University of Florida Thesis Or Dissertation

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

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

To my mother Randy Mogen

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.

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

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

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

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

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

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

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

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

RNS Reactive nitrogen species

ROS Reactive oxygen species

S. aureus Staphylococcus aureus

S. carnosus Staphylococcus carnosus

S. cellulosum Sorangium cellulosum

S. epidermidis Staphylococcus epidermidis

S. saprophyticus Staphylococcus saprophyticus

S. turgidiscabies 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 Tricarboxylic acid

TCS 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

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

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

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.

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

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

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

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.

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,

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

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,

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

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

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

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)

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

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

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

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.

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

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

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

(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)

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-

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

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

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

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

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

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.

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

(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

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

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

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

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

(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

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-

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).

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).

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

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

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

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

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,

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).

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

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

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,

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-ethylisothiourea. 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

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

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.,

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

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

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

2+ the Fenton reaction H2O2 oxidizes free cellular Fe 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.

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-

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.

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).

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

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

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

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

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

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

metabolic adaptations in the nos mutant, Aim 3 sought to determine potential regulators of nos mutant metabolic adaptation.

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).

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.

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.

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).

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.

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.

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.

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

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

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-

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.

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

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

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.

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

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,

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

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

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,

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

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

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

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

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

100

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.

101

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.

102

1e+10 10 A B 1e+9

1 600 1e+8

CFU/ml 1e+7 UAMS-1 UAMS-1 0.1 nos mutant nos mutant nos complement nos complement 1e+6 0 2 4 6 8 0 2 4 6 8 Time (Hours) Time (Hours) 1010 10 UAMS-1 C D nos mutant nos complement

109 1

600

CFU/ml 108 UAMS-1 0.1 nos mutant nos complement 107 0 5 10 15 20 25 0 5 10 15 20 25 Time (Hours) Time (Hours)

10 E 1010 F

109 1

600

CFU/ml 108 UAMS-1 UAMS-1 0.1 nos mutant nos mutant nos complement nos complement 107 0 5 10 15 20 25 0 5 10 15 20 25 Time (Hours) Time (Hours) 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.

103

1010 10 A B

109

600

CFU/ml 108

UAMS-1 UAMS-1 0.1 nos mutant nos mutant nos mutant + NO nos mutant + NO 107 0 2 4 6 8 0 2 4 6 8 Time (Hours) Time (Hours) 1010 10 C D

109

600

CFU/ml 108

0.1 LAC-13C LAC-13C nos mutant nos mutant 107 0 5 10 15 20 25 0 5 10 15 20 25 Time (Hours) Time (Hours)

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.

104

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.

105

Wildtype

D 900 B * 800

nos 700

mutant 600

Cell length (nm) length Cell 500 C 400 nos complement

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.

106

Electron Transport Chain Proteins and Component Biosynthesis Downregulated Pyruvate and Carbohydrate Metabolism Upregulated Amino Acid Metabolism Anaerobic Metabolism and Fermentation Stress Response PTS Systems Two Component Systems ABC Transporters General Signal Transduction Amino Nucleotide/Sugar metabolism Purine and Pyrimidine Degradation Amino Acid Transport Ion Dependant Transporters Other Transport Proteins Other Metabolic Pathways Predicted Small RNAs Hypothetical Proteins DNA Replication and Modification Virulence Lipoproteins TCA cycle and Intermediate Metabolism Transcriptional Regulation Fatty Acid Oxidation Iron Storage and Protein Biosynthesis Mevalonate Pathway Proteases, Protein Folding and Degradation Capsular Biosynthesis Purine and Pyrimidine Biosynthesis Translation Proteins and tRNA

0 10 20 30 40 50 60 Number of Genes 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.

107

Electron Transport Chain Proteins and Component Biosynthesis Downregulated Pyruvate and Carbohydrate Metabolism Upregulated Amino Acid Metabolism Anaerobic Metabolism and Fermentation Stress Response PTS Systems Two Component Systems ABC Transporters General Signal Transduction Amino Nucleotide/Sugar metabolism Purine and Pyrimidine Degradation Amino Acid Transport Ion Dependant Transporters Other Transport Proteins Other Metabolic Pathways Predicted Small RNAs Hypothetical Proteins DNA Replication and Modification Virulence Lipoproteins TCA Cycle and Intermediate Metabolism Transcriptional Regulation Fatty Acid Oxidation Iron Storage and Protein Biosynthesis Mevalonate Pathway Proteases, Protein Folding and Degradation Capsular Biosynthesis Purine and Pyrimidine Biosynthesis Translation Proteins and tRNA

0 5 10 15 20 25 30 35 Number of Genes 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.

108

3000 2500

) ) Wildtype A nos mutant B

600 600 2500 * nos complement 2000 * 2000 1500 1500 * 1000 1000

500 500

Fluorescence (RFU/OD Fluorescence

Fluorescence (RFU/OD Fluorescence 0 0 3 Hours 6 Hours

1000 4x105

)

600 C D 800 ** 5 3x10 **

600 2x105 400

105 200

Fluorescence (RFU/OD Fluorescence 0 0 protein) (U/mg Activity Catalase

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.

109

Wildtype Wildtype + NO A nos mutant nos mutant + NO

nos complement Events

Fluorescence (Red:Green Ratio)

Wildtype nos mutant B

nos complement Events

Fluorescence (Red:Green Ratio)

Wildtype nos mutant C

nos complement Events

Fluorescence (Red:Green Ratio) 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.

110

1.6 1.6 1.4 A * 1.4 B ** 1.2 1.2 1.0 1.0 0.8 0.8 0.6 0.6 0.4 0.4 0.2 0.2 0.0 0.0

Fold-change (Relative to Wildtype) (Relative Fold-change

1.2 1.6 1.4 D ** 1.0 C 1.2 0.8 1.0

0.6 *** 0.8 0.6 0.4 0.4 0.2 0.2

0.0 0.0

Fold-change (Relative to Wildtype) (Relative Fold-change

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.

111

120

100

Relative Oxygen Consumption Rate (%) Rate Consumption Oxygen Relative

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.

112

2500 ) Untreated Thioridizine Treated 600 A 2000 *

1500

1000

500

Fluorescence (RFU/OD Fluorescence 0

30 B 25

10 **

Aconitase Activity Aconitase

(nmol/min/mg protein) (nmol/min/mg 5

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.

113

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.

114

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.

115

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.

116

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.

117

Table 2-1. Generation times for all strains Strain CFU/ml Growth Generation Media Time (Minutes) ± SEM 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

118

Table 2-2. Select genes altered upon nos mutation Category Gene Function Fold-change Fold-change Name (nos/wt) (nos/wt) 4 hrs growth 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 2.1 --- protein 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 Fatty acid degradation operon --- -20.6 SAR2006 NAD biosynthesis operon 49.8 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 ---

119

Table 2-3. qRT-PCR confirmation of select genes Gene RNAseq Real-time PCR (4 hr RNA) Name (Fold-change nos/wt) 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

120

Table 2-4. Select cellular nos mutant metabolites Cellular Metabolite % increase/decrease P-value nos mutant vs wildtype (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

121

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

122

Table 2-6. Select nos srrAB double mutant cellular metabolites Cellular Metabolite % increase/decrease P-value nos/srrAB mutant vs (Two tailed t-test) wildtype Organic Acids Succinate -94 0.014 Malate -87 0.045 Citrate BLOQ --- α-ketoglutarate BLOQ --- Fumarate BLOQ --- Amino Acids Alanine -90 0.002 Arginine BLOQ --- Asparagine -65 0.018 Aspartate -70 0.002 Citrulline 347 0.003 Glutamate -81 <0.001 Glycine BLOQ --- Histidine BLOQ --- Lysine -69 0.042 Methionine -28 0.004 Ornithine -87 <0.001 Proline -82 0.021 Serine BLOQ --- Threonine BLOQ --- Tyrosine -71 0.022 Valine -41 0.003 Adenine Nucleotide NADP -40 0.002 NADPH BLOQ BLOQ NADH -74 0.032 NAD/NADH 252 0.006 ADP 32 0.030 ATP 23 0.578

123

Table 2-7. Select nos srrAB double mutant extracellular metabolites Extracellular % increase/decrease P-value Metabolite nos/srrAB mutant vs (Two tailed t- wildtype 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

124

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 -80C 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

125

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

126

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

127

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

128

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

129

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

130

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 -80C. 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 -80C. 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

131

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

132

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

133

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

134

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.

135

- 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

136

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.

137

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

138

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

139

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.

140

Table 3-1. Bacterial strains and plasmids constructs used in this study Strain or plasmid name Description Reference or source E. coli DH5α Host strain for construction of (Hanahan, recombinant plasmids 1983)

Staphylococcus aureus Easily transformable restriction (Kreiswirth et RN4220 deficient strain al., 1983)

UAMS-1 Osteomyelitis clinical isolate (Gillaspy et al., 1995)

LAC-13C CA-MRSA isolate (Fey et al., 2013)

KR1010 UAMS-1 nos::erm insertion mutant (Sapp et al., 2014)

KR1040 LAC-13C nos::erm insertion mutant Unpublished strain created by K.C. Rice

KB6004 UAMS-1 srrAB deletion mutant (Lewis et al., 2015)

ABM10 UAMS-1 Δnos srrAB::erm double This study mutant pCRBlunt E. coli cloning plasmid; KmR Invitrogen pTR27 nos::erm allele-replacement plasmid; (Sapp et al., ErmR/CmR 2014) pBT2 Temperature-sensitive shuttle vector; (Bruckner, CmR/AmpR 1997) pMKnos nos complementation plasmid (Sapp et al., 2014) pMK4 Shuttle vector; CmR/AmpR (Sullivan et al., 1984)

141

Table 3-2. PCR primers used in this study Primer Purpose Sequence (5-3) Reference or Source sigA-F Real-time CAAGCAATCACTCGTGCAAT (Sapp et al., sigA-R PCR GGTGCTGGATCTCGACCTAA 2014)

SAR2006-RT-F Real-time TCAACCAGCATTAGGTGCAG This study SAR2006-RT-R PCR CTGGCGTAGTAACCTTTTCAGC

SAR0218-RT-F Real-time GCTGTTAAAGCAGCCTACCG This study SAR0218-RT-R PCR AGAAGCATATGCCCCTTCAC

SAR2680-RT-F Real-time CTTGCAGTTTGGTCACAAGC This study SAR2680-RT-R PCR TTCCGCTTTAGCTTCGCTAC

SAR1032-RT-F Real-time ACGCATGGTTGTCACGTATC This study SAR1032-RT-R PCR TGTCTAATCCGCGTCGTTG

SAR2486-RT-F Real-time CGGCAAGAGCAGTTATTTCG This study SAR2486-RT-R PCR GACCCAGGCGTTTGAATATG purH-RT-F Real-time CGAAATAAACCGCAGCATTT (Sapp et al., purH-RT-R PCR TCGTCACATCAGGGTTAGCA 2014) scdA-RT-F Real-time TGCGGCGGACAAGTAAGTAT This study scdA-RT-R PCR GCGAACCTGGTGTATTCGTT hmp-RT-F Real-time AGAGGCATGCAATCTTCAGC This study hmp-RT-R PCR AGTGCGCAGTGTTTATATGC

142

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

143

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

144

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-

145

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.

146

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

147

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

148

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

149

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

150

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

151

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

152

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.

153

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.

154

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.

155

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

156

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

157

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

158

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

159

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

160

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.

161

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).

162

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).

163

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).

164

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).

165

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).

166

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).

167

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

168

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

169

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 2.7 superfamily protein 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

170

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

171

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 2.1 protein 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

172

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

173

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

174

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 -2.5 phosphoribosyltransferase 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 -2.6 GidA 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

175

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

176

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

177

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 -41.8 formyltransferase/IMP cyclohydrolase purF amidophosphoribosyltransferase -42.6 purD phosphoribosylamine--glycine ligase -46.0 purM phosphoribosylaminoimidazole synthetase -46.6 purA adenylosuccinate synthetase -77.1

178

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

179

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

180

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

181

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 -2.1 pyrophosphokinase 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

182

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

183

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

184

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

185

LIST OF REFERENCES

(1999) From the Centers for Disease Control and Prevention. Four pediatric deaths from community-acquired methicillin-resistant Staphylococcus aureus--Minnesota and North Dakota, 1997-1999. Jama 282: 1123-1125.

(2003) Outbreaks of community-associated methicillin-resistant Staphylococcus aureus skin infections--Los Angeles County, California, 2002-2003. MMWR. Morbidity and mortality weekly report 52: 88.

(2012) Active Bacterial Core Surveillance Report, Emerging Infections Program Network, Methicillin - Resistant Staphylococcus aureus. In.: Centers for Disease Control and Prevention, pp.

Abdelnour, A., Arvidson, S., Bremell, T., Ryden, C., and Tarkowski, A. (1993) The accessory gene regulator (agr) controls Staphylococcus aureus virulence in a murine arthritis model. Infect Immun 61: 3879-3885.

Adak, S., Aulak, K.S., and Stuehr, D.J. (2002a) Direct evidence for nitric oxide production by a nitric-oxide synthase-like protein from Bacillus subtilis. J Biol Chem 277: 16167-16171.

Adak, S., Bilwes, A.M., Panda, K., Hosfield, D., Aulak, K.S., McDonald, J.F., Tainer, J.A., Getzoff, E.D., Crane, B.R., and Stuehr, D.J. (2002b) Cloning, expression, and characterization of a nitric oxide synthase protein from Deinococcus radiodurans. Proc Natl Acad Sci U S A 99: 107-112.

Agapie, T., Suseno, S., Woodward, J.J., Stoll, S., Britt, R.D., and Marletta, M.A. (2009) NO formation by a catalytically self-sufficient bacterial nitric oxide synthase from Sorangium cellulosum. Proc Natl Acad Sci U S A 106: 16221-16226.

Agapov, A.A., and Kulbachinskiy, A.V. (2015) Mechanisms of stress resistance and gene regulation in the radioresistant bacterium Deinococcus radiodurans. Biochemistry (Mosc) 80: 1201-1216.

Ahmed, S., Meghji, S., Williams, R.J., Henderson, B., Brock, J.H., and Nair, S.P. (2001) Staphylococcus aureus fibronectin binding proteins are essential for internalization by osteoblasts but do not account for differences in intracellular levels of bacteria. Infect Immun 69: 2872-2877.

Alalwani, S.M., Sierigk, J., Herr, C., Pinkenburg, O., Gallo, R., Vogelmeier, C., and Bals, R. (2010) The antimicrobial peptide LL-37 modulates the inflammatory and host defense response of human neutrophils. Eur J Immunol 40: 1118-1126.

Alderton, W.K., Cooper, C.E., and Knowles, R.G. (2001) Nitric oxide synthases: structure, function and inhibition. Biochem J 357: 593-615.

186

Almand, E., (2010) Characterization of the Staphylococcus aureus nitric oxide synthase gene and its role in biofilm development. In: Microbiology and Cell Science. University of Florida, pp.

Alonzo, F., 3rd, Kozhaya, L., Rawlings, S.A., Reyes-Robles, T., DuMont, A.L., Myszka, D.G., Landau, N.R., Unutmaz, D., and Torres, V.J. (2013) CCR5 is a receptor for Staphylococcus aureus leukotoxin ED. Nature 493: 51-55.

Aly, R., Shirley, C., Cunico, B., and Maibach, H.I. (1978) Effect of prolonged occlusion on the microbial flora, pH, carbon dioxide and transepidermal water loss on human skin. J Invest Dermatol 71: 378-381.

Anderson, C.B., and Witter, L.D. (1982) Glutamine and proline accumulation by Staphylococcus aureus with reduction in water activity. Appl Environ Microbiol 43: 1501-1503.

Angerer, H., Nasiri, H.R., Niedergesass, V., Kerscher, S., Schwalbe, H., and Brandt, U. (2012) Tracing the tail of ubiquinone in mitochondrial complex I. Biochim Biophys Acta 1817: 1776-1784.

Appelbaum, P.C. (2006) The emergence of vancomycin-intermediate and vancomycin- resistant Staphylococcus aureus. Clin Microbiol Infect 12 Suppl 1: 16-23.

Archer, G.L. (1998) Staphylococcus aureus: a well-armed pathogen. Clin Infect Dis 26: 1179-1181.

Archibald, F.S., and Fridovich, I. (1983) Oxygen radicals, oxygen toxicity and the life of microorganisms. Acta Med Port 4: 101-112.

Arner, E.S., and Holmgren, A. (2000) Physiological functions of thioredoxin and thioredoxin reductase. Eur J Biochem 267: 6102-6109.

Arnold, W.P., Mittal, C.K., Katsuki, S., and Murad, F. (1977) Nitric oxide activates guanylate cyclase and increases guanosine 3':5'-cyclic monophosphate levels in various tissue preparations. Proceedings of the National Academy of Sciences of the United States of America 74: 3203-3207.

Arnold, W.V., and Fertala, A. (2013) Skeletal diseases caused by mutations that affect collagen structure and function. Int J Biochem Cell Biol 45: 1556-1567.

Arora, D.P., Hossain, S., Xu, Y., and Boon, E.M. (2015) Nitric Oxide Regulation of Bacterial Biofilms. Biochemistry 54: 3717-3728.

Arroyo, P.L., Hatch-Pigott, V., Mower, H.F., and Cooney, R.V. (1992) Mutagenicity of nitric oxide and its inhibition by antioxidants. Mutation research 281: 193-202.

187

Arruebarrena Di Palma, A., Pereyra, C.M., Moreno Ramirez, L., Xiqui Vazquez, M.L., Baca, B.E., Pereyra, M.A., Lamattina, L., and Creus, C.M. (2013) Denitrification- derived nitric oxide modulates biofilm formation in Azospirillum brasilense. FEMS Microbiol Lett 338: 77-85.

Athanasopoulos, A.N., Economopoulou, M., Orlova, V.V., Sobke, A., Schneider, D., Weber, H., Augustin, H.G., Eming, S.A., Schubert, U., Linn, T., Nawroth, P.P., Hussain, M., Hammes, H.P., Herrmann, M., Preissner, K.T., and Chavakis, T. (2006) The extracellular adherence protein (Eap) of Staphylococcus aureus inhibits wound healing by interfering with host defense and repair mechanisms. Blood 107: 2720-2727.

Atkinson, D.E., and Fall, L. (1967) Adenosine triphosphate conservation in biosynthetic regulation. Escherichia coli phosphoribosylpyrophosphate synthase. J Biol Chem 242: 3241-3242.

Atkinson, D.E., and Walton, G.M. (1967) Adenosine triphosphate conservation in metabolic regulation. Rat liver citrate cleavage enzyme. J Biol Chem 242: 3239- 3241.

Baba, T., Bae, T., Schneewind, O., Takeuchi, F., and Hiramatsu, K. (2008) Genome sequence of Staphylococcus aureus strain Newman and comparative analysis of staphylococcal genomes: polymorphism and evolution of two major pathogenicity islands. J Bacteriol 190: 300-310.

Baba, T., Takeuchi, F., Kuroda, M., Yuzawa, H., Aoki, K., Oguchi, A., Nagai, Y., Iwama, N., Asano, K., Naimi, T., Kuroda, H., Cui, L., Yamamoto, K., and Hiramatsu, K. (2002) Genome and virulence determinants of high virulence community- acquired MRSA. Lancet 359: 1819-1827.

