A Dissertation Entitled Ribonucleic Acids in Disease Etiology and Drug Discovery by Immaculate Sappy Submitted to the Graduate F

Home , Pseudouridine

A Dissertation

entitled

Ribonucleic Acids in Disease Etiology and Drug Discovery

Immaculate Sappy

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the

Doctor of Philosophy Degree in

Medicinal Chemistry

______Amanda C. Bryant-Friedrich, Ph.D., Committee Chair

______Zahoor A Shah, Ph.D., Committee Member

______Steven M Peseckis, Ph.D., Committee Member

______Caren Steinmiller, Ph.D., Committee Member

______Amanda Bryant-Friedrich, PhD, Dean College of Graduate Studies

The University of Toledo

December 2019

This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author.

An Abstract of

Ribonucleic Acids in Disease Etiology and Drug Discovery

Immaculate Sappy

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the Doctor of Philosophy Degree in Medicinal Chemistry

The University of Toledo

December 2019

Pseudouridine (Ψ), the 5-ribosyl isomer of uridine (U) is the most abundant nucleic acid modification found in all domains of life and all types of RNA. Studies have shown that, urinary levels of pseudouridine are higher in Alzheimer’s Disease (AD) patients and that

RNA oxidation is a major component in the pathogenesis of Alzheimer’s Disease (AD) and other neurodegenerative disorders. Therefore, there is a potential correlation between higher urinary levels of pseudouridine in AD patients and oxidative stress. Hence, subjecting pseudouridine to oxidative conditions may provide some key information about the role of this nucleoside in RNA related processes and its role in disease etiology.

Besides neurodegenerative disorders, antibiotic resistance is an additional threat to human health. Analogous to the development and use of nucleoside-analogue inhibitors (NAIs) of viral nucleotide polymerases for treatment of viruses, nucleotide analog inhibitors of bacterial RNA polymerase such as pseudouridimycin are also being investigated. These

NAIs can limit bacterial resistance by mimicking the RNAP nucleoside triphosphate (NTP) binding site.

iii

To investigate these possibilities, the design, synthesis and characterization of pseudouridine analogs will be performed and these nucleosides will be evaluated for their antibacterial properties and their oxidative fate in RNA.

I dedicate my dissertation to the spirit of my beloved dad. I also, dedicate this work to my family and most importantly to my loving and adored children for their patience and tolerance. A special dedication to my best friend turned husband for always being supportive, enduring, encouraging, loving and caring.

Acknowledgements

First, I am grateful to God Almighty for establishing me to complete this program. My sincere gratitude to Dr. Amanda C. Bryant-Friedrich, who believed in me when I did not believe in myself, invested in me and allowed me to enjoy working in her research group and for her valuable scientific and personal experience. Your continued guidance helped me to grow as an individual and also as an outstanding chemist. Thank you for your unceasing mentorship! You created so many opportunities for me and designed tools for me to grow not only as a woman but a mother and a scientist. I will be forever grateful.

To my committee members Dr. Peseckis, Dr. Shah and Dr. Steinmiller thank you for your support and involvement.

To Dr. Bedi, I am truly thankful for all your help and valuable advice. Thank you for walking with me spiritually during this Journey.

To my lab members past and present, thank you for making the Amanda Bryant-Friedrich

Lab home and maintaining the sense of family.

To my Husband, thank you for being so patient with me. May God bless you.

To my kids, not all heroes wear capes. Thank you for all your encouragement, reassurances and support. May God Almighty richly bless and reward you.

To my family and friends, may God bless you for all your prayers and support.

Table of Contents

ACKNOWLEDGEMENTS ...... V LIST OF TABLES ...... IX LIST OF FIGURES ...... X LIST OF SCHEMES ...... XIII LIST OF ABBREVIATIONS ...... XIV LIST OF SYMBOLS ...... XVI 1 INTRODUCTION ...... 1 1.1 OVERVIEW ...... 1 1.2 RNA DAMAGE IN DISEASE ETIOLOGY ...... 3 1.3 RIBONUCLEOSIDES IN DRUG DISCOVERY ...... 4 2 BACKGROUND ...... 5 2.1 ROLE OF PSEUDOURIDINE IN DISEASE ETIOLOGY ...... 5 2.2 RNA OCCURRENCE IN LIVING ORGANISMS ...... 6 2.2.1 Bacteria ...... 6 2.2.2 Viruses ...... 7 2.2.3 Mammalian cells ...... 8 2.3 FUNCTIONS ...... 8 2.4 MODIFIED RIBONUCLEIC ACIDS ...... 9 2.4.1 Pseudouridine Occurrence and Disease Relevance ...... 12 2.5. RNA OXIDATION IN HUMAN CELLS ...... 14 2.5.1 Oxidative Stress ...... 14 2.5.2 Mechanisms of Oxidative Damage to RNA ...... 17 2.5.3 Role of RNA Oxidation in Disease Processes ...... 19 3 SYNTHESIS OF A C5´-PSEUDOURIDINYLRADICAL PRECURSOR ...... 22 3.1 SYNTHESIS OF MODIFIED RIBONUCLEOSIDES AS C5´-RADICAL PRECURSORS ...... 22 3.3 PHOTOCHEMICAL GENERATION AND STUDY OF THE C5´-PSEUDOURIDINYL RADICAL...... 47 3.4 CONCLUSION AND FUTURE DIRECTION ...... 48 4 EXPERIMENTAL PROCEDURE SYNTHESIS OF C5´-RADICAL PRECURSOR 13 ...... 50 4.1 METHODS AND MATERIALS ...... 50 Method ...... 50 Material ...... 50 4.2 SYNTHESIS OF PSEUDOURIDINE ...... 51 4.2.1 Synthesis of (3aR,6R,6aR)-6-(((tert-butyldiphenylsilyl) oxy)methyl)-2,2- dimethyldihydrofuro[3,4-d][1,3]dioxol-4(3aH)-on (62) ...... 51 vi

4.2.2 Synthesis of (3aR,6R,6aR)-6-(((tert-butyldiphenylsilyl)oxy)methyl)-4-(2,4-di- tert-butoxypyrimidin-5-yl)- 2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-ol (80) 34 ...... 53 4.2.3 Synthesis of 2-((tert-butyldiphenylsilyl)oxy)-1-((4S,5R)-5-((R)-(2,4-di-tert- butoxypyrimidin-5-yl)(hydroxy)methyl)-2,2-dimethyl-1,3-dioxolan-4-yl)ethan-1-ol (81) 34 ...... 54 4.2.4 Synthesis of 2,4-di-tert-butoxy-5-((3aS,4S,6R,6aR)-6-(((tert- butyldiphenylsilyl)oxy)methyl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4- yl)pyrimidine (82) 34 ...... 55 4.3 SYNTHESIS OF C5´- PSEUDOURIDINYLRADICAL PRECURSOR ...... 56 4.3.1 Synthesis of ((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)methanol (60) 34 ...... 56 4.3.2 Synthesis of (3aS,4S,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxole-4-carbaldehyde (86) ...... 57 4.3.3 Synthesis of 2-((tert-butyldimethylsilyl)oxy)-2-((3aR,4S,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4- yl)acetonitrile (87) ...... 58 4.3.4 (R)-1-((tert-butyldimethylsilyl)oxy)-1-((3aR,4S,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)-3,3- dimethylbutan-2-imine (88) ...... 59 4.3.5 (S)-1-((tert-butyldimethylsilyl)oxy)-1-((3aR,4S,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)-3,3- dimethylbutan-2-one (89) ...... 59 4.3.6 (S)-1-((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)-1-hydroxy-3,3-dimethylbutan-2-one (90) ...... 60 4.3.7 5-((2S,3R,4S,5S)-3,4-dihydroxy-5-((S)-1-hydroxy-3,3-dimethyl-2- oxobutyl)tetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione (13) ...... 61 4.4 PHOTOCHEMICAL GENERATION OF THE C5´- PSEUDOURIDINYLRADICAL ...... 62 5 PSEUDOURIDIMYCIN AND ITS ANALOGS ...... 76 5.1 DISCOVERY OF PSEUDOURIDIMYCIN ...... 77 5.2 DERIVATIVES OF PSEUDOURIDIMYCIN ...... 85 5.3 NOVEL SYNTHESIS OF PSEUDOURIDIMYCIN ANALOG ...... 91 5.4 CONCLUSION AND FUTURE DIRECTION ...... 99 6 EXPERIMENTAL PROCEDURE: PSEUDOURIDIMYCIN AND ITS ANALOGS ...... 101 6.1 MATERIALS ...... 101 6.2 ANALYSIS METHOD ...... 101 6.3 SYNTHESIS OF 5´-AMINOPSEUDOURIDINE ...... 102 6.3.1 ((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d] [1,3] dioxol-4-yl) methyl methane sulfonate (106) 102 6.3.2 5-((3aS,4S,6R,6aR)-6-(azidomethyl)-2,2-dimethyltetrahydrofuro[3,4-d] [1,3] dioxol-4-yl)-2,4-di-tert-butoxypyrimidine (107) ...... 103 6.3.3 ((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d] [1,3] dioxol-4-yl) methanamine (108) ...... 104 6.3.4 Synthesis of desoxypseudouridimycin ...... 105 vii

REFERENCES ...... ERROR! BOOKMARK NOT DEFINED. APPENDIX A ...... 117 SPECTRA DATA ...... 117

viii

List of Tables

Table 1: LCMS conditions used for the photolysis of 13 ...... 68 Table 2: Mass balances of products from photolysis of 6 in the absence of GSH ...... 70 Table 3: Mass balance from the photolysis of compound 13 in the presence of GSH (4 eq) ...... 71 Table 4: Mass balance from the photolysis of compound 13 (7 nmol) in the presence of GSH ( 2mM) GSH, in Ammonium Acetate buffer (5mM) pH 7.0 ...... 75 Table 5: Spontaneous Mutation Rates of PUM and Rif41 [Reprinted with Permission] . 80

List of Figures

Figure 1: Building Blocks of RNA ...... 2 Figure 2: Structures of Uridine and Pseudouridine ...... 5 Figure 3: Various Types of RNA Modifications ...... 10 Figure 4: The Fate of the C5'-pseudouridinyl Radical Precursor ...... 13 Figure 5: Causes of Oxidative Stress. [ Reprinted with permission] ...... 14 Figure 6: Mechanisms of Free Radical Formation ...... 16 Figure 7: Radical Species in RNA ...... 18 Figure 8:Oxidized Nucleosides ...... 20 Figure 9: Photolabile Radical Precursors ...... 23 Figure 10: C5´ tert-Butyl Ketone Derivatives of Thymidine 26 and 2´-deoxyguanosine 27.(32) ...... 24 Figure 11: Pathway for the Stereoselective reduction of lactol 6 and 72 by Hanessian et. 39 al. A shows the pathway in the presence of ZnCl2 and B shows the pathway in the absence of ZnCl2...... 38 Figure 12: Numbering System for Nucleosides ...... 51 Figure 13: Mass spectral analysis of pseudouridine 6 (Independently synthesized) ...... 64 Figure 14: Fragmentation patterns of Pseudouridine by Edward Dudley [Reprinted with Permission] ...... 65 Figure 15: Fragmentation patterns of the Mass spectral analysis of independently synthesized pseudouridine 6...... 66 Figure 16: Fragmentation patterns of the Mass spectral analysis of independently synthesized pseudouridine 6...... 67 Figure 17: Chromatogram for the photolysate of 6 (10 mM) at 15 min, 30min, and 45min...... 69 Figure 18: Chromatogram for the photolysate of 6 (10 mM) at 60 min with/without GSH...... 70 Figure 19: Calibration curve of compound 6 ...... 71 Figure 20: Calibration curve of compound 13 ...... 72 Figure 21: Photolysis of Radical Precursor 13 ...... 73 Figure 22: Radical precursor 13 conversion upon photolysis in the presence of GSH ( 2mM) in Ammonium Acetate buffer ( 5mM) at pH 7.0...... 74 Figure 23: Formation of pseudouridine (6) upon photolysis of 13 in the presence of GSH ( 2mM) in Ammonium Acetate buffer ( 5mM) at pH 7.0 ...... 75 Figure 24: Structure of Pseudouridimycin (PUM) ...... 76

Figure 25: Inhibition of RNAP-Dependent RNA Synthesis.47 [Reprinted with Permission] ...... 79 Figure 26: Target of PUM in the RNA NTP Addition Site. [ Reprinted with Permission] ...... 81 Figure 27: Target of PUM in the RNA NTP Addition Site. [ Reprinted with Permission] ...... 83 Figure 28: Structural basis of Transcription Inhibition by PUM.41 [Reprinted with Permission] ...... 84 Figure 29: Structure-Activity Relation of PUM47 ...... 88 Figure 30: 1H NMR for compound 61 ...... 117 Figure 31: 1H NMR for compound 62 ...... 118 Figure 32: LCMS for Compound 62 ...... 119 Figure 33: 1H NMR for compound 57 ...... 120 Figure 34: 1H NMR for compound 80 ...... 121 Figure 35: LCMS for compound 80 ...... 122 Figure 36: HRMS for compound 80 ...... 123 Figure 37: 1H NMR for compound 81 ...... 124 Figure 38: LCMS for compound 81 ...... 125 Figure 39: 1H NMR for compound 82 ...... 126 Figure 40: LCMS for compound 82 ...... 127 Figure 41: HRMS for compound 82 ...... 128 Figure 42: 1H NMR for compound 60 ...... 129 Figure 43: LCMS for compound 60 ...... 130 Figure 44: HRMS for compound 60 ...... 131 Figure 45: LCMS of compound 6 ...... 132 Figure 46: 1H NMR for compound 86 ...... 133 Figure 47: LCMS for compound 86 ...... 134 Figure 48: 1H NMR for compound 87 ...... 135 Figure 49: 13C NMR for compound 87 ...... 136 Figure 50: 1H NMR for compound 89 ...... 137 Figure 51: 1H NMR for compound 90 ...... 138 Figure 52: HRMS for compound 90 ...... 139 Figure 53: 1H NMR for compound 13 ...... 140 Figure 54: 13C NMR for Compound 13 ...... 141 Figure 55: LCMS for Compound 13 ...... 142 Figure 56: 1H NMR for compound 106 ...... 143 Figure 57: 1H NMR for compound 107 ...... 144 Figure 58: LCMS for compound 107 ...... 145 Figure 59: 1H NMR for compound 108 ...... 146 Figure 60: 1H NMR for compound 108 ...... 147 Figure 61: 13 C NMR for compound 108 ...... 148 Figure 62: COSY for compound 108 ...... 149 Figure 63: HSQC for compound 108 ...... 150 Figure 64: 1H NMR for compound 110 ...... 151 Figure 65: LCMS for compound 112 ...... 152 Figure 66: 1H NMR for compound 111 ...... 153 xi

Figure 67: 13C NMR for compound 111 ...... 154 Figure 68: HRMS for compound 114 ...... 155

xii

List of Schemes

Scheme 1: The Synthesis of C5´ tert-Butyl Ketone 26 as a 5´-Thymidinyl Radical Precursor .32 ...... 25 Scheme 2: Deprotection Strategy for the Synthesis of 26.32 ...... 26 Scheme 3: Generation of the C5' Radical from 26 and 32.32 ...... 27 Scheme 4: The Synthesis of the C5' tert-Butyl Ketone of 2'-deoxyguanosine 27.32 ...... 28 Scheme 5: The Synthesis of the C5' tert-Butyl Ketone of 2'-deoxyguanosine 27.32 ...... 29 Scheme 6: Synthesis of a C5´radical precursor 49.33,34 ...... 31 Scheme 7: Photolysis Studies of 49 at pH 3.5 and 7.0.33,34 ...... 32 Scheme 8: Photolysis of C5'-pseudouridine radical precursor ...... 33 Scheme 9: Synthesis of Pseudouridine and Confirmation of the 5-D-ribofuranosyluracil by Shapiro et al.36 ...... 34 Scheme 10: Synthesis of Pseudouridine as Reported by Brown et al.37 ...... 35 Scheme 11: Synthesis of Pseudouridine by Chow et al.38 ...... 36 Scheme 12: Synthesis of Pseudouridine as reported by Hanessian et al.39 ...... 37 Scheme 13: Synthesis of unmodified pseudouridine ...... 39 Scheme 14: Synthesis and Mechanism of Formation of Fully Protected Ribonolactone 62 ...... 40 Scheme 15: Synthesis of Compound 80 ...... 41 Scheme 16: Synthesis of Compound 81 and 82 ...... 43 Scheme 17: Synthesis of C5'-Radical Precursor of Pseudouridine ...... 44 Scheme 18: Synthesis of C5´-Radical Precursor 13 ...... 47 Scheme 19: Semi-synthesis of PUM Derivatives47 ...... 86 Scheme 20: Total Synthesis of Desoxy-PUM 10547 ...... 90 Scheme 21: Retrosynthetic Analysis ...... 93 Scheme 22: Synthesis of Primary C5'-Aminopseudouridine 108 ...... 94 Scheme 23: Attempted synthesis of C5-aminopseudouridine nucleoside 109 ...... 95 Scheme 24: Synthesis of C5-aminopseudouridin nucleoside 113...... 96 Scheme 25: Synthesis of compound 110 ...... 97 Scheme 26: Synthesis of compound 111 ...... 98 Scheme 27: Synthesis of compound 113 ...... 99

xiii

List of Abbreviations

ACN ...... Acetonitrile AD ...... Alzheimer’s disease

13C NMR…………… Carbon-13 nuclear magnetic resonance spectroscopy C-C ...... Carbon-Carbon C-N ...... Carbon-Nitrogen COSY ...... Correlation spectroscopy

