Antimicrobial Agents Targeting Early-Stage Isoprenoid

ANTIMICROBIAL AGENTS TARGETING EARLY-STAGE ISOPRENOID

BIOSYNTHESIS: DESIGN, SYNTHESIS AND EVALUATION OF

ACETYLPHOSPHONATE INHIBITORS OF DXP SYNTHASE

by Ryan J. Vierling

A dissertation submitted to The Johns Hopkins University in conformity with the requirements of the degree of Doctor of Philosophy

Baltimore, MD October 2014 Abstract

The global threat of antibiotic resistant bacteria necessitates renewed efforts at designing antibiotics to tackle this healthcare challenge. Battling or overcoming this hazard will require the united efforts of chemists and microbiologists to discover new targets for antibacterial agents and design new molecular scaffolds to inhibit these unique targets. In addition to finding these new targets, scientists need to better understand the complicated, endogenous defenses of bacteria, including the up-regulation of promiscuous multi-drug resistant (MDR) efflux pumps.

One potential new target for antibacterial development is early-stage isoprenoid biosynthesis via the methylerythritol phosphate (MEP) pathway essential to bacteria, parasites and some plants but absent in humans which instead utilize the orthogonal mevalonate pathway. The first catalytic step in the MEP pathway involves the condensation of pyruvate and D-glyceraldehyde 3-phosphate (D-GAP) in a thiamin diphosphate (ThDP)-dependent manner, to generate 1-deoxy-D-xylulose 5-phosphate

(DXP) and carbon dioxide. This initial step is catalyzed by the unique biosynthetic enzyme, DXP synthase, and the product represents a metabolic branch point as DXP is the precursor for isoprenoid, thiamin and pyridoxal biosynthesis in bacteria. The importance of these three pathways in bacterial growth and division highlights DXP synthase as an attractive target for antibacterial development.

DXP synthase is promiscuous in reference to acceptor substrates and possesses a unique domain arrangement and catalytic mechanism, which involves formation of a ternary complex, suggesting that selective inhibition should be achievable by using unnatural bisubstrate analogs which mimic potential donor and acceptor substrates. ii

Towards this goal, we successfully designed, synthesized and evaluated a series of acetylphosphonate inhibitors which interact with the ThDP cofactor to form a phosphonolactylthiamin diphosphate (PLThDP) intermediate, which cannot undergo decarboxylation to form product. Two compounds in this series, butylacetylphosphonate

(BAP) and benzylacetylphosphonate (BnAP) were shown to be selective inhibitors of DXP synthase.

Further, antimicrobial studies show that these compounds can weakly inhibit bacterial cell growth in complex media. Rescue experiments with downstream metabolites and target overexpression strains confirm that they restrict bacterial growth by a mechanism involving inhibition of DXP synthase. One potential contributing factor to their weak antimicrobial activity is their susceptibility to efflux by the E. coli MDR pump,

AcrAB-TolC. We report an observed trend with the length of the alkyl chain of the acetylphosphonate and the potential for uptake and efflux towards defining the optimal properties for a future generation of potent antimicrobial agents targeting DXP synthase.

Advisor: Dr. Caren L. Freel Meyers

2nd Reader: Dr. Paul S. Miller

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Acknowledgements

This dissertation represents the culmination of a six year, arduous journey through graduate school which was only possible because of the love and support of my family, friends and coworkers. I have deeply appreciated every opportunity to grow and develop as a scientist, and I have been blessed to be surrounded by positive, astoundingly intelligent and insightful individuals throughout my journey.

First I would like to thank my own research lab, guided and fostered by our advisor,

Professor Caren Freel Meyers. I am grateful for her advice, support and encouragement.

She has pushed me for over five years to be a better scientist and communicator. I strive to emulate her scientific passion and organization, and I will always appreciate her advice and sense of humor. I would also like to thank my coworker, Dr. Jessica Mott Smith for being an excellent collaborator and friend. We make an excellent team in the lab and I always admired her work ethic and focus. I would like to thank Dr. Francine Morris for her collaboration and friendship. I am thankful for Sara Sanders agreeing to take on parts of this project and to Alicia DeColli for reading and editing my thesis, and to both of them for their energy and humor. Drs. Kip Bitok, Marie Webster and Leighanne Basta are great friends and I have enjoyed working with and learning from each of them.

I would like to thank the members of my thesis committee, Professors Phil Cole,

Craig Townsend and Jun Liu for their support, advice and patience. I received so much insight about research and presenting ideas and results from my interactions with them and

I am forever grateful. Professor Paul Miller gave freely of his time to read and make recommendations to this thesis and I am grateful to him and to Professor Sean Taverna for attending my thesis defense on short notice. I would also like to thank my undergraduate iv research advisors, Professor Gordon H. Purser and Dr. David J. Vanderah for their support and wisdom in starting my career in chemical research.

I have to also thank the institutional support which brought me to Baltimore and started my graduate research at Johns Hopkins. The Chemistry-Biology Interface (CBI) program, which grew under the careful attention of the first Director, Professor Marc

Greenberg and continues to flourish under the advisement of the current Director, Professor

Steven Rokita. I was welcomed my first day in Baltimore by the academic coordinator

Lauren McGhee and I am forever grateful for her advice, organization and friendship. I would also like to thank the NIH for their financial support of the CBI program through our training grant (T32-GM08018901) and for their support of the work in the Freel Meyers lab via an R-O1 grant (GM084998).

Baltimore felt much more like home thanks to the friendship of Jessica Popkin,

Paul Peters, Dr. Phillip Flanders, Dr. John Sivey, Kristopher Thornsbury and Shawn

Lowery, among countless others. I am indebted to them for their humor, company and for always making me feel welcomed.

Most of all I want to thank my family for their support and encouragement from the beginning. My sister Kelly and brother Michael are some of my first friends and I could not be more proud of either of them. They are a great support to me and I love them both. My mother and father are truly two of the hardest-working, most loving and supportive parents I could have asked for. Jim Vierling is an excellent role model, father and the epitome of work ethic. Kim Vierling is a tireless mother, ardent supporter of her children and a generally loving and caring individual. I would not be the man I am today without their friendship, love and support. I strive always to live up to their example. v

This thesis is dedicated to Ginny Vierling and Art Murphy.

Table of Contents

Chapter 1. Introduction ...... 1

1.1 Discovery of antibiotics and development of clinical resistance ...... 1

1.1.1 A history of antibiotics and their molecular targets ...... 3

1.1.2 Efflux pump-mediated resistance ...... 8

1.2 Isoprenoid biosynthesis is an underutilized target in antibiotic development ...... 11

1.2.1 The discovery of the non-mevalonate pathway for isoprenoid biosynthesis .... 13

1.2.2 Current efforts to target enzymes of the MEP pathway ...... 15

1.2.3 Regulation in the MEP pathway...... 18

1.2.4 1-Deoxy-D-xylulose 5-phosphate (DXP) synthase is a favorable target for design

of new antibiotics ...... 22

1.3 Reactive substrate mimics as inhibitors of DXP synthase ...... 30

References ...... 32

Chapter 2. Electrophilic phosphonates as selective inhibitors of bacterial DXP synthase ...... 41

Introduction ...... 41

Results ...... 48

2.1 Synthesis of unbranched acetylphosphonates (2.1 – 2.2) ...... 48

2.2 Synthesis of unbranched acetylphosphonates (2.3 – 2.5) ...... 48

2.3 Synthesis of unbranched acetylphosphonates (2.6 – 2.8) ...... 49

2.4 Synthesis of phenyl acetylphosphonates...... 51

2.5 Synthesis of aromatic acetylphosphonates (2.10 – 2.17) ...... 52

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2.6 Evaluation of acylphosphonates (2.1 – 2.8) as inhibitors of E. coli DXP synthase 54

2.7 Evaluation of acylphosphonates (2.1 – 2.8) as inhibitors of the E1 subunit of porcine

heart pyruvate dehydrogenase complex (PDH) ...... 56

2.8 Alkyl branching proximal to the acetylphosphonate scaffold is not tolerated by DXP

synthase ...... 59

2.9 BnAP is the most selective acetylphosphonate inhibitor of DXP synthase discovered

to date ...... 60

2.10 An oxime of methylacetylphosphonate is a weak, time-dependent inhibitor of DXP

synthase ...... 64

2.11 Methyl(chloromethyl)phosphonate is not an inhibitor of DXP synthase ...... 65

Discussion ...... 67

Experimental ...... 70

NMR data ...... 93

References ...... 111

Chapter 3. Antimicrobial activity of acetylphosphonates ...... 115

Introduction ...... 115

Results ...... 117

3.1 BAP exhibits weak antimicrobial activity ...... 117

3.2 BAP inhibition of E. coli growth is rescued by downstream metabolites, and target

overexpression ...... 119

3.3 Other acetylphosphonates inhibit E. coli growth in a dose-dependent manner ..... 122

3.4 DXP synthase overexpression protects E. coli from acetylphosphonate treatment

...... 124

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3.5 Bacterial resistance to alkylacetylphosphonates includes drug efflux by E. coli .. 125

3.6 PAβN enhances activity of BAP and BnAP, but not MAP or OctAP ...... 130

3.7 Growth inhibition caused by fosmidomycin is significantly enhanced when

administered in combination with APs ...... 133

3.8 OctAP is more toxic to Plasmodium falciparum than BAP ...... 135

Discussion ...... 137

Experimental ...... 140

References ...... 142

Curriculum Vitae ...... 146

List of Tables

Chapter 1. Introduction

Table 1-1 Aromatic nitroso substrates are better substrates for DXP synthase than PDH

...... 27

Chapter 2. Electrophilic phosphonates as selective inhibitors of bacterial DXP synthase

Table 2-1 Inhibition of DXP synthase by the alkyl acylphosphonates, 2.1 – 2.8 ...... 55

Table 2-2 Inhibition of the ThDP-dependent E1 subunit of PDH by alkyl

acylphosphonates, 2.1 – 2.8 ...... 59

Table 2-3 Evaluating aromatic acetylphosphonates (2.10 – 2.17) inhibitors of DXP

synthase and PDH ...... 63

Chapter 3. Antimicrobial activity of acetylphosphonates

Table 3-1 The antimicrobial properties of BAP against a panel of bacterial pathogens

...... 119

Table 3-2 BAP synergizes with fosmidomycin and ampicillin against E. coli ...... 122

List of Figures

Chapter 1. Introduction

Figure 1-1 The timeline of discovery of antibiotic classes (top) and observation of

clinical resistance (bottom) ...... 2

Figure 1-2 Summary of the modes of antibiotic resistance employed by bacteria ...... 3

Figure 1-3 The molecular targets of antibiotics are grouped into five primary cellular

processes ...... 4

Figure 1-4 The four primary classes of MDR pump in bacteria, and some observed

substrates ...... 9

Figure 1-5 Molecular model of AcrAB-TolC ...... 10

Figure 1-6 The mammalian mevalonate pathway of isoprenoid biosynthesis ...... 13

Figure 1-7 The non-mammalian MEP pathway generates the precursors, DMADP and

IDP, in many bacteria, plants and apicocomplexan parasites ...... 15

Figure 1-8 Inhibitors of the enzymes of the non-mammalian MEP pathway...... 17

Figure 1-9 Alternatives to DXP synthase for the intracellular formation of DXP ...... 21

Figure 1-10 Low micromolar inhibitors of DXP synthase described to date ...... 22

Figure 1-11 The crystal structures of the related THDP-dependent enzymes, DXP

synthase, TK and PDH ...... 24

Figure 1-12 The calculated active site volumes of DXP synthase, TK and PDH ...... 25

Figure 1-13 Alternative aldehyde substrates for DXP synthase and isolated products 26

Figure 1-14 ThDP-dependent catalysis in DXP synthase ...... 28

Figure 1-15 The random sequential, preferred ordered mechanism of DXP synthase . 29

Chapter 2. Electrophilic phosphonates as selective inhibitors of bacterial DXP synthase

Figure 2-1 The MEP pathway is composed of seven biosynthetic enzymes to generate

the isoprenoid precursors, DMADP and IDP ...... 42

Figure 2-2 The DXP synthase catalytic mechanism requires both substrates to be present

in order for decarboxylation and product formation to occur ...... 43

Figure 2-3 Inhibition of DXP synthase by MAP and selective inhibitor design ...... 45

Figure 2-4 Acylphosphonates synthesized and evaluated as selective inhibitors of DXP

synthase ...... 47

Figure 2-5 Synthetic scheme for alkyl acylphosphonates (2.1 – 2.8) ...... 50

Figure 2-6 The inherent reactivity of the phenylacetylphosphonates favors

bisphosphonate formation...... 51

Figure 2-7 General synthetic route to aromatic acetylphosphonates (2.10 – 2.17) ...... 53

Figure 2-8 Competitive inhibition by compounds 2.1 – 2.6 for DXP synthase ...... 56

Figure 2-9 Competitive inhibition of porcine PDH E1 subunit by compounds 2.1 – 2.5

...... 58

Figure 2-10 Isopropylacetylphosphonate (2.9) is a weak inhibitor of DXP synthase .. 60

Figure 2-11 Evaluation of the inhibition of DXP synthase and PDH by BnAP (2.10) 61

Figure 2-12 The oxime of MAP shows weak, time-dependent inhibition of DXP

synthase ...... 65

Figure 2-13 Irreversible inhibition of ThDP-dependent enzymes by F-pyr and proposed

mechanism for inhibition by halomethylphosphonates ...... 66

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Chapter 3. Antimicrobial activity of acetylphosphonates

Figure 3-1 DXP synthase is a key metabolic intermediate for isoprenoid production via

the MEP pathway and vitamin B1 and B6 production in bacteria ...... 117

Figure 3-2 Overexpression of DXP synthase protects from the antimicrobial effects of

BAP ...... 121

Figure 3-3 In vitro inhibition of DXP synthase and antimicrobial effects of

acetylphosphonates against E. coli (MG 1655) ...... 123

Figure 3-4 Increasing intracellular DXP synthase levels results in partial rescue of E.

coli growth in the presence of alkylacetylphosphonates ...... 125

Figure 3-5 Antimicrobial activity of the acetylphosphonates against E. coli BW25113

...... 127

Figure 3-6 Acetylphosphonates are substrates for efflux via the AcrAB-TolC transporter

...... 128

Figure 3-7 MAP and EAP are inactive against E. coli BW25113 and the ΔtolC variant.

...... 129

Figure 3-8 PAβN, but not MBX2319, potentiates antimicrobial activity of BAP and

BnAP against E. coli ...... 132

Figure 3-9 Potentiation of fosmidomycin activity by acetylphosphonates ...... 134

Figure 3-10 OctAP is more toxic than BAP to P. falciparum ...... 136

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Chapter 1. Introduction

1.1 Discovery of antibiotics and development of clinical resistance

The importance of antibiotics in modern medicine cannot be overstated. Their relative safety, coupled with reduction of mortality rates caused by childbirth, the development of modern surgery and organ transplantation procedures, and even treatment of ulcers and relatively minor skin cuts were all revolutionized by the discovery of antibiotics. The ubiquitous and relatively inexpensive use of antibiotics is so ingrained into modern medicine that it is hard to imagine how physicians and patients would adapt to circumstances when their use is no longer efficacious [1].

Modern antibiotics are predominately natural products or semisynthetic molecules based on a natural product scaffold. The story of Alexander Fleming’s auspicious discovery of penicillin in 1928 [2], immortalized in scientific lore, is the earliest laboratory discovery of these medicines. The existence of natural sources of these molecules with antibacterial properties is evidence of enduring “chemical warfare” between microbes and also provides an evolutionary cause for resistance [3]. Antibiotic molecules produced by a microbial species exert selective pressure on other species to adapt. An amazing aspect of antibiotic resistance is how quickly it develops. Figure 1-1 illustrates how clinical resistance can be observed within a few years of introducing a new antibiotic molecule for general use [4].

Figure 1-1. The timeline of discovery of antibiotic classes (top) and observation of clinical resistance (bottom) [4].

The rapid development of antibiotic resistance is the result of a combination of factors [3, 5-7], including endogenous physiochemical defenses (i.e., the outer membrane and efflux pumps in Gram-negative species), the SOS response pathway which plays a role in the accumulation of mutations leading to a resistance phenotype, and the accumulation of plasmids which can disseminate resistance gene products throughout a population [8].

These origins of resistance are broadly represented in Figure 1-2. The mechanisms of resistance all involve preventing the antibiotic from finding and binding its molecular target, by sequestration, target modification or active efflux. The combination of traits that encompass “antibiotic resistance” ensures that there is no universal or trivial solution to the problem of antibiotic resistance [9]. Tackling this challenging problem is going to require a combination of approaches including discovery and development of new cellular targets and new molecular scaffolds to overcome existing resistance mechanisms, and a better understanding of the endogenous defense mechanisms including small molecule uptake and efflux pumps. 2

Figure 1-2. Summary of the modes of antibiotic resistance employed by bacteria.

1.1.1 A history of antibiotics and their molecular targets

The decades between 1930 and 1960 saw the discovery and deployment of at least

20 antibiotic classes [10], a so called “golden age” of antibiotic discovery, followed by an

“innovation gap” that lasted from the discovery of quinolones (nalidixic acid) in 1962 [11] to the next discovery of a new structural class, the oxazolidinone linezolid 40 years later

[12]. The initial rapid pace of discovery ensured that there was always another drug class in the discovery pipeline so that observed resistance and loss of efficacy in one class was not considered a cause for alarm. Since the 1960’s, medicinal chemistry efforts have allowed us to continually modify our existing scaffolds [4] in order to improve the pharmacokinetic or pharmacodynamics properties of our existing molecules, as well as circumvent modes of resistance as they are observed; however, utilizing the same core scaffolds allows for more rapid development of cross-resistance.

Compounding the lack of new developments in antibiotic discovery between the

1960’s and 2000’s (Figure 1-1), is the conservation of molecular targets within bacteria

that are exploited by clinically used antibiotics. The five primary cellular processes outlined in Figure 1-3 are fundamental to microbial life [4], and medicinal chemists and microbiologists have been very successful in discovering bioactive molecules that disable these processes. Antibiotics can inhibit the DNA replication/RNA transcription machinery, protein synthesis, cell wall biosynthesis, nucleic acid precursor (folate) synthesis, or disrupt bacterial cell walls.

Figure 1-3. The molecular targets of antibiotics are grouped into five primary cellular processes: inhibition of peptidoglycan (cell wall) synthesis, inhibition of protein synthesis, inhibition of the precursor synthesis for DNA/RNA synthesis, inhibition of DNA/RNA synthesis machinery, and disruption of bacterial cell walls [4]

Inhibitors of DNA/RNA precursor (folate) synthesis. One of the earliest discovered antibiotics, Prontosil, was discovered by Gehard Domagk in Germany in 1932 [13].

Originally synthesized as a chemical dye, he discovered that it would clear an infection of haemolytic streptococci in the peritoneum of mice [14]. The inability of the drug to kill bacteria in a petri dish was later attributed to the fact that Prontosil is actually a pro-drug and must be processed by the human host to release p-aminobenzenesulphonamide as the active species and an inhibitor of dihydropteroate synthetase, the enzyme responsible for the synthesis of dihydrofolate from p-aminobenzoic acid (PABA) [15]. This became known as the first member of the sulfa-drugs which target the early steps in tetrahydrofolate synthesis. This cellular process is also targeted by trimethoprim, an inhibitor of the later biosynthetic step, dihydrofolate reductase; the co-administration with a sulfa drug

(sulfamethoxazole) is known to be strongly synergistic [16, 17], as inhibition of this pathway starves the bacteria of thymidine. Folate biosynthesis in bacteria is essential and required for nucleic acid biosynthesis, but eukaryotic cells import folate from the diet, leading to selective toxicity for bacteria. Although humans also possess DHFR (the target of the anticancer drug, methotrexate), the bacterial and human proteins are sufficiently different to allow for selective inhibition [18]. Resistance to both drug classes is usually caused by mutations in their respective target proteins that decrease their affinity for the antibiotic agent.

Inhibitors of DNA/RNA synthesis. A critical aspect of all cellular life is the replication of DNA and its transcription into RNA for eventual translation into proteins.

Although the processes involved are conserved from archaebacterial all the way to mammalian cells, the function and structure of some of the primary players is divergent, and thus the bacterial proteins can be selectively targeted without inhibiting mammalian host processes. This selectivity is the hallmark of successful antibiotic development and

supports the general view that antibiotics are safe and generally well-tolerated, which can also exacerbate their over-use [19]. The cellular machinery that acts on DNA strands

(polymerase, histone-like proteins [20], etc.) can cause twisting of the DNA double helix that leads to “supercoiling” which must be resolved to prevent damage to the integrity of the DNA. This action is performed by a family of enzymes known as the topoisomerases, which includes the bacterial enzymes, DNA gyrase (Gram-negative) and topoisomerase IV

(Gram-positive), the target of the aminocoumarins and quinolone class of antibiotics [21-

23]. Quinolones are able to trap the topoisomerase proteins in complex with the DNA strands. At low concentrations, bacterial growth is inhibited and the SOS response is activated, but at higher concentrations, this causes double strand breaks and cellular death.

Resistance arises from mutations in the target and a lack of intracellular accumulation typically mediated by rapid efflux. RNA transcription from the DNA template by RNA polymerase is the target of the antibiotic rifacmycin (rifampicin) [24, 25] which blocks the elongation of the growing RNA transcript. While it is potent and broad-spectrum against bacterial pathogens, it does not bind to the eukaryotic polymerase. Resistance typically occurs as a result of mutations to the β-subunit of polymerase with amino acid changes leading to a loss of recognition.