Ballal, A., and Manna, A.C. (2009) Regulation of superoxide dismutase (sod) genes by SarA in Staphylococcus aureus. J Bacteriol 191: 3301-3310.

Ballal, A., and Manna, A.C. (2010) Control of thioredoxin reductase gene (trxB) transcription by SarA in Staphylococcus aureus. J Bacteriol 192: 336-345.

Balwit, J.M., van Langevelde, P., Vann, J.M., and Proctor, R.A. (1994) Gentamicin- resistant menadione and hemin auxotrophic Staphylococcus aureus persist within cultured endothelial cells. The Journal of infectious diseases 170: 1033- 1037.

Barraud, N., Hassett, D.J., Hwang, S.H., Rice, S.A., Kjelleberg, S., and Webb, J.S. (2006) Involvement of nitric oxide in biofilm dispersal of Pseudomonas aeruginosa. J Bacteriol 188: 7344-7353.

188

Barrett, J.G., and Scott, I.R. (1983) Pyrrolidone carboxylic acid synthesis in guinea pig epidermis. J Invest Dermatol 81: 122-124.

Baruah, A., Lindsey, B., Zhu, Y., and Nakano, M.M. (2004) Mutational analysis of the signal-sensing domain of ResE histidine kinase from Bacillus subtilis. J Bacteriol 186: 1694-1704.

Bayer, A.S., McNamara, P., Yeaman, M.R., Lucindo, N., Jones, T., Cheung, A.L., Sahl, H.G., and Proctor, R.A. (2006) Transposon disruption of the complex I NADH oxidoreductase gene (snoD) in Staphylococcus aureus is associated with reduced susceptibility to the microbicidal activity of thrombin-induced platelet microbicidal protein 1. J Bacteriol 188: 211-222.

Berube, B.J., and Bubeck Wardenburg, J. (2013) Staphylococcus aureus alpha-toxin: nearly a century of intrigue. Toxins (Basel) 5: 1140-1166.

Bestebroer, J., van Kessel, K.P., Azouagh, H., Walenkamp, A.M., Boer, I.G., Romijn, R.A., van Strijp, J.A., and de Haas, C.J. (2009) Staphylococcal SSL5 inhibits leukocyte activation by chemokines and anaphylatoxins. Blood 113: 328-337.

Bhattacharyya, S., Dutta, D., Ghosh, A.K., and Das, A.K. (2009) Cloning, overexpression, purification, crystallization and preliminary X-ray diffraction analysis of an atypical two-cysteine peroxiredoxin (SAOUHSC_01822) from Staphylococcus aureus NCTC 8325. Acta Crystallogr Sect F Struct Biol Cryst Commun 65: 1113-1115.

Bird, L.E., Ren, J., Zhang, J., Foxwell, N., Hawkins, A.R., Charles, I.G., and Stammers, D.K. (2002) Crystal structure of SANOS, a bacterial nitric oxide synthase oxygenase protein from Staphylococcus aureus. Structure 10: 1687-1696.

Bitoun, J.P., Wu, G., and Ding, H. (2008) Escherichia coli FtnA acts as an iron buffer for re-assembly of iron-sulfur clusters in response to hydrogen peroxide stress. Biometals 21: 693-703.

Bogdan, C., Rollinghoff, M., and Diefenbach, A. (2000) Reactive oxygen and reactive nitrogen intermediates in innate and specific immunity. Curr Opin Immunol 12: 64-76.

Bohn, C., Rigoulay, C., Chabelskaya, S., Sharma, C.M., Marchais, A., Skorski, P., Borezee-Durant, E., Barbet, R., Jacquet, E., Jacq, A., Gautheret, D., Felden, B., Vogel, J., and Bouloc, P. (2010) Experimental discovery of small RNAs in Staphylococcus aureus reveals a riboregulator of central metabolism. Nucleic Acids Res 38: 6620-6636.

189

Boisset, S., Geissmann, T., Huntzinger, E., Fechter, P., Bendridi, N., Possedko, M., Chevalier, C., Helfer, A.C., Benito, Y., Jacquier, A., Gaspin, C., Vandenesch, F., and Romby, P. (2007) Staphylococcus aureus RNAIII coordinately represses the synthesis of virulence factors and the transcription regulator Rot by an antisense mechanism. Genes & Development 21: 1353-1366.

Boles, B.R., and Horswill, A.R. (2008) Agr-mediated dispersal of Staphylococcus aureus biofilms. PLoS pathogens 4: e1000052.

Borisov, V.B., Forte, E., Konstantinov, A.A., Poole, R.K., Sarti, P., and Giuffre, A. (2004) Interaction of the bacterial terminal oxidase cytochrome bd with nitric oxide. FEBS letters 576: 201-204.

Borisov, V.B., Forte, E., Sarti, P., Brunori, M., Konstantinov, A.A., and Giuffre, A. (2006) Nitric oxide reacts with the ferryl-oxo catalytic intermediate of the CuB-lacking cytochrome bd terminal oxidase. FEBS letters 580: 4823-4826.

Bose, J.L. (2014) Genetic manipulation of staphylococci. Methods Mol Biol 1106: 101- 111.

Boucher, H.W., and Corey, G.R. (2008) Epidemiology of methicillin-resistant Staphylococcus aureus. Clin Infect Dis 46 Suppl 5: S344-349.

Bouillaut, L., Dubois, T., Sonenshein, A.L., and Dupuy, B. (2015) Integration of metabolism and virulence in Clostridium difficile. Res Microbiol 166: 375-383.

Boveris, A., Costa, L.E., Poderoso, J.J., Carreras, M.C., and Cadenas, E. (2000) Regulation of mitochondrial respiration by oxygen and nitric oxide. Ann N Y Acad Sci 899: 121-135.

Bowman, L.A., McLean, S., Poole, R.K., and Fukuto, J.M. (2011) The diversity of microbial responses to nitric oxide and agents of nitrosative stress close cousins but not identical twins. Adv Microb Physiol 59: 135-219.

Brekasis, D., and Paget, M.S. (2003) A novel sensor of NADH/NAD+ redox poise in Streptomyces coelicolor A3(2). Embo j 22: 4856-4865.

Brinsmade, S.R. (2016) CodY, a master integrator of metabolism and virulence in Gram-positive bacteria. Curr Genet.

Brown, G.C. (1995) Nitric oxide regulates mitochondrial respiration and cell functions by inhibiting cytochrome oxidase. FEBS letters 369: 136-139.

Brown, G.C., McBride, A.G., Fox, E.J., McNaught, K.S., and Borutaite, V. (1997) Nitric oxide and oxygen metabolism. Biochemical Society transactions 25: 901-904.

190

Bruckner, R. (1997) Gene replacement in Staphylococcus carnosus and Staphylococcus xylosus. FEMS Microbiol Lett 151: 1-8.

Brunelli, L., Yermilov, V., and Beckman, J.S. (2001) Modulation of catalase peroxidatic and catalatic activity by nitric oxide. Free Radic Biol Med 30: 709-714.

Brunori, M., Forte, E., Arese, M., Mastronicola, D., Giuffre, A., and Sarti, P. (2006) Nitric oxide and the respiratory enzyme. Biochimica et biophysica acta 1757: 1144- 1154.

Brunori, M., Giuffre, A., Forte, E., Mastronicola, D., Barone, M.C., and Sarti, P. (2004) Control of cytochrome c oxidase activity by nitric oxide. Biochimica et biophysica acta 1655: 365-371.

Bryk, R., Griffin, P., and Nathan, C. (2000) Peroxynitrite reductase activity of bacterial peroxiredoxins. Nature 407: 211-215.

Burke, F.M., Di Poto, A., Speziale, P., and Foster, T.J. (2011) The A domain of fibronectin-binding protein B of Staphylococcus aureus contains a novel fibronectin binding site. Febs j 278: 2359-2371.

Burke, K.A., and Lascelles, J. (1975) Nitrate reductase system in Staphylococcus aureus wild type and mutants. J Bacteriol 123: 308-316.

Burke, K.A., and Lascelles, J. (1979) Partial purification and some properties of the Staphylococcus aureus cytoplasmic nitrate reductase. J Bacteriol 139: 120-125.

Butler, C., Forte, E., Maria Scandurra, F., Arese, M., Giuffre, A., Greenwood, C., and Sarti, P. (2002) Cytochrome bo(3) from Escherichia coli: the binding and turnover of nitric oxide. Biochemical and biophysical research communications 296: 1272- 1278.

Buttrey, T.V. (2012) Dio, Zonaras and the Value of the Roman Aureus. Journal of Roman Studies 51: 40-45.

Buys, E., and Sips, P. (2014) New insights into the role of soluble guanylate cyclase in blood pressure regulation. Curr Opin Nephrol Hypertens 23: 135-142.

Carroll, M.C. (2004) The complement system in regulation of adaptive immunity. Nat Immunol 5: 981-986.

Carroll, R.K., Weiss, A., Broach, W.H., Wiemels, R.E., Mogen, A.B., Rice, K.C., and Shaw, L.N. (2016a) Genome-wide annotation, identification, and global transcriptomic analysis of regulatory or small RNA gene expression in Staphylococcus aureus. mBio In press.

191

Carroll, R.K., Weiss, A., and Shaw, L.N. (2016b) RNA-sequencing of Staphylococcus aureus messenger RNA. Methods in molecular biology (Clifton, N.J.) 1373: 131- 141.

Caspi, R., Altman, T., Billington, R., Dreher, K., Foerster, H., Fulcher, C.A., Holland, T.A., Keseler, I.M., Kothari, A., Kubo, A., Krummenacker, M., Latendresse, M., Mueller, L.A., Ong, Q., Paley, S., Subhraveti, P., Weaver, D.S., Weerasinghe, D., Zhang, P., and Karp, P.D. (2014) The MetaCyc database of metabolic pathways and enzymes and the BioCyc collection of Pathway/Genome Databases. Nucleic Acids Res 42: D459-471.

Castro, C.E. (1980) Mechanisms of reaction of hemeproteins with oxygen and hydrogen peroxide in the oxidation of organic substrates. Pharmacol Ther 10: 171-189.

Cedergren, L., Andersson, R., Jansson, B., Uhlen, M., and Nilsson, B. (1993) Mutational analysis of the interaction between staphylococcal protein A and human IgG1. Protein Eng 6: 441-448.

Chambers, H., and Neu, H. (2000) Penicillins. Principles and practice of infectious diseases 5: 261-274.

Chang, J.P., and Lascelles, J. (1963) Nitrate reductase in cell-free extracts of a haemin- requiring strain of Staphylococcus aureus. The Biochemical journal 89: 503-510.

Chang, W., Small, D.A., Toghrol, F., and Bentley, W.E. (2006) Global transcriptome analysis of Staphylococcus aureus response to hydrogen peroxide. Journal of bacteriology 188: 1648-1659.

Chapman, G.H. (1945) The significance of sodium chloride in studies of staphylococci. J Bacteriol 50: 201-203.

Chartier, F.J., Blais, S.P., and Couture, M. (2006) A weak Fe-O bond in the oxygenated complex of the nitric-oxide synthase of Staphylococcus aureus. J Biol Chem 281: 9953-9962.

Chartier, F.J., and Couture, M. (2004) Stability of the heme environment of the nitric oxide synthase from Staphylococcus aureus in the absence of pterin cofactor. Biophys J 87: 1939-1950.

Chartier, F.J., and Couture, M. (2007a) Interactions between substrates and the haem- bound nitric oxide of ferric and ferrous bacterial nitric oxide synthases. Biochem J 401: 235-245.

Chartier, F.J., and Couture, M. (2007b) Substrate-specific interactions with the heme- bound oxygen molecule of nitric-oxide synthase. J Biol Chem 282: 20877-20886.

192

Chatterjee, I., Schmitt, S., Batzilla, C.F., Engelmann, S., Keller, A., Ring, M.W., Kautenburger, R., Ziebuhr, W., Hecker, M., Preissner, K.T., Bischoff, M., Proctor, R.A., Beck, H.P., Lenhof, H.P., Somerville, G.A., and Herrmann, M. (2009) Staphylococcus aureus ClpC ATPase is a late growth phase effector of metabolism and persistence. Proteomics 9: 1152-1176.

Chavakis, T., Hussain, M., Kanse, S.M., Peters, G., Bretzel, R.G., Flock, J.I., Herrmann, M., and Preissner, K.T. (2002) Staphylococcus aureus extracellular adherence protein serves as anti-inflammatory factor by inhibiting the recruitment of host leukocytes. Nat Med 8: 687-693.

Chen, Y., and Junger, W.G. (2012) Measurement of oxidative burst in neutrophils. Methods Mol Biol 844: 115-124.

Chen, Y., and Rosazza, J.P. (1994) A bacterial nitric oxide synthase from a Nocardia species. Biochem Biophys Res Commun 203: 1251-1258.

Chevalier, C., Boisset, S., Romilly, C., Masquida, B., Fechter, P., Geissmann, T., Vandenesch, F., and Romby, P. (2010) Staphylococcus aureus RNAIII binds to two distant regions of coa mRNA to arrest translation and promote mRNA degradation. PLoS Pathog 6: e1000809.

Choi, W.S., Chang, M.S., Han, J.W., Hong, S.Y., and Lee, H.W. (1997) Identification of nitric oxide synthase in Staphylococcus aureus. Biochem Biophys Res Commun 237: 554-558.

Choi, W.S., Seo, D.W., Chang, M.S., Han, J.W., Hong, S.Y., Paik, W.K., and Lee, H.W. (1998) Methylesters of L-arginine and N-nitro-L-arginine induce nitric oxide synthase in Staphylococcus aureus. Biochem Biophys Res Commun 246: 431- 435.

Chung, M.C., Narayanan, A., Popova, T.G., Kashanchi, F., Bailey, C.L., and Popov, S.G. (2013) Bacillus anthracis-derived nitric oxide induces protein S-nitrosylation contributing to macrophage death. Biochem Biophys Res Commun 430: 125- 130.

Clauditz, A., Resch, A., Wieland, K.P., Peschel, A., and Gotz, F. (2006) Staphyloxanthin plays a role in the fitness of Staphylococcus aureus and its ability to cope with oxidative stress. Infect Immun 74: 4950-4953.

Clements, M.O., Watson, S.P., and Foster, S.J. (1999) Characterization of the major superoxide dismutase of Staphylococcus aureus and its role in starvation survival, stress resistance, and pathogenicity. J Bacteriol 181: 3898-3903.

Cohen, J.O., (1972) The staphylococci. Wiley-Interscience.

193

Collins, F.M., and Lascelles, J. (1962) The effect of growth conditions on oxidative and dehydrogenase activity in Staphylococcus aureus. J Gen Microbiol 29: 531-535.

Corker, H., and Poole, R.K. (2003) Nitric oxide formation by Escherichia coli. Dependence on nitrite reductase, the NO-sensing regulator Fnr, and flavohemoglobin Hmp. J Biol Chem 278: 31584-31592.

Cosgrove, K., Coutts, G., Jonsson, I.M., Tarkowski, A., Kokai-Kun, J.F., Mond, J.J., and Foster, S.J. (2007) Catalase (KatA) and alkyl hydroperoxide reductase (AhpC) have compensatory roles in peroxide stress resistance and are required for survival, persistence, and nasal colonization in Staphylococcus aureus. J Bacteriol 189: 1025-1035.

Cowan, S.T., Shaw, C., and Williams, R.E.O. (1954) Type Strain for Staphylococcus aureus Rosenbach. Microbiology 10: 174-176.

Crack, J.C., Smith, L.J., Stapleton, M.R., Peck, J., Watmough, N.J., Buttner, M.J., Buxton, R.S., Green, J., Oganesyan, V.S., Thomson, A.J., and Le Brun, N.E. (2011) Mechanistic insight into the nitrosylation of the [4Fe-4S] cluster of WhiB- like proteins. J Am Chem Soc 133: 1112-1121.

Crane, B.R., Sudhamsu, J., and Patel, B.A. (2010) Bacterial nitric oxide synthases. Annu Rev Biochem 79: 445-470.

Crawford, N.M. (2006) Mechanisms for nitric oxide synthesis in plants. J Exp Bot 57: 471-478.

Czapski, G., and Goldstein, S. (1995) The role of the reactions of .NO with superoxide and oxygen in biological systems: a kinetic approach. Free Radic Biol Med 19: 785-794.

Czyz, A., and Wegrzyn, G. (2005) The Obg subfamily of bacterial GTP-binding proteins: essential proteins of largely unknown functions that are evolutionarily conserved from bacteria to humans. Acta Biochim Pol 52: 35-43.

Dantes, R., Mu, Y., Belflower, R., Aragon, D., Dumyati, G., Harrison, L.H., Lessa, F.C., Lynfield, R., Nadle, J., Petit, S., Ray, S.M., Schaffner, W., Townes, J., and Fridkin, S. (2013) National burden of invasive methicillin-resistant Staphylococcus aureus infections, United States, 2011. JAMA Intern Med 173: 1970-1978. de Haas, C.J., Veldkamp, K.E., Peschel, A., Weerkamp, F., Van Wamel, W.J., Heezius, E.C., Poppelier, M.J., Van Kessel, K.P., and van Strijp, J.A. (2004) Chemotaxis inhibitory protein of Staphylococcus aureus, a bacterial antiinflammatory agent. J Exp Med 199: 687-695.

194

de Niederhausern, S., Bondi, M., Messi, P., Iseppi, R., Sabia, C., Manicardi, G., and Anacarso, I. (2011) Vancomycin-resistance transferability from VanA enterococci to Staphylococcus aureus. Curr Microbiol 62: 1363-1367.

Deisenhofer, J. (1981) Crystallographic refinement and atomic models of a human Fc fragment and its complex with fragment B of protein A from Staphylococcus aureus at 2.9- and 2.8-A resolution. Biochemistry 20: 2361-2370.

Deivanayagam, C.C., Wann, E.R., Chen, W., Carson, M., Rajashankar, K.R., Hook, M., and Narayana, S.V. (2002) A novel variant of the immunoglobulin fold in surface adhesins of Staphylococcus aureus: crystal structure of the fibrinogen-binding MSCRAMM, clumping factor A. Embo j 21: 6660-6672.

DeLeo, F.R., Otto, M., Kreiswirth, B.N., and Chambers, H.F. (2010) Community- associated meticillin-resistant Staphylococcus aureus. Lancet 375: 1557-1568.

Derbyshire, E.R., and Marletta, M.A. (2012) Structure and regulation of soluble guanylate cyclase. Annu Rev Biochem 81: 533-559.

Deurenberg, R.H., and Stobberingh, E.E. (2008) The evolution of Staphylococcus aureus. Infect Genet Evol 8: 747-763.

Di Lullo, G.A., Sweeney, S.M., Korkko, J., Ala-Kokko, L., and San Antonio, J.D. (2002) Mapping the ligand-binding sites and disease-associated mutations on the most abundant protein in the human, type I collagen. J Biol Chem 277: 4223-4231.

Diep, B.A., Gill, S.R., Chang, R.F., Phan, T.H., Chen, J.H., Davidson, M.G., Lin, F., Lin, J., Carleton, H.A., Mongodin, E.F., Sensabaugh, G.F., and Perdreau-Remington, F. (2006) Complete genome sequence of USA300, an epidemic clone of community-acquired meticillin-resistant Staphylococcus aureus. Lancet 367: 731- 739.

