THE COMBINED CONTRIBUTIONS OF NITRIC OXIDE SYNTHASE AND THE STAPHYLOCOCCAL RESPIRATORY RESPONSE REGULATOR TO

STAPHYLOCOCCUS AUREUS PHYSIOLOGY

By

AUSTIN BLAKE MOGEN

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2016

© 2016 Austin Blake Mogen

To my mother Randy Mogen

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ACKNOWLEDGMENTS

I would first like to thank my mentor Kelly Rice and my committee members Tony

Romeo, Julie Maupin, Joseph Larkin III, and Jeannine Brady for all of their knowledge

and suggestions that helped me develop my project. Even more so I would like to

specifically thank Kelly for the environment of free thought and patience that she

fostered as my mentor. I certainly would not be where I am if she had not graciously

decided to commit her time to mentoring an inexperienced undergraduate student.

I would also like to acknowledge my wonderful lab mates, both past and present

who made coming to work every day enjoyable. Many have become life-long friends

and I would like to especially thank April Lewis, Erin Almand, Elisha Roberts, Silvia

Orsini, O’neshia Carney, Matt Turner, Adam Grossman, and Hoang Ngyuen. Each one

of you has contributed to my success by either brainstorming, helping with lab work, or

by simply just being there as a friend. Thanks homies. As well, I would like to thank Jeff

Daskin, Casey Johnson, Krista Seraydar, Will Karbaum, Krista Godbey, Jonathan

Orsini, Mike Lewis, and Mike Albiez for being some of the best friends anyone could ask

for.

Last but not least I would like to thank my family for all of the unconditional love

and support they provided me including my mother Randy Mogen, step father Harold

Thomas, and sister Karli Mogen. I am truly blessed with having a family that has both

emotionally and financially supported me throughout my journey.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS .................................................................................................. 4

LIST OF TABLES ............................................................................................................ 8

LIST OF FIGURES .......................................................................................................... 9

LIST OF ABBREVIATIONS ........................................................................................... 11

ABSTRACT ................................................................................................................... 14

CHAPTER

1 LITERATURE REVIEW .......................................................................................... 16

Staphylococcus aureus ........................................................................................... 16

General Characteristics .................................................................................... 16 Emergence of Antibiotic Resistant Staphylococcus aureus .............................. 17

Staphylococcus aureus Virulence Determinants .............................................. 19

Factors for colonization and dissemination ................................................ 20 Strategies for S. aureus immune evasion .................................................. 24 S. aureus toxins ......................................................................................... 26

Regulation of virulence factors and summary of virulence strategies ........ 27 Metabolism of Staphylococcus aureus.................................................................... 29

Overview .......................................................................................................... 29 Low Oxygen Fermentative Metabolism ............................................................ 31 Amino Acid Metabolism .................................................................................... 32

Respiratory Metabolism .................................................................................... 34 General respiratory chain components ...................................................... 34

Aerobic respiratory components ................................................................ 35 Anaerobic respiration ................................................................................. 38

Genetic Regulation of Staphylococcus aureus Metabolism .............................. 40

Metabolism and Virulence ................................................................................ 44 Global metabolic regulators and virulence ................................................. 44 Link between central metabolism and virulence ......................................... 45

Lactate as a central virulence metabolite ................................................... 46

Biochemistry of Reactive Oxygen and Nitrogen Species ........................................ 47 General ROS Characteristsics ......................................................................... 47

ROS chemistry and toxicity ........................................................................ 47 Pathways of ROS generation ..................................................................... 48

Protection from Oxidative Stress in Staphylococcus aureus ............................ 49

Classical ROS detoxification proteins ........................................................ 49 Thiol-specific redox systems in Staphylococcus aureus ............................ 50

Additional oxidative stress resistance mechanisms ................................... 51 Pathways and Targets of RNS ......................................................................... 52

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RNS chemistry and production .................................................................. 52 Cellular targets of RNS .............................................................................. 53

Protection From Nitrosative Stress in Staphylococcus aureus ......................... 55

NO detoxification proteins in S. aureus ...................................................... 55 S. aureus metabolic flexibility in response to nitrosative stress.................. 56

Nitric Oxide Synthase in Mammals and Bacteria .................................................... 58 Mammalian Nitric Oxide Synthase ................................................................... 58

Mammalian NOS structure and chemical reaction ..................................... 58

Mammalian NOS isotypes and their functions ........................................... 59 Bacterial Nitric Oxide Synthase ........................................................................ 60

Bacterial NOS discovery ............................................................................ 60 Bacterial NOS structure ............................................................................. 61

Reductase partner studies for bNOS ......................................................... 62 bNOS inhibitor studies ............................................................................... 63 Functional studies of bNOS proteins .......................................................... 64

Staphylococcus aureus NOS .................................................................................. 69

General Characteristsics .................................................................................. 69 Discovery and structural characterization .................................................. 69 Sequence identity and genomic organization ............................................ 70

Functional Studies on saNOS .......................................................................... 70 Protection from oxidative stress ................................................................. 70

Contribution of saNOS to virulence and antimicrobial resistance ............... 71 Contributions of saNOS to General Physiology ................................................ 74

Hypothesis and Aims .............................................................................................. 75

2 RESULTS ............................................................................................................... 84

Aim 1. Contribution of saNOS to General Physiology ............................................. 84 Growth Phenotypes Upon nos Mutation ........................................................... 84

saNOS Has an Altered Transcriptome. ............................................................ 86 Intracellular and Secreted Metabolite Profiles of the nos Mutant ...................... 88

Aim 2. saNOS Contributes to Endogenous Oxidative Stress and Respiratory Metabolism .......................................................................................................... 91

Mutation of nos Increases Endogenous Oxidative Stress ................................ 91

saNOS Contributes to Respiratory Function..................................................... 92 Inhibition of Ndh Limits Oxidative Stress in a nos Mutant ................................. 96

Aim 3. SrrAB as a Potential Regulator of nos Mutant Metabolic Adaptation ........... 97

Growth Phenotypes of the nos srrAB Double Mutant ....................................... 97 Membrane Potential of the nos srrAB Double Mutant ...................................... 99 Metabolism of the nos srrAB Double Mutant .................................................... 99

3 MATERIALS AND METHODS .............................................................................. 125

Bacterial Strains and Culture Conditions .............................................................. 125 Creation of nos srrAB Double Mutant and Complement ....................................... 125

Growth Curve Analysis ......................................................................................... 126

Colony Size Comparison ...................................................................................... 126

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Transmission Electron Microscopy ....................................................................... 127 Scanning Electron Microscopy .............................................................................. 128 RNAseq Analysis .................................................................................................. 129

Metabolite Analysis Using LC/MS/MS .................................................................. 131 Cell Collection and Metabolite Sample Preparation ....................................... 131 Extraction, Derivatization, and LC/MS/MS Quantitation of Organic Acids

from Cell Homogenate and Extracellular Media .......................................... 132 Extraction, Derivatization, and LC/MS/MS Quantitation of Amino Acids from

Cell Homogenate and Extracellular Media .................................................. 133 Extraction, Derivatization, and LC/MS/MS Quantitation of Pyridine

Nucleotides and Adenosine Phosphates from Cell Homogenate ................ 134 Measurement of Intracellular ROS and O2

- ........................................................... 136

Determination of Catalase Activity ........................................................................ 137 Assessment of Membrane Potential ..................................................................... 138 CTC Staining ........................................................................................................ 138

Oxygen Consumption ........................................................................................... 139

Determination of Aconitase Activity ...................................................................... 139 Statistical Analysis ................................................................................................ 140

4 DISCUSSION ....................................................................................................... 143

5 CONCLUSIONS AND FUTURE DIRECTIONS .................................................... 156

APPENDIX

A ADDITIONAL FIGURES ....................................................................................... 162

B ADDITIONAL TABLES .......................................................................................... 168

LIST OF REFERENCES ............................................................................................. 186

BIOGRAPHICAL SKETCH .......................................................................................... 238

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LIST OF TABLES

Table page 2-1 Generation times for all strains ......................................................................... 118

2-2 Select genes altered upon nos mutation .......................................................... 119

2-3 qRT-PCR confirmation of select genes ............................................................ 120

2-4 Select cellular nos mutant metabolites ............................................................. 121

2-5 Energy charge .................................................................................................. 122

2-6 Select nos srrAB double mutant cellular metabolites ........................................ 123

2-7 Select nos srrAB double mutant extracellular metabolites ................................ 124

3-1 Bacterial strains and plasmids constructs used in this study ............................ 141

3-2 PCR primers used in this study ........................................................................ 142

B-1 List of all genes altered in the nos mutant at 4 hours growth ............................ 168

B-2 List of all genes altered in the nos mutant at 6 hours growth ............................ 179

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LIST OF FIGURES

Figure page 1-1 Fermentation pathways of S. aureus. ................................................................. 77

1-2 Branched respiratory chain of S. aureus.. .......................................................... 78

1-3 Cellular targets of NO. ........................................................................................ 79

1-4 Structure of saNOS. ........................................................................................... 80

1-5 Genomic organization and distribution of saNOS. .............................................. 81

1-6 Contribution of saNOS to H2O2 resistance ......................................................... 82

1-7 saNOS in a sepsis model of infection. ................................................................ 83

2-1 Wildtype and nos mutant growth curves ........................................................... 103

2-2 Growth curves with addition of chemical NO donor and in a MRSA background. ...................................................................................................... 104

2-3 TEM analysis of nos mutant. ............................................................................ 105

2-4 SEM analysis of nos mutant ............................................................................. 106

2-5 Distribution of gene functional categories expressed by the nos mutant in 4 hour cultures ..................................................................................................... 107

2-6 Distribution of gene functional categories expressed by the nos mutant relative to wildtype of 6 hour cultures ............................................................... 108

2-7 Intracellular ROS, superoxide detection, and catalase activity in wildtype and nos mutant cultures .......................................................................................... 109

2-8 Effect of saNOS on membrane potential .......................................................... 110

2-9 Respiration determined by CTC staining .......................................................... 111

2-10 Effect of saNOS on oxygen consumption. ........................................................ 112

2-11 Intracellular ROS upon Ndh inhibition and aconitase activity of the nos mutant .............................................................................................................. 113

2-12 Agar plate growth of the nos srrAB double mutant ........................................... 114

2-13 Quantification of colony size ............................................................................. 115

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2-14 Growth curves of nos and nos srrAB double mutant strains ............................. 116

2-15 Effect of srrAB single and nos srrAB double mutation on membrane potential 117

4-1 Central metabolic mapping of nos mutant transcriptional and metabolic changes ............................................................................................................ 154

4-2 Central metabolic mapping of nos srrAB double mutant metabolic changes .... 155

A-1 Cellular organic acids of the nos, srrAB, and nos srrAB mutant strains. .......... 162

A-2 Extracellular organic acids of the nos, srrAB, and nos srrAB mutant strains. ... 163

A-3 Cellular amino acids of the nos, srrAB, and nos srrAB mutant strains. ............. 164

A-4 Extracellular amino acids of the nos, srrAB, and nos srrAB mutant strains ...... 165

A-5 Cellular NAD nucleotides of the nos, srrAB, and nos srrAB mutant strains ...... 166

A-6 Cellular adenosine nucleotides of the nos, srrAB, and nos srrAB mutant strains ............................................................................................................... 167

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LIST OF ABBREVIATIONS

Agr Accesory gene regulator

Ala Alanine

Arg Arginine

Asn Asparagine

Asp Aspartate

ATP Adenosine triphosphate

B. anthracis Bacillus anthracis

B. subtilis Bacillus subtilis

baNOS Bacillus anthracis nitric oxide synthase

BCAA Branched chain amino acid

BLOQ Below the limit of quantitation

bNOS Bacterial NOS

bsNOS Bacillus subtilis nitric oxide synthase

CFU/ml Colony forming unit per mililiter

Cm Chloramphenicol

CM-H2DCFDA Carboxy-2′,7′-dichlorofluorescein

CTC 5-cyano-2,3-ditolyl tetrazolium chloride

Ctl Citrulline

Cys Cysteine

D. radiodurans Deinococcus radiodurans

DiOC2(3) 3,3′-diethyloxacarbocyanine iodide

DPTA Dipropylenetriamine

E. coli Escherichia coli

Erm Erythromycin

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Gln Glutamine

Glu Glutamate

Gly Glycine

H2O2 Hydrogen Peroxide

His Histidine

Ile Isoleucine

LB Luria-Bertani broth

Leu Leucine

Lqo Lactate quinone oxidoreductase

Lys Lysine

Met Methionine

Mqo Malate quinone oxidoreductase

Ndh NADH dehydrogenase

NO Nitric oxide

NO2- Nitrite

NO3- Nitrate

NOS Nitric oxide synthase

O2 Oxygen

O2- Superoxide

OD Optical density

Orn Ornithine

Phe Phenylalanine

Pro Proline

qRT-PCR Quantitative real-time polymerase chain reaction

Rex Redox response regulator

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RNS Reactive nitrogen species

ROS Reactive oxygen species

S. aureus Staphylococcus aureus

S. carnosus

S. cellulosum

Staphylococcus carnosus

Sorangium cellulosum

S. epidermidis Staphylococcus epidermidis

S. saprophyticus

S. turgidiscabies

Staphylococcus saprophyticus

Streptomyces turgidiscabies

saNOS Staphylococcus aureus nitric oxide synthase

SATMD Staphylococcus aureus transcriptome meta-database

SCV Small colony variant

SEM Scanning electron microscopy

Ser Serine

SOD Superoxide dismutase

SrrAB Staphylococcal respiratory response regulator

TCA

TCS

Tricarboxylic acid

Two-component system

TEM Transmission electron microscopy

Thr Threonine

Trp Tryptophan

TSB Tryptic soy broth

TSB-G Tryptic soy broth without glucose

Tyr Tyrosine

TZ Thioridizine HCl

Val Valine

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

THE COMBINED CONTRIBUTIONS OF NITRIC OXIDE SYNTHASE AND THE

STAPHYLOCOCCAL RESPIRATORY RESPONSE REGULATOR TO STAPHYLOCOCCUS AUREUS PHYSIOLOGY

By

Austin Blake Mogen

December 2016

Chair: Kelly Rice Major: Microbiology and Cell Science

S. aureus is a successful human pathogen notorious for being resistant to

multiple antibiotics. A promising target for drug development is the S. aureus nitric oxide

synthase (saNOS), as a link between NOS inhibition and increased antimicrobial

efficacy is already been established. Although the exact mechanism is unknown,

saNOS contributes to S. aureus virulence and protection against oxidative stress. When

grown aerobically, reactive oxygen species (ROS) were elevated in a S. aureus nos

mutant, independent of catalase activity. Respiratory chain function was altered in a nos

mutant, highlighted by elevated respiratory dehydrogenase activity and membrane

potential, as well as slightly altered O2 consumption. Multiple transcriptional and

metabolic changes were also observed in a S. aureus nos mutant, as assessed by

RNAseq and targeted metabolomics analyses, respectively. Specifically, expression of

genes associated with stress response (msrA1, scdA, ahpF, hmp, trxA),

anaerobic/lactate metabolism (ldh2, nar, pfl, adhA), and cytochrome

biosynthesis/assembly (qox, ctaB, hemA) were increased in the nos mutant relative to

wildtype. Metabolites utilized to produce reducing equivalents by the oxidative branch of

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the TCA cycle were depleted in a nos mutant (citrate and α-ketoglutarate), whereas

fumarate and malate levels were increased relative to wildtype. A significant reduction in

lactate levels was also observed in the nos mutant. The staphylococcal respiratory

response regulator (SrrAB) is a proposed sensor of the reduction state of respiratory

quinones and regulates many of the genes altered in the nos mutant as identified by

RNAseq. Growth phenotypes of a nos srrAB double mutant included a small colony-like

phenotype and altered growth curves. Metabolic analysis of a nos srrAB double mutant

revealed significant decreases in TCA cycle metabolites, cellular amino acids, and

biosynthetic NADPH, as well as a significant increase in lactate secretion. Collectively,

these results support a model in which the absence of saNOS results in ROS

accumulation and altered respiratory chain function, which is sensed by SrrAB and may

signal the cells to switch to an alternative lactate-based fermentative metabolism. This

contribution is the first to describe a bacterial NOS that is central to metabolism,

respiratory function, and endogenous ROS.

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CHAPTER 1 LITERATURE REVIEW

Staphylococcus aureus

General Characteristics

Staphylococcus aureus is an extremely successful human colonizer and

opportunistic pathogen. This organism has closely evolved with the host to fill a

biological niche by fine-tuning virulence determinants, its metabolism, and how it

responds to external stress; allowing it to readily adapt to the dynamic environment of

the human body. As a Gram positive bacterium, phylogenetic classification places S.

aureus in the firmicute phylum, order Bacillales, and the Staphylococcaceae family

(Somerville & Proctor, 2009b, Gibbons & Murray, 1978). Originally named after the

Greek word staphyle, meaning "bunch of grapes" and coccus meaning "grain or berry",

organisms in this group often present with a grapelike cluster formation when viewed

under the microscope (Somerville & Proctor, 2009b). The designation "aureus" can be

traced to Latin roots as "aurei" constituted the Roman word for "golden", and an aureus

was the name given to a commonly minted solid gold coin (Buttrey, 2012, Scheidel,

2010, West, 1916). When first isolated in pure culture by Rosenbach in 1884 (Cowan et

al., 1954), this gold pigmented bacterium was given the name S. aureus. Since then the

pigmentation has been attributed to a series of complex biosynthetic pathways that

produce carotenoid molecules, the most prevalent of which is designated

staphyloxanthin (Marshall & Wilmoth, 1981b, Marshall & Wilmoth, 1981a). Not all S.

aureus strains are pigmented, with some notable exceptions being lab generated small

colony variant (SCV) mutants (McNamara & Proctor, 2000) and both clinical and lab

isolates with naturally-occuring mutations in regulation of SigB, an alternative sigma

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factor (Karlsson-Kanth et al., 2006). Examination of S. aureus genomes shows that it is

comprised of approximately 2.6-2.9 million bps (depending on the strain), which

translates to between 2600 and 2800 genes (Lindsay, 2008, Sassi et al., 2015). An

important characteristic of staphylococcal genomes is the low G+C content, which is

generally observed to be ~32% (Lindsay, 2008, Sassi et al., 2015). In addition to a

single circular chromosome, the totality of the S. aureus genome also contains

prophages, plasmids, and transposons (Lindsay, 2008), which vary between strains

(Deurenberg & Stobberingh, 2008). Many of the non-chromosomal genetic constituents

have been directly responsible for the emergence of multiple drug resistant strains

(discussed below).

Emergence of Antibiotic Resistant Staphylococcus aureus

Under most conditions S. aureus is a non-virulent colonizer of the nasal cavity,

specifically the anterior nares (Williams, 1963). Up to 30% of humans are predicted to

be asymptomatic carriers of S. aureus (Rim & Bacon, 2007), but when the right

conditions are present these carriers are at a higher risk of infection and are presumed

to be an important source of S. aureus strains that spread among the population

(Gorwitz et al., 2008, Kluytmans et al., 1997). A large epidemiological study conducted

in the U.S. concluded that 11.6 million outpatient and emergency room visits, and nearly

500,000 hospital admissions per year, are attributed to S. aureus skin infection (McCaig

et al., 2006). The history of S. aureus epidemiology is dominated by the emergence of

multiple drug resistant strains including methicillin resistant S. aureus (MRSA),

vancomycin intermediate resistant S. aureus (VISA)(Appelbaum, 2006), and more

recently, fully vancomycin resistant S. aureus (VRSA)(Rodvold & McConeghy, 2014).

While many S. aureus strains are opportunistic pathogens, the emergence of highly

18

virulent MRSA strains has led to this organism becoming one of the leading causes of

human bacterial infections and death worldwide (DeLeo et al., 2010, Monaco et al.,

2016). Classically viewed as a nosocomial infection found in patients with other risk

factors, hospital acquired MRSA (HA-MRSA) is endemic to the healthcare setting,

highlighted by high morbidity and mortality (Boucher & Corey, 2008). Of increasing

importance is the emergence of community acquired MRSA (CA-MRSA) that can be

transmitted person-to-person and infect individuals with no apparent risk factors

(Boucher & Corey, 2008). While distinct CA-MRSA isolates likely originated separately

from HA-MRSA (Udo et al., 1993, 1999), the difference between these isolates has

become blurred in the hospital setting (Boucher & Corey, 2008). According to the CDC,

MRSA is among the most common causes of infections in the U.S. and is responsible

for approximately 80,000 infections per year, with an incidence of 25 per 100,000

population (2012, Dantes et al., 2013). It is also estimated that close to 19,000

hospitalized American patients are killed by MRSA infections each year, similar to the

number of deaths from AIDS, tuberculosis, and viral hepatitis combined (Boucher &

Corey, 2008). The success of MRSA as a pathogen is thought to be in part due to

acquisition of additional virulence factors or adaptation of gene expression (Otto, 2012),

but antibiotic resistance is arguably still one of the main contributors to the dominance

of this pathogen. More than 90% of S. aureus strains are resistant to penicillin (Lowy,

2003, Chambers & Neu, 2000), which is conferred by beta lactamase (BlaZ)(East &

Dyke, 1989). Mechanistically, these enzymes act by hydrolyzing the β-lactam ring,

rendering the β-lactam antibiotic inactive (Massova & Kollman, 2002). Resistance to

methicillin is attributed to the mecA gene, part of a mobile genetic element (Katayama et

19

al., 2000), which codes for an alternative penicillin binding protein (PBP 2′)(Matsuhashi

et al., 1986). PBP 2' works by decreasing the affinity of penicillin binding protein for β-

lactam antibiotics, therefore limiting their ability to inhibit cell wall biosynthesis (Hartman

& Tomasz, 1984, Utsui & Yokota, 1985). The exact mechanism of intermediate

vancomycin resistance has not been conclusively determined, but it is thought to be

associated with cell wall thickening, ultimately reducing the diffusion of vancomycin to

the division septum active site (Howden et al., 2010, Pfeltz et al., 2000, Sieradzki &

Tomasz, 2003). Full vancomycin resistance is caused by alteration in the peptidoglycan

biosynthetic pathway by replacement of the D-Ala-D-Ala dipeptide with D-Ala-D-Lac

(Gonzalez-Zorn & Courvalin, 2003, Severin et al., 2004). This resistance is encoded by

the vanA operon, which was acquired by horizontal gene transfer from enterococcus (de

Niederhausern et al., 2011, Zhu et al., 2008, Zhu et al., 2013). Antibiotic resistance has

clearly added to the success of S. aureus as a pathogen, but other dominant

contributors include its plethora of virulence factors and metabolic versatility, as

discussed in the following sections.

Staphylococcus aureus Virulence Determinants

While S. aureus is primarily considered a nasal colonizer, it has the ability to

infect most tissue and organ systems including skin and soft tissue (impetigo, folliculitis,

abscess), blood (bacteremia), heart valve (endocarditis), lungs (pneumonia), bone/bone

marrow (osteomyelitis), and the central nervous system (meningitis)(Archer, 1998,

Richardson). To be an effective pathogen S. aureus has evolved to deal with the

prodigious immune onslaught present in its natural environment. This bacterium

synthesizes a large number of cell surface and secreted virulence proteins that allow it

to successfully colonize and infect the host.

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Factors for colonization and dissemination

Overcoming the skin barrier. The skin is the primary barrier against bacterial

infection, but S. aureus is particularly successful at causing a variety of skin infections

including impetigo, cellulitis, folliculitis, subcutaneous abscesses, and infected ulcers

and wounds (McCaig et al., 2006, Miller & Kaplan, 2009). The upper epidermal layers

act as a physical barrier consisting of a microbial-limiting high-salt concentration, and

low temperature and pH (Grice et al., 2009, van der Merwe et al., 2002). Staphylococci

encounter salt concentrations up to 60 mM in sweat (van der Merwe et al., 2002), but

are well known for their salt tolerant nature and are routinely isolated in the clinical lab

on selective media containing 7.5% (1.3 M) NaCl (Parfentjev & Catelli, 1964, Chapman,

1945). The mechanism of their salt tolerance hinges on preserving a high intracellular

potassium concentration, allowing the bacteria to maintain osmotic homeostasis (Price-

Whelan et al., 2013, Gries et al., 2013). Potassium uptake systems appear to be

essential for S. aureus to cope with osmotic stress caused by NaCl, although S. aureus

can still grow in high salt upon deletion of the two main potassium uptake systems, ktr

and kdp (Price-Whelan et al., 2013, Gries et al., 2013). Another innate factor that S.

aureus must overcome during skin colonization is acidic pH. On the skin surface,

fillaggrin is naturally broken down into urocanic acid and pyrrolidone carboxylic acid

(Barrett & Scott, 1983) leading to a localized decrease in pH (Miajlovic et al., 2010).

Breakdown to pyrrolidone carboxylic acid occurs by a non-enzymatic process; whereas,

urocanic acid is catalytically generated by histidase (Scott, 1981). S. aureus is likely

able to circumvent this pH barrier under certain conditions, as prolonged skin covering

(i.e., by wound dressings) results in elevated pH, favoring S. aureus growth (Aly et al.,

1978). Some portions of the skin are partially occluded naturally, such as the groin,

21

axillary vault (armpit), and toe web; providing these sites with the ideal temperature,

humidity, and pH for S. aureus growth (Grice et al., 2009, Roth & James, 1988).

Adherence and MSCRAMMs. The host extracellular matrix (ECM) is a complex,

biologically active tissue that serves in a structural capacity, while also promoting cell-

cell adhesion, migration, proliferation, and differentiation of host cells (reviewed in

(Halper & Kjaer, 2014, Mienaltowski & Birk, 2014)). This substrate not only provides a

surface for host cell adhesion, but also for the attachment of microorganisms. As the

name implies, microbial surface components recognizing adhesive matrix molecules

(MSCRAMMs) have classically been described as adhesion molecules that promote

binding to components of the ECM including fibrinogen, fibronectin, and collagen (Patti

et al., 1994a). These proteins contain two adjacent IgG containing subdomains in their

N-terminal region, which allow for a common mechanism of ligand binding

(Deivanayagam et al., 2002, Zong et al., 2005). Arguably the most well studied

MSCRAMMs include the fibrinogen-binding clumping factors (CflA and CflB), fibronectin

binding proteins (FnBPA and FnBPB), and the collagen adhesin protein (Cna). ClfB

specifically contributes to nasal colonization as it was confirmed to bind the structural

protein components of squamous epithelial cells, cytokeratin 10 and loricrin (Walsh et

al., 2004, Mulcahy et al., 2012). ClfA and B were both shown to act as important factors

in S. aureus-associated endocarditis due to their ability to bind thrombi, a blood clot

commonly formed in response to injury (Moreillon et al., 1995, Entenza et al., 2000). In

addition to binding fibronectin, FnBPA and FnBPB can also bind both the C-terminal γ

region of fibrinogen, and elastin (Keane et al., 2007a, Keane et al., 2007b, Peacock et

al., 1999, Burke et al., 2011). FnBPs mediate adherence to the host epithelium, but

22

have also been shown to play a major role in host cell invasion. Internalization by FnBP-

host cell interaction has been confirmed for epithelial cells, endothelial cells, fibroblasts,

osteoblasts, and keratinocytes by binding to the host cell receptor, integrin α5β1 (Sinha

et al., 1999, Kintarak et al., 2004, Ahmed et al., 2001). Collagen is the most abundant

protein found in humans and acts as the main component of connective tissue, while

providing structural support and scaffolding for ECM assembly (reviewed here)(Arnold &

Fertala, 2013, Di Lullo et al., 2002). The S. aureus Cna binds collagen by a unique

"collagen hug" mechanism in which multi-domain collagen binding proteins are able to

bind the extended rope-like collagen ligand (Zong et al., 2005). As would be expected,

Cna was found to be a virulence determinant in infections where collagen is abundant,

such as septic arthritis and osteomyelitis (Patti et al., 1994b). MSCRAMM proteins

provide S. aureus with a promiscuous ability to bind host factors for colonization,

internalization, and/or infection.

Surviving nutritional immunity. In an attempt to limit bacterial infection, the

host sequesters essential nutrients in a process termed "nutritional immunity". Iron

sequestration is the most well characterized example of nutritional immunity during

staphylococcal infection in which iron is maintained by host binding proteins

intracellularly (ferritin, hemoglobin, heme-containing enzymes), or complexed with

secreted factors such as transferrin and lactoferrin (Reviewed in (Ong et al., 2006,

Wooldridge & Williams, 1993, Otto et al., 1992)). NEAr iron Transport (NEAT) family

proteins IsdA, IsdB, and IsdH are heme binding and transport proteins that contain

characteristic hemoglobin and/or heme-binding near iron transporter motifs (Grigg et al.,

2007, Vermeiren et al., 2006, Gaudin et al., 2011, Torres et al., 2006, Pilpa et al., 2009,

23

Fonner et al., 2014, Pishchany et al., 2014). Full internalization of iron by S. aureus is

thought to require the iron surface determinant (Isd) system, encoded by up to 5

operons: isdA, isdB, isdCDEFsrtBisdG, isdH, and orfXisdI (Skaar & Schneewind, 2004,

Maresso & Schneewind, 2006). In brief, the current model for Isd-mediated heme import

proposes that IsdA, IsdB, and IsdH are cell wall associated protein receptors that bind

heme, pass it to IsdC, where IsdC then transports heme through the cell wall to the

membrane localized IsdDEF ABC transport system (Muryoi et al., 2008, Liu et al., 2008,

Grigg et al., 2007, Mazmanian et al., 2003, Hammer & Skaar, 2011). Once heme is

inside the cytoplasm, IsdG and IsdI are hemeoxygenases that cleave the tetrapyrrol ring

structure of heme and release free iron to be used in cellular processes (Skaar et al.,

2004, Wu et al., 2005).

Dissemination. Transition from colonization and the primary infection site to full

bacteremia and secondary infection sites requires specific virulence mechanisms.

Secreted hemolysins, toxins, and enzymes facilitate tissue destruction and

dissemination, but secreted proteases and phagocytosis by host macrophages are

thought to be the primary contributors to S. aureus dissemination (Koziel & Potempa,

2013, Kubica et al., 2008). Proteases control this transition by affecting the stability and

and/or processing of bacterial cell surface proteins. A classic example is cleavage of

FnBPs by the S. aureus V8 serine protease, leading to loss of adhesion and

contributing to deeper invasion of tissues (McGavin et al., 1997). Alternatively, S.

aureus can persist for several days within macrophages and, therefore, hitchhike to

various sites within the host (Kubica et al., 2008). This ability to survive within

24

macrophages is thought to be a primary contributor to S. aureus systemic

dissemination.

Strategies for S. aureus immune evasion

Limitation of immune cell recruitment. A hallmark of S. aureus skin infection,

and requirement for bacterial clearance, is the recruitment of neutrophils to the site of

infection (Molne et al., 2000, Kim et al., 2011). Neutrophils and other leukocytes (white

blood cells) are recruited to infection sites following production of cytokines (immune

signaling peptides) or chemokines (chemo-attractant cytokines)(Griffith et al., 2014).

Interference with host chemokine functions is the primary way that S. aureus subverts

neutrophil recruitment. This leads to disruption of innate immune response kinetics,

delaying the immune response, and favoring bacterial survival. Many S. aureus strains

inhibit neutrophil recruitment via the chemotaxis inhibitory protein of staphylococci

(CHIPS)(de Haas et al., 2004, Veldkamp et al., 2000). CHIPS acts by binding to the

C5aR and formyl peptide (FPR) receptors present on leukocytes (Postma et al., 2004).

Binding blocks signal transduction and leads to a decrease in neutrophil migration.

Staphylococcal superantigen-like (SSL) proteins are a family of exoproteins that share

structural similarity with staphylococcal superantigens, but exhibit no superantigenic

activity. The staphylococcal superantigen-like protein 5 (SSL5) can also limit leukocyte

activation/migration by chemokines, via competitive binding to multiple chemokine

receptors (Bestebroer et al., 2009). Binding has potent downstream anti-inflammatory

effects, leading to decreased leukocyte extravasation. The extracellular adherence

protein (Eap) is another protein produced by S. aureus that interferes with leukocyte

migration, and in turn promotes impaired wound healing at the site of S. aureus infection

(Athanasopoulos et al., 2006). Neutrophil adhesion to endothelial cells, transendothelial

25

migration, and overall inflammation are inhibited by staphylococcal Eap

(Athanasopoulos et al., 2006, Haggar et al., 2004, Chavakis et al., 2002). While

neutrophil recruitment is of utmost importance for bacterial clearance during S. aureus

infection, this bacterium has developed multiple proteins that disrupt the critical

chemokine circuitry necessary for recruitment of cell-mediated innate immunity.

Inhibition and evasion of complement. The human complement system is a

powerful tool that allows immediate recognition of invading pathogens, leading to cell-

mediated innate immune responses as well as activation of adaptive immune

components (Kemper & Atkinson, 2007, Carroll, 2004, Zipfel, 2009). Complement can

be considered a bridge between innate and adaptive immunity as it is composed of

constitutively circulating proteins, but commonly requires recognition by antibodies, a

component of the adaptive immunity. Control of the complement system by S. aureus

occurs at each step of the pathway, but the primary interference occurs by inhibiting

complement activation. Inhibition of complement activation is conferred by various

proteins and primarily acts by binding human immunoglobulins, thereby inhibiting

classical pathway activation. IgG binding proteins include staphylococcal protein A

(Spa), second binder of immunoglobulin (Sbi), and staphylococcal superantigen-like

protein 10 (SSL10)(Zhang et al., 1998, Itoh et al., 2010, Hartleib et al., 2000). Protein A

is the first and arguably most well studied staphylococcal surface protein that is nearly

ubiquitous in S. aureus strains (Peacock et al., 2002, Shakeri et al., 2014, Mitani et al.,

2005). In addition to IgG, Spa has been found to bind a multitude of Igs including the

heavy chain constant region of IgG antibodies (Fc), as well as the Fab regions of VH3

type receptors, which are present on approximately 30-50% of circulating B cells in

26

humans (Gouda et al., 1998, Hillson et al., 1993, Sasso et al., 1989, Sasso et al., 1991,

Sasano et al., 1993, Roben et al., 1995, Lindmark et al., 1983, Deisenhofer, 1981,

Cedergren et al., 1993). Binding of Spa to the Fc region of immunoglobulins inhibits

complement activation and opsonization, whereas binding to the Fab region of B cell

receptors (BCRs) leads to B cell superantigen activity.