Inhibition of protein synthesis. The translation of mRNA to protein product is accomplished by the ribosome, a complicated cellular machine composed of a large and a small unit (in bacteria: 50S and 30S, respectively) composed of protein and RNA. The ribosome is responsible for matching the codons on mRNA with the appropriate tRNA and assembling a growing peptide chain. A diverse array of structurally unrelated antibiotics are known to bind to the subunits [26]. The 30S subunit is the target of the aminoglycosides

(streptomycin), spectinomycin, and the tetracyclines. The 50S subunit is the target of the oxazolidinone (linezolid [27]), macrolides (erythromycin), lincosamides (clindamycin) and streptogramin B. Binding of these antibiotics prevents the normal functions of the ribosome including GTP hydrolysis (30S), synthesizing peptide bonds (30S) and passing the growing peptide chain through the exit tunnel (50S). Amino acid substitutions and nucleoside methylations can confer resistance by interfering with molecular recognition of the ribosome binding unit.

Inhibition of peptidoglycan synthesis. The bacterial cell wall provides structural support, osmotic regulation and physical protection from exterior threats. It is composed of pentapeptides (L-Ala-γ-D-Glu-X-D-Ala-D-Ala) covalently linked to sugar repeats

(alternating N-acetylmuramic acid and N-acetylglucosamine) that form a layer known as the peptidoglycan [28]. The wall is extensively remodeled by a family of enzymes known as the transpeptidases and the transglycosylases. The transpepidases (also known as penicillin-binding proteins, PPB) are the targets of the β-lactam class of antibiotics

(penicillins, cephalosporins, etc.), and the glycopeptides (vancomycin and teicoplanin) targets the and sequesters the building blocks (peptidyl-D-Ala-D-Ala) of the growing cell wall of Gram-positive bacteria, preventing the transpeptidases from accessing them and assembling them into the peptidoglycan network. β-lactams are inactivated by a class of enzymes known as the β-lactamases, which catalytically destroy the β-lactam ring and thereby prevent β-lactam binding to PPBs. Vancomycin resistance can arise from a thickening of the cell wall [29] or by a change in the structure of the components of the cell wall, namely changing the peptidyl-D-Ala-D-Ala to peptidyl-D-Ala-X. These

modifications can prevent hydrogen bonds between vancomycin and the substrate or cause steric clashes that result in a loss of affinity of vancomycin for its substrates.

Disruption of inner membrane integrity. The most recently utilized target in antimicrobial discovery is the Gram-positive bacterial inner membrane. The membrane provides protection from the extracellular environment and establishes a chemiosmotic gradient. The integrity of this important biological barrier is disrupted by the lipopeptide antibiotic, daptomycin, produced by Streptomyces roseoporus, and discovered by Eli Lilly in the 1980s[30]. The mechanism of action [31] is believed to be a calcium ion-dependent oligomerization in the Gram-positive cell membrane which leads to potassium ion leakage, membrane depolarization and cell death. Some resistance is observed as a result of mutations changing the lipid composition of the bacterial membrane outer leaflet. There is also tissue-specific resistance for infections in the lung, as it is hypothesized that the host lung surfactant can sequester daptomycin and prevent fusion with the bacterial membrane

[32].

1.1.2 Efflux pump-mediated resistance. As discussed above, each of the classes of antibiotics utilized in the clinic can lose efficacy as a result of bacterial resistance pathways. These are often the result of mutations/modifications to the enzyme targets

(methylation of the 23S rRNA leads to erythromycin resistance, modifying the D-Ala-D-

Ala peptide prevents vancomycin activity), or sequestration/inactivation by the bacteria (β- lactamase degrades penicillin) [33]. Another, general mode of resistance is the endogenous multidrug efflux pumps. They fall into 5 main classes: 1) the MATE (multidrug and toxic compound extrusion) family, 2) the MFS (major facilitator superfamily) pumps, 3) the 8

RND (resistance nodulation, and division) family and 4) the ABC (ATP binding cassette) family (Figure 1-4). The pumps are active transporters, powered by ATP hydrolysis in the case of the ABC family, the proton-motive force powers the RND and MFS families, while the MATE family utilizes a sodium gradient.

Figure 1-4. The four primary classes of MDR pump in bacteria, and some observed substrates. [7]

The RND class is responsible for the efflux of a wide variety of structurally unrelated antibiotics from Gram-negative bacteria. The prototypical member of this family is the tripartite AcrAB-TolC pump in E. coli, and the P. aeruginosa homolog, MexAB-

OprM [6]. They are composed of three components, the periplasmic membrane fusion protein (MFP), AcrB, an outer-membrane factor (OMF), TolC, and the periplasmic adapter protein, AcrA. A model structure is shown in Figure 1-5. AcrB, a functional trimer, binds substrates in the periplasmic space delivers them to TolC, which pumps them out via peristaltic action. The trimers rotate through three conformations: loose, tight, and open

[34]. The loose and tight conformations allow for binding of substrate. The loose conformation allows the substrate access to the low-affinity proximal pocket in the

transmembrane region. The substrate is then passed to the high affinity distal pocket in the tight conformation, and then pushed towards the TolC access channel in the open conformation [35].

Figure 1-5. Molecular model of AcrAB-TolC. AcrB is buried in the inner membrane and colored green. AcrA is colored blue and TolC passes through the outer membrane and is colored red. [36] Reprinted with permission.

Knocking out or inhibiting the RND class of pumps in P. aeruginosa, A. baumannii and E. coli is known to restore susceptibility to a variety of structurally diverse antibiotics. 10

The AcrB binding pocket is notoriously non-specific, binding and extruding a wide variety of substrates (dyes, cephalosporins, fluoroquinolones, etc.) [6]. As such, in recent years, there has been an effort to design pharmacological inhibitors of these pumps, the two predominant being phenyl-arginine-β-naphthylamide (PAβN) [37] and the pyranopyridine

MBX 2319 [38, 39], as well as the proton-motive force decoupler, carbonyl cyanide m- chlorophenylhydrazone (CCCP) [7], although none have made it to the clinic yet.

1.2 Isoprenoid biosynthesis is an underutilized target in antibiotic development. Isoprenoids are the largest class of natural products, with estimates over

55,000 members [40], all constructed via condensation of two branched C5 building blocks, dimethylallyl diphosphate (DMADP) and its isomer, isopentenyl diphosphate

(IDP) [41]. They have central roles in secondary metabolism leading to biosynthesis of important natural products (artemisinin [42], taxol [43]), as well as bacterial cell wall components (Lipid II [44]), electron transport chain constituents (ubiquinone [45]) as well as signal transduction via protein prenylation in eukaryotic cells (apicocomplexan parasites

[46]). The ubiquity of terpenes highlights their importance and supports a strategy to target their biosynthesis in antimicrobial development.

Until the 1990’s, it was believed that all isoprenoids were synthesized from the same biosynthetic pathway, known as the mevalonate (MVA) pathway (Figure 1-6, [41]).

This process begins with iterative condensation of three units of acetyl-CoA into 3- hydroxy-3-methylgutanyl-CoA (HMG-CoA). This is the substrate for HMG-CoA reductase which forms the committed intermediate, mevalonic acid, in the rate-limiting step of the pathway. HMG-CoA reductase is also the target of the statin class of drugs 11

which are effective cholesterol-lowering medications [47]. The MVA pathway produces

IDP which is isomerized to DMADP by isopentenyl-diphosphate isomerase (IPI). These two C5 isomers are known as the hemiterpenes and are the building blocks of the entire terpene class of natural products. They are condensed by downstream enzymes in a “head to tail” synthesis to create growing chains of C5 units. The pyrophosphate of acts as the leaving group, and the downstream enzymes of the terpene biosynthesis machinery catalyze the formation of an allylic carbocation electrophile on DMADP (“tail”) followed by subsequent attack of the nucleophilic olefin of IDP (“head”) to form a new carbon- carbon bond and a growing carbon chain. Geranyl diphosphate synthase catalyzes the formation of the C10 unit (monoterpene) known as geranyl diphosphate. A subsequent equivalent of IDP can be condensed by farnesyl diphosphate synthase to form a C15 sesquiterpene. Farnesyl diphosphate synthase is the molecular target of the nitrogen- containing bisphosphonates [48] which are medications for bone cancers. The cytotoxic effects on osteoclasts of inhibiting farnesyl diphosphate synthase with bisphosphonates again supports the biological importance of terpene synthesis.

Figure 1-6. The mammalian mevalonate (MVA) pathway of isoprenoid biosynthesis. It begins with the condensation of three acetyl-CoA units and leads to the formation of the universal isoprenoid precursors, isopentenyl diphosphate (IDP) and dimethylallyl diphosphate (DMADP), in mammals. The intermediate enzyme, HMG reductase is the target of the statin class of cholesterol-lowering medications [49].

1.2.1 The discovery of the non-mevalonate pathway for isoprenoid biosynthesis. As early as the 1980’s, labelling studies undertaken by Pandian, et al. utilizing [1-14C] and [2-14C] acetate in bacteria hinted at a distinct pathway for isoprenoid biosynthesis. The experiments revealed an incorrect labelling scheme for the downstream polyprenyl side-chain of ubiquinones in bacteria if the MVA pathway was the exclusive source of these intermediates [50]. They originally postulated an alternative “acetolactate pathway” that eventually passed through HMG-CoA and mevalonic acid on the way to

IDP. In the 1990’s, a series of labelling studies further supported that another pathway

13 was utilized by bacteria. Zhou et al. studied the incorporation of [1,2- C2] acetate and [U-

13 C6] glucose into both bacterial fatty acid units and ubiquinone [51]. When cells were treated with labeled acetate, but unlabeled glucose, they observed the label incorporation

in downstream fatty acid products but not the prenyl chain of ubiquinone. However, when the cells were treated with the labeled glucose but unlabeled acetate, they were able to observe the label incorporation into both fatty acid products and the prenyl chain of ubiquinone. This suggested the precursors of the prenyl chain were derived from pyruvate and not acetate. They further concluded that the lack of E. coli growth inhibition by a statin inhibitor of HMG-CoA (mevanolin) supported an alternative to the mevalonate pathway in bacteria. Finally in subsequent papers in 1993 and 1996, Rohmer et al. [45, 52] concluded that the starting materials of the alternate pathway were derived from glyceraldehyde 3- phosphate and pyruvate based on evidence from incorporation studies with 13C-labeled glycerol or pyruvate in E. coli strains lacking enzymes of the triose phosphate metabolic pathway required for inter-conversion of glycerol and pyruvate. From these studies, they concluded that of the C5 skeleton of isopentenyl diphosphate, two carbons were derived from pyruvate and three carbons were derived from glyceraldehyde 3-phosphate. They hypothesized this reaction would be catalyzed by a thiamin diphosphate (ThDP)-dependent enzyme to form 1-deoxyxylulose 5-phosphate (DXP); however, the gene product that catalyzed this reaction would not be discovered until 1997 when the dxs gene encoding

DXP synthase was identified due to its homology to another ThDP-dependent enzyme, transketolase [53-55].

We now know that this “transketolase-like” enzyme, DXP synthase catalyzes the first step in a pathway that is orthogonal to the mammalian, mevalonate-dependent pathway. DXP, the product of this enzyme, is also a precursor for thiamine biosynthesis and pyridoxal biosynthesis. The subsequent formation of 2-C-methyl-D-erythritol 4- phosphate (MEP) by the enzyme IspC is the first committed step of the pathway, and the

name of the pathway. The MEP pathway consists of seven enzymes (Figure 1-7) with no mammalian homologs, and which form the same final products of the mevalonate pathway,

IDP and DMADP. This is exciting from a drug discovery effort as inhibition of the enzymes should have no deleterious effects on a human host. It is interesting from an evolutionary standpoint as nature essentially developed two answers (MEP vs. MVA pathway) to the question of how to generate the essential precursors, IDP and DMADP.

The pathway is essential in a wide variety of important human pathogens, including

Mycobacterium tuberculosis [56, 57] and Plasmodium falciparum [58, 59] making it a promising target for a number of critical human diseases.

Figure 1-7. The non-mammalian MEP pathway generates the precursors, DMADP and

IDP, in many bacteria, plants and apicocomplexan parasites.

1.2.2 Current efforts to target enzymes of the MEP pathway. Fosmidomycin

(formerly, FR-31564, 1.1) is a natural product isolated from Streptomyces lavendulae in 15

1980 and determined to have antimicrobial properties [60]. In 1998, Kuzuyama, et al. discovered that its mechanism of action was inhibition of DXP reductoisomerase (IspC) the first committed step of the MEP pathway, and it showed efficacy against a P. falciparum murine model of infection[61]. In recent years, it progressed to Phase II clinical trials in combination with clindamycin as a treatment for uncomplicated malaria [62], but it had inadequate efficacy. It is currently in a phase IIa clinical trial in combination with piperaquine to treat uncomplicated P. falciparum infections [63].

While inhibitors of other enzymes in the pathway are being designed and considered, no other compound has made it to clinical trials and most suffer from being weak binders or non-specific inhibitors. From a medicinal chemistry standpoint, the active sites of many of the MEP pathway enzymes are difficult to target due to their highly polar nature and their ability to bind polyanionic phosphate species. Charged species tend to make poor drugs due to poor bioavailability and cell permeability. However, some inhibitors of the MEP pathway enzymes are promising leads for drug design (Figure 1-8)

[64]. Inhibitors of DXP synthase will be discussed further in Section 1.2.4. IspD is inhibited by 7-hydroxy-[1,2,4]triazolo[1,5-α]pyrimidine (1.2) [65] which was discovered as a nanomolar, allosteric inhibitor of the Arabidopsis thaliana enzyme, and possesses herbicidal activity. Recently a class of highly halogenated marine natural products, the pseudilins (i.e., 1.3), were discovered to be divalent-metal dependent allosteric inhibitors of the A. thaliana and P. falciparum IspD enzymes [66]. IspE is moderately inhibited by the alkynyl sulfonamide 1.4, synthesized as the product of structure-based inhibitor design

[67]. Kip Bitok, Ph.D. performed extensive work with IspF in our own lab, synthesizing and testing a series of bisphosphonate substrate analogs as potential inhibitors of the

enzyme. Interestingly, these were weak inhibitors at best, and in some cases acted to enhance IspF activity [68, 69]. Geist, et al. designed a family of thiazolopyrimidine inhibitors (i.e., 1.5) of IspF based on the hits from a high-throughput screen, and these show low micromolar inhibition of IspF. Some of the most potent inhibitors of the [4Fe-

4S] cluster-utilizing proteins, IspG and IspH have been designed by Eric Oldfield’s group.

They have had success with alkyne inhibitors designed to interact with the electron donating character of the Fe-S cluster to form a metallocycle, such as propargyl diphosphate (1.6) which is a nanomolar inhibitor of IspG [70] and a slightly weaker inhibitor of IspH [71]. Inserting an additional methylene into the structure of 1.6 generated a more potent inhibitor (1.7) of IspH [72].

Figure 1-8. Inhibitors of the enzymes of the non-mammalian MEP pathway [64].

1.2.3 Regulation in the MEP pathway. Until recently, not much was known about regulation of the MEP pathway. The genes encoding the MEP pathway enzymes do not cluster in most bacterial genomes, and not much is known about transcriptional regulators of the individual components. Brown, et al. discovered that flux through the

Mycobacterium tuberculosis MEP pathway could be increased by over-expression of the genes encoding DXP synthase and IspG [57], suggesting that they play some role as rate- limiting steps in the synthesis of IDP and DMADP. Many studies in plants point to DXP synthase [73] and IspC [74, 75] overexpression correlating with increased production of downstream terpene products.

X-ray crystallography studies have suggested that IspF is regulated via feedback inhibition by downstream isoprenoids. Crystal structures of the IspF trimer suggested that downstream metabolites (IDP/DMADP, geranyl diphosphate or farnesyl diphosphate) may bind in a hydrophobic pocket at the interface of the monomers [76-79]. Work in the Freel

Meyers laboratory has shed light on this hypothesis and revealed an additional putative regulatory mechanism via feed-forward activation of IspF by MEP. These discoveries were made by J. Kipchirchir Bitok [68, 69]. While studying IspF activity, he determined that MEP, the first committed step of the pathway, could bind to IspF and improve its catalytic activity. Additionally, the IspF-MEP complex retained activity for over 24 hours, while IspF alone lost activity within 30 minutes [69]. This was an unprecedented discovery in this pathway and is evidence of a “feed-forward” mechanism at play in the MEP pathway.

Recently, Odom et al. have determined that flux through the MEP pathway in

Plasmodium falciparum is regulated in part by a sugar phosphatase, a homologue of the

haloacid dehydrogenase family they refer to as PfHAD1 [80]. They were exploring fosmidomycin-resistant mutants of P. falciparum and locating gene mutations not directly related to IspC or other members of the MEP pathway. They determined that resistant mutants had a loss-of function mutation in the gene for PfHAD1 resulting in higher concentrations of intermediates in the MEP pathway. Their results support the idea that

PfHAD1 dephosphorylates intermediates of the MEP pathway and can thus lower flux and potentially divert resources into different pathways.

In addition to the proposed regulatory mechanisms for isoprenoid biosynthesis, there appear to be alternative pathways to access these key building blocks, hinting at potential mechanisms of resistance to agents that target this pathway (Figure 1-9) [81].

The lethal phenotype of deleting the genes encoding DXP synthase or IspC can be circumvented by mutations in the aceE and ribB genes. The aceE gene encodes for the

ThDP-dependent pyruvate dehydrogenase (PDH) protein, and the mutations E636Q/G,

Q408R, or L633R allow for the PDH protein to synthesize DX from pyruvate and D- glyceraldehyde [82, 83]. DX can be phosphorylated to DXP by the enzyme D-xylulose kinase (DXK). The product of the ribB gene is 3,4-dihydroxy-2-butanone-4-phosphate synthase (DHBPS) [84]. It is hypothesized that the mutations observed (G108S and

D113G) allow the enzyme to produce DXP by accepting the alternative substrates D- ribulose 5-phosphate (RP) or D-xylulose 5-phosphate (XP).

In 2012, Erb et al. reported the discovery of an alternative pathway to DXP in a photosynthetic anaerobe, Rhodospirillum rubum [85]. In this organism, polyamine biosynthesis is linked to the MEP pathway by an unusual RubisCO-like protein (RLP) that converts 5-methylthio-d-ribulose-1-phosphate (a product of the degradation of 5-

methylthioadenosine) to 1-methylthio-d-xylulose 5-phosphate (MXTP). This intermediate is a substrate for MTXP methylsulfurylase (MMS) which generates DXP. While genetic analysis revealed the existence of these genes in additional bacterial species (Methylocella,

Rhodomicrobium, etc.), they have not yet discovered its existence in pathogenic bacteria, and it is not clear to what extent this pathway can operate to rescue DXP synthase inhibition. Development of resistance to antimicrobials is inevitable, and these represent the first hints to possible resistance mechanisms against DXP synthase inhibitors, and emphasize the recognized need to use antimicrobials in combination.

Figure 1-9. Alternatives to DXP synthase for the intracellular formation of DXP. A)

Polyamine biosynthesis is linked to the MEP pathway via the RubisCO-like protein (RLP) catalyzed formation of MXTP [85]. B) DHBPS catalyzes the formation of DXP from the isomers, RP and XP [82]. C) Mutations to PDH allow for the synthesis of DX from pyruvate and D-glyceraldehyde, which can be phosphorylated by DXK [86].

1.2.4 1-Deoxy-D-xylulose 5-phosphate (DXP) synthase is a favorable target for design of new antibiotics. As mentioned above, the MEP pathway begins with the condensation of pyruvate and D-GAP to generate DXP and CO2 in a ThDP-dependent process catalyzed by DXP synthase (Figure 1-7) [53, 54]. It is important to mention that this is not a committed step for isoprenoid biosynthesis, as DXP synthase represents a branch point in bacterial biosynthesis. The sugar DXP is also a precursor for thiamin diphosphate (ThDP, vitamin B1) and pyridoxal phosphate (PLP, vitamin B6). It is especially interesting to note that DXP synthase catalyzes the first step in the formation of its own cofactor, ThDP. In addition to this central location in bacterial metabolism, other unique characteristics of DXP synthase make it an attractive target, which we believe can be selectively targeted. Previous attempts at designing inhibitors of DXP synthase are summarized in Figure 1-10.

Figure 1-10. Low micromolar inhibitors of DXP synthase described to date.

Mao et al. attempted to design selective inhibitors of DXP synthase from the thiamin-based transketolase inhibitor, 3-(4-chloro-phenyl)-5-benzyl-4H-pyrazolo[1,5- a]pyrimidin-7-one [87]. Although they designed a DXP synthase inhibitor with an IC50 of

7.6 μM against M. tuberculosis cultures (IC50 = 10.9 μM against purified enzyme, in vitro)

(1.8), they also observed considerable toxicity against mammalian cells, presumably due to off-target inhibition of other ThDP-dependent enzymes. This study highlights the difficulty in designing selective inhibitors of DXP synthase by targeting the conserved

ThDP cofactor binding site [88].

The herbicide, ketoclomazone (1.9) has been shown to act via inhibition of DXP synthase [89] and has some antibacterial activity against Haemophilus influenza. A later study showed inhibitory activity by the product formed by hydrolysis of ketoclomazone, an N-(2-chlorobenzyl)-substituted hydroxymate (1.10) [90]. A high-throughput screen looking for new herbicidal agents discovered the hydrazine, 1.11, as another low micromolar inhibitor of DXP synthase [91]. Additionally, the pyruvate analogs, β- fluoropyruvate and methylacetylphosphonate were observed to inhibit DXP synthase activity [92, 93]. Although both are nanomolar inhibitors of DXP synthase, they are also inhibitors of other ThDP-dependent pyruvate-decarboxylating enzymes. This thesis describes our efforts to generate selective inhibitors of DXP synthase based on the following unique attributes.