Dinges, M.M., Orwin, P.M., and Schlievert, P.M. (2000) Exotoxins of Staphylococcus aureus. Clinical Microbiology Reviews 13: 16-34.

Durand, S., Braun, F., Lioliou, E., Romilly, C., Helfer, A.C., Kuhn, L., Quittot, N., Nicolas, P., Romby, P., and Condon, C. (2015) A nitric oxide regulated small RNA controls expression of genes involved in redox homeostasis in Bacillus subtilis. PLoS Genet 11: e1004957.

Dzal, Y.A., Jenkin, S.E., Lague, S.L., Reichert, M.N., York, J.M., and Pamenter, M.E. (2015) Oxygen in demand: How oxygen has shaped vertebrate physiology. Comp Biochem Physiol A Mol Integr Physiol 186: 4-26.

195

East, A.K., and Dyke, K. (1989) Cloning and sequence determination of six Staphylococcus aureus β-lactamases and their expression in Escherichia coli and Staphylococcus aureus. Microbiology 135: 1001-1015.

Eckmann, C., and Dryden, M. (2010) Treatment of complicated skin and soft-tissue infections caused by resistant bacteria: value of linezolid, tigecycline, daptomycin and vancomycin. Eur J Med Res 15: 554-563.

Efremov, R.G., and Sazanov, L.A. (2011) Structure of the membrane domain of respiratory complex I. Nature 476: 414-420.

Emmett, M., and Kloos, W.E. (1975) Amino acid requirements of staphylococci isolated from human skin. Can J Microbiol 21: 729-733.

Entenza, J.M., Foster, T.J., Ni Eidhin, D., Vaudaux, P., Francioli, P., and Moreillon, P. (2000) Contribution of clumping factor B to pathogenesis of experimental endocarditis due to Staphylococcus aureus. Infect Immun 68: 5443-5446.

Falkowski, P.G., and Godfrey, L.V. (2008) Electrons, life and the evolution of Earth's oxygen cycle. Philos Trans R Soc Lond B Biol Sci 363: 2705-2716.

Fast, B., Lindgren, P., and Gotz, F. (1996) Cloning, sequencing, and characterization of a gene (narT) encoding a transport protein involved in dissimilatory nitrate reduction in Staphylococcus carnosus. Arch Microbiol 166: 361-367.

Feng, Y., Li, W., Li, J., Wang, J., Ge, J., Xu, D., Liu, Y., Wu, K., Zeng, Q., Wu, J.W., Tian, C., Zhou, B., and Yang, M. (2012) Structural insight into the type-II mitochondrial NADH dehydrogenases. Nature 491: 478-482.

Ferreira, M.T., Manso, A.S., Gaspar, P., Pinho, M.G., and Neves, A.R. (2013) Effect of oxygen on glucose metabolism: utilization of lactate in Staphylococcus aureus as revealed by in vivo NMR studies. PLoS One 8: e58277.

Fey, P.D., Endres, J.L., Yajjala, V.K., Widhelm, T.J., Boissy, R.J., Bose, J.L., and Bayles, K.W. (2013) A genetic resource for rapid and comprehensive phenotype screening of nonessential Staphylococcus aureus genes. mBio 4: e00537-00512.

Finocchietto, P.V., Franco, M.C., Holod, S., Gonzalez, A.S., Converso, D.P., Antico Arciuch, V.G., Serra, M.P., Poderoso, J.J., and Carreras, M.C. (2009) Mitochondrial nitric oxide synthase: a masterpiece of metabolic adaptation, cell growth, transformation, and death. Exp Biol Med (Maywood) 234: 1020-1028.

Firoved, A.M., Wood, S.R., Ornatowski, W., Deretic, V., and Timmins, G.S. (2004) Microarray analysis and functional characterization of the nitrosative stress response in nonmucoid and mucoid Pseudomonas aeruginosa. J Bacteriol 186: 4046-4050.

196

Flatley, J., Barrett, J., Pullan, S.T., Hughes, M.N., Green, J., and Poole, R.K. (2005) Transcriptional responses of Escherichia coli to S-nitrosoglutathione under defined chemostat conditions reveal major changes in methionine biosynthesis. J Biol Chem 280: 10065-10072.

Flint, D.H., Tuminello, J.F., and Emptage, M.H. (1993) The inactivation of Fe-S cluster containing hydro-lyases by superoxide. J Biol Chem 268: 22369-22376.

Flowers, R.S., Martin, S.E., Brewer, D.G., and Ordal, Z.J. (1977) Catalase and enumeration of stressed Staphylococcus aureus cells. Appl Environ Microbiol 33: 1112-1117.

Follmann, M., Griebenow, N., Hahn, M.G., Hartung, I., Mais, F.J., Mittendorf, J., Schafer, M., Schirok, H., Stasch, J.P., Stoll, F., and Straub, A. (2013) The chemistry and biology of soluble guanylate cyclase stimulators and activators. Angew Chem Int Ed Engl 52: 9442-9462.

Fonner, B.A., Tripet, B.P., Eilers, B.J., Stanisich, J., Sullivan-Springhetti, R.K., Moore, R., Liu, M., Lei, B., and Copie, V. (2014) Solution structure and molecular determinants of hemoglobin binding of the first NEAT domain of IsdB in Staphylococcus aureus. Biochemistry 53: 3922-3933.

Forstermann, U., and Sessa, W.C. (2012) Nitric oxide synthases: regulation and function. Eur Heart J 33: 829-837, 837a-837d.

Foti, J.J., Schienda, J., Sutera, V.A., Jr., and Lovett, S.T. (2005) A bacterial G protein- mediated response to replication arrest. Mol Cell 17: 549-560.

Fraser, J., Arcus, V., Kong, P., Baker, E., and Proft, T. (2000) Superantigens - powerful modifiers of the immune system. Mol Med Today 6: 125-132.

Fraser, J.D., and Proft, T. (2008) The bacterial superantigen and superantigen-like proteins. Immunol Rev 225: 226-243.

Fuchs, S., Pane-Farre, J., Kohler, C., Hecker, M., and Engelmann, S. (2007) Anaerobic gene expression in Staphylococcus aureus. Journal of bacteriology 189: 4275- 4289.

Fuller, J.R., Vitko, N.P., Perkowski, E.F., Scott, E., Khatri, D., Spontak, J.S., Thurlow, L.R., and Richardson, A.R. (2011) Identification of a lactate-quinone oxidoreductase in Staphylococcus aureus that is essential for virulence. Front Cell Infect Microbiol 1: 19.

Gardner, P.R. (2002) Aconitase: sensitive target and measure of superoxide. Methods in enzymology 349: 9-23.

197

Gardner, P.R., and Fridovich, I. (1991a) Superoxide sensitivity of the Escherichia coli 6- phosphogluconate dehydratase. J Biol Chem 266: 1478-1483.

Gardner, P.R., and Fridovich, I. (1991b) Superoxide sensitivity of the Escherichia coli aconitase. The Journal of biological chemistry 266: 19328-19333.

Gardner, P.R., and Fridovich, I. (1992) Inactivation-reactivation of aconitase in Escherichia coli. A sensitive measure of superoxide radical. J Biol Chem 267: 8757-8763.

Gardner, P.R., Gardner, A.M., Martin, L.A., and Salzman, A.L. (1998) Nitric oxide dioxygenase: an enzymic function for flavohemoglobin. Proc Natl Acad Sci U S A 95: 10378-10383.

Gaudin, C.F., Grigg, J.C., Arrieta, A.L., and Murphy, M.E. (2011) Unique heme-iron coordination by the hemoglobin receptor IsdB of Staphylococcus aureus. Biochemistry 50: 5443-5452.

Gaupp, R., Schlag, S., Liebeke, M., Lalk, M., and Gotz, F. (2010) Advantage of upregulation of succinate dehydrogenase in Staphylococcus aureus biofilms. J Bacteriol 192: 2385-2394.

Gautier, C., Mikula, I., Nioche, P., Martasek, P., Raman, C.S., and Slama-Schwok, A. (2006) Dynamics of NO rebinding to the heme domain of NO synthase-like proteins from bacterial pathogens. Nitric Oxide 15: 312-327.

Geary, L.E., and Meister, A. (1977) On the mechanism of glutamine-dependent reductive amination of alpha-ketoglutarate catalyzed by glutamate synthase. J Biol Chem 252: 3501-3508.

Geissmann, Q. (2013) OpenCFU, a new free and open-source software to count cell colonies and other circular objects. PLoS One 8: e54072.

Gerschman, R., Gilbert, D., Nye, S.W., Dwyer, P., and Fenn, W.O. (2001) Oxygen poisoning and X-irradiation: a mechanism in common. 1954. Nutrition 17: 162.

Ghosh, D.K., Wu, C., Pitters, E., Moloney, M., Werner, E.R., Mayer, B., and Stuehr, D.J. (1997) Characterization of the inducible nitric oxide synthase oxygenase domain identifies a 49 amino acid segment required for subunit dimerization and tetrahydrobiopterin interaction. Biochemistry 36: 10609-10619.

Gibbons, N.E., and Murray, R.G.E. (1978) Proposals Concerning the Higher Taxa of Bacteria. International Journal of Systematic and Evolutionary Microbiology 28: 1-6.

198

Gilberthorpe, N.J., and Poole, R.K. (2008) Nitric oxide homeostasis in Salmonella typhimurium: roles of respiratory nitrate reductase and flavohemoglobin. J Biol Chem 283: 11146-11154.

Gill, S.R., Fouts, D.E., Archer, G.L., Mongodin, E.F., Deboy, R.T., Ravel, J., Paulsen, I.T., Kolonay, J.F., Brinkac, L., Beanan, M., Dodson, R.J., Daugherty, S.C., Madupu, R., Angiuoli, S.V., Durkin, A.S., Haft, D.H., Vamathevan, J., Khouri, H., Utterback, T., Lee, C., Dimitrov, G., Jiang, L., Qin, H., Weidman, J., Tran, K., Kang, K., Hance, I.R., Nelson, K.E., and Fraser, C.M. (2005) Insights on evolution of virulence and resistance from the complete genome analysis of an early methicillin-resistant Staphylococcus aureus strain and a biofilm-producing methicillin-resistant Staphylococcus epidermidis strain. J Bacteriol 187: 2426- 2438.

Gillaspy, A.F., Hickmon, S.G., Skinner, R.A., Thomas, J.R., Nelson, C.L., and Smeltzer, M.S. (1995) Role of the accessory gene regulator (agr) in pathogenesis of staphylococcal osteomyelitis. Infect Immun 63: 3373-3380.

Giuffre, A., Borisov, V.B., Mastronicola, D., Sarti, P., and Forte, E. (2012) Cytochrome bd oxidase and nitric oxide: from reaction mechanisms to bacterial physiology. FEBS Lett 586: 622-629.

Giulivi, C. (2003) Characterization and function of mitochondrial nitric-oxide synthase. Free Radic Biol Med 34: 397-408.

Giulivi, C., Kato, K., and Cooper, C.E. (2006) Nitric oxide regulation of mitochondrial oxygen consumption I: cellular physiology. American journal of physiology. Cell physiology 291: C1225-1231.

Goldstein, S., and Czapski, G. (2000) Reactivity of peroxynitrite versus simultaneous - generation of NO and O2 toward NADH. Chemical research in toxicology 13: 736-741.

Goncalves, V.L., Nobre, L.S., Vicente, J.B., Teixeira, M., and Saraiva, L.M. (2006) Flavohemoglobin requires microaerophilic conditions for nitrosative protection of Staphylococcus aureus. FEBS Lett 580: 1817-1821.

Gong, J., Li, D., Yan, J., Liu, Y., Dong, J., Gao, Y., Sun, T., and Yang, G. (2014) The accessory gene regulator (agr) controls Staphylococcus aureus virulence in a murine intracranial abscesses model. Braz J Infect Dis 18: 501-506.

Gonzalez-Zorn, B., and Courvalin, P., (2003) VanA-mediated high level glycopeptide resistance in MRSA. In: Lancet Infect Dis. United States, pp. 67-68.

Gopal, T., Nagarajan, V., and Elasri, M.O. (2015) SATRAT: Staphylococcus aureus transcript regulatory network analysis tool. PeerJ 3: e717.

199

Gorwitz, R.J., Kruszon-Moran, D., McAllister, S.K., McQuillan, G., McDougal, L.K., Fosheim, G.E., Jensen, B.J., Killgore, G., Tenover, F.C., and Kuehnert, M.J. (2008) Changes in the prevalence of nasal colonization with Staphylococcus aureus in the United States, 2001-2004. J Infect Dis 197: 1226-1234.

Gotz, F., and Mayer, S. (2013) Both terminal oxidases contribute to fitness and virulence during organ-specific Staphylococcus aureus colonization. MBio 4: e00976-00913.

Gouda, H., Shiraishi, M., Takahashi, H., Kato, K., Torigoe, H., Arata, Y., and Shimada, I. (1998) NMR study of the interaction between the B domain of staphylococcal protein A and the Fc portion of immunoglobulin G. Biochemistry 37: 129-136.

Grice, E.A., Kong, H.H., Conlan, S., Deming, C.B., Davis, J., Young, A.C., Bouffard, G.G., Blakesley, R.W., Murray, P.R., Green, E.D., Turner, M.L., and Segre, J.A. (2009) Topographical and temporal diversity of the human skin microbiome. Science 324: 1190-1192.

Gries, C.M., Bose, J.L., Nuxoll, A.S., Fey, P.D., and Bayles, K.W. (2013) The Ktr potassium transport system in Staphylococcus aureus and its role in cell physiology, antimicrobial resistance and pathogenesis. Mol Microbiol 89: 760- 773.

Griffith, J.W., Sokol, C.L., and Luster, A.D. (2014) Chemokines and chemokine receptors: positioning cells for host defense and immunity. Annu Rev Immunol 32: 659-702.

Griffith, O.W., and Stuehr, D.J. (1995) Nitric oxide synthases: properties and catalytic mechanism. Annu Rev Physiol 57: 707-736.

Grigg, J.C., Vermeiren, C.L., Heinrichs, D.E., and Murphy, M.E. (2007) Haem recognition by a Staphylococcus aureus NEAT domain. Mol Microbiol 63: 139- 149.

Grinblat, L., Sreider, C.M., and Stoppani, A.O. (1991) Superoxide anion production by lipoamide dehydrogenase redox-cycling: effect of enzyme modifiers. Biochem Int 23: 83-92.

Grosser, M.R., Weiss, A., Shaw, L.N., and Richardson, A.R. (2016) Regulatory Requirements for Staphylococcus aureus nitric oxide resistance. J Bacteriol 198: 2043-2055.

Guillet, V., Keller, D., Prevost, G., and Mourey, L. (2004) Crystallization and preliminary crystallographic data of a leucotoxin S component from Staphylococcus aureus. Acta Crystallogr D Biol Crystallogr 60: 310-313.

200

Gusarov, I., Gautier, L., Smolentseva, O., Shamovsky, I., Eremina, S., Mironov, A., and Nudler, E. (2013) Bacterial nitric oxide extends the lifespan of C. elegans. Cell 152: 818-830.

Gusarov, I., and Nudler, E. (2005) NO-mediated cytoprotection: instant adaptation to oxidative stress in bacteria. Proc Natl Acad Sci U S A 102: 13855-13860.

Gusarov, I., and Nudler, E. (2012) S-nitrosylation signaling in Escherichia coli. Sci Signal 5: pe26.

Gusarov, I., Shatalin, K., Starodubtseva, M., and Nudler, E. (2009) Endogenous nitric oxide protects bacteria against a wide spectrum of antibiotics. Science 325: 1380-1384.

Gusarov, I., Starodubtseva, M., Wang, Z.Q., McQuade, L., Lippard, S.J., Stuehr, D.J., and Nudler, E. (2008) Bacterial nitric-oxide synthases operate without a dedicated redox partner. J Biol Chem 283: 13140-13147.

Haggar, A., Ehrnfelt, C., Holgersson, J., and Flock, J.-I. (2004) The Extracellular Adherence Protein from Staphylococcus aureus Inhibits Neutrophil Binding to Endothelial Cells.

Halper, J., and Kjaer, M. (2014) Basic components of connective tissues and extracellular matrix: elastin, fibrillin, fibulins, fibrinogen, fibronectin, laminin, tenascins and thrombospondins. Adv Exp Med Biol 802: 31-47.

Hammer, N.D., Reniere, M.L., Cassat, J.E., Zhang, Y., Hirsch, A.O., Indriati Hood, M., and Skaar, E.P. (2013) Two heme-dependent terminal oxidases power Staphylococcus aureus organ-specific colonization of the vertebrate host. MBio 4.

Hammer, N.D., and Skaar, E.P. (2011) Molecular mechanisms of Staphylococcus aureus iron acquisition. Annu Rev Microbiol 65: 129-147.

Hanahan, D. (1983) Studies on transformation of Escherichia coli with plasmids. J Mol Biol 166: 557-580.

Handke, L.D., Shivers, R.P., and Sonenshein, A.L. (2008) Interaction of Bacillus subtilis CodY with GTP. J Bacteriol 190: 798-806.

Hartleib, J., Kohler, N., Dickinson, R.B., Chhatwal, G.S., Sixma, J.J., Hartford, O.M., Foster, T.J., Peters, G., Kehrel, B.E., and Herrmann, M. (2000) Protein A is the von Willebrand factor binding protein on Staphylococcus aureus. Blood 96: 2149- 2156.

201

Hartman, B.J., and Tomasz, A. (1984) Low-affinity penicillin-binding protein associated with beta-lactam resistance in Staphylococcus aureus. J Bacteriol 158: 513-516.

Hartmann, T., Zhang, B., Baronian, G., Schulthess, B., Homerova, D., Grubmuller, S., Kutzner, E., Gaupp, R., Bertram, R., Powers, R., Eisenreich, W., Kormanec, J., Herrmann, M., Molle, V., Somerville, G.A., and Bischoff, M. (2013) Catabolite control protein E (CcpE) is a LysR-type transcriptional regulator of tricarboxylic acid cycle activity in Staphylococcus aureus. J Biol Chem 288: 36116-36128.

Healy, F.G., Wach, M., Krasnoff, S.B., Gibson, D.M., and Loria, R. (2000) The txtAB genes of the plant pathogen Streptomyces acidiscabies encode a peptide synthetase required for phytotoxin thaxtomin A production and pathogenicity. Mol Microbiol 38: 794-804.

Henry, Y.A. (2015) EPR Characterization of Nitric Oxide Binding to Hemoglobin. 61-86.

Hillson, J.L., Karr, N.S., Oppliger, I.R., Mannik, M., and Sasso, E.H. (1993) The structural basis of germline-encoded VH3 immunoglobulin binding to staphylococcal protein A. J Exp Med 178: 331-336.

Hochgrafe, F., Wolf, C., Fuchs, S., Liebeke, M., Lalk, M., Engelmann, S., and Hecker, M. (2008) Nitric oxide stress induces different responses but mediates comparable protein thiol protection in Bacillus subtilis and Staphylococcus aureus. J Bacteriol 190: 4997-5008.

Holden, J.K., Dejam, D., Lewis, M.C., Huang, H., Kang, S., Jing, Q., Xue, F., Silverman, R.B., and Poulos, T.L. (2015a) Inhibitor bound crystal structures of bacterial nitric oxide synthase. Biochemistry 54: 4075-4082.

