounts of the hydrate (26:1 for 61) (Figure 4.8). No evidence of the hydrate was observed in the 1H or 13C NMR. Comparatively, in the case of (S)-2-amino-8,8,8- trifluoro-7-oxo-octanoic acid, a potential arginase inhibitor, a 15:1 ketone/hydrate ratio was 160 observed in D2O.

19 Figure 4.8 F NMR of compound 61 in D2O. Peak at -74.2 ppm represents the ketone. Peak at -84.4 ppm represents the hydrate.

It has been shown that an electron withdrawing group opposite of the trifluoro group is required for significant hydration of the ketone (Figure 4.9). 161 In the cases described here, the carbohydrate ring may not be inductively electron-withdrawing to shift the equilibrium towards the hydrate.

Figure 4.9 Hydration equilibrium of trifluoromethyl ketones in aqueous media.

4.2.3.6 Chemoenzymatic Approach: Pentasaccharide Inhibitor

The chemoenzymatically-synthesized pentasaccharide inhibitor 64 was tested against

PgaB and found to have a Ki of 280 µM (Table 4.8). This value represents the best inhibitor against PgaB found to date. Compared to its chemically-similar monosaccharide thiol- inhibitor 23, 64 was found to have a two-fold increase in inhibitory efficacy. A larger gain in affinity was expected however, given the length dependence of PgaB, which demonstrates higher activity with pentasaccharide PNAG over monosaccharide GlcNAc. Further work is required to determine with confidence the reason behind this limited improvement. Unfortunately, only enough inhibitor was synthesized to test it against PgaB only, and no experiments were performed with IcaB.

Structure

Residual PgaB 12 ± 3 Activity (%)a IcaB -

PgaB 280 µM

Ki (µM) IcaB -

Table 4.8 Inhibition of PgaB and IcaB by polysaccharide inhibitor 64. a Remaining PgaB enzyme activity in the presence of 1 mM inhibitor. bInhibitor preincubated with 1.1 eq DTT prior to assay onset.

4.3 Conclusions

In this chapter, a new method of monitoring PgaB and IcaB activity was explored using chromogenic acetyl substrates. With this method, it was possible to test the inhibitory efficacy of the array of synthesized inhibitors outlined in chapter 3. The thiol-inhibitor 23 was found to be the best monosaccharide inhibitor of PgaB, whereas the C-2 inhibitor 59 was found to be the best inhibitor towards IcaB. Finally, the chemoenzymatically-synthesized pentasaccharide inhibitor

64 was found to be the best inhibitor of PgaB, with a Ki value of 280 µM.

5 Future Perspectives 5.1 Substrate development for hexosaminidases

The availability of substrates to monitor hexosaminidase and glycosidase activity is of great interest to the scientific community.108 Novel glycosyl substrates could be of great value in helping determine the substrate specificity of glycosidase enzymes. As demonstrated with DspB and the carbamate substrate 2, this compound could be used to further elucidate whether other enzymes can act on a 1,6 linkage without consuming potentially-valuable native polysaccharide substrates. Among family 20 glycosyl hydrolases in the CAZy database, only DspB from A. actinomycetemcomitans is classified as a β-1,6-N-acetylglucosaminidase. Considering the prevalence of the β-1,6 linkage of GlcNAc in nature, especially among PNAG-producing microbes, it is unlikely that DspB is the only β-1,6 specific glycosidase. Thus, assays with substrate 2 in conjunction with the commonly-used pNP-GlcNAc, could shed light on the function of other newly-isolated β-hexosaminidases.

Alternatively, despite the inactivity of the acetal substrate 1, its conceptual utility has yet to be disproven. If our hypothesis is correct that its ineffectiveness is due to a non-reactive conformation, exchange of the p-nitrophenol group for another chromophoric analyte may resolve the issue. Particularly, substitution of the 5- or 7-hydroxyquinolinium chromophores appears promising. Due to the charged quinolinium nitrogen, these compounds would have a vastly different electrostatic profile than the pNP acetal substrate counterpart, and such a change may also induce a change in conformation (Figure 5.1). Additionally, their chromogenic properties make them ideal analytes, as a drastic increase in absorbance at 460 nm and 464 nm is observed upon hydrolysis to 5- and 7-hydroxyquinolinium, resulting from the newly possible hydroxyl conjugation. This shift in absorbance wavelengthcould be used to monitor reaction progress.162

Figure 5.1 Potential novel acetal substrates.

Finally, β-Hexosaminidases have been implicated in a number of disease states. Human lysosomal β-hexosaminidases catalyze the hydrolysis of GlcNAc or GalNAc residues from the reducing end of a number of biologically-significant glycoconjugates. Three isoforms are known to exist: β-hexosaminidase A (HexA), β-hexosaminidase B (HexB), and β-hexosaminidase S (HexS).163 These isoforms play an important role in the lysosomal catabolism of glycosphingolipids (GSLs) (Figure 5.2). Defects in any of the three β-hexosaminidase genes can result in the accumulation of non-degraded glycolipids within the lysosomal compartments and leading to severe neurodegenerative storage diseases including Tay-Sachs disease (HexA mutation) and Sandhoff disease (HexB mutation), which are devastating neurological disorders that usually lead to death before the age of four.164

Figure 5.2 HexA enzymatic hydrolysis of GalNAc residue on GM2 ganglioside to produce GM3 ganglioside.

Positive diagnoses of these diseases are dependent on the reliability of hexosaminidase assays of patient blood samples. Serum or leukocytes are most commonly used as HexA/B/S sources, however dried blood spots on filter paper has also been evaluated, with the latter being effective only with homozygote carriers.165 Unfortunately, with the current methodology, false positives can range from 2% to 35% depending on the ethnicity of the individual.166 This is a result of certain variations of HexA having reduced activity towards artificial substrates, but no impairment in its ability to degrade the physiological substrate GM2. Although false negative results are rare in comparison, 10% of the cases result in an inconclusive result among the general population, resulting in the need for more expensive genetic mutation analysis.167

Therefore, novel glycoside substrates hold great potential for improving the reliability of measuring HexA/B/S activity in human patient samples, ultimately improving the clinical standard of care. Increased sensitivity and specificity could reduce the number of false positives, providing greater respite to the affected patients and decreasing the burden on the medical

91 system. With the goal of improving this pharmological assay, novel N-acetylgalacto- carbamate or acetal substrates could be tested with human HexA/B/S for glycosidic activity.

5.2 Inhibitor development for PgaB and IcaB

Strains of bacteria lacking the pgaB or icaB genes produce only the PNAG polysaccharide show limited ability to form biofilms and demonstrate compromised antibiotic resistance. Successful development of an effective inhibitor for PgaB and/or IcaB could lead to the development of a treatment strategy towards the dispersal of bacterial biofilms. Unfortunately, when compounds 23, 35, and 38 were tested against S. epidermidis 1457 in biofilm inhibition assays at a concentration of 1 mM, no inhibition was observed.139 This is likely due to the weak activity of these compounds. Based on these findings, tighter-binding inhibitors are likely required for future utility of this strategy.

One approach that may aid in inhibitor development is further crystallographic analysis of the target enzyme. Although the structure of PgaB has been obtained through X-ray crystallography, attempts to co-crystallize PgaB with either an inhibitor or a PNAG polysaccharide substrate have been unsuccessful.65a Such data would offer valuable information regarding the binding mode of potential inhibitors and could aid in structure-based drug design.168 This includes important points of contact made between the inhibitors and the enzyme, and the alignment of available binding pockets surrounding the active site which can be taken advantage of with further functionalization of the inhibitor candidates.

Another experimental method that could offer information about inhibitor binding would be isothermal titration calorimetry (ITC). As mentioned previously, it was observed that in the case of an MMP-9 inhibitor, the metal-chelating hydroxamic acid moiety accounted for 96% of the binding energy.126 To test the assumption that the inhibitors are acting through a metal- chelating mechanism in the active sites of PgaB and IcaB, utilizing an array of similar metal- chelating inhibitors and obtaining the binding energy of each could offer insight. For example, using an array of N-(methyl)thioglycolamide, N-(cyclohexyl)thioglycolamide, and the successful thiol inhibitors 23 and 64, it may be possible to differentiate the binding energies resulting from the metal-chelating thioglycolamide moieties (Figure 5.3).

Figure 5.3 Array of compounds to test using isothermal calorimetry

Another area to be explored could be the use of non-carbohydrate base inhibitors. As observed with lipopolysaccharide deacetylase LpxC, despite the tight binding of the carbohydrate inhibitor Tu-514, with a Ki of 640 nM, a series of small molecule aryl compounds were found to be over 100-fold more effective, with the best one, CHIR-090, demonstrating a Ki of 2 nM (Figure 5.4a).125, 169 Additionally, utilizing a computational molecule library screen, a small molecule aryl inhibitor of the CE4 family pneumococcal peptidoglycan deacetylase PgdA was discovered, exhibiting an IC50 value of 130 µM under the reported reaction conditions (Figure 5.4b). Computational docking studies suggested that it bound to the metal site in a bidentate fashion, utilizing the carboxylic acid moiety and the ketone oxygen.153 Common features of these non-carbohydrate inhibitors include metal-binding moieties to bind the metal centers and aryl groups which can mimic the carbohydrate rings of native substrates.

Figure 5.4 A) Inhibitors developed against LpxC. B) Small molecule inhibitor developed against pneumococcal peptidoglycan deacetylase PgdA.

Finally, due to the low affinity of PgaB and IcaB to oligosaccharides of their native substrate (refer to table 4.1), it may ultimately prove perplexing to find effective carbohydrate- based inhibitors. Additionally, due to their high polarity, carbohydrate-based drugs have limited capability to cross the enterocyte layer of the small intestine when administered orally, and suffer from fast metabolic renal extraction.170 In principle, given our access to the structures of PgaB and IcaB, using a computational structure-based virtual screening followed by a biochemical evaluation of a small molecule library may unveil novel small molecule inhibitors to these enzymes. This virtual screening method has seen success with several enzyme targets, including the previously-mentioned PgdA.171

6 Experimental 6.1 General Methods

Reagents were obtained from Sigma-Aldrich or Acros Organics and were used without further purification. If appropriate, reactions were carried out under an inert atmosphere of nitrogen. Standard syringe techniques were applied for the transfer of dry solvents and air- or moisture- sensitive reagents. Reactions were monitored by TLC using Silica Gel 60 F254 (EMD Science) with detection by quenching of fluorescence and/or by visualization with phosphomolybdic acid 1 in ethanol (0.5% w/v), methanolic H2SO4 (10% v/v), or ninhydrin in ethanol (0.2% w/v). H and 13C NMR spectra were recorded at 25 °C with a Mercury 300 MHz or a Varian 400 MHz (75 MHz or 100 MHz for 13C, respectively) spectrometer. 1H-NMR chemical shifts are reported in parts per million (ppm) relative to tetramethylsilane (TMS) as an internal standard, or a residual proton peak of the solvent: δ = 7.26 ppm for CDCl3, δ = 4.81 ppm for D2O, δ = 3.31 ppm for

CD3OD. Multiplicities are reported as: s (singlet), d (doublet), t (triplet), dd (doublet of doublets), or m (multiplet). Broad peaks are indicated by br. Coupling constants are reported as J-values in Hz. The number of protons (n) for a given resonance is indicated as nH, and is based on spectral integration values. 13C-NMR chemical shifts are reported in parts per million (ppm) relative to tetramethylsilane (TMS) as an internal standard, or a carbon peak of the solvent: δ =

77.0 for CDCl3, δ = 49.0 for CD3OD. Multiplicities in carbon represent different rotamers and are reported as: d (doublet). Flash chromatography was performed on Silica-P Flash Silica Gel 60 (40–63 μm particle size, Silicycle). High-resolution mass spectra were obtained from an ABI/Sciex QStar mass spectrometer (ESI). Absorption spectra were measured on a Schimadzu UV-2401PC spectrophotometer. Fluorescence measurements were obtained on a TECAN Safire2 plate reader. HPLC was performed on a Waters 1525 binary HPLC pump with a Waters 2487 dual k absorbance detector or a Gilson 321 HPLC pump with a Gilson UV–Vis 156 dual k absorbance detector.

6.2 Procedures

4-nitrophenoxymethoxy 2-pthalimido-3,4,6-tri-O-acetyl-2-deoxy-β-D-glucopyranoside (5)

3,4,6-triacetyl-2deoxy-2-phthalimido-D-glucose (500 mg,1.15 mmol) was dissolved in acetonitrile (5 ml) and to it K2CO3 (238 mg, 1.72 mmol), was added followed by the addition of 4-nitrophenoxymethoxychloride (430 mg, 2.3 mmol), and NaI (258 mg, 1.72 mmol). The reaction was allowed to proceed overnight at room temperature. TLC analysis showed generation of a new spot (EtOAC:n-pentane, 1:2, Rf 0.5). Solvent was evaporated and the crude mixture was diluted with dichloromethane (25 ml), and washed with water (50 ml), the organic phase was separated, dried over MgSO4, filtered and evaporated to syrup in vacuo. The crude product thus obtained was purified by flash chromatography eluting with (EtOAC:n- pentane, 1:3) to furnish 4-nitrophenoxymethoxy 2-pthalimido-3,4,6-tri-O-acetyl-2-deoxy-β-D- 1 glucopyranoside 4 (269 mg, 40%), as colourless paste. H NMR (CDCl3, 300 MHz) δ: 7.68-7.53 (m, 6H), 6.79-6.72 (m, 2H), 5.82 (t, 1H, J = 9.2 Hz,), 5.71 (d, 1H, J = 8.6 Hz), 5.44-5.37 (m, 2H), 5.18 (t, 1H, J = 9.1 Hz), 4.39-4.28 (m, 2H), 4.22 (dd, 1H, J = 2.1 Hz, 10.2 Hz), 3.97-3.92 13 (m, 1H), 2.13 (s, 3H), 2.02 (s, 3H), 1.81 (s, 3H). C NMR (75 MHz, CDCl3) δ: 170.6(2), 170 (2), 169.4, 160.1, 141.6, 134 (2), 125.3 (2), 123.3 (4), 115.2 (2), 94.1, 88.1, 72.2, 70.1, 68.5, 61.7, 54, + 20.7, 20.6, 20.4. HRMS cald for C27H26N2O13Na(M+Na) : 609.1333; found 609.1337.

4-nitrophenoxymethoxy 2-acetamido-3,4,6-tri-O-acetyl-2-deoxy-β-D-glucopyranoside (6)

Compound 5 (200 mg, 0.34 mmol) was dissolved in butanol (5 ml) under Nitrogen and to it ethylene diamine (1 ml) was added. The solution was heated to 120 0C for 20 h, after which the solvent was evaporated in a rotavapour and the crude was dried under pump. To the crude pyridine (3 ml) and Acetic anhydride (2 ml) was added and the solution was stirred at room temperature overnight. The solvent was evaporated and the crude mixture was diluted with dichloromethane (25 ml), and washed with 1N HCl (25 ml), water (50 ml), and sodium bicarbonate (25 ml), the organic phase was separated, dried over MgSO4, filtered and evaporated to syrup in vacuo. The crude product thus obtained was purified by flash chromatography eluting with ( EtOAC:n-pentane, 1:1) to furnish 4-nitrophenoxymethoxy 2- acetamido-3,4,6-tri-O-acetyl-2-deoxy-β-D-glucopyranoside 5 (132 mg, 78%), as colourless

1 paste. H NMR (CDCl3, 300 MHz) δ: 8.16-8.11(m, 2H), 7.07-7.02 (m, 2H), 5.44 (d, 1H, J = 7.6 Hz,), 5.34 (d, 1H, J = 7.6 Hz), 5.23 (t, 1H, J = 8.9 Hz), 5.13-5.05 (m, 2H), 4.85 (d, 1H, J = 8.5 Hz), 4.23 (dd, 1H, J = 4.5 Hz, 12 Hz), 4.12 (dd, 1H, J =2.3 Hz, 10 Hz), 3.97-3.93 (m, 1H), 3.70- 13 3.66 (m, 1H), 2.04 (s, 3H), 1.96 (s, 3H), 1.94 (s, 3H), 1.46 (s, 3H). C NMR (75 MHz, CDCl3) δ: 171, 170.7, 169.8, 161.6, 142.5, 125.7 (2), 116.1 (2), 97.3, 89.1, 72.2, 72, 68, 61.7, 53.9, 22.8, + 20.7, 20.6 (2). HRMS cald for C21H26N2O12Na(M+Na) : 521.1383; found 521.1386.