DCM ...... Dichloromethane DIAD ...... Diisopropyl azodicarboxylate DMSO ...... Dimethyl sulfoxide DNA ...... Deoxyribonucleic acid

GSH ...... Glutathione

HPLC-ECD ...... High performance liquid chromatography-electrochemical detection HRMS ...... High resolution mass spectrometry HSQC ...... Heteronuclear single-quantum correlation experiment

IRES ...... Internal ribosomal entry sight

L-selectride ...... Lithium tri-sec-butylborohydride LC/MS ...... Liquid chromatography/mass spectroscopy min ...... Minutes mRNA………………Messenger ribonucleic acid

NADPH ...... Nicotinamide adenine dinucleotide phosphate NMR ...... Nuclear magnetic resonance spectroscopy

PUS ...... Pseudouridine synthase PUM…………………Pseudouridimycin

RBF ...... Round-bottom flask RNA ...... Ribonucleic acid

xiv rRNA ...... Ribosomal ribonucleic acid RT ...... Room temperature ROS ...... Reactive oxygen species snRNA ...... Small nuclear ribonucleic acid snoRNA ...... Small nucleolar RNA TBAF ...... Tetra-n-butylammonium fluoride TBDMS ...... Tert-butyldimethylsilyl TBDMSCN ...... Tert-butyldimethylsilyl cyanide TBDPSCl ...... Tert-butyldiphenylsilyl Chloride THF ...... Tetrahydrofuran TLC ...... Thin layer chromatography tRNA ...... Transfer ribonucleic acid

U ...... Uridine UV ...... Ultraviolet

WHO ...... World Health Organization

List of Symbols

%...... Percent (-)………………….Minus (+)………………….Plus

Ar…………………. Argon

13C-NMR…………Carbon nuclear magnetic resonance 1H-NMR…………..Proton nuclear magnetic resonance

CD3OD ...... Deuterated methanol CDCl3 ...... Deuterated chloroform CeCl3 ...... Cerium chloride CN- ...... Cyanide anion CH2Cl2……………. Dichloromethane CH3CN……………. Acetonitrile CH3COOH…………. Acetic acid

DMSO ...... dimethyl sulfoxide et al ...... And others Et3N ...... Triethyl amine EtOAc ...... Ethyl acetate

F- ...... Fluoride ion

GSH………………. Glutathione reduced

H•…………………. Hydrogen atom H2O………………...Water HO• ………………. Hydroxyl radical

Hν…………………. Energy

IBX ...... 2-iodoxybenzoic acid i-Pr ...... Isopropyl xvi

KCN ...... Potassium cyanide

M…………………...Molar Mg2+ ...... Magnesium ion MgSO4 ...... Magnesium sulfate MeOH……………...Methanol mM………………. millimolar

NH3………………. Ammonia NaCl ...... Sodium Chloride NaHCO3 ...... Sodium bicarbonate NaOH ...... Sodium hydroxide

OH ...... Hydroxyl t ...... Triplet TBAF ...... tetrabutylammonium fluoride PPh3………………. Triphenylphosphine TMSCN ...... Trimethyl cyanide TFA…………………Trifluoroacetic acid

Ψ………………. ……Pseudouridine α ...... Alpha β ...... Beta δ ...... Delta π ...... Pi

xvii

Chapter 1

1 Introduction

1.1 Overview

Nucleic acids are a class of biomolecules found in all living organism and are responsible for the storage of genetic information and protein synthesis. The two types of nucleic acids are DNA (deoxynucleic acid) and RNA (ribonucleic acid). Nucleic acids are biopolymers made up of monomers called nucleotides. These monomers consist of three components; a five-carbon sugar, a phosphate functional group, and a nitrogenous base. The five-carbon sugar present in DNA is deoxyribose, and the four nitrogenous bases are adenine (A), guanine (G), thymine (T), and cytosine (C). Oligonucleotides are biosynthesized by polymerization reactions between the sugar of one nucleotide and the phosphate group of the subsequent nucleotide. In this process, a sugar-phosphate backbone is formed linking the nucleotides together.

DNA is composed of two antiparallel strands held together by hydrogen bonding interactions between the bases forming a double helix. A bonds with T and G bonds with

C. This complementary base pairing links strands together and plays an essential role in the stability of the DNA molecule.

RNA has some properties that differ from DNA. The pentose sugar present in

RNA is ribose whereas DNA contains a deoxyribose. Additionally, the nitrogenous base uracil (U) is present in RNA in lieu of thymine (T).1

Figure 1: Building Blocks of RNA

Contrary to DNA, RNA is predominantly observed endogenously as single- stranded and does not typically form a helix. RNA is less stable than DNA: the presence

of the 2´ -hydroxyl group on the pentose renders it more susceptible to alkaline hydrolysis. DNA stores the information that is needed to construct a protein and RNA functions in regulating the expression of the information during the protein synthesis process. RNAs are present in the nucleus and the cytoplasm. RNA is synthesized from

DNA by an enzyme known as RNA polymerase during a process called transcription where the DNA unwinds, and the RNA is made from the 5´ to 3´ direction. The transcript sequences are complementary to their DNA template, then translated to proteins by the ribosomes. RNAs are classified according to their role in the translation process: messenger RNA (mRNA), transfer RNA (tRNA), and ribosomal RNA (rRNA), among others. Even though some RNA molecules are passive copies of DNA, many play crucial active roles in the cell. For instance, some RNA molecules participate in switching genes on and off while others make up the critical protein synthesis machinery in ribosomes.2

In this dissertation, our ever-broadening insights into disease processes that are related to

RNA damage and RNA modifications will be utilized to furnish an opportunity for clinical laboratories to develop and implement novel approaches to the diagnostic assessment and treatment of human disease.

1.2 RNA Damage in Disease Etiology

Recent studies have shown that damage caused to RNA increases under oxidative stress and in patients with neurodegenerative diseases. This has drawn attention to the consequences of RNA oxidation at a molecular and cellular level. In comparison, the levels of oxidative damage in RNA are usually higher than in DNA, and this can impair

RNA function and protein synthesis. During RNA synthesis, oxidized ribonucleotides are hydrolyzed or discriminated from intact ribonucleotides during transcription, preventing their incorporation into RNA.3 Collective evidence suggests that RNA oxidative damage is a challenging and persistent problem frequently controlled through RNA surveillance mechanisms critical to maintaining cellular health and preventing diseases.

1.3 Ribonucleosides in Drug Discovery

Drug that target DNA and RNA can either act on the nucleic acids by compromising their structural integrity or inhibit their synthesis. Inhibitors of nucleic acid synthesis usually act as enzyme inhibitors or antimetabolites. Inhibitors that act on existing nucleic acid molecules can be classified broadly into alkylating agents, intercalating agents, and chain-terminating agents. However, these classifications are not rigid, as the drugs can act by more than one mechanism. Inhibitors acting on existing

DNA usually inhibit transcription while inhibitors on existing RNA normally act on translation. The net result in both cases is the prevention of cell growth and cell division.

Consequently, the discovery of new therapeutics that target existing DNA and RNA is a significant consideration when discovering drugs for bacterial, infections due to micro- organism or cancer.

Chapter 2

2 Background

2.1 Role of Pseudouridine in Disease Etiology

Pseudouridine (Ψ), a ubiquitous constituent of ribonucleic acid (RNA), was the first naturally modified nucleoside to be discovered. Ψ is the most abundant modification that has been identified in RNA. Hence it is termed the “fifth nucleoside”. 4

Figure 2: Structures of Uridine and Pseudouridine

Pseudouridine is an isomer of uridine which contains a C-C glycosidic bond whereas uridine has a C-N glycosidic bond. This posttranscriptional modification in humans is 5

performed by the enzyme pseudouridine synthase through the isomerization of uridine to pseudouridine in oligonucleotides.4,5 The function of this modification is not fully understood; however, there is evidence to support the theory that, Ψ could make the RNA backbone more rigid, enhancing the stability of secondary RNA structures, while affecting base stacking and conformational switches in a different way than uridine.6

Earlier reports correlating uridine modifications and human diseases concentrated on urinary metabolites in cancer patients.7,8 Ψ, like other modified nucleosides, cannot be reused and is excreted with urine. Levels of Ψ in urine are dependent on the glomerular filtration rate and RNA turnover. These levels are often found to be higher in cancer patients.9 Hence, the evaluation of urinary Ψs levels has been suggested as a potential tumor marker 10 but the assessment of urinary Ψ levels is currently not incorporated in routine diagnostics.

2.2 RNA Occurrence in Living Organisms

RNA is found mainly in the cytoplasm and the nucleolus. When RNA is inside the cytoplasm, it occurs freely and within the ribosomes. RNA is also detectable in the mitochondria and chromosomes in eukaryotic cells.

2.2.1 Bacteria

Bacteria are prokaryotic organisms without a membrane-bound nucleus. Their nuclear components are either found scattered in the cytoplasm or the nucleoid. They have a primitive DNA which is a single circular chromosome. Bacteria have both DNA and 6

RNA which obey the central dogma where the DNA genome is transcribed to RNA then translated to protein.

2.2.2 Viruses

Viruses are intracellular parasites that cannot reproduce by themselves but can replicate only after penetrating and infecting a susceptible host cell. Once inside the cell, the virus can direct the cell machinery to produce more copies of its genome and package them into new viruses. Viral infection occurs when proteins on the surface of a virion bind to specific receptors on the surface of the host cells. The specificity of this binding interaction is what determines the host range of a virus. Viruses naturally consist of nucleic acids, RNA or DNA as their genetic material and are covered by an outer shell protein coating. RNA viruses that causes human diseases include Hepatitis C (HCV),

Ebola, Severe Acute Respiratory Syndrome (SARS), influenza, polio, measles, and human immunodeficiency virus (HIV). In these viruses, RNA serves as the genetic material and may be single or double-stranded.1 The viruses usually exploit the presence of the enzyme RNA-dependent RNA polymerases for the replication process of their genomes. Retroviruses are a notable exception to the central dogma.

Retroviruses contain two copies of single-strand RNA genomes; reverse transcriptase produces the viral DNA that is integrated into the host DNA through its integrase function.

2.2.3 Mammalian cells

RNA is several times more abundant in mammalian cells. In these cells, RNA is present in the nucleus and largely in the cytoplasm. In addition to the nucleus, mitochondria contain DNA that encodes for RNA and proteins unique to the organelle. Also, RNA occurs in multiple copies and various forms; tRNA, mRNAs, rRNA, and small nuclear

RNA (snRNA) among others.

2.3 Functions

Within the nucleus, DNA is transcribed into mRNA which contains the codons that code for proteins. The mRNA once formed can be modified in several important ways and ultimately ends up in the cytoplasm of the cell, where it will be read by the ribosomes to enable protein synthesis. In eukaryotes, mRNA only codes for one protein but in prokaryotes, a single mRNA molecule can be used to synthesize many different proteins.

Ribosomal RNA is synthesized in the nucleolus of the nucleus and eventually travels to the cytoplasm of the cell. In the cytoplasm, rRNA combines with proteins to form ribosomes which are the machinery for protein synthesis. mRNA molecules contain regions of codons which designate the amino acid that should be added next on to the growing polypeptide chain. tRNAs are supercoiled molecules that collect the appropriate amino acid and bring it to the ribosome, where the amino acid is then attached to the growing protein.

Hence all types of these different RNA molecules are synthesized inside the nucleus and involved in the process of the synthesis of proteins. Each serves its unique function.

2.4 Modified Ribonucleic Acids

Posttranscriptional RNA modifications occur via a complex mechanism and have the potential to alter its function and stability. These changes to the chemical structure of

RNA occur in the nucleus where DNA is transcribed before the mature mRNA is transported to the cytoplasm. Post-transcriptional modifications include 5´modification or capping where a methylated guanosine monophosphate (GMP) is added at the 5´-5´ triphosphate linkage of the pre-mRNA, along with the addition of multiple adenosine monophosphates (poly(A) tail) at the 3´ terminus and splicing of the introns and reordering of the RNA by joining of the exons. In recent years, many naturally occurring modifications to nucleosides have been identified with more than 100 documented in all domains of life and 66 occurring in eukaryotic cells.11 The modifications of adenosine, cytidine, guanosine, and uridine frequently occur by processes such as methylation in tRNA, mRNA, rRNA snRNA and other types of RNA (Figure 3).12

Figure 3: Various Types of RNA Modifications

Although the function of some of these modifications are well known, many remain a mystery. In rRNA and tRNA, these post-transcriptional modifications could contribute significantly to the structure and stability of the nucleic acid as well as provide enhanced accuracy and efficiency of translation and localization.5 The functional dynamics of

RNA, its synergistic nature and the trafficking regulatory roles of these posttranscriptional modifications within the cells are not well characterized. This is because many modified RNA bases are recognized by reverse transcriptase the same way as their unmodified counterparts. Because a routine step in many RNA experiments is to reverse transcribe the RNA into complementary DNA (cDNA), this effectively erases any

information concerning the types and locations of RNA modifications that might have been present. A second reason is that the technical ability to detect and quantitate RNA modifications has been limited until recently. Both issues have severely impaired the ability to systematically characterize the epitranscriptome, which is all the chemical modifications of RNA molecules, that is both coding and noncoding. Hence, the functional roles of many post-transcriptional RNA modifications remain unknown, although they could potentially influence frameworks, such as RNA stability, translation, trafficking, localization, enzymatic activity, or patterns of interactions with other molecules.12,5 The most frequent modifications are 2´-O-methylation and pseudouridinylation. 12 Other modifications include 6-methyl adenosine, 5- methylcytosine, and inosine (Figure 3).13

Pseudouridine (Ψ) 6 as mentioned earlier, (Figure 2) is the most abundant, with around

5% cellular abundance and the first modified nucleoside to be discovered and identified.

Chemically, Ψ possesses an extra hydrogen bonding opportunity at the secondary N-1 position, that can potentially participate in novel base-pairing interactions.5

Transfer RNAs are the most modified RNAs. In eukaryotic cells, about 15%-25% of all tRNAs are modified nucleosides.13 These modifications are thought to be essential for translational efficiency and tRNA analysis. Therefore, translation can be directly affected by tRNA modifications.

Ribosomal RNA modifications include three basic types: base methylation, ribose methylation, and pseudouridylation (Figure 3). Base methylation is the most conserved in total number and position among all species, with bacteria containing slightly more than the ten commonly found in eukaryotes and occurs only in highly conserved rRNA 11

sequences.14 Ribose methylation always occurs at the 2´ hydroxyl position on the sugar backbone, frequently in eukaryotes but limited in bacteria. One proposed theory as to the function of 2´-O methylation is that it assists with tRNAs interactions.14 Interestingly, ribose methylation occurs in highly evolutionary conserved regions of rRNA, in many cases co-clustering with Ψ residues. 15 Because most ribose methylation takes place early in rRNA processing, it is hypothesized to be essential for rRNA folding or association with chaperone proteins that may aid in folding.15

Besides the 5´ cap and inosine that are naturally occurring modifications in eukaryotic mRNA, N6-methyladenosine (m6A), pseudouridine (Ψ), 5-methylcytosine (m5C) and N1- methyladenosine (m1A) are also included (Figure 3).16 m6A is the most abundant and the best-characterized internal mRNA modification, and is involved in various biological functions ranging from splicing to regulation of translation to mRNA decay.17 Although pseudouridine is dynamically deposited in mRNA transcripts, its functions within the mRNA are still not clearly understood. 17–20

2.4.1 Pseudouridine Occurrence and Disease Relevance

Studies have shown that urinary levels of pseudouridine are higher in Alzheimer’s

Disease (AD) patients and that RNA oxidation plays a significant role in the pathogenesis of Alzheimer’s Disease (AD) and other neurodegenerative diseases. 20 Pseudouridine, a post-transcriptional modified nucleoside, can be assumed to correlate with oxidative stress and higher urinary levels in AD.20 Hence, understanding the role of pseudouridine by subjecting it to oxidative conditions may provide some vital pieces of information

about the role of this substrate in RNA processes. There is a possibility that under oxidative conditions, there is an increase in the biosynthesis of pseudouridine; this will explain the high urinary levels in AD. Another scenario may be that, under oxidative stress, uridine is isomerized to pseudouridine by pseudouridine synthetase or some other mechanism. Pseudouridine has also been shown to play a regulatory role in maintaining the various structures of RNA. Hence, it can be hypothesized that isomerization of U to Ψ may be a defense mechanism for the cell to protect itself from oxidative stress.21 In order to study and understand the mechanisms involved in the damage caused to RNA, pseudouridine will be synthesized and subjected to oxidative damage. The first part of this study will involve the synthesis of a C5´-pseudouridinyl radical precursor and an investigation into the fate of the photochemically generated pseudouridine C5´-ribosyl radical (Figure 4).