Holden, J.K., Kang, S., Beasley, F.C., Cinelli, M.A., Li, H., Roy, S.G., Dejam, D., Edinger, A.L., Nizet, V., Silverman, R.B., and Poulos, T.L. (2015b) Nitric oxide synthase as a target for methicillin-resistant Staphylococcus aureus. Chem Biol 22: 785-792.

Holden, J.K., Kang, S., Hollingsworth, S.A., Li, H., Lim, N., Chen, S., Huang, H., Xue, F., Tang, W., Silverman, R.B., and Poulos, T.L. (2015c) Structure-based design of bacterial nitric oxide synthase inhibitors. J Med Chem.

Holden, J.K., Lewis, M.C., Cinelli, M.A., Abdullatif, Z., Pensa, A.V., Silverman, R.B., and Poulos, T.L. (2016) Targeting bacterial nitric oxide synthase with aminoquinoline- based inhibitors. Biochemistry.

Holden, J.K., Li, H., Jing, Q., Kang, S., Richo, J., Silverman, R.B., and Poulos, T.L. (2013) Structural and biological studies on bacterial nitric oxide synthase inhibitors. Proc Natl Acad Sci U S A 110: 18127-18131.

202

Holden, J.K., Lim, N., and Poulos, T.L. (2014) Identification of redox partners and development of a novel chimeric bacterial nitric oxide synthase for structure activity analyses. J Biol Chem 289: 29437-29445.

Holden, M.T., Feil, E.J., Lindsay, J.A., Peacock, S.J., Day, N.P., Enright, M.C., Foster, T.J., Moore, C.E., Hurst, L., Atkin, R., Barron, A., Bason, N., Bentley, S.D., Chillingworth, C., Chillingworth, T., Churcher, C., Clark, L., Corton, C., Cronin, A., Doggett, J., Dowd, L., Feltwell, T., Hance, Z., Harris, B., Hauser, H., Holroyd, S., Jagels, K., James, K.D., Lennard, N., Line, A., Mayes, R., Moule, S., Mungall, K., Ormond, D., Quail, M.A., Rabbinowitsch, E., Rutherford, K., Sanders, M., Sharp, S., Simmonds, M., Stevens, K., Whitehead, S., Barrell, B.G., Spratt, B.G., and Parkhill, J. (2004) Complete genomes of two clinical Staphylococcus aureus strains: evidence for the rapid evolution of virulence and drug resistance. Proc Natl Acad Sci U S A 101: 9786-9791.

Holmgren, A. (2000) Antioxidant function of thioredoxin and glutaredoxin systems. Antioxid Redox Signal 2: 811-820.

Holtfreter, S., and Broker, B.M. (2005) Staphylococcal superantigens: do they play a role in sepsis? Arch Immunol Ther Exp (Warsz) 53: 13-27.

Hong, I.S., Kim, Y.K., Choi, W.S., Seo, D.W., Yoon, J.W., Han, J.W., Lee, H.Y., and Lee, H.W. (2003) Purification and characterization of nitric oxide synthase from Staphylococcus aureus. FEMS Microbiol Lett 222: 177-182.

Horsburgh, M.J., Clements, M.O., Crossley, H., Ingham, E., and Foster, S.J. (2001a) PerR controls oxidative stress resistance and iron storage proteins and is required for virulence in Staphylococcus aureus. Infect Immun 69: 3744-3754.

Horsburgh, M.J., Ingham, E., and Foster, S.J. (2001b) In Staphylococcus aureus, fur is an interactive regulator with PerR, contributes to virulence, and Is necessary for oxidative stress resistance through positive regulation of catalase and iron homeostasis. J Bacteriol 183: 468-475.

Howden, B.P., Davies, J.K., Johnson, P.D., Stinear, T.P., and Grayson, M.L. (2010) Reduced vancomycin susceptibility in Staphylococcus aureus, including vancomycin-intermediate and heterogeneous vancomycin-intermediate strains: resistance mechanisms, laboratory detection, and clinical implications. Clin Microbiol Rev 23: 99-139.

Hromatka, B.S., Noble, S.M., and Johnson, A.D. (2005) Transcriptional response of Candida albicans to nitric oxide and the role of the YHB1 gene in nitrosative stress and virulence. Mol Biol Cell 16: 4814-4826.

203

Hsia, C.C., Schmitz, A., Lambertz, M., Perry, S.F., and Maina, J.N. (2013) Evolution of air breathing: oxygen homeostasis and the transitions from water to land and sky. Compr Physiol 3: 849-915.

Hughes, M.N. (1999) Relationships between nitric oxide, nitroxyl ion, nitrosonium cation and peroxynitrite. 1411: 263-272.

Huie, R.E., and Padmaja, S. (1993) The reaction of NO with superoxide. Free radical research communications 18: 195-199.

Huntzinger, E., Boisset, S., Saveanu, C., Benito, Y., Geissmann, T., Namane, A., Lina, G., Etienne, J., Ehresmann, B., Ehresmann, C., Jacquier, A., Vandenesch, F., and Romby, P. (2005) Staphylococcus aureus RNAIII and the endoribonuclease III coordinately regulate spa gene expression. The EMBO Journal 24: 824-835.

Hurshman, A.R., Krebs, C., Edmondson, D.E., Huynh, B.H., and Marletta, M.A. (1999) Formation of a pterin radical in the reaction of the heme domain of inducible nitric oxide synthase with oxygen. Biochemistry 38: 15689-15696.

Husain, M., Bourret, T.J., McCollister, B.D., Jones-Carson, J., Laughlin, J., and Vazquez-Torres, A. (2008) Nitric oxide evokes an adaptive response to oxidative stress by arresting respiration. The Journal of biological chemistry 283: 7682- 7689.

Imlay, J.A. (2003) Pathways of oxidative damage. Annu Rev Microbiol 57: 395-418.

Imlay, J.A. (2013) The molecular mechanisms and physiological consequences of oxidative stress: lessons from a model bacterium. Nat Rev Microbiol 11: 443-454.

Inoue, S., and Kawanishi, S. (1995) Oxidative DNA damage induced by simultaneous generation of nitric oxide and superoxide. FEBS letters 371: 86-88.

Ischiropoulos, H., Zhu, L., and Beckman, J.S. (1992a) Peroxinitrite formation from macrophage-derived nitric-oxide. Archives of Biochemistry and Biophysics 298: 446-451.

Ischiropoulos, H., Zhu, L., Chen, J., Tsai, M., Martin, J.C., Smith, C.D., and Beckman, J.S. (1992b) Peroxynitrite-mediated tyrosine nitration catalyzed by superoxide dismutase. Archives of biochemistry and biophysics 298: 431-437.

Itoh, S., Hamada, E., Kamoshida, G., Yokoyama, R., Takii, T., Onozaki, K., and Tsuji, T. (2010) Staphylococcal superantigen-like protein 10 (SSL10) binds to human immunoglobulin G (IgG) and inhibits complement activation via the classical pathway. Mol Immunol 47: 932-938.

204

Jakubowski, W., and Bartosz, G. (2000) 2,7-dichlorofluorescin oxidation and reactive oxygen species: what does it measure? Cell biology international 24: 757-760.

Jeandroz, S., Wipf, D., Stuehr, D.J., Lamattina, L., Melkonian, M., Tian, Z., Zhu, Y., Carpenter, E.J., Wong, G.K., and Wendehenne, D. (2016) Occurrence, structure, and evolution of nitric oxide synthase-like proteins in the plant kingdom. Sci Signal 9: re2.

Jett, B.D., Hatter, K.L., Huycke, M.M., and Gilmore, M.S. (1997) Simplified agar plate method for quantifying viable bacteria. Biotechniques 23: 648-650.

Ji, G., Beavis, R., and Novick, R.P. (1997) Bacterial interference caused by autoinducing peptide variants. Science 276: 2027-2030.

Ji, G., Beavis, R.C., and Novick, R.P. (1995) Cell density control of staphylococcal virulence mediated by an octapeptide pheromone. Proceedings of the National Academy of Sciences of the United States of America 92: 12055-12059.

Jiang, J., Jordan, S.J., Barr, D.P., Gunther, M.R., Maeda, H., and Mason, R.P. (1997) In vivo production of nitric oxide in rats after administration of hydroxyurea. Mol Pharmacol 52: 1081-1086.

Johnson, E.G., Sparks, J.P., Dzikovski, B., Crane, B.R., Gibson, D.M., and Loria, R. (2008) Plant-pathogenic Streptomyces species produce nitric oxide synthase- derived nitric oxide in response to host signals. Chem Biol 15: 43-50.

Jonsson, T.J., Ellis, H.R., and Poole, L.B. (2007) Cysteine reactivity and thiol-disulfide interchange pathways in AhpF and AhpC of the bacterial alkyl hydroperoxide reductase system. Biochemistry 46: 5709-5721.

Juarez, O., and Barquera, B. (2012) Insights into the mechanism of electron transfer and sodium translocation of the Na(+)-pumping NADH:quinone oxidoreductase. Biochim Biophys Acta 1817: 1823-1832.

Junemann, S., and Wrigglesworth, J.M. (1996) Binding of NO to the oxygen reaction site of cytochrome bd from Azotobacter vinelandii. Biochemical Society transactions 24: 38s.

Justino, M.C., Vicente, J.B., Teixeira, M., and Saraiva, L.M. (2005) New genes implicated in the protection of anaerobically grown Escherichia coli against nitric oxide. J Biol Chem 280: 2636-2643.

Kaiser, J.C., Omer, S., Sheldon, J.R., Welch, I., and Heinrichs, D.E. (2015) Role of BrnQ1 and BrnQ2 in branched-chain amino acid transport and virulence in Staphylococcus aureus. Infect Immun 83: 1019-1029.

205

Kaiser, J.C., Sen, S., Sinha, A., Wilkinson, B.J., and Heinrichs, D.E. (2016) The role of two branched-chain amino acid transporters in Staphylococcus aureus growth, membrane fatty acid composition and virulence. Mol Microbiol.

Kaneko, J., and Kamio, Y. (2004) Bacterial two-component and hetero-heptameric pore- forming cytolytic toxins: structures, pore-forming mechanism, and organization of the genes. Biosci Biotechnol Biochem 68: 981-1003.

Karavolos, M.H., Horsburgh, M.J., Ingham, E., and Foster, S.J. (2003) Role and regulation of the superoxide dismutases of Staphylococcus aureus. Microbiology 149: 2749-2758.

Karlsson-Kanth, A., Tegmark-Wisell, K., Arvidson, S., and Oscarsson, J. (2006) Natural human isolates of Staphylococcus aureus selected for high production of proteases and alpha-hemolysin are sigmaB deficient. Int J Med Microbiol 296: 229-236.

Katayama, Y., Ito, T., and Hiramatsu, K. (2000) A new class of genetic element, staphylococcus cassette chromosome mec, encodes methicillin resistance in Staphylococcus aureus. Antimicrob Agents Chemother 44: 1549-1555.

Keane, F.M., Clarke, A.W., Foster, T.J., and Weiss, A.S. (2007a) The N-terminal A domain of Staphylococcus aureus fibronectin-binding protein A binds to tropoelastin. Biochemistry 46: 7226-7232.

Keane, F.M., Loughman, A., Valtulina, V., Brennan, M., Speziale, P., and Foster, T.J. (2007b) Fibrinogen and elastin bind to the same region within the A domain of fibronectin binding protein A, an MSCRAMM of Staphylococcus aureus. Mol Microbiol 63: 711-723.

Kemper, C., and Atkinson, J.P. (2007) T-cell regulation: with complements from innate immunity. Nat Rev Immunol 7: 9-18.

Kers, J.A., Wach, M.J., Krasnoff, S.B., Widom, J., Cameron, K.D., Bukhalid, R.A., Gibson, D.M., Crane, B.R., and Loria, R. (2004) Nitration of a peptide phytotoxin by bacterial nitric oxide synthase. Nature 429: 79-82.

Kerscher, S., Drose, S., Zickermann, V., and Brandt, U. (2008) The three families of respiratory NADH dehydrogenases. Results Probl Cell Differ 45: 185-222.

Keshive, M., Singh, S., Wishnok, J.S., Tannenbaum, S.R., and Deen, W.M. (1996) Kinetics of S-nitrosation of thiols in nitric oxide solutions. Chem Res Toxicol 9: 988-993.

Keyer, K., and Imlay, J.A. (1996) Superoxide accelerates DNA damage by elevating free-iron levels. Proc Natl Acad Sci U S A 93: 13635-13640.

206

Kielian, T., Cheung, A., and Hickey, W.F. (2001) Diminished virulence of an alpha-toxin mutant of Staphylococcus aureus in experimental brain abscesses. Infect Immun 69: 6902-6911.

Kim, M.H., Granick, J.L., Kwok, C., Walker, N.J., Borjesson, D.L., Curry, F.R., Miller, L.S., and Simon, S.I. (2011) Neutrophil survival and c-kit(+)-progenitor proliferation in Staphylococcus aureus-infected skin wounds promote resolution. Blood 117: 3343-3352.

Kim, N.H., Kang, Y.M., Han, W.D., Park, K.U., Park, K.H., Yoo, J.I., Lee, D.G., Park, C., Song, K.H., Kim, E.S., Park, S.W., Kim, N.J., Oh, M.D., and Kim, H.B. (2016) Small-colony variants in persistent and recurrent Staphylococcus aureus bacteremia. Microb Drug Resist 22: 538-544.

Kinkel, T.L., Roux, C.M., Dunman, P.M., and Fang, F.C. (2013) The Staphylococcus aureus SrrAB two-component system promotes resistance to nitrosative stress and hypoxia. MBio 4: e00696-00613.

Kintarak, S., Whawell, S.A., Speight, P.M., Packer, S., and Nair, S.P. (2004) Internalization of Staphylococcus aureus by human keratinocytes. Infect Immun 72: 5668-5675.

Kluytmans, J., van Belkum, A., and Verbrugh, H. (1997) Nasal carriage of Staphylococcus aureus: epidemiology, underlying mechanisms, and associated risks. Clin Microbiol Rev 10: 505-520.

Knowles, R.G., and Moncada, S. (1994) Nitric oxide synthases in mammals. Biochemical Journal 298: 249-258.

Kohanski, M.A., Dwyer, D.J., Hayete, B., Lawrence, C.A., and Collins, J.J. (2007) A common mechanism of cellular death induced by bactericidal antibiotics. Cell 130: 797-810.

Kohler, C., von Eiff, C., Peters, G., Proctor, R.A., Hecker, M., and Engelmann, S. (2003) Physiological characterization of a heme-deficient mutant of Staphylococcus aureus by a proteomic approach. Journal of bacteriology 185: 6928-6937.

Kolar, S.L., Ibarra, J.A., Rivera, F.E., Mootz, J.M., Davenport, J.E., Stevens, S.M., Horswill, A.R., and Shaw, L.N. (2013) Extracellular proteases are key mediators of Staphylococcus aureus virulence via the global modulation of virulence- determinant stability. Microbiologyopen 2: 18-34.

Kong, K.-F., Vuong, C., and Otto, M. (2006) Staphylococcus quorum sensing in biofilm formation and infection. Quorum sensing in human pathogens 296: 133-139.

207

Korshunov, S., and Imlay, J.A. (2010) Two sources of endogenous hydrogen peroxide in Escherichia coli. Mol Microbiol 75: 1389-1401.

Koziel, J., and Potempa, J. (2013) Protease-armed bacteria in the skin. Cell Tissue Res 351: 325-337.

Kreiswirth, B.N., Lofdahl, S., Betley, M.J., O'Reilly, M., Schlievert, P.M., Bergdoll, M.S., and Novick, R.P. (1983) The toxic shock syndrome exotoxin structural gene is not detectably transmitted by a prophage. Nature 305: 709-712.

Kriegeskorte, A., Grubmuller, S., Huber, C., Kahl, B.C., von Eiff, C., Proctor, R.A., Peters, G., Eisenreich, W., and Becker, K. (2014) Staphylococcus aureus small colony variants show common metabolic features in central metabolism irrespective of the underlying auxotrophism. Front Cell Infect Microbiol 4: 141.

Krisko, A., and Radman, M. (2013) Biology of extreme radiation resistance: the way of Deinococcus radiodurans. Cold Spring Harb Perspect Biol 5.

Kristian, S.A., Lauth, X., Nizet, V., Goetz, F., Neumeister, B., Peschel, A., and Landmann, R. (2003) Alanylation of teichoic acids protects Staphylococcus aureus against Toll-like receptor 2-dependent host defense in a mouse tissue cage infection model. J Infect Dis 188: 414-423.

Kubica, M., Guzik, K., Koziel, J., Zarebski, M., Richter, W., Gajkowska, B., Golda, A., Maciag-Gudowska, A., Brix, K., Shaw, L., Foster, T., and Potempa, J. (2008) A potential new pathway for Staphylococcus aureus dissemination: the silent survival of S. aureus phagocytosed by human monocyte-derived macrophages. PLoS One 3: e1409.

Kuroda, M., Ohta, T., Uchiyama, I., Baba, T., Yuzawa, H., Kobayashi, I., Cui, L., Oguchi, A., Aoki, K., Nagai, Y., Lian, J., Ito, T., Kanamori, M., Matsumaru, H., Maruyama, A., Murakami, H., Hosoyama, A., Mizutani-Ui, Y., Takahashi, N.K., Sawano, T., Inoue, R., Kaito, C., Sekimizu, K., Hirakawa, H., Kuhara, S., Goto, S., Yabuzaki, J., Kanehisa, M., Yamashita, A., Oshima, K., Furuya, K., Yoshino, C., Shiba, T., Hattori, M., Ogasawara, N., Hayashi, H., and Hiramatsu, K. (2001) Whole genome sequencing of meticillin-resistant Staphylococcus aureus. Lancet 357: 1225-1240.

Kussmaul, L., and Hirst, J. (2006) The mechanism of superoxide production by NADH:ubiquinone oxidoreductase (complex I) from bovine heart mitochondria. Proc Natl Acad Sci U S A 103: 7607-7612.

Lacza, Z., Snipes, J.A., Zhang, J., Horvath, E.M., Figueroa, J.P., Szabo, C., and Busija, D.W. (2003) Mitochondrial nitric oxide synthase is not eNOS, nNOS or iNOS. Free radical biology & medicine 35: 1217-1228.

208

Lam, M.A., Pattison, D.I., Bottle, S.E., Keddie, D.J., and Davies, M.J. (2008) Nitric oxide and nitroxides can act as efficient scavengers of protein-derived free radicals. Chem Res Toxicol 21: 2111-2119.

Lancaster, J.R., Jr. (1997) A tutorial on the diffusibility and reactivity of free nitric oxide. Nitric Oxide 1: 18-30.

Larsen, F.J., Schiffer, T.A., Weitzberg, E., and Lundberg, J.O. (2012) Regulation of mitochondrial function and energetics by reactive nitrogen oxides. Free radical biology & medicine 53: 1919-1928.

Lascelles, J., and Burke, K.A. (1978) Reduction of ferric iron by L-lactate and DL- glycerol-3-phosphate in membrane preparations from Staphylococcus aureus and interactions with the nitrate reductase system. Journal of bacteriology 134: 585-589.

LeBel, C.P., Ischiropoulos, H., and Bondy, S.C. (1992) Evaluation of the probe 2',7'- dichlorofluorescin as an indicator of reactive oxygen species formation and oxidative stress. Chemical research in toxicology 5: 227-231.