DXP synthase is structurally unique. Xiang, et al. successfully solved the crystal structure of DXP synthase from Deinococcus radiodurans with the ThDP cofactor present at a resolution of 2.9 Å [88]. They attempted to solve the structure of the E. coli enzyme but discovered a contaminating fungal protease had cleaved the protein before crystallization, leading to the deletion of key loops in the structure. They were able to construct a homology model for the E. coli enzyme based on their D. radiodurans structure.

They compared the structure of DXP synthase to the weakly homologous (~20% identity) 23

ThDP-dependent enzymes, transketolase (TK) [94] and PDH [95] (Figure 1-11). They noted that although all 3 are homodimers, the organization of the monomers is unique in

DXP synthase, related to the flexible loop region (AA residues 305-324) in DXP synthase being considerably shorter than in the other two enzymes. The active site falls at the interface of Domain I and Domain II within one monomer, as opposed to the monomer interface between Domain I of one monomer and Domain II of another monomer, as is the case in the other two enzymes. This finding suggests that the arrangement of the DXP synthase active site is unique when compared to the other two enzymes and may be selectively targeted.

Figure 1-11. The crystal structures of the related ThDP- dependent enzymes, A) DXP synthase [88], B) yeast transketolase [94], and C) E. coli pyruvate dehydrogenase (PDH)

[95]. Reprinted with permission.

The active site of DXP synthase is comparatively large. By utilizing the crystal structures of DXP synthase (2O1X), TK (3MOS) and PDH (3EXE) published in the PDB

[88, 94, 95], our collaborators Jürgen Bosch and his student Lauren Boucher, were able to 24

model the active site of each enzyme and calculate the relative volumes using Pocket-

Finder [96] (Figure 1-12). They determined that the DXP synthase active site volume

(1295 Å3) is at least twice as large as either TK (539 Å3) or PDH (597 Å3). This suggests that DXP synthase may be able to accommodate larger molecules which would not fit into the active sites of PDH or TK, allowing for selective inhibition.

Figure 1-12. The calculated active site volumes of DXP synthase (teal) and two related mammalian ThDP-dependent enzymes [transketolase (purple), PDH (red)] is compared.

The cofactor is shown in the “V-conformation” as a stick model with the diphosphate moiety in complex with a divalent cation (purple sphere). [96]

DXP synthase is promiscuous toward alternative acceptor substrates. Work pioneered by Leighanne Brammer Basta in the Freel Meyers lab showed that DXP synthase can utilize linear, aliphatic aldehydes in the absence of its natural acceptor substrate, D-

GAP [97]. Utilizing pyruvate as a donor substrate, DXP synthase catalyzes the formation of the corresponding α-hydroxyketones (Figure 1-13). She also determined that the

catalytic efficiency of DXP synthase acting on the des-phosphoryl analog of D-GAP, D- glyceraldehyde (D-GA) to generate DX, decreased by approximately 400-fold. This observation supports that the phosphoryl group of the natural acceptor substrate is an important binding and/or recognition element for turn-over, but it is interesting to note that the linear aldehydes acetaldehyde through butanal had catalytic efficiencies within 10-fold of D-GA, suggesting that once the phosphoryl recognition motif is lost, the other aldehydes are turned over with similar efficiencies.

Figure 1-13. Alternative aldehyde substrates for DXP synthase and isolated products.

Building on this discovery, Francine Morris determined that DXP synthase could also process aromatic aldehydes as alternative acceptor substrates, although the turnover efficiency was poor. She tested the hypothesis that this low turnover efficiency was a consequence of the reduced electrophilicity of the aromatic aldehyde, through exploration of more reactive aromatic nitroso compounds as an alternative substrates and discovered

that DXP synthase could utilize these molecules as substrates, sometimes with KM values comparable to the natural substrate, D-GAP (Figure 1-14) [96]. Interestingly, these substrates are not accommodated as well by PDH, suggesting they could be selective agents for the DXP synthase active site. Unfortunately, further studies by Francine confirmed that aromatic nitroso analogs were poor inhibitors of the natural DXP synthase reaction.

Table 1-1. Aromatic nitroso compounds are better substrates for DXP synthase than PDH.

They also make poor inhibitors of the natural DXP synthase reaction. The aromatic nitroso compounds have similar KM values to D-GAP for DXP synthase but significantly higher

KM values for PDH. However, their IC50 values for DXP synthase are also substantially higher than their KM values [96].

DXP synthase is a mechanistically unique ThDP-dependent enzyme. Evidence suggests that most ThDP-dependent enzymes follow a classical ping-pong bi-bi catalytic 27

mechanism. In this mechanism, the free enzyme can only recognize the donor substrate and must undergo a catalytic event to release the first product before it can recognize the second substrate and release the final product [98] (Figure 1-15 A). However, Eubanks and Poulter provided compelling evidence that DXP synthase is mechanistically unique.

They proposed, on the basis of kinetic analysis and CO2 trapping experiments that DXP synthase follows a strictly ordered mechanism which passes through a ternary complex with both substrates present before it is able to catalyze the release of the first product [99].

This suggested that during the mechanism of DXP synthase, it forms a unique intermediate enzyme form which is able to accommodate both of its substrates in the active site at the same time (Figure 1-15 B), a property which, to date, has not been observed for any other members of this class of enzyme.

Figure 1-14. ThDP-dependent catalysis in DXP synthase. A) A hypothetical model of ping-pong catalysis, typical to most ThDP-dependent enzymes, and B) the ordered

mechanism proposed by Eubanks & Poulter [99]. E-LThDP-GAP = ternary complex formed with the lactyl-thiamine diphosphate and D-GAP.

Poutler’s model proposing the requirement for ternary complex formation was expanded by studies in the Freel Meyers lab in collaboration with the Jordan lab to support a model of random-sequential, preferred ordered catalysis for DXP synthase catalysis

(Figure 1-16) [92, 100, 101]. Either substrate can apparently bind reversibly to free enzyme, although pyruvate is preferred as the first substrate. The rate of decarboxylation of the lactyl-thiamine diphosphate intermediate in the absence of D-GAP is 600-fold slower

(k = 0.07 ± 0.006 s-1) than in the ternary complex when the acceptor substrate is present (k

= 42.0 ± 1.0 s-1). Additionally, studies initiated by Leighanne Brammer Basta [97] and continued by Alicia DeColli support an alternative role for DXP synthase as an acetolactate synthase in which it can utilize pyruvate as both a donor and acceptor substrate.

Figure 1-15. The random sequential, preferred order mechanism of DXP synthase. This mechanistic model is supported by studies in the Freel Meyers lab. The free enzyme (E) can bind either substrate, although pyruvate is preferred, and then binds a second substrate to form the ternary complex (E-LThDP-GAP) which is triggers decarboxylation and formation of DXP [92, 100, 101].

1.3 Reactive substrate mimics as inhibitors of DXP synthase. One route to inactivation of DXP synthase involves the use of a mimic of the donor substrate pyruvate which cannot undergo decarboxylation. The enzyme-bound cofactor, ThDP, will react with the mimic in the same way that it would react with its natural substrate, by addition to a reactive carbonyl group, but the tetrahedral covalent adduct formed would not be a suitable substrate for the next step in the enzymatic mechanism, decarboxylation.

Pioneering work in this field was performed by Westerick and Wolfenden [102] who recognized the role that electrophilic species could play in inhibiting proteolytic enzymes.

Specifically, they investigated the enzymatic mechanism of a cysteine protease, papain, with aldehyde mimics of natural peptide substrates. The aldehydes proved to be potent competitive inhibitors of the natural substrates, with Ki in the nanomolar range.

Contemporaneously, Thompson performed studies with similar peptide aldehydes and elastase [103]. The peptide aldehydes bound with roughly 5,000-fold greater affinity than the natural substrates and were again potent, competitive inhibitors of enzyme activity.

Similarly to the nucleophilic side chains of these proteases, the ylid of the ThDP cofactor in the ThDP-dependent enzyme class is a potential target for inhibition by reactive, electrophilic donor substrate mimics. This was first appreciated by Kluger who evaluated methyl acetylphosphonate (MAP) as a competitive pyruvate mimic of the ThDP-dependent enzyme pyruvate dehydrogenase (PDH) [104, 105]. Kluger hypothesized that a covalent intermediate in PDH catalysis could be trapped by use of a mimic unable to undergo decarboxylation. X-ray crystallography, NMR and CD studies confirm that the electrophilic acetyl group forms a covalent adduct with the cofactor, while the stable

phosphoryl group is incapable of decarboxylation to activate substrate, and the single charge on the phosphonate is hypothesized to mimic the charge on the carboxylic acid of pyruvate.

It was observed that this single negative charge is critical for inhibition, as the doubly charged acetylphosphonate has negligible inhibitory activity against the protein, and the neutral pyruvate analog, pyruvamide is also inactive. An important point to consider is that while the carboxylic acid possesses trigonal planar geometry, the phosphonate adopts a tetrahedral geometry suggesting there may be inherent differences in how the charge of the phosphonate and the carboxylate are positioned within the active site. Regardless, methyl acetylphosphonate was confirmed as a potent inhibitor of the reaction, trapping the enzyme in a phosphonolactyl-thiamine diphosphate (PLThDP) conformation that mimics a “pre-decarboxylation” form of PDH [106].

Work in our own lab with methylacetylphosphonate showed it is also a pyruvate competitive inhibitor of DXP synthase [92]. We sought to incorporate this into a selective inhibitor design along with an acceptor substrate mimic to generate “unnatural bisubstrate inhibitors”. We aimed to take advantage of the large active site of DXP synthase, described above, in order to generate a class of selective inhibitors of DXP synthase, described in detail in Chapter 2. With selective inhibitors in hand, we sought to explore the antimicrobial properties of these inhibitors, and these studies are outlined in Chapter 3.

Although the progenitor of this class, methylacetylphosphonate, does not possess antimicrobial properties, some of our unique compounds do, which has allowed us to explore the antimicrobial activities in this new class in more detail, toward the development of a novel antimicrobial class.

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Chapter 2. Electrophilic phosphonates as selective inhibitors of bacterial DXP synthase

Introduction

The 2-C-methylerythritol 4-phosphate (MEP) pathway provides an attractive target for anti-infective drug development as it is a key source of the essential isoprenoid precursors, dimethylallyl diphosphate (DMADP) and isopentenyl diphosphate (IDP)

(Figure 2-1) in a variety of human pathogens including important bacterial targets such as

Mycobacterium tuberculosis [1] and protozoan parasites such as Plasmodium falciparum

[2]. However, there has been limited success with discovery and development of clinically- used inhibitors of this pathway. Fosmidomycin, a natural product isolated in 1980 and determined to have antimicrobial properties [3, 4] via inhibition of IspC [5] has recently been submitted to a phase IIa clinical trial in combination with the antimalarial drug, piperaquine, as a treatment for adults and older children infected with P. falciparum [6].

This is the only inhibitor of the MEP pathway that has progressed to clinical trials to date.

While IspC produces MEP [7, 8], the first committed intermediate in the pathway, its substrate, 1-Deoxy-D-xylulose 5-phosphate (DXP) is produced via the condensation of pyruvate and D-glyceraldehyde 3-phosphate (GAP) by the thiamin diphosphate (ThDP) - dependent enzyme, DXP synthase [9]. DXP synthase makes an especially attractive target for antimicrobial drug efforts due not only to its central, regulatory role in the MEP pathway [10, 11], but also in the biosynthesis of vitamin B1 and vitamin B6 [9, 12, 13].

Theoretically, small molecule inhibitors of DXP synthase would shut down three critical cellular biosynthetic pathways. However, outside of our research group, little work has

been done towards this goal. Conceivably, this is due to the perceived difficulty of selectively inhibiting a ThDP-dependent enzyme in pathogens

Figure 2-1. The MEP pathway is composed of seven biosynthetic enzymes to generate the isoprenoid precursors, DMADP and IDP.

However, as previously stated in Chapter 1, DXP synthase is distinctive among

ThDP-dependent enzymes in that it follows a random sequential, preferred order mechanism that proceeds via a ternary complex [14-16]. Since most other ThDP- dependent enzymes follow a ping-pong bi-bi mechanism, the ternary complex is a unique enzyme intermediate in a ThDP-dependent mechanism and thus provides an opportunity for selective targeting (Figure 2-2).

Figure 2-2. The DXP synthase catalytic mechanism requires both substrates to be present in order for decarboxylation and product formation to occur. This unique ternary complex is shown as the E-LThDP-GAP “pre-decarboxylation” complex.

Leighanne Brammer Basta showed that DXP synthase can tolerate unbranched, long-chain aliphatic aldehydes and successfully catalyzes their conversion into the corresponding α-hydroxyketones in the presence of the donor substrate, pyruvate (Figure

1-13) [17]. Additionally, Francine Morris’s work with aromatic aldehydes and nitroso compounds (summarized in Table 1-1) showed that DXP synthase could turn over these substrates, sometimes with a Michaelis constant on the order of the acceptor substrate, D-

GAP, suggesting they have reasonably high affinity for the enzyme and could potentially compete with D-GAP in the active site. The aromatic compounds also display significantly higher KM values for PDH, suggesting that they do not fit into the PDH active site and could possibly be starting points for selective inhibitors of DXP synthase. Our collaborators Jürgen Bosch, Ph.D. and Lauren Boucher (Johns Hopkins School of Public

Health) correlated this observation to a calculation of the DXP synthase active site volume, which predicts the DXP synthase active site to be approximately twice as large as the

related mammalian ThDP-dependent enzymes, PDH and TK (Figure 1-12) [18].

However, the IC50 values that Francine recorded for these compounds are high, suggesting that they do not significantly block the progress of the natural reaction. Their low KM values on DXP synthase and high KM values on PDH still suggested that they could be incorporated into an inhibitor scaffold that would have affinity and could impart selectivity for DXP synthase. From this evidence, we postulated that selective inhibition of DXP synthase could be achieved by designing inhibitors possessing sterically demanding hydrophobic structures that the DXP synthase active site would tolerate, but which closely related, yet mechanistically distinct, mammalian ThDP-dependent enzymes would not tolerate.

During studies to elucidate the mechanism of DXP synthase, the Freel Meyers lab established that methylacetylphosphonate (MAP) is a potent inhibitor of the enzyme

(Figure 2-3A) [15]. Further, it was confirmed by CD spectroscopy in collaboration with

Frank Jordan, Ph.D. (Rutgers University) that this inhibition was due, as presumed, by formation of a stable PLThDP intermediate (unpublished results). Our findings were in line with previous work, by Kluger and coworkers describing MAP as a pyruvate analog and a potent inhibitor of pyruvate dehydrogenase complex [19]. In our hands, MAP is a

PDH DXP synthase reasonably selective inhibitor of DXP synthase versus PDH (Ki /Ki = 30) [20], and given its role as a pyruvate mimic suggests a starting point when considering the design of selective inhibitors of DXP synthase. We envisaged a series of selective inhibitors that would be potent and selective from the combination of the reactive pyruvate analog, acetylphosphonate, and sterically demanding aliphatic chains that our previous work had determined the DXP synthase active site could selectively accommodate (Figure 2-3B)

Figure 2-3. Inhibition of DXP synthase by MAP and selective inhibitor design. A)

Inhibition of DXP synthase by MAP showing the ternary complex formed in the presence of D-GAP, and B) proposed unnatural bisubstrate inhibitor design linking the pyruvate analog, acetylphosphonate, to a hydrophobic element inspired by the aliphatic aldehydes tolerated as alternative acceptor substrates by DXP synthase.

This chapter will describe the design, synthesis and evaluation of selective acetylphosphonate inhibitors of DXP synthase. We envisaged two series of molecules which could test our hypothesis concerning selective inhibition of DXP synthase by alkylacylphosphonates (Figure 2-4). Considering the pyruvate mimic MAP, there are two points on the molecule where alterations could be made to create a more sterically demanding inhibitor. Modification of either the phosphonoester (Figure 2-4, compounds

2.1 – 2.5) or acyl (Figure 2-4, compounds 2.6 – 2.8) portion of the molecule could 45

potentially mimic a ternary complex between the donor substrate pyruvate and an unnatural, aldehyde accepter substrate. We hypothesized Series I would be more potent, based on evidence from earlier substrate specificity studies that showed DXP synthase does not readily accept α-ketoacids with modifications at the acyl position [17]. Further, as

DXP synthase does not efficiently turn over α-branched alkyl aldehydes [17], we suspected that compound 2.9 would not be a potent inhibitor. Lastly, we synthesized a preliminary series of aromatic acetylphosphonates based on the evidence that aromatic nitroso compounds are poor substrates for PDH but high affinity substrates for DXP synthase [18], and therefore may impart potency and selectivity of inhibition against DXP synthase.

Figure 2-4. Acylphosphonates synthesized and evaluated as selective inhibitors of DXP synthase. Two possible orientations (2.1 – 2.5, 2.6 – 2.8) of the acylphosphonates were evaluated as mimics of the ternary complex formed by the enzyme pyruvate and nonpolar acceptor substrates. Additionally, we designed the acylphosphonate, 2.9, to determine if

DXP synthase would tolerate branched groups. The aromatic rings of the nitroso and aldehyde alternative substrates of DXP synthase are incorporated into compounds 2.10 –

2.17.

Results

2.1 Synthesis of unbranched acetylphosphonates (2.1 – 2.2). The synthetic scheme for this series of compounds is shown (Figure 2-5A). Procedures for the synthesis of compounds in this study were modified from a published route by Fang, et al. [21]

Methylacetylphosphonate (MAP) had been synthesized previously [15] as a sodium salt and was evaluated in this form. Commercially available trialkyl phosphites were subjected to an equimolar amount of acetyl chloride in a neat reaction under an atmosphere of argon.

The Michaelis-Arbuzov rearrangement yielded the corresponding dialkyl acetylphosphonate as a pale yellow oil in each case. Complete consumption of the starting phosphite occurred within 30 minutes and was confirmed by 31P NMR spectroscopy. The reaction product, (2.18 – 2.19), was isolated by vacuum distillation and carried forward without further purification. The acetylphosphonate was next subjected to mono- dealkylation by lithium bromide. After dissolving the acetylphosphonate in anhydrous acetonitrile, 1.5 equiv. of lithium bromide was added to the theoretical yield of the previous reaction. The reaction mixture was heated to 70 oC overnight, and the product precipitated from solution. Filtration and subsequent rinses with excess acetonitrile and ether yielded pure products, 2.1 – 2.2, as was confirmed by 31P and 1HNMR spectroscopy.

2.2 Synthesis of unbranched acetylphosphonates 2.3 – 2.5. The scheme follows that shown in Figure 2-5B and is closely related to the synthesis of 2.1 – 2.2, but with the variation that phosphites were dissolved in anhydrous CH2Cl2 (~0.1 M) and added to a flask charged with 5 equivalents of neat acetyl chloride. The order of addition ensured that the phosphite was exposed to a large excess of acetyl chloride, favoring the desired reaction to produce acetylphosphonate and avoiding an unintended side reaction in which the

phosphite reacts with in situ generated acetylphosphonate to produce bisphosphonates in the presence of catalytic acid, such as nascent water [22]. After the addition, a needle connected to an argon line was inserted into the reaction mixture to bubble argon through the mixture, in order to help remove byproduct HCl which was generated as a side-product from the reaction between acetyl chloride and water, and which could catalyze the formation of bisphosphonate.

Reaction progress was determined by 31P NMR; when the starting phosphite had been consumed to generate dialkylacetylphosphonate, the mixture was concentrated in vacuo. The pale oil containing the dialkylacetylphosphonate was then dissolved in anhydrous acetonitrile and subjected to three equivalents of lithium bromide and elevated temperature (65 – 70 oC) to remove a single alkyl group and yield the lithium salt of the acetylphosphonate as a product that could be isolated by filtration.

2.3 Synthesis of unbranched acylphosphonates 2.6 – 2.8. The synthetic scheme for this series of compounds is shown (Figure 2-5C). Compound 2.7 had been synthesized previously and spectral data matched previously published characterization data [21].

Briefly, trimethyl phosphite was subjected to a slight excess of commercially available acyl chlorides in a neat reaction under an atmosphere of argon. The Michaelis-Arbuzov rearrangement yielded the corresponding dimethyl acylphosphonate as a pale yellow oil.

Complete consumption of the starting phosphite occurred within 30 minutes and was confirmed by 31P NMR spectroscopy. The reaction product was carried forward without further purification. The dimethyl acylphosphonate was next subjected to mono- dealkylation by lithium bromide. After dissolving dimethyl acylphosphonate in anhydrous acetonitrile, 1.5 equivalents of lithium bromide was added to the theoretical yield of the

previous reaction. The reaction mixture was heated to 70 oC overnight, and the product precipitated from solution. Filtration and subsequent rinses with excess acetonitrile and ether yielded pure products 2.6 – 2.8, as was confirmed by 31P- and 1H-NMR spectroscopy.

