TESTING OXIDATIVE-STRESS HYPOTHESES GENERATED BY SYSTEMS-BIOLOGY ASSAYS

BY

YUANYUAN LIU

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, 2013

Urbana, Illinois

Doctoral Committee:

Professor James A. Imlay, Chair

Associate Professor Andrei Kuzminov

Professor John E. Cronan

Professor Peter A. Orlean

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ABSTRACT

Role of yaaA in hydrogen peroxide stress response in Escherichia coli

Hydrogen peroxide (H2O2) is commonly formed in microbial habitats by either

chemical oxidation processes or host defense responses. H2O2 can penetrate

cell membranes and damage key intracellular biomolecules, including DNA and

certain groups of iron-dependent enzymes. Bacteria defend themselves against

this H2O2 stress by inducing a set of genes that engages multiple defensive

strategies. In Escherichia coli, this defense is regulated by a transcriptional factor,

OxyR. A previous microarray study suggested that yaaA, an uncharacterized

gene found in many bacteria, was induced by H2O2 in Escherichia coli as part of

its OxyR regulon. Here I confirm that yaaA is a key element of H2O2 stress

response. In a catalase/peroxidase-deficient (Hpx-) background, yaaA deletion

mutants grew poorly, filamented extensively and lost substantial viability when

they were cultured in aerobic LB medium. The results from a thyA forward

mutagenesis assay and the growth defect of the yaaA deletion in DNA repair-

deficient background indicated that yaaA mutants accumulated high levels of

DNA damage. The growth defect of yaaA mutants was suppressed by either the

addition of iron chelators or by mutations that slow iron import, suggesting that

the DNA damage was caused by the Fenton reaction. Spin-trapping experiments

confirmed that Hpx- yaaA cells had a higher hydroxyl radical level than yaaA+

strains. EPR analysis showed that the proximate cause was an unusually high

level of intracellular unincorporated iron. These results demonstrate that during

 

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periods of H2O2 stress the induction of YaaA is a critical device to reduce

intracellular iron levels; it thereby attenuates the Fenton reaction and the DNA

damage that would otherwise result. The molecular mechanism of how YaaA

does it remains unknown.

Do antibiotics cause oxidative stress?

Since 2007, several studies have reported that classic bactericidal antibiotics

triggered the oxidation of an intracellular fluorescein probe, and this effect was

attributed to hydroxyl radicals. Therefore, investigators hypothesized that

different classes of antibiotics share a common bactericidal mechanism:

antibiotics stimulate the formation of superoxide and H2O2, which damage certain

iron-dependent enzymes, releasing iron to the cytosol. As a consequence, the

higher levels of H2O2 and free iron drive the Fenton reaction to generate hydroxyl

radicals which cause lethal DNA damage. If this model is true, then it might guide

new antibiotic therapies. Therefore, I evaluated its possibility by testing specific

steps of this hypothesis in Escherichia coli. I used three different antibiotics--

kanamycin, ampicillin and norfloxacin--all of which have been shown to increase

fluorescein-probe oxidation. However, contrary to the hypothesis, these

antibiotics neither induced the peroxide-responsive OxyR regulon, nor

accelerated endogenous H2O2 production. Furthermore, the metallo-enzymes

that are known to be sensitive to superoxide and H2O2 were not damaged, and

the level of intracellular free iron did not increase. Finally, the lethal effect of

antibiotic persisted in the absence of oxygen, whereas DNA repair mutants were

not hypersensitive, challenging the idea that toxicity arose from oxidative DNA

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lesions. Taken together, I conclude that bactericidal antibiotics did not generate

reactive oxygen species to kill bacteria.

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To my family

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ACKNOWLEDGEMENTS

I would like to thank all my colleagues, friends, and family members for their help

and support in my graduate school years. I specially want to thank my advisor Dr.

James A. Imlay for his excellent guidance. He is a very good scientist as well as

a very good teacher. The work presented here would not have been possible

without his advice and help.

I would like to thank my thesis committee Dr. John E. Cronan, Dr. Andrei

Kuzminov, and Dr. Peter A. Orlean for providing me truly helpful insight into my

research directions.

I would like to thank Dr. Ian Gut for his kindly help in differential interference

contrast microscopy, and Dr. Mark Nilges for his help in electron paramagnetic

resonance spectroscopy studies. I would like to thank Sarah C. Bauer, who

discovered the phenotype of yaaA and started my first project.

I would like to thank all the people in Imlay lab: Dr. Adil Anjem, Dr. Mianzhi Gu,

Dr. Soojin Jang, Dr. Sergei Korshunov, Dr. Jia Liu, Dr. Zheng Lu, Dr. Lee

Macomber, Dr. Stefano Mancini, Dr. Surabhi Mishra, Dr. Fangfang Xu, Dr. Yuan

Sui, Kari Imlay, Maryam Khademian, Julia Martin, and Jason Sobota. They are

all smart, friendly, and wonderful people, who make my laboratory work an

enjoyable experience every day.

Finally, I would like to thank my family and friends for their love and support. I

appreciate Dylan Dodd’s for his help and support during my graduate school. His

companionship kept me positive and made my life pleasurable all these years.

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TABLE OF CONTENTS

CHAPTER ONE: INTRODUCTION ...................................................................... 1 

1.1 Oxidative Stress ............................................................................................. 1 

1.2 Damage Caused by Reactive Oxygen Species .............................................. 2 1.2.1 Damage of solvent exposed [4Fe-4S]2+ cluster containing enzymes ............................................................................................................ 3 1.2.2 Damage of mononuclear iron containing enzymes ................................... 4 1.2.3 Damage of DNA ....................................................................................... 5 

1.3 Sources of Reactive Oxygen Species and Defense Systems in E. coli .......... 6 1.3.1 Endogenous ROS and basal defense systems ........................................ 6 1.3.2 Exogenous H2O2 and inducible OxyR response ....................................... 7 1.3.3 Redox-cycling drugs and SoxRS response ............................................ 10 

1.4 Methods to Detect ROS Formation ............................................................... 13 1.4.1 Induction of stress-responsive regulon ................................................... 13 1.4.2 Detection of biomarkers of oxidative damage......................................... 14 1.4.3 Trapping of O2

- and hydroxyl radicals ..................................................... 16 1.4.4 Protection mediated by ROS quenching ................................................ 17 1.4.5 Chemiluminescent /fluorescent probes for ROS detection ..................... 18 

1.5 Bactericidal Antibiotics and Oxidative Stress ................................................ 20 1.5.1 -lactams ................................................................................................ 21 1.5.2 Aminoglycosides .................................................................................... 22 1.5.3 Fluoroquinolones .................................................................................... 23 1.5.4 Hypothesis of antibiotic-mediated oxidative stress ................................. 25 

1.6 Tables ........................................................................................................... 29 

1.7 Figures.......................................................................................................... 31 

1.8 References ................................................................................................... 33 

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CHAPTER TWO: THE YAAA PROTEIN OF THE ESCHERICHIA COLI OXYR REGULON LESSENS HYDROGEN PEROXIDE TOXICITY BY DECREASING INTRACELLULAR UNINCORPORATED IRON CONCENTRATION ............................................................................................ 43 

2.1 Introduction ................................................................................................... 43 

2.2 Materials and Methods ................................................................................. 43 2.2.1 Reagents ................................................................................................ 43 2.2.2 Strain construction .................................................................................. 44 2.2.3 Plasmid construction .............................................................................. 44 2.2.4 Bacterial growth ...................................................................................... 45 2.2.5 Cell viability ............................................................................................ 46 2.2.6 Differential interference contrast (DIC) microscopy ................................ 46 2.2.7 thyA forward mutagenesis assay ............................................................ 47 2.2.8 H2O2 killing assay ................................................................................... 47 2.2.9 EPR spin trapping of hydroxyl radicals ................................................... 48 2.2.10 EPR measurement of intracellular unincorporated iron ........................ 48 2.2.11 -Galactosidase activity ....................................................................... 49 2.2.12 H2O2 measurement .............................................................................. 49 2.2.13 Visualization of protein carbonylation ................................................... 50 2.2.14 Rapid amplification of cDNA ends (5-RACE) ....................................... 50 2.2.15 Real-time PCR ..................................................................................... 51 

2.3 Results.......................................................................................................... 52 2.3.1 The yaaA gene is induced by OxyR ....................................................... 52 2.3.2 Hpx- yaaA cells show a growth defect in LB during H2O2 stress ............ 53 2.3.3 Hpx- yaaA cells lose viability during H2O2 stress .................................... 54 2.3.4 yaaA mutants show higher levels of DNA damage and polypeptide damage during H2O2 stress ............................................................................. 55 2.3.5 The Fenton reaction causes injuries in yaaA mutants during H2O2 stress ............................................................................................................... 57 2.3.6 YaaA protects cells by attenuating the Fenton reaction ......................... 58 2.3.7 YaaA does not affect H2O2 levels ........................................................... 59 2.3.8 YaaA does not inhibit ferric iron reduction .............................................. 60 2.3.9 Hpx- yaaA mutants have higher unincorporated iron levels than the Hpx- parent strain ............................................................................................ 61 2.3.10 YaaA does not repress expression of iron importers ............................ 62 2.3.11 Hpx- mntH yaaA mutants show a strong growth defect in defined medium ........................................................................................................... 63 2.3.12 Regulation of YaaA .............................................................................. 64 

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2.4 Discussion .................................................................................................... 65 

2.5 Acknowledgments ........................................................................................ 68 

2.6 Tables ........................................................................................................... 69 

2.7 Figures.......................................................................................................... 72 

2.8 References ................................................................................................... 90 

CHAPTER THREE: CELL DEATH FROM ANTIBIOTICS WITHOUT THE INVOLVEMENT OF REACTIVE OXYGEN SPECIES ........................................ 94 

3.1 Introduction ................................................................................................... 94 

3.2 Materials and Methods ................................................................................. 95 3.2.1 Reagents ................................................................................................ 95 3.2.2 Strain construction .................................................................................. 96 3.2.3 Bacterial growth ...................................................................................... 97 3.2.4 Cell viability ............................................................................................ 98 3.2.5 O2 consumption ...................................................................................... 98 3.2.6 Real-time PCR ....................................................................................... 99 3.2.7 -Galactosidase activity ....................................................................... 100 3.2.8 H2O2 production measurement ............................................................. 100 3.2.9 Electron paramagnetic resonance (EPR) measurement of intracellular unincorporated iron .................................................................... 100 3.2.10 Enzyme assays .................................................................................. 101 3.2.11 H2O2 killing assay ............................................................................... 103 3.2.12 Hydroxyphenyl fluorescein (HPF) assay............................................. 103 

3.3 Results........................................................................................................ 104 3.3.1 Antibiotic killing does not require oxygen ............................................. 104 3.3.2 H2O2-stressed mutants have similar sensitivity to ampicillin or kanamycin as wild type cells ......................................................................... 104 3.3.3 Antibiotic treatments did not accelerate the formation of hydrogen peroxide ........................................................................................................ 105 3.3.4 Antibiotic treatments did not increase the pool of unincorporated iron ................................................................................................................ 107 3.3.5 General DNA repair pathways had little effect upon antibiotic sensitivity ....................................................................................................... 108 

3.4 Discussion .................................................................................................. 109 

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3.5 Acknowledgments ...................................................................................... 111 

3.6 Tables ......................................................................................................... 112 

3.7 Figures........................................................................................................ 113 

3.8 References ................................................................................................. 128 

CHAPTER FOUR: CONCLUSIONS ................................................................. 131 

4.1 Conclusions from Current Work .................................................................. 131 4.1.1 YaaA plays a role in peroxide defense by controlling iron levels .......... 131 4.1.2 The lethal effect of classic bactericidal antibiotics does not involve oxidative stress ............................................................................................. 131 

4.2 Possible Future Work ................................................................................. 132 4.2.1 What causes the growth defect of Hpx- yaaA mntH mutants in glucose defined medium? ............................................................................. 132 4.2.2 What genes are induced in yaaA mutants? .......................................... 134 4.2.3 What is the three-dimensional structure of YaaA? ............................... 134 4.2.4 Does YaaA bind iron? .......................................................................... 135 4.2.5 Are there any functionally critical residues in YaaA? ............................ 135 4.2.6 What percentage of the HPF is oxidized in antibiotic treated cells? ..... 136 4.2.7 Can KatG oxidize the HPF dye? ........................................................... 136 

4.3 References ................................................................................................. 138 

APPENDIX A: DELETION OF OXYS DOES NOT GENERATE A PHENOTYPE IN CATALASE/PEROXIDASE-DEFICIENT MUTANTS ............. 140 

A.1 Material and Methods ................................................................................. 141 A.1.1 Strain construction ............................................................................... 141 A.1.2 Bacterial growth ................................................................................... 141 

A.2 Table .......................................................................................................... 143 

A.3 Figure ......................................................................................................... 144 

A.4 References ................................................................................................. 145  

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CHAPTER ONE: INTRODUCTION

1.1 Oxidative Stress

During Earth’s early existence, the atmosphere was full of reducing volcanic

gases and had no free oxygen, and the ocean contained abundant dissolved

ferrous iron and sulfur species. Microbial life emerged and subsequently evolved

photosynthetic mechanisms for splitting water to produce oxygen. Despite the

presence of photosynthetic organisms, the Earth’s atmosphere remained

anaerobic for nearly two billion years until sufficient oxygen was produced to

titrate the ferrous iron and sulfur species in the ancient oceans (9,53). It then took

about another billion years for the oxygen level to slowly build up in the

atmosphere and reach the present-day concentration. The anaerobic earth

gradually turned into an aerobic world (Figure 1.1). During this period, microbial

organisms evolved mechanisms to exploit the metabolic potential of oxygen.

However, they also had to develop strategies to prevent its toxicity.

Molecular oxygen (O2) is a triplet species with two spin-aligned, unpaired

electrons (47,83). Because of this orbital structure, O2 can only accept one

electron at a time. The unpaired electrons of O2 allow it to rapidly react with

transition metals and organic radicals (47). Therefore, once aerobic

environments appeared, ancient free-radical-based catalytic mechanisms were

either dispensed with or were sequestered in the interiors of enzymes, where

oxygen could not penetrate.

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Another threat comes from univalent electron transfer feature of oxygen.

Molecular oxygen can be partially reduced to superoxide (O2-), hydrogen

peroxide (H2O2) and hydroxyl radicals (HO) (Scheme 1.1) (47,48). These

oxygen derivatives, also called reactive oxygen species (ROS), are stronger

oxidants than oxygen itself, and therefore can react with certain biological

molecules of fundamental biochemical mechanisms and damage metabolic

pathways inherited from the ancient reducing world. As a result, living organisms

developed approaches to diminish ROS and thereby protect their vulnerable

targets.

Scheme 1.1 The redox states of oxygen with standard reduction potentials.

1.2 Damage Caused by Reactive Oxygen Species

Biochemical and physiological studies of O2- or H2O2 scavenging-deficient

mutants have identified some of the targets that O2- or H2O2 most readily reacts

with. In aerobic Escherichia coli cultures, both superoxide dismutase-deficient

mutants (SOD) and catalase/peroxidase-deficient mutants (Hpx-) demonstrate

growth defects, develop aromatic amino acid auxotrophy, and suffer high levels

of mutagenesis (13,93). These phenotypes are correlated with damage of iron-

dependent enzymes and damage of DNA (5,30,53,67,88,98).

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1.2.1 Damage of solvent exposed [4Fe-4S]2+ cluster containing enzymes

Being small molecules, O2- and H2O2 can penetrate into the active sites of certain

iron-dependent enzymes and cause enzyme inactivation. Aconitase-class

dehydratases are one of the well-known targets for O2- and H2O2 (30,33,53,67).

This group of enzymes contain solvent-exposed, catalytically active [4Fe-4S]2+

clusters, which use an undercoordinated iron atom to bind substrate and catalyze

the reaction. A list of these enzymes includes: aconitase B, fumarase A and

fumarase B in tricarboxylic acid cycle (TCA cycle); 6-phosphogluconate

dehydratases (Edd) in the Entner-Doudoroff pathway; isopropylmalate isomerase

(IPMI) in leucine biosynthesis; dihydroxy-acid dehydratases in branched chain

amino acid synthesis; and L-serine deaminase in L-serine degradation

(12,33,34,53). During oxidative stress, O2- takes one electron from the [4Fe-4S]2+

clusters, is thereby reduced to H2O2 and leaves the active site; the unstable [4Fe-

4S]3+ state cluster then releases the catalytic iron in the ferrous form, resulting in

an inactive enzyme having a [3Fe-4S]+ cluster (30). H2O2, on the other hand,

extracts two electrons from the native cluster, probably through a ferryl-like

intermediate, and is reduced to H2O; the oxidized cluster releases a ferric iron to

generate the [3Fe-4S]+ form (53). In both cases, the enzyme is inactivated, and

an iron atom is released to the cytosol. The rate constants of inactivation by O2-

is about 106 M-1 s-1 (30,33,34), and the inactivation rate of H2O2 is 104 M-1 s-1 (53).

Hence, these solvent-exposed [4Fe-4S]2+ clusters are very susceptible to

oxidative stress. When the [4Fe-4S]2+ clusters are buried inside the enzymes, like

succinate dehydrogenase and NADH dehydrogenase, they are not vulnerable to

oxidative damage (53).

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1.2.2 Damage of mononuclear iron containing enzymes

The mononuclear iron enzymes represent another group of targets for O2- and

H2O2. These enzymes catalyze diverse categories of reaction types, but they all

employ ferrous iron to stabilize an oxyanion intermediate during the reaction

(5,98). Several enzymes in this group already have been demonstrated to be

sensitive to both O2- and H2O2, including ribulose-5-phosphate 3-epimerase

(RPE) in the pentose phosphate pathway; peptide deformylase (PDF) that

removes the formyl group from the initiation methionine of nascent proteins;

threonine dehydrogenase (TDH) in threonine metabolism; cytosine deaminase

(CDA) in the pyrimidine salvage pathway; and 2-dehydro-3-

deoxyphosphoheptonate aldolase (DAHPC) in aromatic amino acid synthesis.

Over 100 enzymes in E. coli use transition metals (i.e. Mn2+, Zn2+, Co2+, Ni2+ and

Fe2+) as prosthetic groups, and a couple dozen of them are able to use ferrous

iron (57). However, the usage of ferrous iron is often under recognized due to

uses of aerobic purification process or assay conditions. Therefore, it is possible

that there are unrecognized enzymes that actually use ferrous iron in

physiological conditions, and might be vulnerable to oxidative stress (5). During

O2- stress, the solvent-exposed ferrous iron cofactor is oxidized to the ferric form

and then dissociates from the active site, while O2- is reduced to H2O2. The apo-

enzymes thus formed can be mis-metallated by other metals (i.e. Zn), and

eventually lose activity (Gu and Imlay, unpublished data). H2O2 oxidizes the

catalytic iron cofactor in a Fenton reaction (see below), producing a ferryl radical.

The ferryl radical can oxidize cysteine residues (if any) or other residues in the

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active site. Oxidation of cysteine to sulfenic acids is reversible, while oxidation of

other amino acid residues can cause irreversible damage (5,98). In both cases,

the catalytic iron cofactor is oxidized to the ferric form and dissociates from the

active site, leaving an inactive enzyme (5,98). Similar to aconitase-type

dehydratases, the rate constants of inactivation of mononuclear iron enzymes is

also high, about 105 to 106 M-1 s-1 by O2- (Gu and Imlay, unpublished data) and

103 M-1 s-1 by H2O2 (5,98).

1.2.3 Damage of DNA

Both H2O2 stress and O2- stress can cause DNA damage. H2O2 reacts with

ferrous iron in the Fenton reaction (Scheme 1.2), generating a ferryl radical

intermediate (FeIV=O)2+, which ultimately decomposes to ferric iron and hydroxyl

radical (49,88). O2- indirectly increases Fenton-mediated hydroxyl radical

production by releasing iron atoms from sensitive enzymes (aconitase-class

dehydratases or mononuclear iron enzymes) (58,59). In vivo ferric iron can be

directly reduced back to the ferrous form by cysteine and FADH2. This

establishes a redox cycle pathway that drives the hydroxyl radical production

(87,111). Hydroxyl radical is an extremely strong oxidant (+2.33V). It reacts with

all types of macromolecules at close to the diffusion-limited rate (4). When

ferrous iron binds to DNA, a Fenton-mediated hydroxyl radical can be generated

on the surface of DNA, subsequently oxidizing nucleic acid bases or ribose

moieties, causing mutation and even cell death (49,88).

Fe2+ + H2O2 → (FeIV=O)2+ Fe3+ + OH- +HO

Scheme 1.2 The Fenton reaction

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1.3 Sources of Reactive Oxygen Species and Defense Systems in E. coli

1.3.1 Endogenous ROS and basal defense systems

O2- and H2O2 are routinely generated as by-products of aerobic metabolism,

apparently when molecular oxygen steals electrons from certain reduced

biomolecules. As a triplet species, molecular oxygen can only accept one

electron at a time from strong univalent electron donors, including menaquinones

or reduced flavins of redox enzymes and generate O2- as the initial product

(65,74). If O2- obtain another electron from flavosemiquinone radicals before it

diffuses out of the active site, then H2O2 is formed (77,78). H2O2 can also be

produced when two O2- dismutate spontaneously or during reactions catalyzed

by superoxide dismutase. Moreover, H2O2 is a stoichiometric reaction product of

certain oxidases, such as L-aspartate oxidase (64,77) and copper-containing

amine oxidase (79). Fortunately, the flux of aspartate oxidase is restrained, and

only limited amounts of H2O2 are produced. In the case of copper-containing

amine oxidase, the periplasmic location of this enzyme lessens the likelihood of

H2O2 interacting with cellular biomolecules (15). As the result, the H2O2 generated

by these enzymes accounts for only a small proportion of total endogenous H2O2

production and thus is tolerated by the cell. Although the respiratory chain is the

major source of O2- in the periplasm, it is not the predominant source of

endogenous H2O2 (<20%) in E. coli (65). To date, no single enzyme has yet been

identified as the major ROS producer.

In wild-type log-phase growing E. coli, the rate of O2- formation inside the cells is

about 5 M/s in fully aerated medium, while the rate of H2O2 formation is about

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15 M/s (48,93). Nevertheless, the potential toxicity of these ROS is diminished

by the high titers of the scavenging enzymes--superoxide dismutases, catalases,

and peroxidases. The levels of scavenging enzymes are sufficient to drive the

steady-state concentration of O2- to 0.1 nM and that of H2O2 to ~20 nM (50,94).

