The Iron-Dependent Cyanide and Hydrogen Peroxide Co-Toxicity in Escherichia Coli and Its Catastrophic Consequences for the Chromosome

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THE IRON-DEPENDENT CYANIDE AND HYDROGEN PEROXIDE CO-TOXICITY IN ESCHERICHIA COLI AND ITS CATASTROPHIC CONSEQUENCES FOR THE CHROMOSOME

TULIP MAHASETH

DISSERTATION

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Microbiology in the Graduate College of the University of Illinois at Urbana-Champaign, 2015

Urbana, Illinois

Doctoral Committee:

Professor Andrei Kuzminov, Chair and Director of Research Professor John E. Cronan Professor Jeffrey F. Gardner Associate Professor Carin K. Vanderpool ABSTRACT

2+ Hydrogen peroxide (H2O2) can oxidize cytoplasmic ferrous ions (Fe ) to produce highly reactive hydroxyl radicals (•OH) via Fenton’s reaction that can damage various biomolecules causing oxidative stress. Even though at concentrations higher than 20 mM H2O2 by itself can efficiently kill micro-organisms, it is metabolically impossible for eukaryotic cells to generate H2O2, an uncharged molecule, in such large quantities inside the cell. We propose that potentiation of physiologically relevant amounts of H2O2 by various small molecules serves as a more feasible and safe mechanism to combat invading microbes. NO potentiation of H2O2 toxicity is a known bactericidal weapon employed by macrophages. In fact, in human neutrophils activated by bacterial infection, the myeloperoxidase enzyme catalyzes the formation of hydrogen cyanide (HCN) from serum thiocyanate (SCN-). In the past, researchers have reported that a combination of low millimolar doses of H2O2 and cyanide (CN), which are individually bacteriostatic, caused rapid synergistic killing in Escherichia coli. Our aim is to understand the immune cells antimicrobial responses by investigating the mechanism of CN potentiation of H2O2 toxicity and its chromosomal consequences.

We have found that the ability of CN to recruit iron from intracellular depots such as ferritin contributes to its potentiation of H2O2 toxicity, whereas the major stationary phase intracellular iron depot protein, Dps, can sequester this iron, thereby quelling

Fenton's reaction. Our work has also demonstrated that this synergistic toxicity is associated with catastrophic chromosomal fragmentation, which is blocked by iron chelators. Moreover, the CN + H2O2 treatment also resulted in destruction of ribosomal

RNA, indicating that RNA is equally susceptible to the oxidative damage induced. The

ii catastrophic chromosomal fragmentation induced by CN and H2O2 was found to be cell density-dependent, but replication- and translation-independent. We propose that disrupting intracellular iron trafficking to cause catastrophic chromosomal fragmentation is a common strategy employed by the immune system to kill invading microbes. We also showed that the base excision repair pathway plays the major role in preventing this catastrophic chromosomal fragmentation, which indicates that the double-strand breaks induced by CN + H2O2 are preceded by a significant number of base modifications, removed by base-excision repair via the abasic site intermediates. These lesions can be converted into double-strand breaks that are repairable via the recombinational repair system. On the other hand, termination of the CN + H2O2 treatment, which results in termination of the CN-induced block of respiration and ATP production, was found to trigger significant linear DNA degradation and disintegration of the nucleoid structure.

We have shown that this disassembly of the nucleoid structure following the removal of

CN and H2O2 is affected by the nucleoid-associated proteins and depends on ongoing transcription, thereby revealing the role of transcription in the nucleoid dynamics.

Therefore, we conclude that cyanide potentiates hydrogen peroxide toxicity by recruiting iron from intracellular depots directly onto the DNA, where the DNA-iron complexes catalyze self-targeted Fenton's reaction leading to catastrophic DNA damage.

This massive DNA damage is actively countered by base-excision repair mechanisms, but it eventually saturates the cellular DNA repair capacity and causes disintegration of the trancriptionally-active nucleoids.

iii To my family and loved ones

iv ACKNOWLEDGEMENTS

This work would not have been possible without Dr. Andrei Kuzminov’s incredible guidance and support. Andrei is a brilliant scientist, from whom I have learned so much. I am extremely grateful to have the opportunity to work alongside him for the past several years. Thank you for always steering me in the right direction, in science and otherwise.

I would like to thank Dr. James Imlay and all the members of my thesis committee – Dr. John Cronan, Dr. Jeffrey Gardner and Dr. Carin Vanderpool, for their advice and invaluable feedback throughout graduate school. Jim, an expert on hydrogen peroxide toxicity, has been especially patient and supportive of my work, which was, in fact, pioneered by him during his time as a graduate student at Berkeley. I am thankful to him for all the times he has listened to the several obstacles I faced with this project and provided his invaluable insights on them.

I would also like to thank all the members of the Kuzminov Lab – Dr. Elena

Kouzminova, Pritha Rao, Dr. Sharik Khan, Dr. Glen Cronan and Dr. Ka wai Kuong, for all their help and suggestions over the years, and for making the lab a fun and wonderful place to work.

I am also thankful to Micah Hodosh for his love and companionship, and for seeing me through the ups and downs of graduate school. Last, but certainly not the least,

I would like to thank my family – Dada, Mamma and Neil, for their love, support, encouragement and patience. Without them I would not be where I am today.

v TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION...... 1 1.1. BIOLOGICAL SIGNIFICANCE OF HYDROGEN PEROXIDE...... 1

1.l.1. Relevance of H2O2 at Physiological Levels...... 1

1.1.2. Biological Sources of H2O2 and Their Roles ...... 2 1.2. HYDROGEN PEROXIDE TOXICITY IN ESCHERICHIA COLI ...... 4 1.2.1. Oxidative Stress and Reactive Oxygen Species...... 4 1.2.2. Sources of ROS in the Cell ...... 5

1.2.3. Targets of H2O2...... 6

1.2.3.1. DNA damage caused by H2O2 ...... 6

1.2.3.2. RNA damage caused by H2O2 ...... 8 1.2.3.3. Protein oxidation...... 10 1.2.3.4. Lipid peroxidation...... 11 1.3. IRON METABOLISM AND TOXICITY...... 11 1.3.1. Iron-Containing Enzymes and Their Distribution in Metabolism ...... 11 1.3.2. The Iron Paradox...... 12 1.3.3. Iron Scavenging in E. coli...... 13 1.3.4. The Iron Depots of E. coli...... 14 1.3.4.1. The DNA binding, iron detoxification and storage protein, Dps...... 14 1.3.4.2. Ferritin - The primary cellular iron storage protein...... 16 1.3.5. Regulation of Iron Homeostasis by Fur ...... 17 1.4. FREE RADICAL STRESS RESPONSES AND SCAVENGERS...... 19 1.4.1. The OxyR Response ...... 19

1.4.1.1. H2O2 scavengers: catalases and peroxidases...... 21 1.4.2. The SoxRS Response...... 23 − 1.4.2.1. The O2 scavengers: SodA, SodB and SodC ...... 24 1.5. MECHANISMS OF CHROMOSOMAL FRAGMENTATION AND REPAIR ...... 27

vi 1.5.1. Mechanisms of Replication-Dependent Chromosomal Fragmentation ...... 28 1.5.2. Mechanisms of DNA Repair...... 31 1.5.2.1. Recombinational repair pathway ...... 31 1.5.2.2. Base excision repair pathway...... 32 1.5.2.3. Nucleotide excision repair pathway...... 32 1.6. POTENTIATED HYDROGEN PEROXIDE TOXICITY ...... 33 1.6.1. The Concept of Potentiated Toxicity ...... 33

1.6.2. Mechanism of NO + H2O2 Co-Toxicity...... 36

1.6.3. Mechanism of Cysteine-Enhanced H2O2 Toxicity ...... 37

1.6.4. The Cyanide + H2O2 Co-Toxicity...... 38

1.6.4.1. The potential of cyanide + H2O2 co-toxicity as a cytotoxic weapon in activated immune cells...... 40 1.7. SCOPE OF THIS THESIS...... 41

1.7.1. Cyanide Potentiates H2O2 Toxicity by Causing Catastrophic Chromosomal Fragmentation via Recruitment of Iron from Intracellular Depots...... 41 1.7.2. The Nature, Consequences and Attempted Repair of the Catastrophic Chromosomal Fragmentation Induced by the Synergistic Cyanide and Hydrogen Peroxide Combination ...... 43 1.8. FIGURES...... 47 1.9. REFERENCES ...... 49

CHAPTER 2: CYANIDE POTENTIATES HYDROGEN PEROXIDE TOXICITY BY RECRUITING IRON FROM INTRACELLULAR DEPOTS TO CAUSE CATASTROPHIC CHROMOSOMAL FRAGMENTATION ...... 64 2.1. INTRODUCTION ...... 64 2.2. MATERIALS AND METHODS...... 67 2.2.1. Strains and Plasmids ...... 67 2.2.2. Enzymes and Reagents ...... 68

vii 2.2.3. Growth Conditions and Viability Assay...... 68 2.2.4. Measuring Chromosomal Fragmentation via Pulsed-Field Gel Electrophoresis...... 68 2.2.5. In vitro Non-DNA Fenton's Reaction ...... 69 2.2.6. Plasmid Relaxation Assay...... 69 2.2.7. Southern Hybridization...... 70 2.3. RESULTS ...... 71

2.3.1. CN + H2O2 Treatment Causes Catastrophic Chromosomal Fragmentation ...... 71

2.3.2. The Nature of CN and H2O2 Co-Toxicity...... 72

2.3.3. Genetic Analysis of CN Potentiation of the H2O2 Killing: Expected Phenotypes and Their Interpretation ...... 73

2.3.4. H2O2 Scavengers Counteract CN Potentiation...... 74 2.3.5. The Electron Transport Connection...... 76

2.3.6. The Role of Iron in CN + H2O2 Co-Toxicity...... 79 2.3.7. Phenotypes of Mutants with Increased "Free" Cytoplasmic Iron...... 80 2.3.8. The Role of Iron-Depository Dps ...... 82 2.3.9. The Role of Iron-Depot Ferritin...... 83 2.4. DISCUSSION...... 85 2.4.1. Iron Distribution Inside the Cell and its Subversion by CN ...... 87 2.4.2. Catastrophic Chromosomal Fragmentation ...... 88 2.4.3. Potentiated Toxicity...... 90 2.4.4. Conclusion ...... 91 2.5. TABLES ...... 93 2.6. FIGURES...... 94 2.7. REFERENCES ...... 104

viii CHAPTER 3: THE MECHANISM, CONSEQUENCES AND ATTEMPTED REPAIR OF THE CATASTROPHIC CHROMOSOMAL FRAGMENTATION INDUCED BY THE BACTERICIDAL COMBINATION OF CYANIDE AND HYDROGEN PEROXIDE IN ESCHERICHIA COLI ...... 110 3.1. INTRODUCTION ...... 110 3.2. MATERIALS AND METHODS...... 113 3.2.1. Strains and Plasmids ...... 113 3.2.2. Reagents...... 114 3.2.3. Growth Conditions and Viability Assay...... 114 3.2.4. Measuring Chromosomal Fragmentation via Pulsed-Field Gel Electrophoresis...... 114 3.2.5. Southern Hybridization of Chromosomal Fragmentation...... 114 3.2.6. Isolation of rRNA ...... 115 3.2.7. Measuring Linear DNA Degradation...... 115 3.2.8. Measuring Nucleoid Disassembly ...... 116 3.3. RESULTS ...... 117 3.3.1. Nucleoid Organization Plays a Small Role in Protecting the

Chromosome from CN + H2O2-Induced Fragmentation, while Bulky DNA Lesions are not Involved ...... 117 3.3.2. Base-Excision Repair and One-Strand Gap Repair Mend the

Bulk of CN + H2O2-Induced DNA Lesions...... 119

3.3.3. Recombinational Repair Mends H2O2-Induced Double-Strand Breaks but is Paralyzed by CN...... 120

3.3.4. CN + H2O2-Induced Double-Strand Breaks do not Depend on DNA Replication and are Uniformly Distributed Over the Chromosome...... 122 3.3.5. Double-Strand Breaks are not Associated with DNA Found in Bayer’s Patches or Other Chromosomal DNA Attachment Sites...... 124

ix 3.3.6. Chromosomal DNA and rRNA Degradation After

CN + H2O2 Treatment ...... 127 3.3.7. Nucleoid Dynamics of Broken DNA Results in Partial Nucleoid Disintegration...... 128 3.3.8. A Comparison of Chromosomal Fragmentation Induced by

the CN + H2O2 with Other Treatments that Potentiate H2O2 Toxicity...... 131 3.4. DISCUSSION...... 132 3.5. TABLES ...... 134 3.6. FIGURES...... 136 3.7. REFERENCES ...... 146

CHAPTER 4: CHROMOSOMAL OVER-REPLICATION AND ITS UNUSUAL CONSEQUENCES IN THE ESCHERICHIA COLI SEQA MUTANT ...... 150 4.1. INTRODUCTION ...... 150 4.2. MATERIALS AND METHODS...... 153 4.2.1. Strains and Plasmids ...... 153 4.2.2. Growth Conditions and Viability Assay...... 153 4.2.3. Color Screen for Identifying SeqA-Dependent Mutants ...... 153 4.2.4. Measuring Rate of DNA Replication...... 154 4.2.5. Flow Cytometry ...... 154 4.2.6. Southern Hybridization...... 155 4.3. RESULTS ...... 155 4.3.1. Identification and Characterization of SeqA-Dependent Mutants ...... 155 4.3.2. seqA Mutants Require Linear DNA Degradation for Survival...... 155 4.3.3. Over-Replication and Problems with Replication Fork Progression in the seqA, rep and seqA rep Mutants...... 156

x 4.3.4. seqA Mutants are Sensitive to Rifampicin and are Unable to Finish Half of Their On-Going Replication Rounds in its Presence ...... 158 4.4. DISCUSSION...... 160 4.4.1. Lack of Sister Chromatid Cohesion in seqA Mutants Could be the Cause of its Dependence on Linear DNA Degradation ...... 160 4.4.2. Possible Mechanism of Replication Fork Inhibition in the seqA Mutant ...... 161 4.5. TABLES ...... 163 4.6. FIGURES...... 165 4.7. REFERENCES ...... 177

CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS ...... 179 5.1. REFERENCES ...... 182

xi CHAPTER 1: INTRODUCTION

1.1. BIOLOGICAL SIGNIFICANCE OF HYDROGEN PEROXIDE

1.1.1. Relevance of H2O2 at Physiological Levels

In aerobic cells, adventitious partial reduction of molecular oxygen (O2) gives rise to highly reactive compounds known as reactive oxygen species (ROS). These by- products of aerobic metabolism can damage a variety of biomolecules in the cells.

However, at physiological levels in eukaryotes, some of these ROS serve as signaling molecules involved in redox homeostasis and in regulation of several biological pathways including stress, growth, immune and hormonal responses and energy metabolism (168).

H2O2, an abundant form of ROS in the cells, due to its relative stability, uncharged nature and small size, can readily diffuse through membranes, and therefore, is a particularly good candidate to serve as a messenger for signaling. H2O2 has been found to play a role in regulating platelet-derived growth factor and epidermal growth factor signaling by inhibiting protein tyrosine phosphatases that attenuate signal transduction from these activated growth factor receptors via reversible oxidation of their redox-sensitive cysteine residues (123). The physiological levels of intracellular H2O2 is strikingly conserved among organisms. In aerobically growing exponential Escherichia coli cells, the intracellular H2O2 concentration is maintained at ~0.02 μM, while being toxic at levels higher than ~0.5 μM (136). Physiological concentrations of intracellular H2O2 lie in the same range for eukaryotic cells, but may vary somewhat depending on factors like cell age and oxidative stress.

1.1.2. Biological Sources of H2O2 and Their Roles

The electron transport chain in mitochondria facilitates ATP synthesis by producing a proton gradient across the membrane via transfer of electrons through a series of redox reactions, in which molecular oxygen serves as the final electron acceptor and is reduced to water. Mitochondria are also the major source of H2O2 in cells, where the superoxide generated by one-electron reduction of oxygen during aerobic respiration can be converted into H2O2 by dismutation (61). An increase in proton-motive force

(PMF) caused by increased proton pumping or by decreased proton return slows down electron flow, aiding in interaction of free electrons with oxygen, resulting in an increase

− in production of the superoxide anion (O2 ). Therefore, when the PMF is enhanced under

− hyperoxic conditions, it causes the mitochondrial O2 levels to rise (177). Conversely, a

− decrease in PMF due to an increase in return of protons reduces O2 generation (103).

− However, hypoxic conditions also stimulate O2 formation in mitochondria, which helps in adaptation, but its mechanism is not yet clear (187). The NADH dehydrogenase

(Complex I) and cytochrome bc1 (Complex III) are the major sites that leak electrons to

− oxygen, thereby resulting in O2 formation (109, 176). Several other mitochondrial enzymes, like α-ketoglutarate dehydrogenase and electron transfer flavoprotein-

− ubiquinone oxidoreductase, are also capable of generating O2 (157, 173). H2O2 generated by mitochondria is important for a variety of biological processes including regulation of the immune response, differentiation, autophagy and apoptosis (151).

I will discuss how, in macrophages, a combination of NO and NADPH-oxidase- generated H2O2 functions as a lethal weapon against pathogens later (in section 1.6.2.).

Here I am going to examine how H2O2 production is triggered by this family of NADPH

2 oxidase enzymes known as NOX enzymes. The NOX enzymes were first identified in activated neutrophils. Neutrophils can provide protection from pathogens by ingesting them via phagocytosis, which allows their subsequent inactivation by a combination of secreted ROS and enzymes in the phagosome. In the absence of infection, the NADPH oxidase complex is latent in neutrophils. Neutrophil stimulation leads to recruitment of the six subunits of the NADPH oxidase complex, comprising of the 5 NADPH phagocyte oxidase cytosolic factors and RAC1GTPase, in the membranes that induce a rapid release of ROS against the pathogens, a response known as respiratory burst. These flavin- and heme-containing NADPH oxidase complexes transfer electrons from cytosolic NADPH

− to O2 to generate O2 , which can be further reduced to H2O2 and generate hydroxyl

• radicals ( OH) (112). The importance of H2O2 production from NOX enzymes in mounting an immune response against invading pathogens is demonstrated by the fact that mutations in the catalytic domain of these enzymes, which appreciably decrease the

− ability to generate O2 , result in chronic granulomatous diseases, a genetic condition characterized by recurring infections (10). Researchers have also shown that a number of growth factors and cytokines, like tumor necrosis factor, epidermal growth factor, platelet-derived growth factor and angiotensin II, are capable of inducing a rapid increase in intracellular ROS in a variety of non-phagocytic cells through activation of NADPH oxidases in these cells (73). In addition to phagocytic NADPH oxidases, there are, in fact,

7 NOX homologues in humans that play a role in a range of host defenses and signaling, including angiogenesis and insulin signaling (114).

Besides mitochondria and the NOX enzymes, several other organelles such as peroxisome, endoplasmic reticulum, as well as enzymes, such as cyclooxygenases,

3 lipoxygenases, xanthine oxidase and the cytochrome P450 enzymes can generate H2O2 in cells, which can perform a myriad of functions (27). For example, H2O2 generated by the cytochrome P450 terminal oxidases, which utilize heme as a co-factor to catalyze oxidation of various metabolic intermediates, is crucial for regulating the diurnal variation of corticosteroid production in the adrenal cortex via its participation in distinct redox signaling pathways (42).

1.2. HYDROGEN PEROXIDE TOXICITY IN ESCHERICHIA COLI

1.2.1. Oxidative Stress and Reactive Oxygen Species

The world in which microbial life first evolved had no oxygen. It was not until a billion years later that photosynthetic life forms caused oxygenation, which brought a new set of problems with it. Although molecular oxygen is not very reactive itself, partially reduced forms of oxygen are more so. O2 has an even number of electrons, but in its π antibonding molecular orbitals, two of them have parallel spins, so they remain unpaired. Therefore, O2 cannot accept a pair of electrons with anti-parallel spins and is a weak oxidizing agent. On the other hand, the same paramagnetic nature of O2 makes it very likely to undergo a single electron reduction and become a more reactive superoxide anion. Reactive oxygen species are formed when molecular oxygen is partially reduced, as shown in the reaction below (91).

Reaction 1.1.

− • ROS include O2 , H2O2 and the highly-reactive hydroxyl radical ( OH). The

1 excited singlet oxygen ( O2, generated in photosynthetic organisms when subjected to 4 photo-oxidative stress) and hydroperoxyl radicals (•OOH) are some other forms of reactive oxygen species. ROS are much better oxidants than O2 and can readily oxidize and damage a wide range of biomolecules in the cell, leading to oxidative stress. The term oxidative stress can be defined as an imbalance between ROS exposure and the detoxification capacity of the cellular antioxidant system (77). The aerobic metabolism generates ROS as by-products, making oxidative stress an inevitable result of life in an oxygen-rich environment. In humans, oxidative stress is thought to be linked to aging

(79), cancer (32), and several other important health conditions, such as Alzheimer’s disease (117, 192) and Parkinson’s disease (194). Therefore, for organisms that depend on oxygen functioning as the terminal electron acceptor in oxidative phosphorylation, oxygen is both required for their existence and is detrimental to their well-being (the

“Oxygen Paradox”).

1.2.2. Sources of ROS in the Cell

Electrons that inadvertently escape from the electron transport chain can reduce

− O2 to form O2 and H2O2. Moreover, hydrogen peroxide can oxidize cytoplasmic ferrous ion, Fe2+, to generate the •OH radical via Fenton’s reaction (57, 100) (Reaction 1.2.).

2+ 3+ • Fe + H2O2 —> Fe + OH + OH–

Reaction 1.2.

• In fact, in vitro OH and H2O2 are produced in respiring E. coli membrane preparations when incubated with NADH (92, 124). In vitro, flavins were found to be the primary sites

− of H2O2 and O2 formation in the respiratory chain and a few other pathways (124).

Surprisingly, it was later discovered that respiratory flavoproteins do not play a major role in generation of H2O2 in vivo (102). Imlay and colleagues, have found that the flavin-

5 dependent dehydrogenase, NadB, which is involved in nicotinamide biosynthesis, accounted for a quarter of endogenous H2O2 (102). They also showed that menaquinone autoxidation was the source of another 5–10% of endogenous H2O2 (102). However, the sources for the remaining two-thirds of H2O2 have not been identified yet. Similarly,

− respiration was found to not be a significant source of the intracellular O2 , but auto- oxidation of the flavin-containing enzyme, fumarate reductase, was identified as the major contributor (90).

1.2.3. Targets of H2O2

Due to its uncharged nature, H2O2 easily passes through the cell membranes

• (150). Virtually all modes of toxicity of H2O2 is via the production of OH by the Fenton reaction (Reaction 1.2.). These hydroxyl radicals are powerful oxidizing agents and can attack a variety of biomolecules at nearly diffusion-limited rates (9). Besides damaging

DNA, •OH causes oxidation of guanosine in RNA (58), damages proteins containing Fe-

S clusters (97) and induces cell membrane damage by means of lipid peroxidation in higher organisms (142).

1.2.3.1. DNA damage caused by H2O2

The structure of the DNA bases, with their labile hydrogens that can be abstracted

(rate constant of the reaction is 2 × 109 M−1 s−1) and double bonding, which is prone to free radical addition (occurring at diffusion-controlling rates with rate constants 3 - 10 ×

109 M−1 s−1), make them particularly susceptible to •OH mediated damage (185).

Damaged bases are removed by DNA N-glycosylases, leading to formation of abasic sites. The •OH radical can also abstract hydrogens from the deoxyribose sugar units leading to the formation of carbon-centered sugar radicals (185). Oxygen can add to these

OH-adduct base radicals and carbon-centered sugar radicals to give peroxyl radicals, at rates limited by diffusion (185). These radicals can readily attack DNA leading to the formation of modified sugars, abasic sites and breaks in the sugar-phosphate backbone by a variety of mechanisms.

•OH radicals yield C5-OH- and C6-OH-adduct radicals, by adding to the C5- and

C6-positions of pyrimidines, respectively (48, 185). Cytosine glycol and thymine glycol can be generated from the C5-OH-adduct radicals in two ways. Oxidation of these adducts to give 5-hydroxy-6-peroxyl radicals and subsequent addition of OH−, or addition of water and subsequent deprotonation can lead to the formation of cytosine and thymine glycols (48, 185). Thymine glycol in the template has been demonstrated to block DNA synthesis in vitro (88). Cytosine products can undergo dehydration or deamination, depending on reaction conditions. Cytosine glycol can be dehydrated to form 5- hydroxycytosine (5-OH-Cyt), or deaminated to yield uracil glycol (48, 49, 185). When cytosine glycol undergoes deamination followed by dehydration, it yields 5- hydroxyuracil (48, 49, 185). These uracil derivatives preferentially basepair with adenine instead of guanine, causing G:C to T:A transversions.

If oxygen is not present, reduction of 5-OH- and 6-OH-adduct radicals of pyrimidines, followed by protonation, leads to formation of 5-hydroxy-6-hydro- and 6- hydroxy-5-hydropyrimidines instead (38). The allyl radical of thymine is formed via the

•OH radical mediated abstraction of hydrogen from the methyl group of thymine (48, 49,

185). Oxidation of this radical yields 5-(hydroxymethyl)uracil and 5-formyluracil (38).

Addition of hydroxyl radicals to purines generates C4-OH-, C5-OH- and C8-OH- adduct radicals (132, 183). Upon dehydration, C4-OH- and C5-OH-adduct radicals

7 produce oxidizing purine(−H) • radicals (132, 183). The C8-OH-adduct can undergo one- electron oxidation or one-electron reduction, yielding 8-hydroxypurines or formamidopyrimidines, respectively (132, 183). Guanine is especially vulnerable to oxidation, due to its low redox potential, which gives rise to products like 7,8-dihydro-8- oxo-2′-deoxyguanosine (8-oxo-dG) (128). In the syn conformation, 8-oxo-dG can functionally mimic thymine, therefore, if not removed it results in G:C to T:A transversions.

•OH attack on the phosphodiester backbone can cause base displacement, along with oxidation and/or fragmentation of the deoxyribose sugar. DNA bases can be displaced by •OH attack at C1', which oxidizes C1' to form deoxyribonic acid (50). This can also happen if C4' is attacked instead, forming 4-keto-deoxyribonate (16, 50). C4' oxidation is also implicated in fragmentation of the deoxyribose sugar (44).

In the absence of oxygen, a unique reaction is observed with the C5’-centered radical of the sugar moiety in DNA, which is formed by the •OH induced abstraction of H atom at C5’. This C5’-centered radical can undergo addition to the C8-position of the purine ring in the same nucleoside causing intramolecular cyclization and yielding 8,5′- cyclopurine-2′-deoxynucleosides (46, 47, 143). These compounds represent tandem lesions, as they arise due to concomitant damage to both base and sugar. However, prior oxidation of the C5′-centered sugar radical prevents this cyclization (46, 47, 143).

1.2.3.2. RNA damage caused by H2O2

Similar to DNA, RNA oxidative damage can lead to formation of modified bases and sugar, abasic sites and strand breaks (96, 115, 144). 8-hydroxyguanosine (8-oxo-G) is the most common oxidized base found in the RNA, because of this and its mutagenic

8 nature, it has been the focus of studies on oxidative stress induced RNA damage. Cells have vastly more RNA than DNA, which makes RNA a major target of hydroxyl radical mediated oxidative damage. In fact, in eukaryotic cells, the normalized 8-oxo-G content in RNA was up to 20 times that of 8-oxo-dG in DNA (85). However, compared to oxidative DNA damage, information on RNA oxidation is scarce, perhaps, owing to the misconception that because RNA is turned over rather quickly, oxidized RNA should not pose much of a threat. Unfortunately, when subjected to oxidative stress, oxidation of

RNA is induced within a few minutes, whereas the half-lives of most human mRNAs is about 10 hours, which could provide the oxidized RNA plenty of time to exert its deleterious effects. Moreover, ribosomal and transfer RNAs, which when taken together form the majority of cellular RNA species, are not even degraded during exponential phase of growth in micro-organisms (191). Having recently recognized this error in reasoning, researchers have conducted a number of studies on oxidation of RNA in eukaryotes and have found a correlation between high levels of RNA oxidation and age- related neurodegenerative disorders like Alzheimer's disease, Parkinson's disease and amyotrophic lateral sclerosis (129).

Still, our understanding of RNA damage under oxidative stress conditions in E. coli remains lacking. E. coli cells challenged with H2O2 exhibit a swift rise in the level of

8-oxo-G in RNA, in a dose-dependent manner (116). Under normal conditions (low

H2O2), about three times as much 8-oxo-G was found to be present in non-ribosomal

RNAs than in ribosomal RNA, suggesting that the ribosomal RNA is better protected from oxidization (116). Nevertheless, when exposed to high concentrations of exogenous

H2O2, the ribosomal RNA was found to be equally susceptible to oxidation, since the

H2O2 induced 8-oxo-G was found to be distributed equally across all RNA species (116).

The E. coli RNA polymerase can discriminate between 8-oxo-G and normal G and incorporates it about ten times slower, thereby reducing the possibility of its incorporation into RNA (184). Another means of defense against 8-oxo-G is the enzyme polynucleotide phosphorylase, which is a 3’-5’ exoribonuclease involved in degradation of mRNA and stable RNA (34). It exhibits higher binding affinity to 8-oxo-G containing

RNA than to undamaged RNA, and therefore, has been suggested to preferentially degrade oxidized RNA. The MutT protein was also shown to protect RNA against 8-oxo-

G incorporation (164). The MutT protein is well-known for interception of 8-oxo-dGTP in order to prevent its incorporation into DNA (119). However, MutT degrades 8-oxo-

GTP even faster than 8-oxo-dGTP (164). These mechanisms of cleaning the nucleotide pools decreases the possibility of transcription errors due to RNA or RNA precursor oxidation.