S. aureus toxins

A well studied group of pore-forming toxins in S. aureus are the hemolysins (Hla

(α-toxin), Hlb, Hld, which bind to the host cell surface, forming a β-barrel

transmembrane pore, allowing for uncontrolled ion transport and cell death through an

aqueous channel (Menestrina, 1986, Menestrina et al., 2001, Song et al., 1996). This

effect appears to be promiscuous for most host cells. Leukotoxins are another type of

pore-forming toxin produced by S. aureus characterized by their canonic bi-component

nature, in which two proteins oligomerize to form a β-barrel pore structure (Kaneko &

Kamio, 2004, Nguyen et al., 2002). Characterization has mostly been completed on

Panton-Valentine leukocidin (PVL), but the pore-forming ability and general structure

are thought to be similar for all leukotoxins (Guillet et al., 2004, Pedelacq et al., 1999,

Olson et al., 1999). PVL, composed of the LukF-PV and LukS-PV subunits, was found

to be specifically associated with recurrent skin and soft tissue infections as well as

necrotizing pneumonia (Masiuk et al., 2010, Monecke et al., 2007, Lina et al., 1999). Hla

and leukocidins directly and indirectly limit the amount of circulating cells by forming

pores in T lymphocytes, causing lysis and death (Berube & Bubeck Wardenburg, 2013,

Alonzo et al., 2013, Nygaard et al., 2012). Leukotoxin ED (LukED) can also bind to the

CCR5 receptor on T cells, macrophages, and dendritic cells leading to destruction of

these cell lines (Alonzo et al., 2013). S. aureus produces multiple superantigen (SAg)

27

proteins that elicit an enhanced immune response by inducing non-specific activation of

T cells, resulting in polyclonal T cell activation and massive cytokine release (Xu &

McCormick, 2012). A canonic SAg produced by S. aureus is toxic shock syndrome toxin

1 (TSST-1), which upon exposure often leads to toxic shock syndrome and death

(Fraser & Proft, 2008, Holtfreter & Broker, 2005, McCormick et al., 2001). SAgs are also

commonly named enterotoxins for their ability to cause staphylococcal food poisoning,

marked by excessive vomiting and diarrhea. The dramatic response of SAgs is likely

due to the conserved structure of superantigen targeting to MHC molecules and T cell

receptors on the surface of T lymphocytes. SAgs activate a large proportion of the T

lymphocyte pool simultaneously, resulting in a "cytokine storm". Although the effects of

T cell antigens on T cell activation are well studied, the evolutionary advantage for S.

aureus is still not clear. The current opinion suggests that there is a refractory period

after the "cytokine storm" in which T cells cannot be activated and many of them die.

Therefore, T cells that would normally be activating the B cell response are eliminated

by the pathogen, creating what has been deemed as an immunogenic "smoke screen"

(Fraser et al., 2000).

Regulation of virulence factors and summary of virulence strategies

Virulence factor gene expression is controlled by a complex array of regulators

and two-component systems, but the most characterized is the accessory gene

regulator (Agr) quorum sensing locus (Wang & Muir, 2016). The Agr quorum sensing

system is comprised of the agrBDCA operon and divergent RNAIII-encoding gene.

Transcription of the agr operon and RNAIII is driven by the P2 and P3 promoters,

respectively. The autoinducing peptide is cleaved from AgrD (Ji et al., 1995, Thoendel &

Horswill, 2009) and exported by AgrB to form a thiolactone ring (Ji et al., 1997). AgrAC

28

makes up a two-component system where AgrC senses the thiolactone ring (Ji et al.,

1995), phosphorylates the AgrA response regulator, and ultimately leads to induction of

both P2 and P3 promoter expression (Novick et al., 1993, Novick et al., 1995). This

further promotes expression of its own genes as well as expression of the RNAIII

transcript. The regulatory effector of this system is the RNAIII molecule, which primarily

controls expression of virulence factor genes by base-pairing to the 5' end of virulence

factor mRNAs (Novick et al., 1995, Novick et al., 1993, Huntzinger et al., 2005,

Chevalier et al., 2010, Boisset et al., 2007). Upon entrance into late exponential phase

S. aureus secretes proteases, hemolysins, exoenzymes and superantigens, while

down-regulating cell wall associated factors; many of these processes are controlled by

Agr (Dinges et al., 2000, Rothfork et al., 2003, Wright & Holland, 2003). Essentially Agr

mediates a density dependent phenotype conversion from tissue-adhering to tissue-

damaging, while regulating genes for immune cell evasion during both phases of

growth. The Agr sysem is critical for pathogenesis (Abdelnour et al., 1993, Gong et al.,

2014, Kielian et al., 2001, Schwan et al., 2003) and, interestingly, Agr-mediated

expression of secreted proteases is thought to somewhat account for the elevated

virulence of some CA-MRSA strains (Kolar et al., 2013). Not only is Agr critical for

virulence factor regulation, but this system also controls the time-dependent expression

of genes associated with biofilm development and dispersal. Specifically, Agr controls

attachment and evasion genes for promotion of biofilm development during lag and

exponential phases of growth, while promoting biofilm dispersal via expression of

proteases during later growth phases (Boles & Horswill, 2008, Yarwood et al., 2004,

Kong et al., 2006). As well, the agr locus is clinically important for promoting the

29

spontaneous development of virulence factor variants in S. aureus biofilms (Yarwood et

al., 2007)

Metabolism of Staphylococcus aureus

Overview

While virulence factors are paramount to S. aureus pathogenesis, the diverse

metabolism of this organism is also critical for its success as a pathogen. As a

facultative anaerobe, the metabolism of S. aureus is highly fluid, retaining the ability to

fluctuate between aerobic and anaerobic respiration and/or mixed acid fermentation

(Liebeke & Lalk, 2014, Somerville & Proctor, 2009a). In fact, S. aureus metabolism is

predicted to be one of the most complex in terms of estimated metabolite numbers

(Liebeke & Lalk, 2014). Using genome-scale reconstruction of metabolic networks

comparing 13 S. aureus strains, researchers predicted approximately 1250 potential

metabolic reactions and 1400 metabolites (Lee et al., 2009). In comparison, small

genome containing bacteria such as Mycoplasma pneumonia and Mycoplasma

genitalium are predicted to produce only 150 and 270 metabolites, respectively (Suthers

et al., 2009, Maier et al., 2013). S. aureus can utilize several major metabolic pathways

including complete glycolytic (Embden-Meyerhof-Parnas), pentose phosphate, and

tricarboxylic acid (TCA) pathways. This bacterium also has major metabolic pathways

for fermentation as well as a complex branched respiratory chain with a variety of

components.

Carbohydrates are primarily catabolized through the glycolytic and pentose

phosphate pathways, with every molecule of glucose producing two molecules each of

NADH and pyruvate (Cohen, 1972). The fate of pyruvate is then determined by the

growth conditions, phase of growth, and is particularly dependent on the availability of

30

O2 (Somerville & Proctor, 2009a). Under glucose rich aerobic conditions, S. aureus

utilizes catabolite repression and suppresses the TCA cycle and pentose phosphate

pathway (discussed below)(Somerville et al., 2002, Strasters & Winkler, 1963,

Somerville et al., 2003b). These conditions induce a fermentative metabolism where

pyruvate is primarily converted to acetate using the acetate kinase pathway, producing

ATP and helping to maintain redox balance (Collins & Lascelles, 1962, Somerville et al.,

2002, Somerville et al., 2003b, Strasters & Winkler, 1963, Sadykov et al., 2013). Once

glucose becomes limiting, acetate is shuttled back through the TCA cycle to generate

reduced dinucleotide cofactors (NADH, FADH2, and NADPH), further supporting aerobic

respiration and biosynthetic pathways (Somerville et al., 2003b). Transition into

late/post-exponential phase growth induces the TCA cycle, which dramatically alters the

metabolome and leads to increased availability of biosynthetic precursors and maximal

expression of virulence factors (Novick, 2000, Vandenesch et al., 1991, Ji et al., 1995).

When O2 is present, aerobic respiration can be supported by glycolysis or TCA

cycle-generated NADH; but when O2 is limiting and nitrate (NO3-) is present S. aureus

can anaerobically respire using a nitrate reductase (Chang & Lascelles, 1963, Burke &

Lascelles, 1975). In the absence of an alternative terminal electron acceptor, anaerobic

growth conditions primarily promote pyruvate reduction to lactic acid, which helps to

maintain redox balance (Pagels et al., 2010, Ferreira et al., 2013). Multiple links

between metabolism and virulence have been described in S. aureus (discussed

below), therefore understanding these processes may lead to novel treatments for S.

aureus infection.

31

Low Oxygen Fermentative Metabolism

Metabolism is often dictated by the presence of final electron acceptors such as

O2 and NO3-, which are required for proper respiratory chain function. When there are

low levels of both of these molecules, S. aureus employs a fermentative metabolism

which ultimately dictates the fate of pyruvate conversion to one or more end-products

(Figure 1-1). Under these conditions the bacterium primarily produces L-lactate, but also

small amounts of D-lactate, acetate, ethanol, formate, and 2,3-butanediol (Ferreira et

al., 2013, Richardson et al., 2008). Combined with genome examination of potential

fermentation pathways, this suggests that when grown without O2, S. aureus undergoes

a mixed acid fermentative metabolism, allowing ATP production by substrate level

phosphorylation while also maintaining redox balance via fermentation. In support of

this, anaerobic gene and protein expression was previously examined in S. aureus

using both proteomic and transcriptomic approaches. In the absence of O2 and NO3-

(alternative terminal electron acceptor), an induction of glycolytic enzymes, combined

with low levels of TCA cycle proteins, was observed (Fuchs et al., 2007). Fermentation

enzymes such as both lactate dehydrogenase 1 and 2 (Ldh1 and Ldh2), alcohol

dehydrogenases (AdhE and Adh), acetolactate decarboxylase (BudA1), acetolactate

synthase (BudB), and acetoin reductase were all present at higher levels under these

anaerobic growth conditions. Additionally, genes associated with lactate and formate

secretion, as well as expression of pyruvate formate lyase (pfl) were more highly

expressed when O2 was limited. Pfl reversibly converts pyruvate and coenzyme-A

(CoA) into formate and acetyl-CoA, with acetyl-CoA catabolism able to promote acetate

and ethanol fermentation (Leibig et al., 2011)(Figure 1-1). Another important

contribution of Pfl under anoxic conditions is generation of formate, which is needed for

32

biosynthesis of formyl-tetrahydrofolate and subsequent purine and protein synthesis

(Leibig et al., 2011). Curiously, even when NO3- was not present, genes for both nitrite

(NO2-) reduction (nirD) and NO3

- respiration (narH, narI, narJ) were upregulated

suggesting a role for these genes in general response to low O2 conditions (Fuchs et

al., 2007).

While S. aureus employs multiple fermentation pathways under low O2 growth, L-

lactate production is the major metabolic pathway and functions in both redox balance

and energy production (Sun et al., 2012, Pagels et al., 2010, Richardson et al., 2008,

Richardson et al., 2006). S. aureus codes for 3 lactate dehydrogenases (ldh1, ldh2,

ddh) that can interconvert pyruvate and lactate, with direction of lactate production

being favored (Richardson et al., 2008). Conversion of pyruvate to lactate regenerates

NAD from NADH, but it is predicted that reversal of the reaction could allow for lactate

utilization as a carbon source (Fuller et al., 2011). Indeed, production and utilization of

lactate is a common theme in S. aureus metabolism and the efficient catabolism of

lactate is thought to endow S. aureus with a metabolic advantage in its ecological niche

(Ferreira et al., 2013).

Amino Acid Metabolism

In comparison to glucose metabolism, fewer studies have been published on

specific amino acid catabolic pathways in S. aureus. However, phenotypic studies using

chemically-defined media have been undertaken to determine the absolute amino acid

requirements for S. aureus (Emmett & Kloos, 1975, Mah et al., 1967, Nychas et al.,

1991, Onoue & Mori, 1997, Taylor & Holland, 1989). These studies provided mixed

results, but it was ultimately determined that S. aureus possesses multiple in vitro

auxotrophies that vary between 3 and 12 amino acids; the most frequently required

33

being Pro, Arg, Val, and Cys. Interestingly, these auxotrophies are likely not due to the

absence of biosynthetic pathways because whole-genome sequencing has confirmed

that biosynthetic pathways exist for all of these amino acids (Baba et al., 2008, Baba et

al., 2002, Diep et al., 2006, Gill et al., 2005, Holden et al., 2004, Kuroda et al., 2001).

This discrepancy between in vitro auxotrophies and the presence of all biosynthetic

pathways is likely due to differences in metabolic requirements between in vitro and in

vivo growth conditions.

For central amino acid catabolism, amino acids enter the TCA cycle via

metabolic intermediates, producing biosynthetic precursors and reducing equivalents for

aerobic respiration. Mutation of the TCA cycle aconitase enzyme shows that a

functioning TCA cycle is required for amino acid utilization (Somerville et al., 2002).

Disruption of TCA cycle activity by an aconitase mutation halted growth, and prevented

both ammonia accumulation and depletion of free amino acids when other carbon

sources had become limiting (Somerville et al., 2002). Most amino acids, including Arg

enter the TCA cycle through biosynthetic intermediates. Studies on Arg biosynthesis in

S. aureus have uncovered two important catabolic pathways that can feed the urea

cycle and then the TCA cycle. Specifically, in silico analysis predicted Arg biosynthesis

from catalysis of either Glu or Pro, with genes for each pathway present in S. aureus

(Nuxoll et al., 2012). Under in vitro growth conditions S. aureus preferentially utilizes a

novel Pro catabolic pathway (via PutA and RocD), whereas Glu catabolism (via

ArgBCDJ) was found to be critical in a mouse kidney abscess model of infection (Nuxoll

et al., 2012). The importance of Arg metabolism to the success of S. aureus as a

pathogen is underscored by the contribution of the arginine catabolic mobile element

34

(ACME) to the pathogenesis of CA-MRSA strains. ACME codes for an arginine

deiminase system (Arc) and was found to be particularly important for survival of

USA300 (CA-MRSA lineage) in acidic environments that mimic human skin (Thurlow et

al., 2013). The branched chain amino acids (BCAAs) Leu,Ile, and Val are also critical for

S. aureus growth and virulence by playing important roles in protein synthesis,

precursors for branched-chain fatty acids, and as co-regulators (with CodY, described

below) of virulence factor synthesis (Majerczyk et al., 2010, Pohl et al., 2009, Shivers &

Sonenshein, 2004, Kaiser et al., 2016, Kaiser et al., 2015). While S. aureus has genes

for biosynthesis of these amino acids, it prefers to repress Leu and Val biosynthesis

pathways and import BCAAs from the extracellular amino acid pool (Kaiser et al., 2016).

The primary BCAA transporter able to transport all three representative metabolites is

BrnQ1, with BrnQ2 and BcaP having subsidiary roles (Kaiser et al., 2015, Kaiser et al.,

2016).

Respiratory Metabolism

General respiratory chain components

As mentioned above, S. aureus has access to a complex respiratory chain

allowing it to respire on multiple electron donors and can utilize both aerobic (O2) and

anaerobic (NO3-, NO2

-, NO) final electron acceptors (Figure 1-2). The purpose of the

respiratory chain is to generate a proton motive force that provides energy for synthesis

of ATP and transport processes. Similar to most organisms, the TCA cycle of S. aureus

generates large amounts of reduced cofactors that provide electrons for translocation of

protons across the membrane. These diffusible carriers (NADH, lactate, succinate,

malate)(discussed below) donate electrons to the respiratory chain, which is composed

of a range of electron transferring redox cofactors such as flavins, iron-sulfur (Fe-S)

35

clusters, heme and copper centers, all of which are bound to integral membrane or

membrane associated protein complexes (Simon et al., 2008). Shuttling of electrons

between protein complexes requires membrane associated quinone molecules (Simon

et al., 2008), of which S. aureus only synthesizes menaquinone for all of its quinone

requirements (Wakeman et al., 2012). As the electrons move down the respiratory

chain they start with a low electrochemical potential and flow "downhill" in terms of

energy, producing free energy (ΔG) that is used to translocate protons across the

membrane (Mitchell, 1961, Mitchell, 2011). This generates a transmembrane

electrochemical gradient, or proton motive force (pmf) characterized by both a chemical

(ΔpH) and electrical (ΔΨ) component, which is ultimately harnessed by ATP synthase to

produce ATP.

Aerobic respiratory components

As described above, aerobic respiration requires reducing equivalents to donate

electrons to the respiratory chain, generating a pmf, and ending with reduction of O2 as

the final electron acceptor. NADH is generally considered the primary product of the

TCA cycle and therefore the primary donor to drive respiration through oxidation by

NADH dehydrogenase (Ndh). In general, bacteria have access to three distinct

NADH:quinone oxidoreductases including complex I, NDH-2, and a Na+ pumping Nqe

complex (Angerer et al., 2012, Feng et al., 2012, Juarez & Barquera, 2012, Efremov &

Sazanov, 2011). Complex I NADH:quinone oxidoreductases (nuoAB, nuoCEF,

nuoGHIJKL) are the classic example observed in E. coli which contains 6 subunits and

55 transmembrane helices (Efremov & Sazanov, 2011). While E. coli synthesizes both

complex I and NDH-2 (Villegas et al., 2011), genomic examination of S. aureus

suggests that it does not have the cellular machinery for a full complex I (Schurig-

36

Briccio et al., 2014). Potential partial complex I homologs present in S. aureus are

decribed below. NDH-2 is a single 50 kDa protein with a non-covalently-bound FAD

cofactor. This family of enzymes transfers electrons to FAD and then to membrane

bound quinones, ultimately helping to maintain cellular redox balance and indirectly

contributing to the pmf (Kerscher et al., 2008).

Two NDH-2 proteins were recently characterized in S. aureus denoted NdhC and

NdhF, with NdhC determined to be the dominant NADH:oxidoreductase in S. aureus

(Schurig-Briccio et al., 2014). A separate study confirmed that activity of the S. aureus

NDH-2 is rate limited by quinone reduction, effectively confirming that electrons are

donated to the quinone pool for NADH-driven respiration (Sena et al., 2015). While a full

complex 1 homolog has not been found in S. aureus, two separate studies have

identified “NuoL-like” proteins with homology to the type 1 NADH dehydrogenase of E.

coli (Mayer et al., 2015, Bayer et al., 2006). The mnhABCDEFG operon was found to

contribute to membrane potential in S. aureus, but NADH oxidation was not confirmed

(Bayer et al., 2006). Alternatively, mutagenesis studies confirmed a role for the mpsABC

operon in NADH oxidation as well as maintenance of membrane potential and O2

consumption (Mayer et al., 2015). Overall, S. aureus likely synthesizes multiple proteins

that can oxidize NADH for respiration, although the specific biological relevance for

each has not been fully determined.

Other electron donors can also drive respiration when their cognate respiratory

oxidoreductase proteins are present. Genomic examination shows that S. aureus has

genes encoding succinate (sdhCAB/complex II) and malate (mqo1) dehydrogenases, as

well as a lactate quinone oxidoreductase (lqo). A staphylococcal protein with electron

37

paramagnetic resonance (EPR) relaxation and redox properties similar to mitochondrial

succinate dehydrogenases was discovered in 1991 (Solozhenkin et al., 1991). It wasn’t

until 2010 that biochemical characterization was completed on SdhCAB showing that

this protein contributed to TCA cycle function and was upregulated in S. aureus biofilms

(Gaupp et al., 2010). As well, respiration can be driven in membrane fractions using

succinate as the primary electron donor (Schurig-Briccio et al., 2014). Until recently the

S. aureus genome was annotated with two malate quinone oxidoreductases (mqo1,

mqo2)(Fuller et al., 2011). Mqo1 was confirmed to be a malate oxidizing enzyme and

required for maximal growth on amino acids (Fuller et al., 2011, Spahich et al., 2016).

This underscores its significance in assimilation of amino acids through the TCA cycle,

and furthermore, an mqo1 mutant secreted excess lactate and acetate, suggestive of

overflow metabolism (Spahich et al., 2016). However, it has not been confirmed

whether Mqo1 directly donates electrons to the respiratory chain. In 1969 NAD-

independent L-lactate dehydrogenase activity was confirmed in S. aureus (Stockland &

San Clemente, 1969). Given that malate and L-lactate are structurally similar, Fuller et.

al., predicted that the misannotated mqo2 may code for a lactate oxidizing enzyme; and

indeed the misannotated mqo2 was in fact a lactate quinone oxidorductase (Lqo) (Fuller

et al., 2011). This enzyme specifically oxidizes L-lactate to pyruvate while subsequently

reducing the quinone pool (Fuller et al., 2011). Lqo supports the respiratory chain by

providing electrons for proton translocation (Fuller et al., 2011) and can contribute to the

terminal reduction of O2, ferric iron, or NO3-(Theodore & Weinbach, 1974, Lascelles &

Burke, 1978, Tynecka & Malm, 1995, Fuller et al., 2011). Moreover, the type aa3 quinol

38

oxidase (Qox) was shown to be required for L-lactate driven respiration (Tynecka et al.,

1999).

The final step in the aerobic respiratory chain is the electron mediated reduction

of O2 to H2O, mediated by heme-dependent terminal oxidases (White et al., 1995). In

many biological systems complex III (the quinone:cytochrome c oxidoreductase or bc1

complex) is the intermediary complex between the upstream respiratory

dehydrogenases and terminal oxidase. S. aureus does not appear to encode a complex

III (Schurig-Briccio et al., 2014), but instead has two menaquinol terminal oxidases, a

type aa3 (qoxABCD) and bd-type (cydAB)(Hammer et al., 2013). Mutation of either of

these enzymes decreases membrane potential, with a double mutant almost completely

eliminating the membrane potential of S. aureus (Hammer et al., 2013). This finding

implies a branched respiratory chain and supports the hypothesis that the individual

oxidases may be important under different growth conditions (Hammer et al., 2013).

Anaerobic respiration

The general principles of the respiratory chain also hold true when S. aureus

respires under anaerobic conditions, with the exception of certain specific contributing

respiratory proteins and terminal electron acceptor. In B. subtilis, both NO3- and NO2

-

reductases can promote respiration (Nakano & Zuber, 1998). Few studies have been

completed on the anaerobic respiratory chain proteins in S. aureus, with some evidence

suggesting that S. aureus can respire using respiratory NO3- (Nar) and nitric oxide

reductases (Nor)(Lewis et al., 2015, Burke & Lascelles, 1975). The narGHJI operon is

predicted to code for the S. aureus nitrate reductase, which likely accounts for the

observed reduction of NO3- to NO2

- (Chang & Lascelles, 1963). While predicted, the nar

39

gene product has not been cloned and no structure-function studies have been

completed in S. aureus. With that said, O2 was found to suppress activity and gene

expression of the NO3- reductase (Chang & Lascelles, 1963, Fuchs et al., 2007), while

NO3- promoted activity and expression of the nar operon (Chang & Lascelles, 1963,

Niemann et al., 2014). As well, both dissimilatory NO3- and NO2

- reduction to ammonia

was also confirmed in S. aureus, but the contribution of this process to respiration was

not elucidated (Schlag et al., 2008). Characterization of NO3- reduction is much more

complete in Staphylococcus carnosus and many of the principals likely hold true for S.

aureus. In S. carnosus NO3- uptake is promoted under anoxic conditions and conferred

by narT, the gene coding for a NO3- transporter (Fast et al., 1996). Moreover,

mutagenesis studies confirmed the narGHJI operon functions as a NO3- reductase, with

transcription promoted by anaerobiosis, NO3-, and NO2

- (Pantel et al., 1998).

Interestingly, in the presence of O2 and NO3-, high transcriptional expression of nar was

observed, but cells presented with low NO3- reducing activity (Pantel et al., 1998).

Nitrate reductase was found to be insensitive to O2, therefore, other O2 sensitive steps

such as post-transcriptional mechanisms or molybdenum cofactor biosynthesis must be

affected (Pantel et al., 1998).

S. aureus also contains genes that code for NO2- (nirBD) and NO (nor)

reductases (Schlag et al., 2008, Lewis et al., 2015). Similar to nar, the nir gene has not

been cloned in S. aureus. However, some potential insight can be obtained by looking

at studies in S. carnosus where a nirBD-encoded cytosolic NO2- reductase oxidized

NADH for reduction of NO2- to ammonia (Neubauer & Gotz, 1996, Neubauer et al.,

1999, Pantel et al., 1998). This enzyme was determined to be cytosolic and not

40

contribute to respiration, with its primary functions being 1) detoxification of Nar-derived

NO2- and 2) maintenance of cellular redox status. S. aureus is not predicted to have a

respiratory NO2- reductase, but there is some evidence that Nar can reduce NO2

- and

sustain respiration in S. carnosus (Neubauer & Gotz, 1996). Alternately, respiratory Nor

proteins appear to contribute to respiration in S. aureus. A subset of S. aureus strains

(~37%) contain the nor gene, with evidence that this protein can promote respiration

upon challenge with excess NO (Lewis et al., 2015).

As mentioned above, lactate is one of the main products of fermentation in S.

aureus and, thus, likely promotes anaerobic respiration via Lqo under anoxic conditions.

Evidence for this was observed by confirmation of NO3- reductase activity using lactate

as an electron donor and addition of menaquinone (Lascelles & Burke, 1978, Sasarman

et al., 1974). Further support for lactate driven anaerobic respiration was observed

when S. aureus was unable to grow anaerobically on L-lactate without NO3- addition to

the media (Fuller et al., 2011). Taken together, these data suggest that lactate donates

its electrons to the respiratory chain where they are transferred between protein

complexes by menaquinones and then are finally used to reduce NO3- to NO2

- by Nar.

Genetic Regulation of Staphylococcus aureus Metabolism

Metabolic regulation in S. aureus is complex, but can be generally understood

by examination of a few central regulators. The carbon catabolite protein A (CcpA) is a

well studied mediator of catabolite repression in S. aureus, where the presence of

glycolytic intermediates such as glucose-6-phosphate and fructose-1,6-bisphosphate

leads to repression of a wide variety of genes (Lopez & Thoms, 1977, Schumacher et

al., 2007). These include virulence genes, in which CcpA acts indirectly through RNAIII

and the agr system (Seidl et al., 2009, Seidl et al., 2008b, Seidl et al., 2008a). In

41

addition to virulence genes, CcpA also regulates central metabolic genes including

those associated with glycolytic pathways and the TCA cycle (Seidl et al., 2009).

Another recently studied mediator of carbon catabolite repression is S. aureus is the

carbon catabolite protein E (CcpE). CcpE is a positive regulator of TCA cycle genes

such as citrate synthase (citZ) and aconitase (citB), the first two steps of the TCA cycle

(Hartmann et al., 2013). Inactivation of ccpE represses amino acid catabolism/TCA

cycle activity, and at the same time significantly increases intracellular lactate levels.

Upon TCA cycle inhibition, L-lactate was is an apparent overflow product (Hartmann et

al., 2013).

The redox sensing regulator (Rex) is considered a central regulator of anaerobic

metabolism that is present in many gram positive bacteria (Brekasis & Paget, 2003).

This family of transcriptional regulators sense the NAD/NADH ratio present in the cell

and alter gene expression accordingly (Sickmier et al., 2005, Pagels et al., 2010).

Binding of NADH to Rex de-represses transcription by preventing the association of the

Rex-NADH complex with Rex regulated transcriptional operators. Dissociation of Rex-

NADH from repressor sites allows transcription of genes for electron transport chain

synthesis (hemE, narG), regulators of anaerobic and nitrogen metabolism (nirR, vicR,

srrA), as well as fermentative and anaerobic metabolism genes (adhE, adh1, pflB, arcA,

ldh1, ldh2)(Pagels et al., 2010). Aerobic and anaerobic growth conditions have vastly

different redox environments, and therefore Rex allows indirect sensing of O2

availability. Responding to the redox status of the cell not only allows S. aureus to

respond to O2 limitation, but also to alterations in overall metabolism such as nutrient

limitation, membrane disruption, or oxidative damage.

42

Regulation of the full metabolic network for amino acid metabolism is currently a

topic where more research is required. While not well studied in S. aureus, the

glutamine synthetase repressor (GlnR) is a conserved regulator in Gram positive

bacteria, including S. aureus (Schreier et al., 1989, Schreier et al., 2000). GlnR

regulates expression of glutamine synthetase (glnA), which synthesizes Gln from Glu

and ammonia. This metabolic reaction is key for ammonia assimilation as both Glu and

Gln are major donors of intracellular nitrogen (Anderson & Witter, 1982). Another

conserved regulator for amino acid metabolism in low G+C content Gram positive

bacteria is the GTP and BCAA transcriptional repressor, CodY (Shivers & Sonenshein,

2004, Sonenshein, 2005). Affinity of CodY for DNA increases with both GTP and BCAA

binding (Handke et al., 2008). Val and Leu biosynthesis requires both pyruvate and the

amino group from Glu, each of which are derived from glycolysis and the TCA cycle,

respectively. Therefore, sensing of BCAAs through CodY allows S. aureus to sample

the overall carbon and nitrogen metabolism. While not directly linked, there is some

evidence that lactate metabolism in S. aureus is also associated with TCA cycle activity

and amino acid catabolism. For example, Ldh2 was found to be directly repressed by

CodY, although the biological relevance is currently not understood (Majerczyk et al.,

2010).

Originally discovered as a low O2 responsive two-component system (TCS)

(Throup et al., 2001, Yarwood et al., 2001), the staphylococcal respiratory response

(SrrAB) regulator is homologous to the ResDE system in B. subtilis (Nakano et al.,

1996). SrrB is the membrane bound sensor histidine kinase that phosphorylates the

DNA-binding response regulator (SrrA), leading to transcriptional control of target genes

43

(Pragman et al., 2004, Ulrich et al., 2007). In S. aureus, SrrAB responds to nitrosative

stress and hypoxia, likely by sensing impaired electron flow through the respiratory

chain (Kinkel et al., 2013, Richardson et al., 2006). SrrB has been postulated to be a

direct sensor of respiratory function by sensing the reduction state of the quinone pool

(Kinkel et al., 2013). The exact mechanism of sensing is unknown, but SrrB is predicted

to have a heme-containing PAS domain, which are well characterized to be internal

sensors for O2 and redox potential (Taylor & Zhulin, 1999). In Bacillus, the PAS domain

of ResE is critical for NO-induced signal transduction and gene expression, but it is still

unknown whether NO is directly interacting with the PAS domain or if sensing an

indirect signal (Baruah et al., 2004). Studies on a srrAB mutant showed that this TCS

affects expression of genes involved in cytochrome biosynthesis and assembly

(qoxABCD, cydAB, hemABCX), anaerobic metabolism (pflAB, adhE, nrdDG), iron-sulfur

cluster repair (scdA), and NO detoxification (hmp) during nitrosative stress in S. aureus

(Kinkel et al., 2013). As well, SrrAB regulates both virulence gene expression (via RNA

III)(Pragman et al., 2004, Pragman et al., 2007) and biofilm formation (Ulrich et al.,

2007, Wu et al., 2015). Recently, SrrA was also found to regulate expression of the

RsaE/RoxS small RNA in both B. subtilis and S. aureus (Durand et al., 2015).

Regulation of this sRNA was determined to be important for redox homeostasis in these

bacteria.

Regulation of nitrogen respiratory genes is completed by the NreABC two-

component system, which contains an oxygen sensitive histidine sensor kinase (NreB)

(Schlag et al., 2008). Aerobic conditions cause the reversible loss of the O2 sensitive

Fe-S cluster contained by NreB, preventing phosphorylation. Under anaerobic

44

conditions NreB becomes autophosphorylated, transfers the phosphate to the response

regulator, NreC, which in turns activates transcription of select genes for NO3- and NO2

-

metabolism (nir and nar operons) (Schlag et al., 2008). Loss of the NreABC two-

component system is critical for S. aureus anaerobic respiration and mutation of this

regulatory system removed the ability of the cells to respire on NO3-, and forced S.

aureus into fermentative metabolism (Schlag et al., 2008).

Metabolism and Virulence

Global metabolic regulators and virulence

Multiple studies have implicated a direct link between S. aureus metabolic

activity, virulence, survival, and persistence, making determination of metabolic

mechanisms critical for the development of novel treatments (Somerville & Proctor,

2009a, Chatterjee et al., 2009, Zhu et al., 2009). Many of the major metabolic regulators

that have been characterized in S. aureus contribute to either virulence factor regulation

and/or in vivo virulence. These regulators include the catabolite control protein (CcpA)

(Seidl et al., 2008b, Seidl et al., 2009, Seidl et al., 2008a), CodY (Pohl et al., 2009,

Waters et al., 2016, Roux et al., 2014), Rex (Pagels et al., 2010), and SrrAB (Pragman

et al., 2004, Pragman et al., 2007, Ulrich et al., 2007). Although an extensive body of

published literature links metabolism to S. aureus virulence, review of this topic will

focus on examples specifically relevant to this study such as the role of SrrAB. In

addition to metabolic and stress response genes, SrrA has been shown to regulate

transcription of virulence factors both directly and indirectly. SrrA binds to the promoter

region of icaADBC operon (coding for biosynthesis of polysaccharide intercellular

adhesion, PIA), activating transcription under anaerobic conditions (Ulrich et al., 2007).

Inactivation of PIA biosynthesis increases expression of TCA cycle enzymes such as

45

aconitase, succinate dehydrogenase, fumarase, and NADH dehydrogenase (Throup et

al., 2001). Furthermore, PIA synthesis is associated with decreased TCA cycle activity

(Vuong et al., 2005, Sadykov et al., 2008). Therefore, SrrAB may link virulence factor

synthesis to central metabolism. Another role for SrrAB in virulence factor regulation

occurs by regulating levels of RNAIII, in which RNAIII acts through the Agr system to

control expression of TSST-1 and Spa (Pragman et al., 2007, Pragman et al., 2004).

Link between central metabolism and virulence

Global metabolic regulators are clearly important for virulence of S. aureus, but

more specific genes involved with central metabolism and the TCA cycle have also

been linked to virulence. For example, a signature-tagged mutagenesis screen

identified genes required for survival in a murine bacteriemia model of infection, which

included multiple metabolism-related genes such as those involved in amino acid

biosynthesis (trpABD, lysA, thrB), purine biosynthesis (purL), and the TCA cycle (citB,

odhB) (Mei et al., 1997). In a separate study, mice infected with an aconitase mutant

(acnA) took longer to develop lesions relative to mice infected with the isogenic wildtype

strain (Somerville et al., 2002). A decrease in production of certain virulence factors (α

and β toxins, lipase, type C enterotoxin) was also observed in this acnA mutant strain

(Somerville et al., 2002). Further evidence for the critical role of the TCA cycle in S.

aureus virulence stems from the loss of virulence observed in a mqo1 mutant (Fuller et

al., 2011, Spahich et al., 2016). Disruption of respiratory chain components also causes

some unique virulence phenotypes. Studies on cytochrome mutants (qoxB, cydB) show

differential colonization between organs (Hammer et al., 2013). For example, a cydB

mutant is impaired in murine heart colonization whereas a qoxB mutant is deficient in

murine liver colonization (Hammer et al., 2013). The importance of the respiratory chain

46

to S. aureus virulence is exemplified by studies on respiratory chain mutants in which

mutation of heme or menaquinone biosynthesis genes (von Eiff et al., 1997, Proctor et

al., 1994, Balwit et al., 1994), or mutation of both terminal oxidases (Gotz & Mayer,

2013) leads to a small-colony variant (SCV) phenotype. SCV isolates survive better

within host cells and are associated with altered metabolism and persistent infections

(von Eiff et al., 2001, von Eiff et al., 1997, Proctor et al., 1995).