These concentrations are low enough to maintain function of vulnerable enzymes,

however, residual oxidative DNA damage still occurs. The evidence comes from

mutants deficient in both recombinational repair and base-excision repair

systems (recA xthA or polA recB strains). While these mutants grow well in the

absence of oxygen, they are inviable aerobically (17,48,51,82). Thus the basal

defenses of the scavenging enzymes and DNA repair system appear calibrated

to avoid problems from endogenous ROS in E. coli.

1.3.2 Exogenous H2O2 and inducible OxyR response

H2O2 can also come from exogenous sources. H2O2 forms at the interface

between aerobic and anaerobic habitats when oxygen reacts with metals and

thiols. H2O2 also accumulates in sterile lab medium (1-20 M) when O2 gets

electrons from flavin by photochemical mechanisms, or from glucose in the

presence of metals (48). Furthermore, H2O2 is generated by natural antimicrobial

systems because of its toxicity. Indeed, plants, amoebae, and mammals all

elaborate NADPH oxidase to kill invading bacteria. This enzyme transfers

electrons from NADPH to molecular oxygen, generating a mixture of O2- and

H2O2 in an effort to kill pathogens (10,36,76). Similarly, lactic acid bacteria

produce abundant H2O2 to suppress competitors (38,95,99).

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Unlike O2-, which cannot cross membranes because it is an anion at

physiological pH, H2O2 penetrates membranes at a rate similar to that of water.

Therefore, when bacteria enter an environment that contains H2O2, they must

activate defensive responses—for example, the OxyR system in E. coli. OxyR is

a transcription factor that can sense as little as 0.1 M of intracellular H2O2 and

induce gene expression (7). An activated thiolate residue (Cys199) on the OxyR

protein is oxidized by H2O2 to a sulfenic acid, which then reacts with a second

thiol group (Cys208) nearby to form a stable disulfide bond. This locks OxyR in

an activated form (68,116). The activated OxyR protein directly stimulates

transcription of at least two dozen genes involved in defense (Table 1.1) (118).

These genes presumably represent a list of cellular strategies to minimize the

toxicity of H2O2. Since Fenton-generated hydroxyl radicals are the most deadly

factors during H2O2 stress, cells induce genes to prevent the Fenton reaction by

scavenging H2O2 and by diminishing the amount of intracellular iron. Catalase

(katG) and peroxidase (ahpCF) are induced to scavenge H2O2 (93). At the same

time, the Fur repressor is up-regulated to diminish iron import (105,117); whereas

Dps, a ferritin-like protein, is induced to sequester unincorporated iron

(39,45,88,115). Maintenance of H2O2-sensitive enzyme function is also an

important goal. A secondary Fe-S cluster assembly system encoded by the Suf

operon is induced by OxyR to replace the H2O2-sensitive housekeeping Isc

system (52,69,85). Suf induction is critical for both synthesis of de novo Fe-S

clusters and repair of damaged clusters during peroxide stress (52). The

induction of MntH, a manganese importer, enables manganese to replace

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catalytic iron in mononuclear metalloenzymes which otherwise would be

inactivated by H2O2 (6,56,98). The significance of these adaptations has been

shown in catalase/peroxidase-deficient mutants (katG katE ahpCF triple mutant,

named Hpx-). During aerobic growth, these strains accumulate approximately 1

M H2O2, which is ~10 times over the concentration sufficient to induce the OxyR

regulon (7,94). Mutants having gene deletions (mutation of suf, mntH, dps or fur)

in the Hpx- background grow poorly in aerobic medium and the phenotypes are

correlated with their cellular function in H2O2 defense (6,52,88).

Thioredoxin C (trxC), glutaredoxin A (grxA), and glutathione reductase (gorA) are

also part of the OxyR regulon (118). The glutathione/glutaredoxin or thioredoxin

systems might be induced to repair oxidized cysteine residues. Although H2O2

efficiently oxidizes sulfhydryls on OxyR and Ahp peroxidase (>107 M-1 s-1) (7,90),

it is a relatively poor oxidant of typical cysteine residues (<60 M-1 s-1) (18,31,113).

Therefore, the importance of the glutathione/glutaredoxin or thioredoxin systems

to remove cysteine oxidation during peroxide stress is not clear. Indeed, deletion

of these genes did not exhibit growth defects in Hpx- cells (Imlay lab, unpublished

data). Interestingly, in many bacteria, glutaredoxin or thioredoxin is not part of the

H2O2-response system. In Streptomyces, the expression of thioredoxin is

activated by disulfide bond formation on an anti-sigma factor protein (48). These

suggest that OxyR in E. coli might also act as a sensor for disulfide stress, and

glutathione/glutaredoxin or thioredoxin are induced to reduce disulfide bonds (48).

OxyR also induces a specificity adapter for the ClpA-ClpP protease complex,

ClpS (yljA) (118). ClpS directs ClpAP towards aggregated proteins and N-end

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rule substrates, facilitating their degradation (24,28). This might be important

during H2O2 stress because hydroxyl radicals can cause multiple types of protein

damage.

There are several members in the OxyR regulon with uncharacterized functions,

including YaaA, YaiA, and YbjM (118). Presumably, the function of these proteins

will fall into one of the defensive categories mentioned above. In principle, if

deletion of these genes in Hpx- background creates a phenotype, one can

characterize the gene function by establishing which type of H2O2-mediated

damage is worsened in the mutant.

1.3.3 Redox-cycling drugs and SoxRS response

Oxidative stress can also arise from redox-cycling drugs, such as phenazines,

viologens, and quinones. It is well known that both plants and microbes use

these small organic compounds to suppress the growth of their competitors. For

example, plumbagin and juglone are naturally synthesized quinones from plants

(42). They are effective herbicides that allow the producing plants to take over a

habitat. Phenazine is excreted by Pseudomonas spp. and a few other bacterial

genera, to allow the producing strains to outcompete with the resident microbes

(75). These redox-cycling compounds may have other functions (19). Some

studies suggest that they act as extracellular electron shuttles in biofilms,

delivering electrons to terminal oxidants that are not easily accessible (either

insoluble or diffusion-limited), such as iron mineral and molecular oxygen

(44,107,108). Since they can mediate the reduction of ferric iron to biologically

available ferrous iron, they may also contribute to iron acquisition (109).

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Most studies have focused on the toxicity of these compounds. Redox-cycling

drugs are well known for poisoning cells by generating reactive oxygen species,

but their toxicity can also come from ROS-independent mechanisms. After the

drugs penetrate into the cell, they can undergo addition reactions with available

thiols, and form covalent adducts on proteins (11,81). More importantly, the

univalent redox activity of these drugs allows them to abstract an electron from

the reduced flavin or the metal center of redox enzymes. This action not only

directly inactivates aconitase-type Fe-S clusters dehydratases and NADPH-

reduced enzymes (42), it can also inactivate NADH dehydrogenase II and block

the cellular metabolism (46). The reduced drug is then re-oxidized by an oxidized

quinone pool. When terminal electron acceptors-such as molecular oxygen,

nitrate or fumarate-are available, the drug can undergo redox-cycling multiple

times, transfer electrons to terminal acceptors to produce water, nitrite or

succinate, respectively (42). The reduced drug is also able to directly transfer

electrons to oxygen. As a result, when molecular oxygen is present, the redox-

cycling reactions not only generate water, but also large amounts of O2- and

H2O2, leading to oxidative damage (42,75).

In E. coli, the SoxRS regulon mediates the response to redox-cycling compounds

and nitric oxide. The transcriptional factor SoxR has an exposed [2Fe-2S]+

cluster, which is oxidized to [2Fe-2S]2+ by univalent oxidants, such as redox-

cycling compounds. The oxidized cluster triggers a large conformational change

in the protein, shifting it to a transcriptionally active form (22,35). Nitric oxide can

also activate SoxR by direct nitrosylation of the [2Fe-2S]+ cluster, generating a

12  

disrupted cluster with dinitrosyl-iron-dithiol complex (21). In E. coli, the activated

SoxR stimulates transcription of only one known gene, the transcriptional

activator SoxS. SoxS then activates about two dozen genes to protect the cell

(Table 1.2) (41,91,101).

Because redox-cycling drug can cause damage without the involvement of

reactive oxygen species, it is not surprising that the major members of the

SoxRS regulon are involved in controlling the intracellular concentration of redox-

cycling drug (42). OmpF antisense sRNA (micF) and lipopolysaccharide (LPS)

modification genes (waaY and waaZ) are induced to reduce drug entry by

changing the permeability of the cell envelope. Genes encoding drug efflux

pumps (acrAB and tolC), drug detoxification (nfsA, ygfZ and nfnB), and multiple

antibiotic resistance systems (marAB) are also up-regulated. Since redox-cycling

drugs can produce O2- in the presence of oxygen, SoxS also induces genes to

defend against potential O2- toxicity. Manganese-containing superoxide

dismutase (sodA) is up-regulated to scavenge O2-. The oxidant-resistant

dehydratase-isoforms, fumarase C (fumC) and aconitase A (acnA), and the [Fe-S]

cluster repair protein YggX are expressed to deal with the Fe-S cluster damage

(70,89,92,104). Fur repressor is induced to decrease iron import, and thereby

reduce its role in Fenton-mediated DNA damage; while endonuclease IV (nfo) is

induced to enhance the repair of oxidative DNA lesions. Furthermore, glucose-6-

phosphate dehydrogenase (zwf), flavodoxin NADP+ reductase (fpr), and

flavodoxins (fldA and fldB) are induced as well, likely to restore the NADPH pool

which may be exhausted by redox-cycling.

13  

1.4 Methods to Detect ROS Formation

Because of their toxicity, reactive oxygen species produced by host cells are

widely believed to be correlated with various disease states, including

neurodegeneration, diabetes, atherosclerosis, and cancer (2). It is also argued

that ROS play a significant role in the aging process, though convincing evidence

to support this notion is not available.

One essential problem in evaluating the relationship of ROS with diseases and

aging is how to detect and measure ROS accurately. These molecules exhibit

very short half-lives which makes detection difficult. In general, there are five

approaches to detect ROS, which include: (a) monitoring induction of stress-

responsive regulons; (b) detecting biomarkers of oxidative damage; (c) detecting

O2- or hydroxyl radical by electron paramagnetic resonance (EPR) spin trapping;

(d) establishing a protective role of ROS quenchers; (e) measuring ROS by

chemiluminescent/fluorescent probes.

1.4.1 Induction of stress-responsive regulon

Activation of inducible stress response systems is a good indicator of oxidative

stress, as these regulons play a specific role in sensing stress at physiologically

relevant concentrations. The OxyR regulon is triggered by ~0.1 M H2O2 (7), and

therefore the induction of OxyR regulated genes gives an excellent readout for

physiological H2O2 stress. Gene expression can be measured by transcriptional-

lacZ fusions, real-time reverse transcription PCR (qRT-PCR), microarrays or

RNA-seq. However, the level of activation may vary between members under the

14  

same conditions, and some members may have more than one regulator. For

example, dps is induced by OxyR as well as RpoS, while being repressed by

MntR (3,112). As a result, it is better to use experimental conditions in which

OxyR is the only activated regulator or to measure the expression of more than

one gene in the regulon.

In contrast to the OxyR regulon which responds to H2O2, the SoxRS system is

not directly triggered by O2-. Rather, SoxRS is induced by redox-cycling drugs,

which can occur even under anaerobic conditions (42). As a result, induction of

SoxRS regulated genes, such as superoxide dismutase (sodA), does not

necessarily correlate with O2- stress, and the data should be interpreted with

caution.

1.4.2 Detection of biomarkers of oxidative damage

ROS can react with biomolecules, generating unique and specific types of

damage. These scars can be used as evidence of oxidative damage. Activity of

ROS-sensitive enzymes as well as levels of oxidative protein damage and DNA

damage have all been utilized as biomarkers to indicate oxidative stress.

Damage of ROS-sensitive enzymes can be evaluated by enzyme assays. In

order to avoid variations between different samples, the actual enzyme activity is

normalized to the total enzyme activity (see below), and the fraction is compared

to that in the absence of oxidative stress. The damaged [3Fe-4S]+ clusters in

aconitase-class dehydratases can be repaired by incubation with ferrous iron and

dithiothreitol (DTT); if the Fe-S cluster is further degraded, it can be rebuilt by

15  

addition of the IscS Fe-S cluster assembly system and cysteine to the repair

reaction (52,53). In this scenario, the activity observed post repair or post

rebuilding is defined as the total enzyme activity. Similarly, the total activity of

mononuclear iron enzymes is determined after remetallating the apo-enzymes

with ferrous iron under anaerobic conditions (5,98). However, there are factors

that limit the use of sensitive enzymes as ROS fingerprints: (a) a lot of these

enzymes require specific growth conditions for protein expression; (b) the

damage of their cofactors might due to reasons other than oxidative stress (i.e.

iron depletion).

Polypeptides can also be oxidized by hydroxyl radicals and undergo different

types of oxidative modifications, such as carbonylation, a product from direct

oxidation of amino-acid side chains. Carbonyl derivatives often serve as markers

of oxidative stress because of their chemical stability. However, they can be also

generated by protein glycation inside the cell. Therefore, the levels of carbonyl

groups in cellular proteins are not restricted to oxidative damage (43).

During oxidative stress, the most disease-related damage to cells is hydroxyl

radical-mediated DNA damage, which can cause spontaneous mutation or even

cell death. Hydroxyl radicals oxidize the first encountered nucleic acid base or

ribose moiety; the initial radical products then attack nearby bases, sugars or

cellular reductants, and eventually produce numerous stable end products after

series of reactions (23). The relative amounts of these end products are

determined by reaction conditions, such as redox environment, association of

transition metals with DNA, etc (23). Hence, it is not appropriate to rely on a

16  

single product as an index of oxidative DNA damage. Such reliance is not very

sensitive, and can lead to false results. For example, 8-hydroxyguanine (8-OHdG)

is a common product of guanine in DNA oxidation. Because of its easy

measurement and strong mutagenicity, the level of 8-OHdG is often used as an

indicator of oxidative DNA damage. However, 8-OHdG is just one of the many

important products generated from oxidative DNA damage (23). Moreover, it can

also be formed during the process of DNA isolation and assay preparation, which

adds uncertainty and potential artifacts to the result (43). A better way to evaluate

DNA damage is to measure the mutation rate or cell viability, as they reveal the

overall effects of critical DNA damage that are most threatening and disease

related.

In general, there is no perfect biomarker for oxidative stress. The ones suggested

above can be used as a support of oxidative damage, but careful controls and

follow up experiments are required.

1.4.3 Trapping of O2- and hydroxyl radicals

Electron paramagnetic resonance (EPR) or electron spin resonance (ESR) is a

technique to detect species with unpaired electrons (paramagnetic species), like

flavin radicals, quinone radicals, O2- radicals and hydroxyl radials (29). In

biological systems, ROS exhibit even shorter lifetimes because of the various

antioxidant mechanisms. As a result, spin traps are employed to react with the

radicals produced in the cell. The reactions produce stable radical adducts, which

can be subsequently detected by EPR.

17  

5,5-Dimethyl-1-pyrroline-N-oxide (DMPO) and 2,5,5-trimethyl-1-pyrroline-N-oxide

(TMPO) are commonly used for O2- detection in EPR spin trapping. However, the

rate constants of these spin traps with O2- are very slow (2-70 M-1s-1), making it

difficult to capture O2- before it is detoxified by scavenging enzymes or reductants

inside the cell (29).

DMPO, hydroxyl-2,2,6,6-tetramethylpiperidine-N-oxyl (TEMPO), -phenyl-N-tert-

butylnitrone (PBN) and-(4-pyridyl N-oxide)-N-tert-butylnitrone (POBN) are

widely used spin traps for hydroxyl radicals. Though the hydroxyl radical reacts

with these compounds rapidly (>109 M-1s-1) (29,43), it also reacts with many

biomolecules at diffusion limited rates (4). Furthermore, the spin trap-radical

adducts can be quickly removed by enzymes or reductants inside the cell (43).

Therefore, though EPR spin trapping can be used for O2- and hydroxyl radical

detection in vivo, it is only a qualitative method.

1.4.4 Protection mediated by ROS quenching

Iron chelators prevent the Fenton reaction by sequestering iron. However, it is

more complicated when it comes to a cellular system. Addition of iron chelator

can cause iron depletion, which represses the TCA cycle and the respiratory

chain, thereby altering cellular metabolism. Therefore, the protective effect of iron

chelators may merely arise from changes in metabolism and not be related to the

Fenton chemistry. Similar reasons apply to hydroxyl radical quenchers, which

may also have pleiotropic effects other than hydroxyl radical scavenging inside

the cells. As a result, protection from ROS quenchers is not sufficient to confirm

18  

ROS production. In order to obtain meaningful results, it requires understanding

of the fundamental cellular physiology, careful data interpretation and more

important, followed up experiments.

1.4.5 Chemiluminescent /fluorescent probes for ROS detection

Chemiluminescent and fluorescent probes are very popular for reactive oxygen

species detection. Compared to other approaches, these dyes are sensitive,

convenient and inexpensive. Yet, caution must be taken when using these dyes,

because many of them are nonspecific and can give rise to artifacts. Listed below

are some examples of commonly used dyes.

Luminol and lucigenin (Figure 1.2A-B) are often employed for O2- detection.

However, neither compound directly reacts with O2-. Luminol must first undergo

one electron oxidation by peroxynitrite, hydroxyl radicals, or high-valent iron to

generate luminol radical. The resulting luminol radical then react with O2- to

produce a light emitting molecule. In the case of lucigenin, the dye must first be

reduced to lucigenin radical cation by the cellular reducing system (i.e. electron

transport chain), before it reacts with O2-. Unfortunately, both luminol radicals and

lucigenin radical cations are also able to react with oxygen to generate O2-,

leading to artificial elevated O2- production. Therefore, luminol and lucigenin are

not reliable probes for O2- detection (43). Actually, lucigenin is now suggested for

Cl- detection by Invitrogen.

Amplex Red is frequently used for H2O2 measurements. In the presence of

horseradish peroxidase, this dye is oxidized by H2O2 to produce a highly

fluorescent product, resorufin (Figure 1.2C). The assay is highly sensitive, and

19  

can detect as little as 50 nM H2O2. However, this dye is not suitable for H2O2

detection inside the cell. In the presence of NADH and oxygen, the fluorescent

product resorufin can undergo redox cycling upon visible light irradiation,

generating O2- and H2O2. This gives self-amplification of fluorescence signal.

Therefore, Amplex Red is only a reliable method to measure extracellular H2O2

formation, such as to detect H2O2 released from the cell or in isolated

mitochondrial preparations (54,114).

Dichlorofluorescein diacetate (DCFH2-DA) is another commonly used probe for

H2O2 and oxidative stress detection (Figure 1.2D). This cell-permeable probe is

deacetylated by cellular esterases to dichlorofluorescin (DCFH2). The

nonfluorescent product DCFH2 does not react with O2- or H2O2 directly; instead, it

is oxidized by one electron oxidants to form an intermediate radical (DCF-).

These one electron oxidants include peroxyl, alkoxyl, peroxynitrite, carbonate

and hydroxyl radicals, as well as high valence iron center in redox enzymes (i.e.

peroxidase with H2O2), cytochrome c, chromium, pyocyanin, mitoxantrone, and

ametantrone. The intermediate radical DCF- rapidly reacts with O2 to form a

fluorescent product, as well as O2- and H2O2. This reaction establishes a redox

cycling pathway, which artificially amplifies the fluorescent signal. The DCF-

intermediate radical can also react with other cellular reductants (i.e. glutathione,

ascorbate, and NADH) to generate more free radicals, further complicating

interpretation of assay results. Therefore, DCFH2-DA is not a reliable probe for

intracellular ROS (20,43,55).

20  

In summary, none of the ROS detection approaches is perfect and can serve as

a ‘gold standard’. Rather, each has specific advantages and disadvantages that

must be carefully considered. Measure the induction of stress response system

(a) and the biomarkers of oxidative damage (b), as well as EPR spin trapping for

O2- and hydroxyl radical formation (c) all can be used to detect ROS formation

inside the cell, though none of these is quantitative; whereas ROS quenchers

and ROS probes are only reliable in cell free systems. ROS detection requires a

complete understanding of the methods employed, careful design of controls,

cautious interpretation of the data, and follow-up experiments to confirm the

results.

1.5 Bactericidal Antibiotics and Oxidative Stress

The discovery and clinical application of antimicrobial reagents has provided the

greatest benefit to human health in the twentieth century. Most clinically used

antibiotics target one of three essential cellular processes: cell-wall assembly (i.e.

-lactams), protein synthesis (i.e. aminoglycosides), or nucleic acid metabolism

and repair (i.e. fluoroquinolones). However, recent reports have raised the

possibility that although these antibiotics block growth by directly inhibiting the

targets mentioned above, they may owe their lethal effects to the indirect creation

of reactive oxygen species which then damage bacterial DNA. In order to

evaluate the possibility of the oxidative stress hypothesis, it is important to

understand the mode of action of these antibiotics.

21  

1.5.1 -lactams

In 1929, Scottish microbiologist Alexander Fleming discovered penicillin, the first

antibiotic in history. It is a -lactam antibiotic that specifically targets the bacterial

wall assembly system (114).

The rigid cell wall of bacteria is critical for maintaining cell shape and integrity, as

well as preventing cell lysis from the high internal osmotic pressure. All bacterial

cell walls have a common structural component, peptidoglycan (murein). It is a

polymer consisting of disaccharide subunits, substituted with tripeptide chains

with crosslinks, thereby establishing a rigid mesh-like network on the outside of

the plasma membrane. During bacterial growth and division, the cell wall is

constantly broken down, new peptidoglycan units inserted and then crosslinked

(63,86). -Lactam antibiotics target this cross-linking step. All -lactam drugs

contain a -lactam ring that has a reactive CO-N bond lying in the same position

as the one in the D-alanyl-D-alanine portion of the peptidoglycan unit. This D-

alanyl-D-alanine moiety is the substrate of penicillin-binding proteins (PBPs),

which include a host of related enzymes (transpeptidases, carboxypeptidases,

transglycosylase, and endopeptidases) involved in peptidoglycan cross-link

formation (63). As a result, -lactams act as substrate analogs of PBPs, inhibiting

their function by forming a covalent bond with the catalytic serine residue of the

enzyme active site. The action of -lactams disrupts the balance between cell

wall degradation and reproduction in growing cells, weakening cell wall structure,

and ultimately causing cytolysis due to osmotic pressure.

