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SbnI is a free serine kinase and heme-sensing regulator required for staphyloferrin B biosynthesis in… Verstraete, Meghan Marie 2018

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SbnI is a free serine kinase and heme-sensing regulator required for staphyloferrin B biosynthesis in Staphylococcus aureus   by Meghan Marie Verstraete  B.Sc., The University of British Columbia, 2012  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Microbiology and Immunology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  June 2018  © Meghan Marie Verstraete, 2018    ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled:  SbnI is a free serine kinase and heme-sensing regulator required for staphyloferrin B biosynthesis in Staphylococcus aureus   submitted by Meghan Marie Verstraete  in partial fulfillment of the requirements for  the degree of Doctor of Philosophy  in Microbiology and Immunology  Examining Committee: Dr. Michael Murphy, Microbiology and Immunology Supervisor  Dr. Rachel Fernandez, Microbiology and Immunology Supervisory Committee Member   Supervisory Committee Member Dr. Katherine Ryan, Chemistry University Examiner Dr. Filip Van Petegem, Biochemistry University Examiner   Additional Supervisory Committee Members: Dr. Jim Kronstad, Microbiology and Immunology Supervisory Committee Member Dr. Lawrence McIntosh, Biochemistry Supervisory Committee Member   iii Abstract Staphylococcus aureus is a common member of the human microbiome, but is an opportunistic pathogen that can cause a variety of infections. Critical to the growth and survival of S. aureus during infection is acquisition of iron from the host. However, host iron availability is restricted to effectively suppress microbial growth as a type of innate, nutritional immunity. Mechanisms used by S. aureus to access the host iron pool include lysing erythrocytes to liberate hemoglobin for heme uptake and through the secretion of staphyloferrins, which are iron-chelating siderophores to scavenge iron from the host. The multiplicity of iron uptake systems S. aureus possesses is likely a reflection of the varied host environments that S. aureus can colonize or infect. However, the spatiotemporal regulatory mechanisms by which S. aureus adapts to changing iron availability over the course of infection are ill-defined. SbnI is a heme-dependent regulator of staphyloferrin B (SB) biosynthesis suggested to mediate between iron-uptake modes. In this thesis, study of the structure SbnI revealed homology to a free L-serine kinase, SerK, from Thermococcus kodakarensis. Biochemical assays and characterization of a serC mutant of S. aureus showed that SbnI is an ATP dependent L-serine kinase required for production of the SB precursor O-phospho-L-serine. SbnI kinase activity enables SB biosynthesis in environments where S. aureus catabolism is primarily reliant on amino acids, as in abscesses. Characterization of heme binding by SbnI and heme transfer reactions with IsdI and IsdG, two heme degrading enzymes, were used to construct a model of heme-binding by SbnI for regulating heme-SB uptake. This model is consistent with a modest effect of heme-binding on SbnI kinase activity. Heme transfer rates were measured from ChdC, the terminal enzyme in heme biosynthesis, to SbnI, IsdG, and IsdI to delineate an intracellular network of heme sensing and trafficking proteins that are likely required for regulation and adaptation to the dynamic host environment.    iv Lay Summary Staphylococcus aureus is a prominent human pathogen and a common cause of bacterial infections ranging in severity from minor skin and soft tissue infections to more serious conditions like infectious endocarditis and sepsis. The increased frequency of S. aureus infections in hospital settings among immunocompromised patients and in community settings affecting healthy individuals has prompted research to investigate the molecular mechanisms that enable it to be a formidable human pathogen. A critical requirement for S. aureus to successfully establish infection is the uptake of nutrient iron from the host, rendering iron metabolic pathways as potential drug targets. One method of iron acquisition by S. aureus is through the secretion of siderophores, which are iron–chelating molecules used to retrieve host iron. The work presented in this thesis investigated the biosynthesis and regulation of the siderophore staphyloferrin B and revealed how it can be produced by S. aureus in abscesses.     v Preface Part of the work presented in this thesis is published or drawn from a manuscript under preparation. The following contributions were made by fellow scientists and collaborators: Chapter 3 A version of Chapter 3 is published.  