Thesis Template

Novel Synthesis of Phosphonamidated Carbohydrates Designed as Transition-State Analog Inhibitors of CE4 De-N-acetylases

Rishikesh Ariyakumaran

A thesis submitted in conformity with the requirements for the degree of Master of Science Department of Chemistry University of Toronto

Novel Synthesis of Phosphonamidated Carbohydrates Designed as Transition-State Analog Inhibitors of CE4 De-N-acetylases

Rishikesh Ariyakumaran

Master of Science

Department of Chemistry University of Toronto

2015 Abstract

Carbohydrate Esterase Family 4 enzymes include de-N-acetylases that are essential bacterial virulence factors. They include enzymes that deacetylate PNAG, a glycan composed of repeating units of β-1,6 linked N-acetylglucosamine residues, to form polysaccharide intercellular adhesion (PIA or dPNAG), which is an essential component of the biofilm matrix of many pathogens. They also include de-N-acetylases that act on β-1,4 linked N- acetylglucosamine residues present in bacterial peptidoglycan, which allows evasion of innate immune defenses. To target these enzymes for development of antimicrobial agents, transition- state analog inhibitors containing a phosphonamidate moiety and various specificity groups around a central glucose scaffold were synthesized using a novel direct Staudinger-phosphonite reaction. The inhibitors were tested via in vitro inhibition assays with purified PgaB, the PNAG de-N-acetylase from Escherichia coli, and SpPgdA, the peptidoglycan de-N-acetylase from streptococcus pneumonia. This resulted in identification of the most potent inhibitor of peptidoglycan de-N-acetylases reported to date, with an inhibition constant of 80 µM.

Acknowledgments

I would like to start by first thanking Professor Mark Nitz for accepting me into his lab, both as an undergraduate and graduate student. I would also like to thank him for being a wonderful supervisor, for always having the door open for discussions and ideas, for providing valuable guidance and for helping me learn about the frustrating world of carbohydrate chemistry. Thank you to Professor Drew Woolley for both being a great 4th year undergraduate thesis supervisor and for taking the time to read this thesis. Also, thank you Professor Deborah Zamble for being very helpful and giving me my first opportunity in a research lab as a 2nd year undergraduate summer student.

A special cheer goes out to all the current and past members of the Nitz lab for creating such a wonderful environment to work in every day. A special thanks to Varvara for working with me in this project and to Ben for helping me. Thanks to Matt for both being an awesome friend and, along with all the other members of room 473 (Rohan, Landon, Hanuel and Jason), for all the help with chemistry as well as the lengthy and time-wasting conversations about all sorts of topics (including cricket!). Also thanks to Nesrin, Lisa, Yedi, Peng-peng and adam for creating an engaging and helpful environment. I would also like to thank all the undergraduate students that have passed through the Nitz lab for infusing new blood into the lab. I would also like to thank Andrew Sydor, Landon and Anil for the direct teachings and supervision during my undergraduate research adventures. An addition salutation goes to all the members of the Zamble and Woolley labs for making my time there enjoyable and memorable. I would also like to acknowledge everyone in the Graduate Office, NMR (especially, Darcy and Dmitry) and AIMS facilities and Chem store (especially, Ken) for all their expertise. One of the most important thanks goes to the foosball table in the graduate lounge for all the countless hours of entertainment.

I am most grateful to both my parents for being who they are and for the support, sacrifice and love they have shown while raising me and to my sister for all her help and friendship.

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Table of Contents

Abstract ...... ii Acknowledgments ...... iii Table of Contents ...... iv List of Abbreviations ...... vi List of Tables ...... viii List of Figures ...... ix 1 Introduction ...... 1 1.1 Bacterial Virulence Factors ...... 1 1.1.1 Biofilms...... 1 1.1.2 Cell Wall ...... 4 1.2 Bacterial Glycans ...... 6 1.2.1 Exopolysaccharides in Biofilms ...... 6 1.2.2 Peptidoglycan Modifications ...... 8 1.2.3 CE4 de-N-acetylases: PgaB and SpPgdA ...... 9 1.3 Strategies for Inhibition of CE4 De-N-acetylases ...... 13 1.3.1 Inhibitors of CE4 de-N-acetylases ...... 13 1.3.2 Phosphonamidates as Potential Transition-State Analog Inhibitors ...... 13 1.3.3 Inhibitor Designs ...... 14 1.4 Phosphonamidates ...... 16 1.4.1 Methods of Synthesis and Challenges ...... 16 1.4.2 Strategy for Synthesis of Phosphonamidated Carbohydrates ...... 18 2 Synthesis...... 20 2.1 Monomeric Inhibitors ...... 20 2.1.1 Synthesis of Intermediates ...... 20 2.1.2 Staudinger reaction and Deprotection of Phosphonamidate Esters ...... 23 2.1.3 Synthesis Scheme ...... 31 2.2 Trisaccharide Inhibitor ...... 32 2.3 Discussion and Conclusions ...... 34

3 Enzymology ...... 36 3.1 Protein expression and purification ...... 36 3.1.1 SpPgdA ...... 36 3.1.2 PgaB...... 37 3.2 Procedures for Analysis of Enzyme Kinetics ...... 38 3.2.1 Fluorogenic Substrate Assay ...... 38 3.2.2 Fluorescamine Assay ...... 39 3.2.3 Data Analysis ...... 40 3.3 Results ...... 41 3.4 Discussion and Conclusions ...... 47 Supplementary Information ...... 51 General Procedures ...... 51 Synthesis Procedures ...... 51 NMR Spectra ...... 64 MALDI-MS Analysis ...... 101 Inhibition Curves from Fluorescamine assays with SpPgdA ...... 103 Inhibition Curves from Fluorogenic assays with PgaB ...... 105 References ...... 108

List of Abbreviations

AcOH Acetic Acid

Ac2O Acetic anhydride

ACN acetonitrile

Bn benzyl

NBS N-bromosuccinimide

ACC 3-carboxyumbelliferyl acetate

Cu(OTf)2 Copper(II) trifluoromethanesulfonate

ºC degree Celsius

CDCl3 deuterated chloroform

CD3OD deuterated methanol

D2O deuterated water

DCM dichloromethane

Et2O diethyl ether

DMF dimethylformamide

EDTA Ethylenediaminetetraacetic acid

Hz hertz

Hr hours

HRMS high Resolution Mass Spectrometry

J coupling constant m/z mass per charge

MALDI-MS matrix-assisted laser desorption/ionization-Mass Sprectrometry (or Spectra or Spectrum)

MHz megahertz

Me Methyl vi

Min minutes nm nanometer

Ph phenyl r.t. room temperature

THF tetrahydrofuran

TLC thin layer chromatography p-TSA p-Toluenesulfonic acid

TsH p-Toluenesulfonyl hydrazide

TfN3 Trifluoromethanesulfonyl azide

Et3N Triethylamine

Tris tris(hydroxymethyl)aminomethane

vii

List of Tables

Table 1: Examples of genes involved in peptidoglycan modification in different pathogenic bacterial species and their phenotypic impact. Table adapted from Weiser, N et al. Table 2: Michaelis–Menten kinetic parameters for various de-N-acetylase homologs. Accurate measurements of KM and kcat for PgaB were not reported due to lack of substrate solubility and significantly weak inherent enzyme activity. Table 3: Conditions explored for the regioselective ring-opening of benzylidene acetals in 4, 5 and 11 to synthesize their corresponding 6-O-benzyl derivatives. Table 4: Strategies investigated to obtain pure inhibitors without formation of impurity that has 1 a H NMR chemical shift at 1.94 ppm) or the additional P-CH3 doublet. Table 5: Michaelis-Menten kinetic parameters measured for the various enzyme and substrate combinations used in the study. aValues previously obtained in the lab for Maltose-binding protein-PgaB fusion protein (MBP-PgaB).

Table 6: Ki values in units of µM obtained for inhibitors 15a-f using fluorescamine assay for SpPgdA and fluorogenic substrate assay with ACC for PgaB. aValue obtained using fluorogenic substrate assay with ACC for SpPgdA (data calculated from experiments with inhibitor concentrations ≤ 500 µM).

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List of Figures

Figure 1: Steps involved in biofilm maturation (top) accompanied by photomicrographs of a developing P. aeruginosa biofilm (bottom). Image adapted from Monroe, D. Figure 2: Examples of modifications to peptidoglycan made by bacteria to evade host immune responses. Image adapted from Vollmer, W. Figure 3: Structure of PIA (Top). Pathway responsible for synthesis and secretion of PIA in the Gram-positive bacteria Staphylococcus epidermidis (bottom left) and in the Gram-negative bacteria E. coli (bottom right). Image adapted from Otto, M. et al., Itoh, Y. et al and Otto, M. Figure 4: Structure of E. coli PgaB (left) and structure of SpPgdA (right). The deacetylase activity is catalyzed by the N-terminal domain of PgaB and the C-terminal domain of SpPgdA. Image adapted from Little, D. J. et al. and Blair, D. E. et al. Figure 5: Superposition of deacetylase domains (top left) and active site residues (top right) of PgaB (green) and SpPgdA (magenta) show their strong structural similarities. The major difference is that an aspartic acid (D391) in SpPgdA is replaced by a water molecule in PgaB. Deacetylation in SpPgdA follows an acid/base mechanism (bottom). A similar mechanism is proposed for PgaB. Image adapted from Little, D. J. et al. and Blair, D. E. et al. Figure 6: Structural comparison between the transition-state formed during deacetylation (left) and methylphosphonamidate (right). Figure 7: Structures of transition-state analog Inhibitors containing the phosphonamindate moreity designed for SpPgda (15a-c) and PgaB (15d-f). Figure 8: Surface structures of PgaB (left) and SpPgda (right) displaying the surface exposed hydrophobic residues (green) surrounding the active sites. Image obtained via PDB structures 4F9D (PgaB) and 2C1G (SpPgdA). Figure 9: Structure of trisaccharide inhibitor (21). Figure 10: Synthetic routes developed for obtaining alkyl-phosphonamidates (R =alkyl group). Nucleophilic substitution reaction with phosphonochloridites (Top) and phosphonochloridates (middle) and Staudinger reaction using silylated phosphinic esters (bottom). Figure 11: Mechanism of direct Staudinger-phosphonite reaction by which phosphonamidated carbohydrates were to be synthesized. It is analogous to Staudinger reaction of azides and

phosphines. Hydrolysis of phosphonamidate esters would lead to phosphonamidates. (R, R’= alkyl group). Figure 12: Intermediates required for the synthesis of inhibitors 15a-f. Figure 13: Scheme for the synthesis of intermediates 4 and 5. Intermediate 5 is synthesized from 4 by methylation of the 3’-OH group. Figure 14: Scheme for the synthesis of intermediate 11. Figure 15: Schematic representation of conditions utilized for the regioselective ring-opening of benzylidene acetals in 4, 5 and 11 to give either their 4- or 6-O-benzyl derivatives. R1 = Me or Bn and R2 = OH or Me. Figure 16: Conditions required for Staudinger reaction of azido sugars and diethyl methylphosphonite. Figure 17: 1H-NMR of 15a showing protons fundamental for verifying product formation. Figure 18: 1H-31P-gHMBC spectrum showing the cross peak resulting from correlation between the phosphonamidate phosphorus nuclei and the proton at C2 located three bonds away in inhibitor 15b. Figure 19: 1H-NMR of 15a highlighting the extra peak at 1.94 ppm due to an unknown impurity and the extra P-CH3 doublet signal that appears on occasion. Figure 20: 1H-NMR spectrum of completely purified 15b. Figure 21: Optimized conditions for phosphonamidate ester deprotection. Excess sodium hydroxide and salt are subsequently removed by size-exclusion chromatography. Figure 22: 1H-NMR of 15a taken immediately after (top) and 22 hours after (bottom) being in solution in phosphate buffer under ambient conditions at pH 6.0, 7.5 and 9.0. Minor impurities were present in all samples and their concentrations did not change throughout the 24 hours. Figure 23: Synthesis Schemes A) Scheme for 4,6-O-benzilidine Intermediates utilized for syntehesis of Inhibitors 15a,b,d and e. B) Scheme for 4,6-O-benzilidine Intermediates utilized for syntehesis of Inhibitors 15c and f. C) Scheme for final synthesis of inhibitors. a,d: R1 = Me, R2

= H/Ac, b,e: R1 = Me, R2 = Me, c,f: R1 = Bn, R2 = H/Ac Figure 24: Scheme for synthesis of trisaccharide inhibitor (21). Figure 25: MALDI-MS of 20 (Calculated m/z [M+Na+] = 1051.4). The spectrum also shows the presence of a diphosphonamidate ester (Calculated m/z [M+Na+] = 1115.4).

Figure26: Amino acid sequence of His6-SpPgdA232-431 utilized in this study (Top) and SDS-PAGE gel of purified His6-SpPgdA232-431 (bottom). Red corresponds to amino acid residues 232 to 431 of full-length SpPgdA. Figure 27: Assays used for kinetic analysis of PgaB and SpPgda. ACC substrate becomes fluorescently active upon deacetylation by either PgaB or SpPgdA (TOP). Following deacetylation of chitotriose by SpPgdA, the generated free amine is reacted with fluorescamine to produce a fluorescently active compound (bottom). Figure 28: Michaelis-Menten curve for SpPgdA-catalyzed ACC hydrolysis (left). Note: Assay was performed in 25 mM Tris/HCl, pH 7.0 buffer. SpPgdA-catalyzed hydrolysis of 1 mM ACC in the presence of varying concentrations of inhibitor 15c as measured by the increase in fluorescence due to product formation (right). Figure 29: Hydrolysis of 1 mM ACC in the presence of varying concentrations of inhibitor 15c. Raw rates of ACC hydrolysis by SpPgda without subtracting for autohydrolysis (left). Autohydrolysis rates in samples without any SpPgda (middle). SpPdgA-catalyzed hydrolysis obtained by subtracting fluorescence due to autohydrolysis form raw data (right). Figure 30: Michaelis-Menten curves for SpPgdA-catalyzed ACC hydrolysis at varying concentrations of inhibitor 15c. Figure 31: Michaelis-Menten curve for PgaB-catalyzed ACC hydrolysis. Figure 32: Reaction progress curves for SpPgdA-catalyzed de-N-acetylation at various substrate (chitotriose) concentrations. (left). Michaelis-Menten curve for SpPgdA-catalyzed chitotriose de-N-acetylation (right). Data measured via fluorescamine assays. Figure 33: Michaelis-Menten curves for SpPgdA-catalyzed chitotriose de-N-acetylation in the presence of varying concentrations of inhibitor 15c. Curves were obtained by using the fluorescamine assay.

1 Introduction

1.1 Bacterial Virulence Factors

Pathogenic bacteria represent a huge concern for healthcare. They have evolved numerous mechanisms for evading the host immune system, which lead to serious infectious diseases. Fortunately, antibiotics targeting bacteria have been the most successful form of chemotherapy in human history. Ever since the discovery of sulfonamides and β-lactams in the 1930s, many novel clinically successful classes of antibiotics were discovered during the so called golden era of antibiotic research, which lasted until the early 1970’s. However, since then the rate of antibiotic discovery has significantly fallen off. From the early 1970’s to 2013, only six new classes have been launched for public use and these are only effective against Gram- positive bacteria1. In addition, bacterial resistance mechanisms have rendered many of these common drugs less effective. For example, 40–60% of nosocomial S. aureus strains in the developed world are methicillin-resistant (MRSA) and three other S. aureus strains that display full resistance to vancomycin have been discovered2–4. Certain members of Enterobacteriaceae display resistance to even the newest generation of β-lactams and cephalosporins. A recent study estimates that antimicrobial resistance can currently cause up to $30 billion dollars a year in healthcare costs5. These realities demonstrate the importance of designing new antimicrobial agents which target resistance factors that help bacteria evade either the immune system response or antibiotic treatment.

