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Title Exploring biomolecular interactions : : New ligands and novel assays

Permalink https://escholarship.org/uc/item/62r0b1pd

Author McCoy, Lisa Christine Sator

Publication Date 2014

Peer reviewed|Thesis/dissertation

eScholarship.org Powered by the California Digital Library University of California UNIVERSITY OF CALIFORNIA, SAN DIEGO

Exploring biomolecular interactions: New ligands and novel assays

A dissertation submitted in partial satisfaction of the requirements for the degree Doctor of Philosophy

Chemistry

Lisa Christine Sator McCoy

Committee in Charge:

Professor Yitzhak Tor, chair Professor Michael Burkart Professor Steven Dowdy Professor Thomas Hermann Professor William Trogler

2014

Lisa Christine Sator McCoy, 2014

The dissertation of Lisa Christine Sator McCoy is approved, and it is acceptable in quality and form for publication on microfilm and electronically:

Chair

University of California, San Diego

2014

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DEDICATION

I would like to dedicate my thesis to my husband Aaron for his unending

support. Aaron has been very kind, loving, and completely selfless throughout my

graduate school career. I am extremely thankful for his sacrifice and compassion,

which helped me tremendously. He brings joy to my life, and I appreciate and

love him deeply.

I would also like to dedicate my thesis to my family. My parents, Frank and

Cynthia, have always been unconditionally loving and encouraging in my

endeavors. My brother, Karl, and his wife, Jennifer, have also been very

supportive. Aaron’s family, especially his parents Joe and Susie, has always

been understanding. I would also like to thank my grandparents, Bob and

Roberta King, and Karl and Edith Sator, as they have taught me to be hard- working and strive to be the best that I can be. I would also like to thank my other

family and friends for showing such wonderful support and generosity.

I very much appreciate my advisor, Dr. Yitzhak Tor, for his mentorship and

encouragement, as he has greatly contributed to my scientific development by

giving me guidance and freedom in my research. I would like to thank him for his patience, support and kindness.

Many members of the Tor lab have been extremely helpful and supportive.

I am very appreciative to have shared a lab with such wonderful people. I am

incredibly grateful to have met Patrycja Hopkins, who has become a meaningful

and lifelong friend during our time spent in and out of the lab. Rich Fair, Ryan

Weiss and Kristina Hamill have also been great friends and labmates, as we enjoyed many lunches and time together.

I would also like to thank my professors at Point Loma Nazarene

University who sparked my passion for science, and especially Dr. Vic Heasley, who introduced me to organic chemistry research. Finally, I would like to thank

God for all of this.

TABLE OF CONTENTS

Signature Page ...... iii

Dedication ...... iv

Table of Contents ...... vi

List of Figures ...... ix

List of Schemes ...... xii

List of Table ...... xiii

List of Spectra ...... xiv

Acknowledgements ...... xv

Vita ...... xvii

Abstract of the Dissertation ...... xviii

Introduction ...... 1

I.1 Prologue ...... 1

I.2 Antibiotics that target protein synthesis ...... 3

I.3 Ribosomal Decoding Site ...... 9

I.4 Peptidyl Transferase Center ...... 18

I.5 Peptide Exit Tunnel ...... 24

I.6 Targeting Other Sites ...... 30

I.7 Summary and Outlook ...... 32

I.8 References ...... 36

Chapter 1: Singly Modified Amikacin and Tobramycin Derivatives Show Increased A-site Binding and Higher Potency against Resistant Bacteria ...... 51

1.1 Introduction ...... 51

1.2 Results ...... 54

1.3 Discussion ...... 66

1.4 Conclusion ...... 69

1.5 Experimental ...... 69

1.6 References ...... 94

Chapter 2: Polymyxins and analogs bind to ribosomal RNA and interfere with eukaryotic translation in vitro ...... 97

2.1 Introduction ...... 97

2.2 Results and Discussion ...... 101

2.3 Conclusion ...... 110

2.4 Future Directions...... 110

2.5 Experimental ...... 112

2.6 References ...... 121 Chapter 3: Synthesis of meso- 2-deoxystreptamine mimetics ...... 127 3.1 Introduction ...... 127

3.2 Results and Discussion ...... 131

3.3 Conclusion ...... 136

3.4 Future Directions...... 137

3.5 Experimental ...... 139

3.6 References ...... 169

Chapter 4: Toward the development of fluorescently labeled RNA: Enzymatic incorporation of thGTP, an emissive GTP surrogate ...... 172

4.1 Introduction ...... 172

4.2 Results ...... 179

4.3 Discussion ...... 189

4.4 Conclusion ...... 192

4.5 Future Directions...... 192

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4.6 Experimental ...... 193

4.7 References ...... 199

Chapter 5: Enzymatic transformation of a fluorescent adenosine analogue into an inosine analogue: Development of a high-throughput assay ...... 203

5.1 Introduction ...... 203

5.2 Results and Discussion ...... 205

5.3 Conclusions ...... 214

5.4 Future Directions...... 215

5.5 Experimental ...... 216

5.6 References ...... 220

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LIST OF FIGURES

Figure I.1: Central Dogma of molecular biology ...... 1

Figure I.2: Main steps of bacterial translation ...... 5

Figure I.3: Crystal structure of the 30S ribosome showing the A-, P-, and E-sites ...... 6

Figure I.4: Section of a crystal structure of the 30S ribosomal subunit depicting the A-site ...... 10

Figure I.5: Aminoglycosides, and derivatives and mimics ...... 13

Figure I.6: Conserved contacts of neamine ...... 14

Figure I.7: Antibiotics that bind the PTC ...... 20

Figure I.8: Crystal structures of linezolid ...... 22

Figure I.9: Macrolides and ketolides ...... 25

Figure I.10: Crystal structures of erythromycin and telithromycin ...... 27

Figure I.11: Antibiotics that target other sites on the ribosome ...... 31

Figure 1.1: Tobramycin, amikacin, and derivatives ...... 53

Figure 1.2: Crystal structures of tobramycin and amikacin with the A-site ...... 55

Figure 1.3: Kanamycin-coumarin, Neomycin-coumarin, and Dy547-16S A-site construct ...... 59

Figure 1.4: Representative displacement curves of aminoglycoside -coumarins ...... 61

Figure 1.5: Kanamycin-coumarin displacement curves ...... 92

Figure 1.6: Neomycin-coumarin displacement curves ...... 93

Figure 2.1: Polymyxin B1 and lipid A ...... 98

Figure 2.2: Polymyxins and analogs ...... 99

Figure 2.3: Kanamycin, kanamycin-coumarin, and Dy547- 16S and 18S A-site constructs ...... 101

Figure 2.4: 16S A-site displacement curves ...... 102

Figure 2.5: 18S A-site displacement curves ...... 103

Figure 2.5: Bacterial in vitro translation assay data ...... 104

Figure 2.7: Eukaryotic in vitro translation assay data ...... 105

Figure 2.8: Comparison of kanamycin and analog 2 A-site binding curves and in vitro translation data ...... 106

Figure 2.9: Titration of 2 with the 16S A-site ...... 108

Figure 2.10: Comparison of 16S, 18S, and mitochondrial A-sites ...... 111

Figure 3.1: Structures of select aminoglycosides ...... 128

Figure 3.2: 2-DOS mimetics ...... 130

Figure 3.3: Comparison of 2-DOS and potential analogs ...... 131

Figure 4.1: T7 RNA polymerase general transcript yields ...... 175

Figure 4.2: Fluorescent nucleosides incorporated by T7 RNA polymerase ...... 177

Figure 4.3: Natural nucleosides and emissive RNA alphabet ...... 178

Figure 4.4: Enzymatic incorporation of thGTP using template 15 ...... 180

Figure 4.5: Small scale and large scale transcription results using thGTP ...... 181

Figure 4.6: Enzymatic incorporation of thGTP using templates 18 and 19 ...... 182

Figure 4.7: Large scale transcription using templates 18 and 19 ...... 183

Figure 4.8: Hammerhead ribozymes ...... 184

Figure 4.9: Hammerhead ribozyme cleavage reactions and kinetic data ...... 185

Figure 4.10: Fluorescence data of ribozyme cleavage ...... 186

Figure 4.11: Gel of thG-S & E cleavage reaction ...... 187

Figure 4.12: Digestion HPLC traces ...... 188

Figure 5.1: ADA catalyzed conversion of A and thA ...... 204

Figure 5.2: Crystal structures of A, thA and I, thI overlaid ...... 206

Figure 5.3: Spectral data of thA and thI and ADA with thA ...... 207

Figure 5.4: HPLC traces showing the conversion of thA to thI by ADA ...... 210

Figure 5.5: Predicted mechanism of deamination by ADA ...... 212

Figure 5.6: Structures of EHNA and Pentostatin and inhibition of ADA ...... 213

Figure 5.7: Negative controls of the thA ADA reaction ...... 214

LIST OF SCHEMES

Scheme 1.1: Synthesis of key tobramycin intermediates ...... 56

Scheme 1.2: Substitution reactions of 6’’-deoxy-6’’-triisopropylbenzylsulfonyl- (Boc)5tobramycin ...... 57

Scheme 1.3: Synthesis of 6’’-deoxy-6’’-ureidotobramycin ...... 58

Scheme 1.4: Synthesis of 6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)- (Boc)6tobramycin ...... 58

Scheme 1.5: Synthesis of key amikacin intermediates ...... 80

Scheme 1.6: Synthesis of 6’’-deoxy-6’’-aminoamikacin ...... 81

Scheme 1.7: Synthesis of 6’’-deoxy-6’’-methylaminoamikacin ...... 82

Scheme 1.8: Synthesis of 6’’-deoxy-6’’-dimethylaminoamikacin...... 84

Scheme 1.9: Synthesis of 6’’-doexy-6’’-(2-(aminoethyl)amino)amikacin ...... 85

Scheme 1.10: Synthesis of 6’’-deoxy-6’’-ureidoamikacin ...... 87

Scheme 1.11: Synthesis of 6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)- amikacin ...... 89

Scheme 2.1: Polymyxin analogs synthesis ...... 113

Scheme 3.1: Synthesis of 19 and 20 ...... 132

Scheme 3.2: Synthesis of 24 and 25 ...... 133

Scheme 3.3: Synthesis of 31, 32*, 24, and 35* ...... 134

Scheme 3.4: Proposed mechanism resulting in 31 and 32* ...... 135

Scheme 3.5: Proposed mechanism resulting in 34 and 35 ...... 136

Scheme 3.6: Possible reactions of 32 ...... 138

Scheme 3.7: Potential products of the building blocks ...... 139

Scheme 4.1: Synthesis the triphosphate of thGTP ...... 179

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LIST OF TABLES

Table 1.1: IC50 values for competing off kanamycin-coumarin ...... 60

Table 1.2: IC50 values for competing off neomycin-coumarin ...... 62

Table 1.3: Antibacterial activities of tobramycin and derivatives ...... 63

Table 1.4: Antibacterial activities of amikacin and derivatives ...... 65

Table 2.1: IC50 values for the 16S and 18S A-sites, and bacterial and eukaryotic IVT assays ...... 107

Table 5.1: IC50 values for the 16S and 18S A-sites, and bacterial and eukaryotic IVT assays ...... 207

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LIST OF SPECTRA

Spectrum 3.1: Triethyl-cis,cis-1,3,5-cyclohexanetricarboxylate (15) ...... 153

Spectrum 3.2: Cis,cis-(5-ethylcarboxylate)cyclohexane-1,3-dicarboxylic acid (16) ...... 154

Spectrum 3.3: Ethyl-cis,cis-(3,5-Cbz2diamine)cyclohexane-1-carboxylate (17) ...... 155

Spectrum 3.4: Cis,cis-(3,5-ethylcarboxylate)cyclohexane-1,3-diammonium acetate (18) ...... 156

Spectrum 3.5: Ethyl-cis,cis-(3,5-Boc2diamine)cyclohexane-1-carboxylate (19) ...... 157

Spectrum 3.6: Cis,cis-(3,5-Boc2diamine)cyclohexane-1-carboxylic acid (20) ... 158

Spectrum 3.7: Cis,cis-3,5-diazidocyclohexane-1-amine (24) ...... 159

Spectrum 3.8: Cis,cis-5-diazidocyclohexane-1,3-Boc2amine (25) ...... 160

Spectrum 3.9: (1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-diol (28) ...... 161

Spectrum 3.10: (1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3-diol (29)...... 162

Spectrum 3.11: (1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3- Tos2diol (30) ...... 163

Spectrum 3.12: (1S,3R,4R,6S)-4,6-diazidocyclohexane-1,3-Boc2diamine (31) ...... 164

Spectrum 3.13: (1S,3R,4S)-4,6-diazidocyclohex-5-ene-1,3-Boc2diamine (32*) ...... 165

Spectrum 3.14: (1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-Tos2diol (33) ...... 166

Spectrum 3.15: (1S,3R,4S,6R)-4,6-diazidocyclohexane-1,3-Boc2diamine (34) ...... 167

Spectrum 3.16: (1S,3R,4R,6R)-4,6-diazidocyclohexane-1,3-Boc2diamine (35*) ...... 168

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ACKNOWLEDGEMENTS

I acknowledge Professor Yitzhak Tor, as he has always been available for scientific questions and discussions. He has helped me tremendously with my research, for which I am extremely grateful.

I greatly appreciate the many members of the Tor lab, as they have helped me immensely in the lab during my graduate career. I would like to acknowledge Dr. Yun Xie for her help on FRET based A-site binding assay. I would like to thank Dr. Rich Fair for our collaboration on the aminoglycoside project, and for his help and support in the lab, as he has become a great friend.

Additionally, I acknowledge Dr. Renatus Sinkledam and Dr. Dongwon Shin for their help and collaboration on the fluorescent nucleoside projects. Additionally, I appreciate the many undergraduates who have helped me with my work.

I am grateful to Professor Thomas Hermann and his lab members for their help with the in vitro translation assays. I would also like to thank the Hermann lab for their scientific discussions that have helped me in my work.

I would also like to acknowledge Professor Victor Nizet and the members of his laboratory for their help in conducting the MIC assays.

I appreciate Dr. Yongxuan Su (UCSD MS Facility) for the mass spectrometry measurements. I also thank Dr. Anthony Mrse for his availability and assistance with the NMR instruments (UCSD NMR Facility).

The introduction, except for the prologue, is a full reprint from: McCoy, L.

S.; Xie, Y.; Tor, Y. Antibiotics that target protein synthesis. Wires RNA 2011, 2,

209. The dissertation author is the main author of this work.

Chapter 1 is in full currently being prepared for submission: Fair, R. J.;

McCoy, L. S.; Hensler, M. E.; Nizet, V.; Tor, Y. Singly Modified Amikacin and

Tobramycin Derivatives Show Increased A-site Binding and Higher Potency against Resistant Bacteria. The dissertation author is the co-main author and researcher of this work.

Chapter 2 is a full reprint from: McCoy, L. S.; Roberts, K. D.; Nation, R. L.;

Thompson, P. E.; Velkov, T.; Li, J.; Tor, Y. Polymyxins and analogues bind to ribosomal RNA and interfere with eukaryotic translation in vitro. Chembiochem

2013, 14, 2083. The dissertation author is the main author and researcher of this work.

Chapter 5 is a partial reprint from: Sinkeldam, R. W.; McCoy, L. S.; Shin,

D.; Tor, Y. Enzymatic Interconversion of Isomorphic Fluorescent Nucleosides:

Adenosine Deaminase Transforms an Adenosine Analogue into an Inosine

Analogue. Angewandte Chemie International Edition 2013, 52, 14026. The dissertation author is the second author and researcher of this work.

xvi

VITA

2005–2007 Teaching Assistant, Point Loma Nazarene University

2007 B.S. in Chemistry, Point Loma Nazarene University, San Diego

2007–2013 Teaching Assistant, University of California, San Diego

2009 M.S. in Chemistry, University of California, San Diego

2014 Ph.D. in Chemistry, University of California, San Diego

PUBLICATIONS

McCoy, L. S.; Xie, Y.; Tor, Y. Antibiotics that target protein synthesis. Wires RNA 2011, 2, 209. McCoy, L. S.; Roberts, K. D.; Nation, R. L.; Thompson, P. E.; Velkov, T.; Li, J.; Tor, Y. Polymyxins and analogues bind to ribosomal RNA and interfere with eukaryotic translation in vitro. Chembiochem 2013, 14, 2083. Sinkeldam, R. W.; McCoy, L. S.; Shin, D.; Tor, Y. Enzymatic Interconversion of Isomorphic Fluorescent Nucleosides: Adenosine Deaminase Transforms an Adenosine Analogue into an Inosine Analogue. Angewandte Chemie International Edition 2013, 52, 14026. McCoy, L. S.; Fair, R. J.; Hensler, M. E.; Nizet, V.; Tor, Y. Singly Modified Amikacin and Tobramycin Derivatives Show Increased A-site Binding and Higher Potency against Resistant Bacteria, In Preparation.

xvii

ABSTRACT OF THE DISSERTATION

Exploring biomolecular interactions: New ligands and novel assays

Lisa Christine Sator McCoy

Doctor of Philosophy in Chemistry

University of California, San Diego, 2014

Professor Yitzhak Tor, Chair

Many antibiotics bind the bacterial ribosome, the only validated RNA target. Derivatizing or mimicking these natural products is a potential way to create new RNA binders or antibacterials overcoming bacterial resistance. This motivated this work, which addressed the preparation of new RNA binders and the development of new assays.

xviii

Semi-synthetic derivatives of clinically useful aminoglycosides, tobramycin and amikacin, were prepared by selectively modifying their 6’’ positions. The binding to the rRNA A-site was probed by an in vitro Förster resonance energy transfer (FRET)-based assay, and antibacterial activity was quantified by determining minimum inhibitory concentrations (MICs). Most analogs displayed greater affinities for the bacterial A-site compared to the parent compounds.

Several amikacin analogs showed potent and broad-spectrum antibacterial activity against resistant bacteria, suggesting they are overcoming resistance mechanisms. However, tobramycin analogs exhibited overall poor antibacterial activity. As an alternative approach to potentially new RNA binders, mimetics of the aminoglycoside pharmacophore, 2-deoxystreptamine, were synthesized. In noting structural similarities to aminoglycosides, polymyxin antibiotics, which target the cell wall, were examined as rRNA binders using a FRET assay. The polymyxins showed significant affinity for to the bacterial and eukaryotic A-sites.

Additionally, in vitro translation assays showed all polymyxins interfered with eukaryotic translation, but not with bacterial, which could account for toxicity effects.

Fluorescent molecules and labeled oligonucleotides are valuable tools to probe small molecule, RNA, and protein interactions. We have developed assays utilizing isomorphic fluorescent nucleosides to study RNA catalysis and small molecule–enzyme interactions. A fluorescent guanosine analog, thG triphosphate, was incorporated into oligonucleotides by T7 RNA polymerase. Additionally,

xix

modified transcripts were used to assemble a hammerhead ribozyme containing

enzyme and substrate strands with thG replacing guanosine. The thG modified

substrate was effectively cleaved by the natural enzyme. However, the thG

modified enzyme showed no cleavage ability, suggesting the modifications likely

disrupted the catalytic center.

Toward exploring protein–nucleoside interactions with fluorescent

nucleoside analogs, a fluorescent adenosine derivative, thA, was found to be

deaminated by the enzyme adenosine deaminase (ADA) into the corresponding fluorescent inosine derivative, thI. A high-throughput assay utilizing the

fluorescent properties of these molecules to discover ADA inhibitors was

developed.

Introduction

I.1 Prologue

The sequential flow of genetic information from DNA to RNA to protein is

known as the central dogma of molecular biology (Figure I.1). Containing the

primary copy of genes, DNA is transcribed by RNA polymerase into messenger

RNA (mRNA) or non-coding RNA. Then mRNA is translated by the ribosome into

proteins, which ultimately carries out cellular functions. However, this is a very

simplistic view, as many other factors are in involved gene expression.1

Specifically, RNA, previously believed to be a dormant messenger, has emerged

as a very important processor and regulator of genetic information.2 For example,

non-coding RNA gives rise to important molecules that can catalyze reactions

(also known as ribozymes), such as the ribosome, group I and group II introns

(self-splicing RNAs), small nuclear RNAs involved in post-transcriptional mRNA splicing, among others.3-5 Other regulatory properties of RNA include guiding

chemical modifications of rRNA by small nucleolar RNAs,6,7 and riboswitches

found in mRNA regulate gene expression by binding of small molecules to the

aptamer domain.8-11

Figure I.1: Central Dogma of molecular biology.

Riboswitches highlight an important missing piece of the central dogma:

small molecules.12,13 Within an organism, small molecules participate in

processes interacting with and resulting from the main players of central dogma,

including riboswitches. Secondary metabolites are synthesized by modular

enzymes that have been used as therapeutics, which include many antibiotics.14-

16 Many of these small molecule antibiotics bind to rRNA and interfere with

translation.17 Additionally, natural and synthetic small molecules have been used

extensively as probes and drugs to manipulate and study these systems.

Specifically, fluorescent molecules have been an especially powerful tool

to study biological systems.18-20 A very sensitive method, fluorescence also

benefits from being non-invasive and allows for real-time reporting. Although

limited in number, some biomolecules are intrinsically fluorescent, such as green

fluorescent protein, tryptophan, NADH, chlorophyll, and others.21 However, since

the native nucleobases found in nucleic acids are almost non-fluorescent,

fluorophore reporters need to be appended to the structure. Extrinsic

fluorophores, often commercially available, tethered to biomolecules have been

exceptionally useful.19 However, a trade-off for stability and brightness is their

sometimes large size and they are often remote to the site of interest.22

Fluorescent nucleosides can be advantageous intrinsic probes, and particularly

isomorphic structures avoid perturbations as they retain Watson-Crick base

pairing and are similar in size to the natural counterparts.22,23 However, internal

or external fluorophore modifications can be extremely useful and provide

3 extensive information, and should be carefully chosen depending of the system of study.

Fluorescent molecules, including extrinsic probes and fluorescent nucleosides, have been used extensively to study RNA structure and function.18,19,22,24 Specifically, many methods using fluorescence have been used to monitor RNA-ligand binding interactions, and particularly with rRNA.23,25-35

Therefore it has been shown that many of these fluorescent tools can be used in the search for new RNA binders. Also, as inspiration for new RNA ligands, one can look to natural product interactions with the only validated RNA target, the ribosome. Thus, a thorough discussion of small molecule antibiotics that target rRNA will be discussed.

I.2 Antibiotics that target protein synthesis

The key function and abundance of the bacterial ribosome make it an obvious target for antibacterials. Indeed, a large number of clinically useful antibiotics, mostly of natural origin, target this translational ribonucleoprotein machinery. Alarmingly, there was a forty year hiatus in the discovery of new antibiotic classes, coinciding with a decrease in antibiotics approved for human use.36 This represents a formidable challenge, as the surge seen in the appearance of resistant bacteria, especially gram-negative bacilli, has not been met by a parallel development of effective and broad-spectrum new antibiotics.

This dire public health concern has revived an interest in the discovery and

development of new antibiotics, a research field that has been dormant for years.

There have been a few new classes of antibiotics introduced into the clinic in

recent years: oxazolidanones,37 lipopeptides,38 diarylquinolines,39 and

macrocycles,40 but all target gram-positive bacteria. In this chapter, we discuss

the major classes of antibiotics that target the bacterial ribosome and classify them according to their respective target. For clarity, we open with a brief

overview of the ribosome, its components and the major steps involved in

translation, also referred to as ribosome-mediated protein synthesis.

The prokaryotic ribosome is comprised of a small subunit (30S), which

consists of the 16S rRNA chain and 20 proteins, and a large subunit (50S) that

contains the 23S and 5S rRNA chains and 34 proteins.41 These constituents,

together with diverse factors, operate in concert to translate mRNA into a

polypeptide chain in three major stages: initiation, elongation, and termination

(Figure I.2). Initiation factors first assist the binding of the 5’ region of the mRNA

to the 30S subunit and bring the initiator fMet-tRNAfmet to the P-site of the 30S subunit. Following initiation, elongation of the peptide chain proceeds by transporting the proper aminoacyl-tRNA, charged with a cognate amino acid, to the A-site with the help of elongation factors. The tRNA anticodon must match the mRNA codon at the A-site for high fidelity synthesis. The peptidyl transferase center (PTC) in the 50S subunit then catalyzes peptide bond formation between the growing polypeptide chain on the peptidyl tRNA and the amino acid attached

Figure I.2: The main steps of bacterial translation. to the incoming aminoacyl tRNA. Finally, with the assistance of an elongation factor, the growing polypeptide chain and its aminoacyl tRNA are translocated from the A-site to the P-site, while the deacylated peptidyl tRNA moves to the E- site, and the mRNA moves by one codon (Figure I.3). Elongation continues until a stop codon is reached and termination takes place, where release factors aid in moving the mRNA stop codon into the P-site, which transfers the last tRNA into the E-site and off the ribosome. Simultaneously, peptidyl transferase catalyzes

the hydrolysis of the tRNA-peptide bond and releases the polypeptide chain

through the peptidyl exit tunnel. As discussed in this chapter, small molecules

can disrupt these intricate processes by interacting with key ribosomal

components.

Figure I.3: Crystal structure of three tRNA molecules in the A-, P-, and E-sites interacting with an mRNA molecule and the 16s rRNA in the 30S ribosome (PDB: 1GIX, 5.50 Å).

Although other RNA targets have been explored as potential drug targets,

the bacterial ribosome remains the only validated one with clinically approved

drugs.42,43 Multiple fundamental and practical reasons account for this. The

ribosome encompasses a large portion of the cellular RNA and contains distinct

7 and well defined binding sites for small molecules, which is not necessarily common with less structured cellular RNA targets.44 Small molecules with moderate affinity may exhibit practical potency due to the sheer volume to rRNA in the cell and the intricate function of its components. From a medicinal chemistry perspective, the bacterial ribosome is an attractive target since its RNA components are often coded by multiple operons, circumstances that may slow down the evolution of resistance.

The fundamental forces dictating RNA–ligand interactions have been discussed elsewhere.45,46 We note that the major groove of RNA duplexes, associated with a large negative electrostatic potential, can be rendered more accessible to small molecules by bulges and loops. This further exposes the hydrogen bonding face and the aromatic surfaces of the nucleobases for additional intermolecular interactions with potential binders.47 The minor groove of RNA duplexes frequently accommodates ligands through hydrophobic interactions. On a more global level, diverse models have been offered to explain the dynamic and structural characteristics of RNA–ligand recognition.

Conformational capture assumes RNA, being structurally dynamic, populates a variety of conformational states, which may or may not be related to the ultimate biological function.48-54 Ligand binding locks a specific conformation.

Alternatively (although the boundaries between these models are clearly vaguely defined), ligand binding may induce structural transformations, a process classically referred to as induced fit.55,56 The binding of highly cationic ligands to

RNA has been described by a spatial electrostatic complementarity model, where

the drugs’ covalently-linked three-dimensional array of positively charged groups

complements in space the negative electrostatic potential created by the RNA

fold.51-54 Frequently, cations natively bound in the highly negative potential

cavities of RNA can be displaced by positively charged ligands.53,54

Prior to the mid-1990s, the binding sites of antimicrobial agents were elucidated by ribosomal mutations that conferred resistance, as well as cross- linking and footprinting experiments.57 Modern biophysical methods, such as

NMR and X-ray crystallography, dramatically enhanced our knowledge of the

structure, function, and dynamics of the ribosome. Structural information at the

atomic level now provides a window into the actual interactions between

antibiotics and the targeted ribosomal constituents.58-81 These contemporary

techniques and the data they produce are being used for a structure-based

design of new candidate antibacterials.82 Although computational methods for structure-based drug design have shown promise, their application for addressing RNA targets remains limited. In particular, scoring functions for molecular docking have been primarily developed for ligand–protein complexes,

which do not always translate well for describing ligand–RNA interactions.52 The

flexibility of the phosphate backbone, hydration shell, and divalent ions, which

are all important for RNA–ligand interactions, represent challenging parameters

to capture in RNA-friendly computational programs.83,84 It is also important to

note that, while crystal structures can significantly aid in small molecule drug

9 design, the ribosome is dynamic, occupying different conformations throughout translation. This has been illustrated by the differences in the crystal structures of ribosomes in the apo form and when bound to tRNA mimics.85 Targeting highly dynamic domains with small molecules is likely to remain challenging.

Antibiotics that target the ribosome almost exclusively bind to one of the three key sites: the decoding (or A-site) on the 30S, the PTC on the 50S, and the peptide exit tunnel on the 50S. Antibiotics that bind the A-site, such as the aminoglycosides, interfere with codon recognition and translocation. Peptide bond formation is inhibited when small-molecules like oxazolidinones bind at the

PTC. Finally, macrolides tend to block the growth of the amino acid chain at the peptide exit tunnel. Notably, these antibiotics almost solely interact with the RNA components of the bacterial ribosome. We therefore classify the antibacterial agents discussed below according to their cognate ribosomal targets. We note, however, that this crude categorization is put in place for organizational purposes only, as the specific sites as well as the mode of binding and actual molecular contacts of individual antibiotics, even when grouped together, may vary.

I.3 Ribosomal Decoding Site

Codon–anticodon recognition and discrimination, performed at the A-site located on the 30S subunit, is critical for translation. Crystal structures of the 30S ribosomal subunit and the whole ribosome of T. thermophilus with mRNA and cognate tRNAs show the universally conserved bases A1492, A1493, and G503

interacting with the codon–anticodon minihelix,60,62 which was originally proposed

based on NMR structures of the A-site86 (Figure I.4A and I.4B, Note Escherichia

coli’s nomenclature is applied throughout). As the cognate tRNA enters the A-

site, conformational changes occur in the rRNA to initiate peptide bond formation.

A1492 and A1493 flip from a syn to an anti conformation and interact in the minor

groove of the first two base pairs in the anticodon–codon helix to scrutinize

hydrogen bonding patterns. G503 also flips into an anti conformation and

interacts with the second anticodon and the third codon base pairs (Figure I.4A

and I.3B). It tolerates non-Watson-Crick hydrogen base-pairs, consistent with the

degeneracy in the genetic code. Lowered stability of the codon–anticodon mini-

helix effectively assists in the rejection of non-cognate tRNA. This efficient

decoding process occurs with extremely low error rates (3 × 10–3 for E. coli).87

Figure I.4: A) Crystal structure of a section of the 30S ribosomal subunit (PDB: 1J5E, 3.05 Å). B) Crystal structure of a portion of the 30S ribosomal subunit complexed with an mRNA fragment and cognate tRNA anticodon stem-loop bound at the A-site (PDB: 1IBM, 3.31 Å). C) Crystal structure of a portion of the 30S ribosomal subunit complexed with an mRNA fragment and cognate tRNA anticodon stem-loop bound at the A-site with paromomycin (PDB: 1IBL, 3.11 Å).

As originally probed by biochemical and footprinting techniques, the

bacterial rRNA decoding site is the natural target of most aminoglycosides.88

The majority of these natural products, isolated from Streptomyces, contain the

11 highly conserved 2-deoxystreptamine (2-DOS) aminocyclitol, which is glycosylated with aminosugars at the 4-, 4,6-, or 4,5-positions (Figure I.5). The amine groups on the aminoglycosidic scaffold possess a range of pKb values, but they are mostly protonated at physiological pHs.89,90 Thus, electrostatic and hydrogen bonding interactions contribute to their RNA binding.91 While aminoglycoside antibiotics have evolved to bind the bacterial decoding site and meddle with protein biosynthesis, their highly cationic character and conformational flexibility facilitates binding to diverse RNA targets.53 In this respect, they can be viewed as rather universal or somewhat promiscuous RNA binders.53,54

The A-site is an autonomous RNA domain capable of mimicking the function and aminoglycosides recognition features of the entire 16S rRNA.92,93

This has paved the way for the use of short RNA constructs for both biochemical as well as NMR and X-ray crystallography studies.94-109 Additionally, structures with various aminoglycosides bound to the 30S ribosomal subunit and the entire bacterial ribosome were solved.58-62,76,79 Although there are inherent differences in resolution and scope among the solved structures, they provide a consistent molecular picture on the overall interactions of aminoglycosides. It is now established that aminoglycoside antibiotics bind the decoding site and stabilize an RNA conformation similar to the one induced by the cognate acyl-tRNA– mRNA. Aminoglycosides cause nucleobases A1492 and A1493 flip outside the helix to a locked anti position (Figure I.4C). This disrupts the decoding process,

lowers its fidelity and ultimately increases the misincorporation of amino acids;110-

112 however, evidence also suggests that aminoglycosides interfere with

translocation.113-118 It is worth noting that streptomycin (1), the only guanidinium- containing aminoglycoside as well as the first to be isolated and clinically used, displays a distinct binding site and mode of action; it interferes with initial tRNA selection.59

Crystal structures revealed that the paromamine/neamine/gentamicin

cores exhibit highly conserved interactions in paromomycin (6),59,100 neomycin B

(5), kanamycin A (13), gentamicin C1A (11), ribostamycin (3), lividomycin A

(7),103 and amikacin (16).106 Positions N3 and N1 hydrogen bond to N7(G1494)

and O4(U1495), respectively. Additionally, N3 forms a salt bridge with

O1P(A1493) and O2P(G1494). Ring II stacks against G1491, and O5’ bonds

with N6(A1408), and the 6’ functional group, a hydroxyl in paromomycin and an

amine in the others listed, interacts with N1(A1408) (Figure I.6A and I.6B).

These observations suggest that rings I and II represent the key pharmacophore

of the aminoglycosides. The 2-DOS ring is now viewed as a privileged RNA

binding scaffold.119,120

Figure I.5: Aminoglycosides, and derivatives and mimics of aminoglycosides.

Aminoglycosides typically display specificity toward prokaryotes over eukaryotes. This is facilitated by both differences in membrane permeability and intrinsic target specificity.121-124 In contrast to prokaryotic ribosomes, which contain an adenosine at 1408 and guanine at 1491, eukaryotic rRNA contains a guanine and an adenine, respectively, at these positions, which are essential for aminoglycoside recognition.103 Point mutations in bacteria, such as A1408G, lowers sensitivity to many 2-DOS containing aminoglycosides, and when these bacteria contain only one copy of rRNA operon, total resistance ensues.122,125

Importantly, human mitochondrial ribosomes that have A1408 and G1491 at analogous positions exhibit higher resemblance to their bacterial counterparts.

This similarity is likely responsible for some of the adverse effects shown by aminoglycosides (e.g., ototoxicity).121,126,127

Figure I.6: A) An overlap of the crystal structures of neomycin (PDB: 2ET4, 2.40 Å), tobramycin (PDB: 1LC4, 2.54 Å), and paromomycin (PDB: 1J7T, 2.50 Å) in an A-site model construct. B) Conserved contacts of neamine (2).