Lactate as a central virulence metabolite

A common theme in all types of S. aureus metabolism (fermentation, aerobic and

anaerobic respiration) appears to be production and consumption of lactate. Described

in more detail below, the inducible lactate dehydrogenase (Ldh1) is critical for

resistance to nitrosative stress in S. aureus (Richardson et al., 2008). Briefly, Ldh1

provides metabolic flexibility and redox balance when the respiratory chain is inhibited

by host-derived NO. As such, mutation of ldh1 causes reduced mortality and a decrease

in renal lesion size when tested in a murine sepsis model (Richardson et al., 2008).

Moreover, a ldh1/ldh2 double mutant is almost completely avirulent. The respiratory Lqo

enzyme has also been linked to virulence and is believed to work in concert with Ldh to

resemble an alternative (non-proton pumping) NADH dehydrogenase, helping to

maintain redox balance (Fuller et al., 2011). In this scenario Ldh converts pyruvate to L-

lactate, L-lactate is oxidized back to pyruvate by Lqo, and redox balance is maintained

by the net consumption of NADH (Fuller et al., 2011). Similar to Ldh1, Lqo is required

for full virulence in a murine sepsis model (Fuller et al., 2011). SrrAB is probably the

best example of metabolic adaptation in response to host stressors. Upon NO

challenge, SrrAB is predicted to sense the altered respiratory state due to NO inhibition

of cytochromes. To maintain redox balance and energy production, SrrAB turns on

47

genes associated with anaerobic metabolism, including those involved in with

metabolism of lactate (Kinkel et al., 2013, Richardson et al., 2006). This response is

important in vivo as a srrAB mutant kills 70% less mice after 10-days intravenous

infection (Richardson et al., 2006). Importantly, mice lacking inducible NOS (NOS-/-) are

still less susceptible to infection by a srrAB mutant, suggesting that SrrAB contributes to

virulence by additional mechanisms seperate from NO protection (Richardson et al.,

2006). While S. aureus clearly has an adaptive metabolism, it must also have

mechanisms in place to respond to, and protect itself, from reactive metabolic by-

products.

Biochemistry of Reactive Oxygen and Nitrogen Species

General ROS Characteristsics

ROS chemistry and toxicity

Much has been written on the biochemical importance of O2 for complex life, the

evolution of organisms to rely on O2, and the efficiency of energy generating systems

that require O2 (Hsia et al., 2013, Dzal et al., 2015, Archibald & Fridovich, 1983,

Falkowski & Godfrey, 2008). Evolution of microbes to utilize O2 as a part of their

metabolism is critical for the energy demands of complex biological systems. The

chemical properties of O2 make it a useful molecule for many metabolic processes, but

its chemical derivatives are highly toxic to cells (Imlay, 2013). O2 itself is unreactive with

major structural molecules in biology such as amino acids, carbohydrates, lipids, and

nucleic acids. Indeed, the actual toxicity of O2 is derived from formation of partially

reduced ROS (Gerschman et al., 2001). As molecular O2 gains electrons it is first

converted to superoxide (O2-), then hydrogen peroxide (H2O2), hydroxyl radicals (HO),

and finally to water (H2O). Most organisms have methods to detoxify both (O2-) and

48

(H2O2)(discussed below). Molecular O2 contains an even number of electrons, with the

final two residing in discrete orbitals as unpaired, spin-aligned electrons (Imlay, 2003,

Imlay, 2013). These properties make molecular O2 a poor univalent electron acceptor.

Therefore, O2 can only take electrons from strong univalent electron donors such as

metal centres, flavins, and respiratory quinones.

Pathways of ROS generation

Many of the electron donors for ROS generation are prominent respiratory

components, and in fact the flavins of respiratory dehydrogenases appear to be the

primary sources of O2- and H2O2 in bacteria (Minghetti & Gennis, 1988, Messner &

Imlay, 1999, Messner & Imlay, 2002, Kussmaul & Hirst, 2006). While respiratory

cythochromes can generate O2- in some biological systems, this mechanism is generally

refuted as a primary O2- production site in bacteria (Minghetti & Gennis, 1988). Auto-

oxidation of non-respiratory chain flavoproteins is also found to be an additional source

of O2- and H2O2 in bacteria, including glutathione reductase, lipoamide dehydrogenase

glutamate synthase, and flavohemoprotein (Hmp)(Korshunov & Imlay, 2010, Seaver &

Imlay, 2004, Massey et al., 1969, Geary & Meister, 1977, Grinblat et al., 1991, Oogai et

al., 2016, Membrillo-Hernandez et al., 1996, McLean et al., 2010). The exact protein

source(s) of S. aureus ROS production are currently not well defined, but it is assumed

that they are created by similar mechanisms. While both O2- and H2O2 are generated by

flavoproteins, H2O2 can also be produced by the enzymatic detoxification of O2- by

superoxide dismutase (SOD) (Miller, 2012), and is then free to generate HO radicals via

the Fenton reaction. Fenton chemistry produces HO radicals by H2O2 interaction with

free ferrous Iron (Fe2+), and Fenton chemistry has been confirmed in S. aureus (Repine

et al., 1981). Thus, levels of free intracellular iron directly contribute to ROS levels. An

49

exogenous source of ROS encountered by S. aureus in its natural environment is the

oxidative burst produced by phagocytic immune cells such as macrophages and

neutrophils (Slauch, 2011, Chen & Junger, 2012). Oxidative damage occurs when these

reactive molecules attack lipids, proteins, nucleic acids, and Fe-S containing proteins

(Imlay, 2003). A specific example of ROS sensitive proteins are the Fe-S cluster

containing dehydratases which are generally sensitive due to their chemical structure. In

E. coli, multiple key metabolic TCA cycle enzymes are deactivated by O2- including

aconitase A, aconitase B, fumarase A and fumarase B (Gardner, 2002, Gardner &

Fridovich, 1991b, Gardner & Fridovich, 1991a, Liochev & Fridovich, 1992, Flint et al.,

1993). With this in mind, S. aureus has developed multiple mechanisms of resistance to

oxidative stress.

Protection from Oxidative Stress in Staphylococcus aureus

Classical ROS detoxification proteins

All organisms have evolved methods of ROS detoxification and prevention of

oxidative damage. In S. aureus, ROS generation pathways are complemented with

mechanisms for detoxification. Superoxide dismutases are metalloproteins that catalzye

the dismutation of O2- to H2O2 (Karavolos et al., 2003). S. aureus has two SOD

encoding genes, sodA (Clements et al., 1999) and sodM (Valderas & Hart, 2001), with

both being manganese (Mn) dependent enzymes. In vitro data currently supports SodA

as being responsible for the majority of SOD activity (Valderas & Hart, 2001). While the

relevance of SodM is not fully understood, the presence of this protein is somewhat

unique to S. aureus, as coagulase-negative staphylococci do not synthesize SodM

(Valderas et al., 2002). Another primary ROS detoxification protein is catalase, which

converts H2O2 to the biologically inert O2 and H2O (Castro, 1980). Catalase is a well-

50

studied detox protein that is pervasive throughout most biological systems and has

been studied since the early 1900s (Nicholls, 2012). S. aureus contains a single

catalase gene (katA), which is important for survival, persistence, and nasal colonization

(Cosgrove et al., 2007, Martin & Chaven, 1987, Flowers et al., 1977).

Thiol-specific redox systems in Staphylococcus aureus

Thiol-specific redox systems are important for maintaining the intracellular thiol-

disulfide balance and for protecting many organisms from toxic oxygen species (Lu &

Holmgren, 2014, Holmgren, 2000). The two main proteins of this type found in biology

are thioredoxin and glutaredoxin. Although each of these contains a pair of redox-active

cysteines (Cys), S. aureus only synthesizes proteins of the thioredoxin system (Newton

et al., 1996, Uziel et al., 2004). In S. aureus, both O2 concentration and oxidative stress

induces thioredoxin expression (Uziel et al., 2004). The thioredoxin system is comprised

of 3 components including NADPH, thioredoxin reductase (TrxB), and thioredoxin

(TrxA), with TrxB maintaining the reduced form of TrxA using electrons from NADPH.

The reduced form of TrxA is able to donate electrons to a large range of enzymes in an

attempt to defend against oxidative stress (Holmgren, 2000, Arner & Holmgren, 2000).

One example is the donation of electrons to peroxiredoxins (Prx) which can then directly

detoxify H2O2 (Pannala & Dash, 2015). Not all Prx proteins require donation of electrons

from TrxA, with some able to use NAD(P)H to drive their reaction. In other bacterial

systems the main Prx is the alkyl hydroperoxide reductase (AhpFC)(Parsonage et al.,

2008, Poole et al., 2000), and indeed a homolog of this protein has also been confirmed

in S. aureus (Bhattacharyya et al., 2009, Cosgrove et al., 2007). Moreover, expression

of ahpF is induced upon peroxide challenge in S. aureus (Nobre & Saraiva, 2013).

51

Additional oxidative stress resistance mechanisms

MgrA is a staphylococcal protein with homology to Dps (DNA-binding protein

from starved cells), a ferritin-like DNA binding and Fe2+ storage protein (Ohniwa et al.,

2011, Martinez & Kolter, 1997, Nair & Finkel, 2004). Due to its elevated expression

under oxidative stress conditions it is thought that MgrA is thought to protect against

oxidative stress by DNA nucleoid condensation and binding of free Fe2+ in S. aureus

(Horsburgh et al., 2001a, Horsburgh et al., 2001b). S. aureus also has genes for up to 4

methionine sulfoxide reductases (msrA1, msrA2, msrA3, msrB), of which only MsrA1 is

specifically attributed to oxidative stress resistance (Singh et al., 2015, Singh &

Moskovitz, 2003). In general, the Msr system reduces oxidized Met-O residues back to

their un-oxidized form, thus repairing proteins after damage from ROS (Sasindran et al.,

2007). A role for S. aureus carotenoids in oxidative stress resistance has also been

described. Staphyloxanthin is the main membrane-associated carotenoid in S. aureus,

giving it its characteristic golden pigment (Marshall & Wilmoth, 1981b). These

carotenoids can act as direct antioxidants, providing protection from H2O2, O2-, HO,

hypochloride, and neutrophil killing (Liu et al., 2005, Clauditz et al., 2006). Regulation of

oxidative stress resistance (katA, ahpCF, trxB) and iron storage (ftn, mgrA) genes in S.

aureus are controlled in part by the peroxide response regulator (PerR) (Horsburgh et

al., 2001a), which encodes a metal-dependent sensor that directly responds to peroxide

stress. Importantly, regulation of kat and oxidative stress resistance occurs at multiple

levels, with katA expression being co-regulated by the ferric uptake repressor

(Fur)(Horsburgh et al., 2001b). Other regulatory pathways for oxidative stress

resistance include the SarA transcriptional regulator, which controls expression of trxB

and both sod transcripts (Ballal & Manna, 2010, Ballal & Manna, 2009).

52

Pathways and Targets of RNS

RNS chemistry and production

Similar to ROS, RNS are highly reactive small molecules that can drastically

affect the biology of most organisms. The major chemical sources of nitrosative stress

in biological systems are the nitroxyl anion (NO-), nitric oxide (NO), the nitrosium cation

(NO+), and peroxinitrite (ONOO-). The reactivity of these compounds lies in the positive

formal oxidation state of the nitrogen atom, of which they have +I, II, III, and III (Hughes,

1999). Arguably the most well studied of the RNS is NO, a small free radical gas that is

easily diffusible across membranes (Lancaster, 1997, Liu & Zweier, 2013). Often times

the literature does not effectively differentiate the specifics between the NO radical and

other RNS; therefore, vagueness can exist with respect to direct NO interaction or

interaction of its by-products (Bowman et al., 2011). The major source of NO in

mammals is enzymatic synthesis by NOS (reviewed here)(Alderton et al., 2001).

Mammalian NOS and its bacterial counterparts will be discussed in more detail below.

In some bacteria, NO is produced by enzymatic reduction of NO2- to NO by respiratory

NO2- reductases and the periplasmic cytochrome c NO2

- reductase (Nrf), but examples

of this have not been demonstrated in S. aureus (Watmough et al., 1999, Van Alst et al.,

2007, Arruebarrena Di Palma et al., 2013, Corker & Poole, 2003). Potential sources of

NO relevant to S. aureus include the membrane bound NO3- reductase (although this

has not been experimentally observed), host-derived NO, and bacterial NOS-derived

NO. An interesting case was observed in Salmonella typhimurium where NO was

produced by the membrane-bound NO3-reductase (narGHI), but only in the presence of

added NO2- as a substrate (Gilberthorpe & Poole, 2008). S. aureus has both NO2

- (Nir)

and NO3- (Nar) reductases, but their potential contribution to NO production is currently

53

unknown (Schlag et al., 2008, Burke & Lascelles, 1979, Burke & Lascelles, 1975). The

primary source of nitrosative stress encountered by S. aureus is the nitrosative burst

generated by human monocytes and macrophages (Nathan & Shiloh, 2000, Bogdan et

al., 2000, MacMicking et al., 1997). Activated leukocytes can produce NO using the

inducible nitric oxide synthase (iNOS) in the micromolar range (Lewis et al., 1995,

Nalwaya & Deen, 2005).

Cellular targets of RNS

Once produced, the highly reactive NO and its derivatives can interact with a

multitude of cellular targets (Figure 1-3)(reviewed here)(Toledo & Augusto, 2012).

Common targets of NO and its RNS by-products include non-organic molecules such as

molecular O2 (Czapski & Goldstein, 1995, Wink et al., 1993a), O2- (Czapski & Goldstein,

1995), lipid and protein-derived radicals (Rubbo et al., 2000, O'Donnell et al., 1997, Lam

et al., 2008), and various cellular targets including lipid membranes (Moller et al., 2007),

heme cofactors (Winger et al., 2007, Stone et al., 1995, Henry, 2015, Gardner et al.,

1998, Olson et al., 2004, Brown, 1995, Boveris et al., 2000), Fe-S clusters (Crack et al.,

2011, Tinberg et al., 2010), cysteine thiols (Gusarov & Nudler, 2012, Keshive et al.,

1996), and DNA (Salgo et al., 1995b, Salgo et al., 1995a, Tamir et al., 1996). NO can

also indirectly modify proteins via NO by-products (Radi, 2013, Radi, 2004, Wong et al.,

2001). Some specific modifications include nitration, nitrosation, and nitrosylation,

where NO can directly or indirectly modify various proteins within the cell by addition of

nitrogen side groups. A common protein modifier is the highly reactive ONOO- anion,

produced by interaction of NO with O2- (Huie & Padmaja, 1993), that not only modifies

proteins but can damage the cell in many ways. Nitrosylation occurs when a nitrosyl ion

or group (NO-) is added to a transition metal or thiol group. The RNS intermediate

54

ONOO- can undergo S-nitrosylation with thiol groups of Cys residues yielding S-

nitrosothiols (Radi et al., 1991a, Wink et al., 1997). Another protein modification

conferred by RNS is nitration, where a nitro group (NO2+) is added to an amine, thiol, or

hydroxy aromatic group. Nitration can also be completed by ONOO- by modification of

Tyr residues, forming a nitrotyrosine (Ischiropoulos et al., 1992b). Finally, nitrosation is

the addition of a nitronium ion (NO+) to an amine, leaving the molecule with a nitroso

group (NO). ONOO- is a powerful one and two electron oxidizing agent which appears

to be a primary contributor to the cytotoxic/cytostatic action of macrophages (Zingarelli

et al., 1996, Xia & Zweier, 1997, Ischiropoulos et al., 1992a). In addition to protein

modification, this molecule can damage DNA by nitration of guanine nucleotides to yield

nitroguanine (Yermilov et al., 1995). Additionally, ONOO- causes mutations and DNA

breakage in both humans and bacteria (Salgo et al., 1995b, Salgo et al., 1995a, Tamir

et al., 1996, Arroyo et al., 1992, Nguyen et al., 1992, Inoue & Kawanishi, 1995). The

autoxidation of NO can also produce nitrous anhydride (N2O3), another RNS that can

damage DNA by deamination of amines, a process that replaces these side chains with

hydroxyl groups (Wink et al., 1991, Nguyen et al., 1992). Damage of membrane lipids

can occur by the interaction of the conjugate acid of ONOO-, peroxynitrous acid

(ONOOH), leading to peroxidation (Radi et al., 1991b). Finally, NO itself has a distinct

relationship with many respiratory chain components and can both interfere with

incorporation of heme groups into respiratory proteins (Waheed et al., 2010) and

compete with O2 at terminal oxidases (Brown et al., 1997). The interaction of NO with

cytochrome heme forms a stable nitrosyl metal complex, and this is well documented

with the cytochrome p450 oxidase (Wink et al., 1993b). This interaction slows

55

respiration, but inhibition of terminal oxidases is not permanent and can be reversed

once NO is removed (Waheed et al., 2010). Depending on the concentration, NO can

act as a signaling molecule at low levels (Arora et al., 2015), and promotes nitrosative

stress when NO levels are high (Poole, 2005).

Protection From Nitrosative Stress in Staphylococcus aureus

NO detoxification proteins in S. aureus

S. aureus has developed a repertoire of proteins and metabolic adaptations

devoted to nitrosative stress resistance. In relation to other bacterial species, S. aureus

is particularly adept at resisting nitrosative stress (Richardson et al., 2008, Richardson

et al., 2006). In general, bacterial responses to nitrosative stress often include similar

responses to oxidative stress such as replenishment of cytoslic thiol pools, altered metal

homeostasis, activation of DNA repair processes, and induction of NO detoxification

pathways (Moore et al., 2004, Mukhopadhyay et al., 2004, Flatley et al., 2005,

Hromatka et al., 2005, Justino et al., 2005, Ohno et al., 2003, Firoved et al., 2004).

Examination of the nitrosative stress response in S. aureus shows that multiple genes

classically associated with oxidative stress resistance (ahpCF, katA, ftnA and mrgA) as

well as some metabolic genes (ldh, hmp, fdaB, nrdDG and cydAB) are upregulated

during nitrosative stress, suggesting an overlap between these two stress responses

(Richardson et al., 2006). Overlap induction of similar genes in response to oxidative

and nitrosative stress makes sense when understanding that both ROS and RNS affect

similar cellular processes. Hmp is a NO detoxification protein that directly converts NO

to NO3- using NAD(P)H in both E. coli and S. aureus (Poole et al., 1996, Goncalves et

al., 2006). In fact, Hmp is the major source of NO detoxification in S. aureus, with this

protein being responsible for ~90% of the NO detoxification under nitrosative stress

56

conditions (Richardson et al., 2006). The S. aureus strains containing Nor (~37%) utilize

this protein in a complementary role to Hmp-mediated NO detoxification (Lewis et al.,

2015). Nor contributes to cell respiration, suggesting that NO detoxification is not the

primary function of S. aureus Nor. Ferritin A (FtnA) has yet to be characterized in S.

aureus, but is expressed under nitrosative stress (Richardson et al., 2006). While not

biochemically characterized in S. aureus, an FtnA homolog in E. coli acts as an iron

buffer for re-assembly of Fe-S clusters upon H2O2 challenge (Bitoun et al., 2008). This

is also likely relevant during nitrosative stress conditions as NO can also damage Fe-S

clusters (Crack et al., 2011, Tinberg et al., 2010). While S. aureus employs

detoxification proteins to relieve nitrosative stress, it also utilizes metabolic flexibility to

survive these stress conditions.

S. aureus metabolic flexibility in response to nitrosative stress

The remarkable ability of S. aureus to replicate in the presence of NO

(Richardson et al., 2008) and recover from NO challenge (Richardson et al., 2006)

appears to be somewhat unique to this pathogenic species, as other commensal

bacteria such as Staphylococcus epidermidis, Staphylococcus saprophyticus, E. coli,

and B. subtilis do not share these capabilities (Richardson et al., 2008). This superior

ability to adapt to NO stress stems in part to the upregulation of genes for fermentation

and lactate metabolism, pathways which are less likely to be damaged by NO/RNS

(Hochgrafe et al., 2008, Richardson et al., 2006). Specifically, NO inhibits both pyruvate

formate lyase and pyruvate dehydrogenase, altering the redox status of the cell and

preventing acetate and ethanol production (Richardson et al., 2008). Additionally, NO is

well established to form cytochromal NO-heme complexes, effectively outcompeting O2

and inhibiting respiration (Giuffre et al., 2012, Sarti et al., 2003, Brunori et al., 2006,

57

McCollister et al., 2011). Five central regulons (SarA, CodY, Rot, Fur, and SrrAB) are

established via combined transposon screens and RNAseq analysis to be important for

resistance of S. aureus to nitrosative stress (Grosser et al., 2016).

A well-studied NO-response regulator in Gram-positive bacteria is the S. aureus

SrrAB TCS and its homologues (i.e., ResDE in Bacillus subtilis). SrrAB is required for

effective response to nitrosative stress in S. aureus, presumably sensing this signal via

impaired electron flow through the respiratory chain (Kinkel et al., 2013, Richardson et

al., 2006). Many of the genes (hmp, cydAB, nrdDG) induced upon RNS challenge are

controlled by SrrAB. With this said, additional regulatory components involved in the

metabolic response to NO exist, since NO-induction of ldh1 expression is not controlled

by SrrAB (Kinkel et al., 2013, Richardson et al., 2006). S. aureus utilizes an inducible L-

lactate dehydrogenase (Ldh1) to maintain redox homeostasis and substrate level

phosphorylation under NO-stress conditions (Richardson et al., 2008). The L-lactic acid

produced by Ldh1 can also promote respiration by donating electrons to the L-lactate-

quinone oxidoreductase (Lqo) (Fuller et al., 2011). Both Lqo and Mqo1 are critical

during nitrosative stress when cells are grown on L-lactate and peptides (Spahich et al.,

2016). Mqo1 is needed for proper TCA cycle function, and Lqo is required for

regeneration of pyruvate from L-lactate, a reaction that is critical for ATP formation

when respiration is inhibited by NO. While high levels of NO can inhibit bacterial growth,

some bacteria have evolved to use NO-mediated respiratory inhibition to their

advantage. For example, NO protects S. aureus from gentamicin by blocking respiration

and limiting the energy-dependent phases of drug uptake (McCollister et al., 2011).

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Nitric Oxide Synthase in Mammals and Bacteria

Mammalian Nitric Oxide Synthase

Mammalian NOS structure and chemical reaction

Nitric oxide synthase-like (NOS) homologs are present in all 6 kingdoms of life

including the Animalia (Knowles & Moncada, 1994), Plantae (Jeandroz et al., 2016),

Fungi (Ninnemann & Maier, 1996), Protista (Malvin et al., 2003), Archaeabacteria

(Sudhamsu & Crane, 2009), and Eubacteria (Sudhamsu & Crane, 2009); with the

biological function often being unique to the individual organism. A vast amount of work

has been completed on the structure and function of mammalian NOS enzymes, but a

review of the literature reveals that research on other NOS proteins is just beginning to

scratch the surface. Mammalian NOS (mNOS) proteins contain both oxygenase and

reductase domains, which catalyze the 2-step oxidation of L-arginine to L-citrulline and

NO, with intermediate formation of Nω-hydroxy-L-arginine (NOHA) (Griffith & Stuehr,

1995, Alderton et al., 2001, Stuehr et al., 2004b, Moore et al., 2004, Mukhopadhyay et

al., 2004, Flatley et al., 2005, Hromatka et al., 2005, Justino et al., 2005, Marletta,

1994). The mNOS is a homodimer containing a C-terminal flavoprotein reductase

(NOSred) and an N-terminal oxygenase domain (NOSox) (Stuehr, 1999). NOSred is the

flavoprotein containing NADH oxidase that has homology to the p450 NADH

oxidoreductase of the respiratory chain (Nishida et al., 2002). This domain has binding

sites for flavin adenine dinucleotide (FAD), flavin mononucleotide (FMN) and NADPH,

allowing it to act as a source of reducing equivalents for O2 binding and activation. The

catalytic domain is contained within NOSox which binds L-arginine and contains heme,

as well as a redox-active 6R-tetrahydrobiopterin (H4B) cofactor. Although NOSox and

NOSred domains are encoded as a single polypeptide, a regulatory calmodulin-binding

59

motif brings both domains together upon calcium (Ca2+) binding (Smith et al., 2013,

Piazza et al., 2012). Electrons flow from NADH to the FAD and FMN cofactors, where

they oxygenate L-arginine to NOHA. Transfer of electrons to the heme-containing active

site of the oxygenase domain catalyzes the final conversion of NOHA to L-citrulline and

NO (Alderton et al., 2001). Both steps of O2 activation notably require H4B to be a

transient electron donor to heme (Stuehr et al., 2004a, Hurshman et al., 1999).

Mammalian NOS isotypes and their functions

A great deal of research is complete on the three mammalian NOS isotypes:

endothelial (eNOS), neuronal NOS (nNOS), and inducible (iNOS). These NOS enzymes

contribute to many critical biological functions, including but not limited to, regulation of

blood pressure (eNOS), nervous system signaling (nNOS), and protection against

pathogens (iNOS)(Sudhamsu & Crane, 2009, Crane et al., 2010, Forstermann & Sessa,

2012, Crawford, 2006, Alderton et al., 2001, Yun et al., 1996, Lipton, 2001). Due to the

vast potential of cellular targets, it is no surprise that NO acts as a signaling molecule

for multiple processes. Arguably the best studied NOS signaling pathway in mammals is

activation of guanylate cyclase by eNOS (Buys & Sips, 2014, Derbyshire & Marletta,

2012, Follmann et al., 2013). Activated guanylate cyclase produces the well-known

cyclic GMP (cGMP) second messenger (Arnold et al., 1977), leading to vasodilation and

regulation of blood pressure. The action of NO on guanylate cyclase occurs through

binding of NO to the heme-NO (H-NOX) binding domain, stimulating the catalytic

domain of this enzyme (Underbakke et al., 2014). Bacteria also contain H-NOX proteins

and, therefore, sense NO in a signaling capacity (Nisbett & Boon, 2016, Plate &

Marletta, 2013). However, a majority of bacteria do not synthesize NOS, therefore the

60

contribution of NO signaling in these bacteria is often from environmental signals and/or

NOS-independent NO production (i.e. from denitrification).

A less well studied putative fourth NOS isoform is described and designated

mitochondrial NOS (mtNOS)(Giulivi, 2003). While it is well accepted that mNOS

proteins regulate mitochondrial respiration, there is controversy in the field over whether

this function is due to a unique mtNOS isoform (Finocchietto et al., 2009). With that

said, mtNOS is thought to specifically modulate the respiration of mitochondria by

forming cytochromal NO-heme complexes, effectively outcompeting O2 and inhibiting

respiration (reviewed here)(Giuffre et al., 2012, Sarti et al., 2003, Brunori et al., 2006). A

second mechanism is also postulated where the mtNOS functionally associates with the

NADH dehydrogenase (complex I) and accepts electrons from this protein complex

(Parihar et al., 2008a). A role for NOS in respiratory modulation has not yet been

described in non-mammalian organisms.

Bacterial Nitric Oxide Synthase

Bacterial NOS discovery

The first bacterial NOS (bNOS) enzyme was discovered nearly 20 years ago in

Nocardia, a strictly aerobic, Gram positive bacterium (Chen & Rosazza, 1994). Most

bacterial species containing NOS are Gram positive obligate aerobes or facultative

anaerobes, with some exceptions (Sudhamsu & Crane, 2009). Biochemical and/or

functional characterization is completed on NOS proteins from Deinococcus,

Streptomyces, Bacillus, and Staphylococcus (Sudhamsu & Crane, 2009). Additional

genomic analyses suggest that NOS proteins are also present in Exiguobacterium,

Geobacillus, Lysinibacillus, Oceanobacillus, Paenibacillus, Rhodococcus, and

Sorangium, as well as the archaeal genus Natronomonas (Sudhamsu & Crane, 2009,

61

Gusarov et al., 2008). In general, a majority of bNOS proteins are present in the

Firmicutes, with examples also observed in the Actinobacteria. Biochemical and

crystallographic studies are completed on multiple bNOS proteins and generally follow

the workflow of genomic identification, cloning, and recombinant expression. For most

cases, bNOS proteins are very similar to the oxygenase domain of mNOS (Adak et al.,

2002b, Adak et al., 2002a, Pant et al., 2002, Salard-Arnaud et al., 2012, Midha et al.,

2005, Chartier & Couture, 2007a, Chartier & Couture, 2007b, Gautier et al., 2006,

Santolini et al., 2006, Sudhamsu & Crane, 2006, Montgomery et al., 2010), including S.

aureus NOS (saNOS)(Bird et al., 2002, Chartier et al., 2006, Salard et al., 2006).

Confirmation of structural similarity upon substrate binding has also been determined in

B. subtilis (Pant et al., 2002), S. aureus (Pant et al., 2002), and G. stearothermophilus

(Sudhamsu & Crane, 2006) with either L-arginine or the L-arginine analog S-ethyl-

isothiourea. In vitro production of NO has been confirmed by biochemical

characterization of purified Bacillus (Adak et al., 2002b, Adak et al., 2002a),

Staphylococcus (Hong et al., 2003), and Deinococcus (Reece et al., 2009) NOS

proteins.

Bacterial NOS structure

The structure of bNOS enzymes are closely related to mNOSoxy domains, with a

few notable differences. Most bacterial NOS lack an attached NOSred, zinc-

coordinating N-terminal hook, and calmodulin binding motif (Sudhamsu & Crane, 2009).

The dimer interface of the oxygenase domain catalyzes the enzyme activity and

comparison of this interface to mNOS shows a high degree of sequence conservation

(Lustig et al., 2011). Notable differences between these oxygenase domains include the

absence of a 50-residue amino terminal hook in bNOS, which provides H4B

62

coordination to an interfacial zinc ion in mNOSs (Raman et al., 1998). This hook

functions by both providing a binding site for the cofactor (H4B) and stabilizing the

dimer, particularly for iNOS isotypes (Ghosh et al., 1997). The second notable

difference lies is the cofactor required for stabilization of the reaction. Mammalian NOS

contain the H4B cofactor in their active site, whereas tetrahydrofolate (H4F), which

contains the same pteridine ring structure as H4B, has been found to act as a functional

replacement in vitro, for Bacillus NOS proteins (Pant et al., 2002). The absence of the

terminal zinc hook likely provides less steric hindrance for the larger H4F molecules

(Pant et al., 2002, Adak et al., 2002b). Aside from these differences, comparison of

mammalian and bacterial NOSoxy domains shows a high degree of structural and

sequence conservation at the cofactor binding site, dimer interface, and heme centers

(Pant et al., 2002, Bird et al., 2002). Crystallographic analysis of heme centers reveals

only minor differences between mNOS and bNOS proteins, but NO release rates from

bacterial homologs are considerably lower (Adak et al., 2002a). These kinetic

differences are due to an Ile substitution in bNOS for the Val residue that normally

resides over the O2 binding site in mNOS (Pant et al., 2002). This was found to account

for slight differences in the kinetic profile of the reactions, but the overall catalytic

mechanism of the oxygenase domain remains the same (Wang et al., 2004, Wang et

al., 2010, Gautier et al., 2006).

Reductase partner studies for bNOS

In contrast to mammalian NOS, most bacterial NOS only contain an oxygenase

domain, with the specific cellular reductase partner yet to be determined in most

bacteria, including S. aureus (Bird et al., 2002, Sudhamsu & Crane, 2009). Lack of an

N-terminal reductase domain makes it necessary for bacterial NOSs to utilize alternative

63

reductase proteins to provide electrons for the reaction. Recently, native flavodoxins

YkuN and YkuP, as well as the YumC ferredoxin reductase were shown to support NOS

oxygenase activity in B. subtilis (Holden et al., 2014, Wang et al., 2007). These

experiments were completed with purified protein in vitro and therefore the biological

relevance is unclear. Isolated YkuN and YkuP also supported similar NO synthesis by

NOS isolated from D. radiodurans (Wang et al., 2007). In a separate study, deletion of

ykuN and ykuP did not cause loss of NOS activity in Bacillus, and the same effect was

seen after deletion of other predicted reductase partners (Gusarov et al., 2008). This

data suggests that there is not one dedicated redox partner for Bacillus NOS and that it

likely “hijacks” cellular redox partners that are not normally dedicated to NO production.

Other studies suggest that the bNOS oxygenase domain may be promiscuous in its

ability to receive electrons from a non-dedicated redox partner (Gusarov et al., 2008). It

is not yet known if this is a common occurrence for all bacterial NO synthases, but it is

important to note that utilization of specific reductase partners may vary and be

dependant on cellular growth conditions. One notable exception to this is the presence

of an attached reductase domain that was found upon genome sequencing of S.

cellulosum (Schneiker et al., 2007) and characterization of scNOS (Agapie et al., 2009).

scNOS is unique among all characterized NOS proteins because it contains an N-

terminal domain of unknown function, a C-terminal NOSox, and an Fe-S cluster which

replaces the FMN binding module.

bNOS inhibitor studies

In 2013, an inhibitor screen for B. subtilis NOS (bsNOS) inhibitors uncovered two

potential inhibitors with antimicrobial capabilities (Holden et al., 2013). Select inhibitors

by themselves, or when combined with antibiotics, killed B. subtilis cells (Holden et al.,

64

2013, Holden et al., 2015c). Due to high sequence conservation between bNOS and

mNOS proteins, the pterin binding site was targeted, which is not as conserved across

bacterial and eukaryotic domains (Holden et al., 2013). Inhibitor bound crystal structure

studies comparing bsNOS and mNOS further confirmed multiple compounds that bind

either the active site, pterin cofactor site, or a unique binding pocket of bsNOS (Holden

et al., 2015a). Active site examination shows an Ile substitution for Val in the bsNOS

protein compared to mNOS (Holden et al., 2015a), which may provide enough

differences in hydrophobicity and steric hindrance to provide selectivity. The pterin

binding site of bNOS proteins is more solvent exposed than mNOS and, therefore, may

be more selective for larger pharmacophore inhibitors (Holden et al., 2015a). Finally, a

unique binding pocket in bsNOS is found to interact with one of the inhibitors and may

provide additional selectivity (Holden et al., 2015a). Further studies elucidated a class of

NOS inhibitors that employ an aminoquinoline scaffold to bind a hydrophobic patch that

is unique to bNOS proteins (Holden et al., 2016). In summary, studies on bNOS

inhibitors have been fruitful, with bsNOS (Holden et al., 2013, Holden et al., 2015c) and

saNOS (described below)(Holden et al., 2015b) inhibitors appearing to be able to limit

growth when combined with other stressors such as antibiotics and peroxide.

Functional studies of bNOS proteins

In addition to biochemical and crystollographic studies, there has been an

emerging focus on the functional role of bacterial NOS proteins. Actual in vivo

production of NOS-derived NO is demonstrated for S. turgidiscabies, D. radiodurans

(Patel et al., 2009), B. subtilis (Gusarov & Nudler, 2005, Schreiber et al., 2011), B.

anthracis (Shatalin et al., 2008), and S. aureus (Sapp et al., 2014). Examination of the

literature reveals unique functions for each NOS protein that depend on the species

65

studied, with some similarities only observed between Bacillus and Staphylococcus

NOS (discussed below)(Patel et al., 2009, Kers et al., 2004, Wach et al., 2005, Gusarov

et al., 2009, Gusarov & Nudler, 2005).