1.1.1 Biofilms

An important target for the development of original antimicrobials is the pathway utilized by bacteria to form biofilms. A Biofilm is a community of microorganisms adhered to a surface and encapsulated within a matrix of self-produced extracellular polymeric substances (EPS)6. The impact of biofilms cannot be underestimated. The National Institute of Health estimates that 80% of human bacterial infections involve biofilm formation7. Common diseases

such as dental caries and periodontitis as well as persistent tissue infections such as chronic wound infections, chronic osteomyelitis, recurrent urinary tract infections, endocarditis and cystic fibrosis-associated lung infections are associated with biofilms7. Additionally, the high incidence of biofilm formation on artificial devices such as catheters, stents, implants, contact lenses and implantable electronic devices are causing major health concerns. Biofilms have even been shown to play a part in acute infections8.

Many species of bacteria tend to form biofilms and many biofilms are polymicrobial, being composed of more than just one bacterial species. They include Gram-positive pathogens like Staphylococcus epidermidis, Staphylococcus aureus and Streptococcus pneumoniae as well as Gram-negative ones such as Pseudomonas aeruginosa and Escherichia coli. Biofilms confer many advantages to microorganisms since they allow them to temporarily exist in a multicellular state. This allows for survival in fluctuating and hostile environments9.

The encapsulating matrix of a biofilm usually accounts for up to 90% of its biomass. It physically and chemically protects bacteria from dehydration, mechanical stress, predation, oxidizing agents, radiation, and other harmful agents, which include immune defenses and antibiotics9. It allows them to share scarce resources and help contain them in close proximity to one another. This in turn permits exchange of genetic material, which can lead to the spreading of traits, including drug resistance markers and other virulence factors. The presence of environmental gradients within biofilms leads to the formation of subpopulations of bacteria with different metabolic and structural morphologies, which may further confer antibiotic resistance. These include non-dividing persister cells. These bacteria are tolerant to most antibiotics and allow for the reestablishment of biofilms after treatment in clinical settings. This fundamentally leads to the chronic nature of bacterial infections9.

Biofilm formation involves a complex series of steps involving numerous biochemical pathways and interactions (Figure 1) and can be segregated into five stages. When the bacteria encounter a surface, they reversibly adhere to it using a combination of extracellular organelles and proteins. Next, if conditions are suited for biofilm formation, the bacteria attach to the

surface irreversibly through the secretion of EPS. EPS consists of a highly hydrated mixture of polysaccharides, proteins, lipids and DNA. During the third stage, the adsorbed bacteria replicate and grow into microcolonies. They continue the secretion of EPS, which forms a layer of hydrogel that encapsulates and physically separates the growing community from the surrounding environment. Subsequently, the biomass matures into a fully-formed three- dimensional biofilm through further replication, functional cellular differentiation and further accumulation of EPS. The final step involves detachment and dispersal of some bacterial cells into the environment where they propagate and form additional biofilms10.

5. Dispersion

4. Maturation II

3. Maturation I 1. Initial Attachment

2. Irreversible Attachment

Figure 1: Steps involved in biofilm maturation (top) accompanied by photomicrographs of a developing P. aeruginosa biofilm (bottom). Image adapted from Monroe, D.12

The description outlined above is rather simplistic when compared to the detailed molecular changes that actually occur during the biofilm lifecycle. The transformation from planktonic cells to a mature biofilm requires extensive changes in gene regulation and protein expression, cell-surface properties and intercellular communication capabilities. The bacteria must coordinate to constantly remodel the composition of EPS depending on changing temporal, special and environmental conditions. They must form adequate channels that penetrate the entire structure for sufficient nutrient sharing. Additionally, this mechanism can

vary substantially depending on the species under consideration. Even the same species can utilize substantially differing processes under different environmental conditions. Multispecies biofilms can create an even more complex picture. In addition, the key component of biofilms, the EPS, can also differ considerably in terms of structural composition. This complexity and variability makes both studying biofilms as well as designing antibiofilm conjugates extremely difficult. Current biofilm models are too simplistic to mimic actual biofilms and potential universal targets within the biofilm forming pathway are rare and hard to identify11,12. Thus, new tools for inhibiting this process can be extremely valuable for studying this process, in addition to their potential as an antimicrobial agent.

1.1.2 Cell Wall

Another major resistance mechanism employed by bacteria involves modification of their cell wall structure in order to evade host immune responses. A major component of the innate immune system is lysozyme, an enzyme present in most bodily fluids and tissues. It is secreted in large quantities at sites of infection by cells of the immune system13. Lysozyme is a glycosyl hydrolase and catalyzes hydrolysis of β-1,4-linkages between N-acetylmuramic acid and N-acetyl-D-glucosamine residues, which makeup the backbone of cell wall peptidoglycan. This damages the cell wall and leads to bacterial cell death14. To evade recognition by lysozyme, other immune response pathways and antibiotics that target the cell wall, pathogenic bacteria have evolved mechanisms to modify the structure of peptidoglycan (Figure 2)15. These mechanisms utilize unique enzymes (Table 1), which are potential targets for the development of potent antimicrobial agents16.

Figure 2: Examples of modifications to peptidoglycan made by bacteria to evade host immune responses. Image adapted from Vollmer, W.15

Table 3: Examples of genes involved in peptidoglycan modification in different pathogenic bacteria and their phenotypic impact. Table adapted from Weiser, N et al.16

1.2 Bacterial Glycans

1.2.1 Exopolysaccharides in Biofilms

An important aspect of the virulence factors mentioned above is the involvement of glycans. Glycans are the major component of biofilm EPS matrix. They are crucial for adhesion, aggregation of bacterial cells, cohesion of biofilms, retention of water, sorption of organic molecules and inorganic ions, physical protection, binding and stabilization of EPS proteins and energy balance since they act as sinks for excess energy17. Owing to these functions, genetic disruption of EPS glycan synthesis pathways leads to abolition of bacterial biofilm forming phenotypes18.

Exopolysaccharide content in biofilms can vary depending on the species. They can be homopolysaccharides, such as sucrose-derived glucans and fructans. More commonly, they are heteropolysaccharides containing a mixture of neutral and charged residues. Examples of these include alginate, xanthan and colonic acid17. Although this variability is common, a particularly abundant polysaccharide in both Gram-negative and Gram-positive bacteria is polysaccharide intercellular adhesion (PIA or dPNAG for deacetylated poly-N-acetylglucosamine). PIA is a glycan composed of β-(1→6)-N-acetylglucosamine residues, which are partially deacetylated to form a polycationic polymer (Figure 3)19. The percent deacetylation varies depending on the species and strain under consideration20. PIA is produced by important nosocomial pathogens such as Staphylococcus aureus and Staphylococcus epidermidis17. It is found in many other species of Staphylococci, Actinobacillai, Bordetella, and Yersinia, as well as in Escherichia coli, Pseudomonas fluorescens, and Aggregatibacter actinomycetemcomitans21.

PIA is synthesized and exported out of Gram-negative bacterial cells via a synthase- dependent pathway (Figure 3). In E. coli, the pgaABCD locus codes for the required genes. PgaC is an inner transmembrane protein with a glycosyltransferase domain on its cytosolic side. It catalyzes the formation of PNAG from UDP-N-acetylglucosamine (UDP-GlcNAc) and while doing so, it also transports the developing polymer across the inner membrane into the periplasm.

PgaD is an integral inner membrane protein that regulates the activity of PgaC by binding to the bacterial second messenger c-di-GMP. PgaA is a porin located on the outer membrane and is predicted to contain a β-barrel domain that is responsible for exporting PIA into the extracellular matrix. The other member of this locus is PgaB, which is a periplasmic outer membrane enzyme that catalyzes the partial de-N-acetylation of PNAG to form PIA18,22,23. Similar arrangements are also found in other PIA synthesizing bacteria. For example, the Gram (+) pathogen Staphylococcus epidermidis contains an analogous icaABCD locus where IcaA and IcaD produce PNAG from UDP-GlcNAc. IcaC is predicted to be a succinyl transferase and may play a role in PNAG export. IcaB is the de-N-acetylase and is found in the extracellular space24– 26.

A vital part of the above machinery is the de-N-acetylase. The positive charge introduced by partial de-N-acetylation aids in solubilizing the exopolysaccharide while also contributing to its other functions within the matrix. This is further validated by the observation that disruption of E. coli PgaB results in accumulation of PNAG in the periplasm and these bacteria are unable to form biofilms18. Similarly, S. epidermidis IcaB knockouts produce fully acetylated PNAG polymers that are incapable of attaching to the bacterial cell surface. These knockouts cannot form biofilms, colonize host, or resist neutrophil phagocytosis and human antibacterial peptides27.

Figure 3: Structure of PIA (Top). Pathway responsible for synthesis and secretion of PIA in the Gram-positive bacteria S. epidermidis (bottom left) and in the Gram-negative bacteria E. coli (bottom right). Image adapted from Otto, M. et al.23, Itoh, Y. et al.18 and Otto, M.24

1.2.2 Peptidoglycan Modifications

` Similar to the deacetylases involved in biofilm formation, many bacteria utilize deacetylases to modify peptidoglycan. The most common of these are enzymes that catalyze de-N-acetylation of N-acetylglucosamine (GlcNAc) residues in the peptidoglycan backbone. The N-acetyl group is a key motif used by lysozyme to bind to peptidoglycan and de-N-acetylation significantly reduces the affinity for this interaction. In these bacteria, anywhere from 40-88% of these residues are deacetylated15. Peptidoglycan de-N-acetylases are expressed in many pathogens, including Streptococcus pneumoniae, Listeria monocytogenes, Helicobacter pylori and Enterococcus faecalis.16. Bacteria belonging to the group bacillus cereus code for up to five 8

different peptidoglycan de-N-acetylases. Interestingly, in Bacillus anthracis, the causative agent of the highly lethal anthrax disease, it has been shown that amine groups generated by deacetylation are necessary for anchoring the bacterial capsule to its cell wall. Without these enzymes, the capsule, a major virulence factor that is necessary for immune system evasion, does not form properly and the bacteria become susceptible to phagocytosis28. In this case, inhibiting these de-N-acetylases can prevent pathogenesis by blocking both resistance to lysosome and capsule formation.

1.2.3 CE4 de-N-acetylases: PgaB and SpPgdA

Streptococcus pneumoniae peptidoglycan deacetylase A (SpPgdA), a member of peptidoglycan GlcNAc deacetylases, and E.coli PgaB, a member of PNAG deacetylases, are the most structurally and functionally characterized enzymes in the carborhydrate esterase family 4 (CE4) enzyme class. The CE4 enzyme family, classified in the CAZy database, contain other de-N- acetylases including chitin de-N-acetylases and peptidoglycan N-acetylmuramic acid de-N- acetylases29. Like other members of this family, these two deacetylases are metalloenzymes. They contain an active site NodB homology domain, in which the metal ion is coordinated by two conserved histidine and one aspartate residues22,30.

SpPgdA has a three domain structure (Figure 4). The N-terminal domain (residues 46– 160) and the middle domain (residues 161–268) are unique in that they are not found in other enzymes and their functions are unknown. The C-terminus contains the catalytic domain (residues 269–463), which has a fold similar to other members of the CE4 family. The active site metal ion is enclosed in an octahedral coordination environment, where a catalytically important water molecule occupies one of the coordination sites. SpPgdA is most active when bound to Cobalt (II) in the active site followed by Zinc (II), which is the metal ion most likely bound under physiologically relevant conditions30.

Figure 4: Structure of E. coli PgaB (left) and structure of SpPgdA (right). The deacetylase activity is catalyzed by the N-terminal domain of PgaB and the C-terminal domain of SpPgdA. Image adapted from Little, D. J. et al.22 and Blair, D. E. et al. 30

PgaB contains two domains (Figure 4). The N-terminus (residues 42–310) contains the catalytic domain, which displays strong conservation of both fold and active site structure when compared to SpPgdA (Figure 5). The C-terminal domain is structurally similar to many glycoside hydrolases. However, it has not been shown to display any hydrolase activity. The observation that deacetylation does not occur in PgaB without the C-terminal domain suggests that it may be important for substrate binding. Unlike SpPgda, PgaB displays a preference for octahedral coordination around Nickel (II) as opposed to Zinc (II). However, the identity of the metal ion has no effect on protein structure22.

Figure 5: Superposition of deacetylase domains (top left) and active site residues (top right) from PgaB (green) and SpPgdA (magenta) show their strong structural similarities. The major difference being an aspartic acid (D391) in SpPgdA is replaced by a water molecule in PgaB. Deacetylation follows an acid/base mechanism in both enzymes (bottom). Image adapted from Little, D. J. et al.22 and Blair, D. E. et al. 30

Although both SpPgdA and PgaB have significant differences in topology, they do have a highly conserved active site and by extension similar mechanisms have been proposed for both (Figure 5). Binding of substrate involves coordination of active site metal ion by the carbonyl oxygen of the substrate’s acetate moiety. Other hydrophobic interactions, electrostatic interactions and hydrogen bonding confer substrate specificity. Subsequently, a conserved aspartic acid acts as a base to deprotonate the water molecule at the active site. Then, a nucleophilic addition reaction occurs between the newly generated hydroxide ion and the nearby acetate carbon on the substrate to form a tetrahedral oxyanion intermediate. This intermediate is stabilized by metal coordination and electrostatic interactions. A conserved histidine residue acts as an acid and protonates the nitrogen atom. This coincides with bond breaking between this nitrogen atom and the carbonyl carbon, which generates the deacetylated substrate and acetic acid as products22,30. 11

The only major difference between the two active sites is that SpPgdA contains an aspartic acid residue (D391) that is replaced by a water molecule in PgaB. This is proposed to be one of the reasons for why PgaB is the much less efficient enzyme of the two (Table 2), since this residue is believed to be responsible for orienting and modifying the pKa of the catalytic acid (H417) in SpPgdA22. The proposed acid/base mechanism is consistent with all known structures of de-N-acetylases in this family and has been proposed for IcaB, B. subtilis peptidoglycan deacetylase A (BsPdaA) and Vibrio cholera chitin de-N-acetylase (VcCDA)26,31,32. Despite having highly conserved active sites, these de-N-acetylases display striking differences in substrate specificity and activity (Table 2) and the structural basis for this is poorly understood (SpPgdA, like other peptidoglycan deacetylases, can deacetylate β-1,4-linked GlcNAc polymers and does not require other peptidoglycan components for activity).

Table 2: Michaelis–Menten kinetic parameters for various de-N-acetylase homologs. Accurate measurements of KM 22,30,32,33 and kcat for PgaB were not reported due to lack of substrate solubility and weak inherent enzyme activity .

1.3 Strategies for Inhibition of CE4 De-N-acetylases

1.3.1 Inhibitors of CE4 de-N-acetylases

Potent inhibitors of CE4 de-N-acetylases could be tremendously valuable for more than their potential as effective antimicrobial and antibiofilm agents. They can be used to obtain structural and mechanistic details via crystallography, which can help elucidate the reasons for the differing activities and substrate specificities exhibited by these enzymes. This can allow for further engineering of deacetylases, especially chitin deacetylases, for biotechnological purposes and for synthesis of unique polysaccharides. These inhibitors can be utilized for studying the details of biofilm formation in numerous different bacteria. They can be an extremely valuable alternative to genetic methods of disrupting biofilm formation. They can provide temporal information that otherwise cannot be obtained.