Figure 4: The Fate of the C5'-pseudouridinyl Radical Precursor

2.5. RNA Oxidation in Human Cells

2.5.1 Oxidative Stress

Oxidative stress is the result of the generation of highly reactive small molecules by endogenous and exogenous sources. These molecules are nitrous oxide, superoxide and the hydroxyl radical to name a few. Due to their high reactivity and size, they can play a vital role in damaging biological molecules such as nucleic acids when they are generated in the cell (Figure 5). An increase in the level of reactive oxygen species (ROS) and a decrease in antioxidant levels is the true nature of oxidative stress.3

Figure 5: Causes of Oxidative Stress. [ Reprinted with permission]

Free radicals that cause oxidative stress are generated via a variety of mechanisms. They can be generated during the reduction-oxidation (redox) reactions that occur during 14

normal metabolic processes, by absorption of radiant energy and enzymatic metabolism of certain exogenous chemicals or drugs. Some transition metals, for example, iron and copper, are also responsible for the generation of free radicals. During normal metabolic redox processes, cells generate ATP. During this reaction, four electrons are donated in four steps. The production of partially reduced intermediates are what becomes the free radicals.

The formation of superoxide occurs by the transfer of one electron by ortho oxidation.

Ortho oxidation can be seen in the electron transport reaction that occurs inside the mitochondria. Also, superoxide can be produced by enzymatic reactions that include specific enzymes such as xanthine oxidases and cytochrome p450s that are in the cytosol and mitochondria. The catabolism of superoxide produces hydrogen peroxide through the transfer of two electrons by the enzyme superoxide dismutase (SOD). SODs include both manganese superoxide located in the mitochondrial and copper-zinc superoxide located in the cytosol. Hydroxyl radicals are generated by the transfer of three electrons. This radical can be produced in two ways, first from hydrogen peroxide through the Fenton

22 reaction and secondly from H2O during radiolysis. (Figure 6).

Figure 6: Mechanisms of Free Radical Formation

The generation of free radicals can also occur by the absorption of radiant energy, for example, absorption of ultraviolet (UV) light and X-rays resulting in hydrolysis of H2O into hydroxyl ions and hydrogen free radicals. More so, during inflammation, leukocytes can become activated and carry out precise reactions in the plasma membrane multi- protein complex. These reactions involve the use of NADPH oxidase enzymes and xanthine oxidases that can generate superoxide anions as well.22 Metabolism of exogenous chemicals and drugs can also generate free radicals — for example, enzymatic metabolism of carbon tetrachloride (CCl4) produces changes in metabolic activity of cytochrome P450 the enzyme responsible for the oxidation of xenobiotic chemicals. Iron and copper can either donate or accept free electrons during various intracellular

reactions and catalyze the formation of free radicals through the Fenton reaction.

Another mechanism of radical formation involves nitric oxide (NO) generation by neurons, macrophages, and endothelial cells. Nitric oxide acts as a free radical and can be converted to a more reactive free radical when combined with superoxide and forms peroxynitrite (ONOO-). 22 Hence, oxidative damage is a constant occurrence of biomolecules. Increased generation of these ROS causes oxidative stress which in turn plays a significant role in the pathology of several diseases including cancer and neurological disorders. It is crucial to understand these mechanisms of the formation of radical derived intermediates and their association with outcomes in biological systems.

2.5.2 Mechanisms of Oxidative Damage to RNA

The human body has various intrinsic mechanisms to counteract oxidative damage by producing antioxidants such as glutathione. Additionally, antioxidants can be obtained through the diet. However, whenever there is an excess of free radicals their accumulation in the body leads to a phenomenon called oxidative stress (Figure 5). Of great significance, under conditions of oxidation in living cells is the creation of the highly reactive hydroxyl radical (OH•). Hydroxyl radicals could well be directly responsible for a significant amount of the oxidative insults in biological macromolecules, including RNA. This highly ROS can abstract a hydrogen atom from any molecule that it encounters. Its reactivity is limited only by distance and it tends to

abstract hydrogen atoms from any position of a molecule that is exposed to solvent.

When the target is RNA, nucleic acid radicals are produced through the abstraction of hydrogen atoms from both the sugar and base moieties of RNA (Figure 7)

Figure 7: Radical Species in RNA

This can lead to the impaired physiological function of RNA. The resulting cellular damage and impact on specific signaling pathways can lead to aging, disease development and in many cases cell death.

2.5.3 Role of RNA Oxidation in Disease Processes

As mentioned earlier, oxidative stress and oxidative damage to RNA have been shown to be significant factors in the pathology of many neurodegenerative diseases such as

Alzheimer’s Disease, Parkinson’s Disease, Huntington’s Disease, vascular dementia, and amyotrophic lateral sclerosis (ALS).23–25. Oxidative damage due to ROS has remained associated with the pathogenesis of cancer, Down syndrome, diabetes, and aging. There is evidence, that oxidative stress could also result in cardiovascular issues such as myocardial infarction and angina.22 Because the brain has a high oxygen consumption rate, and is rich in peroxidizable unsaturated fatty acids and possesses insufficient levels of antioxidants, it is more susceptible to the effects of oxidative stress. Although it is not clear if oxidative stress is a cause or effect of neurodegenerative diseases, it is indeed clear that it plays a major role in the events leading to neuronal death.23 In Alzheimer’s disease, for example, ROS generated in neuronal cell bodies have a low rate of diffusion and are localized in the neurons where they are responsible for oxidative stress which in turn leads to neuron death.

Figure 8:Oxidized Nucleosides

Recent studies observed that there was an increase in oxidative DNA and RNA damage in the post mortem brain of Alzheimer's Disease and Parkinson's Disease patients. Also, alteration of ribosomal functions were seen.26 With the use of specific antibodies, 8- hydroxyguanosine (8-OHG) and 8-hydroxydeoxyguanosine (8-OHdG) (Figure 8) were detected in the cytoplasm of vulnerable neurons. Treatment of 8-OHG and 8-OHdG with

DNases and RNases abolished the immunoreactivity of 8-hydroxydeoxyguanosine; and

8-hydroxyguanosine was greatly reduced when RNase was used, indicating these oxidized nucleosides were more often associated with RNA than DNA.27 In these experiments, when 8-OHG/8-OHdG was pretreated with RNase, the majority of the oxidized 8-OHG were mostly localized in the ribosome.

RNA oxidative damage is regionally distributed in the brain based on the neurological disease. In brain samples of patients with AD, there was an increased level of 8-OHG detected in the hippocampus and cerebral neocortex.28 Increased RNA oxidation was 20

observed in the substantia nigra in PD disease and the motor cortex and spinal cord of patients with amyotrophic lateral sclerosis (ALS).28

Increased levels of RNA damage are a common symptom seen in neurodegenerative disorders and has exhibited disastrous effects on the neurons. Oxidative stress has also been shown to play a vital role in the reduction of protein expression, protein misfolding, and protein aggregation seen in these diseases. The exact role of RNA oxidation is not clear in neurodegenerative diseases, but the mechanisms leading to neuron death are defined.

Chapter 3

3 Synthesis of a C5´-Pseudouridinylradical Precursor

3.1 Synthesis of Modified Ribonucleosides as C5´-Radical Precursors

Modified nucleosides have been synthesized by different groups such as Greenberg,3

Giese,29,30Chatgilialoglu,31,32 and Bryant-Friedrich.33 Chatgilialoglu’s group synthesized

C5´-thymidine and C2´guanosine radical precursors.32 C6 and C2´-uridine radical precursors were synthesized by Greenberg’s group, and Giese’s worked on a 3´ and 4´- thymidine precursor. The Bryant-Friedrich group published a C3´-thymidinyl and C5´- uridinyl precursor. As this dissertation is focused on modifications on the ribose at the

C5´ position, synthetic methods to obtain these modified nucleosides will focus on this position. The tert-butyl ketone group has been established as a suitable photolabile moiety to generate radicals at a specific site in nucleosides and nucleotides. Several examples of modified nucleosides utilizing this group are shown in Figure 9. Radical precursors using this group have been synthesized at different positions on nucleosides.29,33 These modified nucleosides were also incorporated into oligonucleotides to study the radical induced products under more biologically relevant conditions. In

addition to the tert-butyl ketone moiety, other groups such as methyl ketones 31,32 and phenyl selenides 29,32,32 have been utilized for selective generation of radicals (Figure 9).

Figure 9: Photolabile Radical Precursors

These groups were, however, found to be less efficient compared to the tert-butyl ketone.

Tert-butyl ketones smoothly undergo Norrish type I photocleavage to generate radicals at the desired position without forming by-products which interfere with studies. On the other hand, methyl ketones can undergo Norrish type II reactions which lead to the formation of undesirable by-products.32 Similarly, phenyl selenides showed the formation of multiple photoproducts from different pathways which lowers the chemical yield of the desired radical. 29,31,32 Therefore tert-butyl ketones are more commonly used for the synthesis of radical precursors. Through careful comparison of these and other 23

precursors, we decided that the tert-butyl ketone group would likely be most efficient as a radical precursor in our system to study the damage products obtained from the generation of the C5´-pseudouridinyl radical.

Many studies have focused on DNA damage or oxidation both at the nucleobase and the sugar, but the fate of C5´-radicals in DNA sugars have not been extensively explored.

Studies on the generation of radicals at all other positions on the sugar lead to the elimination of the nucleobase. Most of the research on the C5´-radical in nucleosides through site-specific generation comes from the Chatgilialoglu and the Bryant- Friedrich groups.

Figure 10: C5´ tert-Butyl Ketone Derivatives of Thymidine 26 and 2´-deoxyguanosine 27.(32)

Chatgilialoglu’s group synthesized both C5´pivaloyl thymidine and 2´-deoxyguanosine radical precursors (Figure 10) as a tool to study DNA damage through the formation of a ribose radical.

The synthesis of C5´ thymidine radical precursor 26 began with C5´ aldehyde 28

(Scheme 1).32

Scheme 1: The Synthesis of C5´ tert-Butyl Ketone 26 as a 5´-Thymidinyl Radical Precursor .32

The aldehyde was converted into cyanohydrin 29 as a mixture of isomers. This mixture was also observed in work leading to the 3´-thymidine radical precursors by Giese.34

Reaction of cyanohydrins with tert-butyllithium lead to the formation of a tert-butyl imine 30. Hydrolysis of the imine in THF/water/2N HCl solution gave the corresponding fully protected ketone 26 in 82% yield.32

Compound 26 was treated with tetra-n-butylammonium fluoride (TBAF) and THF to afford 31, and final treatment of 31 with TBAF and methanol under reflux furnished fully deprotected precursor 32 (Scheme 2).32

Scheme 2: Deprotection Strategy for the Synthesis of 26.32

Photolysis of thymidine derivatives (scheme 3) 26 and 232in the presence of reducing agents led to the formation of the expected thymidine derivatives. Photolysis of the protected thymidine radical precursor 26 in the presence of 1-butanethiol as a hydrogen donor produced 36 and photolysis of unprotected 32 biologically relevant reducing agent glutathione produced 34.

Scheme 3: Generation of the C5' Radical from 26 and 32.32

Synthesis of the guanosine derivative was performed in a similar a manner as that of thymidine (Scheme 4). Through Moffatt oxidation using EDC, TFA, pyridine, and toluene/DMSO (2:1), the formation of the C5´ aldehyde was achieved in 95%. The products of the oxidation were determined to be a mixture of three diastereomeric hemiacetals — the diastereomers formed due to hydration of the aldehyde in aqueous solution. Treatment of the mixture 37 with TBDMS-CN in the presence of ZnI2 yielded

5´-cyanohydrins 38 as a mixture of diastereomers. These isomers where not separated.

The reaction of the cyanohydrins with TBDMSCl and imidazole in THF gave fully protected cyanohydrin 38. Reaction of 38 with tert-butyllithium produce imine 39.

Chatgilialoglu’s group found that 39 was stable to the conditions of column

32 chromatography CHCl3/MeOH, 97:3. Hence, it was isolated and fully characterized.

The reaction of the imine with acetic acid/THF/water (4:1:1) furnished the fully protected radical precursor 27 in 78% yield.

Scheme 4: The Synthesis of the C5' tert-Butyl Ketone of 2'-deoxyguanosine 27.32

Upon completion of the synthesis of 27 photolysis studies were carried out on the substrate in the presence of tert-Butylthiol (tBuSH) as a hydrogen atom donor (Scheme

5). The study resulted in the formation of two distinct products; guanosine derivative 40 in 27% yield and (5´S)-5´,8-cyclo-2´-deoxyguanosine 41 in 20% yield. It must be noted that, no other diastereomeric products were detected. The competitive pathways which led to the formation of two products both involve radical formation from the photolytic cleavage of the pivaloyl group. The mechanism proposed by Chatgilialoglu can be seen in Scheme 5. The cyclization product 41 forms through a chair transition state of intermediate radical 42 followed by aromatization of the base 43. 32 The stereoselectivity

seen in production formation was attributed to the steric hindrance introduced by the 5´-

O-TBDMS and the guanine moiety.32

Scheme 5: The Synthesis of the C5' tert-Butyl Ketone of 2'-deoxyguanosine 27.32

Chatgilialoglu’s group used similar synthetic methods for the synthesis of the guanosine and thymidine radical precursors to those used by the Giese group.29,32 The approach used for the synthesis of the guanosine substrate established a new route to synthesize the cyanohydrin compound. They also reported the excellent stability of the imine.

Photochemical generation of the radicals established the validity of these substrates as radical precursors for the study of radical damage on nucleosides at the C5´ site of the ribose. Chatgilialoglu has concluded that the fate of a C5´ radical is reliant on the structure of the base.32

The Bryant- Fredrich group was the first to look at the fate of a C5´ radical formation in

RNA through the synthesis of a suitable C5´-uridinyl radical precursor (49) to understand the fate of the C5´radical and its contributions to RNA oxidative damage. Here, uridine was chosen because it is the only nucleoside found in RNA that is not present in DNA.

Hence, understanding the damage done on this nucleoside will give a better understanding of RNA damage.

The pathway to synthesize modified nucleoside 49 was achieved as shown in Scheme

6.33,34 Methods similar to those used by Giese29,30and Chatgilialoglu35were employed.

Literature procedures were followed to attain the C5´ aldehyde 44 using Dess-Martin periodinane for the oxidation of a protected uridine derivative (not shown). No aqueous workup was done to prevent the formation of the corresponding hydrate. TBDMS- protected cyanohydrin 45 was formed using 18-Crown-6, KCN, and TBDMSCN in anhydrous THF in 73% yield. Subsequently, alkylation of 45 with tert-BuLi formed imine 46 followed by hydrolysis to furnish t-butyl ketone 39 in 46% yield over two steps.

Deprotection of compound 47 using TBAF to remove the TBDMS protecting group delivered 48 in 84% yield. Finally, alcohol 48 was refluxed in 25% acetic acid giving the fully deprotected final product, radical precursor 49 in 75% yield.33,34

Scheme 6: Synthesis of a C5´radical precursor 49.33,34

Photolysis of radical precursor 49 was carried under anaerobic conditions in the presence of excess glutathione as a hydrogen atom donor (Scheme 7). Products formed were separated by HPLC and their molecular weights determined by mass spectrometry. The identity of the products was confirmed through the comparison of these products to independently obtained samples of uridine and uracil. Along with the reduction product uridine, the base elimination product uracil was detected under both acidic and physiological conditions. It was proposed that the pH of the solution plays a dominant role in product distribution.33 31

Scheme 7: Photolysis Studies of 49 at pH 3.5 and 7.0.33,34

To determine the similarities in the reactivity of C5´-ribosyl radical in a nucleoside containing a C-C vs. C-N glycosyl bond in RNA, a C5´ pseudouridinyl radical precursor will be synthesized and studied (Figure 4). Our group found that, the C-N glycosyl bond was vulnerable upon generation of the C5´-radical in uridine through the observance of the base release upon photolysis. The proposed mechanism for the cleavage of the C-N glycosyl bond involved a β-CO bond cleavage that led to base release. Because of the presence of the C-C glycosyl bond in pseudouridine, base elimination is not expected in the C5´-Pseudouridinyl radical studies. To our knowledge, the synthesis of pseudouridine radical precursor will be the first that has been performed, and the fate of the radical damage to this substrate determined. 32

To validate the suitability of the C5´-pseudouridinyl radical precursor for radical generation, we photolyzed compound 13 (50:50 acetonitrile/water) under anaerobic conditions in the presence of an excess of GSH a hydrogen atom donor. GSH should reduce the radical upon formation, to deliver pseudouridine (6) as proof that the radical can be photochemically generated at the desired location. Pseudouridine was indeed successfully generated upon photolysis of compound 13 in the presence of an excess of

GSH (Scheme 8).

Scheme 8: Photolysis of C5'-pseudouridine radical precursor

3.2 Synthesis of Pseudouridine and a C5´-Pseudouridinyl Radical Precursor

The eminence of pseudouridine 6 (Figure 2) and related derivatives for the study of RNA and drug development, played a vital role to inspire scientists to develop effective routes to synthesize this substrate. Various methods were established for the synthesis of pseudouridine. We introduced these routes to our approach for the synthesis of pseudouridine and separate modification at the C5´ position to introduce the final compound, C5´-pseudouridine radical precursor. Many routes for the synthesis of pseudouridine yielded a mixture of α and β isomers while the physiological

pseudouridine required for biological studies is exclusively the β-isomer.36 Ψ was first synthesized in 1961 by Shapiro et al. and delivered pseudouridine in 3.5 % yield as a mixture of isomers along with a six-membered heterocyclic ring the 5-D- ribofuranosyluracil 55 substrate as a products (Scheme 9). 36

Scheme 9: Synthesis of Pseudouridine and Confirmation of the 5-D-ribofuranosyluracil by Shapiro et al.36

This study concluded that Ψ could not tolerate strong acidic conditions. However, the synthesis opened doors for synthesizing pseudouridine more efficiently through the coupling of a protected sugar and a lithiated base.