Streptomyces and Deinococcus NOS. Often times in bacteria, the position of a

gene on the chromosome gives insight into its function, where surrounding genes can

be grouped into a certain pathway or mechanism. The genomic organization of bNOS is

highly variable (Sudhamsu & Crane, 2009), but one clear example of the relevance of

genomic organization is with S. turgidiscabies NOS (stNOS). In the plant pathogen S.

turgidiscabies, nos is present on a pathogenicity island with phytotoxins, and NOS-

derived NO is required for synthesis of these molecules (Johnson et al., 2008, Wach et

al., 2005, Kers et al., 2004). Specifically, thaxtomin A is a nitrated phytotoxin that

inhibits plant cell wall synthesis (Healy et al., 2000) and requires nitration by NOS.

Induction of NOS activity occurs in the presence host cellobiose, a plant cell wall

component (Johnson et al., 2008), therefore providing the first evidence of an inducible

bNOS function. NOS-derived NO production is also induced in in D. radiodurans, which

is so named for its extreme resistance to ionizing radiation (Agapov & Kulbachinskiy,

2015, Krisko & Radman, 2013). Specifically, UV radiation both induces nos gene

expression and cellular NO production in this organism (Patel et al., 2009). Loss of NOS

activity also limits the ability of D. radiodurans to recover from UV radiation damage

(Patel et al., 2009). In this case, NO protection relies on NO-induced upregulation of

obgE transcription, a gene involved in stress response and growth proliferation (Patel

et al., 2009, Czyz & Wegrzyn, 2005, Foti et al., 2005). S. turgidiscabies, a species

66

phylogenetically related to D. radiodurans, also produces NOS-derived NO in response

to plant host signals (Johnson et al., 2008).

Bacillus NOS. Similar to D. radiodurans, NOS proteins in Bacillus are found to

play an important role in resistance to external stress. While nos deletion does not

increase sensitivity to oxidative damage and H2O2 stress in D. radiodurans (Patel et al.,

2009), an important role for bNOS in oxidative stress resistance is found in both B.

subtilis and B. anthracis (Gusarov & Nudler, 2005, Shatalin et al., 2008). These bNOS

proteins have been well-characterized in the model soil bacterium B. subtilis and the

human pathogen B. anthracis, with some similarities and differences. Protection from

oxidative stress is conferred by a proposed dual mechanism that interrupts H2O2 toxicity

by 1) directly activating Kat and 2) depleting free Cys, thereby limiting the Fenton

reaction (Gusarov & Nudler, 2005, Shatalin et al., 2008). Free Cys inhibits Kat activity

and NO is believed to directly activate Kat by preventing the Kat-Cys interaction via an

S-nitrosylation mechanism (Gusarov & Nudler, 2005). This mechanism is proposed, but

direct binding of NO to Kat has yet to be confirmed. As an abundant low-molecular

weight thiol in Gram-positive bacteria (Newton et al., 1996), Cys is associated with

promoting oxidative stress by driving the Fenton reaction (Park & Imlay, 2003). During

the Fenton reaction H2O2 oxidizes free cellular Fe2+ to yield toxic HO radicals. At the

same time, Cys reduces Fe3+ back to Fe2+ and forms cystine. To continue driving the

Fenton reaction, the Trx/TrxR system must reduce cystine back to Cys. NO is thought to

interrupt this process by inhibiting the thioredoxin system through direct interaction

(Gusarov & Nudler, 2005). Therefore, NO likely limits the amount of free intracellular

Cys that is regenerated. This can also control the amount of Cys available to inhibit Kat.

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Due to the known oxidative burst of macrophages during infection, B. anthracis

NOS has proven essential for virulence and survival in macrophages (Shatalin et al.,

2008). An additional study also shows that B. anthracis NOS-derived NO is produced as

a toxin, which contributes to macrophage death by S-nitrosylation of intracellular host

proteins (Chung et al., 2013). The supernatants of nos mutant cells are also

substantially less toxic to epithelial cells, and this effect is dependent on epithelial cell

membrane permeability (Popova et al., 2015). Several lines of evidence suggest that

NO-mediated protection from oxidative stress is unique to Gram-positive bacteria,

including: 1) treatment with NO did not provide immediate protection from H2O2 in E. coli

(Gusarov & Nudler, 2005), 2) the major catalase KatA of E. coli is inhibited by NO, in

contrast to B. subtilis (Gusarov & Nudler, 2005, Brunelli et al., 2001), 3) Cys does not

inhibit E. coli catalase as it does in B. subtilis (Switala & Loewen, 2002) and 4) Free Cys

is not a prominent thiol in E. coli, but is prominent in Bacilli and S. aureus (Newton et al.,

1996, Park & Imlay, 2003). Alternatively, the most prominent thiol in E. coli is

glutathione, which was not found to support the Fenton reaction (Park & Imlay, 2003).

Overall, a clear link has been established between oxidative stress resistance and

Bacillus NOS proteins.

Protection against specific antibiotics in B. subtilis and B. anthracis by NOS was

conferred by both direct chemical modification and through alleviating oxidative stress

generated by some antibiotics (Gusarov et al., 2009). For example, acriflavine is a DNA

intercalator/acridine antibiotic containing two aromatic amino groups necessary for

toxicity (Wainwright, 2001). NO products can nitrosate arylamino moieties, rendering

them less effective (Gusarov et al., 2009, Nedospasov et al., 2000). Moreover, pre-

68

treatment of cells with the iron chelator bipyridyl (suppresses the Fenton reaction), or

the radical scavenger thiourea, conferred resistance to antibiotic-induced oxidative

stress, similar to pre-treatment with NO (Gusarov et al., 2009). These results together

suggest that NOS-derived NO can protect against some antibiotics by direct

detoxification and/or limiting oxidative stress generated by the Fenton reaction (Gusarov

& Nudler, 2005, Shatalin et al., 2008).

bNOS has broad potential for affecting multiple cellular targets, and this is further

demonstrated by the additional contribution of NOS to B. subtilis physiology. Many B.

subtilis strains exhibit multicellular traits and form structurally complex biofilms.

Regulating biofilm dispersal is another role that is suggested for NOS-derived NO in B.

subtilis (Schreiber et al., 2011), where a Δnos strain and wildtype treated with NOS

inhibitors both exhibited strongly enhanced biofilm dispersal. While the exact

mechanism is not yet elucidated, NO may signal the transition from oxic to anoxic

conditions (as the biofilm develops), similar to a proposed role for NO in other bacteria

(Zumft, 2002, Spiro, 2007, Barraud et al., 2006). Another contribution of bsNOS to

physiology is elucidated by studies of B. subtilis in the gut of the model worm

Caenorhabditis elegans. Specifically, bsNOS enhances longevity and stress resistance

of the worms by a mechanism that was dependant on the DAF-16 and HSF-1 C.

elegans transcription factors (Gusarov et al., 2013). Overall it seems that bacterial NOS

proteins contribute to specific cellular processes, many of which are unique to the

organism, its environment, and/or the specific biological processes it needs to survive.

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Staphylococcus aureus NOS

General Characteristsics

Discovery and structural characterization

In 1997, the first S. aureus NOS protein (saNOS) was confirmed by Western

blotting analysis using an iNOS antibody and biochemical assays of crude lysates (Choi

et al., 1997). Original biochemical characterization of saNOS was completed by simply

mixing predicted reaction components with cell lyates and measuring both NO and

radiolabeled L-citrulline (Choi et al., 1997). Recombinant saNOS comprises a

homodimer (Bird et al., 2002), similar to other bNOS proteins (Adak et al., 2002b, Adak

et al., 2002a). In this study (Bird et al., 2002), recombinant saNOS is found to require

addition of a reductase partner for activity. In addition, recombinant saNOS was

determined to be a heme containing homodimer that was crystallized with NAD+ bound

to the interface ligand binding site (Bird et al., 2002)(Figure 1-4). Additional important

findings regarding the structural properties of saNOS include determination that neither

H4B nor H4F is required for stability of the catalytic heme in vitro (Chartier & Couture,

2004). Only one structural mutagenic study has been completed on saNOS, where

conserved Trp resides (position 314 and 316) at the pterin binding site/dimer interface

were changed to various alternative amino acids (Lustig et al., 2011). Variants of the

Trp-314 residue presented a “loose” conformation, suggesting that this residue is

important for proper dimerization. Overall, structural and sequence similarity appear to

be relatively conserved between saNOS and other NOS isotypes (Pant et al., 2002, Bird

et al., 2002, Salard et al., 2006, Salard-Arnaud et al., 2012, Wang et al., 2004, Wang et

al., 2010, Gautier et al., 2006).

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Sequence identity and genomic organization

Genomic examination of the S. aureus nos locus has revealed a 1,077 bp open

reading frame coding for a predicted 41.7 kDa protein (Sapp et al., 2014)(Figure 1-5).

Further inspection showed that nos is separated by only 19 bps from the downstream

pdt gene, which encodes a 29.5 kDa predicted prephenate dehydratase (saPDT)(Sapp

et al., 2014). Indeed, co-transcription PCR analysis on cDNA demonstrated nos and pdt

co-transcription (Sapp et al., 2014). The nos-pdt operon is flanked upstream by the NAD

synthetase (nadDE) operon and downstream by a predicted sodium:sulfate symporter.

The nos-pdt operon organization is highly conserved and unique to staphylococci, but is

thus far not present in other bacteria containing nos (Figure 1-5). Important for

phenylalanine biosynthesis, the 795 bp pdt gene product catalyzes the formation of

phenylpyruvate from prephenate (Tan et al., 2008). This finding was confirmed in S.

aureus, as saPDT is required for auxotrophic growth without phenylalanine (Sapp et al.,

2014). At present, functional interactions of saPDT and saNOS have not been

determined.

Functional Studies on saNOS

Protection from oxidative stress

The first functional studies on saNOS were performed by Gusarov & Nudler who

studied the role of bsNOS in resistance to oxidative stress (Gusarov & Nudler, 2005).

Supplemental data in this study showed that exogenous NO could also protect S.

aureus against H2O2 challenge, but the direct contribution of saNOS was not

determined. Since then, three separate studies have confirm a significant role for

saNOS in resistance to oxidative stress (van Sorge et al., 2013, Sapp et al., 2014, Vaish

& Singh, 2013). In each, mutation of S. aureus nos made the cells more sensitive to

71

killing by H2O2, including the Sapp et al. publication by our research group (Figure 1-6).

In one of these studies, SOD activity was measured in a nos mutant by two separate

methods, negative staining on polyacrylamide gel and a colorimetric activity assay (van

Sorge et al., 2013). The gel method showed slightly lower SOD activity at 3 hours, but

not at 2 and 5 hours growth. Activity measured by the colorimetric assay revealed a ~5-

8% decrease in SOD activity due to nos mutation at 2 and 3 hours growth, but no

obvious difference at later time points (4-6 hours). Taken together, these results imply

that there may be a slight decrease in SOD activity that could account for loss of

resistance to oxidative stress, but it seems unlikely that the drastic killing from oxidative

stress observed in S. aureus nos mutants results solely from this subtle change in SOD

activity. As described above, the Fenton reaction is the primary generator of HO within

cells. It is important to note that addition of free iron (to drive the Fenton reaction) did

not affect S. aureus nos mutant growth, providing indirect evidence that saNOS does

not confer resistance to oxidative stress via protection from the Fenton reaction (van

Sorge et al., 2013), as was previously suggested for bNOS in Bacillus (Gusarov &

Nudler, 2005). Therefore, the exact mechanism of saNOS-related resistance to

oxidative stress in S. aureus remains unclear.

Contribution of saNOS to virulence and antimicrobial resistance

As an extremely successful pathogen, physiological studies in S. aureus often

have the overall goal of elucidating novel targets for treatment. Indeed, saNOS appears

to play an important role during infection as noted by its contribution to virulence in both

murine abscess (van Sorge et al., 2013) and sepsis models (Sapp et al., 2014). Murine

abscess size and bacterial counts were significantly lower in the nos mutant infected

abscess model relative to wildtype (van Sorge et al., 2013). Additionally, co-infection

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with S. aureus wildtype and nos mutant in vivo shows that the mutant cells become

outcompeted by wildtype within the abscess (van Sorge et al., 2013). When systemic

infection was studied using a murine sepsis model (Sapp et al., 2014), mice infected

with nos mutant cells presented with statistically-significant decreases in bacterial loads

in the kidneys, lungs, and liver (Figure 1-7). Furthermore, a significant increase in nos

mutant-infected mouse survival was observed in this study (Sapp et al., 2014) and

Figure 1-7. A third study recorded no significant differences in nos mutant infection

when measured in a intraperitoneal infection model (Vaish & Singh, 2013), but this may

be due to strain differences and/or a lack of relevance of saNOS in this specific infection

model. It is quite possible that the decreased virulence associated with the nos mutant

is due to its sensitivity to oxidative stress. As described above, neutrophils and

macrophages both generate an oxidative burst in an attempt to combat invading

pathogens. With this in mind, a nos mutant also presented with increased sensitivity to

killing by human neutrophils (Vaish & Singh, 2013, van Sorge et al., 2013), neutrophil

extracellular traps (van Sorge et al., 2013), and intracellular killing by macrophages (van

Sorge et al., 2013). It is unlikely that saNOS-derived NO directly affects host

components because the oxidative burst of human neutrophils, NO production by

neutrophils, neutrophil lysis, and production of neutrophil extracellular traps were shown

to be unaffected by exposure to a S. aureus nos mutant (van Sorge et al., 2013).

Host cathelicidins are cationic host antimicrobial peptides that cause pore

formation in bacterial cell membranes (Nizet & Gallo, 2003, Nizet et al., 2001). Van

Sorge et. al., showed that a nos mutant is more sensitive to the murine cathelicidin

antimicrobial peptide (mCRAMP)(van Sorge et al., 2013). Bacterial resistance to

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cathelicidins is generally conferred by increasing the positive charge or decreasing the

hydrophobicity of the cell wall (Kristian et al., 2003, Peschel et al., 1999), but no

difference in surface charge or hydrophobicity were observed in a nos mutant (van

Sorge et al., 2013). Protease activity is also associated with cathelicidin resistance and

indeed the nos mutant is less able to degrade mCRAMP. Cathelicidins have been

shown to enhance ROS production by phagocytes (Zheng et al., 2007, Alalwani et al.,

2010) and induce oxidative stress in bacteria (Peters et al., 2010), therefore NOS-

mediated resistance to these peptides is likely due to both limiting oxidative stress and

elevated protease activity.

Multiple studies have demonstrated a role for saNOS in resistance to certain

antibiotics. Pyocyanin, a ROS generating antimicrobial produced by P. aeruginosa, was

slightly more effective at limiting growth of a S. aureus nos mutant during post-

exponential growth phase (Gusarov et al., 2009). Antibiotics that are more relevant in

treating MRSA infections (Eckmann & Dryden, 2010) were also tested including

daptomycin, vancomycin, streptomycin, and gentamicin, with mixed results (van Sorge

et al., 2013). The nos mutant was slightly more sensitive to daptomycin and

vancomycin, but resistant to streptomycin and gentamycin. Moreover, the cell wall

antibiotic vancomycin induced NO production by saNOS in this study. Vancomycin

increases intracellular HO formation in S. aureus (Kohanski et al., 2007), therefore, the

mechanism of saNOS mediated vancomycin resistance may be linked to oxidative

stress. While the reason for elevated resistance to streptomycin and gentamycin (30s

ribosomal inhibitor) is not known, it appears to be unique to this class of

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aminoglycosides because resistance to linezolid (50s ribosomal inhibitor) was not

altered in the nos (van Sorge et al., 2013).

Contributions of saNOS to General Physiology

Considering the current literature on saNOS, this protein is clearly important in

resistance to oxidative stress and antimicrobials, which translates to a role in virulence.

Although work has been done on characterizing saNOS upon external challenge, little

has been completed on the endogenous role of saNOS in general physiology. An

obvious phenotype that was observed by our lab upon nos mutation is elevated

carotenoid pigment production when cultured on agar plates (Sapp et al., 2014).

Elevated pigmentation on agar plate growth was found not to be due to altered

transcription of genes (crtN, purH, asp23) associated with pigment production in S.

aureus (Sapp et al., 2014). No obvious growth defect was observed in the nos mutant

when grown aerobically in complex media containing glucose, but a conserved very

minor OD600 decrease has been noted upon transition of the nos mutant into stationary

phase growth (Gusarov et al., 2009, Almand, 2010). In some earlier studies,

researchers found that methanol treatment elevates saNOS protein levels and

enzymatic activity (Hong et al., 2003, Choi et al., 1998). The relevance of this is

currently unknown, but is predicted to be related to a general stress response. In fact,

published microarray data (Chang et al., 2006) as well as preliminary data from our lab

(Almand, 2010) showed that nos expression is elevated upon challenge with H2O2. NO

production by saNOS was confirmed in live cells from two different studies, using both

the NO-specific probe copper fluorescein (Cu-FL) (van Sorge et al., 2013) and DAF-FM

diacetate, a general RNS stain (Sapp et al., 2014). Staining with each probe confirmed

NO production in wildtype S. aureus during aerobic exponential growth and agar plate

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growth, respectively. A role for saNOS in endogenous physiology also stems from

studies of nos gene expression, which demonstrated elevated nos RNA levels at late

exponential growth phase and low-oxygen growth, relative to aerobic growth (Sapp et

al., 2014). These results are consistent with a separate study which showed that relative

gene expression of nos was highest during early-exponential phase aerobic growth

(100%) and declines to 4.9% by stationary phase growth (Vaish & Singh, 2013).

Overall, studies on the contribution of saNOS to general physiology are mostly

descriptive. There is an obvious need in the field for mechanistic studies on how saNOS

affects the biology of this bacterium under both “normal” and stress growth conditions.

Hypothesis and Aims

The few studies that have been thus far completed on saNOS have primarily

looked at the role of this protein under exogenous stress conditions and/or during

infection. Due to the large amount of literature suggesting multiple targets and roles for

NO in cellular systems, the overall hypothesis of this dissertation was that saNOS plays

a role in modulating general S. aureus physiology, and was tested by three

experimental aims. Aim 1 sought to determine the contribution of saNOS to S. aureus

growth, gene expression, and metabolism. The previously-described role of saNOS in

resistance to exogenous oxidative stress, combined with the transcriptomic data from

Aim 1 led to a secondary hypothesis that saNOS contributes to endogenous oxidative

stress and respiration. Therefore Aim 2 focused on the contribution of saNOS to

endogenous oxidative stress and respiratory phenotypes. Measurements of

endogenous ROS and respiration were completed in an attempt to determine the

source of altered gene expression and metabolism. Due to multiple genetic and

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metabolic adaptations in the nos mutant, Aim 3 sought to determine potential regulators

of nos mutant metabolic adaptation.

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Figure 1-1. Fermentation pathways of S. aureus. Pyruvate is the central metabolite of

fermentation and can be generated from glucose via glycolysis. The fate of pyruvate is determined by various enzymatic reactions including oxidation to D or L lactate and/or fermentation to ethanol, acetate, or 2,3-butanediol. DDH, D-lactate dehydrogenase; LDH, L-lactate dehydrogenase; PFL, pyruvate formate lyase; PDH, pyruvate dehydrogenase; PTA, phosphotransacetylase; AK, aceate kinase; ADH, alcohol dehydrogenase; ALS, α-acetolactate synthase. Adapted from (Ferreira et al., 2013).

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Figure 1-2. Branched respiratory chain of S. aureus. Respiration can be driven using

NADH, other reducing equivalents from the TCA cycle, or L-lactate. Succinate dehydrogenase (Sdh), NADH dehydrogenase (Ndh), nuol-like NADH dehydogenase (Mps/Mnh), or lactate quinone oxidoreductase (Lqo) can all accept electrons from electron donors to promote respiration. Electrons are then shuttled through the membrane by menaquinone (MQ) to generate a membrane potential for ATP synthesis. S. aureus can use O2 (Qox/Cyd), NO3

(Nar), or NO (Nor) as final electron acceptors. Dotted Nor indicates it is only found in a subset of S. aureus strains.

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Figure 1-3. Cellular targets of NO. Common cellular targets of NO include oxygen

species, membrane lipids, DNA, heme and non-heme iron cofactors, Fe-S clusters, and cysteine thiols.

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Figure 1-4. Structure of saNOS. The dimer structure is made of individual monomers

indicated with purple and blue. Rods represent α-helices and ribbons represent β-sheets. Dark red spheres indicate heme molecules. Crystal structure determination required NAD (gray and green molecules) as well as s-ethylisothiourea (gray and red molecules). Structure obtained from NCBI Structure database (MMDB ID: 21756; PDB ID: IMJT) and images were generated in Cn3D 4.3.1 program (Madej et al., 2014).

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Figure 1-5. Genomic organization and distribution of saNOS. This figure was originally

published in (Sapp et al., 2014), and is reproduced here under the Creative Commons Attribution (CC BY) license policy of PLOS ONE.

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Figure 1-6. Contribution of saNOS to H2O2 resistance. This figure was originally published in (Sapp et al., 2014), and is reproduced here under the Creative Commons Attribution (CC BY) license policy of PLOS ONE.

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Figure 1-7. saNOS in a sepsis model of infection. This figure was originally published in

(Sapp et al., 2014), and is reproduced here under the Creative Commons Attribution (CC BY) license policy of PLOS ONE.

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CHAPTER 2 RESULTS

Aim 1. Contribution of saNOS to General Physiology

Growth Phenotypes Upon nos Mutation

Although many research groups have studied saNOS with a focus on its

relevance during infection (Vaish & Singh, 2013, van Sorge et al., 2013, Sapp et al.,

2014), little is known about the potential effects of this enzyme on S. aureus physiology

in the absence of exogenous stress. While optimizing a previously described oxidative

stress assay (Gusarov & Nudler, 2005), it was noted that a previously-published S.

aureus nos::erm mutant (Sapp et al., 2014) consistently displayed a decreased optical

density (OD600) phenotype when grown aerobically in either LB medium or in TSB

lacking glucose (TSB-G)(Figure 2-1). Starting at 2 hours growth (exponential phase) in

LB or 4 hours growth (late-exponential phase) in TSB-G, the OD600 measurements of

the nos mutant in both media lacking glucose were slightly lower than the wildtype

(clinical MSSA strain UAMS-1) and nos complement cultures (Figure 2-1). While not a

drastic decrease, statistical analysis confirmed a significant difference in both LB (P <

0.001, Holm-Sidak method) and TSB-G (P = 0.001, Holm-Sidak method) at 6 hours

growth. Cell viability may account for the decrease in OD600 when grown in LB, as

corresponding CFU/ml counts were slightly lower in the UAMS-1 nos mutant compared

to wildtype. In the TSB-G growth condition, corresponding CFU/ml counts were

comparable between wildtype, nos mutant, and complement strains at all time points;

suggesting that the decreased OD600 was not due to decreased viability of the nos

mutant (Figure 2-1). As well, generation time for all growth conditions was calculated

using a previously-described formula (Todar, 2006). Measurements of generation time

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by CFU/ml in the TSB-G condition show no difference between the nos mutant (43 ± 2

minutes) versus wildtype (42 ± 2 minutes) and nos complement (37 ± 3 minutes)

cultures (Table 2-1). No measurable differences were observed in pH and wet weight

between UAMS-1 wildtype and nos mutant cells grown in TSB-G (data not shown).

Interestingly, the decrease in nos mutant OD600 appeared to be specific to aerobic

growth in media lacking glucose, a growth condition in S. aureus that promotes

exponential-phase aerobic respiration fueled by amino acid catabolism and the TCA

cycle (Somerville et al., 2002). When grown aerobically in TSB containing 14 mM

glucose (TSB)(a growth condition that promotes exponential-phase glycolysis of

glucose to acetate and repression of TCA cycle activity)(Somerville et al., 2002,

Somerville et al., 2003b), OD600 and CFU/ml growth curves of the wildtype, nos mutant,

and complement strains were almost identical to each other (Figure 2-1). This

corresponded with almost identical generation times (Table 2-1). Interestingly, when

graphed in linear scale (data not shown), the nos mutant showed a slight decrease in

OD600 starting at post-exponential phase growth, a condition where the TCA cycle

begins to function. Low O2 growth curves of UAMS-1 wildtype and nos mutant cultures

in TSB did not show an obvious difference in growth between the strains (data not

shown).

To verify that loss of saNOS-derived NO production was responsible for the

lower OD600 in the nos mutant, chemical NO donor (DPTA NONOate) was added to nos

mutant TSB-G cultures at the time of inoculation (Figure 2-2). As expected,

exogenously added NO was able to complement the OD (Table 2-1) phenotype of the

nos mutant to wildtype values without affecting cell viability. To confirm that this nos

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mutant OD effect was not a phenomenon specific to the clinical MSSA strain UAMS-1,

aerobic TSB-G growth curves were also completed in LAC-13C, a plasmid-cured

derivative of community-acquired methicillin resistant S. aureus (CA-MRSA) strain LAC

(Fey et al., 2013, 2003), and its isogenic nos::erm mutant (Figure 2-2). Similar to the

UAMS-1 nos mutant, the OD phenotype was also observed in the LAC-13C nos mutant

(Figure 2-2). Unlike the UAMS-1 nos mutant grown in TSB-G, slightly decreased

CFU/ml values and slightly increased generation time were observed in the LAC-13C

nos mutant relative to the parental strain (Figure 2-2 and Table 2-1).

To determine if alterations in cell morphology accounted for the decreased OD600

phenotype observed in the nos mutant, both scanning electron microscopy (SEM) and

transmission electron microscopy (TEM) were performed on cells isolated from UAMS-1

wildtype, nos mutant, and complement strains grown to stationary phase in TSB-G

(Figs. 2-3 and 2-4). Although TEM analysis of wildtype and nos mutant did not reveal

any apparent differences in cell wall structure or presence of intracellular inclusion

bodies (Figure 2-3), SEM analysis revealed that the nos mutant cells were found to

have an elongated shape relative to wildtype and nos complement cells (Figs. 2-4). This

qualitative observation was confirmed by measuring the length of each whole cell (from

its longest point) in 12-14 fields of view per strain, which verified that the nos mutant

cells were significantly longer than those of the wildtype and complement strains (P

<0.05 Holm-Sidak test; Figure 2-4).

saNOS Has an Altered Transcriptome.

To tease out how saNOS may be affecting S. aureus cell physiology during

aerobic respiration, RNAseq was performed using the IonTorrent PGM platform on RNA

isolated from aerobic TSB-G cultures of UAMS-1 and its isogenic nos mutant at 4 (late-

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exponential phase) and 6 hours (stationary phase). Expression changes (≥ 2-fold) in the

nos mutant were observed for 403 genes at 4 hours growth and 226 genes at 6 hours

growth. Strikingly, expression of multiple genes associated with oxidative and nitrosative

stress resistance (trxA, SAR1984, SAR1492, ahpF, msrA1, qoxC, ldh1, hmp, and scdA)

were altered in the nos mutant at 4 hours growth (Figure 2-5, Table 2-2). In addition,

several metabolic genes, including those associated with anaerobic

metabolism/fermentation (pfl, narG, SAR2013, SAR2210, nrdG, and ldh2, ackA),

pyruvate and carbohydrate metabolism (pyk, lac operon, nanA, fda, gap, and pgi),

amino acid metabolism (SAR1143, otc, SAR1836, and lysA), and cytochrome

biosynthesis/assembly (hemA, cta and qox operons), were all expressed at higher

levels in the nos mutant at this time point. Other notable expression changes in the nos

mutant at 4 hours growth included highly down-regulated expression of purine (pur; -3.2

to -77.1 fold) and pyrimidine (pyr; -2.5 to -7.5 fold) biosynthesis operon genes, as well

as decreased expression of multiple virulence genes (geh, capG, and dltD), ribosome

and translation machinery genes (rpm, rps, rbf, rpl, infA, and gidB), and components of

the fatty acid degradation (fad) operon. Furthermore, 88 hypothetical proteins and 40

predicted small, non-coding sRNAs presented with altered expression (Figure 2-5), with

sRNAs being predicted by a previously published method (Carroll et al., 2016a). Similar

patterns of gene expression changes also occurred in the nos mutant at 6 hours growth

(Figure 2-6 and Table 2-2). Additional gene expression changes not observed at 4

hours growth included decreased expression of the perR peroxide operon regulator

gene (-5.1 fold), highly decreased expression of the fad operon genes (-18.2 to -20.6

fold), and increased expression (3.9 fold) of the alcohol dehydrogenase (adhA) gene.

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Highly decreased expression of the pur and pyr operons was not observed at 6 hours

growth. All genes altered in the nos mutant relative to wildtype that fit the cut-off criteria

(Fold-change greater than 2, percent unique reads greater than 80% in both samples,

expression value greater than 50 in at least one sample) are presented in Appendix B.

RNAseq data for a subset of the differentially-expressed genes at 4 hours growth

was confirmed by qRT-PCR on RNA isolated from wildtype, nos mutant, and

complement strains (Table 2-2). Fold-change expression levels were restored to near-

wildtype levels in the nos complement strain for all tested genes, with the exception of

SAR2006, which encodes an NAD synthetase and is divergently transcribed from the

nos gene (SAR2007). Analysis of the RNAseq reads aligned to SAR2006 in the nos

mutant suggests that these divergent transcripts originate near the insertion of the Erm

cassette, located at nucleotide 232 bp downstream of the nos start codon. However, the

nos mutant was complemented for all other phenotypes by supplying the nos gene in

trans on a plasmid, indicating that increased transcription of SAR2006 is not having an

effect on the nos mutant phenotypes presented in this study.

Intracellular and Secreted Metabolite Profiles of the nos Mutant

As indicated by the 4-hour RNAseq data described in Figure 2-5 and Table 2-2,

multiple genes associated with anaerobic metabolism and/or fermentation were

upregulated in the nos mutant when grown aerobically without glucose, an amino-acid

based growth condition that promotes exponential phase TCA cycle activity and cell

respiration. To confirm that the nos mutant has an altered metabolism relative to

wildtype and complement strains in this growth condition, we performed quantitative

targeted metabolomics analysis on 4 hour (late-exponential) cultures to detect both

cellular and secreted metabolites (organic acids, amino acids, nucleotides) using

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LC/MS/MS analysis (Table 2-4 and Figure A1-6). Interestingly, cellular lactate levels

were significantly reduced by 49% in the nos mutant relative to the wildtype and

complement strains (Table 2-4). Furthermore, comparison of the intracellular organic

acid composition of wildtype and nos mutant suggested that metabolites produced by

the oxidative branch of the TCA cycle (citrate, α-ketoglutarate) were decreased in the

nos mutant, whereas metabolites associated with the reductive branch of the TCA cycle

(fumarate, malate) were increased (Table 2-4). Specifically, compared to the nos

mutant, wildtype cells showed 30% and 54% decreases in citrate and α-ketoglutarate

levels, respectively; whereas fumarate and malate levels were increased in the nos

mutant by 158% and 62%, respectively. Interestingly, extracellular levels of α-

ketoglutarate were also significantly lower in the nos mutant, suggesting that the nos

mutant may be importing and/or consuming α-ketoglutarate at a higher rate.

Amino acid composition of the wildtype, nos mutant, and complement strains was

also quantified by LC/MS/MS analysis. Because saNOS catalyzes the two-step

oxidation of L-arginine (Arg) to L-citrulline (Ctl) and NO, we expected Ctl levels to be

lower in the nos mutant. Surprisingly, cellular Ctl levels were significantly higher in the

nos mutant, whereas Arg levels were similar between wildtype and nos mutant cells

(Table 2-4). Statistically-significant decreases in nos mutant cellular amino acids were

also observed for glutamate (Glu) and all branched-chain amino acids (Leu, Ile, Val),

whereas histidine (His) levels were significantly increased (Table 2-4). Although not

statistically significant, glutamine (Gln) and ornithine (Orn) levels were also decreased

in the nos mutant cells relative to wildtype and complement strains. There were no

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significant differences in extracellular levels of amino acids were observed, suggesting

that amino acid transport is not altered in the nos mutant.

To determine if redox balance and ATP levels were altered in the nos mutant,

nicotinamide nucleotides and adenosine phosphates levels were also measured by

LC/MS/MS analysis. NADH levels (reduced 65%) were significantly lower in the nos

mutant (P =0.015 Two-tailed t-test) whereas NAD+ levels were only 23% lower and not

statistically significant (Table 2-4). This result translated to a higher, but not statistically

significant, NAD+ to NADH ratio in the nos mutant. ATP levels were similar between

wildtype and nos mutant strains (Table 2-4 and Figure A-6). A pattern of lower AMP and

ADP levels in the nos mutant was also observed, which may be related to the

decreased expression of purine biosynthesis genes observed in this strain. Energy

charge, an index based on concentrations of ATP, ADP, and AMP used to measure the

energy status of biolological cells, was determined for each strain (Atkinson & Fall,

1967, Atkinson & Walton, 1967). Theoretically, these values range from 0 (all AMP) to 1

(all ATP), with ATP generating catabolic pathways found to be inhibited at a higher

energy charge (Atkinson & Walton, 1967). For an unknown reason, biological replicate

two of our panel of wildtype, nos mutant, and complement strains was a clear outlier for

only the adenosine nucleotides and is the obvious contributor to the large error bars

observed (Figure A-6). Therefore, calculations of energy charge were completed without

including this outlier, and showed no clear difference between wildtype and nos single

mutant. Therefore, no overall change in cellular energy metabolism was observed in the

nos mutant.

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Aim 2. saNOS Contributes to Endogenous Oxidative Stress and Respiratory Metabolism

Mutation of nos Increases Endogenous Oxidative Stress

In B. subtilis, B. anthracis, and S. aureus, a hallmark of nos mutation is an

increased sensitivity to exogenous oxidative stress (van Sorge et al., 2013, Sapp et al.,

2014, Gusarov & Nudler, 2005, Shatalin et al., 2008). Although S. aureus is subjected to

exogenous sources of oxidative stress from the host immune system, ROS are also

naturally produced during respiration by the bacterium's own metabolism. In line with

this, the RNAseq data described in Aim 1 suggested that the nos mutant may be

subjected to increased endogenous oxidative stress. Multiple genes associated with

oxidative stress (msrA1, ahpF, trxA), heme biosynthesis (hemA), as well as iron storage

and iron-sulfur cluster repair (scdA, SAR1492, SAR1984) presented with increased

expression at 4 hours growth. Likewise, expression of perR, a negative regulator of

oxidative stress genes such as catalase (katA), alkyl hydroperoxide reductase (ahpCF),

and thioredoxin reductase (trxB)(Horsburgh et al., 2001a), was decreased in the nos

mutant at 6 hours growth. To determine if the nos mutant indeed accumulates more

intracellular ROS, cells collected from mid-exponential (3 hours growth) and stationary

phase (6 hours growth) aerobic cultures of wildtype, nos mutant, and nos complement

strains were subjected to staining with the fluorescent cell-permeable general ROS

indicator carboxy- 2′,7′-dichlorofluorescein (CM-H2DCFDA) (Jakubowski & Bartosz,

2000, LeBel et al., 1992). By this approach the nos mutant was found to accumulate

significantly (P <0.001 Tukey test) increased levels of intracellular ROS relative to the

wildtype and complement strains in both TSB-G and TSB cultures, conditions that

promote and inhibit the TCA cycle, respectively (Figure 2-7). There are multiple

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potential sources of endogenous ROS within S. aureus cells undergoing respiration, but

a likely candidate is O2-, a natural by-product of aerobic respiration (Messner & Imlay,

1999). The O2- specific stain MitoSOX Red (Robinson et al., 2006) was therefore

employed to determine if O2- levels were altered in the nos mutant when grown

aerobically in TSB-G (Figure 2-7). Similar to the general intracellular ROS levels, nos

mutant cells also demonstrated increased intracellular O2- levels relative to the wildtype

and nos complement strains.