Verstraete, M.M., Perez-Borrajero, C., Brown, K.L., Heinrichs, D.E., and Murphy, M.E.P. (2018) SbnI is a free serine kinase that generates O-phospho-L-serine for staphyloferrin B biosynthesis in Staphylococcus aureus. Journal of Biological Chemistry. In Press. C. Perez-Borrajero performed the 31P NMR study and K.L. Brown conducted HPLC analysis. I optimized the protein expression and purification for SbnI, performed site-directed mutagenesis, crystallography experiments, UV-visible spectroscopic analyses, kinetic assays, bacterial growth curves, and siderophore bioassay experiments. I wrote the first draft of the manuscript. Dr. D.E. Heinrichs contributed to the initial concept of the project and provided manuscript edits. Dr. M.E.P. Murphy was the principal investigator and was involved throughout the project in concept formation and manuscript edits. Chapter 4 Chapter 4 is a version of a manuscript in preparation.  Verstraete, M.M., Kobylarz M.J., Loutet, S.A., Laakso, H.A., Heinrichs, D.E., and Murphy, M.E.P. SbnI has two distinct roles in staphyloferrin B biosynthesis in Staphylococcus aureus. Manuscript in preparation.  Dr. M.J. Kobylarz and Dr. S.A. Loutet provided advice to guide UV-visible spectroscopy experiments with IsdI. H.A. Laakso produced the SpSbnI expression plasmid. I performed protein expression and purification, site-directed mutagenesis, crystallography experiments,   vi stopped-flow kinetic analyses, UV-visible spectroscopy, and fluorescence spectroscopy. I wrote the first draft of the manuscript. Dr. M.E.P. Murphy was the principal investigaor and was involved throughout the project in concept formation.  Chapter 5 I performed all experiments and analysis of the research data described in Chapter 5. Dr. M.E.P. Murphy was the principal investigator and was involved throughout the project in concept formation.  This project required Biohazard Approval for the handling of Staphylococcus aureus and Escherichia coli and was issued by the UBC Biosafety Committee, Certificate number B13-0096.    vii Table of Contents  Abstract ..................................................................................................................................... ii Lay Summary ........................................................................................................................... iv Preface ....................................................................................................................................... v Table of Contents .................................................................................................................... vii List of Tables .......................................................................................................................... xiii List of Figures ........................................................................................................................ xiv List of Abbreviations ............................................................................................................ xvii Acknowledgements ................................................................................................................. xx Chapter 1: Introduction............................................................................................................ 1 1.1 Staphylococcus aureus ............................................................................................. 1 1.2 Iron as a nutritional requirement .............................................................................. 4 1.2.1 Nutritional immunity ........................................................................................ 5 1.2.2 S. aureus iron-sparing response ........................................................................ 6 1.3 Iron and heme uptake by S. aureus........................................................................... 7 1.3.1 S. aureus heme acquisition ............................................................................... 8 1.3.2 S. aureus non-heme iron uptake systems ........................................................ 11 1.4 Staphyloferrin biosynthesis in S. aureus ................................................................. 13 1.4.1 Staphyloferrin A ............................................................................................ 13 1.4.2 Staphyloferrin B ............................................................................................. 14 1.5 S. aureus iron homeostasis and regulation .............................................................. 17 1.5.1 Intracellular iron metabolism and management .............................................. 17   viii 1.5.2 S. aureus iron management in response to oxidative stress ............................. 18 1.6 S. aureus heme homeostasis and regulation ............................................................ 20 1.6.1 Heme biosynthesis ......................................................................................... 20 1.6.2 Intracellular heme metabolism and management ............................................ 24 1.6.3 Heme degradation .......................................................................................... 25 1.7 Spatiotemporal expression of iron uptake systems by S. aureus in the host ............ 26 1.7.1 S. aureus iron source preference ..................................................................... 28 1.8 Objectives .............................................................................................................. 29 Chapter 2: Methods ................................................................................................................ 