1.2.3.3. Protein oxidation

The level of oxidized proteins in the cells increases with age. Protein oxidation comprises a variety of nonspecific damage: from modification of amino acids and carbonylation to cleavage of the polypeptide chain and formation of cross-linked protein- protein, protein-lipid and even protein-carbohydrate aggregates (26). In E. coli, exposed iron-sulphur clusters in certain proteins like dihydroxyacid dehydratases are especially vulnerable to •OH attack (107). These Fe-S clusters are responsible for substrate binding and stabilization. Therefore, when the •OH radicals oxidize these iron atoms, they inactivate the enzymes (59). While most Fe-S proteins do not contain exposed Fe-S

10 clusters, even such proteins have been demonstrated to be prone to H2O2 damage if subjected to prolonged treatment (127). This is attributed to the ability of •OH to damage the Isc system, which is involved in assembly of iron-sulfur clusters in the cell. H2O2 has also been shown to inactivate mononuclear iron enzymes, like deaminases, deformylases, dehydrogenases and epimerases (8, 156). Apart from directly damaging proteins,

H2O2 was found to inactivate the Fur (ferric uptake regulator) regulon under certain conditions (180). Although the mechanism behind it remains unclear, since Fur is responsible for maintaining iron homeostasis in the cell, its repression would contribute significantly to the aggravation of H2O2 toxicity in the cell.

1.2.3.4. Lipid peroxidation

Lipids can also serve as targets of H2O2 induced oxidative stress. Lipid peroxidation can be initiated by •OH radicals attacking the polyunsaturated fatty acids in the phospholipid bilayer membrane (134). It causes decrease in membrane fluidity, which can disrupt the membrane-bound proteins. Lipid peroxidation can also lead to the formation of reactive aldehydes with long half-lives that can exacerbate the situation by damaging other biomolecules (134). However, most bacteria, including E. coli, contain only monounsaturated and saturated fatty acids in their membranes (20), therefore, making them immune to lipid peroxidation.

1.3. IRON METABOLISM AND TOXICITY

1.3.1. Iron-Containing Enzymes and Their Distribution in Metabolism

Iron is a ubiquitous metal in the cell, found as a critical cofactor in a variety of proteins. It participates in various critical metabolic processes such as the trichloroacetic acid cycle, oxidative phosphorylation, or DNA synthesis. Iron, being a transition metal

11 with an unfilled d-orbital, can exist in several oxidation states, from -2 to +6. This property enables it to act as a prosthetic group in various enzymes catalyzing redox reactions in the cell. In heme-based enzymes like catalase, iron aids in catalysis by being oxidized to porphyrin- Fe4+ from the porphyrin- Fe3+ before being recycled back (74). In the mononuclear enzymes, such as the Fe-superoxide dismutase (Fe-SOD) or ribonucleotide reductase, NrdAB, which depend on iron for catalytic activity, iron functions as a prosthetic group by cycling between the ferric (Fe3+) and ferrous (Fe2+) forms (39, 178). In addition, the Fe-S clusters of the enzymes in the electron transport chain, such as NADH dehydrogenase II, facilitate electron transfer due to the difference in their reduction potentials (89). Since iron also has a coordination number of six, it is capable of binding to a wide variety of ligands (up to six at a time). This property enables it to participate in non-redox-based reactions as well in enzymes such as dehydratases, by assisting in substrate binding and coordination, and serving as an electron sink by providing a local positive charge (89).

1.3.2. The Iron Paradox

When ancient life evolved on an anaerobic Earth about 4 billion years ago, the reduced ferrous form of iron was abundant. In combination with the properties of iron mentioned above, this lead to the wide-spread use of iron in various enzymes. However, oxygenation of the atmosphere lead to iron becoming both poorly available outside the cell and potentially toxic inside the cell. Ferrous iron levels in the environment became limiting due to their oxidation to the ferric form, which is almost insoluble under neutral pH. At the same time, iron-containing proteins became natural targets of oxidative attack.

ROS-induced oxidation of iron not only inactivates iron-containing proteins, but also aids

12 in generation of more ROS species via the Fenton reaction via release of iron, thereby aggravating and multiplying the damage. In fact, the toxicity of hydrogen peroxide (at concentrations less than 20 mM) depends on the presence of iron in the cells. H2O2 cannot directly damage DNA, to do this it needs to be converted into the highly-reactive hydroxyl radical species via the iron-dependent Fenton's reaction.

Since iron is both essential for survival, and at the same time, potentially dangerous, organisms have evolved elaborate ways to handle and distribute intracellular iron. The need to efficiently scavenge iron from the environment to maintain proper cellular functioning must be balanced with management of cellular iron levels to protect the cells from iron-induced ROS toxicity. There are three strategies employed by E. coli that ensure effective iron homeostasis:

1. High-affinity iron import systems and siderophores that aid in iron scavenging

from the environment.

2. Maintaining intracellular iron depots, which provide cellular processes with iron

when it is limiting in the surroundings, while collecting the surplus iron for

storage in a chemically-inert Fe3+ form.

3. A comprehensive iron-responsive regulatory system that manages the

mechanisms described above according to the iron needs of the cell and iron

availability in the environment.

1.3.3. Iron Scavenging in E. coli

E. coli excrete siderophores (highly specific and efficient low molecular mass

Fe3+ chelators) to scavenge and solubilize mineralized ferric iron in their environment

(186). Aerobically growing E. coli can employ at least five different siderophores for this

13 purpose: enterobactin, aerobactin, ferrichrome, coprogen, and citrate (186). These iron- bound siderophores are recognized and transported into the cell via specific membrane transport systems. Since siderophores have significantly lower affinity for Fe2+, the iron from the Fe3+-siderophore complex can be released via its reduction to the ferrous form inside the cells. Interestingly, reduced flavins, produced via the flavin reductase enzyme,

Fre, perform this function quite efficiently by transferring its electrons to Fe3+- ferric citrate complexes, or even to the iron depots like ferritins (40, 62, 122).

1.3.4. The Iron Depots of E. coli

The iron storage proteins maintain intracellular depots of iron which can be accessed to promote growth when iron is scarce in the environment. In E. coli, there are three classes of these proteins: ferritins that are found in almost all living organisms including higher eukaryotes, the heme-containing bacterioferritins and the much smaller

Dps proteins. These iron storage proteins contain 12-24 subunits, each folded together into four α helix bundles that can assemble into a hollow spherical protein shell (7). This spherical shell takes up ferrous iron and oxidizes it via its ferroxidase activity to store it in its central cavity in the ferric form, building up a ferrihydrite, or an amorphous ferric phosphate core, depending on phosphate availability (7). From an evolutionary stand- point, these structural and functional similarities between the iron storage proteins suggest that they share a common ancestry.

1.3.4.1. The DNA binding, iron detoxification and storage protein, Dps

The compact spherical shell-like structure of E. coli Dps, with its hollow cavity at the center (approximately 40–50 Å in diameter), which can complex about 500 molecules of iron at a time, is assembled from 12 identical monomers, each of them containing a

14 protruding lysine-rich N-terminus (76). The ferrihydrate core is connected to the surface of this sphere by means of several hydrophilic and hydrophobic channels that form at the junctions between subunits. Fe2+ ions utilize the histidine and glutamic acid residues in the hydrophilic channels to enter the core (76). There are also twelve putative ferroxidase sites found in Dps, which are located at the six interfaces of its adjacent subunits (76).

Each ferroxidases site is thought to contain a high-affinity iron-binding site A along with a low-affinity site B (76). Unlike ferritin, at these sites in Dps, Fe2+ ions are oxidized by reducing H2O2, thereby consuming H2O2 in the process and making it unavailable for the

•OH generating Fenton’s reaction. Dps can use oxygen for this process, like ferritins, but this O2-driven oxidation of iron occurs at a much slower rate than that in the presence of

H2O2 (198). Therefore, Dps hinders the DNA-damaging Fenton’s reaction in two ways: by limiting the Fe2+ ions available via oxidation and storage and by its ability to reduce hydrogen peroxide in its core.

Additionally, unlike most iron storage proteins, Dps binds DNA non-specifically

(188). Researchers have demonstrated that the formation of large DNA–Dps crystal lattice complexes in the stationary phase, some of which have been found to be several hundred nanometers long, is driven largely by self-aggregation of Dps molecules (188).

During this process, the three adjacent dodecamers create holes lined by their lysine-rich

N-terminal domains, which are positively charged at physiological pH, and therefore, are believed to aid in threading the DNA through the dodecamer holes (31). Stationary E. coli cells can accumulate up to ~180,000 Dps molecules (~6000 in exponential phase), which bind chromosomal DNA to form these stable Dps-DNA co-crystals, thereby protecting the DNA from diverse environmental assaults, including oxidative damage (3,

68). Besides the stationary phase, Dps production is induced under conditions of oxidative stress and starvation (4).

1.3.4.2 Ferritin - The primary cellular iron storage protein

Ferritin A (FtnA) serves as the major iron reservoir in E. coli and is capable of sequestering up to 4,000 iron atoms in its center (87). Structurally, its hollow spherical shell, with its iron storage central cavity, is composed of 24 subunits of four α-helix bundles each (87). It contains three iron binding sites per subunit: two of these sites are found in the ferroxidase center, which uses oxygen to oxidize Fe2+ ions to store it as ferric phosphate, whereas the third site appears to modulate Fe2+ binding to the ferroxidase center (87). In the ferritin-deficient strain, ftnA, cellular iron content in the stationary phase was reduced by about 50%, while the growth-rate was reduced under iron-limiting conditions (2). These observations are in line with the putative role of FtnA as an iron distributor, accumulating excessive iron, so as to serve as an intracellular iron reserve, while releasing the stored iron to promote cell growth in an iron-deficient environment.

In addition, ferritin has been demonstrated to be capable of binding DNA, both in vivo and in vitro (163, 170).

E. coli possesses an additional heme-containing 24 subunit ferritin called bacterioferritin, which is distantly related to ferritin, and therefore, exhibits several structural and functional similarities with it. In bacterioferritin, the 12 protoporphyrin IX heme groups are found within the inner surface of the protein shell at each of the 12 interfaces between the 24-mer subunits (193). Each of the 24 subunits contain a dinuclear-iron binding ferroxidase center (113). Recent work has shown that, like Dps,

2+ bacterioferritin uses H2O2, not O2, to oxidize Fe ions, thus preventing toxic hydroxyl

16 radical generation via Fenton chemistry (21). However, bacterioferritin proteins do not appear to play a major role in iron storage. The bfr mutant was found to be phenotypically the same as wild-type cells, so the physiological significance of bacterioferritin remains unknown (2).

1.3.5. Regulation of Iron Homeostasis by Fur

In E. coli, the ferric uptake regulator protein, Fur, regulates iron homeostasis by controlling the expression of over 90 genes in an iron-dependent fashion (78). Fur is a homodimer composed of 17-kDa subunits capable of binding a ferrous ion each (41). The

N-terminal of each subunit is involved in DNA-binding, while the histidine-rich C- terminus is believed to bind Fe2+ ions and facilitate dimerization (41, 161). Fur acts as a positive repressor, that is, when iron is abundant in the environment, Fur complexed with

Fe2+ represses transcription of genes involved in iron acquisition, whereas when iron is insufficient, Fur is inactivated and the repression subsides (7). Although other transition metal ions, such as Mn2+, can also bind Fur in vitro, this is not physiologically relevant, because of their lower intracellular concentrations (11). Fur is self-regulating; in exponential phase cells, up to 5,000 copies of Fur can be found, while stationary cells contain approximately 10,000 copies (200).

Under iron-limiting conditions, Fur also causes replacement of some Fe- containing proteins with alternatives that do not require iron to function, to conserve iron usage. For example, Fe2+–Fur can repress the Mn- superoxide dismutase (SOD) gene, sodA and it can indirectly induce the Fe-SOD, sodB, therefore, when iron is restricted, the

Mn-SOD replaces Fe-SOD (51). Similarly, fumarases A and B, both of which contain four Fe-S clusters and are inducible by activated Fur, can be replaced by the non-iron-

17 metallated fumarase C, which is otherwise repressed by Fe2+–Fur in iron-sufficient conditions (137). Additionally, Fur can also aid in combating iron deficiency by relieving the Fe2+–Fur triggered repression of the only manganese transporter in E. coli, mntH

(138). The imported Manganese can be used to synthesize alternatives to Fe-containing enzymes, such as the Mn-SOD and the Mn-dependent ribonucletide reductase, NrdEF

(181).

Besides regulating iron uptake genes, Fur transcriptionally regulates a host of other activities including methionine biosynthesis (160), oxidative stress response (167), the tricarboxylic acid cycle (175), glycolysis and gluconeogenesis (181), respiration and purine metabolism (160). Moreover, several genes like acnA, bfr, ftnA, sdhCDAB, fumABC and sodB, promoters of which do not contain a Fur binding site (with the exception of ftnA), can be induced by activated Fe2+–Fur indirectly via the small regulatory RNA, RyhB (120). The RyhB sRNA promotes degradation of mRNAs of non- essential iron-containing enzymes so as to redirect iron for use in cellular activities that are indispensable for survival and growth, such as dNTP synthesis, when conditions are iron-restricting (121). This activity has been termed “iron sparing”. RyhB, in turn, requires the RNA chaperone, Hfq, which prevents its degradation by RNase E (121).

Activated iron-bound Fur has been found to repress the ryhB gene, which prevents the mRNA of such non-essential iron-containing proteins from being degraded, therefore, up- regulating their expression. RyhB can also repress Fe-S cluster assembly by the Isc system by targeting iscSUA mRNA for degradation (121). The resulting lower Fe-S cluster pool causes the sensor IscR (containing no Fe-S clusters itself) to promote transcription from the suf operon, which is more efficient at synthesizing Fe-S clusters

18 when iron is insufficient (121). Therefore, by repressing rhyB Fe2+–Fur can indirectly promote assembly of Fe-S clusters by the Isc system under iron-replete conditions, in addition to being able to directly repress transcription of the suf genes (139).

1.4. FREE RADICAL STRESS RESPONSES AND SCAVENGERS

E. coli has evolved sophisticated mechanisms to monitor the levels of intracellular

ROS so as to activate global antioxidant OxyR and SoxRS responses when required, when ROS levels rise. The transcription factor, OxyR, which is the primary global H2O2 sensor, upon activation induces H2O2 scavenging enzymes, like catalases and alkyl- hydroxy peroxidases, as well as iron homeostasis proteins such as Dps and Fur (43). The

SoxRS regulatory system mounts a defense against redox-cycling drugs (like paraquat) and against nitric oxide (NO) when activated via oxidation of the two Fe-S clusters in

SoxR (71, 174). The genes induced by SoxRS response include sodA, which codes for the

Mn-SOD, nfo, which synthesizes endonuclease IV that participates in base excision repair, and fur (162).

1.4.1. The OxyR Response

The transcriptional oxidative stress regulator, OxyR, is a 34kD protein that belongs to the LysR family of transcription factors (35). In solution, OxyR exists as a homotetramer (106). Its N-terminal domain, which contains a helix-turn-helix motif for

DNA binding, is connected by a flexible linker to the regulatory C-terminal domain, which is responsible for both oligomerization and activation since it contains the two redox active cysteines. OxyR, is activated by low as 100 nM H2O2 via oxidation of these two Cysteine residues, Cys199 and Cys208, to form a disulfide bond (106). These residues enable OxyR to act as a reversible redox switch. Additionally, OxyR can also be

19 turned on by nitric oxide (NO) released from nitrosothiol compounds via S-nitrosylation

(81). When turned on, OxyR induces several scavenger proteins (KatG, KatE, AhpC), iron homeostasis proteins (Dps, Fur) and manganese uptake by the MntH importer, which aids in Mn replacement of Fe in mononuclear enzymes, consequently protecting them from Fenton damage (141). In these genes, OxyR recognizes and binds to a DNA motif containing four ATAG nucleotide elements spaced 10-bp apart from each other and interacts with the C-terminal domain of the α-subunit of RNA polymerase to stimulate transcription (172). OxyR also induces synthesis of glutathione and glutaredoxin that can reduce the disulfide bond in the activated OxyR and return it to the inactive form, making the system auto-regulatory (199). However, oxidation of OxyR is much faster than its reduction, which permits transient activation in the reducing environment of the cell.

Interestingly, OxyR can also act as its own repressor in both oxidized and reduced forms

(35).

OxyR activates oxyS as well, which codes for the small noncoding OxyS RNA

(5). OxyS expression reduces spontaneous or chemically induced mutagenesis (5). It is also involved in regulating the metabolically-generated, endogenous H2O2 levels (65).

OxyS represses a number of genes including the alternative sigma factor, RpoS and fhlA, which stimulates synthesis of the formate hydrogenlyase complex containing several metal cofactors that are thought to be capable of mediating the H2O2-driven oxidative damage (6, 197). Repression of fhlA is at the translation level, since OxyS binds to fhl mRNA and prevents ribosome assembly on it, inhibiting protein synthesis as a result (6).

However, mechanism of rpoS repression remains unclear, even though recent findings

20 have revealed that there is a weak yet direct interaction between OxyS and the rpoS mRNA (60).

The level of oxyR expression and activity varies with endogenous H2O2 levels, which depend on the growth rate of the cell. During the early exponential phase of growth, aerobic respiration increases the intracellular H2O2 levels, this activates the

OxyR regulon and results in synthesis of scavenging catalases and peroxidases that can keep H2O2 levels in check (66). Additionally, oxyR is upregulated by the cyclic-AMP- activated protein, Crp, during exponential phase, while being downregulated by RpoS in the stationary phase (66).

1.4.1.1. H2O2 scavengers: catalases and peroxidases

E. coli has two distinct catalases- KatG and KatE. The bifunctional hydroperoxidase I (HPI), KatG, is both a catalase and a peroxidase, while the KatE enzyme (or HPII) has only catalase activity (36, 37). These two enzymes are the primary

H2O2 scavengers of the cell, specifically, when the intracellular H2O2 concentrations are high (> 20 μM and above) (150). In the catalase reaction, two H2O2 molecules are broken down into oxygen and water. In order to achieve this, first the ferric heme reduces

H2O2 to H2O, generating the ferryl porphyrin cation radical, which is returned to its ferric form via oxidation of the second H2O2 molecule to O2 (Reaction 1.3.) (84).

3+ 4+ H2O2 + Fe -E → H2O + O= Fe -E(.+)

4+ 3+ H2O2 + O= Fe -E(.+) → H2O + Fe -E + O2

Reaction 1.3. (Here Fe-E represents the iron center of the heme group.)

In addition, if H2O2 concentrations are low the peroxidase activity of KatG can also catalyze the first step of the reaction producing H2O from H2O2, while using a

21 suitable donor other than H2O2 for the second step (84). The H2O2 produced endogenously (if [H2O2] < 20 μM) as a result of aerobic metabolism is scavenged by the alkyl-hydroxy peroxidase complex AhpCF (149). All three scavengers are upregulated by

OxyR during oxidative stress (64). The Hpx- mutants lacking all the H2O2 scavengers

(ahpCF katG katE) struggled to grow under aerobic conditions (149).

The KatG enzyme contains two large peroxidase-like domains that are believed to be a consequence of gene duplication (195). Its bifunctional heme-bound active site is found in the N-terminal domain, whereas the C-terminus, despite its inability to bind heme or catalyze the disproportionation of H2O2, is indispensable for KatG functioning

(13, 195). Recent findings suggest that the C-terminal domain may act as a scaffold, on which its I'-helix can direct the folding of the N-terminal domain (12). In addition to being stimulated by OxyR, katG expression increases as the cells enter stationary phase in an RpoS-independent manner (126). Indeed, overexpression of katG has been found to suppress H2O2 sensitivity in oxyR mutants, while ahpCF katG mutants were found to be hyper-sensitive to increased endogenous H2O2 levels (70, 136).

The crystal structure of KatE has revealed it to be a homotetrameric enzyme with each subunit containing a unique cis-heme d group (131). This heme d is generated by

KatE-catalyzed hydroxylation of proheme IX prosthetic group using H2O2 as a substrate

(131). Similar to KatG, both C and N-terminal domains are required for the enzyme to be functional (153). The C-terminal domain of KatE lacks enzymatic activity but has been shown to contribute to its stability by offering increased resistance to proteolytic cleavage

(33). The N-terminus comprises of the heme d groups containing active sites, which are buried deep within the enzyme, and therefore, require various channels for substrate and

22 product transport (152). KatE synthesis is stimulated in stationary phase in an RpoS- dependent fashion and by hyperosmotic stress along with H2O2-induced OxyR response

(166). katE over-expression has also been demonstrated to suppress H2O2 sensitivity of the oxyR mutant (70).

In the two-component alkylhydroperoxide reductase system, AhpCF, AhpC acts as the peroxidase, while the NADH-reducible flavin component, AhpF, reduces AhpC back (91, 149). Endogenous H2O2 is scavenged by AhpC, resulting in the oxidation of its

Cys46 residue in the N-terminus and the subsequent formation of a disulfide bond between Cys46 and Cys165 (54, 149). The Cys46 is suggested to be the peroxidatic

Cysteine, since if it is mutated to serine, the enzyme loses its peroxidase activity (54).

1.4.2. The SoxRS Response

The SoxRS response protects E. coli from superoxide-generating drugs, such as paraquat and menadione and nitric oxide via two distinct mechanisms (69, 130). The

SoxR dimer contains four Fe-S clusters at its C-terminal domains, oxidation of which, by the redox-cycling drugs mentioned above, activates the SoxR protein (82). The activated

SoxR can upregulate transcription of the soxS gene (83). The NO dependent activation of

SoxR, on the other hand, depends on nitrosylation of its Fe-S clusters (45). EPR analysis has revealed that NO reacts with SoxR to form dinitrosyl– Fe-S clusters (45). This nitrosylated SoxR is equally capable of inducing soxS transcription (45).

The SoxS protein activates transcription of over a hundred target genes by aiding in the recruitment of RNA polymerase (RNAP) to their promoters by binding to the C- terminal domain of the α-subunit of RNAP (154). The SoxRS regulated proteins defend the cells from Fenton’s reaction mediated oxidative damage through a variety of

23 mechanisms including: induction of Fur, turning on efflux pumps to excrete the superoxide-generating drugs out (acrAB), reducing back the oxidized iron in various enzymatic prosthetic groups (fldA, fldB, fpr), stimulating repair of damaged bases via endonuclease IV (nfo) (141). In fact, the SoxRS response confers an inducible resistance to superoxide-generating drugs if cells are exposed to sub-lethal doses prior to exposure to killing doses (71, 174). When the drug (or NO) treatments are abated, the SoxRS response ends by reduction of SoxR and by proteolytic degradation of the SoxS protein

(72).

− 1.4.2.1. The O2 scavengers: SodA, SodB and SodC

Superoxide dismutases (SODs) can detoxify endogenous superoxide by catalyzing dismutation of superoxide into hydrogen peroxide and molecular oxygen (Reaction 1.4.).

(n+1)+ − n+ M -SOD + O2 → M -SOD + O2

n+ − + (n+1)+ M -SOD + O2 + 2H → M -SOD + H2O2.

Reaction 1.4. (Here M represents a transition metal that is either Mn, Fe, Cu or Ni.)

E. coli contains three different SODs, each of them has a different transition metal cofactor at its active site: SodA is the manganese-containing SOD, SodB contains iron and SodC has both copper and zinc (23). The MnSOD and the FeSOD are structurally very similar. They are both dimer proteins composed of 23kDa subunits with their active sites located at the dimer interface, which is accessible to substrates by means of channels

(52). Their amino acid sequence is 43% identical, this includes the metal binding residues: three histidines and one aspartic acid and a conserved glutamic acid residue near the active site with which the metal-bound solvent molecule forms hydrogen bonds

(52). However, both the MnSOD and FeSOD are highly specific for their respective

24 metal cofactors: if Mn is substituted with Fe, or vice versa, the enzymes are catalytically inactive (179). Moreover, there are functional differences between how MnSOD and

− FeSOD combat O2 stress. MnSOD can bind DNA non-specifically and is more effective than FeSOD in protecting DNA from oxidative damage (86). On the other hand, FeSOD was found to be better at protecting the cytoplasmic enzyme, 6-phosphogluconate

− dehydrogenase, against O2 attack, while they were both equally effective at protecting the four Fe-S clusters containing class of enzymes, dihydroxy-acid dehydratases (24, 86).

Additionally, both the MnSOD and FeSOD were also found to be a part of the ribonucleoside-diphosphate-reductase-activating system (39, 53).

Both soda and sodB mutants exhibit higher sensitivity and an increased mutation

− rate in response to the O2 generating drug, paraquat, than wild-type cells (29, 55). The sodA sodB double mutant is even more so (94). Under aerobic conditions, this double mutant can grow slowly on rich medium, but not on minimal glucose medium, unless it is supplemented with the 20 amino acids (29). This amino acid auxotrophic phenotype is

− observed because several amino acid biosynthetic pathways depend on O2 sensitive enzymes such as dihydroxy acid dehydratases and transketolases (18, 24). Additionally, the sodA sodB mutant was also shown to be more susceptible to H2O2 induced DNA damage (98).

sodA expression is tightly regulated by the aerobic-anaerobic gene regulation system controlled by the FNR and ArcA transcription regulators and the Fur regulon (80).

FNR aids in transition from aerobic to anaerobic growth by activating genes involved in anaerobic metabolism and repressing genes involved in aerobic metabolism (148). Under anaerobic conditions, ArcA, when phosphorylated, primarily acts as a negative regulator

25 by repressing transcription of several genes including those involved in respiration and carbon oxidation pathways (75). Both FNR and ArcA actively repress sodA expression under anaerobic conditions (80). However, since under aerobic conditions both FNR and phosphorylated ArcA are inactivated, sodA is activated (178). In addition, as discussed previously, both Fur and SoxRS can regulate transcription of SodA. When intracellular iron levels rise, the activated Fe2+–Fur represses transcription of SodA; this repression is lost if iron is deficient in the cells, thereby inducing sodA, while the SoxRS response upregulates sodA upon exposure to redox-cycling drugs or NO (56, 162). On the contrary, sodB is constitutively expressed under aerobic and anaerobic conditions, in addition to being induced by Fe2+–Fur via RyhB (121). Therefore, under conditions of iron sufficiency the cell has more FeSOD than MnSOD.

SodC, is the CuZnSOD and it is the only periplasmic SOD of E. coli, as both the

MnSOD and FeSOD are cytoplasmic (17). It has been reported that SodC dismutates the

− O2 produced in the periplasm as a result of adventitious oxidation of dihydromenaquinone in the cytoplasmic membrane, thereby protecting the periplasmic proteins from oxidative damage (101). SodC is a monomeric protein, a reason for this could be the presence of charged residues instead of hydrophobic ones in the structure of the protein, which would typically act as a dimer interface (15). Crystal structure of SodC revealed it to have a distorted antiparallel β-barrel fold with Cu2+–Zn2+ connected to the bridging His61 residue by an unusually long bond at its active site (140).

SodC expression is induced in the stationary phase by RpoS, while its expression is repressed during anaerobic growth by FNR (67, 95). In fact, when wild-type cells enter stationary phase become acutely sensitive to H2O2 for the next several hours and this

26 sensitivity is further enhanced in a sodC mutant, suggesting that SodC plays a role in

− protecting the cells from O2 induced damage and death during this period (95).

Interestingly, since the SodC enzyme was found to be resistant to hydrogen peroxide inactivation, it has also been implicated in protecting E. coli from macrophage attack

(14).

1.5. MECHANISMS OF CHROMOSOMAL FRAGMENTATION AND REPAIR

Oxidative damage is known to cause chromosomal fragmentation, indicating formation of double-strand DNA breaks. As already discussed above (in 1.2.3.1), oxidative damage to DNA results in various base modifications, generation of abasic sites and single-strand interruptions. The base modifications and abasic sites and are substrates for the base excision repair (see below), both of which go through one-nucleotide single- strand gap intermediate. In other words, the majority of oxidative DNA lesions are, or are processed via, single-strand DNA interruptions. Therefore, occasional double-strand breaks induced by oxidative damage were for a long time explained as a result of clustering of single-strand lesions (coinciding nicks in the opposite strands of the DNA duplex). Eventually, these simplistic models were surpassed by other models that took into account the fact that non-dividing cells are resistant to oxidative damage and experience no double-strand breaks. The new models linked the exclusively single-strand nature of oxidative DNA lesions with replication fork passage through the damaged region to propose formation of double-strand breaks as a result of replication fork disintegration at single-strand lesions.

1.5.1. Mechanisms of Replication-Dependent Chromosomal Fragmentation

There are three confirmed models of replication fork disintegration events that can lead to chromosomal fragmentation in E. coli via formation of single-strand gaps: (1) replication fork collapse, which results from a replication fork running into a single- strand interruption in the template DNA, (2) replication fork regress-split, in which the two template strands reanneal, forcing the two nascent strands into a duplex of their own and forming a Holliday junction that is resolved by RuvABC and (3) replication fork rear-ending that results from a replication fork running into a stalled replication fork of the previous round (Fig. 1.1.) (147).

The conditional polA and ligA mutants, due to their compromised abilities to repair single-strand gaps and nicks, are susceptible to chromosomal fragmentation due to replication fork collapse (99, 104, 110). We have found that the dut mutants also exhibit single-strand interruptions due to increased uracil incorporation and subsequent excision, which lead to double-strand breaks (DSBs) when encountered by replication forks (171).

Excision repair of DNA damage also increases the overall level of single-strand interruptions. For example, during nucleotide excision repair of UV-lesions transient single-strand gaps appear in the DNA (111). If the DNA is replicated while repair is taking place, these lesions lead to double-strand breaks due to replication fork collapse and can be detected as formation of subchromosomal fragments in neutral sucrose gradients, especially in repair-deficient cells (111). In both cases, the conversion of single-strand interruptions into DSBs depends on DNA replication, in the absence of which no chromosomal fragmentation is detectable (111). Importantly, the chromosomal fragments released as a consequence of replication fork collapse are susceptible to

RecBCD exonuclease V (ExoV) degradation. ExoV, is the primary exonuclease responsible for digesting duplex DNA, which has at least one double-strand end unprotected (111). Therefore, if recombinational repair is blocked in mutants of polA, ligA or dut by blocking RecA, or if a recA mutant is exposed to UV and then allowed to grow, substantial ExoV-dependent degradation of chromosomal DNA is observed along with loss of viability (99, 104, 111, 171).