Lee, D.S., Burd, H., Liu, J., Almaas, E., Wiest, O., Barabasi, A.L., Oltvai, Z.N., and Kapatral, V. (2009) Comparative genome-scale metabolic reconstruction and flux balance analysis of multiple Staphylococcus aureus genomes identify novel antimicrobial drug targets. Journal of bacteriology 191: 4015-4024.

Leibig, M., Liebeke, M., Mader, D., Lalk, M., Peschel, A., and Gotz, F. (2011) Pyruvate formate lyase acts as a formate supplier for metabolic processes during anaerobiosis in Staphylococcus aureus. J Bacteriol 193: 952-962.

Lewis, A.M., Matzdorf, S.S., Endres, J.L., Windham, I.H., Bayles, K.W., and Rice, K.C. (2015) Examination of the Staphylococcus aureus nitric oxide reductase (saNOR) reveals its contribution to modulating intracellular NO levels and cellular respiration. Molecular microbiology 96: 651-669.

Lewis, A.M., and Rice, K.C. (2016) Quantitative Real-Time PCR (qPCR) workflow for analyzing Staphylococcus aureus gene expression. Methods in molecular biology (Clifton, N.J.) 1373: 143-154.

Lewis, R.S., Tamir, S., Tannenbaum, S.R., and Deen, W.M. (1995) Kinetic analysis of the fate of nitric oxide synthesized by macrophages in vitro. J Biol Chem 270: 29350-29355.

Liebeke, M., and Lalk, M. (2014) Staphylococcus aureus metabolic response to changing environmental conditions - A metabolomics perspective. Int J Med Microbiol 304: 222-229.

209

Lieberman, M., Marks, A.D., Smith, C.M., and Marks, D.B., (2007) Marks' Essential Medical Biochemistry. Lippincott Williams & Wilkins.

Lina, G., Piemont, Y., Godail-Gamot, F., Bes, M., Peter, M.O., Gauduchon, V., Vandenesch, F., and Etienne, J. (1999) Involvement of Panton-Valentine leukocidin-producing Staphylococcus aureus in primary skin infections and pneumonia. Clin Infect Dis 29: 1128-1132.

Lindmark, R., Thoren-Tolling, K., and Sjoquist, J. (1983) Binding of immunoglobulins to protein A and immunoglobulin levels in mammalian sera. J Immunol Methods 62: 1-13.

Lindsay, J.A., (2008) Staphylococcus: Molecular Genetics. Caister Academic Press.

Liochev, S.I., and Fridovich, I. (1992) Fumarase C, the stable fumarase of Escherichia coli, is controlled by the soxRS regulon. Proc Natl Acad Sci U S A 89: 5892- 5896.

Lipton, S.A., (2001) Physiology. Nitric oxide and respiration. In: Nature. England, pp. 118-119, 121.

Liu, G.Y., Essex, A., Buchanan, J.T., Datta, V., Hoffman, H.M., Bastian, J.F., Fierer, J., and Nizet, V. (2005) Staphylococcus aureus golden pigment impairs neutrophil killing and promotes virulence through its antioxidant activity. J Exp Med 202: 209-215.

Liu, M., Tanaka, W.N., Zhu, H., Xie, G., Dooley, D.M., and Lei, B. (2008) Direct hemin transfer from IsdA to IsdC in the iron-regulated surface determinant (Isd) heme acquisition system of Staphylococcus aureus. J Biol Chem 283: 6668-6676.

Liu, X., and Zweier, J.L. (2013) Application of Electrode Methods in Studies of Nitric Oxide Metabolism and Diffusion Kinetics. J Electroanal Chem (Lausanne) 688: 32-39.

Lopez, J.M., and Thoms, B. (1977) Role of sugar uptake and metabolic intermediates on catabolite repression in Bacillus subtilis. J Bacteriol 129: 217-224.

Lowy, F.D. (2003) Antimicrobial resistance: the example of Staphylococcus aureus. J Clin Invest 111: 1265-1273.

Lu, J., and Holmgren, A. (2014) The thioredoxin antioxidant system. Free Radic Biol Med 66: 75-87.

Lustig, D.B., Kempt, C., Alam, S., Clancy, J., Yee, J., and Rafferty, S.P. (2011) Mutation of conserved tryptophan residues at the dimer interface of Staphylococcus aureus nitric oxide synthase. Arch Biochem Biophys 506: 165-172.

210

Machado, P., Felix, R., Rodrigues, R., Oliveira, S., and Rodrigues-Pousada, C. (2006) Characterization and expression analysis of the cytochrome bd oxidase operon from Desulfovibrio gigas. Curr Microbiol 52: 274-281.

MacMicking, J., Xie, Q.W., and Nathan, C. (1997) Nitric oxide and macrophage function. Annu Rev Immunol 15: 323-350.

Madej, T., Lanczycki, C.J., Zhang, D., Thiessen, P.A., Geer, R.C., Marchler-Bauer, A., and Bryant, S.H. (2014) MMDB and VAST+: tracking structural similarities between macromolecular complexes. Nucleic Acids Res 42: D297-303.

Mah, R.A., Fung, D.Y., and Morse, S.A. (1967) Nutritional requirements of Staphylococcus aureus S-6. Appl Microbiol 15: 866-870.

Maier, T., Marcos, J., Wodke, J.A.H., Paetzold, B., Liebeke, M., Gutierrez-Gallego, R., and Serrano, L. (2013) Large-scale metabolome analysis and quantitative integration with genomics and proteomics data in Mycoplasma pneumoniae. Molecular BioSystems 9: 1743-1755.

Majerczyk, C.D., Dunman, P.M., Luong, T.T., Lee, C.Y., Sadykov, M.R., Somerville, G.A., Bodi, K., and Sonenshein, A.L. (2010) Direct targets of CodY in Staphylococcus aureus. J Bacteriol 192: 2861-2877.

Malvin, G.M., Cecava, N., and Nelin, L.D. (2003) Nitric oxide production and thermoregulation in Paramecium caudatum. Acta protozoologica 42: 259-268.

Maresso, A.W., and Schneewind, O. (2006) Iron acquisition and transport in Staphylococcus aureus. Biometals 19: 193-203.

Marletta, M.A. (1994) Nitric oxide synthase: aspects concerning structure and catalysis. Cell 78: 927-930.

Marshall, J.H., and Wilmoth, G.J. (1981a) Pigments of Staphylococcus aureus, a series of triterpenoid carotenoids. J Bacteriol 147: 900-913.

Marshall, J.H., and Wilmoth, G.J. (1981b) Proposed pathway of triterpenoid carotenoid biosynthesis in Staphylococcus aureus: evidence from a study of mutants. J Bacteriol 147: 914-919.

Martin, S.E., and Chaven, S. (1987) Synthesis of catalase in Staphylococcus aureus MF-31. Appl Environ Microbiol 53: 1207-1209.

Martinez, A., and Kolter, R. (1997) Protection of DNA during oxidative stress by the nonspecific DNA-binding protein Dps. J Bacteriol 179: 5188-5194.

211

Masiuk, H., Kopron, K., Grumann, D., Goerke, C., Kolata, J., Jursa-Kulesza, J., Giedrys- Kalemba, S., Broker, B.M., and Holtfreter, S. (2010) Association of recurrent furunculosis with Panton-Valentine leukocidin and the genetic background of Staphylococcus aureus. J Clin Microbiol 48: 1527-1535.

Massey, V., Strickland, S., Mayhew, S.G., Howell, L.G., Engel, P.C., Matthews, R.G., Schuman, M., and Sullivan, P.A. (1969) The production of superoxide anion radicals in the reaction of reduced flavins and flavoproteins with molecular oxygen. Biochem Biophys Res Commun 36: 891-897.

Massova, I., and Kollman, P.A. (2002) pKa, MM, and QM studies of mechanisms of beta-lactamases and penicillin-binding proteins: acylation step. J Comput Chem 23: 1559-1576.

Mastronicola, D., Arcuri, E., Arese, M., Bacchi, A., Mercadante, S., Cardelli, P., Citro, G., and Sarti, P. (2004) Morphine but not fentanyl and methadone affects mitochondrial membrane potential by inducing nitric oxide release in glioma cells. Cell Mol Life Sci 61: 2991-2997.

Matsuhashi, M., Song, M.D., Ishino, F., Wachi, M., Doi, M., Inoue, M., Ubukata, K., Yamashita, N., and Konno, M. (1986) Molecular cloning of the gene of a penicillin-binding protein supposed to cause high resistance to beta-lactam antibiotics in Staphylococcus aureus. J Bacteriol 167: 975-980.

Mayer, S., Steffen, W., Steuber, J., and Gotz, F. (2015) The Staphylococcus aureus NuoL-like protein MpsA contributes to the generation of membrane potential. J Bacteriol 197: 794-806.

Mazmanian, S.K., Skaar, E.P., Gaspar, A.H., Humayun, M., Gornicki, P., Jelenska, J., Joachmiak, A., Missiakas, D.M., and Schneewind, O. (2003) Passage of heme- iron across the envelope of Staphylococcus aureus. Science 299: 906-909.

McCaig, L.F., McDonald, L.C., Mandal, S., and Jernigan, D.B. (2006) Staphylococcus aureus-associated skin and soft tissue infections in ambulatory care. Emerg Infect Dis 12: 1715-1723.

McCollister, B.D., Hoffman, M., Husain, M., and Vazquez-Torres, A. (2011) Nitric oxide protects bacteria from aminoglycosides by blocking the energy-dependent phases of drug uptake. Antimicrob Agents Chemother 55: 2189-2196.

McCormick, J.K., Yarwood, J.M., and Schlievert, P.M. (2001) Toxic shock syndrome and bacterial superantigens: an update. Annu Rev Microbiol 55: 77-104.

McGavin, M.J., Zahradka, C., Rice, K., and Scott, J.E. (1997) Modification of the Staphylococcus aureus fibronectin binding phenotype by V8 protease. Infect Immun 65: 2621-2628.

212

McLean, S., Bowman, L.A., and Poole, R.K. (2010) Peroxynitrite stress is exacerbated by flavohaemoglobin-derived oxidative stress in Salmonella typhimurium and is relieved by nitric oxide. Microbiology 156: 3556-3565.

McNamara, P.J., and Proctor, R.A. (2000) Staphylococcus aureus small colony variants, electron transport and persistent infections. Int J Antimicrob Agents 14: 117-122.

Mei, J.M., Nourbakhsh, F., Ford, C.W., and Holden, D.W. (1997) Identification of Staphylococcus aureus virulence genes in a murine model of bacteraemia using signature-tagged mutagenesis. Mol Microbiol 26: 399-407.

Membrillo-Hernandez, J., Ioannidis, N., and Poole, R.K. (1996) The flavohaemoglobin (HMP) of Escherichia coli generates superoxide in vitro and causes oxidative stress in vivo. FEBS Lett 382: 141-144.

Menestrina, G. (1986) Ionic channels formed by Staphylococcus aureus alpha-toxin: voltage-dependent inhibition by divalent and trivalent cations. J Membr Biol 90: 177-190.

Menestrina, G., Serra, M.D., and Prevost, G. (2001) Mode of action of beta-barrel pore- forming toxins of the staphylococcal alpha-hemolysin family. Toxicon 39: 1661- 1672.

Messner, K.R., and Imlay, J.A. (1999) The identification of primary sites of superoxide and hydrogen peroxide formation in the aerobic respiratory chain and sulfite reductase complex of Escherichia coli. J Biol Chem 274: 10119-10128.

Messner, K.R., and Imlay, J.A. (2002) Mechanism of superoxide and hydrogen peroxide formation by fumarate reductase, succinate dehydrogenase, and aspartate oxidase. J Biol Chem 277: 42563-42571.

Miajlovic, H., Fallon, P.G., Irvine, A.D., and Foster, T.J. (2010) Effect of filaggrin breakdown products on growth of and protein expression by Staphylococcus aureus. J Allergy Clin Immunol 126: 1184-1190.e1183.

Midha, S., Mishra, R., Aziz, M.A., Sharma, M., Mishra, A., Khandelwal, P., and Bhatnagar, R. (2005) Cloning, expression, and characterization of recombinant nitric oxide synthase-like protein from Bacillus anthracis. Biochem Biophys Res Commun 336: 346-356.

Mienaltowski, M.J., and Birk, D.E. (2014) Structure, physiology, and biochemistry of collagens. Adv Exp Med Biol 802: 5-29.

213

Miller, A.F. (2012) Superoxide dismutases: ancient enzymes and new insights. FEBS Lett 586: 585-595.

Miller, L.G., and Kaplan, S.L. (2009) Staphylococcus aureus: a community pathogen. Infect Dis Clin North Am 23: 35-52.

Minghetti, K.C., and Gennis, R.B. (1988) The two terminal oxidases of the aerobic respiratory chain of Escherichia coli each yield water and not peroxide as a final product. Biochem Biophys Res Commun 155: 243-248.

Mitani, N., Koizumi, A., Sano, R., Masutani, T., Murakawa, K., Mikasa, K., and Okamoto, Y. (2005) Molecular typing of methicillin-resistant Staphylococcus aureus by PCR-RFLP and its usefulness in an epidemiological study of an outbreak. Jpn J Infect Dis 58: 250-252.

Mitchell, P. (1961) Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191: 144-148.

Mitchell, P. (2011) Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. 1966. Biochim Biophys Acta 1807: 1507-1538.

Mohanty, J.G., Jaffe, J.S., Schulman, E.S., and Raible, D.G. (1997) A highly sensitive fluorescent micro-assay of H2O2 release from activated human leukocytes using a dihydroxyphenoxazine derivative. Journal of immunological methods 202: 133- 141.

Moller, M.N., Li, Q., Lancaster, J.R., Jr., and Denicola, A. (2007) Acceleration of nitric oxide autoxidation and nitrosation by membranes. IUBMB Life 59: 243-248.

Molne, L., Verdrengh, M., and Tarkowski, A. (2000) Role of neutrophil leukocytes in cutaneous infection caused by Staphylococcus aureus. Infect Immun 68: 6162- 6167.

Monaco, M., Pimentel de Araujo, F., Cruciani, M., Coccia, E.M., and Pantosti, A. (2016) Worldwide Epidemiology and Antibiotic Resistance of Staphylococcus aureus. Curr Top Microbiol Immunol.

Monecke, S., Slickers, P., Ellington, M.J., Kearns, A.M., and Ehricht, R. (2007) High diversity of Panton-Valentine leukocidin-positive, methicillin-susceptible isolates of Staphylococcus aureus and implications for the evolution of community- associated methicillin-resistant S. aureus. Clin Microbiol Infect 13: 1157-1164.

Montgomery, H.J., Dupont, A.L., Leivo, H.E., and Guillemette, J.G. (2010) Cloning, Expression, and Purification of a Nitric Oxide Synthase-Like Protein from Bacillus cereus. Biochem Res Int 2010: 489892.

214

Moore, C.M., Nakano, M.M., Wang, T., Ye, R.W., and Helmann, J.D. (2004) Response of Bacillus subtilis to nitric oxide and the nitrosating agent sodium nitroprusside. J Bacteriol 186: 4655-4664.

Moreillon, P., Entenza, J.M., Francioli, P., McDevitt, D., Foster, T.J., Francois, P., and Vaudaux, P. (1995) Role of Staphylococcus aureus coagulase and clumping factor in pathogenesis of experimental endocarditis. Infect Immun 63: 4738-4743.

Mukhopadhyay, P., Zheng, M., Bedzyk, L.A., LaRossa, R.A., and Storz, G. (2004) Prominent roles of the NorR and Fur regulators in the Escherichia coli transcriptional response to reactive nitrogen species. Proc Natl Acad Sci U S A 101: 745-750.

Mulcahy, M.E., Geoghegan, J.A., Monk, I.R., O'Keeffe, K.M., Walsh, E.J., Foster, T.J., and McLoughlin, R.M. (2012) Nasal colonisation by Staphylococcus aureus depends upon clumping factor B binding to the squamous epithelial cell envelope protein loricrin. PLoS Pathog 8: e1003092.

Muryoi, N., Tiedemann, M.T., Pluym, M., Cheung, J., Heinrichs, D.E., and Stillman, M.J. (2008) Demonstration of the iron-regulated surface determinant (Isd) heme transfer pathway in Staphylococcus aureus. J Biol Chem 283: 28125-28136.

Nair, S., and Finkel, S.E. (2004) Dps protects cells against multiple stresses during stationary phase. J Bacteriol 186: 4192-4198.

Nakano, M.M., and Zuber, P. (1998) Anaerobic growth of a "strict aerobe" (Bacillus subtilis). Annu Rev Microbiol 52: 165-190.

Nakano, M.M., Zuber, P., Glaser, P., Danchin, A., and Hulett, F.M. (1996) Two- component regulatory proteins ResD-ResE are required for transcriptional activation of fnr upon oxygen limitation in Bacillus subtilis. J Bacteriol 178: 3796- 3802.

Nalwaya, N., and Deen, W.M. (2005) Nitric oxide, oxygen, and superoxide formation and consumption in macrophage cultures. Chem Res Toxicol 18: 486-493.

Nathan, C., and Shiloh, M.U. (2000) Reactive oxygen and nitrogen intermediates in the relationship between mammalian hosts and microbial pathogens. Proc Natl Acad Sci U S A 97: 8841-8848.

Nedospasov, A., Rafikov, R., Beda, N., and Nudler, E. (2000) An autocatalytic mechanism of protein nitrosylation. Proc Natl Acad Sci U S A 97: 13543-13548.

Neubauer, H., and Gotz, F. (1996) Physiology and interaction of nitrate and nitrite reduction in Staphylococcus carnosus. Journal of bacteriology 178: 2005-2009.

215

Neubauer, H., Pantel, I., and Gotz, F. (1999) Molecular characterization of the nitrite- reducing system of Staphylococcus carnosus. Journal of bacteriology 181: 1481- 1488.

Newton, G.L., Arnold, K., Price, M.S., Sherrill, C., Delcardayre, S.B., Aharonowitz, Y., Cohen, G., Davies, J., Fahey, R.C., and Davis, C. (1996) Distribution of thiols in microorganisms: mycothiol is a major thiol in most actinomycetes. J Bacteriol 178: 1990-1995.

Nguyen, T., Brunson, D., Crespi, C.L., Penman, B.W., Wishnok, J.S., and Tannenbaum, S.R. (1992) DNA damage and mutation in human cells exposed to nitric oxide in vitro. Proceedings of the National Academy of Sciences of the United States of America 89: 3030-3034.

Nguyen, V.T., Higuchi, H., and Kamio, Y. (2002) Controlling pore assembly of staphylococcal gamma-haemolysin by low temperature and by disulphide bond formation in double-cysteine LukF mutants. Mol Microbiol 45: 1485-1498.

Nicholls, P. (2012) Classical catalase: ancient and modern. Arch Biochem Biophys 525: 95-101.

Niemann, V., Koch-Singenstreu, M., Neu, A., Nilkens, S., Gotz, F., Unden, G., and Stehle, T. (2014) The NreA protein functions as a nitrate receptor in the staphylococcal nitrate regulation system. J Mol Biol 426: 1539-1553.

Ninnemann, H., and Maier, J. (1996) Indications for the occurrence of nitric oxide synthases in fungi and plants and the involvement in photoconidiation of Neurospora crassa. Photochemistry and Photobiology 64: 393-398.