32 2.1 Cloning, expression, and protein purification for biochemical assays and structure determination ................................................................................................................... 32 2.1.1 Cloning, expression, and purification of SbnI and SbnI variants ..................... 33 2.1.2 Cloning, expression, and purification of S. pseudintermedius SbnI ................. 35 2.1.3 Cloning, expression, and purification of ChdC ............................................... 36 2.1.4 Cloning, expression, and purification of IsdI .................................................. 37 2.1.5 Expression and purification of SbnA .............................................................. 38 2.1.6 Expression and purification of IruO ............................................................... 38 2.1.7 Expression and purification of IsdG ............................................................... 39 2.2 Crystallization, data collection, and structure determination ................................... 39 2.2.1 SbnI1-240 structure determination .................................................................... 39 2.2.2 SpSbnI structure determination ...................................................................... 40 2.2.3 ChdC structure determination ......................................................................... 43 2.3 Bioinformatic analysis ........................................................................................... 45   ix 2.3.1 Genomic neighborhood analysis of SbnI and homologs .................................. 45 2.3.2 SbnI1-240 structure and conservation analysis .................................................. 45 2.3.3 SpSbnI conservation and molecular surface electrostatics analyses ................ 46 2.3.4 Generation of model of full-length S. aureus SbnI and heme-binding ............. 46 2.3.5 ChdC structural superimposition and analysis ................................................ 46 2.4 Heme reconstitution of proteins ............................................................................. 47 2.5 Determination of SbnI oligomeric state in solution................................................. 47 2.6 UV-visible spectroscopic analysis of SbnI OPS production using SbnA ................. 48 2.7 Heme-binding by SbnI and variants ....................................................................... 48 2.8 UV-visible spectroscopic analysis of oxidized, reduced, and CO bound forms of SbnI and SpSbnI bound to heme ...................................................................................... 48 2.9 UV-visible spectroscopic analysis of heme transfer reactions ................................. 49 2.9.1 Heme transfer from IsdI to SbnI and variants ................................................. 49 2.9.2 Heme transfer from IsdI to SbnI in the presence of IruO................................. 49 2.9.3 Heme transfer from ChdC to IsdI, IsdG, and SbnI .......................................... 50 2.9.4 Heme transfer from IsdG to SbnI ................................................................... 50 2.10 IsdI heme transfer to SbnI pulldown assay ......................................................... 50 2.11 Stopped-flow kinetic analysis of enzyme heme off-rate ...................................... 51 2.12 Stopped-flow kinetic analysis of heme transfer reactions.................................... 52 2.12.1 IsdI heme transfer to SbnI .............................................................................. 52 2.12.2 IsdG heme transfer to SbnI ............................................................................. 52 2.12.3 ChdC heme transfer to SbnI, IsdI, and IsdG ................................................... 53 2.13 Fluorescence quenching of SbnI......................................................................... 53   x 2.14 HPLC analysis of SbnI kinase activity ............................................................... 54 2.15 31P NMR spectra of SbnI kinase reaction ........................................................... 54 2.16 Steady-state kinetic analysis of SbnI serine kinase activity ................................. 55 2.16.1 Pyruvate kinase/lactate dehydrogenase coupled assay .................................... 55 2.16.2 SbnA and phosphate release coupled assay .................................................... 56 2.16.3 Software for kinetic analysis .......................................................................... 57 2.17 S. aureus bacterial strains ................................................................................... 57 2.18 S. aureus bacterial growth curve to assess serine auxotrophy ............................. 58 2.19 Disc diffusion bioassays to detect siderophore production .................................. 58 Chapter 3: SbnI is a free serine kinase that generates O-phospho-L-serine for staphyloferrin B biosynthesis in S. aureus ............................................................................. 60 3.1 Introduction ........................................................................................................... 