Previous attempts to inhibit these de-acetylases have not been highly successful. The strongest inhibitor of PgaB reported to date is a PNAG pentasaccharide in which the N-acetyl group of the middle sugar is replaced by a thioglycolyl amide group, which can coordinate to 34 the active site metal ion. It has an inhibition constant (Ki) of only 280 µM . The only known compounds to inhibit SpPgdA are a set of small molecules, which also display only moderate inhibition35.

1.3.2 Phosphonamidates as Potential Transition-State Analog Inhibitors

To probe the de-N-acetylases, development of inhibitors that mimic the transition-state formed during deacetylation were investigated. This type of approach has been extremely effective for inhibiting a host of enzymes. For example, amino acid sulfonamides and hydrated boronic acid analogues of L-arginine were successfully used to inhibit arginases. They mimicked the tetrahedral intermediate formed during the arginase catalytic process36. Picomolar inhibition constants have been reported for molecules that mimic the charge states present in the early and late transition-states of 5’-methylthioadenosine nucleosidases37.

In these CE4 enzymes, deacetylation proceeds via a tetrahedral oxyanion intermediate. According to Hammond's postulate, the structure of their transition-states should resemble this intermediate. To mimic this state, methylphosphonamidates were chosen. This functionality is ideal since it has the same tetrahedral sp3 configuration, negatively charged oxygen atom and methyl group as the oxyanion transition-state. The fundamental difference is that the acetyl carbon is replaced by phosphorus. This substitution should also make the inhibitor resemble the transition-state more accurately since P-O bonds are longer than C-O bonds. Thus, it is an excellent mimic of a partially formed C-O bond. This structural comparison is shown in Figure 6. The substituents around the central glucose scaffold can be varied to confer specificity for different de-N-acetylases.

Figure 6: Structural comparison between the transition-state formed during deacetylation (left) and methylphosphonamidate (right).

1.3.3 Inhibitor Designs

Inhibitors designed for peptidoglycan and PNAG deacetylases are shown in Figure 7. Benzyl groups at either the 4’- or 6’-hydroxyl groups of the pyranose ring were conceived to be the specificity groups that mimic either the β-1,4- linkage in peptidoglycan or β-1,6- linkage in PNAG, respectively. Inhibitors containing benzyl groups at the 1’ position were also envisioned to further imitate these linkages. These benzyl groups, in addition to mimicking the β-glycosidic bond, provide further affinity for the enzyme. Both deacetylases contain surface exposed hydrophobic pockets surrounding the active site (Figure 8). The phenyl rings can form hydrophobic interactions with these residues. In addition, phenyl groups are acceptable substituents for sugar rings. Crystal structures of carbohydrates bound to enzymes and theoretical calculations show that sugarrings are stabilized by hydrophobic binding pockets 14

made up of aromatic residues. These residues stack against the faces of these sugar rings32,38. Thus, upon binding to these inhibitors, any protein structural changes should naturally place these phenyl groups within hydrophobic pockets, which should significantly increase the affinity for this interaction.

Figure 7: Structures of transition-state analog Inhibitors designed for SpPgda (15a-c) and PgaB (15d-f).

Figure 8: Surface structures of PgaB (left) and SpPgda (right) displaying the surface exposed hydrophobic residues (green) surrounding their active sites (active site metal ions are shown in blue). Image obtained via PDB structures 4F9D (PgaB) and 2C1G (SpPgdA).

Inhibitors containing 3’-Me groups were conceived to probe the role of the 3’-hydroxyl group in the binding process. Both docking studies using crystal structures obtained without the natural substrate bound and mechanistic proposals for SpPgdA, PgaB and IcaB suggest that the

3’-OH group of the sugar being deacetylated may coordinate to the metal ion during binding and throughout the reaction22,26,30. This implies that this hydroxyl group may be a major driving force for substrate binding in this enzyme family. If this is accurate, comparatively these should be much weaker inhibitors and provide experimental evidence for this model.

To further extend this approach, synthesis of a polysaccharide version of the phosphonamidate-containing inhibitor was also designed to target SpPgdA (Figure 9). Its structure is that of a trisaccharide of β 1,4-linked GlcNAc, with the middle N-acetyl group replaced by the methylphosphonamidate moiety (21). This should be a potent inhibitor of SpPgdA since it essentially mimics the transition state of one of its native substrates (β 1,4- linked GlcNAc polymers). The middle residue was chosen to be phosphonamidated since tandem mass spectrometry results have shown that when β-1,4-(GlcNAc)3 is used as a substrate, SpPgdA deacetylates the middle residue first 30.

Figure 9: Structure of trisaccharide inhibitor (21).

1.4 Phosphonamidates

1.4.1 Methods of Synthesis and Challenges

Phosphonamidates and phosphoramidates have gained significant interest in the fields of both modern organic chemistry and chemical biology. These include applications in asymmetric catalysis, bioconjugation chemistry, bioorganic research and pharmaceutical research39–41. Unlike phosphoramidates, synthetic routes for obtaining phosphonamidates are especially challenging due the acid liability of the P-N bonds. The most common methods involve nucleophilic substitution on a protected phosphonochloridate or phosphonochloridite using a free amine. The required phosphonamidates would then be obtained after deprotection, and if necessary an oxidation step. (Figure 10)41–45. These methods have several

drawbacks. They require harsh reaction conditions, synthesis of protected phosphonochloridates or phosphonochloridites as precursors and produce numerous side products that make purification challenging. They are also not compatible in the presence of other nucleophiles, which necessitates the development of complex protecting group chemistry. In addition, these reactions require the presence of a free amine, which cannot be carried through a multistep synthesis without being protected. These drawbacks add more steps to synthesis schemes and may limit their versatility as conditions required for protecting group modifications may be incompatible with other functional groups.

The other reported method for synthesizing phosphonamidates is using a Staudinger reaction involving an azide and a phosphonite. In addition to being a powerful bioorthogonal click reaction for labelling, this chemistry has been used to synthesize phosphonamidate ester- containing proteins, peptides and small organic molecules40,45,46. However, these reactions have not been extended to generate phosphonamidates. More importantly, they have only been shown to work with bulky aryl-phosphonites, which are less prone to oxidation and more stable than alkyl-phosphonites. One reported synthesis of alkyl-phosphonamidates via a Staudinger reaction uses silylated phosphinic esters, and it was employed to synthesize phosphonamidated peptides and small hydrophobic molecules (Figure 10)47. This procedure requires additional steps for synthesis of silylated phosphinic esters. Furthermore, in addition to ester hydrolysis, an extra step is required for deprotection of the silyl functional group.

Figure 10: Synthetic routes developed for obtaining alkyl-phosphonamidates (R =alkyl group). Nucleophilic substitution reaction with phosphonochloridites (Top) and phosphonochloridates (middle) and Staudinger reaction using silylated phosphinic esters (bottom)41–45.

1.4.2 Strategy for Synthesis of Phosphonamidated Carbohydrates

To synthesize phosphonamidates, the direct Staudinger reaction of alkyl-phosphonites and carbohydrates containing an azido functional group were investigated. The proposed mechanism for this reaction is shown in Figure 11. It is analogous to the bioorthogonal Staudinger reaction of azides and phosphines, except instead of hydrolysis of the azaylide intermediate via P-N bond breaking, the P-O bond to one of the OR’ groups breaks resulting in an alcohol leaving group and phosphonamidate ester formation. Subsequent hydrolysis of the ester would give the final phosphonamidate. All the listed methods above have mainly been utilized for synthesizing phosphonamidated peptides and proteins. In addition, carbohydrates containing a phosphonamidate moiety have only been synthesized via the nucleophilic substitution reaction using phosphonochloridates48. If developed, this method could be very useful. The azide functional group can tolerate a range of conditions and is inert in comparison to amines in terms of reactivity. This allows for the ability to carry the azido group throughout a multistep synthetic route and therefore create a large variety of carbohydrates, where a phosphonamidate moiety could be easily installed. The procedure could provide a way to install

different alkyl-phosphonamidate groups at various positions in a multitude of sugars. It could be extended to synthesize phosphonamidated polysaccharides and it can lead to the development of a library of compounds that may have usefulness for a variety of applications, including as inhibitors of enzymes that utilize a mechanism involving tetrahedral oxyanion formation.

Figure 11: Mechanism of direct Staudinger-phosphonite reaction by which phosphonamidated carbohydrates were to be synthesized. It is analogous to Staudinger reaction of azides and phosphines49. Hydrolysis of phosphonamidate esters would lead to phosphonamidates. (R, R’= alkyl group).

2 Synthesis

2.1 Monomeric Inhibitors

In order to synthesize inhibitors 15a-f, three key intermediates were conceived (Figure 12). These intermediates are either methyl or benzyl glycosides containing the azido group required for Staudinger reaction at the 2’ position, either a hydroxyl or methoxy group at the 3’ position and a 4,6-O-benzylidene acetal, which can subsequently be converted into either a 4’ or 6’-O-benzyl group through a regioselective ring-opening reaction. This would produce the six azido sugars necessary for synthesizing the six inhibitors.

Figure 12: Intermediates required for the synthesis of inhibitors 15a-f.

2.1.1 Synthesis of Intermediates

The scheme for synthesis of compound 4 is shown in Figure 13 and is based on previous literature reports50,51. In the first reaction, 2-azido-1-nitrate addition products (2) of tri-O- acetyl-D-glucal (1) are obtained using excess ceric ammonium nitrate and sodium azide in ACN. Methanolic sodium methoxide was utilized to obtain the corresponding fully deacetylated methyl glycosides as a mixture of diastereomers (3). Introduction of a benzylidene acetal to the unpurified mixture using benzaldehyde dimethyl acetal and p-toluenesulfonic acid in DMF, followed by flash chromatography afforded 4 in 21% overall yield over three steps. Compound 5 was synthesized in 96% yield by methylation of 3’-OH of 4 using methyl iodide and sodium hydride in DMF.

Figure 13: Scheme for the synthesis of intermediates 4 and 5. Intermediate 5 is synthesized from 4 by methylation of the 3’-OH group.

Compound 11 was synthesized using the scheme shown in figure 14. First, the hydrazide donor of GlcNAc (2) was synthesized from GlcNAc (6) using p-toluenesulfonyl hydrazide and acetic acid in DMF. Subsequently, the donor (2) was activated with NBS and then glycosylated using benzyl alcohol to afford compound 8 as previously reported52. Compound 8 was chemically deacetylated using hydrazine to afford 9. The amino group in 9 was converted to an azide to form 10 using copper (II) sulfate and triflic azide, which was generated in situ from sodium azide and triflic anhydride as previously reported53. Addition of 4,6-O-benzylidene group to 10 afforded 11 in an overall yield of 28% over five steps.

Figure 14: Scheme for the synthesis of intermediate 11.

With these intermediates in hand, conditions for regioselective ring-opening of benzylidene acetals were investigated. Initial attempts to form the 4-O-benzyl derivatives utilized borane tetrahydrofuran complex as a hydride donor and cobalt (II) chloride as a Lewis acid catalyst. However, this reaction was unsuccessful. Subsequently, another previously

reported procedure, which utilized copper (II) triflate instead of cobalt (II) chloride, was found to be very effective and exclusively produced the required regioisomers in 85-93% yields54. Numerous conditions were explored to synthesize the 6-O-benzyl derivatives. These are summarized in Table 3. The most successful method utilized triethylsilane as a reducing agent and boron trifluoride diethyl etherate as a lewis acid. This reaction mostly produces the 6-O- benzyl regioisomers in 67-76% yields. Figure 15 schematically summarizes these two reaction conditions.

Condition Result

Triethylsilane and Copper (II) Triflate in DCM No reaction

Triethylsilane and p-Toluenesulfonic acid in DCM, -78 ºC Low yield (~25 %)

Reaction does not proceed to Triethylsilane and trifluoroacetic acidtrifluoroacetic acid in DCM completion, low yield (~35%)

Sodium cyanoborohydride and Iodine in CAN low yield (~40%)

Triethylsilane and boron trifluoride diethyl etherate in DCM Moderate yields (67-76%)

Table 3: Conditions explored for the regioselective ring-opening of benzylidene acetals in 4, 5 and 11 to synthesize their corresponding 6-O-benzyl derivatives.

Figure 15: Schematic representation of conditions utilized for regioselective ring-opening of benzylidene acetals in

4, 5 and 11 to give either their 4- or 6-O-benzyl derivatives. R1 = Me or Bn and R2 = OH or Me.

2.1.2 Staudinger reaction and Deprotection of Phosphonamidate Esters

Staudinger reaction was investigated using diethyl methylphosphonite and the synthesized azido sugars. This reaction produces the ethyl ester of methylphosphonamidate, which can subsequently be hydrolyzed to form methylphosphonamidate, and ethanol as a side product. Initially, the reaction was performed in DCM followed by the addition of H2O when the reactant had been fully consumed (monitored by TLC for disappearance of azide). However, the reaction did not produce the expected product and instead formed multiple unidentified adducts. Hypothesizing that free hydroxyl groups may be interfering with the reaction, peracetylation of the sugar followed by Staudinger reaction was attempted. This was successful and produced the expected esters in reasonable yields of 54-78%. In the optimized condition, the azido sugar and 10 equivalences of phosphonite are combined in DCM and stirred at r.t. for

1 hr. Subsequently, H2O is added and the reaction is left stirring overnight. Solvent evaporation and flash chromatography afforded the product as a mixture of two diastereomers due to the phosphorus being chiral (Figure 16).

Figure 16: Conditions required for Staudinger reaction of azido sugars and diethyl methylphosphonite. Hydroxyl groups in the sugar must be protected.

Deprotection of phosphonamidate esters and acetyl protecting groups were initially attempted using sodium hydroxide in a solution of MeOH and water. This reaction was successful in producing the expected product. Figure 17 shows the 1H-NMR spectrum of one of the inhibitors (15a). The methyl protons attached to the phosphorus appear at 1.36 ppm as a doublet with a large coupling constant (16.0 Hz) due to being coupled with phosphorus. The C2 proton peak is also shifted more upfield than others, as would be expected upon phosphonamidation. To further confirm the successfulness of this synthesis, 1H-31P gradient-

selected Heteronuclear Multiple Bond Coherence (gHMBC) experiments were conducted to detect the long-range through-bond coupling between the C2 proton and phosphorus. Figure 18 shows this spectrum for inhibitor 15b. As, expected this coupling produces a cross peak at 2.94 ppm (corresponding to 1D 1H-NMR signal of the C2 proton) and 24.8 ppm (corresponding to 1D 31P-NMR signal of the phosphorus nucleus). As expected, the 1D 31P-NMR of these compounds show only one peak, which corresponds to the lone phosphorus nucleus in the final products (supplementary information).

Figure 17: 1H-NMR of 15a showing protons fundamental for verifying product formation.

Figure 18: 1H-31P-gHMBC spectrum of 15b showing a single cross peak resulting from the correlation between the phosphonamidate phosphorus nucleus and the proton at C2 of the pyranose ring located three bonds away.