Not unlike many other antibiotics, the clinical use of aminoglycosides has

been compromised by the emergence of resistant bacteria. The prominent

resistant mechanisms rely on enzymatic modifications of the antibiotics, including

O-adenylation, O-phosphorylation, and N-acetylation;128 all lower their affinity to

the rRNA target. The innate flexibility of the glycosidic bonds in aminoglycosides

promotes their binding to resistance enzymes,129 which likely evolved to kill competitor and predator organisms and to provide self immunity to the producing organism.130-132 Through horizontal gene transfer, this self-preserving trait could

have passed onto other bacteria.131,132 Many other resistant mechanisms exist, as well, such as the operation of efflux pumps, rRNA modifications, and mutations in ribosomal proteins.133 The removal of aminoglycosides through

efflux pumps was first reported in B. pseudomallei and has been observed in many nosocomial infections, such as P. aeruginosa.134,135 Modifications of rRNA include the methylation of N1 in A1408 and N7 in G1405, and it results in resistance to some 4,6-substituted 2-DOS aminoglycosides.130,136

To combat the rise in bacterial resistance, while improving the efficacy and

circumventing the toxicity of known drugs, new aminoglycoside derivatives have

been sought.137-139 Early as well as more recent efforts involved diverse

modifications of the natural products.140-144 For example, amikacin (Figure I.5,

16), containing an acylated 2-DOS ring, is a semi-synthetic aminoglycoside that is highly effective against bacterial inactivating enzymes. The presence of the L-

hydroxyaminobutyramide residue (AHB) at N1 lowers its susceptibility towards

acylation and phosphorylation.133,145 A more recent compound is plazomicin (18)

(ACHN-490), a derivative of sisomycin (17), which is currently in phase III clinical trials.146 Other approaches have been rather diverse and include conformational

restriction of aminoglycosides,105,147,148 the functionalization of 2-DOS with

carbohydrate free moieties,149-154 replacement of the 2-DOS core with mimetic

scaffolds,155 or creation of totally synthetic molecules.156

As previously mentioned, aminoglycosides exhibit some degree of

flexibility around their glycosidic bonds, allowing them to conformationally adapt

and bind diverse targets.53 To explore specificity, conformationally restricted neomycin (19) and paromomycin (20, Figure I.5) derivatives were synthesized to study A-site binding and antibacterial activity.105,147 The crystal structure of the

conformationally locked neomycin derivative bound to the A-site exhibited the

identical intermolecular interactions between rings I and II as neomycin.105 The affinity and antibacterial activity decreased, however, when compared to the natural product. This has been attributed to the impact this intramolecular cyclization had on basicity of the amine at position 2’.157 Intriguingly, this

modification has also been examined in the context of antibiotic resistance and

the susceptibility to resistance enzymes of the restricted neomycin derivative was evaluated.148 Crystal structures of certain aminoglycosides bound to resistance enzymes exhibit significantly different binding modes than the bound A-site conformation, inspiring the design for the restricted neomycin analogue.120 The

restriction of the derivative prevents adenylation by Staphylococcus aureus

ANT(4’) and Mycobacterium tuberculosis AAC(2’), and accordingly retains

antibacterial activity in resistant strains containing these enzymes.148

To maintain conserved contacts, attempts have been made at derivatizing

neamine (2), the key aminoglycoside pharmacophore. Based on molecular

modeling, Mobashery and coworkers prepared neamine analogues with AHB on

N1, similar to amikacin, and aminoaliphatic chains on O6.151 The crystal structure

of the A-site bound derivative 21 shows that both arms provide additional

contacts to RNA. Superimposition of this compound with the neamine core of

kanamycin bound to the phosphotransferase resistance enzyme, APH(3’)-IIIa,

suggested that the AHB substitution hinders binding.158,159

Aminoglycoside mimetics containing non-carbohydrate moieties in place

of the natural sugars rings (e.g., aminoalkyl chains) have been synthesized.149-151

Another approach reported by Griffey and coworkers involved replacing Ring II

on aminoglycosides cores with aromatic groups.152,153 The two approaches were combined to create carbohydrate free aminoglycoside mimetics containing 2-

DOS, with compound 22 showing inhibition of bacterial translation.154

As 2-DOS appears to be a privileged RNA binding scaffold,42 DOS-

mimetics can, in principle, maintain essential A-site contacts. Hermann and coworkers replaced 2-DOS in neamine with azepane glycosides. Compound 23

showed moderate activity against S. aureus and other aminoglycoside resistant

strains.155 Another set of derivatives, based on 3,5-diamimino-piperidinyl triazines

(DAPT) as 2-DOS mimetics, was shown to inhibit protein synthesis, but had

weak antibacterial activity. Further structural refinement generated derivative 24

that demonstrated increased potency against some Gram-negative bacteria.156

To facilitate the discovery of A-site targeting agents and new antibacterials, numerous approaches have been explored.160,161 In addition to the

use of mass spectrometry and NMR techniques, simpler biophysical assays have

been developed.162-166 Fluorescent A-site constructs, which contain 2-

aminopurine at positions 1492 or 1493, have shown great promise, as they rely

on changes in the location and dynamics of these residues.25,167 A robust

analysis platform for antibiotics targeting the bacterial A-site has also been

developed by incorporating a new emissive uridine surrogate into the RNA and

labeling the aminoglycosides with a coumarin. Detection of antibiotic binding and

displacement is dependent on the interaction between the two chromophores

acting as a FRET pair.29 This approach has recently been further advanced to facilitate the rapid determination of the prokaryotic vs. eukaryotic selectivity of A-

site binders.168

I.4 Peptidyl Transferase Center

The PTC connects all functional cores in the ribosome, including the tRNA

entrance and exit regions, as well as the A-site. In essence, it is the ribosomal

active site, catalyzing peptide bond formation by meticulously positioning the

reactive partners.169-171 Not surprisingly, numerous antibiotics target this key site,

including chloramphenicol (25), clindamycin (26), tiamulin (27), sparsomycin (28) and the streptogramins (29−32), as well as the oxazolidinones (33–35), the only fully synthetic class of antibiotics (Figure I.6). These antibiotics either hinder tRNA substrate binding or disrupt peptide bond formation by binding to the PTC itself.

Chloramphenicol (25), one of the first naturally occurring antibiotics to be produced synthetically, interferes with A-site and tRNA binding, preventing peptide bond formation.64,67,172,173 On the other hand, clindamycin (26) and

tiamulin (27) interfere with both A-site and P-site tRNA substrate binding.

Overlapping the chloramphenicol (25) binding site, clindamycin (26), a lincosamide, binds at the PTC and peptide exit tunnel.65,67 It inhibits the peptidyl

transferase reaction and causes dissociation of peptidyl tRNA from the

ribosome.174,175 Tiamulin (27), a member of the pleuromutilin class of antibiotics,

inhibits peptide bond formation.176-178 It competes with chloramphenicol (25),

clindamycin (26), and the saccharide branch of carbomycin A (41) for

binding.63,67,74,178,179

Figure I.7: Antibiotics that bind to the PTC.

Sparsomycin (28), a uracil derivative, is an effective protein synthesis inhibitor of both bacterial and eukaryotic ribosomes.180,181 It has been shown to be necessary for a tRNA to be bound in the P-site for sparsomycin to bind,182 and crystal structures with the H. marismortui 50S subunit (H50S) suggest extensive

interactions between sparsomycin and the CCA 3’-end of the P-site tRNA

substrate analogue.64 It can bind, however, to D. radiodurans 50 S subunit

(D50S) alone and causes a great conformational change in the PTC, which could

be the origin of its inhibitory activity on peptide bond formation.71 Although sparsomycin competes with chloramphenicol (23), it does not competitively

inhibit A-site tRNA binding.180

The streptogramin A antibiotics (29, 30), a family of unsaturated

macrocyclic depsipeptides (Figure I.7), typically bind at the PTC and overlap the

A-site and P-site tRNA, likely interfering with tRNA binding.64,65,73 In contrast, the streptogramin B antibiotics (31, 32), a family of macrocyclic depsipeptides

(Figure I.7), bind in the peptide exit tunnel in the macrolide binding region,65,73

possibly impeding peptide elongation. Notably, while individual members of each

family show modest bacteriostatic activity, the two families act synergistically and

display greater bactericidal effects.183-185

Importantly, the PTC is targeted by the oxazolidinones, a family of

synthetic derivatives representing one of the very few success stories of

antibiotic development. Exploring these derivatives as orally available

antimicrobial agents started in the 1970s, ultimately leading to the introduction of

linezolid (33) into the clinic a decade ago. It is active against many gram-positive

bacteria, including methicillin resistant Staphylococcus aureus (MRSA) and vancomycin-resistant enterococci (VRE).186,187

Early studies suggested linezolid (33, Figure I.7) bound to the 30S and

50S subunits,188 but newer cross linking studies have indicated binding only at the PTC.189,190 Crystal structures of linezolid (33) bound to H50S with and without

CCA-N-acetylphenylalanine (CCA-Phe) show a drug molecule in the PTC and a

CCA-Phe molecule in the P-site.66 It appears likely that oxazolidinones compete

with the A-site substrate, binding near the bases A2451, U2585, and U2506

(Figure I.8A). These bases are shown to be important in A-site−tRNA binding

and the positioning of the substrates for the peptidyl transferase reaction.171

Although alternative structures show slightly distinct contacts, the general

location of linezolid (33) bound at the PTC is similar (Figure I.8C).66,80

Figure I.8: A) Crystal structure of linezolid bound to the H50S (PDB: 3CPW, 2.70 Å). B) Crystal structure of linezolid bound to the H50S (PDB: 3CPW, 2.70 Å) overlapped with the crystal structure of native H50S (PDB: 3CC2, 2.40 Å). C) Crystal structure of linezolid bound to the H50S (PDB: 3CPW, 2.70 Å) and D50S overlapped (PDB: 3DLL, 3.50 Å).

The binding specificity of linezolid (33) may be explained by comparing the

50S subunit of the bacteria E. coli (E50S) and archaeon H. marismortui.66 In

H50S, when linezolid (33) is bound, the conformation of U2504 is nearly identical to the homologous base in E50S with no ligand (Figure I.8B),62 indicating that the

binding pocket for the drug is prearranged in bacterial ribosomes and more rigid.

Oxazolidinones have been shown, however, to bind to and possibly inhibit

archaeal, bacterial, and mitochondrial ribosomes, but not those from human

cytoplasm.190,191 It is also suggested that the position of U2504 in eukaryotic

ribosomes could possibly be incompatible with linezolid (33) binding.80 In addition, cross-resistance between PTC binders occurs due to their overlapping binding sites near U2504.75 Almost all nucleotides conferring resistance are clustered on one side of the PTC near nucleotide U2504. About half of the nucleotides mediating resistance do not directly interact with bound PTC antibiotics, but reshape the binding pocket, often by affecting the conformation and/or flexibility of U2504.75

Bacterial resistance to linezolid (33) is currently minimal,192 but as for all

other antibacterials, it is expected to escalate. The only mechanism of clinical

resistance recorded so far is the target modification of the PTC, with a G2576U

mutation found in Enterococci and Staphylococcus aureus.193 Although

resistance is currently uncommon for linezolid (33), as recognition by efflux

pumps and resistance enzymes of a completely synthetic molecule should not

exist, search is underway for novel oxazolidinones. One example worth

mentioning is hybrid molecules inspired by the partial overlap seen in the binding

sites of linezolid (33) and sparsomycin (28).64,66,194,195 Radezolid (RX-1741) (34)

and Torezolid (TR-700) (35) represent two derivatives that are in advanced

clinical trials (Figure I.7).196,197

I.5 Peptide Exit Tunnel

Towards the end of protein synthesis, the newly synthesized peptide chain

travels through a dynamic tunnel below the PTC, lined with mostly rRNA, and

emerges approximately 100 Å out of the 50S subunit. While the exit tunnel is

primarily straight, there is a bend 20–35 Å away from the PTC. In this bent

section, L22 and L4, two ribosomal proteins, constrain the tunnel into its

narrowest part.198 At this region, the peptide sequence can influence gating, or

pause elongation and termination.199-201 Antibiotics, such as the macrolides,

which bind this domain, impede the progression of the nascent peptides.

Macrolides, a family of naturally occurring 12- to 18-membered

macrocyclic lactones, typically contain one or more deoxy monosaccharides

(Figure I.9). Erythromycin (36) is a prototypical example, as it was the first to be

clinically used. Semi-synthetic derivatives of erythromycin (36) include 14- and

15- membered ring macrocycles (37−40), as well as ketolides (44−46).

Macrolides and ketolides bind to the peptide exit tunnel and prevent elongation,

63,65,67,69,70,72 consequently inhibiting protein synthesis.175 This results in

premature dissociation of short peptidyl-tRNAs from the ribosome,175 which was originally misinterpreted as inhibition of peptidyl transferase or translocation.202

The length of the resulting short terminated peptides depends on the individual

antibiotic.175,203,204

Figure I.9: 14-, 15-, and 16-membered ring macrolides and ketolides.

Macrolides and ketolides have been crystallized with eubacterium D. radiodurans, archaeon H. marismortui, and a H. marismortui mutant (G2058A

H50S).63,65,67,69,70,72 D. radiodurans and pathogenic bacteria contain an adenine

at position 2058, whereas the naturally resistant wild-type H50S contain G2058.

This motivated Steitz to use the G2058A 50S mutant to facilitate a comparison

D50S, since both could be co-crystallized under physiologically relevant drug

concentrations (~3μM).63,65,67,69,70,72 While some discrepancies have been seen in

the crystal structures of ribosome-bound macrolides and ketolides, general

binding patterns emerge. The lactone ring appears to hydrophobically interact

with A2058 and A2069.63,65,67,69,70,72 The 2’ hydroxyl of the aminosugar desosamine, or mycaminose in the 16-membered ring macrolides, hydrogen bonds to N1 of A2058.63,65,67,69,70,72 In addition, consistency in the X-ray

structures has been seen between erythromycin (36) and its semisynthetic

derivatives clarithromycin (37) and roxithromycin (38).67 The structure of

erythromycin (36) bound to H50S is different than the D50S-bound structure

(Figure I.10A).63,65,67 Acetylation of hydroxyl groups, altering their H-bonding

capabilities, impacts the binding mode, as seen for troleandomycin (39). It binds

further into the tunnel and directly contacts the ribosomal protein L22, eliciting the

β-hairpin to move across the tunnel. Perhaps the configuration in L22 shows the conformation dynamics involved in gating, as this protein is likely involved in controlling elongation of certain amino acid sequences.70

Figure I.10: A) Overlap of the crystal structure of erythromycin (orange) in the H50S (PDB: 1YI2, 2.65 Å) with telithromycin in the D50S (PDB: 1P9X, 3.40 Å). B) Overlap of the crystal structure of telithromycin (magenta) in the H50S (PDB: 1YIJ, 2.60 Å) with telithromycin in the D50S (PDB: 1P9X, 3.40 Å).

Crystal structures of azithromycin (40) bound to H50S show one binding

site with the macrocyclic lactone ring adopting a folded out conformation, similar

to the one seen for erythromycin (36).63,65 Two distinct binding sites have been

observed with D50S.72 The primary binding site is in the same pocket as

erythromycin (36), but the macro-lactone ring adopts a different conformation.72

The secondary binding site is adjacent to the other bound azithromycin (40), and

may be specific to H. marismortui.72

A unique feature of the 16-membered ring macrolides (41–43) has been

observed with wild-type H50S, where the aldehyde at position 6 forms a

hemiaminal with the N6 exocyclic amine of A2026.63 This suggests a key role for

this functionality, which is supported by the observed decreased activity of

derivatives with modifications at this position in a wide variety of Gram-positive

bacteria.205,206 In addition, the saccharide moiety attached to C5 extends towards

the peptidyl transferase center, which may explain why carbomycin A (41)

(Figure I.9) strongly inhibits formation of the first peptide bond and why tylosin

and spiramycin allow the formation of very short peptides.175,203,204

Ketolides (44–46), derived from erythromycin (36), contain a 3-keto moiety in place of the L-cladinose. This confers higher stability in acidic media and increased oral bioavailability.207 A cyclic carbamate and an additional

heterocyclic-containing side chain are incorporated (Figure I.9). Crystal

structures of D50S ribosome-bound ketolides show an extended conformation for

the lactone ring and the long side chain protruding down the tunnel (Figure

I.10).69 The G2058A H50S crystal structure shows, however, a different

orientation of telithromycin (45), with its lactone and desosamine rings almost in

the exact orientation as erythromycin (36) (Figure I.10A).65 Stacking interactions

of the side chain may explain the 10-fold increase in binding affinity for E. coli

ribosomes over erythromycin (36).65,208

Macrolides and ketolides selectively target the prokaryotic ribosome. One

factor responsible for this selectivity is the guanine residue at position 2058 in

eukaryotes, which parallels an adenine in prokaryotic ribosomes. Bacteria appear

to mimic this source of eukaryotic resistance by alteration of position 2058.

Resistance can be conferred through methylation of N6 at A2058 by

methyltransferases. Monomethylation has a moderate effect on macrolides and

no effect on ketolides.209 Dimethylation, however, is the most effective form of

resistance for macrolides and ketolides.209 Point mutations on the ribosome,

which lower antibiotic affinity, can also occur (e.g., A2058G and A2059G).

Telithromycin (45) binding is reduced in E. coli A2058G, but it is still twenty times

higher than erythromycin (36) and clarithromycin (37). Mutating A2062 to C

confers resistance to many 16-membered macrolides, but not to the 14- or 15-

membered antibiotics.210 This could be due to the inability to form the hemiaminal

linkage discussed above. Efflux is a main source of resistance for macrolides, as

with many antibiotics,134,211 and certain enzymes that can modify the macrolides

have also been reported.212 In addition, mutations in ribosomal proteins L22 and

L4, which are close to the binding site, can result in resistance to

macrolides.213,214 It is important to note that cross resistance is prevalent

between some peptide exit tunnel and PTC binders, specifically the macrolide,

lincosamide and streptogramin B derivatives, due to their overlapping binding

sites. As mentioned for macrolides, methylation of A2058 or the mutation of the

adenosine to guanosine also confers resistance to lincosamides and

streptogramin Bs.215,216

While the macrolides and ketolides have had and still have significant

clinical utility, new derivatives are clearly needed to combat bacterial resistance.

A new ketolide showing promising activity is cethromycin (ABT-773 or

RestanzaTM) (44) (Figure I.9).217 Its potency is similar to that of telithromycin,

showing a broader range of activity against Gram-positive bacteria and efficacy

against bacteria resistant to macrolides.217 The crystal structure of cethromycin

bound to D50S is comparable to erythromycin, but different from telithromycin.72

Another new agent is Modithromycin (EDP-420) (46), a ketolide that is currently

undergoing clinical trials (Figure I.9).218 It shows activity against many resistant

strains of S. pneumoniae, making it a potential drug candidate.

I.6 Targeting Other Sites

Although most antibiotics inhibit translation by binding to the A-site, PTC, and peptide exit tunnel, there are ribosome-targeting antibacterials that do not fit

into these categorical binding sites. Certain classes of antibiotics bind to the 30S

ribosomal subunit near the A-site, at the P-site and at the E-site. Tetracyclines

(47) and glycylcyclines, derivatives of tetracycline that include tigecycline (48)

(Figure I.11), bind near the A-site and prevent aminoacyl-tRNA from binding.58,68,219-221 Hygromycin B (49) and spectinomycin (50), aminocyclitol

antibiotics related to aminoglycosides (Figure I.11), inhibit translocation by

binding near the A-site.58,78,79,222 Edeine A (51), shown in Figure I.11, inhibits

initiation by binding at the E-site, with its spermidine moiety docked in the P-site

blocking the initiator tRNA.68,223,224 Kasugamycin (52), binding at the P-site and

E-site, perturbs the mRNA tunnel, which likely prevents initiator tRNA

binding.81,223 Pactamycin (53) binds at the E-site, distorting the mRNA path and

subsequently inhibiting translocation.58,223,224 Negamycin (54) inhibits protein

synthesis termination and induces miscoding.139 Its crystal structure, when bound

to the 50S ribosomal subunit, shows binding in the peptide exit tunnel, closer to

the exit than the macrolides.77 Evidence suggests, however, that negamycin (54)

binds to or near the A-site as well.225 Thiostrepton (55) inhibits translocation by

binding at or near protein L11 on the 50S subunit.223,226

Figure I.11: Antibiotics that target the ribosome near the A-site, and at the P- and E-sites.

I.7 Summary and Outlook

Despite its enormous size and intricate function, the ribosome has so far been shown to include only few validated antibiotic targets. While these may represent the most functionally sensitive sites, critical for high fidelity and efficient translation, it is conceivable that additional targets and novel approaches to translational interference exist. Intriguingly, many of the clinically relevant antibiotics exclusively bind the RNA components. Efforts to find new potential antibiotic binding sites on ribosomal RNA have therefore utilized deleterious mutations to identify RNA domains of essential significance to ribosomal function.227 Such putative targets have yet to be validated. It is worth noting here that RNA targeting with small molecules, as a research discipline, is still in its early stages.42,43,228-230 The ability of ribosomal RNA to bind molecules of extremely different molecular structures is remarkable, demonstrating its versatility and potential as a target for antibiotics. While the ribosome and the antibiotics targeting it have provided the main impetus and inspiration for the recent evolution of this field and its expansion into many other RNA targets (even before high resolution structures became available), ribosomal RNA is again becoming a target of critical consequences.

The significant health threats of resistant bacteria have revitalized the search for new antibacterial agents. Researchers have to respond to the increasing abundance of serious infections by identifying new targets and novel agents to selectively destroy existing and resistant bacteria. Facilitating the

discovery of new antibiotics, numerous approaches have been recently

discussed, including diversity and screening based approaches,231,232 chemical genetics and metabolic engineering-based protocols,232,233 as well as the

discovery of new naturally occurring agents,233 the preparation of semi-synthetic derivatives, and the identification of small molecules that can re-sensitize bacteria in resistant biofilms to existing antibacterial agents.234

The challenge in developing new antibiotics that selectively target the

bacterial ribosome becomes apparent when one reviews Paul Ehrlich’s proposal

for Magic Bullets (1906): “It will obviously be easy to effect a cure if substances

have been discovered which have an exclusive afﬁnity for bacteria and act

deleteriously or lethally on these alone, whilst at the same time, they possess no

afﬁnity whatever for the normal constituents of the body and cannot therefore

have the least harmful or other effect on that body.”235 The ribosome, while being

a highly efficient yet susceptible and delicate protein synthesis machine, is a

“universal” apparatus, which is shared, albeit with certain differences, by

numerous low and high organisms. While many factors affect the efficacy and

adverse effects of antibiotics, their inherent affinity to competing targets is of

fundamental significance. The ability of a candidate antibacterial to distinguish

between eukaryotic, prokaryotic and mitochondrial rRNA is likely to impact its

therapeutic window. Although not without side effects, this specific element is

immaterial when antibiotics that act on cell wall biosynthesis, for example, are

concerned. Nevertheless, the plethora of potential small molecule binding sites

on the ribosome and its vulnerability to even minor alterations are likely to

provide valuable opportunities to discover new antibiotics, some possibly with

novel mechanisms of action.

In designing new antibiotics for RNA targets, one can either modify current

scaffolds or synthesize scaffolds mimicking key pharmacophores.

Aminoglycosides, as previously discussed, are known RNA binders that have

been studied extensively. Derivatizing aminoglycosides with minimal

modifications, which are synthetically accessible, could potentially overcome

current resistance mechanisms. Alternatively, purely synthetic mimics could

perhaps avoid some of these resistance mechanisms altogether, as there is no

inherent resistance genes as there is with natural product antibiotics, although

resistance always eventually develops. There is no guarantee, of course, that the

compounds will exhibit the same mode of action or kill bacteria. In our search for

new RNA binders and potential antibiotics, we have tried several approaches.

We made singly modified derivatives of two aminoglycosides, tobramycin and

amikacin. Additionally, we have synthesized mimics of the key pharmacophore of

aminoglycosides, 2-DOS, which could be used as potential RNA binding scaffolds. Additionally, we explored a possible new RNA target for old antibiotics, polymyxins.

In order to assess RNA affinity, we use a binding assay that takes advantage of small molecule fluorophores. The development of these assays is critical for studying biological processes or screening or for new drugs. Classical

techniques, such as footprinting236 and NMR spectroscopy,237 in probing small molecule binding to nucleic acids, or any target such as proteins, have not been as amenable to high-throughput assays.238-240 In order to establish fundamental

structure–activity relationships and to facilitate target-specific drug discovery, assays using fluorescent probes have been developed. Specifically, fluorescent nucleosides are an excellent tool to study DNA, RNA, and any interactions they have with high or low molecular weight ligands, such as proteins or small molecules. Therefore, we aspired to advance the utility of some of the

nucleosides in a fluorescent alphabet recently synthesized in our lab.241

Specifically we sought to enzymatically incorporate thG, a guanosine analog, into oligonucleotides. We have assessed thGTP as a surrogate for GTP in transcription by T7 RNA polymerase. Additionally, we transcribed oligonucleotides to create a modified ribozyme containing strands with thG

replacing G, and tested its function. Furthermore, we have tested the ability of thA

to be deaminated by the enzyme adenosine deaminase (ADA) to develop a high-

throughput assay to discover ADA inhibitors, as an entry into the rather

unexplored area where fluorescent nucleoside analogs interact with proteins.

Acknowledgements

The introduction, except for the prolog, is a full reprint from: McCoy, L. S.;

Xie, Y.; Tor, Y. Antibiotics that target protein synthesis. Wires RNA 2011, 2, 209.

The dissertation author is the main author of this work.

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Chapter 1

Singly Modified Amikacin and Tobramycin Derivatives Show Increased A-site Binding and Higher Potency against Resistant Bacteria

1.1 Introduction

The discovery of penicillin, a β-lactam, and streptomycin, an aminoglycoside, in the 1940s launched the golden age of antibiotics. Many of the antibiotics discovered in the 1940s through the 1970s are used in the clinic today.1 However, the flood of antibiotics into the environment via feedstock and human use has contributed to the increase in resistant pathogens. Horizontal gene transfer between bacteria via plasmids and other methods has played a significant role in conferring resistance.2,3 Drug resistance bacteria, especially the

ESKAPE pathogens (Enterococcus faecium, Staphylococcus aureus, Klebsiella

pneumoniae, Acinetobacter baumannii, Pseudomonas aeruginosa, and

Enterobacter species), Clostridium difficile, and Escherichia coli, have been commonly infecting not only immunocompromised hospital patients, but also otherwise healthy individuals.4-6 This has led to rising healthcare costs, often due

to length of stay in the hospital, and increased mortality.7 Problematically, the number of new antibiotics approved by the US Food and Drug Administration has been steadily decreasing, and concurrently many pharmaceutical companies

have been abandoning or downsizing their antibacterial research and

development.8,9

On a positive note, there have been a few new classes of antibiotics in

recent years, all of which target gram-positive bacteria.10-13 Nevertheless, the

emergence of multi-drug resistant bacteria, especially gram negative bacilli with

no new treatment options, has led to reexamination of drugs from the early years

of antibiotic discovery.14-17 Aminoglycosides are effective against a broad range

of bacteria, although the advent of safer, less toxic antibiotics resulted in their

declined use. However, with the increase in resistant pathogens, especially

severe gram negative infections, aminoglycosides remain useful for specific

infections in the clinic.18,19 Tobramycin (1a) is specifically used for P. aeruginosa

infection in cystic fibrosis patients, amikacin (2a) is used for highly resistant gram

negative infections, and gentamicin is used for more generally for preventative

measures, as well as for sepsis (Figure 1.1).18

Most aminoglycosides bind to the ribosomal RNA (rRNA) A-site, the site of mRNA decoding, and cause translation infidelity.20-22 The mode of action and resistance mechanisms have been well studied and the aminoglycoside scaffold has been established to bind RNA.23 With this as a starting point, derivatives

could lead to compounds that bind the A-site, and show activity against drug resistant bacteria by potentially evading resistance mechanisms. Additionally, modifications could possibly diminish toxicity effects. With this in mind, we have pursued the preparation and evaluation of minimally modified aminoglycoside in

order to test their A-site affinity and, importantly, evaluate their effectiveness as

potential antibiotics against many resistance bacterial strains.

Here we selectively modify two of the most common clinically used

aminoglycoside antibiotics, amikacin and tobramycin. The primary alcohol in the

6” position on these molecules is accessible to modification, so we substituted it

for a variety of hydrogen bond donors and acceptors of different sizes (Figure

1.1). Most of the compounds show an increase in vitro affinity to the A-site as determined by a fluorescence resonance energy transfer (FRET) binding assay.

Additionally, some of the derivatives show equal to or better potency against

certain resistant bacterial strains.

Figure 1.1: Tobramycin (1a), amikacin (2a) and derivatives prepared and studied. The 2- deoxystreptamine (2-DOS) ring is in pink. The 6’’ modification position is in green.

1.2 Results

Design Strategy

The 6’’ hydroxyl group is one of the few functional groups that appears to

form no hydrogen bonds to the A-site RNA, neither direct or water mediated, in the crystal structures of tobramycin (1a) and amikacin (2a), though both are in

close proximity to U1406 and C1407 (Figure 1.2).24,25 Analogs with guanidinium

groups replacing the 6’’ hydroxyl have shown to display increased A-site affinity

and in some cases superior antibacterial activity.26 This suggests that certain

modifications to the 6’’ position may show increased affinity for the A-site and desirable antibacterial efficacy. We set out to test this hypothesis by making derivatives of both 1a and 2a with a variety of substituents differing in size, basicity, and in number of hydrogen bond donors and acceptors. More basic functional groups could increase the overall positive charge of the analogs, creating favorable electrostatic interactions with the polyanionic A-site rRNA.

Hydrogen bond donors and acceptors could create new contacts to the A-site not observed in the parent compounds. Beyond imparting greater affinity for the A- site, some modifications could potentially lead to decreased recognition by aminoglycoside modifying enzymes, the most common form of aminoglycoside resistance, and therefore derivatives may exhibit greater antibacterial potency against resistant bacteria.

Figure 1.2: A) Crystal structure of tobramycin (1a) with A-site rRNA. B) Crystal structure of amikacin (2a) with A-site rRNA. RNA is in orange with U1406 and C1407 highlighted in green. Aminoglycosides are in magenta with 6’’ alcohols highlighted in light blue. Figures were adapted from PDB files: tobramycin (1LC4), amikacin (2GSQ).21

Synthesis

The parent aminoglycosides were converted into three key intermediates

using known procedures.26,27 The synthetic approach for the conversion of the

parent aminoglycosides into these intermediates is illustrated using tobramycin

(1a) as an example (Scheme 1.1). First, all amines were globally tert-

butyloxycarbonyl (Boc)-protected using di-tert-butyl dicarbonate. The single primary alcohol of (Boc)5tobramycin (3) was then selectively converted to a

sterically demanding sulfonate by treatment with 2,4,6-

triisopropylbenzenesulfonyl chloride (TPSCl) in pyridine. Reflux in methanolic

ammonia afforded 6’’-deoxy-6’’-amino(Boc)5tobramycin (5). Alternatively, the

TPS derivative could be converted to 6’’-deoxy-6’’-azido(Boc)5tobramycin (6) by treating it with sodium azide.

Scheme 1.1: Synthesis of key intermediates 4, 5, and 6. Reagents and conditions: a) Boc2O, Et3N, H2O, DMF, 55 °C; b) TPSCl, pyridine, RT; c) NH3, MeOH, 80 °C; d) NaN3, DMF, 55 °C.

6’’-Deoxy-6’’-triisopropylbenzylsulfonyl(Boc)5tobramycin (4) can also undergo substitution reactions with a variety of other nucleophiles (Scheme 1.2).

Reflux in ethanolic methylamine yielded 6’’-deoxy-6’’- methylamino(Boc)5tobramycin (7). Reflux with dimethylamine in tetrahydrofuran

(THF) and dimethylformamide (DMF) mixture gave 6’’-deoxy-6’’- dimethylamino(Boc)5tobramycin (8). 6’’-Deoxy-6’’-(2-(aminoethyl)amino)-

(Boc)7tobramycin (9) was obtained by heating with ethylene diamine in methanol,

followed by Boc protection using di-tert-butyl dicarbonate to facilitate purification of this intermediate.

Scheme 1.2: Substitution reactions of 6''-deoxy-6''-triisopropylbenzylsulfonyl(Boc)5tobramycin (4). Reagents and conditions: a) Methylamine, EtOH, 80 °C; b) Dimethylamine, THF, DMF, 80 °C; c) Ethylene diamine, MeOH, 80 °C; d) Boc2O, Et3N, H2O, DMF, 55 °C.

The free amine of 6’’-deoxy-6’’-amino(Boc)5tobramycin (5) was used

nucleophilically to react with 2,4-dimethoxybenzyl isocyanate in the presence of

pyridine to give a 2,4-dimethoxybenzyl (DMB) protected urea. The DMB and Boc

protecting groups were concurrently removed using a one to one mixture of

trifluoroacetic acid (TFA) and dichloromethane with a tri-iso-propyl silane (TIPS)

cation scavenger. HPLC purification afforded the analytically pure 6’’-deoxy-6’’-

ureidotobramycin (1f) (Scheme 1.3).

6’’-Deoxy-6’’-azido(Boc)5tobramycin (6) was used in a cycloaddition

reaction with propargyl (Boc)amine catalyzed by copper sulfate in the presence

of a sodium ascorbate reductant to give 6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-

triazol-1-yl)-(Boc)6tobramycin (10) (Scheme 1.4). The intermediates 5, 7, 8, 9,

and 10 were all be deprotected using the aforementioned acidic conditions and

HPLC purified to yield the tobramycin analogs 1b, 1c, 1d, 1e, and 1g. All the

amikacin derivatives were made using the same reagents as the tobramycin

analogs.

Scheme 1.3: Synthesis 6''-deoxy-6''-ureidotobramycin (1f). Reagents and conditions: a) 2, 4- Dimethoxybenzyl isocyanate, pyridine, RT; b) TFA, TIPS, CH2Cl2, RT.

Scheme 1.4: Synthesis 6''-deoxy-6''-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)-(Boc)6tobramycin (10). Reagents and conditions: a) Propargyl (Boc)amine, CuSO4·5 H2O, sodium ascorbate, THF, t-BuOH, H2O, RT.

Affinity for the bacterial 16S A-site RNA construct

To determine the affinity of all derivatives to the bacterial 16S A-site, we used a modified version of a FRET-based assay that was previously developed in our lab.28,29 This modified version has been previously used to measure A-site

affinities of modified aminoglycosides.26 It consists of an aminoglycoside-

coumarin conjugate (FRET donor), which binds to a Dy-547 labeled 16S or 18S

A-site RNA hairpin construct (FRET acceptor) (Figure 1.3). The affinity of

unlabeled ligands for the A-site can be measured in a competition experiment,

where the compound of interest is titrated in and displaces the coumarin–

aminoglycoside, resulting in a decreased emission of the FRET acceptor, Dy-

547. Different coumarin–aminoglycoside conjugates can be used to cover distinct

affinity ranges of A-site ligands. Plotting the fractional fluorescent saturation

versus compound concentration generates titration curves.