60 3.2 Results ................................................................................................................... 62 3.2.1 Structure determination of SbnI ...................................................................... 62 3.2.2 SbnI is a dimer in solution .............................................................................. 69 3.2.3 SbnI is a serine kinase that uses L-serine and ATP to generate OPS ............... 70 3.2.4 SbnI active site variants.................................................................................. 74 3.2.5 Kinetic analysis of SbnI kinase activity .......................................................... 74 3.2.6 Physiological function of an ATP-dependent serine kinase activity in S. aureus .   ...................................................................................................................... 77 3.3 Discussion ............................................................................................................. 79 Chapter 4: SbnI has two distinct roles in staphyloferrin B biosynthesis in S. aureus ........... 85 4.1 Introduction ........................................................................................................... 85   xi 4.2 Results ................................................................................................................... 86 4.2.1 Heme binding by SbnI does not greatly hinder L-serine kinase activity .......... 86 4.2.2 Structure of SpSbnI and SpSbnI bound to ATP .............................................. 87 4.2.3 Conservation and molecular surface electrostatics analyses of the SpSbnI structure ...................................................................................................................... 93 4.2.4 Spectroscopic characterization of heme coordination structure of S. aureus SbnI and SpSbnI .................................................................................................................. 96 4.2.5 SbnI heme affinity and heme off-rate ............................................................. 99 4.2.6 Oligomeric state of heme-bound SbnI .......................................................... 101 4.2.7 Site-directed mutagenesis to probe SbnI heme-binding mode ....................... 102 4.2.8 Model of heme-bound SbnI .......................................................................... 104 4.2.9 IsdI can catalytically transfer heme to SbnI .................................................. 108 4.2.10 SbnI1-240, SbnI H3A, SbnI C244A, and SbnI H3A/C244A are deficient in accepting heme transferred from IsdI compared to wildtype SbnI .............................. 109 4.2.11 IsdI heme transfer to SbnI outcompetes heme degradation ........................... 113 4.3 Discussion ........................................................................................................... 115 Chapter 5: Role of the terminal heme biosynthetic enzyme, ChdC, in heme homeostasis and heme-trafficking .................................................................................................................... 121 5.1 Introduction ......................................................................................................... 121 5.2 Results ................................................................................................................. 123 5.2.1 Structure of S. aureus ChdC ......................................................................... 123 5.2.2 Structural comparison of ChdC homologs .................................................... 129 5.2.3 GsChdC structure forms a 60-mer by crystallographic symmetry ................. 132   xii 5.2.4 Low resolution crystal structure of SaChdC crystallized in the presence of heme  .................................................................................................................... 134 5.2.5 Determination of ChdC:heme off-rate .......................................................... 136 5.2.6 ChdC can actively transfer heme to SbnI and IsdG, but not IsdI ................... 137 5.2.7 IsdG can transfer heme to SbnI .................................................................... 139 5.3 Discussion ........................................................................................................... 141 Chapter 6: Conclusion .......................................................................................................... 148 6.1 SbnI provides an alternate route for OPS synthesis in S. aureus ........................... 149 6.2 Defining the function of SbnI is critical to understanding the spatiotemporal production of SB............................................................................................................ 150 6.3 ChdC participates in an intracellular heme transfer network ................................. 153 6.4 Concluding remarks ............................................................................................. 155 References ............................................................................................................................. 157    xiii List of Tables  Table 1-1 Distribution of iron transport related genes identified in eight staphylococcal speciesa. 