Although the reaction was successful, an impurity with a 1H-NMR shift of 1.94 ppm was also produced and it could not be removed by size-exclusion chromatography. Additionally, certain inhibitor fractions contained an extra, more upfield P-CH3 doublet peak in addition to all the other chemical shifts (Figure 19). Postulating that these are due to an acetate salt and migration of the phosphorus to the 3’-OH to form a phosphonate during deacetylation, alternate conditions for purification and deprotection were investigated (Table 4). These conditions were unsuccessful for the most part in removing the impurity. Additionally, the product hydrolyses to the corresponding amine and methylphosphonic acid during any instance where it binds to a resin.

1 Figure 19: H-NMR of 15a highlighting the extra peak at 1.94 ppm due to an unknown impurity and the extra P-CH3 doublet signal that appears on occasion.

To solve these problems, other protecting group strategies were also examined instead of peracetylation. Benzoate and hexanoate protecting groups were utilized due to the possibility that any of their corresponding salts may bind to C18 resin. However, this was not observed and they eluted alongside the product. Additionally, upon deprotection of the hexanoate containing phosphonamidate ester, the same impurity with the 1H-NMR signal at 1.94 ppm was also observed, which suggests that it is due to something other than an acetate salt. The protecting group p-toluenesulfonylcarbamate was also explored, which has been shown to be stable in strong alkaline solutions while being labile under mild alkaline conditions55. This allows selective deprotection of phosphonamidate esters before hydroxyl group deprotection. If successful, this would prevent participation of the 3’-OH in the ester deprotection process and, thus may prevent the phenomenon responsible for the second P-CH3 1H-NMR signal. However, this was unsuccessful, because the final deprotection step produced 26

multiple side products. Removing of these impurities was not possible due to the charged final product’s high polarity, and poor stability under different purification conditions.

Hydroxyl Protecting Groups / Deprotection Conditions Result /Purification Methods

Purification using C18 resin Product/impurity does not bind to resin

HPLC purification Product deterioration/hydrolysis

Purification using activated Product binds to resin but deterioration/hydrolysis upon charcoal resin elution with ACN/MeOH (Only exceptions are inhibitors with a methoxy group at C3 of the sugar ring)

1 THF/H2O Singlet at 1.94 still present / No additional P-CH3 H-NMR signal

Et3N, MeOH, H2O Singlet at 1.94 still present, triethyl ammonium salt forms and is difficult to remove

Benzoate protecting group Cannot separate benzoate salt by size-exclusion chromatography / C18 cartridge

Hexanoate protecting group Singlet at 1.94 still present p-toluenesulfonylcarbamate Deprotection leads to formation of many side Products protecting group

Table 4: Strategies investigated to obtain pure inhibitors without formation of impurity that has a 1H-NMR chemical shift at 1.94 ppm) or the additional P-CH3 doublet.

Interestingly, the inhibitors with the 3’-OMe were able to be completely purified using an activated charcoal cartridge (Figure 20). Additionally, these inhibitors never displayed the 1 additional P-CH3 H-NMR peak. This indicates that the 3’-OH participates in the deprotection

process. Additionally, it may also contribute to the mechanism by which phosphonamidates hydrolyse to amines and phosphonic acids since the 3’-OMe derivatives are more stable towards purification by activated charcoal resin.

Figure 20: 1H-NMR spectrum of completely purified 15b.

Of all the conditions investigated, the best method was peracetylation followed by deprotection using sodium hydroxide in a 1:1 solution of THF:H2O (Figure 21). Under these conditions, formation of the additional P-CH3 signal is not observed unless excessive purification steps are attempted. Ostensibly, MeOH may be aiding the mechanism by which this phenomenon occurs. Thus, all inhibitors were synthesized this way and excess salts were removed by size-exclusion chromatography. All biological studies were performed with the unknown impurity showing a resonance at 1.94 ppm present. ESI-MS was used to further verify the identities of the synthesized molecules. 28

Figure 41: Optimized conditions for phosphonamidate ester deprotection. Excess sodium hydroxide and salts are subsequently removed by size-exclusion chromatography.

To further test the stability of these inhibitors, 1H-NMR spectra of 15a were taken immediately after and 22 hours after being in solution in aqueous conditions at pH 6.0, 7.5 and 9.0 (Figure 22). The compound was stable under all these conditions, which was surprising since phosphonamidates have been shown to be susceptible to acid-catalyzed hydrolysis47. None of the spectra significantly differed from one another and no degradation products were observed. Spectra taken after one week under the same conditions also exhibited no deterioration. However, hydrolysis is observed after a few weeks in solution.

Figure 23 shows the full scheme for synthesis of 15a-f. The characterization data (1H-, 13C- and 31P-NMR and HRMS) and detailed synthesis procedures for all synthesized compounds are presented in the supplementary information section.

Figure 22: 1H-NMR of 15a taken immediately after (top) and 22 hours after (bottom) being in solution in phosphate buffer under ambient conditions at pH 6.0, 7.5 and 9.0. Minor impurities were present in all samples and their concentrations did not change throughout the 24 hours.

2.1.3 Synthesis Scheme

Figure 23: Synthesis Schemes A) Scheme for 4,6-benzilidine Intermediates 4 and 5 B) Scheme for 4,6-benzilidine Intermediate 11 C) Scheme for final synthesis of inhibitors. a,d: R1 = Me, R2 = H/Ac, b,e: R1 = Me, R2 = Me, c,f: R1 = Bn, R2 = H/Ac.

2.2 Trisaccharide Inhibitor

For the synthesis of Inhibitor 21, a chemoenzymatic approach was undertaken (Figure 24). Chitin, polymer of β 1,4-linked GlcNAc, was initially hydrolyzed into polysaccharides of various lengths according to a previously reported protocol56. Briefly, chitin (1 g) was dissolved in 11 M HCl (50 mL) by stirring for 2 hours at 0 ºC. The reaction was warmed to r.t. and stirred for an additional 4 hours before neutralization with concentrated sodium hydroxide. Then, trisaccharides (chitotriose, 16) was purified using size-exclusion chromatography on a BioGel P- 4 column (5-8% yield).

Figure 24: Scheme for synthesis of trisaccharide inhibitor (21).

To synthesize 17, deacetylation of the middle residue of 16 was performed using purified SpPgdA (Protein expression and purification protocols are presented below). The 32

procedure involved incubating 2 mM chitotriose in 1 µM SpPgdA at 37 ºC for 4.5 hours, followed by size-exclusion chromatography to purify the polysaccharide content (30 mL total reaction volume, buffer: 25 mM Tris/HCl, 100 mM NaCl, 5 µM CoCl2, pH 7.5). The mono- deacetylated product was predominantly observed in MALDI-MS following purification.

Synthesis of 18 utilized the same conditions as the diazo transfer reaction used to synthesize 10. Next, the reducing end of the sugar was protected using N,O- dimethylhydroxylamine via previously reported procedures57. Briefly, to a solution of 18 dissolved in a minimal amount of H2O at 0 ºC, sodium acetate (1.1 equivalents) and N,O- dimethylhydroxylamine hydrochloride (1.1 equivalents) were added. The solution was allowed to stir at r.t. for 26 hours and then washed three times with ethyl acetate. Size-exclusion chromatography was used to desalt and obtain 19. Then, peracetylation followed by Staudinger reaction with diethyl methylphosphonite afforded 20. These reactions were performed using the same protocols as ones used for the synthesis of monomeric inhibitors. Figure 25 shows the MALDI-MS of 20. In addition to the monophosphanomidated product, it also shows the presence of diphosphanomidated compounds as minor products. This is due to SpPgdA catalyzing multiple de-N-acetylations on one molecule of chitotriose. MALDI-MS data for 16-19 are presented in the supplementary information section.

Figure 25: MALDI-MS of 20 (Calculated m/z [M+Na+] = 1051.4). The spectrum also shows the presence of a diphosphonamidated product (Calculated m/z [M+Na+] = 1115.4).

The final step of the synthesis was deprotection using alkaline conditions, followed by size-exclusion chromatography. However, this reaction appeared to be unsuccessful as multiple products were observed by MALDI-MS analysis, none of which were the expected phosphonamidate. Additionally, concurrent synthesis in our lab of an analogous pentameric inhibitor based on a PNAG scaffold showed similar difficulties and may suggest that these polymeric compounds are more prone to hydrolysis than their monosaccharide counterparts.

2.3 Discussion and Conclusions

The direct Staudinger-phosphonite reaction investigated in this study produced interesting results. The identity of the impurity produced by this reaction remains unknown. It is even present when hexanoate is used as the protecting group, which suggests that it is not an acetate salt. This is surprising since only the 1H-NMR peak at 1.94 ppm and 13C-NMR peak at 34

23.2 ppm can be assigned to it, which are diagnostic of an acetate salt. A possible explanation for this is the presence of a contaminant. However, this is highly unlikely since a variety of conditions were explored and the impurity was present under all conditions. Instead of base- sensitive protecting groups, other strategies should be explored. This would allow selective deprotection of the phosphonamidate ester before removal of hydroxyl protecting groups. This may allow complete purification of final products or at least help elucidate the source of this impurity.

1 An additional issue with this reaction is the appearance of a second P-CH3 peak in the H- NMR spectra of the fully deprotected product. Surprisingly, no other peaks were concurrently produced. Thus, it cannot be due to hydrolysis as this produces observable shifts in all the other sugar peaks. One explanation could be the migration of the phosphorus to the 3’-OH group. This could explain why this peak is not observed when this position is methylated. However, if this was the case, there would be significant shifts in 1H-NMR peaks of both the 2’ and 3’ protons, which are also not observed.

There are a few different aspects to be gleaned in terms of the stabilities of these phosphonamidated carbohydrates. The 3’-OMe derivatives were the most stable during purification and the much more polar trisaccharide inhibitor was unattainable. This could be due variability in the number of hydroxyl groups in these molecules, which may assist the mechanism by which they deteriorate. Elucidating the reasons for this trend may allow the synthesis of the trisaccharide inhibitor as well as a multitude of other more complex molecules. It may also permit phosphonamidate synthesis exclusively under aqueous conditions without the need for hydroxyl protecting groups.

In conclusion, phosphonamidated monosaccharides were synthesized in reasonable yields via a direct Staudinger-phosphonite reaction. A detailed study of this reaction and the mechanisms by which phosphonamidates hydrolyze may allow the synthesis of phosphonamidated oligosaccharides.

3 Enzymology

3.1 Protein expression and purification

3.1.1 SpPgdA

A pET28b plasmid containing the gene for the C-terminal de-N-acetylase domain of

SpPgda (residues 232-431) attached to a Histidine-tag via an eleven residue long linker (His6-

SpPgdA232-431) was synthesized (Bio Basic Inc.). The tag was inserted at the N-terminus in order to resemble the full-length protein, which has the catalytic domain at the C-terminus. The gene was inserted between NdeI and XhoI restriction sites. The plasmid was used to transform E. coli BL21 (DE3) cells to obtain colonies. A single colony was used to grow a 25 mL bacterial culture overnight at 37 ºC in Luria–Bertani (LB) broth containing 50 µg ml-1 kanamycin. The 25 mL culture was used to grow a 1L culture in LB broth supplemented with 50 µg ml-1 kanamycin at 37 ºC until OD600 reached 0.6. Protein expression was initiated by the addition of isopropyl-d- thiogalactopyranoside (IPTG) such that its final concentration was 1 mM. Expression was allowed to occur for 3 h at 37°C and then the cells were harvested by centrifugation at 3,750 g for 60 min. The cell pellet was resuspended in 10 ml of lysis buffer A (25 mM Tris·HCl, 100 mM NaCl, pH 7.5) and a tablet of Complete Mini, EDTA-free Protease Inhibitor Cocktail (Roche) was added. Cells were lysed by sonication and the lysate was centrifuged at 16,000 g for 30 min to remove any solid cell debris and then filtered through a 0.45-mm filter.

Protein purification was performed with HIS-Select® Nickel Affinity resin (Sigma-Aldrich). The resin (1 mL for one liter of culture) was initially washed with lysis buffer A. The filtered supernatant was flown through the column to allow the protein to bind to the resin. Subsequently, the resin was washed with 10 column volumes (CV) of lysis buffer A followed by 5 column volumes of lysis buffer A supplemented with 5 mM Imidazole. Protein was eluted using lysis buffer A supplemented with increasing concentrations of imidazole until a concentration of 250 mM was reached. The eluded fractions were screened using SDS-PAGE gel electrophoresis and fractions containing pure His6-SpPgdA232-431 were combined. The protein 36

was dialyzed three times in the required buffer, concentrated with a 10,000 Da MW spin filter and stored at 4ºC. To obtain Co2+ loaded protein, the stock was incubated overnight with 5 equivalences of CoCl2 followed by dialysis to remove the excess CoCl2. Protein concentration was quantitated using a 280 nm extinction coefficient of 25440 M-1 cm-1. From this point forward, His6-SpPgdA232-431 will be referred to as SpPgdA. Figure 26 shows the amino acid sequence of His6-SpPgdA232-431 along with the SDS-PAGE gel of the purified protein.

3.1.2 PgaB

PgaB fused to a Histidine-Tag (His6-PgaB) was expressed using previously published procedures22,58. Briefly, E. coli BL21 (DE3) cells transformed with a pET28 vector containing the

His6-PgaB gene was used to grow a 10 mL overnight culture at 37 ºC in LB broth supplemented with 100 µg ml-1 streptomycin. The 10 mL small culture was used to grow a 1L culture at 37 ºC in LB broth supplemented with 100 µg ml-1 streptomycin. At approximately 30 min before expression, NiCl2 was added to a final concentration of 1 mM. and the temperature was reduced 10 ºC. Protein expression was induced by the addition of IPTG to a final concentration of 0.5 mM and expression was allowed to occur overnight. The cells were harvested by centrifugation at 3,750 g for 60 min. The cell pellet was resuspended in 10 ml of lysis buffer B (50 mM phosphate, 300 mM NaCl, pH 8) and a tablet of Complete Mini, EDTA-free Protease Inhibitor Cocktail (Roche) was added. Cells were lysed by sonication and the lysate was centrifuged at 16,000 g for 30 min to remove any solid cell debris. From this point forward, His6- PgaB will be referred to as PgaB.

Protein purification was performed with HIS-Select® Nickel Affinity resin. The resin (600 µL for one liter of culture) was initially washed with lysis buffer. The filtered supernatant was added to the resin and incubated at 4 ºC for 1 hr while rotating on a neutator. The resin was spun down at 4,000 rpm for 10 min and washed with 10 mL of lysis buffer B. Protein was eluted using lysis buffer B supplemented with increasing concentrations of imidazole until a concentration of 100 mM was reached. The eluded fractions were screened using SDS-PAGE gel electrophoresis and fractions containing pure His6-PgaB were combined and dialyzed three 37

times in lysis buffer B. The protein was concentrated with a 30,000 Da MW spin filter and stored at 4ºC. Protein concentration was quantitated using a 280 nm extinction coefficient of 152,000 M-1 cm-1.