22  

In recent years, the prevalence of bacterial resistance to -lactams has increased

considerably, leading to an enormous concern in the health care setting. One

major mechanism of -lactam resistance comes from -lactamases, which

hydrolyze the cyclic amide bond of the -lactam rings, thereby inactivating the

drug (63). Using an alternative PBP that is less reactive with -lactams also

represents a common strategy for -lactam resistance. In Gram-negative

bacteria (which have an outer membrane surrounding the peptidoglycan), porin

modification and induction of efflux pumps can minimize the concentration of -

lactam in the periplasm, and thereby also contributing to -lactam resistance (71).

1.5.2 Aminoglycosides

Aminoglycosides were first discovered in 1944. These amino-modified sugars are

positively charged at neutral pH because of the protonated amine groups. These

drugs bind electrostatically to the 16s rRNA, at the tRNA acceptor A site in the

30S ribosomal subunits. This binding interferes with the decoding process,

causing misreading and mistranslation, inhibiting protein synthesis, and

eventually leading to cell death (66,72).

The lethal activity of aminoglycosides is dependent upon transport across the

bacterial membrane, which can be summarized in three steps (100,102). (a) The

electrostatic binding of cationic drugs to anionic sites on the cell surface. This

step is rapid, reversible, concentration dependent, and energy-independent. (b)

Energy-dependent phase I. This is a slow step that requires a dose-dependent

threshold proton motive force. In this step, a small amount of drug enters the

23  

cytoplasm, causing mistranslation and producing misfolded proteins.

Subsequently, some of the misread proteins are incorporated into the membrane,

increasing membrane permeability and triggering step (c)-energy-dependent

phase II. In step (c), large quantities of the drug gets into the cell, irreversibly

binding to ribosomes, inhibiting protein synthesis, and ultimately causing cell

death.

There are four major mechanisms of aminoglycoside resistance (72,102): (a)

inactivation of aminoglycosides by enzymatic modification, such as N-acetylation,

adenylylation or O-phosphorylation; (b) inhibiting the drug-target interaction by

16S rRNA methylation or ribosomal alteration; and minimizing intracellular

concentration of the drug by (c) reducing drug uptake or (d) increasing drug efflux.

Anaerobiosis is known to be intrinsically resistant to low doses of

aminoglycosides, because anaerobic fermentation usually generates a lower

proton motive force that impairs aminoglycoside uptake. The sensitivity can be

restored by using higher concentrations of aminoglycosides to reduce the

threshold proton motive force, or by supplying nitrate as an alternative respiratory

substrate to generate higher proton motive force under anaerobic conditions

(102).

1.5.3 Fluoroquinolones

The first quinolone, nalidixic acid, was discovered in 1962. Fluoroquinolones are

synthesized by substitution of a fluoride atom at position C-6 on nalidixic acid,

and this modification greatly improves the biological activity of the drug.

Additional structural modification further broadens the antibacterial spectrum,

24  

making fluoroquinolones one of the most wildly used antibiotic agents in clinical

settings (26).

Quinolones inhibit two essential type II topoisomerases in bacterial: DNA gyrase

is their primary target in Gram-negative bacteria, while topoisomerase IV is the

one in Gram-positive bacteria (26,60).

During DNA replication, rapid rotation of the DNA at the replication fork

generates positive supercoils, which will block replication fork movement if they

accumulate. DNA gyrase acts in front of the replication fork to relieve positive

supercoils and introduce negative supercoils. Topoisomerase IV is involved in

decatenation of two daughter molecules after DNA replication. Both enzymes

induce transient staggered, single strand DNA breaks during their function, and

then reseal the phosphodiester bonds after strand passage (26,60).

Fluoroquinolones enter the cell by passive diffusion. These drugs then quickly

bind to type II topoisomerase-DNA complex and trap the enzymes on DNA. After

DNA cleavage occurs in the drug-enzyme-DNA complex, the 5’ ends of the

broken DNA are covalently attached to the enzyme, but re-ligation is inhibited by

the drug. This complex is called cleaved complex, which is reversible if quinolone

is removed and thus is not lethal by itself. At bacteriostatic concentrations, the

formation of cleaved complexes blocks the movement of replication forks,

induces SOS response and inhibits cell growth. But the broken DNA is locked in

the complexes, and the chromosome remains intact. However, at bactericidal

concentrations, the broken DNA ends are released from the cleaved complexes,

25  

causing chromosome fragmentation and cell death. Depending on the drug

structure, lethal effects of fluoroquinolones can be oxygen dependent, protein

synthesis dependent, or both oxygen and protein synthesis independent

(25,26,60,73).

The mechanisms of resistance to quinolone come from three common antibiotic-

resistance strategies: (a) decrease the drug uptake by porin deletion to reduce

outer-membrane permeability; (b) increase the drug export by expression of

multidrug efflux pumps; (c) alter the drug targets by mutations in DNA gyrase and

topoisomerase IV (103).

1.5.4 Hypothesis of antibiotic-mediated oxidative stress

Unlike redox cycling drugs, the action of classic antibiotics does not produce O2-

or H2O2, and their bacterial resistance mechanisms do not involve defense

systems against oxidative stress. However, about ten years ago, several studies

reported that by using nitroblue tetrazolium reaction (NBT) and lucigenin,

intracellular O2- formation were detected after incubation with -lactams or

fluoroquinolones. This raised the possibility that oxidative stress could be

induced in antibiotic treated bacterial cells (1,8). A few years later, researchers

found that addition of glutathione or ascorbic acid, and overexpression of

superoxide dismutase prevented fluoroquinolone killing (37). Then several other

groups did systematic studies and reported that classic bactericidal antibiotics,

including aminoglycosides, -lactams, and fluoroquinolones, triggered oxidative

stress that led to cell death (16,27,32,61,62). In these later studies, E. coli cells

were treated with low doses of antibiotics in LB broth under aerobic conditions. A

26  

redox sensitive, cell-penetrating fluorescent probe (hydroxyphenyl fluorescein,

HPF) gave higher fluorescent signals in antibiotic-treated cells, and this signal

was attributed to hydroxyl radicals. Iron chelators (dipyridyl or o-phenanthroline),

which suppress hydroxyl radical-generating Fenton reaction, and thiourea, a

potent quencher of hydroxyl radicals, improved the survival rate and decreased

the fluorescent signal. The SoxRS system but not the OxyR regulon was induced

after antibiotic treatment. Mutants in the tricarboxylic acid cycle (TCA) were more

resistant to antibiotics. NADH concentration greatly dropped upon antibiotic

exposure. Furthermore, DNA repair-deficient mutants (recA) were somewhat

more sensitive to low doses of antibiotics. Therefore, the authors proposed that

bactericidal antibiotics were able to cause oxidative damage, and that was a

common mechanism of their lethal effect. In this model, bactericidal antibiotics,

irrespective of their drug-targets, stimulate NADH consumption via the electron

transport chain. Hyperactivation of the electron transport chain accelerates

superoxide formation. Superoxide damages solvent exposed [Fe-S] cluster

containing enzymes, releasing iron into the cytoplasm, and driving hydroxyl

radical formation via the Fenton reaction. Then Fenton-mediated DNA damage

contributes to cell death. This scenario could explain the observed oxidation of

intracellular dyes, protection by scavengers and chelators, a requirement for

respiration, and the sensitivity of DNA-repair mutants.

This is a very exciting idea and quickly became highly cited work. These

approaches for ROS detection (using ROS scavenger or HPF probe) were also

widely used in various studies(14,40,80,84,97,106,110). If the hypothesis is true,

27  

it could change our strategy for antimicrobial therapies. However, the available

data are suggestive but not conclusive. It is unclear how different drug-target

interactions stimulate metabolic feedback and NADH consumption. Though

aminoglycosides were suggested to stimulate reactive oxygen species formation

by triggering envelope stress response (62), there was no model for -lactams

and fluoroquinolones. Furthermore, there are no direct data about the respiration

rate, the levels of superoxide or hydrogen peroxide, the oxidant-sensitive enzyme

activities and the intracellular iron concentrations. Also, induction of the SoxRS

system but not the OxyR regulon makes one concerned about whether oxidative

stress really occurs, since activation of the SoxRS system is not directly

correlated with O2- stress (42). Adding iron chelators or thiourea to the cell may

alter cellular metabolism, and their protective effects may not be related to the

Fenton chemistry. At last, chemiluminescent or fluorescence probes were used in

these studies to detect ROS, which might raise some problems. For examples,

NBT and lucigenin were used in cell systems for O2- measurement in the first

studies (1,8). However, NBT is only designed for cell free systems, and lucigenin

is not a reliable probe for O2-. In the later studies, HPF probes were widely

applied for hydroxyl radical detection (16,27,32,61,62). Though HPF probes can

be oxidized in a Fenton system (96), there are no direct data showing that it is

specific for hydroxyl radicals inside the cell. HPF has a structure similar to

DCFH2-DA (Figure 1.2E); therefore, the oxidation of HPF may also be

complicated as that of DCFH2 inside a biological system, and the intensity of

fluorescence signals may not represent the level of hydroxyl radicals. Hence,

28  

more work need to be done to evaluate this hypothesis and my second project is

to directly test the molecular mechanisms underlying this hypothesis.

29  

1.6 Tables

Table 1.1 H2O2-inducible defense system-OxyR regulon.

Role OxyR regulated gene

Defend against the Fenton chemistry

H2O2 scavenging ahpCF

katG

Fe-import control fur

Fe scavenging dps

Maintain enzyme function

Fe-S cluster assembly & repair sufABCDE

Mn transporter mntH

Others

Disulfide reduction

trxC

grxA

gor

Protein degradation clpS

Unknown function

yaaA

yajA

ybjM

   

30  

Table 1.2 Redox cycling drug-induced defense system-SoxRS regulon.

Role SoxS regulated gene

Control intracellular drug concentration

Cell envelop permeability

micF

waaY

waaZ

Drug efflux pumps acrAB

tolC

Drug detoxification

nfsA

ygfZ

nfnB

Multiple antibiotic resistance marAB

Reduce O2- damage

Scavenging O2- sodA

Fe-import control fur

DNA repair nfo

Fe-S cluster repair yggX

Oxidant-resistant dehydratases-isoform acnA

fumC

Restore NADPH pool

Restore NADPH pool

zwf

fpr

fldA

fldB

31  

1.7 Figures

 

 

 

Figure 1.1 Life evolved in an anaerobic environment. (Adapted from Alberts, et al. Molecular Biology of the Cell, 4th edition, 2002)

   

32  

Figure 1.2 Common ROS detection probes

33  

1.8 References

 

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3. Altuvia, S., M. Almiron, G. Huisman, R. Kolter, and G. Storz. 1994. The dps promoter is activated by OxyR during growth and by IHF and sigma S in stationary phase. Mol. Microbiol. 13:265-272.

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5. Anjem, A. and J. A. Imlay. 2012. Mononuclear iron enzymes are primary targets of hydrogen peroxide stress. J. Biol. Chem. 287:15544-15556.

6. Anjem, A., S. Varghese, and J. A. Imlay. 2009. Manganese import is a key element of the OxyR response to hydrogen peroxide in Escherichia coli. Mol. Microbiol. 72:844-858

7. Aslund, F., M. Zheng, J. Beckwith, and G. Storz. 1999. Regulation of the OxyR transcription factor by hydrogen peroxide and the cellular thiol-disulfide status. Proc. Natl. Acad. Sci. U. S. A. 96:6161-6165.

8. Becerra, M. C. and I. Albesa. 2002. Oxidative stress induced by ciprofloxacin in Staphylococcus aureus. Biochem. Biophys. Res. Commun. 297:1003-1007.

9. Bjerrum, C. J. and D. E. Canfield. 2002. Ocean productivity before about 1.9 Gyr ago limited by phosphorus adsorption onto iron oxides. Nature. 417:159-162.

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11. Bolton, J. L., M. A. Trush, T. M. Penning, G. Dryhurst, and T. J. Monks. 2000. Role of quinones in toxicology. Chem. Res. Toxicol. 13:135-160.

12. Burman, J. D., R. L. Harris, K. A. Hauton, D. M. Lawson, and R. G. Sawers. 2004. The iron-sulfur cluster in the L-serine dehydratase TdcG from Escherichia coli is required for enzyme activity. FEBS Lett. 576:442-444.

13. Carlioz, A. and D. Touati. 1986. Isolation of superoxide dismutase mutants in Escherichia coli: is superoxide dismutase necessary for aerobic life? EMBO J. 5:623-630.

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14. Choung, Y. H., A. Taura, K. Pak, S. J. Choi, M. Masuda, and A. F. Ryan. 2009. Generation of highly-reactive oxygen species is closely related to hair cell damage in rat organ of Corti treated with gentamicin. Neuroscience. 161:214-226.

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CHAPTER TWO: THE YAAA PROTEIN OF THE ESCHERICHIA

COLI OXYR REGULON LESSENS HYDROGEN PEROXIDE

TOXICITY BY DECREASING INTRACELLULAR

UNINCORPORATED IRON CONCENTRATION

2.1 Introduction

Microarray data from a previous study indicated that the peroxide responsive

OxyR regulon includes several genes whose roles are not obvious (40). One of

these, yaaA, is predicted to encode a 29.6 kDa cytoplasmic protein that belongs

to a protein family domain of unknown function (DUF328). Members of the

DUF328 family are found in 1, 392 species, including 1, 377 Bacteria and 15

Eukaryotes, but they are not found in Archea currently in Pfam database (8).

Since YaaA is conserved in a large number of bacteria, it appears that

maintaining this gene may provide these organisms with a selective advantage.

Exploring the function of YaaA may provide insight into the H2O2 defense

systems that are used by other organisms.

2.2 Materials and Methods

2.2.1 Reagents

-Mercaptoethanol, bovine liver catalase, chelex 100, L-cystine dihydrochloride,

deferoxamine mesylate (DFO), diethylenetriaminepentaacetic acid (DTPA), ferric

chloride, type II horseradish peroxidase, 30% hydrogen peroxide, -nitropheny--

D-galactopyranoside (ONPG), thiamine, and trimethoprim were purchased from

Sigma. Potassium cyanide and -(4-pyridyl-1-oxide)-N-tert-butylnitrone (4-POBN)

44  

were from Aldrich. Hanks balanced salt solution without CaCl2, MgCl2, MgSO4,

and phenol red (HBSS) was obtained from GIBCO. Coomassie Protein Assay

Reagent and Albumin Standard were purchased from Thermo Scientific. Hy-

Case Amino and 2, 4-dinitrophenylhydrazine (DNPH) were from Fluka, and

Amplex Red was purchased from Molecular Probes. SuperScript II reverse

transcriptase was from Invitrogen. Standard reagents for RNA work and DNase I

were obtained from Ambion. iQTM SYBR Green Supermix was from BioRad, and

real-time PCR plates were purchased from Eppendorf.

2.2.2 Strain construction

The strains used in this study are listed in Table 2.1. Deletions were constructed

by the Red recombination deletion method (7) and were transferred to other

strains by P1 transduction (22). The antibiotic resistant cassettes were

subsequently removed by FLP-mediated excision (7). Gene fusions between

yaaA upstream region (-290 bp to -1 bp) and lacZ were created by standard

methods and integrated into the lambda phage attachment site (13), preserving

the wild-type yaaA locus at its native chromosomal locus.

2.2.3 Plasmid construction

The yaaA coding sequence was amplified by PCR from E. coli MG1655 genomic

DNA using the forward primer 5-TTTCAGAATTCTAAATTTCCTGCAAGGACTG-

3 and the reverse primer 5-GATCCTCTAGACAAACTTAACGCTGCTCGTA-3.

These oligonucleotide primers were designed to engineer EcoRI and XbaI sites

at the 5 and 3 ends of the gene, respectively. The PCR products were purified,

45  

digested with EcoRI and XbaI and inserted into pWKS30 vector (34) so that yaaA

expression was controlled by the lac promoter. The correctness of the plasmid

construct was confirmed by sequencing; it successfully complemented the

phenotype of yaaA deletion in H2O2 stress cells without causing growth defect in

yaaA+ strains.

2.2.4 Bacterial growth

Complex growth medium was LB (10 g BactoTM tryptone, 5 g yeast extract, and

10 g NaCl per liter). Defined glucose/amino acids medium contained minimal A

salts (22), 0.2% glucose, 0.2% casamino acids, 0.5 mM tryptophan, 0.02%

MgSO47H2O, and 5 g/ml thiamine. When antibiotic selection was required,

media were supplemented with 100 g/ml ampicillin sodium salt, 20 g/ml

chloramphenicol, 30 g/ml kanamycin sulfate, 100 g/ml spectinomycin

dihydrochloride, or 12 g/ml tetracycline HCl.

Experimental protocols were designed to ensure that measurements were

performed upon exponentially growing cells. For growth studies, anaerobic

overnight cultures were diluted into anaerobic media and grown for 4 to 5

generations to early log phase (OD600nm of 0.15 to 0.20). The cultures were then

diluted into aerobic medium of the same composition to an OD600nm of 0.0005-

0.010. All cultures were grown at 37 C. Aerobic LB medium was made one day

previously and stored in the dark to avoid the photochemical generation of H2O2.

Aerobic defined medium was prepared immediately prior to use. Anaerobic

medium was transferred immediately after autoclaving to an anaerobic chamber

46  

(Coy Laboratory Products), where it was stored under an atmosphere of 5%

CO2/10% H2/85% N2 for at least one day prior to use. When used, DFO was

made fresh and added into the media at a final concentration of 2 mM before

inoculation.

2.2.5 Cell viability

To determine cell viability, cell samples were collected at designated time points

and mixed with catalase (final 250 U/ml) to scavenge residual H2O2. Samples

were then serially diluted, mixed with top agar, and spread onto the surface of

anaerobic plates. Colony-forming unit (CFU) enumeration was performed after

24-48 h anaerobic incubation.

For co-culture experiments, anaerobic exponential-phase cells were mixed at 9:1

and 1:9 ratio of Hpx- yaaA+ (chloramphenicol sensitive) to Hpx- yaaA-

(chloramphenicol resistant) in aerobic LB. Mixed cultures were aerated by

vigorous shaking and viability was followed over time. The viability of Hpx- yaaA-

cells was determined by plating on LB + 20 g/ml chloramphenicol plates,

whereas the total viability of co-cultured cells was determined by plating on LB

plates.

2.2.6 Differential interference contrast (DIC) microscopy

Cultures were grown in anaerobic LB medium to exponential-phase before

dilution into aerobic LB medium to an OD600nm of 0.0005. They were then grown

for 5 hours. Images were captured by DIC as described by Gut, et. al (12).

47  

2.2.7 thyA forward mutagenesis assay

In order to determine the frequency of mutagenesis, Thy- mutants were selected

using trimethoprim (22). Strains were transferred from anaerobic LB to aerobic

LB as described above. At designated intervals, aliquots were removed, mixed

with catalase to stop oxidative stress, and washed with minimal glucose/amino

acids medium at room temperature three times. The cells were placed in an

anaerobic chamber and diluted into minimal A salts. The number of viable cells

was determined by dilution and plating in 2 ml F-top agar containing 1 mg/ml

thymidine on defined glucose/amino acids plates. thyA mutants were selected for

by including 0.2 mg/ml trimethoprim in the F-top agar and CFU were determined

after anaerobic incubation at 37 °C for 48 h. For each experiment, a subset of the

trimethoprim-resistant colonies were tested and confirmed to be Thy- by

examining their growth on minimal glucose/amino acids plates with and without

thymidine supplementation.

2.2.8 H2O2 killing assay

Cells were grown in minimal A glucose medium containing 0.5 mM tryptophan,

phenylalanine, and tyrosine to an OD600nm of 0.15 as described above. The

cultures were then diluted to an OD600nm of 0.025 in the same medium and

incubated for 5 min with either 3.3 mM potassium cyanide (37) or with 0.5 mM

cystine (23). H2O2 (final 2.5 mM) was then added. At time points, aliquots were

removed, and catalase (final 2500 U/ml) was added in order to terminate the

H2O2 stress. After serial dilution, samples were mixed with top agar and spread

48  

on minimal A defined medium plates. CFUs were determined after anaerobic

incubation at 37 °C for 48 h.

2.2.9 EPR spin trapping of hydroxyl radicals

Cells were cultured in aerobic LB as described above. The cells were then

harvested at an OD600nm of 0.2, when the growth of Hpx- yaaA cells arrested. The

cells were washed twice with 25 ml of ice-cold chelex-treated HBSS and

resuspended in 0.5 ml of ice-cold HBSS. The cell suspension was added to a

room temperature mixture (total volume 1 ml) containing 100 mM DTPA, 10 mM

4-POBN, 170 mM ethanol, 1 mM H2O2, and HBSS and incubated at room

temperature for 12 min. The spectra of POBN ethanol radical adducts were

measured by EPR at room temperature with the following settings: field center,

3,393 G; field sweep, 100 G; modulation frequency, 100 kHz; modulation

amplitude, 1 G; time constant, 0.064; receiver gain, 63,000; and power, 20 mW

(20). The final cell densities were equivalent in all samples (~20 OD600nm).

2.2.10 EPR measurement of intracellular unincorporated iron

The cells were cultured in 600-1000 ml of aerobic LB as described above. Cells

were centrifuged and resuspended at 100-fold higher cell density in LB medium

containing 10 mM DTPA (pH 7.0) and 20 mM DFO (pH 8.0). The suspension was

incubated at 37 C aerobically for 15 min. The purpose of the cell-impermeable

chelator, DTPA, was to block further iron import, while and the cell-penetrating

iron chelator, DFO, facilitated oxidation of unincorporated ferrous iron to EPR-

detectable ferric iron, as described previously (36). The cell pellet was washed

twice with 5 ml of ice-cold 20 mM TrisHCl (pH 7.4) and finally re-suspended in

49  

200 l of ice-cold 20 mM TrisHCl-30% glycerol (pH 7.4). EPR analyses were

performed as described previously (20). EPR signals were measured at a

temperature of 15 K in a Varian Century E-112 X-band spectrophotometer

equipped with a Varian TE102 cavity and temperature controller, with the

following settings: field center, 1,570 G; field sweep, 400 G; modulation

frequency, 100 kHz; modulation amplitude, 12.5 G; time constant, 0.032; receiver

gain, 4,000; and power, 10 mW. The measured EPR signals were normalized by

cell density and converted to intracellular concentrations using the following

conversion: 1 ml of E. coli culture at OD600nm of 1 comprises 0.52 l of

intracellular volume (15).

2.2.11 -Galactosidase activity

Cell extracts were prepared in ice-cold 50 mM TrisHCl (pH 8.0) by passage

through a French pressure cell (SIM Aminco FA-003). Cell debris was removed

by centrifugation, and -galactosidase activity in the supernatant was measured

by standard procedures (22). Protein concentrations were determined by the

Bradford assay using bovine serum albumin as the standard.