Given that B. subtilis NOS-derived NO is implicated in the direct activation of

catalase after it has been naturally inhibited by free Cys (Gusarov & Nudler, 2005), the

increased intracellular ROS observed in the S. aureus nos mutant may have been an

indirect result of impaired catalase activity. Therefore, catalase activity in cytosolic

proteins extracted from wildtype, nos mutant, and nos complement strains grown in

aerobic TSB-G cultures was quantified by measuring the amount of unreacted H2O2

using Amplex Red in the presence of horseradish peroxidase (Zhou et al., 1997,

Mohanty et al., 1997). As indicated in Figure 2-7, catalase activity was not decreased in

the nos mutant, and in fact was measurably increased relative to wildtype and

complement strains, possibly in response to increased endogenous ROS accumulation.

Collectively, these results demonstrate that, in addition to promoting resistance to

exogenous oxidative stress, saNOS helps curtail the production or accumulation of

endogenous ROS during aerobic growth.

saNOS Contributes to Respiratory Function

A defined relationship between NO, NOS, and modulation of cell respiration is

well established in mammals (Giulivi et al., 2006, Parihar et al., 2008a, Larsen et al.,

2012). Although NOS has not been previously found to modulate respiration in bacteria,

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exogenously added NO can slow bacterial respiration by competing with O2 at the final

step of the electron transport chain (Junemann & Wrigglesworth, 1996, Borisov et al.,

2004, Butler et al., 2002, McCollister et al., 2011). These published studies, combined

with the gene expression profiles and increased intracellular ROS observed in the nos

mutant (Figs. 2-5 and 2-7), led us to hypothesize that loss of saNOS activity in the nos

mutant affects some aspect of aerobic respiration. To test this, the membrane potential

of wildtype, nos mutant, and complement strains was measured in aerobic TSB-G

cultures using the carbocyanine dye DiOC2(3) and a previously-described flow-

cytometry method (Novo et al., 1999, Lewis et al., 2015, Shapiro & Nebe-von-Caron,

2004). This dye first stains all the cells green and then aggregates within the cell in a

membrane potential dependant manner. Once in the cell the stain fluoresces red and

the red:green ratio can be determined by flow cytometry. At both 3 and 6 hours growth

(corresponding to mid-exponential and stationary growth phase, respectively), the nos

mutant presented with an increased membrane potential (as reflected by an increased

red:green fluorescence ratio) relative to the wildtype and complement strains (Figure 2-

8). A chemical NO donor (DPTA NONOate) was also employed in these experiments to

determine if NO itself could complement the membrane potential phenotype of the nos

mutant. Addition of 100 µM DPTA NONOate at time of inoculation of aerobic TSB-G

cultures was able to restore the membrane potential of the nos mutant near to wildtype

levels, but had a minimal effect on the membrane potential of the wildtype strain (Figure

2-8). Since TCA cycle activity of S. aureus is inhibited during exponential growth in the

presence of glucose (Somerville et al., 2002), membrane potential was also assessed in

the wildtype, nos mutant, and complemented strains during exponential growth phase in

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TSB. Interestingly, membrane potential was increased in TSB cultures of nos mutant

relative to wildtype, but the degree of shift was slightly smaller (Figure 2-8), supporting

the idea that NADH levels generated by glycolysis likely support some respiratory

activity under this condition. Interestingly, in mammals a direct relationship between

NO-mediated respiratory inhibition and a decrease in membrane potential has been

observed (Mastronicola et al., 2004).

Cell respiratory activity in aerobic TSB-G cultures was also measured by staining

cells with 4.5 mM 5-Cyano-2,3-ditolyl tetrazolium chloride (CTC), a compound that can

be reduced by respiratory dehydrogenases into a insoluble highly-fluorescent CTC

formazan product (Smith & McFeters, 1997). After 3 hours growth (mid-exponential

growth phase), increased fluorescence was observed in the nos mutant relative to the

wildtype strain (Figure 2-9). This nos mutant phenotype was also observed during

growth in TSB (Figure 2-9), and CTC fluorescence was restored to wildtype levels under

both growth conditions in the complement strain. To determine if NO itself could

complement the nos mutant CTC phenotype, DPTA NONOate was also added to

cultures at time of inoculation, and was shown to restore the level of nos mutant CTC

reduction to wildtype levels at 3 hours growth (Figure 2-9). Interestingly, when this

experiment was repeated in TSB-G at 6 hours growth (stationary phase), the opposite

effect was observed in the nos mutant, whereby decreased CTC reduction (decreased

fluorescence) was observed relative to the wildtype strain (Figure 2-9). Again, this

phenotype was complemented by adding NO donor to nos mutant cultures at the time of

inoculation. It is possible that by 6 hours growth the nos mutant has switched to an

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alternative electron donor to drive respiration, possibly accounting for the different nos

mutant CTC staining patterns between these growth phases.

Elevated membrane potential and CTC staining in the nos mutant suggested to

us that respiration may be altered in this strain. It is well established that inhibition of

cytochrome oxidase by NO or KCN can slow respiration, similar to what may be

occuring in wildtype cells containing NOS (Messner & Imlay, 1999, Pearce et al., 2008).

With this in mind, O2 consumption was measured with an O2 Clark-type electrode

attached to a free radical analyzer (TBR-4100, World Precision Instruments) in cells

harvested from aerobic TSB-G cultures. Contrary to what was expected, comparison of

wildtype and nos mutant respiratory rates using this method showed that nos mutant O2

consumption trended towards a non-statistically significant decrease relative to wildtype

and nos complement strains (Figure 2-10). It is possible that this method is not sensitive

enough to measure more subtle differences in respiratory rates and therefore the

measured decrease may actually be biologically relevant. Nevertheless, this O2

consumption pattern was the opposite of what was expected and suggests that altered

respiratory phenotypes in the nos mutant are not likely due to NOS-derived NO

cytochrome inhibition. Although the mechanism behind these respiratory phenotypes is

unknown, it is possible that the elevated CTC staining and membrane potential are a

result of increased "proton backpressure" (the passive movement of protons from

outside to inside the cell membrane independent of the action of ATP synthase), which

can occur when respiration is inhibited and could increase the membrane potential

and/or backup electrons onto respiratory dehydrogenases (van Rotterdam et al., 2001,

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Lieberman et al., 2007). Taken together, these results indicate that saNOS influences

some aspect of the respiratory chain that is required for proper respiratory function.

Inhibition of Ndh Limits Oxidative Stress in a nos Mutant

Endogenously produced ROS in the form of O2- occurs naturally during aerobic

respiration by accumulation of electrons on respiratory chain flavoproteins, which

incompletely reduce O2 to O2- (Minghetti & Gennis, 1988, Messner & Imlay, 1999,

Messner & Imlay, 2002). In an attempt to determine if elevated ROS levels in the nos

mutant were due to disruption of proper respiratory function, an Ndh inhibitor was

employed. Thioridizine HCl (TZ) specifically inhibits S. aureus Ndh activity and was

found to not affect respiration by alternative electron donors such as succinate, malate,

and lactate (Schurig-Briccio et al., 2014). Addition of TZ to wildtype and nos mutant

cultures at time of inoculation substantially decreased overall ROS levels in aerobically

growing cultures (Figure 2-11). While TZ decreased ROS in both wildtype (-26%) and

nos mutant (-40%) cultures, the magnitude of this decrease was greater when the nos

mutant was treated with this Ndh inhibitor. While elevated ROS in the nos mutant could

be due to a variety of factors, these data suggest that increased endogenous ROS is

likely due, in part, to disruption of proper respiratory function.

Superoxide is well established to attack the Fe-S cluster of aconitase, often

making aconitase enzymatic activity an indirect measurement of cellular oxidative stress

(Gardner & Fridovich, 1991b, Gardner, 2002). Therefore, aconitase activity was also

determined in wildtype, nos mutant, and complement cells by a coupled reaction, which

measure the rate of NADPH (340 nm) production generated by isocitrate

dehydrogenase (Rose & O'Connell, 1967). As predicted, aconitase enzymatic activity of

the nos mutant was significantly lower than that of the wildtype and nos complement

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strains (Figure 2-11). In an attempt to promote stability of aconitase in cell lysate

preparations, samples were also isolated in anaerobic vials in a parallel experiment, but

failed to improve the sensitivity of this assay, as aconitase activity in wildtype samples

was comparable between both isolation methods (data not shown). Lower aconitase

activity in the nos mutant could be due to altered gene expression or protein

levels/stability. Examination of the RNAseq gene expression data showed no altered

expression of aconitase transcripts (Appendix B). Western blotting analysis to determine

aconitase protein levels was also attempted with an anti-aconitase antibody against the

eukaryotic aconitase most similar to bacterial aconitase, but it was unsuccesful. While

TZ-treated nos mutant cells presented with decreased accumulation of ROS (Figure 2-

11), aconitase activity in the nos mutant was not restored when cultures were grown

with TZ (Figure 2-11). These combined data support the assertion that Ndh contributes

to elevated levels of ROS in the nos mutant, but this increased endogenous ROS is

likely not causing the observed decrease in aconitase activity.

Aim 3. SrrAB as a Potential Regulator of nos Mutant Metabolic Adaptation

Growth Phenotypes of the nos srrAB Double Mutant

Global transcriptional and metabolic responses were observed in response to S.

aureus nos mutation (Aim. 1), many of which were related to anaerobic and respiratory

metabolism, as well as response to radical stress. Two major metabolic regulators of

anaerobic metabolism and stress response in S. aureus are SrrAB and Rex (See

Chapter 1). Previously-published RNA microarray analysis of S. aureus wild-type and

srrAB mutants exposed to nitrosative stress show that SrrAB regulates anaerobic

metabolism (narG, pflB, and nrdG), nitrosative stress (scdA and hmp), and cytochrome

biosynthesis genes (qox, cta) (Kinkel et al., 2013), all of which presented with altered

98

expression in the nos mutant (Table 2-2). SrrAB is also thought to sense the reduction

state of the quinone pool (Kinkel et al., 2013), which is possibly altered in the nos

mutant due to elevated membrane potential and altered Ndh activity (Figs. 2-8 and 2-9).

Rex responds to the NAD/NADH ratio and controls expression of anaerobic metabolism

genes, including lactate dehydrogenase (ldh1 and ldh2), pyruvate formate lyase (pflB),

nitrate reductase (narG), and flavohemoprotein (hmp) genes (Pagels et al., 2010).

Additionally, NADH levels were 65% lower in the nos mutant, whereas the NAD/NADH

ratio was increased 122%, but not statistically significant compared to wildtype (Table 2-

4). Given that many of the genes regulated by SrrAB and/or Rex showed increased

expression in the nos mutant, nos srrAB and nos rex double mutants were generated in

the UAMS-1 strain background to determine if these regulatory systems contribute to

the altered metabolism and growth phenotypes observed in the nos mutant. Basic

growth assays (agar plate, growth curves) showed that the nos rex double mutant did

not present with any obvious growth phenotypes differing from the nos single mutant

(data not shown). On the other hand, the nos srrAB double mutant was characterized by

smaller colonies on agar plates when compared to the wildtype and nos and srrAB

single mutant strains (Figure 2-12 and 2-13). Further focus was therefore placed on the

nos srrAB double mutant. When grown aerobically in TSB-G, a nos srrAB double

mutant presented with a drastically lower OD600 when compared to wildtype and both

nos and srrAB single mutants (Figure 2-14). Complementation of this phenotype was

completed by adding nos back to the double mutant. Generation time of the nos srrAB

double mutant (85 ± 21) was much higher than wildtype (42 ± 2 minutes) (Table 2-1)

and the double mutant also presented with an altered growth curve compared to all

99

other strains (Figure 2-14). Compared to other strains, slight decreases in OD600 and

CFU/ml were also observed when the nos srrAB double mutant when grown in TSB with

glucose (Fig 2-14), a growth condition where the cells are primarily undergoing

glycolysis during aerobic exponential growth. This corresponded with a slightly altered

TSB growth curve and an increase in generation time of the nos srrAB double mutant

(Figure 2-14 and Table 2-1). Notably, none of the growth characteristics of the nos

srrAB double mutant were observed in the single srrAB mutant.

Membrane Potential of the nos srrAB Double Mutant

Predicted to sense the reduction state of the respiratory chain (Kinkel et al.,

2013), SrrAB is closely related to the respiratory metabolism of S. aureus. Altered

respiratory phenotypes in the nos mutant suggested that SrrAB may sense altered

respiration and regulate genes in response to this metabolic signal. Therefore

membrane potential was measured in the nos srrAB double mutant and each single

mutant strain. Mutation of srrAB alone showed an obvious decrease in the membrane

potential relative to wildtype (Figure 2-15). However, additional mutation of nos in the

srrAB mutant background caused an increase in membrane potential relative to wildtype

(Fig 2-15). Therefore, nos and srrAB likely contribute to membrane potential in opposing

ways. Trans complementation of the nos srrAB double mutant with nos only partially

restored membrane potential to near wildtype levels, possibly because the high copy

nos complementation plasmid is somehow preventing complete reduction in membrane

potenital as seen in the single srrAB mutant.

Metabolism of the nos srrAB Double Mutant

Examination of the nos srrAB double mutant growth curves (Figure 2-14)

suggests that this strain may be undergoing a fermentative/glycolysis-based

100

metabolism, similar to what is typically seen in the wild-type strain during aerobic growth

in the presence of glucose (Figure 2-14) (Somerville et al., 2002). This observation,

combined with the fact that SrrAB regulates multiple fermentative and anaerobic

metabolism genes, led to the hypothesis that metabolism may be altered in the nos

srrAB double mutant. Therefore, targeted metabolomics (LC/MS/MS) was completed on

wildtype, nos mutant, srrAB mutant, and nos srrAB double mutant strains; as well as the

double mutant complemented with nos from 4-hour cultures. This time point was chosen

to match the nos mutant/wild-type RNAseq data, but the nos srrAB double mutant

cultures are in a slightly different growth phase, therefore the following data needs to be

interpreted with this caveat in mind. Both intracellular and extracellular metabolites were

tested for organic acid (Figure A-1 and A-2) and amino acid (Figure A-3 and A-4)

composition, as well as intracellular NAD and ATP nucleotides (Figure A-5 and A-6).

The srrAB single mutant presented with significantly elevated cellular fumarate and

malate, with both of these organic acids being higher in the extracellular media as well.

Similar to the nos single mutant, a significant decrease in intracellular lactate was also

observed in the srrAB single mutant. Amino acid profiles of the srrAB single mutant

showed significant increases in cellular BCAAs (Ile, Leu, Val) as well as a decrease in

Glu. Aside from a decrease in cellular NADP, the srrAB single mutant presented with no

other significant differences in NAD or ATP nucleotides.

Combined mutation of srrAB and nos caused markedly different metabolite

profiles than each single mutation. Levels of intracellular organic acids generated by the

TCA cycle were significantly lower in the nos srrAB double mutant compared to wildtype

(Table 2-6). These differences in metabolites included succinate and malate, with levels

101

of citrate and fumarate being below the limit of quantitation (BLOQ). At the same time,

extracellular levels of α-ketoglutarate were significantly lower, similar to what was seen

in the nos single mutant. Extracellular lactate levels of aerobically growing nos srrAB

double mutant cells were significantly higher, with a 8593% increase compared to

wildtype (Table 2-7). This drastic increase in extracellular lactate supports the assertion

that these cells may be undergoing a fermentative metabolism with subsequent lactate

secretion. Other notable differences include significantly higher levels of extracellular

pyruvate and malate, suggesting that the nos srrAB double mutant is either secreting

these organic acids or is impaired in uptake pathways.

Drastic changes in the intracellular and extracellular amino acid profile of the nos

srrAB double mutant were also observed. The nos srrAB double mutant presented with

significant decreases in the cellular levels of multiple amino acids including Ala, Asn,

Asp, Glu, Lys, Pro, and Val, with Arg being BLOQ (Table 2-6). At the same time,

extracellular levels of these same amino acids were significantly higher (Table 2-7).

Thus, the nos srrAB double mutant has an apparent shut down of multiple amino acid

transport pathways. Levels of intracellular Met, Orn and Tyr were also significantly lower

in the nos srrAB double mutant, while extracellular levels of these metabolites were

unaffected. A generalized decrease in amino acid transport was further supported by

the significant increase in extracellular Gln, Leu, Ser, and Thr relative to wildtype (Table

2-7). Similar to the nos single mutant, a significant (347%) increase in intracellular Ctl

levels was also observed in the double mutant (Table 2-6). The nos srrAB double

mutant has therefore retained some of the metabolic properties observed in the nos

single mutant.

102

A characteristic difference between TCA cycle metabolism and fermentative

metabolism is the redox status of the cell. With this in mind, NADH levels were

significantly lower in the nos srrAB double mutant relative to wildtype, which translated

to a significant increase in the NAD/NADH ratio (Table 2-6). Additionally, molecules

required to provide reducing equivalents for biosynthetic pathways such as NADP were

significantly lower or BLOQ (NADPH). Calculations of energy charge showed no

difference between wildtype and srrAB single mutant, but a slight decrease in energy

charge was observed for the nos srrAB double mutant (Table 2-5). These combined

results support a metabolic situation where the nos srrAB double mutant is undergoing

a fermentative metabolism with shutdown of the TCA cycle and biosynthetic pathways,

and a slight decrease in the overall energy status of the cell.

103

Figure 2-1. Wildtype and nos mutant growth curves. A-B: UAMS-1 wildtype, nos mutant,

and complement strains were inoculated to an OD600 = 0.05 in LB media, and grown with aeration (250 RPM; 1:12.5 volume to flask ratio) at 37°C. Growth over a 8 hour period was monitored by OD600 measurements (A) and CFU/ml by serial dilution plating (B). C-D: UAMS-1 wildtype and nos mutant cultures were grown in TSB-G as described above. OD600 measurements (C) and CFU/ml (D) were determined over a 24 hour period. E-F: UAMS-1 wildtype and nos mutant cultures were grown in TSB as described in C-D. OD600 measurements (E) and CFU/ml (F) were determined. Data points represent the average of 3 independent experiments, error bars = SEM.

Time (Hours)

0 5 10 15 20 25

OD

600

0.1

1

10

UAMS-1nos mutantnos complement

Time (Hours)

0 5 10 15 20 25

CF

U/m

l

107

108

109

1010

UAMS-1nos mutantnos complement

E F

Time (Hours)

0 5 10 15 20 25

CF

U/m

l

107

108

109

1010

UAMS-1nos mutantnos complement

Time (Hours)

0 5 10 15 20 25

OD

600

0.1

1

10

UAMS-1nos mutantnos complement

C D

Time (Hours)

0 2 4 6 8

OD

600

0.1

1

10

UAMS-1nos mutantnos complement

Time (Hours)

0 2 4 6 8

CF

U/m

l

1e+6

1e+7

1e+8

1e+9

1e+10

UAMS-1nos mutantnos complement

A B

104

Figure 2-2. Growth curves with addition of chemical NO donor and in a MRSA background. A-B: UAMS-1 wildtype and nos mutant cultures were inoculated to an OD600 = 0.05 in TSB-G media, and grown with aeration (250 RPM; 1:12.5 volume to flask ratio) at 37°C for 8 hours. Chemical NO donor was added at the time of inoculation to indicated cultures followed by determination OD600 (A) and CFU/ml (B). C-D: LAC-13C wildtype and nos mutant cultures were grown in TSB-G for 24 hours with subsequent OD600 measurements (C) and CFU/ml (D) determination. Data points represent the average of 3 independent experiments, error bars = SEM.

Time (Hours)

0 5 10 15 20 25

CF

U/m

l

107

108

109

1010

LAC-13Cnos mutant

Time (Hours)

0 5 10 15 20 25

OD

600

0.1

1

10

LAC-13Cnos mutant

Time (Hours)

0 2 4 6 8

CF

U/m

l

107

108

109

1010

UAMS-1nos mutantnos mutant + NO

Time (Hours)

0 2 4 6 8

OD

600

0.1

1

10

UAMS-1nos mutantnos mutant + NO

C D

BA

105

Figure 2-3. TEM analysis of nos mutant.: Cells were harvested from 6 hour TSB-G

cultures of wildtype (A and C) and nos mutant (B and D) strains, and samples of each were prepared for TEM. Images are at 30,000X A-B) or 100,000X (C-D) magnification and are representative of 16 random fields of view/condition and 1 biological replicate. White scale bar = 1 µM (A-B) or 0.2 µM (C-D). Photo courtesy of author.

106

Figure 2-4. SEM analysis of nos mutant. A-C: Cells were harvested from 6 hour TSB-G

cultures of wildtype A), nos mutant B), and nos complement C) strains, and samples of each were prepared for SEM. Images are at 50,000X magnification and are representative of 2 stubs and 12-14 random fields of view/condition. Dotted white scale bar = 0.6 µM. D: Cell length (in nm) was measured using ImageJ by measuring the largest diameter for all measurable cells in all fields of view. * statistical significance (P <0.05, Holm-Sidak method) relative to wildtype. Line in box = median; lower and upper box lines = 25th and 75th percentiles; whiskers = error bars (10th and 90th percentiles); dots = outliers (5th/9th percentiles). Photo courtesy of author.

Ce

ll len

gth

(n

m)

400

500

600

700

800

900

D

*

Wildtype

nos

mutant

nos

complement

A

B

C

107

Figure 2-5. Distribution of gene functional categories expressed by the nos mutant in 4

hour cultures. RNA isolated from 4 hour aerobic wildtype and nos mutant TSB-G cultures was subjected to RNAseq transcriptome profiling. Differential expression analysis and cutoff criteria were applied as described in the Materials and Methods. Only genes that met the cutoff criteria were included in this analysis. Functional classification was completed by NCBI gene annotations and pathway analysis using BioCyc software (www.biocyc.org). Total number of down-regulated genes (fold-change > 2.0; black bars) = 199; total number of up-regulated genes (fold-change > 2.0; grey bars) = 204.

Number of Genes

0 10 20 30 40 50 60

Translation Proteins and tRNA

Purine and Pyrimidine Biosynthesis

Capsular Biosynthesis

Proteases, Protein Folding and Degradation

Mevalonate Pathway

Iron Storage and Protein Biosynthesis

Fatty Acid Oxidation

Transcriptional Regulation

TCA cycle and Intermediate Metabolism

Lipoproteins

Virulence

DNA Replication and Modification

Hypothetical Proteins

Predicted Small RNAs

Other Metabolic Pathways

Other Transport Proteins

Ion Dependant Transporters

Amino Acid Transport

Purine and Pyrimidine Degradation

Amino Nucleotide/Sugar metabolism

General Signal Transduction

ABC Transporters

Two Component Systems

PTS Systems

Stress Response

Anaerobic Metabolism and Fermentation

Amino Acid Metabolism

Pyruvate and Carbohydrate Metabolism

Electron Transport Chain Proteins and Component Biosynthesis Downregulated Upregulated

108

Figure 2-6. Distribution of gene functional categories expressed by the nos mutant

relative to wildtype of 6 hour cultures. RNA isolated from 6 hour aerobic wildtype and nos mutant TSB-G cultures was subjected to RNAseq transcriptome profiling. Differential expression analysis and cutoff criteria were applied as described in the materials and methods. Only genes that met the cutoff criteria were included. Functional classification was completed by NCBI gene annotations and pathway analysis using MetaCyc software. Total number of down-regulated genes (fold-change > 2.0; black bars) = 106; total number of up-regulated genes (fold-change > 2.0; grey bars) = 118.

Number of Genes

0 5 10 15 20 25 30 35

Translation Proteins and tRNA

Purine and Pyrimidine Biosynthesis

Capsular Biosynthesis

Proteases, Protein Folding and Degradation

Mevalonate Pathway

Iron Storage and Protein Biosynthesis

Fatty Acid Oxidation

Transcriptional Regulation

TCA Cycle and Intermediate Metabolism

Lipoproteins

Virulence

DNA Replication and Modification

Hypothetical Proteins

Predicted Small RNAs

Other Metabolic Pathways

Other Transport Proteins

Ion Dependant Transporters

Amino Acid Transport

Purine and Pyrimidine Degradation

Amino Nucleotide/Sugar metabolism

General Signal Transduction

ABC Transporters

Two Component Systems

PTS Systems

Stress Response

Anaerobic Metabolism and Fermentation

Amino Acid Metabolism

Pyruvate and Carbohydrate Metabolism

Electron Transport Chain Proteins and Component Biosynthesis Downregulated Upregulated

109

Figure 2-7. Intracellular ROS, superoxide detection, and catalase activity in wildtype and

nos mutant cultures. Wildtype, nos mutant, and complement strains were inoculated to an OD600 = 0.05 in TSB-G A) or TSB B) and grown aerobically at 37°C (for 3 and 6 hours in A, and 3 hours in B), followed by CM-H2DCFDA staining to detect intracellular ROS. After staining, 200 µl aliquots of each cell suspension were immediately transferred in triplicate to a 96-well plate, and incubated at 37°C in a Synergy HT fluorescent plate reader. Fluorescence and OD600 measurements were recorded, and data were reported as relative fluorescent units (RFU) per OD600. Cultures for superoxide staining C) were grown in TSB-G for 3 hours as above and then subjected to MitoSOX Red staining to detect O2

-. Catalase D) activity of protein isolated from 3 hour TSB-G cultures was measured using the Amplex Red Catalase Activity Kit (Life Technologies), respectively. 1 mg/ml porcine heart aconitase was included as a positive control. All data represent the average of n = 3 independent experiments and error bars = SEM. *statistical significance (P <0.001, Tukey test) relative to wildtype; **statistical significance (P <0.05, Holm-Sidak method) relative to wildtype.

Flu

ore

scen

ce (

RF

U/O

D600)

0

200

400

600

800

1000

3 Hours 6 Hours

Flu

ore

scen

ce (

RF

U/O

D600

)

0

500

1000

1500

2000

2500

3000Wildtypenos mutantnos complement

Cata

lase A

cti

vit

y (

U/m

g p

rote

in)

0

105

2x105

3x105

4x105

**

*A

DC**

*

Flu

ore

scen

ce (

RF

U/O

D600)

0

500

1000

1500

2000

2500

B*

110

Figure 2-8. Effect of saNOS on membrane potential. Aerobic TSB-G cultures of

wildtype, nos mutant, and nos complement strains were grown for 3 hours A) with addition of NO donor to wildtype and nos mutant cultures at time of inoculation, as indicated. Cells pellets were then harvested and stained with 30 µM of the membrane potential stain 3,3’-diethyloxacarbocyanin iodide (DiOC2(3)), and subjected to flow cytometry to detect the ratio of red to green fluorescence. Histograms represent the ratio of red to green fluorescence (X axis) plotted against the number of events (Y axis). A shift to the right of the vertical black line indicates an increase in membrane potential. B) TSB-G cultures were harvested at 6 hours and treated as above. C) Samples were grown for 3 hours in TSB and treated as above. Data are representative of n = 6 (A), n = 4 B), or n = 6 C) biological replicates.

Even

tsE

ven

ts

Wildtype

nos mutant

nos complement

Wildtype

Wildtype + NO

nos complement

nos mutant

nos mutant + NO

A

B

Fluorescence (Red:Green Ratio)

Fluorescence (Red:Green Ratio)

Even

ts

Wildtype

nos mutant

nos complement

C

Fluorescence (Red:Green Ratio)

111

Figure 2-9. Respiration determined by CTC staining. Aerobic cultures of wildtype, nos

mutant, and complement strains were grown in TSB-G for 3 hours without NO donor (A). Separate experiments were completed with addition of NO donor and growth for 3 (B) or 6 (C) hours. Cell pellets were isolated and stained with 4.5 mM CTC. Fluorescence (RFU) was measured after 70 minutes of CTC staining with a Biotek Synergy microplate reader, and normalized to the initial OD600 reading of each sample. Fold-change was determined by dividing the RFU/OD600 of each condition by the average of wildtype RFU/OD600. (D) CTC staining was completed as above after growth for 3 hours in TSB. *statistical significance (P <0.005 Tukey test) relative to wildtype. **statistical significance (P <0.001 Holm-Sidak method) relative to untreated wildtype. ***statistical significance (P <0.001 Dunn’s method) relative to untreated wildtype. Data represent the average of n = 3 (A and D) or n = 4 (B-C) biological replicates. Error bars = SEM.

Fo

ld-c

han

ge (

Rela

tive t

o W

ild

typ

e)

0.0

0.2

0.4

0.6

0.8

1.0

1.2

C

***F

old

-ch

an

ge (

Rela

tive t

o W

ild

typ

e)

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

D

Fo

ld-c

han

ge (

Rela

tive t

o W

ild

typ

e)

0.0

0.2

0.4

0.6

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1.0

1.2

1.4

1.6F

old

-ch

an

ge (

Rela

tive t

o W

ild

typ

e)

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

A * **B

**

112

Figure 2-10. Effect of saNOS on oxygen consumption. Oxygen consumption of cultures

grown for 3 hours followed by resuspension in fresh air-saturated TSB-G. Oxygen consumption rate (%) was determined using a Clark type electrode by measuring the slope of the curve and normalizing to CFU/ml. Data is representative of n = 8 independent experiments. Error bars = SEM.

Rela

tive O

xyg

en

Co

nsu

mp

tio

n R

ate

(%

)

0

20

40

60

80

100

120

113

Figure 2-11. Intracellular ROS upon Ndh inhibition and aconitase activity of the nos

mutant. Wildtype and nos mutant strains were inoculated to an OD600 = 0.05 in TSB-G and grown aerobically at 37°C in the presence of 15 µM Thioridizine HCl as indicated. At 3 hours growth cells were A) stained with CM-H2DCFDA staining to detect intracellular ROS or B) isolated for aconitase activity. After staining CM-H2DCFDA (A), 200 µl aliquots of each cell suspension were immediately transferred in triplicate to a 96-well plate, and incubated at 37°C in a Synergy HT fluorescent plate reader. Fluorescence and OD600 measurements were recorded, and data were reported as relative fluorescent units (RFU) per OD600. Aconitase activity of cell lysates (B) was measured using the Aconitase Assay Kit (Cayman Chemical). 1 mg/ml porcine heart aconitase was included as a positive control. Data represents an n = 5 (A) and n = 4 (B) independent experiments. Error bars = SEM. *statistical significance (P <0.001, Paired t-test) relative to wildtype; **statistical significance (P <0.005, Hold-Sidak method) relative to wildtype.

Aco

nit

ase A

cti

vit

y(n

mo

l/m

in/m

g p

rote

in)

0

5

10

15

20

25

30

Flu

ore

scen

ce (

RF

U/O

D600

)

0

500

1000

1500

2000

2500UntreatedThioridizine Treated A*

**

B

114

Figure 2-12. Agar plate growth of the nos srrAB double mutant. Overnight cultures (~16

hours) of wildtype (1), nos mutant (2), nos complement (3), srrAB mutant (4), nos srrAB double mutant (5), and double mutant nos complement (6) were grown in TSB at 37°C and 250 rpms. 1 ml of fresh TSB was inoculated to an OD600 = 0.05 followed by dilutions and and track plating on TSB containing 5 µg/ml chloramphenicol. Plates were allowed to grow at 37°C for 24 hours before imaging. Images are representative of n = 3 biological replicates. Photo courtesy of author.

115

Figure 2-13. Quantification of colony size. Overnight cultures were grown in TSB at

37°C and 250 rpms. 1 ml of fresh TSB was inoculated to an OD600 = 0.05 followed by dilutions and and track plating on TSB containing 5 µg/ml chloramphenicol. Plates were allowed to grow at 37°C for 24 hours before imaging. Images were analyzed using OpenCFU software (Geissmann, 2013). Radius values are unitless and were determined "per object" by the software. Only colonies that were not clumped and clearly round were included in calculations. *statistical significance (P <0.001, Dunn’s method) relative to wildtype.

116

Figure 2-14. Growth curves of nos and nos srrAB double mutant strains. A-B: UAMS-1

wildtype, nos mutant, srrAB mutant, nos srrAB double mutant, and nos srrAB complement strains were inoculated to an OD600 = 0.05 in TSB-G media and grown with aeration (250 RPM; 1:12.5 volume to flask ratio) at 37°C. Growth over a 24 hour period was monitored by OD600 measurements (A) and CFU/ml by serial dilution plating (B). C-D: UAMS-1 wildtype and nos mutant cultures were grown in TSB as described above. OD600 measurements (C) and CFU/ml (D) were determined. Data points represent the average of 5 (A-B) and 4 (C-D) independent experiments, error bars = SEM.

117

Figure 2-15. Effect of srrAB single and nos srrAB double mutation on membrane

potential. Aerobic TSB-G cultures of wildtype, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and the double mutant complemented with nos were grown as described. Cells pellets were then harvested and stained with 30 µM of the membrane potential stain DiOC2(3), and subjected to flow cytometry to detect the ratio of red to green fluorescence. Histograms represent the ratio of red to green fluorescence (X axis) plotted against the number of events (Y axis). A shift to the right of the vertical black line indicates an increase in membrane potential.