Figure 1.3: A) Kanamycin-coumarin and neomycin-coumarin conjugate (donors). Coumarin is colored in green B) Fluorescently labeled with Dy-547 (acceptor) 16S and 18S A-site RNA constructs. Dy-547 is colored in pink.

Amikacin has a much lower affinity to the A-site as compared to

tobramycin, so initial titrations on amikacin analogs were performed with a

coumarin–kanamycin derivative, the lowest affinity aminoglycoside conjugate

(Table 1.1). Tobramycin derivatives and higher affinity amikacin analogs were

titrated against a coumarin–neomycin derivative (Table 1.2). In all cases, binding curves were generated by plotting the fractional fluorescence saturation of the

FRET acceptor against the concentration of the molecule of interest.

Representative curves of kanamycin-coumarin and neomycin-coumarin are shown in Figure 1.4.

Table 1.1: IC50 Values for Competing Off Kanamycin-Coumarin. [a] Conditions: A-site RNA (1 μM), kanamycin-coumarin (0.53 μM), cacodylate buffer pH 7.0 (20 mM), NaCl (100 mM), EDTA (0.5 mM).

Compound IC50 (μM)

Tobramycin (1a) 1.5 ± 0.2

Amikacin (2a) 6.7 ± 0.7

6’’-Deoxy-6’’-aminoamikacin (2b) 2.1 ± 0.2

6’’-Deoxy-6’’-methylaminoamikacin (2c) 1.5 ± 0.2

6’’-Deoxy-6’’-dimethylaminoamikacin (2d) 2.2 ± 0.2

6’’-Deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e) 1.7 ± 0.03

6’’-Deoxy-6’’-ureidoamikacin (2f) 50.7 ± 5.5

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g) 2.2 ± 0.1

All amikacin derivatives showed improved A-site binding with the exception of 6’’-deoxy-6’’-ureidoamikacin (2f), which had a much lower affinity

than any other aminoglycoside tested. All amikacin analogs with modifications

containing a single amine moiety: 2b-e and 2g showed similar binding to each

other and were also comparable to tobramycin (1a). 6’’-Deoxy-6’’-(2-

(aminoethyl)amino)amikacin (2e) showed binding superior to any of the other

amikacin derivatives.

Figure 1.4: Representative displacement curves of A) Kanamycin-Coumarin by 2a (grey solid) and 2c (grey dashed), with IC50 values of 6.7 ± 0.7 and 1.5 ± 0.2, respectively. B) Neomycin- Coumarin by 1a (black solid) and 1b (black dashed) with IC50 values of 53.0 ± 6.0 and 4.7 ± 0.4, respectively.

All tobramycin analogs showed improved binding over tobramycin (1a).

Like the amikacin derivatives, the urea modification resulted in the weakest

binders. This urea tobramycin analog (1f) was the only one that was not superior to all of the amikacin derivatives. In contrast to the amikacin analogs, the tobramycin modifications showed substantial variability in their affinity for the A-

site. 6’’-Deoxy-6’’-aminotobramycin (1b) and 6’’-deoxy-6’’-(2-

(aminoethyl)amino)tobramycin (1e) showed the highest affinities of all derivatives

tested. The methylamino (1c) and dimethylamino (1d) modified derivatives were

the next best binders. 6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-

yl)tobramycin (1g) was worse than these, but still significantly better than the

urea modified analog (1f).

Table 1.2: IC50 Values for Competing Off Neomycin-Coumarin. ] Conditions: A-site RNA (1 μM), neomycin-coumarin (0.53 μM), cacodylate buffer pH 7.0 (20 mM), NaCl (100 mM), EDTA (0.5 mM)

Compound IC50 (μM)

Tobramycin (1a) 53.0 ± 6.0

6’’-Deoxy-6’’-aminotobramycin (1b) 4.7 ± 0.4

6’’-Deoxy-6’’-methylaminotobramycin (1c) 7.4 ± 0.6

6’’-Deoxy-6’’-dimethylaminotobramycin (1d) 6.8 ± 0.8

6’’-Deoxy-6’’-(2-(aminoethyl)amino)tobramycin (1e) 5.3 ± 0.5

6’’-Deoxy-6’’-ureidotobramycin (1f) 30.0 ± 4.0

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)tobramycin (1g) 9.8 ± 1.0

Amikacin (2a) >100

6’’-Deoxy-6’’-aminoamikacin (2b) 46.7 ± 1.5

6’’-Deoxy-6’’-methylaminoamikacin (2c) 45.7 ± 5.8

6’’-Deoxy-6’’-dimethylaminoamikacin (2d) 46.4 ± 5.4

6’’-Deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e) 20.2 ± 2.6

6’’-Deoxy-6’’-ureidoamikacin (2f) >100

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g) 47.6 ± 2.6

Antibacterial activities

To assess the relative antibacterial activities of the synthetic derivatives, minimum inhibitory concentration (MIC) values of both the modified and parent antibiotics were determined against an array of bacterial strains (Tables 1.3 and

1.4). Multiple gram positive and gram negative strains were chosen to establish a broad spectrum representation of antibacterial activity. The compounds were first tested against the antibacterial susceptible control E. coli strain ATCC25922. No derivatives showed improvement against this strain and only one compound, 6’’-

Deoxy-6’’-aminoamikacin (2b), even showed equal activity to its parent aminoglycoside with an MIC value of 6.25 – 12.5 µg/mL.

Table 1.3: Inhibitory activities of tobramycin (1a) and derivatives against a panel of bacterial strains (μg ml-1). Minimum inhibitory concentration (MIC) values [mg mL-1]. MIC value equal to tobramycin (italics); MIC value lower than tobramycin (bold).

Bacterial Strain 1a 1b 1c 1d 1e 1f 1g

E. coli 3.125 25-50 ≥50 ≥50 6.25-12.5 6.25 6.25 (ATCC25922)

P. aeruginosa 3.125- 3.125- 0.78 25 >50 >50 1.56 (P4) 6.25 6.25

P. aeruginosa 0.39 12.5 50 50 1.56 0.78 0.39 (PA01)

P. aeruginosa 0.78- 0.39 12.5-50 50 >50 3.125 0.78 (ATCC27853) 1.56

K. pneumoniae 6.25 12.5 25 25-50 6.25 12.5 12.5 (ATCC700603)

K. pneumoniae >50 >50 >50 >50 >50 >50 >50 (GNR1100)

MRSA 0.78- 6.25- 3.125- 12.5-25 25 6.25 6.25 (TCH1516) 1.56 12.5 6.25

MRSA 3.125- 0.78- 3.125 6.25 12.5-25 25 3.125 (ATCC33591) 6.25 1.56

MRSA >50 >50 >50 >50 >50 >50 >50 (Sanger 252)

The aminoglycosides were tested against three P. aeruginosa strains, P4,

PA01, and ATCC27853. Tobramycin (1a) shows much better activity than amikacin (2a) against these P. aeruginosa strains. Unfortunately, only one tobramycin derivative, 6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1- yl)tobramycin (1g), showed even equal activity to tobramycin (1a) against any of these strains. Both had MIC values of 0.39 µg/mL against PA01. However, the

amikacin derivatives, 6’’-deoxy-6’’-aminoamikacin (2b), 6’’-deoxy-6’’- methylaminoamikacin (2c), and 6’’-deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e)

showed improved activity. 6’’-Deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e) showed superior activity against all three strains including a four-fold improvement to 6.25 µg/mL against P4. 6’’-Deoxy-6’’-aminoamikacin (2b) was equal to amikacin (2a) with MIC values of 1.56 – 3.125 µg/mL against PA01, but was slightly improved against ATCC27853 with an MIC value of 1.56 – 3.125

µg/mL compared to a parent MIC value of 3.125 µg/mL. It also showed a four- fold improvement against P4.

The aminoglycosides were also tested against two K. pneumoniae strains,

ATCC700603 and the highly drug resistant, K. pneumoniae carbapenemase producer GNR1100. Amikacin (2a) shows better activity than tobramycin (1a) against these strains. Again, the tobramycin derivatives were disappointing with only 6’’-deoxy-6’’-(2-(aminoethyl)amino)tobramycin (1e) showing even equal activity to the parent. Both had MIC values of 6.25 µg/mL against ATCC700603.

6’’-Deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e) and 6’’-deoxy-6’’-(4-

(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g) showed equal activity to amikacin (2a) against ATCC700603 with MIC values of 0.781 µg/mL.

Interestingly, they both improved from 50 µg/mL to 12.5 – 25 µg/mL against

GNR1100 also.

Table 1.4: Inhibitory activities of amikacin (2a) and derivatives against a panel of bacterial strains (μg ml-1). Minimum inhibitory concentration (MIC) values [mg mL-1]. MIC value equal to amikacin (italics); MIC value lower than amikacin (bold).

Bacterial Strain 2a 2b 2c 2d 2e 2f 2g

E. coli 6.25 - 12.5 - 6.25 - 12.5 25 - 50 12.5 50 12.5 (ATCC25922) 12.5 25

P. aeruginosa 25 6.25 12.5 >50 6.25 >50 25 - 50 (P4)

P. aeruginosa 1.56 - 1.56 - 12.5 - 1.56 - 3.125 1.56 25 - 50 (PA01) 3.125 3.125 25 3.125

P. aeruginosa 1.56 - 3.125 - 3.125 25 1.56 25 - 50 3.125 (ATCC27853) 3.125 6.25

K. pneumoniae 0.781 - 6.25 - 0.781 1.56 3.125 0.781 0.781 (ATCC700603) 1.56 12.5

K. pneumoniae 12.5 - 12.5 - 50 50 >50 >50 >50 (GNR1100) 25 25

MRSA 12.5 - 12.5 - 6.25 - 12.5 12.5 - 25 50 >50 >50 (TCH1516) 25 25

MRSA 25 12.5 - 25 25 >50 12.5 >50 12.5 (ATCC33591)

MRSA 12.5 - 25 >50 >50 >50 25 - 50 >50 25 - 50 (Sanger 252)

To test efficacy against gram positive bacteria, the aminoglycosides were tested against MRSA strains TCH1516, ATCC33591, and Sanger 252. No amikacin or tobramycin derivatives showed any improvements or even equal activity to their parents against TCH1516 or Sanger 252. There were several compounds that showed improved activity against ATCC33591, however. 6’’-

Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)tobramycin (1g) improved to an

MIC value of 0.78 – 1.56 µg/mL from a parent value of 3.125 µg/mL. Several amikacin derivatives showed increased potency compared to the parent MIC value of 25 µg/mL for amikacin (2a). These included 6’’-deoxy-6’’-aminoamikacin

(2b), with a slight improvement to 12.5 – 25 µg/mL and 6’’-deoxy-6’’-(4-

(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g) and 6’’-deoxy-6’’-(2-

(aminoethyl)amino)amikacin (2e), which both had more significant improvements to 12.5 µg/mL.

1.3 Discussion

Tobramycin and amikacin analogs modified at the 6’’ position were

synthesized to evaluate their A-site affinities and antibacterial activity compared

to their parent compounds. All tobramycin analogs showed superior affinity for

the A-site as compared to tobramycin (1a). There was more variation in A-site

affinity among the tobramycin analogs compared to the amikacin derivatives. The

tightest binders were 6’’-deoxy-6’’-aminotobramycin (1b) and 6’’-deoxy-6’’-(2-

(aminoethyl)amino)tobramycin (1e) and the worst tobramycin analog was 6’’-

deoxy-6’’-ureidotobramycin (1f). The general trend among the tobramycin

analogs suggests that binders with smaller steric bulk or with greater overall

potential charge show higher affinity.

All amikacin analogs showed improved A-site binding with the exception of

6’’-deoxy-6’’-ureidoamikacin (2f), which had by far the lowest A-site affinity of any

compound tested. It is the only modification made without a basic functionality,

which likely contributed to its lack of affinity. The amikacin analogs with one

additional basic functional group showed similar IC50 values including the bulky

6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g). 6’’-Deoxy-6’’-

(2-(aminoethyl)amino)amikacin (2e) has two additional basic amines as

compared to amikacin (2a) and it was the highest affinity amikacin analog. In

contrast to the tobramycin analogs, amikacin analogs appear to exhibit affinities

based solely on electrostatic effects with no apparent steric preference among

the analogs tested.

When analyzing MIC values, it is important to remember that

aminoglycoside affinities to the A-site do not correlate with antibacterial

potency.30 Interestingly, all but one derivative, 6’’-deoxy-6’’-aminoamikacin (2b),

showed inferior antibacterial activity against the control E. coli strain

ATCC25922. This suggests that improvements in activity against resistant strains are at least partially due to overcoming bacterial resistance mechanisms.

The tobramycin analogs generally showed disappointing antibacterial

activity. The most successful analog was 6’’-deoxy-6’’-(4-(aminomethyl)-1H-

1,2,3-triazol-1-yl)tobramycin (1g). It showed better activity than tobramycin (1a)

against a MRSA strain and equal activity against one P. aeruginosa strain. In

most other cases its MIC values were two fold worse. This was also one of the

more successful modifications among the amikacin analogs. 6’’-Deoxy-6’’-(4-

(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g) showed equal or better activity than amikacin (2a) against five out of nine strains tested against and in all other cases its MIC was within one serial dilution. It is intriguing that this modification was so efficacious since it was the most structurally significant alteration made.

6’’-Deoxy-6’’-aminoamikacin (2b) was also promising with equal or improved activity against six out of nine strains, including all three P. aeruginosa strains. It is interesting to note that antibacterial activity was reduced across the entire panel for 6’’-deoxy-6’’-methylaminoamikacin (2c) and even more so for 6’’- deoxy-6’’-dimethylaminoamikacin (2d). This trend was also present in the tobramycin derivatives. This suggests hydrogen bonding may be playing a role in the increased activity of 6’’-deoxy-6’’-aminoamikacin (2b).

The most successful derivative made, however, was 6’’-deoxy-6’’-(2-

(aminoethyl)amino)- amikacin (2e). This compound showed increased activity against five strains and equal activity against one. It was universally better against the P. aeruginosa strains and it showed equal or better activity against both K. pneumoniae strains including an improvement against GNR1100. This makes the broad spectrum improvement of some of the amikacin derivatives particularly fascinating given that amikacin itself is a semi-synthetic aminoglycoside with an amino 2-hydroxybutyryl (AHB) side chain, which lowers its susceptibility to aminoglycoside-modifying enzymes.31 It is possible that the

AHB and 6’’ modifications operate synergistically to further decrease its affinity for these enzymes. This is a hypothesis that we have previously posited when we observed increased antibacterial activity in an analog with a guanidinium group in this position.26

1.4 Conclusion

A series of 6’’ modified tobramycin and amikacin analogs were

synthesized. In all cases the derivatives showed improved A-site affinity compared with their parent antibiotics when tested in an in vitro FRET-based assay with the exception of 6’’-deoxy-6’’-ureidoamikacin (2f), which showed greatly decreased binding. Tobramycin analogs generally showed disappointing antibacterial activity. In contrast, several amikacin analogs exhibited promising antibacterial potency. Most notably 6’’-deoxy-6’’-(2-(aminoethyl)amino)amikacin

(2e) showed greater potency than amikacin (2a) against the majority of strains that were tested in MIC assays.

1.5 Experimental

Materials

Unless otherwise specified, materials purchased from commercial suppliers were used without further purification. Tobramycin (1a) and amikacin

(2a) were obtained from Sigma–Aldrich as their free bases. Propargyl

(Boc)amine was synthesized according to an established procedure.32

Anhydrous NH3 was purchased from Airgas. All other anhydrous solvents and

reagents, and ion exchange resins were purchased from Sigma–Aldrich. NMR

solvents were purchased from Cambridge Isotope Laboratories (Andover, MA,

USA).

The Dy-547-labeled A-site construct was purchased from Thermo

Scientific and purified by gel electrophoresis. Kanamycin–coumarin and

neomycin–coumarin conjugates were synthesized and purified according to established procedures. Chemicals for preparing buffer solutions (enzyme grade)

were purchased from Fisher Biotech. Autoclaved water was used in all

fluorescence titrations.

Mueller–Hinton broth used for sensitivity testing was obtained from Hardy

Diagnostics (Santa Maria, CA, USA). Polystyrene 96-well microplates for MIC

testing were purchased from Corning Inc. (Corning, NY, USA). Bacterial strains

for sensitivity testing included five strains from the American Type Culture

Collection (Manassas, VA, USA): hospital-associated MRSA strain 33591

rendered resistant to rifampicin by serial passage, USA300 MRSA strain

TCH1516 (BAA-1717), K. pneumoniae strain 700603, P. aeruginosa strain

27853, and E. coli strain 25922. P. aeruginosa strain PA01 was used as a general antibiotic-sensitive P. aeruginosa strain.33 USA200 MRSA strain Sanger

252 was obtained from the Network on Antimicrobial Resistance in S. aureus

(NARSA) program supported under NIAID/NIH contract number

HHSN272200700055C. Other Gram-negative strains used were clinical isolates

obtained from a tertiary academic hospital in the New York metropolitan area;

these were: K. pneumoniae strain GNR1100 (respiratory isolate) and P.

aeruginosa strain P4 (sputum isolate).

Instrumentation

NMR spectra were recorded on Varian Mercury 300 and 400 MHz, Varian

VX 500 MHz, and Jeol ECA 500 MHz spectrometers. Mass spectra (MS) were

recorded at the University of California, San Diego Chemistry and Biochemistry

Mass Spectrometry Facility, utilizing an Agilent 6230 HR-ESI-TOF mass

spectrometer. Reverse-phase HPLC (Vydac C18 column) purification and

analysis were carried out using an Agilent 1200 series instrument. Products were

lyophilized utilizing a Labconco FreeZone 2.5 freeze drier. Steady-state

fluorescence experiments were carried out in a microfluorescence cell with a

path length of 1.0 cm (Hellma GmH & Co KG, Mullenheim, Germany) on a Jobin

Yvon Horiba FluoroMax-3 luminescence spectrometer. A background spectrum

(buffer) was subtracted from each sample. A VersaMax plate reader (Molecular

Devices, Mountain View, CA, USA) set at 600 nm wavelength was used for MIC

assays.

Synthesis

(Boc)5tobramycin (3), 6’’-deoxy-6’’-triisopropylbenzylsulfonyl(Boc)5- tobramycin (4), 6’’-deoxy-6’’-amino(Boc)5tobramycin (5), (Boc)4amikacin (11), 6’’- deoxy-6’’-triisopropylbenzylsulfonyl(Boc)4amikacin (12), and 6’’-deoxy-6’’-

26,27 amino(Boc)4amikacin (13) were previously synthesized.

Tobramycin Derivatives

6’’-Deoxy-6’’-azido-(Boc)5tobramycin (6). DMF (8.3 mL) was added to 6’’-

27 deoxy-6’’-triisopropylbenzylsulfonyl-(Boc)5tobramycin (4) (818 mg, 0.663

mmol). Sodium azide (344 mg, 5.30 mmol) was added. The yellow solution was

heated to 65 °C and stirred for 1 day. The solvent was removed under reduced

pressure and the resulting solid was dissolved in DCM and washed with water.

The organic layers were dried with sodium sulfate and the solvent was removed

under reduced pressure. The product was isolated by flash chromatography (3.5,

4, and 5% methanol in DCM). Product: White solid (469 mg, 0.472 mmol, 71%

1 yield). H NMR (400 MHz, CD3OD): δ 5.10 (brs, 1H), 5.09 (brs, 1H), 4.21 – 4.10

(m, 1H), 3.81 – 3.27 (m, 15H), 2.19 – 1.95 (m, 2H), 1.65 (q, J = 12 Hz, 1H), 1.60

+ – 1.20 (m, 46H); HR-ESI-MS calculated for C43H76N8O18Na [M+Na] 1015.5159,

found 1015.5161.

6’’-Deoxy-6’’-aminotobramycin (1b). DCM (3.0 mL) and TIPS (180 µL) were

added to 6’’-Deoxy-6’’-amino-(Boc)5tobramycin (5) (180 mg, 0.182 mmol). TFA

(3.0 mL) was added. The yellow solution was stirred for 2.5 hours. The solvent

was removed under reduced pressure. The remaining white solid was dissolved

in water and purified by reverse phase HPLC [0 – 1.5% ACN in water (0.1% TFA)

over 7 mins, flow rate is 3 mL / min, eluted after 5.6 min], then lyophilized and

desalted. Product: White solid (65.1 mg, 0.140 mmol, 75% yield). 1H NMR (500

MHz, CD3OD): δ 5.14 (d, J = 4.0 Hz, 1H), 5.02 (d, J = 4 Hz, 1H), 3.85 (m, 1H),

3.62 – 3.58 (m, 2H), 3.52 – 3.46 (m, 2H), 3.30 (t, J = 9.5 Hz, 1H), 3.22 (t, J = 5

Hz, 1H), 3.19 (t, J = 10 Hz, 1H), 3.00 – 2.92 (m, 4H), 2.89 – 2.82 (m, 2H), 2.74

(td, J1 = 13.5 Hz, J2 = 7 Hz, 2H), 2.00 (dt, J1= 12 Hz, J2 = 4.5 Hz, 1H), 1.92 (dt,

J1= 13 Hz, J2 = 4.5 Hz, 1H), 1.59 (q, J = 12 Hz, 1H), 1.20 (q, J = 12.5 Hz, 1H);

13 C NMR (125 MHz, D2O): δ 100.74, 100.24, 88.51, 87.20, 75.00, 74.01, 72.46,

72.38, 71.35, 66.80, 54.71, 51.27, 50.00, 49.70, 42.11, 41.72, 36.06, 35.45; HR-

+ ESI-MS calculated for C18H39N6O8 [M+H] 467.2828, found 467.2828.

6’’-Deoxy-6”-methylamino-(Boc)5tobramycin (7). A solution of 33%

methylamine in ethanol (10 mL) was added to 6’’-deoxy-6’’-

27 triisopropylbenzylsulfonyl-(Boc)5tobramycin (4) (0.143 g, 0.116 mmol) in a

pressure tube. The vessel was capped and heated to 80 °C overnight. The

vessel was cooled to 0 °C and opened, and let warm to room temperature. The

solvent was removed under reduced pressure. The remaining residue was

redissolved in methanol (5 mL) and DOWEX® Monosphere® 550A ion exchange

resin (OH– form) was added. The reaction was stirred for 12 hours at rt and filtered. The solvent was removed under reduced pressure. The product was

isolated by flash chromatography (7% methanol, then 12% methanol and 0.5%

TEA in DCM). Product: White solid (0.100 mg, 0.102 mmol, 88% yield). 1H-NMR

(300 MHz, CD3OD): δ 5.16 (brs, 2H), 4.33 – 4.22 (m, 1H), 3.71 (t, J=10.2 Hz,

1H), 3.66 – 3.33 (m, 11H), 3.21 (t, J = 9.8 Hz, 1H) 3.14– 3.07 (m, 1H), 2.86 (p, J

= 6.9 Hz, 1H), 2.77 (s, 3H), 2.02 (brs, 2H), 1.65 (q, J = 12.4 Hz, 1H), 1.55 – 1.35

+ (m, 46H). HR-ESI-MS calculated for C44H81N6O18 [M+H] 981.5602, found

981.5598.

6’’-Deoxy-6’’-methylaminotobramycin (1c). DCM (1.7 mL) and TIPS (0.10 mL)

were added to 6’’-Deoxy-6’’-methylamino-(Boc)5tobramycin (7) (100 mg, 0.101

mmol). TFA (1.7 mL) was added. The yellow solution was stirred for 2.5 hours.

The solvent was removed under reduced pressure. The remaining white solid

was dissolved in water and purified by reverse phase HPLC [0 – 0.1% ACN in

water (0.1% TFA) over 10 mins, flow rate is 3 mL / min, eluted after 5.3 min],

then lyophilized and desalted. Product: White solid (21.5 mg, 0.045 mmol, 44%

1 yield). H NMR (500 MHz, D2O): δ 5.13 (d, J = 4.0 Hz, 1H), 5.03 (d, J = 4.0 Hz,

1H), 4.00 (m, 1H), 3.64 – 3.58 (m, 2H), 3.55 – 3.46 (m, 2H), 3.31 (t, J = 9.5 Hz,

1H), 3.24 (t, J = 9.5 Hz, 1H), 3.18 (t, J = 9.5 Hz, 1H) , 3.03 – 2.93 (m, 4H), 2.92 –

2.82 (m, 2H), 2.81 – 2.72 (m, 2H), 2.40 (s, 3H), 2.00 (dt, J1= 11.5 Hz, J2 = 4.5

Hz, 1H), 1.92 (dt, J1= 13 Hz, J2 = 4.5 Hz, 1H), 1.59 (q, J = 11.5 Hz, 1H), 1.20 (q,

13 J = 13 Hz, 1H); C NMR (125 MHz, D2O): δ 100.57, 88.31, 87.65, 75.04, 72.38,

71.96, 70.30, 66.78, 54.68, 51.70, 51.28, 49.99, 49.86, 49.71, 42.13, 36.08,

+ 35.51, 35.26; HR-ESI-MS calculated for C19H41N6O8 [M+H] 481.2980, found

481.2983.

6’’-Deoxy-6’’-dimethylamino-(Boc)5tobramycin (8). Anhydrous DMF (4 mL)

27 was added to 6’’-deoxy-6’’-triisopropylbenzylsulfonyl-(Boc)5tobramycin (4) (320

mg, 0.26 mmol) in a pressure tube. A 2 M solution of dimethylamine in THF (4

mL) was added. The vessel was capped and heated to 80 °C overnight. The

vessel was cooled to 0 °C and opened. The solvent was removed under reduced

pressure. The remaining residue was redissolved in methanol (6.2 mL) and

DOWEX® Monosphere® 550A ion exchange resin (-OH form) was added. The

reaction was stirred for 12 hours at rt and filtered. The solvent was removed

under reduced pressure. Product: White solid (235 mg, 0.246 mmol, 91% yield).

1 H NMR (400 MHz, CD3OD): δ 5.28 (br s, 1H), 5.05 (br s, 1H), 4.10 – 3.96 (m,

1H), 3.77 – 3.28 (m, 11H), 3.16 – 3.07 (m, 1H), 2.65 – 2. 56 (m, 1H), 2.53 – 2.41

(m, 1H), 2.32 (s, 6H), 2.46 (t, J = 7.6 Hz, 1H), 2.17 – 1.90 (m, 2H), 1.74 – 1.22

+ (m, 47H); HR-ESI-MS calculated for C45H83N6O18 [M+H] 995.5758, found

995.5756.

6’’-Deoxy-6’’-dimethylaminotobramycin (1d). DCM (3.3 mL) and TIPS (199

µL) were added to 6’’-Deoxy-6’’-dimethylamino-(Boc)5tobramycin (8) (198 mg,

0.200 mmol). TFA (3.3 mL) was added. The yellow solution was stirred for 2.5

hours. The solvent was removed under reduced pressure. The remaining white

solid was dissolved in water and purified by reverse phase HPLC [0 – 1.5% ACN

in water (0.1% TFA) over 7 mins, flow rate is 3 mL / min) eluted after 5.3 min],

then lyophilized and desalted. Product: White solid (72.8 mg, 0.147 mmol, 74%

1 yield) H NMR (500 MHz, D2O): δ 5.10 (d, J = 3.5 Hz, 1H), 5.05 (d, J = 4 Hz,

1H), 4.02 – 3.96 (m, 1H), 3.67 – 3.60 (m, 2H), 3.59 – 3.52 (m, 1H), 3.51 (dd, J1 =

10 Hz, J2 = 4 Hz, 1H), 3.31 (t, J = 9.5 Hz, 1H), 3.28 (t, J = 9.5 Hz, 1H), 3.14 (t, J =

9.5 Hz, 1H), 3.06 – 2.94 (m, 3H), 2.94 – 2.84 (m, 2H), 2.76 (dd, J1 = 14 Hz, J2 =

7.5 Hz, 1H), 2.68 (dd, J1 = 16 Hz, J2 = 2 Hz, 1H), 2.56 (dd, J1 = 13.5 Hz, J2 = 8.5

Hz, 1H), 2.28 (s, 6H), 2.05 (dt, J1 = 12 Hz, J2 = 4.5 Hz, 1H), 1.97 (dt, J1 = 13.5

13 Hz, J2 = 4.5 Hz, 1H), 1.62 (q, J = 12 Hz, 1H); 1.24 (q, J = 12.5 Hz, 1H); C NMR

(125 MHz, D2O): δ 101.02, 100.46, 88.67, 88.08, 75.24, 74.30, 72.67, 72.49,

70.11, 66.84, 60.87, 54.70, 51.14, 50.29, 49.89, 45.65, 42.22, 36.21, 35.35; HR-

+ ESI-MS calculated for C20H43N6O8 [M+H] 495.3139, found 495.3139.

6’’-Deoxy-6’’-(2-(aminoethyl)amino)-(Boc)5tobramycin (9). Anhydrous

methanol (15 mL) was added to 6’’-deoxy-6’’-triisopropylbenzylsulfonyl-

27 (Boc)5tobramaycin (4) (0.170 g, 0.138 mmol) in a pressure tube. Ethylene diamine (0.4 mL, 6 mmol) was added. The vessel was capped and heated to 80

°C for 2 days. The vessel was cooled to 0 °C and opened. The solvent was removed under reduced pressure. The remaining residue was suspended in toluene and the solvent was removed under reduced pressure. This was repeated three more times. DCM (17.5 mL) and methanol (1 mL) were added to the remaining pale yellow solid. TEA (0.66 mL, 4.8 mmol) and di-tert-butyl dicarbonate (2.62 g, 12 mmol) were added. The orange solution was stirred overnight. The solvent was removed under reduced pressure. The product was isolated by automated flash chromatography (15 - 80% ethyl acetate in hexanes over 18 mins) eluted after 13 min. Product: Light yellow solid (96.6 mg, 0.080

1 mmol, 58% yield). H NMR (500 MHz, CD3OD): δ 5.17 (br s, 1H), 5.10 (br s, 1H),

4.14 – 4.06 (m, 1H), 3.85 – 3.07 (m, 19H), 2.14 – 1.95 (m, 2H), 1.72 – 1.60 (m,

+ 1H), 1.54 – 1.37 (m, 64H); HR-ESI-MS calculated for C55H99N7O22Na [M+Na]

1232.6735, found 1232.6733.

6’’-Deoxy-6’’-(2-(aminoethyl)amino)tobramycin (1e). DCM (1.68 mL) and TIPS

(120 µL) were added to 6’’-Deoxy-6’’-(2-(aminoethyl)amino)-(Boc)5tobramycin (9)

(96.6 mg, 0.16 mmol). TFA (1.68 mL) was added. The yellow solution was stirred

for 2.5 hours. The solvent was removed under reduced pressure. The remaining

white solid was dissolved in water and purified by reverse phase HPLC [0 – 0.8%

ACN in water (0.1% TFA) over 8 mins, flow rate is 3 mL / min), eluted after 5.0

min], then lyophilized and desalted. Product: Light yellow solid (16.1 mg, 0.032

1 mmol, 40% yield). H NMR (500 MHz, D2O): δ 5.07 – 5.03 (m, 1H), 5.00 – 4.94

(m, 1H), 3.95 – 3.83 (m, 1H), 3.62 –3.50 (m, 2H), 3.50 – 3.38 (m, 2H), 3.27 –

3.20 (m, 1H), 3.20 – 3.14 (m, 1H), 3.14 – 3.07 (m, 1H), 2. 97 – 2.95 (m, 6H), 2.69

– 2.52 (m, 6H), 2.21 – 2.11 (m, 1H), 1.92 – 1.84 (m, 1H), 1.60 – 1.49 (m, 1H),

13 1.22 – 1,09 (m, 1H); C NMR (125 MHz, D2O): δ 133.30, 129.84, 100.70, 88.52,

87.91, 74.94, 74.53, 72.46, 72.24, 71.19, 66.79, 52.64, 51.63, 51.29, 50.17,

50.07, 49.79, 42.24, 40.34, 36.14, 35.57; HR-ESI-MS calculated for C20H44N7O8

[M+H]+ 510.3046, found 510.3243.

6’’-Deoxy-6’’-ureidotobramycin (1f). Anhydrous pyridine (1.78 mL) was added

26 to 6’’-deoxy-6’’-amino-(Boc)5tobramycin (5) (200 mg, 0.207 mmol).

Dimethoxybenzyl isocyanate (59 µL, 0.197 mmol) was added. The yellow

solution was wrapped in tinfoil and stirred overnight. The solvent was removed

under reduced pressure. The product was isolated by flash chromatography (6%

methanol in DCM). The pale yellow solid product was carried on without further

purification. DCM (2.24 mL) and TIPS (134 µL) were added to the crude product.

TFA (2.24 mL) was added. The yellow solution was stirred for 3.5 hours. The

solvent was removed under reduced pressure. The remaining white solid was

dissolved in water and purified by reverse phase HPLC [0 – 1% ACN in water

(0.1% TFA) over 7 min, eluted after 5.3 min], then lyophilized and desalted.

1 Product: White solid (53 mg, 0.104 mmol, 53%). H NMR (500 MHz, D2O): δ 5.13

(d, J = 3.5 Hz, 1H), 4.98 (d, J = 4.0 Hz, 1H), 3.94 – 3.87 (m, 1H), 3.61 – 3.52 (m,

2H), 3.51 – 3.44 (m, 2H), 3.40 (dd, J1 = 14 Hz, J2 = 3 Hz, 1H), 3.31 – 3.22 (m,

2H), 3.19 (t, J = 10 Hz, 1H), 3.13 (t, J = 10 Hz, 1H), 3.00 – 2.88 (m, 3H), 2.88 –

2.79 (m, 2H), 2.72 (dd, J1 = 14 Hz, J2 = 7.5 Hz, 1H), 1.99 (dd, J1 = 12 Hz, J2 = 4.5

Hz, 1H), 1.89 (dd, J1 = 13 Hz, J2 = 4 Hz, 1H), 1.57 (q, J = 12 Hz, 1H), 1.17 (q, J =

13 12.5 Hz, 1H); C NMR (125 MHz, D2O): δ 162.21, 100.71, 100.10, 88.49, 87.07,

74.83, 74.03, 72.35, 71.46, 71.17, 88.82, 54.48, 51.18, 50.01, 49.71, 42.11,

+ 40.93, 35.97, 35.44; HR-ESI-MS calculated for C19H39N7O9Na [M+Na] 532.2704,

found 532.2701

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)-(Boc)6tobramycin (10).

A 3:1:1 of THF: tert-butanol: water solution (2.94 mL) was added to 6’’-Deoxy-6’’-

azido-(Boc)5tobramycin (6) (86.6 mg, 0.148 mmol) and N-Boc-propargylamine

(27.2mg, 0.174 mmol). The solution was degassed by bubbling through argon for

25 minutes. A 1 M aqueous sodium ascorbate solution (94 µL) was added. Then a 7.5% weight / volume aqueous copper sulfate pentahydrate solution (38 µL) was added. The solution turned brown and then yellow overnight. The solvent was removed under reduced pressure. The product was isolated by flash chromatography (5% methanol in DCM). Product: Light yellow solid (80.7 mg,

1 0.090 mmol, 81% yield). H NMR (400 MHz, CD3OD): δ 7.92 (s, 1H), 5.23 (s,

2H), 4.76 – 4.22 (m, 5H), 3.82 – 3.22 (m, 12H), 3.00 (t, J = 9.6Hz, 1H), 2.01 (brs,

2H), 2.68 (q, J = 12Hz, 1H), 1.60 – 1.20 (m, 55H); HR-ESI-MS calculated for

+ C51H89N9O20Na [M+Na] 1170.6116, found 1170.6112.