2.2.12 H2O2 measurement

Cells were cultured in defined glucose/amino acids medium under anaerobic

conditions to log phase, and then they were diluted into aerobic medium to an

OD600nm of 0.005 and grown to an OD600nm of ~ 0.15. These cells were

centrifuged, suspended in fresh defined medium at an OD600nm of 0.020, and

incubated aerobically at 37 °C with vigorous aeration. At time points, cells were

50  

removed by centrifugation, and H2O2 concentrations were measured in the

supernatants using the Amplex Red/horseradish peroxidase method (28) in a

Shimadzu RF Mini-150 fluorometer. The rate of H2O2 formation was normalized

by cell density and converted to intracellular concentrations using the following

conversion: 1 ml of E. coli culture at OD600nm of 1 comprises 0.52 l of

intracellular volume (15).

2.2.13 Visualization of protein carbonylation

Cells were cultured in aerobic LB as described above. Hpx- yaaA cells were

harvested by centrifugation at the beginning (0.2 OD600nm) and in the middle of

the growth lag (~0.25 OD600nm). Other strains were collected at an OD600nm of

~0.2. Cell pellets were washed with ice-cold 50 mM potassium phosphate buffer,

pH 7.0, and re-suspended in the same buffer with 5 mM DTPA, which prevents

further protein oxidation in cell extracts. After lysis in a French pressing cell,

cellular debris was removed by centrifugation, and protein concentrations in the

cell extracts were determined using the Bradford assay. All samples were then

diluted to the same protein concentration (6 mg/ml). Protein carbonyl groups

were derivatized with DNPH, and the adducts were visualized by western blotting

(anti-DNP) (2).

2.2.14 Rapid amplification of cDNA ends (5-RACE)

MG1655 cells and Hpx- cells were cultured in LB as described above and

harvested at an OD600nm of ~0.2. RNA was extracted by the hot phenol method.

Contaminating DNA was removed by DNase I digestion, and the DNA-free RNA

samples were converted to cDNA by SuperScript II Reverse Transcriptase

51  

according to the manufacturer’s instructions. The 5-end of yaaA transcripts was

determined by FirstChoice RLM-RACE Kit (Applied Biosystems). The PCR

primers used were: yaaA outer 5-race, 5-CTCGTTCAGCTTGTTGGTGAT-3;

yaaA inner 5-race, 5-AGCCGGTGTAGACATCACCTTT-3. Thermal cycling was

performed for 35 cycles in 3 steps: 94 C for 30 s, 65 C for 30 s and 72 C for 1

min.

2.2.15 Real-time PCR

Cells were cultured in LB as described above. When cultures achieved an

OD600nm of ~0.2, they were treated with H2O2 (0, 0.25, 1, and 2 mM) for 10 min at

room temperature. mRNA was harvested and converted to cDNA as described

above. The following primers were used to analyze expression of yaaA and the

housekeeping gene gapA: yaaA forward, 5-GCTGTTAGACAATTCCCAGCAG-3;

yaaA reverse, 5-AAGCCGGTGTAGACATCACCTT-3; gapA forward, 5-

CGTTCTGGGCTACACCGAAGATGACG-3; gapA reverse, 5-

AACCGGTTTCGTTGTCGTACCAGGA-3. The 25 l real-time PCR mixture

contained 1 ng of cDNA, 12.5 l of SYBR Green Supermix, and 1.6 M of each

primer. Thermal cycling was performed in Mastercycler ep realplex (Eppendorf)

for 40 cycles in 3 steps: 95 C for 20 s, 55 C for 20 s and 68 C for 20 s.

52  

2.3 Results

2.3.1 The yaaA gene is induced by OxyR

The yaaA gene is predicted to encode a 29.6 kDa conserved cytoplasmic protein

that belongs to family domain of unknown function DUF328 (8). YaaA does not

possess amino acid similarity to proteins that have defined function, and it does

not have any recognizable motifs. DNA microarray data from Zheng et al.

showed that yaaA was induced in wild-type E. coli cells in an OxyR-dependent

manner when 1 mM H2O2 was applied (40). Our analysis of the DNA sequence

upstream of yaaA revealed a potential OxyR binding site approximately 74 bp

upstream of the yaaA start codon (Figure 2.1). To verify these observations, we

constructed a yaaA-lacZ transcriptional fusion and measured the expression of

yaaA in different strains.

When transferred from anaerobic to aerobic conditions, the expression of yaaA

was not induced in wild-type cells (E. coli strain MG1655). However, in an

aerated catalase/peroxidase-deficient mutant (Hpx-), which accumulates ~1 M

endogenous H2O2 and in which the OxyR response is fully activated (3,28), the

expression of yaaA was induced 10-fold (Figure 2.2A). Induction did not occur if

an oxyR deletion was present (Figure 2.2B). Rapid amplification of cDNA ends

(5-RACE) revealed that the 5-end of yaaA transcripts were identical for both

wild-type cells and Hpx- cells, mapping to 22 bp upstream of the yaaA start

codon (Figure 2.1). When wild-type cells were transformed with the plasmid

pGSO58, which contains a mutated oxyR allele (A233V) that constitutively

53  

expresses OxyR-regulated genes even in the absence of H2O2 stress (19), the

expression of yaaA was induced under both anaerobic (data not shown) and

aerobic conditions (Figure 2.2A). These results confirm that yaaA is a gene of the

OxyR regulon.

Interestingly, in ahp- (catalase-proficient) mutants that experience minor H2O2

stress (29), the expression of yaaA was at the basal level under aerobic

conditions (Figure 2.2A), even though katG, another member of OxyR regulon,

was strongly induced (data not shown). The result suggests either that the yaaA

promoter region has a lesser affinity for oxidized OxyR, or alternatively that a

modifying effect is imposed by another regulator that is affected by more-severe

H2O2 stress.

2.3.2 Hpx- yaaA cells show a growth defect in LB during H2O2 stress

Since yaaA was induced in Hpx- strains during aerobic culture, we constructed a

yaaA deletion in the catalase/peroxidase-deficient background (Hpx- yaaA). The

Hpx- yaaA cells grew normally under anaerobic conditions, but they stopped

growing about five generations after shift to aerobic conditions, lagging for

several hours before recovering (Figure 2.3A).

The aerobically grown Hpx- yaaA cultures filamented profusely (Figure 2.4). Both

anaerobic cultures and aerobic Hpx+ yaaA cultures appeared as normal unit-

sized cells (data not shown), confirming that H2O2 was needed to create this

phenotype. Filamentation is a hallmark of difficulty in DNA replication, and it

54  

commonly results when chemical stress damages DNA but does not substantially

disrupt protein or membrane biogenesis.

A chromosomal insertion of the yaaA+ allele at the phage attachment site

suppressed the growth defect and the filamentation of Hpx- yaaA cells (data not

shown). Deletion of yaaJ, which terminates 70 bp upstream of yaaA, in the Hpx-

strain does not result in a growth defect during H2O2 stress (data not shown).

These results indicated that the growth defect of the Hpx- yaaA mutant was due

to the loss of YaaA function and that YaaA is involved in the cellular response to

H2O2 stress.

The yaaA expression was also strongly induced when Hpx- strains were aerated

in defined glucose medium (data not shown). However, in contrast to the results

in LB broth, the Hpx- yaaA mutant did not exhibit any phenotype in defined

glucose medium (data not shown). The growth defect reemerged when yeast

extract was added to the defined medium (see Discussion).

2.3.3 Hpx- yaaA cells lose viability during H2O2 stress

The viability of Hpx- yaaA strains and Hpx- parent strains were monitored upon

aeration (Figure 2.5). Samples were harvested at different time points and colony

forming units (CFUs) were determined by plating in LB top agar. After a brief lag,

the number of viable cells increased steadily in the Hpx- parent strain. In contrast,

the viable cell count of the Hpx- yaaA strain did not change during the first two

hours of aerobic culture, confirming that the increase in OD600nm was due to

filamentation rather than cell division. Approximately three hours after aeration,

55  

concomitantly with the onset of the growth lag, the viability of the Hpx- yaaA

mutant decreased by one order of magnitude.

Ultimately, the number of viable cells began to increase again, albeit at a slower

rate than in the yaaA+ parent. This recovery was not due to suppressor mutations,

since the isolated outgrowers reproduced the same growth pattern when the

experiment was repeated. On the other hand, a lag resumed when the

outgrowing cells were centrifuged and then diluted into fresh aerobic medium,

which suggested that some of the outgrowth depended upon conditioning of the

medium. Sterile lab medium can accumulate 1-20 M H2O2 which causes a

temporary H2O2 stress at the beginning of culture. Although Hpx- strains lose

major H2O2 scavenging enzymes, they retain ~5% scavenging activity because

the cytochrome oxidases in the respiratory chain can use H2O2 as a weak

substrate (28) (Woodmansee and Imlay, unpublished data). Therefore, Hpx-

mutants can slowly degrade H2O2 and condition the medium. A similar recovery

has been observed in Hpx- strains that lack fur or dps, two other genes that are

critical during H2O2 stress (24,33).

2.3.4 yaaA mutants show higher levels of DNA damage and polypeptide

damage during H2O2 stress

Dysfunction of YaaA generates a phenotype in H2O2 stress cells, which provides

an opportunity to gain insight into the function of YaaA by identifying the cellular

injury that arrests the growth of Hpx- yaaA mutants.

H2O2 can directly inactivate enzymes that employ solvent-exposed iron atoms

either in Fe-S clusters or as mononuclear cofactors (17,30). Further, if H2O2

56  

reacts with ferrous irons associated on DNA or proteins, the resulting hydroxyl

radicals will subsequently create DNA or polypeptide damage (2,14,24).

The filamentation and cell death of Hpx- yaaA strains suggested DNA damage.

Indeed, a thyA forward mutagenesis assay revealed that the mutation frequency

of aerobically grown Hpx- yaaA cells was 4-8 fold higher than that of the Hpx-

strain (Figure 2.6). The effect depended upon oxidative stress, since the yaaA

mutant in the wild-type (scavenger-proficient) background exhibited a mutation

frequency similar to that of the parent strain. The yaaA mutation did not sensitize

cells to UV radiation, an alternative source of DNA damage (data not shown).

In a scavenger-proficient background the yaaA mutation did create a growth

defect in recA56 strains (Figure 2.7A-B), which are defective in the repair of

oxidative DNA lesions (1). This defect was apparent, once again, only in LB

medium and only under aerobic conditions. The defect was blocked when

catalase was added to the medium (Figure 2.7C), which implied that the damage

was due to the trace H2O2 that is present in aerobic LB medium. A similar result

was obtained with polA1 mutants (Figure 2.8). The defect is transient due to the

decomposition of H2O2 by cellular enzymes. These data indicate that (a) YaaA

has a role independent of the DNA recombination pathway or the SOS response,

which are already absent from recA56 mutants, and (b) even very low levels of

H2O2 can cause debilitating DNA damage in the yaaA mutant. The data, however,

do not resolve whether YaaA has a role in preventing DNA damage or in

repairing it or both.

57  

During H2O2 stress, the Fenton-mediated hydroxyl radicals can also damage

polypeptides and generate various oxidative modifications. One of these is

carbonyl adducts, which arise when hydroxyl radicals oxidize side chains of

amino acids. An ELISA-based carbonylation assay did not detect any difference

in the extent of protein carbonylation in Hpx- yaaA and Hpx- yaaA+ mutants when

they were grown in defined glucose medium (data not shown). However, when

they were cultured in aerobic LB medium, the Hpx- yaaA cells gradually

accumulated higher levels of protein carbonyls than did the Hpx- parent strain.

These modifications were apparent at the later time point that coincides with the

growth lag (Figure 2.9). While the most straightforward interpretation is that the

dysfunction of YaaA directly causes the higher carbonylation, it remains possible

that the carbonylation is an indirect consequence of the growth defect.

We also evaluated whether YaaA has a role in protecting mononuclear iron

containing enzymes or aconitase-class dehydratases from H2O2 stress. The

activities of ribulose-5-phosphate epimerase and isopropylmalate isomerase,

respectively, were compared in Hpx- and Hpx- yaaA mutants. Both enzymes

exhibited a reduction in activities relative to scavenger-proficient strains, but the

yaaA allele had no further impact (data not shown).

2.3.5 The Fenton reaction causes injuries in yaaA mutants during H2O2

stress

The preceding results suggested that yaaA might suppress cellular injuries that

arise from the Fenton reaction. Consistent with this idea, the growth defect of

Hpx- yaaA mutants was suppressed by conditions that diminish the amount of

58  

intracellular iron. The exogenous addition of deferoxamine (DFO), a cell-

permeable iron chelator that blocks Fenton chemistry (14), eliminated the growth

lag (Figure 2.3B). A similar effect was obtained by a tonB mutation, which

diminishes iron import by the siderophore and ferric-citrate systems (35) (Figure

2.3B). Furthermore, DFO also eliminated the growth defect of recA yaaA mutants

(Figure 2.7D). These results confirm that the key injury in the yaaA mutant during

H2O2 stress is due to the Fenton reaction.

2.3.6 YaaA protects cells by attenuating the Fenton reaction

In principle YaaA might protect cells either by reducing Fenton-mediated hydroxyl

radical formation or by helping cells to tolerate the damage caused by hydroxyl

radicals. In order to distinguish between these possibilities, we employed

electron paramagnetic resonance spectroscopy (EPR) to evaluate the amount of

hydroxyl radicals formed in yaaA mutants.

Intracellular hydroxyl radicals were trapped by ethanol/POBN and detected at

room-temperature by EPR. Wild-type cells exhibited a vanishingly small signal

(data not shown), whereas the level of hydroxyl-radical formation was substantial

in Hpx- mutants, consistent with prior reports (24). In the early period after

aeration (before the growth lag), the Hpx- yaaA cells exhibited a signal similar to

that of Hpx- cells (data not shown). However, the signal continually increased in

the Hpx- yaaA cells, ultimately becoming about twice that of the Hpx- strain when

the growth arrested (Figure 2.10). Pretreatment with 2 mM dipyridyl (an iron

chelator) for 5 min eradicated the EPR signal (data not shown), confirming that it

59  

represented hydroxyl radicals produced by the Fenton reaction. These results

indicated that YaaA protects cells by attenuating the Fenton reaction.

2.3.7 YaaA does not affect H2O2 levels

The ferric iron produced by the Fenton reaction (reaction 1) can be re-reduced by

cellular reductants (reaction 2), continuously driving the hydroxyl radical

formation as long as peroxide stress exists.

Fe2+ + H2O2 Fe3+ + OH- + HO (Reaction 1)

Fe3+ + red Fe2+ + ox (Reaction 2)

Both intracellular cysteine and reduced flavins have been shown to serve as the

predominant reductants in reaction 2, depending upon the growth conditions

(23,37). To minimize oxidative injuries, cells must control the levels of H2O2,

unincorporated iron, and cysteine/flavin. The spin-trapping data suggested that

YaaA suppresses the level of one of these reactants.

The rate of intracellular H2O2 formation depends upon the titers of autoxidizable

enzymes in the cell, and in Hpx- strains the steady-state level of H2O2 also

depends upon minor scavenging mechanisms. We sought to test whether YaaA

might affect either of these processes. Because H2O2 can diffuse across the

membrane, in a peroxide-scavenging deficient mutant (i.e. Hpx- strains), the

H2O2 concentration rapidly equilibrates between the cytoplasm and the external

medium. Therefore, co-cultured Hpx- strains experience equivalent intracellular

concentrations of H2O2, and a strain that efficiently scavenges H2O2 can

suppress the growth defects that would otherwise arise in a co-cultured strain

60  

that cannot (29). To this end, we co-cultured Hpx- cells and Hpx- yaaA cells

together in LB medium under aerobic conditions, and analyzed the viability of the

two individual strains, which were distinguished upon plating by drug-resistance

markers. In mixed culture, the Hpx- yaaA strain continued to exhibit its full

aerobic viability defect, while the Hpx- strain did not. This result indicates that

YaaA does not alter the intracellular levels of H2O2 (data not shown).

In Hpx- strains, the rate of cellular H2O2 formation equals the rate of H2O2

releasing to external medium, which can be directly measured by Amplex Red.

These experiments were performed in glucose defined media, because yeast

extract in LB interferes with H2O2 measurements. In these conditions, the rate of

H2O2 accumulation was 11 1 nM/min for Hpx- strains, versus 11 0.3 nM/min

for Hpx- yaaA mutants (both at OD600 of 0.020). After normalizing to the

intracellular volume, the H2O2 accumulation rate was about 18 M/s for both Hpx-

and Hpx- yaaA strains, similar to what was previous reported (28). This result

further indicates that YaaA does not affect the rate of H2O2 accumulation.

2.3.8 YaaA does not inhibit ferric iron reduction

FADH2 and cysteine can each reduce ferric iron and drive the Fenton reaction

forward in vivo. Reduced free flavin concentrations are elevated by respiratory

blocks (i.e. cyanide treatment), through the action of an NADH-dependent flavin

reductase (Fre). Under these conditions H2O2 creates Fenton-driven DNA lesions

at a very high rate (37). However, in defined medium neither deletion nor

induction of the yaaA allele affected the rate at which cyanide/H2O2 killed cells

(data not shown). The same was true when cells were shifted from sulfate to

61  

cystine medium (data not shown), a protocol that elevates intracellular cysteine

levels and similarly amplifies H2O2 sensitivity (23). These results suggest that

YaaA is unlikely to attenuate the Fenton reaction by inhibiting the rate of ferric

iron reduction (reaction 2).

2.3.9 Hpx- yaaA mutants have higher unincorporated iron levels than the

Hpx- parent strain

The rate of oxidative DNA damage is directly determined by the H2O2

concentration as well as the amount of unincorporated iron inside the cell (18,31).

To test whether or not YaaA controls iron levels, we measured intracellular

unincorporated iron by whole-cell EPR. Samples were cultured aerobically in LB

and collected at OD600nm of 0.1, 0.2 and 0.3; these time points represented Hpx-

yaaA cells prior to the growth lag, at its onset, and at its midpoint. The data

showed that deletion of yaaA in unstressed Hpx+ cells had no effect on the iron

levels (~ 65 M) (Figure 2.11A). The level of unincorporated iron in H2O2-

stressed Hpx- strains was substantially higher (150 M), an effect that may reflect

the oxidative degradation of Fe-S clusters. The key observation, however, was

that the Hpx- yaaA cells consistently accumulated two-fold higher levels of

unincorporated iron (300 M) by the beginning (0.2 OD600nm) of the growth lag

(Figure 2.11A). This result matches the two-fold effect of yaaA upon hydroxyl-

radical formation. The iron returned to Hpx- yaaA+ levels when yaaA was

expressed in trans on a plasmid (Figure 2.11B).

62  

Induction of plasmid-borne yaaA anaerobically also protected recA56 against

H2O2 killing (Figure 2.12A), though it had no impact on the unincorporated iron

level in anaerobic cells (~310 M) (Figure 2.12B).

Hpx- yaaA cells do not show a growth defect compared to Hpx- parent strains in

defined medium during aerobic culture. Consistent with the growth curve results,

the EPR data showed that the level of intracellular free iron in Hpx- yaaA cells

was similar to that of Hpx- cells in defined medium, approximately 100 M for

both strains at 0.2 OD600nm (data not shown).

These results indicate that YaaA suppresses hydroxyl-radical formation, and thus

the rate of oxidative DNA damage by diminishing the amount of unincorporated

iron inside oxidatively stressed cells.

2.3.10 YaaA does not repress expression of iron importers

The factors controlling iron import, trafficking, and disposition within the cell are

largely mysterious. The best-understood facet of iron regulation is the

transcriptional control upon the synthesis of iron importers and of iron-containing

enzymes (4,21). The primary iron importers active in lab cultures of E. coli are

the ferric enterobactin uptake (Ent) system, the ferric citrate uptake (Fec) system,

and the ATP-dependent ferrous import (Feo) system. We observed that Hpx- fec

yaaA, Hpx- entA yaaA and Hpx- feo entA yaaA mutants all exhibited aerobic

growth defects compared to their yaaA+ counterpart strains (data not shown),

which implied that the yaaA phenotype did not depend on a specific iron importer.

However, all three systems are controlled at the transcriptional level by the Fur

63  

repressor, and it seemed plausible that YaaA might perturb the function of this

regulator. More generally, in order to evaluate whether YaaA controls iron levels

by controlling transcription of the importer(s), we constructed feo entA

derivatives which rely upon the Fec system as the sole mechanism of iron uptake.

Using a fecA-lacZ transcriptional fusion integrated at the lambda attachment site,

we found that mutation of the yaaA allele had no impact upon fecA transcription

(Figure 2.13). Thus YaaA does not control the expression of iron importers. Other

possible points of iron control exist, such as influence on transporter activity.

2.3.11 Hpx- mntH yaaA mutants show a strong growth defect in defined

medium

YaaA may work with other proteins to control the iron level during peroxide stress.

Candidates for these proteins include the iron uptake regulator (Fur), iron

scavenging protein (Dps), and manganese transporter (MntH); all of them are

involved in iron homeostasis during the OxyR response. The yaaA deletion

exacerbated the growth defect and viability defects of Hpx- fur and Hpx- dps

strains in LB (Figure 2.14A-B), but not in defined medium (data not shown).

However, Hpx- mntH yaaA mutants showed a strong growth defect in both LB

and defined glucose medium (Figure 2.15). During H2O2 stress, manganese is

imported via MntH to replace the iron cofactor in certain peroxide-sensitive

enzymes (2,30). In defined medium, Hpx- mntH yaaA cells stopped growing

almost immediately after aeration, and supplementation with manganese had

little effect (Figure 2.15B-C). However, the number of viable cells remained

constant (Figure 2.15D), which suggested that this particular phenotype might be

64  

caused by inhibition of metabolism instead of DNA damage. Collectively, these

data show that YaaA had functions independent of Fur, Dps and MntH.

Additional experiments revealed that Hpx- ftn and Hpx- bfr mutants grow as well

as Hpx- strains, indicating that under our conditions ferritin and bacterioferritin do

not play a key role during H2O2 stress and are not involved in the critical function

of YaaA (data not shown). Similarly, although FieF is suspected to act as an iron-

efflux system (9), an fieF deletion did not diminish the growth of an Hpx- strain,

and the further addition of a yaaA mutation recapitulated the Hpx- yaaA

phenotype (Figure 2.16).

2.3.12 Regulation of YaaA

To obtain further insight into the function of YaaA, we measured expression of a

yaaA-lacZ fusion under different conditions (Table 2.2). Though previous DNA

microarray data showed that yaaA was induced 4-fold by treatment with 1 mM

H2O2 in an oxyR strain (40), we did not observe induction using either -

galactosidase assay or real-time PCR measurements (data not shown). The

disagreement may due to different H2O2 concentrations used in experiments.