Figure26: Amino acid sequence of His6-SpPgdA232-431 utilized in this study (Top) and SDS-PAGE gel of purified His6- SpPgdA232-431 (bottom). Red corresponds to amino acid residues 232 to 431 of full-length SpPgdA. Each lane on the gel corresponds to a different fraction collected during purification (increasing imidazole concentrations from left to right)

3.2 Procedures for Analysis of Enzyme Kinetics

3.2.1 Fluorogenic Substrate Assay

The activities of PgaB and SpPgdA were analyzed using the fluorogenic Substrate 3- carboxyumbelliferyl acetate (ACC), which was synthesized according to previously published procedures (Figure 27)34,59. In a standard assay, the enzyme (final concentration of 1 µM Co2+- loaded SpPgdA or 10 µM PgaB) and if present the inhibitor (dissolved in H2O) were combined with buffer (final concentration of 25 mM HEPES, pH 7.0 for SpPgdA and 100 mM HEPES, pH 7.5 for PgaB) in a black 96-well plate and incubated at 37 ºC for 10 min. Subsequently, ACC dissolved in DMSO was added to obtain the required substrate concentration and fluorescence (excitation: 386 nm, emission: 446 nm) was immediately monitored. Fluorescence readings were taken every minute for 10 min for PgaB and every 5 min for 30 min for SpPgdA. Each 38

reaction contained a final DMSO concentration of 10% and a total volume of 50 µL for PgaB and 100 µL for SpPgdA. Each reaction was performed in triplicates and each replicate was accompanied by an analogous control, which did not contain enzyme. These controls gave the substrate autohydrolysis rates, which were subtracted from rates obtained from enzyme- containing reactions in order to measure the enzyme-catalyzed rate. Standard curves attained using the fluorogenic product of ACC hydrolysis (7-hydroxycoumarin-3-carboxylic acid) were used to convert fluorescence values to product concentrations. All assays were performed under these conditions unless stated otherwise. Error bars represent standard deviations calculated from three separate biological replicates.

3.2.2 Fluorescamine Assay

Fluorescamine assays with SpPgdA were performed using chitotriose (synthesized as in Figure 24) as the substrate (Figure 27). A standard optimized reaction consisted of 2 µM

SpPgdA in 25 mM HEPES, pH 7 and 5 µM CoCl2. The enzyme, buffer, CoCl2 and inhibitor if present (dissolved in H2O) were combined in a microcentrifuge tube and incubated at 30 ºC for

5 min. Next, the required amount of chitotriose dissolved in H2O was added to obtain the desired final substrate concentration. The total reaction volume was 50 µL. Immediately, the sample was incubated at 30 ºC. Aliquots of 10 µL were removed from the reaction every 75 or 100 seconds for up to 5 min. Each aliquot was immediately added to a 20 µL solution of 0.4 mM Borate Buffer, 100 mM EDTA, pH 9. To this solution, 10 µL of a freshly prepared 2 mg ml-1 solution of fluorescamine in DMF was added. This labelling reaction was allowed to occur for 10 min at r.t. before addition of 65 µL of a 1:1 mix of DMF:H2O. Afterwards, 100 µL of this solution was transferred to a black 96-well plate and its fluorescence was measured (excitation: 360 nm, emission: 460 nm). A standard curve attained using glucosamine was used to convert fluorescence values to product concentrations. All assays were performed under these conditions unless stated otherwise. Error bars represent standard deviations calculated from three separate biological replicates.

Figure 27: Assays used for kinetic analysis of PgaB and SpPgda. ACC substrate becomes fluorescently active upon deacetylation by either PgaB or SpPgdA (TOP). Following deacetylation of chitotriose by SpPgdA, the generated free amine is reacted with fluorescamine to produce a fluorescently active compound (bottom).

3.2.3 Data Analysis

All data were graphed and fitted using Microsoft Excel 2010. Michaelis–Menten parameters (KM and kcat) for each enzyme-substrate combination were obtained by fitting the initial reaction rates at various substrate concentrations to the Michaelis–Menten equation (Equation 1). The initial rate data from enzymatic reactions performed with various inhibitor Apparent concentrations were also fitted to the Michaelis–Menten equation to obtain (KM ). These were subsequently utilized to obtain inhibition constants (Ki) via Equation 2, where [I] is the inhibitor concentration. All Michaelis–Menten curves are averages of three independent replicates and all reported Ki values are averages calculated from at least three independent experiments using at least three different inhibitor concentrations. ACC and chitotrose were synthesized according to previously reported procedures56,59.

3.3 Results

To probe the activity and to develop an assay for screening inhibitors of SpPgdA, the fluorogenic substrate ACC was initially investigated. ACC was previously found in our lab to be a 34 suitable substrate for PgaB . SpPgdA is able to catalyze the hydrolysis of ACC with a similar kcat and a smaller KM than PgaB (Table 5). The Michaelis-Menten curve for this enzyme-catalysis is shown in Figure 28. It also shows the reaction progress (as measured by the increase in fluorescence due to product formation) in the presence of varying concentrations of inhibitor 15c. The reaction displays an inhibitor concentration-dependent decrease in initial rate. From this data an approximate IC50 of 50 µM was calculated. However, when the assay was repeated with additional inhibitor concentrations, a complex result was observed.

k /K (M- -1 cat M Enzyme Substrate KM (mM) kcat (s ) 1S-1)

SpPgdA ACC 0.26 ± 0.09 0.01 ± 0.002 40 ± 20

β-1,4-(GlcNAc)3 5.9 ± 0.3 1.13 ± 0.07 190 ± 20

PgaB ACC 1.4 ± 0.2 0.0087 ± 0.008 6 ± 6

1.2 ± 0.2a 0.013 ± 0.002a 11 ± 2a

Table 5: Michaelis-Menten kinetic parameters measured for the various enzyme and substrate combinations used in the study. aValues previously obtained in the lab for Maltose-binding protein-PgaB fusion protein (MBP-PgaB)34

Figure 28: Michaelis-Menten curve for SpPgdA-catalyzed ACC hydrolysis (left). Note: Assay was performed in 25 mM Tris/HCl, pH 7.0 buffer. SpPgdA-catalyzed hydrolysis of 1 mM ACC in the presence of varying concentrations of inhibitor 15c as measured by the increase in fluorescence due to product formation (right).

Figure 29 shows the raw fluorescence data obtained during the reaction, fluorescence from ACC autohydrolysis in analogous controls with no enzyme, and SpPgdA-catalyzed hydrolysis determined by subtracting the fluorescence due to autohydrolysis from raw fluorescence. At high inhibitor concentrations, the autohydrolysis rate increases dramatically suggesting that the inhibitor itself is able to weakly catalyze substrate hydrolysis. This is conceivable as ACC is not particularly stable as observed by its non-negligible autohydrolysis rate under aqueous conditions and the inhibitor itself is ionic and any interaction between the inhibitor and substrate can potentially lead to an increase in the autohydrolysis rate. Additionally, inhibition of SpPgdA-catalyzed hydrolysis completely disappears at high inhibitor concentrations. This is particularly noticeable at 2 mM inhibitor as the initial rate is nearly identical to the rate observed without any inhibitor present. This effect is also seen in the Michaelis-Menten curves measured at different inhibitor concentrations (Figure 30) as the enzyme’s affinity for substrate (KM) at 2 mM inhibitor is nearly identical to its affinity when the inhibitor is absent (the kcat is constant at all inhibitor concentrations since these are competitive inhibitors as proven by experiments presented below). This is also in direct contradiction to the decrease in affinity that was observed at low inhibitor concentrations. These results indicate that at high concentrations, the inhibitor also activates the enzyme to some degree for ACC hydrolysis, which results in the cancelation of its inhibitory effect. This outcome was also qualitatively observed with inhibitors 15a,b,d-f (unpublished results).

Figure 29: Hydrolysis of 1 mM ACC in the presence of varying concentrations of inhibitor 15c. Raw rates of ACC hydrolysis by SpPgda without subtracting for autohydrolysis (left). Autohydrolysis rates in samples without any SpPgda (middle). SpPdgA-catalyzed hydrolysis obtained by subtracting fluorescence due to autohydrolysis form raw data (right).

Figure 30: Michaelis-Menten curves for SpPgdA-catalyzed ACC hydrolysis at varying concentrations of inhibitor 15c.

The above observed phenomenon is rather surprising. It may be due to enzyme instability at high inhibitor and substrate concentrations as reproducibility diminishes at these conditions. For example, at an inhibitor concentration of 1 mM, stronger inhibition is observed than at 100 µM in the first experiment (Figure 28). The opposite was detected in the second experiment (Figure 29). In addition, it was observed that SpPgdA activity is completely lost at ACC concentrations above 1.5 mM. This is not the case for PgaB, which is fully active even at 43

ACC’s solubility limit of 10 mM in 10% DMSO (Figure 31). This further suggests that SpPgdA is less stable in conditions where these small molecule concentrations are high and therefore the results become complex and unpredictable.

This effect may also be due to substrate-inhibitor interactions that might aid in funneling the substrate to the enzyme and thereby facilitating enzyme-substrate binding. This may be the most probably reason as ACC contains a hydrophobic coumarin moiety that may interact with the hydrophobic regions of the inhibitor (for example, the benzyl rings). In addition, ACC does not remotely resemble the natural substrate of the enzyme. Thus, both the inhibitor and substrate may be able to bind to different areas of the relatively large substrate biding pocket of the enzyme at the same time. The binding pocket is designed for large glycans and ACC is a much smaller molecule. This effect would predictably, and as observed, be more prevalent at high inhibitor concentrations if the inhibitor-substrate interaction is relatively weak. Two small molecules interacting through a few hydrophobic interactions would likely be a weak interaction. In addition, the observation that the inhibitor itself is able to weakly catalyze substrate hydrolysis further supports this model as this requires that the two molecules interact in solution.

Figure 31: Michaelis-Menten curve for PgaB-catalyzed ACC hydrolysis.

Since ACC was not a suitable substrate, fluorescamine assays were utilized for kinetic analysis of SpPgdA. Fluorescamine reacts quantitatively with primary amines, which allows quantitation of free amines generated by de-N-acetylation (Figure 27). Chitotriose, which contains the β 1,4-linked GlcNAc residues that mimic the peptidoglycan backbone, was previously shown to be de-N-acetylated by SpPgdA30. Thus, it was utilized as the substrate for the reaction. Figure 32 displays the time-course for SpPgdA-catalyzed de-N-acetylation at various chitotriose concentrations. It displays linear initial rates that increase in a concentration-dependent manner. The figure also displays the Michaelis-Menten curve for this catalytic process. The measured kcat and KM values (Table 5) are in good agreement with previously reported values for full-length SpPgdA (Table 2)30. Michaelis-Menten curves measured at different inhibitor concentrations are shown in Figure 33 for inhibitor 15c. The corresponding Lineweaver–Burk plots for all inhibitors screened against SpPgdA are presented in the supplementary information. As expected, these curves show that these are competitive inhibitors as the KM for substrate increases with increasing inhibitor concentration (x-intercept of Lineweaver–Burk plots increase with inhibitor concentration) while the kcat remains constant (y-intercept of Lineweaver–Burk plots remain constant).

Figure 32: Reaction progress curves for SpPgdA-catalyzed de-N-acetylation at various substrate (chitotriose) concentrations. (left). Michaelis-Menten curve for SpPgdA-catalyzed chitotriose de-N-acetylation (right). Data measured via fluorescamine assays.

PgaB was analyzed using fluorogenic substrate assays with ACC as this was found to be a suitable method in this case. The problems encountered with SpPgdA at high inhibitor 45

concentrations were not observed in this case. The Michaelis-Menten curve for PgaB-catalyzed

ACC hydrolysis is shown in Figure 31 and the calculated kcat and KM values are in good agreement with previously reported results in our lab (Table 5)34. Lineweaver–Burk plots for all inhibitors screened against PgaB are also presented in the supplementary information.

Figure 33: Michaelis-Menten curves for SpPgdA-catalyzed chitotriose de-N-acetylation in the presence of varying concentrations of inhibitor 15c. Curves were obtained by using the fluorescamine assay.

The calculated Ki values for all inhibitors screened against PgaB and SpPgdA are shown in Table 6. The most potent SpPgdA inhibitor was 15c, which contains benzyl groups at positions

1’ and 4’ of the glucose ring. Ki values calculated for this inhibitor using the two different assays show good agreement with each other. This validates the utilization of ACC for kinetic analysis of these enzymes at least when the concentrations of these specific inhibitors are relatively not too high (≤ 500 µM). The inhibitor containing the methylated 3’-OH shows a significantly lower

Ki than the one without this modification, which suggests that this hydroxyl group is vital for binding. Unfortunately, the inhibitors designed for PNAG deacetylases are not able to inhibit PgaB. Interestingly, this lack of inhibition may account for why unlike for SpPgdA, no problems were encountered when using ACC as the substrate.

15a 15b 15c 15d 15e 15f

80 ± 20 SpPgdA 170 ± 30 720 ± 150 580 ± 40 - 220 ± 40 40 ± 20a

PgaB > 1000 - > 1000 > 1000 > 1000 > 1000

3.4 Discussion and Conclusions

The molecules designed for SpPgdA exhibited reasonable inhibition. The most potent inhibitor (15c) is the most hydrophobic one and contains two benzyl groups that mimic sugar rings. It has a Ki of 80 µM, which makes it the most potent inhibitor reported to date for any peptidoglycan de-N-acetylase. It was also twice as effective as the mono-benzylated inhibitor (15a), thus providing evidence for our premise that these benzyl rings bind to the hydrophobic pockets that surround the active site with reasonable affinity. As hypothesized, the inhibitor with the 3’-OMe group (15b) was a much weaker inhibitor. It was more than four times as weak as its 3’-OH counterpart. This provides the first experimental evidence for previously reported docking studies and proposals for many of the enzymes in this family, which suggested that the 3’-OH of all substrates make essential contacts within the active site and likely coordinate to the metal ion22,26,30. Therefore, any future designs of inhibitors for this group of enzymes should try to mimic this interaction. As expected, inhibitors with the 6’-benzyl group designed for PgaB (15d,f) display three to four times weaker inhibition of SpPgdA than their 4’-benzyl counterparts. The ability to distinguish between β-1,4 and β-1,6-linkages plays a major role in determining the substrate preferences exhibited by these enzymes.

All of the designed molecules show complete lack of PgaB inhibition. This is surprising since previous reports in our lab showed that adding metal-coordinating motifs at the 2’ position of a glucose scaffold results in inhibitors that have at least weak levels of inhibition34. There are several possible reasons for these results. For one, PgaB may be a very selective enzyme. The benzyl groups may not be an adequate replacement for sugar rings and may in fact cause steric hindrances that may prevent binding. However, this would be unlikely since PgaB can accommodate these benzyl groups in the hydrophobic pockets that surround the active site. These 1,6-linkages are flexible and can adapt a large array of conformations since they are attached to primary carbons on either side. Also, PgaB has a large and relatively open substrate-binding cleft. This is in contrast to SpPgdA, which contains a much more narrow and pinched off binding cleft. PgaB’s catalytic efficiency and substrate affinity are the lowest reported for any enzyme in this class. This probably contributes to the ineffectiveness of these inhibitors. An enzyme that weakly binds its substrate while also having a low turnover rate is also most likely to poorly bind competitive inhibitors that resemble the substrate.