Figure 2-5. Synthetic scheme for alkyl acylphosphonates (2.1 – 2.8). A) Synthetic route to alkylacetylphosphonates of Series I (2.1 – 2.2) [20]. B) Modified scheme to yield acetylphosphonates 2.3 – 2.5. C) Synthetic route to methylacylphosphonates of Series II

(2.6 – 2.8) [20]

2.4 Synthesis of phenyl acetylphosphonates. Initially, we proposed to synthesize and evaluate phenyl acetylphosphonate as a possible inhibitor of DXP synthase. Towards this end, the mixed phosphite, 2.20, was subjected to the Arbuzov reaction with acetyl chloride (Figure 2-6). However, the extended conjugation between the aromatic ring and the phosphonate caused in situ generated acetylphosphonate to be apparently more electrophilic than acetyl chloride [23], and as a result the primary product observed was assigned as a bisphosphonate (Figure 2-6, B) resulting from the acetylphosphonate

(Figure 2-6, A) reacting with a second equivalent of the phosphite 2.20. Product ratios were determined by integrating 31P NMR signals of crude reaction products (δ = -28 ppm

(A), -6 ppm (B, tentative assignment for bisphosphonate side product)). Even dissolving the starting phosphite in CH2Cl2, and adding it to neat acetyl chloride did not improve the yields above 1:4 (majority undesired bisphosphonate).

Figure 2-6. The inherent reactivity of the phenylacetylphosphonates favors bisphosphonate formation. Ratios were determined by integration of 31P NMR signals of crude reaction products.

We next tested if increasing the electron density of the ring by starting with the mixed phosphite 2.21 would decrease the electrophilicity of the nascent acetylphosphonate and improve yield of the desired product (Figure 2-6, A). As determined by 31P NMR, the product ratio did improve to 2:1 (majority desired acetylphosphonate). However this is a low yield for an intermediate step, and we anticipated that the inherent reactivity of this class of compounds would limit stability in solution. We thus pursued the more stable benzyl acetylphosphonate scaffold.

2.5 Synthesis of aromatic acetylphosphonates 2.10 – 2.17. In order to limit the reactivity of the phenyl acetylphosphonate scaffold, we sought to break up the conjugation between the aromatic ring and phosphonyl group by inserting a methylene spacer. Even with this modification, the electron-withdrawing groups on the aromatic rings can facilitate the formation of bisphosphonates through inductive effects [24]; there was a limit to which scaffolds we could access, as electron-rich aromatic rings are capable of releasing acetylphosphonate via elimination through the ring [25]. The following synthesis was based on a synthesis published by Saady et al [26] (Figure 2-7). Tribenzyl phosphite was prepared by Dr. J. Kipchirchir Bitok. Compounds 2.10 – 2.17 were generated starting from the reaction of the appropriate benzyl alcohol (2.10 – 2.15) or phenethyl alcohol (2.16 –

2.17) with phosphorus trichloride and Hünig’s base dissolved in a 1:1 mixture of diethyl ether and tetrahydrofuran (THF), starting at -40 oC and warming slowly to room temperature. Reaction progress was followed by 31P NMR and deemed complete when

PCl3 (δ ~ 180 ppm) was consumed, and the major peak was the desired phosphite (δ ~ 115-

120 ppm). The reaction mixture was then washed with cold, alkaline aqueous solution

(KOH or NaHCO3) to remove the acidic byproduct, phosphorus acid (tautomerizes to H- phosphonate). The organic layer and aqueous layers were separated, and the organic layer concentrated in vacuo to yield the phosphite of interest.

Figure 2-7. General synthetic route to aromatic acetylphosphonates 2.10 – 2.17.

The phosphite was dissolved in anhydrous CH2Cl2 to a concentration of ~ 0.1 M and added to an excess of neat acetyl chloride. After the addition, a needle connected to an argon line was immersed in the reaction mixture and argon bubbled through to remove

HCl from the reaction mixture that was produced and improve yields by preventing side reactions catalyzed by the acid [22]. Again, reaction progress was monitored using 31P

NMR, and the reaction was considered complete after consumption of the starting phosphite and conversion to the desired acetyl phosphonate (δ ~ -29 ppm). Volatiles were quickly removed, the product was dissolved in anhydrous acetonitrile to a concentration of

~0.6 M, and 1.5 equivalents of lithium bromide was added before the reaction was heated to 45-50 oC with stirring overnight. The product precipitated from solution as a lithium

salt and was separated from the reaction mixture by filtration. The filter cake was washed with cold ether, acetonitrile and CH2Cl2. In some cases, further purification via reverse- phase HPLC was required. Based on the 31P NMR spectra and work by Eli Breuer, et al.,

[27] we assigned common contaminants to be the tautomer of dialkyl phosphite, H- phosphonate (δ ~ -16 – 19 ppm), and the bisphosphonate (δ ~ -3 – 8 ppm). We were also able to observe the hydrate of the product in equilibrium with the acetylphosphonate form

(δ ~ -9 – 10 ppm).

2.6 Evaluation of acylphosphonates 2.1 – 2.8 as inhibitors of E. coli DXP synthase. Dr. Jessica Mott Smith evaluated alkylacylphosphonates 2.1 – 2.2 and 2.6 – 2.8 as inhibitors of DXP synthase in a validated in vitro coupled assay using the DXP reductoisomerase, IspC [15, 28]. DXP synthase turns over pyruvate and D-GAP to yield

DXP which in turn is turned over by IspC along with a stoichiometric equivalent of nicotinamide adenine dinucleotide phosphate (NADPH) to yield 2C-methylerythritol-4- phosphate (MEP). The progress of the reaction was monitored by measuring the consumption of NADPH (340 nm) as a function of time. The concentration of NADPH can be determined using its extinction coefficient at that wavelength, 6220 M-1 cm-1, and the initial reaction velocity can be calculated by determining the change in NADPH concentration over time.

MAP Previously, MAP (Ki = 0.96 ± 0.3 μM, [15]) was shown to be a competitive

pyruvate pyruvate analog (KM = 40.8 ±4.6 μM) and potent inhibitor of DXP synthase [15].

Double-reciprocal plots were used to determine which mode of inhibition best fit the data and solve for Ki for compounds 2.1 – 2.2 and 2.6 (GraFit by Erithracus Software). The Ki values for compounds 2.3 – 2.5 were determined by a non-linear regression model 54

(Graphpad Prism). There was not significant inhibition by compounds 2.7 – 2.8, so Ki values could not be calculated. Compounds 2.2 – 2.5 had Ki values within 7-fold of MAP

EAP BAP PentAP HexAP (Ki = 6.7 ± 0.03 μM, Ki = 5.6 ± 0.08 μM, Ki = 9.9 ± 0.4 μM, Ki = 8.9 ± 0.5

OctAP μM, Ki = 6.0 ± 0.2 μM) (Table 2-1). Additionally, all were shown to fit the profile of a competitive mode of inhibition with pyruvate (Figure 2-8). Compound 2.6 displayed over 250-fold reduction in inhibitory activity compared to MAP.

DXPS Compound Ki (μM) MAP 0.96 ± 0.3 2.1 (EAP) 6.7 ± 0.03 2.2 (BAP) 5.6 ± 0.8 2.3 (PentAP) 9.9 ± 0.4 2.4 (HexAP) 8.9 ± 0.5 2.5 (OctAP) 6.0 ± 0.2 2.6 (MPP) 258.4 ± 67.8 2.7 > 1 mM 2.8 > 1 mM

Table 2-1. Inhibition of DXP synthase by the acylphosphonates 2.1 – 2.8. [20]

Figure 2-8. Competitive inhibition by compounds 2.1 – 2.6 for DXP synthase.

Lineweaver-Burke analysis of A) EAP (2.1); B) BAP (2.2); C) MPP (2.6,); D) PentAP

(2.3); E) HexAP (2.4) and F) OctAP (2.5); showing competitive inhibition with respect to pyruvate. The concentration of pyruvate was varied with increasing acetylphosphonate: 0

μM (○), 10 μM (1-5) or 250 μM (6) (●), 25 μM (2.1 – 2.5) or 500 μM (2.6) (□) and 50 μM

(2.1 – 2.5) or 750 μM (2.6) (). [20]

2.7 Evaluation of acylphosphonates 2.1 – 2.8 as inhibitors of the E1 subunit of porcine heart pyruvate dehydrogenase complex (PDH). The goal of this work was to demonstrate selective inhibition of DXP synthase over another member of the ThDP- dependent class of enzymes, PDH. Pyruvate dehydrogenase E1 subunit (PDH) uses

pyruvate pyruvate as its natural substrate (KM = 54.0 ±5.3 μM), and the pyruvate analogs MAP and the related phosphonates are known inhibitors of this enzyme [29, 30]. However, the kinetic mechanism of PDH is distinct from DXP synthase [14, 15] and small hydrogen and 56

methyl phosphonates show a marked increase in inhibitory activity against PDH compared to MAP [30].

We hypothesized that EAP (2.1), BAP (2.2) and compounds 2.3 – 2.5 could exhibit high affinity for DXP synthase, because it is known to accommodate both substrates in its active site prior to decarboxylation of pyruvate; however, these sterically demanding analogs should be less potent inhibitors of PDH. Indeed, we determined that all five of these acetylphosphonates are less potent inhibitors of porcine PDH compared to DXP synthase (work performed by Jessica Mott Smith), with Ki values against PDH of 29.9 ±

12.6 μM (MAP), 46.6 ± 1.9 μM (2.1), 335.4 ± 7.9 μM (2.2), 250 ± 31 μM (2.3), 117 ± 9

μM (2.4), and 154 ± 14 μM (2.5) (Table 2-2, Figure 2-9). Even the smallest alkylacetylphosphonate, MAP, exhibits selective inhibition towards DXP synthase, expressed as a ratio of the Ki values for inhibition of PDH to the inhibition of DXP synthase. Selectivity of inhibition increases with increasing chain length from MAP

PDH DXPS PDH DXPS (Ki /Ki = 31) to 2.2 (Ki /Ki = 60) (Table 2-2). Interestingly, as the alkyl chain length increases beyond the C4 of 2.2, the degree of selectivity decreases, as shown by 2.3

PDH DXPS PDH DXPS PDH DXPS (Ki /Ki = 25), 2.4 (Ki /Ki = 13) and 2.5 (Ki /Ki = 26). Compounds 2.6

– 2.8 bearing modified acyl groups are not inhibitors of porcine PDH up to 1 mM (Table

2-2).

Figure 2-9. Competitive inhibition of porcine PDH E1 subunit by compounds 2.1 – 2.5.

Pyruvate concentration was varied from 12.5 - 250 μM and acetylphosphonate concentration was varied: 0 μM (○), 250 μM (2.1, 2.3) or 200 μM (2.2, 2.4, 2.5) (●), 500

μM (□) and 750 μM (2.1) or 1000 μM (2.2 – 2.4) (). Experiments were performed in triplicate, and data were subjected to non-linear regression analysis for Ki determinations for compounds 2.3 – 2.5 (GraphPad Prism) and GraFit from Erithacus Software for 2.1 &

2.2. Representative Lineweaver-Burk plots (GraFit) are shown to illustrate the competitive inhibition mode with respect to pyruvate.

PDH PDH DXPS Compound Ki (μM) Ki /Ki MAP 29.9 ± 12.6 31 1. (EAP) 46.6 ± 1.9 7 2. (BAP) 335.4 ± 7.9 60 3. (PentAP) 250 ± 31 25 4. (HexAP) 117 ± 9 13 5. (OctAP) 154 ± 14 26 6. (MPP) > 1 mM N.D. 7. > 1 mM N.D. 8. > 1 mM N.D. Table 2-2. Inhibition of the ThDP-dependent E1 subunit of PDH by alkyl acylphosphonates 2.1 – 2.8 [20]. The selectivity of inhibition is shown as a ratio of the inhibition constant for PDH vs. the inhibition constant for DXP synthase (Table 2-1, N.D.

= not determined).

2.8 Alkyl branching proximal to the acetylphosphonate scaffold is not tolerated by DXP synthase. In contrast to the previous studies with long chain alkyl acetylphosphonates, isopropylacetylphosphonate (iPrAP, 2.9) exhibits weak inhibition of

iPrAP DXP synthase with an IC50 an order of magnitude higher than that of BAP (IC50 = 250

BAP ± 70 μM; IC50 = 24 ± 4 μM, Figure 2-10). The results suggest that while acetylphosphonates bearing long alkyl chains are readily accommodated in the DXP synthase active site, analogs bearing branched alkyl groups adjacent to the acetylphosphonate moiety exhibit lower affinity for DXP synthase, presumably as a result of unfavorable steric interactions near the cofactor binding site.

Figure 2-10. iPrAP (2.9) is a weak inhibitor of DXP synthase. IC50 determinations for

BAP () and iPrAP () were carried out in the presence of pyruvate (95 μM) and D-GAP

BAP (56 μM). Acetylphosphonate concentration was varied from 0 to 250 μM. IC50 = 24

iPrAP ± 4 μM, IC50 = 250 ± 70 μM (GraphPad Prism, error represents 95% Confidence

Interval). Experiments were performed in triplicate for BAP and in duplicate for iPrAP.

2.9 BnAP is the most selective acetylphosphonate inhibitor of DXP synthase discovered to date. The aromatic acetylphosphonates 2.10 – 2.17 were designed to test the hypothesis that the sterically demanding aromatic rings would be tolerated by the DXP synthase active site and not by the smaller active sites of related ThDP-dependent enzymes.

DXPS Francine Morris tested the simplest molecule in this series, BnAP (Ki = 10.4 ±1.3 μM)

DXPS (2.10) and measured a comparable Ki, only ~2-fold higher than BAP (Ki = 5.6 ± 0.8

PDH μM) (2.2); however, it is a weaker inhibitor of PDH (Ki = 882 ± 78 μM) and is therefore

PDH DXPS a more selective inhibitor of DXP synthase (Ki /Ki = 85) [18] (Figure 2-11).

Figure 2-11. Evaluation of the inhibition of A) DXP synthase, and B) PDH by BnAP

(2.10). Representative Lineweaver-Burke plots showing BnAP competing with pyruvate for inhibition of both enzymes. A) The concentration of pyruvate was varied (20-200 μM) at several fixed concentrations of BnAP (0 (○), 15 (●), 30 (□) and 60 () μM and 100 μM

D-GAP. B) The concentration of pyruvate was varied (20-200 μM) at several fixed concentrations of BnAP (0 (○), 0.5 (●), 1 (□), 2.25 () mM). [18]

Encouraged by this preliminary success, we designed, synthesized and evaluated compounds 2.11 – 2.17 as inhibitors of DXP synthase (summarized in Table 2-3).

Francine Morris tested the para-bromo analog, 2.11, and showed that it inhibits DXP

DXPS synthase (Ki = 9.9 ± 1.3 μM) at a potency comparable to compound 2.10; however, it

PDH is also a more potent inhibitor of PDH (Ki = 364 ± 20 μM), and thus not as selective as

PDH DXPS 2.10 (Ki /Ki = 36). This was an unexpected result, as we had hypothesized that larger inhibitors should have lower affinity for the smaller PDH active site. Perhaps, as in the case of long-chain alkylacetylphosphonates (2.3 – 2.5), the more hydrophobic bromophenyl makes favorable hydrophobic contacts at a distal region in the PDH active site.

Francine Morris determined that naphthylacetylphosphonate 2.15 is a comparable

DXPS inhibitor of DXP synthase (Ki ~ 25 μM) with 2.10 and 2.11, and displays mixed-type inhibition rather than the competitive inhibition against pyruvate, as we have observed for all other acetylphosphonates tested. Stability problems with the compound limited testing before we could determine inhibitory activity against PDH. Incorporation of an extra methylene into the scaffold, as in compound 2.16, does not significantly change the activity against DXP synthase (Ki ~ 18 μM), and the other naphthyl variant, 2.14, is comparably potent, with an IC50 ~ 72 μM (compared to IC50 = 108 M for compound 2.15). Like 2.10,

DXP the biphenyl analog 2.12 is promising as a selective inhibitor of DXP synthase (IC50 synthase PDH = 17 μM, IC50 > 500 μM) but instability of 2.12 when stored in pure form limited our studies of the compound. The methoxy analog 2.13 also inhibits DXP synthase comparably to 2.10 (IC50 = 37 μM) while 2.17 is the weakest inhibitor of DXP synthase tested in this series (IC50 = 190 μM). Further SAR analysis is planned on aromatic scaffolds, and given time constraints was not pursued further here. Given the higher potency of BnAP relative to other aromatic analogs tested here, BnAP was the only aromatic pursued further in the aromatic acetylphosphonate class (Chapter 3).

Table 2-3. Evaluating aromatic acetylphosphonates (2.10 – 2.17) as inhibitors of DXP synthase and PDH.

2.10 An oxime of methylacetylphosphonate is a weak, time-dependent inhibitor of DXP synthase. Realizing that methylacetylphosphonate is an inhibitor of

DXP synthase [15, 20], and considering the stability problems that we had with some of the aromatic acetylphosphonates in the previous section and with diester precursors, we hypothesized that if we decreased the electrophilicity of the acetyl group in this scaffold, we would still have inhibitors of DXP synthase that would be more stable in solution, and could be good candidates for prodrug approaches to mask the phosphonate charges. Thus, we tested a simple oxime of methylacetylphosphonate, (2.22) synthesized previously [31],

Figure 2-12A, as an inhibitor of DXP synthase. It does not inhibit DXP synthase up to a concentration of 500 μM. We next sought to study if it inhibits DXP synthase in a time- dependent fashion, in line with its role as a less-reactive analog of MAP and time that may be required for reaction with the ThDP cofactor. A pre-incubation of DXP synthase for 15 minutes with the inhibitor at 500 μM reduces DXP synthase activity by 64% (Figure 2-

MAP 12B). However, given that MAP is a potent inhibitor of DXP synthase (Ki = 0.7 ± 0.1

μM, [15]), and the oxime can react with water to regenerate MAP and hydroxylamine, it is possible that this inhibition is due to in situ generation of MAP. Although we could not observe the formation of MAP by 31P NMR after a 30 minute incubation in the reaction mixture, it is possible that even a small population of MAP (below the limits of detection by 31P NMR) could account for the inhibition.

Figure 2-12. The oxime of MAP (2.22) shows weak, time-dependent inhibition of DXP synthase. A) Synthetic scheme to generate the oxime (2.22) [31], and B) time-dependent inhibition of DXP synthase by 2.22. The reaction was initiated with pyruvate after 15 minute incubation with or without the inhibitor; conditions are 0.15 μM DXP synthase, 50

μM D-GAP, and 100 μM pyruvate.

2.11 Methyl(chloromethyl)phosphonate is not an inhibitor of DXP synthase.

As expected from other studies with ThDP-dependent enzymes [32, 33], β-fluoropyruvate

F-Pyr (F-Pyr, 2.23) is an inhibitor of DXP synthase (Ki = 430 ±135 μM) [15], theoretically by the mechanism shown in Figure 2-13A, although it was never confirmed that this inhibition was irreversible in the case of DXP synthase. Toward identifying other, novel 65

modes of inhibition of DXP synthase, we sought to test the hypothesis that an electrophilic halomethylphosphonate (2.24) could act as an irreversible inhibitor of DXP synthase by a mechanism involving nucleophilic attack at the methylene and displacement of chloride ion (Figure 2-13B). We wanted to assess whether this compound is electrophilic enough to bind the DXP synthase active site and react irreversibly, to offer new inhibitors with novel modes of inhibition. To this end, methyl(chloromethyl)phosphonate (2.24) was synthesized from the dimethyl(chloromethyl)phosphonate, 2.25, which was prepared by J.

Kipchirchir Bitok, by the synthetic scheme shown in Figure 2-13C. Inhibition studies show, however, that 2.24 does not inhibit DXP synthase at concentrations up to 250 μM, with or without pre-incubation of the analog with DXP synthase, invalidating our hypothesis and further supporting the acetyl group as an important component of acylphosphonate DXP synthase inhibitors competitive with pyruvate.

Figure 2-13. Irreversible inhibition of ThDP-dependent enzymes by F-pyr and proposed mechanism for inhibition by halomethylphosphonates. A) F-pyr (2.23) is an irreversible

inhibitor of ThDP-dependent enzymes. B) The halomethylphosphonate (2.24) is hypothesized to be an irreversible inhibitor by the mechanism shown, and C) 2.24 was generated via pictured the synthetic route.

Discussion

Alkylacetylphosphonates 2.1 – 2 .5 were designed, synthesized and evaluated as potential selective inhibitors of E. coli DXP synthase using the previously mentioned spectrophotometric IspC-coupled assay. While MAP has previously been shown to act competitively against pyruvate as an inhibitor of DXP synthase with a Ki of 0.7 ± 0.1 μM

[15], it also displays inhibitory activity against the related homolog, PDH, with a Ki of 29.9

± 12.6 μM under our assay conditions [20].

Unfortunately, our inhibitor design could not be guided by the published crystal structure of DXP synthase, showing only the cofactor bound [34]. It would be advantageous to know the active site architecture at the pre-decarboxylation (ternary complex) step of the reaction coordinate. Conceivably, this structure could differ from the existing structure and may better inform our inhibitor design as a goal of our studies is to selectively target this form.

Without this information, we embarked to gain selectivity against DXP synthase by increasing the steric bulk of our inhibitors and thereby take advantage of the large active site volume presumably required for ternary complex formation on DXP synthase [16, 17,

35]. In accordance with our hypothesis, addition of steric bulk to the phosphonoester portion of the acetylphosphonate scaffold (compounds 2.1 – 2.5) was tolerated by the active

site of DXP synthase. These inhibitors exhibit low micromolar Ki values for DXP synthase, within 5-fold of MAP. They are also competitive against pyruvate. Branching immediately proximal to the acetylphosphonate is not tolerated, as compound 2.9 inhibits

DXP synthase with a 10-fold higher IC50 than BAP (2.2). Variations on the acetyl portion of the scaffold are also not well-tolerated, as had been observed with the ketoacids evaluated by Leighanne Brammer Basta [17] and further confirmed by the negligible inhibitory activity of compounds 2.6 – 2.8. Similarly, the acetyl portion seems to be required for efficient inhibition as neither the oxime nor chloromethyl analogs (2.22 &

2.24, respectively) are effective inhibitors of DXP synthase. Further work could be done to look at other acetyl isosteres, such as hydrazine, to see if they have inhibitory activity, although it is possible that there is not sufficient space in the enzyme active near the cofactor to accommodate larger isosteres. This seems to be supported by the lack of inhibitory activity by compounds 2.6 – 2.8, suggesting changes to the acetyl group are not as well tolerated by the enzyme.