8 Table 2-1 E. coli strains and plasmids used in this study............................................................ 32 Table 2-2 Primers used for site-directed mutagenesis of S. aureus sbnI active site and putative heme-binding site. ..................................................................................................................... 34 Table 2-3 X-ray diffraction data collection and refinement statistics for SbnI1-240. ..................... 40 Table 2-4 X-ray diffraction data collection and refinement statistics for SpSbnI structures. ....... 42 Table 2-5 X-ray diffraction data collection and refinement statistics for ChdC structures. ......... 44 Table 2-6 S. aureus strains used in this study. ........................................................................... 58 Table 3-1 Apparent steady-state kinetic parameters of SbnI and SbnI1-240. ................................. 75 Table 4-1 Apparent steady-state kinetic parameters of the L-serine kinase reaction by heme-bound SbnI................................................................................................................................ 87 Table 5-1 ChdC structures, oligomerizations state, and area of protomer interfaces. ................ 126 Table 5-2 Kinetic parameters of ChdC:heme off-rate and ChdC:heme transfer to SbnI, IsdG, and IsdI. ........................................................................................................................................ 138 Table 5-3 Comparison of kinetic parameters of IsdG and IsdI heme off-rates and heme transfer to SbnI. ....................................................................................................................................... 140    xiv List of Figures  Figure 1-1 Schematic of heme uptake and degradation by the Isd system in S. aureus. .............. 10 Figure 1-2 Physical map of the sbn genetic locus and SB biosynthetic pathway. ....................... 16 Figure 1-3 Bacterial heme biosynthesis. .................................................................................... 22 Figure 1-4 Schematic of SbnI regulation of SB biosynthesis and SB-mediated iron acquisition. 29 Figure 2-1 31P NMR spectra of ATP and OPS standards............................................................ 55 Figure 3-1 Metabolic pathways for the production of L-Dap and a-KG from glucose or L-serine in S. aureus. .............................................................................................................................. 61 Figure 3-2 Structure of SbnI1-240. ............................................................................................... 63 Figure 3-3 Sequence alignment of SbnI with SerK and SbnI homolog proteins generated using T-Coffee Expresso. ....................................................................................................................... 66 Figure 3-4 Illustration of gene neighborhoods containing SbnI homologs from diverse species from Firmicute and Proteobacteria phyla. .................................................................................. 68 Figure 3-5 Conservation of surface residues of SbnI1-240 generated using ConSurf. ................... 69 Figure 3-6 Representative DLS results for analysis of SbnI and SbnI1-240 oligomerization state. 70 Figure 3-7 Detection of O-phospho-L-serine produced by SbnI via reaction with SbnA. ........... 71 Figure 3-8 HPLC analysis of the nucleotide reaction products. .................................................. 72 Figure 3-9 A stack plot of 31P NMR spectra for a single reaction of SbnI mediated conversion of L-serine and ATP to OPS and ADP. .......................................................................................... 73 Figure 3-10 Plots of initial velocities used for determination of kinetic constants for SbnI and SbnI1-240. ................................................................................................................................... 76 Figure 3-11 S. aureus serC mutant is a serine auxotroph but can produce SB. ........................... 78   xv Figure 4-1 Plots of initial velocities versus substrate concentration used for determination of kinetic constants for SbnI bound to equimolar heme.................................................................. 86 Figure 4-2 Overall structure of SpSbnI. ..................................................................................... 88 Figure 4-3 SpSbnI disulfide bonds and conservation with S. aureus SbnI. ................................. 90 Figure 4-4 Structure of SpSbnI bound to ADP........................................................................... 92 Figure 4-5 Active site of SpSbnI and ADP co-crystal structure with OPS modelled. ................. 92 Figure 4-6 Amino acid conservation and surface electrostatics of SpSbnI.................................. 94 Figure 4-7 Multiple sequence alignment of staphylococcal SbnI homologs. .............................. 95 Figure 4-8 UV-visible spectra of heme-bound SbnI and heme-bound SpSbnI in the oxidized, reduced, and CO bound forms. .................................................................................................. 