In replication fork regress-split, inhibition of replication fork progress causes regression of the inhibited replication fork, which leads to its splitting via Holliday junction resolution and formation of double-strand break (111, 125). Unlike replication fork collapse, this mechanism does not depend on the presence of pre-existing lesions in the template DNA, as replication fork progress can be blocked by DNA-bound proteins, malfunctioning of the replisome enzymes, or by accumulation of positive supercoiling ahead of the fork (111, 125). Upon replication fork stalling and regression, annealing of nascent and template DNA strands gives rise to a Holliday junction (with an open-ended arm susceptible to ExoV degradation), resolution and cleavage of which by the RuvABC resolvasome leads to DSBs (111, 125). Since fragmentation here depends on RuvABC, no chromosomal fragmentation is observed in ruv mutants (125). Replication forks can be impeded by DNA gyrase inhibitors such as ciprofloxacin and nalidixic Acid, or by inactivating the primary and secondary replication fork helicases, DnaB and Rep, respectively (125). Therefore, when dnaB(Ts) mutants are deleted for recBC and shifted to semi-permissive temperature, thereby hindering replication fork progress, significant chromosomal fragmentation is observed (125). Similarly, in the absence of Rep, which facilitates the replication fork progress by removing proteins tightly-bound to template

DNA, the slower replication forks are prone to chromosomal fragmentation, which is revealed in the absence of double-strand break repair (125). These breaks can be repaired via the RecBCD-mediated RecA-dependent homologous recombination pathway in which the Holliday junction intermediates are resolved by RuvABC. Alternatively, in the absence of RecA, the exonuclease activity of the RecBCD complex can eliminate the

Holliday junction by degrading the open double-strand end of the regressed fork. Thus, cells suffering chromosomal fragmentation due to replication fork regress-split do not strictly require the recombination repair protein, RecA, for their survival (125). In contrast to RecA, RecBCD is essential in such cells since, in its absence, Holliday junction resolution of the reversed fork by RuvABC leads to generation of double-strand breaks in the chromosome, which require RecBCD for repair (125).

The third mechanism of fragmentation, replication fork rear-ending, requires a replication fork to run into a stalled replication fork from the previous round, thereby causing the newly synthesized DNA to be processed into sub-chromosomal fragments

(147, 155). This mechanism of fragmentation is observed in the dnaA(cos) mutants expressing constitutively-activated DnaA, the E. coli replication initiator protein, and therefore, are hyper-initiating, which results in multiple replication forks running into stalled forks (155). Additionally, in strains, in which Ter sites were placed in the blocking orientation in the middle of replichores, the bacterial chromosome were also found to exhibit chromosomal fragmentation due to replication fork rear-ending, as a consequence of newer replication forks colliding with stalled replication forks from previous rounds at these sites (155). It is interesting to note that, since the replication fork broken double- strand ends produced as a result of the collision can be effectively degraded by RecBC,

30 this mechanism of chromosomal fragmentation does not lead to DSBs, which could be fatal if unrepaired. Therefore, these mutants’ dependence on recombinational repair and the replication fork restart protein, PriA, for survival is puzzling (155).

1.5.2. Mechanisms of DNA Repair

1.5.2.1. Recombinational repair pathway

There are two ATP-dependent RecA mediated pathways of homologous recombination in E. coli: the RecBCD pathway, which is involved in the repair of double- strand breaks and the RecFOR pathway, responsible for repairing persisitent single-strand gaps (Fig. 1.2.). In the RecBCD pathway, recombination is initiated by binding of

RecBCD, the ATP-dependent helicase/nuclease complex, to the exposed ends of the

DSBs. This results in unwinding and subsequent degradation of the duplex DNA via the helicase activities of RecB (3' to 5') and RecD (5' to 3') and the exonuclease activity of

RecBC, respectively. As it progresses along the DNA, the RecBCD complex eventually comes to a pause upon encountering a Chi site, GCTGGTGG, recognized by RecC and then RecBCD begins to preferentially degrade the strand with the 5’ end, so as to generate a long, single-stranded 3’ overhang. In the next step, the RecBCD complex promotes the assembly of the RecA filament on the 3’ overhang. This RecA filament, upon searching and locating a homologous sequence in the chromosome, catalyzes strand invasion, in which the invading 3' overhang causes the displacement of the intact duplex

DNA strand and gives rise to the X-shaped Holliday junction structure. These Holliday junctions are primarily resolved via the RuvABC resolvasome complex through RuvAB mediated branch migration followed by RuvC catalyzed symmetrical double incisions to resolve the junction. In addition, translocation of Holliday junctions by the RecG helicase

31 also leads to resolution. Double-strand break repair is completed when DNA polymerase

I (PolA) fills in the remaining gaps and its ends are joined via the DNA ligase (LigA).

1.5.2.2. Base excision repair pathway

As the name suggests, the base excision repair pathway (BER) is involved in repair of non-canonical, modified, or mismatched bases in the DNA, which are generated as a result of attack by reactive chemical species or by high energy radiation. The first step in BER is excision of these damaged bases from the DNA by DNA glycosylases that cleave their N-glycosidic bonds producing abasic sites. Incisions are then made by abasic site endonucleases at the 5’-side of these abasic sites to initiate removal of these sites and subsequent repair via DNA polymerase I and ligase follows.

Oxidative damage gives rise to a host of modified DNA bases including 5,6- dihydrothymine and formamidopyrimidine, which are recognized and excised by the

DNA glycosylases Endo III and Endo VIII, as well as 8-oxo-G, which is recognized and cleaved by MutM (for a detailed discussion of ROS induced DNA lesions please refer to section 1.2.3.1.) (44). The abasic sites generated as a result of excision of these oxidized bases are nicked at their 5’ site by Exo III (XthA), the major 5' AP endonuclease in E. coli, or by Endo IV (NthA), which is followed by removal of these abasic sites by deoxyribophosphodiesterases in order to provide an appropriate substrate for DNA pol I.

1.5.2.3. Nucleotide excision repair pathway

In E. coli, nucleotide excision repair (NER) is performed by the UvrABCD complex, which removes a variety of DNA lesions including bulky adducts, DNA cross- links and UV irradiation-induced lesions such as pyrimidine dimers (PDs), 6-4 photoproducts etc. Repair is initiated by the heterotetrameric UvrA2-UvrB2 complex that

32 recognizes lesions like PDs and deposits the UvrB protein on them. UvrC recognizes this

UvrB-PD complex and forms the UvrBC complex. UvrC then nicks the PD-containing

DNA strand five nucleotides downstream of the PD and then makes a second nick eight nucleotides upstream of the PD. Finally, this PD-containing 13 bp DNA segment is unwound by the UvrD helicase, leaving the resulting gap to be filled in by DNA polymerase I and joined by DNA ligase.

1.6. POTENTIATED HYDROGEN PEROXIDE TOXICITY

1.6.1. The Concept of Potentiated Toxicity

When two viable single mutations are combined in a double mutant, the double mutant usually exhibits an additive combination of their phenotypes, but could also behave like one of the single mutants (epistasis). However, occasionally the double mutant turns out to be inviable and this phenomenon is referred to as co-lethality or synthetic lethality. Co-lethality is commonly attributed to enzymatic redundancy. In this scenario, there are two different proteins that can perform the same essential transaction in the cell, which is required for survival. Therefore, in the case of single mutants where only one such protein-coding gene is deleted, the protein synthesized by the other can substitute for it and perform the said transaction, preventing lethality. However, in the double mutant, where both genes are deleted, the essential transaction can no longer be carried out and the cells die. There are several examples of co-lethality due to an apparent enzymatic redundancy in E. coli such as uvrD rep, skp surA, degP surA and suf isc (1,

145, 165). In the case of uvrD rep co-lethality, both genes code for a related 3’ 5'

DNA helicases that aid in DNA replication or repair, hence the loss of one is compensated by the other helicase in the single uvrD or rep mutant, but a double mutant,

33 which lacks both, is dead (1). Similarly, there are two parallel pathways for chaperone activity in the periplasm, SurA is a component in one of them and both Skp and DegP are part of the other, which is why the double mutants, skp surA and degP surA, are not viable (145). Finally, as previously mentioned, both the Suf and Isc systems are involved in the synthesis and assembly of iron-sulfur clusters, so in the absence of both of these pathways the suf isc double mutant is unable to survive (165).

However, in the past, our lab has proposed an alternate explanation for the phenomenon of co-lethality, which holds that if the function of one protein is to avoid a potentially lethal damage, whereas that of the other is to repair this damage, then a double mutant which lacks both of these functions would not be able to survive (105).

Remarkably, the two participants of this co-lethal combination have no relationship to one another, as they belong to distinct areas of metabolism. In fact, the majority of co- lethal combinations fall under this category, which we have termed “avoidance-repair couples”, two examples of which are dut recA and seqA recA (105). The Dut gene codes for dUTPase, which degrades dUTP and keeps its levels within check in the cells, so as to reduce its incorporation into DNA. Therefore, mutating dut increases uracil incorporation in the DNA, which leads to recombinational repair and base excision repair dependence

(105, 171). Consequently, in the double mutants of dut with either recA, recBC or xthA, polA and ligA, absence of both mechanisms to prevent dUTP incorporation, on the one hand, and DNA damage repair, on the other, leads to co-lethality (171). Similarly, SeqA, which prevents over-initiation by sequestering the replication origin, oriC, between consecutive initiation rounds and also plays a role in chromosome segregation, was found to protect the chromosomal DNA from fragmentation (146). Deleting seqA leads to

34 increased chromosomal fragmentation and the recombination repair enzymes become essential for survival of the seqA mutants (105, 146). Thus, in co-lethality, one mutation may potentiate the effect of the other, thereby leading to cell death.

The phenomenon of synergistic toxicity or co-toxicity, in which the combination of two treatments that are weakly toxic individually is lethal to the cells, is analogous to co-lethality. Likewise, co-toxicity can be explained by either redundancy or potentiation mechanisms. If both treatments induce the same cellular damage, then their combination could kill the cells by saturating their capacity to counteract or repair the damage. At the same time, it could also be the case that one treatment potentiates the damage caused by the other, either by indirectly amplifying the damage caused, or by preventing repair of the damage, both of which could lead to cell death. One way to distinguish between these two mechanisms of toxicity would be to test if either of the treatments is individually toxic at higher doses. If the synergistic toxicity is due to redundancy, then either treatment would eventually overwhelm the cellular capacity to withstand the damage at sufficiently high doses. However, in the case of potentiated toxicity, only one treatment would be able to kill cells at higher doses, whereas the other, which functions as the potentiator, would be unable to do so. In the past, researchers have found that several individually bacteriostatic treatments such as nitric oxide, cyanide (CN) or an excess of cysteine when combined with a low millimolar growth-inhibiting concentrations of hydrogen peroxide kill E. coli. In the upcoming sections, I will discuss the mechanisms of their synergistic toxicities.

1.6.2. Mechanism of NO + H2O2 Co-Toxicity

In the human immune system, the phagocytic macrophages that engulf and destroy the invading pathogens form a crucial part of the innate immune response, which is the first line of defense. Phagocytosis of these pathogens induces two important antimicrobial pathways in macrophages: the NADPH phagocyte oxidase pathway that catalyzes the univalent reduction of molecular oxygen to generate superoxide, which can subsequently be reduced to H2O2 and the inducible nitric oxide synthase pathway, responsible for producing nitric oxide. NO can synergistically potentiate H2O2 toxicity, making their combination lethal for invading pathogens (133, 189). In fact, mutant mice, in which these pathways were inactivated, were killed by otherwise non-lethal doses of

Salmonella typhimurium (182). Moreover, macrophages isolated from these mutant mice were unable to suppress intracellular bacterial proliferation (182).

In E. coli, exposure to low millimolar concentrations of NO or H2O2, which are bacteriostatic individually, were found to cause two to four orders of magnitude killing in combination (133). This killing was accompanied by significant DNA damage, which was blocked by prior exposure of the cells to iron chelators, indicating that the DNA damage and the following cell death were mediated by Fenton’s reaction (133). Initially it was proposed that NO’s ability to damage enzymes containing Fe-S clusters could cause release of iron atoms, thereby increasing the intracellular levels of free iron, which can fuel the •OH production (133). However, the Imlay lab observed no increase in the free intracellular iron levels upon exposing cells to NO and H2O2. In fact, they found that NO, which blocks bacterial respiration via inactivation of quinol oxidases, causes NADH to accumulate in the cells (189). This increase in NADH levels could promote reduction of

36 free flavins to FADH2 via the flavin reductase enzyme, Fre, which may accelerate

Fenton's reaction by reducing the ferric iron, thereby fueling the generation of DNA damaging •OH radicals (Reaction 1.5.) (189).

+ Fre + H + NADH + FAD NAD + FADH2

3+ 2+ + FADH2+ Fe FADH + Fe + H

2+ 3+ Fe + H2O2 Fe + •OH + OH–

Reaction 1.5.

This proposed mechanism was supported further by the fact that the fre mutant was more resistant to the synergistic killing of NO + H2O2, whereas strains that overproduced Fre were hypersensitive (189). In addition, mutants lacking the terminal respiratory quinol oxidases, cyo cyd, which would accumulate NADH were found to be hypersensitive to the H2O2 only treatment, while a combination of NO with H2O2 failed to further enhance the killing in these mutants (189).

1.6.3. Mechanism of Cysteine-Enhanced H2O2 Toxicity

Several studies observed that cystine exposure after growing cells on poor sulfur sources transiently potentiated bacteriostatic low millimolar doses of H2O2 to cause up to four orders of magnitude killing in E. coli (19, 28, 30). This killing was also found to be associated with DNA damage and dependent on iron, again suggesting that it is mediated by •OH generation via Fenton chemistry (135). Growth on poor sulfur sources followed by cystine exposure leads to a temporary disruption in cysteine homeostasis in the cells, resulting in up to eight fold increase in the intracellular cysteine levels (135). The Imlay lab found that in vitro cysteine could reduce ferric iron to the ferrous form and drive

Fenton’s reaction to generate •OH, which caused plasmid nicking when glutathione was

37 provided in the reaction (135). They observed that mutants deficient in glutathione synthesis or unable to reduce glutathione disulfide did not demonstrate an increase in their intracellular cysteine pools when exposed to cystine post-starvation, and therefore, they were also resistant to cysteine-driven H2O2 toxicity (135). However, the mechanism by which glutathione impacts cysteine homeostasis remains to be determined.

Under normal growth conditions, the cells maintain a much lower level of intracellular cysteine, which varies between 0.1 to 0.2 mM and is insufficient to drive

Fenton’s reaction to cause DNA damage (135). Therefore, to avoid ROS-mediated oxidative damage it is critical to tightly regulate cysteine levels in aerobically respiring cells .This may be the reason why glutathione rather than cysteine serves as the thiol buffer in E. coli, despite their similar reduction potentials and pKas (135). Unlike cysteine, glutathione is markedly inefficient at reducing iron to the ferrous form, therefore, cells can maintain up to 10 to 20 times more glutathione than cysteine without risking the possibility of oxidative damage (135).

1.6.4. The Cyanide + H2O2 Co-Toxicity

A combination of low millimolar doses of cyanide and hydrogen peroxide, both of which are bacteriostatic by themselves, was also found to cause up to four orders of magnitude synergistic killing in E. coli (93, 190). This synergistic treatment was also demonstrated to be iron-dependent and DNA-damaging, and hence, dependent on the

Fenton reaction (93). In the past, the Imlay lab has proposed that, by blocking respiration,

CN could accelerate the killing and DNA damage caused by bacteriostatic doses of H2O2

(190). The mechanism proposed for this co-toxicity was similar to that of NO-driven

H2O2 poisoning. CN blocks respiration by binding to the terminal electron acceptor

38 cytochrome c oxidase complex and prevents it from being reduced. This causes an increase in the intracellular NADH levels, which could, in turn, catalyze the reduction of flavins via Fre (190). Reduced flavins (FADH2) can act as electron donors and reduce iron from its ferric form to its ferrous form, thereby recycling it and enabling to participate the in the toxic •OH-generating Fenton’s reaction again (Reaction 1.5.) (190).

Indeed, Woodmansee and Imlay found the ndh mutant, which is blocked for respiration and consequently has an elevated NADH pool, to be just as sensitive to H2O2 alone as it was to the combined treatment of CN and H2O2, whereas the fre mutant, lacking the flavin reductase, was resistant to this synergistic treatment (190). However, since reduced flavins are known to facilitate release of iron from the ferritin core (40, 62, 122), this result could be interpreted differently. Fre could promote CN and H2O2 co-toxicity by aiding CN in supplying iron to the Fenton reaction, therefore, we decided to investigate the role of iron depots in mediating this co-toxicity. Furthermore, from the perspective of

DNA damage and repair, a lot of questions remain unanswered, for example: 1) Does the

DNA damage inflicted by CN and H2O2 co-treatment depend on processes like DNA replication, transcription or translation? 2) Is the damage uniformly distributed over the chromosome, or is it concentrated around certain loci like the origin? 3) Are the double- strand breaks produced directly, or are they preceded by primary single-strand DNA lesions? 4) How much of this damage is in fact reparable? 5) What roles do recombinational repair, base excision repair, nucleoid excision repair and nucleoid associated proteins play in repairing this DNA damage? 6) Does the chromosomal damage lead to disassembly of the nucleoid structure? Our work aims to provide answers to these crucial questions.

1.6.4.1. The potential of cyanide + H2O2 co-toxicity as a cytotoxic weapon in activated immune cells

Myeloperoxidase (MPO) is a heme-containing peroxidase enzyme found in azurophilic granules of neutrophils and lysosomes of monocytes. During respiratory burst, MPO aids in combating invading microorganisms by catalyzing the generation of hypochlorous acid (HOCl) from H2O2 or other similarly highly-reactive molecules such as tyrosyl radicals using H2O2 to oxidize tyrosine residues, which are extremely toxic to pathogens. MPO can also convert the abundant thiocyanate (SCN-) found in blood serum into hypothiocyanite (OSCN-), which is also bactericidal. In addition to catalyzing these reactions, MPO catalyzes the formation of hydrogen cyanide (HCN) from serum thiocyanate during neutrophil activation on encountering bacterial infection (25, 63, 159,

169). HCN production during phagocytosis of penicillin treated Staphylococcus epidermis was found to be higher compared to when undamaged bacteria was phagocytosed (158). Part of the HCN produced came from MPO-H2O2 catalyzed oxidation of plasma thiocyanate (158). Additionally, MPO-H2O2 induced chlorination of glycine released due to the disintegration of the bacterial cell walls also contributed to

HCN generation by stimulated neutrophils (196).

However, the implications of the fact that MPO also catalyzes the formation of

HCN are not well appreciated. Our lab has previously reported that a mixture of NO,

HCN and H2O2 is tremendously bactericidal, and therefore, could potentially be another cytotoxic agent in the arsenal of our immune cells (108). As mentioned earlier, the NO potentiation of H2O2 toxicity is a known bactericidal weapon employed by macrophages, the mechanism of which has been well studied.

Even though at concentrations higher than 20 mM H2O2 alone can efficiently kill microorganisms, it is metabolically not possible for eukaryotic cells to generate H2O2, an uncharged molecule, at such high concentrations inside the cell, in addition to it being potentially toxic. Therefore, potentiation of physiologically relevant amounts of H2O2 by low concentrations of CN and NO may serve as a more feasible and safe mechanism to combat invading microbes. Therefore, we propose that the toxic combination of HCN,

NO and H2O2 could function as an efficient lethal agent against invading pathogens in activated neutrophils or phagocytizing monocytes. Our aim is to better understand the immune cells’ antimicrobial responses by investigating the metabolic and chromosomal aspects of CN potentiation of H2O2 toxicity.

1.7. SCOPE OF THIS THESIS

1.7.1. Cyanide Potentiates H2O2 Toxicity by Causing Catastrophic Chromosomal

Fragmentation via Recruitment of Iron from Intracellular Depots

Low millimolar concentrations of hydrogen peroxide or cyanide are individually bacteriostatic, while the double treatment kills exponentially growing Escherichia coli by

3-4 orders of magnitude in 15 minutes (118, 190). We have found that CN by itself does not kill even at a 100-fold higher concentration, whereas H2O2 can kill to completion at only 10-fold higher concentration (118). This indicates that the nature of synergistic toxicity is cyanide potentiation of H2O2 poisoning. We have also observed that this synergistic toxicity is associated with catastrophic chromosomal fragmentation: a violent, rapid and complete chromosome demise in 15 minutes of the treatment (118). Both the killing and fragmentation are blocked by iron chelators, suggesting that DNA damage and cell death are mediated by Fenton's reaction (118, 190).

Moreover, we have found that in vitro CN promotes the DNA-self-targeted

Fenton's reaction (since DNA, by binding iron readily, serves as both a platform for

Fenton's reagent and a target for the generated hydroxyl radicals). Remarkably, when cyanide joins this DNA-iron complex, it further potentiates Fenton's reaction (118).

However, sensitivity to the CN + H2O2 treatment of mutants with elevated intracellular iron levels (fur), or those unable to scavenge H2O2 (katEG) is increased by about the same magnitude as their sensitivity to the H2O2 only treatment, suggesting existence of two subpopulations of iron in the cell: one which CN can utilize to potentiate H2O2 toxicity and the other that it cannot, but which can nevertheless promote Fenton’s reaction (118). Through in vivo and in vitro experiments, we have found that the iron depot/distribution protein, ferritin, which can also bind DNA, directly transfers iron to

DNA in the presence of CN to provide at least a part of the first sub-population of iron that is involved in this potentiation (118). Since reduced flavins facilitate iron release from ferritin, this suggests that the action of flavin reductase would aid in CN meditated iron recruitment from ferritins, thereby fueling the Fenton reaction (118). We also found the resistance of the double fre ftnA double mutant to the CN and H2O2 treatment to be the same as that of the single mutants, which indicates that both of these enzymes function in the same pathway to promote the Fenton’s reaction, consistent with our prediction (118).

At the same time, we show that the stationary phase and starvation-induced protein, Dps, plays a protective role by sequestering this DNA-bound iron and by binding to the DNA itself (118). Dps protection against the synergistic treatment of CN and H2O2 is partial during the exponential phase, but absolute in stationary phase, when DNA is

42 complexed with up to 180,000 Dps molecules (118). Therefore, we propose that CN potentiation of H2O2 toxicity depends on CN-induced transfer of iron from intracellular iron depots, like Ferritin, directly to DNA and on further promotion of the DNA-self- targeted Fenton's reaction, both efficiently counteracted by Dps.

1.7.2. The Nature, Consequences and Attempted Repair of the Catastrophic

Chromosomal Fragmentation Induced by the Synergistic Cyanide and Hydrogen

Peroxide Combination

We have found that the catastrophic chromosomal fragmentation induced by CN and H2O2 is replication and translation independent and uniformly distributed over the chromosome. The catastrophic fragmentation was observed in non-replicating dnaA and dnaC mutants, or cells pretreated with chloramphenicol. However, the fragmentation does depend on cell density during treatment, since both saturated non-replicating dnaA and dnaC cells and dps overnight cultures, failed to show any chromosomal fragmentation, even though they were still fully sensitive to the CN + H2O2 treatment.

However, these stationary cultures showed normal levels of fragmentation if diluted 10- fold before the treatment (even though metabolism cannot really start in the presence of cyanide, due to the blocked ATP production). Regardless, since the presence of even a single double-strand break in the circular E. coli chromosome can be fatal, the lack of

~60% chromosomal fragmentation above the background in these cells does not necessarily mean that the loss in viability is not due to DNA damage.

We have also demonstrated that the DNA damage induced in wild-type cells treated with CN and H2O2 for 5 minutes is completely repairable, whereas recombinational repair or excision repair deficient mutants could not repair their

43 chromosomes. In fact, the wild-type cells showed a dramatic revival in cell titer when CN and H2O2 was removed from their cultures (up to 3 orders of magnitude in half an hour), while the repair deficient cells failed to recover from the treatment. Moreover, we show that the base excision repair-deficient mutant, xthA nfo, was more sensitive to CN + H2O2 and exhibited substantially more chromosomal fragmentation than the recombinational repair mutants, indicating that oxidized DNA bases are the primary DNA lesions, which eventually get converted into double-strand breaks. Additionally, the polA and ligA mutants (defective in completion of any kind of DNA repair in which DNA strands are broken) were found to be acutely sensitive to both the CN + H2O2 treatment and the

H2O2-only treatment and showed massive DNA damage. This suggests that the H2O2 treatment itself generates several single-strand breaks, which DNA pol I and DNA ligase promptly repair. On the other hand, mutants in nucleotide excision repair were found to exhibit wild-type like sensitivity to the CN + H2O2 treatment, indicating that they do not play a role in repair of this catastrophic fragmentation. Furthermore, mutants in nucleoid associated proteins showed a minor enhancement in their sensitivities to CN and H2O2 as compared to wild-type cells, suggesting that the nucleoid structure does not offer significant protection from CN and H2O2-induced chromosomal fragmentation.

Initially, we were surprised that the CN + H2O2-induced catastrophic chromosomal fragmentation was not accompanied by massive degradation of the generated linear DNA, as seen after gamma irradiation (22). A potential reason for this was that the fragmentation we observe in pulsed-field gels does not happen in vivo, but is rather an in vitro artifact of cell lysis or gel electrophoresis. However, the answer turned out to be much simpler: since cyanide blocks ATP production of the cell it freezes all

ATP-dependent processes, including linear DNA degradation by the RecBCD helicase/exonuclease, the main linear DNA degradation pathway in bacteria. In fact, over-expression of RecBCD in wild-type cells resulted in an enhancement in the degradation observed, suggesting that the RecBCD activity in wild-type cells treated with

CN and H2O2 was saturated. To study ATP-dependent consequences of CN +

H2O2treatment, we resuspended cells in a fresh medium without the two chemicals.

Removal of cyanide revealed at least three major post-treatment phenomena: 1) degradation of the linearized DNA, 2) degradation of stable RNA and 3) nucleoid disintegration. Mutants lacking nucleoid-associated proteins showed a minor enhancement in their sensitivities to CN and H2O2 as compared to wild-type cells.

We have found that the synergistic CN + H2O2 treatment not only causes catastrophic fragmentation but also leads to significant loss of ribosomal RNA (rRNA) in

E. coli. 45 minutes of treatment lead to degradation of ~40% of the rRNA. However, we show that this measurement underestimated the damage inflicted on rRNA, since removal of CN and H2O2 from the cells followed by resuspension results in ~80% of the rRNA being degraded, suggesting that the process of complete degradation of the damaged rRNA species required ATP.

DNA association with the nucleoid is transient, via the point of rotation and the broken chromosome dissociates from the nucleoid as a result of normal nucleoid dynamics once the cells are resuspended in fresh medium after the treatment. Therefore, resumption of the DNA-nucleoid rotation after CN removal resulted in release of more broken DNA from the nucleoid or nucleoid disintegration. Indeed, mutants lacking nucleoid associated proteins such as StpA and Fis were found to exhibit significantly

45 more nucleoid disassembly than the wild-type cells. On the other hand, H-NS, another nucleoid associated protein, was found to promote this disintegration along with processes like transcription.

1.8. FIGURES

Figure 1.1. A schematic representation of the three confirmed models of replication fork (RF) disintegration. The parental DNA is shown in blue, while the newly synthesized strands are in red. The bricked circle represents a block in replication progression such as replication fork inhibition or DNA-bound proteins. (Left to Right) 1) Replication fork running into a stalled replication fork of the previous round results in replication fork rear-ending 2) Replication fork collapse is caused by a replication fork running into a single-strand interruption in the template DNA 3) In replication fork regress-split the two template strands reanneal, forcing the two nascent strands into a duplex of their own and forming a Holliday junction that is cleaved by RuvABC.

Figure 1.2. A schematic of the recombinational repair pathway. Recombinational repair is initiated by binding of RecBCD to the exposed ends of the double-strand breaks, which unwinds and degrades the duplex DNA via the helicase/nuclease activities to give a long, single- stranded 3’ overhang for the assembly of the RecA filament. Subsequently, RecA catalyzes strand invasion and homologous recombination giving rise to the X-shaped Holliday junction structure, which are resolved via the RuvABC complex. The remaining gaps are filled and ligated by DNA polymerase I and ligase, respectively.

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CHAPTER 2: CYANIDE POTENTIATES HYDROGEN PEROXIDE TOXICITY

BY RECRUITING IRON FROM INTRACELLULAR DEPOTS TO CAUSE

CATASTROPHIC CHROMOSOMAL FRAGMENTATION

2.1. INTRODUCTION

Hydrogen peroxide (H2O2) is an uncharged stable molecule in which oxygen is reduced halfway between the ground state molecular oxygen O2 and the fully reduced oxygen of water (H2O). Due to its uncharged character and generally low reactivity with organic compounds, H2O2 moves freely from outside to inside the cell. However, the cell tries to keep intracellular concentrations of H2O2, as well as superoxide O2– (a reactive oxygen species halfway between O2 and H2O2 and easily reducible to H2O2) as low as possible, because intracellular iron makes H2O2 extremely toxic (31).

For E. coli, H2O2 saturates cellular defenses at low millimolar concentrations (1-

10 mM), becoming bacteriostatic during short treatments (34, 35), while bactericidal and clastogenic (degradation of chromosomal DNA) upon prolonged exposures (8). H2O2 shows rapid toxicity in E. coli at concentrations higher than 20-30 mM (34, 35). In fact, the famous 3% hydrogen peroxide first aid antiseptic (~1 M concentration) kills any kind of bacteria within a few minutes (21, 66).

The extreme intracellular H2O2 toxicity is attributed to its catalytic cycle with free cytoplasmic iron that produces hydroxyl radicals (first used in the famous Fenton's

2+ 3+ 3+ 2+ reagent (22, 46): Fe + H2O2  Fe + OH· + OH– ( while Fe is then returned to Fe by metabolites like free flavins). The generated hydroxyl radicals react with organic compounds at diffusion rates (12). Because of this amplified reactivity in the presence of

64 free cytoplasmic iron, H2O2 and superoxide are classified as "reactive oxygen species", even though they themselves do not show reactivity with typical organic biomolecules.