While MAP is already a less potent inhibitor of the related homolog, PDH, the analogs 2.2 and 2.3 are significantly less potent against PDH compared to MAP.

Interestingly, although we expected this trend to continue with increasing alkyl chain length, the weakest inhibitor of PDH (and therefore the most selective for DXP synthase)

PDH DXP synthase in the alkylacetylphosphonates series is BAP (the ratio of Ki /Ki for BAP is 60- fold, compared to 30-fold for MAP). The longer chain alkyl compounds 2.3 – 2.5 all display comparable or stronger inhibition of PDH compared to BAP. It is possible that increasing the hydrophobicity of the acetylphosphonate scaffold increases affinity for the

PDH active site by a mode as yet uncharacterized; however further modeling studies would be required to explore this hypothesis.

None of the compounds bearing short alkyl chains (2.1 – 2.2, 2.6 – 2.8) tested are inhibitors of TK, a ThDP-dependent enzyme which does not utilize pyruvate as a substrate.

This observation further supports our hypothesis that acylphosphonates are acting as pyruvate mimics in the DXP synthase and PDH active sites. This is corroborated by the kinetics data which shows they inhibit the other two enzymes in a competitive manner with pyruvate [20].

Following on the discovery by Francine Morris [18] that DXP synthase can successfully utilize aromatic aldehydes and nitroso compounds as substrates (Table 2-1), we sought to evaluate their incorporation into the acetylphosphonate scaffold. Our rationale was that the aromatic rings could impart further selectivity into our inhibitors, as they would hypothetically not be able to fit into the active site of PDH or other related

ThDP-dependent enzymes. This was supported by the observation that although the aromatic nitroso compounds are also substrates for PDH, they have significantly higher KM values. To pursue these studies, aromatic compounds 2.10 – 2.18 were synthesized and evaluated as potential DXP synthase inhibitors. All showed inhibitory activity against

DXP synthase which provided evidence that although the DXP synthase active site does not tolerate branching directly proximal to the phosphonate, as in the case of 2.9, it can accommodate steric bulk one or two methylene units removed from this position. The most potent inhibitors of DXP synthase in this series are 2.10 and 2.11. Although the para- bromo substituent of 2.11 is tolerated by DXP synthase without a loss of inhibitory activity, the compound is also more potent against PDH, making it a less selective inhibitor.

Although this discovery was unexpected, it is consistent with our discovery in the alkylacetylphosphonate series suggesting long-chain alkyl groups display higher affinity for PDH, perhaps a result of favorable hydrophobic interactions in the PDH active site.

Further SAR studies are needed to optimize the alkylacetylphosphonates scaffold.

It is interesting that biphenyl analog 2.12 and naphthylethyl analog 2.16 show comparable activity against DXP synthase which seems to further support a model that branching is better tolerated further away from the phosphonyl scaffold. The other naphthyl (2.14 and 2.15) and the meta-methoxybenzyl (2.13) compounds show slightly weaker inhibitory activity against DXP synthase, suggesting para-substituents may be better tolerated in the enzyme active site.

Experimental

Enzyme kinetic analyses were performed by myself and Jessica Smith (2.1 – 2.2 &

2.6 – 2.8). Suppression of DXP synthase activity was measured spectrophotometrically using IspC as a coupling enzyme and monitoring NADPH consumption by IspC [15, 28].

The concentration of D-GAP in stock solutions was determined by using the GAPDH assay

[17]. DXP synthase reaction mixtures containing 2-[4-(2-hydroxyethyl)piperazin-1- yl]ethanesulfonic acid (HEPES) (100 mM, pH 8.0), 1 mg/ml bovine serum albumin (BSA),

2 mM MgCl2, 2.5 mM tris(2-carboxyethyl)phosphine (TCEP), 1 mM ThDP, 100 μM

NADPH, 1 μM IspC, D,L-GAP, and pyruvate were preincubated at 37 °C for 5 min. DXP synthase (100 nM) was added to initiate each reaction, and the rate of NADPH depletion in the coupled step was monitored spectrophotometrically at 340 nm at 37 °C. The rate of

NADPH depletion was used to calculate initial rates of DXP formation. All reactions were performed in triplicate.

DXP synthase reaction mixtures containing 100 mM HEPES buffer, pH 8.0

(described above) were pre-incubated at 37°C for 5 minutes with varying inhibitor concentrations. The reaction was initiated with enzyme. Inhibition of DXP synthase by compounds 2.7 – 2.8 was not observed. Inhibition of the coupling enzyme (IspC) by the inhibitors was not observed. Inhibition experiments for (2.1 – 2.2 & 2.6 – 2.8) were performed in duplicate and all others were performed in triplicate. Double reciprocal analysis of data was carried out using GraFit version 7 from Erithacus Software for compounds 2.1 – 2.2 & 2.6 – 2.8 (Jessica Mott Smith) and 2.10 – 2.12 (Francine Morris).

All other data was evaluated via non-linear regression analysis by GraphPad Prism.

Pyruvate dehydrogenase activity was measured spectrophotometrically as previously reported [36] by measuring changes in absorbance at 340 nm. The assay medium (30°C) contained 100 mM HEPES (pH 8.0), 1 mg/mL BSA, 0.2 mM ThDP, 0.1 mM coenzyme A, 1 mM MgCl2, 2 mM cysteine, 0.3 mM TCEP. The reaction was initiated with enzyme. Suppression of pyruvate dehydrogenase activity was measured spectrophotometrically by measuring changes in optical density at 340 nm. Pyruvate dehydrogenase reaction mixtures containing 100 mM HEPES buffer, pH 8.0 (described above) were pre-incubated at 30°C for 5 minutes with varying inhibitor concentrations.

Inhibition of porcine pyruvate dehydrogenase by compounds 2.6 – 2.8 was not observed up to 1 mM. Inhibition experiments of PDH by (2.1 – 2.2 & 2.6 – 2.8) were performed in duplicate, while all other experiments were performed in triplicate. Double reciprocal analysis of data was carried out using GraFit version 7 from Erithacus Software for 2.1 –

2.2 & 2.6 – 2.8 (Jessica Mott Smith) and 2.10 – 2.12 (Francine Morris). All other data was evaluated via non-linear regression analysis by GraphPad Prism.

Transketolase activity was measured as previously reported by measuring changes in optical density at 340 nm (Jessica Mott Smith) [37]. The basic assay medium (37°) contained 50 mM glycylgylcine (pH = 7.7), 10 mM sodium arsenate, GADPH, 3.2 TCEP,

2.5 CaCl2, 80 μM ThDP, 70 μM xylulose-5-phosphate, 0.5 mM ribose-5-phosphate, 0.4 mM β-nicotinamide adenine dinucleotide phosphate sodium salt hydrate (NAD+). The reaction was initiated with enzyme. Inhibition of transketolase from baker’s yeast by compounds 2.1 – 2.2 & 2.6 – 2.8 was not observed up to 1 mM. All experiments were performed in duplicate.

Synthesis of acetylphosphonates

General Experimental. Trialkyl phosphites, acyl chlorides, benzyl alcohols and lithium bromide were obtained from commercial sources and used without further purification. Methylene chloride, acetonitrile and diisopropylethylamine (DIPEA, Hünig’s base) were distilled over calcium hydride and collected under argon. All reactions were carried out in oven-dried glassware under an inert argon atmosphere. NMR spectra were recorded on a Bruker 400 MHz spectrometer for dialkyl acyl phosphonate intermediates, or a Varian 500 MHz spectrometer for final compounds, and processed via the ACD/NMR

Processor Academic Edition. Chemical shifts are reported in units of parts per million

(ppm), relative to a standard reference. 1H NMR chemical shifts are reported relative to the residual 1H signal of the deuterated solvent as an internal reference (CDCl3 δ = 7.27 ppm;

31 D2O δ = 4.75 ppm). P chemical shifts are reported relative to triphenylphosphine oxide

(TPPO, δ = 0 ppm) as an external standard. Mass spectrometry analysis was carried out at

University of Illinois at Urbana-Champagne, School of Chemical Sciences, Mass

Spectrometry Laboratory. Chemical synthesis and characterization of MAP was reported previously [17]. Compound 2.7 was prepared according to published protocol [21].

Tribenzyl phosphite (reported previously [26]) and methyl (chloromethyl)phosphonate were prepared by J. Kipchirchir Bitok.

Synthesis of ethylacetylphosphonate (2.1). An oven-dried flask was equipped with a stir-bar and charged with 0.21 mL of acetyl chloride (0.24g, 3.0 mmol). Triethyl phosphite (0.52 mL, 3.0 mmol) was added drop-wise at room temperature. The solution was stirred at room temperature for about 1 hour, until triethyl phosphite had disappeared

(as indicated by 31P NMR analysis). Diethyl acylphosphonate (2.18) was isolated as a pale yellow oil following vacuum distillation (241.6mg, 97% pure as determined by 31P NMR,

31 45% yield) and was carried on without further purification. P-NMR (CDCl3): δ -27.84 (s)

1 H-NMR (CDCl3): δ 1.08 (t, 6H), 2.182 (d, 3H), 4.02 (m, 4H).

Diethyl acetylphosphonate (2.18) was dissolved in 1.7 mL of anhydrous acetonitrile. Lithium bromide (106.8 mg, 1.2 mmol) was added in one portion, and the reaction mixture was heated to 70 oC and stirred overnight. The resulting white solid was filtered and washed with two 10 mL portions of anhydrous acetonitrile followed by two 10 mL portions of diethyl ether. Ethyl acetylphosphonate (1) was isolated as a white solid 73

(132.5 mg, 69% yield). 31P-NMR (D2O): δ -27.08 (s); 1H-NMR (D2O): δ 1.17 (t, 3H), 2.34

(d, 3H), 3.84 (m, 2H). HRMS (ESI), calculated m/z for C4H9LiO4P (free acid form),

[M+H]+ = 159.0399; observed: 159.0398.

Synthesis of butylacetylphosphonate (2.2). An oven-dried flask was equipped with a stir-bar and charged with 0.40 mL of acetyl chloride (0.44g, 5.6 mmol). Tributyl phosphite (1.5 mL, 6.0 mmol) was added drop-wise at room temperature. The solution was stirred at room temperature for about 1 hour, until tributyl phosphite had disappeared (as indicated by 31P NMR analysis). Dibutyl acylphosphonate (2.19) was isolated as a pale yellow oil following vacuum distillation (754mg, 90% pure as determined by 31P NMR,

64% yield) and was carried on without further purification. 31P-NMR (CDCl3): δ -28.75

(s) 1H-NMR (CDCl3): δ 0.94 (t, 6H), 1.42 (m, 4H), 1.71 (m, 4H), 2.49 (d, 3H), 4.16 (m,

4H).

Dibutyl acetylphosphonate (2.19) was dissolved in 4 mL of anhydrous acetonitrile.

Lithium bromide (269.7 mg, 3.1 mmol) was added in one portion, and the reaction mixture was heated to 70 oC and stirred overnight. The resulting white solid was filtered and washed with two 10 mL portions of anhydrous acetonitrile followed by two 10 mL portions of diethyl ether. Butyl acetylphosphonate (2.2) was isolated as a white solid (261.1 mg, 44%).

31P 1 -NMR (D2O): δ -27.01(s); H-NMR (D2O): 0.76 (t, 3H), 1.22 (m, 2H), 1.47 (m, 2H),

2.29 (d, 3H), 3.76 (m, 2H). HRMS (ESI), calculated m/z for C6H13LiO4P (free acid form),

[M+H]+ = 187.0712; observed: 187.0709.

Synthesis of pentylacetylphosphonate (2.3). A flame-dried flask, cooled under argon, was charged with acetyl chloride (0.7 mL, 10 mmol). Tripentyl phosphite (0.98 g,

3.4 mmol) was dissolved in anhydrous CH2Cl2 (32 mL), and the resulting solution was added drop-wise to the stirring acetyl chloride. Following addition of the trialkyl phosphite, argon was bubbled through the reaction mixture to remove HCl byproduct. The progress of the reaction was monitored by 31P NMR spectroscopy, and the complete conversion of tripentyl phosphite (δ = 113 ppm) to dipentylacetylphosphonate (δ = - 28 ppm) was observed within 1 h. Volatiles were removed in vacuo, and the crude material was used without further purification. Dipentylacetylphosphonate was dissolved in anhydrous acetonitrile (5.6 mL), and lithium bromide (0.38 g, 4.4 mmol) was added in one portion. The reaction mixture was heated to 65 oC and stirred overnight. The lithium salt of pentylacetylphosphonate precipitated from solution and was removed by filtration. The filter cake was washed successively with cold acetonitrile, diethyl ether and methylene chloride (30 mL portions of). Lithium pentylacetylphosphonate was isolated as a white

1 powder (177 mg, 26% yield). H NMR (D2O): δ = 0.81 ppm (t, 3H), 1.26 ppm (m, 4H),

31 1.53 ppm (m, 2H), 2.37 ppm (d, 3H), 3.82 ppm (m, 2H); P NMR (D2O): δ = - 27.6 ppm

+ + (s); HRMS (ESI): m/z calcd for C7H16O4P (H form): 195.0786 [M+H] , found: 195.0788

Synthesis of hexylacetylphosphonate (2.4). A flame-dried flask, cooled under argon, was charged with acetyl chloride (1.1 mL, 15 mmol). Trihexyl phosphite (1.68 g,

5.0 mmol) was dissolved in anhydrous CH2Cl2 (50 mL), and the resulting solution was added drop-wise to the stirring acetyl chloride. Following addition of the trialkyl phosphite, argon gas was bubbled through the reaction mixture to remove HCl byproduct.

The progress of the reaction was monitored by 31P NMR spectroscopy, and the complete conversion of trihexyl phosphite (δ = 117 ppm) to dihexylacetylphosphonate (δ = - 28 ppm) was observed within 2 h. Volatiles were removed in vacuo, and the crude material was used without further purification. Dihexylacetylphosphonate was dissolved in anhydrous acetonitrile (8.3 mL), and lithium bromide (0.65 g, 7.5 mmol) was added in one portion.

The reaction mixture was heated to 65 oC and stirred overnight. The lithium salt of hexylacetylphosphonate precipitated from solution and was removed by filtration. The filter cake was washed successively with cold acetonitrile, diethyl ether and methylene chloride (30 mL portions). Lithium hexylacetylphosphonate was isolated as a white solid

1 (130 mg, 12% yield over two steps). H NMR (D2O): δ = 0.77 ppm (t, 3H), 1.20 ppm (m,

31 6H), 1.56 ppm (m, 2H), 2.35 ppm (d, 3H), 3.83 ppm (m, 2H); P NMR (D2O): δ = - 27.6

+ + ppm (s); HRMS (ESI): m/z calcd for C8H18O4P (H form): 209.0943 [M+H] , found:

209.0944

Synthesis of octylacetylphosphonate (2.5). A flame-dried flask, cooled under argon, was charged with acetyl chloride (1.1 mL, 15 mmol). Trioctyl phosphite (2.1 g, 5.0 mmol) was dissolved in anhydrous CH2Cl2 (50 mL), and the resulting solution was added drop-wise to the acetyl chloride. Following addition of the trialkyl phosphite, argon was bubbled through the reaction mixture. The progress of the reaction was monitored via 31P

NMR spectroscopy, and the complete conversion of trioctyl phosphite (δ = 118 ppm) to dioctylacetylphosphonate (δ = -28 ppm) was observed within 1 h. Volatiles were removed in vacuo, and the crude material was used without further purification.

Dioctylacetylphosphonate was dissolved in anhydrous acetonitrile (8.3 mL), and lithium bromide (0.65 g, 7.5 mmol) was added in one portion. The reaction was heated to 60 oC and stirred overnight. The lithium salt of octylacetylphosphonate precipitated from solution and was removed by filtration. The filter cake was washed successively with cold acetonitrile and diethyl ether (30 mL portions of). Lithium octylacetylphosphonate was

1 isolated as a white powder (290 mg, 24 % over two steps). H NMR (D2O): δ = 0.7 ppm

(m 7H), 0.93 ppm (m, 2H), 1.2 ppm (m, 4H), 1.5 ppm (m, 2H), 2.36 ppm (d, 3H),3.85 ppm

31 + (m, 2H); P NMR (D2O): δ = - 27.7 ppm (s); HRMS (ESI): m/z calcd for C10H22O4P (H form): 237.1256 [M+H]+, found: 237.1255.

Synthesis of methylpropionylphosphonate (2.6). An oven-dried flask was equipped with a stir-bar and charged with 0.50 mL of propionyl chloride (0.53g, 5.5 mmol).

Trimethyl phosphite (0.60 mL, 5.1 mmol) was added drop-wise at room temperature. The solution was stirred at room temperature for about 1 hour, until trimethyl phosphite had disappeared (as indicated by 31P NMR analysis). Dimethylpropionylphosphonate was isolated as a pale yellow oil following vacuum distillation (542.3 mg, 98% pure as determined by 31P NMR, 64% yield) and was carried on without further purification. 31P-

NMR (CDCl3): δ -26.77 (s)

Dimethylpropionylphosphonate was dissolved in 4 mL of anhydrous acetonitrile.

Lithium bromide (269.7 mg, 3.1 mmol) was added in one portion, and the reaction mixture was heated to 70 oC and stirred overnight. The resulting white solid was filtered and washed with two 10 mL portions of anhydrous acetonitrile followed by two 10 mL portions of diethyl ether. Methylpropionylphosphonate 2.6 was isolated as a white solid (241.0 mg,

31 1 47%). P-NMR (D2O): δ -27.01; H-NMR (D2O): 0.76 (t, 3H), 1.22 (m, 2H), 1.47 (m, 2H),

2.29 (d, 3H), 3.76 (m, 2H). ESI (M/S) calculated m/z for C4H9LiO4P (free acid form),

[M+H]+ = 159.0403; observed: 159.0399.

Synthesis of methylvaleroylphosphonate (2.8). An oven-dried flask was equipped with a stir-bar and charged with 0.3 mL of valeroyl chloride (0.3g, 2 mmol), to which 0.3mL of trimethyl phosphite (0.3g, 2 mmol) was added. After one hour, reaction mixture was diluted with 3.1 mL of anhydrous acetonitrile. To this solution, 220 mg of lithium bromide (2.53 mmol) was added in one portion. The reaction mixture was warmed to 70oC overnight. The solid white precipitate was filtered with two 10 mL portions of anhydrous acetonitrile, followed by two 10 mL portions of diethyl ether.

Methylvaleroylphosphonate was isolated as a white solid (110.3mg, 23% overall yield).

31 1 P-NMR (D2O): δ -25.89 (s); H-NMR (D2O): 0.79 (t, 3H), 1.18 (m, 2H), 1.46 (m, 2H),

2.73 (t, 2H), 3.52 (d, 3H). HRMS (ESI), calculated m/z for C6H13LiO4P (free acid form),

[M+H]+ = 187.0712; observed: 187.0708.

Synthesis of isopropylacetylphosphonate (2.9). A flame-dried flask, cooled under argon, was charged with acetyl chloride (0.3 mL, 4 mmol). Triisopropyl phosphite

(0.9 mL, 4 mmol) was added neat to the acetyl chloride. The progress of the reaction was monitored via 31P NMR spectroscopy, and the complete conversion of the trialkyl phosphite (δ = 113 ppm) to diisopropylacetylphosphonate (δ = -27 ppm) was observed within 1 h. The product was purified by vacuum distillation and isolated as a pale yellow oil (471 mg, 59% yield). Diisopropylacetylphosphonate was dissolved in anhydrous acetonitrile (2.8 mL), and lithium bromide (0.18 g, 2.0 mmol) was added in one portion.

The reaction was heated to 70 oC and stirred overnight. The lithium salt of

isopropylacetylphosphonate precipitated from solution and was removed by filtration. The filter cake was washed successively with cold acetonitrile and diethyl ether (15 mL portions of). Lithium isopropylacetylphosphonate was isolated as a white powder (57 mg, 15%

1 31 yield). H NMR (D2O) 1.21 ppm (d, 6H), 2.37 ppm (d, 3H), 4.36 ppm (m, 1H); P NMR

+ (D2O): δ = - 28.3 ppm (s); HRMS (ESI): m/z calcd for C5H12O4P (H form): 167.0473

[M+H]+, found: 167.0469.

Synthesis of benzylacetylphosphonate (2.10). Tribenzyl phosphite was generated from benzyl alcohol, diisopropylethylamine, and phosphorous trichloride according to

Saady et al [26] and was supplied by J. Kipchirchir Bitok. The spectral properties of the compound were identical to published values. For the preparation of benzylacetylphosphonate (BnAP), a flame-dried flask, cooled under argon, was charged with acetyl chloride (0.32 mL, 4.5 mmol). Tribenzyl phosphite (0.46 g, 1.3 mmol) was dissolved in anhydrous CH2Cl2 (13 mL), and the resulting mixture was added dropwise to acetyl chloride. The progress of the reaction was monitored through 31P NMR spectroscopy, and complete conversion of tribenzyl phosphite (δ = 113 ppm) to dibenzylacetyl-phosphonate (δ = ~26 ppm) was observed within 1 h. Volatiles were removed in vacuo, and the crude material was used without further purification.