97 Figure 4-9 SbnI heme affinity and heme off-rate. .................................................................... 100 Figure 4-10 Representative DLS results for analysis of heme-bound SbnI oligomerization state. ............................................................................................................................................... 101 Figure 4-11 UV-visible spectroscopic analysis of heme binding by SbnI variants. ................... 103 Figure 4-12 Models of heme binding by SpSbnI and a full-length model of S. aureus SbnI dimer. ............................................................................................................................................... 106 Figure 4-13 Superimposition of SpSbnI models bound to heme, ADP, and OPS. .................... 107 Figure 4-14 Stopped-flow kinetic data of IsdI heme transfer to SbnI and measurement of the IsdI heme off-rate........................................................................................................................... 109 Figure 4-15 IsdI heme transfer to SbnI and SbnI variants. ....................................................... 110 Figure 4-16 Quantification of IsdI heme transfer to SbnI and SbnI variants. ............................ 112 Figure 4-17 IsdI heme transfer to SbnI and SbnI variants using the pull-down assay. .............. 113   xvi Figure 4-18 IsdI heme transfer or degradation competition experiment between SbnI and IruO. ............................................................................................................................................... 115 Figure 5-1 Structure of S. aureus ChdC. .................................................................................. 124 Figure 5-2 Superimposition of apo and ligand-bound forms of G. stearothermophilus ChdC with S. aureus ChdC. ...................................................................................................................... 128 Figure 5-3 Conservation of SaChdC residues generated using ConSurf. .................................. 129 Figure 5-4 Structural overview of ChdC protomers. ................................................................ 131 Figure 5-5 Structure of GsChdC 60-mer and 60-mer multimerization interface. ...................... 133 Figure 5-6 GsChdC 60-mer multimerization encloses the active site. ...................................... 134 Figure 5-7 Structure of S. aureus ChdC co-crystallized with heme .......................................... 136 Figure 5-8 ChdC:heme transfer to SbnI, IsdG, and IsdI. .......................................................... 137 Figure 5-9 IsdG heme transfer to SbnI. .................................................................................... 140 Figure 5-10 Model of how ChdC, SbnI, IsdI, and IsdG participate in heme trafficking in S. aureus. .................................................................................................................................... 146    xvii List of Abbreviations  a-KG ACEGA Acetyl-CoA ADP ALA AMP AmpR ATP CA-MRSA  CDMG CDMG-L-Ser  CLS DLS EDDHA EDTA Fur His6 Isd a-ketoglutarate N-(1-amino-1-carboxyl-2-ethyl)-glutamic acid Acetyl coenzyme A Adenosine diphosphate d-amino-levulinic acid Adenosine monophosphate Ampicillin resistance Adenosine triphosphate Community-associated methicillin resistant Staphylococcus aureus Chemically-defined medium with 0.4% (w/v) glucose Chemically-defined medium with 0.4% (w/v) glucose lacking L-serine  Canadian Light Source Dynamic light scattering Ethylenediamine-N,N’-bis(2-hydroxyphenylacetic acid) Ethylenediaminetetraacetic acid Ferric uptake regulator Hexahistidine affinity purification tag Iron-regulated surface determinant   xviii KmR HEPES HPLC IPTG L-DAP MRSA NAD+ NADH NADPH NIS NMR OPS PDB PEG PK/LDH PLP RMSD ROS SA SB SCV SDS-PAGE Kanamycin resistance 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid High performance liquid chromatography Isopropyl β-D-thiogalactopyranoside L-2,3-diaminopropionic acid Methicillin resistant Staphylococcus aureus Nicotinamide adenine dinucleotide in the oxidized form Nicotinamide adenine dinucleotide in the reduced form Nicotinamide adenine dinucleotide phosphate in the reduced form Non-ribosomal peptide synthetase-independent siderophore  Nuclear magnetic resonance O-phospho-L-serine Protein data bank Poly(ethylene glycol) Pyruvate kinase lactate dehydrogenase Pyridoxal 5’-phosphate Root mean square deviation Reactive oxygen species Staphyloferrin A Staphyloferrin B Small colony variant Sodium dodecyl sulfate polyacrylamide gel electrophoresis   xix SSRL TCA TCEP TMS 2xYT Stanford Synchrotron Radiation Lightsource Tricarboxylic acid Tris(2-carboxyethyl)phosphine Tris minimal succinate 2x yeast extract tryptone     xx Acknowledgements  I thank Dr. Michael Murphy for giving me the opportunity to be a graduate student in his laboratory. Thank you for your support throughout the ups and downs of my thesis project and reminding me that in the words of Louis Pasteur “in the realm of scientific observation, luck is granted only to those who are prepared”. Thank you to my committee members, Dr. Rachel Fernandez, Dr. Jim Kronstad, and Dr. Lawrence McIntosh, for your guidance on my project and encouragement during our meetings.  