Ψ was also successfully synthesized by Brown’s group 37and Chow’s group.38 Brown’s group enhanced the yield of Ψ from Shapiro et al. by introducing benzyl protecting groups on the sugar moiety instead of benzoyl and replacing methoxy protecting groups on the base with tert-butoxy groups. These base protecting groups were chosen due to their acid lability. The yield was improved to 42% but, the stereoselectivity remained poor with a ratio of 5:2 β/α isomers (Scheme 10). 37

Scheme 10: Synthesis of Pseudouridine as Reported by Brown et al.37

The stereoselectivity of the coupling was enhanced by Chow’s group to get a ratio of 8:1

β/α isomers (Scheme 11). Brown’s synthesis started with a benzoyl protected ribose, but

Chow employed the 2', 3' isopropylidene ribonolactone. Coupling resulted in a 1:8

mixture of α/ β isomers, higher than Shapiro’s and Brown's method. BF3.Et2O/ Et3SiH was used to reduce the C1' alcohol via two separate approaches with varying results, followed by deprotection in two steps in overall 20 % yield.38

Scheme 11: Synthesis of Pseudouridine by Chow et al.38

However, Hanessian’s group improved the yield and stereoselectivity by introducing stereoselective reduction to obtain the desired β anomer in less than five steps in 46% overall yield (Scheme 12).39

Scheme 12: Synthesis of Pseudouridine as reported by Hanessian et al.39 37

The protected D-lactone and the protected base were coupled. The most critical part of this synthesis is formation of the β-anomer in >20/1 ratio β/α was a stereoselective reduction of the ring with L-selectride in the presence of zinc chloride. Zinc chloride chelates with the carbonyl of the ketone and the oxygen of the acetal. Through this method the D-altro-hexitol was obtained as a single isomer in 85% yield. Zinc chelation allowed the delivery of the hydride from the Si-face of the molecule (Figure 11). 39 In the absence of ZnCl2, the α anomer was shown to be the dominant anomer. A Mitsunobu reaction was used to recyclize the ring and to get the protected β-pseudouridine 6.

Figure 11: Pathway for the Stereoselective reduction of lactol 6 and 72 by Hanessian et. 39 al. A shows the pathway in the presence of ZnCl2 and B shows the pathway in the absence of ZnCl2.

To synthesize the unmodified 5´-pseudouridinyl precursor, we chose to modify

Hanessian’s approach based on the fact that it gave the best yield of the β-anomer

(Scheme 13).

Scheme 13: Synthesis of unmodified pseudouridine

We maintained the use of the acetonide group for protection of the 2 and 3 positions of lactone 83 followed by introduction of the tert-butyldiphenylsilyl (tBuPh2Si) protecting group at the 5´ position of the lactone to obtain 62 (Scheme 14). As a result, selective deprotection of the 5´-hydroxyl group could be achieved while keeping the 2 and 3- hydroxyl acetonide groups in place to allow for specific modifications at the C5´- hydroxyl position.

Scheme 14: Synthesis and Mechanism of Formation of Fully Protected Ribonolactone 62

We chose a traditional isopropylidene reaction to obtain 83 employing acetone with hydrochloric acid as the catalyst to obtain the desired compound in a 94% yield (Scheme

14). The 5´-hydroxyl was protected with a tert-butyldiphenylsilyl (TBDPS) group to give

62 in 96% yield. The use of the TBDPS provided a means to maintain a fully protected pseudouridine derivative near the end of the synthesis. This group undergoes deprotection under mild conditions using TBAF while all other protecting groups on the nucleoside needs more acidic approaches. Also, the protecting group of the pyrimidine base was changed to a t-butyl group. Compound 84 is commercially available and undergoes facile ethers protection with the tert-butyl in the presence of potassium tert-

Butoxide (KOtBu) and THF in 98% yield (Scheme 15). Treatment of 85 with tert- butyllithium and THF at -78 0C facilitates a lithium-halogen exchange to give the lithiated pyrimidinyl salt 57. Fully protected ribonolactone 62 in THF was cannulated to lithiated pyrimidinyl base 57 to achieve 80 (Scheme 15) a mixture of diastereomers.

Product formation of 80 was confirmed by spectral analysis (Figure 33). The protons for

C6 were observed at 8.51 and 8.42 ppm as singlets for both isomers. Signals for the phenyl groups were also identified in the aromatic region between 7.66 to 7.40 ppm, and sugar protons were observed between 4.93 and 3.78 ppm. the isopropylidene and tert- 40

butyl groups were observed between 1.66 to 1.01 ppm. The formation of 80 was further supported by LCMS and HRMS analysis (Figure 34 and Figure 35).

Scheme 15: Synthesis of Compound 80

A higher yield was achieved when the di-t-butyl protected pyrimidine was introduced in the lactone with the bulkier silyl protecting group compared to Hanessian. This indicates that steric hindrance may not have a significant impact on this reaction. Compound 80 was stereoselectively reduced using L-selectride in the presence of zinc chloride which favors the formation of β-isomer, 81. The product was confirmed by NMR and HRMS

(Figure 36). A signal for H-6 was observed at 8.28 ppm as a singlet. Moreover, a doublet at 5.25 ppm for H-1' visibly indicated the formation of 81. In parallel, signals for the

sugar protons were viewed between 4.34 to 3.84 ppm. And the signals for all protecting groups were observed in the region between 1.61 to 1.09 ppm.

An intramolecular Mitsunobu reaction of 81 with DEAD and Ph3P for cycloetherification formed the sugar ring to generate α and β- pseudouridine compound 82 (Scheme 16). In the early stages of our attempts to obtain this product, the reaction did not go to completion as reported by Hanessian, and previous work from our group also afforded low yields. We optimized the synthesis by changing the order of addition from that reported by Hanessian where they added Ph3P to alcohol 82 in THF followed by the addition of DIAD at 0 °C. In our current work, in addition to the altered order of addition, we switched DIAD to DEAD and DEAD was added to Ph3P first at 0 °C for 30 minutes then alcohol 73 in THF was cannulated to the reaction mixture. Although this new order of addition provided us with a higher yield, the reaction did not go to complete conversion to fully protected β- pseudouridine 82. 74 was confirmed by NMR (Figure

38) signals for H-6 observed at 8.34 ppm as a singlet with a shift from 4.99 ppm for the

H-1'. Signals for all protons of the sugar were identified from 4.0 to 3.87 ppm. Moreover,

HRMS gave additional support (Figure 40).

Scheme 16: Synthesis of Compound 81 and 82

Subsequently, treatment of 82 with tetrabutylammonium fluoride (TBAF) in THF deprotect the 5´-silyl ether and furnished the desired C5´-alcohol 60 in 98% yield

(Scheme 13). Signals for the phenyl groups in the aromatic region between 7.66 to 7.40 ppm and the tert-butyl group disappeared (Scheme 41). High-resolution mass spectrometry further confirmed product formation (Figure 43).

To produce a C5´-pseudouridinyl radical precursor (Scheme 17), a C5´ pivaloyl substrate was chosen as a suitable compound. The pivaloyl moiety can undergo homolytic photocleavage via a Norrish Type I mechanism generating a radical at the desired C5´- position.

Scheme 17: Synthesis of C5'-Radical Precursor of Pseudouridine

Known approaches established by the Bryant-Friedrich group for the synthesis of the

5´uridinyl radical precursor were employed to install the same photolabile group at the 5´ positions of Ψ (Scheme 6).33 This route takes extra steps, but gives the desired radical precursor as reported by our group. Therefore, with some enhancements to the overall

scheme, we were able to obtain the final C5´pseudouridine radical precursor 13 in higher yields.

A synthetic approach was utilized focusing on the C5´position. The synthesis would begin with fully protected Ψ 82 that would be deprotected to yield alcohol 60. This alcohol would be converted to aldehyde 86 after oxidation with 2-iodoxybenzoic acid

(IBX). Oxidation of the 5´-hydroxyl of 60 to aldehyde 86 was carried out using IBX which was synthesized in house from iodobenzoic acid and potassium peroxymonosulfate

(Oxone). Studies show that the insolubility of IBX in traditional solvents can be overcome at elevated temperatures and also works as a solid-phase reagent for oxidation in most organic solvents.40 Hence, we employed this protocol to our synthesis and used

IBX in the presence of ACN at 70 0C, and improving the yield to 96% (Scheme 17).

Initially, isolating the pure aldehyde 86 was very problematic, and the purity of this compound was essential for the subsequent steps. However, through filtration of the reaction mixture with iced cold ACN, we were able to isolate the pure aldehyde 86 in

96% yield. Formation of 86 was confirmed by a strong signal for the aldehyde proton at

9.70 ppm in the NMR (Figure 45). When LCMS was carried out by dissolving the aldehyde 86in methanol, we also observed the formation of hemiacetal and diol in m/z

427 and 413 respectively (Figure 46).

Compound 86 was immediately treated with a catalytic amount of potassium cyanide and

18- Crown-6 followed by the addition of tert-butyldimethylsilyl cyanide to form protected cyanohydrin 9788 as a mixture of isomers (1:1) in 73% yield (Scheme 17). For the optimization of this step, potassium cyanide and 18- Crown-6 were co-evaporated three times with anhydrous dichloromethane (DCM) under strictly inert conditions. More 45

extended purging of the reaction flask with argon was employed, and very dry THF was used. The reaction mixture was stirred for 10 minutes before the addition of tert- butyldimethylsilyl cyanide followed by cannulation of 86 in anhydrous THF into the reaction flask. Using these strict synthetic protocols, we were able to attain high yields of the protected cyanohydrin. We also observed that compound 87 was very stable.

Compound 87 was subjected to alkylation with t-BuLi for 3 minutes forming unstable imine 88 which was immediately hydrolyzed by the addition of 1N HCl to reach pH 3 to afford fully protected ketone 89. We postulated that at lower pH, we would observe the deprotection of the acetonide and TBDMS protecting groups but that was not the case.

Deprotection of 89 with TBAF and hydrolysis of 90 provided the fully deprotected target radical precursor 13 (Scheme 18).

Scheme 18: Synthesis of C5´-Radical Precursor 13

3.3 Photochemical Generation and Study of the C5´-Pseudouridinyl radical.

Studies have utilized an approach to the study of RNA oxidation that include the generation of individual radicals at specific sites on the base or ribose of nucleosides using synthetic approaches to install distinct photolabile groups. Phenyl selenium or pivaloyl groups have been used because they can be photolyzed using UV light. Pivaloyl groups undergo a Norrish Type I photocleavage to generate the radical of interest.

Studies on RNA propose that, the C5´is a suitable position to abstract a hydrogen atom by a hydroxyl radical due to its solvent accessibility. It is essential to synthesize a C5´ - radical precursor as a tool for the studies of oxidative damage. The Bryant-Friedrich group previously synthesized and explored radical formation in RNA at the C5´-position of uridine using a pivaloyl containing radical precursor that undergoes Norrish type I photocleavage. Upon exposure to uv light at ≥ 320 nm. We have explored the mechanisms of formation of the C5´-radical derived products.33

A similar synthetic approach for pseudouridine as that used to install the t-butyl keto group to the 5´-carbon of uridine (Scheme 6) will be used followed by the photochemical generation of the C5´-radical in a site-specific manner.

3.4 Conclusion and Future Direction

The focus of this study was to understand oxidative damage to RNA through the studies of the C-glycoside analog pseudouridine 6. We aimed at generating a C5'-radical on the nucleoside and understand the fate and mechanism of the product derived. The synthetic strategy to obtain a C5'-pseudourinalradical precursor was successfully established. The radical precursor was subjected to a Norrish type I photocleavage, and the product obtained through the generation of the C5'-pseudourinalradical was analyzed by LCMS.

The C5'-pseudourinalradical precursor was photolyzed under anaerobic conditions in the presence of the biologically relevant hydrogen donor glutathione. In this study, the formation of the reduction product, pseudouridine (6) was observed in the presence of

glutathione. In the absence of glutathione, no product was identified. These results conclude that C5'- RNA radical pseudouridine, may lead to less damage than the C-N nucleosides by providing the protective role of pseudouridine towards RNA oxidative damage. However, further studies are essential before coming to any conclusion.

Moreso, 6 was successfully synthesized by altering the protecting group strategy previously used by other groups. This route was found to be beneficial for the later steps of the synthesis when introducing the protected cyanohydrin at the C5'-position. Some of these reactions need to be optimized for higher yields.

Finally, our study was done at the monomer level and provided insights on the fate of the radical and the mechanism of the radical reaction. However, to get the complete picture, incorporation of the radical precursor into RNA oligomers and study the fate of the radical under physiologically relevant conditions will give a better understanding of the oxidative damage to RNA under oxidative stress.

Chapter 4

4 Experimental Procedure Synthesis of C5´-Radical Precursor 13

4.1 Methods and Materials

Method

Unless stated otherwise, all organic reactions were carried out under inert standard laboratory conditions using magnetic stirring with oven-dried glassware. For oxygen or water sensitive reactions, reagents were cannulated through rubber septa via a syringe.

Material

All chemicals were commercially obtained and were of the highest grade and used without further purification unless otherwise specified. Thin layer chromatography was carried out on Silicycle® silica gel 60 F254 aluminum-backed plates. Product spots were visualized by UV light at 254 nm and/or staining of the TLC plate with permanganate dip made in house. All synthesized products were characterized by NMR using an Inova-600 or Avance-600 Bruker Avance III 600Mhz spectrometer in CDCl3 or CD3OD as the

solvent. Additional MS spectra were obtained using Shimazu LCMS 2020. All chemical shift values and, are reported in parts per million and coupling constants in Hertz.

Figure 12: Numbering System for Nucleosides

4.2 Synthesis of Pseudouridine

4.2.1 Synthesis of (3aR,6R,6aR)-6-(((tert-butyldiphenylsilyl) oxy)methyl)-2,2- dimethyldihydrofuro[3,4-d][1,3]dioxol-4(3aH)-on (62)

This procedure was modified from Chow et al1 and Hanessian et al2. D-ribonolactone

(66) (3.5 g, 23.7 mmol,) was co-evaporated with DCM and dissolved in acetone (150 ml)

and H2SO4 (1.5 ml, 1.0 eq) was added and stirred at 0°C to room temperature overnight.

The reaction was quenched with 6% NaHCO3 to basic pH and the product extracted with ethyl acetate to obtain the protected acetonide 83 in 94% yield. Compound 83 used in the next reaction without further purification

1 H NMR (600 MHz, CDCl3) δ 7.37 (d, J = 82.6 Hz, 4H), 7.27 (s, 23H), 7.09 (s, 2H), 4.55

(dd, J = 7.6, 4.8 Hz, 63H), 4.40 (dd, J = 47.1, 34.3 Hz, 3H), 4.13 (q, J = 7.1 Hz, 6H), 4.00

(ddd, J = 12.4, 5.6, 3.0 Hz, 33H), 3.84 (ddd, J = 12.4, 5.9, 2.7 Hz, 33H), 2.90 – 2.59 (m,

5H), 2.70 (s, 4H), 2.13 (d, J = 36.0 Hz, 39H), 2.05 (s, 7H), 1.77 (t, J = 17.6 Hz, 4H), 1.72

– 1.58 (m, 133H), 1.45 (d, J = 11.8 Hz, 126H), 1.44 – 1.19 (m, 75H).

Acetonide 83 (1.5 g, 7.95 mmol) was dissolved in anhydrous DCM (15.0 ml). To this was added imidazole (1.2 g, 17.5 mmol, 2.2 eq) followed by tert-butyl-diphenyl-silyl chloride (2.19 mL, 8.64 mmol, 1.1 eq) which was added dropwise to a stirred solution at

0°C. The reaction mixture stirred overnight, and the reaction was then quenched with 6%

NaHCO3, extracted with ethyl acetate. The organic extract was dried over Na2SO4, concentrated in vacuo and the isolated product purified on silica using a gradient of 5 –

15% EtOAc in hexanes to give 62 as a white solid in 96% yield.

1 H NMR (600 MHz, CDCl3) δ 8.58 (s, 2H), 8.43 (d, J = 105.5 Hz, 4H), 8.72 – 8.16 (m,

6H), 8.26 (dd, J = 203.5, 92.3 Hz, 8H), 8.72 – 7.83 (m, 10H), 8.72 – 7.78 (m, 11H), 8.72

– 7.73 (m, 12H), 8.72 – 7.54 (m, 54H), 8.72 – 7.55 (m, 53H), 8.72 – 7.32 (m, 115H), 8.72

– 7.08 (m, 140H), 8.72 – 7.22 (m, 138H), 8.72 – 7.29 (m, 116H), 5.21 (d, J = 10.5 Hz,

6H), 5.12 (d, J = 10.5 Hz, 5H), 4.53 (s, 5H), 4.48 (s, 6H), 4.26 (d, J = 10.5 Hz, 6H), 4.21

(s, 6H), 4.14 (s, 5H), 3.93 – 3.62 (m, 30H), 2.18 (s, 6H), 1.70 – 1.60 (m, 3H), 1.56 (d, J =

8.3 Hz, 26H), 1.53 – 0.95 (m, 181H), 1.08 (d, J = 16.4 Hz, 86H), 1.08 (d, J = 16.4 Hz,

87H), 1.03 – 0.95 (m, 3H), 1.04 – 0.83 (m, 12H), 0.26 – 0.04 (m, 5H).