4-nitrophenoxymethoxy 2-acetamido-2-deoxy- β-D-glucopyranoside (1).

Compound 6 (100 mg, 0.2 mmol) was dissolved in MeOH (5mL) followed by addition of NaOMe (0.5 M in MeOH, 0.1 mL) PH Paper alkaline and the solution was allowed to stir at room temperature for 6 hours. The solution was then neutralized with Amberlite IR 120 H+ resin and filtered. The filtrate was evaporated to afford pure compound 4- nitrophenoxymethoxy 2-acetamido-2-deoxy- β-D-glucopyranoside 1 (67 mg, 90%) as amorphous white solid. 1H NMR (MeOD, 300 MHz) δ: 8.24-8.19 (m, 2H), 7.21-7.15 (m, 2H), 5.55 (d, 1H, J = 7.6 Hz), 5.46 (d, 1H, J = 7.6 Hz), 4.75 (d, 1H, J = 8.5 Hz), 3.89 (dd, 1H, J = 1.7 Hz, 10.4 Hz), 3.72-3.63 (m, 2H), 3.47-3.41 (ddd, 1H, J = 8.5 Hz, 1.5 Hz, 8.7 Hz), 3.36-3.27 (m, 3H), 1.49 (s, 3H).13C NMR (75 MHz, MeOD) δ: 173.5, 163.6, 143.7, 126.6 (2), 117.6 (2), 99.3, + 90.5, 78.4, 75.5, 72, 62.8, 57, 22.7. HRMS cald for C15H20N2O9Na(M+Na) : 395.1067; found 395.1064.

4-Nitrophenyl carbamoyl 2-acetamido-3,4,6-tri-O-acetyl-2-deoxy-β-D-glucopyranoside (8)

Compound 7 (200 mg, 0.58mmol) was dissolved in toluene (10 mL) followed by the addition of a catalytic amount of triethylamine (0.05 mL) and p-nitrophenyl isocyanate (190mg, 1.16mmol).The solution was stirred at room temperature for 6 hours, at which point a yellow solid had precipitated. The solution was chilled in an ice bath and pure 4-nitrophenyl carbamoyl 2-acetamido-3,4,6-tri-O-acetyl-2-deoxy- 1 β-D-glucopyranoside was isolated by filtration (253 mg, 86% yield) H NMR (CDCl3, 400 MHz) δ: 8.22 (d, 2H, J = 9.2 Hz), 7.51 (d, 2H, J = 9.2 Hz), 5.61 (d, 1H, J = 8.8 Hz,), 5.58 (bd, 1H, J = 5.2) 5.13-5.05 (m, 2H), 4.45 (d, 1H, J = 8.5 Hz), 4.35 (dd, 1H, J = 4.5 Hz, 12 Hz), 4.15 (dd, 1H, J =2.3 Hz, 12 Hz), 3.86-3.72 (m, 1H), 2.06 (s, 3H), 2.04 (s, 3H), 2.03 (s, 3H), 1.98 (s, 3H). 13C

NMR (75 MHz, CDCl3) δ: 171.4, 170.6, 169.8, 161.2, 148.5, 141.2, 130.7 (2), 116.1 (2), 102.3, + 89.2, 73.1, 71.9, 68, 61.4, 53.7, 22.4, 20.8, 20.6 (2). HRMS cald for C21H26N2O12Na(M+Na) : 534.1374; found 534.1379.

4-Nitrophenyl carbamoyl 2-acetamido-2-deoxy-β-D-glucopyranoside (2)

Compound 8 (200 mg, 0.4mmol) was stirred as a slurry in MeOH (10 mL) and to it was added hydrazine monohydrate (0.05ml 0.13 mmol). After 4 hours, the reaction mixture was then evaporated under reduced pressure, and the remaining residue was purified by flash chromatography eluting with MeOH : DCM (1:20) to afford 4-nitrophenyl carbamoyl 2-acetamido-2-deoxy-β-D-glucopyranoside (80 mg, 48% yield) as a faint yellow powder.1H NMR (MeOD, 400 MHz) δ: 8.22 (d, 2H, J = 9.2Hz), 7.51 (d, 2H, J = 9.2Hz), 5.61 (d, 1H, J = 8.8 Hz), 3.95 (m, 2H), 3.75 (dd, 1H, J = 1.7 Hz, 10.4 Hz), 3.62-3.31 (m, 3H), 2.04 (s, 3H).13C NMR (75 MHz, MeOD) δ: 173.5, 143.6, 141.7, 129.2 (2), 113.6 (2), 105.2, 90.5, 78.1, 75.2, 72.3, 62.4, 57.3, 22.4. HRMS cald for + C15H19N3O9Na(M+Na) : 408.1062; found 408.1068.

Methyl 2-ammonium-2-deoxy-β-D-glucopyranoside acetate140, 142 (15)

N-Acetylglucosamine (5 g, 25 mmol) and tosylhydrazide (5.5 g, 25 mmol) in a mixture of DMF (20 mL), water (4.5 mL) and acetic acid (0.5 mL) was stirred at 37 °C for 24 h. The solution was then poured into diethyl ether (1 L) and stirred vigorously to promote formation of a white precipitate, 13. Vacuum filtration of the suspension yielded the glycosyltosylhydrazide donor (10.2 g, 92%). The glycosyl donor (5 g, 12 mmol) was then dissolved in DMF (150 mL) containing MeOH (10 mL, 240 mmol). N-Bromosuccinimide (2.2 g, 26 mmol) was then added to the solution portion-wise over 5 min 0 °C. Upon completion of the addition, the solution was allowed to stir for 30 min at room temperature. The solution was then evaporated under reduced pressure, and the resulting residue was dissolved in hot EtOH (100 mL), and chilled to -20 °C to allow crystallization of the succinimide. Filtration and subsequent evaporation of the EtOH filtrate followed by flash chromatography (10% MeOH in DCM) of the resulting residue yielded 14 (1.8 g 62%) as a white solid. 1 (1.8 g, 8 mmol) was then dissolved in hydrazine monohydrate (10 mL) and allowed to

98 stir at 110 °C for 48 h. The solution was then evaporate under reduced pressure, followed by the addition of 1M AcOH (5 mL) and a subsequent evaporation under reduced pressure to form the acetate salt. Recrystallization of the resulting residue with MeOH/EtOAc yielded 15 (1.85 g, 96%).

Methyl 2-(2-benzyloxy)acetamido-2-deoxy-β-D-glucopyranoside (18)

The title compound was prepared by dissolving compound 15 (200 mg, 0.93 mmol), benzyloxyacetic acid (220 mg, 0.93 mmol) and triethylamine (55 µL, 1 mmol) in DMF. HBTU (355 mg, 0.93 mmol) was then added portion wise to the mixture at 0 °C over 30 min and allowed to stir overnight at room temperature. The solution was then evaporated under reduced pressure and the resulting residue was purified by flash chromatography (8% MeOH in 1 DCM) to afford 18 (198 mg, 84%) as a white solid. H NMR (400 MHz, D2O) δ 7.56 – 7.41 (m,

5H, Ar-H), 4.69 (s, 2H, CH2), 4.54 (d, J = 8.5 Hz, 1H, H-1), 4.14 (d, J = 5.3 Hz, 2H, CH2Ar), 3.97 (dd, J = 12.1 Hz, 1.8 Hz, 1H, H-6a), 3.84 – 3.73 (m, 2H, H-6b, H-5), 3.64 (dd, J = 10.3 Hz, 13 8.7 Hz, 1H, H-2), 3.54 (s, 3H, CH3), 3.52 - 3.47 (m, 2H, H-3, H-4). C NMR (75 MHz, D2O) δ 173.37, 136.72, 128.99, 128.79, 128.76, 101.93, 76.07, 73.80, 73.37, 70.18, 68.60, 60.90, 57.28, + 55.32. HRMS m/z calcd. for C16H24NO7 (M+H ) 342.1536, found 342.1547.

Methyl 2-(2-hydroxy)acetamido-2-deoxy-β-D-glucopyranoside (19)

The title compound was prepared by dissolving compound 18 (100 mg, 0.41 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10% Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. The suspension was filtered through celite, and the resulting filtrate evaporated under reduced pressure to afford 19 1 (82 mg, 89%) as a colorless oil. H NMR (400 MHz, D2O) δ 4.57 (d, J = 8.4 Hz, 1H, H-1), 4.16

(s, 2H, CH2), 3.98 (dd, J = 12.3 Hz, 1.9 Hz, 1H, H-6a), 3.84 – 3.76 (m, 2H, H-6b, H-5), 3.67 (dd, 13 J = 10.3 Hz, 8.6 Hz, 1H, H-2), 3.55 (s, 3H, CH3), 3.53 – 3.45 (m, 2H, H-3, H-4). C NMR (75

MHz, D2O) δ 175.77, 101.93, 76.08, 73.86, 70.15, 61.17, 60.91, 57.24, 55.30. HRMS m/z calcd. + for C9H16NO7Na (M+Na ) 274.0906, found 274.0921

Methyl 2-(2-benzylcarboxyamino)acetamido-2-deoxy-β-D-glucopyranoside (20)

The title compound was prepared by dissolving compound 15 (200 mg, 0.93 mmol), Cbz-Gly-OH (180 mg, 0.93 mmol) and triethylamine (55 µL, 1 mmol) in DMF. HBTU (355 mg, 0.93 mmol) was then added portion wise to the mixture at 0 °C over 30 min and allowed to stir overnight at room temperature. The solution was then evaporated under reduced pressure and the resulting residue was purified by flash chromatography (8% MeOH in 1 DCM) to afford 20 (178 mg, 76%) as a white solid. H NMR (400 MHz, D2O) δ 7.58 – 7.39 (m,

5H, Ar-H), 4.52 (d, J = 8.4 Hz, 1H, H-1), 4.22 (d, J = 5.3 Hz, 2H, CH2Ar), 3.97 (dd, J = 12.0

Hz, 1.9 Hz, 1H, H-6a), 3.90 – 3.70 (m, 4H, H-6b, H-5, CH2), 3.66 – 3.58 (m, 1H, H-2), 3.58 – 13 3.42 (m, 5H, H-3, H-4, CH3). C NMR (75 MHz, D2O) δ 175.74, 160.93, 137.29, 130.29, 129.38, 129.17, 104.55, 84.41, 78.61, 76.49, 72.70, 70.12, 63.48, 59.92, 58.30. HRMS m/z calcd + for C17H25N2O8 (M+H ) 385.1554, found 385.1561 Methyl 2-(2-amino)acetamido-2-deoxy-β-D-glucopyranoside (21)

The title compound was prepared by dissolving compound 20 (100 mg, 0.40 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10% Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. The suspension was filtered through celite, and the resulting filtrate evaporated under reduced pressure to 1 afford 21 (72 mg, 81%) as a colourless oil. H NMR (400 MHz, D2O) δ 4.57 (d, J = 8.4 Hz, 1H,

H-1), 3.98 (dd, J = 12.3 Hz, 1.9 Hz, 1H, H-6a), 3.91 (d, J = 2.8 Hz, 2H, CH2), 3.84 – 3.76 (m,

2H, H-6b, H-5), 3.67 (dd, J = 10.3 Hz, 8.6 Hz, 1H, H-3), 3.55 (s, 3H, CH3), 3.53 – 3.45 (m, 2H, 13 H-4, H-2). C NMR (75 MHz, D2O) δ 175.77, 101.93, 76.08, 73.86, 70.15, 61.17, 60.91, 57.24, + 55.30. HRMS m/z calcd. For C9H18N2O6Na (M+Na ) 273.0900, found 273.0918

100

Methyl 2-(2-thioacetyl)acetamido-2-deoxy-β-D-glucopyranoside (22)

The title compound was prepared by dissolving compound 15 (200 mg, 0.93 mmol) and triethylamine (55 µL, 1 mmol) in DMF. Pentafluorophenyl 2-(acetylthio)acetate (310 mg, 0.93 mmol) was then added to the mixture and allowed to stir overnight at room temperature. The solution was evaporated under reduced pressure and the resulting residue was purified by flash chromatography (8% MeOH in DCM) to afford 22 (160 mg, 78%) as a white solid. 1H

NMR (400 MHz, D2O) δ 4.49 (d, J = 8.3 Hz, 1H, H-1), 3.96 (dd, J = 12.2 Hz, 1.6 Hz, 1H, H-

6a), 3.74 - 3.64 (m, 4H, H-6b, H-2, CH2), 3.60 (dd, J = 9.8 Hz, J = 8.6 Hz, 1H, H-3), 3.52 (s, 13 3H, OCH3), 3.51 – 3.44 (m, 2H, H-3, H-4), 2.45 (s, 3H, CH3). C NMR (100 MHz, D2O) δ 199.48, 171.71, 102.14, 76.04, 73.77, 70.16, 60.90, 57.40, 56.12, 33.24, 29.67. HRMS m/z calcd. for C11H19NO7SNa (M+Na+) 332.0778, found 332.0774

2,2'-disulfanediylbis(N-(Methyl-2-deoxy-β-D-glucopyranos-2-yl)acetamide) (23)

The title compound was prepared by dissolving compound 22 (80 mg, 0.3 mmol) in MeOH (5 mL) containing a catalytic amount of NaOMe and the solution was stirred for 10 min. Amberlite resin (H+) was added to the mixture until the solution was neutral. Filtration and evaporation under reduced pressure yielded the symmetrical disulfide 23 (65 mg, 1 92%) as a white powder. H NMR (400 MHz, D2O) δ 4.54 (d, J = 8.4 Hz, 1H, H-1), 3.98 (d, J =

11.9 Hz, 1H, H-6a), 3.78 (d, J = 9.1 Hz, 2H, CH2), 3.68 – 3.43 (m, 8H, H-6b, H-5, H-3, H-4, 13 OCH3, H-2). C NMR (100 MHz, D2O) δ 175.23, 100.27, 76.34, 72.38, 69.93, 60.60, 57.58, + 55.94, 23.11. C18H33N2O14S2 (M+H ) 533.1484, found 533.1469

101

Methyl 2-(3-hydroxy)ureayl-2-deoxy-β-D-glucopyranoside (24)

The title compound was prepared by dissolving compound 15 (200 mg, 0.93 mmol) in MeOH with triethylamine (55 µL, 1 mmol). 4-nitrophenyl carboxamate (385 mg, 1.5 mmol),4 was subsequently added portion wise over 30 min at 0 ºC and the mixture was allowed to stir overnight. Following complete consumption of the glycosyl starting material, as judged by TLC, the reaction mixture was evaporated under reduced pressure, and the resulting residue was dissolved in 1M AcOH (25 mL) and washed with 3x50 mL EtOAc. Collection and evaporation of the aqueous layer under reduced pressure, followed by flash chromatography of the resulting residue (8% MeOH in DCM) yielded 24 (110 mg, 54%) as a white solid. 1H NMR

(400 MHz, D2O) δ 4.54 (d, J = 8.1 Hz, 1H, H-1), 3.98 (dd, J = 12.3 Hz, 1.8 Hz, 1H, H-6a), 3.79 13 (dd, J = 12.3 Hz, 5.6 Hz, 1H, H-6b), 3.62 – 3.42 (m, 7H, H-2, H-3, CH3, H-4, H-5). C NMR