High level of H2O2 challenge may induce yaaA by other regulators, but under

physiological relevant H2O2 stress, OxyR is the sole transcriptional regulator of

yaaA.

The expression of yaaA was unaltered (< 2-fold induction/repression) under other

tested conditions. It did not respond to manganese availability; it was not induced

65  

by redox-cycling drugs, which indicates that it is not regulated by SoxRS

(10,11,32); and it was not affected by Fur, IscR, Hfq, RpoS, or YaaA itself.

2.4 Discussion

In some natural environments bacteria must cope with extended periods of

exposure to low levels of H2O2, an experience that was recreated in this study

through the use of strains that lack H2O2 scavenging enzymes. The H2O2

concentration that accumulates in the cytoplasm of aerobic Hpx- mutants—

approximately 1 M—is expected to be reached in the cytoplasm of wild-type

cells when they enter environments that contain approximately 5 M H2O2 (29).

The primary effect of these doses upon E. coli is to block growth. The particular

damage mechanisms that have been identified thus far all appear to involve

Fenton-type reactions between iron and H2O2: the creation of DNA lesions and

the inactivation of mononuclear iron enzymes, of Fe-S dehydratases, of the Isc

Fe-S assembly system, and of the Fur repressor (2,2,14,16,17,30,33). The

second-order rate constants for these reactions are high—ca. 103-104 M-1 s-1—

which means that when H2O2 is 1 M they occur with half-times on a minutes

time scale. No other target molecules are known that react with H2O2 so readily,

and so it seems fitting that many of the genes controlled by OxyR serve to avoid

or to alleviate the damage done by Fenton reactions. YaaA now joins Dps (24,38)

and Fur (33) as mediators that minimize the amount of unincorporated iron inside

H2O2-stressed cells. Like Dps and Fur, YaaA has the effect of minimizing

damage both to DNA and to proteins. Like Dps—but unlike Fur—YaaA seems

not to play much of a role during routine growth; it is only during H2O2 stress,

66  

when YaaA levels rise, that its influence is apparent. For this reason yaaA

mutants resemble dps mutants in that they do not exhibit unusual sensitivity in

experiments that involve a sudden bolus of H2O2—a period of OxyR regulon

expression is required for it to become functional. Indeed, mntH and suf mutants

only evince a phenotype during protracted, low-grade stress, too (2,16).

Little is known about the DUF328 family of proteins. They do not have

recognizable motifs or absolutely conserved residues, and no crystal structures

are available. YaaA homologues are found in only a few eukaryotes, all of which

are photosynthetic (four green algae and Ricinus communis). YaaA genes are

widely distributed among bacteria, including both anaerobes and aerobes. Their

presence in anaerobes does not necessarily indicate that they have a non-

oxidative-stress role, since even anaerobes are periodically exposed to

oxygenated waters and customarily encode antioxidant enzymes, including

scavengers of H2O2. A yaaA gene is identifiable in Lactobacillus gasseri, which

uses H2O2 as a chemical weapon to inhibit competitors; but we did not find any

homologue in Borrelia burgdorferi, a bacterium that seems not to have iron-

dependent enzymes or to import iron (26). Thus the circumstantial data are

broadly consistent with a role in iron metabolism and antioxidant activity.

The yaaA phenotype was pronounced in medium that included yeast extract.

EPR measurements show that E. coli generally contains higher levels (~3 fold) of

unincorporated iron when yeast extract is in the medium. This effect cannot be

achieved simply by adding high levels of iron to defined medium, which leads us

to suspect that a component of yeast extract makes iron more available for

67  

import. Interestingly, yaaA was not as easily induced by H2O2 as was katG, as

katG was induced in ahp- mutants, while yaaA was not up-regulated in ahp-

mutant unless the cells were iron starved (Figure 2.17). We do not know whether

the mechanistic basis of this difference stems from a difference in OxyR affinity,

or from an unknown, iron related regulator—but the end effect resembles the

graded responses of members of the SOS regulon, some of which are turned on

rapidly, and others of which are activated later (25). Thus catalase synthesis

might comprise the first response to slight H2O2 stress, whereas alterations in

iron metabolism are engaged only if stress is severe.

All the data suggest that YaaA affects iron levels, so how does YaaA do that?

This question is challenging, because the details of iron metabolism remain foggy.

In principle, intracellular iron levels might be suppressed in a handful of ways: by

slowing iron import, speeding delivery to iron enzymes, sequestering iron in

storage proteins, preventing the leakage of iron from labile enzymes, or pumping

excess iron from the cell (Figure. 2.18). However, YaaA is not an integral

membrane protein, meaning that it could at best be an accessory to efflux; it was

effective no matter which iron import system was active; it did not seem to affect

the damage to aconitase-class dehydratases or mononuclear iron containing

enzymes; and it retained a phenotype whether or not cells contained Dps, the

primary iron-storage system during oxidative stress. Although one can contrive

arguments around these facts, they collectively hint that perhaps the remaining

option—that YaaA is somehow involved in iron trafficking—is the most likely one.

Pull-down experiments did not identify any convincing partner proteins (data not

68  

shown). As our understanding of iron metabolism progresses, perhaps a specific

hypothesis for YaaA mechanism will become apparent, as will experimental

methods that would test it.

2.5 Acknowledgments

We thank Ian Gut (University of Illinois) for assistance with DIC microscopy,

Jason Sobota (University of Illinois) for assistance with the ribulose-5-phosphate

epimerase assay, and Mark Nilges (the Illinois EPR Research Center) for

assistance with EPR experiments.

This work was supported by GM049640 from the National Institutes of Health.

69  

2.6 Tables

Table 2.1 Strains and plasmids used in this study.

Name Genotype/characteristics Source/Ref.

Strains

AA30 As LC106 plus mntH2::cat (2)

AL145 As LC106 plus (lacZ1::cat)1 att::[pSJ501:: yaaA-lacZ+]~cat This study

AL154 As AL157 plus (ahpC-ahpF) kan:: ahpF This study

AL157 As MG1655 plus (lacZ1::cat)1 att::[pSJ501:: yaaA-lacZ+]~cat This study

AL194 As LC106 plus (yaaA1::cat)1 fur::kan zbf-507::Tn10 This study

AL221 As LC106 plus (yaaA1::cat)1 mntH2::cat This study

AL223 As MG1655 plus recA56 srl300::Tn10 (yaaA1::cat)1 This study

AL245 As MG1655 plus (yaaA1::cat)1 This study

AL273 As LC106 plus (feoABC::cat)1 entA1::cat (lacZ1::cat)1

att::[pSJ501::fecA-lacZ+]~cat

This study

AL285 As LC106 plus (yaaA1::cat)1 ∆dps::cat This study

AL293 As AL273 plus (yaaA1::cat)1 This study

AL300 As AL145 plus (tonB1::cat)1feoABC::cat This study

AL302 As AL154 plus (tonB1::cat)1feoABC::cat This study

AL304 As 157 plus (tonB1::cat)1feoABC::cat This study

AL344 As AL145 plus oxyR::spec This study

AL435 As LC106 plus ∆fieF1::cat This study

AL437 As LC106 plus ∆(yaaA1::cat)1 ∆fieF1::cat This study

BW25113 lacIq rrnBT14 lacZWJ16 hsdR514 araBADAH33 rhaBADLD78 (7)

LC106 As MG1655 plus (ahpC-ahpF’) kan:: ahpF (katG17::Tn10)1

(katE12::Tn10)1

(27)

LEM17 As MG1655 plus recA56 srl300::Tn10 Lab stock

70  

Table 2.1 (cont.)

MG1655 Wild type Lab Stock

SB53 As LC106 plus (yaaA1::cat)1 This study

SP66 As LC106 plus mhpC281::Tn10 lacY1 dps::cat (24)

Plasmid

pAH125 A plasmid for lacZ transcriptional fusion construction (13)

pCP20 Temperature-sensitive Flp expression plasmid (6)

pKD3 FRT-flanked cat (7)

pKD46 -Red recombinase expression plasmid (7)

pACYC184 Cmr Tetr (5)

pGSO58 oxyR A233V in pACYC184 (19)

pSJ501 pAH125 derivative with FRT-flanked cat (16)

pWKS30 Plac polylinker Ampr (34)

pWKyaaA pWKS30 containing yaaA insert This study

71  

Table 2.2 Regulation of yaaA.

Test condition Mutants (media) Regulator Resulta

H2O2 stress Hpx- (LB and defined medium) OxyR Yes

Redox-cycling drug WT (LB + 200 M paraquat) SoxR No

Iron rich fur (LB) Fur No

Iron starved tonB feo (defined media + DFO) No

Mn rich mntR (LB) No

Mn starved mntH (defined media) No

Small RNA hfq (LB) Small RNAs No

Stationary phase WT (LB and defined media) RpoS No

iscR- iscR (LB) IscR No

Autoregulation Hpx- yaaA (LB) YaaA No

a Negative results are defined as less than 2-fold changes in the expression of

yaaA.

72  

2.7 Figures

Figure 2.1 Genomic context of yaaA in E. coli. In E. coli, yaaA is located at

0.12 min and is flanked by two genes with undefined functions, yaaJ and yaaX.

The genomic context for yaaA is not conserved amongst bacteria that possess

this gene, and no obvious gene partners were found by computational analyses.

A computational search using an OxyR consensus sequence (39) identified one

potential OxyR-binding site (-74 bp to -38 bp), and 5-RACE analysis mapped the

5-end of yaaA transcripts to 22 bp upstream of the yaaA start codon. TSS,

transcriptional start site.

73  

Figure 2.2 A transcriptional yaaA-lacZ fusion confirms that yaaA is part of

the OxyR regulon. (A) Cultures were grown in anaerobic or aerobic LB medium

for five generations before collection and β-galactosidase measurement (~0.2

OD600). WT, MG1655; Vector, empty vector control (pACYC184); pGSO58,

plasmid-borne mutant OxyR A233V that constitutively overexpresses OxyR

regulated genes; ahp- (AL154), NADH peroxidase mutant; Hpx- (AL145),

catalase/peroxidase-deficient mutant. (B) Because Hpx- oxyR mutants are lethal

aerobically, anaerobic exponential-phase cultures were harvested at ~0.2

OD600nm (-O2) or were aerated for 30 min before collection (+O2). During the

period of aeration, the cell densities of the Hpx- oxyR+ (AL145) and Hpx- oxyR-

(AL344) strains increased from ~0.2 OD600nm to ~0.35 OD600nm. In this and other

figures, the error bars represent the standard error of the mean.

74  

Figure 2.3 The Hpx- yaaA strain exhibits a growth defect in aerobic LB

medium. Exponential-phase cultures in anaerobic LB medium were diluted into

aerobic LB medium at time zero, and optical density was monitored. (A) Wild

type cells (MG1655), black squares; yaaA (AL245), grey triangles; Hpx-

(catalase/peroxidase-deficient strain, LC106), grey diamonds; Hpx- yaaA (SB53),

black circles. (B) Hpx- yaaA cells with 2 mM DFO in media, open circles; Hpx-

tonB yaaA (AL179), stars.

75  

Figure 2.4 The Hpx- yaaA cells exhibit a severe filamentous phenotype.

Exponentially growing cultures in anaerobic LB medium were diluted into aerobic

LB medium and were grown for 5 hours. Images were captured by differential

interference contrast microscopy (DIC) using a 40X oil-immersion objective.

76  

Figure 2.5 The growth defect of the Hpx- yaaA mutant occurred

concomitantly with cell death. Exponentially growing cultures in anaerobic LB

medium were diluted into aerobic LB medium at time zero. (A) Optical density. (B)

Viable cell counts. Hpx- (LC106), grey diamonds; Hpx- yaaA (SB53), black circles.

77  

Figure 2.6 Aerobic mutation frequencies are elevated in the Hpx- yaaA

mutant compared to its Hpx- parent strain. Exponentially growing cells in

anaerobic LB medium were diluted into aerobic LB medium, and the mutation

frequencies were measured at subsequent time points as described in Materials

and Methods. The zero time point was obtained immediately prior to aeration.

The mutant frequencies in Hpx+ strains (barely visible above) were about 0.5 to 1

mutants/106 cells.

78  

Figure 2.7 Hpx+ recA yaaA mutants exhibited growth and viability defects.

Anaerobic exponential-phase cultures were diluted into aerobic LB medium at

time zero. (A, C, D) Optical density. The recA yaaA (AL223) mutants exhibited a

growth (A) and viability (B) defect that was suppressed by ~50 U/ml catalase (C)

or 2 mM DFO (D).

79  

Figure 2.8 Hpx+ polA1 yaaA mutant exhibited growth and viability defects.

Exponentially growing cultures in anaerobic LB medium were diluted into aerobic

LB medium at time zero. (A) Optical density. (B) Viable cell counts. yaaA (AL245),

grey triangles; polA1 (AL410), black squares; polA1 yaaA (AL412), open squares.

80  

Figure 2.9 Protein carbonylation in Hpx- yaaA cells. Cells were cultured in

aerobic LB medium as described in Materials and Methods. The Hpx- yaaA cells

were harvested at the beginning (0.2 OD600nm, 3 hours) and in the middle of the

growth lag (6 hours). Other strains were collected at ~0.2 OD600nm. Strains used:

WT (MG1655), Hpx- dps (SP66)--a positive control for carbonylation, Hpx-

(LC106), and Hpx- yaaA (SB53).

81  

Figure 2.10 The Hpx- yaaA mutant exhibited higher levels of hydroxyl

radicals. Cells were cultured in aerobic LB as described before. They were

harvested at 0.2 OD600nm, which corresponded to the beginning of the growth lag

of Hpx- yaaA cells. The final cell densities were equivalent in all samples. The six

peaks represented the HO signal of Hpx- cells and Hpx- yaaA cells.

82  

Figure 2.11 Levels of unincorporated iron are elevated in Hpx- yaaA cells.

The strains were cultured in aerobic LB medium, and intracellular unincorporated

iron was determined by whole-cell EPR analysis. Data are reported as the means

and SEM from at least 3 independent experiments. (A) Cells were harvested at

0.1, 0.2, and 0.3 OD600nm. (B) Cells were harvested at 0.2 OD600nm. pWKS30,

empty vector; pWKyaaA, a plasmid expressing yaaA under control of the lac

promoter.

83  

Figure 2.12 Overexpression of YaaA anaerobically protects cells against

H2O2 killing, but does not affect unincorporated iron levels. Cells were grown

in anaerobic LB with 0.2% lactose and 100 g/ml Amp to early log-phase, and

then were treated with 2.5 mM H2O2 (A) or were harvested for EPR to measure

intracellular unincorporated iron concentration (B).

84  

Figure 2.13 fecA-lacZ induction in Hpx- feo entA strains (yaaA+ vs yaaA-).

Cultures were grown in anaerobic LB medium for five generations and either

harvested at ~0.2 OD600nm (-O2) or diluted into aerobic LB medium and grown

aerobically for five more generations before collection at ~0.2 OD600nm (+O2). The

yaaA+ strain is Hpx- feo entA fecA-lacZ (AL273); the yaaA- strain is Hpx- feo entA

yaaA fecA-lacZ (AL293).

85  

Figure 2.14 YaaA has functions independent of Fur and Dps. Exponential-

phase cultures in anaerobic LB medium were diluted into aerobic LB medium to

an OD600 of 0.005 at time zero, and cell viability was monitored. (A) Viable cell

counts for Hpx- fur yaaA cells. (B) Viable cell counts for Hpx- dps yaaA cells. To

reduce lethality, the inoculum was at a higher density than in the experiment of

Figure. 2.7.5.

86  

Figure 2.15 Hpx- mntH yaaA cells exhibited growth defects in both LB and

defined glucose medium. Anaerobic exponential-phase cultures were diluted

into the same aerobic medium at time zero. Hpx- mntH yaaA had a growth defect

in both LB (A) and glucose defined medium with all amino acid supplement (B).

Adding MnCl2 (50 M) rescued Hpx- mntH cells, but had little effect on Hpx- mntH

yaaA mutants (C). (D) The growth defect in defined medium was bacteriostatic.

Strains used: Hpx- (LC106), grey diamonds; Hpx- mntH (AA30), grey triangles;

Hpx- yaaA (SB53), black circles; and Hpx- mntH yaaA (AL221), black squares.

87  

Figure 2.16 YaaA has functions independent of FieF. Exponential-phase

cultures in anaerobic LB medium were diluted into aerobic LB medium to an

OD600 of 0.005 at time zero, and optical density was monitored. Hpx- (LC106),

grey diamonds; Hpx- fieF (AL435), open diamonds; Hpx- yaaA (SB53), black

circles, Hpx- fieF yaaA (AL437), open circles. The growth curves of the Hpx- yaaA

and Hpx- fieF yaaA strains are almost indistinguishable.

88  

Figure 2.17 Expression of yaaA is induced in aerated ahp- mutant during

iron starved conditions. The tonB feo mutants in Hpx+, ahp- or Hpx-

background were grown aerobically in glucose defined medium with (iron

depleted, gray bar) or without (iron sufficient, open bar) 2 mM DFO as described

above. Cells were harvested to measure yaaA-lacZ transcriptional fusion activity.

Strains used: tonB feo (AL304), ahp- tonB feo (AL302), Hpx- tonB feo (AL300).

89  

Figure 2.18 Potential mechanisms by which YaaA might diminish levels of

unincorporated iron. YaaA might slow down the increase of unincorporated

iron by (1) slowing iron import, (2) preventing the leakage of iron from H2O2-

damaged enzymes; or YaaA might accelerate the decrease of unincorporated

iron by (3) sequestering iron, (4) speeding iron delivery to target proteins, or (5)

increasing iron efflux.

90  

2.8 References

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2. Anjem, A., S. Varghese, and J. A. Imlay. 2009. Manganese import is a key element of the OxyR response to hydrogen peroxide in Escherichia coli. Mol. Microbiol. 72:844-858

3. Aslund, F., M. Zheng, J. Beckwith, and G. Storz. 1999. Regulation of the OxyR transcription factor by hydrogen peroxide and the cellular thiol-disulfide status. Proc. Natl. Acad. Sci. U. S. A. 96:6161-6165.

4. Braun, V. 2003. Iron uptake by Escherichia coli. Front Biosci. 8:s1409-s1421.

5. Chang, A. C. and S. N. Cohen. 1978. Construction and characterization of amplifiable multicopy DNA cloning vehicles derived from the P15A cryptic miniplasmid. J. Bacteriol. 134:1141-1156.

6. Cherepanov, P. P. and W. Wackernagel. 1995. Gene disruption in Escherichia coli: TcR and KmR cassettes with the option of Flp-catalyzed excision of the antibiotic-resistance determinant. Gene. 158:9-14.

7. Datsenko, K. A. and B. L. Wanner. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. U. S. A. 97:6640-6645.

8. Finn, R. D., J. Tate, J. Mistry, P. C. Coggill, S. J. Sammut, H. R. Hotz, G. Ceric, K. Forslund, S. R. Eddy, E. L. Sonnhammer, and A. Bateman. 2008. The Pfam protein families database. Nucleic Acids Res. 36:D281-D288.

9. Grass, G., M. Otto, B. Fricke, C. J. Haney, C. Rensing, D. H. Nies, and D. Munkelt. 2005. FieF (YiiP) from Escherichia coli mediates decreased cellular accumulation of iron and relieves iron stress. Arch. Microbiol. 183:9-18.

10. Greenberg, J. T., P. Monach, J. H. Chou, P. D. Josephy, and B. Demple. 1990. Positive control of a global antioxidant defense regulon activated by superoxide-generating agents in Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 87:6181-6185.

11. Gu, M. and J. A. Imlay. 2011. The SoxRS response of Escherichia coli is directly activated by redox-cycling drugs rather than by superoxide. Mol. Microbiol. 79:1136-1150.

12. Gut, I. M., A. M. Prouty, J. D. Ballard, W. A. van der Donk, and S. R. Blanke. 2008. Inhibition of Bacillus anthracis spore outgrowth by nisin. Antimicrob. Agents Chemother. 52:4281-4288.

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13. Haldimann, A. and B. L. Wanner. 2001. Conditional-replication, integration, excision, and retrieval plasmid-host systems for gene structure-function studies of bacteria. J. Bacteriol. 183:6384-6393.

14. Imlay, J. A., S. M. Chin, and S. Linn. 1988. Toxic DNA damage by hydrogen peroxide through the Fenton reaction in vivo and in vitro. Science. 240:640-642.

15. Imlay, J. A. and I. Fridovich. 1991. Assay of metabolic superoxide production in Escherichia coli. J. Biol. Chem. 266:6957-6965.

16. Jang, S. and J. A. Imlay. 2010. Hydrogen peroxide inactivates the Escherichia coli Isc iron-sulphur assembly system, and OxyR induces the Suf system to compensate. Mol. Microbiol. 78:1448-1467.

17. Jang, S. and J. A. Imlay. 2007. Micromolar intracellular hydrogen peroxide disrupts metabolism by damaging iron-sulfur enzymes. J. Biol. Chem. 282:929-937.

18. Keyer, K. and J. A. Imlay. 1996. Superoxide accelerates DNA damage by elevating free-iron levels. Proc. Natl. Acad. Sci. U. S. A. 93:13635-13640.

19. Kullik, I., M. B. Toledano, L. A. Tartaglia, and G. Storz. 1995. Mutational analysis of the redox-sensitive transcriptional regulator OxyR: regions important for oxidation and transcriptional activation. J. Bacteriol. 177:1275-1284.

20. Macomber, L., C. Rensing, and J. A. Imlay. 2007. Intracellular copper does not catalyze the formation of oxidative DNA damage in Escherichia coli. J. Bacteriol. 189:1616-1626.

21. Masse, E., H. Salvail, G. Desnoyers, and M. Arguin. 2007. Small RNAs controlling iron metabolism. Curr. Opin. Microbiol. 10:140-145.

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23. Park, S. and J. A. Imlay. 2003. High levels of intracellular cysteine promote oxidative DNA damage by driving the fenton reaction. J. Bacteriol. 185:1942-1950.

24. Park, S., X. You, and J. A. Imlay. 2005. Substantial DNA damage from submicromolar intracellular hydrogen peroxide detected in Hpx- mutants of Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 102:9317-9322.

25. Patel, M., Q. Jiang, R. Woodgate, M. M. Cox, and M. F. Goodman. 2010. A new model for SOS-induced mutagenesis: how RecA protein activates DNA polymerase V. Crit Rev. Biochem. Mol. Biol. 45:171-184.

26. Posey, J. E. and F. C. Gherardini. 2000. Lack of a role for iron in the Lyme disease pathogen. Science. 288:1651-1653.