4.2.2 Synthesis of (3aR,6R,6aR)-6-(((tert-butyldiphenylsilyl)oxy)methyl)-4-(2,4-di-tert- butoxypyrimidin-5-yl)- 2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-ol (80) 34

A 2.5 M solution of tert-butyllithium (16.1 mL, 40.3 mmol, 1.1 eq) in hexanes was added dropwise to a stirred solution of 5-bromo-2,4-di-tert-butoxypyrimidine 85 (11.1 g, 36.6 mmol, 2.0 eq) in anhydrous THF (200 mL) at -78 °C under argon over 30 min.

Compound 80 (7.80 g, 18.3 mmol, 1 eq) in THF ( 100 mL) at -78 °C was cannulated into the reaction mixture with continued stirring at -78 °C for 2 h. The mixture was quenched with brine (100 mL) and allowed to warm slowly to room temperature over 4 h. The product was extracted with ethyl acetate (3 × 30 mL). The combined organic phases were dried over Na2SO4 and concentrated. The product was purified by column chromatography 5-10% using EtOAc in hexanes to deliver 80 white foam as a mixture of two anomers (α β) 9:1 in 85% yield.

1 H NMR (CDCl3): δ 8.51 (1H, s), 8.42 (1H, s), 7.74 (3H, m), 7.66 (3H, m), 7.61 (2H, m),

7.40 (12H, m), 4.93 (2H, m), 4.86 (1H, dd, J = 5.7 Hz, J = 1.3 Hz), 4.74 (1H, m), 4.52

(1H, m), 4.36 (2H, m), 3.90 (1H, m), 3.80 (1H, d, J = 3.9 Hz), 3.78 (1H, m), 1.66 (9H, s),

1.63 (3H, s), 1.62 (9H, s), 1.61 (3H, s), 1.28 (3H, s), 1.23 (3H, s), 1.11 (12H, s), 1.05 (3H, s), 1.01 (3H, s).

4.2.3 Synthesis of 2-((tert-butyldiphenylsilyl)oxy)-1-((4S,5R)-5-((R)-(2,4-di-tert- butoxypyrimidin-5-yl)(hydroxy)methyl)-2,2-dimethyl-1,3-dioxolan-4-yl)ethan-1-ol (81) 34

A solution of ZnCl2 1.0 M (12.3 mL, 12.3 mmol, 1.5 eq) in ether was added dropwise to a stirred solution of 80 (5.33 g, 8.2 mmol, 1.0 eq) in anhydrous DCM (500 mL) at - 78 °C over 30 min. A 1.0 M solution of L-selectride (28.7 mL, 28.7 mmol, 3.5 eq) in THF was added dropwise over 30 min at the same temperature. The mixture was allowed to warm slowly to room temperature and stirred overnight. EtOH (7 mL), water (7 mL), 30%

H2O2 (7 mL), and 6 N NaOH (7 mL) were added to quench the reaction. The product was extracted into EtOAc and the combined organic phases were washed with brine and dried

over MgSO4. Chromatography using a gradient of 10 – 35% EtOAc in hexanes delivered

81 as a white foam in 92% yield.

1 H NMR (600 MHz, CDCl3) δ 8.28 (1H, s), 7.7-7.68 (4H, m), 7.46 – 7.40 (6H, m), 5.25

(1H, d, J = 6.9 Hz), 4.34 (1H, dd, J = 6.2, 1.5 Hz), 4.26 – 4.20 (2H, m), 3.91 (1H, dd, J =

10.3, 3.0 Hz), 3.84 (1H, dd, J = 10.4, 5.5 Hz), 3.06 (1H, d, OH), 2.85 (1H, d, OH), 1.61

(9H, s), 1.60 (9H, s), 1.50 (3H, s), 1.33 (3H, s), 1.09 (9H, s).

4.2.4 Synthesis of 2,4-di-tert-butoxy-5-((3aS,4S,6R,6aR)-6-(((tert- butyldiphenylsilyl)oxy)methyl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4- yl)pyrimidine (82) 34

To a stirred solution of PPh3 (2.48 g, 9.5 mmol, 2.0 eq) in dry THF (190 ml ), was added

DEAD (1.87 mL, 9.5 mmol, 2.0 eq) at 0 °C. Compound 81 (3.1 g, 4.75 mmol, 1.0 eq ) was cannulated into the reaction flask and the mixture allowed to warm to room temperature slowly and left to stir overnight. Upon completion, the solvent was removed under reduced pressure and purified by column chromatography using a gradient of 15-

35% EtOAc in hexanes to deliver 82 as a white foam in 85% yield.

1H NMR (600 MHz, CDCl3) δ 8.34 (1H, s), 7.74 (4H, m), 7.44 (6H, m), 4.99 (1H, d, J =

4.4 Hz), 4.70 (1H, dd, J = 6.6, 4.7 Hz), 4.61 (1H, dd, J = 6.6, 4.5 Hz), 4.16 (1H, q, J = 4.4

Hz), 3.94 (1H, dd, J = 11.2, 3.8 Hz), 3.87 (1H, dd, J = 11.3, 4.6 Hz), 1.66 (9H, s), 1.66

(9H, s), 1.63 (3H, s), 1.39 (3H, s), 1.10 (9H, s).

4.3 Synthesis of C5´- Pseudouridinylradical Precursor

4.3.1 Synthesis of ((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)methanol (60) 34

Compound 82 (0.670 g, 1.06 mmol) in anhydrous THF (20 ml) was treated with TBAF

(1.0 M solution,1.60 mL, 1.59 mmol, 1.5 eq) at 0 °C. The mixture was allowed to slowly warm up to room temperature and stirred overnight. The solvent was concentrated under reduced pressure and the product purified by column chromatography on silica gel (5-25

% EtOAc/Hex) to deliver 60 as a white foam in 98% yield.

1 H NMR (600 MHz, CDCl3): δ 8.16 (1H, s), 4.81 (2H, d, J = 4.2 Hz), 4.76 (1H, m), 4.73

(1H, m), 4.04 (1H, m), 3.89 (1H, dd, J1 = 11.8 Hz, J2 = 3.1 Hz), 3.76 (1H, m), 2.36 (1H, s,

OH), 1.64 (9H, s), 1.60 (9H, s), 1.58 (3H, s), 1.34 (3H, s).

4.3.2 Synthesis of (3aS,4S,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxole-4-carbaldehyde (86)

Compound 60 (2.5 g, 6.31 mmol) in ACN (40 ml) was treated with IBX (1.17 g, 6.31 mmol ) and the suspension stirred at 70 °C for 3 h under reflux. The mixture was allowed to cool to room temperature over 2 h and filtered. The residue was washed with iced cold

ACN. The filtrate was concentrated under reduced pressure to deliver 86 as a white foam in 94% yield.

1H NMR (CDCl3): δ 9.80 (1H, s), 8.18 (1H, s), 5.03 (1H, d, J = 2.9 Hz), 4.95 (1H, m),

4.83 (1H, m), 4.40 (1H, d, J = 3.9 Hz), 1.62 (9H, s), 1.60 (9H, s), 1.60 (3H, s), 1.35 (3H, s).

4.3.3 Synthesis of 2-((tert-butyldimethylsilyl)oxy)-2-((3aR,4S,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)acetonitrile (87)

Catalytic amounts of 18-Crown-6 (12.9 mg, 48.6 mm, 0.02 eq) and KCN (31.6 mg ,

48.6 mm, 0.20 eq ) were co-evaporated three times with anhydrous DCM under inert conditions. Aldehyde 86 (96 mg, 2.4 mmol) was dissolved in dry THF (8ml) and cannulated to the mixture. To the solution, was added TBDMSCN (80 mg, 0.57 mmol) dropwise at room temperature and the solution stirred for 27 hours. The reaction was quenched with brine 9 ml and extracted with EtOAc (3 × 6 ml). The combined organic phases were dried over MgSO4, and solvent the concentrated. Column chromatography

(0-5 % EtoAc/Hex) provided pure 87 in 73% as a white foam.

1 H NMR (CDCl3): δ 8.29 (1H, s), 8.21 (1H, s), 5.01 (1H, d, J = 4.6 Hz), 4.97 (1H, d, J =

4.6 Hz), 4.75 (2H, m), 4.67 (1H, m), 4.65 (1H, m), 4.63 (1H, m), 4.60 (1H, dd, J = 6.6

Hz, J = 4.6 Hz), 4.15 (2H, m), 1.64 (18H, s), 1.61 (18H, s), 1.60 (6H, s), 1.59 (3H,s), 1.37

(6H, s), 0.92 (9H, s), 0.91 (9H, s), 0.24 (3H, s), 0.22 (3H, s), 0.19 (3H, s), 0.18 (3H, s).

4.3.4 (R)-1-((tert-butyldimethylsilyl)oxy)-1-((3aR,4S,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)-3,3- dimethylbutan-2-imine (88)

Cyanohydrin 87 (0.70 g, 1.31 mmol) was co-evaporated with toluene and dry DCM. The compound was dissolved in anhydrous THF (9.2 ml) and the mixture cooled to -78 °C.

Tert-BuLi (0.12 ml, 1.31 mmol) was added dropwise at -78 °C and the reaction stirred for

4 min. The reaction was quenched with H2O (3ml). The reaction mixture was allowed to warm to room temperature for 30 min and the product was extracted with EtOAc (5 × 5 ml). The combined organic layers were dried over MgSO4 and concentrated under reduced pressure to give the crude product as a brown foam. Without further purification, the crude imine was taken immediately to the next reaction.

4.3.5 (S)-1-((tert-butyldimethylsilyl)oxy)-1-((3aR,4S,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)-3,3- dimethylbutan-2-one (89)

Compound 88 was dissolved in ACN (6 ml), 3 mL of water and 5.0 ml of ACN/H2O/2N

HCl [66:33:1]. After the reaction mixture was refluxed to at 60 °C for one hour. It was allowed to cool and was diluted with water and extracted with EtOAc (4×5 ml). The combined and organic extracts were washed with brine, dried over MgSO4 and concentrated under reduced pressure. Column chromatography with 5-15 % EtoAc/Hex afford 89 in 68% yield.

1H NMR (600 MHz, CDCl3) δ 7.86 (dd, J = 5.9, 1.5 Hz, 1H), 5.03 – 4.70 (m, 6H), 4.69-

4.59 (M, 2H), 4.22 (q, J = 4.0, 3.6 Hz, 1H), 4.15 (q, J = 7.2 Hz, 1H), 3.48 (dd, J = 3.7,

2.2 Hz, 1H), 1.39 (s, 6 H) 1.37 (s 5H), 1.33 (s, 2H), 1.28 – 1.24 (m, 7H), 1.23 – 1.19 (m,

6H), 0.95 – 0.90 (m, 11H), 0.88 (s, 5H).

4.3.6 (S)-1-((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)-1-hydroxy-3,3-dimethylbutan-2-one (90)

Compound 99 (6.4 g, 10.76 mmol 1.0 eq) was dissolved in dry THF (4.0ml) and cooled to 0 °C. TBAF (4.33 ml, 16.14 mmol, 1M in THF, 1.5eq) was added dropwise to the reaction flask. The mixture was allowed to warm to room temperature and stir overnight.

The reaction mixture was concentrated, and the product was purified by column chromatography using 10-40 % EtoAc/Hex to provide 90 in 97% yield.

1 H NMR (600 MHz, D2O) δ 7.53 (2, 2H), 5.12 (s, 1H), 4.88 (d, J =1.9 Hz, 2H), 4.83 –

4.69 (m, 2H), 4.58 (d, J = 3.0 Hz, 1H), 4.43 – 4.24 (m, 1H), 4.21 – 4.03 (m, 3H), 1.40 –

1.33 (m, 4H), 1.29 – 1.21 (m, 18H), 0.89 – 0.83 (m, 9H).

4.3.7 5-((2S,3R,4S,5S)-3,4-dihydroxy-5-((S)-1-hydroxy-3,3-dimethyl-2- oxobutyl)tetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione (13)

Compound 90 (0.30 g, .625 mmol) was dissolved in 3.5 ml of 70% acetic acid (v/v). The reaction mixture was heated to 50 °C and refluxed for 5 h. The mixture was allowed to cool and concentrated in vacuo. The product was purified by column chromatography on silica gel (0-2% MeOH, EtOAc) to give 13 as a white solid 13 mixture of two isomers in a ratio 4:1 in 94% yield.

1H NMR (600 MHz, MeOD): δ 7.57 (1H, d, J=0.90 Hz), 7.56 (1H, d, J=0.54 Hz), 4.75

(1H, d, J=3.90 Hz), 4.69 (1H, d, J=1.80 Hz), 4.54 (1H, dd, J=5.22, 0.90 Hz), 4.50 (1H, dd, J=7.08, 0.54 Hz), 4.39 (1H, dd, J=4.56, 1.80 Hz), 4.27 (1H, dd, J=7.08, 5.28 Hz), 4.24

1 ( H, dd, J=3.90, 3.11 Hz), 4.23-4.18 (3H, m)1.23 (9H, s), 1.21 (9H, s). 13C NMR (600

MHz, MeOD): δ 216.00, 215.35, 166.00, 165.86, 153.26,152.46, 142.41,

141.69, 112.30, 111.61, 86.07, 84.86, 81.67, 80.82, 75.11, 75.00, 74.54, 73.57, 73.30,

71.72, 44.76, 44.20, 27.15, 26.67

4.4 Photochemical Generation of the C5´- Pseudouridinylradical

There is an emerging need to understand the damage caused along with the associated mechanisms to nucleosides through their exposure to ROS. Research groups, including ours, focus on understanding mechanisms related to DNA oxidative damage. However,

RNA has had very little attention. Hence, our group is also interested in studying the damage caused to RNA through the generation of radicals at the C5´ position. In this section, our focus is to understand the fate and mechanism of formation of the products obtained through the generation of the C5´-pseudouridynyl radical.

We aimed at generating a C5´- on our modified nucleoside to understand the fate of the radical formed and the mechanisms of the radical derived products. We successfully optimized the synthetic strategy to obtain Pseudouridine and a novel C5´-pseudouridinyl radical precursor. Photolysis under anaerobic conditions was performed by subjecting the radical precursor to a Norrish type 1 Photocleavage. The products formed were identified using Shimadzu LCMS. The UV radiation on the uridine radical precursor resulted in base elimination in a significant amount. Of great importance, the break at the glycosyl bond as seen in the case of uridine did not occur in pseudouridine because of the presence of the stronger C-C bond.

At a molecular level, monomer studies on the fate and mechanism of the radical reaction could provide some insight on the level of oxidative damage to RNA, but a better understanding can be attained by introducing the radical precursor into an oligomer that represents RNA. Furthermore, in the oligomer studies, physiological conditions should be employed during the radiation to mimic the biological system — this way the level of oxidative insults to RNA under oxidative stress will be understood better.

To begin our investigations on the oxidative insults to substrate 13, initial investigations were carried out on Ψ to study the fragmentation patterns of this C-C nucleoside to establish the identities of Ψ-related ions. The preliminary results and analysis were carried with an independently synthesized pseudouridine and was not further purified.

Inten. (x1,000,000) 245.80

2.25

2.00

1.75

1.50

1.25

1.00

477.05 0.75

0.50 231.95 342.00 455.15 703.85 0.25 284.05 151.30 387.00 682.30

0.00 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 m/z

Figure 13: Mass spectral analysis of pseudouridine 6 (Independently synthesized)

Mass spectra were recorded on an LCMS 8050 mass spectrometer, using a direct insertion probe. High-resolution mass measurements were performed by reference to fragment ions from Edward Dudley (Figure 14).

Figure 14: Fragmentation patterns of Pseudouridine by Edward Dudley [Reprinted with Permission]

The spectrum of pseudouridine (Figure 14) was dominated by an intense peak at m/z

209. The loss of H2O gives an ion of m/z 226 and undergoes further fragmentation, losing an extra H2O in succession for the m/z 209. Another fragment ion was generated at m/z

179, which signifies the loss of two H2O plus CH2O. Fragment ion at m/z 125 which consists of the uracil moiety plus a methylene group derived from the ribose was also observed.

Figure 15: Fragmentation patterns of the Mass spectral analysis of independently synthesized pseudouridine 6.

The sugar nor the base moiety did not experience fragmentation independently. This feature is predicted to be general for the complete stability of the pseudouridine C-C glycosidic bond. According to Gould, 1959, the average C-N bond energy is 62 kcal/mole, and the C-C bond averages approximately 80 kcal/mole. This difference would allow higher activation energy for cleavage of the stronger C-C bond.

Inten. (x100,000) 9.0 209.00 8.5

8.0

7.5

7.0

6.5

6.0

5.5

5.0

4.5

4.0

3.5 3.0 155.00 2.5 179.00 2.0

1.5

1.0

0.5

0.0 155.0 157.5 160.0 162.5 165.0 167.5 170.0 172.5 175.0 177.5 180.0 182.5 185.0 187.5 190.0 192.5 195.0 197.5 200.0 202.5 205.0 207.5 m/z

Figure 16: Fragmentation patterns of the Mass spectral analysis of independently synthesized pseudouridine 6.