I would like to acknowledge funding of this work from a Natural Sciences and Engineering Research Council Post-Graduate Scholarship, as well as project funding from the Canadian Institutes of Health Research held by Dr. Michael Murphy. I offer my enduring gratitude to all of those who contributed to my research. Especially Dr. Anson Chan for tirelessly answering my crystallography questions and Angele Arrieta for maintaining a great laboratory environment to work in. I also want to thank Dr. Lindsay Eltis for access to his spectrophotometers and advice.  Thank you to our research collaborator Dr. David Heinrichs for your advice and technical assistance. The progress I made on my thesis project was greatly helped by your expert contributions. Special thanks to past and present members of the Murphy lab for the scientific and emotional support over the years. My time in the Murphy lab was very enjoyable thanks to you. Last, but not least, I thank my family and friends for their constant love and support. I couldn’t have made it through this process without your words of encouragement, adventures to the mountains, and sunny trips to the desert.     1 Chapter 1: Introduction  1.1 Staphylococcus aureus Staphylococcus aureus is a prominent human pathogen that also asymptomatically colonizes a proportion of the human population (1, 2). Though colonization is typically not harmful to the host, S. aureus is frequently associated with minor skin and soft tissue infections (3). Additionally, this bacteria is capable of breaching host innate immune responses to gain access to deep tissues causing more severe and invasive infections, including bacteremia, endocarditis, osteomyelitis, and necrotizing pneumonia (2, 4). Study of the mechanisms of S. aureus pathogenesis and host evasion have come to the forefront of research due to the emergence of antibiotic-resistant strains. Methicillin-resistant S. aureus (MRSA) and the recent evolution of hypervirulent, community-associated MRSA (CA-MRSA) can be clinically challenging to treat as therapeutic options are more limited (5). Two clear shifts in the epidemiology of S. aureus infections have occurred in the past decade. First, there has been a growing number of health-care associated infections, particularly in infective endocarditis and prosthetic joint infections, and second, an epidemic of skin and soft tissue infections driven by CA-MRSA strains has occurred (6, 7). USA300 is a dominant CA-MRSA lineage that causes the majority of skin and soft tissue infections in North America (5, 8). The rapid increase in prevalence of USA300 lineage strains is thought to be in part due to the carriage of the arginine catabolic mobile element, a distinguishing genomic feature of these strains acquired by horizontal gene transfer from the skin commensal Staphylococcus epidermidis (9). This genetic element promotes survival by providing resistance to polyamines produced on skin that are otherwise toxic to related S. aureus   2 strains and confers acid tolerance during skin colonization (9, 10). This locus likely provides significant selective advantage during skin colonization and infections, explaining the evolutionary success of this lineage. A fundamental biological property of S. aureus is its ability to asymptomatically colonize healthy individuals. S. aureus infections most often originate from colonizing flora present on the infected host (11). Accordingly, S. aureus can exist as a commensal or be responsible for invasive disease. This duplicitous lifestyle requires the controlled expression of a diverse array of virulence factors as it transitions between these radically different states. S. aureus integrates environmental signals, host stimuli, and cell density as determined through a quorum-sensing signal as a means to trigger these phenotypic changes (12). Many staphylococcal virulence determinants are controlled by the interdependent global regulators Agr and SarA, including adhesins, hemolysins, proteases, and superantigenic toxins (13, 14). S. aureus can produce leukotoxins like Panton-Valentine Leukocidin (PVL) and cytolytic peptides including phenol soluble modulins (PSMs) and the pore-forming hemolysin (a-toxin, Hla) to destroy host cells and lead to an exaggerated inflammatory response (15, 16). A molecular mechanism attributed to the increased virulence exhibited by MRSA strains is the increased regulatory control and production of these cytotoxins (17). S. aureus also has mechanisms to evade immune cell clearance and promote bacterial survival. Collectively, many virulence factors facilitate tissue inflammation and destruction, and impair immune cell function to promote S. aureus pathogenesis. S. aureus is a non-motile, facultative anaerobic, Gram-positive coccus belonging to the Firmicute phylum. The coagulase test was historically used to distinguish S. aureus from commensal staphylococci. Coagulase is an extracellular enzyme that enables conversion of   3 fibrinogen to fibrin and promotes fibrin clot formation, which can promote virulence of the bacterium by evasion of phagocytosis and other immune defenses (49). A