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27. Seaver, L. C. and J. A. Imlay. 2004. Are respiratory enzymes the primary sources of intracellular hydrogen peroxide? J. Biol. Chem. 279:48742-48750.

28. Seaver, L. C. and J. A. Imlay. 2001. Alkyl hydroperoxide reductase is the primary scavenger of endogenous hydrogen peroxide in Escherichia coli. J. Bacteriol. 183:7173-7181.

29. Seaver, L. C. and J. A. Imlay. 2001. Hydrogen peroxide fluxes and compartmentalization inside growing Escherichia coli. J. Bacteriol. 183:7182-7189.

30. Sobota, J. M. and J. A. Imlay. 2011. Iron enzyme ribulose-5-phosphate 3-epimerase in Escherichia coli is rapidly damaged by hydrogen peroxide but can be protected by manganese. Proc. Natl. Acad. Sci. U. S. A. 108:5402-5407.

31. Touati, D., M. Jacques, B. Tardat, L. Bouchard, and S. Despied. 1995. Lethal oxidative damage and mutagenesis are generated by iron in delta fur mutants of Escherichia coli: protective role of superoxide dismutase. J. Bacteriol. 177:2305-2314.

32. Tsaneva, I. R. and B. Weiss. 1990. soxR, a locus governing a superoxide response regulon in Escherichia coli K-12. J. Bacteriol. 172:4197-4205.

33. Varghese, S., A. Wu, S. Park, K. R. Imlay, and J. A. Imlay. 2007. Submicromolar hydrogen peroxide disrupts the ability of Fur protein to control free-iron levels in Escherichia coli. Mol. Microbiol. 64:822-830.

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36. Woodmansee, A. N. and J. A. Imlay. 2002. Quantitation of intracellular free iron by electron paramagnetic resonance spectroscopy. Methods Enzymol. 349:3-9.

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38. Zhao, G., P. Ceci, A. Ilari, L. Giangiacomo, T. M. Laue, E. Chiancone, and N. D. Chasteen. 2002. Iron and hydrogen peroxide detoxification properties of DNA-binding protein from starved cells. A ferritin-like DNA-binding protein of Escherichia coli. J. Biol. Chem. 277:27689-27696.

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40. Zheng, M., X. Wang, L. J. Templeton, D. R. Smulski, R. A. LaRossa, and G. Storz. 2001. DNA microarray-mediated transcriptional profiling of the Escherichia coli response to hydrogen peroxide. J. Bacteriol. 183:4562-4570.

 

 

 

 

 

 

 

 

   

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CHAPTER THREE: CELL DEATH FROM ANTIBIOTICS WITHOUT

THE INVOLVEMENT OF REACTIVE OXYGEN SPECIES

3.1 Introduction

In last couple of decades the growing number of antibiotic-resistant pathogens

has urged efforts to further understand and improve the efficacy of the basic

antibiotic classes. The majority of antibiotics with clinical utility achieve their

bactericidal effects by inhibiting bacterial cell wall biosynthesis, protein translation

or DNA replication. However, within the past six years, observations from several

labs raised the possibility that these antibiotics, irrespective of drug targets, kill

bacteria by stimulating the formation of reactive oxygen species that

subsequently cause to DNA lesions and cell death.

The evidence supporting this proposal included the observation that cell-

penetrating fluorescent probes (hydroxyphenyl fluorescein, HPF) gave higher

fluorescent signals inside antibiotic-treated bacteria (5,8,9,19,20,36), and the

signals were ascribed to hydroxyl radicals. Furthermore, iron chelators

(8,9,19,32,33,36), which suppress hydroxyl-radical-generating Fenton chemistry,

and thiourea (5,9,11,19,32,33), a potential scavenger of hydroxyl radicals,

lessened toxicity. Mutations that diminish fluxes through the tricarboxylic acid

cycle (TCA cycle) were protective (8,19,20), suggesting a key role for respiration,

and DNA-repair mutants were somewhat sensitive (9,19). Systems analysis of

aminoglycoside-treated Escherichia coli by microarray suggested a model that

fits these data (20). It was postulated that interference with ribosome progression

95  

would release incomplete polypeptides, some of which are translocated to the

cell membranes where they might trigger envelope stress. The Arc regulatory

system is perturbed, potentially accelerating respiration and thereby increasing

the flux of O2- and H2O2 into the cell interior. These two oxidants have known

sequelae that ultimately lead to DNA damage. Specifically, O2- and H2O2 damage

the Fe-S clusters of aconitase-class dehydratases (17,21), releasing iron atoms

and elevating the pool of intracellular unincorporated iron (18,22). This iron can

then react with H2O2 in the Fenton reaction, generating hydroxyl radicals that

either directly damage DNA (14) or indirectly oxidize the deoxynucleotide pool,

which is subsequently incorporated into DNA (35). This scenario could explain

the observed oxidation of intracellular dyes, protection by scavengers and

chelators, a requirement for respiration, and the sensitivity of DNA-repair mutants.

This model is plausible, and so we devised experiments to directly test the

molecular events that underpin it. The bacterial strain (Escherichia coli MG1655),

growth medium (LB), and antibiotic doses were chosen to match those of

previous studies (19). Kanamycin was used to target translation, ampicillin to

block cell-wall synthesis, and norfloxacin to disrupt DNA replication.

3.2 Materials and Methods

3.2.1 Reagents

-Mercaptoethanol, bovine liver catalase, diethylenetriaminepentaacetic acid

(DTPA), deferoxamine mesylate (DFO), dipyridyl, ferric chloride, type II

horseradish peroxidase, 30% hydrogen peroxide, -nitrophenyl -D-

96  

galactopyranoside (ONPG), K3Fe(CN)3, thiourea, dithionite, 6-phosphogluconate,

lactate dehydrogenase, NADH, cysteine, Fe(NH4)2(SO4)26H2O, dithiothreitol

(DTT), L-malic acid disodium salt, citraconic acid, potassium D-gluconate,

ampicillin, kanamycin, and norfloxacin were purchased from Sigma. KNO3 and D-

glucose were obtained from Fisher ChemAlert Guide. Albumin standard and

Coomassie protein assay reagent were purchased from Thermo Scientific. 3-(p-

hydroxyphenyl) fluorescein (HPF), Amplex Red and SuperScript III Reverse

Transcriptase were purchased from Invitrogen. Standard reagents for RNA work

were obtained from Ambion. DNase I digestion is from Biolabs. iQTM SYBR

Green Supermix was from BioRad, and real-time PCR plates were purchased

from Eppendorf.

3.2.2 Strain construction

The strains used in this study are listed in Table 3.1; they are all congenic

derivatives of MG1655. Deletions were constructed by the Red recombination

deletion method (4), were transferred to other strains by P1 transduction, and

were confirmed by PCR and/or phenotype analyses (25). The antibiotic-

resistance cassettes were subsequently removed by FLP-mediated excision (4).

Gene fusions between the katG upstream region (-160 bp to +16 bp) and lacZ

were created by standard methods and integrated into the lambda phage

attachment site (12), preserving the wild-type katG gene at its native

chromosomal locus.

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3.2.3 Bacterial growth

All cultures were grown in LB (10 g tryptone, 5 g yeast extract, and 10 g NaCl per

liter) except indicated. Experimental protocols were designed to ensure that

measurements were performed upon exponentially growing cells. Overnight

cultures were diluted into fresh media and grown for 5 generations to early log

phase (OD600nm of 0.10 to 0.20) before treatment with antibiotics. All cultures

were grown at 37 C in the dark. Aerobic LB medium was made one day

previously and stored in the dark to avoid the photochemical generation of H2O2.

Anaerobic medium was transferred immediately after autoclaving to an anaerobic

chamber (Coy Laboratory Products), where it was stored under an atmosphere of

5% CO2/10% H2/85% N2 for at least one day prior to use. Where indicated, 40

mM KNO3, 0.2% gluconate, or 0.2% glucose was included in the medium. When

used, dipyridyl was made fresh and added into the media at a final concentration

of 500 M before inoculation.

Defined glucose/amino acids medium contained minimal A salts (25), 0.2%

glucose, 0.2% Casamino Acids, 0.5 mM tryptophan, 0.02% MgSO47H2O, and 5

g/ml thiamine.

Most experiments included in this report employed fixed concentrations of

antibiotics (5 g/ml ampicillin, 250 ng/ml norfloxacin or 5 g/ml kanamycin)

designed to match doses used by other workers (19). However, a higher dose of

kanamycin (30 g/ml) was used in testing the effects of nitrate upon killing so that

the impact of nitrate was clear. In the Edd assay, a higher dose of kanamycin (20

98  

g/ml) was used because supplementation with 0.2% gluconate decreased

sensitivity of the cells to kanamycin. The fur mutant was also more resistant to

kanamycin, and therefore 10 g/ml kanamycin was used to show the killing.

3.2.4 Cell viability

To determine cell viability, cultures were grown to 0.1-0.2 OD600nm (aerobic

cultures) or 0.05 OD600nm (anaerobic cultures) prior to the addition of antibiotics.

The lower density of anaerobic cultures ensured that they did not approach

stationary phase during antibiotic treatment. Cell samples were collected at

designated time points and washed twice with PBS (50 mM potassium

phosphate with 0.9% NaCl, pH 7.2) and then serially diluted, mixed with LB top

agar, and spread onto the surface of LB plates. Colonies were allowed to grow

aerobically (or anaerobically for anaerobic killing) for 24 h and plates that had

30~300 colonies were selected for counting. At least three replicates of the

survival-curve experiments were performed. Day-to-day variations in medium

composition shifted the curves slightly, but the sample-to-sample comparisons

cited in the text were invariable. Therefore, representative survival curves are

presented.

3.2.5 O2 consumption

Cells were cultured in LB and treated with antibiotics as described above. At

designated time points samples were collected and diluted in air-saturated LB to

0.05~0.15 OD600nm, and respiration was measured with a Digital Model 10 Clark-

type oxygen electrode (Rank Brothers Ltd) at 37 C. The machine was calibrated

with air-saturated LB medium and sodium dithionite.

99  

3.2.6 Real-time PCR

Cells were cultured in LB as described above. For the H2O2 control, aliquots of

cultures were treated with 250 M H2O2 at room temperature for 10 min. RNA

was harvested by the hot phenol method and cleaned up using the RNeasy mini

Kit (Qiagen). The DNA-free RNA samples were generated by DNase I digestion

and converted to cDNA by SuperScript III Reverse Transcriptase. The following

primers were used to analyze expression of ahpC, katG, yaaA, and the

housekeeping gene gapA: ahpC forward, 5-GACCCGTAACTTCGACAACA-3;

ahpC reverse, 5-ATGCCTTCAGCGGTAACTTC-3; katG forward, 5-

TACTGGGTGCCAACTTCGAT-3; katG reverse, 5-

GGTCGCTTTCCACTCGTAAC-3; yaaA forward, 5-

GCTGTTAGACAATTCCCAGCAG-3; yaaA reverse, 5-

AAGCCGGTGTAGACATCACCTT-3; gapA forward, 5-

CGTTCTGGGCTACACCGAAGATGACG-3; gapA reverse, 5-

AACCGGTTTCGTTGTCGTACCAGGA-3. The 25 l real-time PCR mixture

contained 50 ng of cDNA, 12.5 l of SYBR Green Supermix, and 400 nM of

ahpC, gapA or yaaA primers, or 1 M of katG primers. Thermal cycling was

performed using a Mastercycler ep realplex (Eppendorf) for 40 cycles in 3 steps:

95 C for 20 s, 58 C for 20 s and 72 C for 20 s. Amplification efficiencies were

determined for each primer set (0.9 to 1.1) and were used in calculating the fold

change. The melting curves were checked for all samples at the end.

100  

3.2.7 -Galactosidase activity

Samples were collected at designated time points, and cells were centrifuged,

washed and resuspended in ice-cold 50 mM TrisHCl (pH 8.0), followed by lysis

with a French pressure cell (SIM Aminco FA-003). Cell debris was removed by

centrifugation, and -galactosidase activity in the supernatant was measured by

standard procedures (25). Protein concentrations were determined by the

Bradford assay using bovine serum albumin as the standard.

3.2.8 H2O2 production measurement

The rate of H2O2 formation by antibiotic-treated cells was measured with Hpx-

(catalase/peroxidase-null) mutants as described (27). Anaerobic overnight

cultures of Hpx- cells were diluted and grown at least five generations in aerobic

LB broth to 0.1~0.2 OD600nm. They were then treated with antibiotics as described

above. At time points the cells were centrifuged, suspended in fresh

tryptone/NaCl (10 g tryptone and 10 g NaCl per liter) at an OD600nm of 0.05~0.1,

and incubated aerobically at 37 °C with vigorous aeration. At intervals, cells were

removed by centrifugation, and H2O2 concentrations were measured in the

supernatants using the Amplex Red/horseradish peroxidase assay (27) in a

Shimadzu RF Mini-150 fluorometer.

3.2.9 Electron paramagnetic resonance (EPR) measurement of intracellular

unincorporated iron

The pool of intracellular chelatable iron was quantified by established EPR

methods (34). The cells were cultured aerobically in 1 L of LB as described

above. At time points during antibiotic treatment, cells were centrifuged and

resuspended at 100-fold higher cell density in LB medium containing 10 mM

101  

DTPA (pH 7.0) to block further iron import, and 20 mM DFO (pH 8.0) to facilitate

oxidation of intracellular unincorporated ferrous iron to EPR-detectable ferric iron.

The suspension was incubated at 37 C aerobically with shaking for 15 min. The

cell pellet was washed once with 5 ml of ice-cold 20 mM TrisHCl-10% glycerol

(pH 7.4) and finally re-suspended in 200 l of the same buffer before freezing.

The EPR analysis was performed as described previously (34). Different

concentrations of FeCl3 in 20 mM TrisHCl-10% glycerol-1 mM DFO (pH 7.4)

were used to make an EPR signal standard curve. The measured EPR signals

were normalized to cell density and converted to intracellular concentrations

using the following conversion: 1 ml of E. coli culture at OD600nm of 1 comprises

0.52 l of intracellular volume (15).

3.2.10 Enzyme assays

6-phosphogluconate dehydratase (Edd) assay

Cells were cultured in LB with 0.2% gluconate as described above. Overnight

cultures of SOD-deficient mutants were grown in anaerobic media to avoid the

selection of suppressor mutations; log-phase cells were then diluted and grown

in aerobic media for 3 hours (to 0.2 OD600) prior to harvesting. Samples were

collected at designated time points, washed with ice-cold 50 mM Tris-HCl (pH

7.65) buffer, and frozen at -80 C. Before the assay, the cell pellets were

resuspended in anaerobic ice-cold 50 mM Tris-HCl (pH 7.65) buffer, and cell

extracts were prepared by anaerobic sonication. The activity of Edd was

determined by incubating the cell lysate with 6-phosphogluconate for 2, 4, 6 and

8 min, then the amount of pyruvate produced was determined by monitoring

102  

NADH consumption catalyzed by lactate dehydrogenase (16). In order to repair

the damaged Fe-S clusters, cell lysates were incubated with 0.5 mM

Fe(NH4)2(SO4)2 and 5 mM DTT at room temperature anaerobically for 45 min

before measuring the activity. Typically a minor fraction of the enzyme population

has [3Fe-4S] clusters that can be reactivated; this becomes the great majority

during oxidative stress (10). The addition of IscS protein and cysteine to the

repair reaction (16) did not further increase the activity, indicating that no

apoprotein was present.

Isopropylmalate isomerase (IPMI) assay and fumarase assay

Cells were cultured in minimal A defined medium with 0.2% glucose, and aerated

log-phase cells (0.1 to 0.2 OD600) were treated with 20 g/ml kanamycin, and

samples were collected at designated time points.

For the IPMI samples, the growth medium was supplemented with 0.5 mM each

of histidine, phenylalanine, tyrosine and tryptophan. Harvested samples were

washed with ice-cold 100 mM Tris-Cl (pH 7.6) buffer, and frozen at -80 C.

Before the assay, the cell pellets were resuspended in anaerobic ice-cold 100

mM Tris-Cl buffer (pH 7.6) with 2 mM EDTA, and cell extracts were prepared by

anaerobic sonication. The activity of IPMI was monitored at A235 by incubating

the cell lysate with 0.4 mM citraconate (17).

For fumarase assay, the growth medium was supplemented with 0.2% casamino

acides and 0.5 mM tryptophan. After harvest, samples were washed with ice-

cold 50 mM sodium phosphate buffer (pH 7.3), and frozen at -80 C. Before the

103  

assay, the cell pellets were resuspended in anaerobic ice-cold sodium phosphate

buffer, and cell extracts were prepared by anaerobic sonication. The activity of

fumarase was monitored at A250nm by incubating the cell lysate with 50 mM

malate (17).

The damaged Fe-S clusters of IPMI and fumarase were repaired by incubation

with ferrous iron and DTT as described above.

3.2.11 H2O2 killing assay

At designed time points, cell samples were diluted to an OD600nm of 0.05 in fresh

LB and treated with 2.5 mM H2O2 at 37 °C. At intervals, aliquots were harvested

and immediately mixed with catalase (final concentration: 2500 U/ml) in order to

terminate H2O2 stress. Cells were serially diluted, mixed with top agar, and then

spread on LB plates. CFUs were determined after aerobic incubation at 37 °C for

24 h.

3.2.12 Hydroxyphenyl fluorescein (HPF) assay

One millimolar H2O2 was added into 0.1 M sodium phosphate buffer, pH 7.4,

containing 100 M Fe(NH4)2(SO4)2 and 10 M 3-(p-hydroxyphenyl) fluorescein

(HPF) (28). The resultant fluorescent signal was measured in a Shimadzu RF

Mini-150 fluorometer using 490/520 nm as excitation/emission wavelengths. To

scavenge the oxidant generated by the Fenton reaction, thiourea or ethanol was

added to the mixture before H2O2 was added. In other experiments, different

amounts of K3Fe(CN)3 were mixed with 10 M HPF in 0.1 M sodium phosphate

buffer, and the fluorescent signal was recorded after 20 min incubation. 100 M

H2O2 and 0.2 g/ml horseradish peroxidase (HRP) were mixed with 10 M HPF,

104  

or 100 M H2O2 and 20 g/ml horseradish peroxidase (HRP) were mixed with 2

M HPF, and the fluorescent signal was monitored over time.

3.3 Results

3.3.1 Antibiotic killing does not require oxygen

Superoxide and hydrogen peroxide are generated inside cells when

flavoenzymes inadvertently transfer a fraction of their electron flux directly to

molecular oxygen (14,27). Thus neither of these reactive oxygen species can be

formed under anoxic conditions. We found, however, that ampicillin and

norfloxacin were as lethal to cells in an anaerobic chamber as to cells in air-

saturated medium (Figure 3.1A-B). Anoxia sharply diminished the toxicity of

kanamycin, but lethal activity was partially restored at high kanamycin doses

when nitrate was provided as an alternative respiratory substrate (Figure 3.1C).

This pattern mirrors the effects of oxygen and nitrate upon kanamycin import,

which is governed by the magnitude of the proton motive force (30,31). Thus

reactive oxygen species were not required for the lethal actions of kanamycin,

and they made no apparent contribution at all to the toxicity of ampicillin and

norfloxacin.

3.3.2 H2O2-stressed mutants have similar sensitivity to ampicillin or

kanamycin as wild type cells

Hydroxyl radicals are formed inside cells when ferrous iron transfers electrons to

H2O2, so hydroxyl-radical stress is greatest when either H2O2 or ferrous

concentrations are high. An E. coli mutant that lacks catalase and peroxidase

activities (denoted Hpx-) cannot scavenge H2O2 and more easily accumulates it

105  

to toxic levels; resulting in substantial damage to proteins and DNA (2,17,26,29).

However, this mutant only showed increased sensitivity to norfloxacin, but not to

ampicillin or kanamycin (Figure 3.2). The elevated sensitivity to norfloxacin could

be due to antibiotic-mediated oxidative stress; yet it is also possible that with the

inhibitory effect of norfloxacin upon DNA metabolism merely exacerbates the pre-

existing DNA damage in the Hpx- mutants (7,26). To further test these

possibilities, we first sought to evaluate the levels of H2O2 production after

antibiotic incubation.

3.3.3 Antibiotic treatments did not accelerate the formation of hydrogen

peroxide

We directly tested the prediction that antibiotic treatment would augment

respiration and thereby increase O2- and H2O2 formation. The respiratory chain is

normally a minor source (< 20%) of these oxidants (27), so a very large flux

increase would be needed to substantially amplify total intracellular oxidant

production. However, measurements of oxygen consumption with a Clark-type

electrode showed that respiration slowly declined, rather than increased, when

cells were treated with kanamycin (Figure 3.3). This trend persisted throughout

the period during which the number of viable cells declined by > 99% (Figure

3.2B). Norfloxacin had little effect on respiration. Ampicillin transiently increased

respiration, perhaps by causing envelope damage that dissipated the

backpressure of the proton motive force, before cell lysis ended metabolism. In

no case did respiration accelerate enough to be a sufficient explanation for

toxicity.

106  

E. coli has a transcription factor, OxyR, that is activated by H2O2 whenever its

levels approach the concentrations (0.5-1 M) sufficient to damage enzymes or

DNA (3). The OxyR regulon includes genes whose products assist in H2O2

scavenging (katG, ahpCF), free-iron control (dps, fur, yaaA), and enzyme

protection (mntH, sufABCDE) (14,23,37). When cells were treated with ampicillin

and norfloxacin, they accumulated lethal damage within one hour (Figure 3.1A-B);

however, real-time PCR analysis indicated that OxyR-controlled genes-katG,

ahpC, and yaaA-were not induced (Figure 3.4A). These antibiotic-treated cells

robustly activated katG, ahpC, and yaaA when exogenous H2O2 was added as a

control (Figure 3.4A). After one hour, kanamycin-treated cells failed to respond

even to authentic H2O2, perhaps because OxyR protein was depleted in these

translationally blocked cells (data not shown). This result was confirmed by

measurements of katG::lacZ expression (Figure 3.4B-C). Thus, lethal doses of

neither ampicillin nor norfloxacin create enough H2O2 stress to trigger the natural

H2O2 sensor within the cell.

The rate of intracellular H2O2 formation can be directly measured using

catalase/peroxidase-deficient mutants. Endogenous H2O2 rapidly diffuses from

these cells out into the growth medium, and the rate of its accumulation in the

medium can be quantified by direct assay (Figure 3.5) (27). When cells were

treated with lethal concentrations of kanamycin and ampicillin, H2O2 production

did not increase (Figure 3.6A-B). In norfloxacin-treated cells H2O2 formation

increased slightly (< 2-fold) after one hour but not at all within the first 30 min

when > 99% of cells accumulated lethal damage (Figure 3.6C and Figure 3.2C).