(75 MHz, D2O) δ 163.57, 100.43, 76.05, 74.14, 70.20, 60.95, 57.30, 56.00. HRMS m/z calcd. for + C8H16N2O7Na (M+Na ) 275.0855, found 275.0849

Methyl 2-methylsulfonamido-2-deoxy-β-D-glucopyranoside (25)

The title compound was prepared by dissolving compound 15 (200 mg, 0.93 mmol) and triethylamine (110 uL, 2 mmol) in DMF. Mesyl chloride (65 uL, 1 mmol) was then added to the mixture slowly and allowed to stir overnight at room temperature. The mixture was evaporated under reduced pressure and the resulting residue was purified by flash chromatography (8% MeOH in 1 DCM) to afford 25 (112 mg, 58%) as a white solid. H NMR (400 MHz, CD3OD) δ 4.37 (d, J = 8.4 Hz, 1H, H-1), 3.89 (dd, J = 12.3 Hz, 1.9 Hz, 1H, H-6a), 3.69 (dd, J = 12.4 Hz, 5.8 Hz, 1H, 13 H-6b), 3.54 (s, 3H, OCH3), 3.46 – 3.34 (m, 4H, H-2, H-3, H-4, H-5), 3.11 (s, 3H, SCH3). C

NMR (100 MHz, D2O) δ 100.83, 77.01, 73.21, 70.50, 60.41, 57.38, 56.02, 39.12 HRMS m/z + calcd. for C8H17NO7SNa (M+Na ) 294.0711, found 294.0713

102

Methyl 2-methylamino-2-deoxy-3,4,6-tri-O-benzyl-β-D-glucopyranoside (29)

The title compound was synthesized in 3 steps following a literature procedure.144 Methyl 2-acetamido-2-deoxy-3,4,6-tri-O-benzyl-β-D- glucopyranoside 26 (5 g, 10 mmol) was dissolved in THF (100 mL). Boc Anhydride (4.4 g, 20 mmol) and DMAP (0.5 g, 0.5 mmol) were then added and the solution was then refluxed for 16 h. The solution was then evaporated to dryness under reduced pressure. The resulting residue was then dissolved in MeOH (100 mL) containing a catalytic amount of NaOMe for 4 h, at which point the Boc protected compound precipitated. Vacuum filtration of resulting solid yielded methyl 2-tert-butylcarboxamido-2-deoxy-3,4,6-tri-O-benzyl-β-D- glucopyranoside 28 (3.8 g, 7.2 mmol). The resulting sugar (28) was then dissolved in THF (100 mL) and to the solution, on ice, was slowly added LiAlH4 (21.6 mmol) a 1M solution in THF.

Following complete addition of LiAlH4, the solution was refluxed for 12 h. The solution was then neutralized with water, washed consecutively with a 200 mL of a 0.1 M solution of sodium potassium tartrate, water, brine, and the resulting organics were dried over magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography (1:1 EtOAc/Pentane) 1 afforded 29 (2.1 g, 4.8 mmol) as a colorless oil. H NMR (400 MHz, CDCl3) δ 7.56 – 7.12 (m,

15H, ArH), 4.95 (d, J = 11.5 Hz, 1H, Ar-CHaHb), 4.79 (d, J = 10.8 Hz, 1H, Ar-CHaHb), 4.74 –

4.55 (m, 4H, Ar-CH2, Ar-CH2), 4.16 (d, J = 7.9 Hz, 1H, H-1), 3.81 – 3.65 (m, 3H, H-6a, H-6b,

H5), 3.57 – 3.44 (m, 5H, H-4, H-3, OCH3), 2.51 – 2.43 (m, 4H, H-2, NCH3), 1.26 (s, 1H, N-H). 13 C NMR (75 MHz, CDCl3) δ 138.53, 138.26, 137.86, 128.64, 128.59, 128.56, 128.49, 128.36, 128.08, 127.93, 127.75, 100.22, 100.08, 79.63, 79.39, 79.26, 78.54, 75.17, 74.96, 73.63, 68.82, + 61.49, 57.01, 42.73, 29.84, 28.56, 28.51. HRMS m/z calcd. for C29H36NO5 (M+H ) 478.2596, found 478.2588.

103

Methyl 2-N-methyl-(2-benzyloxy)acetamido-2-deoxy-3,4,6-tri-O-benzyl-β-D-glucopyranoside (30)

The title compound was prepared by dissolving 29 (250 mg, 0.55 mmol), benzyloxyacetic acid (120 mg, 0.6 mmol) and triethylamine (30 uL, 0.58 mmol) in a 50:50 EtOAc/DCM solution. DCC (140 mg, 0.66 mmol) was then added portion wise to the mixture at 0°C over 30 min and allowed to stir overnight at room temperature. The mixture was then filtered through celite and evaporated under reduced pressure. The resulting residue was then dissolved in EtOAc, washed consecutively with 1M NaOH(aq), water, and brine, and dried with magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography (1:1 EtOAc/Pentane) 1 afforded 15 (260 mg 89%) as a colourless oil. H NMR (400 MHz, CDCl3; mixture of rotamers)

δ 7.40 – 7.04 (m, 20H, Ar-H), 4.81 – 4.63 (m, 2H, Ar-CH2), 4.60 – 4.37 (m, 7H, Ar-CH2, H-1),

4.32 – 4.15 (m, 1.4H, COCH2), 3.97 (d, J = 13.8 Hz, 0.6H, COCH2), 3.73 – 3.55 (m, 5H, H-6a,

H-6b, H-5, H-2, H-4), 3.48 – 3.30 (m, 4H, H-3, OCH3), 2.98 - 2.79 (br s, 2.2H, NCH3), 2.68 (s, 13 1.8H NCH3). C NMR (75 MHz, CDCl3) δ 171.07, 138.65, 138.30, 137.98, 137.96, 137.85, 137.57, 128.66, 128.61, 128.56, 128.52, 128.49, 128.44, 128.39, 128.33, 128.17, 128.15, 128.08, 128.04, 128.00, 127.96, 127.88, 127.78, 127.68, 100.43 (d), 79.05, 78.98, 75.14, 75.07, 74.96, 74.82, 73.72, 73.56, 73.10, 69.67, 68.89, 68.53, 61.31, 57.13, 34.05, 27.83, 25.07. HRMS m/z + calcd. for C38H43NO7Na (M+Na ) 648.2938, found 648.2931.

Methyl 2-N-methyl-(2-benzylcarboxyamino)acetamido-2-deoxy-3,4,6-tri-O-benzyl-β-D- glucopyranoside (31)

The title compound was prepared by dissolving 29 (250 mg, 0.55 mmol), Cbz-Gly-OH (120 mg, 0.6 mmol) and triethylamine (30 µL, 0.58 mmol) in a 50:50 EtOAc/DCM solution. DCC (140 mg, 0.66 mmol) was then added portion wise to the mixture at 0°C over 30 min and allowed to stir overnight at room temperature. The mixture was then filtered through celite and evaporated under reduced pressure. The resulting residue was then dissolved in EtOAc, washed consecutively with 1M NaOH(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography (1:1 EtOAc/Pentane) afforded 31 1 (268 mg, 92%) as a colourless oil. H NMR (400 MHz, CDCl3; mixture of rotamers) δ 7.45 –

104

7.00 (m, 20H, Ar-H), 4.79 – 4.59 (m, 2H, Ar-CH2), 4.57 – 4.40 (m, 7H, Ar-CH2, H-1), 4.35 –

4.15 (m, 1.5H, COCH2), 3.98 (d, J = 11.9 Hz, 0.5H, COCH2), 3.73 – 3.50 (m, 5H, H-6a, H-6b,

H-5, H-2, H-4), 3.45 – 3.26 (m, 4H, H-3, OCH3), 2.95 (br s, 0.9H, NCH3), 2.71 (s, 2.1H, NCH3). 13 C NMR (75 MHz, CDCl3; mixture of rotamers) δ 170.64, 160.38, 138.22, 137.87, 137.56, 137.54, 137.43, 137.15, 128.23, 128.18, 128.14, 128.10, 128.07, 128.02, 127.97, 127.90, 127.75, 127.72, 127.66, 127.62, 127.58, 127.54, 127.45, 127.36, 127.26, 100.00 (d), 78.62 (d), 74.72, 74.65, 74.54, 74.40 (d), 73.13, 72.68, 69.25, 68.46, 68.11 (d), 56.70, 33.63, 27.40. HRMS m/z + calcd. for C38H43NO7Na (M+Na ) 691.8276, found 691.8270.

Methyl 2-N-methyl-(3-hydroxy)ureayl-2-deoxy-3,4,6-tri-O-benzyl-β-D-glucopyranoside (32)

The title compound was prepared by dissolving 29 (250 mg, 0.55 mmol) and triethylamine (30 µL, 0.58 mmol) in DCM. 4-nitrophenyl carboxamate (210 mg, 0.66 mmol) was then added slowly to the mixture at 0 °C over 30 min and allowed to stir overnight at room temperature. The mixture was washed consecutively with 1M NaOH(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography (2:1 EtOAc/Pentane) 1 afforded 32 (204 mg, 72%) as a colourless oil. H NMR (400 MHz, CDCl3) δ 11.01 (br s, 1H,

OH) 7.62 – 6.91 (m, 15H, Ar-H), 4.95 (d, J = 11.3 Hz, 1H, Ar-CHaHb), 4.84 – 4.73 (m, 2H, H-1,

CHaHb), 4.69 – 4.52 (m, 4H, Ar-CH2, Ar-CH2), 4.31 (d, J = 7.7 Hz, 1H, N-H), 3.79 – 3.42 (m, 13 10H, H-6a, H-6b, H-5, H-4, H-2, H-3, OCH3), 2.54 (s, 3H, N-CH3). C NMR (75 MHz, CDCl3) δ 161.03, 138.65, 138.40, 135.86, 129.55, 129.49, 129.42, 129.34, 129.33, 129.27, 129.19, 129.01, 128.84, 128.80, 128.63, 128.59. 99.88, 79.82, 77.38, 77.11, 73.51, 71.60, 67.66, 58.40, + 54.76, 53.46, 26.10. HRMS m/z calcd. for C30H36N2O7Na (M+Na ) 559.6523, found 559.6531.

105

Methyl 2-N-methyl-(methylsulfonamido)-2-deoxy-3,4,6-tri-O-benzyl-β-D-glucopy-ranoside (33)

The title compound was prepared by dissolving 29 (250 mg, 0.55 mmol) and triethylamine (60 µL, 1.15 mmol) in DCM. MsCl (50 µL, 0.82 mmol) was then added slowly to the mixture at 0 °C over 30 min and allowed to stir overnight at room temperature. The mixture was then washed consecutively with 1M NaOH(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography (1:1 EtOAc/Pentane) afforded 33 (232 mg, 88%) as a colourless oil. 1 H NMR (400 MHz, CDCl3) δ 7.40 – 7.10 (m, 15H, ArH), 4.87 – 4.65 (m, 4H, ArCH2, ArCH2),

4.50 – 4.35 (m, 3H, H-1, ArCH2), 3.65 – 3.53 (m, 6H, H-6a, H-6b, H-5, H-3, H-4, H-2), 3.32 (s, 13 3H, OCH3), 2.98 (s, 3H, SCH3), 2.58 (s, 3H, NCH3). C NMR (75 MHz, CDCl3) δ 137.61, 137.51, 134.72, 128.41, 128.35, 128.28, 128.27, 128.23, 128.18, 128.13, 128.05, 127.94, 127.86, 127.69, 127.66, 127.62, 127.49, 127.44, 99.78, 78.48, 74.86, 74.75, 74.62, 73.21, 68.16, 67.77, + 66.67, 56.41, 31.06, 21.72. HRMS m/z calcd. for C30H37NO7SNa (M+Na ) 578.1120, found 578.1131.

Methyl 2-N-methyl-N-benzylcarboxyaminosulfonamido-2-deoxy-3,4,6-tri-O-benzyl-β-D- glucopyranoside (34)

7.42 – 6.99 (m, 20H, ArH), 5.07 (s, 2H, ArCH2), 4.77 – 4.68 (m, 4H, ArCH2, ArCH2), 4.55 –

4.41 (m, 3H, H-1, ArCH2), 4.26 (br s, 1H, NH), 3.69 – 3.56 (m, 5H, H-6a, H-6b, H-5, H-3, H-4), 13 3.32 (m, 4H, OCH3. H-2), 2.74 (s, 3H, NCH3). C NMR (100 MHz, CDCl3) δ 151.55, 137.98, 137.88, 137.63, 136.32, 135.09, 128.78, 128.72, 128.65, 128.64, 128.60, 128.56, 128.50, 128.42,

106

128.31, 128.24, 128.07, 128.03, 127.99, 127.86, 127.82, 100.16, 78.86, 75.23, 75.13, 75.00, + 73.58, 68.54, 68.14, 67.04, 56.78, 22.86. HRMS m/z calcd. for C37H46N3O9S (M+NH4 ) 708.2922, found 708.2928.

Methyl 2-N-methyl-(2-hydroxy)acetamido-2-deoxy-β-D-glucopyranoside (35)

The title compound was prepared by dissolving 30 (200 mg, 0.41 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate was evaporated under reduced 1 pressure to afford 35 (68 mg, 82%) as a white powder. H NMR (400 MHz, D2O; mixture of rotamers) δ 4.89 (d, J = 7.9 Hz, 1H, H-1) 4.39 (br d, J = 5.2 Hz, 2H, CH2), 4.03 – 3.71 (m, 3H,

H-6a, H-6b, H-5), 3.58 – 3.48 (br m, 5H, H-4, H-3, CH3), 3.34 – 3.23 (br m, 1H, H-2), 2.95 (br s, 13 3H, NCH3). C NMR (100 MHz, D2O; mixture of rotamers) δ 175.00, 99.86, 76.03 -75.98 (d), 70.69 - 70.58 (d), 70.36, 61.74, 60.93-60.87 (d), 60.24 - 60.14 (d) 57.48 – 57.36 (d), 27.97. + HRMS m/z calcd. For C10H19NO7Na (M+Na ) 288.1223, found 288.1222

Methyl 2-N-methyl-(2-amino)acetamido-2-deoxy-β-D-glucopyranoside (36)

The title compound was prepared by dissolving 31 (200 mg, 0.41 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate evaporated under reduced pressure 1 to afford 36 (64 mg, 78%) as a white powder. H NMR (400 MHz, CD3OD; mixture of rotamers) δ 4.50 (br d, J = 7.5 Hz, 1H, H-1), 4.00 – 3.76 (br m, 3H, H-6a, H-6b, H-5), 3.63 (br m, 2H,

CH2), 3.52 – 3.18 (br m, 6H, OCH3, H-4. H-2, H-3), 2.95 (br s, 1.2H, NCH3), 2.80 (br s, 1.8H, 13 NCH3). C NMR (100 MHz, CD3OD; mixture of rotamers) δ 172.47, 101.07 - 100.99 (d), 77.69 – 77.56 (d), 72.13, 63.37, 62.50 – 62.42 (d), 57.04, 43.10, 28.09. HRMS m/z calcd. For + C10H21N2O6 (M+H ) 265.1393, found 265.1394

107

Methyl 2-N-methyl-(3-hydroxy)ureayl-2-deoxy-β-D-glucopyranoside (37)

The title compound was prepared by dissolving 32 (200 mg, 0.38 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate evaporated under reduced pressure 1 to afford 37 (54 mg, 75%) as a white powder. H NMR (400 MHz, D2O) δ 4.75 (d, J = 7.3 Hz, 1H, H-1), 3.93 (dd, J = 11.5 Hz, 4.3 Hz, 1H. H-6a), 3.86 (m, 1H, H-5), 3.74 (dd, J = 11.1 Hz,