More importantly, is the fact that cleavage of a C-N glycosidic bond generates a pyrimidine fragment, an unpaired electron, and positive charge are delocalized over the entire ring, while the corresponding C-C cleavage would generate a vinyl radical or cation. We argue that the latter structure is incapable of resonance stabilization, and consequently much more difficult to generate. Resonance stabilization also accounts for the reasonable formation of pseudouridine fragment ions at m/z 209 and 125 ( Figure 15 and Figure 16) . The mechanism of this loss is unknown, and other likely fragmentations may offer different pathways. Thus, this is proposed as a short-term pathway which would require further detailed analysis. These considerations should apply to the mass spectra of C-C nucleosides in general, and strongly suggest that the mass spectra can 67

provide convincing evidence for the existence of a C-C glycosidic bond. To gain further insight into the nature of the radical precursor and understanding the C5ʹ-radical chemistry, radical precursor 13 was subjected to photolysis in the presence of other hydrogen atom donors.

Photolysis was carried out in 4.5 cm × 10 mm quartz cuvettes using an Oriel 500 W high- pressure mercury arc lamp fitted with an IR filter, focusing lens and cut off filter. All photolysis experiments were carried out at ≥ 320 nm and the temperature maintained between 15-25 ºC using a Peltier PTP-1 single cell temperature control system.

Photolysis experiments were carried out at 15 min, 30 min, 45 min and 60 min and the resulting photolysate were directly injected into an LCMS for analysis.

Table 1: LCMS conditions used for the photolysis of 13

Column Hypersil Gold C-18 (50 mm L x 2.1 mm I.D.; 3 µm

Mobile Phase A: 0.1% Acetic Acid in Water B: 15% Acetonitrile

Gradient Program %B 0-9 min 15%; 9-20 min 15%; 20-30 min 100%; 30-40 min 15%

Flow Rate 0.1mL/min

Oven Temperature 21 °C

Injection Volume 1µL

MS interface Electrospray Ionization (ESI)

MS temperature De-solvation line 250 °C, Heating block 399 °C

Figure 17: Chromatogram for the photolysate of 6 (10 mM) at 15 min, 30min, and 45min.

Photolysis was carried out on 6 at 15, 30, and 45 min (Figure 17) in the absence of a hydrogen donor. This resulted in the degradation of pseudouridine from 15 min to 45 min. However, no other product formation was observed.

uV 4000000

3750000

3500000

3250000

3000000

2750000

2500000

2250000

2000000

1750000

1500000

1250000

1000000

750000

500000

250000

0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 min

Figure 18: Chromatogram for the photolysate of 6 (10 mM) at 60 min with/without GSH.

It was noticed that 6 was completely consumed in 60 min without GSH (Figure 18).

However, when the experiment was done in the presence of GSH at 60 min, 6 was still observed. Hence, 60 min was applied to the rest of the photolysis experiments. These photolysis experiments were carried out under anaerobic conditions using an CH3CN:

H2O (1:1) solvent system.

Table 2: Mass balances of products from photolysis of 6 in the absence of GSH

Photolysis (min) Pseudouridine (%)

15 65.5

30 96.9

45 0.86

Table 3: Mass balance from the photolysis of compound 13 in the presence of GSH (4 eq)

Photolysis (min) Pseudouridine % Yield % Remaining of precursor 13 Mass Balance

15 21 71.2 92.3

30 22.5 7.7 30.2

40 35.5 3.9 39.4

60 39.5 2.8 38.3

Photolysis of 13 was performed using glutathione concentrations of 0.8 nmol/µL of GSH under anaerobic conditions. Formation of 6 was confirmed by mass spectroscopy and the yield of 6 slowly increased. The maximum yield of 6 (39.5%) was observed with 0.8 nmol/µL glutathione. Mass balances for the formation of this product was calculated based on calibration curves as seen in tables 3 and 4.

Figure 19: Calibration curve of compound 6

Figure 20: Calibration curve of compound 13

These results were unexpected as it was assumed that the photolysis of 13 would result in the formation of 6 in greater yields. Surprisingly, the reaction did not go to completion.

Figure 21: Photolysis of Radical Precursor 13

It was noticed that radical precursor 13 was not completely consumed after 60 min, and no base release products were observed at the wavelength of detection 320 nm (Figure

19). An intense peak was observed at a retention time of 4.5 min. This retention time matched the retention time of the expected product, pseudouridine. This indicates the necessity to explore the behavior of the radical precursor further and determine efficient conditions for the complete conversion of the radical precursor 13 to pseudouridine (6).

Radical Precursor Conversion 7000000

6000000

5000000

4000000 AUC 3000000

2000000

1000000

0 0 15 30 60 Photolysis Time

Figure 22: Radical precursor 13 conversion upon photolysis in the presence of GSH ( 2mM) in Ammonium Acetate buffer ( 5mM) at pH 7.0.

Photolysis of 13 (7 nmol) was performed using 2 mM of glutathione in ammonium acetate buffer (5mM) at pH 7.0 under anaerobic conditions. Gradual convention of 13 and the formation of 6 were observed under these conditions (Figure 22 and Figure 23).

The mass balance (Table 4) for the formation of product 6 was calculated based on the calibration curve. The yield of 6 increased with increasing photolysis time, and the maximum yield of 6 (70.26%) was observed at 60 min. These results were expected as it was assumed that only the formation of 6 would be observed. Further experiments will be done as a part of future work to investigate conditions more suitable for the complete conversion of the radical precursor 13 to pseudouridine (6).

Radical Precursor Conversion and Pseudouriine Formation 120

100

60 Percentage 40

0 0 15 30 60 Photolysis Time

Radical Precursor Pseudouridine

Figure 23: Formation of pseudouridine (6) upon photolysis of 13 in the presence of GSH ( 2mM) in Ammonium Acetate buffer ( 5mM) at pH 7.0

Photolysis % Radical Precursor % Pseudouridine Total Remaining Formation 15 min 70.73 28.06 99.01

30 min 51.07 50.91 97.67

60 min 26.51 70.26 96.69

Table 4: Mass balance from the photolysis of compound 13 (7 nmol) in the presence of GSH ( 2mM) GSH, in Ammonium Acetate buffer (5mM) pH 7.0

Chapter 5

5 Pseudouridimycin and its analogs

Naturally occurring modifications of nucleic acids are fairly common in all organisms.

Those modifications often are the results of evolutionary requirements for the survival of the organisms. Several of these modifications are seen only in bacteria.41 In this study, a new pseudouridine analog modified at the 5ʹ-carbon will be designed and synthesized.

Our approach will consist in modifying a ribose sugar followed by coupling with the peptide portion. The design of the nucleoside analog inhibitor (NAIs) will be based on the structure of pseudouridimycin (PUM) 101 (Figure 20).41,42 Successful synthesis of the target analog using this approach will allow for the introduction of additional pseudouridimycin analogs with modified sugars.

Figure 24: Structure of Pseudouridimycin (PUM)

5.1 Discovery of Pseudouridimycin

As resistance to antibiotics continues to present a serious threat to human health the number of antibiotics in the pipeline has significantly decreased overtime. Bacteria are prokaryotic microorganisms representing one of the three domains of life. Fewer than 1% of bacteria cause disease in humans.42 Bacterial RNA polymerases, the enzymes that mediate bacterial RNA synthesis, have been proven to be a target for broad-spectrum antibacterial therapy. Pseudouridimycin (PUM) 91 ( Figure 22) a novel natural product from microbial extracts consisting of a formamidinylated, N-hydroxylated Gly-Gln dipeptide conjugated to 6'-amino-pseudouridine, has shown potent inhibitory properties towards bacterial RNAP in vitro, bacterial growth in culture, and cleared a Streptococcus pyogenes peritonitis infection in a mouse model.43,44

From 1950s and 1960s, a prolific number of antibiotics were discovered that are still in clinical use today. With the advances in screening processes and tools, several new compounds have been were screened for anti-infective activity with a relative amount of success. With clinical benefits in mind, newly discovered compounds are expected to possess several advantages over the antibiotics currently in clinical use: this implies a clinically relevant antimicrobial spectrum, low resistance potential and other factors that could influence the drug overall pharmacology such as molecular weight range, solubility, and potential preferred route of administration. Maffioli et al.41 screened a library of over 3000 actinobacteria and extracts from fungal culture for the inhibition of ribonucleic acid polymerase (RNAP). The samples were isolated and purified using techniques best known to the art to produce active strains. The strains that produced the hit extracts were strains IDI38640 and IDI38673. Strains ID38640 and ID38673 were 77

Actinobacterial isolated from soil samples collected in Italy and France, respectively.41

These two hit strains exhibited cell morphologies that were consistent with the genus

Streptomyces and exhibit 16S rRNA gene sequences determined as described in

GenBank with the accession numbers of JQ929050 and JQ92905. The gene sequence of both strains was 99.9% identical over 1.4 kB to each other and highly similar to clusters of closely-related Streptomyces species; S. nigrescens, S. tubercidicus, S. rimosus subsp. rimosus, S. hygroscopicus subsp. angustmyceticus and S. libani subsp. libani.41 Structure elucidation of these two active strains was done by using an Ion-trap ESI-MS which showed a protonated molecular ion at m/z 487 [M+H]+ and a dimerized ion at 973

[2M+H]+. High-resolution mass spectroscopy (HSMS) showed an exact mass of

487.18865, consistent with the molecular formula C17H26N8O9. Reversed-phase HPLC showed a single peak with a retention time of 12 minutes. The UV-absorbance spectrum showed maxima at 200 nm and 262 nm, consistent with the presence of a pyrimidine moiety. The 1H NMR spectrum revealed one olefinic (H6), five amides, four methylene, and five methine signals. The 2D 1H–13CHSQC and -HMBC NMR spectra identified five carboxyl-amide groups, two olefinic carbons, four methine carbons belonging to a sugar ring, one other methine carbon, and four methylene groups. The COSY NMR spectrum identified correlations between the five sugar protons. The chemical shift of C5' and ribose 15N-HSQC correlations between H6' and H5' indicated the presence of 6'-amino- ribose. 13C-NMR spectra and COSY indicated the presence of a glutaminyl moiety.

NOESY NMR spectrum indicated the presence of glycine C-linked to glutamine and indicated the presence of formamidine C-linked to glycine. The stereochemistry of the glutamine residue was determined to be (L), and the ribose sugar was inferred to be D by 78

similarities to the natural occurring pseudouridine.41 The structure elucidation of these two active components by mass spectrometry and multidimensional NMR spectrometry revealed that, the extracts had the same novel active constituent as pseudouridimycin.

The two extracts inhibited bacterial ribonucleic acid polymerase from E. coli RNAP by

≥80%. However, these extracts did not inhibit a structurally distinct bacteriophage

RNAP and have not been previously characterized as an inhibitor of bacterial RNAP. 41

Pseudouridinmycin (PUM) was shown to selectively inhibit bacterial RNAP with an IC50 of 0.1 μM and the selectivity was >4- to >500-fold which means PUM is selective to the bacterial RNAP and not the host. PUM also selectively inhibited bacterial growth with an

IC50 of 2 to 16 μM and selectivity of >6- to >60-fold. Interestingly, the infection was cleared by PUM in Streptococcus pyogenes peritonitis mouse model in vivo. These results correspond to the typical pattern of inhibition of RNA-dependent RNAP synthesis

45–51: that is, slight to no inhibition of DNA synthesis, fast and robust inhibition of RNA synthesis and weaker inhibition of protein synthesis (Figure 25).

Figure 25: Inhibition of RNAP-Dependent RNA Synthesis.47 [Reprinted with Permission]

Studies of antibacterial activity and RNAP-inhibition, PUM showed antibacterial activity in vivo against Gram-positive bacteria, Gram-negative bacteria, drug-sensitive bacteria strains, and drug-resistant bacterial strains. These drug-sensitive and drug-resistant bacterial strains were tested against rifamycin-, fluoroquinolone-, β-lactam-, aminoglycoside-, macrolide-, tetracycline-, lincosamide-, chloramphenicol-, trimethoprim-, oxazolidinone-, glycopeptide-, mupirocin-, and lipopeptide-. 41 This study did not observe any cross-resistance with the typical RNAP inhibitor rifampin (Rif) when

PUM was co-administered. Interestingly, additive antibacterial activity was shown and displayed spontaneous resistance rates an order of magnitude lower than those of Rif

(Table 5).

Table 5: Spontaneous Mutation Rates of PUM and Rif41 [Reprinted with Permission]

Inhibition Concentration Resistance Rate (per generation) (95% confidence interval) PUM 8xMIC 0.4x10-9 (0.06-0.9 x 10-9) 16xMIC 0.5x10-9 (0.1-1 x 10-9) Rif 8xMIC 3x10-9 (1.6x10-9) 16xMIC 4x10-9 (1.7 x 10-9)

This signifies that PUM inhibits RNAP through a binding site mechanism different from

Rif. Further studies of the target of transcription inhibition by gene sequencing indicated that PUM-resistant mutants contain mutations in the rpoB gene which encodes for the

RNAP β subunit or the rpoC gene that encodes for the RNAP β′ subunit. These results confirmed that RNAP is the functional cellular target of PUM (Figure 25 A) which is located within the RNAP active-center and overlaps with the RNAP active-site; the i+1

NTP binding site (Figure 25 A), suggesting that inhibition of RNAP by PUM occurs by mimicking NTP and interfering with the incorporation of UTP at the active site; the i+1.

PUM target is not the same as that of rif and does not overlap with the Rifampin target site (Figure 25 B ). 52–54

Figure 26: Target of PUM in the RNA NTP Addition Site. [ Reprinted with Permission]

This is consistent and accounts for the observation that PUM does not share cross- resistance with Rifampin (Figure 24) and does not overlap with the targets of other

RNAP inhibitors such as lipiarmycin (Lpm), 43,55streptolydigin (Stl)56,57and myxopyronin

(Myx)55,58(Figure 26 B).

The observation that PUM is a nucleoside analog inhibitor with same Watson-Crick base- pairing similar to uridine triphosphate (UTP) (Figure 23) and the observation that the target of PUM overlaps with the RNAP i+1 nucleoside triphosphate ( NTP) binding site

suggest that PUM functions as an NAI by competing with UTP for occupancy of the

RNAP i+1 NTP active site (Figure 26). As mentioned earlier transcription is the process where RNA is synthesized from its template DNA by the enzyme RNA polymerase

(RNAP). The three main steps in the transcription cycle are initiation, elongation, and termination. During initiation, the core RNAP enzyme binds initiation factors. The resulting holoenzyme can bind specifically to the promoter DNA, forming the closed complex. The melted DNA allow the active-site access to the template strand.

Transcription ends when RNAP reaches an intrinsic termination signal, characterized by an RNA hairpin in the nascent transcript.

The i and i + 1 active site of the bacteria RNAP has Mg2+ ions that aids to catalyze phosphodiester bond formation between the 3 prime ends of the nascent transcript and an incoming NTP. Nucleoside analog inhibitors interact with bacterial RNAP through the nucleotide-binding site, the "i+1" nucleotide triphosphate (NTP) binding site and competes with uracil triphosphate (UTP) for occupancy of the "i+1" NTP binding site.

2+ Two Mg ions that chelate the active center are thought to catalyze an SN2 nucleophilic attack of the 3' OH group on the end of the RNA transcript on the alpha phosphate of the incoming NTP by stabilizing the transition state. The newly formed 3' end is positioned in the i + 1 site after the phosphodiester bond formation occurs. A molecule of pyrophosphate (PPi) is produced and for a new round of NTP addition to occur; the PPi must be released from the active center. Nucleoside analog inhibitors can disrupt this catalytic cycle.

Figure 27: Target of PUM in the RNA NTP Addition Site. [ Reprinted with Permission]

Thus, it was concluded that PUM functions as an NAI that competes with UTP at positions that direct incorporation of uridine and not at positions that direct incorporation of guanosine, adenosine.47

To further define the mechanism of transcription inhibition by pseudouridimycin, a crystal structure of a transcription initiation complex containing pseudouridimycin; RPo-

GpA-PUM (Figure 28A ) and, for comparison, a crystal structure of a corresponding transcription initiation complex containing uridine triphosphate UTP ( Figure 28B ) was studied.

Figure 28: Structural basis of Transcription Inhibition by PUM.41 [Reprinted with Permission]

This further concludes that PUM is indeed a nucleoside analog NAI that competes for occupancy of the i+1 active site and binds to this site. The Ψ base of PUM forms

Watson-Crick hydrogen bonds with the DNA template-strand in a fashion corresponding to the base of NTP. The sugar moiety of PUM makes similar interactions within the i+1 active site when compared to the sugar moiety of NTP. PUM’s glutamine moiety mimics the interactions made by the NTP triphosphate; and finally, the guanidinyl moieties and

N-hydroxy moiety interact with the RNA nucleotide base-pair at the previous template position- the RNA 3'-nucleotide.47 The N-hydroxy donates a hydrogen bond to the 3'-OH of the RNA 3'-nucleotide, and the guanidinyl moiety donates one hydrogen bond to the

5´-phosphate of the RNA 3'-nucleotide and another to the base of the RNA 3'-nucleotide

(Figure 28A).

The Watson-Crick pairing observed in the PUM RNAP structure confers absolute specificity for a position directing incorporation of U and preceding position directing incorporation of G, A, or C each of which contains an H-bond acceptor at the proper 84

position (Figure 26 A). The structure also explains the selectivity of transcription inhibition by PUM. All RNAP residues contacted by PUM are highly conserved across

Gram-positive and Gram-negative bacterial RNAP, accounting for the inhibition of both

Gram-positive and Gram-negative bacterial RNAP.

The discovery of this new class of antibiotic from conventional microbial extract screening shows that contrary to the general belief, routine microbial extract screening has not been exhausted as a source of new antibacterial lead compounds.