Dibenzylacetylphosphonate was dissolved in anhydrous acetonitrile (2.2 mL), and lithium

bromide (0.17 g, 0.95 mmol) was added in one portion. The reaction mixture was heated to 50 oC for ~4 h. The lithium salt of benzylacetylphosphonate precipitated from solution and was removed by filtration. The filter cake was washed successively with cold acetonitrile and diethyl ether (20 mL portions of). The crude product was purified by

-1 reversed-phase preparative HPLC. Flow rate= 10 mLmin ; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol, method 5–80% B over 75 min. The purity of fractions was

-1 determined by analytical RP-HPLC. Flow rate=3 mLmin ; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol, method 5–80% B over 12 min. Combined fractions were lyophilized to yield 0.0975 g BnAP as the triethylammonium salt (24% over two steps). 1H

NMR (D2O): δ = 1.20 (t, 9H), 2.31 (d, 3H), 3.11 (m, 6H), 4.91 (d, 2H), 7.35 ppm (m, 5H);

31 P NMR (D2O): δ = -27.43 ppm (s); HRMS (ESI): m/z calcd for C15H27NO4P

(triethylammonium salt): 316.1678 [M+H]+, found: 316.1673.

Synthesis of para-bromobenzylacetylphosphonate (2.11). A flame-dried flask, cooled under argon, was charged with 4-bromobenzyl alcohol (0.65 g, 3.4 mmol) dissolved in 11.5 mL of anhydrous methylene chloride. The flask was cooled to 0 oC, and Hünig’s base (0.7 mL, 4 mmol) was added followed by phosphorus trichloride (0.1 mL, 1 mmol).

31 After 20 minutes, the reaction was deemed complete by P NMR, and PCl3 was consumed while tris(4-bromobenzyl) phosphite was observed (δ = 114 ppm). The reaction mixture

was washed with ice water, and the organic layer was dried over Na2SO4 and volatiles were removed in vacuo to yield 0.65 g of tris(4-bromobenzyl)phosphite as a yellow, flaky solid

(96% yield) which was carried forward without further purification.

The phosphite was dissolved in 11 mL of anhydrous methylene chloride and added to a flask charged with acetyl chloride (0.24 mL, 3.4 mmol) and stirred under argon. After

45 minutes, the reaction progress was assessed via 31P NMR. The phosphite had been consumed and di(4-bromobenzyl)acetylphosphonate was observed (δ = -28 ppm).

Volatiles were removed in vacuo to yield approximately 0.65 g of crude material which was dissolved in 2.3 mL anhydrous acetonitrile and subjected to lithium bromide (0.18 g,

2.1 mmol) at 55 oC. The lithium salt of 4-bromobenzylacetylphosphonte precipitated from solution and was isolated by filtration, followed by successive washes with cold acetonitrile and ether. Approximately 0.19 g (~ 86% purity as determined by 31P NMR) of crude material was isolated and purified via preparative HPLC.

A 0.05 g aliquot of the crude product was purified by reversed-phase preparative

HPLC. Flow rate = 10 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method: 5–80% B over 75 min. The purity of fractions was determined by analytical RP-HPLC. Flow rate = 3 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method: 5–80% B over 12 min. Combined fractions were lyophilized to yield 0.0192 g p-BrBnAP as the triethylammonium salt (4.2% over three

1 steps). H NMR (D2O): δ = 1.19 (t, 9H), 2.31 (d, 3H), 3.09 (m, 6H), 4.85 (d,2 H), 7.27 ppm

31 (d, 2H), 7.52 ppm (d, 2H); P NMR (D2O): δ = -27.45 ppm (s); HRMS (ESI): m/z calcd

+ for C15H26NO4PBr (triethylammonium salt): 394.0783 [M+H] , found: 394.0800.

Synthesis of para-phenylbenzylacetylphosphonate (2.12). Synthesis of 2.12 was performed by Nick Calcaterra. A flame-dried flask, cooled under argon, was charged with

4-phenylbenzyl alcohol (0.44 g, 2.4 mmol)) dissolved in 12 mL of dry methylene chloride.

The flask was cooled to 0 oC and Hünig’s base (0.4 mL, 2.4 mmol) was added followed by the addition of phosphorus trichloride (0.07 mL, 0.8 mmol). Reaction progress was

31 determined by TLC. P NMR analysis confirmed PCl3 was consumed while tris(4- phenylbenzyl) phosphite was observed (δ = 114 ppm). Volatiles were removed in vacuo, and the product was eluted from a silica gel column with 1:1 ethyl acetate/hexanes to yield

0.1 g of tris(4-phenylbenzyl)phosphite which was brought on without further workup.

The phosphite was dissolved in 2 mL of dry methylene chloride and added to a flask charged with acetyl chloride (0.06 mL, 0.8 mmol) and stirred under argon. Reaction progress was checked by 31P NMR analysis. After 30 min, the phosphite had been consumed and di(4-phenylbenzyl)acetylphosphonate was observed (δ = -28 ppm).

Volatiles were removed in vacuo to yield approximately 0.1 g of crude material which was dissolved in 0.3 mL dry acetonitrile and subjected to lithium bromide (0.03 g, 0.3 mmol) at 55 oC. The lithium salt of 4-phenylbenzylacetylphosphonte precipitated from solution and was isolated by filtration, followed by successive washes with cold acetonitrile and

ether. Approximately 0.05 g of crude material was isolated and purified via preparative

HPLC to yield 0.006 g of 4-phenylbenzylacetylphosphonate as a triethylammonium salt

1 (20% yield over 3 steps) H NMR (D2O): δ = 1.19 (t, 9H), 2.30 (d, 3H), 3.09 (m, 6H), 4.94

31 (d,2 H), 7.37 ppm (t, H), 7.46 ppm (m, 4H); 7.64 ppm (m, 4H); P NMR (D2O): δ = -27.67

+ ppm (s); HRMS (ESI): m/z calcd for C15H15O4PNa (sodium salt): 313.0606 [M+H] , found: 313.0609.

Synthesis of meta-methoxybenzylacetylphosphonate (2.13). A flame-dried flask, cooled under argon was charged with 3-methoxybenzyl alcohol (0.66 mL, 5.3 mmol)) dissolved in 17 mL of an anhydrous mixture of 1:1 diethyl ether and tetrahydrofuran. The flask was cooled to -40 oC and Hünig’s base (1.0 mL, 6 mmol) was added followed by phosphorus trichloride (0.15 mL, 1.7 mmol). After 20 minutes, the

31 reaction was deemed complete by P NMR analysis, and PCl3 was consumed while tris(4- bromobenzyl) phosphite was observed (δ = 113 ppm). Solid precipitate was filtered, and the soluble organic layer was washed with 1M KOH followed by cold brine, and the organic layer was dried over sodium sulfate. Volatiles were removed in vacuo to yield

0.62 g of tris(3-methoxybenzyl)phosphite as a yellow oil (82% yield, 85% purity by 31P

NMR) which was carried forward without further purification.

The phosphite (0.3 g) was dissolved in 7 mL of dry methylene chloride and added to a flask charged with acetyl chloride (1 mL, 14 mmol) and stirred under argon. After 45 minutes, reaction progress was checked via 31P NMR. The phosphite had been consumed and di(3-methoxybenzyl)acetylphosphonate was observed (δ = -28 ppm). Volatiles were removed in vacuo to yield crude material which was dissolved in 4.9 mL dry acetonitrile and subjected to lithium bromide (0.16 g, 1.8 mmol) at 50 oC. The lithium salt of 3- methoxybenzylacetylphosphonte precipitated from solution and was isolated by filtration, followed by successive washes with cold acetonitrile and ether. Approximately 0.03 g

(approximately 54% pure by 31P NMR) of crude material was isolated and purified via preparative HPLC.

The crude product was purified by reversed-phase preparative HPLC. Flow rate =

10 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method:

5–80% B over 75 min. The purity of fractions was determined by analytical RP-HPLC.

Flow rate = 3 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method: 5–80% B over 12 min. Combined fractions were lyophilized to yield 0.031 g 3- methoxybenzyl acetylphosphonate as the triethylammonium salt (5.2% over three steps).

1 H NMR (D2O): δ = 1.23 (t, 9H), 2.29 (d, 3H), 3.15 (m, 6H), 5.1 (d, 2H), 6.96 (d, H), 7.56

31 (m, 4H), 7.94 (m, 3H); P NMR (D2O): δ = -27.3 ppm (s); HRMS (ESI): m/z calcd for

+ C10H13O5PNa (sodium salt): 267.0398 [M+H] , found: 267.0397.

Synthesis of 2-naphthylacetylphosphonate (2.14). A flame-dried flask, cooled under argon was charged with 2-naphthalenemethanol (0.33 g, 2.0 mmol) dissolved in 6.7 mL of anhydrous methylene chloride. The flask was cooled to -78 oC, and Hünig’s base

(0.4 mL, 2.2 mmol) was added, followed by the addition of phosphorus trichloride (0.06 mL, 0.7 mmol). After 90 minutes, reaction progress was determined via 31P NMR, and

PCl3 was consumed while tris(2-naphthyl)phosphite was observed (δ = 114 ppm).

Reaction was quenched and washed with saturated ammonium chloride and dried over sodium sulfate. Volatiles were removed in vacuo and eluted from a silica plug with 1:10 ethyl acetate/hexanes to yield 0.08 g of tris(2-naphthyl)phosphite as a yellow oil which was carried forward without further purification.

The phosphite was dissolved in 1 mL of anhydrous methylene chloride and added to a flask cooled to 0 oC and charged with acetyl chloride (0.03 mL, 0.5 mmol). The reaction mixture was warmed to room temperature and reaction progress was assessed by

31P NMR. After 30 minutes, reaction was determined to be complete, as the phosphite was consumed and the product, di(2-naphthyl)acetylphosphonate was observed (δ = -28 ppm).

Volatiles were removed in vacuo and the crude material was dissolved in 0.6 mL dry acetonitrile and subjected to lithium bromide (0.014 g, 0.16 mmol) at 50 oC for 90 minutes.

2-naphthylacetylphosphonate (0.017 g) precipitated from solution as a lithium salt. 86

The crude product was purified by reversed-phase preparative HPLC. Flow rate =

10 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method

5–80% B over 75 min. The purity of fractions was determined by analytical RP-HPLC.

Flow rate = 3 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method: 5–80% B over 12 min. Combined fractions were lyophilized to yield 0.055 g 2- naphthylacetylphosphonate as the triethylammonium salt (2.9% over three steps). 1H NMR

(D2O): δ = 1.19 (t, 9H), 2.24 (d, 3H), 3.12 (m, 6H), 5.04 (d, 2 H), 7.51 (m, 3H), 7.88 (m,

31 4H); P NMR (D2O): δ = -27.3 ppm (s); HRMS (ESI): m/z calcd for C13H13O4PNa (sodium salt): 287.0449 [M+H]+, found: 287.0444.

Synthesis of 1-naphthylacetylphosphonate (2.15). A flame-dried flask, cooled under argon, was charged with 1-naphthalenemethanol (0.56 g, 3.5 mmol) dissolved in

11.5 mL of anhydrous methylene chloride. The flask was cooled to -78 oC and Hünig’s base (0.7 mL, 4.0 mmol) was added followed by phosphorus trichloride (0.1 mL, 1.2 mmol). Reaction progress was monitored via 31P NMR, and considered complete when

PCl3 was consumed while tris(1-naphthyl)phosphite was observed (δ = 115 ppm). The reaction mixture and washed with cold water and the organic layer was dried over sodium sulfate. Volatiles were removed in vacuo to yield 0.42 g of the phosphite, which was carried forward without further purification.

The phosphite was dissolved in 8.3 mL of anhydrous methylene chloride and added to a flask charged with acetyl chloride (0.1 mL, 1.2 mmol) and stirred under argon.

Reaction progress was assessed via 31P NMR at 90 minutes and the phosphite had been consumed and the product, di(1-naphthyl)acetylphosphonate was observed (δ = -28 ppm).

Volatiles were removed in vacuo and the crude product was dissolved in 1.3 mL dry acetonitrile and subjected to lithium bromide (0.11 g, 1.2 mmol) at 50 oC for 4 hours. 1- naphthylacetylphosphonate (0.11 g) precipitated from solution as a lithium salt.

An aliquot of the crude product was purified by reversed-phase preparative HPLC.

Flow rate = 10 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol;

HPLC method: 5–80% B over 75 min. The purity of fractions was determined by analytical

RP-HPLC. Flow rate = 3 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method: 5–80% B over 12 min. Combined fractions were lyophilized to yield 0.043 g 1-naphthylacetylphosphonate as the triethylammonium salt (10% over three

1 steps). H NMR (D2O): δ = 1.19 (t, 9H), 2.16 (d, 3H), 3.10 (m, 6H), 5.39 (d, 2 H), 7.50 (m,

31 4H), 7.93 (m, 2H), 8.15 (d, H); P NMR (D2O): δ = -27.3 ppm (s); HRMS (ESI): m/z calcd

+ for C13H13O4PLi (lithium salt): 271.0712 [M+H] , found: 271.0725.

Synthesis of 2-(naphthalen-1-yl)ethylacetylphosphonate (2.16). A flame-dried flask, cooled under argon was charged with 2-(1-naphthyl) ethanol (0.90 g, 5.2 mmol) 88

dissolved in 17 mL of an anhydrous mixture of 1:1 diethyl ether and tetrahydrofuran. The flask was cooled to -40 oC and Hünig’s base (1.0 mL, 6.0 mmol) was added followed by phosphorus trichloride (0.15 mL, 1.7 mmol). After 60 minutes, the reaction progress was

31 assessed via P NMR, and was deemed complete as PCl3 was consumed while tris(1- naphthylethyl)phosphite was observed (δ = 115 ppm). The reaction mixture and washed with cold 1M KOH, and the organic layer was dried over sodium sulfate. Volatiles were removed in vacuo to yield 0.75 g of the crude phosphite, which was carried forward without further purification.

The phosphite was dissolved in 12 mL of anhydrous methylene chloride and added to a flask charged with acetyl chloride (0.3 mL, 4.2 mmol). At 30 minutes, reaction progress was assessed by 31P NMR. The phosphite had been consumed and the product, di(1-naphthylethyl)acetylphosphonate was observed (δ = -28 ppm). Volatiles were removed in vacuo and the crude product mix was dissolved in 1.4 mL dry acetonitrile and subjected to lithium bromide (0.27 g, 3.1 mmol) at 60 oC for 4 hours. 1- naphthylacetylphosphonate (0.08 g) precipitated from solution as a lithium salt.

An aliquot of the crude product was purified by reversed-phase preparative HPLC.

Flow rat e= 10 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol;

HPLC method: 5–80% B over 75 min. The purity of fractions was determined by analytical

RP-HPLC. Flow rate = 3 mL/min; solvent A: HNEt3OAc (50 mm, pH 6.0), solvent B: methanol; HPLC method 5–80% B over 12 min. Combined fractions were lyophilized to yield 0.035 g 1-naphthylethylacetylphosphonate as the triethylammonium salt (5.4% over

1 three steps). H NMR (D2O): δ = 1.24 (t, 9H), 2.05 (d, 3H), 3.16 (m, 6H), 3.44 (t, 2 H),

31 4.22 (m, 2H), 7.48 (m, 4H), 7.84 (d, H), 7.96 (d, H), 8.14 (d, H); P NMR (D2O): δ = -

+ 27.8 ppm (s); HRMS (ESI): m/z calcd for C14H15O4PNa (Sodium salt): 301.0606 [M+H] , found: 301.0609

Synthesis of 4-methoxyphenethylacetylphosphonate (2.17) ) A flame-dried flask, cooled under argon was charged with 2-(4-methoxyphenyl) ethanol (0.79 g, 5.2 mmol) dissolved in 17 mL of an anhydrous mixture of 1:1 diethyl ether and tetrahydrofuran. The flask was cooled to -40 oC and Hünig’s base (1.0 mL, 6.0 mmol) was added followed by phosphorus trichloride (0.15 mL, 1.7 mmol). After 60 minutes, the

31 reaction progress was assessed via P NMR, and was considered complete as PCl3 was consumed while tris(4-methoxyphenethyl)phosphite was observed (δ = 112 ppm). The crude reaction mixture was washed with cold 1M KOH and the organic layer was dried over sodium sulfate. Volatiles were removed in vacuo to yield 0.75 g of the phosphite, which was carried forward without further purification.

The phosphite was dissolved in 15 mL of dry methylene chloride and added to a flask charged with acetyl chloride (0.16 mL, 2.3 mmol). After 60 minutes, Reaction progress was assessed via 31P NMR and was deemed complete as determined by the formation of di(4-methoxyphenethyl)acetylphosphonate (δ = -29 ppm) and the consumption of the phosphite. The volatiles were removed in vacuo and the crude di(4- methoxyphenethyl)acetylphosphonate was dissolved in dry acetonitrile (2.5 mL) and 90

subjected to lithium bromide (0.21 g, 2.4 mmol) at 50 oC for 3 hours. The product did not precipitate from solution. The reaction mixture was diluted with water and washed with methylene chloride. The aqueous layer was washed with hexanes and then frozen and lyophilized to yield 0.073 g (16% yield over 3 steps) of 4-methoxyphenethyl

1 acetylphosphonate as the lithium salt (white powder). H NMR (D2O): δ = 2.20 (d, 2H),

31 2.87 (t, 2H), 3.81 (s, 3H), 4.05 (m, 2 H), 6.95 (d, 2H), 7.24 (d, 2H); P NMR (D2O): δ = -

+ 27.8 ppm (s); HRMS (ESI): m/z calcd for C11H15O5PNa (sodium salt): 281.0558 [M+H] , found: 281.0555.

Synthesis of MAP oxime (2.22). The synthesis of 2.22 has been reported previously, and product matched published parameters for the E-conformation [31].

Briefly, methylacetylphosphonate (0.2 g, 1.4 mmol) was suspended in 7 mL of methanol.

Ammonium hydroxide (HCl salt, 0.16 g, 2.3 mmol) was added, followed by addition of sodium acetate (0.14 g, 1.7 mmol). Approximately 50 drops of water helped the solubility of the reaction mixture. The reaction was complete with in 10 minutes, as determined by

31P NMR. Methanol and acetate were removed in vacuo and the remaining aqueous solution was frozen and lyophilized to yield 0.25 g of the white solid powder, which is a

1 quantitative yield with some excess acetate. H NMR (D2O): δ = 1.93 (d, 3H), 3.52 (d,

31 3H); P NMR (D2O): δ = -16.7 ppm (s); MS (ESI, negative mode): m/z calcd for

+ C3H7O4NP: 152.01 [M+H] , found: 151.95.

Synthesis of methyl chloromethylphosphonate (2.24) J. Kipchirchir Bitok synthesized dimethyl (chloromethyl)phosphonate as the starting material, which he prepared from (chloromethyl) phosphonic dichloride and methanol. This phosphonate

(0.13 g, 0.8 mmol) was dissolved in anhydrous acetonitrile (0.75 mL) and subjected to lithium bromide (0.092g, 1.0 mmol) at 75 oC overnight. The product precipitated from solution and was filtered and washed with cold acetonitrile and diethyl ether. The isolated methyl (chloromethyl)phosphonate was 0.78 g of a white solid (65% yield). 1H NMR

31 (D2O): δ = 3.42 (d, 2H), 3.54 (d, 3H); P NMR (D2O): δ = -8.62 ppm (s); HRMS (ESI):

+ m/z calcd for C2H7O3PCl: 144.9821 [M+H] , found: 144.9823.

NMR data Note: all 31P NMR are referenced to TPPO (δ = 0 ppm) as an external reference.

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21. Fang, M., et al., Succinylphosphonate esters are competitive inhibitors of MenD that show active-site discrimination between homologous alpha-ketoglutarate- decarboxylating enzymes. Biochemistry, 2010. 49(12): p. 2672-9.

22. Bénech, J.M., D. El Manouni, and Y. Leroux, NOUVELLE MÉTHODE DE PRÉPARATION D'ESTERS HYDROXY BISPHOSPHONIQUES SYMÉTRIQUES. Phosphorus, Sulfur, and Silicon and the Related Elements, 1996. 113(1-4): p. 295- 298.

23. Maeda, H., K. Takahashi, and H. Ohmori, Reactions of acyl tributylphosphonium chlorides and dialkyl acylphosphonates with Grignard and organolithium reagents. Tetrahedron, 1998. 54(40): p. 12233-12242.

24. Benech, J., et al., SYNTHESIS OF NEW α-KETOPHOSPHONATES. Phosphorus, Sulfur, and Silicon and the Related Elements, 1997. 123(1): p. 377-383.

25. Schultz, C., Prodrugs of biologically active phosphate esters. Bioorganic & Medicinal Chemistry, 2003. 11(6): p. 885-898.

26. Saady, M., L. Lebeau, and C. Mioskowski, First Use of Benzyl Phosphites in the Michaelis-Arbuzov Reaction synthesis of mono-, Di-, and triphosphate analogs. Helvetica Chimica Acta, 1995. 78(3): p. 670-678.

27. Katzhendler, J., et al., Acylphosphonate hemiketals-formation rate and equilibrium. The electron-withdrawing effect of dimethoxyphosphinyl group. Journal of the Chemical Society, Perkin Transactions 2, 1997(2): p. 341-350.

28. Altincicek, B., et al., Tools for discovery of inhibitors of the 1-deoxy-D-xylulose 5- phosphate (DXP) synthase and DXP reductoisomerase: an approach with enzymes from the pathogenic bacterium Pseudomonas aeruginosa. FEMS Microbiology Letters, 2000. 190(2): p. 329-333.

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29. Arjunan, P., et al., A thiamin-bound, pre-decarboxylation reaction intermediate analogue in the pyruvate dehydrogenase E1 subunit induces large scale disorder- to-order transformations in the enzyme and reveals novel structural features in the covalently bound adduct. J Biol Chem, 2006. 281(22): p. 15296-303.