Nisbett, L.M., and Boon, E.M. (2016) Nitric Oxide Regulation of H-NOX signaling pathways in bacteria. Biochemistry 55: 4873-4884.

Nishida, C.R., Knudsen, G., Straub, W., and Ortiz de Montellano, P.R. (2002) Electron supply and catalytic oxidation of nitrogen by cytochrome P450 and nitric oxide synthase. Drug Metab Rev 34: 479-501.

Nizet, V., and Gallo, R.L. (2003) Cathelicidins and innate defense against invasive bacterial infection. Scand J Infect Dis 35: 670-676.

Nizet, V., Ohtake, T., Lauth, X., Trowbridge, J., Rudisill, J., Dorschner, R.A., Pestonjamasp, V., Piraino, J., Huttner, K., and Gallo, R.L. (2001) Innate antimicrobial peptide protects the skin from invasive bacterial infection. Nature 414: 454-457.

216

Nobre, L.S., and Saraiva, L.M. (2013) Effect of combined oxidative and nitrosative stresses on Staphylococcus aureus transcriptome. Appl Microbiol Biotechnol 97: 2563-2573.

Novick, R.P. (2000) Pathogenicity factors and their regulation. Gram-positive pathogens. ASM Press, Washington, DC: 392-407.

Novick, R.P., Projan, S.J., Kornblum, J., Ross, H.F., Ji, G., Kreiswirth, B., Vandenesch, F., and Moghazeh, S. (1995) The agr P2 operon: an autocatalytic sensory transduction system in Staphylococcus aureus. Molecular & general genetics : MGG 248: 446-458.

Novick, R.P., Ross, H.F., Projan, S.J., Kornblum, J., Kreiswirth, B., and Moghazeh, S. (1993) Synthesis of staphylococcal virulence factors is controlled by a regulatory RNA molecule. The EMBO journal 12: 3967-3975.

Novo, D., Perlmutter, N.G., Hunt, R.H., and Shapiro, H.M. (1999) Accurate flow cytometric membrane potential measurement in bacteria using diethyloxacarbocyanine and a ratiometric technique. Cytometry 35: 55-63.

Nuxoll, A.S., Halouska, S.M., Sadykov, M.R., Hanke, M.L., Bayles, K.W., Kielian, T., Powers, R., and Fey, P.D. (2012) CcpA regulates arginine biosynthesis in Staphylococcus aureus through repression of proline catabolism. PLoS Pathog 8: e1003033.

Nychas, G.J., Tranter, H.S., Brehm, R.D., and Board, R.G. (1991) Staphylococcus aureus S-6: factors affecting its growth, enterotoxin B production and exoprotein formation. J Appl Bacteriol 70: 344-350.

Nygaard, T.K., Pallister, K.B., DuMont, A.L., DeWald, M., Watkins, R.L., Pallister, E.Q., Malone, C., Griffith, S., Horswill, A.R., Torres, V.J., and Voyich, J.M. (2012) Alpha-toxin induces programmed cell death of human T cells, B cells, and monocytes during USA300 infection. PLoS One 7: e36532.

O'Donnell, V.B., Chumley, P.H., Hogg, N., Bloodsworth, A., Darley-Usmar, V.M., and Freeman, B.A. (1997) Nitric oxide inhibition of lipid peroxidation: kinetics of reaction with lipid peroxyl radicals and comparison with alpha-tocopherol. Biochemistry 36: 15216-15223.

Ohniwa, R.L., Ushijima, Y., Saito, S., and Morikawa, K. (2011) Proteomic analyses of nucleoid-associated proteins in Escherichia coli, Pseudomonas aeruginosa, Bacillus subtilis, and Staphylococcus aureus. PLoS One 6: e19172.

Ohno, H., Zhu, G., Mohan, V.P., Chu, D., Kohno, S., Jacobs, W.R., Jr., and Chan, J. (2003) The effects of reactive nitrogen intermediates on gene expression in Mycobacterium tuberculosis. Cell Microbiol 5: 637-648.

217

Olson, J.S., Foley, E.W., Rogge, C., Tsai, A.L., Doyle, M.P., and Lemon, D.D. (2004) No scavenging and the hypertensive effect of hemoglobin-based blood substitutes. Free Radic Biol Med 36: 685-697.

Olson, R., Nariya, H., Yokota, K., Kamio, Y., and Gouaux, E. (1999) Crystal structure of staphylococcal LukF delineates conformational changes accompanying formation of a transmembrane channel. Nat Struct Biol 6: 134-140.

Ong, S.T., Ho, J.Z., Ho, B., and Ding, J.L. (2006) Iron-withholding strategy in innate immunity. Immunobiology 211: 295-314.

Onoue, Y., and Mori, M. (1997) Amino acid requirements for the growth and enterotoxin production by Staphylococcus aureus in chemically defined media. Int J Food Microbiol 36: 77-82.

Oogai, Y., Kawada-Matsuo, M., and Komatsuzawa, H. (2016) Staphylococcus aureus SrrAB affects susceptibility to hydrogen peroxide and co-existence with Streptococcus sanguinis. PLoS One 11: e0159768.

Otto, B.R., Verweij-van Vught, A.M., and MacLaren, D.M. (1992) Transferrins and heme-compounds as iron sources for pathogenic bacteria. Crit Rev Microbiol 18: 217-233.

Otto, M. (2012) MRSA virulence and spread. Cell Microbiol 14: 1513-1521.

Overton, T.W., Justino, M.C., Li, Y., Baptista, J.M., Melo, A.M., Cole, J.A., and Saraiva, L.M. (2008) Widespread distribution in pathogenic bacteria of di-iron proteins that repair oxidative and nitrosative damage to iron-sulfur centers. J Bacteriol 190: 2004-2013.

Owen, O.E., Kalhan, S.C., and Hanson, R.W. (2002) The key role of anaplerosis and cataplerosis for citric acid cycle function. J Biol Chem 277: 30409-30412.

Pagels, M., Fuchs, S., Pane-Farre, J., Kohler, C., Menschner, L., Hecker, M., McNamarra, P.J., Bauer, M.C., von Wachenfeldt, C., Liebeke, M., Lalk, M., Sander, G., von Eiff, C., Proctor, R.A., and Engelmann, S. (2010) Redox sensing by a Rex-family repressor is involved in the regulation of anaerobic gene expression in Staphylococcus aureus. Mol Microbiol 76: 1142-1161.

Pannala, V.R., and Dash, R.K. (2015) Mechanistic characterization of the thioredoxin system in the removal of hydrogen peroxide. Free Radic Biol Med 78: 42-55.

Pant, K., Bilwes, A.M., Adak, S., Stuehr, D.J., and Crane, B.R. (2002) Structure of a nitric oxide synthase heme protein from Bacillus subtilis. Biochemistry 41: 11071- 11079.

218

Pantel, I., Lindgren, P.E., Neubauer, H., and Gotz, F. (1998) Identification and characterization of the Staphylococcus carnosus nitrate reductase operon. Molecular & general genetics : MGG 259: 105-114.

Parfentjev, I.A., and Catelli, A.R. (1964) Tolerance of Staphylococcus aureus to sodium chloride. J Bacteriol 88: 1-3.

Parihar, M.S., Nazarewicz, R.R., Kincaid, E., Bringold, U., and Ghafourifar, P. (2008a) Association of mitochondrial nitric oxide synthase activity with respiratory chain complex I. Biochem Biophys Res Commun 366: 23-28.

Parihar, M.S., Parihar, A., Chen, Z., Nazarewicz, R., and Ghafourifar, P. (2008b) mAtNOS1 regulates mitochondrial functions and apoptosis of human neuroblastoma cells. Biochim Biophys Acta 1780: 921-926.

Parihar, M.S., Parihar, A., Villamena, F.A., Vaccaro, P.S., and Ghafourifar, P. (2008c) Inactivation of mitochondrial respiratory chain complex I leads mitochondrial nitric oxide synthase to become pro-oxidative. Biochem Biophys Res Commun 367: 761-767.

Park, S., and Imlay, J.A. (2003) High levels of intracellular cysteine promote oxidative DNA damage by driving the fenton reaction. J Bacteriol 185: 1942-1950.

Parsonage, D., Karplus, P.A., and Poole, L.B. (2008) Substrate specificity and redox potential of AhpC, a bacterial peroxiredoxin. Proc Natl Acad Sci U S A 105: 8209- 8214.

Patel, B.A., Moreau, M., Widom, J., Chen, H., Yin, L., Hua, Y., and Crane, B.R. (2009) Endogenous nitric oxide regulates the recovery of the radiation-resistant bacterium Deinococcus radiodurans from exposure to UV light. Proc Natl Acad Sci U S A 106: 18183-18188.

Patti, J.M., Allen, B.L., McGavin, M.J., and Hook, M. (1994a) MSCRAMM-mediated adherence of microorganisms to host tissues. Annu Rev Microbiol 48: 585-617.

Patti, J.M., Bremell, T., Krajewska-Pietrasik, D., Abdelnour, A., Tarkowski, A., Ryden, C., and Hook, M. (1994b) The Staphylococcus aureus collagen adhesin is a virulence determinant in experimental septic arthritis. Infect Immun 62: 152-161.

Patton, T.G., Yang, S.J., and Bayles, K.W. (2006) The role of proton motive force in expression of the Staphylococcus aureus cid and lrg operons. Molecular microbiology 59: 1395-1404.

219

Peacock, S.J., Foster, T.J., Cameron, B.J., and Berendt, A.R. (1999) Bacterial fibronectin-binding proteins and endothelial cell surface fibronectin mediate adherence of Staphylococcus aureus to resting human endothelial cells. Microbiology 145 ( Pt 12): 3477-3486.

Peacock, S.J., Moore, C.E., Justice, A., Kantzanou, M., Story, L., Mackie, K., O'Neill, G., and Day, N.P. (2002) Virulent combinations of adhesin and toxin genes in natural populations of Staphylococcus aureus. Infect Immun 70: 4987-4996.

Pearce, L.L., Lopez Manzano, E., Martinez-Bosch, S., and Peterson, J. (2008) Antagonism of nitric oxide toward the inhibition of cytochrome c oxidase by carbon monoxide and cyanide. Chem Res Toxicol 21: 2073-2081.

Pedelacq, J.D., Maveyraud, L., Prevost, G., Baba-Moussa, L., Gonzalez, A., Courcelle, E., Shepard, W., Monteil, H., Samama, J.P., and Mourey, L. (1999) The structure of a Staphylococcus aureus leucocidin component (LukF-PV) reveals the fold of the water-soluble species of a family of transmembrane pore-forming toxins. Structure 7: 277-287.

Peschel, A., Otto, M., Jack, R.W., Kalbacher, H., Jung, G., and Gotz, F. (1999) Inactivation of the dlt operon in Staphylococcus aureus confers sensitivity to defensins, protegrins, and other antimicrobial peptides. J Biol Chem 274: 8405- 8410.

Peters, B.M., Shirtliff, M.E., and Jabra-Rizk, M.A. (2010) Antimicrobial peptides: primeval molecules or future drugs? PLoS Pathog 6: e1001067.

Pfeltz, R.F., Singh, V.K., Schmidt, J.L., Batten, M.A., Baranyk, C.S., Nadakavukaren, M.J., Jayaswal, R.K., and Wilkinson, B.J. (2000) Characterization of passage- selected vancomycin-resistant Staphylococcus aureus strains of diverse parental backgrounds. Antimicrob Agents Chemother 44: 294-303.

Piazza, M., Futrega, K., Spratt, D.E., Dieckmann, T., and Guillemette, J.G. (2012) Structure and dynamics of calmodulin (CaM) bound to nitric oxide synthase peptides: effects of a phosphomimetic CaM mutation. Biochemistry 51: 3651- 3661.

Pilpa, R.M., Robson, S.A., Villareal, V.A., Wong, M.L., Phillips, M., and Clubb, R.T. (2009) Functionally distinct NEAT (NEAr Transporter) domains within the Staphylococcus aureus IsdH/HarA protein extract heme from methemoglobin. J Biol Chem 284: 1166-1176.

Pishchany, G., Sheldon, J.R., Dickson, C.F., Alam, M.T., Read, T.D., Gell, D.A., Heinrichs, D.E., and Skaar, E.P. (2014) IsdB-dependent hemoglobin binding is required for acquisition of heme by Staphylococcus aureus. J Infect Dis 209: 1764-1772.

220

Plate, L., and Marletta, M.A. (2013) Nitric oxide-sensing H-NOX proteins govern bacterial communal behavior. Trends Biochem Sci 38: 566-575.

Pohl, K., Francois, P., Stenz, L., Schlink, F., Geiger, T., Herbert, S., Goerke, C., Schrenzel, J., and Wolz, C. (2009) CodY in Staphylococcus aureus: a regulatory link between metabolism and virulence gene expression. J Bacteriol 191: 2953- 2963.

Poole, L.B., Reynolds, C.M., Wood, Z.A., Karplus, P.A., Ellis, H.R., and Li Calzi, M. (2000) AhpF and other NADH:peroxiredoxin oxidoreductases, homologues of low Mr thioredoxin reductase. Eur J Biochem 267: 6126-6133.

Poole, R.K. (2005) Nitric oxide and nitrosative stress tolerance in bacteria. Biochem Soc Trans 33: 176-180.

Poole, R.K., Anjum, M.F., Membrillo-Hernandez, J., Kim, S.O., Hughes, M.N., and Stewart, V. (1996) Nitric oxide, nitrite, and Fnr regulation of hmp (flavohemoglobin) gene expression in Escherichia coli K-12. J Bacteriol 178: 5487-5492.

Popova, T.G., Teunis, A., Vaseghi, H., Zhou, W., Espina, V., Liotta, L.A., and Popov, S.G. (2015) Nitric oxide as a regulator of Bacillus anthracis pathogenicity. Front Microbiol 6: 921.

Postma, B., Poppelier, M.J., van Galen, J.C., Prossnitz, E.R., van Strijp, J.A., de Haas, C.J., and van Kessel, K.P. (2004) Chemotaxis inhibitory protein of Staphylococcus aureus binds specifically to the C5a and formylated peptide receptor. J Immunol 172: 6994-7001.

Pragman, A.A., Ji, Y., and Schlievert, P.M. (2007) Repression of Staphylococcus aureus SrrAB using inducible antisense srrA alters growth and virulence factor transcript levels. Biochemistry 46: 314-321.

Pragman, A.A., Yarwood, J.M., Tripp, T.J., and Schlievert, P.M. (2004) Characterization of virulence factor regulation by SrrAB, a two-component system in Staphylococcus aureus. J Bacteriol 186: 2430-2438.

Precit, M.R., Wolter, D.J., Griffith, A., Emerson, J., Burns, J.L., and Hoffman, L.R. (2016) Optimized in vitro antibiotic susceptibility testing method for small-colony variant Staphylococcus aureus. Antimicrob Agents Chemother 60: 1725-1735.

Price-Whelan, A., Poon, C.K., Benson, M.A., Eidem, T.T., Roux, C.M., Boyd, J.M., Dunman, P.M., Torres, V.J., and Krulwich, T.A. (2013) Transcriptional profiling of Staphylococcus aureus during growth in 2 M NaCl leads to clarification of physiological roles for Kdp and Ktr K+ uptake systems. MBio 4.

221

Proctor, R.A., Balwit, J.M., and Vesga, O. (1994) Variant subpopulations of Staphylococcus aureus as cause of persistent and recurrent infections. Infectious agents and disease 3: 302-312.

Proctor, R.A., van Langevelde, P., Kristjansson, M., Maslow, J.N., and Arbeit, R.D. (1995) Persistent and relapsing infections associated with small-colony variants of Staphylococcus aureus. Clinical infectious diseases : an official publication of the Infectious Diseases Society of America 20: 95-102.

Radi, R. (2004) Nitric oxide, oxidants, and protein tyrosine nitration. Proc Natl Acad Sci U S A 101: 4003-4008.

Radi, R. (2013) Protein tyrosine nitration: biochemical mechanisms and structural basis of functional effects. Acc Chem Res 46: 550-559.

Radi, R., Beckman, J.S., Bush, K.M., and Freeman, B.A. (1991a) Peroxynitrite oxidation of sulfhydryls. The cytotoxic potential of superoxide and nitric oxide. The Journal of biological chemistry 266: 4244-4250.

Radi, R., Beckman, J.S., Bush, K.M., and Freeman, B.A. (1991b) Peroxynitrite-induced membrane lipid peroxidation: the cytotoxic potential of superoxide and nitric oxide. Archives of biochemistry and biophysics 288: 481-487.

Raman, C.S., Li, H., Martasek, P., Kral, V., Masters, B.S., and Poulos, T.L. (1998) Crystal structure of constitutive endothelial nitric oxide synthase: a paradigm for pterin function involving a novel metal center. Cell 95: 939-950.

Reece, S.Y., Woodward, J.J., and Marletta, M.A. (2009) Synthesis of nitric oxide by the NOS-like protein from Deinococcus radiodurans: a direct role for tetrahydrofolate. Biochemistry 48: 5483-5491.

Repine, J.E., Fox, R.B., and Berger, E.M. (1981) Hydrogen peroxide kills Staphylococcus aureus by reacting with staphylococcal iron to form hydroxyl radical. J Biol Chem 256: 7094-7096.

Richardson, A.R., Dunman, P.M., and Fang, F.C. (2006) The nitrosative stress response of Staphylococcus aureus is required for resistance to innate immunity. Mol Microbiol 61: 927-939.

Richardson, A.R., Libby, S.J., and Fang, F.C. (2008) A nitric oxide-inducible lactate dehydrogenase enables Staphylococcus aureus to resist innate immunity. Science 319: 1672-1676.

Richardson, J.F. The Staphylococci in Human Disease. Eds. K. B. Crossley and G. L. Archer. Churchill Livingstone 1997. Pp. 682. £90.00. ISBN 0 4430 7644 8. Epidemiology and Infection 120: 111-114.

222

Rim, J.Y., and Bacon, A.E., 3rd (2007) Prevalence of community-acquired methicillin- resistant Staphylococcus aureus colonization in a random sample of healthy individuals. Infect Control Hosp Epidemiol 28: 1044-1046.

Roben, P.W., Salem, A.N., and Silverman, G.J. (1995) VH3 family antibodies bind domain D of staphylococcal protein A. J Immunol 154: 6437-6445.

Robinson, K.M., Janes, M.S., Pehar, M., Monette, J.S., Ross, M.F., Hagen, T.M., Murphy, M.P., and Beckman, J.S. (2006) Selective fluorescent imaging of superoxide in vivo using ethidium-based probes. Proceedings of the National Academy of Sciences of the United States of America 103: 15038-15043.

Rodvold, K.A., and McConeghy, K.W. (2014) Methicillin-resistant Staphylococcus aureus therapy: past, present, and future. Clin Infect Dis 58 Suppl 1: S20-27.

Rose, I.A., and O'Connell, E.L. (1967) Mechanism of aconitase action. I. The hydrogen transfer reaction. The Journal of biological chemistry 242: 1870-1879.

Roth, R.R., and James, W.D. (1988) Microbial ecology of the skin. Annu Rev Microbiol 42: 441-464.

Rothfork, J.M., Dessus-Babus, S., Van Wamel, W.J., Cheung, A.L., and Gresham, H.D. (2003) Fibrinogen depletion attenuates Staphylococcus aureus infection by preventing density-dependent virulence gene up-regulation. J Immunol 171: 5389-5395.