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)tobramycin (1g). DCM

(1.8 mL) and TIPS (90 μL) were added to 6’’-Deoxy-6’’-(4-(aminomethyl)-1H-

1,2,3-triazol-1-yl)-(Boc)6tobramycin (10) (62 mg, 0.054 mmol). TFA (1.8 mL) was

added. The yellow solution was stirred for 2.5 hours. The solvent was removed

under reduced pressure. The remaining white solid was dissolved in water and

purified by reverse phase HPLC [0 – 0.1% ACN in water (0.1% TFA) over 6.5

mins, eluted after 5.6 min], then lyophilized and desalted. Product: White solid

1 (27.5 mg, 0.050 mmol, 82% yield). H NMR (500 MHz, D2O): δ 7.85 (s, 1H), 5.06

(d, J = 2.5 Hz, 1H), 4.98 (d, J = 3.0 Hz, 1H), 4.62 (d, J = 3 Hz, 1H), 4.26 – 4.22

(m, 1H), 3.82 (s, 2H), 3.58 – 3.52 (m, 1H), 3.51 – 3.44 (m, 1H), 3.35 (dd, J2 = 3

Hz, J1 = 10 Hz, 1H), 3.21 (t, J = 9.5 Hz, 1H), 3.14 (t, J = 9.5 Hz, 1H), 3.01 – 2.87

(m, 4H), 2.83 – 2.74 (m, 2H), 2.74 – 2.65 (m, 1H), 2.04 – 1.97 (m, 1H), 1.91 –

1.83 (m, 1H), 1.56 (q, J = 11.5 Hz, 1H), 1.14 (q, J = 12.5 Hz, 1H); 13C NMR (125

MHz, D2O): δ 148.86, 125.17, 100.51, 100.10, 88.05, 86.93, 74.77, 74.09, 72.16,

70.93, 70.73, 66.81, 54.68, 51.29, 51.16, 49.96, 49.66, 42.15, 36.26, 36.03,

+ 35.53; HR-ESI-MS calculated for C21H41N9O8Na [M+Na] 570.2970, found

570.2971.

Amikacin Derivatives

Scheme 1.5: Synthesis of key intermediates 12, 13, and 14. Reagents and conditions: a) Boc2O, Et3N, H2O, DMF, 55 °C; b) TPSCl, pyridine, RT; c) NH3, MeOH, 80 °C; d) NaN3, DMF, 55 °C

6’’-Deoxy-6’’-azido-(Boc)4amikacin (14). DMF (48.6 mL) was added to 6’’-

26 deoxy-6’’-triisopropylbenzylsulfonyl-(Boc)4amikacin (12) (2.5 g, 2 mmol).

Sodium azide (520 mg, 8 mmol) was added. The yellow solution was heated to

55 °C and stirred for 2 days. The solvent was removed under reduced pressure and the resulting solid was dissolved in DCM and washed with water. The organic layers were dried with sodium sulfate and the solvent was removed under reduced pressure. The product was isolated by automated flash chromatography (0 - 20% methanol in DCM over 25 mins) eluted after 16 min.

1 Product: White solid (1.7 g, 1.7 mmol, 85% yield). H NMR (500 MHz, CD3OD): δ

5.09 (d, J = 2 Hz, 1H), 5.01 (d, J = 3.5 Hz, 1H), 4.13 – 4.08 (m, 1H), 4.01 – 3.93

(m, 1H), 3.68 – 3.57 (m, 4H), 3.51 – 3.34 (m, 6H), 3.28 – 3.10 (m, 8 H), 2.06 –

2.00 (m, 2H), 1.94 – 1.86 (m, 1H), 1.81 – 1.77 (m, 1H), 1.47 – 1.37 (m, 36H);

+ HR-ESI-MS calculated for C50H87N9O22Na [M+Na] 1188.5858, found 1188.5859.

Scheme 1.6: Synthesis of 6''-deoxy-6''-aminoamikacin (2b). Reagents and conditions: a) TFA, TIPS, CH2Cl2, RT

6’’-Deoxy-6’’-aminoamikacin (2b). DCM (3.9 mL) and TIPS (0.23 mL) were

26 added to 6’’-deoxy-6’’-amino-(Boc)4amikacin (13) (160 mg, 0.16 mmol). TFA

(3.9 mL) was added. The yellow solution was stirred for 2.5 hours. The solvent

was removed under reduced pressure. The remaining white solid was dissolved

in water and purified by reverse phase HPLC [0 – 0.1% ACN in water (0.1% TFA)

over 10 mins, flow rate is 2 mL / min] eluted after 7.8 min, then lyophilized and

desalted. Product: White solid (82 mg, 0.14 mmol, 86% yield). 1H NMR (400

MHz, CD3OD): δ 5.28 (d, J = 4.0 Hz, 1H), 5.05 (d, J = 3.6 Hz, 1H), 4.15 (dd, J1 =

9.2 Hz, J2 = 3.6 Hz, 1H), 4.01 – 3.89 (m, 1H), 3.77 – 3.66 (m, 3H), 3.57 (dd, J1 =

9.6 Hz, J2 = 4.0 Hz, 1H), 3.36 – 3.27 (m, 3H), 3.16 (t, J = 9.6 Hz, 2H), 2.98 – 2.84

(m, 5H), 2.76 – 2.67 (m, 3H), 1.95 – 1.82 (m, 2H), 1.74 – 1.64 (m, 1H), 1.39 (q, J

13 = 12.4 Hz, 1H); C NMR (125 MHz, D2O): δ 174.89, 97.92, 96.19, 85.10, 77.96,

72.58, 71.14, 71.10, 70.27, 70.09, 69.80, 69.12, 68.89, 67.99, 52.12, 47.80,

46.69, 39.60, 39.37, 35.41, 34.45, 32.36; HR-ESI-MS calculated for

+ C22H44N6O12Na [M+Na] 607.2909, found 607.2914.

Scheme 1.7: Synthesis of 6''-Deoxy-6''-methylaminoamikacin (2c). Reagents and conditions: a) Methylamine, EtOH, 80 °C; b) TFA, TIPS, CH2Cl2, RT.

6’’-Deoxy-6’’-methylamino-(Boc)4amikacin (15). A solution of 33%

methylamine in ethanol (22 mL) was added to 6’’-deoxy-6’’-

26 triisopropylbenzylsulfonyl-(Boc)4amikacin (12) (320 mg, 0.26 mmol) in a

pressure tube. The vessel was capped and heated to 80 °C overnight. The

vessel was cooled to 0 °C and opened. The solvent was removed under reduced

pressure. The remaining residue was redissolved in methanol (15 mL) and

DOWEX® Monosphere® 550A ion exchange resin (OH– form) was added. The

reaction was stirred for 12 hours at rt and filtered. The solvent was removed

under reduced pressure. The product was isolated by flash chromatography

(10% methanol, 1% TEA in DCM). Product: White solid (230 mg, 0.23 mmol,

1 90% yield). H NMR (400 MHz, CD3OD): δ 5.08 (br s, 1H), 5.00 (s, 1H), 4.14 –

4.06 (m, 1H), 3.99 – 3.93 (m, 1H), 3.84 – 3.75 (m, 1H), 3.75 – 3.39 (m, 5H), 3.20

– 3.05 (m, 3H), 2.99 – 2.52 (m, 6H), 2.40 – 2.30 (m, 3H), 2.25 (s, 3H), 1.95 –

1.87 (m, 2H), 1.77 – 1.65 (m, 1H), 1.41 (br s, 37H); HR-ESI-MS calculated for

+ C43H78N6O20Na [M+Na] 1021.5163, found 1021.5165.

6’’-Deoxy-6’’-methylaminoamikacin (2c). DCM (2.1 mL) and TIPS (0.13 mL)

were added to 6’’-Deoxy-6’’-methylamino-(Boc)4amikacin (15) (85 mg, 0.085

mmol). TFA (2.1 mL) was added. The yellow solution was stirred for 2.5 hours.

The solvent was removed under reduced pressure. The remaining white solid

was dissolved in water and purified by reverse phase HPLC [0 – 0.1% ACN in

water (0.1% TFA) over 10 mins, flow rate is 2 mL / min, eluted after 8.0 min],

then lyophilized and desalted. Product: White solid (40 mg, 0.067 mmol, 79%

1 yield). H NMR (500 MHz, D2O): δ 5.28 (d, J = 4.0 Hz, 1H), 5.03 (d, J = 3.5 Hz,

1H), 4.13 (dd, J1 = 9.3 Hz, J2 = 3.5 Hz, 1H), 4.03 – 3.92 (m, 2H), 3.72 – 3.65 (m,

4H), 3.57 (dd, J1 = 9.8 Hz, J2 = 4.0 Hz, 1H), 3.34 – 3.26 (m, 3H), 3.15 – 3.07 (m,

1H), 2.96 – 2.87 (m, 3H), 2.81 – 2.79 (m, 1H), 2.75 – 2.60 (m, 4H), 2.29 (s, 3H),

1.92 – 1.83 (m, 2H), 1.71 – 1.65 (m, 1H), 1.38 (q, J = 12.5 Hz, 1H); 13C NMR

(125 MHz, D2O): δ 174.77, 98.06, 96.17, 85.15, 77.81, 72.84, 71.29, 71.10,

70.16, 69.85, 69.83, 69.19, 68.60, 68.09, 52.17, 49.72, 47.93, 46.95, 39.74,

+ 35.51, 34.37, 33.26, 32.49; HR-ESI-MS calculated for C23H47N6O12 [M+H]

599.3246, found 599.3245.

Scheme 1.8: Synthesis of 6''-deoxy-6''-dimethylaminoamikacin (2d). Reagents and conditions: a) Dimethylamine, THF, DMF, 80 °C; b) TFA, TIPS, CH2Cl2, RT.

6’’-Deoxy-6’’-dimethylamino-(Boc)4amikacin (16). Anhydrous DMF (3.1 mL)

26 was added to 6’’-deoxy-6’’-triisopropylbenzylsulfonyl-(Boc)4amikacin (12) (320

mg, 0.26 mmol) in a pressure tube. A 2 M solution of dimethylamine in THF (3.1

mL) was added. The vessel was capped and heated to 80 °C overnight. The

vessel was cooled to 0 °C and opened. The solvent was removed under reduced

pressure. The remaining residue was redissolved in methanol (6.2 mL) and

DOWEX® Monosphere® 550A ion exchange resin (OH– form) was added. The

reaction was stirred for 12 hours at rt and filtered. The solvent was removed

under reduced pressure. The product was isolated by flash chromatography

(10% methanol, 1% TEA in DCM). Product: White solid (160 mg, 0.16 mmol,

1 62% yield). H NMR (400 MHz, CD3OD): δ 5.08 (br s, 2H), 3.99 (dd, J1 = 8.4 Hz,

J2 = 3.6 Hz, 1H), 3.84 – 3.78 (m, 1H), 3.72 – 3.56 (m, 3H), 3.52 – 3.34 (m, 3H),

3.24 – 3.16 (m, 2H), 3.07 – 2.82 (m, 9H), 2.69 (s, 6H), 2.46 (t, J = 7.6 Hz, 1H),

1.85 – 1.77 (m, 2H), 1.50 – 1.41 (m, 38H); HR-ESI-MS calculated for

+ C44H80N6O20Na [M+Na] 1035.5320, found 1035.5322.

6’’-Deoxy-6’’-dimethylaminoamikacin (2d). DCM (2.1 mL) and TIPS (0.13 mL) were added to 6’’-Deoxy-6’’-dimethylamino-(Boc)4amikacin (16) (85 mg, 0.085

mmol). TFA (2.1 mL) was added. The yellow solution was stirred for 2.5 hours.

The solvent was removed under reduced pressure. The remaining white solid

was dissolved in water and purified by reverse phase HPLC [0 – 0.1% ACN in

water (0.1% TFA) over 14 mins, flow rate is 2 mL / min, eluted after 7.9 min],

then lyophilized and desalted. Product: White solid (46 mg, 0.076 mmol, 89%

1 yield) H NMR (300 MHz, D2O): δ 5.26 (d, J = 3.9 Hz, 1H), 5.03 (d, J = 3.9 Hz,

1H), 4.12 (dd, J1 = 9.3 Hz, J2 = 3.8 Hz, 1H), 4.07 – 3.89 (m, 2H), 3.76 – 3.62 (m,

4H), 3.53 (dd, J1 = 9.6 Hz, J2 = 4.2 Hz, 1H), 3.34 – 3.20 (m, 3H), 3.07 – 2.84 (m,

4H), 2.76 – 2.43 (m, 5H), 2.21 (s, 6H), 1.95 – 1.79 (m, 2H), 1.74 – 1.64 (m, 1H),

13 1.35 (q, J = 12.6 Hz, 1H); C NMR (125 MHz, D2O): δ 174.13, 97.92, 95.68,

84.63, 77.32, 72.61, 70.89, 70.77, 70.01, 69.87, 69.39, 68.78, 67.68, 67.03,

58.21, 51.60, 47.49, 46.85, 39.35, 36.64, 35.03, 33.14, 32.85, 32.06; HR-ESI-MS

+ calculated for C24H49N6O12 [M+H] 613.3403, found 613.3407.

Scheme 1.9: Synthesis of 6''-deoxy-6''-(2-(aminoethyl)amino)amikacin (2e). Reagents and conditions: a) Ethylene diamine, MeOH, 80 °C; a) Boc2O, Et3N, H2O, DMF, 55 °C; c) TFA, TIPS, CH2Cl2, RT.

6’’-Deoxy-6’’-(2-(aminoethyl)amino)-(Boc)6amikacin (17). Anhydrous methanol

(75 mL) was added to 6’’-deoxy-6’’-triisopropylbenzylsulfonyl-(Boc)4amikacin

(12)26 (1.5 g, 1.2 mmol) in a pressure tube. Ethylene diamine (4 mL, 60 mmol)

was added. . The vessel was capped and heated to 80 °C for 2 days. The vessel

was cooled to 0 °C and opened. The solvent was removed under reduced

pressure. The remaining residue was suspended in toluene and the solvent was

removed under reduced pressure. This was repeated three more times. DCM

(17.5 mL) and methanol (1 mL) were added to the remaining pale yellow solid.

TEA (0.66 mL, 4.8 mmol) and di-tert-butyl dicarbonate (2.62 g, 12 mmol) were added. The orange solution was stirred overnight. The solvent was removed under reduced pressure. The product was isolated by automated flash chromatography (0 – 20% methanol in DCM over 30 mins) eluted after 13 min.

Product: Light yellow solid (905 mg, 0.74 mmol, 61% yield). 1H NMR (400 MHz,

CD3OD): δ 5.09 (br s, 1H), 5.01 (d, J = 3.6 Hz, 1H), 4.19 – 4.11 (m, 1H), 3.99

(dd, J1 = 9.0 Hz, J2 = 3.8 Hz, 1H), 3.84 (t, J = 5.4 Hz, 1H), 3.77 – 3.60 (m, 5H),

3.51 – 3.32 (m, 6H), 3.25 – 3.08 (m, 4H), 3.00 – 2.63 (m, 6H), 2.16 – 2.07 (m,

1H), 1.99 – 1.91 (m, 1H), 1.81 – 1.72 (m, 1H), 1.63 – 1.55 (m, 1H), 1.51 – 1.25

+ (m, 54H); HR-ESI-MS calculated for C54H97N7O24Na [M+Na] 1250.6477, found

1250.6483.

6’’-Deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e). DCM (3.9 mL) and TIPS

(0.23 mL) were added to 6’’-Deoxy-6’’-(2-(aminoethyl)amino)-(Boc)6amikacin (17)

(200 mg, 0.16 mmol). TFA (3.9 mL) was added. The yellow solution was stirred

for 2.5 hours. The solvent was removed under reduced pressure. The remaining

white solid was dissolved in water and purified by reverse phase HPLC [0 – 0.1%

ACN in water (0.1% TFA) over 10 mins, flow rate is 2 mL / min, eluted after 7.5 min], then lyophilized and desalted. Product: White solid (75 mg, 0.12 mmol, 74%

1 yield). H NMR (400 MHz, D2O): δ 5.47 (d, J = 3.2 Hz, 1H), 5.17 (d, J = 3.6 Hz,

1H), 4.38 – 4.34 (m, 1H), 4.24 (dd, J1 = 9.2 Hz, J2 = 4.2 Hz, 1H), 4.11 – 3.98 (m,

2H), 3.88 – 3.73 (m, 4H), 3.65 (dd, J1 = 10.2 Hz, J2 = 3.8 Hz, 1H), 3.60 – 3.34 (m,

12H), 3.21 (q, J = 6.8 Hz, 1H), 3.14 (t, J = 7.2 Hz, 2H), 2.21 – 2.11 (m, 2H), 1.97

13 – 1.88 (m, 1H), 1.77 (q, J = 12.6 Hz, 1H); C NMR (125 MHz, D2O): δ 174.68,

98.22, 96.10, 85.30, 77.62, 72.76, 71.16, 70.95, 69.89, 69.69, 69.04, 68.41,

67.94, 51.99, 48.87, 47.80, 46.83, 42.88, 40.86, 39.62, 37.85, 35.37, 34.28,

+ 32.35; HR-ESI-MS calculated for C24H49N7O12Na [M+Na] 650.3331, found

650.3326.

Scheme 1.10: Synthesis 6''-deoxy-6''-ureidoamikacin (2f). Reagents and conditions: a) 2, 4- Dimethoxybenzyl isocyanate, pyridine, RT; b) TFA, TIPS, CH2Cl2, RT.

6’’-Deoxy-6’’-ureidoamikacin (2f). Anhydrous pyridine (1.78 mL) was added to

6’’-deoxy-6’’-amino-(Boc)4amikacin (13) (266 mg, 0.27 mmol). Dimethoxybenzyl isocyanate (82 µL, 0.27 mmol) was added. The yellow solution was wrapped in tinfoil and stirred overnight. The solvent was removed under reduced pressure.

The product was isolated by automated flash chromatography (0 – 20%

methanol in DCM over 11 mins) eluted after 7.5 min. The pale yellow solid

product was carried on without further purification. DCM (6.4 mL) and TIPS (0.38

mL) were added to the crude product. TFA (6.4 mL) was added. The yellow

solution was stirred for 3.5 hours. The solvent was removed under reduced

pressure. The remaining white solid was dissolved in water and purified by

reverse phase HPLC [0 – 1% ACN in water (0.1% TFA) over 8 mins, eluted after

5.1 min], then lyophilized and desalted. Product: White solid (95 mg, 0.15 mmol,

1 56%). H NMR (500 MHz, D2O): δ 5.28 (s, 1H), 4.98 (d, J = 3.5 Hz, 1H), 4.09 (dd,

J1 = 9.5 Hz, J2 = 3.8 Hz, 1H), 4.02 – 3.89 (m, 2H), 3.79 – 3.60 (m, 4H), 3.55 (dd,

J1 = 9.4 Hz, J2 = 4.2 Hz, 1H), 3.42 – 3.16 (m, 6H), 3.14 – 3.02 (m, 3H), 2.97 –

2.82 (m, 3H), 1.98 – 1.84 (m, 2H), 1.74 – 1.62 (m, 1H), 1.36 (q, J = 12.4 Hz, 1H);

13 C NMR (125 MHz, D2O): δ 173.42, 158.43, 96.03, 95.36, 79.63, 77.04, 71.45,

71.22, 69.71, 68.71, 68.68, 67.95, 67.37, 66.00, 57.18, 51.26, 46.59, 45.48,

42.47, 38.29, 34.97, 31.15, 26.42; HR-ESI-MS calculated for C23H45N7O13Na

[M+Na]+ 650.2968, found 650.2974.

Scheme 1.11: Synthesis of 6''-deoxy-6''-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g). Reagents and conditions: a) Propargyl (Boc)amine, CuSO4 5 H2O, sodium ascorbate, THF, tBuOH, H2O, RT; b) TFA, TIPS, CH2Cl2, RT.

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)-(Boc)5amikacin (18). A 3

THF: 1 tert-butanol: 1 water solution (5.1 mL) was added to 6’’-deoxy-6’’-azido-

(Boc)4amikacin (14) (150 mg, 0.148 mmol) and N-Boc-propargylamine (27 mg,

0.174 mmol). The solution was degassed by bubbling through argon for 25 minutes. A 7.5% weight / volume aqueous copper sulfate pentahydrate solution

(64 µL) was added. Then a 1 M aqueous sodium ascorbate solution (156 µL)

was added. The solution turned from light blue to orange. The solution was

stirred overnight during which time it became a yellow mixture. The solvent was

removed under reduced pressure. The product was isolated by automated flash

chromatography (0 – 20% methanol in DCM over 11 mins) eluted after 7.5 min.

Product: Light yellow solid (120 mg, 0.103 mmol, 70% yield). 1H NMR (400 MHz,

CD3OD): δ 6.89 (s, 1H), 5.00 (d, J = 3.6 Hz, 1H), 4.97 (s, 1H), 4.67 – 4.39 (m,

6H), 4.34 – 4.24 (m, 1H), 3.67 – 3.43 (m, 6H), 3.19 – 2.79 (m, 8H), 2.68 (s, 1H),

2.01 – 1.85 (m, 2H), 1.75 – 0.97 (m, 56H); HR-ESI-MS calculated for

+ C42H74N8O20Na [M+Na] 1033.4912, found 1033.4913.

6’’-Deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g). DCM (2.45

mL) and TIPS (140 μL) were added to 6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3- triazol-1-yl)-(Boc)5amikacin (18) (110 mg, 0.094 mmol). TFA (2.45 mL) was

added. The yellow solution was stirred for 2.5 hours. The solvent was removed

under reduced pressure. The remaining white solid was dissolved in water and

purified by reverse phase HPLC [0 – 1.5% ACN in water (0.1% TFA) over 7 mins, eluted after 5.0 min], then lyophilized and desalted. Product: White solid (51 mg,

1 0.077 mmol, 82% yield). H NMR (400 MHz, D2O): δ 7.89 (s, 1H), 5.17 (d, J = 2.4

Hz, 1H), 5.02 (d, J = 4.0 Hz, 1H), 4.68 – 4.52 (m, 2H), 4.41 – 4.34 (m, 1H), 4.26

(t, J = 11.2 Hz, 1H), 4.15 (d, J = 8.8 Hz, 1H), 3.96 – 3.88 (m, 2H), 3.77 – 3.53 (m,

5H), 3.33 – 3.25 (m, 2H), 3.15 – 3.07 (m, 2H), 2.98 – 2.95 (m, 2H), 2.89 – 2.68

(m, 4H), 1.91 – 1.81 (m, 2H), 1.73 – 1.59 (m, 1H), 1.34 (q, J = 12.6 Hz, 1H); 13C

NMR (125 MHz, D2O): δ 172.7, 147.18, 123.23, 99.32, 96.97, 86.43, 78.81,

78.06, 77.15, 74.63, 73.15, 72.03, 70.32, 69.92, 69.41, 69.05, 68.80, 53.10,

49.77, 48.52, 47.58, 40.47, 36.25, 34.93, 34.64; HR-ESI-MS calculated for

+ C25H47N9O12Na [M+Na] 688.3236, found 688.3234.

Desalting

Aminoglycoside·TFA (up to 40 mg) was dissolved in autoclaved H2O (0.6

mL) in a sterile eppendorf tube. Dowex Monosphere 550 A (100 mg) was added,

and the suspension was shaken lightly on a Fisher Vortex Genie 2 overnight. The

resin was removed by centrifugal filtration and washed twice with autoclaved

H2O. The desalted solutions were lyophilized, and the removal of TFA counterions was confirmed by 13C NMR spectroscopy.

A-site Binding Assay

All titrations were performed with working solutions of 1 μM Dy-547 labeled A-site in 20 mM cacodylate buffer (pH = 7.0, 100 mM NaCl, 0.5 mM

EDTA). The solutions were heated to 75 °C for 5 min, cooled to room temperature over 2 h, cooled to 0 °C for 30 min, then allowed to warm back to room temperature. Kanamycin-coumarin or neomycin-coumarin was added, to give a working concentration of 0.53 μM, just prior to aminoglycoside titrations.

Steady state fluorescence experiments were carried out at ambient temperature

(20 °C). Excitation and emission slit widths were 9 nm for kanamycin-coumarin experiments and 7 nm for neomycin-coumarin. The system was excited at 400 nm and changes in Dy-547 emission were monitored at 561 nm. Errors were generated from three sets of measurements. IC50 values were calculated using

OriginPro 8.5 software by fitting a dose response curve (eq. 1) to the fractional fluorescence saturation (Fs) plotted against the log of antibiotic (A) concentration.

n n n (1) Fs = F0 + (F∞[A] )/([IC50] + [A] ) (1)

Fs is the fluorescence intensity at each titration point. F0 and F∞ are the fluorescence intensity in the absence of aminoglycoside or at saturation, respectively, and n is the Hill coefficient or degree of cooperativity associated

92 with binding. The binding curves for the displacement of kanamycin-coumarin

(Figure 1.5) and neomycin-coumarin (Figure 1.6) are pictured.

Figure 1.5: Kanamycin-Coumarin displacement curves for tobramycin (1a), amikacin (2a), 6’’- deoxy-6’’-aminoamikacin (2b), 6’’-deoxy-6’’-methylaminoamikacin (2c), 6’’-deoxy-6’’- dimethylaminoamikacin (2d), 6’’-deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e), 6’’-deoxy-6’’- ureidoamikacin (2f), and 6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g).

Figure 1.6: Neomycin-Coumarin displacement curves for tobramycin (1a), 6’’-deoxy-6’’- aminotobramycin (1b), 6’’-deoxy-6’’-methylaminotobramycin (1c), 6’’-deoxy-6’’- dimethylaminotobramycin (1d), 6’’-deoxy-6’’-(2-(aminoethyl)amino)tobramycin (1e), 6’’-deoxy-6’’- ureidotobramycin (1f), 6’’-deoxy-6’’-(4-(aminomethyl)-1H-1,2,3-triazol-1-yl)tobramycin (1g), 6’’- deoxy-6’’-aminoamikacin (2b), 6’’-deoxy-6’’-methylaminoamikacin (2c), 6’’-Deoxy-6’’- dimethylaminoamikacin (2d), 6’’-deoxy-6’’-(2-(aminoethyl)amino)amikacin (2e), and 6’’-deoxy-6’’- (4-(aminomethyl)-1H-1,2,3-triazol-1-yl)amikacin (2g).

Minimum inhibitory concentration (MIC) determinations

MIC values for aminoglycosides were determined using broth microdilution

in accordance with Clinical Laboratory Standards Institute guidelines.34

Parent Aminoglycoside Crystal Structures

Crystal structure representations (Figure 1.2) were made using PyMOL

Molecular Graphics Systems, Version 1.4.1, Schrödinger, LLC. All structures

were adapted from PDB files: tobramycin (1LC4), amikacin (2GSQ).

Acknowledgements

Chapter 1 is in full currently being prepared for submission: Fair, R. J.;

McCoy, L. S.; Hensler, M. E.; Nizet, V.; Tor, Y. Singly Modified Amikacin and

Tobramycin Derivatives Show Increased A-site Binding and Higher Potency against Resistant Bacteria. The dissertation author is the co-main author and researcher of this work.

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(10) Barbachyn, M. R.; Ford, C. W. Angew Chem Int Edit 2003, 42, 2010.

(11) Carpenter, C. F.; Chambers, H. F. Clin Infect Dis 2004, 38, 994.

(12) Andries, K.; Verhasselt, P.; Guillemont, J.; Gohlmann, H. W. H.; Neefs, J. M.; Winkler, H.; Van Gestel, J.; Timmerman, P.; Zhu, M.; Lee, E.; Williams, P.; de Chaffoy, D.; Huitric, E.; Hoffner, S.; Cambau, E.; Truffot-Pernot, C.; Lounis, N.; Jarlier, V. Science 2005, 307, 223.

(13) Mullane, K. M.; Gorbach, S. Expert Rev Anti-Infe 2011, 9, 767.

(14) Falagas, M. E.; Grammatikos, A. P.; Michalopoulos, A. Expert Rev Anti-Infe 2008, 6, 593.

(15) Hermann, T. Cell. Mol. Life Sci. 2007, 64, 1841.

(16) Falagas, M. E.; Kasiakou, S. K. Clin Infect Dis 2005, 40, 1333.

(17) McCoy, L. S.; Roberts, K. D.; Nation, R. L.; Thompson, P. E.; Velkov, T.; Li, J.; Tor, Y. Chembiochem 2013, 14, 2083.

(18) Avent, M. L.; Rogers, B. A.; Cheng, A. C.; Paterson, D. L. Intern Med J 2011, 41, 441.

(19) Jackson, J.; Chen, C.; Buising, K. Curr Opin Infect Dis 2013, 26, 516.

(20) Moazed, D.; Noller, H. F. Nature 1987, 327, 389.

(21) Carter, A. P.; Clemons, W. M.; Brodersen, D. E.; Morgan-Warren, R. J.; Wimberly, B. T.; Ramakrishnan, V. Nature 2000, 407, 340.

(22) Ogle, J. M.; Brodersen, D. E.; Clemons, W. M.; Tarry, M. J.; Carter, A. P.; Ramakrishnan, V. Science 2001, 292, 897.

(23) Becker, B.; Cooper, M. A. Acs Chem Biol 2013, 8, 105.

(24) Vicens, Q.; Westhof, E. Chemistry & Biology 2002, 9, 747.

(25) Kondo, J.; Francois, B.; Russell, R. J. M.; Murray, J. B.; Westhof, E. Biochimie 2006, 88, 1027.

(26) Fair, R. J.; Hensler, M. E.; Thienphrapa, W.; Dam, Q. N.; Nizet, V.; Tor, Y. Chemmedchem 2012, 7, 1237.

(27) Michael, K.; Wang, H.; Tor, Y. Bioorgan Med Chem 1999, 7, 1361.

(28) Xie, Y.; Dix, A. V.; Tor, Y. Journal of the American Chemical Society 2009, 131, 17605.

(29) Xie, Y.; Dix, A. V.; Tor, Y. Chemical Communications 2010, 46, 5542.

(30) Kaul, M.; Barbieri, C. M.; Pilch, D. S. Journal of the American Chemical Society 2006, 128, 1261.

(31) Kawaguchi, H.; Naito, T.; Nakagawa, S.; Fujisawa, K. J. Antibiot. 1972, 25, 695.

(32) Denton, T. T.; Zhang, X. D.; Cashman, J. R. J Med Chem 2005, 48, 224.

(33) Huston, W. M.; Jennings, M. P.; McEwan, A. G. Mol Microbiol 2002, 45, 1741.

(34) Performance Standards for Antimicrobial Susceptibility Testing, 19th Informational Supplement M100-S19, Clinical and Laboratory Standards Institute, Wayne, PA, USA, 2008.

Chapter 2

Polymyxins and analogs bind to ribosomal RNA and interfere with eukaryotic translation in vitro

2.1 Introduction

Many antibiotics bind to the ribosome, validating it as an RNA target.

These molecules have inspired the search for new molecules that bind to RNA,

whether semi- or totally synthetic. However, in taking into consideration the

properties of the molecules that are known to bind to RNA, especially those as

well studied as aminoglycosides, we can look at “old compounds” for new

insights. The structural properties of aminoglycosides have led us to look more

closely at polymyxin antibiotics.

The renewed interest in the clinical use of polymyxin (PMB) and colistin, is

due to bacterial resistance to current antibiotics and the lack of new antibiotics to

treat multi-drug resistant infections, especially by Gram-negative bacteria.1-3

Indeed, although abandoned as antibiotics in the 1970s due to toxicity concerns,

the polymyxins have been recently resurrected as last-line treatments for otherwise untreatable multi-drug resistant infections.4-6 The search for new

polymyxins derivatives is also underway.7-10

Polymyxins are penta-cationic cyclic lipodecapeptides. Their amphiphillic nature is the presumed basis for the proposed mechanism of action against

Gram-negative bacteria, which involves outer membrane (OM) and inner

membrane (IM) permeation. It has been suggested that polymyxins first bind to

the lipid A core of the lipopolysaccharide component of the OM through

electrostatic attractions between the positively charged γ-L-diaminobutyric acid

(Dab) residues with the negatively charged lipid A phosphate groups (Figure

2.1).11-15 This then allows the hydrophobic fatty acyl tail and D-Phe6-Leu7

hydrophobic motif to insert into the OM, destabilizing the packing of the fatty acyl

chains of lipid A.11-13,15,16 Subsequently, through a self-promoted uptake mechanism, polymyxins pass through the OM.

Figure 2.1: Polymyxin B1 (PMB1) and E. coli lipid A. The functional groups that are reported to interact are color coded. The phosphate groups and Dab residues PMB1 that interact are colored in purple. The hydrophobic groups of Polymyxin B that interact with lipid A fatty acyl chains are colored orange. The blue groups interact with the aqueous environment outside the cell.

The exact mechanism relating IM permeation to bactericidal activity

remains unclear. 11,12,14,17,18 The amphiphillic character alone likely does not

account for this activity, as common cationic detergents function as antibacterial

agents at much higher concentrations than the polymyxin’s MICs.12,13,15,19,20

These observations have led to the hypotheses that polymyxins act on the IM by

permeating the phospholipids bilayer or by pore formation.12,14,17 An alternative

hypothesis suggests that polymyxins may act by contacting the periplasmic

leaflets of the OM and IM, which facilitates phospholipid exchange through

vesicle-vesicle contacts.11,18 This could create an osmotic imbalance, leading to

bacteriolytic events and eventually cell death.21,22

Figure 2.2: Structures of PMB, colistin, and two new synthetic analogs. Primary amines, likely to be protonated at physiological pHs, are colored blue.

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The diminished clinical use of polymyxins attributed to their neurotoxicity

and nephrotoxicity and, less commonly, ototoxicity and pruritus,23 although the

underlying mechanisms responsible for these adverse effects are not known. We

noted that aminoglycoside antibiotics,24-26 exhibit similar side effects, such as

nephrotoxicity and ototoxicity.27,28 Like polymyxins, aminoglycosides are highly

cationic, with typically four to six protonated amines at physiological pH. Another

similarity is that aminoglycosides are proposed to have a self-promoted uptake

mechanism across the OM in P. Aeruginosa.29-31 We hypothesized that the

similar toxicity profiles along with the overall chemical characteristics of

aminoglycosides and polymyxins could be due to shared targets and molecular

mechanisms.