30. Nemeria, N.S., et al., Acetylphosphinate is the most potent mechanism-based substrate-like inhibitor of both the human and Escherichia coli pyruvate dehydrogenase components of the pyruvate dehydrogenase complex. Bioorg Chem, 2006. 34(6): p. 362-79.

31. Breuer, E., et al., [alpha]-Oxyiminophosphonates: chemical and physical properties. Reactions, theoretical calculations, and X-ray crystal structures of (E) and (Z)-dimethyl [alpha]-hydroxyiminobenzylphosphonates. Journal of the Chemical Society, Perkin Transactions 1, 1988(11): p. 3047-3057.

32. Gish, G., T. Smyth, and R. Kluger, Thiamin diphosphate catalysis. Mechanistic divergence as a probe of substrate activation of pyruvate decarboxylase. Journal of the American Chemical Society, 1988. 110(18): p. 6230-6234.

33. Bisswanger, H., Fluoropyruvate: A potent inhibitor of the bacterial and the mammalian pyruvate dehydrogenase complex. Biochemical and Biophysical Research Communications, 1980. 95(2): p. 513-519.

34. Xiang, S., et al., Crystal Structure of 1-Deoxy-d-xylulose 5-Phosphate Synthase, a Crucial Enzyme for Isoprenoids Biosynthesis. Journal of Biological Chemistry, 2007. 282(4): p. 2676-2682.

35. Patel, H., et al., Observation of Thiamin-Bound Intermediates and Microscopic Rate Constants for Their Interconversion on 1-Deoxy-d-xylulose 5-Phosphate Synthase: 600-Fold Rate Acceleration of Pyruvate Decarboxylation by d- Glyceraldehyde-3-phosphate. Journal of the American Chemical Society, 2012. 134(44): p. 18374-18379.

36. Strumilo, S., J. Czerniecki, and P. Dobrzyn, Regulatory effect of thiamin pyrophosphate on pig heart pyruvate dehydrogenase complex. Biochem Biophys Res Commun, 1999. 256(2): p. 341-5.

37. Sprenger, G.A., et al., Transketolase a of Escherichia coli K12. European Journal of Biochemistry, 1995. 230(2): p. 525-532.

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Chapter 3. Antimicrobial activity of acetylphosphonates

Introduction:

As new insights emerge about the distinctive characteristics of DXP synthase, so will new opportunities for the development of selective inhibitors. DXP synthase is a metabolic branch point in bacterial biosynthesis and therefore an attractive target for inhibitor design (Figure 3-1A). Evidence suggests that electrophilic acetylphosphonates interact with the nucleophilic thiamin diphosphate (ThDP) cofactor to generate a phosphonolactylthiamin diphosphate (PLThDP) which cannot de-carboxylate and therefore inhibits catalysis (Figure 3-1B). Our previous work has established that sterically demanding alkylacetylphosphonates, exemplified by butylacetylphosphonate

(BAP) and benzylacetylphosphonate (BnAP), are selective inhibitors of DXP synthase

(Chapter 2) [1, 2]. Conceivably, they do so by acting as unnatural bisubstrate inhibitors and taking advantage of the unique ternary complex of DXP synthase, which is distinctively accommodating towards these substrates.

Here, we explore the antimicrobial activity of the some of the acetylphosphonates introduced in Chapter 2. Jessica Mott Smith, Ph.D. and our collaborator Andrew

Koppisch, Ph.D. (Northern Arizona University) showed that butylacetylphosphonate

(BAP) shows weak antimicrobial activity against a wide variety of bacterial targets possessing the MEP pathway. Further, BAP inhibits growth of E. coli in a dose-dependent manner via a mechanism involving inhibition of DXP synthase, which is confirmed by downstream metabolite rescue and target overexpression [3]. Given that achieving cell permeability of bacterial enzyme inhibitors [4] is a major barrier to new antibiotic development, our preliminary findings in this respect are encouraging. 115

Building on these studies, we determined that evaluating the antimicrobial properties of additional long-chain alkylacetylphosphonates (APs, discussed in Chapter

2) against the Gram-negative bacteria E. coli reveals a trend of increasing cell permeability with increasing alkyl chain length, which is offset by efflux of APs bearing the longest alkyl chains. The primary cause of acetylphosphonate efflux appears to be mediated by the endogenous AcrAB-TolC multidrug resistance efflux pump. Overall, our results suggest that modification of acetylphosphonates significantly influences uptake and efflux.

This is evidence that additional medicinal chemistry efforts could lead to substantial improvement in the antimicrobial activity of this new inhibitor class. Finally, we show that

APs can act in concert with other drugs via our studies with the efflux pump inhibitors

(EPI) PAβN and MBX 2319, and the IspC inhibitor fosmidomycin, a promising lead for future studies in combination therapies [5].

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Figure 3-1. A) DXP is a key metabolic intermediate for isoprenoid production via the MEP pathway and vitamin B1 and B6 production in bacteria. B) Alkylacetylphosphonates (alkyl

AP) inhibit DXP synthase via formation of a phosphonolactyl thiamin diphosphate intermediate (PLThDP) which competes with pyruvate binding.

Results:

3.1 BAP exhibits weak antimicrobial activity. While MAP does not have antimicrobial properties [6] presumably as a consequence of poor cellular uptake, we

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wanted to evaluate if some of our larger acetylphosphonates could inhibit bacterial growth.

As a starting point, Jessica Mott Smith, Ph.D. first evaluated the antimicrobial activity of

BAP against a common laboratory strain for susceptibility testing, Escherichia coli ATCC strain MG 1655 via the macrodilution method [7, 8]. She observed that BAP inhibits 85% of E. coli growth (MIC85) at a concentration of ~1,000 μg/mL in cation-adjusted Müeller

Hinton Broth, but a significantly lower concentration (122 μg/mL) in M9 minimal media achieved the same effect. Our collaborator, Andrew Koppisch, Ph.D. (Northern Arizona

University) further evaluated the compound against a larger panel of clinically isolated pathogens. He observed an MIC for BAP of 4,000 μg/mL for Salmonella enterica serovar

Typhimurium, Pseudomonas aeruginosa and Micrococcus sp., and 1,000 μg/mL against

Bacillus antracis Sterne. The summary of these results is shown in Table 3-1.

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MIC Organism (µg/mL) Bacillus anthracis Sterne 1 1000 Bacillus subtillis > 2000 Bacillus thuringiensis HD34 1 > 4000 Enterobacter cloacae > 4000 Escherichia coli 1000 Klebsiella oxytoca > 4000 Klebsiella pneumonia > 4000 Micrococcus sp. 2000 Mycobacterium smegmatis > 2000 Pseudomonas aeruginosa 4000 Pseudomonas fluorescens > 4000 Rhizobium radiobacter 1 > 4000 Salmonella typhimurium 4000 Staphylococcus aureus2 > 4000

Table 3-1. The antimicrobial properties of BAP against a panel of bacterial pathogens.

1The slower growth of these species required fractional growth determination with slightly longer incubations (22-24 hrs) at 37 °C. 2 Does not use the MEP pathway. [3]

3.2 BAP inhibition of E. coli growth is rescued by downstream metabolites, and target overexpression. Although weak, the antimicrobial activity of BAP against E. coli allowed us to evaluate the specific cause of growth inhibition. We confirmed that a target of BAP was intracellular DXP synthase by rescuing with downstream metabolites

(1-deoxy-D-xylulose (DX) and thiamine), as well as by overexpressing the proposed target.

Although DX requires an intracellular phosphorylation to become DXP, previous studies utilizing isotopically labelled DX have shown it enters bacterial cells and undergoes intracellular phosphorylation [9, 10]. E. coli growth is suppressed by over 70% by 0.66 mM BAP, but in the presence of 125 μM DX, the growth is only suppressed by 40%.

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We were also able to observe near complete rescue of the growth inhibitory effects of low concentrations of BAP on E. coli in M9 minimal media (which does not contain any thiamin) by supplementing the media with ≥ 62.5 μM thiamin; however, the rescue was not as pronounced at higher concentrations of BAP, suggesting that inhibition of DXP synthase could interfere with thiamin uptake at higher concentrations, or other non-specific effects of BAP at high concentration. Under these conditions, 0.165 mM BAP causes

~80% reduction in E. coli growth at 16 hours, but there is almost complete rescue by 16 hours when the media is supplemented with 125 μM thiamin, and this concentration of thiamin does not significantly alter or improve the growth curve of E. coli in the absence of BAP [3]. Other intermediates of the MEP pathway and pyridoxal pathway were not able to rescue the antimicrobial effects of BAP. This could be due to a lack of permeability or uptake of these compounds, or the absence of intracellular enzymes to activate the precursors into bioactive species.

The most compelling evidence that BAP inhibits E. coli growth by DXP synthase inhibition is the observed protective effects on cells overexpressing DXP synthase.

Previous work by Eric Brown and colleagues has shown that overexpression of MEP pathway enzymes can overcome inhibition and rescue growth [11]. Thus, four strains of

BL21 E. coli cells were treated with varying concentration of BAP. The BL21 parent strain, a strain harboring the empty pET37b expression vector, a strain transformed with the pET37b/dxs plasmid and a strain with the pET37b/dxsE70A plasmid which overexpresses a catalytically incompetent form of DXP synthase [12]. As summarized in

Figure 3-2, only the BL21 strain containing the catalytically active form of DXP synthase is able to protect E. coli from growth inhibitory activity of low levels of BAP.

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Figure 3-2. Overexpression of DXP synthase protects from the antimicrobial effects of

BAP. Four BL21 strains were exposed to increasing concentrations of BAP. Only the catalytically competent form of DXP synthase protected against the antimicrobial effects of BAP. [3]

Finally, the observed weak antibacterial activity of BAP is a little disappointing; thus, we evaluated the potential for synergy of BAP in combination with inhibitors of downstream pathway enzymes. The synergy between different classes of compounds that target enzymes in the same pathway is well established, such as with the antifolate agents and sulfa antibiotics [13]. Synergy between inhibitors has also been shown within the MEP pathway itself: the IspC inhibitor, fosmidomycin, synergizes with the bisphosphonate inhibitors of the downstream isoprenoid biosynthetic enzyme, farnesyl diphosphate synthase [14], and conditional knockouts of the MEP pathway enzyme IspF lead to synthetic lethality in combination with fosmidomycin and cell wall biosynthesis inhibitors

[11]. The results of our synergy experiments between BAP and antimicrobial agents are

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summarized in Table 3-2. BAP in combination with fosmidomycin, the inhibitor of the subsequent enzyme in the MEP pathway (IspC), has an FIC index of 0.25, predicting a strongly synergistic relationship. A combination of BAP and ampicillin has an FIC index of 0.31, also suggestive of synergy, but deemed additive upon closer inspection of the checkerboard assay results [3]. Finally, the effects of BAP combined with tetracycline is additive (FIC ~0.5). The discovery of synergy between BAP and fosmidomcyin is encouraging and supports continued exploration of inhibitors DXP synthase as a potential members of a combination drug therapy.

organism MIC BAP (µg/mL) combination FIC index BAP-fosmidomycin 0.25 Escherichia coli MG1655 1000 BAP-ampicillin 0.31 BAP-tetracycline 0.53 Table 3-2 BAP synergizes with fosmidomycin and ampicillin against E. coli [3]

3.3 Other acetylphosphonates inhibit E. coli growth in a dose-dependent manner. Given the antimicrobial activity of BAP, we investigated if the other acetylphosphonates (synthesis and in vitro evaluation described in Chapter 2) also display antimicrobial activity. Despite the observation that acetylphosphonate analogs tested here exhibit comparable low micromolar inhibitory activity against DXP synthase (Figure 3-

3A), only BAP (3.3), PentAP (3.4) and HexAP (3.5), and BnAP (3.7) exert a dose- dependent delay in the growth of E. coli (Figure 3-3B); OctAP (3.6) is inactive against wild-type K-12 E. coli up to 5 mM (1210 ug/mL). In terms of minimum inhibitory

PentAP concentration (MIC90), PentAP (MIC = 1.25 mM) displays somewhat more potent antimicrobial activity compared to BAP (MICBAP = 2.5 mM), whereas BnAP exerts the most potent antimicrobial activity in this series (MICBnAP = 0.63 mM), and HexAP displays 122

markedly less potent antimicrobial effects (MICHexAP = 5mM) against E. coli compared to

BnAP, BAP or PentAP.

In addition to OctAP, the shortest acetylphosphonates, MAP (3.1) and EAP (3.2) also lack antimicrobial activity up to 5 mM. While unexpected, this data argues against general cell disruption by amphipathic molecules as a mode of action of this class of compounds, as conceivably, at 5 mM we would observe general cell disruption from all members of this class if APs also act by that non-specific mechanism. In fact, we went on to confirm that the intracellular targets of the active members of this class are shown to possess a mode of action involving inhibition of DXP synthase.

Figure 3-3. A) In vitro inhibition of E. coli DXP synthase and B) antimicrobial effects of acetylphosphonates against E. coli (MG1655) grown in CAMHB. OctAP (○) is inactive while BnAP (■), PentAP (●), BAP (♦) and HexAP (▲) exert dose-dependent inhibition of

E. coli. Concentrations of acetylphosphonate were tested from 0.08 to 5 mM and fractional growth was determined at 16 hours as compared to no drug control.

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3.4 DXP synthase overexpression protects E. coli from acetylphosphonate treatment. One possibility for the differences in antimicrobial activity of the acetylphosphonates observed in Figure 3-3B is that some of the compounds have alternative targets. We employed our discovery that DXP synthase over expression is protective against BAP treatment in E. coli [3] as a tool to determine if the acetylphosphonates share this intracellular target.

As expected, DXP synthase over expression is protective for BL21 (DE3) cells treated with BnAP, PentAP, HexAP and to a lesser degree, OctAP (Figure 3-4).

Additionally, it is only the active form of DXP synthase that is protective. Neither an empty expression vector (pET37b), nor a strain overexpressing a catalytically inactive variant of the enzyme (dxs–E370A) [12] protects E. coli from the acetylphosphonates. The data suggest that DXP synthase is a target of the acetylphosphonates; however at high AP concentration complete rescue is not observe, suggesting other targets cannot be ruled out.

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Figure 3-4. Increasing intracellular DXP synthase levels results in partial rescue of E. coli growth in the presence of alkylacetylphosphonates. Rescue is observed in the presence of

BnAP (0.16 mM), PentAP (0.31 mM), HexAP (0.63 mM), and to some extent OctAP (5 mM).

3.5 Bacterial resistance to alkylacetylphosphonates includes drug efflux by E. coli. As summarized in Figure 3-3, the alkylacetylphosphonates tested as antimicrobial agents (3.3 – 3.7) were all comparably low micromolar inhibitors of DXP synthase in vitro, although they seem to show weak (millimolar) and diverse potencies as antimicrobial agents. We have also shown that compounds with antimicrobial activity against E. coli share a mode of action that includes inhibition of cellular DXP synthase; thus, the difference in their potencies may be due to different intracellular concentrations. This could be the result of differences in efficiency of uptake via porins or efflux via multi-drug resistance (MDR) pumps. 125

Initially, we sought to determine the mode of entry of the acetylphosphonate inhibitors. The difficulty in designing antibiotics that enter Gram-negative cells, and the negatively charged phosphonate argue against passive diffusion as the likely mode of entry.

This conclusion is supported by the observation that the shortest acetylphosphonates (MAP and EAP), as well as the longest acetylphosphonate (OctAP) are not active as antimicrobial agents (Figure 3-2b), although the other compounds in this class are. It is more likely that these molecules are actively transported into the cell. One possible point of entry is the E. coli glycerol 3-phosphate transporter (glpT), known to be responsible for the active uptake of the negatively charged phosphonate antibiotic, fosmidomycin [15, 16]. However,

Jessica Mott Smith showed that deletion of this gene in E. coli does not provide protection from BAP treatment, whereas deletion of this gene does protect E. coli from fosmidomycin treatment, as expected [3]. Further experiments are needed to determine the mode of entry.

It is possible that the general antibiotic uptake porins, OmpF and OmpC [17, 18] are responsible for the entry of these compounds into Gram-negative cells.

Next, we sought to explore differences in efficiency of efflux by studying the susceptibility of E. coli strains either possessing or deficient in the canonical MDR pump, the tripartite AcrAB-TolC transporter [19], a member of the ubiquitous Gram- resistance- nodulation-division (RND) family. It is composed of AcrB, a periplasmic membrane fusion protein (MFP), AcrA, a periplasmic adapter protein, and the porin TolC, an outer membrane factor (OMF). Recent studies have shown that substrates are selected by asymmetric trimer, AcrB, from the periplasmic space or the outer leaflet of the inner membrane and pumped via a proton motive-coupled peristaltic action [20, 21] towards an access channel to TolC to be extruded from the cell [22, 23]. While multiple component

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combinations can be assembled, TolC is the outer OMF common to almost all E. coli RND pumps responsible for resistance to endogenous toxins and clinically used agents [24].

The susceptibilities to acetylphosphonates of a genetic variant of E. coli lacking the tolC gene (ΔtolC) and the corresponding parent E. coli strain, BW25113, were evaluated in CAMHB medium. Again, BnAP displays the most potent antimicrobial activity against the parent strain, BW25113 (MICBnAP = 1.25 mM, Figure 3-5), followed by PentAP

(MICPentAP = 2.5 mM), BAP (MICBAP = 5 mM) and HexAP (MICHexAP = 5 mM); OctAP is inactive up to 5 mM.

Figure 3-5. Antimicrobial activity of acetylphosphonates against E. coli BW25113.

Standardized cell cultures (approximately ~105 CFU mL-1) were exposed to increasing concentrations of acetylphosphonate. Fractional growth was determined at 16 hours as a percentage of the growth of no drug control. BAP (), PentAP () and HexAP () were varied from 0.31 to 5 mM. OctAP () was varied from 0.63 to 5 mM, and BnAP () was varied from 0.16 mM to 5 mM. Each data point is the average of 3 independent experiments and the error bars are standard error.

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E. coli cells lacking the tolC gene (ΔtolC) indeed exhibit increased susceptibility to

BAP, PentAP, HexAP, OctAP and BnAP relative to the parent strain (Figure 3-6 A – E), illustrated by the shift in dose response curves and indicating these acetylphosphonates can act as substrates for efflux via the AcrAB-TolC transporter. However, excluding OctAP which displays a substantially lower MIC against the ΔtolC strain relative to wild type E. coli, a significant decrease in the MIC was not observed for the other compounds tested.

Methylacetylphosphonate (MAP) and ethylacetylphosphonate (EAP) are inactive against both E. coli BW25113 and the tolC variant at concentrations up to 5 mM (Figure 3-7).

Figure 3-6. Acetylphosphonates are substrates for efflux via the AcrAB-TolC transporter.

E. coli BW25113 (●) and the ΔtolC E. coli variant (▲) were treated with A) BAP, B)

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PentAP, C) HexAP, D) OctAP and E) BnAP. F) Fractional growth of E. coli BW25113

(black bars) and the ΔtolC E. coli variant (gray bars) in the presence of 1.25 mM alkylacetylphosphonate.

Figure 3-7. MAP and EAP are inactive against E. coli BW25113 and the ΔtolC variant.

Standardized innocula of E. coli BW25113 (approximately ~105 CFU mL-1) were treated with varying concentrations of either MAP (blue) or EAP (orange) from 0.63 to 5 mM for

16 hours, and fractional growth was determined by comparison to a no drug control. The

ΔtolC variant of BW25113 was also treated with varying concentrations of MAP (grey) or

EAP (yellow) from 0.63 to 5 mM for 16 hour, and fractional growth was determined by comparison to no drug controls. Experiments were performed in duplicate, and error bars represent standard deviation.

Taken together, the data suggest that small acetylphosphonates MAP and EAP are not cell permeable, and increasing alkyl chain length (BAP and PentAP) appears to enhance permeability and antimicrobial activity somewhat. However, these antimicrobial 129

effects are increasingly offset by efflux (HexAP and OctAP) which lowers the intracellular acetylphosphonate concentration (Figure 3-6F). The most dramatic effect of deleting the transporter is observed with OctAP (Figure 3-6D), supporting the idea that increasing hydrophobicity of the acetylphosphonates seems to increase susceptibility to efflux.

Nevertheless, deletion of the tolC component of the AcrAB-TolC transporter fails to restore antimicrobial activity to micromolar levels observed for these compounds in biochemical inhibition experiments (Figure 3-2), indicating that cellular uptake is inefficient or additional mechanisms of acetylphosphonate efflux exist. It is also possible that deletion of TolC negatively impacts import of APs via altering the ratio of two key E. coli porins,

OmpC and OmpF [44], therefore more experiments are required to understand the complex nature of cellular uptake and efflux and how it relates to the antimicrobial activity of APs.

We next evaluated whether we could recapitulate these results using a pharmacological tool, an efflux-pump inhibitor (EPI).

3.6 PAβN enhances activity of BAP and BnAP, but not MAP or OctAP.

Alkylacetylphosphonates appear to act as substrates for the AcrAB-TolC transporter. Thus, as a starting point we have evaluated the antimicrobial activities of three acetylphosphonates with varied alkyl chain lengths, MAP, BAP and OctAP, and the most potent acetylphosphonate, BnAP, in combination with two efflux pump inhibitors (EPI), phenylalanine-arginine β-naphthylamide (PAβN) and MBX 2319. PAβN is a well-studied

EPI and was originally found to increase P. aeruginosa susceptibility to efflux-prone fluoroquinolones [25]. Subsequently, PAβN has been shown to decrease bacterial (i.e. E. coli, Acinetobacter, Klebsiella and Psuedomonas) resistance to multiple classes of 130

antibiotics (i.e. macrolides, aminoglycosides, and β-lactams) [26-28]. However, recent reports suggest that caution should be used in interpreting results from PAβN co-treatment experiments due to the additional function of PAN as an outer-membrane permeabilizing agent [29, 30].