Roux, A., Todd, D.A., Velazquez, J.V., Cech, N.B., and Sonenshein, A.L. (2014) CodY- mediated regulation of the Staphylococcus aureus Agr system integrates nutritional and population density signals. J Bacteriol 196: 1184-1196.

Rubbo, H., Radi, R., Anselmi, D., Kirk, M., Barnes, S., Butler, J., Eiserich, J.P., and Freeman, B.A. (2000) Nitric oxide reaction with lipid peroxyl radicals spares alpha-tocopherol during lipid peroxidation. Greater oxidant protection from the pair nitric oxide/alpha-tocopherol than alpha-tocopherol/ascorbate. J Biol Chem 275: 10812-10818.

Sadykov, M.R., Olson, M.E., Halouska, S., Zhu, Y., Fey, P.D., Powers, R., and Somerville, G.A. (2008) Tricarboxylic acid cycle-dependent regulation of Staphylococcus epidermidis polysaccharide intercellular adhesin synthesis. J Bacteriol 190: 7621-7632.

Sadykov, M.R., Thomas, V.C., Marshall, D.D., Wenstrom, C.J., Moormeier, D.E., Widhelm, T.J., Nuxoll, A.S., Powers, R., and Bayles, K.W. (2013) Inactivation of the Pta-AckA pathway causes cell death in Staphylococcus aureus. J Bacteriol 195: 3035-3044.

223

Salard, I., Mercey, E., Rekka, E., Boucher, J.L., Nioche, P., Mikula, I., Martasek, P., Raman, C.S., and Mansuy, D. (2006) Analogies and surprising differences between recombinant nitric oxide synthase-like proteins from Staphylococcus aureus and Bacillus anthracis in their interactions with l-arginine analogs and iron ligands. J Inorg Biochem 100: 2024-2033.

Salard-Arnaud, I., Stuehr, D., Boucher, J.L., and Mansuy, D. (2012) Spectroscopic, catalytic and binding properties of Bacillus subtilis NO synthase-like protein: comparison with other bacterial and mammalian NO synthases. J Inorg Biochem 106: 164-171.

Salgo, M.G., Bermudez, E., Squadrito, G.L., and Pryor, W.A. (1995a) Peroxynitrite causes DNA damage and oxidation of thiols in rat thymocytes. Arch Biochem Biophys 322: 500-505.

Salgo, M.G., Stone, K., Squadrito, G.L., Battista, J.R., and Pryor, W.A. (1995b) Peroxynitrite causes DNA nicks in plasmid pBR322. Biochem Biophys Res Commun 210: 1025-1030.

Santolini, J., Roman, M., Stuehr, D.J., and Mattioli, T.A. (2006) Resonance Raman study of Bacillus subtilis NO synthase-like protein: similarities and differences with mammalian NO synthases. Biochemistry 45: 1480-1489.

Sapp, A.M., Mogen, A.B., Almand, E.A., Rivera, F.E., Shaw, L.N., Richardson, A.R., and Rice, K.C. (2014) Contribution of the nos-pdt operon to virulence phenotypes in methicillin-sensitive Staphylococcus aureus. PloS one 9: e108868.

Sarti, P., Giuffre, A., Barone, M.C., Forte, E., Mastronicola, D., and Brunori, M. (2003) Nitric oxide and cytochrome oxidase: reaction mechanisms from the enzyme to the cell. Free Radic Biol Med 34: 509-520.

Sasano, M., Burton, D.R., and Silverman, G.J. (1993) Molecular selection of human antibodies with an unconventional bacterial B cell antigen. J Immunol 151: 5822- 5839.

Sasarman, A., Purvis, P., and Portelance, V. (1974) Role of menaquinone in nitrate respiration in Staphylococcus aureus. J Bacteriol 117: 911-913.

Sasindran, S.J., Saikolappan, S., and Dhandayuthapani, S. (2007) Methionine sulfoxide reductases and virulence of bacterial pathogens. Future Microbiol 2: 619-630.

Sassi, M., Sharma, D., Brinsmade, S.R., Felden, B., and Augagneur, Y., (2015) Genome sequence of the clinical isolate Staphylococcus aureus subsp. aureus strain UAMS-1. In: Genome Announc. pp.

224

Sasso, E.H., Silverman, G.J., and Mannik, M. (1989) Human IgM molecules that bind staphylococcal protein A contain VHIII H chains. J Immunol 142: 2778-2783.

Sasso, E.H., Silverman, G.J., and Mannik, M. (1991) Human IgA and IgG F(ab')2 that bind to staphylococcal protein A belong to the VHIII subgroup. J Immunol 147: 1877-1883.

Scheidel, W. (2010) Coin quality, coin quantity, and coin value in early China and the Roman world. 22: 93-118.

Schlag, S., Fuchs, S., Nerz, C., Gaupp, R., Engelmann, S., Liebeke, M., Lalk, M., Hecker, M., and Gotz, F. (2008) Characterization of the oxygen-responsive NreABC regulon of Staphylococcus aureus. J Bacteriol 190: 7847-7858.

Schneider, C.A., Rasband, W.S., and Eliceiri, K.W. (2012) NIH Image to ImageJ: 25 years of image analysis. Nature methods 9: 671-675.

Schneiker, S., Perlova, O., Kaiser, O., Gerth, K., Alici, A., Altmeyer, M.O., Bartels, D., Bekel, T., Beyer, S., Bode, E., Bode, H.B., Bolten, C.J., Choudhuri, J.V., Doss, S., Elnakady, Y.A., Frank, B., Gaigalat, L., Goesmann, A., Groeger, C., Gross, F., Jelsbak, L., Kalinowski, J., Kegler, C., Knauber, T., Konietzny, S., Kopp, M., Krause, L., Krug, D., Linke, B., Mahmud, T., Martinez-Arias, R., McHardy, A.C., Merai, M., Meyer, F., Mormann, S., Munoz-Dorado, J., Perez, J., Pradella, S., Rachid, S., Raddatz, G., Rosenau, F., Ruckert, C., Sasse, F., Scharfe, M., Schuster, S.C., Suen, G., Treuner-Lange, A., Velicer, G.J., Vorholter, F.J., Weissman, K.J., Welch, R.D., Wenzel, S.C., Whitworth, D.E., Wilhelm, S., Wittmann, C., Blocker, H., Puhler, A., and Muller, R. (2007) Complete genome sequence of the myxobacterium Sorangium cellulosum. Nat Biotechnol 25: 1281- 1289.

Schoen, C., Kischkies, L., Elias, J., and Ampattu, B.J. (2014) Metabolism and virulence in Neisseria meningitidis. Front Cell Infect Microbiol 4: 114.

Schreiber, F., Beutler, M., Enning, D., Lamprecht-Grandio, M., Zafra, O., González- Pastor, J.E., and de Beer, D. (2011) The role of nitric-oxide-synthase-derived nitric oxide in multicellular traits of Bacillus subtilis 3610: biofilm formation, swarming, and dispersal. BMC Microbiol 11: 111.

Schreier, H.J., Brown, S.W., Hirschi, K.D., Nomellini, J.F., and Sonenshein, A.L. (1989) Regulation of Bacillus subtilis glutamine synthetase gene expression by the product of the glnR gene. J Mol Biol 210: 51-63.

Schreier, H.J., Caruso, S.M., and Maier, K.C. (2000) Control of Bacillus subtilis glutamine synthetase expression by glnR from Staphylococcus aureus. Curr Microbiol 41: 425-429.

225

Schumacher, M.A., Seidel, G., Hillen, W., and Brennan, R.G. (2007) Structural mechanism for the fine-tuning of CcpA function by the small molecule effectors glucose 6-phosphate and fructose 1,6-bisphosphate. J Mol Biol 368: 1042-1050.

Schurig-Briccio, L.A., Yano, T., Rubin, H., and Gennis, R.B. (2014) Characterization of the type 2 NADH:menaquinone oxidoreductases from Staphylococcus aureus and the bactericidal action of phenothiazines. Biochim Biophys Acta 1837: 954- 963.

Schwan, W.R., Langhorne, M.H., Ritchie, H.D., and Stover, C.K. (2003) Loss of hemolysin expression in Staphylococcus aureus agr mutants correlates with selective survival during mixed infections in murine abscesses and wounds. FEMS Immunol Med Microbiol 38: 23-28.

Scott, I.R. (1981) Factors controlling the expressed activity of histidine ammonia-lyase in the epidermis and the resulting accumulation of urocanic acid. Biochem J 194: 829-838.

Scovill, W.H., Schreier, H.J., and Bayles, K.W. (1996) Identification and characterization of the pckA gene from Staphylococcus aureus. Journal of bacteriology 178: 3362-3364.

Seaver, L.C., and Imlay, J.A. (2004) Are respiratory enzymes the primary sources of intracellular hydrogen peroxide? J Biol Chem 279: 48742-48750.

Seidl, K., Bischoff, M., and Berger-Bachi, B. (2008a) CcpA mediates the catabolite repression of tst in Staphylococcus aureus. Infect Immun 76: 5093-5099.

Seidl, K., Goerke, C., Wolz, C., Mack, D., Berger-Bachi, B., and Bischoff, M. (2008b) Staphylococcus aureus CcpA affects biofilm formation. Infect Immun 76: 2044- 2050.

Seidl, K., Muller, S., Francois, P., Kriebitzsch, C., Schrenzel, J., Engelmann, S., Bischoff, M., and Berger-Bachi, B. (2009) Effect of a glucose impulse on the CcpA regulon in Staphylococcus aureus. BMC Microbiol 9: 95.

Sena, F.V., Batista, A.P., Catarino, T., Brito, J.A., Archer, M., Viertler, M., Madl, T., Cabrita, E.J., and Pereira, M.M. (2015) Type-II NADH:quinone oxidoreductase from Staphylococcus aureus has two distinct binding sites and is rate limited by quinone reduction. Mol Microbiol 98: 272-288.

Severin, A., Tabei, K., Tenover, F., Chung, M., Clarke, N., and Tomasz, A. (2004) High level oxacillin and vancomycin resistance and altered cell wall composition in Staphylococcus aureus carrying the staphylococcal mecA and the enterococcal vanA gene complex. J Biol Chem 279: 3398-3407.

226

Shakeri, F., Ghaemi, E.A., and Babai Kochkaksaraei, M. (2014) Staphylococcus aureus typing by digestion of protein A coding gene using Bsp143I. Jundishapur J Microbiol 7: e10320.

Shapiro, H.M., and Nebe-von-Caron, G. (2004) Multiparameter flow cytometry of bacteria. Methods in molecular biology (Clifton, N.J.) 263: 33-44.

Shatalin, K., Gusarov, I., Avetissova, E., Shatalina, Y., McQuade, L.E., Lippard, S.J., and Nudler, E. (2008) Bacillus anthracis-derived nitric oxide is essential for pathogen virulence and survival in macrophages. Proc Natl Acad Sci U S A 105: 1009-1013.

Shi, L., Sohaskey, C.D., Kana, B.D., Dawes, S., North, R.J., Mizrahi, V., and Gennaro, M.L. (2005) Changes in energy metabolism of Mycobacterium tuberculosis in mouse lung and under in vitro conditions affecting aerobic respiration. Proc Natl Acad Sci U S A 102: 15629-15634.

Shivers, R.P., and Sonenshein, A.L. (2004) Activation of the Bacillus subtilis global regulator CodY by direct interaction with branched-chain amino acids. Mol Microbiol 53: 599-611.

Sickmier, E.A., Brekasis, D., Paranawithana, S., Bonanno, J.B., Paget, M.S., Burley, S.K., and Kielkopf, C.L. (2005) X-ray structure of a Rex-family repressor/NADH complex insights into the mechanism of redox sensing. Structure 13: 43-54.

Sieradzki, K., and Tomasz, A. (2003) Alterations of cell wall structure and metabolism accompany reduced susceptibility to vancomycin in an isogenic series of clinical isolates of Staphylococcus aureus. J Bacteriol 185: 7103-7110.

Simon, J., van Spanning, R.J., and Richardson, D.J. (2008) The organisation of proton motive and non-proton motive redox loops in prokaryotic respiratory systems. Biochim Biophys Acta 1777: 1480-1490.

Singh, V.K., and Moskovitz, J. (2003) Multiple methionine sulfoxide reductase genes in Staphylococcus aureus: expression of activity and roles in tolerance of oxidative stress. Microbiology 149: 2739-2747.

Singh, V.K., Vaish, M., Johansson, T.R., Baum, K.R., Ring, R.P., Singh, S., Shukla, S.K., and Moskovitz, J. (2015) Significance of four methionine sulfoxide reductases in Staphylococcus aureus. PLoS One 10: e0117594.

Sinha, B., Francois, P.P., Nusse, O., Foti, M., Hartford, O.M., Vaudaux, P., Foster, T.J., Lew, D.P., Herrmann, M., and Krause, K.H. (1999) Fibronectin-binding protein acts as Staphylococcus aureus invasin via fibronectin bridging to integrin alpha5beta1. Cell Microbiol 1: 101-117.

227

Skaar, E.P., Gaspar, A.H., and Schneewind, O. (2004) IsdG and IsdI, heme-degrading enzymes in the cytoplasm of Staphylococcus aureus. J Biol Chem 279: 436-443.

Skaar, E.P., and Schneewind, O. (2004) Iron-regulated surface determinants (Isd) of Staphylococcus aureus: stealing iron from heme. Microbes Infect 6: 390-397.

Slauch, J.M. (2011) How does the oxidative burst of macrophages kill bacteria? Still an open question. Mol Microbiol 80: 580-583.

Smith, B.C., Underbakke, E.S., Kulp, D.W., Schief, W.R., and Marletta, M.A. (2013) Nitric oxide synthase domain interfaces regulate electron transfer and calmodulin activation. Proc Natl Acad Sci U S A 110: E3577-3586.

Smith, J.J., and McFeters, G.A. (1997) Mechanisms of INT (2-(4-iodophenyl)-3-(4- nitrophenyl)-5-phenyltetrazolium chloride), and CTC (5-cyano-2,3-ditolyl tetrazolium chloride) reduction in Escherichia coli K-12. Journal of Microbiological Methods 29: 161-175.

Solozhenkin, I.P., Kukushkina, N.V., Kutsemako, R.T., Kotelevets, L.M., Ermachenko, V.A., Lukoianova, M.A., and Burbaev, D. (1991) Iron-sulfur centers in the Staphylococcus aureus respiratory chain. Biokhimiia 56: 1259-1263.

Somerville, G.A., Chaussee, M.S., Morgan, C.I., Fitzgerald, J.R., Dorward, D.W., Reitzer, L.J., and Musser, J.M. (2002) Staphylococcus aureus aconitase inactivation unexpectedly inhibits post-exponential-phase growth and enhances stationary-phase survival. Infect Immun 70: 6373-6382.

Somerville, G.A., Cockayne, A., Durr, M., Peschel, A., Otto, M., and Musser, J.M. (2003a) Synthesis and deformylation of Staphylococcus aureus delta-toxin are linked to tricarboxylic acid cycle activity. J Bacteriol 185: 6686-6694.

Somerville, G.A., and Proctor, R.A. (2009a) At the crossroads of bacterial metabolism and virulence factor synthesis in Staphylococci. Microbiol Mol Biol Rev 73: 233- 248.

Somerville, G.A., and Proctor, R.A., (2009b) The Biology of Staphylococci. In: Staphylococci in human disease. Wiley-Blackwell, pp. 1-18.

Somerville, G.A., Said-Salim, B., Wickman, J.M., Raffel, S.J., Kreiswirth, B.N., and Musser, J.M. (2003b) Correlation of acetate catabolism and growth yield in Staphylococcus aureus: implications for host-pathogen interactions. Infection and immunity 71: 4724-4732.

Sonenshein, A.L. (2005) CodY, a global regulator of stationary phase and virulence in Gram-positive bacteria. Curr Opin Microbiol 8: 203-207.

228

Song, L., Hobaugh, M.R., Shustak, C., Cheley, S., Bayley, H., and Gouaux, J.E. (1996) Structure of staphylococcal alpha-hemolysin, a heptameric transmembrane pore. Science 274: 1859-1866.

Spahich, N.A., Vitko, N.P., Thurlow, L.R., Temple, B., and Richardson, A.R. (2016) Staphylococcus aureus lactate- and malate-quinone oxidoreductases contribute to nitric oxide resistance and virulence. Mol Microbiol 100: 759-773.

Spiro, S. (2007) Regulators of bacterial responses to nitric oxide. FEMS Microbiol Rev 31: 193-211.

Stockland, A.E., and San Clemente, C.L. (1969) Multiple forms of lactate dehydrogenase in Staphylococcus aureus. J Bacteriol 100: 347-353.

Stone, J.R., Sands, R.H., Dunham, W.R., and Marletta, M.A. (1995) Electron paramagnetic resonance spectral evidence for the formation of a pentacoordinate nitrosyl-heme complex on soluble guanylate cyclase. Biochem Biophys Res Commun 207: 572-577.

Strasters, K.C., and Winkler, K.C. (1963) Carbohydrate metabolism of Staphylococcus aureus. J Gen Microbiol 33: 213-229.

Stuehr, D.J. (1999) Mammalian nitric oxide synthases. Biochim Biophys Acta 1411: 217-230.

Stuehr, D.J., Santolini, J., Wang, Z.Q., Wei, C.C., and Adak, S. (2004a) Update on mechanism and catalytic regulation in the NO synthases. J Biol Chem 279: 36167-36170.

Stuehr, D.J., Wei, C.C., Santolini, J., Wang, Z., Aoyagi, M., and Getzoff, E.D. (2004b) Radical reactions of nitric oxide synthases. Biochem Soc Symp: 39-49.

Sudhamsu, J., and Crane, B.R. (2006) Structure and reactivity of a thermostable prokaryotic nitric-oxide synthase that forms a long-lived oxy-heme complex. J Biol Chem 281: 9623-9632.

Sudhamsu, J., and Crane, B.R. (2009) Bacterial nitric oxide synthases: what are they good for? Trends Microbiol 17: 212-218.

Sullivan, M.A., Yasbin, R.E., and Young, F.E. (1984) New shuttle vectors for Bacillus subtilis and Escherichia coli which allow rapid detection of inserted fragments. Gene 29: 21-26.

Sun, J.L., Zhang, S.K., Chen, J.Y., and Han, B.Z. (2012) Metabolic profiling of Staphylococcus aureus cultivated under aerobic and anaerobic conditions with (1)H NMR-based nontargeted analysis. Can J Microbiol 58: 709-718.

229

Surdel, M.C., Dutter, B.F., Sulikowski, G.A., and Skaar, E.P. (2016) Bacterial nitric oxide synthase is required for the Staphylococcus aureus response to heme stress. ACS Infect Dis 2: 572-578.

Suthers, P.F., Dasika, M.S., Kumar, V.S., Denisov, G., Glass, J.I., and Maranas, C.D. (2009) A Genome-Scale Metabolic Reconstruction of Mycoplasma genitalium, iPS189. PLoS Comput Biol 5: e1000285.

Switala, J., and Loewen, P.C. (2002) Diversity of properties among catalases. Arch Biochem Biophys 401: 145-154.

Tamir, S., deRojas-Walker, T., Wishnok, J.S., and Tannenbaum, S.R. (1996) DNA damage and genotoxicity by nitric oxide. Methods Enzymol 269: 230-243.