7.1 Hz, 1H, H-6b), 3.60 (s, 3H, OCH3), 3.50 – 3.40 (m, 3H, H-4, H-2, H-3), 2.62 (s, 3H, NCH3). 13 C NMR (75 MHz, D2O) δ 161.43, 100.33, 73.97, 72.05, 68.12, 58.86, 55.21, 53.91, 25.26. + HRMS m/z calcd. for C8H16N2O7Na (M+Na ) 289.0735, found 289.0747

Methyl 2-N-methyl-(methylsulfonamido)-2-deoxy-β-D-glucopyranoside (38)

The title compound was prepared by dissolving 33 (200 mg, 0.38 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10% Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate evaporated under reduced pressure to afford 38 1 (71 mg, 75%) as a colourless paste. H NMR (400 MHz, D2O) δ 4.67 (d, J = 8.4 Hz, 1H, H-1), 3.92 (dd, J = 11.3 Hz, 1.9 Hz, 1H, H-6a), 3.81 (m, 1H, H-5) 3.69 (dd, J = 11.4, 5.8 Hz, 1H, H-

6b), 3.59 (s, 3H, OCH3), 3.46 – 3.34 (m, 3H, H-2, H-3, H-4), 3.11 (s, 3H, SCH3), 2.74 (s, 3H, 13 NCH3). C NMR (100 MHz, D2O) δ 101.98, 76.34, 72.58, 69.72, 60.24, 57.58, 55.12, 40.21, + 23.31. HRMS m/z calcd. for C8H17NO7SNa (M+Na ) 289.0859, found 289.0851

108

Methyl 2-N-methyl-N-aminosulfonamido-2-deoxy-β-D-glucopyranoside (39)

The title compound was prepared by dissolving 34 (200 mg, 0.42 mmol) in MeOH (5 mL), followed by the addition of a catylitic amount of 10% Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate evaporated under 1 reduced pressure to afford 39 (74 mg, 75%) as a colourless oil. H NMR (400 MHz, D2O) δ 4.61 (d, J = 8.1 Hz, 1H, H-1) 3.93 (dd, J = 7.8 Hz, 4.6 Hz, 1H, H-6a), 3.88 – 3.84 (m, 1H, H-5), 3.78

– 3.71 (m, 1H, H-6b), 3.60 (s, 3H, OCH3), 3.48 – 3.44 (m, 3H, H-4, H-2, H-3), 2.80 (s, 3H, 13 NCH3). C NMR (100 MHz, D2O) δ 99.69, 75.60, 70.51, 70.32, 63.12, 60.63, 56.76, 21.46. + HRMS m/z calcd. for C8H18N2O7NaS (M+Na ) 309.0726, found 309.0726

Methyl 2-benzylcarboxyaminosulfonamido-2-deoxy-3,4,6-tri-O-benzyl-β-D- glucopyranoside (41)

The title compound was prepared by first dissolving chlorosufonyl isocyanate (0.80 µL, 0.75 mmol) and benzyl alcohol (95 µL, 0.75 mmol) in DCM, and allowed to stir for 30 min at 0 oC. Following this, the reaction mixture was combined with a DCM solution containing 40 (240 mg, 0.55 mmol) and triethylamine (60 uL, 1.2 mmol), and allowed to stir for 3 h. The mixture was then washed consecutively with 1M NaOH(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography (1:1 1 EtOAc/Pentane) afforded 41 (232 mg, 88%) as a colourless oil. H NMR (400 MHz, CDCl3) δ 7.40 – 7.04 (m, 20H), 4.88 – 4.61 (m, 3H), 4.60 – 4.37 (m, 6H), 4.35 – 4.15 (m, 2H), 3.73 – 3.55 13 (m, 3H), 3.48 – 3.30 (m, 4H), C NMR (100 MHz, CDCl3) δ 156.78, 151.43, 137.87, 137.77, 137.52, 136.20, 134.97, 128.67, 128.61, 128.54, 128.53, 128.48, 128.44, 128.38, 128.30, 128.20, 128.12, 127.95, 127.91, 127.88, 127.74, 127.70, 100.04, 78.74, 75.12, 75.01, 74.88, 73.46, 68.42, + 68.02, 66.92, 56.66. HRMS m/z calcd. for C37H42N2O9S (M+Na ) 699.2385, found 694.2388.

109

Methyl 2-aminosulfonamido-2-deoxy-β-D-glucopyranoside (42)

The title compound was prepared by dissolving 41 (200 mg, 0.42 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10% Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate evaporated under reduced pressure to afford 42 (74 mg, 75%) as a colourless oil. 1H NMR (400

MHz, D2O) δ 4.53 (d, J = 8.4 Hz, 1H, H-1), 3.78 (dd, J = 12.3 Hz, 2.0 Hz, 1H H-6a ), 3.59 - 13 3.54 (m, 2H H-6b, H-5), 3.47 – 3.25 (m, 6H, H-4, H-2, H-3, CH3) C NMR (100 MHz, D2O) δ + 102.65, 76.60, 74.51, 70.32, 60.12, 57.63, 56.76. HRMS m/z calcd. for C8H18N2O7NaS (M+Na ) 294.0825, found 294.0820

Methyl 2-methylamino 2-deoxy-β-D-glucopyranoside (43)

The title compound was prepared by dissolving 29 (200 mg, 0.41 mmol) in aldehyde free MeOH (5 mL) followed by the addition of a catalytic amount of 10% Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate evaporated under reduced 1 pressure to afford 43 (64 mg, 78%) as a colorless film. H NMR (400 MHz, D2O) δ 4.47 (d, J = 8.5 Hz, 1H, H-1), 3.77 (dd, J = 12.6 Hz, 4.3 Hz, 1H, H-6a), 3.60 (dd, J = 12.4 Hz, 6.4 Hz, 1H,

H-6b), 3.50 (dd, J = 10.6, 8.4 Hz, 1H, H-3), 3.43 (s, 3H, OCH3), 3.37 – 3.27 (m, 2H, H-5, H-4), 13 2.84 (dd, J = 10.6 Hz, 8.5 Hz, 1H, H-2), 2.51 (s, 3H, NCH3). C NMR (100 MHz, D2O) δ + 100.27, 76.34, 72.38, 69.93, 60.60, 57.58, 55.94, 23.34. HRMS m/z calcd. for C8H19NO5 (M+H ) 206.1014, found 206.1022

110

Methyl 2-N-methyl-(2-thioacetyl)acetamido-2-deoxy-β-D-glucopyranoside (44)

The title compound was prepared by dissolving compound 43 (50 mg, 0.23 mmol) and triethylamine (12 µL, 0.25 mmol) in DMF. Pentafluorophenyl 2-(acetylthio)acetate (75 mg, 0.23 mmol) was then added to the mixture and allowed to stir overnight at room temperature. The mixture was evaporated under reduced pressure. The resulting residue was then purified by flash chromatography (8% MeOH in DCM) to afford 44 (35 mg, 69%) as a 1 white solid. H NMR (400 MHz, D2O) δ 4.72 (d, J = 8.3 Hz, 1H, H-1), 3.94 (dd, J = 12.2 Hz,

1.6 Hz, 1H, H-6a), 3.74 – 3.44 (m, 9H, H-6b, H-5, H-2, CH2, H-4, H-3, OCH3), 2.78 (s, 3H, N- 13 CH3), 2.32 (s, 3H, CH3). C NMR (100 MHz, D2O) δ 197.04, 169.27, 99.70, 73.61, 71.34, + 67.72, 58.47, 54.97, 53.68, 30.80, 29.09, 27.23. HRMS m/z calcd. for C11H19NO7SNa (M+Na ) 323.0978, found 323.0981

Methyl 3,4,6-tri-O-benzyl-2-deoxy-2-C-[bis-(methoxycarbonyl)methyl]-β-D- glucopyranoside (52a) The title compound was synthesized in two steps following a literature procedure.148 D-glucal (1 g, 7.5 mmol) was dissolved in DMF (150 mL) in the presence of BnBr (5.8 g, 0.0340 mol) and cooled to 0 ºC. Sodium hydride (0.55 g, 0.024 mol) was then slowly added and the solution allowed to rise to room temperature and stirred overnight. The mixture was then diluted with

EtoAc and subsequently washed with 1 M NaOH(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure, yielded per-O-benzylated D-glucal which was then dissolved in MeOH (200 mL) containing NaHCO3. Dimethyl malonate (9.5 mL, 0.075 mol) and cerium ammonium nitrate (8 g, 0.015 mol) and stirred for 3 hours. The resulting solution was neutralized with water and evaporated to dryness. The resulting suspension was dissolved in

EtOAc and washed with 1 M HCl(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure, followed by flash chromatography yielded 52a (48% over 2 steps, 1.89 g) as a clear oil. All acquired spectral data were in agreement with literature values.148

111

Methyl 3,4,6-tri-O-benzyl-2-deoxy-2-C-[malonic acid]-β-D-glucopyranoside (53) The title compound was prepared by dissolving 52a (0.2 g, 0.35 mmol) in a 1:1 solution of 2M NaOH/dioxane and stirred overnight at room temperature. The solution was then diluted with DCM and washed with

1M HCl(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure yielded 53 (86%, 0.16 g) as a clear oil. All acquired spectral data were in agreement with literature values.148

Methyl 2-deoxy-2-C-[malonic acid]-β-D-glucopyranoside (54) The title compound was prepared by dissolving 53 (0.1 g, 0.18 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate was evaporated under reduced pressure to afford 54 (35 mg, 77%) as a white powder. All acquired spectral data were in agreement with literature values.148

Methyl 3,4,6-tri-O-benzyl-2-deoxy-2-C-[1,1,1,5,5,5-hexafluoropentane-2,4-dione]-β-D- glucopyranoside (55) The title compound was prepared by dissolving 52a (0.2 g, 0.35mmol) in ethylene glycol dimethyl ether (5 mL) in the presence of TMS-CF3 (0.55 mL, 3.5 mmol) and a catalytic amount of cesium fluoride and allowed to stir overnight at room temperature. The solution was then diluted with

DCM, and washed consecutively with HCl(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure followed by flash chromatography (8:1 pentane/EtOAc) 1 yielded 55 (71%, 0.14 g,) as a colorless oil. H NMR (400 MHz, D2O) δ 4.61 (d, J = 8.2 Hz, 1H,

H-1), 3.96 – 3.40 (m, 5H, H-6b, H-6a, H-5, H-4, H-3), 3.31 (s, 3H, OCH3), 3.13 (m, 1H, H-7), + 2.34 (m, 1H, H-2). HRMS m/z calcd. for C33H32F6O7Na (M+Na ) 677.2093, found 677.2101.

112

Methyl 2-deoxy-2-C-[1,1,1,5,5,5-hexafluoropentane-2,4-dione]-β-D-glucopyranoside (56) The title compound was prepared by dissolving 53 (0.12 g, 0.18 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate was evaporated under reduced pressure to afford 54 (42 1 mg, 72%) as a white powder. H NMR (400 MHz, CDCl3) δ 7.42 – 7.20 (m, 15H, ArH), 4.95 (m,

2H, ArCH2) 4.70– 4.52 (m, 5H, ArCH2, ArCH2, H-1), 4.21 (m, 1H, H6a) 4.09 (m, 1H, H-6b), 13 3.90-3.70 (m, 3H, H-5, H-3, H-4), 3.51 (s, 3H, OCH3, 3.35 (m, 1H, H-7) 2.35 (m, 1H, H-2). C

NMR (100 MHz, D2O) δ 160.70, 160.41, 119.20, 118.75 103.42, 76.31, 74.92, 70.18, 61.14, 19 57.87, 56.12, 42.21. F NMR (376 MHz, D2O) δ -75.2 (2) (COCF3), -84.9 (2) (C(OH)2CF3) + HRMS m/z calcd. for C12H14F6O7Na (M+Na ) 407.0601, found 407.0605.

Methyl 3,4,6-tri-O-benzyl-2-deoxy-2-C-[(methoxycarbonyl)methyl]-β-D-glucopyranoside (57) The title compound was prepared following a procedure developed by Yin et al148. Briefly, 52a (1 g, 1.7 mmol) was dissolved in DMSO (10 mL) in the presence of excess lithium iodide (2.2 g, 17 mmol) and submitted to microwave irradiation (200 W, 10 bar, 100 ºC, 20 min). The solution was then diluted with EtOAc and washed 5 times with water, followed by brine and drying with magnesium sulphate, and evaporated to dryness. Flash chromatography of the resulting mixture (10:1 pentane/EtOAc) yielded 57 (62%, 0.51 g) as a colorless oil. All acquired spectral data were in agreement with literature values.148

Methyl 3,4,6-tri-O-benzyl-2-deoxy-2-C-[acetic acid]-β-D-glucopyranoside (58) The title compound was prepared by dissolving 57 (0.15 g, 0.28 mmol) in a 1:1 solution of 2M NaOH/dioxane and stirred overnight at room temperature. The solution was then diluted with DCM and washed with

1M HCl(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure yielded 53 (82%, 0.11 g) as a clear oil. All acquired spectral data were in agreement with literature values.148

113

Methyl 2-deoxy-2-C-[acetic acid]-β-D-glucopyranoside (59) The title compound was prepared by dissolving 58 (0.1 g, 0.2 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate was evaporated under reduced pressure to afford 54 (72%, 30 mg) as a white powder. All acquired spectral data were in agreement with literature values.148

Methyl 3,4,6-tri-O-benzyl-2-deoxy-2-C-[1,1,1-trifluoropropan-2-one]-β-D-glucopyranoside (60) The title compound was prepared by dissolving 57 (0.15 g, 0.28 mmol) in ethylene glycol dimethyl ether (5 mL) in the presence of TMS-CF3 (0.26 ml, 0.175 mmol) and a catalytic amount of cesium fluoride and allowed to stir overnight at room temperature. The solution was then diluted with

DCM, and washed consecutively with HCl(aq), water, and brine, dried with magnesium sulphate. Evaporation under reduced pressure followed by flash chromatography (8:1 pentane/EtOAc) 1 yielded 55 (64%, 96 mg ) as a colorless oil. H NMR (400 MHz, CDCl3) δ 7.42 – 7.10 (m, 15H,

ArH), 4.95 – 4.60 (m, 6H, ArCH2), 4.4 (d, 1H, H-1, J = 8.1 Hz), 3.75 – 3.40 (m, 8H, H-6a, H-6b,

H-5, H-3, H-4, OCH3), 2.50 (m, 2H, CH2) 2.18 (m, 1H, H-2) HRMS m/z calcd. for + C31H33F3O6Na (M+Na ) 581.2219, found 581.2213.

Methyl 2-deoxy-2-C-[1,1,1-trifluoropropan-2-one]-β-D-glucopyranoside (61) The title compound was prepared by dissolving 53 (75 mg, 0.13 mmol) in MeOH (5 mL), followed by the addition of a catalytic amount of 10%

Pd/C. The resulting suspension was stirred vigorously under H2 gas (1 atm) for 12 h. Upon reaction completion, the suspension was filtered through celite, and the resulting filtrate was evaporated under reduced pressure to afford 54 (81%, 30 mg) 1 as a white powder (ketone/hydrate 26:1). H NMR (400 MHz, D2O) δ 4.40 (d, J = 8.3 Hz, 1H, H- 1), 3.94 (dd, J = 12.2 Hz, 1.6 Hz, 1H, H-6a), 3.71 (dd, J = 12.2 Hz, 6.6 Hz, 1H, H-6b), 3.61 (dd,

114

J = 9.4 Hz, 8.7 Hz, 1H, H-3), 3.52 (s, 3H, OCH3), 3.42 (m, 2H, H-4. H-5), 2.55 (m, 2H, CH2), 13 2.19 (m, 1H, H-2). C NMR (100 MHz, D2O) δ 162.33, 119.4, 100.42, 75.3, 71.24, 68.80, 19 57.40, 54.81, 49.68, 35.02. F NMR (376 MHz, D2O) δ -74.2 (COCF3), -84.4 (C(OH)2CF3) + HRMS m/z calcd. for C10H15F3O6Na (M+Na ) 311.0713, found 311.0720.