The cell minimizes Fenton's reaction by actively reducing intracellular concentrations of H2O2, superoxide and free iron. The efficiency of H2O2 scavenging is demonstrated by the fact that cells completely deficient in it cannot grow once the endogenous H2O2 reaches ~0.5 µM (60), while the wild type (WT) cells can still grow even in the presence of 1 mM exogenous H2O2 (8). In E. coli, H2O2 is efficiently degraded by a two-step enzyme system. At concentrations up to 20 µM in E. coli, H2O2 is mostly degraded by the copious alklyperoxidase, AhpCF (31), while higher H2O2 concentrations are scavenged by induction of two catalases, encoded by katE and katG genes in E. coli (28). At the same time, cytoplasmic superoxide is efficiently removed by two superoxide dismutases, SodA and SodB (31). Finally, the amount of free cytoplasmic iron is tightly controlled by the ferric uptake regulator Fur (29). Because of low availability of soluble iron (Fe2+) in the oxidizing environment, the cells have to secrete various iron- Fe3+-chelating molecules, called siderophores, and then collect their iron- loaded forms back, retrieving the iron for internal use (75). Since iron is expensive to procure, and frequently limited in the environment, surplus cytoplasmic iron is not excreted into the environment, but is stored in association with the specialized proteins of the ferritin superfamily (68). Typical ferritins function as iron distribution centers, taking in or releasing iron according to metabolic needs of the cell, while the mini-ferritin Dps is induced in response to H2O2-stress and in the stationary cells, to sequester "reactive" iron from iron- H2O2 complexes.

A peculiar phenomenon is potentiation of H2O2 toxicity by some simple substances, which by themselves do not kill, and some of them are not even bacteriostatic. For example, amino acids cysteine and histidine in high, but still physiological, concentrations potentiate the effect of bacteriostatic H2O2 concentrations, killing the cells (4). Ascorbic acid has also been long known to potentiate H2O2 toxicity

(56). Nitric oxide (NO) is by itself bacteriostatic in high enough concentrations (69), but addition of NO to otherwise bacteriostatic concentrations of H2O2 makes it strongly bactericidal (43). In fact, synergistic toxicity of NO + H2O2 is used by our immune cells to kill invading microbes (9, 58), but the mechanisms behind the microbial death are not entirely clear (77).

Synergistic toxicity of H2O2 with cyanide (CN) was also noted before (33) and explored by Woodmansee and Imlay, who proposed that CN-block of respiration leads to an increase in Fe3+-reduction potential of the major siderophore reductase Fre, resulting in faster Fe3+  Fe2+ cycling, thereby accelerating Fenton's reaction (78). The proposed mechanism was later confirmed for a similar case of NO + H2O2 co-toxicity (77). Even though CN may seem like an artificial potentiator, it is in fact produced by many plants, most fungi and even by some microbes (44), in addition to being generated by our activated immune cells (11, 27, 70, 73).

We encountered the co-toxicity with H2O2 while characterizing the surprising killing potential of old hydroxyurea solutions, that turned out to be due to a mixture of

NO, CN and H2O2 (50). Here we are reporting our investigation on the nature of the CN

+ H2O2 co-toxicity, of its potential targets and of its chromosomal effects. Theoretically, there are two possible explanations of a co-toxicity. The most obvious one is

"redundancy"; according to the redundancy explanation, both CN and H2O2 are toxic, and both toxicities are counteracted by the same mechanism. Therefore, adding them together inactivates an essential cellular function or saturates anti-toxicity mechanisms, killing the cell. For example, both CN (37) and H2O2 (5) are known to bind cytochrome oxidase enzymes, so it is possible that this inactivation could be the cause of death by the CN +

H2O2 treatment.

The second explanation of co-toxicity is "potentiation", according to which only one of the two chemicals is toxic, but normally there are efficient mechanisms to prevent or neutralize its toxicity, while the second chemical targets these anti-toxicity mechanisms without being toxic itself. Since H2O2 is known to be toxic in high concentrations (34, 35), its toxicity at low concentrations could be potentiated by CN, for example, by inactivating catalases, which are heme-containing enzymes (heme iron is a well-known intracellular CN target). One of the initial objectives of this work was to distinguish between these two explanations for CN + H2O2 co-toxicity. The second objective was to identify targets of CN and H2O2 poisoning. The third objective was to explore chromosomal aspects of CN + H2O2 toxicity.

2.2. MATERIALS AND METHODS

2.2.1. Strains and Plasmids

Escherichia coli strains used are all K-12 BW25117 derivatives (3). Alleles were moved between strains by P1 transduction (55). The mutants were all deletions from the

Keio collection (3), purchased from the E. coli Genetic Stock Center, and were verified by PCR (and, whenever possible, phenotypically). For double mutants construction, the resident kanamycin-resistance cassette was first removed by transforming the strain with

67 pCP20 plasmid (20). The plasmid pMTL20 (14) was used for all in vitro plasmid relaxation assays.

2.2.2. Enzymes and Reagents

Ferritin and apoferritin from equine spleen, catalase from bovine liver, hydrogen peroxide, deferoxamine mesylate, 2,2′-Bipyridyl and N,N-Dimethyl-4-nitrosoaniline (p- nitrosodimethylaniline) were purchased from Sigma. Potassium Cyanide was purchased from Fisher-Scientific. Yeast chromosome pulsed-field gel electrophoresis markers were from New England Biolabs.

2.2.3. Growth Conditions and Viability Assay

To generate killing kinetics, fresh overnight cultures were diluted 500-fold into

LB medium (10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, 250 μl of 4 M NaOH per liter (55)) and were shaken at 37°C for about two and a half hours or until they reached exponential phase (OD600 approximately 0.3). At this point, the cultures were made 3 mM for CN and/or 2 mM for H2O2 (or the indicated treatment) and the shaking at 37°C was continued. Viability of cultures was measured at the indicated time points by spotting

10 μL of serial dilutions in 1% NaCl on LB plates (LB medium supplemented with 15 g of agar per liter). The plates were incubated overnight at 28 °C, the next morning colonies in each spot were counted under the stereomicroscope. All titers have been normalized to the titer at time 0 (before addition of the treatment).

2.2.4. Measuring Chromosomal Fragmentation via Pulsed-Field Gel Electrophoresis

This generally follows our previous protocols (42, 49). All strains were grown in

LB medium; overnight cultures were diluted 500-fold and grown with 1-10 μCi of 32P- orthophosphoric acid per ml of culture for 2.5 h at 37 °C (OD600 approximately 0.3)

68 before addition of 3 mM CN + 2 mM H2O2 (or the indicated treatment). The reactions were stopped by addition of 315 µg of catalase, and aliquots of the culture were taken at the indicated times to make plugs. Cells of the aliquot were spun down, resuspended in

60 μl of TE buffer and put at 37°C. 2.5 μl of proteinase K (5 mg/ml) was added, immediately followed by 63 μl of molten 1.2% agarose in the lysis buffer (1% sarcosine,

50 mM Tris-HCl, and 25 mM EDTA) held at 70°C. The mixture was pipetted a couple of times before being poured into a plug mold and let solidify for 2 minutes at room temperature. The plugs were then pushed out of the molds and incubated overnight at

60°C in 1 ml of the lysis buffer. Half plugs were loaded into a 1.0% agarose gel in 0.5X

Tris-borate-EDTA buffer and run at 6.0 V/cm with the initial and the final switch times of 60 and 120 s, respectively, at 12°C in a Bio-Rad CHEF-DR II PFGE system for 20-

22 hours. The gel was vacuum-dried at 80°C and then exposed to a PhosphorImager screen overnight. The resulting signals were measured with a PhosphorImager (Fuji Film

FLA-3000).

2.2.5. In vitro Non-DNA Fenton's Reaction

The reaction containing 40 µM N,N-dimethyl-4-nitrosoaniline, 2 mM H2O2 and/or

3 mM CN in 2 ml water (33) was supplemented with various concentrations of up to 1 mM of stable salts of the indicated transition metals. The reaction was followed spectrophotometrically by measuring absorbance at 440 nm.

2.2.6. Plasmid Relaxation Assay

About 100-200 ng of plasmid (1-2 μl) was incubated with 5 µM of either ferritin or apoferritin with 3 mM CN and 2 mM H2O2 in 1X TAE buffer (40 mM Tris-HCl, 20 mM acetic acid, 1 mM EDTA, pH 8) at 37°C. At specified times, aliquots were removed

69 and the reaction was stopped by adding 45 µg of catalase. Since apoferritin was in 50% glycerol (an inhibitor of Fenton's reagent), both apoferritn and ferritin preparations were first changed into 1% NaCl solution by using 100K Amicon ultra-0.5 ml centrifugal filter and spin column units.

When using iron instead of enzymes, the plasmid was incubated for only 2 minutes at room temperature before addition of catalase. These samples were then run on a 1.1% agarose gel at 3 V/cm before being transferred to a nylon membrane and hybridized with pMTL20-specific radioactive probe to calculate the percentage of the relaxed plasmid form in the total plasmid.

2.2.7. Southern Hybridization

The agarose gels were washed with 0.25 M HCl, followed by 0.5 M NaOH and, finally, with 1 M Tris HCl pH 8.0. Each wash was 40 minute long. The treated gels were then placed on Amersham Hybond N+ (GE Healthcare) nylon membrane, covered with

Saran wrap, and DNA was transferred by vacuum for 1-2 hours. After that the DNA was

UV-crosslinked to the membranes and probed with 32P-labeled pMTL20-specific probe.

Hybridization was carried out overnight at 63°C in a 0.5M Sodium Phosphate (pH 7.4) and 5% SDS hybridization buffer. In the morning, the membranes were washed thrice with 1% hybridization buffer and rinsed with water just before covering them with Saran wrap and exposing to a PhosphorImager screen. The resulting signals were measured with a PhosphorImager (Fuji Film FLA-3000).

2.3. RESULTS

2.3.1. CN + H2O2 Treatment Causes Catastrophic Chromosomal Fragmentation

The CN(3)+H2O2 (2) killing in our standard conditions (3 mM CN, 2 mM H2O2) is at least four orders of magnitude within one hour, in line with the previous report (78)

(Fig. 2.1. ABC). At the same time, CN (3)-alone or H2O2 (2)-alone treatments are bacteriostatic (Fig. 2.1. BC). The CN (3) + H2O2 (2) toxicity is robust, as demonstrated by step-wise lowering concentrations of one reagent, while keeping the other one constant: the extent of the killing is significantly reduced only by 10-fold reduction of either reagent. In fact, the killing of the wild-type strain used in these experiments can be stopped immediately by diluting the treated culture 10-fold, a useful stopping technique during a short time course.

Woodmansee and Imlay detected significant DNA damage in CN (3)+H2O2 (2.5)- treated cells by quantitative PCR (78), whereas we have earlier reported detectable chromosomal fragmentation caused by unspecified low levels of CN+NO+H2O2 (50).

Therefore, we expected to observe comparatively higher levels of chromosomal fragmentation after our standard CN (3)+H2O2 (2) treatment. Nevertheless, we were still surprised to find that CN+H2O2 killing in our conditions is associated with extraordinary levels of chromosomal fragmentation, detected by pulsed-field gel electrophoresis (Fig.

2.1. DE). We use the term "catastrophic fragmentation" to represents the violent, rapid and complete chromosome demise that CN+H2O2 treatment induces. Since massive fragmentation is already observed after only one minute of treatment (see below), it is likely the cause, rather than the consequence, of death. As a negative control, we found no chromosomal fragmentation after treatment with lethal concentrations of transcription

71 inhibitor rifampicin or translation inhibitor kanamycin (Fig. 2.1. DE), this also contradicts the reports of massive oxidative damage associated with various mechanisms of antibiotic-caused bacterial cell death (45).

After reaching a maximum of 65% (signal over background) at 15 minutes, the apparent level of CN+H2O2 caused fragmentation goes down at the later time point (Fig.

2.1. E). Since the average molecular weight of the chromosomal fragments continues to decrease with time of the treatment (Fig.2.1. D), the observed decrease in fragmentation must be an artifact of the smallest DNA fragments migrating out of the gel. We conclude that, over the period of 45 minutes, CN+H2O2 treatment kills E. coli cells continuously by inducing double-strand breaks in the chromosomal DNA. Moreover, since in other experimental systems, strong lethality is already observed with chromosomal fragmentation levels of 15-20% (see 2.4. Discussion), the catastrophic fragmentation induced by CN+H2O2 treatment should leave cells no chance of survival, an expectation supported by the 10,000-fold drop in the viable counts (Fig. 2.1. BC).

2.3.2. The Nature of CN and H2O2 Co-Toxicity

One could distinguish between the two explanations for CN+H2O2 co-toxicity

(redundancy vs potentiation) by checking whether CN alone or H2O2 alone would kill at higher concentrations. If both single treatments are toxic at high concentrations, then the redundancy explanation would be possible (potentiated toxicity is always a possibility).

However, if only one of the single treatments is toxic at high concentrations, then the corresponding substance is the poison, while the other one acts as a potentiator, by enabling the poisoning mechanism or by disabling a resistance mechanism.

We found that CN alone, even at very high concentration of 300 mM, neither kills, nor causes any chromosomal fragmentation (Fig. 2.1. FG). In contrast, H2O2 alone kills at concentrations of 15 mM (H2O2 (15)) or higher, by inducing chromosomal fragmentation (Fig. 2.1. FG). Remarkably, H2O2 (10) does not kill or cause chromosomal fragmentation (Fig. 2.1. H), whereas H2O2 (20) already shows significant killing/fragmentation potential. In other words, transition H2O2 (10) —> H2O2 (20) does not translate into a simple 2-fold increase in DNA damage; rather, it brings about a qualitative shift, as if a potentiator like CN has been added. Apparently, at high concentrations, H2O2 functions as a self-potentiator, for example, by partially converting into a substance that acts like CN. The kinetics of killing by H2O2 (20) is similar to the one by the standard CN(3)+H2O2(2) co-treatment (compare Fig. 2.1. I versus 2.1. C) and is accompanied by a similar magnitude of catastrophic chromosomal fragmentation (Fig.

2.1. J). Therefore, we conclude that 1) H2O2 kills by causing catastrophic chromosomal fragmentation, while CN potentiates this killing by increasing the effective intracellular

H2O2 concentrations at least 10-fold; 2) at high concentrations, H2O2 may self-potentiate its own poisoning; 3) the killing target of H2O2 is the chromosomal DNA, while CN targets are unclear.

2.3.3. Genetic Analysis of CN Potentiation of the H2O2 Killing: Expected Phenotypes and Their Interpretation

To identify the targets of CN potentiation, as well as CN potentiation counteracting processes, if any, we quantified H2O2 alone and CN+H2O2 sensitivity of select mutants. Formally, the original phenotypes of WT cells are: “H2O2 alone” treatment is bacteriostatic, while CN+H2O2 treatment is strongly bactericidal (Fig. 2.1. C

73 and 2.2. A). Four distinct changes from these WT effects are possible in mutants, with specific interpretations (Fig. 2.2. B-E)

The first (perhaps the most intuitive) mutant phenotype is similarly increased sensitivity to both H2O2 alone and CN+H2O2 (Fig. 2.2. B). Interpretation: the corresponding protein counteracts H2O2 toxicity without affecting CN potentiation of it.

In other words, such a mutant identifies an independent branch of H2O2 toxicity that acts in parallel with the CN potentiated branch.

Another mutant phenotype is a similar sensitivity to both H2O2 alone and

CN+H2O2 (that is, H2O2 alone sensitivity drops to the level of CN+ H2O2 sensitivity)

(Fig. 2.2. C). Interpretation: the corresponding protein counteracts H2O2 toxicity and is a target of CN potentiation. Identification of such targets was the primary objective of our mutant search.

A third expected mutant phenotype is no H2O2 alone sensitivity (like in WT), but a deeper CN+H2O2 sensitivity (Fig. 2.2. D). Interpretation: the corresponding protein counteracts CN potentiation without affecting H2O2 toxicity.

Finally, the fourth effect is again no sensitivity to H2O2 alone (like in WT), but this time a shallower sensitivity to CN+H2O2 (Fig. 2.2. E). Interpretation: the corresponding protein promotes CN potentiation of H2O2 toxicity.

2.3.4. H2O2 Scavengers Counteract CN Potentiation

Consideration of the scheme of H2O2 toxicity/detoxification (Fig. 2.1. A) for possible targets of CN potentiation leads to a perplexing possibility that tight CN binding to the heme iron inhibits both sides of the process, resulting in a larger intracellular H2O2 pool on the one hand, yet a less efficient hydroxyl radical production on the other.

Indeed, the CN vulnerability of the hydroxyl-radical-generating side of the scheme (Fig.

2.1A right) is due to the fact that CN blocks the electron transport chain, via binding to hemes of cytochrome oxidases (37), and at the same time, via complexing highly reactive free iron into mildly reactive ferrocyanide. However, the "catalase" side of the scheme

(Fig. 2.1. A left) should be equally CN vulnerable, as both E. coli catalases are also heme-containing enzymes (15), making them ideal targets for CN potentiation of H2O2 toxicity.

There is a disagreement over whether inactivation of catalases sensitizes cells to

H2O2 alone treatment in E. coli: Pueyo and colleagues report that the double ∆katEG mutant is extremely sensitive to 20' treatment with H2O2 (61), while Imlay and colleagues claim their double catalase mutant is not generally sensitive to H2O2 (~10-fold deeper viability loss compared to WT after 15' treatment) (35). In our hands, the ∆katEG mutant is killed by CN+H2O2 faster and deeper than WT cells (Fig. 2.2. F), but it is the unusual kinetics of H2O2 alone sensitivity, which fails to plateau within 45 minutes of treatment, that makes the overall pattern of the double catalase deletion mutant complex. Indeed, at

15 minutes it corresponds best to Fig. 2.2. B, but then matches Fig. 2.2. C pattern at 45 minutes. If we are to ignore the earliest time point, catalases counteract H2O2 toxicity (as they should), without affecting CN potentiation at first, but then they gradually become targets of CN inhibition. Still, the dramatic difference between the first 15 minutes of

CN+H2O2 versus the H2O2 alone sensitivity curves excludes catalases from being the potential targets of immediate CN inhibition.

While catalases are responsible for defending cells against millimolar concentrations of exogenous H2O2, E. coli employs alkylperoxidase, AhpCF, to remove

75 micromolar concentrations of endogenous H2O2 (31). Alkylperoxidase does not have any metal that cyanide could complex and is too weak to counter 2 mM H2O2 of our treatment anyway, so the ahpC mutant was expected to be killed similar to the WT with the

CN+H2O2 treatment. Surprisingly, we observed increased CN+H2O2 sensitivity in the

∆ahpC mutant with no increase in H2O2 alone sensitivity (Fig. 2.2. G). Thus, unexpectedly, if the first 5 minutes of the two treatments are considered, both ∆ahpC and

∆katEG results are generally consistent with the "Fig. 2.2. D" scenario, suggesting that both enzymes somehow counteract CN potentiation of H2O2 toxicity, with either no or delayed effect on H2O2 alone toxicity. Since the existence of a specific intracellular pool of H2O2 that is preferentially used for CN potentiation of Fenton's reagent is unlikely

(uncharged and small H2O2 molecules diffuse freely), the unexpected phenotype of the

∆katEG and ∆ahpC mutants suggest tight trafficking of iron inside the cells.

2.3.5. The Electron Transport Connection

Interestingly, Woodmansee and Imlay have already reported a major target of CN potentiation of H2O2 toxicity, working in the "electron transport" area in the Fig. 2.1. A scheme (expanded in Fig. 2.3. A) (78). Nicotinamide adenine dinucleotide (phosphate)

(NAD(P)H) is the main reducing power of E. coli cells grown aerobically in rich medium, its electrons being funneled into the aerobic respiratory chain by two NADH dehydrogenase (NDH) enzymes (one coded by the nuo gene cluster, the other by the ndh gene) through quinones to the three terminal cytochrome oxidases, bo, bd-I and bd-II

(Fig. 2.3. A) (7). Woodmansee and Imlay confirmed an earlier report (33) that the ndh mutants are sensitive to H2O2 alone treatment, and this sensitivity is not further increased by CN (78). At the same time, the nuo mutants behaved like WT in these assays. The

76 equal and high sensitivity of ∆ndh mutant to both H2O2 alone and CN+H2O2 ("Fig. 2.2.

C" pattern) was a clear genetic indication that the electron transport chain is a target of cyanide potentiation of the H2O2 killing (78).

Woodmansee and Imlay also reported that loss of fre, the flavin mononucleotide

(FMN) reductase, has an opposite effect, by mostly preventing CN potentiation of the

H2O2 killing ("Fig. 2.2. E" pattern) (78). The FMN reductase Fre reduces free flavins using excess of NAD(P)H (23). Interestingly, it is the major ferrisiderophore reductase activity in E. coli, reducing ferric citrate or other iron-siderophores so that they release ferrous iron (18). Woodmansee and Imlay proposed that Ndh action prevents buildup of excess of NAD(P)H that would otherwise be used by Fre to reduce ferric iron (Fe3+ 

Fe2+), thereby promoting Fenton's reaction (Fig. 2.3. A) (78).

We have confirmed that the ∆fre mutant has a reduced sensitivity to CN+H2O2 treatment, whereas Fre overproduction dramatically increases this sensitivity (Fig. 2.3.

B). Thus, Fre is indeed partially responsible for fueling Fenton’s reagent in E. coli cells, probably via reduction of insoluble Fe3+ to soluble Fe2+. We have also confirmed the WT behavior of the nuo mutant (Fig. 2.3. C). However, using ndh alleles both from Imlay's collection and from the E. coli Genetic Stock Center, we failed to observe the expected sensitivity of ∆ndh mutants to H2O2 alone (Fig. 2.3. C). Even more surprisingly, while being resistant to H2O2 alone, the ∆ndh mutants, in our hands, turned out to be hypersensitive to CN+H2O2 (Fig. 2.3. C). This opposite effect of ndh inactivation, compared to the effect of fre inactivation, leads to faster chromosomal fragmentation in the ndh mutants and slower fragmentation in the ∆fre mutant (Fig. 2.3. D). Remarkably, the fact that Ndh, like H2O2 targeting enzymes above, counteracts CN potentiation without

77 affecting H2O2 toxicity ("Fig. 2.2. D" pattern) means that Ndh counteracts CN potentiation directly, rather than through the elevated NADH concentration boost to the overall Fenton's reaction, otherwise, the ∆ndh mutant would have increased sensitivity to both H2O2 alone and CN+H2O2 treatments (Fig. 2.2. B). The general idea of tight intracellular iron control and its possible disruption by CN is consistent with this overall interpretation.

The possibility of NADH-independent nature of the ∆ndh mutant effect was further supported by the phenotypes of the ∆cyo, ∆cyd and ∆app mutants, deficient in the three cytochrome oxidases that catalyze the final step in electron transport from NADH to O2 in aerobic respiration (7). According to the scheme (Fig. 2.3. A), inactivation of these enzymes should lead to accumulation of NADH, elevating sensitivity to both H2O2 alone and CN+H2O2. However, from a different perspective, we can also imagine that since all cytochrome oxidases have heme as a critical cofactor that binds CN tightly, they should be targets of CN potentiation, meaning that all three mutants should be equally sensitive to H2O2 alone or CN+H2O2 ("Fig. 2.2. C" pattern). However, contrary to this strong prediction, all three cytochrome oxidase mutants showed the phenotype of the ∆ fre mutant: no H2O2 alone sensitivity and at the same time a shallower CN+H2O2 sensitivity (Fig. 2.3. E), as if CN utilizes the cytochrome oxidase molecules themselves to promote H2O2 toxicity.

The disagreement among these results within the confines of the NADH-centered scheme (Fig. 2.3. A) suggests that the overall aerobic respiratory chain has no direct role in CN potentiation of H2O2 toxicity, even though individual activities of (or around) this chain do contribute to the phenomenon. Specifically, if cyanide potentiates formation of

Fenton's reagent in vivo, NADH dehydrogenase II (Ndh) counteracts this potentiation, while flavin reductase and cytochrome oxidases promote this potentiation.

2.3.6. The Role of Iron in CN + H2O2 Co-Toxicity

Our simplistic in vitro modeling of CN potentiation of H2O2 toxicity, by measuring the effect of cyanide on the standard in vitro Fenton's reaction (33), expectedly revealed a complete block, the stoichiometry of inhibition suggesting formation of

Fe(CN)6 as the inert species (Fig. 2.4. A). The iron-cyanide complexes Fe(CN)6 are known to be significantly less reactive than free iron, and this inhibition of Fenton's reaction in vitro emphasizes the paradoxical phenomenon of the CN+H2O2 toxicity in vivo (Fig. 2.1.), indicating complicated nature of the underlying mechanisms. At face value, the complete shutdown of Fenton's reaction by CN in vitro predicts that complexing of iron by CN in vivo should similarly prevent production of hydroxyl radicals and should save the cells from H2O2 toxicity. To make sure that CN+H2O2 killing is due to Fenton's reaction in vivo, we have confirmed the previous report (78) that the presence of in vivo iron chelators, either deferoxamine or dipyridyl, completely blocks CN+ H2O2 killing (Fig. 2.4. B). The two iron chelators also block chromosomal fragmentation (Fig. 2.4. C). The dramatic effect of iron chelators implies that, in contrast to the formation of inert iron-cyanide complexes in vitro, the intracellular iron is somehow recruited by CN to promote Fenton's reaction.

This surprising discrepancy could be explained if, instead of iron, another common transition metal in the cell is responsible for CN potentiation of Fenton's reaction. The result with iron chelators would still be consistent with this idea, as both deferoxamine and dipyridyl are known to chelate other transition metals (17, 39). To test

79 the involvement of other transition metals in CN potentiation of in vivo Fenton's reagent, we assembled standard Fenton reactions in vitro with 2 mM H2O2 and 250 µM of the following transition metals: chromium, manganese, (iron as a positive control), cobalt, nickel, copper and zinc. We found that some of the metals did promote weaker Fenton- like reactions, but these reactions were all inhibited by addition of 3 mM CN, just like the iron-promoted Fenton's reaction (Fig. 2.4. D), making contribution of other transition metals to CN potentiation of H2O2 toxicity unlikely.

Since iron was the best transition metal to promote Fenton's reaction in vitro and apparently in vivo, we tested the possibility that CN stimulation of Fenton's reaction in vivo was specific to our readout, which was (chromosomal) DNA. Plasmid relaxation is a convenient and sensitive DNA-based readout for Fenton's reaction in vitro (38, 59, 72).

Remarkably, in contrast to CN inhibition of the classic in vitro Fenton's reaction (Fig.

2.4. A), we have observed a robust CN stimulation of Fe+H2O2 promoted plasmid relaxation in vitro (Fig. 2.4. EF). Since DNA readily binds both Fe2+ and Fe3+ iron (57),

DNA-iron complexes can act as a self-targeting Fenton's reagent (52) ("self-targeting" in the sense that DNA, by binding iron readily, serves not only as a platform for Fenton's reagent, but also as a proximal target for the generated hydroxyl radicals). Remarkably, when cyanide joins this DNA-iron complex, it further potentiates Fenton's reaction, instead of inhibiting it.

2.3.7. Phenotypes of Mutants with Increased "Free" Cytoplasmic Iron

With this better understanding of iron's role in the DNA-self-targeted Fenton's reaction in vivo, we tested the effect of increased free intracellular iron in certain mutants on the CN potentiation of H2O2 toxicity. The Fur protein is a ferric ion uptake regulator

80 that functions as either a repressor or activator (or sometimes both) for a number of genes

(29). In aerobically-growing ∆fur mutant cells, the amount of free cytoplasmic iron is elevated ~7-fold (40). We found that the ∆fur mutant shows delayed, yet significant sensitivity to H2O2 and a matching additional sensitivity to CN+H2O2 (Fig. 2.4. G). In other words, potentiation of H2O2 alone toxicity by fur inactivation is independent of CN potentiation of the same toxicity. Thus, the Fur regulator is not a target of CN potentiation, but counteracts H2O2 toxicity directly ("Fig. 2.2. B" pattern). Apparently, the extra free cytoplasmic iron in ∆fur mutants neither contributes to CN potentiation of

H2O2 killing, nor is "neutralized" by CN, like it is in vitro (Fig. 2.4. A). In other words, there appear to be at least two subpopulations of iron in the cell: the one that can be recruited by CN for potentiation of H2O2 toxicity, and the other that is neither

"recruitable", nor "neutralizable" by CN, yet fuels its own Fenton's reagent.

Besides fur, a similar ~8-fold increase in pool of free intracellular iron in E. coli is detected in the double ∆sodAB mutant lacking cytoplasmic superoxide dismutases (40,

53). Surprisingly, in contrast to the fur mutant behavior (Fig. 2.4. G), the ∆sodAB mutant showed only a slight sensitivity to H2O2-alone treatment and at the same time a greatly- reduced sensitivity to CN + H2O2 (Fig. 2.4. H), as if the "increased free iron" in this mutant is not only poorly-recruited by CN to fuel Fenton's reagent, but also is not accessible to H2O2-alone for the same purpose. At a minimum, these unexpected effects in mutants with increased free cytoplasmic iron argue against the existence of a significant pool of DNA-accessible cytoplasmic iron outside the control of the iron- trafficking system (Fig. 2.4. I, top). Instead, the bulk of cytoplasmic iron must be

81 distributed among specific depots (Fig. 2.4. I, bottom), with CN apparently recruiting it from a subset of these depots.

2.3.8. The Role of Iron-Depository Dps

We tested the idea that CN recruits iron for Fenton's reagent from two major intracellular depots of the bound iron. One of such iron depository is Dps, a major nucleoid-organizing protein of stationary cells. Its dodecamers form ball-like structures that bind DNA non-specifically and organize it in a paracrystalline form for preservation during non-growth state (25), as well as offering long-term storage of iron by converting it into the oxidized form within the balls (16) (Fig. 2.5. A). Although accumulated in massive numbers only in stationary cells, Dps is also present in 5-10-times fewer copies in exponentially-growing cells (1), while its synthesis is induced in growing cells by oxidative damage or starvation (2). We found that exponential cultures of ∆dps mutants are not different from WT in their lack of sensitivity to H2O2 alone (suggesting no general increase in free iron), but they are killed to a greater extent by CN+H2O2 (Fig.

2.5. B), showing that Dps counters CN potentiation in exponential cells ("Fig. 2.2. D" pattern), perhaps by sequestering the CN recruited iron.