Another possible explanation for the observed lack of PgaB inhibition is the enzyme’s mechanism of substrate binding. Recent structural, functional and molecular dynamics simulations indicate that association of PgaB’s N- and C-terminal domains leads to the formation of a cleft where PNAG binds. In addition, they suggest that PgaB associates with PNAG continuously while being transported through the periplasm. The continuous binding interaction extends from the active site within the N-terminal domain, all the way to an electronegative groove on the C-terminal domain. The N-terminal domain has a binding preference for GlcNAc residues and the C-terminal domain has a binding preference for glucosammonium. These data suggest a processive mechanism in which newly synthesized PNAG enters the periplasm and traverses through it by binding to low affinity binding sites located throughout the surface of PgaB. Along this path, PNAG is de-N-acetylated. This mechanism indicates that the major affinity for PNAG binding by PgaB results from these numerous low affinity interactions and the fact that PgaB is located in close proximity to the PNAG synthase and transporters60. Thus, inhibitors designed by only considering the interactions that occur within the binding site may not be effective. 48

The structural basis for the differences in substrate specificity and catalytic efficiency displayed by CE4 de-N-acetylases remains elusive. Our data suggest that these enzymes have significant structural differences and rely on different aspects of a substrate for affinity. SpPgdA behaves like a conventional enzyme in terms of binding its substrate using predominately an independent active site and considerably stabilizing the transition-state. On the contrary, PgaB appears to behave like other glycan binding proteins and enzymes in terms of depending on a close cluster of multiple low affinity interactions to drive substrate binding and the subsequent reaction. It would be worthwhile to screen these inhibitors against PNAG deacetylases, such as IcaB, from Gram-positive bacteria. These deacetylases are extracellular proteins and as such they cannot rely as much on local concentrations to drive reactivity, even if they are associated with the cell membrane. In fact, the only available structure of a PNAG deacetylase from Gram- positive bacteria, IcaB from Ammonifex degensii, shows that these enzymes have loop architectures and narrow binding clefts similar to PgdA. According to docking studies, they also contain three conserved residues not present in PgaB that make vital interactions with the substrate sugar rings. They also don’t require the additional C-terminal domain for activity. These observations indicate that these deacetylases may behave in a similar manner as SpPgdA and be susceptible to these inhibitors.

It would be extremely valuable to obtain crystal structures of these enzymes bound to transition-state analogs. It would clarify many of these mechanistic differences and allow for rational design of inhibitors tailored to each specific enzyme. In fact, the only available crystal structures of one these enzymes bound to its natural substrates, an inactive mutant of VcCDA bound to either chitobiose or chitotriose, show that binding leads to significant conformational changes in loops that surround the active site. These structural changes differ depending on the lengths of the oligosaccharide substrate. These details are in support of an induced-fit mechanism in which specific loops and their inherent dynamics both shape and differentially block available subsites within these enzymes. By doing so, these loops essentially modulate substrate specificity and define deacetylation patterns of CE4 enzymes. It will be interesting to find out whether and how this applies for the various PNAG and peptidoglycan deacetylases

and also whether formation of the transition-state is accompanied by further structural changes.

In conclusion, the designed transition-state analog inhibitors for peptidoglycan de-N- acetylases show low micromolar inhibition of SpPgdA. They verify that the 3’-OH of the sugar being deacetylated is crucial for substrate binding. Inhibitors designed for PNAG deacetylases show no inhibition of PgaB. Thus, alternate strategies for the development of inhibitors of this enzyme must be devised. In the future, the structure of the most potent inhibitor of SpPgdA that was identified (15c) should be further optimized to obtain a stronger inhibitor.

Supplementary Information

General Procedures

Reagents were obtained from Acros Organics or Sigma-Aldrich and were used without further purification. Standard syringe techniques were utilized for the transfer of dry solvents and air-sensitive reagents. Reactions were monitored by TLC using Silica Gel 60 F254 (EMD Science). 1H-, 13C- and 31P-NMR spectra were recorded at 25 °C on Varian 400, Agilent DD2 500 or 600 MHz (100, 126 or 151 MHz for 13C and 162, 202 or 243 MHz for 31P, 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), ddd (doublet of doublet of doublets), dt (doublet of triplets) or m (multiplet). Broad peaks are indicated by the abbreviation br. Coupling constants are reported as J-values in Hz. For a given resonance, the number of protons (n) 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. Flash chromatography was performed on Silica-P Flash Silica Gel 60 (40–63 μm particle size, Silicycle). High-resolution mass spectra were obtained from a JEOL AccuTOF model JMS-T1000LC (DART) or AB Sciex QStarXL (ESI) mass spectrometer. Inhibitor stock concentrations were determined by 1H-NMR using imidazole as an internal standard. All fluorescence measurements were performed on either a Tecan Safire 2 or BMG LABTECH CLARIOstar® plate reader. Protein concentrations were measured using a NanoDrop® ND-1000 Spectrophotometer.

Synthesis Procedures

Methyl 2-azido-2-deoxy-4,6-O-benzylidene-β-D-glucopyranoside

The following procedure was based on a previously reported procedure50. To a mixture of NaN3 (0.18 g, 2.8 mmol) and ceric (IV) ammonium nitrate (3.03 g, 5.5 mmol) cooled to -30 °C, a solution of tri-O-acetyl-D-glucal (0.5 g, 1.8 mmol) in CH3CN (10 mL) was added dropwise.

Following vigorous stirring under N2 for 8 h at -20 °C, the reaction was warmed to 0 °C, poured into ice water and extracted with EtOAc (3 x 35 mL). The organic extracts were combined and washed with H2O (1 x 50 mL) and brine (1 x 50 mL) and dried with Na2SO4. The solution was concentrated in vacuo and dried overnight to obtain 2 as a clear syrup.

To a solution of 2 in dry MeOH (10 mL) at 0 °C, Na° (0.254 g, 11.0 mmol) was added in portions and stirred for 30 min. The reaction was neutralized with Dowex 50W-X8 cation exchange resin. The resin was removed by filtration and the filtrate was concentrated in vacuo. The resulting syrup was redissolved in MeOH:acetone (1:3) and cooled to –20 °C. The precipitate (NaNO3) that results was removed by filtration and the filtrate was concentrated in vacuo and dried overnight to afford 3 as a clear syrup.

The mixture 3 was dissolved in dry DMF (19 mL) and benzaldehyde dimethyl acetal (0.42 mL, 7.1 mmol) and catalytic p-toluenesulfonic acid monohydrate were added. The reaction was stirred under N2 at r.t. overnight and then neutralized with Et3N. The solution was concentrated in vacuo and purified by flash chromatography (6:2:2, Toluene:Pentanes:EtOAc) to afford 1 isomer 4 (147 mg, 21%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.53 – 7.43 (m, 2H), 7.44 – 7.32 (m, 3H), 5.55 (s, 1H), 4.41 – 4.27 (m, 2H), 3.85 – 3.73 (t, J = 10.5 Hz, 1H), 3.69 (dt, J = 2.1, 9.4 Hz, 1H), 3.58 (s, 3H), 3.56 (t, J = 9.2 Hz, 1H), 3.48 – 3.34 (m, 2H), 2.59 (d, J = 2.0 Hz, 1H). 13C

NMR (126 MHz, CDCl3) δ 136.7, 129.4, 128.4, 126.2, 103.5, 102.0, 80.6, 76.7, 72.1, 68.5, 66.4, + 66.1, 57.5. HRMS m/z calcd. C14H17N3O5 (M+H ) 308.1246, found 308.1253.

Methyl 2-azido-2-deoxy-3-O-methyl-4,6-O-benzylidene-β-D-glucopyranoside (5)

To a stirring solution of 4 (80 mg, 0.27 mmol), CH3I (19 µL, 0.30 mmol) and dry DMF (2 mL) under N2 at 0 ºC, NaH (14 mg, 0.34 mmol) was added as a suspension in dry DMF (2 mL) and stirred for 4 hr. Reaction was quenched by addition of H2O and extracted with EtOAc (3 x

15 mL). The combined organic extracts were washed with brine (1 x 15 mL), dried with MgSO4, concentrated in vacuo and purified by flash chromatography (2:1 Pentanes:EtOAc) to obtain 5 1 (83 mg, 96%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.52 – 7.41 (m, 2H), 7.43 – 7.30 (m, 3H), 5.56 (s, 1H), 4.35 (dd, J = 10.5, 5.0 Hz, 1H), 4.34 – 4.22 (m, 1H), 3.80 (t, J = 10.3 Hz, 1H), 3.65

(s, 3H), 3.65 – 3.60 (m, 1H), 3.58 (s, 3H), 3.62 – 3.50 (m, 1H), 3.42– 3.30 (m, 3H). 13C NMR (126

MHz, CDCl3) δ 137.06 , 129.04 , 128.25 , 125.97 , 103.37 , 101.26 , 81.57 , 81.25 , 68.54 , 66.12 , + 66.08 , 65.79 , 60.98 , 57.43. HRMS m/z calcd. C15H19N3O5 (M+H ) 322.1403, found 322.1397.

N’-(2-acetamido-2-deoxy-β-D-glucopyranosyl)-p-toluenesulfono-hydrazide (7)

To a suspension of N-acetylglucosamine (5 g, 23 mmol) and p-tosylhydrazide (5.5 g, 30 mmol) in DMF (15 mL), acetic acid (1 mL) was added and the mixture was incubated at 37 °C for

48 hrs. The solution was poured into Et2O (700 mL) and stirred overnight. The resulting white 1 residue was collected by filtration and washed with Et2O and DCM to obtain 7 (8.2 g, 91%). H

NMR (500 MHz, D2O) δ 7.74 (d, J = 8.3 Hz, 2H), 7.48 (d, J = 8.3 Hz, 2H), 3.93 (d, J = 9.2 Hz, 1H), 3.89 (dd, J = 11.7, 1.7 Hz, 1H), 3.70 (m, 1H), 3.52 (t, J = 10.1, 9.2 Hz, 1H), 3.49-3.44 (m, 1H), 3.40- 13 3.28 (m, 2H), 2.46 (s, 3H), 2.05 (s, 3H). C NMR (126 MHz, D2O) δ 174.14, 145.53, 132.97, 129.77, 127.71, 89.67, 76.54, 74.09, 69.72, 60.66, 53.12, 22.10, 20.66. HRMS m/z calcd. + C15H23N3O7S (M+H ) 390.1329, found 390.1329.

Benzyl 2 -acetamido-2-deoxy-β-D-glucopyranoside (8)

To a solution of 7 (500 mg, 1.2 mmol) and benzyl alcohol (1.3 mL, 12.5 mmol) dissolved in anhydrous DMF (10 mL), N-bromosuccinimide (530 mg, 3.0 mmol) was added in portions. After 15 min of stirring, amberlite resin (-OH) was added and the solution was stirred at r.t. for 2 hr. The resin was removed by filtration. The filtrate was concentrated in vacuo and purified by flash chromatography (1:9, MeOH:DCM) to afford 8 (300 mg, 77%) as a white solid. 1H NMR

(500 MHz, CD3OD) δ = 7.30-7.25 (m, 4H), 7.24-7.18 (m, 1H), 4.83 (d, J = 12.2, 1H), 4.55 (d, J = 12.2 Hz, 1H), 4.41 (d, J = 8.5 Hz, 1H), 3.86 (dd, J = 12.1, 2.1 Hz, 1H), 3.70-3.63 (m, 2H), 3.39 (dd, J = 10.4, 8.4 Hz, 1H), 3.29 (dd, J = 9.7, 8.4 Hz, 1H), 3.22 (ddd, J = 9.7, 5.7, 2.1 Hz, 1H), 1.90 (s, 3H). 13 C NMR (126 MHz, CD3OD) δ 172.29, 137.77, 127.95, 127.88, 127.37, 100.40, 76.63, 74.53, + 70.75, 70.12, 61.43, 55.93, 21.55. HRMS m/z calcd. for C15H21NO6 (M+Na ) 334.1261, found 334.1270.

Benzyl 2 -amino-2-deoxy-β-D-glucopyranoside (9)

Hydrazine monohydrate (2 mL) was added to 8 (180 mg, 0.58 mmol) and stirred for 48 hr at 110 °C. The solvent was evaporated in vacuo to obtain 9 (100 mg, 64%) as an oil. 1H NMR

(500 MHz, D2O) δ 7.49 – 7.45 (m, 5H), 4.97 (d, J = 11.5 Hz, 1H), 4.75 (d, J = 11.4 Hz, 1H), 4.54 (d, J = 8.2 Hz, 1H), 3.96 (dd, J = 12.4, 2.2 Hz, 1H), 3.79 – 3.75 (m, 1H), 3.52 – 3.38 (m, 3H), 2.79 – 13 2.71 (m, 1H), 1.96 (s, 3H). C NMR (126 MHz, D2O) δ 136.34, 128.83, 128.71, 128.53, 100.94, + 76.06, 74.79, 71.53, 69.74, 60.69, 56.19, 19.64. HRMS m/z calcd. C13H19NO5 (M+H ) 270.1342, found 270.1348.

Benzyl 2 -azido-2-deoxy-β-D-glucopyranoside (10)

To a solution of NaN3 (650 mg, 10 mmol) in H2O (2 mL) and DCM (4 mL) at 0 ºC, triflic anhydride (335 µL, 2.0 mmol) was added and stirred for 2 hr. The solution was then extracted with DCM (3 x 5 mL). The organic layers were combined, washed with NaHCO3 (1 x 15 mL) and added to a stirring solution of 9 (280 mg, 1.0 mmol), K2CO3 (210 mg, 1.5 mmol) and catalytic amount of CuSO4 in 3 mL of H2O and 6 mL of MeOH. To the resulting mixture MeOH was added until the solution became homogeneous and it was left stirring for 2 hr. The solution was then concentrated in vacuo and purified by flash chromatography (1:19, MeOH: DCM) to afford 10 1 (266 mg, 90%) as a white solid. H NMR (600 MHz, CD3OD) δ 7.42 – 7.28 (m, 5H), 4.94 (d, J = 11.8 Hz, 1H), 4.70 (d, J = 11.8 Hz, 1H), 4.41 (d, J = 7.9 Hz, 1H), 3.89 (dd, J = 12.0, 2.2 Hz, 1H), 3.70 13 (dd, J = 11.9, 5.9 Hz, 1H), 3.39 – 3.19 (m, 4H). C NMR (151 MHz, CD3OD) δ 138.64, 129.37, 129.09, 129.06, 128.86, 101.87, 78.04, 76.35, 71.84, 71.57, 68.26, 62.59. HRMS m/z calcd. + C13H17N3O5 (M+NH4 ) 313.1512, found 313.1512.

Benzyl 2-azido-2-deoxy-4,6-O-benzylidene-β-D-glucopyranoside (11)

To a solution of 10 (300 mg, 1.0 mmol) and benzaldehyde dimethyl acetal (230 µL, 1.5 mmol) dissolved in dry DMF (6 mL), catalytic amount of p-toluenesulfonic acid monohydrate was added. The reaction was stirred under N2 at 60 °C for 8 hr and then at r.t. overnight. The solution was concentrated in vacuo, dissolved in DCM (20 mL) and washed with NaHCO3 (2 x 20

mL) and brine (1 x 20 mL). The resulting organic layer concentrated in vacuo and purified by flash chromatography (7:3, Pentanes:EtOAc) to afford 11 (270 mg, 70%) as a white solid. 1H

NMR (500 MHz, CDCl3) δ 7.52 – 7.43 (m, 2H), 7.44 – 7.29 (m, 8H), 5.55 (s, 1H), 4.95 (d, J = 11.7, 1H), 4.71 (d, J = 11.7, 1H), 4.52 (d, J = 8.0 Hz, 1H), 4.37 (dd, J = 10.5, 5.0 Hz, 1H), 3.82 (t, J = 10.3, 1H), 3.66 (dt, J = 9.3, 2.3 Hz, 1H), 3.58 (t, J = 9.3 Hz, 1H), 3.50 – 3.39 (m, 2H), 2.61 (d, J = 2.5 Hz, 13 1H). C NMR (126 MHz, CDCl3) δ 136.75, 136.29, 129.38, 128.55, 128.38, 128.19, 128.08,

126.23, 102.00, 101.06, 80.60, 72.04, 71.46, 68.52, 66.47, 66.19. HRMS m/z calcd. C20H21N3O5 (M+H+) 384.1560, found 384.1575.