107  

Therefore, the moderately increased of H2O2 generation in later time points was

not the primary bactericidal mechanism for norfloxacin. In contrast, when cells

were treated with a non-lethal dose of the redox-cycling drug, paraquat, H2O2

production was stimulated ~ 7-fold (Figure 3.6D).

Taken together, we conclude that these classic bactericidal antibiotics did not

promote H2O2 formation.

3.3.4 Antibiotic treatments did not increase the pool of unincorporated iron

There is one other way that hydroxyl-radical formation might be accelerated: by

increases in the amount of unincorporated iron. Iron can accumulate inside cells

under conditions of superoxide stress, due to the destruction of labile enzymic

Fe-S clusters (18,22), and this mechanism was postulated to occur during

antibiotic exposure (8,19). We monitored the status of 6-phosphogluconate

dehydratase, an aconitase-class Fe-S cluster containing enzyme that quickly

loses activity in superoxide-stressed cells (10). We observed that the overall

enzyme activity diminished during hours of antibiotic treatment, but the fraction of

enzymes with incomplete clusters did not significantly rise (Figure 3.7A-B). In

contrast, in control mutants that lack superoxide dismutase, the entire enzyme

population was inactive due to cluster damage (Figure 3.7C). Similarly, lethal

doses of kanamycin did not damage the Fe-S clusters in fumarase and isopropyl

isomerase (Figure 3.8), another two aconitase-class enzymes that are known to

be sensitive to oxidative stress (17).

Levels of intracellular unincorporated iron were directly imaged by whole-cell

electron paramagnetic resonance (EPR) spectroscopy (34). These levels did not

108  

rise in kanamycin- or norfloxacin-treated cells (Figure 3.9A-B). (Levels could not

be measured after ampicillin treatment, because lysing cells could not be

concentrated for the measurement.) In contrast, free-iron levels were increased

4-fold in fur regulatory mutants that over-synthesize iron import systems (Figure

3.9C). Notably, while fur mutants were slightly more sensitive to ampicillin, they

were actually more resistant to kanamycin and norfloxacin (Figure 3.10). This

result, also observed by others (8), indicates that intracellular free-iron levels do

not correlate with antibiotic sensitivity.

3.3.5 General DNA repair pathways had little effect upon antibiotic

sensitivity

DNA is substantially damaged when hydroxyl radicals are rapidly formed inside

cells, and mutants that are deficient in the excision or recombinational repair of

oxidative DNA lesions are particularly vulnerable (14). The extreme sensitivity of

recA and polA mutants to exogenous H2O2 is illustrated in Figure 3.11A.

However, recA strains exhibited little or no hypersensitivity to killing doses of

kanamycin or ampicillin, suggesting that these antibiotics have little effect upon

the rate of DNA damage (Figure 3.11C-D). The recA mutants were

hypersensitive to norfloxacin (Figure 3.11E), as expected, since this antibiotic

directly introduces DNA strand breaks by interrupting the catalytic cycle of

topoisomerases. The polA mutants were actually more resistant to kanamycin

and ampicillin than were wild-type strains (Figure 3.11B), a phenomenon that

likely reflects the slower growth rate of this mutant strain. In sum, these data

indicate that, unlike H2O2 stress, these antibiotics do not introduce lethal damage

into the DNA of cells.

109  

Other workers noted that lower doses (1-2 g/ml) of ampicillin slowly kill recA

mutants while being merely bacteriostatic to wild-type strains (9,19). We

reproduced that result (Figure 3.12A). However, this behavior also occurred

under anoxic conditions, when reactive oxygen species are non-existent (Figure

3.12B). The slight sensitivity of recA mutants may reflect defects in septation

from their innate replication problems. At higher doses of ampicillin, i.e., at doses

sufficient to kill wild-type bacteria, recombination proficiency had little or no

impact upon death rate (Figure 3.11C and 3.12A).

Collectively, these data indicate that the known mechanisms of oxidative stress

by reactive oxygen species were not involved in causing the death of antibiotic-

treated cells. Oxygen was dispensable for toxicity, and neither OxyR activity nor

direct measurements of H2O2 formation indicated that reactive oxygen species

were formed at accelerated rates. Further, no standard marker of oxidative

damage—enzyme damage, increases in free-iron levels, or hypersensitivity of

DNA-repair mutants—was detected.

3.4 Discussion

Fluorescein-based dyes such as HPF shows higher fluorescent signal within

antibiotic-treated cells than in control cells, and this fact has been regarded as

evidence that hydroxyl radicals are present (28). Indeed, thiourea, a presumptive

scavenger of hydroxyl radicals, diminished the antibiotic-augmented fluorescence

(5,19). However, we observed that although thiourea efficiently blocked dye

oxidation in a simple in vitro Fenton system, ethanol—another hydroxyl-radical

scavenger of similar efficiency (1)—failed to do so, even at far higher doses

110  

(Figure 3.13A-B). This result implies that the dye was actually oxidized by the

high-valence iron (FeIV=O)2+ initially formed by the Fenton reaction, rather than

by the hydroxyl radical into which it decomposes; thiourea, a sulfur compound,

may have re-reduced the metal before it oxidized the dye. Indeed, the dye was

very rapidly oxidized by horseradish peroxidase (Figure 3.13C), which has an

analogous high-valence metal intermediate but does not release hydroxyl

radicals (13). The dye was even oxidized by ferricyanide, a ferric-iron chelate of

moderate oxidizing potential (+ 0.44 V) (Figure 3.13E). Finally, the fluorescence

of the oxidized dye product was itself quenched after the fact by addition of the

same doses of thiourea that were used in antibiotic studies (Figure 3.13D); this

effect cannot be due to hydroxyl-radical scavenging activity. Thus it is plausible

that fluorescein-based dyes are predominantly oxidized in vivo by oxidized

enzymic metal centers, which may be more prominent in antibiotic-treated cells

as metabolism fails. In any case, one must be cautious in interpreting the

oxidation of these dyes inside cells.

Finally, cell protection by either thiourea or iron chelators cannot be regarded as

diagnostic of lethal Fenton chemistry. Both agents suppressed antibiotic

sensitivity even under anaerobic conditions (Figure 3.14), when death occurs

through pathways that do not involve reactive oxygen species. We conjecture

that they did so by reducing growth and/or metabolic rates. In general, it is

unlikely that exogenous agents like thiourea could substantially diminish the

lifetimes of hydroxyl radicals in vivo, since these radicals already react at nearly

diffusion-limited rates with intracellular biomolecules that are collectively at molar

111  

concentrations (6). Iron chelators have pleiotropic effects, including the

repression of TCA-cycle and respiratory enzymes (24), and may thereby diminish

the membrane potential that drives aminoglycoside uptake. Synthesis of these

enzymes is also repressed when cells are fed glucose, and this treatment

analogously diminished kanamycin sensitivity (Figure 3.15). Thus it is advisable

to follow up experiments that rely upon these agents with ones that employ more-

direct markers of oxidative stress.

We conclude that under our experimental conditions these major classes of

antibiotics did not exert their lethal actions through the known mechanisms of

oxidative stress. Of course, this does not preclude the possibility that antibiotics

trigger novel mechanisms of stress that do not involve the reactive oxygen

species that were tested here.

3.5 Acknowledgments

We thank Dan Hassett (University of Cincinnati) for helpful conversations about

this topic, and Jim Collins, Graham Walker, and members of their laboratories for

generously sharing ideas, unpublished data, and strains. We also thank Mark

Nilges of the Illinois EPR Research Center for assistance. This work was

supported by GM49060 from the National Institutes of Health.

112  

3.6 Tables

Table 3.1 Strains used in this study.

Name Genotype/characteristics

AA16 As MG1655 plus gltA::cat~Tn10

AA98 As MG1655 plus icd1::cat

AL410 As MG1655 plus polA1~zih-102::Tn10

AL427 As MG1655 plus (ahpCF::cat)1 (katG17::Tn10)1 (katE12::Tn10)1

AL441 As MG1655 plus (lacZ1::cat)1 att::[pSJ501:: katG-lacZ+ cat+]

AL486 As MG1655 plus (recA774::kan)1

AL494 As MG1655 plus (ahpC-ahpF’) kan:: ahpF (katG17::Tn10)1 (katE12::Tn10)1 (lacZ1::cat)1

att::[pSJ501:: katG-lacZ+ cat+]

JEM913 As MG1655 plus (lacZ1::cat)1 (fur-731::kan)1

KI232 As MG1655 plus (sodB-kan)1-2 (sodA::cat)1

LEM17 As MG1655 plus recA56 srl300::Tn10

MG1655 Wild type

113  

3.7 Figures

Figure 3.1 Antibiotic efficacy does not require oxygen. (A-C) Wild type cells

were treated with ampicillin (A), norfloxacin (B) or kanamycin (C) in the presence

or absence of oxygen. In panel C anoxic killing was also tested in the presence

of 40 mM KNO3 (gray squares).

114  

Figure 3.2 Antibiotic efficacy does not require H2O2. Wild-type cells (solid

squares, MG1655) and congenic Hpx- cells (open diamonds, AL427) were

treated with antibiotics. (A-C) Cells were treated with one dose of antibiotics, and

viability was monitored overtime. (D-F) Cells were treated with different doses of

antibiotics for designated time period, and viable cells were determined.

115  

Figure 3.3 O2 consumption rates were not substantially elevated by

antibiotic treatment. Oxygen consumption of wild type cells (MG1655) was

measured by a Clark-type electrode before and during exposure to 5 g/ml

kanamycin, 5 g/ml ampicillin, or 250 ng/ml norfloxacin. Respiration rates were

normalized to optical density.

116  

Figure 3.4 The OxyR regulon was not induced by ampicillin (5 g/ml) or

norfloxacin (250 ng/ml). (A) mRNA was collected from naïve and antibiotic-

exposed cells (WT, MG1655), and the expression of OxyR-responsive genes

was quantified by RT-PCR. Signals were normalized to that of the housekeeping

gene gapA, and fold inductions were calculated. Where indicated, cultures were

additionally treated with exogenous H2O2 at room temperature for 10 min prior to

mRNA collection. (B-C) Expression of a katG::lacZ fusion was measured in

exponential-phase wild-type cells (AL441) before and after antibiotic incubations

(B). As a positive control, expression was measured in wild-type and Hpx- cells

(AL494) before (-O2) and after 1 hour aeration (+O2) (C).

117  

Figure 3.5 Sample H2O2 production time course. Naïve and 1 h antibiotic-

treated Hpx- cells (AL427) were collected for H2O2 measurement. The redox-

cycling compound paraquat was included as a positive control. The suspension

of untreated Hpx- cells at 0.05 OD600nm excreted 16 nM H2O2/min into growth

medium (~10 M H2O2/s if normalized to cell volume).

118  

Figure 3.6 Classic antibiotics did not promote H2O2 formation. Aerobic Hpx-

cells (AL427) were treated with antibiotic, and at designated time points samples

were collected and the ongoing rate of H2O2 production was measured. The

redox-cycling compound paraquat (D) was included as a positive control; note

that this dose was insufficient to cause cell death (viable counts increased for > 3

hours). Data in this figure are reported as the means and SEM from at least 3

independent experiments.

119  

Figure 3.7 The superoxide-sensitive enzyme Edd was not damaged by cell

exposure to kanamycin or norfloxacin. Cells growing in aerobic LB with 0.2%

gluconate were incubated with lethal doses of kanamycin (20 g/ml, A) or

norfloxacin (250 ng/ml, B), and Edd activity was measured before (open bar) and

after (closed bar) in vitro cluster repair with ferrous iron/DTT. Panel C represents

control data from an SOD-deficient strain (KI232). The star denotes < 4% normal

activity (the sensitivity of the assay).

120  

Figure 3.8 The peroxide-sensitive enzyme fumarase and IPMI were not

damaged by cell exposure to kanamycin. (A) Wild type cells (MG1655) were

grown in aerobic glucose defined medium with aromatic amino acids to early log

phase, and then were incubated with lethal doses of kanamycin (B), and IPMI

activity was measured before (open bar) and after (closed bar) in vitro cluster

repair with ferrous iron/DTT. (C) Early log phase fumC (AL482) cells growing in

aerobic glucose defined medium with case amino acid were incubated with lethal

doses of kanamycin (D); and fumarase activity was measured before (open bar)

and after (closed bar) in vitro cluster repair with ferrous iron/DTT.

121  

Figure 3.9 Norfloxacin and kanamycin did not increase intracellular iron

levels. The levels of intracellular unincorporated iron were determined by whole-

cell EPR analysis before and during antibiotic exposure. Gray bars represent iron

levels in untreated cells at similar optical densities. (C) Levels of unincorporated

iron were measured in OD600nm of 0.15 untreated congenic wild type cells

(MG1655) and fur mutants (Jem913).

122  

Figure 3.10 fur mutants were invariably more resistant to kanamycin and

norfloxacin. fur mutants were treated with ampicillin (A), kanamycin (B) or

norfloxacin (C).

123  

Figure 3.11 DNA-repair mutants are not hypersensitive to antibiotic doses

that kill wild-type cells. (A) recA mutants and polA1 mutants are very sensitive

to H2O2 challenge. (B) Aerobic log-phase wild type cells and polA1 mutants were

treated with 100 ng/ml norfloxacin for 1 hour or with 5 g/ml ampicillin or 5 g/ml

kanamycin for 2 hours. The error bars represent the standard error of the mean

(SEM). (C-E) Wild-type cells (solid squares) and recA mutants (open circles)

were treated with the indicated doses of ampicillin (C) or kanamycin (D) for 2

hours, or norfloxacin (E) for one hour. Data in this figure are reported as the

means and SEM from at least 3 independent experiments. WT, MG1655; recA,

AL486; recA56, Lem17; polA1, AL410.

124  

Figure 3.12 A recA mutant is slightly sensitive to doses of ampicillin that

are bacteriostatic to wild-type cells, under both aerobic and anaerobic

conditions. Aerobic (A) or anaerobic (B) exponential-phase wild type cells and

recA mutants were treated with 1, 1.5 and 2 g/ml Amp, and survival was

monitored over time. Data are representative of at least three independent

experiments.

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Figure 3.13 HPF can be oxidized by oxidants other than hydroxyl radical.

(A) HPF dye was oxidized in vitro by a Fenton system consisting of 100 M

ferrous iron plus 1 mM H2O2. (B) Thiourea inhibited the oxidation of the dye, but

ethanol did not. (C) Horseradish peroxidase (HRP) plus H2O2 rapidly oxidized

HPF (10 M HPF, 100 M H2O2, and 0.2 g/ml HPR). (D) The HRP-oxidized

fluorescence (2 M HPF, 100 M H2O2, and 20 g/ml HPR) was immediately

suppressed by the subsequent addition of 100 mM thiourea, added after the

completion of the reaction. (E) K3Fe(CN)3 directly oxidized HPF. The signals

were recorded after 20 min incubation. Data from single experiments are shown;

they are representative of at least three independent replicates.

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Figure 3.14 The protective effects of thiourea and dipyridyl persist when

cells are treated with antibiotics in the absence of oxygen. (A) Exponential-

phase wild type cells were incubated with 150 mM thiourea and 5 g/ml ampicillin,

under both anoxic and aerated conditions. (B, C) Aerobic (B) or anaerobic (C)

wild-type cells were incubated with 5 g/ml ampicillin or 250 ng/ml norfloxacin.

Where indicated, 500 M dipyridyl (DP) was included. Thiourea and dipyridyl

concentrations matched those used in previous studies (19).

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Figure 3.15 Diminution of TCA-cycle flux protects cells from kanamycin.

Wild-type cells and TCA-cycle mutants (gltA and icd, lacking citrate synthase

and isocitrate dehydrogenase, respectively) were treated with 5 g/ml kanamycin,

and survival was monitored over time. Where indicated, 0.2% glucose was

included in the LB medium. In response to glucose, the expression of TCA-cycle

enzymes is repressed. The results suggest that a shift towards fermentative

metabolism suppresses kanamycin toxicity.

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3.8 References

1. Anbar, M. and P. Neta. 1967. A Compilation of Specific Biomolecular Rate Constants for the Reactions of Hydrated Electrons, Hydrogen Atoms and Hydroxyl Radicals with Inorganic and Organic Compounds in Aqueous Solutions. International Journal of Applied Radiation and Isotopes 18:493-523.

2. Anjem, A. and J. A. Imlay. 2012. Mononuclear iron enzymes are primary targets of hydrogen peroxide stress. J. Biol. Chem. 287:15544-15556.

3. Aslund, F., M. Zheng, J. Beckwith, and G. Storz. 1999. Regulation of the OxyR transcription factor by hydrogen peroxide and the cellular thiol-disulfide status. Proc. Natl. Acad. Sci. U. S. A. 96:6161-6165.

4. Datsenko, K. A. and B. L. Wanner. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. U. S. A. 97:6640-6645.

5. Davies, B. W., M. A. Kohanski, L. A. Simmons, J. A. Winkler, J. J. Collins, and G. C. Walker. 2009. Hydroxyurea induces hydroxyl radical-mediated cell death in Escherichia coli. Mol. Cell. 36:845-860.

6. Davies, M. J. 2005. The oxidative environment and protein damage. Biochim. Biophys. Acta. 1703:93-109.

7. Drlica, K., H. Hiasa, R. Kerns, M. Malik, A. Mustaev, and X. Zhao. 2009. Quinolones: action and resistance updated. Curr. Top. Med. Chem. 9:981-998.

8. Dwyer, D. J., M. A. Kohanski, B. Hayete, and J. J. Collins. 2007. Gyrase inhibitors induce an oxidative damage cellular death pathway in Escherichia coli. Mol. Syst. Biol. 3:91.

9. Foti, J. J., B. Devadoss, J. A. Winkler, J. J. Collins, and G. C. Walker. 2012. Oxidation of the guanine nucleotide pool underlies cell death by bactericidal antibiotics. Science. 336:315-319.

10. Gardner, P. R. and I. Fridovich. 1991. Superoxide sensitivity of the Escherichia coli 6-phosphogluconate dehydratase. J. Biol. Chem. 266:1478-1483.

11. Grant, S. S., B. B. Kaufmann, N. S. Chand, N. Haseley, and D. T. Hung. 2012. Eradication of bacterial persisters with antibiotic-generated hydroxyl radicals. Proc. Natl. Acad. Sci. U. S. A. 109:12147-12152.

12. Haldimann, A. and B. L. Wanner. 2001. Conditional-replication, integration, excision, and retrieval plasmid-host systems for gene structure-function studies of bacteria. J. Bacteriol. 183:6384-6393.

129  

13. Ikemura, K., M. Mukai, H. Shimada, T. Tsukihara, S. Yamaguchi, K. Shinzawa-Itoh, S. Yoshikawa, and T. Ogura. 2008. Red-excitation resonance Raman analysis of the nu(Fe=O) mode of ferryl-oxo hemoproteins. J. Am. Chem. Soc. 130:14384-14385.

14. Imlay, J. A. 2008. Cellular Defenses against Superoxide and Hydrogen Peroxide. Annu. Rev. Biochem. 77:755-776.

15. Imlay, J. A. and I. Fridovich. 1991. Assay of metabolic superoxide production in Escherichia coli. J. Biol. Chem. 266:6957-6965.

16. Jang, S. and J. A. Imlay. 2010. Hydrogen peroxide inactivates the Escherichia coli Isc iron-sulphur assembly system, and OxyR induces the Suf system to compensate. Mol. Microbiol. 78:1448-1467.

17. Jang, S. and J. A. Imlay. 2007. Micromolar intracellular hydrogen peroxide disrupts metabolism by damaging iron-sulfur enzymes. J. Biol. Chem. 282:929-937.

18. Keyer, K. and J. A. Imlay. 1996. Superoxide accelerates DNA damage by elevating free-iron levels. Proc. Natl. Acad. Sci. U. S. A. 93:13635-13640.

19. Kohanski, M. A., D. J. Dwyer, B. Hayete, C. A. Lawrence, and J. J. Collins. 2007. A common mechanism of cellular death induced by bactericidal antibiotics. Cell. 130:797-810.

20. Kohanski, M. A., D. J. Dwyer, J. Wierzbowski, G. Cottarel, and J. J. Collins. 2008. Mistranslation of membrane proteins and two-component system activation trigger antibiotic-mediated cell death. Cell. 135:679-690.

21. Kuo, C. F., T. Mashino, and I. Fridovich. 1987. alpha, beta-Dihydroxyisovalerate dehydratase. A superoxide-sensitive enzyme. J. Biol. Chem. 262:4724-4727.

22. Liochev, S. I. and I. Fridovich. 1994. The role of O2.- in the production of HO.: in

vitro and in vivo. Free Radic. Biol. Med. 16:29-33.

23. Liu, Y., S. C. Bauer, and J. A. Imlay. 2011. The YaaA protein of the Escherichia coli OxyR regulon lessens hydrogen peroxide toxicity by diminishing the amount of intracellular unincorporated iron. J. Bacteriol. 193:2186-2196.

24. Masse, E., C. K. Vanderpool, and S. Gottesman. 2005. Effect of RyhB small RNA on global iron use in Escherichia coli. J. Bacteriol. 187:6962-6971.

25. Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.

26. Park, S., X. You, and J. A. Imlay. 2005. Substantial DNA damage from submicromolar intracellular hydrogen peroxide detected in Hpx- mutants of Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 102:9317-9322.

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27. Seaver, L. C. and J. A. Imlay. 2004. Are respiratory enzymes the primary sources of intracellular hydrogen peroxide? J. Biol. Chem. 279:48742-48750.

28. Setsukinai, K., Y. Urano, K. Kakinuma, H. J. Majima, and T. Nagano. 2003. Development of novel fluorescence probes that can reliably detect reactive oxygen species and distinguish specific species. J. Biol. Chem. 278:3170-3175.

29. Sobota, J. M. and J. A. Imlay. 2011. Iron enzyme ribulose-5-phosphate 3-epimerase in Escherichia coli is rapidly damaged by hydrogen peroxide but can be protected by manganese. Proc. Natl. Acad. Sci. U. S. A. 108:5402-5407.

30. Taber, H. W., J. P. Mueller, P. F. Miller, and A. S. Arrow. 1987. Bacterial uptake of aminoglycoside antibiotics. Microbiol. Rev. 51:439-457.

31. Vakulenko, S. B. and S. Mobashery. 2003. Versatility of aminoglycosides and prospects for their future. Clin. Microbiol. Rev. 16:430-450.

32. Wang, X. and X. Zhao. 2009. Contribution of oxidative damage to antimicrobial lethality. Antimicrob. Agents Chemother. 53:1395-1402.