118

Table 2-1. Generation times for all strains

Strain CFU/ml Generation Time (Minutes) ± SEM

Growth Media

Wildtype UAMS-1 pMK4 32 ± 2 LB

UAMS-1 nos::erm pMK4 37 ± 4 LB

UAMS-1 nos::erm pMKnos 42 ± 9 LB

Wildtype UAMS-1 pMK4 42 ± 2 TSB-G

UAMS-1 nos::erm pMK4 43 ± 2 TSB-G

UAMS-1 nos::erm pMKnos 37 ± 3 TSB-G

Wildtype UAMS-1 pMK4 33 ± 3 TSB

UAMS-1 nos::erm pMK4 33 ± 3 TSB

UAMS-1 nos::erm pMKnos 29 ± 2 TSB

Wildtype LAC-13C 52 ± 2 TSB-G

LAC-13C nos::erm 56 ± 1 TSB-G

UAMS-1 nos::erm pMK4 + NO 39 ± 1 TSB-G

UAMS-1 ΔsrrAB pMK4 46 ± 2 TSB-G

UAMS-1 nos::erm ΔsrrAB pMK4 85 ± 21** TSB-G

UAMS-1 nos::erm ΔsrrAB pMKnos 41 ± 4 TSB-G

UAMS-1 ΔsrrAB pMK4 38 ± 2 TSB

UAMS-1 nos::erm ΔsrrAB pMK4 47 ± 6* TSB

UAMS-1 nos::erm ΔsrrAB pMKnos 32 ± 1 TSB

119

Table 2-2. Select genes altered upon nos mutation

Category Gene Name

Function Fold-change (nos/wt) 4 hrs growth

Fold-change (nos/wt) 6 hrs growth

Oxidative Stress:

trxA Thioredoxin 3.2 --- SAR1984 Ferritin -4.1 -3.1 msrA1 Methionine sulfoxide reductase A 3.1 --- perR Peroxide operon regulator --- -5.1 ahpF Alkyl hydroperoxide reductase subunit F 2.1 --- SAR1492 Ferredoxin -2.8 Nitrosative Stress:

hmp Flavohemoprotein 5.6 9.5 scdA Putative Iron sulfur cluster repair protein 3.8 7.1 ldh1 L-lactate dehydrogenase 1 -3.4 --- Anaerobic Metabolism:

pfl Pyruvate formate lyase 5.8 45.0 narG Nitrate reductase operon 3.8 --- ldh2 L-lactate dehydrogenase 2 8.1 3.3

ackA Acetate kinase 2.1 ---

SAR2013 Aldehyde dehydrogenase 4.1 --- SAR2210 Aldehyde dehydrogenase 2.2 --- adhA Alcohol dehydrogenase --- 3.9

nrdG Anaerobic ribonucleotide reductase activating protein

2.1 ---

Other Metabolic Genes:

ctaB Cytochrome bd oxidase 2.8 3.3

qoxC Putative quinol oxidase polypeptide III 5.5 5.3

hemA Heme biosynthesis 2.3 ---

pyk Pyruvate kinase 2.3 --- lacE PTS system, lactose-specific IIBC component 2.9 --- purH Purine biosynthesis operon -41.8 --- pyrG Pyrimidine biosynthesis -6.5 --- fadB

SAR2006 Fatty acid degradation operon NAD biosynthesis operon

--- 49.8

-20.6 21.0

Virulence: geh Lipase precursor -2.7 -2.0 czrB Zince resistance protein --- -3.8 capG Capsular biosynthesis operon -4.3 -3.0 dltD Lipoteichoic acid biosynthesis protein -3.6 -2.3 spa Protein A 4.0 ---

120

Table 2-3. qRT-PCR confirmation of select genes

Gene Name

RNAseq (Fold-change nos/wt)

Real-time PCR (4 hr RNA) Wildtype nos mutant nos complement

hmp

5.6

1.1

2.7

0.9

scdA

3.8

1.1

11.8

1.2

ldh2

8.1

1.1

4.5

1.6

qoxC

5.5

1.0

4.0

1.1

pflB 5.8 1.1 4.0 1.2

narG

3.8

1.0

2.0

0.8

purH

-41.8

1.1

-46.0

1.1

SAR2006 49.8 1.0 24.0 33.9

121

Table 2-4. Select cellular nos mutant metabolites

Cellular Metabolite % increase/decrease nos mutant vs wildtype

P-value (Two tailed t-test)

Organic Acids

Lactate -49 0.010 Citrate -30 0.151

α-ketoglutarate -54 0.037 Fumarate 158 0.060 Malate 62 0.195 Pyruvate 27 0.663 Amino Acids Citrulline 149 0.002 Glutamine -63 0.212 Glutamate -57 0.004 Ornithine -30 0.059 Leucine -38 0.007 Isoleucine -41 0.018 Valine -40 0.005 Histidine 228 0.044 Arginine 2 0.936 Adenine Nucleotide NADH -65 0.015 NAD+ -23 0.084 NAD/NADH 122 0.106 ATP -10 0.830

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Table 2-5. Energy charge

Strain Energy charge (ATP + 1/2ADP)/ (AMP + ADP + ATP)

Wildtype 0.72 nos mutant 0.75 nos complement 0.72 srrAB mutant 0.76 nos srrAB double mutant 0.68 Double mutant nos complement 0.74

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Table 2-6. Select nos srrAB double mutant cellular metabolites

Cellular Metabolite % increase/decrease nos/srrAB mutant vs wildtype

P-value (Two tailed t-test)

Organic Acids

Succinate -94 0.014 Malate -87 0.045

Citrate BLOQ ---

α-ketoglutarate BLOQ ---

Fumarate BLOQ ---

Amino Acids Alanine Arginine

-90 BLOQ

0.002 ---

Asparagine -65 0.018 Aspartate -70 0.002 Citrulline 347 0.003 Glutamate Glycine

-81 BLOQ

<0.001 ---

Histidine Lysine

BLOQ -69

--- 0.042

Methionine -28 0.004 Ornithine -87 <0.001 Proline Serine Threonine

-82 BLOQ BLOQ

0.021 --- ---

Tyrosine -71 0.022 Valine -41 0.003 Adenine Nucleotide NADP NADPH

-40 BLOQ

0.002 BLOQ

NADH -74 0.032 NAD/NADH 252 0.006 ADP 32 0.030 ATP 23 0.578

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Table 2-7. Select nos srrAB double mutant extracellular metabolites

Extracellular Metabolite

% increase/decrease nos/srrAB mutant vs wildtype

P-value (Two tailed t-test)

Organic Acids Lactate 8593 <0.001 Pyruvate 201 0.007 Malate 359 0.006

α-ketoglutarate -82 0.006

Amino Acids Alanine 120 0.009 Asparagine 568 <0.001 Aspartate 118 0.005 Glutamate 226 0.007 Glutamine 173 <0.001 Leucine 15 0.021 Lysine 47 0.002 Proline 28 0.008 Serine 2966 <0.001 Threonine 1616 <0.001 Valine 9 0.027

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CHAPTER 3 MATERIALS AND METHODS

Bacterial Strains and Culture Conditions

All strains and primers used in this study are indicated in Tables 3-1 and 3-2,

respectively. Generation of the nos::erm mutation in both UAMS-1 (Sapp et al., 2014)

and LAC-13C (this study) strains was performed as previously described by inserting an

erythromycin resistance cassette 232 bp downstream of the nos ATG start site (Sapp et

al., 2014). Prior to each experiment, fresh cultures of S. aureus were streaked from -80

°C frozen stocks on tryptic soy agar (TSA) containing antibiotic (as required, Table 2-1),

and grown for 24 hours. A single isolated colony was used to inoculate overnight

cultures of S. aureus grown in tryptic soy broth containing 14 mM glucose (TSB) with

antibiotic selection (as appropriate) at 37 °C and shaking at 250 RPM. Unless otherwise

noted, for aerobic growth conditions, 40 ml (500 ml flask, 1:12.5 volume:flask ratio) of

TSB or TSB lacking glucose (TSB-G) was inoculated to an OD600 = 0.05 and grown at

37 °C and 250 RPM. For all chemical complementation experiments, DPTA NONOate

(Cayman) was used as the NO donor. A 150 mM stock solution of DPTA NONOate was

made by dissolving 10 mg in 0.01 M NaOH, and aliquots were stored at -80C for no

more than two weeks. For each experiment, DPTA NONOate was added to a final

concentration of 100 µM in sterile media just prior to bacterial inoculation.

Creation of nos srrAB Double Mutant and Complement

For generation of the srrAB nos double mutant, the temperature sensitive allele

replacement vector pTR27 (Sapp et al., 2014) was phage transduced from S. aureus

RN4220 into the unmarked srrAB mutant, KB6004 (Lewis et al., 2015, Bose, 2014).

Once confirmed in the target strain, a temperature sensitive allele replacement event

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was initiated by growth at 43°C on TSA + 10 mg/ml Erm (non-permissive temperature

for plasmid replication) to promote chromosomal integration via homologous

recombination at the nos locus. A second recombination event was induced by growing

a single isolated colony in TSB (no antibiotic) for 5 days at 30°C with sub-culturing every

24 hours. Screening for nos insertion and loss of the vector was completed on both TSA

+ 2 µg/ml Erm and TSA + 10 µg/ml Cm. PCR was used to confirm nos and srrAB

mutations. Complementation of the nos srrAB double mutant with pMKnos and

generation this strain containing empty pMK4 vector was completed by phage-

transducing each plasmid into the nos srrAB double mutant.

Growth Curve Analysis

For each growth curve experiment, LB, TSB-G, or TSB was inoculated to an

OD600 = 0.05 from fresh (approximately 15 hours growth) overnight cultures and grown

aerobically for 24 hours at 37 °C and 250 RPM in 500 ml Erlenmeyer flasks (1:12.5

volume:flask ratio). Samples (1 ml) were withdrawn from each culture every two hours

and serial diluted, followed by track plating (Jett et al., 1997) to determine CFU/ml.

OD600 readings were also acquired at each time point. For NO complementation growth

curves, experiments were performed as described above in TSB-G, except that cultures

were grown for only 8 hours in 250 ml Erlenmeyer flasks at a 1:12.5 volume:flask ratio.

Colony Size Comparison

For comparison of colony sizes, fresh overnight cultures of each strain were

diluted in 1 ml of sterile TSB to an OD600 of 0.05. Serial dilutions and track plating of

each diluted culture were then completed to bring colony counts in the observable range

(Jett et al., 1997). For track plating, 10 µl of diluted culture was placed in one lane of the

square track plate and the plate was tilted at a 45˚ angle to allow the culture to run down

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the plate. All cultures were plated on TSB containing 5 µg/ml chloramphenical and

pictures were taken after 24 hours of incubation at 37°C. Colony size was quantified

using the OpenCFU software (Geissmann, 2013) with parameters line width = 1,

Threshold = inverted (Auto), radius = 1 (Auto-max). Radius values are unitless and were

determined "per object" by the software. Only colonies that were not clumped and

clearly round were included in calculations.

Transmission Electron Microscopy

Bacterial cultures were grown for 6 hours in TSB-G, at which point 10 ml was

collected from each culture and centrifuged at 3901 x g for 3 minutes at room

temperature. Supernatants from each tube were discarded and cell pellets resuspended

in 1 ml of 0.1 M cacodylate buffer (pH 7.2). The suspension was then centrifuged at

17,000 x g for 3 minutes, supernatant was discarded, and cell pellets were suspended

in 1 ml Trumps fixative (4% formalin and 2% glutaraldehyde) containing 0.2 M

cacodylate (Electron Microscopy Sciences, Hatfield PA) and placed overnight at 4˚C.

Subsequent washes were completed with 0.1 M cacodylate buffer to remove the

Trumps fixative before further fixation in a solution of 2% glutaraldehyde, 50 mM lysine,

500 ppm ruthenium red in 0.1 M cacodylate buffer (pH 7.2) for 1 hour at room

temperature. Once fixed, cells were again washed with 0.1 M cacodylate buffer. The

suspension was then centrifuged to form a pellet and encapsulated in 3% low-

temperature gelling agarose type VII (Sigma-Aldrich). The following steps were

processed with the aid of a Pelco BioWave Pro laboratory microwave (Ted Pella,

Redding, CA, USA). Fixed cells were post-fixed with 2% buffered osmium tetroxide 1’ in

hood followed by microwave for 45 seconds at 100 W under vacuum and finally 3’ in

hood. Post-fixed cells were then water washed and dehydrated in a graded ethanol

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series (25%, 50%, 75%, 95%, 100%) followed by 100% acetone, once each for 45

seconds at 220 W. Dehydrated samples were then infiltrated in a graded

acetone/Spurrs epoxy resin (30%, 50%, 70%, 100%, 100%), once each for 3 minutes at

220 W under vacuum, followed by 10 minutes at room temperature on the bench. Resin

infiltrated cells were cured at 60ºC for 2 days. Cured resin blocks were trimmed, thin

sectioned and collected on Formvar copper 100 mesh grids, post-stained with 2%

aqueous. uranyl acetate and Reynold’s lead citrate. Sections were examined with a

Hitachi H-7000 TEM (Hitachi High Technologies America, Inc. Schaumburg, IL) and

digital images acquired with a Veleta 2k x 2k megapixels side-mount camera and iTEM

software (Olympus Soft-Imaging Solutions Corp, Lakewood, CO). White scale bars on

images indicate 1 micrometer whereas black bars indicate 0.2 micrometer.

Scanning Electron Microscopy

Aerobic bacterial cultures were grown for 6 hours in TSB-G, followed by

harvesting of 10 ml from each culture by centrifugation at room temperature for 3

minutes at 3901 xg. Data is representative of one biological replicate mounted on two

individual stubs for imaging of 12-14 random fields of view. Cell pellets were washed in

10 ml sterile 1X PBS, centrifuged as described above, and resuspended in 1.2 ml of

Trumps fixative (4% formalin and 2% glutaraldehyde in 0.2 M sodium cacodylate buffer)

followed by incubation at room temperature for 15 minutes. Cells were stored at 4 °C

before being processed using a microwave-assisted methodology (Pelco BioWave Pro,

Ted Pella, Redding, CA, USA). Fixed cells were washed in 1X PBS, pH 7.24, post fixed

with 2% buffered osmium tetroxide, water washed, dehydrated in a graded ethanol

series 25%, 50%, 60%, 75%, 95%, 100% and subjected to critical point drying

(Autosamdri 815, Tousimis, Rocksville, MD USA). Cells on Millipore filters were then

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mounted onto aluminum specimen mounts with double sided adhesive tabs, and sputter

coated with Gold/Palladium (DeskV, Denton Vacuum, Moorestown, NJ USA). Samples

were then imaged with a field-emission scanning electron microscope (S-4000, Hitachi

High Technologies America, Inc. Schaumburg, IL,USA). Cell lengths from 12-14 fields

of view were measured using the ImageJ software program (Schneider et al., 2012).

RNAseq Analysis

RNAseq analysis was performed on total RNA isolated from 4 and 6 hour aerobic

TSB-G cultures of wildtype UAMS-1 and the nos::erm mutant as previously-described

for S. aureus (Carroll et al., 2016b, Carroll et al., 2016a). All reagents used were

dedicated RNase free and care was taken to process isolated RNA as quickly as

possible. In brief, isolation of RNA was first completed using an RNeasy kit (Qiagen)

followed by DNAse treatment (Ambion Turbo DNA-free kit) of the purified RNA. DNAse-

treated RNA was immediately analyzed on a Bioanalyzer (Agilent RNA 6000 nano chip)

to determine RNA integrity based on the RNA integrity number (RIN). To ensure

minimal rRNA degradation, an RIN number of 9.9 out of 10 was confirmed before

proceeding. Removal of rRNA was then completed using both the Ribo-Zero Magnetic

Kit (Epicentre) for Gram-positive bacteria, followed by a second round of purification

using the MicrobExpress Bacterial mRNA Enrichment Kit (Life technologies). Wildtype

or nos mutant RNA isolated from 3 independent experiments was pooled before

proceeding with RNAseq analysis. RNAseq was carried out using the IonTorrent PGM

platform, with library construction first being generated by Ion Total RNAseq v2 Kits

(Life Technologies). Template positive Ion Sphere™ Particles (ISPs) were generated

using an Ion PGM™ Template OT2 200 Kit, followed by sequencing on an Ion 318™

Chip v2 using Ion PGM™ 200 Sequencing Kits. Read alignment and data analysis was

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completed by using the CLC Genomics Workbench platform (Qiagen) as previously

described (Carroll et al., 2016b). Alignment was completed by mapping all reads to the

MRSA252 genome (NC_002952.2). First, raw data files were imported in .sff format to

the software platform, and any residual reads corresponding to rRNA were filtered out

(due to the fact that rRNA was physically removed prior to RNAseq). Expression values

for S. aureus genes were calculated as RPKM values (reads per kilobase material per

million reads), according to the CLC Genomics Workbench protocols. Data sets were

normalized by quantile normalization (1). To identify genes demonstrating meaningful

differences in expression, the following cut off criteria were applied to the data: (1) To

reduce the impact of non-unique reads that can map to multiple locations, the percent

unique reads mapping to genes had to exceed 80%. (2) To eliminate lowly expressed

genes we imposed a cut-off whereby the RPKM expression value of a gene must be

greater than or equal to 50 in at least one data set. (3) A cut-off of 2-fold or higher was

applied to identify genes showing differential expression. Differential expression

analysis was conducted using the CLC Genomics Workbench software platform. All raw

RNAseq data has been deposited to the GEO database. The UAMS-1 4 hour RNAseq

data was previously reported in another publication (Carroll et al., 2016a), and the data

is available through GEO accession number GSE74936 (sample GSM1938000). The

nos mutant 4 hour sample, UAMS-1 6 hour sample, and nos mutant 6 hour sample are

available through GEO accession number GSE77400 (samples GSM2051351,

GSM2051352, and GSM2051353). Culture and RNA isolations were completed by the

author whereas library preparation, RNAseq, and data analysis were completed by our

collaborator, Ronan Carroll. Gene expression was confirmed by qRT-PCR on RNA

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isolated from 3 ml of culture after growth for 4 hours in TSB-G using our previously-

published methods for S. aureus (Lewis & Rice, 2016). For all qRT-PCR reactions, 10

µM stock solutions of each forward and reverse primer was used. All qRT-PCR

reactions were performed on RNA from 3 individual biological replicates.

Metabolite Analysis Using LC/MS/MS

Cell Collection and Metabolite Sample Preparation

To isolate cell pellets and extracellular media (EXM) for targeted metabolite

analysis, 40 ml of each TSB-G culture was harvested by 10 minutes centrifugation at

3901 x g and 4 °C. After centrifugation, 2 x 1 ml aliquots of each supernatant (EXM)

were removed and immediately frozen in liquid nitrogen and stored at -80C. Cell pellets

were quickly resuspended in 2 ml 1X PBS, centrifuged at 3901 x g and 4 °C for 3

minutes, and immediately resuspended in fresh 2 ml PBS. Aliquots (at 1 ml) were

separated into two microcentrifuge tubes and centrifuged at 13,000 x g and 4 °C for 3

minutes. The supernatant was saved for extracellular metabolite analysis and pellets

were immediately frozen in liquid nitrogen and stored at -80C. One tube was

subsequently processed for metabolite analysis (see below) and the other tube was

used to determine the protein concentration using the Pierce™ BCA protein

quantification assay. All samples were kept on ice throughout the entire procedure

before being flash frozen in liquid nitrogen and stored at -80 °C. Cell pellets and EXM

were lyophilized to dryness overnight. Lyophilized cell pellets were homogenized in 400

μL of 50/50 acetonitrile/0.3% formic acid using a Precellys (bead-beating) system

maintained at 4 °C. The lyophilized EXM samples were reconstituted in 400 μL of 50/50

acetonitrile/0.3% formic acid and vortexed thoroughly. For the pyridine nucleotide and

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adenosine phosphate samples, homogenization with the Precellys system was

completed in the presence of 18O2-labeled NADH as an internal standard. For NADH

and NADPH sample preparation only, a 100µL aliquot of homogenate was immediately

treated with 50/50 methanol/0.2 M NaOH. The resulting samples were aliquoted and

stored at -80 °C. Sample isolation was completed by the author whereas

homogenization and further sample preparation were completed by our collaborators

Christopher Petucci and Jeffrey Culver.

Extraction, Derivatization, and LC/MS/MS Quantitation of Organic Acids from Cell Homogenate and Extracellular Media

A 50µL aliquot of either cell homogenate or EXM was spiked with a 10µL mixture

of heavy isotope-labeled organic acid internal standards (lactate, pyruvate, 3-

hydroxybutyrate, succinate, fumarate, malate, α-ketoglutarate, and citrate; Sigma-

Aldrich, St Louis, MO; Cambridge Isotopes, Cambridge, MA; CDN Isotopes, Quebec,

Canada). This was followed by the addition of 50 μL of 0.4 M O-benzylhydroxylamine

and 10 μL of 2 M 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide. Samples were

vortexed thoroughly and derivatized at room temperature for 10 min. The derivatized

organic acids were then extracted from the homogenate by liquid-liquid extraction using

100 μL of water and 600 μL of ethyl acetate. Samples were vortexed for 5 seconds and

then centrifuged at 18,000 x g for 5 min at 10 °C. A 100µL aliquot of the ethyl acetate

layer was dried under nitrogen and reconstituted in 1 mL of 50/50 methanol/water prior

to LC/MS/MS analysis. Derivatized organic acids were separated on a 2.1 x 100 mm,

1.7 μm Waters Acquity UPLC BEH C18 column (T = 45 °C) using a 7.5-min linear

gradient with 0.1% formic acid in water and 0.1% formic acid in acetonitrile at a flow rate

of 0.3 mL/min. Quantitation of derivatized organic acids was achieved using multiple

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reaction monitoring on a Dionex UltiMate 3000 HPLC/Thermo Scientific Quantiva triple

quadrupole mass spectrometer (Thermo Scientific, San Jose, CA). A standard

calibration curve (1-5000 μM for lactate; 0.2-1000 μM for 3-hydroxybutyrate; 0.05-250

μM for pyruvate, succinate, fumarate, malate, and citrate; 0.02-100 μM for α-

ketoglutarate) for derivatized organic acids was prepared by spiking 10µL aliquots of

organic acids (Sigma-Aldrich, St. Louis, MO) and internal standards (Sigma-Aldrich, St

Louis, MO; Cambridge Isotopes, Cambridge, MA; CDN Isotopes, Quebec, Canada) into

50µL aliquots of a 50/50 acetonitrile/0.3% formic acid solution. Calibration samples were

derivatized and extracted similarly to organic acids in cell homogenate and EXM

(above). Data for cell samples were normalized to protein whereas EXM concentrations

were given in µM. This expriment was completed by our collaborators Christopher

Petucci and Jeffrey Culver.

Extraction, Derivatization, and LC/MS/MS Quantitation of Amino Acids from Cell Homogenate and Extracellular Media

A 100µL aliquot of either cell homogenate or reconstituted EXM was spiked with

a 10µL mixture of heavy isotope-labeled amino acid internal standards (Sigma-Aldrich,

St Louis, MO; Cambridge Isotopes, Cambridge, MA; CDN Isotopes, Quebec, Canada).

This was followed by the addition of 800 μL of ice-cold methanol. Samples were

vortexed thoroughly and then centrifuged at 18,000 x g for 5 min at 10 °C. A 100µL

aliquot of the methanolic extract was dried under nitrogen and reconstituted in 80 μL of

borate buffer and 20 μL of MassTrak AAA Reagent (both provided in MassTrak AAA

Derivatization Kit; Waters Corp., Milford, MA). The samples were then derivatized at 55

°C for 10 minutes prior to LC/MS/MS analysis. Derivatized amino acids were separated

on a 2.1 x 100 mm, 1.7 μm Waters AccQ·Tag column (T = 55 °C) using a 9.55 min

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linear gradient with eluents proprietary to Waters Corp. at a flow rate of 0.7 mL/min.

Quantitation of derivatized amino acids was achieved using multiple reaction monitoring

on an Agilent 1290/6490 HPLC/triple quadrupole mass spectrometer (Waters Corp.,

Milford, MA). A standard calibration curve (1-1000 μM for Gly, Ala, Pro, Val, Arg, Thr,

Lys and Gln; 0.5-500 μM for Ser, Leu, Ile, Met, His, Phe, Tyr, Asn, Asp, Gly, Orn and

Cit; 0.25-250 μM for Trp) for derivatized amino acids was prepared by spiking 10µL

aliquots of amino acids (Sigma-Aldrich, St. Louis, MO) and internal standards (Sigma-

Aldrich, St Louis, MO; Cambridge Isotopes, Cambridge, MA; CDN Isotopes, Quebec,

Canada) into 100 µL aliquots of a 50/50 acetonitrile/0.3% formic acid solution.

Calibration samples were derivatized and extracted similarly to organic acids in cell

homogenate and EXM (above). Data for cell samples were normalized to protein

whereas EXM concentrations were given in µM. This expriment was completed by our

collaborators Christopher Petucci and Jeffrey Culver.

Extraction, Derivatization, and LC/MS/MS Quantitation of Pyridine Nucleotides and Adenosine Phosphates from Cell Homogenate

For the extraction of NMN, NAD, NADP, and all adenosine phosphates, a 100 µL

aliquot of cell homogenate was spiked with a 10µL mixture of heavy isotope-labeled

internal standards (18O2-labeled NMN and NAD, synthesized by the Sanford-Burnham

Medicinal Chemistry Core; AMP and ADP, Sigma-Aldrich, St. Louis, MO). This was

followed by the addition of 100 μL of 1 M ammonium formate to adjust the homogenate

pH to ~4. Samples were vortexed thoroughly and centrifuged at 18,000 x g for 5 min at

10 °C. The clarified homogenates were passed through an AcroPrep Advance 3K

Omega Filter Plate (Pall Corporation, Port Washington, NY) prior to LC/MS/MS

analysis. For NADH and NADPH extraction, the 200µL aliquot of cell homogenate

135

prepared above was vortexed thoroughly and centrifuged at 18,000 x g for 5 min at 10

°C. The clarified homogenates were passed through an AcroPrep Advance 3K Omega

Filter Plate (Pall Corporation, Port Washington, NY) prior to LC/MS/MS analysis. Select

pyridine nucleotides (NMN, NAD, and NADP) and all adenosine phosphates were

separated on a 2.1 x 50 mm, 3 μm Thermo Hypercarb column (T = 30 °C) using a 2.1-

min linear gradient with 10 mM ammonium acetate, pH 9.5 and acetonitrile at a flow rate

of 0.65 mL/min. Quantitation of these analytes was achieved using multiple reaction

monitoring on a Dionex UltiMate 3000 HPLC/Thermo Scientific Quantiva triple

quadrupole mass spectrometer (Thermo Scientific, San Jose, CA). For adenosine

phosphates and NMN, NAD, and NADP determination, a standard calibration curve

(0.625-500 μM for adenosine phosphates, 0.25-200 μM for NAD, 0.025-20 μM for

NADP, 0.0025-2 μM for NMN) was prepared by spiking 10µL aliquots of pyridine

nucleotides (Sigma-Aldrich, St. Louis, MO) and internal standards (synthesized by the

Sanford Burnham Prebys Medicinal Chemistry Core) into 100µL aliquots of 0.5 M

perchloric acid. Calibration samples were extracted similarly to adenosine phosphates

and pyridine nucleotides in cell homogenate. Data for cell samples were normalized to

protein. NADH and NADPH were separated on a 2.1 x 50 mm, 1.8 μm HSS T3 column

(Waters Corp., Milford, MA) at 40 °C using a 2.2-min linear gradient with 5 mM

ammonium acetate, pH 6 and acetonitrile at a flow rate of 0.54 mL/min. Quantitation of

pyridine nucleotides and adenosine phosphates was achieved using multiple reaction

monitoring on a Dionex UltiMate 3000 HPLC/Thermo Scientific Quantiva triple

quadrupole mass spectrometer (Thermo Scientific, San Jose, CA). This expriment was

completed by our collaborators Christopher Petucci and Jeffrey Culver.

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Measurement of Intracellular ROS and O2-

Total ROS and O2- were measured with the cell-permeable fluorescent stains

CM-H2DCFDA (Life technologies) and MitoSOX™ Red (Life technologies), respectively.

For intracellular ROS detection, 19 ml of culture was removed from aerobic TSB-G

cultures after 3 or 6 hours of growth (n=3 independent experiments per strain). Addition

of 15 µM Thioridizine HCl was completed at time of inoculation for treated cultures. Cell

pellets were harvested by centrifugation at room temperature and 3901 x g, washed in 1

ml Hanks Balanced Salt Solution (HBSS), and then resuspended in 1 ml HBSS

containing 10 µM final concentration of CM-H2CFDA. A 10 mM stock solution of CM-

H2DCFDA was freshly prepared for each experiment by dissolving the appropriate mg of

the dried stain in DMSO per the manufacturer's instructions. Cell suspensions were

incubated at 37 °C for 60 minutes, followed by an additional HBSS wash step (as

above) and resuspension in 1 ml HBSS (3 hour culture samples) or 1.2 ml HBSS (6

hour culture samples). Triplicate aliquots (200 µl) of each stained cell suspension were

then transferred to wells of a 96-well optically clear black tissue culture plate (Costar

3904), and the relative fluorescence units (RFU) and OD600 of each well was measured

(EX: 485±20 nm, EM: 516±20 nm) after plate incubation for 15 minutes at 37 °C using a

Synergy HT plate reader (Biotek). RFU were normalized to the OD600 of each well. For

MitoSOX™ superoxide staining, 10 ml of each culture was isolated after 3 hours aerobic

growth in TSB-G, centrifuged at 3901 x g for 5 minutes, and resuspended in MitoSOX™

reagent (prepared by diluting a freshly-made 5 mM stock solution to 5 µM in 1x PBS,

according to manufacturer’s protocols). Cells were then incubated for 10 minutes at 37

°C and washed once with 1x PBS. Triplicate aliquots (200 µl) of each stained cell

suspension were then transferred to wells of a Costar 3904 plate, and the RFU and

137

OD600 of each well was immediately measured (EX: 500±27 nm, EM: 600±40 nm) using

a Synergy HT plate reader (Biotek). RFU were normalized to the OD600 of each well.

Determination of Catalase Activity

Cell pellets from 5 ml of 3 hour TSB-G cultures were isolated by centrifugation at

3901 x g for 5 minutes at 4 °C. The supernatants were removed, and cell pellets were

immediately stored at -80 °C until protein isolation and catalase activity determination.

Cytosolic protein preparations were acquired by resuspending thawed pellets in 1 ml of

reaction buffer (0.1 M Tris-HCl pH 7.0) followed by mechanical disruption with lysing

matrix B tubes (0.1 mm silica spheres, MP Biomedicals) and separation of cellular

debris by centrifugation at 13,000 x g at 4 °C. Total cytosolic protein was determined

using the BCA protein quantification assay, followed by catalase activity measurements

using the Amplex Red catalase activity kit (Life Technologies) following manufacturers

protocols. In brief, both 1:1000 diluted cytosolic protein samples and a series of known

purified catalase concentrations (used to generate a standard curve) were treated with

hydrogen peroxide (H2O2), followed by addition of Amplex Red and horseradish

peroxidase to final concentrations of 50 µM and 0.2 U/ml, respectively. Fluorescence

(EX: 540±25 nm, EM: 600±40 nm) of each sample (caused by residual H2O2 reaction

with Amplex Red) was then recorded after 1 hour of fluorescent measurements (RFU)

using a Synergy HT microplate reader (Biotek). A standard curve was generated by

plotting the RFU measurement of each catalase standard (first subtracted from the no

catalase control) on the y axis against the amount (units) of each catalase standard on

the x axis. The catalase activity of each unknown sample was then extrapolated from

the standard curve, and normalized to the total cytosolic protein.

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Assessment of Membrane Potential

Aerobic cultures were grown in TSB or TSB-G for 3 or 6 hours, then cells from 1

ml of culture were harvested by centrifugation and stained with the BacLight™ Bacterial

Membrane Potential Kit (Invitrogen) as previously described for S. aureus (Lewis et al.,

2015, Novo et al., 1999, Patton et al., 2006). In brief, cell pellets from 1 ml of each

culture were washed with once with 1x PBS and 25 µl of each washed cell suspension

was added to 2 ml 1x PBS containing 30 µM 3,3′-diethloxacarbocyanine iodide

(DiOC2(3)). Carbonyl cyanide 3-chlorophenylhydrazone (CCCP), a depolarizing agent,

was added to one tube of diluted wildtype cells during staining to a final concentration of

5 µM as a positive control for membrane depolarization. After staining, cells were

subjected to flow cytometry analysis with the FACSort flow cytometer (BD Biosciences

San Jose CA, USA) using published specifications (Lewis et al., 2015). For each

sample, 50,000 events were monitored for red and green fluorescence and then a

red/green ratio was calculated. In select experiments, 100 µM of DPTA NONOate was

included at time of inoculation as described above. Histograms were generated using

the FCS Express 4 Flow Cytometry Software (DeNovo).

CTC Staining

Respiration was measured using a 5-Cyano-2,3-ditolyl tetrazolium chloride (CTC)

stain as previously described for S. aureus (Lewis et al., 2015). In brief, 2 ml of each

culture was centrifuged to collect cell pellets, which were washed and centrifuged with 1

ml of 1x PBS and then resuspended in 650 µl of PBS containing 4.5 mM CTC. Triplicate

aliquots (200 µl) of each stained cell suspension were then transferred to wells of a

Costar 3904 plate. The RFU and OD600 of each well was measured (EX: 485±20 nm,

EM: 645±40 nm) at 10 minute intervals for 120 minutes at 37 °C using a Synergy HT

139

plate reader (Biotek). RFU readings collected at 70 minutes were normalized to the first

(t=0) OD600 reading of each well. Data is listed as fold-change relative to wildtype due to

variability in absolute numbers from day to day experiments. To prevent clumping the

plate was shaken for 5 seconds before each read. For DPTA complementation

experiments, DPTA NONOate (100 µM) was added to TSB-G as above and 1 ml of

culture was isolated for staining at 3 or 6 hours growth for all samples.

Oxygen Consumption

Oxygen consumption measurements were completed using a 4-channel free

radical analyzer (TBR-4100, World Precision Instruments) and Clark type electrode

(ISO-Oxy-2, World Precision instruments) on 3 hour aerobically grown cultures in TSB-

G. 15 ml of culture was re-suspended in fresh air saturated TSB-G and then diluted 1:2

before measurements were taken. Directly before measuring consumption, 3 ml of

mineral oil was placed over the top of the fresh re-suspended culture and then the

change in voltage was measured over 5 minutes. Data was only used in the linear

portion of the consumption curves (approximately 2 minutes) and relative rate (%) of

Oxygen consumption to wildtype was determined by measuring the slope of the

consumption curve and normalizing to cell counts. To assure proper function of the

electrode, calibration curves were completed each day before experimentation.

Determination of Aconitase Activity

Whole cytosolic protein was isolated from 18 ml of aerobic TSB-G cultures grown

for 3 hours. Addition of 15 µM Thioridizine HCl was completed at time of inoculation for

treated cultures. Cells were isolated by centrifugation at 3901 x g for 10 minutes at 4 °C

followed by washing with 1 ml of cold 1X PBS. Cell lysis was completed in 1X aconitase

assay buffer (50 mM Tris-HCl, pH 7.4) containing 100 µg/ml lysostaphin (Sigma) and

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27.3 Kunitz units DNase I (Qiagen). After 30 minutes of incubation at 37˚C, cell debris

were removed by centrifugation at 13,000 x g (4˚C) and proteins were immediately

frozen at -80˚C for future analysis. Aconitase activity was quantified using the Cayman

Chemicals Aconitase Assay Kit, following the manufacturer’s recommended protocols.

Assays were performed on 1:4 dilutions of each protein sample, and optical density

measurements at 340 nm were taken every 1 minute for 1 hour with incubation at 37˚C.

Sample background wells were included for each sample, which did not receive

substrate. Enzyme activity was determined by measuring the reaction rate (ΔA340/min.)

and using the NADPH extinction coefficient (0.00313 µM-1, adjusted for the 0.503 cm

path length of the well). All sample activity was normalized to total cytosolic protein as

determined by the Pierce™ BCA protein quantification assay (Life Technologies).

Statistical Analysis

Statistical analysis was completed with Sigmaplot software version 13, build

13.0.0.83 (Systat). Data were tested for normality and equal variance prior to choosing

the appropriate parametric or non-parametric test, respectively.