Methyl 2-azido-2-deoxy-4-O-benzyl-β-D-glucopyranoside (12a)

BH3-THF (1 M, 1.0 mL) was stirred with 4 (60 mg, 0.20 mmol) under N2 at r.t. for 10 min.

To a solution Cu(OTf)2 (11 mg, 0.03 mmol) was added and stirred for a further 5 hr. The reaction was quenched by sequential additions of Et3N (0.5 mL) and MeOH (0.5 mL). The resulting solution was concentrated in vacuo and purified by flash chromatography (3:2, 1 Pentanes:EtOAc) to afford 12a (51 mg, 85%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.43 – 7.26 (m, 5H), 4.81 (d, J = 11.4 Hz, 1H) 4.74 (d, J = 11.4 Hz, 1H), 4.25 (d, J = 8.1 Hz, 1H), 3.91 (ddd, J = 12.2, 5.3, 2.6 Hz, 1H), 3.77 (ddd, J = 12.2, 8.2, 4.1 Hz, 1H), 3.57 (s, 3H), 3.55 – 3.47 (m, 2H), 3.36 – 3.32 (m, 1H), 3.29 – 3.24 (m, 1H), 2.39 (d, J = 2.7 Hz, 1H), 1.87 (dd, J = 8.2, 5.4 Hz, 1H). 13C

NMR (126 MHz, CDCl3) δ 137.83, 128.68, 128.19, 128.11, 102.95, 77.04, 75.23, 75.13, 74.89, + 66.15, 61.72, 57.29. HRMS m/z calcd. C14H19N3O5 (M+NH4 ) 327.1668, found 327.1669.

Methyl 2-azido-2-deoxy-3-O-methyl-4-O-benzyl-β-D-glucopyranoside (12b)

Procedure identical to the one used for the synthesis of 12a was utilized using 5 (55 mg, 0.17 mmol) as starting material and the product was purified by flash chromatography (2:3, 1 Pentanes:EtOAc) to afford 12b (44 mg, 85%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.39 – 7.29 (m, 5H), 4.84 (d, J = 11.0 Hz, 1H) 4.66 (d, J = 11.0 Hz, 1H), 4.18 (d, J = 8.0 Hz, 1H), 3.86 (ddd, J = 12.0, 5.4, 2.7 Hz, 1H), 3.76 – 3.69 (m, 1H), 3.66 (s, 3H), 3.56 (s, 3H), 3.53 – 3.47 (m, 1H), 3.33 13 – 3.24 (m, 2H), 3.21 – 3.15 (m, 1H). C NMR (126 MHz, CDCl3) δ 137.73, 128.54, 128.12, 128.05,

102.86, 85.12, 77.19, 75.10, 74.99, 65.99, 61.69, 61.04, 57.27. HRMS m/z calcd. C15H21N3O5 + (M+NH4 ) 341.1825, found 341.1817.

Benzyl 2-azido-2-deoxy-4-O-benzyl-β-D-glucopyranoside (12c)

Procedure identical to the one used for the synthesis of 12a was utilized using 11 (83 mg, 0.18 mmol) as starting material and the product was purified by flash chromatography (1:1, 1 Pentanes:EtOAc) to afford 12c (77 mg, 93%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.38 – 7.31 (m, 10H), 4.90 (d, J = 11.4 Hz, 1H), 4.81 (d, J = 11.4 Hz, 1H), 4.78 – 4.68 (m, 2H), 4.43 (d, J = 8.0 Hz, 1H), 3.94 – 3.86 (m, 1H), 3.74 (ddd, J = 12.1, 7.7, 4.4 Hz, 1H), 3.59 – 3.46 (m, 2H), 3.40 – 13 3.29 (m, 2H), 2.41 (br. d, J = 2.5 Hz, 1H), 1.82 (br. t, J = 6.9 Hz, 1H). C NMR (151 MHz, CDCl3) δ 137.85, 136.59, 128.64, 128.51, 128.14, 128.10, 128.08, 127.95, 100.78, 77.06, 75.19, 74.83, + 71.50, 66.25, 61.78, 29.68. HRMS m/z calcd. C20H23N3O5 (M+NH4 ) 403.1981, found 403.1997.

Methyl 2-azido-2-deoxy-6-O-benzyl-β-D-glucopyranoside (12d)

To a stirring solution of 4 (27 mg, 0.09 mmol) in DCM (1 mL) at 0 ºC, Et3SiH (176 µL, 1.1 mmol) and BF3·Et2O (23 µL, 0.18 mmol) were added. The solution was allowed to warm to r.t. over 4 hours and left to stir overnight. The reaction was subsequently quenched with MeOH, concentrated in vacuo and purified by flash chromatography (3:2, Pentanes:EtOAc) to afford 1 12d (21 mg, 76%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.41 – 7.22 (m, 5H), 4.63 (d, J = 11.9 Hz, 1H), 4.57 (d, J = 11.9 Hz, 1H), 4.23 (d, J = 8.0 Hz, 1H), 3.76 (qd, J = 10.2, 5.0 Hz, 2H), 3.63 13 – 3.51 (m, 4H), 3.49 – 3.37 (m, 2H), 3.29 (dd, J = 10.0, 8.0 Hz, 1H). C NMR (126 MHz, CDCl3) δ 137.48, 128.54, 127.99, 127.79, 102.96, 74.95, 73.78, 73.44, 72.31, 70.22, 65.56, 57.15. HRMS + m/z calcd. C14H19N3O5 (M+NH4 ) 327.1668, found 327.1669.

Methyl 2-azido-2-deoxy-3-O-methyl-6-O-benzyl-β-D-glucopyranoside (12e)

Procedure identical to the one used for the synthesis of 12d was utilized using 5 (82 mg, 0.26 mmol) as starting material and the product was purified by flash chromatography (2:1, 1 Pentanes:EtOAc) to afford 12e (56 mg, 67%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.41 – 7.28 (m, 5H), 4.62 (d, J = 12.0 Hz, 1H), 4.57 (d, J = 12.0 Hz, 1H), 4.17 (d, J = 8.1 Hz, 1H), 3.82 – 56

3.71 (m, 2H), 3.72 – 3.49 (m, 7H), 3.47 – 3.36 (m, 1H), 3.28 (dd, J = 9.5, 8.1 Hz, 1H), 3.04 (dd, J = 13 10.2, 9.0 Hz, 1H), 2.86 (br. s, 1H). C NMR (126 MHz, CDCl3) δ 137.54, 128.49, 127.90, 127.75,

102.90, 84.40, 73.75, 73.64, 72.28, 70.24, 65.28, 60.92, 57.10. HRMS m/z calcd. C15H21N3O5 + (M+NH4 ) 341.1825, found 341.1828.

Benzyl 2-azido-2-deoxy-6-O-benzyl-β-D-glucopyranoside (12f)

Procedure identical to the one used for the synthesis of 12d was utilized using 11 (101 mg, 0.26 mmol) as starting material and the product was purified by flash chromatography (7:3, 1 Pentanes:EtOAc) to afford 12f (67 mg, 67%) as a white solid. H NMR (500 MHz, CDCl3) δ 7.40 – 7.32 (m, 5H), 4.92 (d, J = 11.6 Hz, 1H), 4.68 (d, J = 11.6 Hz, 1H), 4.64 (d, J = 12.0 Hz, 1H), 4.58 (d, J = 12.0 Hz, 1H), 4.40 (d, J = 7.8 Hz, 1H), 3.82 – 3.74 (m, 2H), 3.68 – 3.55 (m, 1H), 3.48 – 3.32 (m, 13 3H), 2.93 (br. s, 1H), 2.65 (br. s, 1H). C NMR (126 MHz, CDCl3) δ 137.49, 136.57, 128.54, 128.46, 128.00, 127.98, 127.77, 127.71, 100.57, 74.91, 73.77, 73.55, 72.22, 71.07, 70.22, 65.64. + HRMS m/z calcd. C20H23N3O5 (M+NH4 ) 403.1981, found 403.1987.

General Procedure for Peracetylation (13a-f)

To a solution of 12 (0.2 mmol) in pyridine (1 mL), acetic anhydride (0.5 mL) was added and stirred at r.t. overnight. The solution was concentrated in vacuo and co-evaporated with toluene to obtain 13 in quantitative yield.

Methyl 2-azido-2-deoxy-3,6-di-O-acetyl-4-O-benzyl-β-D-glucopyranoside (13a)

1 H NMR (400 MHz, CDCl3) δ 7.38 – 7.28 (m, 3H), 7.26 – 7.21 (m, 2H), 5.13 – 5.02 (m, 1H), 4.57 (d, J = 11.1 Hz, 1H), 4.53 (d, J = 11.1 Hz, 1H), 4.40 – 4.33 (m, 1H), 4.28 (d, J = 8.2 Hz, 1H), 4.26 – 4.20 (m, 1H), 3.63 – 3.51 (m, 5H), 3.47 – 3.32 (m, 1H), 2.14 – 2.01 (m, 6H). 13C NMR (151

MHz, CDCl3) δ 170.52, 169.69, 137.03, 128.55, 128.16, 128.00, 102.77, 75.79, 74.67, 74.11, + 72.92, 64.15, 62.54, 57.32, 20.89, 20.81. HRMS m/z calcd. C18H23N3O7 (M+NH4 ) 411.1880, found 411.1891.

Methyl 2-azido-2-deoxy-3-O-methyl-4-O-benzyl-6-O-acetyl-β-D-glucopyranoside (13b)

1 H NMR (600 MHz, CDCl3) δ 7.37 – 7.30 (m, 5H), 4.84 (d, J = 10.8 Hz, 1H), 4.58 (d, J = 10.8 Hz, 1H), 4.32 (dd, J = 12.0, 1.6 Hz, 1H), 4.24 – 4.18 (m, 1H), 4.14 (s, J = 8.1 Hz, 1H), 3.67 (s, 3H), 3.55 (s, 3H), 3.48 – 3.41 (m, 2H), 3.30 (dd, J = 9.9, 8.1 Hz, 1H), 3.22 – 3.14 (m, 1H), 2.04 (d, J = 13 0.7 Hz, 3H). C NMR (126 MHz, CDCl3) δ 170.69, 137.45, 128.55, 128.20, 128.12, 102.77, 85.34, + 77.17, 74.96, 72.89, 65.91, 62.79, 61.07, 57.18, 20.84. HRMS m/z calcd. C17H23N3O6 (M+NH4 ) 383.1931, found 383.1937.

Benzyl 2-azido-2-deoxy-4-O-benzyl-3,6-di-O-acetyl-β-D-glucopyranoside (13c)

1 H NMR (500 MHz, CDCl3) δ 7.43 – 7.29 (m, 8H), 7.25 – 7.21 (m, 2H), 5.04 (dd, J = 10.4, 8.8 Hz, 1H), 4.92 (d, J = 11.7 Hz, 1H), 4.69 (d, J = 11.7 Hz, 1H), 4.57 (d, J = 11.1 Hz, 1H), 4.53 (d, J = 11.1 Hz, 1H), 4.44 (d, J = 8.0 Hz, 1H), 4.39 (dd, J = 12.0, 2.1 Hz, 1H), 4.22 (dd, J = 12.0, 4.6 Hz, 1H), 3.62 – 3.51 (m, 2H), 3.45 (dd, J = 10.4, 8.0 Hz, 1H), 2.09 (s, 3H), 2.06 (s, 3H). 13C NMR (126

MHz, CDCl3) δ 170.57, 169.72, 137.03, 136.19, 128.57, 128.53, 128.18, 128.13, 127.98, 113.90, 100.23, 75.80, 74.67, 74.07, 72.99, 71.22, 64.17, 62.60, 20.93, 20.87. HRMS m/z calcd. + C24H27N3O7 (M+NH4 ) 487.2193, found 487.2209.

Methyl 2-azido-2-deoxy-3,4-di-O-acetyl-6-O-benzyl-β-D-glucopyranoside (13d)

1 H NMR (500 MHz, CDCl3) δ 7.39 – 7.26 (m, 5H), 5.08 – 4.93 (m, 2H), 4.59 (d, J = 12.1 Hz, 1H) 4.49 (d, J = 12.1 Hz, 1H), 4.28 (d, J = 8.0 Hz, 1H), 3.65 – 3.60 (m, 1H), 3.60 (s, 3H), 3.58 – 3.52 13 (m, 2H), 3.49 – 3.45 (m, 1H), 2.07 (s, 3H), 1.89 (s, 3H). C NMR (151 MHz, CDCl3) δ 170.07, 169.65, 137.59, 128.38, 127.86, 127.78, 102.78, 73.57, 73.23, 72.80, 69.16, 68.43, 62.75, 57.33, + 29.68, 20.71. HRMS m/z calcd. C18H23N3O7 (M+NH4 ) 411.1880, found 411.1890.

Methyl 2-azido-2-deoxy-3-O-methyl-4-O-acetyl-6-O-benzyl-β-D-glucopyranoside (13e)

1 H NMR (600 MHz, CDCl3) δ 7.37 – 7.27 (m, 5H), 4.96 – 4.89 (m, 1H), 4.53 (s, 2H), 4.18 (d, J = 8.1 Hz, 1H), 3.59 – 3.48 (m, 9H), 3.41 – 3.32 (m, 1H), 3.19 (t, J = 9.6 Hz, 1H), 1.99 (s, 3H). 13C

NMR (151 MHz, CDCl3) δ 169.59, 137.73, 128.35, 127.80, 127.71, 102.68, 82.35, 73.64, 73.41,

+ 70.81, 69.36, 65.17, 60.15, 57.15, 20.84. HRMS m/z calcd. C17H23N3O6 (M+NH4 ) 383.1931, found 383.1945.

Benzyl 2-azido-2-deoxy-3,4-di-O-acetyl-6-O-benzyl-β-D-glucopyranoside (13f)

1 H NMR (500 MHz, CDCl3) δ 7.38 – 7.28 (m, 10H), 5.05 – 5.00 (m, 1H), 4.97 – 4.92 (m, 2H), 4.71 (d, J = 11.9 Hz, 1H), 4.61 (d, J = 11.9 Hz, 1H), 4.53 – 4.49 (m, 1H), 4.44 (d, J = 8.1 Hz, 13 1H), 3.63 – 3.51 (m, 4H), 2.07 (s, 3H), 1.90 (s, 3H). C NMR (126 MHz, CDCl3) δ 170.05, 169.65, 137.64, 136.27, 128.52, 128.40, 128.14, 128.10, 127.82, 127.78, 73.57, 73.29, 72.72, 71.17, + 69.18, 68.53, 63.75, 20.71, 20.58. HRMS m/z calcd. C24H27N3O7 (M+NH4 ) 487.2193, found 487.2204.

General Procedure for Phosphonamidate Synthesis (14a-f)

To a solution of 13 (0.2 mmol) in dry DCM (1 mL), diethyl methylphosphonite (2 mmol) was added and stirred at r.t. for 1 hr. To the resulting solution H2O (1 mL) was added and left to stir overnight. The solvent and excess reagent were evaporated in vacuo and purified by flash chromatography (2:3, EtOAc:Toluene) to obtain 14 as a pair of diastereomers.