Since the major role of Dps is during the stationary phase, we next tested the sensitivity to the two treatments of wild-type cultures sampled at different densities from rapid growth all the way to complete saturation. We found that, as cells enter stationary phase, their sensitivity to CN+H2O2 treatment completely disappears (Fig. 2.5. C, the green curve), in parallel with disappearance of chromosomal fragmentation (Fig. 2.5. C, the purple curve and Fig. 2.5. D), suggesting that CN+H2O2 induced chromosomal fragmentation requires active metabolism. Similarly, starving the exponentially growing

82 cultures for two hours in M9 salts protects them against the killing. In contrast, stationary

∆dps cultures, while still resistant to H2O2, remain fully sensitive to CN+H2O2 (Fig. 2.5.

EF), while CN+H2O2 still induces catastrophic chromosomal fragmentation in ∆dps mutants in early stationary phase (Fig. 2.5. G), further stressing the link between chromosomal fragmentation and the loss of viability. At first, we detected no CN+H2O2 induced fragmentation in the fully stationary ∆dps mutant cells (Fig. 2.5. G), even though they were killed by CN+H2O2 just like exponential ∆dps mutant cells (Fig. 2.5. E), suggesting a major discrepancy between the two readouts. However, diluting fully- stationary cultures 10-fold into a fresh medium just before the treatment makes ∆dps mutants again susceptible to CN+H2O2 induced fragmentation (wild-type stationary cells continue to be resistant) (Fig. 2.5. HI), suggesting an activation step. Since the energy production is blocked by CN in these cells, we are not sure about the nature of this activation, but the chromosomal fragmentation is blocked by iron chelator deferoxamine, suggesting participation of free iron. Diluting stationary cultures 10-fold into a

"conditioned" medium failed to revive fragmentation in the ∆dps mutant, ruling out cell density as the fragmentation-inhibiting factor. We conclude that the resistance of stationary WT cells to CN+H2O2 is due to Dps organizing the chromosome and/or sequestering CN mobilizable iron, but even in exponentially-growing cells Dps already offers considerable protection against CN+H2O2 treatment.

2.3.9. The Role of Iron-Depot Ferritin

In fact, Dps is a member of the ferritin family of proteins, present in all organisms from bacteria to mammals. All ferritins have a highly-conserved structure of thick-walled spheres accumulating insoluble ferric iron. Besides Dps, there are two more ferritins in E.

83 coli: the ferritin proper (homologous to all ferritins from other kingdoms) and bacterioferritin (a heme-containing ferritin found only in bacteria) (68). In contrast to the smaller 12-mer Dps spheres, which hold up to 500 iron atoms, the larger 24-mer ferritin spheres can hold up to 4,500 iron atoms (68). Another important difference from Dps, which functions as a long-term iron depository, is that ferritins act as dynamic iron distribution centers, both accumulating and releasing iron (Fig. 2.6. A), prompted by interactions with iron-containing proteins or by some simple chemical cues (63).

Although CN was not reported to release iron from ferritin, we decided to use ferritin mutants to test if CN could be one of those "iron-release" cues.

The ∆bfr mutant lacking bacterioferritin shows essentially the wild-type behavior in H2O2 or CN+H2O2 treatments, but the∆ ftnA mutant lacking regular ferritin shows the

"Fig. 2.2. E" pattern of reduced sensitivity to CN+H2O2 (Fig. 2.6. B), suggesting that ferritin does participate in CN potentiation of H2O2 toxicity, perhaps by releasing its iron upon CN cue. The ∆ftnA ∆bfr double mutant shows an intermediate phenotype of some

CN+H2O2 resistance. Although no difference is seen in the level of CN+H2O2 induced fragmentation at 5 or 15 minutes of treatment (see above), the level of chromosomal fragmentation at 1 or 2 minutes of the treatment in all the mutants showing resistance to

CN+H2O2 (∆ftnA, ∆fre, ∆sodAB) is only half of the level in wild-type cells (Fig. 2.6.

CD), corroborating our earlier conclusion that the early fragmentation is the cause of death.

In order to use the plasmid relaxation assay to monitor ferritin-driven Fenton's reaction in vitro, we introduced EDTA in the reaction buffer to block the background reaction with free iron (Fig. 2.6. EFG). We found that iron-loaded (horse spleen) ferritin

84 does not promote Fenton's reaction with H2O2 alone, but does promote it with CN+H2O2

(Fig. 2.6. EF). Apoferritin (the same protein shell, but supposedly lacking iron) is much less active in this nicking reaction (Fig. 2.6. EF). Moreover, addition of iron chelators completely eliminate this residual activity of apoferritin (Fig. 2.6. G), indicating that this activity is due to traces of iron still present in apoferritin preparations. Not only Fenton's reagent is not formed by free iron under these reaction conditions, but also it is not stimulated by CN (Fig. 2.6. EG). Thus, the in vitro evidence with horse spleen ferritin is consistent with the genetic evidence that CN recruits iron from ferritin to fuel Fenton's reagent.

The partial resistance of the ftnA mutants to CN+H2O2 is not further increased by additional fre defect (Fig. 2.6. H), suggesting that ferritin and flavin reductase work together in CN mobilization of reduced iron. On the other hand, the double ∆ftnA ∆dps mutant is as sensitive to CN+H2O2 as the single dps mutant (Fig. 2.6. I), suggesting that, in the absence of the iron depository Dps, CN recruited iron from sources other than ferritin could still kill. We conclude that CN potentiation of H2O2 toxicity is in part due to iron recruitment from ferritins by CN, perhaps even with direct delivery of the recruited iron to DNA. The CN recruited iron then either fuels the DNA-self-targeting Fenton's reagent or is removed from DNA by Dps, the long-term iron depository of E. coli.

2.4. DISCUSSION

Our investigation into the spectacular co-toxicity of cyanide and hydrogen peroxide, which by themselves are bacteriostatic at the corresponding concentrations, yielded the following original findings: 1) rapid exponential cell death is accompanied by catastrophic chromosomal fragmentation; 2) CN potentiates H2O2 killing, but does not

85 kill by itself in any concentration, while H2O2 kills in 10-fold higher concentrations, apparently via self-potentiation; 3) in vitro CN blocks non-DNA Fenton's reaction, but promotes DNA-self-targeting Fenton's reaction, while iron chelators block CN+H2O2 induced chromosomal fragmentation in vivo, suggesting importance of iron-DNA complexes in CN+H2O2 toxicity; 4) the weak or no sensitivity to H2O2 alone of mutants that cannot remove H2O2 (katEG, ahpC) or that elevate "free" cytoplasmic iron (fur, sodAB) suggests a strict system of iron distribution inside the cell, while frequent hyper- sensitivity of these mutants to CN+H2O2 suggests that CN targets the iron-distribution system; 5) the iron depository Dps partially protects against CN+H2O2 killing during growth and offers absolute protection during stationary phase, indicating that Dps neutralizes the CN recruited iron; 6) at the same time, the iron depot ferritin may be one source of CN recruited iron that works together with flavin reductase to fuel Fenton's reagent in vivo.

Our overall results are surprising not only in what we have found, but also in what we failed to find. One of the original objectives of this study was to find targets of CN inhibition that would explain CN potentiation of H2O2 toxicity. Such targets would be identified by mutants that would show a characteristic pattern of similar toxicities of

H2O2 alone treatment versus CN+H2O2 treatment (Fig. 2.2. C). Remarkably, we failed to find a single example of Fig. 2.2. C pattern, instead encountering several examples of

Fig. 2.2. D pattern (Figs.2. 2. F (5 minute time point), 2.2. G, 2.3. C, 2.5. B). This means that CN potentiation does not increase the intracellular concentrations of the two components of Fenton's reagent, H2O2 and iron (Fig. 2.7. A, II), but stimulates Fenton's reaction directly (Fig. 2.7. A, IV). This stimulation could be either at the level of

86 hydroxyl radical formation, or at the level of bringing these radicals to DNA, or both. In fact, CN potentiation could accomplish both by promoting formation of iron-DNA complexes.

With this clarification in mind and on the basis of our new findings, we propose that CN potentiates H2O2 toxicity by fueling Fentons' reagent in at least two ways: 1) by recruiting iron from depots like ferritin directly to DNA; 2) by further activating iron-

DNA complexes for DNA-self-targeting Fenton's reaction. At least two activities, NADH dehydrogenase II and the iron depository Dps, counteract this CN potentiation, the former probably indirectly, via decreasing the pool of reduced flavins, the latter apparently by sequestering CN recruited iron. The proposed hypothetical interplay of ferritin and Dps in iron sequestration and CN recruitment vis-à-vis chromosomal fragmentation is schematically presented in Fig. 2.7. B. We will discuss our specific findings below.

2.4.1. Iron Distribution Inside the Cell and its Subversion by CN

One general conclusion from our results is that distribution of iron is tightly controlled inside the cell. The pool of free cytoplasmic iron in the cell that mixes with freely-diffusing H2O2 and forms hydroxyl radicals throughout the cytoplasm must be limited by iron sequestration in depots (Fig. 2.4. I, bottom). At the least the pool of free cytoplasmic iron is limited in the vicinity of DNA. This conclusion mostly follows from our frequent finding that a particular mutant, expected to be sensitive to H2O2 alone treatment, either due to higher intracellular H2O2 concentrations or increased free cytoplasmic iron, failed to show increased sensitivity for at least the first 10 minutes of

H2O2 treatment (Fig. 2.2. FG, 2.3. C, 2.4. GH and 2.5. B). Since most of these mutants show increased sensitivity to CN+H2O2, the interpretation is that CN disrupts the overall

87 distribution of iron or the restrictions on iron movement around DNA, allowing Fenton's reaction to happen in the vicinity of DNA.

One specific mechanism we propose is direct iron recruitment to DNA from ferritin depots that are themselves in the physical proximity to DNA. Direct iron-DNA interactions are important for DNA damage during in vivo Fenton's reaction (52), but is there any evidence that ferritins are close to DNA inside the cell? It is not a widely- appreciated fact that ferritin readily binds DNA, both in vivo (74) and in vitro (72).

Moreover, in vitro in the presence of ferritin, supercoiled plasmid DNA is gradually nicked, this nicking being dependent on the internal iron, because it is blocked by iron chelators (72). Interestingly, reduced flavin mononucleotides facilitate iron release from ferritin in vitro (26, 54), consistent with our (indirect) conclusion about the role of FMN reductase in the ferritin-mediated fueling of Fenton's reagent in vivo.

2.4.2. Catastrophic Chromosomal Fragmentation

Chromosomal fragmentation does not have to be dramatic to kill. In fact, a single unrepaired double-strand break kills organisms of various levels of complexity, from phages and bacteria to lower and higher eukaryotes (6, 19, 24, 30, 47, 64). Interestingly, one break per chromosome in growing bacterial cultures, where chromosome replicates continuously, will not be even detectable by our physical analysis. Indeed, the chromosomal fragmentation detectable in growing bacterial cells (cultures) by pulsed- field gels as a smear below the compression zone reflects far more than a single break, because the detected sub-chromosomal fragments must have no branches to be able to enter the gel. In other words, in order for random double-strand breaks to generate detectable sub-chromosomal fragments in a theta-replicating chromosome, more than one

88 break needs to happen in the same chromosomal arc. Therefore, quantifying chromosomal fragmentation from the smear within pulsed-field gels seriously underestimates the true density of double-strand breaks in the chromosome. In support of this argument, we reported before that even a 5% level of spontaneous fragmentation corresponds to severe growth inhibition in strains that experience it (49), with 10% spontaneous fragmentation usually meaning complete block of growth (10, 49, 65). In fact, the maximal levels of spontaneous fragmentation of ~15% in the ligA recBC mutants (47) and ~20% in the dut recBC mutants (48, 49) are associated with a high level of lethality.

Even higher levels of fragmentation can be achieved in E. coli cells after DNA damaging treatments — up to 25% fragmentation during thymineless death (51), up to

30% fragmentation after UV irradiation (41), up to 40% fragmentation after phleomycin treatment (42). In all cases, cells were excision-repair-proficient, and there were essentially no survivors. Finally, the maximum level of induced fragmentation approaching 50% was observed in repair-defective polA mutants after UV treatment (42).

But never before have we encountered such a swift and utter chromosome demise as in the case of CN + H2O2 treatment, which we therefore termed "catastrophic chromosomal fragmentation". The actual fragmentation over the background peaks around 70% in this case, but only because some chromosomal DNA is chopped to pieces small enough to migrate out of the gel, while ~10% of the total orthophosphate label in the well is a non-

DNA species (probably fragments of the cell wall) (50). Such a local catastrophic chromosomal fragmentation within an eukaryotic nucleus could be the initial event

89 behind chromothripsis, which is a complete randomization of a particular chromosome or one of its arms, observed with some frequency in certain cancers (36).

There are currently more questions than answers about the possible mechanisms behind this catastrophic fragmentation. Is it dependent on DNA replication, transcription or translation? Does it happen randomly over the chromosome or is concentrated around the origin, for example? Is it associated with Bayer's patches (chromosome's attachments to the envelope) or with the proteinaceous core of the nucleoid (its central part)? Does the nucleoid structure collapse or does it mostly survive during fragmentation? Are double- strand breaks direct or are they preceded by primary (one-strand) DNA lesions? Are these primary DNA lesions repaired by excision repair? Are the double-strand breaks repaired by recombinational repair? Finally, why do fragmentation levels in various mutants poorly correlate with differential survival of these mutants? We would be addressing these questions in our future studies.

2.4.3. Potentiated Toxicity

Potentiated toxicity is a useful practical concept, because it allows an organism to reduce dangers associated with generation and handling of immediately toxic concentrations of a poisonous substance, like H2O2. The transition from bacteriostatic (10 mM) to strongly bactericidal (20 mM) concentrations of H2O2 in our experiments is quite abrupt (Fig. 2.1FG), immediately justifying development of ways to potentiate this toxicity as a strategy to keep the working concentrations of the poison, H2O2, in the static range for the organism that produces it. As mentioned in the introduction, one well- studied example of how H2O2 toxicity is potentiated in real life is offered by immune cells, that use nitric oxide to potentiate killing of pathogenic bacteria with H2O2 (9, 58).

The proposed mechanism of this potentiated toxicity is similar to the one proposed before

3+ for CN+H2O2 toxicity: block of respiration increases NADH pools, accelerating Fe 

Fe2+ reduction by Fre (77). In the light of this work, it would be interesting to test if NO potentiates H2O2 induced catastrophic chromosomal fragmentation by the same kind of iron recruitment from intracellular depots, especially since NO is known to cause iron release from ferritins (63).

Another potentiator of H2O2 toxicity is the amino acid cysteine, the only redox- active amino acid. Cysteine is always present in the cell, but its normal levels are not enough to cause problems in combination with low millimolar H2O2 treatment. However, when cells are forced to take in too much cysteine from the environment, they become transiently sensitive to low mM H2O2 concentrations (59). Cysteine directly stimulates

Fenton's reaction in vitro (59), but the in vivo effectors of cysteine potentiation may also include hydrogen sulfide (4). Interestingly, sulfide is known to release iron from mammalian ferritins (13). Besides cysteine, another amino acid that potentiates H2O2 toxicity is histidine (4). Finally, the fact that 10 mM H2O2 is bacteriostatic, while 20 mM is already strongly bactericidal, suggests that H2O2 self-potentiates at higher concentrations. However, our preliminary experiments show that 20 mM H2O2 still kills wild-type cells in the presence of iron-chelator deferoxamine, suggesting a distinct, iron- unrelated mechanism in this case.

2.4.4. Conclusion

Concentrations of H2O2 in excess of 20 mM efficiently kill by causing catastrophic chromosomal fragmentation, yet it is metabolically hard and physiologically dangerous to produce high concentrations of H2O2 as a bio-weapon. Potentiation with an

91 unrelated substance, like CN described in this work, allows organisms to kill their enemies using physiologically-achievable concentrations of H2O2, which may be indeed quite modest inside the cells (32, 67, 76). Cells of our immune system use NO as such a potentiator, but bacteria scavenge NO with several dedicated enzymes (71), decreasing efficiency of this potentiation route. At the same time, CN is readily generated from serum thiocyanate by myeloperoxidase of our immune cells (11, 27, 70, 73), and bacteria typically have no capacity to degrade CN. Therefore, as we have argued before, our immune cells may also use CN as another or co-potentiator of H2O2 toxicity (50). This makes CN+H2O2, NO+H2O2 and similar mixtures the binary bio-weapons. We propose that the iron depot ferritin plays a dual role in CN potentiation: first, ferritin releases small amount of iron on CN cue; second, since ferritin is a DNA-associated protein, the released iron is instantly complexed by DNA, which allows CN to potentiate DNA-self- targeting Fenton's reaction. The overall validity and the details of this complicated potentiation scheme will have to be addressed in future studies.

2.5. TABLES

Table 2.1. E. coli strains and plasmids (all strains except CP909 are in the BW25113 background).

Strain name Relevant Genotype Source/ CGSC# or Construction CP909 nuoG::Tn10 purF (62) BW25113 F-, Δ(araD-araB)567, ΔlacZ4787(::rrnB- 7636, (3) 3), λ-, rph-1, Δ(rhaD-rhaB)568, hsdR514 JW1721-1 ∆katE731::kan 9453 JW3914-1 ∆katG729::kan 10827 TM10 ∆katE732 JW1721-1 –> pCP20, 42°C, screen for KnS TM12 ΔkatG729::kan ΔkatE732 TM10 x P1 JW3914-1 JW3820-1 Δfre-784::kan 10763 JW0598-2 ΔahpC744::kan 8713 JW1095-1 Δndh-771::kan 11791 TM17 nuoG::Tn10 purF BW2113 x P1 CP909 JW0960-1 ΔappC721::kan 8956 JW0421-1 ΔcyoB788::kan 8585 JW0723-2 ΔcydB782::kan 8790 JW0669-2 Δfur-731::kan 8758 JW3879-1 ΔsodA768::kan 10798 JW1648-1 ΔsodB734::kan 9402 TM11 ΔsodB735 JW1648-1–> pCP20, 42°C, screen for KnS TM13 ΔsodA768::kan ΔsodB735 TM11 x P1 JW3879-1 JW0797-1 Δdps-784::kan 8844 JW1893-1 ΔftnA-755::kan 9575 JW3298-1 Δbfr-746::kan 10467 TM12 ΔftnA-756 JW1893-1–> pCP20, 42°C, screen for KnS TM14 ΔftnA-756 Δbfr-746::kan TM12 x P1 JW3298-1 TM15 ΔftnA-756 Δfre-784::kan TM12 x P1 JW3820-1 TM16 ΔftnA-756 Δdps-784::kan TM12 x P1 JW0797-1 Plasmids pCP20 Flp recombinase gene on a temperature- (20) sensitive replicon pMTL20 Cloning vector (14) pES1 fre overproducer (78)

2.6. FIGURES

Figure 2.1. A. A scheme of in vivo H2O2 toxicity and detoxification. Big balls in shades of blue, oxygen atoms (the lighter the shade, the more reactive the compound), small yellow or orange balls, hydrogen atoms. B. A representative spot-titers of treated WT cultures. Time is in minutes. Here and everywhere: unless indicated otherwise, "CN" means 3 mM KCN, while "HP" means 2 mM hydrogen peroxide. C. Kinetics of CN + H2O2 killing (and H2O2-only stasis) of exponential WT cultures. Here and for the rest of the paper, the values are means of 3 or more independent measurements ± SEM. D. Catastrophic chromosomal fragmentation triggered by CN + H2O2 treatment, as detected by pulsed-field gels. Left, ethidium bromide-stained gel (reversed image) to show the size distribution of the fragmentation smear (M, the yeast chromosome markers with their 50 µg/ml kanamycin for 45 minutes (survival 1.5x10-5). Center and right, 94

Figure 2.1. (cont.) 32 scans of P-labeled chromosomal DNA. Time(min) is duration of CN + H2O2 treatment (center) or of 100 µg/ml rifampicin treatment (survival 2.8 x 10-3) (right). E. Quantification of CN + H2O2- or kanamycin- or rifampicin-induced fragmentation from several gels like in "D-right". Here and for the rest of the paper, chromosomal fragmentation is quantified within a particular lane by dividing the signal in the gel by the combined signal in the gel plus well (total signal) and multiplying the product by 100. F. The nature of CN + H2O2 toxicity revealed by CN-alone or H2O2-alone dose- dependence of survival. All treatments were carried out for 45 minutes. Note that the upper X axis (CN-alone) is logarithmic, and the starting point of it is, actually, "0.1". G. A representative pulsed-field gel demonstrating the catastrophic nature of H2O2-alone- induced chromosomal fragmentation. All treatments were carried out for 45 minutes. Concentrations in parentheses are in mM. H. Quantification of the dose-dependence of chromosomal fragmentation by H2O2-alone, from several gels like in "G". H2O2 concentrations are in mM. I. Kinetics of killing and of chromosomal fragmentation by H2O2(20). The latter values are from several gels like in "J". J. A representative pulsed-field gel showing kinetics of fragmentation by H2O2(20).

Figure 2.2. A. WT kinetics (cf. Fig. 1C). B. Increased sensitivity to both H2O2 and CN + H2O2 treatments. C. Increased sensitivity to H2O2 only, approaching the unchanged sensitivity to CN + H2O2. D. Increased sensitivity to CN + H2O2 only. E. Decreased sensitivity to CN + H2O2. F. Kinetics of killing by the two treatments of the double ∆katEG catalase-deficient mutant, compared to WT. G. Kinetics of killing by the two treatments of the ∆ahpC mutant in alkylperoxidase, compared to WT.

Figure 2.3. A. The expanded view of the "electron transport" part of the Fig. 1A scheme. B. Kinetics of CN+ H2O2 killing: the ∆fre mutant shows partial resistance, while Fre overproduction elevates CN+ H2O2 sensitivity of WT cells. The pFre+ plasmid is pES1. C. Kinetics of killing by the two treatments of ∆ndh and nuo mutants. The nuo mutant behaves like WT, whereas ∆ndh is hyper-sensitive to CN + H2O2, but shows no sensitivity to H2O2-alone. D. Comparison of WT chromosomal fragmentation kinetic pattern to the ones in the ∆ndh and ∆fre mutants. E. The two treatment survival of the cytochrome oxidase mutants. For clarity, sensitivity is shown at a single time point of 60 minutes (H2O2-alone) or 45 minutes (CN + H2O2).

Figure 2.4. A. CN inhibits the standard Fenton reaction in vitro via formation of Fe(CN)6. The reaction, which is started by addition of FeSO4 to the indicated concentration to solution containing 2 mM H2O2, is followed spectrophotometrically (absorbance at 440 nm), by disappearance of a colored substance, p-nitrosodimethylaniline (p-NDA). When added, CN is at 3 mM. B. Pre-treatment of the cultures with iron chelators, such as 20 mM deferoxamine or 2 mM dipyridyl, for five minutes before the treatment blocks CN + H2O2 killing. C. Pre-treatment with the iron chelators like in "B" similarly blocks CN + H2O2-induced chromosomal fragmentation. D. CN inhibits Fenton-like in vitro reactions with other transition metals. The reactions contained 50 µM EDTA and 250 µM of the indicated metal and were ran as in "A". The values are means of two independent repetitions. E. CN stimulates in vitro Fenton's reaction with free iron in water or a Tris buffer and plasmid relaxation as a readout. SCD, supercoiled dimer; SCM, supercoiled monomer; RCD, relaxed circular dimer; RCM, relaxed circular monomer. Concentrations were: 250 µM FeSO4, 2 mM H2O2, 3 mM CN, 40 mM Tris-HCl pH 8.0. F. Quantification of several gels like in "E". The values are (RCM/RCM+SCM)x100. G. Kinetics of killing by the two treatments of the ∆fur mutant. H. Kinetics of killing by the two treatments of the ∆sodAB double mutant. I. Our original idea about free cytoplasmic iron (top) and its current evolution (bottom). Small green circles, DNA-accessible Fe2+:orange and brown hexagons, Fe3+-depots.

Figure 2.5. A. Dodecameric Dps protein forms spheres that bind DNA non-specifically and accumulate up to 500 iron atoms per sphere. Orange/brown small circles, Dps dodecamers; black line in the shape of a star, chromosomal (duplex) DNA. B. Kinetics of killing by the two treatments of ∆dps mutant. WT curves from Fig. 1C are shown in grey for comparison. C. OD-dependence of CN + H2O2 killing and fragmentation of wild-type cells. Aliquots of the same culture were challenged with standard CN + H2O2 treatment at various ODs of the culture, and their normalized survival or fragmentation were plotted as a function of OD. In our growth conditions, the maximal density (fully stationary cultures) of WT cells does not exceed 4.0. D. A representative pulsed-field gel of OD-dependence of fragmentation in WT cells after CN + H2O2 treatment. E. Kinetics of CN + H2O2 killing of exponential vs stationary cultures of wild-type cells and ∆dps mutant. F. Stationary cultures killing by H2O2-alone or CN + H2O2 45 minute treatment.

Figure 2.5. (cont.) G. OD-dependence of fragmentation in ∆dps mutants: a representative gel, similar to the one in (D). Cultures were treated without prior dilution. H. A representative pulsed-field gel of CN + H2O2 treatment kinetics to show that ∆dps stationary cultures undergo robust chromosomal fragmentation if diluted 10x into a fresh medium right before the treatment. I. Kinetics of chromosomal fragmentation in wild-type and ∆dps cells, either in growing or stationary/diluted cultures. This is quantification of several independent runs like in "H".

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Figure 2.6. A. Ferritin spheres function as iron-distribution centers. Small bright-green circles, Fe2+ iron; small orange circle, Fe3+ iron. B. Kinetics of killing by the two treatments of the ∆ftnA mutant. C. Early chromosomal fragmentation is slower in the ∆ftnA, ∆fre and ∆sodAB mutants — a representative pulsed-field gel of 1' and 2' CN + H2O2 treatments. D. Quantification of the early fragmentation from three gels like in "C". In this case, fragmentation values at 0 time point are subtracted as a background. E. In vitro Fenton reaction with ferritin as a source of iron and plasmid relaxation as a readout. The left panel, kinetics of ferritin + CN + H2O2, with CN-only and H2O2-only controls (both at 120 minutes). All reaction have 5 µg of ferritin. The right panel, the apoferritin (also 5 µg, 120 minutes) and the pure iron (the indicated amount, 2 minutes) controls. Rm, relaxed monomer; scm, supercoiled monomer. F. Qauntitative plasmid relaxation kinetics from several gels as in "E".

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Figure 2.6. (cont.) G. The influence of iron chelators on the plasmid relaxation by ferritin and apoferritin (120 minutes). DF, deferoxamine; DP, dipyridyl. Plasmid relaxation by pure iron in the reaction conditions (treatment length is 2 minutes) is also shown. H. Kinetics of killing by the two treatments of the ∆ftnA ∆fre double mutant. I. Kinetics of killing by the two treatments of the ∆ftnA ∆dps double mutant.

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Figure 2.7. A. Interpretation of genetic results. The intracellular H2O2, iron and OH· pools are shown by spheres whose size reflect the relative size of the pools. Stack of straight lines at the bottom, chromosomal DNA; broken lines, broken DNA. (I) Our initial H2O2-only treatment scenario, in which both H2O2 and iron pools are actively limited by dedicated cellular functions, genX and genY. (II) Our initial expectation of a typical CN- potentiation route (inactivation of the H2O2- or iron-limiting functions). (III) The expected phenotypes of the corresponding mutants (in functions that are CN-targets) in H2O2-only treatment. (IV) An alternative scenario suggested by the most frequently observed phenotypes of mutants (no H2O2-only sensitivity, increased CN + H2O2 sensitivity), according to which CN stimulates Fenton's reaction near DNA. B. The proposed role of ferritin-like proteins in CN potentiation of H2O2 poisoning via catastrophic chromosomal fragmentation, based on phenotypes of the corresponding mutants. Top row: yellow/orange smaller circles, Dps dodecamers; blue/orange bigger circles, ferritin 24-mers; black line in the shape of a star, chromosomal (duplex) DNA. Middle row: small green circles, released ferrous iron (note a lighter color of the ferritin circles). In the absence of ferritin, some ferrous iron is still coming from other (unknown) sources. Bottom row: broken black lines, chromosome fragments. The released iron is taken into Dps spheres (note their darker color). For clarity, DNA fragments are shown dissociated from ferritin and Dps.

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CHAPTER 3: THE MECHANISM, CONSEQUENCES AND ATTEMPTED

REPAIR OF THE CATASTROPHIC CHROMOSOMAL FRAGMENTATION

INDUCED BY THE BACTERICIDAL COMBINATION OF CYANIDE AND

HYDROGEN PEROXIDE IN ESCHERICHIA COLI

3.1. INTRODUCTION

2+ Hydrogen peroxide (H2O2) can oxidize cytoplasmic ferrous ions (Fe ) to produce toxic hydroxyl radicals (•OH) that can damage a host of biomolecules causing oxidative

2+ 3+ • stress via Fenton’s reaction (Fe + H2O2  Fe + OH + OH–). Cells of our immune system use Fenton's reaction to kill the invading microbes. Indeed, hydrogen peroxide in concentrations higher than 20 mM can efficiently kill microorganisms (18, 32). At the same time, it is metabolically impossible for eukaryotic cells to generate H2O2, an uncharged molecule, in high concentrations inside the cell, besides it being potentially toxic. Therefore, potentiation of physiologically relevant amounts of H2O2 by various small molecules could serve as a more feasible and safe mechanism to combat the invading microbes.

NO potentiation of H2O2 toxicity is a known bactericidal strategy employed by macrophages, the mechanism of which has been well studied (36, 46), but bacteria scavenge NO with several dedicated enzymes (42), decreasing the efficiency of this potentiation route. In the past, we and others have reported that a combination of low millimolar dosages of hydrogen peroxide and cyanide (CN), which are individually bacteriostatic, caused up to four orders of magnitude synergistic killing in E. coli (19, 20,

32, 47). We have shown that CN could promote the release of iron from iron depots like

110 ferritin directly onto the DNA both in vitro and in vivo, thereby fuelling Fenton’s reaction and causing catastrophic chromosomal fragmentation underlying H2O2 toxicity (32).