GlcNAc-GlcNAc-GlcNAcSOc-GlcNAc-GlcNAc (63)

The title compound was prepared by dissolving GlcNAc pentasaccharide (50 mg, 0.045 mmol) in HEPES buffer (50 mM, pH 7.5) to a final concentration of 10 mM, followed by the addition of MBP-PgaB to a final concentration of 50 µM. The solution was then incubated at 37 °C for 48 h. Desalting of the de-N-acetylated pentasaccharide on HPLC p2 size exclusion column yielded a 1:1 mixture of product and starting material (38 mg), which was used without further purification. The obtained powder (38 mg) was then dissolved in water (0.5 mL), and N-hydroxysuccinidyl 2-(octoylthio)acetate (100 mg, 0.55 mmol), dissolved in MeOH (0.5 mL), was then added to the solution and allowed to mix for 2 h. Upon complete consumption of the starting material, as judged by MALDI mass spectrometry, the reaction mixture was then evaporated under reduced pressure, redissolved in water, washed with EtOAc, and the aqueous layer was then subjected to reverse phase HPLC to isolate 63 (7.2 mg, 0.007 1 mmol) as a white foam. H NMR (400 MHz, D2O) δ 5.17 (d, J = 3.5 Hz, 0.4H. H-1α), 4.62 – 4.51 (m, 4H, H-1‘, H-1‘‘, H-1‘‘‘, H-1‘‘‘‘), 4.25 – 3.37 (m, 34H), 2.17 – 1.99 (m, 12H), 1.58 - + 1.52 (m, 2H), 1.30 - 1.22 (m, 8H), 0.86 – 0.78 (m, 3H) CH3C18H33N2O14S2 (M+Na ) 1214.8043, found 1214.8051

115

GlcNAc-GlcNAc-GlcNAcSH-GlcNAc-GlcNAc (64)

The title compound was prepared by dissolving 63 (7.2 mg, 0.007 mmol) in MeOH (0.5 mL) containing NaOMe (1 µM) for 1 hr. Upon complete consumption of the starting material, as judged by MALDI mass spectrometry, amberlite resin (H+) was added to the mixture until the pH was neutral, and the solution was evaporated under reduced pressure. The residue was then redissolved in AcOH (0.5 M, 0.5 mL), and washed 3 times with EtOAc (0.5 mL). The aqueous phase was then freeze-dried, yielding 64 1 (4.6 mg, 0.005 mmol) as a white foam. H NMR (500 MHz, D2O) δ 5.19 (d, J = 3.5 Hz, 0.4H. H-1α), 4.70 (d, J = 8.6 Hz, 0.6H, H-1β), 4.60 – 4.53 (m, 4H, H-1‘, H-1‘‘, H-1‘‘‘, H-1‘‘‘‘), 4.25 – + 3.37 (m, 32H), 2.17 – 1.99 (m, 12H, CH3). C18H33N2O14S2 (M+Na ) 1088.1461, found 1088.1462

116

References

1. Dunne, W. M., Jr., Bacterial Adhesion: Seen Any Good Biofilms Lately? Clin. Microbiol. Rev. 2002, 15 (2), 155-166.

2. (a) Fux, C. A.; Costerton, J. W.; Stewart, P. S.; Stoodley, P., Survival strategies of infectious biofilms. Trends in Microbiology 2005, 13 (1), 34-40; (b) Cerca, N.; Jefferson, K. K.; Maira-Litran, T.; Pier, D. B.; Kelly-Quintos, C.; Goldmann, D. A.; Azeredo, J.; Pier, G. B., Molecular Basis for Preferential Protective Efficacy of Antibodies Directed to the Poorly Acetylated Form of Staphylococcal Poly-N-Acetyl-{beta}-(1-6)-Glucosamine. Infect. Immun. 2007, 75 (7), 3406-3413.

3. Nikolaev, Y.; Plakunov, V., Biofilm—―City of microbes‖ or an analogue of multicellular organisms? Microbiology 2007, 76 (2), 125-138.

4. (a) Palmer, J.; Flint, S.; Brooks, J., Bacterial cell attachment, the beginning of a biofilm. Journal of Industrial Microbiology and Biotechnology 2007, 34 (9), 577-588; (b) van der Wal, A.; Minor, M.; Norde, W.; Zehnder, A. J. B.; Lyklema, J., Conductivity and Dielectric Dispersion of Gram-Positive Bacterial Cells. Journal of Colloid and Interface Science 1997, 186 (1), 71-79.

5. Patti, J. M.; Allen, B. L.; McGavin, M. J.; Hook, M., MSCRAMM-mediated adherence of microorganisms to host tissues. Annu Rev Microbiol 1994, 48, 585-617.

6. Marraffini, L. A.; Dedent, A. C.; Schneewind, O., Sortases and the art of anchoring proteins to the envelopes of gram-positive bacteria. Microbiol Mol Biol Rev 2006, 70 (1), 192- 221.

7. (a) Tormo, M. Á.; Knecht, E.; Götz, F.; Lasa, I.; Penadés, J. R., Bap-dependent biofilm formation by pathogenic species of Staphylococcus: evidence of horizontal gene transfer? Microbiology 2005, 151 (7), 2465-2475; (b) Latasa, C.; Solano, C.; Penadés, J. R.; Lasa, I., Biofilm-associated proteins. Comptes Rendus Biologies 2006, 329 (11), 849-857.

8. (a) Hengge, R., Principles of c-di-GMP signalling in bacteria. Nat Rev Micro 2009, 7 (4), 263-273; (b) Kolter, R.; Greenberg, E. P., Microbial sciences: The superficial life of microbes. Nature 2006, 441 (7091), 300-302.

9. Stoodley, P.; Sauer, K.; Davies, D. G.; Costerton, J. W., Biofilms as complex differentiated communities. Annu Rev Microbiol 2002, 56, 187-209.

10. Berk, V.; Fong, J. C. N.; Dempsey, G. T.; Develioglu, O. N.; Zhuang, X.; Liphardt, J.; Yildiz, F. H.; Chu, S., Molecular Architecture and Assembly Principles of Vibrio cholerae Biofilms. Science 2012, 337 (6091), 236-239.

11. Kaplan, J. B.; Velliyagounder, K.; Ragunath, C.; Rohde, H.; Mack, D.; Knobloch, J. K.- M.; Ramasubbu, N., Genes Involved in the Synthesis and Degradation of Matrix Polysaccharide in Actinobacillus actinomycetemcomitans and Actinobacillus pleuropneumoniae Biofilms. Journal of bacteriology 2004, 186 (24), 8213-8220.

117

12. Boles, B. R.; Thoendel, M.; Singh, P. K., Rhamnolipids mediate detachment of Pseudomonas aeruginosa from biofilms. Molecular Microbiology 2005, 57 (5), 1210-1223.

13. Rohde, H.; Mack, D.; Christner, M.; Burdelski, C.; Franke, G.; Knobloch, J. K. M., Pathogenesis of staphylococcal device-related infections: from basic science to new diagnostic, therapeutic and prophylactic approaches. Reviews in Medical Microbiology 2006, 17 (2), 45-54 10.1097/01.revmedmi.0000244134.43170.83.

14. (a) Morris, N. S.; Stickler, D. J.; McLean, R. J. C., The development of bacterial biofilms on indwelling urethral catheters. World J Urol 1999, 17 (6), 345-350; (b) Hyde, J. A.; Darouiche, R. O.; Costerton, J. W., Strategies for prophylaxis against prosthetic valve endocarditis: a review article. J Heart Valve Dis 1998, 7 (3), 316-26; (c) Gristina, A. G.; Shibata, Y.; Giridhar, G.; Kreger, A.; Myrvik, Q. N., The glycocalyx, biofilm, microbes, and resistant infection. Seminars in arthroplasty 1994, 5 (4), 160-170.

15. ten Cate, J. M., Biofilms, a new approach to the microbiology of dental plaque. Odontology 2006, 94 (1), 1-9.

16. Bjarnsholt, T.; Jensen, P. Ø.; Fiandaca, M. J.; Pedersen, J.; Hansen, C. R.; Andersen, C. B.; Pressler, T.; Givskov, M.; Høiby, N., Pseudomonas aeruginosa biofilms in the respiratory tract of cystic fibrosis patients. Pediatric Pulmonology 2009, 44 (6), 547-558.

17. Hall-Stoodley, L.; Hu, F.; Gieseke, A.; et al., DIrect detection of bacterial biofilms on the middle-ear mucosa of children with chronic otitis media. JAMA 2006, 296 (2), 202-211.

18. Yakandawala, N.; Gawande, P. V.; LoVetri, K.; Cardona, S. T.; Romeo, T.; Nitz, M.; Madhyastha, S., Characterization of the Poly-β-1,6-N-Acetylglucosamine Polysaccharide Component of Burkholderia Biofilms. Applied and environmental microbiology 2011, 77 (23), 8303-8309.

19. Vinogradov, E.; Sadovskaya, I.; Li, J.; Jabbouri, S., Structural elucidation of the extracellular and cell-wall teichoic acids of Staphylococcus aureus MN8m, a biofilm forming strain. Carbohydr. Res. 2006, 341 (6), 738-743.

20. (a) Darby, C., Uniquely insidious: Yersinia pestis biofilms. Trends Microbiol 2008, 16 (4), 158-64; (b) Wang, X.; Preston, J. F., III; Romeo, T., The pgaABCD Locus of Escherichia coli Promotes the Synthesis of a Polysaccharide Adhesin Required for Biofilm Formation. J. Bacteriol. 2004, 186 (9), 2724-2734.

21. Leipold, M. D.; Herrera, I.; Ornatsky, O.; Baranov, V.; Nitz, M., ICP-MS-Based Multiplex Profiling of Glycoproteins Using Lectins Conjugated to Lanthanide-Chelating Polymers. J. Proteome Res. 2009, 8 (2), 443-449.

22. Sloan, G. P.; Love, C. F.; Sukumar, N.; Mishra, M.; Deora, R., The Bordetella Bps Polysaccharide Is Critical for Biofilm Development in the Mouse Respiratory Tract. Journal of bacteriology 2007, 189 (22), 8270-8276.

118

23. Choi, A. H. K.; Slamti, L.; Avci, F. Y.; Pier, G. B.; Maira-Litrán, T., The pgaABCD Locus of Acinetobacter baumannii Encodes the Production of Poly-β-1-6-N-Acetylglucosamine, Which Is Critical for Biofilm Formation. Journal of bacteriology 2009, 191 (19), 5953-5963.

24. Darby, C., Uniquely insidious: Yersinia pestis biofilms. Trends in Microbiology 2008, 16 (4), 158-164.

25. Izano, E. A.; Sadovskaya, I.; Vinogradov, E.; Mulks, M. H.; Velliyagounder, K.; Ragunath, C.; Kher, W. B.; Ramasubbu, N.; Jabbouri, S.; Perry, M. B.; Kaplan, J. B., Poly-N- acetylglucosamine mediates biofilm formation and antibiotic resistance in Actinobacillus pleuropneumoniae. Microbial Pathogenesis 2007, 43 (1), 1-9.

26. Franklin, M. J.; Nivens, D. E.; Weadge, J. T.; Howell, P. L., Biosynthesis of the Pseudomonas aeruginosa Extracellular Polysaccharides, Alginate, Pel, and Psl. Frontiers in Microbiology 2011, 2.

27. Costerton, J. W.; Stewart, P. S.; Greenberg, E. P., Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284 (5418), 1318-1322.

28. Stewart, P. S.; William Costerton, J., Antibiotic resistance of bacteria in biofilms. The Lancet 2001, 358 (9276), 135-138.

29. Darouiche, R. O.; Mansouri, M. D.; Gawande, P. V.; Madhyastha, S., Antimicrobial and antibiofilm efficacy of triclosan and DispersinB combination. J Antimicrob Chemother 2009, 64 (1), 88-93.

30. Maira-Litrán, T.; Allison, D. G.; Gilbert, P., An evaluation of the potential of the multiple antibiotic resistance operon (mar) and the multidrug efflux pump acrAB to moderate resistance towards ciprofloxacin in Escherichia coli biofilms. Journal of Antimicrobial Chemotherapy 2000, 45 (6), 789-795.

31. Grayson, E. J.; Ward, S. J.; Hall, A. L.; Rendle, P. M.; Gamblin, D. P.; Batsanov, A. S.; Davis, B. G., Glycosyl Disulfides: Novel Glycosylating Reagents with Flexible Aglycon Alteration. J. Org. Chem. 2005, 70 (24), 9740-9754.

32. Stewart, P. S., Theoretical aspects of antibiotic diffusion into microbial biofilms. Antimicrobial Agents and Chemotherapy 1996, 40 (11), 2517-22.

33. Anwar, H.; Strap, J. L.; Costerton, J. W., Establishment of aging biofilms: possible mechanism of bacterial resistance to antimicrobial therapy. Antimicrobial Agents and Chemotherapy 1992, 36 (7), 1347-1351.

34. Bjarnsholt, T.; Ciofu, O.; Molin, S.; Givskov, M.; Hoiby, N., Applying insights from biofilm biology to drug development [mdash] can a new approach be developed? Nat Rev Drug Discov 2013, 12 (10), 791-808.

35. Borriello, G.; Werner, E.; Roe, F.; Kim, A. M.; Ehrlich, G. D.; Stewart, P. S., Oxygen Limitation Contributes to Antibiotic Tolerance of Pseudomonas aeruginosa in Biofilms. Antimicrobial Agents and Chemotherapy 2004, 48 (7), 2659-2664.

119

36. Borriello, G.; Richards, L.; Ehrlich, G. D.; Stewart, P. S., Arginine or Nitrate Enhances Antibiotic Susceptibility of Pseudomonas aeruginosa in Biofilms. Antimicrobial Agents and Chemotherapy 2006, 50 (1), 382-384.

37. Driffield, K.; Miller, K.; Bostock, J. M.; O'Neill, A. J.; Chopra, I., Increased mutability of Pseudomonas aeruginosa in biofilms. Journal of Antimicrobial Chemotherapy 2008, 61 (5), 1053-1056.

38. Hausner, M.; Wuertz, S., High Rates of Conjugation in Bacterial Biofilms as Determined by Quantitative In Situ Analysis. Applied and environmental microbiology 1999, 65 (8), 3710- 3713.

39. Branda, S. S.; Vik, A.; Friedman, L.; Kolter, R., Biofilms: the matrix revisited. Trends in Microbiology 2005, 13 (1), 20-26.

40. Wagstaff, J. L.; Sadovskaya, I.; Vinogradov, E.; Jabbouri, S.; Howard, M. J., Poly-N- acetylglucosamine and poly(glycerol phosphate) teichoic acid identification from staphylococcal biofilm extracts using excitation sculptured TOCSY NMR. Molecular BioSystems 2008, 4 (2), 170-174.

41. Whitchurch, C. B.; Tolker-Nielsen, T.; Ragas, P. C.; Mattick, J. S., Extracellular DNA Required for Bacterial Biofilm Formation. Science 2002, 295 (5559), 1487.

42. Karatan, E.; Watnick, P., Signals, Regulatory Networks, and Materials That Build and Break Bacterial Biofilms. Microbiology and Molecular Biology Reviews 2009, 73 (2), 310-347.