We decided to probe the binding of polymyxins to the bacterial 16S

ribosomal RNA decoding site (or A-site),32,33 the cognate target of

aminoglycosides, as well to the corresponding eukaryotic 18S RNA A-site, and

examine their effect on prokaryotic and eukaryotic translation in vitro. Due to the

recent efforts to make novel derivatives, we have expanded our analysis to new

analogs made by Dr. Jian Li’s lab (Monash School of Pharmacy), in which the R1

and R6 positions are modified with different hydrophobic resides (Figure 2.2).

These analogs have been shown to have favorable antibacterial activity.34 We

demonstrate that polymyxin B, colistin and analogs (1 and 2) bind to the 16S and

18S A-site RNA constructs (Figure 2.3), and interfere with eukaryotic translation

in vitro, but not with bacterial translation.

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2.2 Results and Discussion

To examine the affinity of PMB, colistin and analogs to the 16S and 18S

A-sites, we utilized a modified version of a FRET assay originally developed in

our laboratory;35,36 this assay has recently been used for exploring the affinity of

modified aminoglycosides to the 16S RNA construct and is discussed in Chapter

1.37 Titration curves for the 16S and 18S A-sites of all compounds’ displacement of kanamycin-coumarin (Figure 2.3) are in Figure 2.4 and 2.5, respectively.

Figure 2.3: A) Kanamycin and kanamycin-coumarin (donor) conjugate. B) Fluorescently labeled with Dy-547 (acceptor) 16S and 18S A-site RNA constructs. Primary amines, likely to be protonated at physiological pHs, are colored blue.

Polymyxins and the analogs display significant affinity to the 16S A-site

(Figure 2.4 and Table 2.1). PMB and colistin are the weakest binders with IC50

values of 52±2 µM and 77±5 µM, respectively. The analogs 1 and 2 exhibit

higher affinities than PMB and colistin. Notably, 2 with an IC50 value of 4.5±0.2

µM is more potent than kanamycin, which is considered a mediocre A-site binder.

Polymyxin B nonapeptide (PMBN), where the hydrophobic tail-Dab1 has been

proteolytically removed,38 shows a 1.5-fold increase in affinity compared to PMB.

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Colistin methanesulfonate (CMS), a derivative where the primary amino groups

are sulfomethylated, exhibits no displacement up to 500 μM. CMS also acts a

useful negative control, as it does not interfere with the FRET signal, helping to

verify that the polymyxins do not disrupt the A-site construct. Repeating the experiments using the 18S eukaryotic A-site RNA construct yields a similar trend of IC50 values across the polymyxins and analogs to those obtained for the

titrations using the 16S A-site construct (Figure 2.3, Figure 2.4 and Table 2.1).

Figure 2.4: Response of the emissive acceptor (Dy547) on the 16S A-site measured for polymyxin B, colistin, 1, 2, kanamycin, PMBN, and CMS. Titrations were performed with solutions of 1 μM Dy-547 labeled 16S A-site in 20 mM cacodylate buffer (pH = 7.0, 100 mM NaCl, 0.5 mM EDTA).

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Figure 2.5: Response of the emissive acceptor (Dy547) on the 18S A-site measured for polymyxin B, colistin, 1, 2, and kanamycin. Titrations were performed with solutions of 1 μM Dy- 547 labeled 18S A-site in 20 mM cacodylate buffer (pH = 7.0, 100 mM NaCl, 0.5 mM EDTA).

All compounds are examined in vitro for their ability to hamper bacterial

and eukaryotic translation. This coupled transcription/translation assay monitors

the generation of a functional luciferase following the addition of luciferin and

measuring the resulting luminescence. Plotting the relative luminescence versus

compound concentration generates interference curves. Note that this assay

does not discriminate between inhibition of translation (lower protein yield) and

mistranslation (generation of defective proteins), as either results in lower relative

luminescence. We therefore refer to the observed results as translation

interference.

As shown in Figures 2.6 and 2.7, kanamycin (Figure 2.3), used as a

positive control, preferentially hinders bacterial translation over eukaryotic

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translation with IC50 values differing by 250-fold, consistent with previously

published results.39,40 No activity against bacterial translation occurs up to 220

µM for PMB and colistin, and 110 µM for 1 and 2 (Figures 2.6, Figure 2.7 and

Table 2.1). Importantly, PMB, colistin and the analogs 1 and 2 show dose-

dependent interference of eukaryotic translation. All disrupt eukaryotic translation

more potently than kanamycin, with 2 being the most effective with an IC50 value

of 8.8±0.8 µM (Table 1).

Figure 2.6: Bacterial in vitro translation interference data measured for polymyxin B, colistin, 1, 2, and kanamycin. See experimental section for details.

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Figure 2.7: Eukaryotic in vitro translation interference data measured for polymyxin B, colistin, 1, 2, and kanamycin. See experimental section for details.

As illustrated by the results outlined above, the polymyxins show

considerable binding to the bacterial and eukaryotic A-site RNA constructs. This

is likely due to the five protonatable amine groups’ electrostatic interaction with

RNA, as well as the ability of the amides, threonines, and amines to act as

hydrogen bonding donors and acceptors. PMB and colistin are relatively weaker

binders, as compared to the analogs 1 and 2 which incorporate a hydrophobic D-

octylglycine (D-OctGly) substitution at the R6 position (Figure 2.2). Notably, 2,

having a biphenyl in the R1 position, binds to the 16S and 18S A-sites with

greater and equal potency to kanamycin, respectively (Table 2.1). Bacterial

translation is not affected by any of the polymyxins or analogs.41 Intriguingly, all

polymyxins potently prevent eukaryotic translation with the general trend

corresponding to their 18S A-site binding, with 2 being the most potent. Notably,

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the very hydrophobic substitutions at positions R1 and R6 make 2 the most

hydrophobic of the analog series (Table 2.1). These results are highlighted by

comparing kanamycin and 2 in Figure 2.8.

Figure 2.8: A-site RNA binding of kanamycin (●, blue) and 2 (○, red) overlaid for A) 16S and B) 18S A-site constructs, and C) bacterial and D) eukaryotic IVT assays.

The bacterial A-site has been established to be a very selective binding

site for aminoglycoside antibiotics.42 It is intriguing to discover that polymyxins could disrupt the kanamycin–A-site complex. For comparison to the aminoglycosides, under the same experimental conditions for the 16S A-site, the

IC50 value previously observed for neomycin (strong binder) was 1.6±0.2 µM, neamine (moderate binder) was 4.5±0.4 µM, and kanamycin (weak binder) was

7.0±0.7 µM.37 PMB, colistin, and 1 show lower affinities compared to

aminoglycosides. Derivative 2, however, has an affinity comparable to neamine

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for the 16S A-site and to kanamycin for the 18S A-site under the same

conditions. Polymyxins contain five primary ammonium groups that could

participate in hydrogen bonding and can electrostatically interact with the RNA

scaffold. It is therefore interesting that these unrelated antibiotics can be

accommodated by this compact site, although allosteric effects, which have been

previously proposed for other RNA–ligand binding events, where polymyxins

displace kanamycin-coumarin by inducing RNA conformation changes, cannot be

excluded.43

Table 2.1: Binding to the 16S and 18S A-sites and interference with in vitro bacterial and eukaryotic translation.

Compounds 16S A-site 18S A-site Bacterial IVT Eukaryotic IC50 (μM) IC50 (μM) IC50 (μM) IVT IC50 (μM)

PMB 52±2 54±5 >220 46±1 Colistin 77±5 83±5 >220 32±2 1 20±1 19±3 >110 13±1 2 4.5±0.2 4.1±0.1 >110 8.8±0.8 Kanamycin 6.6±0.2 4.5±0.5 0.41±0.02 100 PMBN 32±2 - - - CMS >500 - - -

Polymyxins show little selectivity between the 16S and 18S RNA A-sites.

Between the parent compounds, colistin has a higher IC50 than PMB. The

possible difference in affinity could be due to the D-Phe6 in PMB compared to the

D-Leu6 of colistin, as phenylalanine has been suggested to intercalate in DNA.44

The compound that shows the most significant increase in A-site binding, 2, has

a D-OctGly in place of the D-Phe6 and also a biphenyl group substituting the N-

terminal fatty acyl chain. The modifications in 2 improve the relative affinity for

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the A-site dramatically. Biphenyl is known to base-stack in other systems;45,46

therefore, this dramatic increase could be due to its ability to intercalate. In noting

the fluorescence of 2 due to its biphenyl moiety, we suspected that we could

monitor the direct binding to the 16S A-site RNA using the excitation and

emission of 2 alone. Therefore, we probed the quenching fluorescence of 2 upon

titration with annealed 16S A-site RNA without using FRET. Indeed, monitoring the fluorescence of the biphenyl residue shows dose-dependent quenching upon titration of 16S A-site RNA (Figure 2.9). It should be noted that the FRET assay and this assay were done under different conditions to obtain IC50 values, so they

cannot be directly compared.

Figure 2.9: A) Excitation and B) emission of 2 while titrating in Dy547 16S A-site. Fractional saturation of 2 C) excitation spectra monitored at I281 and D) emission spectra monitored at I363. The IC50 for of Dy547 16S A-site RNA generated from the emission spectra (C) is 0.87±0.01 μM and excitation spectra (D) is 0.88±0.01 μM. Titrations were performed with solutions of 12.4 μM of 2 in 20 mM cacodylate buffer (pH = 7.0, 100 mM NaCl, 0.5 mM EDTA).

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A substantial difference between the natural aminoglycosides and

polymyxins is the presence of a non-polar fatty acyl group in the latter, which

likely has an effect on A-site binding. The likely bound position of the methyl

octanoyl/heptanoyl chain of PMB is closer to the groove wall, in the most

hydrophobic region.47 The apparent increase in affinity for PMBN compared to

PMB could indicate that the hydrophobic tail of PMB may be hindering certain

attractive interactions. In 1, a synthetic derivative, the hydrophobic D-Phe6 is

6 replaced by a D-OctGly . Compared to PMB, the IC50 of 1 decreases by almost four-fold. Perhaps the presence of the two hydrophobic chains causes a reorientation of the molecule to accommodate both octanoyl chains closer to the

RNA skeleton.

Polymyxins and analogs show no effect on bacterial translation. A second antibacterial mode of action of polymyxins by interference of bacterial translation is therefore unlikely, despite their affinity to the 16S RNA A-site, although it should be noted that A-site binding does not necessarily correlate to in vitro translation (IVT) activities, as previously observed for the aminoglycosides.32,39,48

Additionally, although the A-site is considered the cognate target of

aminoglycosides and is thought to contribute to their ability to interfere with

translation, they have been shown to bind to multiple sites on the ribosome.49-51

The polymyxins and analogs do effectively interfere with eukaryotic translation

and the general trend observed correlates with their relative affinity to the 18S A-

site. The comparison of kanamycin and 2 in Figure 2.8 emphasizes these

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similarities and differences. Other targets or modes of action contributing to this

outcome cannot be discounted. Whether or not the ability of polymyxins to

interfere with eukaryotic translation is responsible for the adverse effects seen in

the clinic remains to be explored.

2.3 Conclusion

Polymyxins and analogs displayed considerable affinity to the 16S and 18S

ribosomal A-sites, the natural target of aminoglycoside antibiotics. Most notably,

analog 2 showed the highest affinity, with IC50 values similar to and better than

kanamycin for the 16S and 18S A-sites, respectively. Additionally, all polymyxins

interfered with eukaryotic translation more effectively than kanamycin, but had no

effect on bacterial translation.

2.4 Future Directions

Ototoxicity has been speculated to result from aminoglycoside binding to

the mitochondrial ribosome A-site.52,53 Specifically, individuals with mitochondrial

mutations A1490G and C1490T (E. coli numbering) seem to be hypersensitive to

aminoglycoside induced deafness (Figure 2.10).54-56 Since polymyxins do exhibit ototoxicity,23 albeit rarely, testing their affinity for the 12S (mitochondrial) A-site

and mutants could give insight into this side effect.

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Figure 2.10: Comparison of the 16S (bacterial), 18S (eukaryotic), mitochondrial, mitochondrial mutant, and mitochondrial mutant A-site. E. coli numbering is used for consistency.

Interestingly there have been aminoglycoside conjugated to fatty acids and other non-polar moieties synthesized attempt to improve cellular uptake; however, only the antibacterial activity and not the A-site binding activity was examined.57-59 So as aminoglycosides have been modified to be more structurally similar to polymyxins, one could imagine investigating polymyxins in a similar fashion to aminoglycosides. Aminoglycosides bind to many RNA targets including many catalytic RNAs,60-66 regulatory elements of viral RNA, 67-70 structured region of a human mRNA, tRNA,71 mRNA,72 and others;25,73 we could investigate polymyxin binding to these and other RNA targets.

Guanidinylated aminoglycosides display very efficient cellular uptake, and they are able transport bioactive, high molecular weight cargo.74-78

Guanidinylating the amino groups of polymyxin and testing the uptake activity could also be an interesting endeavor.

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2.5 Experimental

Synthesis (by the Dr. Jian Li Lab)

Materials. Piperidine, diisopropylethylamine (DIPEA), trifluoroacetic acid (TFA)

and 1H-Benzotriazolium-1-[bis(dimethylamino)methylene]-5-chlorohexafluoro-

phosphate-(1-),3-oxide (HCTU) were obtained from Auspep (Melbourne,

Australia). Fmoc-Dab(Boc)-OH and Fmoc-Dab(ivDde)-OH (ivDde = 1-(4,4-

Dimethyl-2,6-dioxocyclohex-1-ylidene)-3-methylbutyl) were obtained from Chem-

Impex International (USA). Fmoc-Thr(tBu)-OH was obtained from Mimotopes

(Melbourne, Australia). Fmoc-D-OctGly-OH was obtained from Try-lead Chem

(China). Dimethylformamide (DMF), methanol (MeOH), diethyl ether, dichloromethane (DCM), hydrochloric acid (HCl) and acetonitrile were obtained from Merck (Melbourne, Australia). Fmoc-Thr(tBu)-TCP-Resin (Intavis

Bioanyltical Instruments, Germany). Triisopropylsilane (TIPS), diphenylphosphorylazide (DPPA) and diisopropylethylamine (DIPEA) were obtained from Sigma-Aldrich (Castle Hill, Australia).

Analytical LC/MS was performed on a Shimadzu 2020 LCMS system, incorporating a photodiode array detector (214 nm) coupled directly to an electrospray ionization source and a single quadrupole mass analyzer. RP-HPLC was carried out employing a Phenomenex column (Luna C8(2), 100 × 2.0 mm

ID) eluting with a gradient of 80% acetonitrile in 0.05% aqueous TFA, over 10 minutes at a flow rate of 0.2 mL/min. Mass spectra were acquired in positive ion mode with a scan range of 200-2000 m/z.

113

Crude cyclic peptides were purified by reversed phase HPLC on a Agilent

1200 quaternary pump system, photodiode array detector (214 nm), employing a

PhenomenexAxia column (Luna C8(2), 50 × 21.3 mm) eluting with a gradient of

60% acetonitrile in 0.1% aqueous TFA, over 60 minutes at a flow rate of 5 mL/min.

Scheme 2.1: Synthesis of 1 and 2.

114

Synthesis of 1 (HCl salt)

Synthesis of the protected linear precursor. Synthesis of the protected linear peptide was carried out using out on a CEM Liberty Microwave automated peptide synthesizer using standard Fmoc solid phase peptide chemistry.

Specifically, synthesis was undertaken using TCP-Resin, pre-loaded with Fmoc-

Thr(tBu)-OH (loading 1.0mmol/g), 0.25 mmol scale (260 mg of resin). Coupling of the Fmoc-amino acids and the N-terminal octanoyl group was performed using the default instrument protocol: 5 molar equivalents (eq.) of Fmoc amino acid and

HCTU with activation using DIPEA in DMF over 2 minutes at room temperature then for 4 mins at 50°C (25W microwave power). Fmoc deprotection was performed using the default instrument protocol: 20% piperidine in dimethylformamide (1 × 30s, 1 × 3 min) at 75°C (35W microwave power). The resin was removed from the instrument and transferred to a synthesis syringe and treated with 2% hydrazine in DMF (4 × 15 min) to remove the ivDde group.

The resin was then washed with MeOH (2 × 2 min) and diethyl ether (1 x 2 min)

115

then air dried under vacuum suction. The protected linear peptide was then

cleaved from the resin by washing the resin with 1% TFA in DCM (1 × 5 min, 3 x

10 min). The resulting residue was dissolved in 50% acetonitrile/water and freeze

dried overnight.34

Cyclization of the protected linear peptide. The protected linear peptide was

dissolved in DMF (10 mL) to which DPPA (3 eq. relative to the loading of the

resin) and DIPEA (6 eq. relative to the loading of the resin) were added. This

solution was stirred at room temperature overnight. The reaction solution was

then concentrated under vacuum overnight.

Removal of the side-chain protecting groups from the crude protected

cyclic peptide. The resulting residue was taken up in a solution of 5% TIPS in

TFA and stirred at room temperature for 2 h. The TFA was removed under a

stream of nitrogen and the crude cyclic peptide precipitated with cold diethyl

ether. The resulting precipitate was collected by centrifugation and air-dried in a

fume hood to give the crude cyclic peptide as a residue. The resulting residue

was taken up in milli-Q water and desalted using a Vari-Pure IPE SAX column.

The resulting solution containing the crude cyclic peptide was subjected to RP-

HPLC purification. Fractions collected were analyzed by LC-MS as described above. Concentrated HCl was added to the combined fractions at a concentration of 40 µL/10 mL of eluent. Combined fraction where the freeze- dried for two days to give the 1 HCl salt as a white solid in a yield of 34 mg .The purity was >95% as estimated by reversed-phase HPLC. The compound was

116

confirmed as having the correct mass by ESI-MS analysis: m/z (monoisotopic)

+ 2+ + calculated; C56H106N16O13 [M+H] 1211.82, [M+2H] 606.42 observed; [M+H]

1211.95, [M+2H]2+ 606.65.

Synthesis of 2 (HCl salt)

As described above, 2 was synthesized as an HCl salt as a white solid in a yield

of 25 mg. The purity was >95% as estimated by reversed-phase HPLC. The

compound was confirmed as having the correct mass by ESI-MS analysis: m/z

+ 2+ (monoisotopic) calculated; C61H100N16O13 [M+H] 1265.77, [M+2H] 633.39 observed; [M+H]+ 1266.00, [M+2H]2+ 633.80.

A-site Binding and In vitro Translation Assays

Materials. The Dy-547-labeled 16S and 18S A-site constructs were purchased from Thermo Scientific Dharmacon®. Kanamycin–coumarin was synthesized and purified according to established procedures.36 Chemicals for buffer solutions

(enzyme grade) were purchased from Fisher Biotech. Autoclaved water was

used for all stock solutions and experiments. Kanamycin was obtained from

117

Sigma Aldrich as the sulfate salt, and was converted into the corresponding

neutral form using Dowex Monosphere 550A (OH) anion exchange resin. CMS

and PMBN were obtained from Sigma Aldrich as sodium and HCl salts,

respectively.

Instrumentation. Chemiluminescence was measured with a Molecular Devices

SPECTRAmax® GEMINI XS plate reader. Steady-state fluorescence

experiments were carried out in a microfluorescence cell with a path length of 1.0

cm (Hellma GmH & Co KG, Mullenheim, Germany) on a Jobin Yvon Horiba

FluoroMax-3 luminescence spectrometer.

Oligonucleotide Purification. The Dy-547-labeled 16S and 18S A-site

constructs were purchased from Thermo Scientific Dharmacon®. The 2’-ACE®

protected RNA was deprotected according to the protocol by Thermo Scientific

Dharmacon®. The deprotection buffer, provided by Thermo Scientific

Dharmacon®, was 100 mM acetic acid, adjusted to pH 3.8 with tetramethylenediamine. After drying with a speed-vac, the oligonucleotides were purified by 20% polyacrylamide gel electrophoresis. The RNA was pink in color and visualized by UV shadowing. The bands were excised from the gel, pulverized, and extracted with 300 mM sodium acetate buffer (pH 7.0) rotating overnight. The resulting solution was filtered (Bio Rad poly-prep chromatography column) and desalted using a Sep-Pak C18 cartridge (Waters Corporation, MA).

The oligonucleotide was eluted with 50% acetonitrile in water. Fractions

118

containing RNA were pooled and evaporated in a speed vac. The purified RNA

was quantified by UV at 260 nm using the nearest-neighbor method.79,80

A-site Binding Assay. All titrations were performed with working solutions of 1

μM Dy-547 labeled A-site in 20 mM cacodylate buffer (pH = 7.0, 100 mM NaCl,

0.5 mM EDTA). The solutions were heated to 80°C for 5 min, cooled to room temperature over 2 h, cooled to 4°C for 30 min, then allowed to warm back to room temperature. Kanamycin-coumarin was added to give a working concentration of 0.55 μM, just prior to aminoglycoside titrations, so the initial I561

was between 5.0 × 106 and 7.0 × 106 cps. Compounds were tested at varying

concentrations from 0.024 μM to 1600 μM. Steady-state fluorescence experiments were carried out at 20°C. Excitation and emission slit widths were 9 nm. The system was excited at 400 nm and changes in Dy-547 emission upon compound titration were monitored at 561 nm. Errors were standard deviations generated from three sets of measurements. IC50 values were calculated using

OriginPro 8 software by fitting a dose response curve (eq. 1) to the fractional fluorescence saturation (FS) plotted against the log of antibiotic (A) concentration.

n n n (1) FS = F0 + (F∞ [A] )/([IC50] + [A] )

FS is the fluorescence intensity at each titration point. F0 and F∞ are the

fluorescence intensity in the absence of aminoglycoside or at saturation,

respectively, and n is the Hill coefficient or degree of cooperativity associated

with binding.

119

In vitro translation assays. Bacterial in vitro translation was quantified using a

coupled transcription translation assay (S30 T7 High-Yield Protein Expression

System). Compounds were tested at varying concentrations from 0.11 μM to 220

μM. A DNA plasmid (100 ng/uL) containing the renilla luciferase gene under

control of a T7 phage RNA polymerase promoter was used. For each reaction

S30 premix plus (3.2 µL) and T7 S30 extract (2.88 µL) were premixed just before

use. The experiment was done in strip tubes with compound or water (1.2 µL),

plasmid DNA (1.2 µL), and the premixed S30 extract mixture (6.08 µL) for a total

reaction volume of 8.48 µL. Reactions were incubated at 37°C for 30 min. in a

thermocycler, and the reaction was cooled to 4°C when completed. Then 5 µL of

each reaction was added to a 96-well plate. Renilla luciferin substrate in buffer

(25 µL; Promega) was added and luminescence was immediately measured with

a plate reader.

Eukaryotic in vitro translation was quantified using a coupled transcription translation assay (TnT® SP6 Coupled Reticulocyte Lysate System). Compounds

were tested at varying concentrations from 2.4 µM to 156 µM. A DNA plasmid

(100 ng/µL) containing the luciferase gene under control of a SP6 phage RNA

polymerase promoter was used. For each reaction, rabbit reticulocyte (3.75 µL),

TnT buffer (0.3 µL), amino acids (0.15 µL), SP6 polymerase (0.15 µL), and an

RNase inhibitor (0.15 µL) were premixed just before use. The experiment was

done in strip tubes with compound or water (1.5 µL), plasmid DNA (1.5 µL), and

the premixed reticulocyte mixture (4.5 µL) for a total reaction volume of 7.5 µL.

120

Reactions were incubated at 30°C for 30 min. in a thermocycler, and the reaction

was cooled to 4°C upon completion. Then 5 µL of each reaction is added to a 96-

well plate. Firefly luciferase substrate in buffer (25 µL; Promega) was added and

luminescence was immediately measured in the same manner as above.

The half-maximum inhibitor concentration (IC50) was determined from

relative luminescence (% of the control) plotted against the log of compound

concentration by fitting a dose-response curve using OriginPro 8 software. Each

concentration was examined in triplicate per experiment. Each data point

represents the average of at least two independent experimental results. Error

bars were generated from the standard deviation.

Dose-dependent response of 2 to 16S A-site RNA. The excitation spectra

monitored at 281 nm and emission spectra monitored at 363 nm obtained upon

addition of the Dy547 16S A-site RNA constructs are IC50 of 0.87±0.01 μM and

0.88±0.01 μM, respectively (Figure 2.9). All titrations were performed with

working solutions of 12.4 μM of 2 in 20 mM cacodylate buffer (pH = 7.0, 100 mM

NaCl, 0.5 mM EDTA). Solutions of 20 μM and 40 μM Dy547 labeled 16S A-site in

20 mM cacodylate buffer (pH = 7.0, 100 mM NaCl, 0.5 mM EDTA) were heated

to 80°C for 5 min, cooled to room temperature over 2 h, cooled to 4°C for 30 min,

and then allowed to warm back to room temperature. The solution of 2 was

titrated with Dy547 16S A-site RNA from 0.16 μM to 2.4 μM. Steady-state fluorescence spectra were taken at 20°C in a 125 µL quartz fluorescence cell with a path length of 1.0 cm. Excitation and emission slit widths were 3 nm. The

121

solution was excited at a wavelength of 273 nm and changes in the emission of 2 upon RNA titration were monitored at 363 nm. Excitation spectra was taken at

363 nm and monitored at 281 nm. Errors were standard deviations generated from two sets of measurements. IC50 values were calculated using OriginPro 8 software by fitting a sigmoidal dose response curve to the fractional fluorescence saturation plotted against the log of Dy547 16S A-site concentration.

Acknowledgements

Chapter 2, is a full reprint from: McCoy, L. S.; Roberts, K. D.; Nation, R.

L.; Thompson, P. E.; Velkov, T.; Li, J.; Tor, Y. Polymyxins and analogues bind to ribosomal RNA and interfere with eukaryotic translation in vitro. Chembiochem

2013, 14, 2083. The dissertation author is the main author and researcher of this work.

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Chapter 3

Synthesis of meso- 2-deoxystreptamine mimetics

3.1 Introduction

An important player in many biological functions, RNA has become a

potential therapeutic target for small molecules to combat human disease and

pathogens.1-3 The ribosome has been substantiated as an RNA target, as many

classes of antibiotics bind to defined regions of this cellular machinery.4-6 This

has set the precedence for the pursuit of small molecules that bind RNA. The

bacterial rRNA decoding site is the natural target of most aminoglycosides,7 but

because of their ability to bind numerous other RNA targets,8,9 they have served as inspiration for creating new RNA binders.8-11

The majority of aminoglycosides contain the highly conserved 2-

deoxystreptamine (2-DOS) ring, typically substituted with aminosugars at the 4,6-

or 4,5-positions (Figure 3.1). At the A-site, codon–anticodon recognition and

discrimination is performed, which is crucial for high fidelity translation.

Aminoglycosides bind to the A-site and disrupt the decoding process, lower its

fidelity and increase misincorporation of amino acids.12 Additionally, some

aminoglycosides are thought to interfere with translocation.13

127

128

Figure 3.1: Structures of select aminoglycosides. 2-DOS is in pink.

Electrostatic and hydrogen bonding interactions contribute to

aminoglycoside RNA binding.14 Aminoglycoside amine groups have a range of

15,16 pKb values, but most are protonated at physiological pHs. Crystal structures

of aminoglycoside–A-site constructs illustrate that the

paromamine/neamine/gentamicin cores are involved in highly conserved

interactions with the RNA.17 This indicates the key pharmacophore of

aminoglycosides are rings I and II, and the 2-DOS is regarded as a privileged

RNA binding scaffold.18,19

As with other antibiotics, the clinical use of aminoglycosides has been

compromised by the emergence of resistant bacteria. Mechanisms of resistance

include efflux pumps, rRNA modifications, mutations in ribosomal proteins, and

enzymatic modifications of the aminoglycosides, which is the most common in the clinical setting.20,21 With the onslaught of resistant bacteria and an effort to

improve toxicity profiles, new aminoglycoside derivatives have been pursued with

129

many approaches. Efforts have included modifications of the natural products,22-

24 the functionalization of 2-DOS with non-carbohydrate moieties,25-29 or replacing

the 2-DOS core with mimetic scaffolds, of which many examples will be outlined

here and shown in Figure 3.2.

There have been many attempts at mimicking the 2-DOS core of

aminoglycosides. Analogs 2-6 preserve the 1,3-diamine moiety. One of the most

similar compounds to 2-DOS is 2,5-dideoxystreptmine30 (2), which has been

used as part of a neamine mimic with a variety of other substitutions; some

showed anti-translational activity but none as potent as parent

aminoglycosides.31 Mimics capable of forming an amide bond have been

synthesized. Compound 3 was used as 2-DOS mimic but the derivatives

synthesized containing 3 exhibited poor activity in an in vitro transcription– translation assay.32 More recently, another 2-DOS mimic was synthesized (4),11

and this inspired an aminoglycoside mimetic where neamine was attached to a

similar 2-DOS mimic (5) by amide connectivity.33 The A-site affinity was

increased compared to neamine, although antibacterial activity was not tested.

Several derivatives containing the 3,5-diaminopiperazine moiety (6) were

synthesized and a few showed potent antibacterial activity, although potency was

significantly reduced in the presence of serum.34,35 Molecules containing a

piperidine core, with 7 being one example of the substitution pattern, inhibited

bacterial translation but not as effectively as parent aminoglycosides.36 A carbohydrate mimetic (8) was used to create in conformationally restricted

130

derivatives, although no activity testing or further manuscripts have been

published.37 Azepane containing compounds showed A-site binding, bacterial

translation inhibition, and moderate antibacterial activity (9).38

Figure 3.2: 2-DOS (1) and previously published mimetics (2-9). The amines that are speculated to mimic the 1,3-cis-diamine functionality of 2-DOS are colored in blue.

As 2-DOS appears to be a privileged RNA binding scaffold,2 DOS- mimetics can, in theory, maintain essential A-site contacts. We have selected to synthesize new 2-DOS mimics that contain simplified structures without extensive functionalization and multiple stereocenters (Figure 3.3). The intrinsic meso characteristic of these compounds will hopefully diminish the number of possible stereoisomers that can form throughout the synthesis. The importance of the amino groups for hydrogen bonding and electrostatic interactions with the

RNA target is reflected in the design of these molecules, as we retain the 1,3-cis- diamine functionality to mimic 2-DOS (Figure 3.3).The versatile scaffolds allow

131

for an amide bond or triazole linkage, which in being more rigid and planar, may

limit the promiscuous binding characteristics of aminoglycosides due to the

flexibility of their glycosidic bonds. Additionally, the amide bond and triazole39

would allow for additional hydrogen bonding. In addition, with a new synthetic

scaffold, resistance enzymes and efflux pumps that recognize aminoglycosides

will likely not identify these molecules. In this chapter we focus on the synthesis

of the 2-DOS mimetics.

Figure 3.3: Comparison of 2-DOS (1) and potential analogs 10, 11, 12, and 13.

3.2 Results and Discussion

To synthesize the protected 3,5-diaminocyclohexane carboxylic acid (10),

we began with the symmetrical precursor cis,cis-1,3,5-cyclohexanetricarboxylic

acid (14), which was reacted with thionyl chloride in ethanol to form the triethyl

ester (15) (Scheme 3.1). Two of the ethyl esters were then hydrolyzed under basic conditions. The resulting dicarboxylic acid (16) was treated with diphenylphosphoryl azide under reflux conditions. Double Curtius rearrangement and subsequent in situ reaction with benzyl alcohol yielded the Cbz protected diamine (17). From intermediate 17, the Cbz groups were deprotected by hydrogenolysis, resulting in compound 18. Alternatively, the Cbz groups of 17

132

were exchanged for Boc groups (19) and subsequently the ethyl ester was

hydrolyzed to the carboxylic acid (20).

Scheme 3.1: Synthesis of 19 and 20.

The next 1,3,5- derivative was synthesized from the precursor cis, cis-

1,3,5-triazidecyclohexane, which was prepared according to published procedure.40 Cis,cis-1,3,5-cyclohexanetriol (21) was activated with tosyl chloride

to give rise to 22 (Scheme 3.2). The sulfonates were displaced to generate the

tri-azido compound 23. Staudinger reduction conditions, either with one

equivalent PPh3 or two equivalents PPh3 in the presence of di-tert-butyl

dicarbonate, converted 23 into 24 or 25, respectively. Triphenylphosphine oxide

often co-elutes with non-polar compounds, which can complicate purification.

133

However, the high polarity of 24 compared to triphenylphosphine oxide allows it

to be separated by flash chromatography. After 25 was subjected to two

equivalents of PPh3, it was then extracted from CH2Cl2 with 0.5 M HCl, which

easily separated the diamine from triphenylphosphine oxide. Then the aqueous layer containing the diamine was evaporated under reduced pressure and Boc protected to give rise to 25.

Scheme 3.2: Synthesis of 24 and 25.

Towards the synthesis of 12 and 13, we chose to start with 2,5- dideoxystreptmine (2) as a precursor, which made using established procedures.30 Epoxidation of 1,4-cyclohexadiene (25) at 0 °C yielded the cis-

bisepoxide 26 (Scheme 3.3). Reaction of 26 with hydrazine formed 27, and

subsequent hydrogenolysis yielded the diamine diol as an acetate salt 2. The

amines of 2 were then Boc-protected to give rise to 28. In order to retain the

relative stereochemistry of 2-DOS, 28 was subjected to Mitsunobu conditions to give diol 29. The diols were tosylated, and then 24 was reacted with an

134

tetrabutylammonium azide to yield 31 and 32* as a racemic mixture. Additionally,

28 was tosylated, and 33 was reacted with sodium azide to give rise to 34 and a racemic mixture of 35*.

Scheme 3.3: Synthesis of 31, 32*, 34, and 35*. A racemic mixture is designated by *.

The ditosylates 30 and 33 reactions with an azide source resulted in different products that can be explained by the potential conformations and mechanisms described in Scheme 3.4 and 3.5. The preferred conformation of 30

135

is likely with the –NHBoc groups being equatorial, as it is more sterically hindering than the –OTos groups. In this conformation, the hydrogen and leaving group are anti, and therefore elimination can lead to 31, or alternatively disubstitution leads to 32* (Scheme 3.4).

Scheme 3.4: Proposed mechanism resulting in 31 and 32*. A racemic mixture is designated by *.

For compound 33, the most stable conformation is with all substituents in an equatorial position (33a) (Scheme 3.5). However, to undergo a displacement reaction, the conformer 33b must be populated. After substitution by one azide, the intermediate 36 is formed, which has been isolated and confirmed by MS.

From 36, another displacement by an azide anion results in 34. To form 35, there must be a double inversion of the stereochemistry. Thus we speculate that from intermediate 36 the Boc protecting group displaces the tosylate resulting in the intermediate 37. Subsequent azide attack of 37 gives rise to 35. Neighboring- group participation by Boc groups and other carboxyl moieties has been previously reported.41-43 With the Boc-group trans to the tosylate leaving group in

36, this attack can occur. However this does not occur with compound 30, likely because the Boc group and -OTos are cis to one another, making such an attack unfavorable.

136

Scheme 3.5: Proposed mechanism resulting in 34 and 35*. The neighboring-group participation by the Boc protecting group is colored purple for clarity.