5.2 Derivatives of Pseudouridimycin

With the discovery of PUM, Maffioli and colleagues wanted to reconfirm the structure of

PUM and understand the structure-activity relationships (SAR). Hence, a semi-synthesis of PUM was performed to corroborate the actual structure of PUM and to provide a synthetic approach to prepare novel derivatives and a better understanding of SAR.

The semi-synthesis of these derivatives started with 1 mg PUM (Scheme 25), which was reacted with TiCl3 in 1 M sodium acetate (NaOAc) (pH 7.0) for 2 h at room temperature to reduce the N-hydroxy moiety, producing a desoxy-PUM 92. 91 was also reacted with

PdCl2 in a 1: 1 acetonitrile: water (ACN: H2O) for 2 h at room temperature which resulted in dehydration of the glutamine sidechain to nitrile 93.

Scheme 19: Semi-synthesis of PUM Derivatives47

Hydrolysis of 91 with 0.1% trifluoro acidic acid (TFA) in H2O for 3 days at room temperature resulted in the carboxy analog 94. A reaction of 94 with benzylamine

(BnNH2), in dimethylformamide (DMF) containing benzotriazole-1-yl- oxytripyrrolidinophosphonium hexafluorophosphate (PyBOP) for 30 min at room temperature resulted in the benzylamide analog 95. Compound 91 was subjected to 2,3-

butanedione in 10 mM NH4OAc (pH 8.0) for 30 min at room temperature and resulted in diol intermediate 96, which was subsequently trapped by treatment with phenylboronic acid (PhB(OH)2), for 2 h at room temperature, giving a phenyl-dioxaborolane analog 97.

Here 95 lacked the PUM N-hydroxy group but had the most activity. Compound 93, 94 and 95 containing alterations at the glutamine sidechain displayed no activity and 96 and

97 with modifications at the guanidinyl sidechain also displayed no activity.

The structure of 92 enables the structure-based design of novel PUM analogs with improved potency and selectivity. These initial lead-optimization efforts corroborated the importance of the PUM N-hydroxy, glutamine, and guanidinyl moieties and demonstrated that the PUM glutamine C(O)NH2 could be replaced by C(O)NHR while retaining RNAP inhibitory and antibacterial activity (Figure 28).

Figure 29: Structure-Activity Relation of PUM47

Structure activity relationship studies indicate that, the 2´-hydroxyl and 3ʹ-hydroxyl of the

NAI ribose is crucial for activity, along with the glutamine and guanidinyl moieties. The base of the NAI makes Watson-Crick hydrogen bonds with the DNA template-strand in the A site in a manner equivalent to the NTP base. The sugar moiety of NAI interacts with the i+1 site of the RNAP NTP addition site equivalent to the NTP sugar. The glutamine moiety makes interactions that mimic NTP triphosphate interactions; and the

N-hydroxy and guanidinyl moieties interact with the RNA nucleotide base-paired to the preceding template position at the RNA 3'-nucleotide, with the N-hydroxy donating an H-

bond to the 3'-OH of the RNA 3'-nucleotide, and the guanidinyl moiety donating one H- bond to the 5´-phosphate of the RNA 3'-nucleotide and another to the base of the RNA 3'- nucleotide (Figure 28).

With the promising results seen in the semi-synthesis of compound 92, and the subsequent assays, the desoxy-PUM, was determined to be a reasonable lead. A total synthetic approach of the desoxy-PUM would provide a better reference molecule that will not only corroborate the structure and stereochemistry of 92 but will provide additional routes to synthesize novel PUM derivatives. Synthesis of desoxy-PUM was attained in eight steps through the coupling of commercially available β-D-pseudouridine and glycyl-L-glutamine, as shown in (Scheme 20).41

Scheme 20: Total Synthesis of Desoxy-PUM 10547

The synthesis started with acetonide protection of β-D-pseudouridine through treatment with 2,2-dimethoxypropane in dimethylformamide in the presence of concentrated HCl at room temperature. The resulting compound was mesylated by treatment with methanesulfonyl chloride to yield compound 98. Azidation of 98 (compound not shown) followed by reduction in THF in the presence of trimethylphosphine delivered compound

99. Compound 100 was then protected with Fmoc in the presence of dioxane and water sodium carbonate, followed by Fmoc chloride to give 101. The protected nucleoside 99 and protected Fmoc dipeptide 101 were coupled by reacting them with dry dimethylformamide, N, N′-dicyclohexylcarbodiimide, and 1-hydroxybenzotriazole to furnish 102. Crude 102 was treated with dimethylformamide and piperidine to give

Fmoc deprotected 103. To a solution of crude 103 in MeOH, 3,5-dimethylpyrazole-1- carboxamidine was used to obtain compound 104. Finally, deprotection of the acetonide by treatment of the crude product with acetic acid: water (7:3; 2 ml) gave final product

105. All products were analyzed by LC-MS (Scheme 20).

5.3 Novel synthesis of Pseudouridimycin Analog

Most antibiotics kill bacteria that are multiplying in infected patients.59 But PUM is also predicted to kill dormant bacteria, by inhibiting the enzyme polymerase required for every function in every organism. As mentioned earlier, polymerase transcribes DNA into RNA, and the RNA guides the synthesis of all cellular proteins.1

For more than a decade, Ebright et al. looked for compounds such as PUM that target polymerase.60 They showed that PUM not only inhibits polymerase, but it does by

mimicking uridine triphosphate (UTP) one of the building blocks of RNA. It binds to the polymerase at the RNAP active-center NDP addition site, thus competing with UTP for binding.41,43 For resistance to evolve, the bacteria would have to change its polymerase just enough to exclude PUM while allowing all other nucleotides to fit. Given the fact that the amino acids that interact with PUM at the binding site are essential, mutations to exclude or prevent binding of PUM would render the polymerase itself inactive. This gives PUM an added advantage that makes it about ten times less likely to trigger antibiotic resistance than traditional antibiotics.

Although human and bacterial RNAP are thought to have similar shapes, PUM explicitly interacts only with bacterial polymerases and not with the human polymerase. This will enhances the potential clinical profile of PUM derivatives by decreasing the likelihood of unwanted adverse reactions.41,43

Analogs that mimic RNA building blocks have been previously used to treat viruses such as human immunodeficiency virus (HIV) and hepatitis C virus (HCV), but this approach has only been recently applied to antibiotics.44 To investigate the mechanism of action of nucleoside inhibitors against bacteria, new 5´-modified nucleoside analogues will be synthesized that will act as NAIs capable of competing with NTPs to occupy the RNAP

NTP addition site as described by Maffioli et al.41 These compounds will target bacterial

RNAP in gram negative and gram-positive organisms while maintaining Watson-Crick base pairing in the active site. It is imperative to design a synthetic pathway for the routine synthesis of these substrates, followed by testing against Gram-positive and

Gram-negative bacteria to determine their antibacterial potency.

Novel derivatives of the NAI will be synthesized based on the structure activity requirements elucidated by investigations into PUM biological activity. A retrosynthetic approach to obtain these derivatives is shown in scheme 21.

Scheme 21: Retrosynthetic Analysis

As mentioned earlier, the synthesis of pseudouridine has been previously described by several laboratories.36–39Most approaches begin with a modified sugar, followed by coupling, oxidative-reduction, condensation and subsequent acid deblocking of the tert- butyl and isopropylidene groups under standard conditions (Scheme 13). Modifications will be introduced at the 5´ position of the nucleoside following synthetic methods developed by the Bryant-Friedrich group (Scheme 18) 33 and Maffioli et al41 (Scheme

20).

The synthesis of the C5¢ aminopseudouridine begins with commercially available ribonolactone 66, a substrate transformed over eight steps, then coupled to the nucleobase to form key intermediate 60 using the previously described approach. (Scheme 18).

Compound 60 was treated with methanesulfonyl chloride in pyridine to yield mesylated

compound 106 in 91 % yield. Conversion of 106 to 108 was done in two steps by azidation of 107 with sodium azide in dimethylformamide, followed by azide reduction to furnish the desired C5¢ primary amine 10841,43,36–39in 46% (Scheme 22).

Scheme 22: Synthesis of Primary C5'-Aminopseudouridine 108

Similar conditions as seen in Scheme 16 were applied to compound 60 to synthesize 109.

Compound 86 was obtained through oxidation of alcohol 60 with IBX in 94% yield.

Formation of cyanohydrin 87 was achieved using similar methods as previously described, using catalytic amounts of 18-Crown-6, KCN and TBDMSCN in 73% yield.

To obtain a primary amine at the 5¢ position, we attempted to reduce 87 to compound 109

using LAH. This approach was not successful. Other reducing agents will be investigated in the future. (Scheme 23).

Scheme 23: Attempted synthesis of C5¢-aminopseudouridine nucleoside 109

To facilitate the peptide coupling to amine 108, an FMOC protected substrate was utilized as seen in Schemes 20 and 21.41

Scheme 24: Synthesis of C5¢-aminopseudouridine nucleoside 113.

The protected glycine-glutamine dipeptide 101 was obtained commercially and coupled to 5´-aminopseudouridine 108 in the presence of N,N'-Dicyclohexylcarbodiimide (DCC) and hydroxybenzotriazole (HOBt) to furnish 110 (Scheme 24). The direct conversion of the carboxylic acid on 101 to an amide is difficult because of the basic nature of amines.

Amines tend to convert carboxylic acids to their highly unreactive carboxylates. In this reaction, the addition of the carboxylic acid to the DCC molecule formed an excellent leaving group which was then displaced by an amine during nucleophilic substitution.

The use of DCC to induce coupling to form an amide linkage is a significant reaction in peptide synthesis. HOBt was also used in this reaction as a dehydrating agent and as an additive to decrease racemization in the carbodiimide peptide coupling (Scheme 24).

Scheme 25: Synthesis of compound 110

Deprotection of the protected amine in the presence of piperidine in DMF gave product

111 (Scheme 25).

Scheme 26: Synthesis of compound 111

Incorporation of the guanidine moiety with 3´5-dimethylpyrazole-1 carboxamidine furnished compound 112. Final deprotection of the 2´3´-isopropylidene group with 7:3 acetic acid: water (AcOH: H2O) obtained the desired nucleoside analog inhibitor 114

(Scheme 27). High-resolution mass spectrometry further confirmed product formation

(Figure 67).

Scheme 27: Synthesis of compound 113

5.4 Conclusion and Future Direction

Studies on the impact of pseudouridine based nucleoside-analog inhibitors (NAIs) of viral nucleotide polymerases as treatments of Human Immunodeficiency Virus (HIV) and

Hepatitis C Virus (HCV) have broadened our understanding of the mechanism of action of RNA polymerases.47 These findings have encouraged the development of pseudouridine NAIs of bacterial RNA polymerases (RNAP) that can limit bacterial resistance by mimicking the RNAP nucleoside triphosphate (NTP).47,54 In this study, we designed and synthesized desoxy-pseudouridymicin. Our future efforts will focus on the full characterization of compounds 110 to 113. The compound synthesized is expected to

interact with the RNAP in a similar manner as PUM. Albeit, a reduction in the binding affinity or potency is expected due to the loss of the hydrogen binding ability caused by the absence of the hydroxyl group (Figure 14)41. The modified C5´ pseudouridine nucleoside 109 was successfully synthesized and characterized. This was used to obtain dipeptide nucleoside analog 113.

100

Chapter 6

6 Experimental Procedure: Pseudouridimycin and its

analogs

6.1 Materials

6.2 Analysis Method

All chemicals were commercially obtained, were of the highest grade and used without further purification unless otherwise specified. Thin layer chromatography was carried out on Silicycle® silica gel 60 F254 aluminum-backed plates. Product spots were visualized by UV light at 254 nm and/or staining of the TLC plate with permanganate or anisaldehyde dip made in house. All synthesized products were characterized by NMR using an Inova-600 or Avance-600 Bruker Avance III 600 Mhz. spectrometer in CDCl3 or

CD3OD as the solvent. Additional MS spectra were obtained using Shimazu LCMS 2020.

All chemical shift values and, are reported in parts per million and coupling constants in

Hertz.

101

6.3 Synthesis of 5´-aminopseudouridine

6.3.1 ((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d] [1,3] dioxol-4-yl) methyl methane sulfonate (106)

To a stirred solution of 60 (.40 g, 1.01 mmol) in pyridine (2 ml), mesylchloride was added dropwise and the reaction mixture left to stir at room temperature overnight until

TLC showed no remaining starting material. The mixture was quenched with water and extracted with DCM. The organic layer was dried with MgSO4 and concentrated.

Purification with column chromatography (5-20% EtoAc/Hex) afforded 117 in 94% yield.

1H NMR (CDCl3): δ 8.16 (d, J = 2.6 Hz, 1H), 4.91 (d, J = 3.8 Hz, 1H), 4.74 (dd, J = 6.8,

3.8 Hz, 1H), ), 4.64 (dd, J = 6.4, 3.9 Hz, 1H), 4.54 – 4.32 (2H, m), 4.74 (m, 2H), 4.18 (dd,

J = 5.5, 3.2 Hz, 1H), 3.04 (m, 3H), 1.65 (m, 5H), 1.64 – 1.56 (m , 13H), 1.35 (d, J = 2.7

Hz 2H), 1.29 – 1.25 (m, OH). 102

6.3.2 5-((3aS,4S,6R,6aR)-6-(azidomethyl)-2,2-dimethyltetrahydrofuro[3,4-d] [1,3] dioxol-4-yl)-2,4-di-tert-butoxypyrimidine (107)

Compound 106 (360 mg, 0.76 mmol) and NaN3 (44.6 mg, 1.06 mmol, 1.4 eq) were dissolved in DMF (4 ml) and the reaction refluxed at 100 °C for 3hrs. The mixture was allowed to cool to room temperature and solvent removed by rotary evaporation to give

107 in 84% yield.

1 H NMR (600 MHz, CD3CN) δ 7.95 (s, 1H), 7.42 – 7.32 (m, 1H), 4.79 – 4.67 (m, 2H),

4.05 (ddd, J = 15.3, 8.6, 5.0 Hz, 1H), 3.57 (ddd, J = 17.2, 14.4, 8.0 Hz, 1H), 3.50 – 3.38

(m, 1H), 2.95 (d, J = 47.9 Hz, 4H), 2.80 (d, J = 0.5 Hz, 4H), 1.66 – 1.51 (m, 4H), 1.36 –

1.25 (m, 4H), 0.09 – 0.00 (m, 12H).

103

6.3.3 ((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d] [1,3] dioxol-4-yl) methanamine (108)

A solution of compound 107 (0.28 g, 0.67 mmol) in THF (12 ml), H2O (13 ml) and 1 M trimethylphosphine in THF (1.34 mml, 1.34 mmol, 2 eq) was stirred for 2 h at room temperature, and solvent removed under reduced pressure. Column chromatography with silica gel using a DCM/MeOH/NH3 solvent system delivered 108 in 46% yield as a brown oil.

1H NMR (600 MHz, DMSO) δ 8.24 (s, 1 H),4.47 (m, 5H) 4.68 – 4.57 (m, 5H), 4.03 (dd,

J = 14.2, 7.1 Hz, 6H), 3.80 (dd, J = 10.1, 5.1 Hz, 6H), 3.55 – 3.49 (m, 2H), 3.23 (d, J =

7.3 Hz, 4H), 2.89 (s, 3H), 2.72 (m, 5H), 2.63 – 2.59 (m, 4H), 1.59 – 1.40 (m, 6H), 1.38 (s,

3H), 1.30 – 1.22 (m, 11H), 1.17 (dd, J = 15.5, 8.4 Hz, 7H), 1.09 (s, 3H), 1.00 – 0.90 (m,

9H), 0.86 (s, 3H).

104

6.3.4 Synthesis of desoxypseudouridimycin

6.3.4.1 (9H-fluoren-9-yl)methyl (2-((5-amino-1-((((3aR,4R,6S,6aS)-6-(2,4-di-tert- butoxypyrimidin-5-yl)-2,2-dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)methyl)amino)-1,5- dioxopentan-2-yl)amino)-2-oxoethyl)carbamate (110)

To a solution of 101 (40 mg, .094 mmol) and primary amine 108 (50 mg, .126 mmol, 1.1 eq.) in dry DMF (3.5 ml), was added N, N′-dicyclohexylcarbodiimide (38 mg, .094 mmol, .094, 1.2 eq.) and 1-hydroxybenzotriazole (40 mg, 094 mmol, 2 eq.). The reaction mixture stirred overnight at room temperature, and the solvent was evaporated under reduced pressure to deliver 110. The compound was used in the next step without purification

6.3.4.2 2-(2-aminoacetamido)-N1-(((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)methyl)pentanediamide (111)

105

To a solution of 110 (60 mg, 0.074 mmol) in DMF (4 ml), piperidine (1 ml) was added, and the reaction mixture was stirred for 10 min at room temperature. The solvent was evaporated under reduced pressure and the compound dried under high vacuum for 24 hours. solid residue was washed with DCM (5 x 3 ml) to obtain crude 111 which was used in the next step without further purification.

6.3.4.3 N1-(((3aR,4R,6S,6aS)-6-(2,4-di-tert-butoxypyrimidin-5-yl)-2,2- dimethyltetrahydrofuro[3,4-d][1,3]dioxol-4-yl)methyl)-2-(2- guanidinoacetamido)pentanediamide (112)

106

To a solution of crude solution of 111 (40 mg, 0.69 mmol) in methanol (500 μl) was added 3,5-dimethylpyrazole-1-carboxamidine (90 mg, 0.69 mmol, 10 eq). The reaction mixture was stirred overnight at 25°C followed by reflux for 8h at 65°C. The solvent was evaporated under reduced pressure and the residual solid washed with DCM (3 x 5 ml) to deliver crude compound 112 without further purification.