43. Colvin, K. M.; Irie, Y.; Tart, C. S.; Urbano, R.; Whitney, J. C.; Ryder, C.; Howell, P. L.; Wozniak, D. J.; Parsek, M. R., The Pel and Psl polysaccharides provide Pseudomonas aeruginosa structural redundancy within the biofilm matrix. Environ Microbiol 2012, 14 (8), 1913-28.

44. Jackson, K. D.; Starkey, M.; Kremer, S.; Parsek, M. R.; Wozniak, D. J., Identification of psl, a Locus Encoding a Potential Exopolysaccharide That Is Essential for Pseudomonas aeruginosa PAO1 Biofilm Formation. Journal of bacteriology 2004, 186 (14), 4466-4475.

45. Whitfield, C., Biosynthesis and Assembly of Capsular Polysaccharides in Escherichia coli. Annual Review of Biochemistry 2006, 75 (1), 39-68.

46. Hay, I. D.; Ur Rehman, Z.; Ghafoor, A.; Rehm, B. H. A., Bacterial biosynthesis of alginates. Journal of Chemical Technology & Biotechnology 2010, 85 (6), 752-759.

47. Remminghorst, U.; Rehm, B. H. A., In Vitro Alginate Polymerization and the Functional Role of Alg8 in Alginate Production by Pseudomonas aeruginosa. Applied and environmental microbiology 2006, 72 (1), 298-305.

48. Bales, P. M.; Renke, E. M.; May, S. L.; Shen, Y.; Nelson, D. C., Purification and Characterization of Biofilm-Associated EPS Exopolysaccharides from ESKAPE Organisms and Other Pathogens. PLoS ONE 2013, 8 (6), e67950.

120

49. Klemm, D.; Heublein, B.; Fink, H.-P.; Bohn, A., Cellulose: Fascinating Biopolymer and Sustainable Raw Material. ChemInform 2005, 36 (36), no-no.

50. Matthysse, A. G.; Thomas, D. L.; White, A. R., Mechanism of cellulose synthesis in Agrobacterium tumefaciens. Journal of bacteriology 1995, 177 (4), 1076-81.

51. Zogaj, X.; Nimtz, M.; Rohde, M.; Bokranz, W.; Römling, U., The multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Molecular Microbiology 2001, 39 (6), 1452-1463.

52. Gal, M.; Preston, G. M.; Massey, R. C.; Spiers, A. J.; Rainey, P. B., Genes encoding a cellulosic polymer contribute toward the ecological success of Pseudomonas fluorescens SBW25 on plant surfaces. Molecular Ecology 2003, 12 (11), 3109-3121.

53. Ross, P.; Mayer, R.; Benziman, M., Cellulose biosynthesis and function in bacteria. Microbiological Reviews 1991, 55 (1), 35-58.

54. Jackson, D. W.; Suzuki, K.; Oakford, L.; Simecka, J. W.; Hart, M. E.; Romeo, T., Biofilm Formation and Dispersal under the Influence of the Global Regulator CsrA of Escherichia coli. J. Bacteriol. 2002, 184 (1), 290-301.

55. Itoh, Y.; Wang, X.; Hinnebusch, B. J.; Preston, J. F., III; Romeo, T., Depolymerization of {beta}-1,6-N-Acetyl-D-Glucosamine Disrupts the Integrity of Diverse Bacterial Biofilms. J. Bacteriol. 2005, 187 (1), 382-387.

56. Vuong, C.; Kocianova, S.; Voyich, J. M.; Yao, Y.; Fischer, E. R.; DeLeo, F. R.; Otto, M., A Crucial Role for Exopolysaccharide Modification in Bacterial Biofilm Formation, Immune Evasion, and Virulence. Journal of Biological Chemistry 2004, 279 (52), 54881-54886.

57. Steiner, S.; Lori, C.; Boehm, A.; Jenal, U., Allosteric activation of exopolysaccharide synthesis through cyclic di‐GMP‐stimulated protein–protein interaction. The EMBO Journal 2012, 32 (3), 354-368.

58. (a) Izano, E. A.; Sadovskaya, I.; Wang, H.; Vinogradov, E.; Ragunath, C.; Ramasubbu, N.; Jabbouri, S.; Perry, M. B.; Kaplan, J. B., Poly-N-acetylglucosamine mediates biofilm formation and detergent resistance in Aggregatibacter actinomycetemcomitans. Microbial Pathogenesis 2008, 44 (1), 52-60; (b) Kaplan, J. B., Biofilm Dispersal: Mechanisms, Clinical Implications, and Potential Therapeutic Uses. Journal of Dental Research 2010, 89 (3), 205-218.

59. Kirillina, O.; Fetherston, J. D.; Bobrov, A. G.; Abney, J.; Perry, R. D., HmsP, a putative phosphodiesterase, and HmsT, a putative diguanylate cyclase, control Hms-dependent biofilm formation in Yersinia pestis. Molecular Microbiology 2004, 54 (1), 75-88.

60. Mack, D.; Fischer, W.; Krokotsch, A.; Leopold, K.; Hartmann, R.; Egge, H.; Laufs, R., The intercellular adhesin involved in biofilm accumulation of Staphylococcus epidermidis is a linear beta-1,6-linked glucosaminoglycan: purification and structural analysis. J. Bacteriol. 1996, 178 (1), 175-183.

121

61. Maira-Litran, T.; Kropec, A.; Abeygunawardana, C.; Joyce, J.; Mark Iii, G.; Goldmann, D. A.; Pier, G. B., Immunochemical Properties of the Staphylococcal Poly-N-Acetylglucosamine Surface Polysaccharide. Infect. Immun. 2002, 70 (8), 4433-4440.

62. Götz, F., Staphylococcus and biofilms. Molecular Microbiology 2002, 43 (6), 1367-1378.

63. Gerke, C.; Kraft, A.; Süßmuth, R.; Schweitzer, O.; Götz, F., Characterization of theN- Acetylglucosaminyltransferase Activity Involved in the Biosynthesis of the Staphylococcus epidermidisPolysaccharide Intercellular Adhesin. Journal of Biological Chemistry 1998, 273 (29), 18586-18593.

64. Little, D. J.; Poloczek, J.; Whitney, J. C.; Robinson, H.; Nitz, M.; Howell, P. L., The Structure- and Metal-dependent Activity of Escherichia coli PgaB Provides Insight into the Partial De-N-acetylation of Poly-β-1,6-N-acetyl-d-glucosamine. Journal of Biological Chemistry 2012, 287 (37), 31126-31137.

65. (a) Little, D. J.; Whitney, J. C.; Robinson, H.; Yip, P.; Nitz, M.; Howell, P. L., Combining in situ proteolysis and mass spectrometry to crystallize Escherichia coli PgaB. Acta Crystallographica Section F 2012, 68 (7), 842-845; (b) Nishiyama, T.; Noguchi, H.; Yoshida, H.; Park, S.-Y.; Tame, J. R. H., The structure of the deacetylase domain of Escherichia coli PgaB, an enzyme required for biofilm formation: a circularly permuted member of the carbohydrate esterase 4 family. Acta Crystallographica Section D 2013, 69 (1), 44-51.

66. Heilmann, C.; Schweitzer, O.; Gerke, C.; Vanittanakom, N.; Mack, D.; Götz, F., Molecular basis of intercellular adhesion in the biofilm-forming Staphylococcus epidermidis. Molecular Microbiology 1996, 20 (5), 1083-1091.

67. Pokrovskaya, V.; Poloczek, J.; Little, D. J.; Griffiths, H.; Howell, P. L.; Nitz, M., Functional Characterization of Staphylococcus epidermidis IcaB, a De-N-acetylase Important for Biofilm Formation. Biochemistry 2013, 52 (32), 5463-5471.

68. (a) Deng, D. M.; Urch, J. E.; ten Cate, J. M.; Rao, V. A.; van Aalten, D. M. F.; Crielaard, W., Streptococcus mutans SMU.623c Codes for a Functional, Metal-Dependent Polysaccharide Deacetylase That Modulates Interactions with Salivary Agglutinin. Journal of bacteriology 2009, 191 (1), 394-402; (b) Meister, A. W. I. Advances in enzymology and related areas of molecular biology. Volume 56 Volume 56. http://dx.doi.org/10.1002/9780470123027.

69. Potera, C., Forging a Link Between Biofilms and Disease. Science 1999, 283 (5409), 1837-1839.

70. (a) Maira-Litran, T.; Kropec, A.; Goldmann, D.; Pier, G. B., Biologic properties and vaccine potential of the staphylococcal poly-N-acetyl glucosamine surface polysaccharide. Vaccine 2004, 22 (7), 872-879; (b) Maira-Litran, T.; Kropec, A.; Goldmann, D. A.; Pier, G. B., Comparative Opsonic and Protective Activities of Staphylococcus aureus Conjugate Vaccines Containing Native or Deacetylated Staphylococcal Poly-N-Acetyl-{beta}-(1-6)-Glucosamine. Infect. Immun. 2005, 73 (10), 6752-6762.

71. Maira-Litrán, T.; Bentancor, L. V.; Bozkurt-Guzel, C.; O'Malley, J. M.; Cywes-Bentley, C.; Pier, G. B., Synthesis and Evaluation of a Conjugate Vaccine Composed of

122

Staphylococcus aureus Poly-N-Acetyl-Glucosamine and Clumping Factor A. PLoS ONE 2012, 7 (9), e43813.

72. Leung, C.; Chibba, A.; Gomez-Biagi, R. F.; Nitz, M., Efficient synthesis and protein conjugation of Î²-(1â†‘6)-D-N-acetylglucosamine oligosaccharides from the polysaccharide intercellular adhesin. Carbohydr. Res. 2009, 344 (5), 570-575.

73. (a) Gening, M. L.; Tsvetkov, Y. E.; Pier, G. B.; Nifantiev, N. E., Synthesis of [beta]-(1-- >6)-linked glucosamine oligosaccharides corresponding to fragments of the bacterial surface polysaccharide poly-N-acetylglucosamine. Carbohydr. Res. 2007, 342 (3-4), 567-575; (b) Yudina, O. N.; Gening, M. L.; Tsvetkov, Y. E.; Grachev, A. A.; Pier, G. B.; Nifantiev, N. E., Synthesis of five nona-[beta]-(1-->6)-d-glucosamines with various patterns of N-acetylation corresponding to the fragments of exopolysaccharide of Staphylococcus aureus. Carbohydr. Res. 2011, 346 (7), 905-913.

74. Gening, M. L.; Maira-Litrán, T.; Kropec, A.; Skurnik, D.; Grout, M.; Tsvetkov, Y. E.; Nifantiev, N. E.; Pier, G. B., Synthetic β-(1→6)-Linked N-Acetylated and Nonacetylated Oligoglucosamines Used To Produce Conjugate Vaccines for Bacterial Pathogens. Infection and immunity 2010, 78 (2), 764-772.

75. Cywes-Bentley, C.; Skurnik, D.; Zaidi, T.; Roux, D.; DeOliveira, R. B.; Garrett, W. S.; Lu, X.; O‘Malley, J.; Kinzel, K.; Zaidi, T.; Rey, A.; Perrin, C.; Fichorova, R. N.; Kayatani, A. K. K.; Maira-Litràn, T.; Gening, M. L.; Tsvetkov, Y. E.; Nifantiev, N. E.; Bakaletz, L. O.; Pelton, S. I.; Golenbock, D. T.; Pier, G. B., Antibody to a conserved antigenic target is protective against diverse prokaryotic and eukaryotic pathogens. Proceedings of the National Academy of Sciences 2013.

76. Pozzi, C.; Wilk, K.; Lee, J. C.; Gening, M.; Nifantiev, N.; Pier, G. B., Opsonic and protective properties of antibodies raised to conjugate vaccines targeting six Staphylococcus aureus antigens. PLoS ONE 2012, 7 (10), e46648.

77. Khoo, X.; Grinstaff, M. W., Novel infection-resistant surface coatings: A bioengineering approach. MRS Bulletin 2011, 36 (05), 357-366.

78. TEBBS, S. E.; SAWYER, A.; ELLIOTT, T. S. J., Influence of surface morphology on in vitro bacterial adherence to central venous catheters. British Journal of Anaesthesia 1994, 72 (5), 587-591.

79. Gottenbos, B.; van der Mei, H. C.; Klatter, F.; Nieuwenhuis, P.; Busscher, H. J., In vitro and in vivo antimicrobial activity of covalently coupled quaternary ammonium silane coatings on silicone rubber. Biomaterials 2002, 23 (6), 1417-23.

80. (a) Wilson, R. M.; Dowker, S. E. P.; Elliott, J. C., Rietveld refinements and spectroscopic structural studies of a Na-free carbonate apatite made by hydrolysis of monetite. Biomaterials 2006, 27 (27), 4682-4692; (b) Lucke, M.; Schmidmaier, G.; Sadoni, S.; Wildemann, B.; Schiller, R.; Haas, N. P.; Raschke, M., Gentamicin coating of metallic implants reduces implant-related osteomyelitis in rats. Bone 2003, 32 (5), 521-31; (c) Adams, C. S.; Antoci, V., Jr.; Harrison, G.; Patal, P.; Freeman, T. A.; Shapiro, I. M.; Parvizi, J.; Hickok, N. J.; Radin, S.; Ducheyne, P.,

123

Controlled release of vancomycin from thin sol-gel films on implant surfaces successfully controls osteomyelitis. J Orthop Res 2009, 27 (6), 701-9.

81. Kwok, C. S.; Wan, C.; Hendricks, S.; Bryers, J. D.; Horbett, T. A.; Ratner, B. D., Design of infection-resistant antibiotic-releasing polymers: I. Fabrication and formulation. Journal of Controlled Release 1999, 62 (3), 289-299.

82. Anagnostakos, K.; Hitzler, P.; Pape, D.; Kohn, D.; Kelm, J., Persistence of bacterial growth on antibiotic-loaded beads: Is it actually a problem? Acta Orthopaedica 2008, 79 (2), 302-307.

83. Anagnostakos, K.; Schr; #xf6; der, K., Antibiotic-Impregnated Bone Grafts in Orthopaedic and Trauma Surgery: A Systematic Review of the Literature. International Journal of Biomaterials 2012, 2012, 9.

84. Chen, X.; Schluesener, H. J., Nanosilver: A nanoproduct in medical application. Toxicology Letters 2008, 176 (1), 1-12.

85. Allaker, R. P., The Use of Nanoparticles to Control Oral Biofilm Formation. Journal of Dental Research 2010, 89 (11), 1175-1186.

86. Yamanaka, M.; Hara, K.; Kudo, J., Bactericidal Actions of a Silver Ion Solution on Escherichia coli, Studied by Energy-Filtering Transmission Electron Microscopy and Proteomic Analysis. Applied and environmental microbiology 2005, 71 (11), 7589-7593.

87. (a) Kalishwaralal, K.; BarathManiKanth, S.; Pandian, S. R. K.; Deepak, V.; Gurunathan, S., Silver nanoparticles impede the biofilm formation by Pseudomonas aeruginosa and Staphylococcus epidermidis. Colloids and Surfaces B: Biointerfaces 2010, 79 (2), 340-344; (b) Secinti, K. D.; Özalp, H.; Attar, A.; Sargon, M. F., Nanoparticle silver ion coatings inhibit biofilm formation on titanium implants. Journal of Clinical Neuroscience 2011, 18 (3), 391-395.

88. Berra, L.; Kolobow, T.; Laquerriere, P.; Pitts, B.; Bramati, S.; Pohlmann, J.; Marelli, C.; Panzeri, M.; Brambillasca, P.; Villa, F.; Baccarelli, A.; Bouthors, S.; Stelfox, H.; Bigatello, L.; Moss, J.; Pesenti, A., Internally coated endotracheal tubes with silver sulfadiazine in polyurethane to prevent bacterial colonization: a clinical trial. Intensive Care Med 2008, 34 (6), 1030-1037.