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Table 3-1. Bacterial strains and plasmids constructs used in this study

Strain or plasmid name Description Reference or source

E. coli DH5α Staphylococcus aureus RN4220 UAMS-1 LAC-13C KR1010 KR1040 KB6004 ABM10 pCRBlunt pTR27 pBT2 pMKnos pMK4

Host strain for construction of recombinant plasmids Easily transformable restriction deficient strain Osteomyelitis clinical isolate CA-MRSA isolate UAMS-1 nos::erm insertion mutant LAC-13C nos::erm insertion mutant UAMS-1 srrAB deletion mutant UAMS-1 Δnos srrAB::erm double mutant E. coli cloning plasmid; KmR nos::erm allele-replacement plasmid; ErmR/CmR

Temperature-sensitive shuttle vector; CmR/AmpR

nos complementation plasmid Shuttle vector; CmR/AmpR

(Hanahan, 1983) (Kreiswirth et al., 1983) (Gillaspy et al., 1995) (Fey et al., 2013) (Sapp et al., 2014) Unpublished strain created by K.C. Rice (Lewis et al., 2015) This study Invitrogen (Sapp et al., 2014) (Bruckner, 1997) (Sapp et al., 2014) (Sullivan et al., 1984)

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Table 3-2. PCR primers used in this study

Primer Purpose Sequence (5-3)

Reference or Source

sigA-F sigA-R SAR2006-RT-F SAR2006-RT-R SAR0218-RT-F SAR0218-RT-R SAR2680-RT-F SAR2680-RT-R SAR1032-RT-F SAR1032-RT-R SAR2486-RT-F SAR2486-RT-R purH-RT-F purH-RT-R scdA-RT-F scdA-RT-R hmp-RT-F hmp-RT-R

Real-time PCR Real-time PCR Real-time PCR Real-time PCR Real-time PCR Real-time PCR Real-time PCR Real-time PCR Real-time PCR

CAAGCAATCACTCGTGCAAT GGTGCTGGATCTCGACCTAA TCAACCAGCATTAGGTGCAG CTGGCGTAGTAACCTTTTCAGC GCTGTTAAAGCAGCCTACCG AGAAGCATATGCCCCTTCAC CTTGCAGTTTGGTCACAAGC TTCCGCTTTAGCTTCGCTAC ACGCATGGTTGTCACGTATC TGTCTAATCCGCGTCGTTG CGGCAAGAGCAGTTATTTCG GACCCAGGCGTTTGAATATG CGAAATAAACCGCAGCATTT TCGTCACATCAGGGTTAGCA TGCGGCGGACAAGTAAGTAT GCGAACCTGGTGTATTCGTT AGAGGCATGCAATCTTCAGC AGTGCGCAGTGTTTATATGC

(Sapp et al., 2014) This study This study This study This study This study (Sapp et al., 2014) This study This study

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CHAPTER 4 DISCUSSION

Since the discovery of bacterial NOS enzymes, many biochemical and

crystallographic characterization studies have been completed, but only a handful of

papers have investigated the functional role of these proteins (Salard-Arnaud et al.,

2012, Pant et al., 2002, Adak et al., 2002a, Choi et al., 1997, Bird et al., 2002, Chartier

et al., 2006). Most examples highlighting the contribution of NOS to bacterial physiology

have been observed by imposing external stressors (Gusarov & Nudler, 2005, van

Sorge et al., 2013, Gusarov et al., 2009, Patel et al., 2009). The nos mutation in both D.

radiodurans (Patel et al., 2009) and B. anthracis (Popova et al., 2015) revealed a

decreased OD600 phenotype; similar to what was seen in S. aureus when grown in the

absence of glucose, (Figure 2-1), a condition promoting aerobic respiration. Given that

the nos mutant OD phenotype was unique to exponential phase growth without glucose,

it is possibly linked to TCA cycle utilization and aerobic respiratory metabolism. In

support of this hypothesis, TSB growth curves (Figure 2-1) showed no apparent OD or

generation time (Table 2-1) phenotype during exponential growth, a growth situation

where S. aureus is primarily fermenting glucose to acetate (Somerville et al., 2002).

Although the exact cause of the OD (Figure 2-1) and cell elongation (Figure 2-3)

phenotypes of the S. aureus nos mutant is unknown, a previous study revealed a

comparable cell elongation phenotype in an S. aureus aconitase mutant (Somerville et

al., 2002). The Fe-S cluster in aconitase is sensitive to attack by ROS (Gardner &

Fridovich, 1991b, Gardner & Fridovich, 1992, Overton et al., 2008). Thus, one possibility

is that the increased ROS observed in the nos mutant (Figure 2-7) may be disabling

aconitase, effectively producing an aconitase mutant. Although aconitase activity was

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significantly lower in the nos mutant (Figure 2-11), ROS damage of this enzyme is

unlikely to be the cause of this decreased activity, since growth of nos mutant cultures

in the presence of TZ decreased ROS levels back to wild-type levels, but did not restore

aconitase activity (Figure 2-11). Another possible explanation for the OD600 phenotype

in the nos mutant condition is that the cells are clumping differently than wildtype.

Examination of cells using light microscopy suggests that clumping is likely not a

contributor to the OD600 phenotype (data not shown). A more likely scenario may be

altered membrane potential in the nos mutant condition, which could account for the

OD600 phenotype. Respiratory phenotypes (Figure 2-8 and 2-9) as well as the published

increase in carotenoid pigmentation ((Sapp et al., 2014) support an altered membrane

composition. Although the mechanism behind the OD600 phenotype is unknown, it

appears to be directly related to the action of NO itself, as addition of exogneous NO

donor complemented the nos mutant growth phenotype (Figure 2-2).

In S. aureus, disruption of preferred metabolic pathways and/or proper

respiratory function by nitrosative stress (Richardson et al., 2006, Richardson et al.,

2008), H2O2 (Chang et al., 2006), or mutation of heme biosynthesis genes (Kohler et al.,

2003) induce expression of genes associated with a lactate based anaerobic

metabolism. With this in mind, the nos mutant showed increased expression of genes

related to a lactate based anaerobic metabolism (ldh2, nar, pflAB, pyk, ackA) when

grown under conditions promoting aerobic respiration (Table 2-2), with a significant

decrease in intracellular lactate levels also being observed (Table 2-4). In S. aureus,

production of lactate occurs via several lactate dehydrogenases (ldh1, ldh2, ddh)

(Richardson et al., 2008), which convert pyruvate to lactate with the subsequent

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recycling of NAD from NADH. Indeed, ldh2 expression was increased 8.1-fold and a

significant decrease in NADH levels was observed in the nos mutant relative to

wildtype. L-lactate can drive respiration by donating its electrons to Lqo, and therefore

the nos mutant may be respiring on lactate in an attempt to compensate for altered

NADH driven respiration. As well, the differential CTC staining patterns (Figure 2-9)

provide additional indirect evidence supporting a model for potential Lqo-driven

respiration in the nos mutant. CTC staining, which was shown to accept electrons from

Ndh in E. coli (Smith & McFeters, 1997), was increased at 3 hours in the nos mutant

relative to wildtype. However, at 6 hours growth, CTC staining was lower in the nos

mutant relative to wildtype, suggesting that cell respiration may be diverted from Ndh to

Lqo or an alternative respiratory dehydrogenase in the nos mutant. Additional evidence

for induction of a stress response in the nos mutant comes from the dramatic decrease

in expression of purine and pyrimidine biosynthesis genes (Figure 2-5). Altered

expression of these genes seems to be a general response in S. aureus, as

transcriptome metadata using the S. aureus transcriptome meta-database (SATMD)

(Gopal et al., 2015) previously described these gene expression changes under multiple

stress conditions including acid shock, DNA damage, antimicrobial challenge, as well as

oxidative and nitrosative stress. A switch to a lactate-based fermentative metabolism by

S. aureus may therefore present a common strategy when it is challenged with

ROS/RNS species that inhibit its preferred metabolic pathways and/or upon disruption

of proper respiratory function.

It is interesting that the nos mutant presents with increased expression of

anaerobic metabolism genes during aerobic respiratory growth (Figure 2-5 and Table 2-

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2). One possible explanation for this may be related to decreased aconitase activity

(Figure 2-11) in the nos mutant. Examination of the metabolomics data suggests a

partial shutdown of the oxidative branch of the TCA cycle in the nos mutant, which

would explain the decreased levels of α-ketoglutarate and citrate, combined with

accumulation of fumarate and malate (Figure 4-1). Fermentation pathways may provide

an outlet for these overflow metabolites as fumarate and malate could be converted to

oxaloacetate, phosphoenolpyruvate (catalyzed by phosphoenolpyruvate carboxykinase

(PckA) (Scovill et al., 1996)), and finally into pyruvate (catalyzed by pyruvate kinase

(Pyk) (Zoraghi et al., 2010) (Figure 4-1). In support of this, pyk expression was

increased 2-fold in the nos mutant relative to wildtype (Figure 4-1). Partial shutdown of

the TCA cycle in the nos mutant may also be contributing to accumulation of Ctl, as Ctl

levels were significantly higher in the nos mutant relative to wildtype (Figure 4-1).

Catalysis of two intermediate reactions (arginosuccinate synthase and arginosuccinate

lyase) (Figure 4-1) can convert Ctl to fumarate, with this pathway potentially adding to

accumulation of these metabolites at this node. Furthermore, higher Ctl levels may be

driven by increased ornithine carbamoyltransferase (otc; 3.01-fold increased expression

(Appendix B)) activity in the nos mutant. This is supported by the fact that Glt, Gln, and

Orn are all decreased in the nos mutant, which could be due to increased consumption

of these metabolites by Otc (Table 2-4). When mapped together (Figure 4-1), the

RNAseq and metabolomics data support a metabolic scenario in which the nos mutant

presents with a partial shutdown of the oxidative branch of the TCA cycle and increased

expression of upstream anaerobic metabolism genes, possibly in an attempt to balance

this partial loss of TCA cycle activity.

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Although bacterial NOS has been found to contribute to oxidative stress

resistance in S. aureus (Sapp et al., 2014, van Sorge et al., 2013, Gusarov & Nudler,

2005), the exact protective mechanism(s) are unknown. In Bacillus, a mechanism of

NOS-derived oxidative stress resistance involving NO mediated activation of catalase

and depletion of free Cys (thereby limiting the Fenton reaction) has been proposed

(Gusarov & Nudler, 2005, Shatalin et al., 2008). It is currently unknown how NOS

promotes oxidative stress resistance in S. aureus (Sapp et al., 2014, van Sorge et al.,

2013, Gusarov & Nudler, 2005), but it is unlikely to function as in Bacillus, given that

catalase activity was not adversely affected in the S. aureus nos mutant (Figure 2-7). At

this time, NO-mediated Cys reduction in S. aureus cannot be ruled out, but a potential

alternative mechanism has been observed in Salmonella, whereby exogenous NO was

found to trigger an adaptive response to oxidative stress by arresting respiration

(Husain et al., 2008). Respiratory inhibition led to accumulation of NADH, with NADH

being able to 1) directly scavenge OH· (Goldstein & Czapski, 2000) 2) promote AhpCF

peroxidactic detoxification of peroxynitrite (Bryk et al., 2000) and 3) fuel detoxification of

H2O2 by AhpCF alkylhydroperoxidase (Jonsson et al., 2007). In a S. aureus nos mutant,

it is therefore possible that altered Ndh activity (Figure 2-9) leads to decreased NADH

levels (Table 2-4), and loss of NADH promoted protection. In fact, NADH levels were

lower in the nos mutant (Table 2-4) and ahpF expression was increased 2.1-fold (Table

2-2), potentially in an attempt to compensate for decreased levels of NADH. An elegant

set of experiments in E. coli confirmed that NDH-II is the primary source of ROS

formation by the respiratory chain (Messner & Imlay, 1999). While S. aureus appears to

have a Nuol-like NADH dehydrogenase subunit (Mayer et al., 2015), the type II NADH

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dehydrogenases are thought to be the primary respiratory driving enzymes (Schurig-

Briccio et al., 2014). Here we showed that chemical inhibition of NDH-II with TZ brought

ROS back down to wildtype levels (Figure 2-11), suggesting that elevated ROS in the

nos mutant is likely due to disruption in Ndh function (Figure 2-9), backup of electrons

onto Ndh, and/or decreased levels of NADH. Furthermore, the increased overall

endogenous ROS experienced by the nos mutant may be challenging its cellular

oxidative defense mechanisms at a level that predisposes the sensitivity of this strain to

external oxidative stress.

In both mammals (Brown, 1995, Giulivi et al., 2006, Brunori et al., 2004) and

bacteria (Borisov et al., 2004, Borisov et al., 2006, Butler et al., 2002, Junemann &

Wrigglesworth, 1996), NO is well established to complex with heme-containing

cytochromes, effectively outcompeting O2 and inhibiting respiration (reviewed in (Giuffre

et al., 2012, Sarti et al., 2003, Brunori et al., 2006)). Regulation of respiration by NOS

has also been observed in mammals by a proposed mitochondrial NOS (mtNOS)

isoform (Lacza et al., 2003, Boveris et al., 2000), and disruption of this NOS-mediated

regulation was found to increase generation of ROS (Parihar et al., 2008c), similar to

what was observed in the S. aureus nos mutant (Figure 2-7). Although it does not

appear that NOS-derived NO inhibits cytochrome-mediated O2 consumption in a

significant way (Figure 2-10), saNOS does seem to affect some currently unknown

component of the respiratory chain. In this respect, studies on bacterial photosynthetic

reaction centers have shown that the passive diffusion of protons across the membrane

(“proton backpressure”) can directly influence the membrane potential of the system

(van Rotterdam et al., 2001). Electron transfer down the respiratory chain requires

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proton movement, therefore these processes are coupled and one cannot occur without

the other (Lieberman et al., 2007). When protons passively diffuse across the

membrane, they block the flow of electrons down the transport chain, leading to a

buildup of membrane potential and electrons on upstream components (Lieberman et

al., 2007). This phenomenon may relate to the elevated membrane potential, CTC

staining, and accumulation of ROS observed in the nos mutant. Nevertheless, it

appears that saNOS influences some aspect of the respiratory chain, which results in

altered CTC staining and membrane potential (Figure 2-8 and 2-9), but does not lead to

measurable differences in overall respiratory rates (Figure 2-10).

In many bacterial species, NO-mediated respiratory inhibition leads to a plethora

of downstream transcriptional and physiological changes (Kinkel et al., 2013, Shi et al.,

2005, Machado et al., 2006, Richardson et al., 2006). While some of the nos mutant

phenotypes may be directly due to increased ROS (Figure 2-7), disruption of proper

respiratory chain function (Figure 2-8 and 2-9) presumably also contributes to the

observed transcriptional (Figure 2-5 and 2-6, Table 2-2) and metabolic changes

(Appendix A). NO-mediated cytochrome inhibition, mutation of the quinol oxidase

(qoxABCD), or low O2 conditions all impair the flow of electrons down the respiratory

chain, and SrrAB has been implicated in sensing each one of these conditions and

responding accordingly (Kinkel et al., 2013, Richardson et al., 2006). Altered membrane

potential (Figure 2-8) and respiratory dehydrogenase activity (Figure 2-9) in the nos

mutant could directly affect quinone pool reduction and would be sensed by SrrAB.

Elevated expression of lactate and fermentative metabolism genes (Table 2-2),

combined with the predicted contribution of Lqo supports a potential regulatory model in

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which SrrAB senses altered respiratory chain reduction in the nos mutant and alters

gene expression accordingly. In the absence of srrAB and nos, the double mutant

exhibits a fermentative metabolism resulting from loss of SrrAB-dependent gene

regulation. While the classic role of SrrAB in S. aureus metabolism has been to regulate

anaerobic pathways (Kinkel et al., 2013), SrrAB was also found to control expression of

virulence factors (Ulrich et al., 2007, Pragman et al., 2007, Pragman et al., 2004), NO

detoxification genes (Kinkel et al., 2013, Lewis et al., 2015), biofilm regulatory genes

(Windham et al., 2016), and most recently a small regulatory RNA (RsaE)(Durand et al.,

2015). RsaE is a small trans-acting sRNA that was found to respond to NO, with its

expression being dependant on SrrAB in S. aureus (Durand et al., 2015). Microarray

analysis showed that RsaE regulates TCA cycle and BCAA biosynthesis genes in S.

aureus (Bohn et al., 2010), which may account for the altered aconitase activity (Figure

2-11), levels of TCA cycle metabolites (Figure A-1), and levels of BCAAs (Figure A-3) in

the nos mutant. While the biological contribution of RsaE has not been extensively

characterized in S. aureus, multiple overlapping genes altered in the nos mutant (Table

2-2) were found to be influenced by RsaE in B. subtilis (Durand et al., 2015); many of

which are associated with oxidative stress and redox balance. Moreover, both the

protein and transcript levels of the promiscuous reductase partners for bsNOS (YkuN,

YkuP)(Holden et al., 2014, Wang et al., 2007) are regulated by B. subtilis RsaE (Durand

et al., 2015). In fact, annotation of our RNAseq data set as described in (Carroll et al.,

2016a) shows that rsaE (SARs051) is upregulated 7.37-fold in the nos mutant

(Appendix B). Ultimately, RsaE may be a good candidate as a small regulatory RNA

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that functions in concert with saNOS to mediate oxidative stress, redox balance, and

TCA cycle activity.

In the aerobic growth condition tested, the srrAB single mutant was not affected

in its ability to grow in either TSB-G or TSB (Figure 2-14). At the same time the single

srrAB mutant presented with some previously unpublished metabolic phenotypes,

including significantly increased levels of intracellular fumarate and malate, as well as a

significant decrease in lactate levels (Appendix A). Mqo oxidizes malate and promotes

TCA cycle function and subsequent generation of reducing equivalents for the

respiratory chain (Spahich et al., 2016). Elevated malate levels in the srrAB single

mutant may be related to decreased respiratory consumption, and indeed respiration as

measured by membrane potential was lower relative to wildtype (Figure 2-8). A similar

metabolic pattern (increased malate/fumarate, decreased lactate) was observed in the

nos single mutant relative to wildtype (Table 2-4). While not seen in the nos single

mutant, extracellur levels of malate and fumarate were higher in the srrAB single mutant

(Figure A-2). Therefore, saNOS and SrrAB may partially affect redundant pathways

associated with organic acid catabolism in S. aureus. Alternatively, BCAA profiles were

not the same in the nos and srrAB single mutants, with Val, Leu, Ile, and His levels

being significantly higher in the srrAB single mutant, as opposed to the nos single

mutant (Figure A-3). Another difference is the significantly lower levels of NADH seen in

the nos mutant but not seen in the srrAB single mutant. Moreover, the NAD/NADH ratio

was 122% (P =0.106) higher in the nos mutant, but not the single srrAB mutant (Figure

A-5). It can be difficult to retain consistency when measuring NAD nucleotides due to

the instability of these metabolites. Although not statistically significant (potentially due

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to large variation), there is a trend towards altered redox status of the nos mutant. This

lends additional evidence to the specific contribution of nos to proper Ndh function, as

altered Ndh function could be causing decreased levels of NADH in the nos mutant,

with this effect not seen in the srrAB single mutant.

Mutation of both srrAB and nos causes multiple significant metabolic changes,

enough to drastically alter the growth of this strain (Figure 2-14). A near-complete

shutdown of the TCA cycle is observed in the nos srrAB double mutant, with levels of all

TCA cycle organic acids either being significantly lower than wildtype or BLOQ (Figs.

Figure A-1 and 4-2). Comparison of Ctl levels between nos single and nos srrAB double

mutants can give additional insight into TCA cycle shutdown. Ctl enters the TCA cycle

via fumarate and the partial shutdown of the nos mutant TCA cycle may cause Ctl to

accumulate (Figure 4-1). As well, full TCA cycle shutdown as observed in the nos srrAB

double mutant, corresponded with almost twice as much Cit accumulation as the single

nos mutant (Figure A-3 and 4-2). Overall, the nos srrAB double mutant also appears to

be limiting amino acid uptake, which is a confirmed characteristic of TCA cycle

shutdown and decreased usage of biosynthetic pathways in S. aureus (Somerville et al.,

2002). Moreover, levels of NADP (-40%, p = 0.002) and NADPH (BLOQ) were

drastically reduced in the double mutant, again suggestive of biosynthetic pathway

shutdown. Overall redox status of the nos srrAB double mutant was also altered, with

the NAD/NADH ratio increased by 252% (p = 0.006) relative to wildtype (Table 2-4,

Figure A-5). All three lactate dehydrogenases consume NADH in the conversion of

pyruvate to lactate (Richardson et al., 2008). The lactate secretion profile, combined

with loss of NADH generation by the TCA cycle likely accounts for the altered redox

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status of the nos srrAB double mutant. Ultimately the overall metabolic profile of the nos

srrAB double mutant is consistent with TCA cycle shutdown, decreased biosynthetic

and amino acid transport pathways, and heightened lactate secretion (Figure 4-2).

In summary, these data suggest that saNOS and/or NOS-derived NO influence

some component(s) of the aerobic respiratory chain. Disruption of this relationship leads

to elevated ROS levels, as well as altered membrane potential and respiratory

dehydrogenase activity. In this model, NO is most likely protecting against damaging

ROS by either: 1) managing appropriate levels of protective NADH, 2) slowing the

production of endogenous ROS by contributing to proper respiratory function or 3) by an

as-yet unidentified mechanism. In any of these scenarios, endogenous NO production

via saNOS plays an important role in S. aureus physiology in the absence of external

stress. Loss of saNOS results in dramatic gene expression and metabolic adaptations

that presumably enable the nos mutant to continue to grow when normal respiratory

function is altered. These data suggest that when saNOS is present, the oxidative

branch of the TCA cycle is fully active, producing NADH and/or reductants to drive

respiration. Upon loss of saNOS, multiple anaerobic metabolism genes present with

increased expression, providing S. aureus with a way to respond to disrupted

respiratory metabolism and decreased TCA cycle activity. SrrAB provides S. aureus

with the metabolic flexibility to continue central metabolism using the TCA cycle, but

upon srrAB mutation, the nos srrAB double mutant is forced into a fermentative-like

metabolism. On-going research seeks to determine exactly how NOS influences the

respiratory chain, the genes regulated by SrrAB in this system, and the contribution of

nos srrAB mutation to virulence phenotypes.

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Figure 4-1. Central metabolic mapping of nos mutant transcriptional and metabolic

changes. Transcriptomic and targeted metabolomics data for the nos mutant relative to wildtype were mapped to select metabolic pathways using Biocyc software (Biocyc.org)(Caspi et al., 2014). Gene expression changes (squares) are indicated as fold-change whereas metabolite levels (circles) are indicated as % increase or decrease. All data was mapped to known and predicted pathways for MRSA252.

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Figure 4-2. Central metabolic mapping of nos srrAB double mutant metabolic changes.

Targeted metabolomics data for the nos mutant relative to wildtype were mapped to select metabolic pathways using Biocyc software (Biocyc.org)(Caspi et al., 2014) and a review of the literature for elucidation of amino acid entrance pathways to the TCA cycle (Owen et al., 2002). Metabolite levels (circles) are indicated as % increase or decrease. All data was mapped to known and predicted pathways for MRSA252.

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CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTIONS

This work presents previously undescribed phenotypes associated with nos

mutation that contribute to the further understanding of S. aureus oxidative stress

resistance, general physiology, and central metabolism. As well, the SrrAB two-

component system has been identified as an important regulator of response to nos

mutation and furthers the already described relationship between NO, SrrAB, and

respiration (Kinkel et al., 2013). Bacterial nitric oxide synthase proteins have only begun

to be characterized when compared to the extensive literature on mammalian NOS

isotypes. A link between NOS, respiratory activity (Figure 2-8 and 2-9), central

metabolism (Appendix A), and SrrAB (Figure 2-14) has not been previously described.

Elucidating these mechanisms may give insight into the evolutionary relationship

between bacteria and other domains of life, as NOS homologs are present in

prokaryotes, archaea, and eukaryotes (Sudhamsu & Crane, 2009).

saNOS appears to confers resistance to oxidative stress by an alternative

mechanism to what has been described in Bacillus. Firstly, saNOS does not activate

catalase in this bacterium (Figure 2-7), and secondly we show that elevated ROS levels

can be reduced upon TZ inhibition of the respiratory NADH dehydrogenase (Figure 2-

11). Loss of NOS regulated respiration of mammalian complex I has been attributed to a

pro-oxidant state, and therefore the relationship between NOS, NADH dehydrogenae

activity and ROS may be conserved between these domains of life (Parihar et al.,

2008c). It is possible that there are other potential mechanisms of NOS contribution to

oxidative stress resistance. These include reduction of free Cys (thereby limiting the

Fenton reaction) and/or modulation of intracellular Iron levels. In fact, a link between

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saNOS and heme stress was recently established (Surdel et al., 2016). Future work in

determining oxidative stress mechanisms by saNOS should focus on quantifying cellular

reduced thiols (Gusarov & Nudler, 2005) and intracellular iron levels (Keyer & Imlay,

1996).

While NO has been well established to influence respiration in both bacteria and

mammals, no previous role for NOS-derived NO has been found to contribute to

respiration in bacteria. Our data suggest that NOS-derived NO is not likely limiting

respiration by compeitive binding to the cytochrome (as evidenced by little change in O2

consumption rates between wild-type and nos mutant; Figure 2-10), but somehow

affects both membrane potential and CTC staining (Figure 2-8 and 2-9). This exact

mechanism has not yet been determined in S. aureus, but will be the focus of future

work. There are a few potential mechanisms that should be explored, including the

contribution of saNOS to membrane permeability, the interaction of NO with other

components of the respiratory chain, and elucidation of the currently unknown saNOS

reductase partner. Another technique combining heavy Nitrogen 15-labeled arginine

and electron paramagnetic resonance could help track NO binding to membrane bound

heme complexes (Jiang et al., 1997), and in parallel, this heavy nitrogen could possibly

be used in conjunction with NMR to monitor the fate of saNOS-derived NO in wildtype

and nos mutant cells. Interestingly, the attached mammalian NOS reductase domain

has homology to the p450 NADH oxidoreductase of the respiratory chain, suggesting

that the currently unknown saNOS reductase partner may have similarities to NADH

dehydrogenase (Nishida et al., 2002). It may not be a coincidence that NADH is the

oxidized metabolite utilized by both proteins. Moreover, the unique predicted mtNOS

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isotype is thought to functionally associate with complex I and accepts electrons for its

own synthesis of NO (Parihar et al., 2008a, Giulivi et al., 2006, Parihar et al., 2008b,

Boveris et al., 2000). Decreased NADH levels in the nos mutant (Table 2-4) provides

additional evidence that NADH oxidation may be altered in the nos mutant. Protein pull

down assays combined with in vitro enzymatic assays could help determine if saNOS is

utilizing the NADH dehydrogenase as its reductase partner. Overall this has laid the

ground work towards discerning a previously unknown contribution of saNOS to the

respiratory chain and has identified a potential reductase partner for bacterial NOS

proteins.

S. aureus is an extremely successful pathogen of humans and livestock, with

many strains being resistant to multiple antibiotics. The described nos mutant

phenotypes lend additional evidence that targeting this protein may be a viable

antimicrobial strategy. Coincidentally, bacterial NOS proteins are a potential novel

therapeutic target, as multiple research groups have already linked NOS inhibition to

increased antimicrobial efficacy (Holden et al., 2013, Holden et al., 2015b, Holden et al.,

2016). This work provies additional mechanistic insight as to why a S. aureus nos

mutant is attenuated in vivo (Sapp et al., 2014, van Sorge et al., 2013) as seen by the

clear alterations in endogenous ROS levels (Figure 2-7), altered respiratory function

(Figure 2-8 and 2-9), altered expression of various metabolic genes (Appendix B) and

changes in metabolite levels associated with central metabolism (Appendix A). There is

a growing body of work in the field of bacterial pathogenesis showing that many

virulence determinants are closely associated with metabolism, including work done in

S. aureus, B. anthracis, Listeria monocytogenes, Clostridium perfringens, Clostridium

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difficile, pathogenic streptococci, Neisseria meningitidis, and many others (Somerville

& Proctor, 2009a, Brinsmade, 2016, Willenborg & Goethe, 2016, Bouillaut et al., 2015,

Schoen et al., 2014). Understanding metabolic pathways in these bacteria can provide

additional insight into pathogenesis and alternative treatment methods. Moreover, future

studies to determine critical points in metabolic response pathways to nos mutation may

provide secondary targets for antimicrobial development. The combined RNAseq

(Figure 2-5 and 2-6) and metabolomic data (Appendix A) described herein point towards

oxidative stress resistance and fermentation pathways as being good candidates for

combined drug therapy. Further studies will focus on measuring ethanol and acetate

levels in the nos single mutant, which could not be determined by the completed

targeted metabolomics experiment. The observed decrease in nos mutant lactate levels

(Table 2-4) in combination with measurements of other fermentation products could

help to underscore S. aureus metabolic flexibility and preference of fermentation when

other metabolic pathways are disrupted. This study also suggests a switch to lactate-

based metabolism in the nos mutant (Table 2-4), building upon a previously described

stress response in S. aureus (Richardson et al., 2006, Richardson et al., 2008).

Alternatively, it may be a good antimicrobial strategy to target a regulatory

system combined with saNOS. In this work, the SrrAB two-component system has been

identified as a potential regulator in response to nos mutation. Clear growth defects

(Figure 2-12, 2-13, and 2-14) and drastic metabolic changes (Appendix A) occurred

when both nos and srrAB are mutated. All previous work in this context was associated

with the role of SrrAB in response to nitrosative stress, but a metabolic interplay

between saNOS-derived NO and SrrAB has not been previously described. One of the

160

first future studies with the nos srrAB double mutant will be qRT-PCR or RNAseq

studies to identify which genes altered in the nos single mutant are regulated by SrrAB.

It would be a great contribution to the understanding of S. aureus physiology and stress

response if SrrAB were determined to regulate a response to disrupted respiratory

metabolism or oxidative stress in the nos single mutant.

Overall, examination of metabolite levels suggests that the TCA cycle,

biosynthetic pathways, and amino acid transport pathways are drastically decreased in

the nos srrAB double mutant (Appendix A). Interestingly, similarities exist between the

metabolic profiles of the nos srrAB double mutant and small colony variants (SCV) of S.

aureus, a biologically unique isolate often associated with respiratory chain or

thymidylate biosynthesis deficiencies (Kriegeskorte et al., 2014). Downregulation of

TCA cycle activity as well as decreased levels of Asp and Glu were observed in 6

clinical SCV strains (Kriegeskorte et al., 2014), similar to what is seen in the nos srrAB

double mutant. Agar plate growth of the nos srrAB nos double mutant shows a clear

decrease in colony size compared to the single nos mutant (Figure 2-12 and 2-13),

lending additional support to this strain having SCV-like properties. Since the TCA cycle

produces biosynthetic precursors required for many virulence factors, both disruption of

TCA cycle activity (Somerville & Proctor, 2009a, Somerville et al., 2003a, Sadykov et

al., 2008) and SCV (TCA deficient) strains (Kriegeskorte et al., 2014) are attenuated in

virulence factor production. SCV strains have important clinical considerations as they

are associated with both persistant infections, intracellular survival, and resistance to

antimicrobials (Kim et al., 2016, Precit et al., 2016). Therefore, further virulence and

antimicrobial studies on the nos srrAB double mutant would have to be completed

161

before determining if targeting both of these proteins using a dual antimicrobial therapy

may be a viable strategy. Overall, this work outlines a central role of saNOS to bacterial

physology and metabolism, while at the same time identifying the SrrAB regulatory two-

component system as an important contributor to metabolic flexibility when NOS activity

is lost.

162

APPENDIX A

ADDITIONAL FIGURES

Figure A-1. Cellular organic acids of the nos, srrAB, and nos srrAB mutant strains. Data

are from cells pellets isolated from 4 hour aerobic cultures of wild-type, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and double mutant nos complement strains grown at 37ºC in TSB-G (n=3 independent experiments). Metabolites were determined by LC/MS/MS. All intracellular organic acids were normalized to total cytosolic protein. Error bars = SEM. *significance (P <0.05 Two-tailed t-test relative to wildtype).

163

Figure A-2. Extracellular organic acids of the nos, srrAB, and nos srrAB mutant strains.

Data are from cells pellets and supernatants isolated from 4 hour aerobic cultures of wild-type, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and double mutant nos complement strains grown at 37ºC in TSB-G (n=3 independent experiments). Metabolites were determined by LC/MS/MS. All extracellular organic acids are given in µM concentrations. Error bars = SEM. *significance (P <0.05 Two-tailed t-test relative to wildtype).

164

Figure A-3. Cellular amino acids of the nos, srrAB, and nos srrAB mutant strains. Data

are from cell pellets isolated from 4 hour aerobic cultures of wild-type, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and double mutant nos complement strains grown at 37ºC in TSB-G (n=3 independent experiments). Metabolites were determined by LC/MS/MS. Intracellular organic acids were normalized to total cytosolic protein. Error bars = SEM. *significance (P <0.05 Two-tailed t-test relative to wildtype).

165

Figure A-4. Extracellular amino acids of the nos, srrAB, and nos srrAB mutant strains.

Data are from supernatants isolated from 4 hour aerobic cultures of wild-type, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and double mutant nos complement strains grown at 37ºC in TSB-G (n=3 independent experiments). Metabolites were determined by LC/MS/MS. Extracellular organic acids are given in µM concentrations with the media control being sterile TSB-G. Error bars = SEM. *significance (P <0.05 Two-tailed t-test relative to wildtype).

166

Figure A-5. Cellular NAD nucleotides of the nos, srrAB, and nos srrAB mutant strains.

Data are from cells pellets and supernatants isolated from 4 hour aerobic cultures of wild-type, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and double mutant nos complement strains grown at 37ºC in TSB-G (n=3 independent experiments). Metabolites were determined by LC/MS/MS. All metabolites were normalized to total cytosolic protein. Error bars = SEM. *significance (P <0.05 Two-tailed t-test relative to wildtype).

167

Figure A-6. Cellular adenosine nucleotides of the nos, srrAB, and nos srrAB mutant

strains. Data are from cells pellets and supernatants isolated from 4 hour aerobic cultures of wild-type, nos mutant, nos complement, srrAB mutant, nos srrAB double mutant, and double mutant nos complement strains grown at 37ºC in TSB-G (n=3 independent experiments). Metabolites were determined by LC/MS/MS. All metabolites were normalized to total cytosolic protein. Error bars = SEM. *significance (P <0.05 Two-tailed t-test relative to wildtype).