Another recently discovered EPI, MBX 2319 [31] improves potency of multiple antibiotic classes by inhibiting the AcrAB-TolC transporter in E. coli. Notably, MBX 2319 does not exhibit the same membrane-permeabilizing properties of PAβN. The recognition subunit of the pump, AcrB, is proposed to have multiple substrate binding sites [32] and

PAN and MBX 2319 are proposed to act by competing with antibiotics at distinct sites of the AcrB recognition subunit [31, 33] Here, addition of PAβN at a concentration that itself does not influence E. coli growth (34g/mL) markedly increases E. coli susceptibility to

BAP (0.63 mM), and to a greater extent than deletion of the tolC gene, as illustrated in

Figure 3-8). The activity of BnAP is also potentiated by PAβN but not MBX 2319. In this case, the increased susceptibility caused by co-administering PAβN is similar to the levels achieved in the absence of tolC. In contrast, addition of MBX 2319 at its upper limit of solubility (10 g/mL) has minimal effects on the antimicrobial activity of BAP or BnAP, suggesting they may bind at a distinct site from MBX 2319 and are instead competitive with PAN.

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Figure 3-8. PAβN, but not MBX2319, potentiates the antimicrobial activity of BAP (0.63 mM) and BnAP (0.31 mM) against E. coli. Activities of the smaller acetylphosphonate

MAP (5 mM) and longer analog OctAP (5 mM) are not potentiated by PAβN or MBX

2319.

Notably, neither EPI potentiates the activity of 5 mM OctAP (Figure 3-8), although at this high concentration of OctAP growth of the ΔtolC mutant strain is inhibited by > 90%.

This observation suggests a different mechanism of OctAP efflux compared to BAP and

BnAP. The longer hydrophobic chain of OctAP could promote binding to a site on AcrB that is distinct from the PAN binding site putatively shared by BAP and BnAP. It is also possible that a different MFP is responsible for the binding and extrusion of OctAP to TolC.

Given recent reports on the outer membrane permeabilizing effects of PAN [25, 29,

30] we considered the possibility that potentiation of BAP and BnAP antimicrobial activity

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by PAN could result in part from increased uptake of BAP and BnAP through the permeabilized outer membrane. We reasoned that if outer membrane permeabilization plays a key role in the potentiating effects of PAN on BAP and BnAP activity, a similar effect could be observed by combining PAN with the structurally related alkylacetylphosphonate MAP, which does not appear to be cell permeable and is inactive against E. coli. Interestingly, PAN does not enhance the antimicrobial activity of MAP

(Figure 3-8) suggesting this AP cannot traverse a destabilized outer membrane in the presence of PAN. Taken together with the observation that OctAP is also inactive, even when combined with PAN, these results suggest the enhanced antimicrobial activity of

BAP and BnAP in the presence of PAN is due primarily to inhibition of the AcrB component of the multi-drug efflux pump.

3.7 Growth inhibition caused by fosmidomycin is significantly enhanced when administered in combination with APs. Continuing our studies toward identifying synergistic combinations of APs with antimicrobials we investigated whether a low, non- toxic level of each acetylphosphonate could potentiate the antimicrobial activity of fosmidomycin, as observed for BAP [3]. For these experiments, we used the BW25113 cell line of E. coli under conditions of varying concentrations of fosmidomycin in the presence or absence of a sub-lethal concentration of acetylphosphonate. Under these conditions, the MIC for fosmidomycin is approximately 3 μg/mL. We then screened the susceptibility of E. coli to fosmidomycin concentrations below and up to the MIC (0.01 to

3 μg/mL) in the presence or absence of acetylphosphonates at a sub-inhibitory

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concentration (Figure 3-9). BAP and PentAP were tested at 0.31 mM (Figure 3-9A and

3-9B, respectively) and clearly improve the activity of fosmidomycin (MIC = 0.19 μg/mL).

The BAP data supports our earlier finding that BAP and fosmidomycin are a synergistic combination against E. coli.

Figure 3-9. Potentiation of fosmidomycin activity by acetylphosphonates. BW25113 cells were grown in CAMHB media for 16 hours with varying concentrations of fosmidomycin

(0.01 to 3 μg/mL), either alone or in addition to A) 0.31 mM BAP, B) 0.31 mM PentAP,

C) 0.31 mM HexAP or D) 0.16 mM BnAP. Data is reported as the optical density (OD600) of the cell culture at 16 hours. BAP and PentAP experiments were performed in triplicate.

HexAP and BnAP experiments were performed in quadruplicate. Error bars represent standard error. 134

It is interesting that the trend of improved activity is not as profound for HexAP

(0.31 mM) and BnAP (0.16 mM) (Figure 3-9C and 3-9D, respectively). BnAP was tested at a lower concentration due to the fact that it is inherently more toxic to the BW25113 cell line. Also, HexAP could potentially have been tested at a higher concentration, as it is not as toxic as BAP and PentAP against BW25113 (Figure 3-6). The differences in potency as a potentiating agent for fosmidomycin could also be attributed to the high degree of variability in experiments of fosmidomycin susceptibility.

3.8 OctAP is more toxic to Plasmodium falciparum than BAP. In addition to bacteria, the apicocomplexan parasites including the causative agent of malaria, P. falciparum, also utilize the MEP pathway to generate essential precursors for isoprenoids.

The apicoplast organelle contains the enzymes of the pathway and evidence suggests that isoprenoid production is an essential role of this organelle [34, 35]. Important cellular processes, such as the synthesis of ubiqunone for the electron transport chain and the prenylation of malarial proteins are all dependent on isoprenoid production [36] and thus the MEP pathway makes an attractive target, and inhibition of this pathway is thought to be a road to new anti-malarial drugs, especially in combination therapies [37]. Validation of this concept is the IspC inhibitor fosmidomycin which is currently undergoing clinical trials for treatment of uncomplicated malarial infection in combination with piperaquine

[38].

Our collaborators, Dr. Theresa Shapiro and Elizabeth Nenortas evaluated the antimalarial properties of the three acetylphosphonates, BAP, OctAP and BnAP against the

P. falciparum NF54 strain. BAP kills approximately 97% of parasites at the maximum concentration tested, 10 mM (Figure 3-10A) and has an EC50 = 4.6 mM. BnAP kills

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approximately 10% of parasites at 1 mM (Figure 3-10 B), but solubility in media precluded experiments at a higher concentration. OctAP kills 100% of parasites at approximately 2.5 mM and has an EC50 = 5.6 mM.

Figure 3-10. OctAP is more toxic than BAP to P. falciparum. P. falciparum NF54 strain was exposed to varying concentrations of acetylphosphonates in Albumax CM and percent of parasites killed was determined at 72 h. A) BAP (20 – 10,000 μM) has an EC50 = 5.6 mM. B) BnAP (blue, 0.5 – 1000 μM) did not saturate, but OctAP (red, 10 – 5000 μM) has an EC50 = 4.6 mM. Credit: Elizabeth Nenortas.

Next, Elizabeth examined the cytotoxicity of BAP and OctAP against the mammalian L1210 cell lines (Mus musculus lymphocytic leukemia cells) to determine if the agents were acting as general cell-disruptors. The highest concentration of BAP tested,

10 mM caused approximately 35 % cell death. 10 mM OctAP causes 100% cell death of the mammalian cells; OctAP has an EC50 = 1.7 mM against this mammalian cell line. This data suggests that there is a small therapeutic window for BAP in which it is effective against the malarial parasite, but not generally toxic to a mammalian cell. Unfortunately,

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the results are discouraging for OctAP as it seems that it is comparably toxic to mammalian cells. The toxicity of the amphipathic OctAP is not overly surprising as the long aliphatic chain and charged phosphonate make the molecule seem especially detergent-like. It is possible that the shorter chain length of BAP limits some of this toxicity against mammalian cells.

Discussion

Each of the seven enzymes in the MEP pathway to isoprenoids represents a potential point of intervention in the development of new antimicrobial agents. Inhibition of DXP synthase could negatively impact multiple metabolic pathways to produce potent antimicrobial effects, highlighting this enzymatic step as an attractive drug target.

Alkylacetylphosphonate inhibitors bearing C4, C5 and C6 alkyl chains exert a dose- dependent delay in the growth of wild-type E. coli by a mechanism that appears to involve, in part, inhibition of DXP synthase, while the smaller inhibitors MAP and EAP and the longest inhibitor OctAP are inactive. The observed lack of antimicrobial activity by MAP is consistent with previous studies to elucidate the mechanism of action of the natural product dehydrophos [6]. This phosphonotripeptide precursor to MAP enters bacterial cells by the action of oligopeptide permeases and is subsequently cleaved intracellularly by peptidases to release MAP. Our results suggest increasing alkyl chain length on the acetylphosphonate scaffold can increase cell permeability of this compound class; however, increased cellular uptake appears to be offset by efflux to reduce the intracellular concentration of inhibitor. The aromatic acetylphosphonate BnAP displays the most potent

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antimicrobial activity in this inhibitor series, and also appears to be a substrate for efflux.

Its more potent activity could suggest it enters E. coli cells more efficiently.

Future work could look at synergy between the acetylphosphonates and an inhibitor of a later stage of the thiamin biosynthetic machinery. However, until recently, little was known about potent inhibitors of these enzymes [39] such as antimetabolites, bacimethrin

[40] and 4-amino-2-trifluoromethyl-5-hydroxymethylpyrimidine (CF3-HMP), as well as the natural product rugulactone [41]. A recent screen targeting the Mycobacterium tuberculosis thiamine phosphate synthase has shed light on leads with impressive anti- mycobacterial activity [42]. These could provide promising starts for further synergy or combination therapy experiments with the acetylphosphonates.

Not surprisingly, overexpression of active DXP synthase in E. coli also results in pronounced rescue of growth inhibitory activity by BAP, and to a lesser extent, AmAP,

HexAP and BnAP. The observation that rescue is less pronounced increasing acetylphosphonate concentration is consistent with DXP synthase inhibition coupled with either a second target or other nonspecific antimicrobial effects of the acetylphosphonates at high concentrations.

Despite the low micromolar inhibitory activity of alkylacetylphosphonates against

DXP synthase, antimicrobial activity is observed at high micromolar/low millimolar concentrations. There are several possible reasons for this observation. First, we have performed antimicrobial assays in the typically-used complex CAMHB medium, which contains vitamins, including thiamin, and protein from beef infusion. Given the pronounced rescue of the antimicrobial effects of BAP by thiamin supplementation to minimal medium, we presume that the low activity of APs against E. coli in CAMHB

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medium is attributed, in part, to rescue by thiamin. AP binding to protein, especially with

APs bearing longer alkyl chains, could also account for weak antimicrobial activity.

Moving forward, these are questions that can be pursued. It is also possible that high intracellular concentrations of inhibitors in this series are not achieved. Low cell permeability of alkylacetylphosphonates and their susceptibility to efflux conjointly contribute to this problem. Jessica Mott Smith determined that BAP is not a substrate for the glycerol 3-phosphate transporter (glpT) known to actively transport fosmidomycin into bacterial cells [3, 15, 16]. More detailed studies are required to understand the mechanisms of AP entry into bacterial cells, to optimize this inhibitor scaffold. Toward understanding the mechanism(s) of inhibitor efflux, we have demonstrated that alkylacetylphosphonates are extruded via the AcrAB-TolC transporter. Further, we propose on the basis of experiments combining BAP with efflux pump inhibitors PAN and MBX 2319, that BAP may bind the AcrB subunit in a binding site that is common to PAN. PAN appears to enhance the antimicrobial activity of BAP to a greater extent than deletion of TolC. Our results indicate the outer membrane destabilizing effect of PAN does not apparently enhance permeability of MAP or the more lipophilic OctAP in E. coli, suggesting enhanced permeability does not contribute significantly to the potentiating effects of PAN on BAP.

It is possible that TolC is not the only OMF at play in the efflux of BAP. Three other OMFs are produced in E. coli, including yohG, yjcP and ylcB [43], and further studies are required to determine their roles in efflux of alkylacetylphosphonates. Additionally, it is conceivable that BAP synergizes with PAβN by an unknown mechanism. Studies could be pursued to explore a potential synergistic relationship between PAβN and inhibitors of DXP synthase or other steps of the MEP pathway (such as fosmidomycin). Future experiments could also

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explore the relationship between the acetylphosphonates and cell membrane disrupting agents, such as colistin. Our observations could influence the design of future acetylphosphonates to resist efflux via AcrB as well as inform potential EPI candidates to co-administer with the acetylphosphonate inhibitors. The results reported here are of significance to the development of novel, much needed antimicrobial drugs to combat drug resistant human pathogens.

Experimental

Antimicrobial susceptibility studies. Using aseptic techniques, 3 isolated colonies were selected from a plate containing ATCC MG 1655 E. coli K-12 and inoculated into 5 mL of cation-adjusted Mueller Hinton Broth (CAMHB; Sigma, St. Louis, MO, USA) at 37oC. The inoculated culture was incubated with shaking until the turbidity matched a

MacFarland standard of 0.5 (~OD600 = 0.1) [7, 8] Colony counts were checked after incubation on CAMHB agar plates for 16 hours at 37oC to confirm consistency between experiments. The standardized inoculums (MacFarland = 0.5) contained approximately 1-

2 x 108 CFU/mL and were diluted 1:100 into sterile CAMHB to yield the experimental inoculum which was mixed 1:1 with CAMHB containing the antimicrobial agent at double the desired concentration. The final concentration in a well was therefore approximately

~105 CFU/mL and each well contained a volume of 200 μL. The 96-well plates were incubated at 37 oC for 16 hours with intermittent shaking. Fractional growth was determined at 16 hours as a ratio to the no drug control. The acetylphosphonates were tested from a concentration of 80 μM to 5 mM.

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E. coli growth rescue studies. Adapted from previously published work [3], DXP synthase overexpression rescue experiments were carried out CAMHB, and the following cell types were treated with 0-5.0 mM acetylphosphonate: BL21 (no vector), pET37b-

BL21, wt E. coli dxs¬pET37b/BL21, and E370A E. coli dxs-pET37b/BL21. Experiments were performed in triplicate. For rescue of growth by BnAP, PentAP, HexAP and OctAP, an overnight starter culture of the appropriate cell line was started by inoculating sterile

CAMHB with 1 colony. After growing to saturation, the culture was diluted 1:100 into fresh CAMHB and grown to an OD600 ~0.45 (approximately 500 CFU mL-1). Cultures were then diluted 1:1 with acetylphosphonate at the indicated concentration, and cultures were grown for 16 hours. Fractional growth (measured at 16 h) was determined relative to the no drug control in each case.

Antimicrobial susceptibility of E. coli BW25113-tolC to alkylacetylphosphonates. Experiments were performed following the same procedure as described above, but starter colonies were picked from plates of either E. coli BW25113

(parent) or BW25113-ΔtolC (JW5503-1). Fractional growth was determined at 16 hours as a ratio to the no drug control in that cell line

Potentiation of alkylacetylphosphonates by efflux pump inhibitors.

Experiments were performed following conditions described above. Starter cultures of

BW25113 were mixed 2:1:1 with sterile CAMHB containing 4 acetylphosphonate (20 mM MAP and OctAP, 2.5 mM BAP) and 4 EPI of interest (100 μM MBX 2319, 308 μM

PAβN), and the final well volume contained 200 μL. Fractional growth was determined at

16 hours as a fraction of growth compared to the control BW25113 containing the appropriate EPI and no acetylphosphonate.

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CURRICULUM VITAE FOR Ph.D. CANDIDATES

The Johns Hopkins University Krieger School of Arts & Sciences

Ryan James Vierling October 24, 2014

Educational History:

Ph.D. expected 2014 Chemistry-Biology Interface The Johns Hopkins (CBI) Program Krieger School of Arts & Sciences Mentor: Dr. Caren L. Freel Meyers B.Sc. 2008 Department of Chemistry and University of Tulsa Biochemistry

Other Professional Experience: Teaching Assistant (2009-10) Organic chemistry lecture, Johns Hopkins University Research Rotation (2009) Lab of Dr. Phil Cole, Johns Hopkins University Research Rotation (2008) Lab of Dr. Blake Hill, Johns Hopkins University OCAST Intern (2007) Winston Chemical Company, Tulsa, OK SURF Student (2007) Lab of Dr. David Vanderah, NIST, Gaithersburg, MD Lab Assistant (2006-7) Organic chemistry I & II lab, University of Tulsa SURF Student (2006) Lab of Dr. David Vanderah, NIST, Gaithersburg, MD CSAS Tutor (2006-8) Student-athlete tutor, University of Tulsa MCAT Instructor (2006-8) Kaplan Test Prep, Inc., Tulsa, OK Student Research (2004-6) Lab of Dr. Gordon Purser, University of Tulsa

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Fellowships and Academic Honors:

2014 Chemistry-Biology Interface Program Travel Grant, Johns Hopkins 2008 Ralph J. Kaufmann Award in Biochemistry, University of Tulsa 2007 Best Undergraduate Poster, ACS Tulsa Chapter “Evening of Chemistry” 2007 Jess Chouteau Outstanding Senior, University of Tulsa 2006 Independent Research Grant, University of Tulsa 2006 Honorable Mention, Barry M. Goldwater Scholarship 2005 Top Ten Freshmen Award, Omicron Delta Kappa, University of Tulsa 2005 President’s List, University of Tulsa 2004-8 Dean’s List, College of Engineering and Natural Sciences, University of Tulsa 2004-8 National Merit Scholar, University of Tulsa 2004-8 Oklahoma Regents Scholarship, University of Tulsa

Publications:

Vierling, R.J., Sanders, S., Freel Meyers, C.L. Enzyme inhibition and antimicrobial activity of long-chain alkylacetylphosphonates targeting DXP synthase. Manuscript in preparation.

Vaish, A., Vanderah, D., Vierling, R., Crawshaw, F., Walker, M. (2014) Impact of Oligo (ethylene oxide) self-assembling monolayer terminal groups on the adsorption of integral membrane proteins. Colloids & Surfaces B: Biointerfaces 122: 552-558

Smith, J.M., Warrington, N.V., Vierling, R.J., Kuhn, M.L., Anderson, W.F., Koppisch, A.T., Freel Meyers, C.L., (2013) Targeting DXP synthase in human pathogens: enzyme inhibition and antimicrobial activity of butylacetylphosphonate. J. Antibiotics 67: 77-83

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Morris, F., Vierling, R., Boucher, L., Bosch, J., Freel Meyers, C.L. (2013) DXP synthase- catalyzed C-N bond formation: nitroso substrate specificity studies guide selective inhibitor design. ChemBioChem 14: 1309-1315

Smith, J.M., Vierling, R.J., Freel Meyers, C.L. (2012) Selective inhibition of E. coli 1- deoxy-D-xylulose 5-phosphate synthase by acetylphosphonates. MedChemComm 3: 65-66

Vanderah, D.J., Vierling, R.J., Walker, M.L. (2009) Oligo(ethylene oxide) self-assembled monolayers with self-limiting packing densities for the inhibition of nonspecific protein adsorption. Langmuir 25: 5026-5030

Posters and Presentations:

Vierling, R.J.. Investigating antimicrobial activity of acetylphosphonate inhibitors of DXP synthase. (2014) Invited speaker, CBI Annual Retreat, Johns Hopkins. (Baltimore, MD). (seminar)

Vierling, R.J., Freel Meyers, C.L.. Acetylphosphonates targeting DXP synthase as potential antimicrobial agents (2014) Frontiers at the Chemistry-Biology Interface Symposium (Baltimore, MD). (poster)

Vierling R.J., Freel Meyers, C.L., Acetylphosphonates targeting DXP synthase as potential antimicrobial agents. (2014) Gordon Research Conference, New Antibacterial Discovery & Development (Ventura, CA). (poster)

Vierling, R.J., Morris, F.M., Smith, J.M., Freel Meyers, C.L. Synthesis and evaluation of aromatic acetylphosphonates as selective inhibitors of DXP synthase Frontiers at the Chemistry-Biology Interface Symposium (Baltimore County, MD) (poster)

Vierling, R.J., Morris, F.M., Smith, J.M., Freel Meyers, C.L. Selective inhibition of E. coli 1-deoxy-D-xylulose 5-phosphate synthase by acetylphosphonates (2012) JHSOM Department of Pharmacology Annual Retreat (Baltimore, MD). (poster)

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Vierling, R.J., Morris, F.M., Smith, J.M., Freel Meyers, C.L. Selective inhibition of E. coli 1-deoxy-D-xylulose 5-phosphate synthase by acetylphosphonates (2012) CBI Program Annual Retreat (Baltimore, MD). (poster)

Vierling, R.J., DiCesare, J.C., Vanderah, D.J. (Oligo)ethylene oxide SAMS on Au: A system to understand protein adsorption on surfaces (2007) 233rd American Chemical Society National Conference (Chicacgo, IL). (poster)

Vierling, R.J., Brumback, K.J., Fry, A.L., DiCesare, J.C., Purser, G.H. Kinetics of the reaction between creatine and hypochlorous acid. (2006) 231st American Chemical Society National Conference (Atlanta, GA). (poster)

Hoppe, T.J., Robinson, D.M., Vierling, R.J., Purser, G.H. Recent findings concerning the kinetics of the reaction of monochloramine and chlorine dioxide. (2005) 229th American Chemical Society National Conference (San Diego, CA) (poster)

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