89. Qin, Z.; Yang, L.; Qu, D.; Molin, S.; Tolker-Nielsen, T., Pseudomonas aeruginosa extracellular products inhibit staphylococcal growth, and disrupt established biofilms produced by Staphylococcus epidermidis. Microbiology 2009, 155 (7), 2148-2156.

90. Pihl, M.; Davies, J. R.; Chávez de Paz, L. E.; Svensäter, G., Differential effects of Pseudomonas aeruginosa on biofilm formation by different strains of Staphylococcus epidermidis. FEMS Immunology & Medical Microbiology 2010, 59 (3), 439-446.

91. Rendueles, O.; Travier, L.; Latour-Lambert, P.; Fontaine, T.; Magnus, J.; Denamur, E.; Ghigo, J.-M., Screening of Escherichia coli Species Biodiversity Reveals New Biofilm- Associated Antiadhesion Polysaccharides. mBio 2011, 2 (3).

124

92. Rendueles, O.; Kaplan, J. B.; Ghigo, J.-M., Antibiofilm polysaccharides. Environmental Microbiology 2013, 15 (2), 334-346.

93. Kostakioti, M.; Hadjifrangiskou, M.; Hultgren, S. J., Bacterial Biofilms: Development, Dispersal, and Therapeutic Strategies in the Dawn of the Postantibiotic Era. Cold Spring Harbor Perspectives in Medicine 2013, 3 (4).

94. (a) Xavier, J. B.; Picioreanu, C.; Rani, S. A.; van Loosdrecht, M. C. M.; Stewart, P. S., Biofilm-control strategies based on enzymic disruption of the extracellular polymeric substance matrix – a modelling study. Microbiology 2005, 151 (12), 3817-3832; (b) Manuel, S. G. A.; Ragunath, C.; Sait, H. B. R.; Izano, E. A.; Kaplan, J. B.; Ramasubbu, N., Role of active-site residues of dispersin B, a biofilm-releasing β-hexosaminidase from a periodontal pathogen, in substrate hydrolysis. FEBS Journal 2007, 274 (22), 5987-5999.

95. Kaplan, J. B.; LoVetri, K.; Cardona, S. T.; Madhyastha, S.; Sadovskaya, I.; Jabbouri, S.; Izano, E. A., Recombinant human DNase I decreases biofilm and increases antimicrobial susceptibility in staphylococci. J Antibiot (Tokyo) 2012, 65 (2), 73-7.

96. Purushottam V. Gawande, N. Y., Karen LoVetri and Srinivasa Madhyastha, In Vitro Antimicrobial and Antibiofilm Activity of DispersinB®-Triclosan Wound Gel against Chronic Wound-associated Bacteria. The Open Antimicrobial Agents Journal 2011, 3 (1), 12-16.

97. (a) Smith, R. S.; Iglewski, B. H., P. aeruginosa quorum-sensing systems and virulence. Curr Opin Microbiol 2003, 6 (1), 56-60; (b) Withers, H.; Swift, S.; Williams, P., Quorum sensing as an integral component of gene regulatory networks in Gram-negative bacteria. Curr Opin Microbiol 2001, 4 (2), 186-93; (c) Xu, L.; Li, H.; Vuong, C.; Vadyvaloo, V.; Wang, J.; Yao, Y.; Otto, M.; Gao, Q., Role of the luxS Quorum-Sensing System in Biofilm Formation and Virulence of Staphylococcus epidermidis. Infect. Immun. 2006, 74 (1), 488-496; (d) Davies, D. G.; Parsek, M. R.; Pearson, J. P.; Iglewski, B. H.; Costerton, J. W.; Greenberg, E. P., The Involvement of Cell-to-Cell Signals in the Development of a Bacterial Biofilm. Science 1998, 280 (5361), 295-298.

98. (a) Beatson, S. A.; Whitchurch, C. B.; Semmler, A. B. T.; Mattick, J. S., Quorum Sensing Is Not Required for Twitching Motility in Pseudomonas aeruginosa. Journal of bacteriology 2002, 184 (13), 3598-3604; (b) Bjarnsholt, T.; Jensen, P. O.; Burmolle, M.; Hentzer, M.; Haagensen, J. A.; Hougen, H. P.; Calum, H.; Madsen, K. G.; Moser, C.; Molin, S.; Hoiby, N.; Givskov, M., Pseudomonas aeruginosa tolerance to tobramycin, hydrogen peroxide and polymorphonuclear leukocytes is quorum-sensing dependent. Microbiology 2005, 151 (Pt 2), 373-83; (c) Rasmussen, T. B.; Skindersoe, M. E.; Bjarnsholt, T.; Phipps, R. K.; Christensen, K. B.; Jensen, P. O.; Andersen, J. B.; Koch, B.; Larsen, T. O.; Hentzer, M.; Eberl, L.; Hoiby, N.; Givskov, M., Identity and effects of quorum-sensing inhibitors produced by Penicillium species. Microbiology 2005, 151 (Pt 5), 1325-40.

99. Waters, C. M.; Bassler, B. L., Quorum sensing: cell-to-cell communication in bacteria. Annu Rev Cell Dev Biol 2005, 21, 319-46.

100. Geske, G. D.; Wezeman, R. J.; Siegel, A. P.; Blackwell, H. E., Small Molecule Inhibitors of Bacterial Quorum Sensing and Biofilm Formation. J. Am. Chem. Soc. 2005, 127 (37), 12762- 12763.

125

101. (a) Cuong, V.; Jovanka, M. V.; Elizabeth, R. F.; Kevin, R. B.; Adeline, R. W.; Frank, R. D.; Michael, O., Polysaccharide intercellular adhesin (PIA) protects Staphylococcus epidermidis against major components of the human innate immune system. Cellular Microbiology 2004, 6 (3), 269-275; (b) Vuong, C.; Kidder, J. B.; Jacobson, E. R.; Otto, M.; Proctor, R. A.; Somerville, G. A., Staphylococcus epidermidis Polysaccharide Intercellular Adhesin Production Significantly Increases during Tricarboxylic Acid Cycle Stress. J. Bacteriol. 2005, 187 (9), 2967-2973.

102. Kolodkin-Gal, I.; Romero, D.; Cao, S.; Clardy, J.; Kolter, R.; Losick, R., D-amino acids trigger biofilm disassembly. Science 2010, 328 (5978), 627-9.

103. Cava, F.; Lam, H.; de Pedro, M. A.; Waldor, M. K., Emerging knowledge of regulatory roles of D-amino acids in bacteria. Cell Mol Life Sci 2011, 68 (5), 817-31.

104. Hochbaum, A. I.; Kolodkin-Gal, I.; Foulston, L.; Kolter, R.; Aizenberg, J.; Losick, R., Inhibitory Effects of d-Amino Acids on Staphylococcus aureus Biofilm Development. Journal of bacteriology 2011, 193 (20), 5616-5622.

105. Rinehart, K. L.; Holt, T. G.; Fregeau, N. L.; Keifer, P. A.; Wilson, G. R.; Perun, T. J.; Sakai, R.; Thompson, A. G.; Stroh, J. G.; Shield, L. S.; Seigler, D. S.; Li, L. H.; Martin, D. G.; Grimmelikhuijzen, C. J. P.; Gäde, G., Bioactive Compounds from Aquatic and Terrestrial Sources. Journal of Natural Products 1990, 53 (4), 771-792.

106. Rogers, S. A.; Huigens Iii, R. W.; Melander, C., A 2-Aminobenzimidazole That Inhibits and Disperses Gram-Positive Biofilms through a Zinc-Dependent Mechanism. J. Am. Chem. Soc. 2009, 131 (29), 9868-9869.

107. (a) Davies, G.; Henrissat, B., Structures and mechanisms of glycosyl hydrolases. Structure 1995, 3 (9), 853-859; (b) Vocadlo, D. J.; Withers, S. G., Detailed Comparative Analysis of the Catalytic Mechanisms of β-N-Acetylglucosaminidases from Families 3 and 20 of Glycoside Hydrolases. Biochemistry 2005, 44 (38), 12809-12818.

108. Wendeler, M.; Sandhoff, K., Hexosaminidase assays. Glycoconj J 2009, 26 (8), 945-952.

109. LEABACK, D. H.; WALKER, P. G., Studies on glucosaminidase. 4. The fluorimetric assay of N-acetyl-beta-glucosaminidase. Biochem. J. 1961, 78, 151-156.

110. Novak, A.; Lowden, J. A.; Gravel, Y. L.; Wolfe, L. S., Preparation of radiolabeled GM2 and GA2 gangliosides. Journal of Lipid Research 1979, 20 (5), 678-81.

111. Fekete, A.; Borbás, A.; Gyémánt, G.; Kandra, L.; Fazekas, E.; Ramasubbu, N.; Antus, S., Synthesis of β-(1→6)-linked N-acetyl-d-glucosamine oligosaccharide substrates and their hydrolysis by Dispersin B. Carbohydr. Res. 2011, 346 (12), 1445-1453.

112. http://www.kanebiotech.com/dispersinb.html (Accesed Janruary 2013)

113. Kerrigan, J.; Ragunath, C.; Kandra, L.; Gyémánt, G.; Lipták, A.; Jánossy, L.; Kaplan, J.; Ramasubbu, N., Modeling and biochemical analysis of the activity of antibiofilm agent Dispersin B. Acta Biologica Hungarica 2008, 59 (4), 439-451.

126

114. Chibba, A.; Dasgupta, S.; Yakandawala, N.; Madhyastha, S.; Nitz, M., Chromogenic Carbamate and Acetal Substrates for Glycosaminidases. Journal of Carbohydrate Chemistry 2011, 30 (7-9), 549-558.

115. (a) Jungheim, L. N.; Shepherd, T. A., Design of Antitumor Prodrugs: Substrates for Antibody Targeted Enzymes. Chemical Reviews 1994, 94 (6), 1553-1566; (b) Leenders, R. G. G.; Ruytenbeek, R.; Damen, E. W. P.; Scheeren, H. W., Highly Diastereoselective Synthesis of Anomeric Î²-O-Glycopyranosyl Carbamates from Isocyanates. Synthesis 1996, 1996 (11), 1309,1312.

116. Erez, R.; Shabat, D., The azaquinone-methide elimination: comparison study of 1,6- and 1,4-eliminations under physiological conditions. Org. Biomol. Chem. 2008, 6 (15), 2669-2672.

117. (a) Mehta, S.; Pinto, B. M., Novel glycosidation methodology. The use of phenyl selenoglycosides as glycosyl donors and acceptors in oligosaccharide synthesis. The Journal of Organic Chemistry 1993, 58 (12), 3269-3276; (b) PavÃ©, G. g.; LÃ©ger, J.-M.; Jarry, C.; Viaud-Massuard, M.-C.; Guillaumet, G. r., Enantioselective Synthesis of Spirocyclic Aminochroman Derivatives According to the CN(R,S) Strategy. The Journal of Organic Chemistry 2003, 68 (4), 1401-1408.

118. Sarkar, A. K.; Brown, J. R.; Esko, J. D., Synthesis and glycan priming activity of acetylated disaccharides. Carbohydr. Res. 2000, 329 (2), 287-300.

119. Li, S.-C.; Li, Y.-T., Studies on the Glycosidases of Jack Bean Meal. Journal of Biological Chemistry 1970, 245 (19), 5153-5160.

120. Caufrier, F.; Martinou, A.; Dupont, C.; Bouriotis, V., Carbohydrate esterase family 4 enzymes: substrate specificity. Carbohydr. Res. 2003, 338 (7), 687-692.

121. Hernick, M.; Fierke, C. A., Zinc hydrolases: the mechanisms of zinc-dependent deacetylases. Archives of Biochemistry and Biophysics 2005, 433 (1), 71-84.

122. (a) Ravanti, L.; Kähäri, V. M., Matrix metalloproteinases in wound repair (review). International journal of molecular medicine 2000, 6 (4), 391-407; (b) Wang, X.; Li, K. F.; Adams, E.; Van Schepdael, A., Matrix metalloproteinase inhibitors: a review on bioanalytical methods, pharmacokinetics and metabolism. Current drug metabolism 2011, 12 (4), 395-410; (c) Whittaker, M.; Floyd, C. D.; Brown, P.; Gearing, A. J., Design and therapeutic application of matrix metalloproteinase inhibitors. (Chem. Rev. 1999, 99, 2735-2776. Published on the web september 8, 1999). Chem Rev 2001, 101 (7), 2205-6; (d) Skiles, J. W.; Gonnella, N. C.; Jeng, A. Y., The design, structure, and clinical update of small molecular weight matrix metalloproteinase inhibitors. Curr Med Chem 2004, 11 (22), 2911-77.

123. (a) Kapustin, G. V.; Fejer, G.; Gronlund, J. L.; McCafferty, D. G.; Seto, E.; Etzkorn, F. A., Phosphorus-Based SAHA Analogues as Histone Deacetylase Inhibitorsâ€ Org. Lett. 2003, 5 (17), 3053-3056; (b) Suzuki, T.; Nagano, Y.; Kouketsu, A.; Matsuura, A.; Maruyama, S.; Kurotaki, M.; Nakagawa, H.; Miyata, N., Novel Inhibitors of Human Histone Deacetylases: Design, Synthesis, Enzyme Inhibition, and Cancer Cell Growth Inhibition of SAHA-Based Non- hydroxamates. Journal of Medicinal Chemistry 2005, 48 (4), 1019-1032; (c) Vigushin, D. M.;

127

Coombes, R. C., Histone deacetylase inhibitors in cancer treatment. Anticancer Drugs 2002, 13 (1), 1-13.

124. (a) Li, X.; Uchiyama, T.; Raetz, C. R. H.; Hindsgaul, O., Synthesis of a Carbohydrate- Derived Hydroxamic Acid Inhibitor of the Bacterial Enzyme (LpxC) Involved in Lipid A Biosynthesis. Org. Lett. 2003, 5 (4), 539-541; (b) Li, X.; McClerren, A. L.; Raetz, C. R. H.; Hindsgaul, O., Mapping the Active Site of the Bacterial Enzyme LpxC Using Novel Carbohydrate‐Based Hydroxamic Acid Inhibitors*. Journal of Carbohydrate Chemistry 2005, 24 (4-6), 583-609; (c) Blair, D. E.; SchÃ¼ttelkopf, A. W.; MacRae, J. I.; van Aalten, D. M. F., Structure and metal-dependent mechanism of peptidoglycan deacetylase, a streptococcal virulence factor. Proceedings of the National Academy of Sciences of the United States of America 2005, 102 (43), 15429-15434.

125. Coggins, B. E.; McClerren, A. L.; Jiang, L.; Li, X.; Rudolph, J.; Hindsgaul, O.; Raetz, C. R.; Zhou, P., Refined solution structure of the LpxC-TU-514 complex and pKa analysis of an active site histidine: insights into the mechanism and inhibitor design. Biochemistry 2005, 44 (4), 1114-26.

126. Raetz, C. R. H.; Whitfield, C., LIPOPOLYSACCHARIDE ENDOTOXINS. Annual Review of Biochemistry 2002, 71 (1), 635-700.

127. Marks, P. A.; Rifkind, R. A.; Richon, V. M.; Breslow, R.; Miller, T.; Kelly, W. K., Histone deacetylases and cancer: causes and therapies. Nat Rev Cancer 2001, 1 (3), 194-202.

128. (a) Stewart, I.; Schluter, P. J.; Shaw, G. R., Cyanobacterial lipopolysaccharides and human health - a review. Environ Health 2006, 5, 7; (b) Ruiz, N.; Kahne, D.; Silhavy, T. J., Transport of lipopolysaccharide across the cell envelope: the long road of discovery. Nat Rev Microbiol 2009, 7 (9), 677-83.

129. Raetz, C. R. H.; Reynolds, C. M.; Trent, M. S.; Bishop, R. E., Lipid A Modification Systems in Gram-Negative Bacteria. Annual Review of Biochemistry 2007, 76 (1), 295-329.