6.3.4.4 N1-(((2R,3S,4R,5S)-5-(2,4-dioxo-1,2,3,4-tetrahydropyrimidin-5-yl)-3,4- dihydroxytetrahydrofuran-2-yl)methyl)-2-(2-guanidinoacetamido)pentanediamide (113)

Crude 112 was treated (33 mg, 0.56 mmol) in acetic acid: water (7:3; 5 ml) and was stirred overnight at 25 °C then heated to 50 °C under reflux for 10 h under argon. After

107

the solvent was evaporated, the solid residue was washed with DCM (4 x 5 ml) and methanol (5 ml), yielding a white solid 113 in 87% crude yield.

108

Work Cited

1. Uzman, A. Molecular Cell Biology (4th edition) Harvey Lodish, Arnold Berk, S.

Lawrence Zipursky, Paul Matsudaira, David Baltimore and James Darnell;

Freeman & Co., New York, NY, 2000, 1084 pp., list price $102.25, ISBN 0-7167-

3136-3. Biochem. Mol. Biol. Educ. 29, 126–128 (2001).

2. Oyejide, L., Mendes, O. R. & Mikaelian, I. Molecular Pathology. A

Comprehensive Guide to Toxicology in Nonclinical Drug Development 407–445

(2017). doi:10.1016/b978-0-12-803620-4.00016-5

3. Kong, Q. & Lin, C. G. Oxidative damage to RNA: mechanisms, consequences,

and diseases. Cell. Mol. Life Sci. 67, 1817–1829 (2010).

4. Durairaj, A. & Limbach, P. A. Mass spectrometry of the fifth nucleoside: A review

of the identification of pseudouridine in nucleic acids. Anal. Chim. Acta 623, 117–

125 (2008).

5. Meier, U. T. Pseudouridylation goes regulatory. EMBO J. 30, 3–4 (2011).

6. Gray Michael W., M. C. Pseudouridine in RNA: What, Where, How, and Why.

IUBMB Life (International Union Biochem. Mol. Biol. Life) 49, 341–351 (2000).

7. Fedorov, N. A., Bogomazov, M. Y. & Kosenko, A. S. Excretion of 5-ribosyluracil

(Pseudouridine) by healthy persons and cancer patients before and after

radiotherapy. Bull. Exp. Biol. Med. 62, 1174–1176 (1966).

8. Waalkes, T. P., Dinsmere, S. R. & Mrochek, J. E. Urinary Excretion by Cancer 109

Patients of the Nucleosides N 2 , N 2 -Dimethylguanosine, 1-Methylinosine, and

Pseudouridine 2. JNCI J. Natl. Cancer Inst. 51, 271–274 (1973).

9. Gehrke, C. W., Kuo, K., Walkes, T. & Borek, E. Patterns of urinary excretion of

modified nucleosides. Cancer Res. 39, 1150–1153 (1979).

10. Seidel, A., Brunner, S., Seidel, P., Fritz, G. I. & Herbarth, O. Modified

nucleosides: an accurate tumour marker for clinical diagnosis of cancer, early

detection and therapy control. Br. J. Cancer 94, 1726–1733 (2006).

11. Li, S. & Mason, C. E. The Pivotal Regulatory Landscape of RNA Modifications.

Annu. Rev. Genomics Hum. Genet. 15, 127–150 (2014).

12. Satterlee, J. S. et al. Novel RNA Modifications in the Nervous System: Form and

Function. J. Neurosci. 34, 15170–15177 (2014).

13. Wang, X. & He, C. Dynamic RNA Modifications in Posttranscriptional

Regulation. Mol. Cell 56, 5–12 (2014).

14. Jack, K. et al. rRNA Pseudouridylation Defects Affect Ribosomal Ligand Binding

and Translational Fidelity from Yeast to Human Cells. Mol. Cell 44, 660–666

(2011).

15. Maden, B. E. H. The Numerous Modified Nucleotides in Eukaryotic Ribosomal

RNA. Progress in Nucleic Acid Research and Molecular Biology 241–303 (1990).

doi:10.1016/s0079-6603(08)60629-7

16. Dominissini, D. et al. Topology of the human and mouse m6A RNA methylomes

revealed by m6A-seq. Nature 485, 201–206 (2012).

17. Carlile, T. M. et al. Pseudouridine profiling reveals regulated mRNA

pseudouridylation in yeast and human cells. Nature 515, 143–146 (2014). 110

18. Schwartz, S. et al. Transcriptome-wide Mapping Reveals Widespread Dynamic-

Regulated Pseudouridylation of ncRNA and mRNA. Cell 159, 148–162 (2014).

19. Lovejoy, A. F., Riordan, D. P. & Brown, P. O. Transcriptome-Wide Mapping of

Pseudouridines: Pseudouridine Synthases Modify Specific mRNAs in S.

cerevisiae. PLoS One 9, e110799 (2014).

20. Hee Lee, S., Kim, I. & Chul Chung, B. Increased urinary level of oxidized

nucleosides in patients with mild-to-moderate Alzheimer’s disease. Clin. Biochem.

40, 936–938 (2007).

21. Shan, X., Tashiro, H. & Lin, C. G. The Identification and Characterization of

Oxidized RNAs in Alzheimer’s Disease. J. Neurosci. 23, 4913–4921 (2003).

22. Abe, J. & Berk, B. C. Reactive Oxygen Species as Mediators of Signal

Transduction in Cardiovascular Disease. Trends Cardiovasc. Med. 8, 59–64

(1998).

23. Luca, M., Luca, A. & Calandra, C. The Role of Oxidative Damage in the

Pathogenesis and Progression of Alzheimer’s Disease and Vascular Dementia.

Oxid. Med. Cell. Longev. 2015, 1–8 (2015).

24. Castellani, R. et al. Sublethal RNA Oxidation as a Mechanism for

Neurodegenerative Disease. Int. J. Mol. Sci. 9, 789–806 (2008).

25. Li, J., O, W., Li, W., Jiang, Z.-G. & Ghanbari, H. Oxidative Stress and

Neurodegenerative Disorders. Int. J. Mol. Sci. 14, 24438–24475 (2013).

26. Zhang, J. et al. Parkinson’s Disease Is Associated with Oxidative Damage to

Cytoplasmic DNA and RNA in Substantia Nigra Neurons. Am. J. Pathol. 154,

1423–1429 (1999). 111

27. Nunomura, A. et al. RNA Oxidation Is a Prominent Feature of Vulnerable

Neurons in Alzheimer’s Disease. J. Neurosci. 19, 1959–1964 (1999).

28. Fimognari, C. Role of Oxidative RNA Damage in Chronic-Degenerative Diseases.

Oxid. Med. Cell. Longev. 2015, 1–8 (2015).

29. Audat, S. A. S., Trzasko Love, C., Al-Oudat, B. A. S. & Bryant-Friedrich, A. C.

Synthesis of C3′ Modified Nucleosides for Selective Generation of the C3′-Deoxy-

3′-thymidinyl Radical: A Proposed Intermediate in LEE Induced DNA Damage. J.

Org. Chem. 77, 3829–3837 (2012).

30. Körner, S., Bryant-Friedrich, A. & Giese, B. C-3’-branched thymidines as

precursors for the selective generation of C-3’-nucleoside radicals. J. Org. Chem.

(1999). doi:10.1021/jo982022y

31. Schiemann, O., Feresin, E., Carl, T. & Giese, B. 4′-Pivaloyl Substituted Thymidine

as a Precursor for the Thymyl Radical: An EPR Spectroscopic Study.

ChemPhysChem 5, 270–274 (2004).

32. Manetto, A. et al. Independent Generation of C5´-Nucleosidyl Radicals in

Thymidine and 2‘-Deoxyguanosine. J. Org. Chem. 72, 3659–3666 (2007).

33. Shaik, R., Ellis, M. W., Starr, M. J., Amato, N. J. & Bryant-Friedrich, A. C.

Photochemical Generation of a C5′-Uridinyl Radical. ChemBioChem 16, 2379–

2384 (2015).

34. Shaik, R. Photochemical Generation of the C5´ -Uridinyl and

Pseudouridinylradical for the Study of Oxidative Damage in RNA. (University of

Toledo, 2013).

35. Menon, R. M. et al. Drug-drug interaction profile of the all-oral anti-hepatitis C 112

virus regimen of paritaprevir/ritonavir, ombitasvir, and dasabuvir. J. Hepatol.

(2015). doi:10.1016/j.jhep.2015.01.026

36. Shapiro, R. & Chambers, R. W. SYNTHESIS OF PSEUDOURIDINE. J. Am.

Chem. Soc. 83, 3920–3921 (1961).

37. Brown, D. M., Burdon, M. G. & Slatcher, R. P. A synthesis of pseudouridine and

of 5-β-D-ribofuranosyluridine. J. Chem. Soc. C 0, 1051–1053 (1968).

38. Grohar, P. J. & Chow, C. S. A practical synthesis of the modified RNA nucleoside

pseudouridine. Tetrahedron Lett. 40, 2049–2052 (1999).

39. Hanessian, S. & Machaalani, R. A highly stereocontrolled and efficient synthesis

of α- and β-pseudouridines. Tetrahedron Lett. 44, 8321–8323 (2003).

40. More, J. D. & Finney, N. S. A Simple and Advantageous Protocol for the

Oxidation of Alcohols witho-Iodoxybenzoic Acid (IBX). Org. Lett. 4, 3001–3003

(2002).

41. Maffioli, S. I. et al. Antibacterial Nucleoside-Analog Inhibitor of Bacterial RNA

Polymerase. Cell 169, 1240-1248.e23 (2017).

42. Frieden, T. Antibiotic Resistance Threats in the United States, 2013. Brochure -

US Centrs for Disease Control and Prevention (2013). doi:CS239559-B

43. Ebright, R. H. RNA-Exit Channel: Target and Method for Inhibition of Bacterial

RNA. Patent (2005). doi:10.1016/j.molstruc.2011.02.014

44. Landwehr, W., Wolf, C. & Wink, J. Actinobacteria and Myxobacteria—Two of

the Most Important Bacterial Resources for Novel Antibiotics. Current Topics in

Microbiology and Immunology 273–302 (2016). doi:10.1007/82_2016_503

45. Degen, D. et al. Transcription inhibition by the depsipeptide antibiotic salinamide 113

A. Elife 3, (2014).

46. Lancini, G. C. & Sartori, G. Rifamycins LXI: In vivo inhibition of RNA synthesis

by rifamycins. Experientia 24, 1105–1106 (1968).

47. Lancini, G., Pallanza, R. & Silvestri, L. G. Relationships between bactericidal

effect and inhibition of ribonucleic acid nucleotidyltransferase by rifampicin in

Escherichia coli K-12. J. Bacteriol. 97, 761–768 (1969).

48. SERGIO, S., PIRALI, G., WHITE, R. & PARENTI, F. Lipiarmycin, a new

antibiotic from Actinoplanes. III. Mechanism of action. J. Antibiot. (Tokyo). 28,

543–549 (1975).

49. IRSCHIK, H., GERTH, K., HÖFLE, G., KOHL, W. & REICHENBACH, H. The

myxopyronins, new inhibitors of bacterial RNA synthesis from Myxococcus

fulvus (Myxobacterales). J. Antibiot. (Tokyo). 36, 1651–1658 (1983).

50. IRSCHIK, H., JANSEN, R., HÖFLE, G., GERTH, K. & REICHENBACH, H. The

corallopyronins, new inhibitors of bacterial RNA synthesis from Myxobacteria. J.

Antibiot. (Tokyo). 38, 145–152 (1985).

51. IRSCHIK, H., AUGUSTINIAK, H., GERTH, K., HÖFLE, G. &

REICHENBACH, H. Antibiotics from gliding bacteria. No. 68. The Ripostatins,

Novel Inhibitors of Eubacterial RNA Polymerase Isolated from Myxobacteria. J.

Antibiot. (Tokyo). 48, 787–792 (1995).

52. Jin, D. J. & Gross, C. A. Mapping and sequencing of mutations in the Escherichia

colirpoB gene that lead to rifampicin resistance. J. Mol. Biol. 202, 45–58 (1988).

53. Campbell, E. A. et al. Structural Mechanism for Rifampicin Inhibition of Bacterial

RNA Polymerase. Cell 104, 901–912 (2001). 114

54. Garibyan, L. Use of the rpoB gene to determine the specificity of base substitution

mutations on the Escherichia coli chromosome. DNA Repair (Amst). 2, 593–608

(2003).

55. Srivastava, A. et al. New target for inhibition of bacterial RNA polymerase:

‘switch region’. Curr. Opin. Microbiol. 14, 532–543 (2011).

56. Tuske, S. et al. Inhibition of Bacterial RNA Polymerase by Streptolydigin:

Stabilization of a Straight-Bridge-Helix Active-Center Conformation. Cell 122,

541–552 (2005).

57. Temiakov, D. et al. Structural Basis of Transcription Inhibition by Antibiotic

Streptolydigin. Mol. Cell 19, 655–666 (2005).

58. Belogurov, G. A. et al. Transcription inactivation through local refolding of the

RNA polymerase structure. Nature 457, 332–335 (2008).

59. Brown, E. D. & Wright, G. D. Antibacterial drug discovery in the resistance era.

Nature 529, 336–343 (2016).

60. Zhang, Y. et al. GE23077 binds to the RNA polymerase ‘i’ and ‘i+1’ sites and

prevents the binding of initiating nucleotides. Elife 3, (2014).

115

116

Appendix A

Spectra Data

Figure 30: 1H NMR for compound 61

117

Figure 31: 1H NMR for compound 62

118

Figure 32: LCMS for Compound 62

119

Figure 33: 1H NMR for compound 57

120

Figure 34: 1H NMR for compound 80

121

Figure 35: LCMS for compound 80

122

GS_SS_050319_ABY_IM_CP_72 120 (2.057) AM (Cen,4, 80.00, Ar,8000.0,556.28,0.70); Cm (117:121) TOF MS ES+ 651.3425 100 6.88e4

% 652.3484

595.2809 561.2029 593.1411 653.3491 596.2842 577.2371 651.1606 654.3513 563.2080 579.2189 601.1353 640.1595 673.3292 689.2869 0 m/z 550 560 570 580 590 600 610 620 630 640 650 660 670 680 690 700

Figure 36: HRMS for compound 80

123

Figure 37: 1H NMR for compound 81

124

Figure 38: LCMS for compound 81

125

Figure 39: 1H NMR for compound 82

126

Figure 40: LCMS for compound 82

127

GS_SS_050319_ABY_IM_CP_74 142 (2.431) AM (Cen,4, 80.00, Ar,8000.0,556.28,0.70); Cm (141:144) 635.2086 100 3.58e4

630.2537

* 556.2771 % 613.2283 636.2144

640.1787 640.6813 557.2850 641.1838 667.2363 503.1367 556.1363 558.2931 612.8043 644.6901 585.2390 699.2009 0 m/z 500 520 540 560 580 600 620 640 660 680 700

Figure 41: HRMS for compound 82

128

Figure 42: 1H NMR for compound 60

129

Figure 43: LCMS for compound 60

130

Figure 44: HRMS for compound 60

131

Figure 45: LCMS of compound 6

132

Figure 46: 1H NMR for compound 86

133

Figure 47: LCMS for compound 86

134

Figure 48: 1H NMR for compound 87

135

Figure 49: 13C NMR for compound 87

136

Figure 50: 1H NMR for compound 89

137

Figure 51: 1H NMR for compound 90

138

GS_SS_050319_ABY_IM_CP_100_1 131 (2.244) AM (Cen,4, 80.00, Ar,8000.0,556.28,0.70); Cm (123:131) 464.1196 100 1.32e3

465.2019 481.1700

466.1888 %

499.1678 482.1721 470.1693 497.1440 481.1165

479.1497 490.2001

0 m/z 465 470 475 480 485 490 495 500

Figure 52: HRMS for compound 90

139

Figure 53: 1H NMR for compound 13

140

Figure 54: 13C NMR for Compound 13

141

Figure 55: LCMS for Compound 13

142

Figure 56: 1H NMR for compound 106

143

Figure 57: 1H NMR for compound 107

144

Figure 58: LCMS for compound 107

145

Figure 59: 1H NMR for compound 108

146

Figure 60: 1H NMR for compound 108

147

Figure 61: 13 C NMR for compound 108

148

Figure 62: COSY for compound 108

149

Figure 63: HSQC for compound 108

150

Figure 64: 1H NMR for compound 110

151

Figure 65: LCMS for compound 112

152

Figure 66: 1H NMR for compound 111

153

Figure 67: 13C NMR for compound 111

154

GS_SS_050319_ABY_IM_CP_124 10 (0.187) AM (Cen,4, 80.00, Ar,8000.0,556.28,0.70); Cm (10:18) TOF MS ES+ 471.1960 100 3.97e4 %

521.2134

472.1976 * 556.2771

522.2136

454.1686 464.1184 523.2125 550.2401 473.1985 486.1970 527.2085 540.2157 497.1452 503.1664511.2126 0 m/z 450 460 470 480 490 500 510 520 530 540 550 560

Figure 68: HRMS for compound 114

155