Although CN may seem like a non-biological potentiator, it is in fact produced by many plants and fungi and even by some bacteria (22). In humans, the myeloperoxidase enzyme catalyzes the formation of hydrogen cyanide (HCN) from serum thiocyanate

(SCN-) upon neutrophil activation upon encountering bacterial infection (6, 14, 41, 44), yet the implications of this capability are yet to be appreciated. As we have previously reported that a mixture of NO, CN and H2O2 is unusually bactericidal (26), we propose that this synergistically toxic combination could function as an efficient lethal agent against invading pathogens in activated neutrophils or phagocytizing monocytes. Our aim is to better understand the power of this antimicrobial response of our immune cells by investigating the mechanism and consequences of this catastrophic chromosomal fragmentation resulting from CN potentiation of H2O2 toxicity in bacterial cells.

Oxidative DNA damage by hydrogen peroxide comprises one-strand DNA breaks, as well as modified DNA bases (hypoxanthine, 5,6-dihydrothymine, fapy-G, 8- oxo-G), that are repaired via excision of the affected stretch of the DNA strand (as described previously in section 1.2.3.1.). Thus, the current chromosomal fragmentation mechanisms must assume one-strand DNA interruptions as a starting point. A confirmed model of one-strand DNA interruptions leading to double-strand breaks and chromosomal fragmentation in E. coli is replication fork collapse, which results from a replication fork running into a one-strand interruption in the template DNA (27).

Replication fork collapse is observed in polA and ligA mutants, which accumulate one- strand gaps in their DNA due to their compromised nick translation and ligation abilities,

111 respectively (21, 23, 27). We reported previously that replication fork collapse is also observed in dut mutants due to one-strand interruptions generated as a consequence of increased uracil incorporation and subsequent excision (24). In both cases, the conversion of one-strand interruptions into DSBs depends on DNA replication, in the absence of which no chromosomal fragmentation is detectable.

In E. coli, the base excision repair pathway (BER) removes base modifications, such as oxidized or methylated bases. The first step in BER is removal of these modified bases from the DNA by DNA glycosylases that cleave the N-glycosidic bonds producing abasic sites. Incisions are then made by the abasic site endonucleases, such as Exo III

(XthA) or Endo IV (NthA), at the 5’-side of these abasic sites to facilitate the removal of the sugar-phosphates and subsequent closure of the one-strand gap via DNA polymerase I and ligase. On the other hand, double-strand breaks are repaired by the ATP-dependent

RecA mediated homologous recombination pathway. In this pathway, recombination is initiated by binding of RecBCD, the ATP-dependent helicase/nuclease complex, to the exposed ends of the DSBs leading to formation of a 3’ overhang to promote assembly of the RecA filament. This RecA filament, upon searching and locating a homologous sequence in the chromosome, catalyzes strand invasion, giving rise to the X-shaped

Holliday junction, which is resolved via the RuvABC resolvasome or via translocation by the RecG DNA pump. Double-strand break repair is completed when DNA polymerase I

(PolA) fills in the remaining gaps and its ends are joined via the DNA ligase (LigA).

We have recently shown that the synergistic CN + H2O2 treatment kills E. coli by inducing catastrophic chromosomal fragmentation (32). However, the DNA-specific mechanism behind this catastrophic chromosome fragmentation as well as its

112 consequences for the nucleoid remained unexplored. Here we seek to answer four major questions: 1) Are the double-strand breaks generated directly or are they formed as a result of replication fork collapse at one-strand interruptions in the template DNA? 2) To what extent is this DNA damage repairable, and what is the role and relative significance of recombinational repair and base excision repair pathways? 3) How is the nucleoid structure affected by the CN + H2O2-induced chromosomal fragmentation? Even today the structure and dynamics of bacterial nucleoid remains a mystery. The DNA in the nucleoid is organized in supercoiled loops, the lengths of which have been reported to be between 10 kb to 100 kb, coiled around a proteinaceous core (17, 37, 38, 48) but whether these loops are mostly static or dynamic is unknown. Here we report how catastrophic

DNA damage affects nucleoid disassembly and what cellular processes or nucleoid associated proteins (if any) contribute to promoting or hindering this unraveling.

3.2. MATERIALS AND METHODS

3.2.1. Strains and Plasmids

Escherichia coli strains used are all K-12 BW25113 derivatives (2) except for dnaA46, dnaC2 and dut recBC (Ts), which are in the AB1157 background (Table 3.1.).

Alleles were moved between strains by P1 transduction (34). The mutants were all deletions from the Keio collection (2) (unless indicated otherwise), purchased from the E. coli Genetic Stock Center, and were verified by PCR or, phenotypically. For double mutant construction, the resident kanamycin-resistance cassette was first removed by transforming the strain with pCP20 plasmid (10).

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3.2.2. Reagents

Hydrogen peroxide and diethylamine NONOate were purchased from Sigma.

Potassium cyanide and sodium azide were purchased from Fisher-Scientific.

3.2.3. Growth Conditions and Viability Assay

To generate killing kinetics, fresh overnight cultures were diluted 500-fold into

LB medium (10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, 250 μl of 4 M NaOH per liter (34)) and were shaken at 37°C for about two and a half hours or until they reached exponential phase (OD600 approximately 0.3). At this point, the cultures were made 3 mM for CN and/or 2 mM for H2O2 (or the indicated treatment) and the shaking at 37°C was continued. In order to measure survival/revival in cells treated with CN and H2O2 for 45 minutes, the cells were spun down and resuspended in fresh LB and allowed to grow at

37°C post-treatment. Viability of cultures was measured at the indicated time points by spotting 10 μL of serial dilutions in 1% NaCl on LB plates (LB medium supplemented with 15 g of agar per liter). The plates were incubated overnight at 28 °C, the next morning colonies in each spot were counted under the stereomicroscope. All titers have been normalized to the titer at time 0 (before addition of the treatment).

3.2.4. Measuring Chromosomal Fragmentation via Pulsed-Field Gel Electrophoresis

This follows our previous protocol and is the same as described in 2.2.4.

3.2.5. Southern Hybridization of Chromosomal Fragmentation

At the indicated time-points, samples were collected in duplicates and non- radioactive plugs were made and run on pulsed field gels, as described above. The plugs were taken out of the wells after the run and transferred to glass tubes and both the pulsed-field gels and the plugs were washed with 0.25 M HCl, followed by 0.5 M NaOH,

114 and finally, with 1 M Tris HCl pH 8.0. Each wash was 40 minutes long. The treated plugs were then placed on Amersham Hybond N+ (GE Healthcare) nylon membrane, covered with Saran wrap, and DNA was transferred by vacuum for 1-2 hours, while the DNA from the treated pulsed-field gels were transferred overnight via capillary transfer. Next, the DNA was UV-crosslinked to the membranes and probed with 32P-labeled ori- or ter- specific probe. Hybridization was carried out overnight at 63°C in a 0.5 M sodium phosphate (pH 7.4) and 5% SDS hybridization buffer. In the morning, the membranes were washed thrice with 1% hybridization buffer and rinsed with water just before covering them with Saran wrap and exposing to a phosphorImager screen. The resulting signals were measured with a PhosphorImager (Fuji Film FLA-3000).

3.2.6. Isolation of rRNA

We followed the RNAsnap procedure (40) to isolate rRNA from 32P labeled exponential cultures of cells (OD600 approximately 0.3) grown at 37°C. The rRNA samples were precipitated out via ethanol reprecipitation ( 0.1X 5 M NaCl and 2X ethanol were added to 1X volume of rRNA sample and mixed vigorously by vortexing.

The samples were then spun down at 16000g for 5 minutes and the supernatant was completely removed via aspiration and the pellet was resuspended in TE) prior to running them on a 1.1% agarose gel at 60V for 2 hours. After the run, the gel was dried and exposed to a phosphorImager screen. The resulting signals were measured with a

PhosphorImager (Fuji Film FLA-3000).

3.2.7. Measuring Linear DNA Degradation

This generally follows our previous protocol (23). Cells were grown overnight in

1 ml of LB and 10 µCi (methyl-3H)-thymidine at 28°C. In the morning, cells were spun

115 down and washed thrice with 1 ml of LB, then diluted 200 times into LB and were allowed to grow at 37°C for 2 hours. Cells were then treated with 3 mM for CN and 2 mM for H2O2 for an hour after which the cultures were split into halves. CN and H2O2 was removed from one half via centrifugation and the cells were resuspended in LB, subsequently both halves were again allowed to grow at 37°C for 2 hours. At the indicated times, 4 ml aliquots were taken and mixed with 4 ml of chilled 10% trichloroacetic acid. Cells were collected by filtration and prepared for scintillation counting as described in our previously published works (1, 23).

3.2.8. Measuring Nucleoid Disassembly

Overnight cultures were diluted 500-fold into 25 ml LB medium containing 25 -

75 µCi 32P and were shaken at 37°C for about two and a half hours or until they reached exponential phase (OD600 approximately 0.3). At this point, total DNA plugs were made of 300 µl aliquots of untreated cultures as described above. The cells were then treated with 3 mM CN and 2 mM H2O2 (or the indicated treatment) and were continued to grow at 37°C for 45 minutes. Following this, CN and H2O2 were removed from these cultures by centrifugation and the cells were resuspended in LB. To isolate nucleoid core free

DNA, we used a modified version of the total plasmid isolation protocol (16). At the indicated time-points, 3 ml aliquots were spun down and resuspended in 50 µl 30%

Sucrose dissolved in 50 mM Tris HCl (pH 8) and 10 mM EDTA. 350 µl of 2% SDS was then added to it, mixed by inversion and placed at 70°C for 5 minutes. Later, 100 µl 5M

NaCl was added to these samples and mixed thoroughly via inversions and allowed to rest on ice for at least an hour. Next, the samples were spun down at 16000 X g for 20 minutes and the supernatant was transferred to a fresh tube. In order to precipitate nucleic

116 acids, 1 ml of ethanol was added and the samples and they were spun down at 16000g for

5 minutes, after which the supernatant was aspirated and the pellet was resuspended in 20

µl TE. To degrade RNA, 30 µl LiCl was added, mixed by vortexing and the samples were allowed to rest on ice for 15 minutes. The samples were then spun down and the supernatant was transferred to a fresh tube. Nucleic acids were again precipitated with

100 µl ethanol as described above and the DNA was resuspended in 20 µl TE. 10 µl of each sample was used to run on a 1.1 % agarose gel run at 60 V for 3 hours. The gel was then dried and exposed to a PhosphorImager screen. The resulting signals were measured with a phosphorImager (Fuji Film FLA-3000).

3.3. RESULTS

3.3.1. Nucleoid Organization Plays a Small Role in Protecting the Chromosome from CN + H2O2-Induced Fragmentation, while Bulky DNA Lesions are not

Involved

To gain insights into the primary DNA lesions and chromosomal consequences of

CN(3) + H2O2(2) treatment (3 mM KCN + 2 mM H2O2), we used survival and chromosomal fragmentation as the two readouts with select mutants lacking either DNA repair enzymes or specific nucleoid-associated proteins. In our previous study, we have reported that the DNA binding protein, Dps, present in a few copies in rapidly-growing cells, but massively-induced in stationary phase cells, protects the wild-type cells from

CN + H2O2 killing by blocking chromosomal fragmentation (32), so we first tested whether mutants in other major nucleoid-associated proteins would have an effect in the two readouts. However, we observed that only the Δfis mutant is slightly more sensitive

117 to the treatment, whereas ΔhupA, Δihf, Δhns and ΔstpA mutants display CN + H2O2 sensitivities similar to the wild-type (WT) (Fig. 3.1. AB).

The two major pathways for repair of one-strand DNA lesions in E. coli are nucleotide excision repair (NER) and base excision repair (BER). In addition, there is also the most basic pathway to close one-strand interruptions, catalyzed by DNA Pol I and DNA ligase that serves both one-strand repair pathways. The NER-deficient uvrA and ΔuvrB mutants behave like the wild-type when treated with CN + H2O2 (Fig. 3.1. C), indicating that NER has no role in mending the CN + H2O2 induced DNA lesions. In other words, CN + H2O2 induced lesions are unlike the bulky or DNA helix-distorting lesions repaired by NER.

In contrast, mutants in the base-excision repair or in one strand-gap closing proved to be so sensitive to CN + H2O2 killing that we had to develop much milder treatment regimens to observe some survivors at the earliest time points. We found that reducing CN concentration 10-fold does not affect the early rate of killing of WT cells, while reducing H2O2 concentration 10-fold decreases the early rate of killing somewhat

(Fig. 3.1. D). At the same time, if the concentration of both CN and H2O2 is reduced by

10-fold, the treatment becomes bacteriostatic for WT cells (Fig. 3.1. D), as we have already reported before (32). Chromosomal fragmentation after 5 minutes of CN + H2O2 treatment is not affected dramatically when 10 times lower CN concentration is used, whereas only a 10% increase in fragmentation over the background is detected when both

CN and H2O2 concentrations are decreased 10 fold (Fig. 3.1. EF). For the sensitive mutants we used 0.3 mM CN + 2 mM H2O2 conditions; we reserved the 0.3 mM CN +

118

0.2 mM H2O2 conditions that do not kill wild-type cells only for the extremely sensitive mutants.

3.3.2. Base-Excision Repair and One-Strand Gap Repair Mend the Bulk of CN +

H2O2-Induced DNA Lesions

We blocked base-excision repair at the critical stage of abasic site nicking, inactivating either the major abasic site endonucleases (XthA) or both abasic site endonucleases (XthA and Nfo) (7). Both xthA and xthA nfo mutants are hypersensitive to the CN + H2O2 treatment (Fig. 3.2. A) and exhibit massive chromosomal fragmentation compared to the wild-type cells under similar conditions (see Fig. 3.2. BC for xthA nfo and Fig. 3.3. BC for xthA alone). Remarkably, these abasic site endonuclease mutants are not sensitive to the corresponding H2O2-alone treatment (Fig. 3.2. A), meaning that abasic sites are not produced by H2O2 acting alone, yet they are massively produced when cyanide potentiates the same H2O2 treatment.

The most sensitive mutants to CN + H2O2 killing turned out to be the one-strand gap repair mutants, polA and ligA (the difference in the treatment doses is because the polA12 mutant has the mildest DNA pol I defect known (43), in contrast to our ligA251 mutant, which has the strongest ligase defect known (23, 29)). Remarkably, unlike the abasic site endonuclease mutants, one-strand gap repair mutants show similar sensitivities to both CN + H2O2 and H2O2 alone treatments (Fig. 3.2. DE) and exhibit similarly enhanced catastrophic chromosomal fragmentation in response to both treatments (Fig.

3.2. FGH). In other words, H2O2 alone treatment is not innocuous and generates a lot of one-strand breaks, which DNA pol I and DNA ligase promptly repair, while CN- potentiation appears to add little to the number of these "direct" one-strand breaks.

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Taken together, the results from the most sensitive mutants suggest that one- strand breaks dominate among H2O2-induced lesions, while CN potentiation of H2O2 treatment leads to the appearance of significant numbers of base modifications, removed by base-excision repair via the abasic site intermediate (Fig. 3.2. I). Perhaps even more surprisingly, without immediate repair, a significant fraction of these one-strand lesions is converted into double-strand breaks, which we detect as catastrophic, "pulverizing" chromosome fragmentation. The mode of distribution of sizes of the chromosomal fragments in ligase mutants after CN(0.3) + H2O2(0.2) treatment is ~50 kbp, translating to at least 100 double-strand breaks per genome-equivalent.

3.3.3. Recombinational Repair Mends H2O2-Induced Double-Strand Breaks but is

Paralyzed by CN

Since the major pathway to repair double-strand DNA breaks in E. coli is recombinational repair (28) (Fig. 3.3. A), recombinational repair mutants were expected to show extreme sensitivity to CN + H2O2 treatment. It is known that recombinational repair mends double-strand breaks induced by hydrogen peroxide alone (30), and we have confirmed these observations for 2 mM H2O2, both genetically (Fig. 3.3. B) and physically (Fig. 3CD). Surprisingly, deficiency in double-strand break repair due to the

ΔrecA, or ΔrecBCD, or the double recG ΔruvABC defects translates into only a minor additional sensitivity to CN + H2O2 treatment compared with the WT cells (Fig. 3.3. E) and to almost no additional effect on chromosomal fragmentation (Fig. 3.3. FG). In fact, recombinational repair mutants are much less sensitive to CN + H2O2 treatment compared to the mutants in BER or one-strand gap repair. It is obvious that the additional killing effect in recombinational repair mutants is entirely due to the modest sensitivity of these

120 mutants to the corresponding H2O2-alone treatment (Fig. 3.3. E). This means that recombinational repair does not participate in mending double-strand breaks in the presence of CN, but it does efficiently repair a small number of double-strand breaks due to H2O2 alone treatment.

Indeed, this conclusion was expected, as CN should completely block recombinational repair enzymes, since all of them require ATP hydrolysis for catalysis

(28), by poisoning ATP production. Moreover, because of this dependence of recombinational repair on ATP-hydrolysis, and the ATP-production inhibition by CN, we originally considered recombinational repair a likely target of CN-potentiation. However, a much higher sensitivity of recombinational repair mutants to CN + H2O2 treatment compared to H2O2 alone treatment (Fig. 3.3. E) argues against this possibility.

To reveal the role of recombinational repair in mending CN + H2O2-induced chromosomal fragmentation, we removed CN by pelleting cells and resuspended them in fresh medium to allow them to resume ATP-production, and then followed both the culture titer and the level of fragmentation. We detected no repair after 45 minute CN +

H2O2 treatment (not shown), apparently due to too many double-strand breaks overwhelming the repair capacity. In contrast, after a 5 minute treatment and removal of

CN + H2O2, there was almost three orders of magnitude recovery in the culture titer (to

30% of the original titer) (Fig. 3.3. H), accompanied by a complete disappearance of chromosomal fragmentation (Fig. 3.3. IJ). The median size of sub-chromosomal fragments is ~500 kbp after 5 minutes of treatment (Fig. 3.3. H), translating into ~10 double-strand breaks per genome-equivalent. There was neither recovery of the culture titer, nor significant repair of chromosomal fragmentation in the recA single mutant or in

121 the recG ΔruvABC mutant (Fig. 3.3. HIJ), indicating that both phenomena are due to recombinational repair. There was also no recovery or repair in the xthA nfo mutant, in which the density of double-strand breaks after even 5 minutes of treatment is much higher and is similar to the density of double-strand breaks in WT cells after a 45 minute treatment (the median size of sub-chromosomal fragments is ~50 kbp (Fig. 3.3. I), translating into ~100 double-strand breaks per genome-equivalent). Therefore, we conclude that when ATP-production resumes, recombinational repair is still capable of reassembling fragmented chromosomes if there are fewer than 10 double-strand breaks per genome equivalent, but its capacity is saturated when the density of double-strand breaks is increased 10-fold.

3.3.4. CN + H2O2-Induced Double-Strand Breaks do not Depend on DNA

Replication and are Uniformly Distributed Over the Chromosome

The extremely high density of double-strand breaks (~100 per genome equivalent) is the unique hallmark of CN + H2O2-induced catastrophic chromosome fragmentation, but the nature of these high density double-strand breaks is perplexing. Indeed, as our genetic analysis suggests, the primary DNA lesions caused by H2O2 are one-strand interruptions, while CN potentiation causes additional base modifications, which eventually also translate into one-strand interruptions. Typically, one-strand interruptions by themselves cannot break DNA; they cause chromosomal fragmentation only during the replication-segregation transition, as either replication fork collapse or segregation fork collapse events. Therefore, the subchromosomal fragments released as a result of fork collapse events in asynchronous cultures have the length distribution from close to zero to the full chromosomal size (24, 25), which is not what we observe after 45 minutes

122 of CN + H2O2 treatment that results in the chromosomal DNA being chopped into uniformly short fragments (Fig. 3.1. E).

To test the replication-dependence of CN + H2O2-induced fragmentation, we utilized dnaA(Ts) and dnaC(Ts) mutants, that do not initiate new replication rounds at

42°C, but finish the ongoing replication rounds, so that after 2 hours of incubation at

42°C, their chromosomes lack replication forks. We found that even after 2 hours at 42°C the dnaA(Ts) and dnaC(Ts) mutants are killed with exactly the same kinetics as the corresponding WT strain (Fig. 3.4. D), while the level of chromosomal fragmentation in them is roughly two times lower (Fig. 3.4. EF), but still, apparently enough to kill them.

Another way to align the chromosome is to block protein synthesis with chloramphenicol for two hours. The chloramphenicol block does reduce the killing of WT cells slightly

(Fig. 3.4. A) and again reduces the overall fragmentation about two-fold (Fig. 3.4. BC).

However, the overall phenomenon looks the same and still kills by several orders of magnitude, demonstrating its general independence of replication/segregation events (and even of protein synthesis).

Furthermore, if the dnaA(Ts) and dnaC(Ts) cultures are not diluted before shifting them to 42°C, they start to saturate by the time of CN + H2O2 treatment and lose the ability to fragment, but they are still killed to the same extent by the treatment. A similar effect is observed in the overnight stationary cultures of Δdps mutants, and again, prior deep dilution before the treatment restores the typical one-to-one relationship between the loss of viability and chromosomal fragmentation.

A strong prediction of the replication-dependent chromosomal fragmentation is a specific format of the resulting subchromosomal fragments: since they result from the

123 demise of replication bubbles, they all must carry the replication origin (Fig. 3.4. G, left)

(24). In contrast, the replication terminus sequence is expected to be only on the chromosome-size fragments (found in the compression zone under our PFGE conditions), but absent on shorter fragments (Fig. 3.4. G, left) (24). This is illustrated by analyzing the chromosomal fragmentation smear in the dut recBC mutant with Southern hybridization: the entire smear hybridizes to the origin-specific probe, whereas the terminus-specific probe shows a much reduced signal (Fig. 3.4. HI). Chromosomal fragmentation in the dut recBC mutant critically depends on the ongoing DNA replication and serves as a positive control for this analysis (24). Remarkably, the uniformly-sized chromosomal fragments after 45 minutes of CN + H2O2 treatment hybridize equally well to both the origin and the terminus-specific probes (Fig 3.4. HI), confirming complete replication-independence and uniform chromosomal distribution of CN + H2O2 -induced double-strand breaks (Fig.

3.4. G, right). The uniform distribution of the double-strand breaks over the chromosome means that some of them happen in the unreplicated part of the chromosome, making them irreparable by definition and explaining cell death.

3.3.5. Double-Strand Breaks are not Associated with DNA Found in Bayer’s Patches or Other Chromosomal DNA Attachment Sites

Next we asked whether these uniformly-distributed double-strand breaks are random, or they are associated with the chromosomal DNA attachment sites. The chromosomal DNA in both bacteria and eukaryotes is organized as rosettes of radial loops (13). Such loops in E. coli start at the nucleoid core in the middle of the cell and extend all the way out to the inner membrane. If not separated from the associated protein, intact chromosomal DNA precipitates as a single nucleoid complex, while

124 smaller DNA molecules (like plasmids) that are protein-free ("naked") stay in solution

(16). We decided to use this protocol of plasmid-like DNA preparation to isolate pieces of the chromosomal DNA that separated from the nucleoid as a result of double-strand breaks and to inquire into a possible relation between these breaks and the nucleoid structure.

At least two types of the nucleoid attachment sites are known in E. coli: the base of the nucleoid loop attachment at the nucleoid core at the perigee vertex of the loops and the loop attachment at the cell envelope at specific sites called Bayer's patches at the apogee vertex of the loops (Fig. 3.5. A) (4) (33). For example, Bayer's patches are absent in stationary cells, and the resistance of stationary phase wild-type cells to the CN + H2O2 treatment and to the accompanied DNA damage maybe a reflection of preferential DNA breakage at Bayer's patches. Remarkably, the three types of double-strand breaks

(random vs. nucleoid core vs. the envelope) predict distinct patterns of fragmented naked

DNA release from the overall nucleoid structure (Fig. 3.5. A). Random breaks predict some DNA will be released from the nucleoid. If double-strand breaks happen only at the nucleoid core instead then a substantial amount of DNA would be released from the nucleoid (because only two breaks per loop is enough to release the entire loop). In contrast, double-strand breaks at Bayer's patches should release no DNA from the nucleoids (Fig. 3.5. A).

To assess the impact of CN + H2O2 treatment on the integrity of the nucleoid structure and to distinguish between the possibilities mentioned above, DNA was isolated from CN + H2O2 treated cells using the total plasmid isolation protocol at the indicated time-points (Fig. 3.5. B) and run on an agarose gel. Total chromosomal DNA isolated in

125 agarose plugs at time 0 is used for normalizing the amount of naked DNA released from the nucleoid. The amount of naked DNA recovered by this protocol from untreated cells is quite stable at 2% of the total (Fig. 3.7. BC). At the same time, after 45 minutes of treatment with CN + H2O2, ~4% of the total genomic DNA is released from nucleoid

(Fig. 3.7. BC), suggesting that ~2% of the chromosomal DNA lost its association with the nucleoid, ostensibly due to double-strand breaks in the DNA loops. Taking into account the above reasoning and at the density of 50-100 double-strand breaks per genome equivalent, this low, but detectable level of released DNA is most consistent with the random break model. The other two models of breaks at the nucleoid DNA attachments, predict either release of "almost everything" (breaks at the nucleoid core) or, quite the contrary, release of "nothing" (breaks at Bayer’s patches).

If breaks are random, increase in their density should release more naked DNA from the nucleoid. This is indeed observed in the Δdps mutant cells (Fig. 3.5. D) and especially in the base-excision-repair-deficient mutant xthA nfo that releases almost 30% of the total DNA as naked fragments (see below). To verify that double-strand break position is not influenced by nucleoid-associated proteins, we also quantified “release of naked DNA” after 45 minutes of CN + H2O2 treatment in the corresponding mutants. We found that ΔhupA and Δihf mutants release the “WT amounts” of naked DNA, whileΔ fis and ΔstpA mutants release significantly more naked DNA, suggesting that the corresponding proteins help protect DNA from breakage (Fig. 3.5. D). Interestingly, the

Δhns mutant shows reduced DNA release, suggesting that CN + H2O2-induced double- strand breaks are targeted to H-NS-DNA binding sites. Therefore, we conclude that CN +

H2O2-induced double-strand breaks are not targeted to chromosomal DNA attachments at

126 either the nucleoid core in the center or at Bayer’s patches at the periphery of the cell, but these breaks may be still targeted to specific proteins bound to DNA, like H-NS.

3.3.6. Chromosomal DNA and rRNA Degradation After CN + H2O2 Treatment

The amount of linear DNA released from the WT nucleoids was surprisingly low, suggesting relatively small size of the nucleoid loops. However, there was also a possibility that the catastrophic chromosomal fragmentation detected by pulsed-field gels was an artifact of the DNA-preparation procedure or of the separation technique themselves, while no breaks actually happened in vivo. Perhaps the most direct confirmation of massive chromosomal fragmentation inside the cell, rather than during subsequent in vitro DNA isolation, would be equally massive subsequent chromosomal

DNA degradation inside the treated cells, like the one observed in gamma-irradiated E. coli cells (5). However, our initial attempts at detecting chromosomal DNA degradation even after 60 minutes of CN + H2O2 treatment were futile (Fig. 3.5. EF), although we quickly realized why this was the case. Linearized DNA with unprotected ends are degraded in E. coli by the ATP dependent RecBCD enzyme complex, and the lack of degradation reflected the fact that cyanide blocks respiration and ATP-production in aerobic cells. Indeed, once we removed CN + H2O2 by resuspending cells into fresh medium, we detected massive chromosomal DNA loss (20% of the chromosome was degraded in two hours) (Fig. 3.5. E), similar in the rate and extent to the chromosomal

DNA degradation after gamma-radiation. Producing additional RecBCD enzyme from a medium-copy-number plasmid pDWS2 further increases the rate and extent of this degradation (35% of the chromosomal DNA lost in two hours) (Fig. 3.5. F), also showing that the reaction is not significantly limited by the availability of the enzyme. Thus,

127 double-strand breaks do form inside the cell as a result of CN + H2O2 treatment, but the fragmented chromosome is stable because cellular metabolism is frozen in the presence of CN.

We have also observed that ribosomal RNA (rRNA) is also significantly damaged during the CN + H2O2 treatment (Fig. 3.5. G). This indicates that CN + H2O2 damages all nucleic acids. We tested whether the remaining rRNA could be further degraded due to the lack of ATP in the presence of CN, and followed rRNA stability in CN + H2O2 treated cells after resuspending them in fresh medium. Indeed, we found that rRNA signal weakens and smears after change of the cells into fresh medium (Fig. 3.5. GH), showing that for their subsequent recycling, ATP-dependent processes are required

(possibly due to dependence on RNA helicases to disassemble the complex ribosome structure), requiring CN removal.

3.3.7. Nucleoid Dynamics of Broken DNA Results in Partial Nucleoid Disintegration

Although the structure of the nucleoid as a rosette of similar-sized radial loops looks static and suggests designated attachment sites on DNA separating independently- supercoiled domains (loops), several attempts to find these designated domain boundaries, or in fact any short-range periodicity in the nucleoid structure, have failed.

However, four separate 0.5-1.0 Mbp-size macrodomains bundling up to a hundred individual DNA loops were identified, which restricteded interaction of macrodomain

DNA with any DNA outside any of these macrodomain, but no internal structure within these macrodomains was revealed (12). The explanation for this highly uniform structure without any determinants of stable periodicity was found to lie in the dynamic nature of the DNA-nucleoid attachments. In fact, the DNA is constantly moving through the

128 nucleoid attachments, like a rope through a pulley, with no lasting contact of any particular sequence with any particular protein structure of the nucleoid (Fig. 3.6. A).

However, no experimental data to support this model was offered. We decided to test this

“rotating contacts” model using broken DNA after CN + H2O2 treatment.

When the CN + H2O2-treated cells are resuspended in fresh LB, they resume ATP production, which will restart, among other processes, their nucleoid dynamics and elusive DNA rotation. Above we have reported that naked DNA release from the wild- type nucleoid is minimal in CN + H2O2-treated cells (a mere 2% (Fig. 3.5. BC)), suggesting random nature of DNA breaks and their low density relative to the average size of the nucleoid loops. We reasoned that resumption of DNA-nucleoid rotation after

CN + H2O2 treatment should release more broken naked DNA from the nucleoids.