33. Wang, X., X. Zhao, M. Malik, and K. Drlica. 2010. Contribution of reactive oxygen species to pathways of quinolone-mediated bacterial cell death. J. Antimicrob. Chemother. 65:520-524.

34. Woodmansee, A. N. and J. A. Imlay. 2002. Quantitation of intracellular free iron by electron paramagnetic resonance spectroscopy. Methods Enzymol. 349:3-9.

35. Yamada, M., M. Shimizu, A. Katafuchi, P. Gruz, S. Fujii, Y. Usui, R. P. Fuchs, and T. Nohmi. 2012. Escherichia coli DNA polymerase III is responsible for the high level of spontaneous mutations in mutT strains. Mol. Microbiol. 86:1364-1375.

36. Yeom, J., J. A. Imlay, and W. Park. 2010. Iron homeostasis affects antibiotic-mediated cell death in Pseudomonas species. J. Biol. Chem. 285:22689-22695.

37. Zheng, M., X. Wang, L. J. Templeton, D. R. Smulski, R. A. LaRossa, and G. Storz. 2001. DNA microarray-mediated transcriptional profiling of the Escherichia coli response to hydrogen peroxide. J. Bacteriol. 183:4562-4570.

 

 

 

 

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CHAPTER FOUR: CONCLUSIONS

4.1 Conclusions from Current Work

4.1.1 YaaA plays a role in peroxide defense by controlling iron levels

The unknown function gene, yaaA, is induced by H2O2 in an OxyR-dependent

manner (24), which suggests that it has functions important during cellular

response to peroxide stress. Indeed, deletion of yaaA caused a growth defect

during peroxide stress in rich medium, which was due to Fenton-mediated DNA

damage. YaaA did not prevent the Fenton reaction by limiting the levels of H2O2,

cysteine or reduced flavin; instead, it protected the cell by controlling iron

concentrations. YaaA had functions independent of other iron homeostasis

proteins--Fur and Dps--in the OxyR regulon, which implied that YaaA played a

role other than inhibiting iron uptake and scavenging iron. We hypothesize that

YaaA is involved in iron trafficking during peroxide stress, but its molecular

mechanism is still unknown.

4.1.2 The lethal effect of classic bactericidal antibiotics does not involve

oxidative stress

For the past six years, several studies raised the hypothesis that different classes

of antibiotics-- including -lactams, aminoglycosides, and fluoroquinolones--

shared a common bactericidal mechanism: they kill bacteria by stimulating the

production of reactive oxygen species (5,6,8,12,13). In the current study, the

proposed events underpinning this model were directly tested. However, in

contrast to the hypothesis, bactericidal antibiotics did not induce the native

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peroxide defense system nor accelerate peroxide formation. Oxidative stress

sensitive enzymes were not damaged, and intracellular unincorporated iron level

did not increase. Oxidative DNA lesions were not important for their bactericidal

effects because the lethality of antibiotics persisted under anoxic conditions. In

addition, DNA repair systems had little effect on the toxicity of -lactams or

aminoglycosides. Taken together, we conclude that classical antibiotics did not

generate reactive oxygen species for their bactericidal effects.

4.2 Possible Future Work

4.2.1 What causes the growth defect of Hpx- yaaA mntH mutants in glucose

defined medium?

In LB medium, deletion of yaaA increased the level of unincorporated iron in

H2O2 stress cells, leading to hydroxyl radical-mediated DNA damage. However,

YaaA dysfunction had no impact on H2O2 stress cells growing in glucose defined

medium unless the manganese importer MntH was deleted.

In aerated glucose defined medium, the number of viable cells of Hpx- mntH

mutants kept increasing for three hours before growth arrested, while in the Hpx-

yaaA mntH strain, growth stopped immediately (Figure 2.15D). Because there

was no loss of viability, it was likely that the damage in yaaA mutants was due to

inhibition of metabolism, for example, poisoning H2O2 sensitive Fe-S cluster

containing enzymes or mononuclear iron enzymes.

The growth defect occurred even when all amino acids were provided (Figure

2.15B-D), which suggested that the damaged target was not in H2O2-sensitive

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amino acid biosynthetic pathways. In other words, the sensitive target was not

isopropylmalate isomerase, dihydroxy acid dehydratases, or 2-dehydro-3-

deoxyphosphoheptonate aldolase.

When high levels of manganese were supplied in the medium, they suppressed

the growth defect of Hpx- mntH cells. We hypothesize this protective effect

occurs because there is enough manganese entering the cell through other metal

importers (i.e. FeoB, ZupT or NikD), replacing the ferrous iron cofactor in

mononuclear iron containing enzymes and restoring their activities (1,2,18).

However, adding manganese made little improvement to the growth of Hpx- yaaA

mntH mutants (Figure 2.15C). There are two possible explanations for this result:

1) the yaaA mutant needs more manganese than non-specific import mechanism

can provide; or 2) the damaged mononuclear enzymes cannot obtain

manganese because either the yaaA deletion causes a block in non-specific

manganese import or it disrupts the intracellular manganese delivery.

Our results show that YaaA controls iron homeostasis during peroxide stress,

and it may do so by acting as an iron chaperone. Therefore, it is possible that

YaaA might also participate in manganese trafficking, since iron and manganese

are very similar metals and they are often exchangeable in biological systems.

Indeed, many mononuclear metalloenzymes can use either iron or manganese to

catalyze their reactions (1,2,18). The only known exceptions in E. coli are

superoxide dismutases and class I ribonucleotide reductases, both of which have

isozymes specifically using iron or manganese (3,4,14,19,20,22,23). Therefore, it

is possible that YaaA can deliver both iron and manganese to the target enzymes,

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and this becomes important in H2O2 stressed cells when manganese is depleted

(mntH mutant).

Most of the growth defects in oxidative stress do not appear until several

generation times after beginning aeration (2,16,18,21). We think this is because it

takes time to accumulate enough damage to overwhelm the repair systems and

stop growth. Hence the immediate growth defect of Hpx- yaaA mntH cells in

defined medium is unique. It suggests that not only is the target highly sensitive

but also the repair system fails rapidly. In order to identify the sensitive targets,

we can use different carbon sources (i.e. glycerol) to pinpoint the susceptible

metabolic pathways, and then screen for potential enzyme targets and the failing

repair systems. These data may provide some additional important information

about the function of YaaA.

4.2.2 What genes are induced in yaaA mutants?

Systems-biology assays are useful to generate new hypothesis that are

unexpected. Genome-wild microarray profiles or RNA-Seq (whole transcriptome

shotgun sequencing) of Hpx- yaaA strains and Hpx- yaaA mntH strains may

produce new hypotheses for YaaA function.

4.2.3 What is the three-dimensional structure of YaaA?

YaaA belongs to protein family domain of unknown function DUF328. This family

of proteins is found in over 1,300 bacterial species, but there is no recognizable

motif or available crystal structure (7). Therefore, solving the structure of YaaA

may provide some insight into the function of YaaA and this family of proteins.

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Indeed, the molecular function of Dps remained mysterious until its crystal

structure revealed that it was a structural analogue of ferritin, and it actually

functioned in iron sequestration (9). If the crystal structure of YaaA resembles

structures of proteins with assigned functions, it could provide insight into the

function of YaaA. This work is underway in other group. Combined with the yaaA

phenotype and the three-dimensional structure, we may be able to identify the

key amino acid residues in YaaA protein.

4.2.4 Does YaaA bind iron?

If YaaA is an iron chaperone, YaaA should be able to bind iron, most likely in the

ferrous form (3). Usually, the interaction between ligands and proteins can be

detected by isothermal titration calorimetry (ITC). ITC is a technique used to

measure the heat released or consumed during the binding events. Theoretically,

it could be employed to detect the binding of iron to YaaA. However, the binding

of ferrous iron to proteins is usually rather weak, according to the Irving-Williams

series, and as a metal chaperone the binding of YaaA and iron would be more

exchange labile, and thus the interaction between YaaA and iron might be below

the detection limit of direct ITC measurement. Thus, a negative result would not

necessarily reject the possibility that YaaA is an iron chaperone.

4.2.5 Are there any functionally critical residues in YaaA?

Cysteine residues form the basis of conserved motifs in Fe-S cluster dependent

proteins; however, there is no conserved binding motif for ferrous iron, which

makes it impossible to predict whether YaaA binds iron by just looking at the

protein sequence. When we compared the protein sequences of DUF328, we

136  

found no absolutely conserved residues. Most of the highly conserved residues

are tyrosine, phenylalanine and glycine residues, which are usually used for

structure reasons. There are also some positive charged and negative charged

residues that are relatively conserved. Mutation of conserved residues in YaaA

could alter the activity of YaaA, and in combination with the crystal structure, this

information will be useful to determine the function of YaaA.

4.2.6 What percentage of the HPF is oxidized in antibiotic treated cells?

Hydroxyphenyl fluorescein (HPF) has been widely used for hydroxyl radical

detection (5,6,10,12,13,15,17). As a cell permeable dye, the amount of HPF

inside the cell depends upon envelope permeability. Therefore, an increased

HPF fluorescent signal inside the cell can be caused in two ways: 1) oxidation of

HPF or 2) larger uptake of un-oxidized dye. Some antibiotics, such as

aminoglycosides, can increase membrane permeability during their action, which

might be the reason for higher HPF signal in antibiotic-treated cells. In order to

distinguish these two possibilities, it is necessary to determine what percentage

of HPF inside the cell is oxidized. One possible way to do that is to use

horseradish peroxidase/H2O2 or K3Fe(CN)6 to completely oxidize HPF in the cell

extract, and compare the ratio of fluorescence before and after oxidation.

4.2.7 Can KatG oxidize the HPF dye?

Although HPF is often used as a hydroxyl radical specific detection probe, my in

vitro studies showed that HPF can be oxidized by high-valent iron centers, such

as the ferryl radical intermediate in the Fenton reaction, or ferryl-porphyrin radical

(Compound I) intermediate of horseradish peroxidase during its catalytic cycle

137  

(11). Thus, we hypothesize that HPF can be oxidized by heme prosthetic groups

in enzymes, if they are accessible.

There are several candidates in aerobic growing E. coli that might oxidize HPF,

including hydroperoxidase I (KatG) and cytochrome oxidases. KatG has both

catalase and peroxidase activity, which means that after it reduce one molecule

of H2O2, the oxidized enzyme intermediate (Compound I) can be re-reduced back

to the resting state by abstracting two electrons from a second H2O2 or an

organic compound. Actually, the organic substrate, o-dianisidine, is used in

hydroperoxidase I assay to measure KatG activity. Therefore, the oxidized KatG

intermediate may be able to abstract electrons from HPF and oxidize the dye to a

fluorescent form. To test this idea, one can measure the HPF fluorescent signal

in katG deletion mutants, wild type cells and cells overexpressing katG to see

whether there is correlation between the signal intensity and the amount of KatG.

Similar experiments can be used to determine whether cytochrome oxidase can

oxidize HPF. These high-valence iron centers may accumulate inside the cell

when metabolism is blocked, a situation occurring in antibiotic-treated cells.

These results might help to explain why the antibiotic treated cells show higher

HPF fluorescent signals. Because of its convenience, HPF is also widely

employed to detect oxidative stress in other scenarios. Therefore, it is important

to find out what this probe really reacts with inside the cell. Based on the results,

we might be able to find a condition under which the probe works more

specifically and provides valid results, or we might have to search for a new way

to detect ROS.

138  

4.3 References

 

  1.   Anjem, A. and J. A. Imlay. 2012. Mononuclear iron enzymes are primary targets of hydrogen peroxide stress. J. Biol. Chem. 287:15544-15556.

2. Anjem, A., S. Varghese, and J. A. Imlay. 2009. Manganese import is a key element of the OxyR response to hydrogen peroxide in Escherichia coli. Mol. Microbiol. 72:844-858.

3. Cotruvo, J. A. and J. Stubbe. 2011. Class I ribonucleotide reductases: metallocofactor assembly and repair in vitro and in vivo. Annu. Rev. Biochem. 80:733-767.

4. Cotruvo, J. A., Jr. and J. Stubbe. 2012. Metallation and mismetallation of iron and manganese proteins in vitro and in vivo: the class I ribonucleotide reductases as a case study. Metallomics. 4:1020-1036.

5. Davies, B. W., M. A. Kohanski, L. A. Simmons, J. A. Winkler, J. J. Collins, and G. C. Walker. 2009. Hydroxyurea induces hydroxyl radical-mediated cell death in Escherichia coli. Mol. Cell. 36:845-860.

6. Dwyer, D. J., M. A. Kohanski, B. Hayete, and J. J. Collins. 2007. Gyrase inhibitors induce an oxidative damage cellular death pathway in Escherichia coli. Mol. Syst. Biol. 3:91.

7. Finn, R. D., J. Tate, J. Mistry, P. C. Coggill, S. J. Sammut, H. R. Hotz, G. Ceric, K. Forslund, S. R. Eddy, E. L. Sonnhammer, and A. Bateman. 2008. The Pfam protein families database. Nucleic Acids Res. 36:D281-D288.

8. Foti, J. J., B. Devadoss, J. A. Winkler, J. J. Collins, and G. C. Walker. 2012. Oxidation of the guanine nucleotide pool underlies cell death by bactericidal antibiotics. Science. 336:315-319.

9. Grant, R. A., D. J. Filman, S. E. Finkel, R. Kolter, and J. M. Hogle. 1998. The crystal structure of Dps, a ferritin homolog that binds and protects DNA. Nat. Struct. Biol. 5:294-303.

10. Grant, S. S., B. B. Kaufmann, N. S. Chand, N. Haseley, and D. T. Hung. 2012. Eradication of bacterial persisters with antibiotic-generated hydroxyl radicals. Proc. Natl. Acad. Sci. U. S. A. 109:12147-12152.

11. Hersleth, H. P., U. Ryde, P. Rydberg, C. H. Gorbitz, and K. K. Andersson. 2006. Structures of the high-valent metal-ion haem-oxygen intermediates in peroxidases, oxygenases and catalases. J. Inorg. Biochem. 100:460-476.

12. Kohanski, M. A., D. J. Dwyer, B. Hayete, C. A. Lawrence, and J. J. Collins. 2007. A common mechanism of cellular death induced by bactericidal antibiotics. Cell. 130:797-810.

139  

13. Kohanski, M. A., D. J. Dwyer, J. Wierzbowski, G. Cottarel, and J. J. Collins. 2008. Mistranslation of membrane proteins and two-component system activation trigger antibiotic-mediated cell death. Cell. 135:679-690.

14. Martin, J. E. and J. A. Imlay. 2011. The alternative aerobic ribonucleotide reductase of Escherichia coli, NrdEF, is a manganese-dependent enzyme that enables cell replication during periods of iron starvation. Mol. Microbiol. 80:319-334.

15. Nguyen, D., A. Joshi-Datar, F. Lepine, E. Bauerle, O. Olakanmi, K. Beer, G. McKay, R. Siehnel, J. Schafhauser, Y. Wang, B. E. Britigan, and P. K. Singh. 2011. Active starvation responses mediate antibiotic tolerance in biofilms and nutrient-limited bacteria. Science. 334:982-986.

16. Park, S., X. You, and J. A. Imlay. 2005. Substantial DNA damage from submicromolar intracellular hydrogen peroxide detected in Hpx- mutants of Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 102:9317-9322.

17. Shatalin, K., E. Shatalina, A. Mironov, and E. Nudler. 2011. H2S: a universal defense against antibiotics in bacteria. Science. 334:986-990.

18. Sobota, J. M. and J. A. Imlay. 2011. Iron enzyme ribulose-5-phosphate 3-epimerase in Escherichia coli is rapidly damaged by hydrogen peroxide but can be protected by manganese. Proc. Natl. Acad. Sci. U. S. A. 108:5402-5407.

19. Stubbe, J. and J. A. Cotruvo, Jr. 2011. Control of metallation and active cofactor assembly in the class Ia and Ib ribonucleotide reductases: diiron or dimanganese? Curr. Opin. Chem. Biol. 15:284-290.

20. Vance, C. K. and A. F. Miller. 1998. Spectroscopic comparisons of the pH dependencies of Fe-substituted (Mn)superoxide dismutase and Fe-superoxide dismutase. Biochemistry. 37:5518-5527.

21. Varghese, S., A. Wu, S. Park, K. R. Imlay, and J. A. Imlay. 2007. Submicromolar hydrogen peroxide disrupts the ability of Fur protein to control free-iron levels in Escherichia coli. Mol. Microbiol. 64:822-830.

22. Whittaker, M. M. and J. W. Whittaker. 1997. Mutagenesis of a proton linkage pathway in Escherichia coli manganese superoxide dismutase. Biochemistry. 36:8923-8931.

23. Yost, F. J., Jr. and I. Fridovich. 1973. An iron-containing superoxide dismutase from Escherichia coli. J. Biol. Chem. 248:4905-4908.

24. Zheng, M., X. Wang, L. J. Templeton, D. R. Smulski, R. A. LaRossa, and G. Storz. 2001. DNA microarray-mediated transcriptional profiling of the Escherichia coli response to hydrogen peroxide. J. Bacteriol. 183:4562-4570.

 

 

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APPENDIX A: DELETION OF OXYS DOES NOT GENERATE A

PHENOTYPE IN CATALASE/PEROXIDASE-DEFICIENT MUTANTS

During H2O2 stress, OxyR induces a 109 nucleotide, non-translated small RNA

oxyS (1). This small RNA is located 95 base pairs upstream of the oxyR gene,

and is divergently transcribed. In H2O2 stressed cells, oxyS is induced within 1

min; the number of its transcripts can reach to ~4,500 copies per cell; the oxyS

RNA is quite stable and it has a half-life of 12-30 min (1). The oxyS RNA

activates or represses the expression of about 40 genes, including the

transcriptional regulator, FhlA, and an alternative sigma factor, RpoS (1). FhlA is

a formate hydrogen lyase activator that controls genes required for formate

metabolism. The oxyS RNA directly binds to FhlA mRNA, interfering with

ribosome binding, reducing the translational level of FhlA (2,3). In the case of

RpoS, oxyS works with the RNA-binding protein Hfq, and negatively regulates

the translation of RpoS (7). A subset of the oxyS regulated genes are RpoS

dependent (i.e. dps), while the others are RpoS independent (1). The regulation

of FhlA and RpoS by oxyS suggests that oxyS might play a role in

communication between the H2O2-stress response and other regulatory networks.

Previous studies reported that oxyS protects cells against spontaneous

mutagenesis induced by alkylating chemicals (i.e. MNNG and ethyl

methanesulfonate) or H2O2. However, in these experiments, millimolar

concentrations of H2O2 were used (1). In order to study the function of oxyS

under physiological H2O2 stress, a gene deletion of oxyS was constructed in

141  

catalase/peroxidase-deficient (Hpx-) mutants, and the resulting Hpx- oxyS strain

was screened for growth defects. However, Hpx- oxyS mutants exhibited similar

growth rates as compared to its Hpx- parent strain during H2O2 stress in both LB

and glucose defined medium (Figure A.1). We conclude that oxyS is not

important for H2O2-stress under the condition we tested.

A.1 Material and Methods

A.1.1 Strain construction

The strains used in this study are listed in Table A.1. Deletions were constructed

by the Red recombination deletion method (4) and were transferred to other

strains by P1 transduction (5). The antibiotic resistant cassettes were

subsequently removed by FLP-mediated excision (4).

A.1.2 Bacterial growth

Complex growth media was LB (10 g tryptone, 5 g yeast extract, and 10 g NaCl

per liter). Defined glucose/amino acids media contained minimal A salts (5), 0.2%

glucose, 0.02% MgSO47H2O, and 5 g/ml thiamine. When indicated, 0.2%

casamino acids and 0.5 mM tryptophan, or 0.5 mM of aromatic amino acids

(histidine, phenylalanine and tyrosine) were added. Experimental protocols were

designed to ensure that measurements were performed upon exponentially

growing cells. For growth studies, anaerobic overnight cultures were diluted into

anaerobic media and grown for 4 to 5 generations to early log phase (OD600nm of

0.15 to 0.20). The cultures were then diluted into aerobic medium of the same

composition to an OD600nm of 0.005. All cultures were grown at 37 C. Aerobic

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defined medium was prepared immediately prior to use. Anaerobic medium was

transferred immediately after autoclaving to an anaerobic chamber (Coy

Laboratory Products), where it was stored under an atmosphere of 5% CO2/10%

H2/85% N2 for at least one day prior to use.

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A.2 Table

Table A.1 Strains used in this study.

Name Genotype/characteristics Source/Ref.

AL107 As LC106 plus oxyS1::cat This study

AL110 As MG1655 plus oxyS1::cat This study

BW25113 lacIq rrnBT14 lacZWJ16 hsdR514 araBADAH33 rhaBADLD78 (4)

LC106 As MG1655 plus (ahpC-ahpF’) kan:: ahpF (katG17::Tn10)1

(katE12::Tn10)1

(6)

MG1655 Wild type Lab stock

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A.3 Figure

Figure A.1 Deletion of small RNA oxyS did not introduce a growth defect in

H2O2 stressed cells. Anaerobic exponential-phase cultures were diluted into the

same aerobic medium at time zero. Hpx- oxyR mutants did not have growth

defect in LB (A), or glucose defined medium with all amino acid supplement (B)

or just aromatic amino acid supplement (C). Strains used: WT (MG1655), AL110

(oxyS), Hpx- (LC106), Hpx- oxyS (AL107).

   

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A.4 References

 

  1. Altuvia, S., D. Weinstein-Fischer, A. Zhang, L. Postow, and G. Storz. 1997. A small, stable RNA induced by oxidative stress: role as a pleiotropic regulator and antimutator. Cell. 90:43-53.

2. Altuvia, S., A. Zhang, L. Argaman, A. Tiwari, and G. Storz. 1998. The Escherichia coli OxyS regulatory RNA represses fhlA translation by blocking ribosome binding. EMBO J. 17:6069-6075.

3. Argaman, L. and S. Altuvia. 2000. fhlA repression by OxyS RNA: kissing complex formation at two sites results in a stable antisense-target RNA complex. J. Mol. Biol. 300:1101-1112.

4. Datsenko, K. A. and B. L. Wanner. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. U. S. A. 97:6640-6645.

5. Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.

6. Seaver, L. C. and J. A. Imlay. 2004. Are respiratory enzymes the primary sources of intracellular hydrogen peroxide? J. Biol. Chem. 279:48742-48750.

7. Zhang, A., S. Altuvia, A. Tiwari, L. Argaman, R. Hengge-Aronis, and G. Storz. 1998. The OxyS regulatory RNA represses rpoS translation and binds the Hfq (HF-I) protein. EMBO J. 17:6061-6068.