Tan, K., Li, H., Zhang, R., Gu, M., Clancy, S.T., and Joachimiak, A. (2008) Structures of open (R) and close (T) states of prephenate dehydratase (PDT)-implication of allosteric regulation by L-phenylalanine. J Struct Biol 162: 94-107.

Taylor, B.L., and Zhulin, I.B. (1999) PAS domains: internal sensors of oxygen, redox potential, and light. Microbiol Mol Biol Rev 63: 479-506.

Taylor, D., and Holland, K.T. (1989) Amino acid requirements for the growth and production of some exocellular products of Staphylococcus aureus. J Appl Bacteriol 66: 319-329.

Theodore, T.S., and Weinbach, E.C. (1974) Respiratory activities associated with mesosomal vesicles and protoplast membranes of Staphylococcus aureus. Journal of bacteriology 120: 562-564.

Thoendel, M., and Horswill, A.R. (2009) Identification of Staphylococcus aureus AgrD residues required for autoinducing peptide biosynthesis. J Biol Chem 284: 21828-21838.

Throup, J.P., Zappacosta, F., Lunsford, R.D., Annan, R.S., Carr, S.A., Lonsdale, J.T., Bryant, A.P., McDevitt, D., Rosenberg, M., and Burnham, M.K. (2001) The srhSR gene pair from Staphylococcus aureus: genomic and proteomic approaches to the identification and characterization of gene function. Biochemistry 40: 10392- 10401.

Thurlow, L.R., Joshi, G.S., Clark, J.R., Spontak, J.S., Neely, C.J., Maile, R., and Richardson, A.R. (2013) Functional modularity of the arginine catabolic mobile element contributes to the success of USA300 methicillin-resistant Staphylococcus aureus. Cell Host Microbe 13: 100-107.

230

Tinberg, C.E., Tonzetich, Z.J., Wang, H., Do, L.H., Yoda, Y., Cramer, S.P., and Lippard, S.J. (2010) Characterization of iron dinitrosyl species formed in the reaction of nitric oxide with a biological Rieske center. J Am Chem Soc 132: 18168-18176.

Todar, K., (2006) Todar's Online Textbook of Bacteriology. University of Wisconsin- Madison Department of Bacteriology.

Toledo, J.C., Jr., and Augusto, O. (2012) Connecting the chemical and biological properties of nitric oxide. Chem Res Toxicol 25: 975-989.

Torres, V.J., Pishchany, G., Humayun, M., Schneewind, O., and Skaar, E.P. (2006) Staphylococcus aureus IsdB is a hemoglobin receptor required for heme iron utilization. J Bacteriol 188: 8421-8429.

Tynecka, Z., and Malm, A. (1995) Intrinsic insensitivity to cadmium of the L-lactate oxidizing system in Staphylococcus aureus. FEMS microbiology letters 129: 11- 15.

Tynecka, Z., Szczesniak, Z., Malm, A., and Los, R. (1999) Energy conservation in aerobically grown Staphylococcus aureus. Res Microbiol 150: 555-566.

Udo, E.E., Pearman, J.W., and Grubb, W.B. (1993) Genetic analysis of community isolates of methicillin-resistant Staphylococcus aureus in Western Australia. J Hosp Infect 25: 97-108.

Ulrich, M., Bastian, M., Cramton, S.E., Ziegler, K., Pragman, A.A., Bragonzi, A., Memmi, G., Wolz, C., Schlievert, P.M., Cheung, A., and Doring, G. (2007) The staphylococcal respiratory response regulator SrrAB induces ica gene transcription and polysaccharide intercellular adhesin expression, protecting Staphylococcus aureus from neutrophil killing under anaerobic growth conditions. Mol Microbiol 65: 1276-1287.

Underbakke, E.S., Iavarone, A.T., Chalmers, M.J., Pascal, B.D., Novick, S., Griffin, P.R., and Marletta, M.A. (2014) Nitric oxide-induced conformational changes in soluble guanylate cyclase. Structure 22: 602-611.

Utsui, Y., and Yokota, T. (1985) Role of an altered penicillin-binding protein in methicillin- and cephem-resistant Staphylococcus aureus. Antimicrob Agents Chemother 28: 397-403.

Uziel, O., Borovok, I., Schreiber, R., Cohen, G., and Aharonowitz, Y. (2004) Transcriptional regulation of the Staphylococcus aureus thioredoxin and thioredoxin reductase genes in response to oxygen and disulfide stress. J Bacteriol 186: 326-334.

231

Vaish, M., and Singh, V.K. (2013) Antioxidant Functions of Nitric Oxide Synthase in a Methicillin Sensitive Staphylococcus aureus. Int J Microbiol 2013: 312146.

Valderas, M.W., Gatson, J.W., Wreyford, N., and Hart, M.E. (2002) The Superoxide Dismutase Gene sodM Is Unique to Staphylococcus aureus: Absence of sodM in Coagulase-Negative Staphylococci.

Valderas, M.W., and Hart, M.E. (2001) Identification and characterization of a second superoxide dismutase gene (sodM) from Staphylococcus aureus. J Bacteriol 183: 3399-3407.

Van Alst, N.E., Picardo, K.F., Iglewski, B.H., and Haidaris, C.G. (2007) Nitrate sensing and metabolism modulate motility, biofilm formation, and virulence in Pseudomonas aeruginosa. Infect Immun 75: 3780-3790. van der Merwe, D.E., Ubbink, J.B., Delport, R., Becker, P., Dhatt, G.S., and Vermaak, W.J. (2002) Biological variation in sweat sodium chloride conductivity. Ann Clin Biochem 39: 39-43. van Rotterdam, B.J., Westerhoff, H.V., Visschers, R.W., Bloch, D.A., Hellingwerf, K.J., Jones, M.R., and Crielaard, W. (2001) Pumping capacity of bacterial reaction centers and backpressure regulation of energy transduction. Eur J Biochem 268: 958-970. van Sorge, N.M., Beasley, F.C., Gusarov, I., Gonzalez, D.J., von Kockritz-Blickwede, M., Anik, S., Borkowski, A.W., Dorrestein, P.C., Nudler, E., and Nizet, V. (2013) Methicillin-resistant Staphylococcus aureus bacterial nitric-oxide synthase affects antibiotic sensitivity and skin abscess development. J Biol Chem 288: 6417- 6426.

Vandenesch, F., Kornblum, J., and Novick, R.P. (1991) A temporal signal, independent of agr, is required for hla but not spa transcription in Staphylococcus aureus. J Bacteriol 173: 6313-6320.

Veldkamp, K.E., Heezius, H.C., Verhoef, J., van Strijp, J.A., and van Kessel, K.P. (2000) Modulation of neutrophil chemokine receptors by Staphylococcus aureus supernate. Infect Immun 68: 5908-5913.

Vermeiren, C.L., Pluym, M., Mack, J., Heinrichs, D.E., and Stillman, M.J. (2006) Characterization of the heme binding properties of Staphylococcus aureus IsdA. Biochemistry 45: 12867-12875.

Villegas, J.M., Volentini, S.I., Rintoul, M.R., and Rapisarda, V.A. (2011) Amphipathic C- terminal region of Escherichia coli NADH dehydrogenase-2 mediates membrane localization. Arch Biochem Biophys 505: 155-159.

232

von Eiff, C., Becker, K., Metze, D., Lubritz, G., Hockmann, J., Schwarz, T., and Peters, G. (2001) Intracellular persistence of Staphylococcus aureus small-colony variants within keratinocytes: a cause for antibiotic treatment failure in a patient with darier's disease. Clinical infectious diseases : an official publication of the Infectious Diseases Society of America 32: 1643-1647. von Eiff, C., Heilmann, C., Proctor, R.A., Woltz, C., Peters, G., and Gotz, F. (1997) A site-directed Staphylococcus aureus hemB mutant is a small-colony variant which persists intracellularly. Journal of bacteriology 179: 4706-4712.

Vuong, C., Kidder, J.B., Jacobson, E.R., Otto, M., Proctor, R.A., and Somerville, G.A. (2005) Staphylococcus epidermidis polysaccharide intercellular adhesin production significantly increases during tricarboxylic acid cycle stress. J Bacteriol 187: 2967-2973.

Wach, M.J., Kers, J.A., Krasnoff, S.B., Loria, R., and Gibson, D.M. (2005) Nitric oxide synthase inhibitors and nitric oxide donors modulate the biosynthesis of thaxtomin A, a nitrated phytotoxin produced by Streptomyces spp. Nitric Oxide 12: 46-53.

Waheed, S.M., Ghosh, A., Chakravarti, R., Biswas, A., Haque, M.M., Panda, K., and Stuehr, D.J. (2010) Nitric oxide blocks cellular heme insertion into a broad range of heme proteins. Free radical biology & medicine 48: 1548-1558.

Wainwright, M. (2001) Acridine—a neglected antibacterial chromophore. Journal of Antimicrobial Chemotherapy 47: 1-13.

Wakeman, C.A., Hammer, N.D., Stauff, D.L., Attia, A.S., Anzaldi, L.L., Dikalov, S.I., Calcutt, M.W., and Skaar, E.P. (2012) Menaquinone biosynthesis potentiates haem toxicity in Staphylococcus aureus. Mol Microbiol 86: 1376-1392.

Walsh, E.J., O'Brien, L.M., Liang, X., Hook, M., and Foster, T.J. (2004) Clumping factor B, a fibrinogen-binding MSCRAMM (microbial surface components recognizing adhesive matrix molecules) adhesin of Staphylococcus aureus, also binds to the tail region of type I cytokeratin 10. J Biol Chem 279: 50691-50699.

Wang, B., and Muir, T.W. (2016) Regulation of virulence in Staphylococcus aureus: molecular mechanisms and remaining puzzles. Cell Chem Biol 23: 214-224.

Wang, Z.Q., Lawson, R.J., Buddha, M.R., Wei, C.C., Crane, B.R., Munro, A.W., and Stuehr, D.J. (2007) Bacterial flavodoxins support nitric oxide production by Bacillus subtilis nitric-oxide synthase. J Biol Chem 282: 2196-2202.

233

Wang, Z.Q., Wei, C.C., Sharma, M., Pant, K., Crane, B.R., and Stuehr, D.J. (2004) A conserved Val to Ile switch near the heme pocket of animal and bacterial nitric- oxide synthases helps determine their distinct catalytic profiles. J Biol Chem 279: 19018-19025.

Wang, Z.Q., Wei, C.C., and Stuehr, D.J. (2010) How does a valine residue that modulates heme-NO binding kinetics in inducible NO synthase regulate enzyme catalysis? J Inorg Biochem 104: 349-356.

Waters, N.R., Samuels, D.J., Behera, R.K., Livny, J., Rhee, K.Y., Sadykov, M.R., and Brinsmade, S.R. (2016) A spectrum of CodY activities drives metabolic reorganization and virulence gene expression in Staphylococcus aureus. Mol Microbiol 101: 495-514.

Watmough, N.J., Butland, G., Cheesman, M.R., Moir, J.W.B., Richardson, D.J., and Spiro, S. (1999) Nitric oxide in bacteria: synthesis and consumption. 1411: 456- 474.

West, L.C.C.F.p.d.J. (1916) The Cost of Living in Roman Egypt. 11: 293-314.

White, D., Drummond, J.T., and Fuqua, C., (1995) The physiology and biochemistry of prokaryotes. Oxford University Press New York.

Willenborg, J., and Goethe, R. (2016) Metabolic traits of pathogenic streptococci. FEBS Lett.

Williams, R.E. (1963) Healthy carriage of Staphylococcus aureus: its prevalence and importance. Bacteriol Rev 27: 56-71.

Windham, I.H., Chaudhari, S.S., Bose, J.L., Thomas, V.C., and Bayles, K.W. (2016) SrrAB modulates Staphylococcus aureus cell death through regulation of cidABC transcription. J Bacteriol 198: 1114-1122.

Winger, J.A., Derbyshire, E.R., and Marletta, M.A. (2007) Dissociation of nitric oxide from soluble guanylate cyclase and heme-nitric oxide/oxygen binding domain constructs. J Biol Chem 282: 897-907.

Wink, D.A., Cook, J.A., Kim, S.Y., Vodovotz, Y., Pacelli, R., Krishna, M.C., Russo, A., Mitchell, J.B., Jourd'heuil, D., Miles, A.M., and Grisham, M.B. (1997) Superoxide modulates the oxidation and nitrosation of thiols by nitric oxide-derived reactive intermediates. Chemical aspects involved in the balance between oxidative and nitrosative stress. The Journal of biological chemistry 272: 11147-11151.

234

Wink, D.A., Darbyshire, J.F., Nims, R.W., Saavedra, J.E., and Ford, P.C. (1993a) Reactions of the bioregulatory agent nitric oxide in oxygenated aqueous media: determination of the kinetics for oxidation and nitrosation by intermediates generated in the NO/O2 reaction. Chem Res Toxicol 6: 23-27.

Wink, D.A., Kasprzak, K.S., Maragos, C.M., Elespuru, R.K., Misra, M., Dunams, T.M., Cebula, T.A., Koch, W.H., Andrews, A.W., Allen, J.S., and et al. (1991) DNA deaminating ability and genotoxicity of nitric oxide and its progenitors. Science 254: 1001-1003.

Wink, D.A., Osawa, Y., Darbyshire, J.F., Jones, C.R., Eshenaur, S.C., and Nims, R.W. (1993b) Inhibition of cytochromes P450 by nitric oxide and a nitric oxide- releasing agent. Archives of biochemistry and biophysics 300: 115-123.

Wong, P.S., Eiserich, J.P., Reddy, S., Lopez, C.L., Cross, C.E., and van der Vliet, A. (2001) Inactivation of glutathione S-transferases by nitric oxide-derived oxidants: exploring a role for tyrosine nitration. Arch Biochem Biophys 394: 216-228.

Wooldridge, K.G., and Williams, P.H. (1993) Iron uptake mechanisms of pathogenic bacteria. FEMS Microbiol Rev 12: 325-348.

Wright, J.D., and Holland, K.T. (2003) The effect of cell density and specific growth rate on accessory gene regulator and toxic shock syndrome toxin-1 gene expression in Staphylococcus aureus. FEMS Microbiol Lett 218: 377-383.

Wu, R., Skaar, E.P., Zhang, R., Joachimiak, G., Gornicki, P., Schneewind, O., and Joachimiak, A. (2005) Staphylococcus aureus IsdG and IsdI, heme-degrading enzymes with structural similarity to monooxygenases. J Biol Chem 280: 2840- 2846.

Wu, Y., Zhu, T., Han, H., Liu, H., Xu, T., Francois, P., Fischer, A., Bai, L., Gotz, F., and Qu, D. (2015) Staphylococcus epidermidis SrrAB regulates bacterial growth and biofilm formation differently under oxic and microaerobic conditions. J Bacteriol 197: 459-476.

Xia, Y., and Zweier, J.L. (1997) Superoxide and peroxynitrite generation from inducible nitric oxide synthase in macrophages. Proceedings of the National Academy of Sciences 94: 6954-6958.

Xu, S.X., and McCormick, J.K. (2012) Staphylococcal superantigens in colonization and disease. Front Cell Infect Microbiol 2: 52.

Yarwood, J.M., Bartels, D.J., Volper, E.M., and Greenberg, E.P. (2004) Quorum sensing in Staphylococcus aureus biofilms. Journal of bacteriology 186: 1838-1850.

235

Yarwood, J.M., McCormick, J.K., and Schlievert, P.M. (2001) Identification of a novel two-component regulatory system that acts in global regulation of virulence factors of Staphylococcus aureus. Journal of bacteriology 183: 1113-1123.

Yarwood, J.M., Paquette, K.M., Tikh, I.B., Volper, E.M., and Greenberg, E.P. (2007) Generation of virulence factor variants in Staphylococcus aureus biofilms. J Bacteriol 189: 7961-7967.

Yermilov, V., Rubio, J., Becchi, M., Friesen, M.D., Pignatelli, B., and Ohshima, H. (1995) Formation of 8-nitroguanine by the reaction of guanine with peroxynitrite in vitro. Carcinogenesis 16: 2045-2050.

Yun, H.Y., Dawson, V.L., and Dawson, T.M. (1996) Neurobiology of nitric oxide. Crit Rev Neurobiol 10: 291-316.

Zhang, L., Jacobsson, K., Vasi, J., Lindberg, M., and Frykberg, L. (1998) A second IgG- binding protein in Staphylococcus aureus. Microbiology 144 ( Pt 4): 985-991.

Zheng, Y., Niyonsaba, F., Ushio, H., Nagaoka, I., Ikeda, S., Okumura, K., and Ogawa, H. (2007) Cathelicidin LL-37 induces the generation of reactive oxygen species and release of human alpha-defensins from neutrophils. Br J Dermatol 157: 1124-1131.

Zhou, M., Diwu, Z., Panchuk-Voloshina, N., and Haugland, R.P. (1997) A stable nonfluorescent derivative of resorufin for the fluorometric determination of trace hydrogen peroxide: applications in detecting the activity of phagocyte NADPH oxidase and other oxidases. Analytical biochemistry 253: 162-168.

Zhu, W., Clark, N., and Patel, J.B. (2013) pSK41-like plasmid is necessary for Inc18-like vanA plasmid transfer from Enterococcus faecalis to Staphylococcus aureus in vitro. Antimicrob Agents Chemother 57: 212-219.

Zhu, W., Clark, N.C., McDougal, L.K., Hageman, J., McDonald, L.C., and Patel, J.B. (2008) Vancomycin-resistant Staphylococcus aureus isolates associated with Inc18-like vanA plasmids in Michigan. Antimicrob Agents Chemother 52: 452- 457.

Zhu, Y., Xiong, Y.Q., Sadykov, M.R., Fey, P.D., Lei, M.G., Lee, C.Y., Bayer, A.S., and Somerville, G.A. (2009) Tricarboxylic acid cycle-dependent attenuation of Staphylococcus aureus in vivo virulence by selective inhibition of amino acid transport. Infect Immun 77: 4256-4264.

236

Zingarelli, B., O'Connor, M., Wong, H., Salzman, A.L., and Szabó, C. (1996) Peroxynitrite-mediated DNA strand breakage activates poly-adenosine diphosphate ribosyl synthetase and causes cellular energy depletion in macrophages stimulated with bacterial lipopolysaccharide. The Journal of Immunology 156: 350-358.

Zipfel, P.F. (2009) Complement and immune defense: from innate immunity to human diseases. Immunol Lett 126: 1-7.

Zong, Y., Xu, Y., Liang, X., Keene, D.R., Hook, A., Gurusiddappa, S., Hook, M., and Narayana, S.V. (2005) A 'Collagen Hug' model for Staphylococcus aureus CNA binding to collagen. Embo j 24: 4224-4236.

Zoraghi, R., See, R.H., Gong, H., Lian, T., Swayze, R., Finlay, B.B., Brunham, R.C., McMaster, W.R., and Reiner, N.E. (2010) Functional analysis, overexpression, and kinetic characterization of pyruvate kinase from methicillin-resistant Staphylococcus aureus. Biochemistry 49: 7733-7747.

Zumft, W.G. (2002) Nitric oxide signaling and NO dependent transcriptional control in bacterial denitrification by members of the FNR-CRP regulator family. J Mol Microbiol Biotechnol 4: 277-286.

237

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 238

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.

239