168

APPENDIX B ADDITIONAL TABLES

Table B1. List of all genes altered in the nos mutant at 4 hours growth Gene name Function Fold Change (nos

mutant/ wild-type)

SAR2006 nicotinate phosphoribosyltransferase 49.8

nadE NAD synthetase 23.1

SAR2003 hypothetical protein 13.7

SAR2004 hypothetical protein 13.1

ldh2 L-lactate dehydrogenase 2 8.1

SARs054 predicted small RNA 7.4

SARs051 predicted small RNA 7.4

sstA FecCD transport family protein 7.2

SAR0218 pyruvate formate-lyase activating enzyme 6.7

SARs052 predicted small RNA 6.5

SARs265 predicted small RNA 5.9

pflB pyruvate formate-lyase B 5.8

SAR2636 hypothetical protein 5.7

hmp flavohemoprotein 5.6

qoxC quinol oxidase polypeptide III 5.5

SARs021 GJA5-1824-RNA 5.2

qoxA quinol oxidase polypeptide II precursor 5.2

qoxD quinol oxidase polypeptide IV 5.1

SAR0310 nucleoside permease 5.0

SAR0309 hypothetical protein 4.9

SAR2529 sodium/hydrogen exchanger family protein 4.8

qoxB quinol oxidase polypeptide I 4.7

SAR1376 4-oxalocrotonate tautomerase 4.6

SAR0312 N-acetylneuraminate lyase 4.5

SAR0311 sodium:solute symporter family protein 4.5

SAR0556 chaperone protein HchA 4.4

SAR0642 ABC transporter permease 4.3

dal alanine racemase 4.3

sstC ABC transporter ATP-binding protein 4.2

SAR0643 ABC transporter ATP-binding protein 4.2

SAR0308 PfkB family carbohydrate kinase 4.1

SAR2013 aldehyde dehydrogenase 4.1

spa immunoglobulin G binding protein A precursor 4.0

glmS glucosamine--fructose-6-phosphate aminotransferase 3.8

ulaA PTS system ascorbate-specific transporter subunit IIC 3.8

169

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

SAR1813 histone deacetylase 3.8

scdA cell wall biosynthesis protein ScdA 3.8

narG nitrate reductase subunit alpha 3.8

lacF PTS system lactose-specific transporter subunit IIA 3.6

SARs049 predicted small RNA 3.6

SAR1796 hypothetical protein 3.6

SAR2635 acetyltransferase 3.6

lrgB antiholin-like protein LrgB 3.5

SAR0641 ABC transporter 3.5

SAR1080 hypothetical protein 3.5

SAR2528 amino acid permease 3.4

SARs113 predicted small RNA 3.4

SARs111 predicted small RNA 3.4

trxA thioredoxin 3.2

cycA D-serine/D-alanine/glycine transporter 3.2

SAR0230 extracellular solute-binding lipoprotein 3.2

SAR0694 hypothetical protein 3.2

SAR0478 hypothetical protein 3.1

SAR0558 hypothetical protein 3.1

SARs073 predicted small RNA 3.1

SARs086 predicted small RNA 3.1

opuCA glycine betaine/carnitine/choline transport ATP-binding protein 3.1

lacD tagatose 1,6-diphosphate aldolase 3.1

msrA1 methionine sulfoxide reductase A 3.1

SAR1730 hypothetical protein 3.1

SARs227 predicted small RNA 3.1

xpt xanthine phosphoribosyltransferase 3.1

otc ornithine carbamoyltransferase 3.0

pbuX xanthine permease 3.0

SAR1063 hypothetical protein 3.0

SARs097 predicted small RNA 3.0

SAR1143 carbamate kinase 3.0

SAR0437 hypothetical protein 3.0

SAR1944 hypothetical protein 3.0

SAR0620 haloacid dehalogenase-like hydrolase 2.9

lacE PTS system lactose-specific transporter subunit IIBC 2.9

SARs132 predicted small RNA 2.9

170

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

ctaB protoheme IX farnesyltransferase 2.8

SARs019 GJA5-1758-RNA 2.8

SAR2596 hypothetical protein 2.8

SAR0232 hypothetical protein 2.8

guaB inosine-5'-monophosphate dehydrogenase 2.8

SAR1930 hypothetical protein 2.8

SAR1945 hypothetical protein 2.8

lacB galactose-6-phosphate isomerase subunit LacB 2.8

trap signal transduction protein 2.8

SAR0231 hypothetical protein 2.8

SAR1035 hypothetical protein 2.7

SAR1586 glyoxalase/bleomycin resistance protein/dioxygenase superfamily protein

2.7

SAR1471 hypothetical protein 2.7

clpL ATP-dependent protease ATP-binding subunit ClpL 2.7

SAR0315 N-acetylmannosamine-6-phosphate 2-epimerase 2.7

SAR0930 fumarylacetoacetate (FAA) hydrolase 2.7

gap2 glyceraldehyde 3-phosphate dehydrogenase 2 2.7

SAR1091 hypothetical protein 2.7

dat D-alanine aminotransferase 2.6

SAR1864 translaldolase 2.6

SARs015 GJA5-1458-RNA 2.6

SAR2021 hypothetical protein 2.6

SAR2407 hypothetical protein 2.6

lytS autolysin sensor kinase 2.6

guaA GMP synthase 2.6

narT nitrite transport protein 2.6

SAR0918 NADH:flavin oxidoreductase / NADH oxidase 2.6

SAR1352 transketolase 2.6

SAR1816 hypothetical protein 2.6

lacG 6-phospho-beta-galactosidase 2.6

acuA acetoin utilization protein 2.5

SAR0235 PTS transport system, IIBC component 2.5

SAR1836 dipeptidase PepV 2.5

SAR1849 proline dehydrogenase 2.5

sstD lipoprotein 2.5

SAR0211 hypothetical protein 2.5

SAR1163 hypothetical protein 2.5

171

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

SAR0336 hypothetical protein 2.5

SARs016 GJA5-1650-RNA 2.5

tag DNA-3-methyladenine glycosylase I 2.5

folP dihydropteroate synthase 2.5

lacA galactose-6-phosphate isomerase subunit LacA 2.5

SAR0929 hypothetical protein 2.5

SAR1328 cardiolipin synthase 2.5

SAR0578 hypothetical protein 2.4

SAR0670 sensor histidine kinase 2.4

SAR0739 MarR family regulatory protein 2.4

tnpR resolvase 2.4

pyn pyrimidine-nucleoside phosphorylase 2.4

SAR0731 hypothetical protein 2.4

SAR0396 hypothetical protein 2.4

SARs024 GJA5-2215-RNA 2.4

SAR0514 O-acetylserine (thiol)-lyase 2.4

opp-1A oligopeptide transporter substrate binding protein 2.4

SAR0335 luciferase-like monooxygenase 2.4

SAR2007 oxygenase 2.4

ebpS cell surface elastin binding protein 2.4

SAR0210 oxidoreductase 2.4

SARs256 predicted small RNA 2.4

SAR0403 hypothetical protein 2.4

SAR1365 hypothetical protein 2.4

hemA glutamyl-tRNA reductase 2.3

SAR1579 pyrroline-5-carboxylate reductase 2.3

agrD autoinducer peptide 2.3

SAR1953 AhpC/TSA family protein 2.3

ubiE ubiquinone/menaquinone biosynthesis methyltransferase 2.3

fda fructose-1,6-bisphosphate aldolase 2.3

lysA diaminopimelate decarboxylase 2.3

pyk pyruvate kinase 2.3

SAR1827 transposase 2.3

SAR2474 MarR family regulatory protein 2.3

nuc thermonuclease precursor 2.3

putP high affinity proline permease 2.3

SAR0781 proton-dependent oligopeptide transport protein 2.3

172

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

mraW S-adenosyl-methyltransferase MraW 2.3

SAR1705 hypothetical protein 2.3

menB naphthoate synthase 2.3

ppaC manganese-dependent inorganic pyrophosphatase 2.3

SAR2171 hypothetical protein 2.3

SAR2549 transporter 2.3

pgi glucose-6-phosphate isomerase 2.3

SAR2268 transport system binding lipoprotein 2.2

SAR2435 hypothetical protein 2.2

SARs018 GJA5-1713-RNA 2.2

ahpF alkyl hydroperoxide reductase subunit F 2.2

SAR0111 myosin-cross-reactive antigen 2.2

SAR2641 aminotransferase 2.2

SAR1335 hypothetical protein 2.2

SAR1610 lipoate-protein ligase A protein 2.2

SAR2210 aldehyde dehydrogenase 2.2

pepB oligopeptidase 2.2

deoC2 deoxyribose-phosphate aldolase 2.2

SAR2228 hypothetical protein 2.2

lacC tagatose-6-phosphate kinase 2.1

SAR1397 peptidase 2.1

ispD_1 2-C-methyl-D-erythritol 4-phosphate cytidylyltransferase 2.1

arlR response regulator protein 2.1

dnaB chromosome replication initiation/membrane attachment protein

2.1

SAR0862 thioredoxin 2.1

scrB sucrose-6-phosphate hydrolase 2.1

deoD purine nucleoside phosphorylase 2.1

mnmA tRNA-specific 2-thiouridylase MnmA 2.1

nrdG anaerobic ribonucleotide reductase activating protein 2.1

SARs125 predicted small RNA 2.1

SAR0560 haloacid dehalogenase-like hydrolase 2.1

SAR2223 hypothetical protein 2.1

mnhA monovalent cation/H+ antiporter subunit A 2.1

SAR0621 hydrolase 2.1

SAR0334 dioxygenase 2.1

SAR0405 hypothetical protein 2.1

ackA acetate kinase 2.1

173

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

glpD aerobic glycerol-3-phosphate dehydrogenase 2.1

SAR0826 hypothetical protein 2.1

nrdR NrdR family transcriptional regulator 2.1

SAR1952 hypothetical protein 2.1

SAR1017 menaquinone biosynthesis bifunctional protein 2.0

spoVG regulatory protein SpoVG 2.0

pfkA 6-phosphofructokinase 2.0

SAR0390 hypothetical protein 2.0

SAR1156 cell division protein 2.0

SAR1438 hypothetical protein 2.0

SAR2264 hypothetical protein 2.0

SAR2421 hypothetical protein 2.0

SAR0473 sugar-specific PTS transport system, IIBC component 2.0

SAR1165 hypothetical protein 2.0

kbl 2-amino-3-ketobutyrate coenzyme A ligase 2.0

SAR0669 response regulator protein 2.0

SAR1066 hypothetical protein 2.0

SAR1704 hypothetical protein 2.0

SAR2009 sodium:sulfate symporter 2.0

SAR2011 isochorismatase 2.0

SAR2413 short chain dehydrogenase 2.0

opuD1 glycine betaine transporter 1 -2.0

SAR0268 sugar transport protein -2.0

SAR2189 hypothetical protein -2.0

SARs214 predicted small RNA -2.0

groEL chaperonin GroEL -2.0

recU Holliday junction-specific endonuclease -2.0

SAR0664 hypothetical protein -2.0

SAR0487 DNA replication intiation control protein YabA -2.0

SARs129 predicted small RNA -2.0

trmD tRNA (guanine-N(1)-)-methyltransferase -2.0

SAR1982 hypothetical protein -2.1

pcrA ATP-dependent DNA helicase -2.1

SARs039 predicted small RNA -2.1

SARs137 predicted small RNA -2.1

SAR2795 DNA-binding protein -2.1

SARt021 tRNA-His -2.1

174

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

tuf elongation factor Tu -2.1

SAR0292 hypothetical protein -2.1

SAR0274 ABC transporter ATP-binding protein -2.1

SAR0800 hypothetical protein -2.1

adk adenylate kinase -2.1

mvaK2 phosphomevalonate kinase -2.1

SAR1222 succinyl-CoA synthetase subunit alpha -2.1

SAR1716 single-stranded-DNA-specific exonuclease -2.1

SAR1488 pyridine nucleotide-disulfide oxidoreductase -2.1

sucC succinyl-CoA synthetase subunit beta -2.1

folC folylpolyglutamate synthase -2.2

SAR0182 hypothetical protein -2.2

SAR0445 lipoprotein -2.2

SAR0630 monovalent cation/H+ antiporter subunit A -2.2

SAR2428 hypothetical protein -2.2

SAR2493 nitrite transporter -2.2

SARs022 GJA5-2092-RNA -2.2

SARt009 tRNA-Arg -2.2

secY_1 preprotein translocase subunit SecY -2.2

rpmG_3 ribosomal protein L33 -2.2

capC capsular polysaccharide synthesis enzyme -2.2

rpmH 50S ribosomal protein L34 -2.3

SAR0170 cation efflux system protein -2.3

SARs128 predicted small RNA -2.3

icaR ica operon transcriptional regulator -2.3

SAR0632 monovalent cation/H+ antiporter subunit C -2.3

SAR2623 hypothetical protein -2.3

prmA 50S ribosomal protein L11 methyltransferase -2.3

SARs003 GJA5-344-RNA -2.3

SAR0634 monovalent cation/H+ antiporter subunit E -2.3

SAR0636 hypothetical protein -2.3

SAR0549 ribosomal protein L7Ae-like -2.3

SARs061 predicted small RNA -2.3

SAR0287 hypothetical protein -2.4

SAR1173 RNA pseudouridylate synthase -2.4

SAR1726 hypothetical protein -2.4

SAR2150 hypothetical protein -2.4

175

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

SAR2295 hypothetical protein -2.4

infC translation initiation factor IF-3 -2.4

rpsG 30S ribosomal protein S7 -2.4

lig DNA ligase -2.4

rpoE DNA-directed RNA polymerase subunit delta -2.4

SAR2217 acetyltransferase -2.4

SAR0856 phosphoglycerate mutase -2.4

SAR1456 hypothetical protein -2.4

mvaS 3-hydroxy-3-methylglutaryl coenzyme A synthase -2.5

rplY 50S ribosomal protein L25/general stress protein Ctc -2.5

pyrR bifunctional pyrimidine regulatory protein PyrR/uracil phosphoribosyltransferase

-2.5

rplA 50S ribosomal protein L1 -2.5

capB capsular polysaccharide synthesis enzyme -2.5

SAR0633 monovalent cation/H+ antiporter subunit D -2.5

capA capsular polysaccharide synthesis enzyme -2.6

SAR1957 hypothetical protein -2.6

SAR2218 pantothenate kinase -2.6

SAR2561 hypothetical protein -2.6

gidA tRNA uridine 5-carboxymethylaminomethyl modification protein GidA

-2.6

SAR2430 permease -2.6

rpsP 30S ribosomal protein S16 -2.7

rpmE2 50S ribosomal protein L31 -2.7

SARs107 predicted small RNA -2.7

SAR2603 hypothetical protein -2.7

SAR2692 hypothetical protein -2.7

geh lipase precursor -2.7

rplS 50S ribosomal protein L19 -2.7

SARs233 predicted small RNA -2.7

SAR2769 hypothetical protein -2.7

SAR1455 hypothetical protein -2.8

rpsT 30S ribosomal protein S20 -2.8

SAR2043 enterotoxin type A precursor -2.8

SARs023 GJA5-2157-RNA -2.8

SAR0546 hypothetical protein -2.8

rplT 50S ribosomal protein L20 -2.8

SAR1100 hypothetical protein -2.8

176

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

SAR0280 hypothetical protein -2.8

rpmI 50S ribosomal protein L35 -2.9

SAR2168 helicase -2.9

capE capsular polysaccharide synthesis enzyme -2.9

grpE heat shock protein GrpE -2.9

SARs168 predicted small RNA -2.9

SAR2238 hypothetical protein -2.9

dltB activated D-alanine transport protein -2.9

hrcA heat-inducible transcription repressor -2.9

SARt031 tRNA-Trp -2.9

dnaJ chaperone protein DnaJ -2.9

rpsD 30S ribosomal protein S4 -2.9

groES co-chaperonin GroES -3.0

SAR0284 hypothetical protein -3.0

infA translation initiation factor IF-1 -3.0

fadA thiolase -3.0

rpsJ 30S ribosomal protein S10 -3.0

SAR2730 hypothetical protein -3.0

dltA D-alanine--poly(phosphoribitol) ligase subunit 1 -3.1

capD capsular polysaccharide synthesis enzyme -3.1

SAR1492 ferredoxin -3.1

SAR1402 phosphate-binding lipoprotein -3.1

ssb single-strand DNA-binding protein -3.1

capL capsular polysaccharide synthesis enzyme -3.1

capN capsular polysaccharide synthesis enzyme -3.1

purB adenylosuccinate lyase -3.2

SAR2610 L-serine dehydratase subunit alpha -3.2

ddh D-lactate dehydrogenase -3.2

capP capsular polysaccharide synthesis enzyme -3.2

SARs060 predicted small RNA -3.3

gidB 16S rRNA methyltransferase GidB -3.3

rplO 50S ribosomal protein L15 -3.3

SARs149 predicted small RNA -3.3

SAR1999 hypothetical protein -3.3

ldh1 L-lactate dehydrogenase -3.4

rplC 50S ribosomal protein L3 -3.4

capO capsular polysaccharide synthesis enzyme -3.4

177

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

SAR0171 hypothetical protein -3.5

rpsF 30S ribosomal protein S6 -3.5

rplW 50S ribosomal protein L23 -3.5

rpmJ 50S ribosomal protein L36 -3.6

dltD lipoteichoic acid biosynthesis protein -3.6

rpsS 30S ribosomal protein S19 -3.6

SAR1083 BipA family GTPase -3.6

SAR2660 hypothetical protein -3.7

SARs074 predicted small RNA -3.7

SARt041 tRNA-Met -3.7

cudA betaine aldehyde dehydrogenase -3.7

SAR0635 monovalent cation/H+ antiporter subunit F -3.8

SAR1348 hypothetical protein -3.8

cudB choline dehydrogenase -3.8

rplD 50S ribosomal protein L4 -3.9

capF capsular polysaccharide synthesis enzyme -3.9

SAR2666 hypothetical protein -3.9

SARs131 predicted small RNA -3.9

SAR0301 hypothetical protein -4.0

dltC D-alanine--poly(phosphoribitol) ligase subunit 2 -4.0

SAR2612 hypothetical protein -4.0

SARs130 predicted small RNA -4.1

rplV 50S ribosomal protein L22 -4.1

SAR1984 ferritin -4.1

capG capsular polysaccharide synthesis enzyme -4.3

rpmC 50S ribosomal protein L29 -4.5

SAR0378 hypothetical protein -4.6

SAR2338 xanthine/uracil permease -4.6

rplF 50S ribosomal protein L6 -4.7

rplB 50S ribosomal protein L2 -4.8

ssaA secretory antigen precursor -4.9

rplP 50S ribosomal protein L16 -4.9

rpsR 30S ribosomal protein S18 -5.0

SARs099 predicted small RNA -5.1

rpsC 30S ribosomal protein S3 -5.1

rpsH 30S ribosomal protein S8 -5.1

rpsE 30S ribosomal protein S5 -5.2

178

Table B-1. Continued

Gene name Function Fold Change (nos mutant/ wild-type)

rplN 50S ribosomal protein L14 -5.2

rplX 50S ribosomal protein L24 -5.4

rplR 50S ribosomal protein L18 -5.5

pyrB aspartate carbamoyltransferase catalytic subunit -5.6

rpsQ 30S ribosomal protein S17 -5.6

pyrP uracil permease -5.7

SARs121 predicted small RNA -5.7

rplE 50S ribosomal protein L5 -5.8

pyrC dihydroorotase -5.9

rpsN_2 ribosomal protein S14p/S29e -5.9

carB carbamoyl phosphate synthase large subunit -6.2

SARs120 predicted small RNA -6.3

pyrG CTP synthetase -6.5

PSMa phenol-soluble modulin alpha -6.7

rplJ 50S ribosomal protein L10 -7.0

pyrAA carbamoyl phosphate synthase small subunit -7.1

rpmD 50S ribosomal protein L30 -7.2

pyrE orotate phosphoribosyltransferase -7.2

pyrF orotidine 5'-phosphate decarboxylase -7.5

SAR1182 hypothetical protein -7.7

cudT choline transporter -7.7

SARt027 tRNA-Gly -8.5

rplL 50S ribosomal protein L7/L12 -9.0

purE phosphoribosylaminoimidazole carboxylase catalytic subunit -11.1

SAR1347 guanosine 5'-monophosphate oxidoreductase -11.5

purK phosphoribosylaminoimidazole carboxylase ATPase subunit -15.6

purC phosphoribosylaminoimidazole-succinocarboxamide synthase -22.3

purQ phosphoribosylformylglycinamidine synthase I -25.1

purL phosphoribosylformylglycinamidine synthase II -34.4

purN phosphoribosylglycinamide formyltransferase -36.8

SAR1041 hypothetical protein -37.6

purH bifunctional phosphoribosylaminoimidazolecarboxamide formyltransferase/IMP cyclohydrolase

-41.8

purF amidophosphoribosyltransferase -42.6

purD phosphoribosylamine--glycine ligase -46.0

purM phosphoribosylaminoimidazole synthetase -46.6

purA adenylosuccinate synthetase -77.1

179

Table B-2. List of all genes altered in the nos mutant at 6 hours growth Gene name Function Fold-change (nos

mutant/ wild-type)

SAR0231 hypothetical protein 26.6

SAR2006 nicotinate phosphoribosyltransferase 21.0

SAR2003 hypothetical protein 11.7

SARs054 predicted small RNA 10.9

nadE NAD synthetase 10.1

hmp flavohemoprotein 9.5

SAR1143 carbamate kinase 9.2

SAR0230 extracellular solute-binding lipoprotein 8.9

SAR2004 hypothetical protein 8.6

glnR glutamine synthetase 8.4

SAR0111 myosin-cross-reactive antigen 7.3

scdA cell wall biosynthesis protein ScdA 7.1

otc ornithine carbamoyltransferase 7.0

clpL ATP-dependent protease ATP-binding subunit ClpL 7.0

SAR0310 nucleoside permease 6.5

SAR0232 hypothetical protein 6.4

SAR1454 hypothetical protein 6.4

glnA glutamine synthetase, type I 6.0

SAR0218 pyruvate formate-lyase activating enzyme 6.0

opp-1A oligopeptide transporter substrate binding protein 5.9

qoxC quinol oxidase polypeptide III 5.3

SAR0308 PfkB family carbohydrate kinase 5.1

qoxD quinol oxidase polypeptide IV 5.1

pflB pyruvate formate lyase 5.0

SAR1402 phosphate-binding lipoprotein 4.9

SAR2681 amino acid permease 4.9

SAR2549 transporter 4.4

opp-1F oligopeptide transporter ATPase 4.4

qoxB quinol oxidase polypeptide I 4.3

SAR0309 hypothetical protein 4.3

qoxA quinol oxidase polypeptide II precursor 4.2

glmS glucosamine--fructose-6-phosphate aminotransferase 4.2

SARs086 predicted small RNA 4.2

cudA betaine aldehyde dehydrogenase 4.1

adhA alcohol dehydrogenase 3.9

SAR1091 hypothetical protein 3.8

fhs formate--tetrahydrofolate ligase 3.7

180

Table B-2. Continued

Gene name Function Fold-change (nos mutant/ wild-type)

SAR2186 hypothetical protein 3.7

fda fructose-1,6-bisphosphate aldolase 3.6

ldh2 L-lactate dehydrogenase 2 3.3

ctaB protoheme IX farnesyltransferase 3.3

glpQ glycerophosphoryl diester phosphodiesterase 3.3

opp-1C oligopeptide transporter membrane permease 3.2

SARs003 GJA5-344-RNA 3.1

opp-1D oligopeptide transporter ATPase 3.0

SAR2669 dihydroorotate dehydrogenase 2 3.0

SARs077 predicted small RNA 3.0

cudB choline dehydrogenase 3.0

SAR2528 amino acid permease 2.9

SAR2232 hypothetical protein 2.9

SAR0390 hypothetical protein 2.8

SAR0918 NADH:flavin oxidoreductase / NADH oxidase 2.8

ctaA heme A synthase 2.7

SAR0859 OsmC-like protein 2.7

SAR0865 hypothetical protein 2.6

kbl 2-amino-3-ketobutyrate coenzyme A ligase 2.6

pheS phenylalanyl-tRNA synthetase subunit alpha 2.6

SAR2228 hypothetical protein 2.6

SARs024 GJA5-2215-RNA 2.5

SARs097 predicted small RNA 2.5

sstD lipoprotein 2.5

opuD2 glycine betaine transporter 2 2.5

SAR0556 chaperone protein HchA 2.5

SAR0110 Na+/Pi-cotransporter protein 2.5

SAR0112 hypothetical protein 2.4

SAR2569 hypothetical protein 2.4

SAR2670 hypothetical protein 2.4

SAR2775 sodium:sulfate symporter family protein 2.4

SAR2245 transcriptional antiterminator 2.4

SAR0574 hexulose-6-phosphate synthase 2.4

SAR2646 phytoene dehydrogenase related protein 2.4

nrdD anaerobic ribonucleoside triphosphate reductase 2.4

SAR2007 oxygenase 2.3

SARs133 predicted small RNA 2.3

181

Table B-2. Continued

Gene name Function Fold-change (nos mutant/ wild-type)

pyrP uracil permease 2.3

cudT choline transporter 2.3

gap2 glyceraldehyde 3-phosphate dehydrogenase 2 2.3

SAR2470 hypothetical protein 2.3

thrS threonyl-tRNA synthetase 2.3

gcvH glycine-cleavage complex H protein 2.2

nupC nucleoside permease 2.2

uhpT sugar phosphate antiporter 2.2

SAR2778 nickel transport protein 2.2

citB aconitate hydratase 2.2

SAR2413 short chain dehydrogenase 2.2

pdhA pyruvate dehydrogenase E1 component subunit alpha 2.2

rpsA 30S ribosomal protein S1 2.2

SAR0559 branched-chain amino acid aminotransferase 2.2

SAR0585 phosphomethylpyrimidine kinase 2.1

SAR2647 hypothetical protein 2.1

SAR0307 hypothetical protein 2.1

SAR1222 succinyl-CoA synthetase subunit alpha 2.1

SAR2290 aldo/keto reductase 2.1

pdhB pyruvate dehydrogenase E1 component subunit beta 2.1

SAR0874 hypothetical protein 2.1

SAR1973 hypothetical protein 2.1

SAR2275 hypothetical protein 2.1

fumC fumarate hydratase 2.1

cycA D-serine/D-alanine/glycine transporter 2.1

SAR0575 6-phospho 3-hexuloisomerase 2.1

SAR2773 hypothetical protein 2.0

SAR2016 hypothetical protein 2.0

SAR2668 hypothetical protein 2.0

sucC succinyl-CoA synthetase, beta subunit 2.0

SAR1335 hypothetical protein 2.0

SAR0392 hypothetical protein 2.0

mscL large-conductance mechanosensitive channel -2.0

SAR1131 hypothetical protein -2.0

SAR0711 hypothetical protein -2.0

geh lipase precursor -2.0

agrD autoinducer peptide -2.0

182

Table B-2. Continued

Gene name Function Fold-change (nos mutant/ wild-type)

SAR0335 luciferase-like monooxygenase -2.0

SAR1600 exodeoxyribonuclease VII small subunit -2.0

dltB activated D-alanine transport protein -2.1

folK 2-amino-4-hydroxy-6- hydroxymethyldihydropteridine pyrophosphokinase

-2.1

SARs132 predicted small RNA -2.1

SAR1672 hypothetical protein -2.1

atl bifunctional autolysin precursor -2.1

dltA D-alanine--poly(phosphoribitol) ligase subunit 1 -2.1

gidB 16S rRNA methyltransferase GidB -2.1

SAR0632 monovalent cation/H+ antiporter subunit C -2.1

SAR2102 hypothetical protein -2.1

blaZ beta-lactamase precursor -2.1

SAR0634 monovalent cation/H+ antiporter subunit E -2.1

saeS histidine kinase -2.1

ruvB Holliday junction DNA helicase RuvB -2.1

SAR0284 hypothetical protein -2.1

est carboxylesterase -2.1

rplW 50S ribosomal protein L23 -2.2

rpsU 30S ribosomal protein S21 -2.2

SAR0437 hypothetical protein -2.2

SAR1857 hypothetical protein -2.2

SAR0179 transporter protein -2.2

SAR0633 monovalent cation/H+ antiporter subunit D -2.2

SAR2015 hypothetical protein -2.2

SARs107 predicted small RNA -2.2

hla alpha-hemolysin precursor -2.3

SARs224 predicted small RNA -2.3

malA alpha-D-1,4-glucosidase -2.3

SAR0286 hypothetical protein -2.3

rpsS 30S ribosomal protein S19 -2.3

SAR1379 peptidase -2.3

dltD lipoteichoic acid biosynthesis protein -2.3

rpsC 30S ribosomal protein S3 -2.3

SAR0763 radical activating enzyme -2.4

rplP 50S ribosomal protein L16 -2.4

rplV 50S ribosomal protein L22 -2.4

SAR2067 hypothetical protein -2.4

183

Table B-2. Continued

Gene name Function Fold-change (nos mutant/ wild-type)

rpmF 50S ribosomal protein L32 -2.5

arsB1 arsenical pump membrane protein 1 -2.5

SAR1079 manganese transport protein MntH -2.5

SARs226 predicted small RNA -2.5

rplN 50S ribosomal protein L14 -2.5

capP capsular polysaccharide synthesis enzyme -2.5

rplO 50S ribosomal protein L15 -2.5

SAR0401 sodium:dicarboxylate symporter protein -2.5

saeR response regulator protein -2.6

SAR0382 terminase small subunit -2.6

SAR0966 adaptor protein -2.6

SAR2056 hypothetical protein -3.0

SAR2052 hypothetical protein -2.6

SAR2062 Clp protease -2.6

SAR2050 hypothetical protein -2.6

infA translation initiation factor IF-1 -2.6

SAR2001 staphopain protease -2.7

SAR2054 hypothetical protein -2.7

SAR0636 hypothetical protein -2.7

sbi IgG-binding protein -2.7

SAR0760 hypothetical protein -2.7

SAR0635 monovalent cation/H+ antiporter subunit F -2.7

SARt041 tRNA-Met -2.7

ipk 4-diphosphocytidyl-2-C-methyl-D-erythritol kinase -2.8

dltC D-alanine--poly(phosphoribitol) ligase subunit 2 -2.8

rplF 50S ribosomal protein L6 -2.8

rpmC 50S ribosomal protein L29 -2.8

arsC arsenate reductase -2.8

rplE 50S ribosomal protein L5 -2.8

SAR1050 ABC transporter ATP-binding protein -2.9

rplR 50S ribosomal protein L18 -2.9

SAR2061 hypothetical protein -2.9

SAR2086 hypothetical protein -2.9

SARs128 predicted small RNA -3.0

cadA cadmium-transporting ATPase -2.9

capG capsular polysaccharide synthesis enzyme -3.0

rpsN_2 30S ribosomal protein S14 -3.0

184

Table B-2. Continued

Gene name Function Fold-change (nos mutant/ wild-type)

SAR1984 ferritin -3.1

capO capsular polysaccharide synthesis enzyme -3.1

SARt027 tRNA-Gly -3.1

SARs061 predicted small RNA -3.2

SAR0378 hypothetical protein -3.2

rpsE 30S ribosomal protein S5 -3.2

rpsH 30S ribosomal protein S8 -3.2

rpmD 50S ribosomal protein L30 -3.3

SAR2600 MarR family regulatory protein -3.3

SAR1378 prephenate dehydrogenase -3.3

SAR2119 membrane anchored protein -3.4

SAR0653 ABC transporter ATP-binding protein -3.5

SAR2048 hypothetical protein -3.5

SARs022 GJA5-2092-RNA -3.5

rpsQ 30S ribosomal protein S17 -3.6

SARt048 tRNA-Lys -3.6

SAR2096 anti repressor -3.6

rplX 50S ribosomal protein L24 -3.7

SAR2060 hypothetical protein -3.8

czrB zinc resistance protein -3.8

SAR2053 hypothetical protein -3.8

SAR2098 hypothetical protein -3.9

SAR0172 hypothetical protein -3.9

SAR2085 hypothetical protein -4.0

capL capsular polysaccharide synthesis enzyme -4.1

acpD azoreductase -4.8

rplJ 50S ribosomal protein L10 -4.8

capN capsular polysaccharide synthesis enzyme -4.8

rplL 50S ribosomal protein L7/L12 -5.0

perR peroxide operon regulator -5.

czrA zinc and cobalt transport repressor protein -5.2

lip lipase precursor -5.2

SAR0546 hypothetical protein -5.3

SARs131 predicted small RNA -5.5

SAR2598 phospholipase/carboxylesterase -5.7

PSMa phenol-soluble modulin alpha -6.3

SAR2227 non-heme iron-containing ferritin -7.2

185

Table B-2. Continued

Gene name Function Fold-change (nos mutant/ wild-type)

SAR1150 anti protein -8.4

SAR1377 ImpB/MucB/SamB family protein -10.6

fadA thiolase -18.2

fadB fatty oxidation complex protein -20.6

186

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BIOGRAPHICAL SKETCH

Austin has always been passionate for the pursuit of knowledge. It wasn’t until

volunteering as an undergraduate researcher did he begin to realize that scientific study

could fulfill that need. The ability to think critically, develop solutions, and then apply

them through experimental methods was something that he found in scientific research.

As an undergraduate Austin moved from various schools pursuing the premedical track

until transferring to the University of Florida with a major focus in microbiology and cell

science.

He began his research career as an undergraduate research assistant in the lab

of Dr. Kelly Rice where he studied Streptococcus mutans biofilms and the effects of a

novel drug delivery system on their growth and physiology. Here he learned how to

cultivate both static and flow cell biofilms as well as common antimicrobial testing

techniques for enumerating bacteria. At the same time, he became proficient on use of

the Zeiss confocal microscope for biofilm imaging and, in fact, became the department

trainer for anyone who wished access to the microscope. This work was published in

2015. While completing undergraduate research he was awarded a University Scholars

fellowship and presented his research at the 111th General Meeting American Society

for Microbiology (New Orleans LA, 2011). In 2011 he was award a Bachelor of Science

and culminated his undergraduate career by graduating with honors.

Thoroughly enjoying research on bacterial pathogens, Austin was accepted to

the Department of Microbiology and Cell Science graduate program where he began

research on the well-known human pathogen Staphylococcus aureus. Upon admission

he was granted a Grinter fellowship from the University of Florida Graduate School.

Graduate student responsibilities included courses, laboratory research, and teaching

239

for two semesters. Austin continued to teach the undergraduate microbiology

laboratories for 6 total semesters and was invited to teach the advanced laboratory, as

well as guest lecture in an undergraduate bioinformatics course. While in the laboratory

Austin also mentored multiple undergraduate students, allowing them to work

independently on research projects. He thoroughly enjoys teaching and seeing students

grow as scientists.

During his tenure as a graduate student, Austin presented his research at various

local and international conferences including the American Society for Microbiology

Southeastern Branch Meeting (Athens GA, 2012 and Gainesville FL, 2011),

International Conference for Gram Positive Pathogens (Omaha NE, 2014 and 2016),

ASM microbe (Boston MA, 2016), and multiple department seminars. As well, he was

awarded a prestigious ASM student travel grant 2016 for outstanding abstract

submission. These experiences further developed Austin as a public speaker where he

honed the art of clearly communicating scientific concepts and ideas.

Austin’s PhD dissertation focused on characterizing the role of the nitric oxide

synthase in general S. aureus physiology. The hope was to uncover the mechanisms

that this bacterium uses for its biological processes in an attempt to discover novel

antimicrobial drug targets. Austin has a co-first author publication on this topic and has

contributed to a second publication on Staphylococcal small RNAs. Currently a third

manuscript is under review that encompasses the bulk of his dissertation work. Austin

will begin his post graduate career as a senior scientist working for Brammer Bio.