3.3 Conclusion

In conclusion, we have synthesized several 2-DOS mimetics that retain the 1,3-cis-diamine moiety. The meso character of these compounds allowed for

137

their straightforward synthesis, curbing the number of stereochemcial isomers

that can form.

3.4 Future Directions

Although most of compounds in the synthesis of the 2-DOS mimetics were

obtained in high yield, there is room to enhance some of the product yields. The

reaction of the ditosylate 30 with tetrabutylammonium azide can be optimized for the production of diazide 31 rather than the elimination product 32*. A preliminary

small scale reaction analyzed by NMR shows that using tetrabutylammonium

azide in dioxane at 80 °C favors formation 31 over 32* by a ratio of 70:30. The same reaction with sodium azide in dioxane showed no reaction, likely because of the lack of solubility of sodium azide. Additionally, the formation of diazides 34 and 35* from the ditosylate 33 should be optimized, as only a total of 49% yield of both products were isolated in our early attempts.

The elimination product 32* could also be further functionalized as a 2-

DOS mimic. Reactions of 32* could include epoxidation (38), Sharpless aminohydroxylation (39), elctrophillic addition (40), and dihydroxylation (41)

(Scheme 3.6). However, the regio- and stereo-chemical outcome of these reactions could be complex. The epoxide 38 could be opened up with a variety of nucleophiles as well.

138

Scheme 3.6: Possible reactions of 32*.

With these building blocks in hand, one could imagine reacting the

building blocks together or with an alkyne-modified sugar44 (42) to create

potential aminoglycoside mimetics, with examples shown in Scheme 3.7. These

building blocks could be used as precursors to make aminoglycoside analogs.

Using an amide or trizaole linkage could limit the conformational freedom of certain functional groups, possibly leading to more selective RNA binders.

Additionally, synthetic molecules could evade resistance mechanisms that

aminoglycosides are plagued by.

139

Scheme 3.7: Potential products that could be formed from the building blocks and an alkyne- modified sugar.

3.5 Experimental

Materials

Unless otherwise specified, materials purchased from commercial suppliers were used without further purification. Cis,cis-1,3,5-

cyclohexanetricarboxylic acid (14) and 1,4-cyclohexadiene (25) was purchased

from Sigma Aldrich. Cis,cis-1,3,5-cyclohexanetriol was purchased from TCI

America. NMR solvents were purchased from Cambridge Isotope Laboratories

(Andover, MA, USA).

Instrumentation

NMR spectra were recorded on Varian Mercury 300 and 400 MHz, Varian

VX 500 MHz, and Jeol ECA 500 MHz spectrometers. Mass spectra (MS) were

140 recorded at the University of California, San Diego Chemistry and Biochemistry

Mass Spectrometry Facility

Synthesis

Derivatives 2, 21, 22, and 23 were synthesized according a previously published procedure. 30,40

Triethyl-cis,cis-1,3,5-cyclohexanetricarboxylate (15). Cis,cis-1,3,5- cyclohexanetricarboyxlic acid (14) (6.00 g, 27.7 mmol) was added to a 500 mL two neck round bottom flask and dried overnight under vacuum. After under argon, one neck was fitted with a dropping funnel with an adapter for argon flow in and the other neck with a condenser with an adapter for outflow into a flask containing a 5% NaHCO3 solution to neutralize expelled HCl gas. Dry absolute ethanol was added and the starting material was dissolved with stirring. Then the reaction was cooled to 0°C in an ice bath. The SOCl2 (19.6 mL, 33.0 g, 277 mmol) was added to the dropping funnel, which was slowly added to the reaction over 30 min. The reaction was then warmed to 50 °C for 6 hours. The reaction was evaporated under reduced pressure resulting in an oil. The flask with the oil was put in an ice bath and hexanes were added. Scratching of the flask also facilitate precipitation the product. The product was filtered and rinsed with cold hexanes. Product: off-white solid (7.25 g, 26.6 mmol, 96% yield). 1H-NMR (400

141

MHz, CDCl3): 4.14 (q, J = 7.2 Hz, 6H), 2.37 (dt, J1 = 12.8 Hz, J2 = 4.2 Hz, 3H),

2.31 – 2.22 (m, 3H), 1.54 (q, J = 12.8 Hz, 3H), 1.26 (t, J = 7.2 Hz, 9H). 13C NMR

(100 MHz, CDCl3): 174.26, 60.75, 42.01, 30.56, 14.27. ESI-MS calculated for

+ C15H25O6 [M+H] 301.17, found 301.00.

Cis,cis-(5-ethylcarboxylate)cyclohexane-1,3-dicarboxylic acid (16). Cis,cis-

1,3,5-cyclohexanetriethylester (15) (4.86 g, 17.8 mmol) was added to a 1 L round

bottom flask fitted with a dropping funnel with 300 mL absolute ethanol. A fresh

solution of KOH (2.00 g, 35.6 mmol) in 89 mL, 33:67 water:abs. EtOH, 0.4 M)

was prepared and added slowly (over 5.5 hrs.) with vigorous stirring. The

reaction was stirred overnight and then most of the solvent was reduced under

pressure to greatly reduce the amount of EtOH. The slurry was acidified to ~pH 2

with HCl and then extracted with EtOAc x 3. The EtOAc was rinsed with brine

and then dried over MgSO4. Then the EtOAc was evaporated under reduced

pressure to give rise to a white solid. The white solid was then rinsed and

sonicated with CH2Cl2 x3 to dissolve the product and the tricarboxylic acid

remained solid. The CH2Cl2 was evaporated to give rise to the product.

Alternatively, flash chromatography could be used to purify the product using gradients from 6-15% EtOH in CH2Cl2. Product: white solid (3.13 g, 3.13 mmol,

1 72% yield). H-NMR (400 MHz, CDCl3): δ 4.14 (q, J = 7.2 Hz, 2H), 2.54 – 2.25

13 (m, 6H), 1.66 – 1.45 (m, 3H), 1.25 (t, J = 7.2 Hz, 3H); C NMR (75 MHz, CDCl3):

142

δ 180.36, 174.06, 61, 41.83, 41.58, 30.23, 30.06, 14.31; ESI-MS calculated for

- C11H15O6 [M-H] 243.09, found 243.15.

Ethyl-cis,cis-(3,5-Cbz2diamine)cyclohexane-1-carboxylate (17). Cis,cis-(5- ethylcarboxylate)cyclohexane-1,3-dicarboxylic acid (16) (1.15 g, 4.72 mmol) was

dried overnight in a 3 neck 100 mL round bottom flask. The flask was flushed

with argon and fitted with a condenser and a stir bar was added. Anhydrous

toluene (16 mL) was added to 8. Then DPPA (2.14 mL, 2.73 g, 9.02 mmol) and

NEt3 (1.45 mL, 1.05 g, 10.4 mmol) was added to the flask. The reaction was then heated to reflux for 1.5 hours and evolution of N2 was observed. Benzyl alcohol

(1.13 mL, 1.17 g, 10.9 mmol) was then added and the reaction was refluxed for

22 hours. The reaction was then cooled and left at 4 °C overnight and crystals formed. Then reaction was filtered and the crystals were pure 9. To increase the yield, the product was further isolated from the residue by flash chromatography

(1–1.5% MeOH). Product: white solid (1.41g, 3.11, 66%, yield) 1H- NMR (400

MHz, CDCl3): δ 7.45 – 7.28 (m, 10H), 5.08 (s, 4H), 4.64 (brd, J = 7.2 Hz, 2H),

4.11 (q, J = 7.2 Hz, 2H), 3.74 – 3.53 (m, 12H), 2.49 (t, J = 11.2 Hz, 1H), 2.30 (t,

J = 12 Hz, 3H), 1.25 – 1.16 (m, 5H), 1.01 (q, J =12 Hz, 1H). 13C-NMR(100 MHz,

CDCl3): δ 174.09, 155.52, 136.5, 128.68, 128.31, 66.87, 60.87, 48.22, 40.22,

143

+ 39.29, 34.76, 14.27. ESI-MS calculated for C25H31N2O6 [M+H] 455.21, found

454.93.

Cis,cis-(3,5-ethylcarboxylate)cyclohexane-1,3-diammonium acetate (18). To

the flask 10% Pd/C (0.108 g Pd/C, 0.101 mmol Pd), 10.2 mL of 90:10 of

EtOH:AcOH, and Ethyl-cis,cis-(3,5-Cbz2diamine)cyclohexane-1-carboxylate (17)

(0.461 g, 1.01 mmol) was added to 100 mL round bottom flask with a stir bar.

The flask was evacuated and then flushed with H2 x3. Then the reaction was left

under H2 for 4 hours. The reaction was then filtered over celite and the celite was rinsed with EtOH. The solvent was evaporated under reduced pressure. Product:

1 white solid (0.642 g, 0.952 mmol, 94% yield). H-NMR (400 MHz, D2O): δ 4.20 (q,

J = 7.2 Hz, 2H), 3.43 (dt, J1 = 12 Hz, J2 = 4 Hz, 2H), 2.72 (dt, J1 = 12.4 Hz, J2 =

3.6 Hz, 1H), 2.49 – 2.33 (m, 3H), 1,91 (s, 6H), 1.65 – 1.47 (m, 3H), 1.26 (t, J =

13 7.2 Hz, 3H). C-NMR (75 MHz, D2O) δ 182.02, 175.52, 63.02, 47.63, 39.42,

+ 34.03, 31.53, 23.99, 13.85. ESI-MS calculated for C9H19N2O2 [M+H] 187.14,

found 187.03.

Ethyl-cis,cis-(3,5-Boc2diamine)cyclohexane-1-carboxylate (19). To the flask

10% Pd/C (0.135 g Pd/C, 0.127 mmol Pd), 15.9 mL of EtOH, Ethyl-cis,cis-(3,5-

144

Cbz2diamine)cyclohexane-1-carboxylate (17) (0.577 g, 1.27 mmol), and Boc2O was added to a 50 mL round bottom flask with a stir bar. The flask was evacuated and then flushed with H2 x3. Then the reaction was left under H2 for 4 hours. The reaction turned into slurry so some CH2Cl2 was added to dissolve the precipitate. Then the reaction was then filtered over celite and the celite was rinsed with a mixture of CH2Cl2 and EtOH. The solvent was evaporated under reduced pressure. Product: white solid (0.437 g, 1.13 mmol, 89% yield). 1H-NMR

(400 MHz, CDCl3): δ 4.83 (brs, 1H), 4.05 (q, J = 7.2 Hz, 2H), 3.46 (s, 2H), 2.83

(s, 1H), 2.39 (t, J = 12.4 Hz, 1H), 2.15 (d, J = 11.2 Hz, 3H), 1.37 (s, 18H), 1.18 (t,

13 J = 7.2 Hz, 3H), 1.10 (q, J = 12.4 Hz, 2H), (q, J = , 1H); C (100 MHz, CDCl3): δ

174.57, 155.23, 79.54, 60.79, 47.55, 40.25, 39.15, 34.76, 28.36, 14.11. ESI-MS

+ calculated for C19H34N2O6Na [M+Na] 409.23, found 409.28.

Cis,cis-(3,5-Boc2diamine)cyclohexane-1-carboxylic acid (20). Ethyl-cis,cis-

(3,5-Boc2diamine)cyclohexane-1-carboxylate (19) (0.527 g, 1.36 mmol) and THF

(13.7 mL) were added to a 100 mL round bottom flask. To the flask 6.81 mL of a

3 M NaOH solution (0.818 g, 20.4 mmol) was added. The reaction was stirred for

2 hours and then evaporated under reduced pressure. Using 0.5 M HCl, the slurry was adjusted to pH 4. The aqueous layer was extracted with EtOAc x3 and then the organic layer was washed with brine. After drying with MgSO4, the

145

EtOAc was evaporated under reduced pressure. . Product: white solid (0.434 g,

1.21 mmol, 89% yield). 1H-NMR (400 MHz, DMSO): δ 12.17 (brs, 1H), 6.86 (brd,

J = 7.6 Hz, 2H), 3.33 – 3.18 (m, 2H), 2.31 (t, J = 12.5 Hz, 1H), 1.95 – 1.79 (m,

3H), 1.37 (s, 18H), 1.15 – 1.94 (m, 3H). 13C-NMR (125 MHz, DMSO): δ 175.46,

154.75, 77.55, 47.23, 38.67, 34.19, 29.31, 28.25. ESI-MS calculated for

- C17H30N2O6 [M-H] 357.20, found. 357.31.

Cis,cis-3,5-diazidocyclohexane-1-amine (24). Cis,cis-1,3,5- triazidocyclohexane (23) (0.169 g, 0.814 mmol) was added to a 25 mL round bottom flask with stir bar under argon. Caution! Polyazides are potentially explosive. THF (4 mL) and H2O (0.44 mL) was added to the starting material.

Triphenylphosphine (0.213 g, 0.814 mmol) was dissolved in THF and added

drop-wise to the reaction flask while stirring. The reaction was stirred overnight and then evaporated under reduced pressure. The product was isolated by flash

chromatography (4% MeOH in CH2Cl2 until all triphenylphosphine oxide is eluted,

and then the product is eluted with 12% MeOH in CH2Cl2). Product: yellow oil (

-1 1 0.093 g, 0.513 mmol, 63% yield). IR (neat): (NH), (N3), cm . H-NMR (400 MHz,

DMSO): δ 3.48 (dt, J1 = 12 Hz, J2 = 4 Hz, 2H), 2.63, (dt, J1 = 11.6 Hz, J2 = 4 Hz,

1H), 2.18 – 2.09 (m, 1H), 2.05 – 1.94 (m, 2H), 1.69 (brs, 2H), 1.16 (q, J = 11.6

13 Hz, 1H), 1.02 (q, J = 11.6 Hz, 2H). C NMR (125 MHz, CDCl3): δ 55.62, 46.29,

+ 40.26, 36.09. ESI-MS calculated for C6H12N7 [M+H] 182.12, found 181.98.

146

Cis,cis-5-diazidocyclohexane-1,3-Boc2amine (25). Cis,cis-1,3,5-

triazidocyclohexane (23) (0.162 g, 0.781 mmol) was added to a 25 mL round

bottom flask with stir bar under argon. Caution! Polyazides are potentially

explosive. THF (3.9 mL) and H2O (0.84 mL) was added to the starting material.

Triphenylphosphine (0.410 g, 1.56 mmol) was dissolved in THF and added drop-

wise to the reaction flask while stirring. The reaction was stirred overnight and

then evaporated under reduced pressure. To the 0.5 M HCl (35 mL) was added

and was washed with 5 mL CH2Cl2 x3. The aqueous layer was evaporated under

reduced pressure. Then to the residue DMF (7.8 mL), NEt3 (0.55 mL, 3.91 mmol,

0.395 g), and Boc2O (0.682 g, 3.13 mmol) was added and stirred overnight. The

reaction was evaporated under reduced pressure, and then CH2Cl2 (40) was added and washed with 5% NaHCO3 (6 mL) x3. The organic layer was evaporated under reduced pressure and the product was isolated by flash chromatography (a gradient of 0.5 – 5% MeOH in CH2Cl2). Product: white solid

(0.175 g, 0.492 mmol, 63% yield). IR (neat): 3355 (NH), 2093 (N3), 1679 (C=O)

-1 1 cm . H-NMR (400 MHz, CDCl3): δ 4.48 (brd, J = 6 Hz, 2H), 3.63 – 3.45 (m, 2H),

3.38 (dt, J1 = 12, J2= 4 Hz, 1H), 2.34 – 2.18 (m, 3H), 1.42 (s, 18H), 1.08 (q, 12

13 Hz, 2H), 0.97 (q, 12 Hz, 1H). C NMR (100 MHz, CDCl3): δ 154.94, 79.83,

+ 56.34, 46.38, 39.21, 38.14, 28.49. ESI-MS calculated for C6H12N7 [M+Na]

378.21, found 378.01.

147

(1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-diol (28). 2,5- dideoxystreptammonium acetate (2) (0.293 g, 1.10 mmol) was added to a 15mL round bottom flask. DMF (4.18 mL), H2O (1.32 mL) (24:76), NEt3 (0.77 mL, 0.557

g, 5.50 mmol), and then di-tert-butyl dicarbonate (0.961 g, 4.40 mmol) was added

to the flask. The reaction was stirred overnight and concentrated. Water and

minimal CH2Cl2 was added, and the solid was sonicated then filtered. The solid

was washed with water several times and then dried. Product: white solid (0.368

g, 1.06 mmol, 96% yield). 1H-NMR (400M Hz, DMSO): δ 6.54 (brd, J=7.2Hz, 2H),

4.56 (d, J=5.6 Hz, 2H), 3.21–3.14 (m, 2H), 3.07 (m, 2H), 1.99 (dt, J1 = 12Hz, J2 =

4Hz, 1H), 1.80 (d, =12 Hz, 1H), 1.37 (s, 18H), 1.23 (q, J=12 Hz, 1H), 1.00 (q,

J=12Hz, 1H); 13C-NMR (75 MHz, APT, DMSO): δ 155.35, 77.39, 68.83, 54.40,

+ 28.35, 28.27; ESI-MS calculated for C16H30N2O6Na [M+Na] 369.20, found

369.09. (Large scale: 8.81 g, 33.1 mmol; yield 76% 8.72 g, 25.14 mmol).

(1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3-diol (29). (1R,3S,4S,6R)- 4,6-

Boc2diaminocyclohexane-1,3-diol (28) (2.02 g, 5.83 mmol), triphenylphosphine

(6.34 g, 23.3 mmol) and p-nitrobenzoic acid (3.89 g, 23.3 mmol) was added to a

250 mL round bottom flask fitted with a reflux condenser under argon. 125 mL of

THF previously passed over basic alumina was added and cooled to 0 °C in an

148

ice bath. Slowly, 4.6 mL (4.71 g, 23.3 mmol) of diisopropyl azodicarboxylate

(DIAD) was added. The reaction was warmed to room temperature, and then

heated to reflux for 16 hours. The reaction was cooled and precipitate was

filtered off. The THF was evaporated under reduced pressure. MeOH (100 mL)

was added to the residue and then K2CO3 (2.67 g, 69.9 mmol) and was stirred for

1.5 hours. The reaction was evaporated under reduced pressure. The product

was purified by automated flash chromatography (30-50% hexanes over 5 min so

the triphenylphosphine was eluted, then 50-60% over 30 min) and the product

eluted after 13 min. 1H-NMR (500 MHz, DMSO): δ 6.37 (d, J = 8 Hz, 2H), 4.79 (d,

J = 6 Hz, 2H), 3.79–3.68 (m, 2H), 3.46–3.33 (m, 2H), 1.92 (dt, J1 = 14.4 Hz, J2 =

3.2 Hz, 1H), 1.73 (q, J = 12.4 Hz, 1H), 1.59 (q, J = 14.8 Hz, 1H), 1.74–1.31 (m,

19H). 13C-NMR (125 MHz, DMSO): δ 154.75, 77.76, 67.61, 51.30, 48.65, 34.68,

28.27, 27.35. Product: white solid (1.30 g, 3.73 mmol, 64% yield). ESI-MS

+ calculated for C16H31N2O6 [M+H] 347.22, found 346.93.

(1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3-Tos2diol (30).

(1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3-diol (28) (0.432 g, 1.25 mmol) was added to a dry Erlenmeyer flask with a septa under argon. Freshly distilled pyridine (9.7 mL) was added to the flask, and it was cooled to 0°C in an ice bath.

Then p-toluenesulfonyl chloride (2.37 g, 12.5 mmol) was added and the reaction was at 4°C for 24 hours. Since there was still starting material by TLC after 24

149

hours, 3.37 g (17.7 mmol) of p-toluenesulfonyl chloride was added was at 4°C for

two more days until all starting material was gone. The reaction was poured into

a 100 g ice bath, filtered and rinsed with cold 0.1 M HCl and water. Product:

white solid (0.546 g, 0.833 mmol, 67% yield). 1H-NMR (500 MHz, DMSO): δ 7.78

(d, J = 8 Hz, 4H), 7.42 (d, J = 8.5 Hz, 4H), 6.87 (d, J = 7.5 Hz, 2H), 4.47 (s, 2H),

3.62–3.53 (m, 2H), 2.41 (s, 6H), 1.99 (d, J = 16 Hz, 1H), 1.94–1.80 (m, 2H),

1.40–1.21 (m, 19H). 13C-NMR (125 MHz, DMSO): δ 154.58, 144.28, 133.17,

129.92, 127.91, 77.811, 76.36, 49.36, 31.44, 27.96, ESI-MS calculated for

+ C30H46N3O10S2 [M+NH4] 672.26, found 672.12.

(1S,3R,4R,6S)-4,6-diazidocyclohexane-1,3-Boc2diamine (31). (1R,3S,4R,6S)-

4,6-Boc2diaminocyclohexane-1,3-Tos2diol (30) (0.868 g, 1.33 mmol) was put into was put into a 50 mL round bottom flask. DMF (13.3 mL) passed through silica was added. Then N(Bu)4N3 (1.89 g, 6.65 mmol) was added and the reaction was

heated to 80 °C. The reaction was stirred overnight. The solvent was evaporated

under reduced pressure. Ethyl acetate was added and was washed with water x2

and brine once. The organic layer was dried over MgSO4, filtered, and

evaporated under reduced pressure. The product was isolated using automated

flash chromatography (0 - 7% dichloromethane in ethyl acetate over 37 mins)

eluted after 18 min. Product: white solid (0.100 g, 0.253 mmol, 19% yield). IR

-1 1 (neat): 3281 (NH), 2097 (N3), 1662 (C=O) cm . H-NMR (400 MHz, CDCl3): δ

150

4.61 (brs, 1H), 3.52 – 3.26 (m, 4H), 2.40 – 2.22 (m, 2H), 1.56 – 1.33 (m, 22H).

+ ESI-MS calculated for C16H28N8O4Na [M+Na] 419.21, found 419.2.

(1S,3R,4S)-4,6-diazidocyclohex-5-ene-1,3-Boc2diamine (32*). The product

was isolated as a racemic mixture from the same reaction conditions for 30. The

product was isolated using automated flash chromatography (0 - 7%

dichloromethane in ethyl acetate over 37 mins) eluted after 25 min. Product:

1 white solid (0.250 g, 0.665 mmol, 50% yield). H-NMR (400 MHz, CDCl3): δ 5.79

(d, J = 10 Hz, 1H), 5.66 (dt, J1 = 10 Hz, J2 = 2 Hz, 1H), 4.73 (s, 1H), 4.50 (s, 1H),

4.34 (s, 1H), 3.93 (s, 1H), 3.74 – 3.58 (m, 1H), 2.41 – 2.27 (m, 1H), 1.59 – 1.35

-1 (m, 19H). IR (neat): 3350 (NH), 2095 (N3), 1678 (C=O) cm . ESI-MS calculated

+ for C16H27N5O4Na [M+Na] 376.20, found 376.2.

(1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-Tos2diol (33).

(1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-diol (28) (0.508 g, 1.47 mmol) was added to a dry 25 mL flask with under argon. Freshly distilled pyridine (10 mL) was added to the flask, and it was cooled to 0°C in an ice bath. Then p- toluenesulfonyl chloride (2.80 g, 14.7 mmol) was added and the reaction was at

4°C for 21 hours. The reaction was poured into a 110 g ice bath, filtered and

151

rinsed with cold 0.1 M HCl and water. Product: white solid (0.802 g, 1.23 mmol,

1 83% yield). H-NMR (400 MHz, CDCl3): δ 7.74 (d, J = 8.4 Hz, 4H), 7.33 (d, J =

8.4 Hz, 4H), 4.57 (s, 2H), 4.36 (s, 2H), 3.62 – 3.46 (m, 2H), 2.45 (s, 6H), 2.35 (dt,

13 J1 = 13.6 Hz, J2 = 4.4 Hz, 1H), 2.31 – 2.21 (m, 1H), 1.45 – 1.35 (m, 19H). C-

NMR (75 MHz, DMSO): δ 155.01, 145.2, 133.47, 130.03, 127.82, 79.88, 77.44,

51.50, 36.13, 28.39, 21.8, 21.77. ESI-MS calculated for C30H42N2O10S2Na

[M+Na]+ 677.22, found 677.2.

(1S,3R,4S,6R)-4,6-diazidocyclohexane-1,3-Boc2diamine (34). (1R,3S,4S,6R)-

4,6-Boc2diaminocyclohexane-1,3-Tos2diol (33) (0.542 g, 0.827 mmol) was put

into a 50 mL round bottom flask. DMF (8.3 mL) passed through silica was added.

Then NaN3 (0.403 g, 6.20 mmol) was added and the reaction was heated to 80

°C. The reaction was stirred for 5 days. The solvent was evaporated under

reduced pressure. Ethyl acetate was added and was washed with water x2 and

brine once. The organic layer was dried over MgSO4, filtered, and evaporated

under reduced pressure. The product was isolated using automated flash

chromatography (0 - 50% ethyl acetate in hexanes over 30 mins) eluted after 14

min. Product: white solid (g, 0.086g, 0.217 mmol, 26% yield). IR (neat): 3350

-1 1 (NH), 2095 (N3), 1684 (C=O) cm . H-NMR (300 MHz, CDCl3): 4.79 (brd, J = 7.2

Hz, 2H), 3.97 (s, 2H), 3.78 – 3.62 (m, 2H), 2.46 – 2.30 (m, 1H), 1.87 (dt, J1 = 15.9

13 Hz, J2 = 3.3 Hz, 1H), 1.71 – 1.62 (m, 2H), 1.42 (s, 18H). C-NMR (75 MHz,

152

CDCl3): δ 154.89, 80.18, 57.85, 50.27, 30.03, 28.82, 28.42. ESI-MS calculated

+ for C16H28N8O4Na [M+Na] 419.21, found 419.2.

(1S,3R,4R,6R)-4,6-diazidocyclohexane-1,3-Boc2diamine (35*). The product

was isolated as a racemic mixture from the same reaction conditions for 34. The

product was isolated using automated flash chromatography (0 - 50% ethyl

acetate in hexanes over 30 mins) eluted after 16 min. Product: white solid (g,

0.076g, 0.191 mmol, 23% yield). IR (neat): 3332 (NH), 2096 (N3), 1683 (C=O)

-1 1 cm . H-NMR (300 MHz, CDCl3) δ 4.67 (brs, 1H), 4.57 (brs, 1H), 4.10 – 4.04 (m,

1H), 3.81 – 3.64 (m, 1H), 3.54 – 3.37 (m, 2H), 3.33 – 3.21 (m, 1H), 2.04 – 1.95

(m, 1H), 1.70 –1.62 (m, 2H), 1.45 (s, 9H), 1.44 (s, 9H). 13C-NMR (100 MHz,

CDCl3) δ 155.24, 154.99, 80.41, 60.58, 59.08, 53.01, 50.30, 33.44, 32.94, 28.54.

+ ESI-MS calculated for C16H28N8O4Na [M+Na] 419.21, found 419.2.

(1S,3R,4R,6R)-4-Toshydroxyl-6-diazidocyclohexane-1,3-Boc2diamine (36*).

+ ESI-MS calculated for C23H35N5O7SNa [M+Na] 548.22, found 548.2.

153

Spectrum 3.1: Triethyl-cis,cis-1,3,5-cyclohexanetricarboxylate (15).

032009 _ LMN 131 113 412 404 395 380 372 364 349 341 333 279 245 590 558 526 494 274 256 239 149 167 260 ...... hg 402 , CDCl 3 4 4 2 2 2 2 2 2 2 2 2 2 2 1 1 1 1 1 1 1 4 4 7 239 256 274 494 113 412 404 395 380 372 364 131 349 526 341 149 333 167 558 279 590 245 648 ...... 1 1 1 1 4 2 2 2 2 2 2 4 2 1 2 4 2 4 1 2 1 2 1 16 26 07 24 00 . . . . . 3 3 9 3 6

4 .1 3 .8 3 .5 3 .2 2 .9 2 .6 2 .3 2 .0 1 .7 1 .4 δ (ppm ) 00 24 16 26 07 . . . . . 6 3 3 3 9

7 .5 7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0 δ (ppm ) 255 . 478 160 842 750 014 559 032109 _ 13 C _ LMN 274 ...... 402 CDCl 3 174 77 77 76 60 42 30 hg , 14

CHCl 3

190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 δ (ppm )

154

Spectrum 3.2: Cis,cis-(5-ethylcarboxylate)cyclohexane-1,3-dicarboxylic acid (16). 231 254 278 527 568 299 347 399 440 107 131 155 178 260 945 ...... 1 1 1 1 1 2 2 2 2 4 4 4 4 7 8 231 254 278 527 568 299 347 399 440 107 131 155 178 ...... 1 1 1 1 1 2 2 2 2 4 4 4 4 041609 _ LMP hg 300 , CDCl 3 40 95 90 95 . . . . 3 2 5 1

4 .2 4 .0 3 .8 3 .6 3 .4 3 .2 3 .0 2 .8 2 .6 2 .4 2 .2 2 .0 1 .8 1 .6 1 .4 1 .2 δ (ppm ) 40 95 90 95 00 . . . . . 3 2 5 1 2

11 .5 10 .5 9 .5 9 .0 8 .5 8 .0 7 .5 7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0 0 .5 0 .0 δ (ppm ) 061 362 . . 313 057 232 580 834 001 736 160 041613 _ 13 C _ LMP 583 ...... 300 CDCl 3 14 30 30 41 41 61 76 77 77 174 180 hg ,

190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 δ (ppm )

155

Spectrum 3.3: Ethyl-cis,cis-(3,5-Cbz2diamine)cyclohexane-1-carboxylate (17).

080409 _ LMT _ recrys 361 349 338 317 260 084 650 632 141 123 105 087 645 515 487 459 325 295 567 256 022 992 962 895 878 861 265 172 ...... 7 7 7 7 7 5 4 4 4 4 4 4 3 2 2 2 2 2 1 1 1 0 0 0 0 0 hg 402 , CDCl 3 2 2 487 459 295 265 172 567 256 220 190 159 022 992 084 650 632 141 123 105 087 645 ...... 2 2 2 2 2 1 1 1 1 1 1 0 5 4 4 4 4 4 4 3 82 03 97 36 00 64 97 28 ...... 0 3 4 1 4 1 1 2

5 .2 4 .9 4 .6 4 .3 4 .0 3 .7 2 .5 2 .3 2 .1 1 .9 1 .7 1 .5 1 .3 1 .1

water

CHCl 3 hexanes

acetone hexanes 31 . 00 64 97 28 82 03 97 36 ...... 10 4 1 1 2 0 3 4 1

7 .5 7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0 0 .5 309 680 504 515 086 . . . . . 271 755 295 216 225 868 871 905 160 414 ...... 14 34 39 40 48 60 66 76 77 77 128 128 136 155 174

061109 _ LMT _ 13 C eca 500 , CDCl 3

190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0

156

Spectrum 3.4: Cis,cis-(3,5-ethylcarboxylate)cyclohexane-1,3-diammonium acetate (18). 243 261 279 496 527 559 585 614 914 367 399 439 688 697 711 720 728 742 751 391 401 411 422 432 441 452 462 472 172 190 208 226 790 ...... 1 1 1 1 1 1 1 1 1 2 2 2 2 2 2 2 2 2 2 3 3 3 3 3 3 3 3 3 4 4 4 4 4

090909 _ LMX hg 400 , D 2 O 679 688 697 711 720 728 742 751 760 391 401 411 422 432 441 452 462 472 ...... 2 2 2 2 2 2 2 2 2 3 3 3 3 3 3 3 3 3 00 11 . . 1 2

3 .50 3 .46 3 .42 3 .38 2 .80 2 .75 2 .70 2 .65 δ (ppm ) δ (ppm ) 19 24 83 24 00 11 13 ...... 3 3 5 3 1 2 2

4 .9 4 .6 4 .3 4 .0 3 .7 3 .4 3 .1 2 .8 2 .5 2 .2 1 .9 1 .6 1 .3 δ (ppm )

016 519 090909 _ LMX _ 13 C _ dioxane . . 846 988 190 018 629 421 027 528 . . hg 402 , D 2 O ...... 13 23 67 63 47 39 34 31 182 175

dioxane

180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10

157

Spectrum 3.5: Ethyl-cis,cis-(3,5-Boc2diamine)cyclohexane-1-carboxylate (19). 943 972 180 367 133 161 356 387 418 829 459 023 041 058 076 826 260 051110 _ LMAD _ 25 mg ...... 0 0 1 1 2 2 2 2 2 2 3 4 4 4 4 4 7 hg 400 , CDCl 3 884 913 943 972 056 087 118 150 180 197 ...... 0 0 0 0 1 1 1 1 1 1 91 07 46 . . . 0 2 3

1 .25 1 .20 1 .15 1 .10 1 .05 1 .00 0 .95 0 .90 0 .85 δ (ppm ) 82 . 91 07 46 15 96 24 85 00 94 ...... 0 2 3 18 3 0 1 1 2 0

7 .2 6 .8 6 .4 6 .0 5 .6 5 .2 4 .8 4 .4 4 .0 3 .6 3 .2 2 .8 2 .4 2 .0 1 .6 1 .2 δ (ppm ) 571 235 . . 535 477 160 839 792 555 246 152 761 356 051110 _ LMAD _ 25 mg _ 13 C _ apt 109 ...... 400 CDCl 3 174 155 79 77 77 76 60 47 40 39 34 28 hg , 14

170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 δ (ppm )

158

Spectrum 3.6: Cis,cis-(3,5-Boc2diamine)cyclohexane-1-carboxylic acid (20).