Transferring WT cells into a fresh medium after CN + H2O2 treatment indeed released an additional 10% of the total DNA within 2 hours, half of this amount being released within the first 15 minutes. One of the processes participating in this DNA-nucleoid rotation is proposed to be transcription by RNA polymerase (11). Indeed, blocking transcription initiation with rifampicin reduces the amount of naked DNA released after ATP restoration to about half of the uninhibited level (Fig. 3.6. BC), confirming the role of transcription in this double-strand break-promoted nucleoid disintegration.

We measured the extent of double-strand break-promoted nucleoid disintegration following the release of naked DNA after CN + H2O2 removal in several mutants that were lacking various nucleoid-associated proteins (9). The only mutant that reduced the

WT amounts of released DNA was Δhns (Fig. 3.6. FG), suggesting H-NS protein participation in DNA-nucleoid rotation. The Δihf and ΔhupA mutants released similar to

129

WT amounts of naked DNA, while the Δdps mutant releases similar amounts, but starting from a much higher background during the treatment itself, indicating more initial breaks

(Fig. 3.6. CF). Interestingly, the ΔstpA andΔ fis mutants not only released more naked

DNA, but also failed to saturate even after two hours after CN + H2O2 removal (Fig. 3.6.

FG), suggesting that both StpA and Fis act to interfere with the DNA-nucleoid rotation in

WT cells. Remarkably, a similar (but moderate) effect was shown by the ΔrecBCD mutants (Fig. 3.6. DE), deficient in the double-strand break repair, even though no repair was detectable either physically or genetically after 45 minute treatment. The limitation of double-strand break-promoted nucleoid disintegration by activities of double-strand break repair can be rationalized by RecBCD-promoted RecA filament assembly at the broken ends in preparation for their subsequent repair: the bulky RecA filament will block the end rotation through the nucleoid pulley (Fig. 3.6. H). Finally, the base-excision repair mutant xthA nfo releases ten-fold more DNA upon CN + H2O2 treatment, but this extremely high nucleoid disintegration is not further enhanced by availability of ATP

(Fig. 3.6. DE), suggesting that DNA with unrepaired abasic sites cannot freely rotate through the nucleoid pulleys. Thus, we have identified two variables (the density of double-strand breaks and transcription) and one nucleoid associated protein, H-NS, that promote nucleoid disintegration. We have also found proteins and processes that interfere with nucleoid disintegration: abasic sites, nicks, StpA, Fis, HU, IHF and recombinational repair of double-strand ends.

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3.3.8. A Comparison of Chromosomal Fragmentation Induced by the CN + H2O2 with Other Treatments that Potentiate H2O2 Toxicity

Finally, we investigated the robustness of death and DNA damage caused by other agents, such as azide and NO, that are capable of potentiating hydrogen peroxide toxicity just like cyanide. We found that the early kinetics of death was similar for NO +

H2O2 and CN + H2O2 (Fig. 3.8. A). However, because NO is actively degraded in the cells by nitric oxide dioxygenase, this allows the surviving cells to recover at later time- points, which is why we actually see an increase in titer at these times. azide + H2O2 , on the other hand, behaves very differently from CN + H2O2, and shows a shoulder of resistance for about half an hour before succumbing to a very sharp death at 45 minutes.

The chromosomal fragmentation observed for these synergistic treatments correspond very well to their killing kinetics. In the case of NO + H2O2, we find that the fragmentation increases steadily at early time-points, like that of CN + H2O2, however, the fragmentation observed in NO + H2O2 is significantly lower (Fig. 3.8. BC). At later times, the chromosomal fragmentation begins to recover for NO + H2O2 because of increased survival. Similarly, azide+ H2O2 shows maximum fragmentation at the last time-point, but again, is much lower than that exhibited by CN + H2O2. This suggest that the mechanism of potentiation of hydrogen peroxide toxicity for these compounds may be significantly different than CN + H2O2. We also tested whether cell death and chromosomal fragmentation were dependent on iron in NO + H2O2 or azide + H2O2, as is the case with CN + H2O2, by carrying out the treatment in the presence of the iron chelator, deferoxamine (DF). We confirmed that NO + H2O2 co-toxicity did depend on iron (or the Fenton chemistry), however, loss in viability and chromosomal fragmentation

131 due to the azide and H2O2 treatment was found to be only partially blocked by DF

(perhaps due shut-down of metabolism) (Fig. 3.8. ABD). This indicates that the synergistic toxicity of azide and H2O2 occurs via a distinct iron-independent mechanism that remains to be studied.

3.4. DISCUSSION

We have shown that the synergistic cyanide and hydrogen peroxide treatment not only induces a robust iron-dependent catastrophic chromosomal fragmentation similar to that induced by NO + H2O2, but also results in a complete destruction of rRNA, indicating the susceptibility of all nucleic acids to this co-toxic regimen. Iron-dependent oxidative damage to chromosomal DNA inside the cell by hydrogen peroxide, thought to mostly affect individual DNA strands, also causes some double-strand breaks that were attributed to replication fork collapse at one-strand DNA lesions and their excision repair intermediates. When studying cyanide potentiated hydrogen peroxide-induced double- strand breaks, we found that their number is amplified by inactivation of base-excision repair, suggesting that unrepaired primary DNA lesions are directly converted into double-strand breaks, while their sheer numbers rule out the involvement of replication forks. Indeed, blocking DNA replication reduces CN + H2O2 -induced fragmentation only slightly, without affecting the survival.

Interestingly, this catastrophic chromosomal fragmentation was found to be completely repairable in wild-type cells treated with CN and H2O2 for 5 minutes, as they exhibited a dramatic 1,000-fold revival upon removal of CN + H2O2, while base excision repair or recombinational repair deficient mutants mutants were unable to survive. Even though recombinational repair can mend the first dozen of the early CN + H2O2 -induced

132 double-strand breaks (once cyanide is removed), it still cannot assemble the chromosome broken into 100 pieces, suggesting that repair is saturated.

Termination of the CN + H2O2 treatment, which allows the cells to synthesize

ATP again, was found to trigger significant linear DNA degradation and disintegration of the nucleoid structure. Our investigation into the consequences of catastrophic chromosomal fragmentation induced by CN + H2O2 on the nucleoid using various repair- and nucleoid associated protein-deficient mutants has allowed us to gain some insights on the nucleoid structure, and particularly, on nucleoid dynamics. Release of broken DNA from the nucleoid is limited, consistent with random position of the breaks and generally short nucleoid loops. In repair deficient mutants, the nucleoid disassembly was exaggerated, which is not surprising, given the fact that these mutants accumulate more

DNA damage than the wild-type cells. The disassembly of the nucleoid structure was also enhanced in all mutants lacking nucleoid associated proteins except Δhns, which, curiously, showed diminished nucleoid disintegration compared to the wild-type.

Inhibition of transcription via rifampicin also diminished nucleoid disintegration, suggesting that the nucleoid structure is dynamic and in part depends on transcription, and while most nucleoid associated proteins prevent the disassembly of nucleoid structure via their binding, H-NS and processes like transcription promote its unravelling.

Therefore, we conclude that double-strand breaks induced by oxidative damage happen at the sites of unrepaired primary one-strand DNA lesions, are evenly-distributed along the chromosome and can provide insights into the dynamic nature of the nucleoid structure.

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3.5. TABLES

Table 3.1. E. coli strains and plasmids (all strains are in the BW25113 background unless indicated otherwise). Strain name Relevant Genotype Source/ CGSC# or Construction AB1157 F− λ− rac- thi-1 hisG4 Δ(gpt-proA)62 argE3 (3) thr-1 leuB6 kdgK51 rfbD1 araC14 lacY1 galK2 xylA5 mtl-1 tsx-33 glnV44 rpsL31. L-216 AB1157 dnaA46(Ts) (15) lac/CE ori::bla Elena Kouzminova L-393 AB1157 dnaC2 (Ts) (45) 10827 AK4 AB1157 ∆(srlR-recA306)::Tn10 (8) JB1 AB1157 ∆recBCD3::kan (35) N2731 AB1157 recG258::Tn10 (31) JJC754 AB1157 ∆ruvABC232::cat (39) AK25 AB1157 polA12(Ts)::Tn10 Lab Collection GR501 Hfr(PO45), λ, ligA251(ts), relA1, spoT1, thiE1 6087 LA20 GR501 ligA251 ΔypeB::kan (1) SX1253 F-, ∆(argF-lac)169, gal-490, ∆(modF- 12808 ybhJ)803, λ[cI857 Δ(cro-bioA)], xthA791- YFP(::cat), IN(rrnD-rrnE)1, rph-1 BW535 F-, thr-1, araC14, leuB6(Am), ∆(gpt- 7047 proA)62, lacY1, tsx-33, glnX44(AS), galK2(Oc), λ-, Rac-0, nth- 1::kan,ble, ∆(xthA- pncA)90, hisG4(Oc), rfbC1, mgl-51, nfo- 1::kan, rpsL31(strR),kdgK51, xylA5, mtl- 1, argE3(Oc), thiE1 N3055 F-, λ-, IN(rrnD-rrnE)1, rph-1, uvrA277::Tn10 6661 JW0762-2 ΔuvrB751::kan 8819 BW25113 F-, Δ(araD-araB)567, ΔlacZ4787(::rrnB-3), λ- (2) , rph-1, Δ(rhaD-rhaB)568, hsdR514 JW0797-1 Δdps-784::kan 8844 TM21 ∆(srlR-recA306)::Tn10 BW2113 x P1 AK4 TM22 ∆recBCD3::kan BW2113 x P1 JB1 TM23 ∆ruvABC232::cat BW2113 x P1 JJC754 TM24 recG258::Tn10 ∆ruvABC232::cat TM23 x P1 N2731 TM25 xthA791-YFP(::cat) BW2113 x P1 SX1253 TM26 xthA791-YFP(::cat) nfo-1::kan TM25 x P1 BW535 TM 27 ligA251 ΔypeB::kan BW2113 x P1 LA20 TM 28 polA12(Ts)::Tn10 BW2113 x P1 AK25

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Table 3.1. (cont.)

TM 29 uvrA277::Tn10 BW2113 x P1 N3055 JW3229-1 Δfis-779::kan 10443 JW1702-1 ΔihfA786::kan 9441 JW1225-2 Δhns-746::kan 9111 JW3964-1 ΔhupA771::kan 10851 JW2644-3 ΔstpA750::kan 10083

Plasmids pCP20 Flp recombinase gene on a temperature- (10) sensitive replicon

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3.6. FIGURES

Figure 3.1. A. Kinetics of death of wild-type (WT) and Δfis mutants when treated with 2 mM H2O2 (HP) alone or with 3 mM KCN and 2 mM H2O2 (CN+HP) in growing cultures. Here and in the rest of the paper, all values are means of 3 or more independent measurements ± SEM. B. Same as in "A", but with the Δhns ΔhupA, Δihf and ΔstpA mutants lacking nucleoid organizing proteins. C. Same as in "A", but with the uvrA and ΔuvrB mutants. D. Kinetics of death of WT cells treated with 2 mM H2O2 in the presence of varying concentrations of CN (3 mM, 0.3 mM and 0.1 mM). Note that no killing is observed when concentrations are reduced 10-fold from the standard concentrations: 0.3 mM CN and 0.2 mM H2O2. E. A representative pulsed-field gel demonstrating chromosomal fragmentation induced in WT treated with 3 mM CN and 2 mM H2O2, 0.3 mM CN and 2 mM H2O2 or 0.3 mM CN and 0.2 mM H2O2. F. Kinetics of chromosomal fragmentation upon treatment with 3 mM CN and 2 mM H2O2, 0.3 mM CN and 2 mM H2O2 or 0.3 mM CN and 0.2 mM H2O2 (from several gels like in "E"). Here and for the rest of the chapter, fragmentation level at any time point is normalized to the fragmentation level at time = 0.

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Figure 3.2. A. Kinetics of death of the xthA nfo double mutant treated with 0.3 mM CN + 0.2 mM H2O2 or 0.2 mM H2O2 alone. B. A representative pulsed-field gel demonstrating chromosomal fragmentation induced in the xthA nfo double mutant treated with 0.3 mM CN + 0.2 mM H2O2 compared to WT cells. C. Quantification of the kinetics of chromosomal fragmentation in xthA nfo and WT cells upon treatment with 0.3 mM CN + 0.2 mM H2O2 (from several gels like in "B"). D. Kinetics of death of the polA12 (Ts) mutant and WT cells treated with 0.3 mM CN + 2 mM H2O2 or 2 mM H2O2 alone at 42°C. E. Kinetics of death of the ligA251(Ts) mutant and WT cells treated with 0.3 mM CN + 0.2 mM H2O2 or 0.2 mM H2O2 alone at 42°C. The ligA251(Ts) mutant and WT cells are pre-grown at 28°C for two and a half hours prior to addition of 0.3 mM CN + 0.2 mM H2O2 or 0.2 mM H2O2, after which they are shifted to 42°C for the duration of the treatment. An untreated ligA control is also included to demonstrate that the mutant does not begin to die due to the ligase defect for up to 15 minutes after being shifted to 42°C.

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Figure 3.2. (cont.) F. A representative pulsed-field gel demonstrating chromosomal fragmentation induced in the ligA mutant treated with 0.3 mM CN + 0.2 mM H2O2 or 0.2 mM H2O2 alone compared to WT at 42°C (along with untreated ligA control), and in the WT and polA mutant treated with 0.3 mM CN + 2 mM H2O2 at 42°C. G. Quantification of the kinetics of chromosomal fragmentation upon treatment with 0.3 mM CN + 2 mM H2O2 in polA and WT cells (from several gels like in "F"). H. Quantification of the kinetics of chromosomal fragmentation upon treatment with 0.3 mM CN + 0.2 mM H2O2 or 0.2 mM H2O2 alone in ligA and WT cells (from several gels like in "F"). I. A hypothetical scheme depicting H2O2-alone or CN + H2O2-induced DNA damage, one-strand gap repair and base-excision repair in E. coli.

138

Figure 3.3. A. A scheme of double-strand breaks repair in E. coli. B. Kinetics of survival of the WT cells, as well as ΔrecA, ΔrecBCD or recG ΔruvABC mutants treated with 2 mM H2O2 alone. C. A representative pulsed-field gel demonstrating chromosomal fragmentation induced in the WT cells, as well as ΔrecA and ΔrecBC mutants with 2 mM H2O2 alone. D. Quantification of the kinetics of chromosomal fragmentation upon treatment with 2 mM H2O2 alone in ΔrecA and ΔrecBC mutants compared to WT cells (from several gels like in "C"). E. Kinetics of death of the ΔrecA, Δ recBCD or recG ΔruvABC mutants treated with 0.3 mM CN + 2 mM H2O2 or 2 mM H2O2 alone. For comparison, death kinetics of WT cells treated with the same concentrations of CN and H2O2 is also plotted. F. A representative pulsed-field gel demonstrating chromosomal fragmentation induced in the ΔrecA and ΔrecBC mutants treated with 0.3 mM CN + 2 mM H2O2 compared to WT and xthA cells.

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Figure 3.3. (cont.) G. Quantification of the kinetics of chromosomal fragmentation upon treatment with 0.3 mM CN + 2 mM H2O2 in ΔrecA and ΔrecBC mutants compared to WT and xthA cells (from several gels like in "F"). H. Kinetics of survival/’revival’ of WT cells treated with 3 mM CN + 2 mM H2O2 for 5 minutes. After 5 minutes of treatment, CN and H2O2 is removed and the cells are resuspended in fresh LB and allowed to recover at 37°C. Repair deficient mutants, such as ΔrecA, recG ΔruvABC and xthA nfo, were included as negative controls. Growth rate of the untreated WT culture was also monitored in parallel. I. A representative pulsed-field gel showing the disappearance of catastrophic chromosomal fragmentation in WT cells induced by 5 minute CN + H2O2 treatment, upon removal of the treatment. In contrast, ΔrecA, recG ΔruvABC and xthA nfo after the same treatment show no rapid decrease in the levels of chromosomal fragmentation. J. Quantification of the disappearance of catastrophic chromosomal fragmentation in WT cells induced by 5 minute CN + H2O2 treatment upon their removal compared to the lack of it in ΔrecA, recG ΔruvABC and xthA nfo (from several gels like in ‘I’).

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Figure 3.4. A. Kinetics of death of non-replicating WT cultures pre-treated with 40 µg/ml chloramphenicol (Cam) for 2 hours before treatment with 3 mM CN and 2 mM H2O2. B. A representative pulsed-field gel demonstrating the catastrophic chromosomal fragmentation induced in chloramphenicol pre-treated non-replicating WT cells by the 3 mM CN and 2 mM H2O2 treatment compared to the fragmentation observed in exponentially growing CN+H2O2 treated WT cells.

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Figure 3.4. (cont.) C. Quantification of the kinetics of chromosomal fragmentation upon treatment with 3 mM CN and 2 mM H2O2 (from several gels like in ‘B’) in chloramphenicol pre-treated non-replicating WT cells. For comparison, chromosomal fragmentation in exponentially growing WT cultures treated with 3 mM CN and 2 mM H2O2 is also plotted. D. Kinetics of death of non-replicating dnaA46(Ts) and dnaC2(Ts) cultures in the AB1157 background (compared to exponential AB1157 cultures) upon treatment with 3 mM CN and 2 mM H2O2. The AB1157, dnaA46(Ts) and dnaC2(Ts) mutants were pre- grown at 28°C for two hours and then shifted to 42°C for two hours, following which the CN+H2O2 treatment was carried out at 42°C. E. A representative pulsed-field gel demonstrating the catastrophic chromosomal fragmentation induced in non-replicating dnaA46(Ts) and dnaC2(Ts) cultures by the 3 mM CN and 2 mM H2O2 treatment (compared to the fragmentation observed in exponentially growing CN+H2O2 treated AB1157 cells). The AB1157, dnaA46(Ts) and dnaC2(Ts) mutants were pre-grown at 28°C for two hours and then diluted 20 times and shifted to 42°C for two hours, following which the CN+H2O2 treatment was carried out at 42°C. F. Quantification of the kinetics of chromosomal fragmentation upon treatment with 3 mM CN and 2 mM H2O2 (from several gels like in E) in non-replicating dnaA46(Ts) and dnaC2(Ts) cultures. For comparison, chromosomal fragmentation in exponentially growing AB1157 cultures treated with 3 mM CN and 2 mM H2O2 is also plotted. G. Schemes of chromosomal fragmentation patterns induced by replication fork collapse (i), or by random replication-independent breaks (ii). H. Blot hybridization of chromosomal fragmentation with origin-specific or terminus- specific probes, of WT cells treated with 3 mM CN and 2 mM H2O2. For efficient transfer, the plugs were taken out of the wells after the run and transferred separately. The dut recBC(Ts) strain was used as the "replication fork collapse" control. I. Quantification of hybridization of the CN+H2O2-fragmented chromosomes to the origin-specific and terminus-specific probes (compared to the dut recBC control) from several gels like in "H".

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Figure 3.5. A. A scheme of double-strand break positioning versus linear DNA release from the nucleoid. A cell cross-section is shown with double-strand breaks (CN + H2O2 hits) indicated by small red stars. The nucleoid-free DNA is isolated and analyzed. B. A representative agarose gel image demonstrating the linear DNA release from nucleoid after 3 mM CN and 2 mM H2O2 treatment in growing WT cultures. To provide a reference for the total chromosomal DNA, at time zero, the total DNA from 1/10th of the culture volume is collected in agarose plugs; the total signal from both the well and the lane is collected. C. Quantitative kinetics of linear DNA release from the nucleoid during CN + H2O2 treatment from several gels like in ‘B’. D. Nucleoid-associated proteins protect DNA from CN + H2O2 -induced breaks. The indicated mutants were treated with CN + H2O2 for 45 minutes, and the level of nucleoid- free linear DNA was determined. E. Kinetics of linear DNA degradation observed upon removal of CN and H2O2 from WT cells treated for 60 minutes. F. Kinetics of linear DNA degradation observed upon removal of CN and H2O2 from WT cells over-expressing RecBCD from the plasmid pDSW2. G. A representative agarose gel image demonstrating the loss in rRNA subsequent to 3 mM CN and 2 mM H2O2 treatment in growing WT cultures, as well as after removal of CN and H2O2 from cultures. H. Quantitative kinetics of rRNA loss from several gels like in ‘G’.

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Figure 3.6. A. A scheme of the nucleoid organization, with pulleys for DNA attached at both the nucleoid core, as well as at the inner membrane. Duplex DNA is shown as the blue cord being threaded through the pulleys. B. A representative agarose gel image showing further release of linear DNA from nucleoid upon removal of CN and H2O2 in WT cells treated for 45 minutes, and the effect of addition of 100 µg/ml Rifampicin for 5 minutes prior to removal of CN and H2O2. C. Quantitative kinetics of the results from several gels like in ‘B’, including the Δdps mutant results. D. A representative agarose gel image showing an increased release of linear DNA from the nucleoid upon removal of CN and H2O2 from cultures of the ΔrecBCD mutant or the xthA nfo mutant treated for 45 minutes. E. Quantitative kinetics from several gels like in ‘D’. F. A representative nucleoid disintegration assay of the Δdps, Δhns, ΔstpA and Δfis mutants. G. Quantitative kinetics of the results from several gels like in ‘F’. H. A scheme showing processes/proteins that enhance linear DNA release from the nucleoid in red and its inhibitors in blue. Dps acts to lower one-strand gap formation.

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Figure 3.7. A. Kinetics of death of WT cells subjected to several synergistic treatments: 3 mM CN + 2 mM H2O2, or 3 mM sodium azide (AZ) + 2 mM H2O2, or 0.3 mM NO + 2 mM H2O2. B. A representative pulsed-field gel showing the kinetics of chromosomal fragmentation of WT cells subjected to 3 mM CN + 2 mM H2O2, or 3 mM AZ + 2 mM H2O2, or 0.3 mM NO + 2 mM H2O2. Cells treated with azide or NO alone were also included as controls. C. Quantitative kinetics of chromosomal fragmentation of WT cells subjected to 3 mM CN + 2 mM H2O2, or 3 mM AZ + 2 mM H2O2, or 0.3 mM NO + 2 mM H2O2 from several gels like in ‘B’. D. Quantification of chromosomal fragmentation of WT cells subjected to 3 mM CN + 2 mM H2O2, or 3 mM AZ + 2 mM H2O2, or 0.3 mM NO + 2 mM H2O2 in the presence or absence of the iron chelator, deferoxamine (DF), from several gels like in ‘B’.

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CHAPTER 4: CHROMOSOMAL OVER-REPLICATION AND ITS UNUSUAL

CONSEQUENCES IN THE ESCHERICHIA COLI SEQA MUTANT

4.1. INTRODUCTION

Faithful replication of the parental chromosome and proper organization, condensation and segregation of the resultant daughter chromosomes is crucial for bacterial cells, as disruption of these processes directly affects viability. Integrity of the chromosomal DNA is continuously compromised by reactive oxygen species, insertion of improper nucleotides, or by mistakes in the complex processes of chromosome replication and segregation (7). The cell employs various strategies to both prevent and correct these aberrations before the damage becomes lethal. There is a variety of repair pathways to mend DNA lesions, including double-strand break repair by homologous recombination. A double-strand break is particularly deleterious, since the cellular nucleases, e.g. RecBCD, can efficiently degrade unprotected double stranded DNA, making chromosome vulnerable to gene loss. The RecA protein is instrumental in repair of such lesions via homologous recombination with intact sister duplexes. If there is a double-strand break in DNA, then the RecBCD helicase catalyzes RecA polymerization at the double-strand ends. RecA performs strand exchange, which is followed by

RuvABC- or RecG-catalyzed resolution of Holliday junctions holding the recombination intermediates together (Fig. 1.2.) (8).

The Escherichia coli SeqA protein negatively regulates initiation of DNA replication by preventing re-initiation at the newly-replicated chromosomal origins, oriC, via binding to pairs of hemimethylated GATC sequences separated by one, two or three

150 helical turns (positioned on the same face of DNA), thereby preventing the replication initiator protein, DnaA, from binding to the origin of replication prematurely (2, 10).

SeqA is a 20 kDa membrane-associated protein with an N-terminal domain (amino acid residues 1-59) facilitating dimerization of the protein and formation of filaments, whereas its C-terminal domain (amino acids 71-181) is involved in DNA-binding (18). Moreover,

SeqA negatively regulates the expression of the DnaA itself, thus playing a role in inhibition of primary initiations as well (4). In addition, SeqA binding sites also transiently occur in the hemimethylated newly-replicated DNA, possibly facilitating organization of the newly-synthesized daughter duplexes. Specifically, SeqA tracts have been proposed to guide the newly-synthesized daughter DNAs to the sites of formation of the daughter nucleoids, thus aiding in chromosome segregation (3). Reflecting these potential multiple roles of the SeqA protein, its inactivation is pleiotropic: seqA mutant cells overinitiate (10), show replication asynchrony (10), have slow forks (14), fragment their chromosomes (13), exhibit an increased rate of chromosome mis-segregation (10), hyper-negative chromosomal DNA supercoiling (19) and cell envelope defects (13).

Over-initiation, slowing down of replication forks (1) and chromosomal fragmentation (16) has been observed in cells overexpressing the replication initiator protein, DnaA. The seqA mutants also have elevated levels of DnaA and similarly exhibit chromosomal fragmentation. We propose that the chromosomal fragmentation in seqA mutants is caused by over-expression of DnaA. This over-expression could inhibit replication forks directly, via increased DnaA-DNA binding inhibiting replication fork progression, or indirectly, by limiting factor(s) essential for fork progression, and thereby slowing down the forks. In the seqA mutant, the presence of numerous replication forks

151 could lead to rear-ending of replication forks into the preceding inhibited forks causing chromosomal fragmentation (Fig. 1.1.). Alternatively, chromosomal fragmentation in the seqA mutant could be caused via the replication fork regress-split model (Fig. 1.1.) (6).

According to this model, in a stalled replication fork, the newly synthesized strands unwind from the daughter duplexes, reverse and instead pair with each other to form a

Holliday junction (HJ), which can be cleaved by RuvABC, resulting in chromosomal fragmentation. A hallmark of the replication fork regress-split is the requirement of linear

DNA degradation, and hence RecBCD for survival, but not for RecA or RuvABC (6).

However, recent work from our lab has shown that replication fork regress-split, in fact, does not contribute to chromosomal fragmentation in the seqA mutants (14).

Moreover, we suspect that SeqA, by binding to the pairs of hemi-methylated

GATC sites in the newly-synthesized daughter DNA, effectively acts like eukaryotic cohesins, and thereby performs sister-chromatid cohesion in E. coli. Disruption of this suspected sister-chromatid cohesion in seqA mutants could lead to problems with proper homologous recombination repair in the newly-synthesized daughter DNA resulting in improper re-attachments of the broken double strand ends to the wrong partners (e.g. the end attaches itself to the homologous sequence in one of the two "cousin" duplexes instead of its own sister duplex) (Fig. 4.1.). This situation could be detrimental to the cell, and in this case, recombinational repair would be poisonous. In contrast, linear DNA degradation capability of the cell could prevent this error in recombination and the associated lethality.

This project aims to investigate the underlying cause and the mechanisms of replication fork inhibition in the seqA mutant. The finding that the seqA mutant depends

152 on linear DNA degradation for viability even in repair-proficient backgrounds is consistent with the idea that SeqA performs sister chromatid cohesion in E. coli, the disruption of which could have grave consequences for the cells that have increased number of replication rounds in their chromosomes, like seqA mutants do.

4.2. MATERIALS AND METHODS

4.2.1. Strains and Plasmids

Escherichia coli strains used are all K-12 in the AB1157 background. Alleles were moved between strains by P1 transduction (11). The mutants were all verified by

PCR or phenotypically. For double mutant construction, the resident kanamycin- resistance cassette was first removed by transforming the strain with pCP20 plasmid (5).

4.2.2. Growth Conditions and Viability Assays

Overnight cultures grown in LB medium (10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, 250 μl of 4 M NaOH per liter (11)) at 28°C were normalized to the same OD, serially diluted and spotted by 10 µl on LB plates (or as indicated). The plates were incubated at 28°C or at the indicated temperature overnight so as to observe and compare surviving colonies.

4.2.3. Color Screen for Identifying SeqA-Dependent Mutants

This screen identifies mutants that depend on a functional seqA gene residing on a plasmid at the temperatures semi-permissive for plasmid replication. We mutagenized the seqA21 lacZ double mutant carrying an ori(Ts) plasmid that complements both the seqA and lacZ defects and identified mutants unable to lose the plasmid (as solidly-colored colonies on the background of sectored colonies). Insertional mutagenesis was carried out using the pRL27 plasmid carrying a hyperactive Tn5 transposase and a separate insertion

153 cassette that comprises a kanamycin resistance marker and the R6K pir-dependent origin of replication, which was generously provided to us by Dr. William Metcalf. These mutants were further streaked at non-permissive temperature for plasmid replication to rule out plasmid multimerization as the cause of solidly colored colonies observed at semi-permissive temperature. To exclude repair deficient mutants from subsequent steps, these mutants were tested for their sensitivity to UV. If not UV sensitive, they were P1 transduced into AB1157 background to confirm SeqA dependence. Mutants that could not grow without the seqA gene in AB1157 background, or those that exhibited UV sensitivity were sequenced.

4.2.4. Measuring Rate of DNA Replication

Overnight cultures were diluted 100-fold and grown to an OD ~ 0.2 in LB at the indicated temperatures. At the indicated time points, 200 µl of this culture was incubated with 200 µl of M9 minimal medium supplemented with 40µg/ml of arginine, histidine, proline, leucine and threonine, 1µCi/ml of 3H-thymidine and 2µg/ml of cold thymidine, for 5 minutes at room temperature. The counts were measured in a liquid scintillation counter after Trichloroacetic acid precipitation and were normalized to time 0 for each strain (taken for "1"). A background control, AB1157 treated with 200J UV radiation was also included; this value was subtracted from all counts as background incorporation.

4.2.5. Flow Cytometry

Overnight cultures were diluted 100-fold and grown to an OD~.2 in LB at 28°C.