130. (a) Crossman, A. T.; Urbaniak, M. D.; Ferguson, M. A. J., Synthesis of 1-d-6-O-[2-(N- hydroxyaminocarbonyl)amino-2-deoxy-[alpha]-d-glucopyranosyl]-myo-inositol 1-(n-octadecyl phosphate): a potential metalloenzyme inhibitor of glycosylphosphatidylinositol biosynthesis. Carbohydr. Res. 2008, 343 (9), 1478-1481; (b) Abdelwahab, N. Z.; Urbaniak, M. D.; Ferguson, M. A.; Crossman, A. T., Synthesis of potential metal-binding group compounds to examine the zinc dependency of the GPI de-N-acetylase metalloenzyme in Trypanosoma brucei. Carbohydr Res 2011, 346 (6), 708-14; (c) Abdelwahab, N. Z.; Crossman, A. T.; Sullivan, L.; Ferguson, M. A.; Urbaniak, M. D., Inhibitors incorporating zinc-binding groups target the GlcNAc-PI de-N- acetylase in Trypanosoma brucei, the causative agent of African sleeping sickness. Chem Biol Drug Des 2012, 79 (3), 270-8.

131. (a) Sandler, M. S. H. J., Design of enzyme inhibitors as drugs. Oxford University Press: Oxford [England]; New York, 1989; (b) Schramm, V. L., Enzymatic Transition States, Transition-State Analogs, Dynamics, Thermodynamics, and Lifetimes. Annual Review of Biochemistry 2011, 80 (1), 703-732.

128

132. Hall, R. S.; Brown, S.; Fedorov, A. A.; Fedorov, E. V.; Xu, C.; Babbitt, P. C.; Almo, S. C.; Raushel, F. M., Structural Diversity within the Mononuclear and Binuclear Active Sites of N- Acetyl-d-glucosamine-6-phosphate Deacetylase†,‡. Biochemistry 2007, 46 (27), 7953-7962.

133. (a) Uehara, T.; Park, J. T., The N-Acetyl-d-Glucosamine Kinase of Escherichia coli and Its Role in Murein Recycling. Journal of bacteriology 2004, 186 (21), 7273-7279; (b) Uehara, T.; Suefuji, K.; Valbuena, N.; Meehan, B.; Donegan, M.; Park, J. T., Recycling of the Anhydro- N-Acetylmuramic Acid Derived from Cell Wall Murein Involves a Two-Step Conversion to N- Acetylglucosamine-Phosphate. Journal of bacteriology 2005, 187 (11), 3643-3649; (c) Uehara, T.; Park, J. T., Growth of Escherichia coli: Significance of Peptidoglycan Degradation during Elongation and Septation. Journal of bacteriology 2008, 190 (11), 3914-3922.

134. Xu, C.; Hall, R.; Cummings, J.; Raushel, F. M., Tight Binding Inhibitors of N-Acyl Amino Sugar and N-Acyl Amino Acid Deacetylases. J. Am. Chem. Soc. 2006, 128 (13), 4244- 4245.

135. Gelb, M. H.; Svaren, J. P.; Abeles, R. H., Fluoro ketone inhibitors of hydrolytic enzymes. Biochemistry 1985, 24 (8), 1813-1817.

136. Anslyn, E. V. D. D. A., Modern physical organic chemistry. University Science: Sausalito, CA, 2006.

137. Christianson, D. W.; Lipscomb, W. N., Complex between carboxypeptidase A and a possible transition-state analog: mechanistic inferences from high-resolution x-ray structures of enzyme-inhibitor complexes. J. Am. Chem. Soc. 1986, 108 (16), 4998-5003.

138. Frey, R. R.; Wada, C. K.; Garland, R. B.; Curtin, M. L.; Michaelides, M. R.; Li, J.; Pease, L. J.; Glaser, K. B.; Marcotte, P. A.; Bouska, J. J.; Murphy, S. S.; Davidsen, S. K., Trifluoromethyl ketones as inhibitors of histone deacetylase. Bioorganic & Medicinal Chemistry Letters 2002, 12 (23), 3443-3447.

139. Chibba, A.; Poloczek, J.; Little, D. J.; Howell, P. L.; Nitz, M., Synthesis and evaluation of inhibitors of E. coli PgaB, a polysaccharide de-N-acetylase involved in biofilm formation. Org. Biomol. Chem. 2012, 10 (35), 7103-7107.

140. Gudmundsdottir, A. V.; Nitz, M., Protecting Group Free Glycosidations Using p- Toluenesulfonohydrazide Donors. Org. Lett. 2008, 10 (16), 3461-3463.

141. Fujinaga, M.; Matsushima, Y., Studies on amino-Hexoses. VII. The N-Deacetylation of Methyl N-Acetyl D-Glucosamine. Bulletin of the Chemical Society of Japan 1964, 37 (4), 468- 470.

142. Billing, J. F.; Nilsson, U. J., Cyclic peptides containing a [delta]-sugar amino acid-- synthesis and evaluation as artificial receptors. Tetrahedron 2005, 61 (4), 863-874.

143. Beier, P.; Mindl, J.; Sterba, V.; Hanusek, J., Kinetics and mechanism of base-catalysed degradations of substituted aryl-N-hydroxycarbamates, their N-methyl and N-phenyl analogues. Org. Biomol. Chem. 2004, 2 (4), 562-569.

129

144. Henry, C.; Joly, J.-P.; Chapleur, Y., Efficient Syntheses of Methyl 2-Amino-2-Deoxy- 3,4,6-Tri-O-Benzyl-Î±-D-Glucopyranoside and its 2-tert-Butoxycarbonylamino- and 2- Methylamino Derivatives from N-Acetyl-D-Glucosamine. Journal of Carbohydrate Chemistry 1999, 18 (6), 689-695.

145. Zhang, J.; Matteucci, M. D., Synthesis of a N-acylsulfamide linked dinucleoside and its incorporation into an oligonucleotide. Bioorganic & Medicinal Chemistry Letters 1999, 9 (15), 2213-2216.

146. Gibbs, T. J. K.; Boomhoff, M.; Tomkinson, N. C. O., A Mild and Efficient Method for the One-Pot Monocarboxymethylation of Primary Amines. Synlett 2007, 2007 (10), 1573-1576.

147. Dinh Thanh, N.; Van Quoc, N., Study on Synthesis of 2-(Substituted Benzylidene)amino- 2-Deoxy-1,3,4,6- Tetra-O-Acetyl-β-D-Glucopyranoses from D-Glucosamine. Letters in Organic Chemistry 2013, 10 (2), 85-90.

148. Yin, J.; Sommermann, T.; Linker, T., Convenient Syntheses and Transformations of 2-C- Malonyl Carbohydrates. Chemistry – A European Journal 2007, 13 (36), 10152-10167.

149. Singh, R. P.; Cao, G.; Kirchmeier, R. L.; Shreeve, J. n. M., Cesium Fluoride Catalyzed Trifluoromethylation of Esters, Aldehydes, and Ketones with (Trifluoromethyl)trimethylsilane. The Journal of Organic Chemistry 1999, 64 (8), 2873-2876.

150. Defaye, J.; Gadelle, A.; Pedersen, C., Hydrogen fluoride-catalyzed formation of glycosides. Preparation of methyl 2-acetamido-2-deoxy-[beta]--gluco- and -[beta]--galacto- pyranosides, and of [beta]-(1-->6)-linked 2-acetamido-2-deoxy--gluco- and --galacto-pyranosyl oligosaccharides. Carbohydr. Res. 1989, 186 (2), 177-188.

151. Blair, D. E.; Hekmat, O.; SchÃ¼ttelkopf, A. W.; Shrestha, B.; Tokuyasu, K.; Withers, S. G.; van Aalten, D. M. F., Structure and Mechanism of Chitin Deacetylase from the Fungal Pathogen Colletotrichum lindemuthianumâ€ ,â€¡. Biochemistry 2006, 45 (31), 9416-9426.

152. Udenfriend, S.; Stein, S.; Böhlen, P.; Dairman, W.; Leimgruber, W.; Weigele, M., Fluorescamine: A Reagent for Assay of Amino Acids, Peptides, Proteins, and Primary Amines in the Picomole Range. Science 1972, 178 (4063), 871-872.

153. Bui, N. K.; Turk, S.; Buckenmaier, S.; Stevenson-Jones, F.; Zeuch, B.; Gobec, S.; Vollmer, W., Development of screening assays and discovery of initial inhibitors of pneumococcal peptidoglycan deacetylase PgdA. Biochemical Pharmacology 2011, 82 (1), 43-52.

154. Koller, E.; Wolfbeis, O., Syntheses and spectral properties of longwave absorbing and fluorescing substrates for the direct and continuous kinetic assay of carboxylesterases, phosphatases, and sulfatases. Monatsh Chem 1985, 116 (1), 65-75.

155. Lauffer, B. E. L.; Mintzer, R.; Fong, R.; Mukund, S.; Tam, C.; Zilberleyb, I.; Flicke, B.; Ritscher, A.; Fedorowicz, G.; Vallero, R.; Ortwine, D. F.; Gunzner, J.; Modrusan, Z.; Neumann, L.; Koth, C. M.; Lupardus, P. J.; Kaminker, J. S.; Heise, C. E.; Steiner, P., Histone Deacetylase (HDAC) Inhibitor Kinetic Rate Constants Correlate with Cellular Histone Acetylation but Not Transcription and Cell Viability. Journal of Biological Chemistry 2013, 288 (37), 26926-26943.

130

156. Hu, X.; Zhang, W.; Carmichael, I.; Serianni, A. S., Amide Cis−Trans Isomerization in Aqueous Solutions of Methyl N-Formyl-d-glucosaminides and Methyl N-Acetyl-d- glucosaminides: Chemical Equilibria and Exchange Kinetics. J. Am. Chem. Soc. 2010, 132 (13), 4641-4652.

157. (a) Maris, A., On the conformational equilibrium of glycolamide: A free jet millimetre- wave spectroscopy and computational study. Physical Chemistry Chemical Physics 2004, 6 (10), 2611-2616; (b) Kuhls, K. A.; Centrone, C. A.; Tubergen, M. J., Microwave Spectroscopy of the Twist Cβ-Exo/Cγ-Endo Conformation of Prolinamide. J. Am. Chem. Soc. 1998, 120 (39), 10194- 10198.

158. Fowler, P.; Bernet, B.; Vasella, A., A 1H-NMR Spectroscopic Investigation of the Conformation of the Acetamido Group in Some Derivatives of N-Acetyl-D-allosamine and -D- glucosamine. Helvetica Chimica Acta 1996, 79 (1), 269-287.

159. Winum, J.-Y.; Scozzafava, A.; Montero, J.-L.; Supuran, C. T., Therapeutic potential of sulfamides as enzyme inhibitors. Medicinal Research Reviews 2006, 26 (6), 767-792.

160. Ilies, M.; Dowling, D. P.; Lombardi, P. M.; Christianson, D. W., Synthesis of a new trifluoromethylketone analogue of l-arginine and contrasting inhibitory activity against human arginase I and histone deacetylase 8. Bioorganic & Medicinal Chemistry Letters 2011, 21 (19), 5854-5858.

161. Kelly, C. B.; Mercadante, M. A.; Leadbeater, N. E., Trifluoromethyl ketones: properties, preparation, and application. Chemical Communications 2013, 49 (95), 11133-11148.

162. Caturla, F.; Enjo, J.; Bernabeu, M. C.; Le Serre, S., New fluorescent probes for testing combinatorial catalysts with phosphodiesterase and esterase activities. Tetrahedron 2004, 60 (8), 1903-1911.

163. Hou, Y.; Tse, R.; Mahuran, D. J., Direct Determination of the Substrate Specificity of the α-Active Site in Heterodimeric β-Hexosaminidase A†. Biochemistry 1996, 35 (13), 3963-3969.

164. (a) Mark, B. L.; Mahuran, D. J.; Cherney, M. M.; Zhao, D.; Knapp, S.; James, M. N. G., Crystal Structure of Human β-Hexosaminidase B: Understanding the Molecular Basis of Sandhoff and Tay–Sachs Disease. J. Mol. Biol. 2003, 327 (5), 1093-1109; (b) Specola, N.; Vanier, M. T.; Goutieres, F.; Mikol, J.; Aicardi, J., The juvenile and chronic forms of GM2 gangliosidosis: clinical and enzymatic heterogeneity. Neurology 1990, 40 (1), 145-50.

165. Civallero, G.; Michelin, K.; de Mari, J.; Viapiana, M.; Burin, M.; Coelho, J. C.; Giugliani, R., Twelve different enzyme assays on dried-blood filter paper samples for detection of patients with selected inherited lysosomal storage diseases. Clinica Chimica Acta 2006, 372 (1–2), 98-102.

166. Sutton, V. R., Tay-Sachs disease: Screening and counseling families at risk for metabolic disease. Obstetrics and gynecology clinics of North America 2002, 29 (2), 287-296.

167. Kaback MM, Desnick RJ. Hexosaminidase A Deficiency. 1999 Mar 11 [Updated 2011 Aug 11]. In: Pagon RA, Adam MP, Bird TD, et al., editors. GeneReviews™ [Internet]. Seattle

131

(WA): University of Washington, Seattle; 1993-2014. Available from: http://www.ncbi.nlm.nih.gov/books/NBK1218/

168. Blundell, T. L., Structure-based drug design. Nature 1996, 384 (6604 Suppl), 23-26.

169. Bodewits, K.; Raetz, C. R. H.; Govan, J. R.; Campopiano, D. J., Antimicrobial Activity of CHIR-090, an Inhibitor of Lipopolysaccharide Biosynthesis, against the Burkholderia cepacia Complex. Antimicrobial Agents and Chemotherapy 2010, 54 (8), 3531-3533.

170. Ernst, B.; Magnani, J. L., From carbohydrate leads to glycomimetic drugs. Nat Rev Drug Discov 2009, 8 (8), 661-677.

171. Sliwoski, G.; Kothiwale, S.; Meiler, J.; Lowe, E. W., Computational Methods in Drug Discovery. Pharmacological Reviews 2014, 66 (1), 334-395.

172. Nicolaou, K. C.; Snyder, S. A.; Longbottom, D. A.; Nalbandian, A. Z.; Huang, X., New Uses for the Burgess Reagent in Chemical Synthesis: Methods for the Facile and Stereoselective Formation of Sulfamidates, Glycosylamines, and Sulfamides. Chemistry – A European Journal 2004, 10 (22), 5581-5606.

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Appendix A

Selected NMR spectra

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Appendix B

Lineweaver-Burk and Michaelis-Menten plots

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0.00012

0.00010

0.00008

0.00006

0.00004 Rate (mM/s) Rate

0.00002

0.00000

0 2 4 6 8 10 [ACC] (mM)

PgaB ACC deacetylation rates at various ACC concentrations. Conditions were as follows: HEPES buffer containing 10% (v/v) DMSO (100 mM, pH 7.5), 10 µM PgaB, ACC (0.5 mM – 10 mM). Assays were done in triplicate. Rates were obtained by measuring the increase of fluorescence emission at 446 nM resulting from the acetylosis of ACC. Vmax, KM and kcat were determined from direct curve fitting of the data.

160

Top – Absorbance spectra of 1 (ACC) and 2 (Deacetylated ACC) at 5mM in 100mM HEPES containing 10% (v/v) DMSO. Bottom – Fluorescence spectra of 2 when excited at either 346 or 388.

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Lineweaver Burke plots – All inhibitors showed competitive enzyme inhibition. The value of α was assessed as the factor increase of the slope in the presence of different inhibitor concentrations (slope = αKm/Vmax). Using each obtained α coefficient, a mean

Ki value was derived from the following formula (α = 1 + [I]/Ki).