166 052610 _ LMAF . 995 026 060 372 757 283 314 500 285 598 850 869 ...... 402 DMSO . 0 1 1 1 1 2 2 2 3 3 6 6 12 hg , 995 026 060 091 122 372 757 819 860 891 283 314 345 500 285 333 ...... 0 1 1 1 1 1 1 1 1 1 2 2 2 2 3 3

DMSO THF water 25 . 21 35 00 05 . . . . 3 18 3 1 2

3 .4 3 .2 3 .0 2 .8 2 .6 2 .4 2 .2 2 .0 1 .8 1 .6 1 .4 1 .2 1 .0

DMSO THF water THF 25 . 21 35 00 05 88 30 ...... 3 18 3 1 2 1 1 12 .5 11 .5 10 .5 9 .5 9 .0 8 .5 8 .0 7 .5 7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0 0 .5 749 459 . . 252 313 194 667 513 228 549 ...... 28 29 34 38 39 47 77 154 175

051510 _ LMAF _ 6 mg _ 13 C _ d 1 = 3 vx 500 , DMSO

180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30

159

Spectrum 3.7: Cis,cis-3,5-diazidocyclohexane-1-amine (24). 123 118 005 976 693 203 173 144 115 063 034 004 975 141 146 500 596 606 616 625 634 663 653 644 492 482 472 462 452 442 329 ...... 2 2 2 1 1 1 1 1 1 1 1 1 0 2 2 2 2 2 2 2 2 2 2 2 3 3 3 3 3 3 3

072810 _ LMAO hg 400 , DMSO 522 512 502 492 482 472 462 452 442 596 606 616 625 634 644 653 663 672 ...... 3 3 3 3 3 3 3 3 3 2 2 2 2 2 2 2 2 2 00 . 94 1 . 1

3 .55 3 .50 3 .45 3 .40 2 .70 2 .65 2 .60 2 .55 δ (ppm ) δ (ppm )

water 86 08 01 02 00 06 02 ...... 1 1 2 2 1 1 2

3 .6 3 .4 3 .2 3 .0 2 .8 2 .6 2 .4 2 .2 2 .0 1 .8 1 .6 1 .4 1 .2 1 .0 δ (ppm ) 620 293 259 022 854 687 520 353 187 020 091 ...... 072813 _ LMAO _ 13 C . 55 46 40 40 39 39 39 39 39 39 vx 500 , DMSO 36

56 55 54 53 52 51 50 49 48 47 46 45 44 43 42 41 40 39 38 37 36 δ (ppm )

160

Spectrum 3.8: Cis,cis-5-diazidocyclohexane-1,3-Boc2amine (25). 920 950 979 009 099 423 207 237 271 301 340 350 361 370 380 390 399 409 535 475 475 490 260 092410 LMAY ...... _ 0 0 0 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 4 4 4 7 hg 400 , CDCl 3 340 350 361 370 380 390 399 409 419 920 950 979 009 039 069 099 129 ...... 3 3 3 3 3 3 3 3 3 0 0 0 1 1 1 1 1 03 00 04 . . . 1 1 2

3 .44 3 .40 3 .36 3 .32 1 .20 1 .10 1 .00 0 .90 δ (ppm ) δ (ppm ) 79 . 00 04 07 03 95 91 ...... 1 2 18 3 1 1 1

7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0 δ (ppm )

943 092410 _ LMAY _ 13 C . 827 478 160 842 339 376 205 138 489 ...... hg 402 , CDCl 3 . 154 79 77 77 76 56 46 39 38 28

155 145 135 125 115 105 95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 δ (ppm )

161

Spectrum 3.9: (1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-diol (28). 951 981 012 185 372 528 782 894 976 007 500 729 889 077 528 565 551 318 547 040411 dibocdiaminediol 6 .4 ...... _ _ mg 0 0 1 1 1 1 1 1 1 2 2 2 2 3 3 6 4 4 6 hg 402 , DMSO

water

DMSO 85 . 00 29 01 04 16 29 13 98 ...... 1 1 18 1 1 2 2 2 1 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0

162

Spectrum 3.10: (1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3-diol (29).

122013 LMBY 14 .7 373 438 463 576 716 893 900 916 922 500 156 167 355 721 727 108 118 129 139 793 806 359 375 _ _ mg ...... 1 1 1 1 1 1 1 1 1 2 3 3 3 3 3 4 4 4 4 4 4 6 6 vx 500 , DMSO

water

DMSO MeOH

MeOH 69 . 00 01 01 90 92 87 68 ...... 18 1 1 1 1 1 1 1

6 .2 5 .8 5 .4 5 .0 4 .6 4 .2 3 .8 3 .4 3 .0 2 .6 2 .2 1 .8 1 .4 δ (ppm )

750 122013 _ LMBY _ 14 .7 _ 13 C . 346 269 680 019 353 687 654 302 605 757 ...... vx 500 , DMSO . 27 28 34 39 39 39 48 51 67 77 154

DMSO

MeOH

155 145 135 125 115 105 95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 δ (ppm )

163

Spectrum 3.11: (1R,3S,4R,6S)-4,6-Boc2diaminocyclohexane-1,3- Tos2diol (30). 783 767 429 412 873 858 764 473 044 030 016 001 574 351 500 412 987 906 881 855 329 289 184 170 155 ...... 7 7 7 7 6 6 5 4 4 4 4 4 3 3 2 2 1 1 1 1 1 1 1 1 1

122013 _ LMCA _ 6 .7 mg _ 1 H DMSO - d 6 , vx 500 Spectra NB # 3 , p .58 D

water

DMSO

EtOAc

EtOAc EtOAc DCM 28 . 00 10 92 13 05 98 44 01 ...... 4 4 1 2 2 5 1 2 19

8 .0 7 .5 7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 1 .0 914 921 171 281 578 415 ...... 148 838 228 963 135 439 019 186 353 520 355 827 361 811 ...... 14 20 21 27 28 31 39 39 39 39 49 59 76 77 127 129 133 144 154 170

DMSO 122013 _ LMCA _ 6 .7 mg _ 13 C DMSO - d 6 , vx 500 Spectra NB # 3 , p .58 E

EtOAc EtOAc EtOAc

170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20

164

Spectrum 3.12: (1S,3R,4R,6S)-4,6-diazidocyclohexane-1,3-Boc2diamine (31). 26 61 39 35 31 27 62 44 ...... 060313 6 .8 frac 47 - 48 . 7 4 3 2 2 2 1 _ LMBZpure _ mg _ 1 hg 402 , CDCl 3

CHCl 3 water 19 . 87 00 15 . . . 1 4 2 22 7 .2 6 .8 6 .4 6 .0 5 .6 5 .2 4 .8 4 .4 4 .0 3 .6 3 .2 2 .8 2 .4 2 .0 1 .6

165

Spectrum 3.13: (1S,3R,4S)-4,6-diazidocyclohex-5-ene-1,3-Boc2diamine (32*). 260 804 779 680 675 670 655 650 645 726 501 344 088 932 668 647 355 324 041 643 453 436 ...... 7 5 5 5 5 5 5 5 5 4 4 4 4 3 3 3 2 2 2 1 1 1

060313 _ LMCBpure _ 7 .4 mg _ frac 37 - 38 402 CDCl 3 804 779 680 675 670 655 650 hg , 645 ...... 5 5 5 5 5 5 5 5 99 00 . . 0 1

5 .80 5 .75 5 .70 5 .65

CHCl 3

EtOAc

EtOAc 71 . 99 00 89 88 10 81 01 05 ...... 0 1 0 0 1 0 1 1 19

7 .2 6 .8 6 .4 6 .0 5 .6 5 .2 4 .8 4 .4 4 .0 3 .6 3 .2 2 .8 2 .4 2 .0 1 .6

166

Spectrum 3.14: (1R,3S,4S,6R)-4,6-Boc2diaminocyclohexane-1,3-Tos2diol (33).

053011 LMCD 8 .8 751 730 345 324 260 571 364 557 532 451 367 356 344 333 323 247 806 776 745 715 _ _ mg 401 ...... 7 7 7 7 7 4 4 3 3 2 2 2 2 2 2 2 1 1 1 1 hg 402 , CDCl 3 1

CHCl 3 37 . 06 13 61 97 00 09 38 18 35 ...... 4 4 1 1 2 6 1 1 1 19

7 .5 7 .0 6 .5 6 .0 5 .5 5 .0 4 .5 4 .0 3 .5 3 .0 2 .5 2 .0 1 .5 δ (ppm ) 013 202 072 474 033 819 ...... 797 773 387 884 584 436 160 736 497 128 ...... 21 21 28 155 145 144 133 130 127 79 77 77 77 76 51 36

072011 _ LMCD _ 13 C _ 59 .8 mg hg 300 , CDCl 3 CHCl 3

155 145 135 125 115 105 95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 20 δ (ppm )

167

Spectrum 3.15: (1S,3R,4S,6R)-4,6-diazidocyclohexane-1,3-Boc2diamine (34).

102411 _ LMCE _ 44 .7 mg 260 804 780 967 705 395 343 898 856 833 643 422 ...... 7 4 4 3 3 2 hg 300 , CDCl 3 2 1 1 1 1 1

CHCl 3 58 . 94 07 00 00 17 03 ...... 1 2 2 1 1 2 18

7 .2 6 .8 6 .4 6 .0 5 .6 5 .2 4 .8 4 .4 4 .0 3 .6 3 .2 2 .8 2 .4 2 .0 1 .6 δ (ppm ) 892

. 102711 _ LMCE _ 44 .7 mg _ apt 180 582 160 734 851 268 033 822 422 ...... hg 300 , CDCl 3 154 80 77 77 76 57 50 30 28 28 033 822 422 . . . 30 28 28

30 .5 30 .0 29 .5 29 .0 28 .5 28 .0 δ (ppm )

CHCl 3

155 145 135 125 115 105 95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 δ (ppm )

168

Spectrum 3.16: (1S,3R,4R,6R)-4,6-diazidocyclohexane-1,3-Boc2diamine (35*). 238 256 438 603 638 989 042 251 287 464 722 062 078 110 128 565 665 260 080911 _ LMCK _ 4 .6 ...... mg 1 1 1 1 1 1 2 2 2 3 3 4 4 4 4 4 4 7 hg 300 , CDCl 3

CDCl 3 H 2 O EtOAc

EtOAc

EtOAc 48 16 15 16 12 04 00 31 01 96 ...... 9 9 2 1 1 2 1 1 1 0

7 .4 7 .0 6 .6 6 .2 5 .8 5 .4 5 .0 4 .6 4 .2 3 .8 3 .4 3 .0 2 .6 2 .2 1 .8 1 .4 δ (ppm )

010313 _ LMCK _ 13 C 240 990 . . 415 574 256 938 583 082 007 302 445 935 536

hg 400 , CDCl 3 ...... 155 154 80 77 77 76 60 59 53 50 33 32 28

CDCl 3

155 145 135 125 115 105 95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 δ (ppm )

169

3.6 References

(1) Ecker, D. J.; Griffey, R. H. Drug Discov Today 1999, 4, 420.

(2) Hermann, T.; Tor, Y. Expert Opin. Ther. Patents 2005, 15, 49.

(3) Thomas, J. R.; Hergenrother, P. J. Chem Rev 2008, 108, 1171.

(4) Hermann, T. Current Opinion in Structural Biology 2005, 15, 355.

(5) Tenson, T.; Mankin, A. Molecular Microbiology 2006, 59, 1664.

(6) Yonath, A. Annual Review of Biochemistry 2005, 74, 649.

(7) Moazed, D.; Noller, H. F. Nature 1987, 327, 389.

(8) Hermann, T. Cell. Mol. Life Sci. 2007, 64, 1841.

(9) Chittapragada, M.; Roberts, S.; Ham, Y. W. Perspectives in Medicinal Chemistry 2009, 2009, 21.

(10) Sutcliffe, J. A. Current Opinion in Microbiology 2005, 8, 534.

(11) Roberts, S.; Chittapragada, M.; Pendem, K.; Leavitt, B. J.; Mahler, J. W.; Ham, Y. W. Tetrahedron Lett 2010, 51, 1779.

(12) The Molecular Basis of Antibiotic Action; Gale, E. F.; Cundliffe, E.; Renolds, P. E.; Richmond, M. H.; Waring, M. J., Eds.; John Wiley & Sons: London, 1981.

(13) Feldman, M. B.; Terry, D. S.; Altman, R. B.; Blanchard, S. C. Nature Chemical Biology 2010, 6, 54.

(14) Wang, H.; Tor, Y. Journal of the American Chemical Society 1997, 119, 8734.

(15) Botto, R. E.; Coxon, B. Journal of the American Chemical Society 1983, 105, 1021.

(16) Szilagyi, L.; Pusztahelyi, Z. S.; Jakab, S.; Kovacs, I. Carbohydrate Research 1993, 247, 99.

(17) Francois, B.; Russell, R. J. M.; Murray, J. B.; Aboul-ela, F.; Masquida, B.; Vicens, Q.; Westhof, E. Nucleic Acids Res. 2005, 33, 5677.

170

(18) Yoshizawa, S.; Fourmy, D.; Eason, R. G.; Puglisi, J. D. Biochemistry 2002, 41, 6263.

(19) Vicens, Q.; Westhof, E. Biopolymers 2003, 70, 42.

(20) Shakya, T.; Wright, G. D. In Aminoglycoside Antibiotics: From Chemical Biology to Drug Discovery; Arya, D. P., Ed.; John Wiley & Sons, Inc.: Hoboken, 2007.

(21) Mingeot-Leclercq, M. P.; Glupczynski, Y.; Tulkens, P. M. Antimicrobial Agents and Chemotherapy 1999, 43, 727.

(22) Umezawa, H.; Umezawa, S.; Tsuchiya, T.; Okazaki, Y. J. Antibiot. 1971, 24, 485.

(23) Umezawa, S.; Tsuchiya, T.; Muto, R.; Nishimur.Y; Umezawa, H. J. Antibiot. 1971, 24, 274.

(24) Aggen, J. B.; Armstrong, E. S.; Goldblum, A. A.; Dozzo, P.; Linsell, M. S.; Gliedt, M. J.; Hildebrandt, D. J.; Feeney, L. A.; Kubo, A.; Matias, R. D.; Lopez, S.; Gomez, M.; Wlasichuk, K. B.; Diokno, R.; Miller, G. H.; Moser, H. E. Antimicrobial Agents and Chemotherapy 2010, 54, 4636.

(25) Greenberg, W. A.; Priestley, E. S.; Sears, P. S.; Alper, P. B.; Rosenbohm, C.; Hendrix, M.; Hung, S. C.; Wong, C. H. Journal of the American Chemical Society 1999, 121, 6527.

(26) Hanessian, S.; Tremblay, M.; Kornienko, A.; Moitessier, N. Tetrahedron 2001, 57, 3255.

(27) Haddad, J.; Kotra, L. P.; Llano-Sotelo, B.; Kim, C.; Azucena, E. F.; Liu, M. Z.; Vakulenko, S. B.; Chow, C. S.; Mobashery, S. Journal of the American Chemical Society 2002, 124, 3229.

(28) Ding, Y. L.; Hofstadler, S. A.; Swayze, E. E.; Griffey, R. H. Chemistry Letters 2003, 32, 908.

(29) Wang, X. J.; Migawa, M. T.; Sannes-Lowery, K. A.; Swayze, E. E. Bioorg. Med. Chem. Lett. 2005, 15, 4919.

(30) Kavadias, G.; Velkof, S.; Belleau, B. Can. J. Chem.-Rev. Can. Chim. 1978, 56, 404.

(31) Vourloumis, D.; Winters, G. C.; Simonsen, K. B.; Takahashi, M.; Ayida, B. K.; Shandrick, S.; Zhao, Q.; Han, Q.; Hermann, T. Chembiochem 2005, 6, 58.

171

(32) Vourloumis, D.; Takahashi, M.; Winters, G. C.; Simonsen, K. B.; Ayida, B. K.; Barluenga, S.; Qamar, S.; Shandrick, S.; Zhao, Q.; Hermann, T. Bioorg. Med. Chem. Lett. 2002, 12, 3367.

(33) Udumula, V.; Chittapragada, M.; Marble, J. B.; Dayton, D. L.; Ham, Y. W. Bioorg. Med. Chem. Lett. 2011, 21, 4713.

(34) Zhou, Y. F.; Chow, C.; Murphy, D. E.; Sun, Z. X.; Bertolini, T.; Froelich, J. M.; Webber, S. E.; Hermann, T.; Wall, D. Bioorg. Med. Chem. Lett. 2008, 18, 3369.

(35) Zhou, Y. F.; Gregor, V. E.; Ayida, B. K.; Winters, G. C.; Sun, Z. X.; Murphy, D.; Haley, G.; Bailey, D.; Froelich, J. M.; Fish, S.; Webber, S. E.; Hermann, T.; Wall, D. Bioorg. Med. Chem. Lett. 2007, 17, 1206.

(36) Simonsen, K. B.; Ayida, B. K.; Vourloumis, D.; Winters, G. C.; Takahashi, M.; Shandrick, S.; Zhao, Q.; Hermann, T. Chembiochem 2003, 4, 886.

(37) Busscher, G. F.; van den Broek, S. A. M. W.; Rutjes, F. P. J. T.; van Delft, F. L. Tetrahedron 2007, 63, 3183.

(38) Barluenga, S.; Simonsen, K. B.; Littlefield, E. S.; Ayida, B. K.; Vourloumis, D.; Winters, G. C.; Takahashi, M.; Shandrick, S.; Zhao, Q.; Han, Q.; Hermann, T. Bioorg. Med. Chem. Lett. 2004, 14, 713.

(39) Angell, Y. L.; Burgess, K. Chem Soc Rev 2007, 36, 1674.

(40) Dube, C. E.; Mukhopadhyay, S.; Bonitatebus, P. J.; Staples, R. J.; Armstrong, W. H. Inorganic Chemistry 2005, 44, 5161.

(41) Langlois, N.; Moro, A. Eur J Org Chem 1999, 3483.

(42) Winstein, S.; Grunwald, E.; Buckles, R. E.; Hanson, C. Journal of the American Chemical Society 1948, 70, 816.

(43) Winstein, S.; Hanson, C.; Grunwald, E. Journal of the American Chemical Society 1948, 70, 812.

(44) Lo Conte, M.; Grotto, D.; Chambery, A.; Dondoni, A.; Marra, A. Chemical Communications 2011, 47, 1240.

Chapter 4

Toward the development of fluorescently labeled RNA: Enzymatic incorporation of thGTP, an emissive GTP surrogate

4.1 Introduction

RNA plays a pivotal role in the central dogma, as mRNA, tRNA and rRNA

are responsible for carrying and translating the genetic message to construct

proteins. However, there are other important roles of RNA that include non-

coding RNA in specialized regulatory functions1,2 and a possible target for small

molecules therapeutics.3,4 To investigate the plethora of RNA structure, function,

dynamics, and binding events, fluorescence-based techniques are of key

significance due to their sensitivity and the ability to monitor events in real-time.5-7

A formidable challenge, however, is the modification of the RNA constructs with

fluorescent probes for diverse biophysical applications, either by end- or site-

specific labeling. Although end-labeling oligonucleotides with fluorophores can be

useful, they may not be sensitive to distant binding events or interactions,

whereas internal modifications can be close to the site of investigation.6 A

powerful internal labeling approach involves the replacement of the native bases,

which are practically non-emissive, with fluorescent nucleosides analogs.8-10

Additionally, specifically regarding the subset of isomorphic fluorescent

172

173

nucleosides, these can be especially powerful as they can be non-perturbing and

retain the properties of the natural corresponding nucleosides.6,11

For incorporation of fluorescent RNA nucleosides into oligonucleotides,

the readily utilized techniques are solid phase synthesis and enzymatic incorporation. Many nucleoside analogs can be incorporated via solid phase synthesis, but this depends on the multi-step synthesis of the corresponding

phosphoramidite, and the nucleoside’s ability to withstand the stringent chemistry

of the synthetic cycles.12-14 For enzymatic incorporation, the requirements and

hurdles are distinct, but of the highest significance is the ability of the RNA

polymerase to recognize the corresponding nucleoside triphosphate, incorporate

it with high fidelity and continue elongation on the modified strand.15-21 If this hurdle can be overcome, there are several benefits to enzymatic incorporation.

The triphosphate can be synthesized in one step, avoiding the multistep syntheses frequently necessary for phosphoramidite synthesis. In addition, small amounts of the triphosphate and other reagents can typically be used to generate relatively large quantities of the RNA due to substantial amplification of the DNA template, as much as 250 fold, by the polymerase, although reaction conditions for each template should be optimized.22-24

Thus, in vitro transcription reactions, particularly those mediated by T7

RNA polymerase, have become a cornerstone of modern RNA biochemistry and biophysics. These cell-free transformations facilitate the preparation of short as

well as long native RNA transcripts using synthetic and plasmid-derived

174

templates. While of rather general utility, T7 RNA polymerase requires a specific

promoter for optimal transcription and tends to also be rather sensitive to

sequence composition, particularly next to its consensus promoter. The majority

of T7 phage promoters begin with guanosine (G), including those used typically

for run-off transcription.25-28 Numerous studies have analyzed the initiation and

elongation stages in this processes and, practically, identified optimal sequences

at the transcript’s 5’-end. Regarding the promoter regions (+1 to +6), sequences

typically suffer from diminished transcription efficiency if G residues at position +1

and +2 are altered, but most significantly when +1 changed.22,24 The yields varied

much less when the +3 to +6 base pairs were altered.22,24 To illustrate, an

example from Uhlenbeck and coworkers is shown in Figure 4.1. In comparing

transcription yields to transcript B3, the yield of B9 is reduced by almost half as

the +2 position is changed, and B10 is reduced five-fold as the +1 position is

modified (Figure 4.1). Although, in general, transcription is less stable during initiation phase, resulting in abortive transcripts typically 2–8 nucleotides

long.22,29,30 However, once the polymerase has reached the elongation phase,

typically after incorporating about 10 nucleotides, complete synthesis of RNA

transcript will occur.

175

Figure 4.1: Adapted from Uhlenbeck and coworkers.22,24 a) Comparison of transcript yields resulting from templates B3, B9, B10. b) Effects on transcript yield when +1, +2, or +3 to +6 were modified.

The enzymatic incorporation of fluorescent nucleosides using T7 RNA

polymerase has been tackled in several ways. Although initiation is very

sensitive, especially to alterations at the +1 position, nevertheless, there are

fluorescent analogs used as alternatives to GTP (Figure 4.2). These include a fluorescein-linked AMP either at the 5’-α-phosphate or N6 position (1 and 2)31

coumarin attached to the γ-phosphate of GTP (3)32 Additionally, priming with the non-fluorescent 5’-azido-5’-dexoyguanosine (4) allows for many commercial

fluorophores to be attached post-transcriptionally via azide/alkyne cycloaddition

(click) reactions.33 However, as the elongation phase is more stable, many more

fluorescent alternatives to the natural NTPs have been explored, with most

incorporations remote to the promoter. There has been development of several

176

orthogonal base pairing systems, which require, however, the synthesis of

modified DNA templates in addition to the necessary modified triphosphates.34-36

Additionally, fluorescent isomorphic nucleoside triphosphates with modifications

to the heterocycle have been used in enzymatic incorporation to replace their corresponding natural base. Several fluorescent uracil analogs (5-9) have been

enzymatically incorporated, although some of modifications very close to the

promoter resulted in reduced transcription yields.37-39 A tricyclic CTP analog (10)

acted as a substrate for T7 polymerase in the initiation and elongation phase.40 A

fluorescent guanosine analogue with a modified heterocycle, 8-aza-G

triphosphate (11), was incorporated in position +10, but not during initiation, as a

GpG dimer was used to prime the transcription reaction.41 With these few

examples, it is apparent there are opportunities for discovering new and

improved fluorescent nucleosides to be incorporated in the initiation and

elongation phases.

177

Figure 4.2: Examples of fluorescent nucleosides incorporated by T7 RNA polymerase a) in the +1 position and b) in the initiation or elongation phases.

The recent completion of an emissive RNA alphabet, a fluorescent

ribonucleoside set comprised of highly emissive purine and pyrimidine analogs,

all derived from thieno[3,4-d]pyrimidine,42 presents unique opportunities for the

178 generation of modified RNA constructs. Synthesized as a member of a fluorescent RNA alphabet, thG (13) is highly isomorphic to G (12) (Figure 4.3).42

Recently, thG was shown to be a responsive reporter in directly monitoring different stages of translation.43 So far the incorporation of thG into oligonucleotides has been done only using solid phase synthesis.

Figure 4.3: a) Natural nucleosides and b) Emissive RNA alphabet. The nucleosides are colored to match their corresponding mimic, and the sulfur is in orange.

Despite the high structural resemblance to their native counterparts, it was unclear whether or not enzymes, particularly polymerases, could accommodate and incorporate the fluorescent nucleoside alphabet into oligonucleotides. This has motivated this study, where thGTP (14), has been explored as an emissive and highly isomorphic GTP surrogate in T7 RNA transcription reactions (Scheme

4.1). In this project we have critically assessed the ability of T7 RNA polymerase to initiate and elongate RNA transcription using thGTP (14) in comparison to

179

GTP. We demonstrate that the former is capable of initiating transcription, leading to the formation of fully modified (and emissive) transcripts. Additionally, to assess the impact of substituting G for thG on RNA structure/function, we have transcribed the modified enzyme and substrate components of a hammerhead ribozyme and evaluated the impact of replacing all G residues with thG in the hammerhead enzyme and its substrate. We further demonstrated that the emissive transcripts can be used to monitor the ribozyme-mediated cleavage reaction in real time.

4.2 Results

th th Scheme 4.1: Syntheses of GTP (14) from G (13). Reagents and conditions: i) POCl3, (MeO)3PO, 0 °C; ii) tributylammonium pyrophosphate, Bu3N, 0–4 °C.

The 5’-triphosphate of thG was synthesized by Dr. Dongwon Shin in our lab from the parent nucleoside using freshly distilled POCl3 and tributylammonium phosphate (Scheme 4.1).44 The triphosphate was purified by ion-exchange chromatography and HPLC. With the analytically pure thGTP (14), transcription reactions with T7 polymerase were performed to analyze its enzymatic incorporation into short RNA oligonucleotides. First, a short DNA promoter–template duplex18,37 was used to discern the ability of thGTP to initiate

180

transcription and be incorporated during the elongation phase (Figure 4.4). The

DNA template contained a single T at the 5’ end so a lone A is directed to the 3’

end of the transcript. When α-32P ATP is used, only successfully transcribed full

length labeled RNA products would be visible, whereas short failed transcripts

would be undetected after PAGE (Figure 4.4).

Figure 4.4: Enzymatic incorporation reaction of thGTP using template 15. The T7 promoter DNA was annealed to the DNA template. thG is underlined and bolded blue in transcript 17.

A phosphorimage revealed a full length 10mer product (17) using thGTP that corresponded to the natural triphosphates transcript 16 (Figure 4.5a).

Compared to the natural transcript 16, the yield of transcript 17, containing four

thGs, was 70±3%; therefore, the average individual incorporation 91±1%. Next a large scale transcription reaction was done and UV shadowing was used to visualize all products. Comparing the transcription reactions with GTP to reactions with thGTP illustrate that the desired product and abortive transcripts

appear almost identical (Figure 4.5b). Importantly, when using a UV wavelength

of 302 nm to visualize the gel, the product and initiation phase truncated

181 transcripts are highly fluorescent (Figure 4.5b). The isolated yield of the thG

10mer (17) was 85% of the natural 10mer (16).

Figure 4.5: Transcription reactions with template 15 in the presence of thGTP. a) Small scale transcription using α-32P ATP. Lane 1: control transcription reaction the presence of natural NTPs. Lane 2: control reaction in the absence of GTP and thGTP. Lane 2: reaction in the presence of equimolar concentration of thGTP and GTP. Lane 3: reaction in the presence of thGTP. Incorporation efficiencies thGTPs are reported with respect to transcription in the presence of GTP. All reactions were performed in triplicate and the standard deviations were ±3%. b) Large scale transcription reaction using template 15 with all natural NTPs (lane 1 and 1’) or ATP, UTP, CTP and thGTP (lanes 2 and 2’) with UV light at 254 nm (on TLC plate) and 302 nm (PL). The reaction was resolved by gel electrophoresis on a denaturing 20% polyacrylamide gel.

To test the run-off transcription of longer constructs and test the function of the resulting transcripts, longer DNA templates (18 and 19) were used to generate hammerhead ribozymes: the natural substrate (S), modified substrate

182

(thG-S), natural enzyme (E), and modified enzyme (thG-E) (Figure 4.6). The

transcription experiments were done only on a large scale and the RNA

transcripts (S, thG-S, E and thG-E) were isolated after polyacrylamide gel

electrophoresis (Figure 4.7). Once again, the full length products and short failed

th transcripts are highly emissive for the reactions using GTP.

Figure 4.6: Enzymatic incorporation reaction of thGTP using templates 18 and 19. The T7 promoter DNA was annealed to the DNA templates 18 and 19. thG is underlined and bolded blue in the transcripts thG-S and thG-E.

After transcription, the natural substrate (S) and thG-substrate (thG-S) were

dephosphorylated with alkaline phosphatase and 5’-labeled with T4

polynucleotide kinase according to standard procedures.45 The oligonucleotides

were then tested for ribozyme activity in all different combinations (Figure 4.8),

using conditions similar to those previously published.46,47 The reactions were

done with excess enzyme to obtain psudeo first order kinetic rate constants

46,47 (k2). Single turn-over reactions at 31 °C contained 0.3 µM substrate (including

a trace of 5’-32P labeled material), 3 µM enzyme, 50 mM Tris-HCl pH 7.0, 200

th mM NaCl, and 10 mM MgCl2. The rate constants for S & E and G-S & E were

0.15±0.1 min-1 and 0.12±0.1 min-1, respectively (Figure 8). Additionally, each

183 substrate showed 87% and 86% cleavage for S and thG-S, respectively, with E at

20 min. The thG-enzyme (thG-E) showed minimal, if any, cleavage of S or thG-S

(Figure 4.9).

Figure 4.7: b) Large scale transcription reaction using templates 18 and 19 with all natural NTPs (lane 1 and 3) or ATP, UTP, CTP and thGTP (lane 2 and 4) with UV light at a) 254 nm (on TLC plate) and b) 302 nm (PL). The reaction was resolved by gel electrophoresis on a denaturing 15% polyacrylamide gel.

184

Figure 4.8: Hammerhead ribozymes and cleavage reactions.

185

Figure 4.9: a) Cleavage reactions results determined by radioactive labeling of substrate strands th th S and G-S. S and P1, and G-S and thG-P1 indicate substrate and product strands. All reactions were conducted at 31 °C and contained 0.3 µM substrate (including a trace of 5’-32P labeled material), 3 µM enzyme, 50 mM Tris pH 7.0, 200 mM NaCl, and 10 mM MgCl2. The reactions were quenched at the given times (t in min) and resolved by 20% PAGE with 7M urea. b) Initial th kinetics of S & E and G-S & E. The pseudo first order rate constant (k2), of the cleavage reactions are determined as the slope of ln(fraction cleaved) versus time. c) Ribozyme mediated cleavage curves as determined by radioactive data for S & E and thG-S & E. The reaction was resolved by gel electrophoresis on a denaturing 20% polyacrylamide gel.

The cleavage of thG-S by E was monitored using steady-state fluorescence spectroscopy, and upon mixing the fluorescence intensity increased during the reaction (Figure 4.10a,c). Alternatively, thG-E mixed with S, which showed no cleavage in the radioactive data, displayed minimal fluorescence intensity changes (Figure 4.10b,d). The fraction cleaved of thG-S from the radioactive and fluorescence experiments was normalized and showed a similar

-1 trend (Figure 4.10e). The initial rate from the fluorescence (0.26±0.1 min ) was about two-fold faster than the radioactive data (0.12±0.1 min-1) (Figure 4.10f).

186

Figure 4.10: The fluorescence spectra (ex 370 nm, em 425-485nm) of a) thG-S & E in blue (slits 8,8) and b) S & thG-E in black (slits 4,4), where t = 0 min and t = 20 min spectra have thicker lines. The fluorescence intensity at 450 nm over time of the c) cleavage of thG-S by E (blue) and d) mixing S and thG-E (black). e) The radioactive fraction cleaved (black) and fluorescence (pink) data was normalized for comparison. f) The rate determined from the normalized radioactive data in black compared to the rate derived from the normalized fluorescence data at 450 nm [1- ln(normalized fluorescence)] in pink.

To summarize, thGTP was accepted by T7 polymerase as a GTP surrogate to initiate transcription, as well as incorporated during the elongation

187

phase in short and longer oligonucleotides. To investigate functionality, we tested

a substrate and enzyme per-modified with thG. The thG-S oligonucleotide was

successfully dephosphorylated with alkaline phosphatase and radiolabeled with

T4 polynucleotide kinase. The modified thG-S, when mixed with E, was found to

be active and undergoes efficient phosphodiester bond cleavage. The thG-E has

very little cleavage with S or thG-S, however. The ribozyme cleavage of thG-S by

E, or lack thereof in the case of S and thG-E, was also monitored by steady-state

fluorescence. Additionally, a gel was run on the finished thG-S and E cleavage

reaction to confirm the fluorescent products (Figure 4.11). Furthermore, all

transcripts were digested and analyzed using HPLC-MS to confirm the presence

of thG in the modified oligonucleotides (Figure 4.12).

Figure 4.11: Lane 1 is thG-S, lane 2 is a cleavage reaction of thG-S & E after 20 minutes, and lane 3 is E. The reaction was conducted at 31 °C and contained 0.3 µM thG-S, 3 µM E, 50mMTris pH 7.0, 200 mM NaCl, and 10 mM MgCl2. Photos were taken of the gel with a) UV light at 254 nm (on TLC plate), b) 302 nm (PL), and c) in a BioRad gel imaging system with 302 nm (PL). The reaction was resolved by gel electrophoresis on a denaturing 20% polyacrylamide gel.

188

Figure 4.12: Digestion HPLC traces of a) the mixture of nucleosides used as a standard. This is in both panels to compare to the natural and modified transcripts. Digestion and HPLC traces of b) transcript 17 and 18, c) S and thG-S, and d) E and thG-E.

189

4.3 Discussion

RNA is involved in many important biological processes. The use of

isomorphic fluorescent can be of significant advantage in the study of

oligonucleotide interactions, as they can be non-perturbing of the native

structure. RNA labeled with thG, with its highly isomorphic and emissive

properties, could be used for variety of studies. The one step synthesis of thGTP

and incorporation by T7 RNA polymerase highlights the accessibility to

fluorescent oligonucleotides containing thG.

With the commonly used T7 promoters beginning with C, we could not design a template to test single incorporation. We therefore decided to initially use a short oligonucleotide that has been previously used in our lab. 16,24 The

transcription reaction using template 15 indicated that a full length product was

formed in the presence of thGTP, CTP, ATP, and UTP (Figure 4.4, lane 3), which

suggested that thGTP initiated the transcription. Additionally, the high

incorporation efficiency of thG modification at an average of 91±1%,

demonstrated its structural and functional similarity to G. Due to the sensitive

nature of initiation, it is likely that the thG at +1 and +3 have lower incorporation

efficiency than those at +6 and +8, although this would need to be experimentally

determined. Nevertheless, capitalizing on our positive results, we scaled up the

transcription reaction to confirm that the full length product contained the

modified nucleoside thG (Figure 4.5). The fluorescent abortive transcripts was

another positive indication that thGTP was initiating transcription. The N7 of G

190

was reported to be important in initiation,28,48 however, thGTP was nonetheless recognized very effectively incorporated into the +1 position. Perhaps the N7 in

transcription initiation is not as important as previously thought, although this

would need to be further explored.

Furthermore, thGTP is incorporated fully during the initiation and

elongation phases using longer templates 18 and 19 (Figure 4.7, 4.8). This exhibited the versatile length of transcripts that can be formed using thGTP. The

thG-E transcript contains thirteen thG modifications in which the T7 RNA

polymerase incorporated. Digestion of all transcripts (Figure 4.12) further

confirmed that thGTP was recognized by T7 RNA polymerase, and incorporated

into the initiation and elongation phases. We believe the synthesis of the

modified transcripts validated thG as an isomorphic nucleoside, as it is

recognized enzymatically to pair with cytosine. A benefit to having many thG

modifications is that less material could be needed for detection compared to

singly modified oligonucleotides, although self-quenching could occur.