These cultures were then diluted to an OD of 0.05 and 150 µg/ml rifampicin was added at time 0. At the indicated time points, 500µl of the cultures were taken, washed with

154 ethanol and Tris-HCl, stained with Cytox Green dye and observed in the FACSCanto

Flow Cytometer.

4.2.6. Southern Hybridization

Same as described previously in section 2.2.7.

4.3. RESULTS

4.3.1. Identification and Characterization of SeqA-Dependent Mutants

The SeqA protein negatively regulates initiation of chromosomal replication preventing both asynchronous initiation and over-initiation from the chromosomal replication origin, oriC (10). We used a color screen, developed in this laboratory (7) to identify SeqA-dependent mutants in E. coli. We screened over 150,000 mutants and sequenced 37 candidates that exhibit synthetic lethality with the ΔseqA defect (Table

4.1.). Of these 37 mutants, we most frequently isolated hits to recB and recC (ten times together) and ruvA (five times). recG was also isolated once, while recA was never isolated, suggesting that the ΔseqA mutants depend more on linear DNA degradation than on recombinational repair for survival, in contrast to what was observed before (7).

4.3.2. seqA Mutants Require Linear DNA Degradation for Survival

The ΔseqA mutant exhibits increased chromosomal fragmentation, and therefore, depends on the recombinational repair proteins RecBCD and RuvABC. However, the

ΔseqA ΔrecA mutant, although severely inhibited, is not completely dead (Fig. 4.2.), suggesting alternatives to recombinational repair of double-strand breaks. We constructed the ΔseqAΔ recD double mutant and found it to be inhibited for growth (Fig. 4.2. A.), which was unexpected, because the ΔrecD mutants generally behave like wild-type cells, with linear DNA degradation reduced only two-fold in them. We then constructed a 155

ΔseqA recA(Cs) ΔrecD triple mutant and confirmed that it was completely dead at the semi-permissive temperature of 28°C (Fig. 4.2. C). This result indicates that inactivation of ExoV linear DNA degradation activity of the RecBCD enzyme along with recBC- dependent homologous recombination activity is responsible for the inviability of the

ΔseqA Δrec mutants, suggesting that recombination is important but not essential for viability unless the RecBCD-dependent linear DNA degradation is also absent.

Furthermore, by transducing the ∆seqA mutation into ΔrecBC sbcA and ΔrecBC sbcBC strains complemented with the seqA gene on a temperature-sensitive plasmid we found that these severely linear-DNA-degradation-impaired strains are inviable in combination with the ∆seqA defect (Fig. 4.3.), despite being fully proficient in recombinational repair of double-strand breaks via the RecF and RecE pathways, confirming that linear DNA degradation is required for the survival of ΔseqA mutant cells.

4.3.3. Over-Replication and Problems with Replication Fork Progression in the seqA, rep and seqA rep Mutants

In the ΔseqA mutant, replication fork inhibition could lead to fork stalling and subsequent disintegration (with eventual repair), further slowing down replication fork progression. We constructed ΔseqA rep, ΔseqA dnaB(Ts) and ΔseqA dnaC(Ts) and found them all to be either severely inhibited or completely dead at temperatures semi- permissive for the single dnaC(Ts) and dnaB(Ts) mutants (Fig. 4.4.). The rep mutant has an elevated ori/ter ratio like ΔseqA mutants. In order to rule out excessive over-initiation as a possible reason for the ΔseqA rep co-inhibition, we measured the ori/ter ratios in seqA rep double mutant cells and found them to be similar to ori/ter ratios in the ΔseqA and rep single mutants (Fig.4.5.), suggesting that the SeqA and Rep proteins work in the

156 same pathway to prevent the ori/ter ratio from being elevated. Since the DnaC protein is involved in replication fork restart (15), we also measured changes in the rate of DNA synthesis upon shift to semi-permissive temperature for AB1157, ΔseqA, dnaC(Ts) and

ΔseqA dnaC(Ts) mutants using 3H-thymidine and observed that the rate of replication of the exponentially growing cells of both single mutants keeps increasing like the wild- type, while in the ΔseqA dnaC(Ts) mutant, the rate of replication levels off upon the shift and continues to stay at the same level as before the shift (Fig. 4.6.). Thus, upon shift to semi-permissive temperature, DNA replication in the ΔseqA dnaC(Ts) double mutant is inhibited, possibly due to its inability to assemble an excess of disintegrated replication forks.

Furthermore, since the ΔseqA mutant is inhibited by sub-lethal concentrations of hydroxyurea (HU), a ribonucleotide reductase inhibitor, that do not affect the wild-type cells (17), we measured the rate of replication in the ΔseqA mutant and the wild-type cells, so as to examine the differences, if any, upon treating them with HU. We found that the rate of DNA synthesis indeed decreases in the ΔseqA mutant upon HU addition and does not recover, unlike the wild-type and rep mutants (Fig. 4.7.), showing that inhibiting replication fork progression in the ΔseqA mutant by either mutating the helicases, dnaB or rep, or via HU-induced ribonucleotide reductase inhibition is not well tolerated. These observations suggest that the replication forks in the absence of SeqA are compromised in a general way that makes them extremely vulnerable to replisome irregularities that are well tolerated by the SeqA expressing cells.

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4.3.4. seqA Mutants are Sensitive to Rifampicin and are Unable to Finish Half of

Their On-Going Replication Rounds in its Presence

As was mentioned before, seqA mutants exhibit replication asynchrony: in contrast to the WT cells, that have 2n number of nucleoids, seqA mutants show any number of nucleoids, from 1 to 16 (10). In order to find conditions relieving the asynchrony phenotype of the seqA mutants, we profiled the kinetics of genome replication of AB1157, seqA and rep through flow cytometry using rifampicin to block transcription and cephalexin to block cell division. We chose to include rep in our analysis because it has been reported to have slower replication forks (9) and has elevated ori/ter ratios like the seqA mutant (Fig. 4.5.), so it would serve as a good negative control for replication asynchrony. The flow cytometry profiles demonstrated that it took wild- type cells 60 minutes to finish its replication rounds, whereas the rep mutant took at least

120 minutes due to its slower replication forks. In contrast to the wild-type and the rep mutant, the seqA mutant appeared to be unable to finish ongoing rounds of replication even four hours after rifampicin addition (Fig. 4.8.). In fact, the seqA mutant seems to make no progress towards finishing replication, as no peaks corresponding to genome equivalents form at any time-point after addition of rifampicin. Expecting a complete block to DNA synthesis by rifampicin in the seqA mutant, we measured the rate of DNA synthesis using 3H-thymidine, with AB1157 (WT control), as well as seqA and rep mutants treated with rifampicin for up to 4 hours. Unexpectedly, we observed that the rate of replication in both seqA and rep mutants did not decrease as rapidly in response to rifampicin as it did in AB1157 (Fig. 4.9. A). In other words, both the seqA and rep mutants continued to replicate for quite some time even though new rounds of initiations

158 were blocked by rifampicin. This was consistent with the presence of extra replication rounds in the chromosome of these mutants compared to WT cells. A somewhat different effect was observed with chloramphenicol, a translation inhibitor that also blocks new replication initiations. Chloramphenicol inhibited replication in the rep mutant and in the wild-type cells to the same extent, but it failed to inhibit replication in the seqA mutant

(Fig. 4.9. B). Thus, surprisingly, the seqA mutants turned out to be quite resistant to replication inhibition via replication initiation block indicating that their inability to form discreet chromosome equivalents (as detected by flow cytometry) was not because they shut down DNA synthesis, but in spite of their inability to stop it.

This inability to finish replication upon transcription inhibition in the seqA mutant is not without consequences, as we have found that the seqA mutant is actually killed by rifampicin at concentrations bacteriostatic to the wild-type (Fig. 4.10.). We also measured the ori/ter ratios upon rifampicin treatment using Southern blot analysis and found that the wild-type ratio decreases from ~2 to ~1, as expected, while the seqA mutant ratio also decreases from 5 to 2.5 (by two-fold, like in wild-type) (Fig. 4.11.).

This result indicates that in the seqA mutant, approximately half of the ongoing multiple replication rounds cannot be completed in the presence of rifampicin. Since the seqA mutant can, in principle, complete replication rounds under the most favorable conditions

(42°C, 300 mM MgSO4) (Fig. 4.12.), we conclude that rifampicin apparently blocks the completion of the remainder of the replication rounds by causing shortage of certain proteins required for fork progression in the seqA mutant.

159

4.4. DISCUSSION

4.4.1. Lack of Sister Chromatid Cohesion in seqA Mutants Could be the Cause of its

Dependence on Linear DNA Degradation

The seqA mutant suffers from increased chromosomal fragmentation, the mechanism of which is consistent with the replication fork breakage scenario (14).

Nevertheless, the recombinational repair protein RecA is not essential for its survival.

The seqA recA double mutant, although severely inhibited, is not completely dead despite being deficient in carrying out recombinational repair, suggesting alternatives to recombinational repair of double-strand breaks in the seqA mutant. At the same time, the seqA recA(Cs) recD triple mutant is completely dead at semi-permissive temperatures, indicating that in the presence of RecBCD-dependent linear DNA degradation, recombination is important but not essential for the viability of the seqA mutant.

Moreover, the inviability of seqA recBC sbcA and seqA recBC sbcBC combinations, which are proficient in recombination via the RecF and RecE pathways, yet grossly deficient in linear DNA degradation, underscores the significance of linear DNA degradation pathways in the seqA mutant. This dependence on linear DNA degradation in the seqA mutant for survival can be explained by disruption of sister-chromatid cohesion leading to improper re-attachments of the broken double strand ends, thus, necessitating the presence of linear DNA degradation pathways for resolution. We propose that SeqA, by binding to pairs of hemi-methylated GATC sites in the newly-synthesized daughter

DNA, effectively acts like eukaryotic cohesins and performs sister-chromatid cohesion in

E. coli. In seqA mutants, disruption of such hypothetical sister-chromatid cohesion could lead to problems with proper homologous recombination repair in the newly synthesized

160 daughter DNA, which coupled with excessive initiation rounds could result in frequent improper re-attachments of the broken double-strand ends to a wrong partner (e.g. the end could attach itself to the homologous sequence in the "cousin" DNA duplexes instead of recombining with its own sister) (Fig. 4.1.). This would explain the inability of the seqA mutant to finish replication rounds in the presence of rifampicin and could even contribute to the rifampicin-induced lethality. However, the precise mechanism and consequences of lack of sister chromatid cohesion in seqA mutants remain to be investigated.

4.4.2. Possible Mechanism of Replication Fork Inhibition in the seqA Mutant

Inhibition of replication fork progression in the seqA mutant by inactivating the replicative helicase, DnaB, or the auxiliary helicase, Rep, is a lethal event. Moreover, inhibiting fork progression by HU-induced ribonucleotide reductase inhibition is also severely inhibitory in the seqA mutant. Inhibition of replication forks leads to chromosomal fragmentation, which could possibly be the cause of lethality in the seqA mutant, because seqA mutant cells may be limited for replication factors(s) and have slower forks more prone to disintegration. Furthermore, inactivation of the DnaC protein, which is involved in the replication fork restart pathway, also causes the seqA mutants to die, suggesting that in the absence of optimal replication fork reloading, the frequent replication fork disintegration leads to DNA fragmentation and lethality. Therefore, replication forks in the absence of SeqA are compromised in a way that makes them extremely vulnerable to replisome irregularities that are otherwise well tolerated by the

SeqA containing cells. DnaA over-expression in the seqA mutant could be responsible for at least some of these phenotypes. The seqA mutants are reported to have twice the

161 amount of DnaA protein found in the wild-type cells, perhaps due to the absence of negative regulation of the dnaA gene by the SeqA protein (12). Elevated DnaA levels have previously been shown to cause chromosomal fragmentation, and remarkably, problems with finishing ongoing rounds of replication in the presence of rifampicin (1).

This DnaA over-expression in seqA mutants could inhibit replication forks directly via increased DnaA-DNA binding inhibiting replication fork progression, or indirectly by limiting factor(s) essential for fork progression, and thereby slowing down the forks. The overall validity and specifics of this proposal will have to be addressed in future studies.

Limitation of factors required for fork progression could also contribute to the seqA mutant’s inability to complete all ongoing replication rounds in the presence of rifampicin, a transcription inhibitor. The inability to complete all of the ongoing replication rounds should promote chromosomal fragmentation and make the seqA mutant susceptible to rifampicin, and we have indeed found that seqA mutant cells are killed by rifampicin at concentrations of the drug bacteriostatic for the wild-type cells.

Therefore, studying the underlying cause(s) of transcription inhibition-induced lethality in the seqA mutant could shed some light on replication factor(s) that maybe limiting in seqA.

162

4.5. TABLES

Table 4.1. Identities of the 37 SeqA-dependent mutants isolated using the color screen. Gene UV Function name Sensitive glnD No Catalyzes the uridylylation as well as the de-uridylylation isolated of the regulatory protein PII, which plays a critical role in thrice the regulation of nitrogen metabolism. acpS No Encodes holo-[ACP] synthase, which converts apo-ACP isolated to form holo-ACP, the active form of the carrier in lipid twice synthesis; essential for viability.

rfaQ No RfaQ is a transferase responsible for addition of the side- isolated branch heptose of the inner core of LPS. twice

ubiG No Codes for O-methyltransferase that catalyzes both O- isolated methylation reactions in the biosynthesis of ubiquinone. twice atpD, atpH Yes Code for subunits of the F-1 complex of ATP synthase & atpF complex; complex consists of five subunits, each of which is required for activity. These mutants have a 3- fold decreased level of ATP. Probably affects recombinational repair machinery. recB Yes A helicase and a component of the RecBCD complex; isolated six essential for recombination and dsDNA repair. times

recC Yes A component of the RecBCD complex; essential for isolated recombination and dsDNA repair. four times ruvA Yes A component of RuvABC resolvasome; involved in isolated resolving Holliday junctions. five times recG Yes RecG is a DNA helicase involved in double-strand break repair and dissociating R-loops. iscS No Cysteine desulfurase (IscS) catalyzes the transfer of isolated sulfur from cysteine; critical for synthesis and repair of twice iron-sulfur (Fe-S) clusters. eno No Codes for enolase that catalyzes the interconversion of 2- phosphoglycerate and phosphoenolpyruvate. ypaB No No function known. However ypaB is located just upstream of the nrdAB operon and the direction of insertion suggests downregulation of the operon. 163

Table 4.1. (cont.) cysB No Controls the transcription of the operon involved in novobiocin resistance and transcription of genes involved in sulfur utilization and sulfonate-sulfur catabolism via cysteine biosynthesis. aspS No Codes for Aspartyl-tRNA synthetase (AspRS). rseA No RseA is an anti-sigma factor that inhibits sigma E. sigma E regulates responses to heat shock and other stresses on membrane and periplasmic proteins. serA No Codes for D-3-phosphoglycerate dehydrogenase, which catalyzes the first step in the biosynthesis of L- serine. rpoN No Codes for the sigma factor controlling nitrogen- regulated and nitrogen-related promoters. ychA No No function known.

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4.6. FIGURES

Figure 4.1. Model for replication in the seqA mutants in linear DNA degradation deficient conditions due to the absence of sister chromatid cohesion. Disruption of this suspected sister- chromatid cohesion leads to problems with proper homologous recombination repair in the newly-synthesized daughter DNA resulting in improper re-attachments of the broken double strand ends to the wrong partners (e.g. the end could attach itself to the homologous sequence in one of the two "cousin" duplexes instead of its own sister duplex).

165

A 28°C

AB1157 seqA recD seqA recD

28°C on LB 42°C on LB+300mM MgSO4

AB1157 AB1157 seqA seqA recA recA seqA recA seqA recA

C 28°C 42°C

AB1157 AB1157

seqA seqA

seqA recA(Cs) seqA recA(Cs)

seqA recA(Cs ) seqA recA(Cs ) recD recD

Figure 4.2. A. to C. AB1157, ΔseqA, ΔrecD, ΔseqA ΔrecD, ΔrecA, ΔseqA ΔrecA, ΔseqA recA(Cs) and ΔseqA recA(Cs) ΔrecD, grown on LB or LB supplemented with 300 mM MgSO4 and incubated at the indicated temperatures. Overnight cultures of these strains were normalized to the same OD, serially diluted and spotted by 10 µl and incubated at 28 °C or 42 °C. 166

42°C 28°C

AB1157 seqA recBC sbcA seqA recBC sbcA (pER16) recBC sbcBC seqA recBC sbcBC (pER16)

Figure 4.3. AB1157, ΔseqA, ΔrecBC sbcA, ΔseqA ΔrecBC sbcA(pER16), ΔrecBC sbcBC and ΔseqA ΔrecBC sbcBC(pER16) grown at the indicated temperatures. Overnight cultures of these strains were normalized to the same OD, serially diluted and spotted by 10 µl on LB plates and incubated at 28 °C or 42 °C. The plasmid pER16 which has a temperature sensitive origin carries the seqA gene on it for complementation.

167

A 28 °C 32 °C 37 °C

AB1157 seqA dnaB (Ts) seqA dnaB (T s)

B 28 °C 37 °C 42 °C AB1157 seqA dnaC (Ts) seqA dnaC (Ts)

C 28 °C 42°C on LB+300mM

AB1157 MgSO4 seqA rep seqA rep

Figure 4.4. A. and B. AB1157, ΔseqA, dnaB(Ts), ΔseqA dnaB(Ts), dnaC(Ts) and ΔseqA dnaC(Ts) under permissive, semi-permissive and non-permissive temperatures of growth for dnaB (Ts) and dnaC(Ts) mutants, respectively. Overnight cultures of these strains were normalized to the same OD, serially diluted and spotted by 10 µl on LB plates. C. AB1157, ΔseqA, rep and ΔseqA rep overnight cultures were normalized to the same OD, serially diluted and spotted by 10 µl on an LB plate at 28 °C and on an LB plate supplemented with 300 mM MgSO4 at 42 °C. 168

Figure 4.5. ori/ter ratios in exponentially growing cultures of AB1157, ΔseqA, rep and ΔseqA rep. Total DNA extracted from exponentially growing cultures of these strains at 28 °C was deposited on a hybridization membrane and hybridized to the origin and terminus- specific probes. The resulting signals were then normalized to the signal from the wild- type overnight culture (taken as "1"). The values are averages of 4-6 independent determinations ± SEM.

169

Figure 4.6. A. to D. Rates of replication in exponentially growing cultures of AB1157, ΔseqA, dnaC(Ts) and ΔseqA dnaC(Ts) at indicated temperatures. The rates of replication were measured by incorporation of 3H-thymidine into acid-insoluble material.

170

Figure 4.7. A. and B. Rates of replication in exponentially growing cultures of AB1157, ΔseqA and rep upon treatment with indicated concentrations of Hydroxyurea (HU). The rates of replication were measured by incorporation of 3H-thymidine into acid-insoluble material. For (B), the values are averages of 3 independent determinations ± SEM.

171

Figure 4.8. A. to O. depict the flow cytometry profiles of exponentially growing AB1157, rep and ΔseqA cultures upon 150 µg/ml rifampicin addition.

172

Figure 4.9. A. and B. Rates of replication in exponentially growing cultures of AB1157, ΔseqA and rep upon treatment with 150 µg/ml rifampicin or 100 µg/ml chloramphenicol. The rates of replication were measured by incorporation of 3H-thymidine into acid-insoluble material. The values are averages of 3-6 independent determinations ± SEM.

173

Rif treated AB1157

75 µg/ml

50 µg/ml

20 µg/ml

10 µg/ml Rif treated seqA 75 µg/ml

Untreated 50 µg/ml

AB1157 20 µg/ml

seqA 10 µg/ml

Figure 4.10. AB1157, ΔseqA treated with different concentrations of rifampicin (Rif) for 24 hours. Overnight cultures were diluted 100 fold and grown to an OD~.2 in LB at 28 °C. These cultures were treated with 10, 20, 50 and 75 µg/ml of Rifampicin for 24 hours. After 24 hours, the cultures were serially diluted and spotted by 10 µl on LB plates at 28 °C.

174

ori/ter ratios upon Treatment with Rifampicin

4 AB1157 3 seqA

Fold Change 2

0 0 20 40 60 80 100 120 140 time(mins) Figure 4.11. Kinetics of absolute ori and ter signals and ori/ter ratios in AB1157 and ΔseqA upon treatment with 150 µg/ml Rifampicin. The resulting signals were normalized to the signal from the wild-type overnight culture (taken for "1"). The values are averages of 3-4 independent determinations ± SEM.

175

1182

887

591

Count

296

0 0 65536 131072 196608 262144 FITC-A

Figure 4.12. Flow cytometry profile of ΔseqA cultures grown overnight at 42°C in LB containing 300 mM MgSO4 treated with 150 µg/ml of rifampicin for 4 hours. Discreet peaks suggest completion of replication.

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4.7. REFERENCES

1. Atlung T, and Hansen FG. Three distinct chromosome replication states are induced by increasing concentrations of DnaA protein in Escherichia coli. Journal of bacteriology 175: 6537- 6545, 1993. 2. Brendler T, and Austin S. Binding of SeqA protein to DNA requires interaction between two or more complexes bound to separate hemimethylated GATC sequences. The EMBO journal 18: 2304-2310, 1999. 3. Brendler T, Sawitzke J, Sergueev K, and Austin S. A case for sliding SeqA tracts at anchored replication forks during Escherichia coli chromosome replication and segregation. The EMBO journal 19: 6249-6258, 2000. 4. Campbell JL, and Kleckner N. E. coli oriC and the dnaA gene promoter are sequestered from dam methyltransferase following the passage of the chromosomal replication fork. Cell 62: 967-979, 1990. 5. Datsenko KA, and Wanner BL. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc Natl Acad Sci USA 97: 6640-6645, 2000. 6. Grompone G, Ehrlich D, and Michel B. Cells defective for replication restart undergo replication fork reversal. EMBO reports 5: 607-612, 2004. 7. Kouzminova EA, Rotman E, Macomber L, Zhang J, and Kuzminov A. RecA- dependent mutants in Escherichia coli reveal strategies to avoid chromosomal fragmentation. Proceedings of the National Academy of Sciences of the United States of America 101: 16262- 16267, 2004. 8. Kuzminov A. Recombinational repair of DNA damage in Escherichia coli and bacteriophage lambda. Microbiology and molecular biology reviews : MMBR 63: 751-813, table of contents, 1999. 9. Lane HE, and Denhardt DT. The rep mutation. IV. Slower movement of replication forks in Escherichia coli rep strains. Journal of molecular biology 97: 99-112, 1975. 10. Lu M, Campbell JL, Boye E, and Kleckner N. SeqA: a negative modulator of replication initiation in E. coli. Cell 77: 413-426, 1994. 11. Miller JH. Experiments in Molecular Genetics. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory, 1972, p. 466. 12. Riber L, and Lobner-Olesen A. Coordinated replication and sequestration of oriC and dnaA are required for maintaining controlled once-per-cell-cycle initiation in Escherichia coli. Journal of bacteriology 187: 5605-5613, 2005. 13. Rotman E, Bratcher P, and Kuzminov A. Reduced lipopolysaccharide phosphorylation in Escherichia coli lowers the elevated ori/ter ratio in seqA mutants. Molecular microbiology 72: 1273-1292, 2009. 14. Rotman E, Khan SR, Kouzminova E, and Kuzminov A. Replication fork inhibition in seqA mutants of Escherichia coli triggers replication fork breakage. Molecular microbiology 93: 50-64, 2014.

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15. Sandler SJ. Multiple genetic pathways for restarting DNA replication forks in Escherichia coli K-12. Genetics 155: 487-497, 2000. 16. Simmons LA, Breier AM, Cozzarelli NR, and Kaguni JM. Hyperinitiation of DNA replication in Escherichia coli leads to replication fork collapse and inviability. Molecular microbiology 51: 349-358, 2004. 17. Sutera VA, Jr., and Lovett ST. The role of replication initiation control in promoting survival of replication fork damage. Molecular microbiology 60: 229-239, 2006. 18. Waldminghaus T, and Skarstad K. The Escherichia coli SeqA protein. Plasmid 61: 141-150, 2009. 19. Weitao T, Nordstrom K, and Dasgupta S. Escherichia coli cell cycle control genes affect chromosome superhelicity. EMBO reports 1: 494-499, 2000.

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CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS

In this work, we have found that CN potentiates H2O2 toxicity by recruiting iron from iron depots like ferritins (FtnA), possibly aided by the flavin reductase enzyme

(Fre), and fuelling the DNA-self-targeting Fenton's reaction to cause catastrophic chromosomal fragmentation. On the other hand, the iron depository, Dps, protects against

CN + H2O2 induced killing and catastrophic chromosomal fragmentation by sequestering the CN-recruited iron (6). The catastrophic chromosomal fragmentation induced by CN +

H2O2 was found to be cell density-dependent, but replication- and translation-independent and uniformly distributed over the chromosome. Our genetic analysis suggests that only the combination of CN + H2O2, but not H2O2 alone, damages bases, which are processed via abasic sites, repaired by the base excision repair pathway.

In addition, termination of the CN + H2O2 treatment, which allows the cells to synthesize ATP again, was found to trigger significant linear DNA degradation and release of linear DNA fragments from the nucleoid. Our investigation into the consequences of the CN + H2O2-induced catastrophic chromosomal fragmentation on the nucleoid provides insights into the nucleoid structure and its dynamics. The release of fragmented DNA from the nucleoid was enhanced in all mutants lacking nucleoid associated proteins except hns, which instead exhibited diminished nucleoid disintegration compared to the wild-type. Moreover, inhibition of transcription also reduced the release of fragmented DNA. Therefore, most nucleoid associated proteins prevent the disassembly of nucleoid structure via their binding, while H-NS and processes like transcription promote its unravelling. 179

However, several questions remain unanswered. To complement our genetic evidence, the role of flavin mononucleotide reductase (Fre) in the CN-promoted recruitment of iron from iron depots like FtnA can be analyzed directly in vitro by measuring plasmid relaxation by CN + H2O2 in the presence of the Fre enzyme and ferritins. Moreover, since our data indicates that FtnA is one of the major sources from which CN recruits iron, other potential sources of iron can be investigated using ftnA mutants additionally lacking, for example, Fe-S cluster containing enzymes, such as dehydratases. At the same time, future work will also focus on elucidating the precise mechanism via which Dps protects the cell from CN + H2O2. The iron depot, Dps, in addition to sequestering iron in its core is also present in abundance in the stationary phase complexed with the chromosomal DNA (5). Using dps mutants that lack the N- terminal DNA binding activity but maintain their ability to store iron would allow us to distinguish the extent to which these two capabilities aid in counteracting the CN + H2O2 co-toxicity (3). The role of iron depots in other potentiated H2O2 toxicities such as NO +

H2O2 and excess cysteine + H2O2 will also be studied using our panel of various mutants as well as the in vitro assays that we have developed.

The roles of electron transport chain proteins in CN + H2O2 co-toxicity appear multi-faceted and complex. Our work indicates that NADH dehydrogenase II (Ndh) counteracts this co-toxicity, while other enzymes, like AppC, CydB and CyoB cytochrome oxidases act to promote this synergistic toxicity. It would be interesting to explore the mechanism by which these electron transport enzymes affect CN + H2O2 co- toxicity. Measuring the NADH levels in mutants lacking these electron transport chain proteins will allow to directly address what effect (if any) elevation of intracellular

180

NADH levels due to CN-induced respiration block has on the CN + H2O2 co-toxicity.

Additionally, since changing the treatment temperature and medium would affect the metabolic state of the cell, this would allow us to observe if these enzymes play similar roles under different treatment condition, which could possibly shed some light on their mechanisms of action.

Furthermore, we would also like to address how CN + H2O2 co-toxicity affects other microorganisms such as Deinococcus radiodurans, which can survive up to 15,000

Gy γ-rays irradiation, lethal to most living organisms, and demonstrates similar resistance to UV irradiation and oxidative DNA damage (1). In Deinococcus radiodurans, the CN +

H2O2-induced DNA damage should be completely repairable upon the removal of CN by their robust recombinational repair system, resulting in complete revival after the treatment. However, unlike E. coli, D. radiodurans has substantially higher intracellular manganese concentrations and this higher Mn/Fe ratio has been reported to contribute to its resistance to the DNA damaging treatments mentioned above (4). Therefore, using D. radiodurans as our target organism we would be able to specifically investigate how important the level of intracellular iron concentrations are to promote the CN + H2O2 co- toxicity and what role (if any) intracellular manganese plays in it. Yet another interesting organism for the study of CN + H2O2-induced chromosomal fragmentation is the budding yeast, Saccharomyces cerevisiae, which is one of those rare organisms that completely lack ferritins and instead use vacuoles to store transition metals (2).

181

5.1. REFERENCES

1. Battista JR, Earl AM, and Park MJ. Why is Deinococcus radiodurans so resistant to ionizing radiation? Trends Microbiol 7: 362-365, 1999.

2. Bleackley MR, and Macgillivray RT. Transition metal homeostasis: from yeast to human disease. Biometals : an international journal on the role of metal ions in biology, biochemistry, and medicine 24: 785-809, 2011.

3. Ceci P, Cellai S, Falvo E, Rivetti C, Rossi GL, and Chiancone E. DNA condensation and self-aggregation of Escherichia coli Dps are coupled phenomena related to the properties of the N-terminus. Nucleic Acids Res 32: 5935-5944, 2004.

4. Daly MJ, Gaidamakova EK, Matrosova VY, Vasilenko A, Zhai M, Venkateswaran A, Hess M, Omelchenko MV, Kostandarithes HM, Makarova KS, Wackett LP, Fredrickson JK, and Ghosal D. Accumulation of Mn(II) in, Deinococcus radiodurans facilitates gamma- radiation resistance. Science 306: 1025-1028, 2004.

5. Haikarainen T, and Papageorgiou AC. Dps-like proteins: structural and functional insights into a versatile protein family. Cell Mol Life Sci 67: 341-351, 2010.

6. Mahaseth T, and Kuzminov A. Cyanide enhances hydrogen peroxide toxicity by recruiting endogenous iron to trigger catastrophic chromosomal fragmentation. Molecular microbiology 2015.

182