N-methylethoxyphosphinyl-1-O-methyl-3,6-di-O-acetyl-4-O-benzyl-β-D-glucosamine (14a)

1 (Yield: 64%) H NMR (500 MHz, CDCl3) δ 7.39 – 7.19 (m, 5H), 5.00 (dd, J = 10.6, 8.9 Hz, 1H), 4.62 – 4.51 (m, 2H), 4.42 – 4.28 (m, 1H), 4.25 – 3.86 (m, 4H), 3.65 – 3.34 (m, 4H), 3.31 – 3.15 (m, 1H), 2.59 – 2.43 (m, 1H), 2.12 – 1.96 (m, 6H), 1.49 – 1.19 (m, 6H). 13C NMR (126 MHz,

CDCl3) δ 171.14, 170.93, 170.62, 170.61, 137.25, 137.23, 128.56, 128.53, 128.12, 127.98, 127.92, 127.86, 104.19, 104.08, 76.29, 76.08, 75.99, 75.89, 74.74, 74.72, 72.87, 72.85, 62.64, 59.78, 59.73, 59.43, 56.87, 56.25, 56.14, 21.13, 21.10, 20.82, 16.49, 16.44, 16.35, 16.29, 15.33, 31 15.01, 14.26, 13.93. P NMR (243 MHz, CDCl3) δ 32.47, 31.68. HRMS m/z calcd. C21H32NO9P (M+H+) 474.1893, found 474.1914.

N-methylethoxyphosphinyl-1,3-di-O-methyl-4-O-benzyl-6-O-acetyl-β-D-glucosamine (14b)

1 (Yield: 61%) H NMR (500 MHz, CDCl3) δ 7.42 – 7.24 (m, 5H), 4.87 – 4.71 (m, 1H), 4.62 – 4.50 (m, 1H), 4.38 – 4.26 (m, 1H), 4.28 – 4.16 (m, 1H), 4.18 – 4.01 (m, 2H), 3.75 – 3.36 (m, 9H), 3.22 – 3.03 (m, 2H), 2.92 – 2.74 (m, 1H), 2.07 – 2.01 (m, 3H), 1.60 – 1.37 (m, 3H), 1.29 (m, 3H). 13 C NMR (126 MHz, CDCl3) δ 170.72, 137.60, 137.58, 128.50, 128.00, 104.24, 104.22, 103.89, 102.75, 86.68, 86.36, 85.32, 78.20, 78.19, 78.08, 77.15, 74.64, 72.70, 65.90, 63.00, 61.44, 61.16, 59.40, 59.35, 59.30, 57.49, 57.32, 57.21, 56.93, 20.86, 16.33, 16.25, 15.02, 14.48, 13.95, 13.33. 31 + P NMR (243 MHz, CDCl3) δ 33.69, 33.55. HRMS m/z calcd. for C20H32N1O8P1 (M+H ) 446.1944, found 446.1941.

N-methylethoxyphosphinyl-1,4-di-O-benzyl-3,6-di-O-acetyl-β-D-glucosamine (14c)

1 (Yield: 78%) H NMR (500 MHz, CDCl3) δ 7.40 – 7.15 (m, 10H), 4.97 (dd, J = 10.5, 8.9 Hz, 1H), 4.89 (m, 1H), 4.64 – 4.53 (m, 3H), 4.39 (m, 1H), 4.34 – 4.16 (m, 2H), 3.98 (m, 1H), 3.94 – 3.77 (m, 1H), 3.62 (m, 1H), 3.57 – 3.48 (m, 1H), 3.43 – 3.25 (m, 1H), 2.45 - 2.28 (m, 1H), 2.10 – 13 2.03 (m, 6H), 1.42 – 1.33 (m, 3H), 1.23 – 1.15 (m, 3H). C NMR (126 MHz, CDCl3) δ 171.13, 170.93, 170.62, 137.26, 136.34, 128.60, 128.56, 127.89, 127.84, 101.27, 101.16, 77.26, 77.21, 76.40, 76.37, 76.19, 76.16, 76.04, 75.98, 74.67, 74.66, 72.98, 70.72, 70.61, 62.75, 62.73, 59.89, 59.84, 59.54, 59.49, 56.08, 56.02, 29.68, 21.18, 21.13, 20.86, 16.37, 16.33, 16.27, 15.36, 14.97, 31 + 14.29, 13.91. P NMR (243 MHz, CDCl3) δ 32.43, 31.57. HRMS m/z calcd. C27H36NO9P (M+H ) 550.2206, found 550.2189.

N-methylethoxyphosphinyl-1-O-methyl-3,4-di-O-acetyl-6-O-benzyl-β-D-glucosamine (14d)

1 (Yield: 54%) H NMR (500 MHz, CDCl3) δ 7.39 – 7.20 (m, 5H), 5.17 – 4.92 (m, 2H), 4.60 – 4.47 (m, 2H), 4.19 (m, 1H), 4.13 – 4.00 (m, 1H), 4.01 – 3.87 (m, 1H), 3.67 – 3.47 (m, 6H), 3.30 – 3.15 (m, 1H), 2.48 - 2.32 (m, 1H), 2.07 – 2.05 (m, 3H), 1.92 – 1.89 (m, 3H), 1.55 – 1.36 (m, 3H), 13 1.30 – 1.25 (m, 3H). C NMR (126 MHz, CDCl3) δ 170.93, 170.78, 169.51, 137.70, 137.68, 128.36, 127.82, 127.73, 104.09, 104.07, 74.28, 73.59, 73.28, 69.61, 68.94, 59.53, 56.95, 56.10,

31 20.88, 20.62, 16.46, 15.12, 14.04. P NMR (243 MHz, CDCl3) δ 32.35, 32.05. HRMS m/z calcd. + C21H32NO9P (M+H ) 474.1893, found 474.1876.

N-methylethoxyphosphinyl-1,3-di-O-methyl-4-O-acetyl-6-O-benzyl-β-D-glucosamine (14e)

1 (Yield: 65%) H NMR (500 MHz, CDCl3) δ 7.36 – 7.24 (m, 5H), 5.00 – 4.88 (m, 1H), 4.52 (s, 2H), 4.18 – 3.98 (m, 3H), 3.56 – 3.45 (m, 8H), 3.29 – 3.17 (m, 2H), 2.98 – 2.85 (m, 1H), 1.98 (s, 13 3H), 1.57 – 1.44 (m, 3H), 1.27 (m, 3H). C NMR (126 MHz, CDCl3) δ 169.68, 137.82, 137.47, 128.33, 128.30, 127.79, 127.67, 104.22, 103.91, 83.70, 83.52, 73.59, 73.22, 71.27, 69.47, 59.73, 59.47, 56.97, 56.76, 55.44, 54.45, 20.93, 16.20, 14.98, 14.39, 13.90, 13.31. 31P NMR (202 MHz, + CDCl3) δ 34.50, 34.42. HRMS m/z calcd. C20H32NO8P (M+H ) 446.1944, found 446.1955.

N-methylethoxyphosphinyl-1,6-di-O-benzyl-3,4-di-O-acetyl-β-D-glucosamine (14f)

1 (Yield: 67%) H NMR (600 MHz, CDCl3) δ 7.49 – 7.30 (m, 5H), 5.13 – 4.99 (m, 1H), 4.97 – 4.87 (m, 2H), 4.63 – 4.51 (m, 3H), 4.37 – 4.30 (m, 1H), 4.04 – 3.78 (m, 2H), 3.63 – 3.53 (m, 3H), 3.36 – 3.23 (m, 1H), 2.48 – 2.34 (m, 2H), 2.07 – 2.02 (m, 3H), 1.92 – 1.88 (m, 3H), 1.41 – 1.35 (m, 13 3H), 1.24 – 1.10 (m, 3H). C NMR (126 MHz, CDCl3) δ 170.87, 170.76, 169.57, 169.53, 137.74, 137.72, 136.32, 136.31, 128.69, 128.58, 128.49, 128.48, 128.38, 128.16, 128.15, 127.79, 127.78, 127.76, 127.75, 127.74, 101.35, 101.21, 74.49, 73.60, 73.58, 73.35, 70.92, 70.85, 69.69, 69.60, 69.01, 59.67, 57.30, 56.87, 55.96, 20.91, 20.62, 16.39, 16.22, 15.15, 14.95, 14.07, 13.89. 31P + NMR (243 MHz, CDCl3) δ 32.29, 31.97. HRMS m/z calcd. C27H36NO9P (M+H ) 550.2206, found 550.2226.

General Procedure for Phosphonamidate Deprotection (15a-f)

To a solution of 14 (0.2 mmol) in THF (2 mL), NaOH (1 mmol) dissolved in H2O (2 mL) was added and stirred at r.t. overnight (14b and 14e required overnight reflux). The solvent was evaporated in vacuo and the resulting residue was redissolved in a minimal amount of H2O and desalted on HPLC p2 size exclusion column. Lyophilization afforded 15 as a white solid.

Sodium N-methylphosphate-1-O-methyl-4-O-benzyl-β-D-glucosamine (15a)

1 (Yield: 85%) H NMR (400 MHz, D2O) δ 7.52 – 7.41 (m, 5H), 4.91 (d, J = 10.6 Hz, 1H), 4.72 (d, J = 10.6 Hz, 1H), 4.32 (d, J = 8.2 Hz, 1H), 3.96 – 3.88 (m, 1H), 3.74 (dd, J = 12.2, 5.2 Hz, 1H), 3.63 – 3.52 (m, 4H), 3.55 – 3.46 (m, 2H), 2.93 – 2.81 (m, 1H), 1.36 (d, J = 16.0 Hz, 3H). 13C NMR

(126 MHz, D2O) δ 137.05, 128.82, 128.66, 128.44, 103.92, 77.96, 76.49, 74.68, 60.60, 58.21, 31 - - 57.15, 15.88, 14.87. P NMR (162 MHz, D2O) δ 26.54. HRMS m/z calcd. [C15H23NO7P] (M ) 360.1218, found 360.1218.

Sodium N-methylphosphate-1,3-di-O-methyl-4-O-benzyl-β-D-glucosamine (15b)

1 (Yield: 95%) H NMR (600 MHz, D2O) δ 7.52 – 7.43 (m, 5H), 4.82 (d, J = 10.5 Hz, 1H), 4.74 (d, J = 10.5 Hz, 1H), 4.28 (d, J = 8.2 Hz, 1H), 3.93 (dd, J = 12.3, 2.2 Hz, 1H), 3.76 (dd, J = 12.3, 5.7 Hz, 1H), 3.69 (s, 3H), 3.57 (s, 3H), 3.55 – 3.50 (m, 1H), 3.48 – 3.42 (m, 1H), 3.31 (dd, J = 9.9, 9.0 13 Hz, 1H), 2.98 – 2.91 (m, 1H), 1.34 (d, J = 16.0 Hz, 3H). C NMR (126 MHz, D2O) δ 136.81, 129.01, 128.70, 128.55, 104.57, 85.93, 77.43, 74.89, 74.68, 60.52, 57.39, 57.23, 16.66, 15.64. 31P NMR - - (243 MHz, D2O) δ 24.83. HRMS m/z calcd. [C16H25NO7P] (M ) 374.1374, found 374.1383.

Sodium N-methylphosphate-1,4-di-O-benzyl-β-D-glucosamine (15c)

1 (Yield: 92%) H NMR (400 MHz, D2O) δ 7.55 – 7.39 (m, 10H), 4.93 (dd, J = 16.9, 11.1 Hz, 2H), 4.80 – 4.69 (m, 2H), 4.49 (d, J = 8.3 Hz, 1H), 3.92 (dd, J = 12.2, 2.1 Hz, 1H), 3.74 (dd, J = 12.3, 13 5.4 Hz, 1H), 3.61 – 3.42 (m, 3H), 2.91 (m, 1H), 1.28 (d, J = 16.0 Hz, 3H). C NMR (126 MHz, D2O) δ 137.06, 136.60, 128.86, 128.82, 128.66, 128.60, 128.44, 128.33, 101.59, 77.94, 76.75, 74.74, 31 74.60, 71.54, 60.64, 58.20, 15.95, 14.94. P NMR (162 MHz, D2O) δ 26.41. HRMS m/z calcd. - - [C21H27NO7P] (M ) 436.1531, found 436.1529.

Sodium N-methylphosphate-1-O-methyl-6-O-benzyl-β-D-glucosamine (15d)

1 (Yield: 87%) H NMR (500 MHz, D2O) δ 7.36 – 7.23 (m, 5H), 4.51 (s, 2H), 4.16 (d, J = 8.3 Hz, 1H), 3.78 (dd, J = 11.3, 2.1 Hz, 1H), 3.56 (dd, J = 11.3, 6.8 Hz, 1H), 3.45 – 3.42 (m, 1H), 3.41 (s, 3H), 3.29 – 3.23 (m, 2H), 2.71 - 2.63 (m, 1H), 1.19 (d, J = 16.0 Hz, 3H). 13C NMR (126 MHz,

D2O) δ 137.18, 128.63, 128.33, 128.21, 103.93, 76.25, 74.36, 73.08, 70.19, 69.15, 57.92, 57.19, 31 - - 15.76, 14.75. P NMR (162 MHz, D2O) δ 26.66. HRMS m/z calcd. [C15H23NO7P] (M ) 360.1218, found 360.1210.

Sodium N-methylphosphate-1,3-di-O-methyl-6-O-benzyl-β-D-glucosamine (15e)

1 (Yield: 92%) H NMR (400 MHz, D2O) δ 7.50 – 7.43 (m, 5H), 4.66 (s, 2H), 4.27 (d, J = 8.2 Hz, 1H), 3.92 (dd, J = 11.3, 2.0 Hz, 1H), 3.74 – 3.42 (m, 9H), 3.20 (dd, J = 10.0, 9.1 Hz, 1H), 2.96 – 13 2.87 (m, 1H), 1.33 (d, J = 16.0 Hz, 3H). C NMR (126 MHz, D2O) δ 162.02, 137.19, 128.64, 128.32, 128.21, 104.65, 85.59, 74.17, 73.08, 69.43, 69.08, 62.40, 59.78, 57.29, 56.82, 16.69, 31 - - 15.67. P NMR (162 MHz, D2O) δ 25.03. HRMS m/z calcd. [C16H25NO7P] (M ) 374.1374, found 374.1372.

Sodium N-methylphosphate-1,6-di-O-benzyl-β-D-glucosamine (15f)

1 (Yield: 91%) H NMR (400 MHz, D2O) δ 7.56 – 7.35 (m, 5H), 4.87 (d, J = 11.6 Hz, 1H), 4.72 (d, J = 11.6 Hz, 1H), 4.64 (s, 2H), 4.62 – 4.58 (m, 1H), 4.46 (d, J = 8.3 Hz, 1H), 3.96 – 3.89 (m, 1H), 3.70 (dd, J = 11.4, 6.8 Hz, 1H), 3.61 – 3.48 (m, 1H), 3.43 – 3.33 (m, 2H), 2.85 (m, 1H), 1.29 – 1.23 13 (m, 3H). C NMR (126 MHz, D2O) δ 137.29, 136.66, 128.80, 128.64, 128.61, 128.59, 128.29, 128.19, 101.65, 76.47, 74.47, 73.07, 71.64, 70.19, 69.15, 57.93, 15.86, 14.85. 31P NMR (243 - - MHz, D2O) δ 26.51. HRMS m/z calcd. [C21H27NO7P] (M ) 436.1531, found 436.1526.

NMR Spectra

100

MALDI-MS Analysis

101

102

Inhibition Curves from Fluorescamine assays with SpPgdA

103

104

Inhibition Curves from Fluorogenic assays with PgaB

105

106

107

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