In addition to thGTP being a substrate for T7 RNA polymerase, we tested

the fully modified oligonucleotides (thG-S and thG-E) ability to function as a

ribozyme. Initially the substrates (S and thG-S) were radioactively labeled to visualize ribozyme cleavage. With this we discovered that the 5’-end thG of thG-S

was recognized by alkaline phosphatase in order for triphosphate cleavage to

occur. Additionally, the 5’-thG also was accepted to be phosphorylated by T4

191

polynucleotide kinase. The versatility of thG to be recognized by several enzymes

is apparent with these results.

The ribozyme cleavage rates of S and thG-S with E are similar, indicating the thG modifications do not interfere greatly with ribozyme catalysis (Figure 4.9).

The ability of thG-S to cleave suggests it forms a duplex with E, further validating

that thG base pairs with C. In contrast, the thG-E showed very little cleavage of S

or thG-S. In the proposed ribozyme cleavage mechanism, the N7 of G10.1

(Figure 4.8) binds a divalent metal ion,49 which seems to be very important for

the catalysis of the cleavage reaction. The absence of the N7 in thG may explain

the lack of cleavage when using thG-E. Although the thG modifications of thG-E

perturbed the ribozyme cleavage, it seemed unproblematic in the thG-S cleavage.

Importantly, the cleavage of thG-S by E can be observed using steady-

state fluorescence spectroscopy, demonstrating the utility of the emissive

transcripts (Figure 4.10). The large increase in fluorescence intensity likely

correlated to the cleavage of thG-S by E because S and thG-E, which showed no

cleavage in the radioactive experiments, exhibited minimal increase in

fluorescence intensity (Figure 4.10a-d). However, it should be noted that the

fluorescence signal from mixing thG-S and E represents an average of

conformational changes, including the annealing and folding of the strands with

Mg2+, as well as strand cleavage and dissociation. The small increase in

fluorescence of S and thG-E could be due to only conformational changes. When

the thG-S and E data from the radioactive and fluorescence experiments are

192

normalized, the overall trend was very similar, which suggests that the

fluorescence experiment can give insight into ribozyme cleavage (Figure 4.10e).

Compared to the radioactive experiments, the derived initial rate of the

fluorescence data of the cleavage of thG-S by E was about twice as fast (Figure

4.10f). This enhanced rate could be due in part to the slight increase in fluorescence that seemed to be occurring in the absence of cleavage. Monitoring

catalytic RNAs using steady-state fluorescence, however, can be an effective

way to probe substrate cleavage and inhibition. It greatly reduced experimental

time compared to the radioactive experiments.

4.4 Conclusion

For the first time, T7 RNA polymerase initiated transcription a fluorescent

isomorphic nucleotide guanosine analog. Thus, short and long transcripts fully

modified with thG were transcribed. Additionally, thG was demonstrated to be a

substrate for other enzymes including alkaline phosphatase and T4

polynucleotide kinase. Although not without limitations, emissive oligonucleotides

containing thG instead of G can be functional and useful in studying oligonucleotide interactions.

4.5 Future Directions

One further application of thG (and possibly thA) is to use them to study of

purine-sensing riboswitches.50 Gene control mediated by riboswitches is especially prevalent in bacterial.50-53 Riboswitches contain an aptamer motif, the

193

RNA region that binds specific ligands, that is primarily found in the 5’-

untranslated region of bacterial mRNA. Once the ligand binds to the aptamer,

typically a conformational change occurs which does not allow translation of the

mRNA to occur, thus regulating gene expression.51-54 Specifically guanine

riboswitches have been looked into as antibacterial drug targets.50,55,56 As binding of thG to the aptamer domain would likely cause a change in fluorescence

intensity, and perhaps wavelength, it could potentially be used in an assay to

study and discover new inhibitors of riboswitches.

4.6 Experimental

Materials

Unmodified DNA oligonucleotides were purchased from Integrated DNA

Technologies, Inc. Oligonucleotides were purified by gel electrophoresis and desalted on Sep-Pak (Waters Corporation). Enzymes were purchased from New

England Biolabs. NTPs and the ribonuclease inhibitor (RiboLock) were obtained from Fermentas Life Science.

Radiolabeled α-32P ATP (10mCi/mL, 3000 Ci/mmol) and γ-32P ATP

(10mCi/mL, 6000 Ci/mmol) was obtained from PerkinElmer. Chemicals for

preparing buffer solutions were purchased from Fisher Biotech (enzyme grade).

Autoclaved 0.1% DEPC treated water was used in all biochemical reactions and

fluorescence titrations.

194

Transcription reactions with α-32P ATP

Single strand DNA templates were annealed to an 18-mer T7 RNA polymerase consensus promoter sequence in TE buffer (10 mM Tris-HCl, 1 mM

EDTA, 100 mM NaCl, pH 7.8) by heating a 1:1 mixture (10 μM) at 90°C for 3 min and cooling the solution slowly to room temperature. Transcription reactions were performed in 40 mM Tris-HCl buffer (pH 7.9) containing 500 nM annealed template, 10 mM MgCl2, 10 mM dithiothreitol (DTT), 10 mM NaCl, 2 mM

spermidine, 1 U/μL RNase inhibitor (RiboLock), 1 mM GTP or 1 mM thGTP, 1 mM

CTP, 1 mM UTP, 20 μM ATP, 2 μCi α-32P ATP and 2.5 U/μL T7 RNA polymerase

(Fermentas) in a total volume of 20 μL. After 3 h at 37°C, reactions were

quenched by adding 10 μL of loading buffer (7 M urea in 10 mM Tris-HCl, 100 mM EDTA, pH 8 and 0.05% bromophenol blue), heated to 75°C for 3 min, and 10

μL was loaded onto an analytical 20% denaturing polyacrylamide gel. The products on the gel were analyzed using a phosphorimager. Transcription efficiencies are reported with respect to transcription in the presence of natural nucleotides . Transcription efficiencies were determined from three independent reactions and the errors are ±3%.

Large scale transcription reactions for Template 15

Large-scale transcription reaction using template 15 was performed in 250

μL reaction volume to isolate RNA and for enzymatic digestion. The reaction

th contained 2 mM ATP, 2 mM CTP, 1 mM GTP, 15 mM MgCl2, 500 nM template,

1500 units T7 RNA polymerase, and 250 units of Ribolock. After incubation for 5

or 6 h at 37°C, the precipitated magnesium pyrophosphate was removed by

195

centrifugation. The reaction was quenched by adding 150 μL of loading buffer.

The mixture was heated at 75°C for 3 min, and loaded onto a preparative 20%

denaturing polyacrylamide gel. The gel was UV shadowed; appropriate bands

were excised, extracted with 0.5 M ammonium acetate and desalted on a Sep-

Pak column. Concentrations of the RNA transcript were determined using

absorption spectroscopy at 260 nm using the following extension coefficients: C,

7200; U, 9900; G; 11500; A, 15400; and thG, 5517 L·mol-1· cm-1.

Large scale transcription reactions for Templates 18 and 19

Large-scale transcription reaction using template 18 and 19 were

performed in 250 μL reaction volume under similar conditions to isolate RNA and

for enzymatic digestion. The reaction contained 2 mM GTP or 2 mM thGTP, 1 mM

CTP, 1 mM UTP, 2 mM ATP, 15 mM MgCl2, 500 nM template, 6 U/μL (1500 U)

T7 RNA polymerase, and 1 U/μL (250 U) of Ribolock. After incubation for 5 or 6 h at 37°C, the precipitated magnesium pyrophosphate was removed by centrifugation. The reaction was speed-vac to reduce half the volume. Then 125

μL of loading buffer was added. The mixture was heated at 75°C for 3 min, and loaded onto a preparative 20% or 15% denaturing polyacrylamide gel. The gel was UV shadowed; appropriate bands were excised, extracted with 0.5 M ammonium acetate and desalted on a Sep-Pak column. Concentrations of the

RNA transcript were determined using absorption spectroscopy at 260 nm using the following extension coefficients: C, 7200; U, 9900; G; 11500; A, 15400; and

thG, 5517 L·mol-1· cm-1.

196

Oligonucleotide Characterization

Digestions: All transcripts (1-2 nmol of 16, 17, S, thG-S, E, thG-E) were

incubated at with S1 nuclease in S1 nuclease reaction buffer (Promega) for two

hours at 37 °C, and then with alkaline phosphatase with dephosphorylation buffer

(Promega) for two hours at 37°C. The ribonucleoside mixture obtained was

analyzed by reverse-phase analytical HPLC by using an Agilent column eclipse

XDB-C18 (5um, 4.6x150 mm. Mobile phase: 0-5% acetonitrile (0.1% formic acid)

in water (0.1% formic acid) over 10 min; flow rate 1 mL/min.

5’ Labeling

S and thG-S (12.9 pmol and 12.3 pmol, respectively) with 3 µL 10x

dephosphorylation buffer, and 1 µL calf intestinal alkaline phosphatase in a total

volume of 30 µL was heated to 37 °C for 2 hrs. 70 µL of water was added and

then extracted with 100 µL phenol:chloroform (CHCl3):isoamyl alcohol (iAA)

25:24:1. The water was extracted with 100 µL CHCl3. The RNA was precipitated

with 6 µL glycoblue™ (Life Technologies), 20 µL 10 M NH4OAc, and 400 µL

EtOH and put on dry ice for 1 hour. Then the eppendorfs were centrifuged at

14,000 rpm for 20 min and the supernatant was removed. The pellet was washed

with 4x with 50 µL of cold 70% EtOH. The pellet was air dried for 30 min and then

dissolved in 38 µL water. 5 µL of 10x kinase buffer, 1 µL dithiothreitol, 5 µL of γ-

32P ATP, and 1 µL of T4 polynucleotide kinase was added and the reaction was heated to 37 °C for 2 hours. The RNA was then precipitated (2 µL glycoblue™,

10 µL 10 M NH4OAc, and 200 µL EtOH) and washed (x1 with 25 µL cold 70%

EtOH) similar to the above procedure. The pellet was dissolved on 1x TBE 7 M

197

urea loading buffer and then the RNA was resolved by gel electrophoresis on a

denaturing 20% polyacrylamide gel. The RNA was cut out and extracted with

water overnight.

Ribozyme reaction conditions

Cleavage reactions were conducted in a total reaction volume of 34 µL for

the natural enzyme and 22 µL for the thG-enzyme, with the substrate or thG- substrate for the radioactive experiments. For the fluorescent experiments, a total volume of 125 µL total was used. The reactions were carried out at 31 °C in a

buffer containing 50mM Tris-HCl (pH 7.0) and NaCl (200 mM). Buffered solutions of the substrate (0.6 µM) and enzyme (6 µM) were denatured separately by heating to 90 °C for 90 s and cooled to room temperature over 10 min to allow for refolding. MgCl2 (10mM) was added to both the enzyme and substrate and were

equilibrated at 31 °C for 10 min. The cleavage reaction was then initiated by

manually mixing equal volumes of the modified or natural substrate (0.6 µM) with

the natural or modified enzyme (6 µM) in a heat block or fluorimeter at 31 °C, to

give final concentrations of 0.3 µM of the substrate and 3 µM of the enzyme at

the total volumes.

Radioactive ribozyme assay

Experiments with radioactively-labeled substrates contained a trace of 5’-

32P labeled substrate. For initial data points (time=0), 2 µL of the substrate were

removed immediately prior to starting the reaction. Following initiation of the

reaction, 4 µL aliquots were removed at designated time periods and quenched

with 12 µL of urea containing loading buffer (7M urea, 1x TBE, and 0.05%

198

bromophenol blue and xylene cyanol). The tubes were heated to 90 °C for 90 s

and 15 µL was loaded on a 20% polyacrylamide with 7M urea gel. Gels were quantified on a Personal Molecular ImagerTM and analyzed with Quantity One software (Biorad).

Radioactive ribozyme cleavage data analysis

When cleavage is conducted under single-turnover conditions, the pseudo

first order rate constant k2 can be extracted from measurements of the ribozyme’s initial rate of cleavage.57 Since the cleavage reaction occurs within

the hammerhead complex, it can be treated as a first order intramolecular

reaction. Rate constants (k2) were calculated as the slope of ln(1-S/S0) versus

time, where S/S0 is the fraction of cleaved substrate. For experiments utilizing a

radioactively labeled substrate, S/S0 was determined by dividing the amount of

cleaved substrate by the sum of the full length and cleaved substrate. Error bars

represent the standard deviation of the numbers averaged for each data point.

Fluorescence ribozyme assay

The reactions were done in a 125 µL microfluorescence cell with a path

length of 1.0 cm (Hellma GmH & Co KG, Mullenheim, Germany) on a Jobin Yvon

Horiba FluoroMax-3 luminescence spectrometer. A background spectrum was

subtracted from each sample. All samples were excited at 370 nm, and the

emission was recorded from 425-485 nm. For samples containing thG-S and thG-

E, slits of 8 and 4 nm were used, respectively.

Following mixing of the ribozyme substrate and enzyme, fluorescence

spectra was taken immediately. Then spectra were taken at designated time

199 periods. Error bars represent the standard deviation of the numbers averaged for each data point. For comparison, the radioactive fraction cleaved and fluorescence data was normalized from 0 to 1 (Figure 4.10e). ate derived from the normalized fluorescence data at 450 nm [1-ln(normalized fluorescence).

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Chapter 5

Enzymatic transformation of a fluorescent adenosine analogue into an inosine analogue: Development of a high-throughput assay

5.1 Introduction

Emissive isomorphic nucleoside analogs, in conjunction with versatile and

sensitive fluorescence spectroscopy techniques,1,2 have shown to be of great

value in the study of nucleic acid structure and their interactions with diverse

ligands.3-12 While of great value within the context of oligomeric structures, much

less is known about the function of such emissive analogs in the “protein world”,

where nucleosides and nucleotides intricately interact with enzymes.13-16

In contrast to the use of fluorogenic enzyme substrates and fluorophore

precursors17-23 as well as the enzymatic unmasking or uncaging of established

fluorophores for biochemical assays or imaging applications,24-28 no examples

exist where isomorphic fluorescent nucleosides are transformed in enzymatically

catalyzed reactions to form new and distinct fluorophores. This is likely due to the

substrate specificity of the enzymes responsible for metabolizing and utilizing

such key nucleoside and nucleotide cellular components.29-35

A catabolically important deamination reaction, whereby adenosine (A) is

converted to inosine (I), is catalyzed by adenosine deamin ase (ADA) (Figure

203

204

5.1a).36-38 We speculated that thA (1), a member of the fluorescent RNA alphabet recently synthesized in our lab by Dr. Dongwon Shin,39 could be a candidate for monitoring the ADA catalyzed deamination reaction (Figure 5.1b).36-38 The underlying hypothesis is that due to its similarity to adenosine, its counterpart, thA will be transformed to thI by ADA (Figure 5.1b). As thA is emissive, thI is likely to be fluorescent as well, and their electronic differences are expected to render the two chromophores distinct. This, in principle, should allow one to monitor the progression of the deamination reaction in real time using fluorescence spectroscopy, an impossible task with the natural nucleobases.

Figure 5.1: ADA catalyzed interconversion of a) A to I and b) thA (1) to thI (2).

Monitoring the ADA catalyzed deamination reaction with fluorescence could be of great benefit, as ADA is critical for regulation of intra- and extracellular biosynthesis, transport, and metabolism of adenosine.40 ADA, a cytosolic and ecto-enzyme, is distributed throughout human tissue, and the highest activity

205

of ADA is found in the lymph system.41 Increased adenosine concentrations

result from many immune responses, and chronic high concentrations can lead

to tissue damage or possible sepsis.42-45 ADA down regulates extracellular

adenosine levels and influences adenosine receptor stimulation, thus controlling

many inflammatory responses. Thus the regulation of ADA could be a potential

treatment for pathologies including inflammation. Additionally, ADA inhibition has

been shown to be effective against cancers affecting the immune system.41,46

If the deamination of thA by ADA is successful, this can provide a new

method for exploring and identifying inhibitors of ADA, small molecules of clinical

utility as chemotherapeutic agents.41,47,48 In this chapter we demonstrate that

ADA converts thA to thI, and this reaction can be monitored by steady-state

emission spectroscopy. The utility of this sensitive fluorescently-monitored transformation is highlighted with the development of a high throughput assay for real-time detection of ADA inhibitors.

5.2 Results and Discussion

To be able to analytically and photophysically verify thI as the product of

the enzymatic deamination of thA, thI was independently synthesized by Dr.

Dongwon Shin in our laboratory.39 X-ray crystallography unequivocally shows their correct anomeric configuration. Overlaying thA (1) and thI (2) crystal structures with the reported structures of their natural counterparts A49 and I,50

respectively, illustrates their truly isomorphic nature (Figure 5.2).51 Importantly,

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thA (1) adopts an anti-conformation having N-ribose (3'-endo) puckering, conformational features known to be preferred by ADA.52

Figure 5.2: Crystal structures of a) A, thA (1), and their nucleobase overlay (RMS: 0.0383), and b) I, thI (2), and their nucleobase overlay (RMS: 0.0330).51

To ensure that the photophysical characteristics of thA (1) and thI (2) are

distinguishable, Dr. Renatus Sinkeldam in our laboratory examined binary

mixtures containing different ratios of the two nucleosides by absorption and

fluorescence spectroscopy in phosphate buffer at pH 7.4, conditions commonly

used for enzymatic deamination reactions (Figure 5.3a). The overlaid absorption

spectra of the mixtures showed a distinct hypsochromic shift of the absorption

maximum from that of thA (1) at 339 nm to that of thI (2) at 315 nm with a

concomitant reduction in the optical density at the lower energy transition >300

nm (Table 5.1). Upon excitation at 318 nm, their isosbestic point, the recorded

emission spectra also presented a blue shift from 410 nm, the emission

207 maximum of thA, to 391 nm, the emission maximum of thI. In contrast to the lower optical density, the increasing thI concentration resulted in an increase of the emission intensity, illustrating the higher fluorescent quantum yield for thI (2) compared to that of thA (1).

Figure 5.3: a) Absorption (dashed lines) and emission (solid lines) spectra of samples prepared with different ratios thA (1) and thI (2) of 11µM in phosphate buffers of pH 7.4; b) correlation between inosine fraction and λabs.max (solid blue triangles) and O.D. (open orange triangles) at 340 nm; c) correlation between λem.max. (solid blue circles) and PLint. at 391 nm (open orange circles); and d) enzymatic deamination of thA (1) to thI (2) with ADA monitored in real-time by absorption at 339 nm (blue) and emission at 391 nm (orange; excitation at 318 nm). Reaction conditions: [thA] =11.7 µM, [ADA] = 27 mU/mL in 50 mM phosphate buffer of pH 7.4 kept at 25 °C for 1h. The normalized kinetic curves yield a t1/2 of 11.5 min.

Table 5.1: Relevant spectroscopic properties of thA (1) and thI (2). ] λ and ε are reported in nm and M–1cm–1, respectively.

rel. Absint. rel. PLint. λabs.max. ε λem.max. @ 339 nm @ 391 nm

thA (1) 339 7.1 × 103 3.9 410 1.0 thI (2) 315 4.8 × 103 1.0 391 3.3

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To gain insight into the spectral changes, two correlation plots have been

constructed (Figures 5.3b and 5.3c). Both the shift in absorption maximum and

the drop in optical density showed a nearly linear dependence on the thA/thI ratio

(Figure 3b). Interestingly, both the emission wavelength and intensity responded

linearly to an increasing thI concentration in binary mixtures with thA (Figure 5.3c).

Clearly, absorption as well as emission spectroscopy revealed significant

spectral changes suitable to study the enzymatic conversion of thA (1) to thI (2).

To investigate thA (1) as a substrate surrogate for adenosine in ADA- mediated deamination reactions, absorption and emission were measured by Dr.

Sinkeldam before and after its reaction with ADA.51 The spectra obtained after

enzymatic treatment perfectly resemble the spectral characteristics of thI (2). 51 A

control experiment using a fresh solution of thI (2) mixed with ADA, established

that the presence of the protein does not immediately affect the spectral

properties of the emissive nucleoside.51 Next, the enzymatic conversion of thA

(1) to thI (2) was followed in real-time using absorption as well as fluorescence

th spectroscopy. Absorption changes were followed at 339 nm, the λabs.max of A,

th and emission changes were followed at 391 nm, the λem.max of I, upon excitation

at 318 nm, their isosbestic point (Figure 5.3d). As expected, the absorption

spectra show a decrease (or an “OFF” signal) in the optical density upon

conversion of thA (1) to thI (2). By exploiting thI’s higher fluorescence intensity compared to thA to follow the enzymatic conversion, intensification of the

emission, or “ON” signal, is obtained. Absorption and emission spectra taken in

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the absence of ADA indicated that there is no conversion without the enzyme

under the experimental conditions used.51

To substantiate the observations listed above, LC-MS analysis was performed on the deamination reaction of thA (1) with ADA under the same

conditions as the fluorescence studies (Figure 5.4). A nucleoside standard

containing thA (1) and thI (2) was utilized to corroborate the identity of the starting

material and product. Taking into consideration the different absorption maxima,

the chromatograms were followed at 250, 260, 315 and 340 nm. Initially the thA

(1) reaction mixture was run prior to the addition of ADA, showing a single peak.

After ADA addition, aliquots were taken from the reaction and filtered to remove the enzyme. Since the reaction was run under the same conditions of the fluorescence studies, the reaction was fairly dilute. Thus, large samples (500 µL) were taken, and concentrated by lyophilization, to visualize the nucleoside absorbances on the HPLC chromatogram. As shown in Figure 5.4, over time the

thA peak (blue) decreases as the thI peak increases (pink) for all wavelengths.

Importantly, mass spectrometry confirmed the identity of thA and thI at the

appropriate retention times for the standard and each time aliquot. In agreement

with the fluorescence studies, thA (1) is quantitatively converted to thI (2) in about

one hour. However, using the HPLC-MS to monitor the reaction was quite

cumbersome, as it took several hours as each sample had to be filtered,

lyophilized and then run on HPLC-MS. Thus, using fluorescence is a much more

210 efficient and sensitive technique, but the confirmation of thI formation from thA plus ADA was essential.

Figure 5.4: HPLC-MS chromatograms showing the enzymatic conversion of thA (1) to thI (2) monitored at 250 nm, b) 260 nm, c) 315 nm, and d) 340 nm (black) against a reference mixture of authentic nucleosides (grey). Each set shows four time points at t= 0, 13, 35, and 65 min following the addition of ADA. The HPLC absorbance peak of thA (blue) and thI (pink) corresponded to the ESI-MS spectra shown in e) [thA+H]+ calculated 284.06, found 284.09 and f) [thI+H]+ calculated 285.05, found 284.97, respectively.

211

The enzyme kinetic parameters of the deamination reactions were

determined for both thA and A by Dr. Renatus Sinkeldam.51 According the Henri-

th Michaelis-Menten kinetics, Km values of 417 and 29 µM are obtained for the A

to thI, and A to I conversion, respectively. The lower conversion rate of thA,

compared to that of adenosine, appeared to be due to the lower affinity of the

th former to ADA. We speculate that the higher Km values observed for A are likely due to the replacement of N7 in adenosine with a CH group in thA, as previous

structural analysis has shown contacts between an aspartic acid side chain

37-40,53-55 residue (D296) and this position of the substrate (Figure 5.5). Nontheless,

thA still acted as a substrate surrogate for ADA-mediated deamination to thI. This can be rationalized as the isomorphic hydrogen bonding face of thA is similar to A

and can react in the same fashion (Figure 5). The mechanism of ADA

deamination begins with the attack of an OH group coordinated to a zinc ion, and

36-38,41,53-55 protonation of the N2 via the glutamic acid residue (E217).

Subsequently, E217 then deprotonates the OH, driving carbonyl formation and

releases of ammonia (Figure 5). With ADA successfully converting thA to thI, this

sets the stage for a potential new method for identifying new inhibitors of ADA.

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Figure 5.5. a) Predicted mechanism of deamination of A by ADA. b) Ability of thA to undergo a similar transition state as A.

We noted that current methods for identifying inhibitors typically rely on

absorption spectroscopy.44,59,60 However, this could be problematic as adenosine

and often the inhibitor molecules show absorbance in similar wavelengths around

260 nm. Thus, the inherent emission of thA, and enhanced and distinct emission upon deamination by ADA to thI, provided a robust foundation for a high throughput assay for inhibitor discovery. To illustrate potential screening for novel

ADA inhibitors, we developed a 96-well plate based assay, exploiting the rapid

and sensitive fluorescence monitoring of the deamination reaction. The emission

enhancement, associated with the conversion of thA (1) to thI (2), was monitored at 391 nm (excitation at 318 nm) over a 60-minute time window with increasing concentrations of EHNA and pentostatin, potent inhibitors of ADA (Figure 5.6a).

As shown in Figures 5.6b and 5.6c, the inhibition of ADA is readily apparent even at low nM concentrations. The data obtained can be quantified by plotting the percent inhibition at 60 minutes against log[inhibitor] and applying a sigmoidal fit.

213

Figure 5.6: a) Structures of ADA inhibitors EHNA and Pentostatin; conversion of thA (1) followed with fluorescence in the presence of b) EHNA and c) Pentostatin (black, purple, blue, cyan, green, orange, and red lines represent 0, 0.1, 1, 2.5, 10, 25, 100 nM [inhibitor], respectively. The grey lines represent the conversion of thA (1) in the absence of ADA. The experiment is performed in triplicate and error bars reflect the standard deviation. d) A plot of % inhibition at 60 min. vs. log[inhibitor] in triplicate (data points) and sigmoidal logistic fits performed in OriginPro (lines) for EHNA (open circles, blue line) and Pentostatin (solid circles, orange line). The grey dashed lines visualize graphical determination of the IC50 values. Actual values have been interpolated using the fit. Assay conditions: [thA] = 11.7 µM, [ADA] = 27 mU/mL, in 50 mM phosphate buffer of pH 7.4 at 21 °C.

This yielded IC50 values of 13.4 ± 1.3, and 1.9 ± 0.1 nM for EHNA and pentostatin, respectively, illustrating the established higher potency of the latter.

Importantly, two negative controls were examined. Guanosine, as expected, had no impact on the deamination reaction up to 100 nM (Figure 5.7a). Additionally, thI mixed with ADA resulted in the same fluorescence intensity of thA plus ADA

214

over the length of the experiment (Figure 5.7b). This effect can likely be attributed to a mild binding of thI to ADA

Figure 5.7: a) ADA conversion of thA (1) to thI (2) followed over 60 min. in the absence (black line) and presence of 0.1-100 nM G (colored lines). For completeness, the conversion of thA in the absence of ADA (grey line) is added. b) The emission of thI (2) followed over 60 min. in the absence (green line) and presence (blue line) of ADA. For comparison, the emission of thA in the absence (grey line) and presence (black line) of ADA are added.

5.3 Conclusions

To summarize, an isomorphic emissive adenosine analogue, thA (1),

serves as a viable substrate for ADA, a nucleoside-modifying enzyme.56-59 The

enzymatic deamination process yields the corresponding emissive inosine

analogue thI (2) confirmed by LC-MS, which possesses distinct spectral features,

allowing one to monitor the enzyme-catalyzed reaction and its inhibition in real time. To demonstrate its practical utility, we formulated a high throughput assay for the discovery and biophysical evaluation of ADA inhibitors, key agents for researchers and clinicians, as the regulation of adenosine is linked to inflammatory diseases and certain cancers. This unique proof-of-principle process, where the nucleobase core of a fluorescent nucleoside analogue is

215

enzymatically transformed into a distinctly emissive product, demonstrates a new

facet for isomorphic nucleoside analogues and expands their utility landscape

beyond their “natural” and typically explored oligonucleotide environments.

5.4 Future Directions

With thA being deaminated by ADA, there is potential for other enzymes to

recognize and perhaps the others of the isomorphic fluorescent alphabet.39

Cyclic guanine monophosphate and cyclic adenosine monophosphate (cGMP and cAMP, respectively) are important cellular secondary messengers. Cyclic nucleotide phosphodiesterases (PDEs) are responsible for the degradation of cGMP and cAMP to their 5’-GMP and 5’-AMP, and have been examined as drug targets.60 The most well-known example is inhibitors for PDE5 marketed for the

treatment of erectile dysfunction and pulmonary artery hypertension,61 and other

therapeutic potentials include treatment for cognitive disorders62 and

vasoconstriction, among others.63 If cthGMP or cthAMP were synthesized, they

could, in principle, be used for monitoring the inhibition of these PDEs. However,

spectral changes from the cyclic monophosphates to the 5’-monophosphates

may not occur. Also interestingly, there is a tryptophan near the active site of

63 many of the PDEs, which could be a potential FRET pair as the λem.max around

th th 350 nm could overlap with the absorbance of A or G (λabsmax and 340 and 320

nm, respectively).

216

5.5 Experimental Section

Spectroscopy

Bovine Intestine ADA was obtained from CalBioChem. The commercial solution (2600 U/ml in 3.2 M (NH4)2SO4 buffer at pH 6.0) was diluted to 4.0 U/mL by dissolving 2.77 µL in 1.8 mL buffer. The enzyme dilution was freshly prepared and kept on ice prior to use. Concentrated 11.3 mM stock solutions of thA (1) and thI (2) were prepared in DMSO. Final spectroscopy samples were prepared in a

4-sided quartz cuvette to arrive at a nucleoside and enzyme concentration of

11.7 µM and 27.5 mU/mL, respectively. All samples were measured in a 50 mM phosphate buffer set to pH 7.4 and measured at 25°C.

Absorption spectroscopy measurements have been performed on a

Shimadzu UV 2450. Real time conversion of thA (1) to thI (2) is followed by monitoring the absorption at 339 nm every 2 seconds for 3600 seconds after addition of ADA.

Fluorescence spectroscopy measurements have been performed on a PTI fluorimeter. Fluorescence was recorded upon excitation at 318 nm (isobestic point of thA−thI mixtures) using slit−widths of 1.6 nm. Excitation spectra were measured by monitoring at 391 nm (emission maximum of thI) using slit−widths of

1.6 nm. Real time conversion of thA (1) to thI (2) was followed continuously every

2 seconds for 3600 seconds immediately after addition of ADA by monitoring the emission at 391 nm (excitation at 318 nm).

217

LC-MS analysis of enzymatic interconversion of thA into thI

HPLC-MS analysis was carried out with an Agilent 1260 system coupled

with a Thermo LCQ deca mass spectrometer using positive ion mode

electrospray ionization as the ion source. Wavelengths recorded were 250nm,

260nm, 315nm, and 340nm. The column used for the HPLC was an Eclipse

XDB-C18 5 µm analytical 4.6x100 mm column. ADA was obtained from

Worthington (calf spleen) as a freeze-dried powder (12.5 mg) with an assayed

activity of 20.2 U/mg. A stable stock solution was prepared and stored as

described under S4.1. A fresh enzyme work solution was prepared daily by

dissolving enzyme stock solution (1 μL) in 50 mM pH 7.4 phosphate buffer (0.099

mL) to arrive at an ADA concentration of 0.1033 mg/mL (2.08 U/mL assumed

activity). This solution was kept on ice prior to use and left over solution was

discarded at the end of the day. Final substrate sample solutions containing thA

(1) were each prepared from a DMSO stock solution of 11.3 mM. ADA was

obtained from Worthington (calf spleen) as a freeze-dried powder (12.5 mg) with

an assayed activity of 20.2 U/mg. A stable stock solution was prepared and

stored as described under S4.1. A fresh enzyme work solution was prepared

daily by dissolving enzyme stock solution (1 μL) in 50 mM pH 7.4 phosphate

buffer (0.099 mL) to arrive at an ADA concentration of 0.1033 mg/mL (2.08 U/mL assumed activity). The samples were filtered using a Millipore amicon ultra,

0.5mL, with a 30 kDa cut-off to remove the ADA enzyme.

218

An 11.7 µM solution of thA was made in 50 mM phosphate buffer, pH 7.4

for a total of 1974 µL. For t=0 min, an aliquot of 493.5 µL was removed. To the

rest of the thA solution (1480.6 µL), 19.4 µL of the 2.08 U/mL solution of enzyme

was added for a final concentration of 27 mU/mL. At 8, 30, and 60 min, a 500 µL

aliquot was removed and then filtered on the spin column for 5 min at 14,000 rcf

for total time points of 13, 35, and 65 min. The flow-through was frozen and

subsequently lyophilized to dryness. Each sample was dissolved in 30 µL of

water of which 20 µL was injected into the HPLC-MS. A run of 8 minutes using a flow of 1.0 mL/min, with 5% acetonitrile (0.1% formic acid) in 0.1% formic acid aq. as the mobile phase, completely separated thA (1) from thI (2) (Fig. S3.1).

Fluorescence-based High-throughput inhibition assay

Pentostatin and EHNA HCl were purchased from R&D Systems and EMD

Millipore, respectively. Both ADA inhibitors were used without further purification.

ADA was obtained from Worthington (calf spleen) as a freeze-dried powder (12.5 mg) with an assayed activity of 20.2 U/mg. A stable stock solution was prepared and stored as described under S4.1. A fresh enzyme work solution was prepared daily by dissolving enzyme stock solution (1 μL) in 50 mM pH 7.4 phosphate buffer (0.099 mL) to arrive at an ADA concentration of 0.1033 mg/mL

(2.08 U/mL assumed activity). This solution was kept on ice prior to use and left over solution was discarded at the end of the day. Final substrate sample solutions containing thA (1) and thI (2) were each prepared from a DMSO stock

solution of 11.3 mM. For the Pentostatin experiments stock solutions of 10 nM,

219

100 nM, 250 nM, and 1 µM Pentostatin in water were used. For the EHNA HCl

experiments stock solutions of 100 nM, 1 µM, 2.5 µM, and 10 µM EHNA HCl in

water were used. All experiments were performed in 50 mM phosphate buffer of

pH 7.4 at 21 °C. For the high-throughput fluorescence assay a 96-well plate was

used in conjunction with a Molecular Devices SPECTRAmax® GEMINI XS plate

reader.

All experiments were performed using a 96-well plate. Each reaction

(well) contained 97.7 µL of an 11.7 µM solution of thA or thI in 50mM phosphate

buffer pH 7.4. When applicable, 1 µL of the inhibitor stock solutions were added

to the reaction. Using a 318 nm excitation the initial thA emission, in the absence

of ADA, was recorded. Using a multichannel pipettor, 1.3 µL of 2.08 U/mL of

enzyme was added to the appropriate wells to give a total of 27 mU/mL per

reaction. Immediately the plate was inserted into the plate reader and automixed

for 10 seconds. The change in emission at 391 nm was recorded for each well

every 30 seconds for 60 minutes. All experiments were done in triplicate;

averages and standard deviations were calculated in MS-Excel. Subsequently,

the data was normalized to the thA fluorescence without the enzyme set as “0”, and the final fluorescence of thA + ADA set as “1”.

Acknowledgements

Chapter 5 is a partial reprint from: Sinkeldam, R. W.; McCoy, L. S.; Shin,

D.; Tor, Y. Enzymatic Interconversion of Isomorphic Fluorescent Nucleosides:

Adenosine Deaminase Transforms an Adenosine Analogue into an Inosine

220

Analogue. Angewandte Chemie International Edition 2013, 52, 14026. The dissertation author is the second author and researcher of this work.

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