hallmark of S. aureus colonies is a characteristic yellow color due to the production of a unique membrane-embedded golden carotenoid, staphyloxanthin, which serves as an important antioxidant (18). S. aureus is predominantly characterized as an extracellular pathogen but it can also adopt an intracellular lifestyle contributing to difficult to treat, recurrent infections (19). Small colony variants (SCVs) are a S. aureus subpopulation that result from a phenotypic switch and are particularly well adapted to the survival in host cells and are clinically associated with persistent infections (20, 21). Most SCVs share common phenotypes, including: small colonies on agar plates, impaired growth rates, increased antibiotic resistance, decreased expression of secreted proteins, and up-regulation of adhesins (22). All these features lead to the attenuated virulence profile of SCVs but with an improved capacity for causing persistent infection. Slow growth and small colony size are due to defects in respiration, usually caused by genetic mutation in the biosynthesis of cofactors for respiration, such as menaquinone or heme (19). Increased resistance to antibiotic treatment is thought to be due to the defect in respiration causing a low membrane potential and reducing antibiotic uptake. Phenotypic switching enables the bacteria to hide inside host cells as an insurance policy against adverse environmental conditions, such as physical stress, nutrient limitation, and/or antibiotic treatment. Both intracellular survival and extracellular growth are important to the pathogenesis of S. aureus infections. A complicating facet of S. aureus research is the use of different strains throughout the research field. Current models of global virulence are based predominantly on methicillin-sensitive S. aureus (MSSA) laboratory strain NCTC 8325-4 (RN450) and widely used derivative RN6390. MSSA strain Newman is currently used extensively in animal infection models because   4 of its robust virulence phenotype (23). Hospital-associated MRSA strain COL has become more popular for in vitro and in vivo studies, along with the more virulent and invasive CA-MRSA strain USA300, because of their prevalence in current clinical infections (5, 24–26). Problems arise due to inconsistencies between the genetic potential of different strains that may not be entirely representative of the events that occur in clinical isolates.  S. aureus is a formidable pathogen with metabolic flexibility that allows it to infect diverse tissues and cause a spectrum of diseases. The continuous evolution and growing prevalence of antibiotic resistant strains underlines the need to define the molecular mechanisms of virulence so that novel therapeutics can be developed. Due to the rapid development and breadth of research, a truly comprehensive review of S. aureus pathogenesis is beyond the scope of this thesis. Instead, the remainder of the Chapter will provide an introduction to the iron uptake, metabolism, and homeostasis strategies used by S. aureus and their contributions during pathogenesis.  1.2 Iron as a nutritional requirement One commonality for all forms of life is the essentiality of iron, with only a few identified exceptions (27, 28). The indispensable nature of iron is due to its ability to participate in single electron transfers as it converts between reduced ferrous (Fe2+) and oxidized ferric (Fe3+) redox states with a reduction potential favourable for carrying out biological enzymatic reactions. Iron can be incorporated into enzymes either as free ions, heme, or iron-sulfur clusters. The biological utility of iron is made evident by its involvement in many key metabolic processes including amino acid synthesis, tricarboxylic acid (TCA) cycle activity, DNA replication, cellular respiration, and electron transport. The benefits of iron must be balanced with potential toxicity   5 as it can catalyze Fenton reactions, generating reactive oxygen intermediates like hydroxyl radicals, hydrogen peroxide, and superoxide anions. These reactive species can cause damage to cellular lipids, proteins, and DNA (29). Due to the potential hazards of this biometal, it is imperative that organisms, microbes and humans alike, maintain iron homeostasis. Bacteria generally require 10-5-10-7 M iron to support growth (30). However, aerobic environments limit the bioavailability of iron due to its propensity to form ferric oxyhydroxide precipitates. Consequently, the concentration of free bioavailable iron in solution is in the range of 10-8-10-9 M, below the concentration necessary to support microbial growth (30). Pathogenic microbes must contend with even stricter iron deprivation imposed by the human host. The host goes to tremendous lengths to control free iron levels not only to suppress the generation of iron-catalyzed reactive oxygen species, but to limit the bioavailability of nutrient iron to invading pathogens as a type of innate, nutritional immunity (31). The concentration of free iron in extracellular fluids in humans is estimated to be ~10-24 M (32, 33).  1.2.1 Nutritional immunity The human body contains about 45 – 75 mg of iron per kg to satisfy its metabolic needs (34). Regulation of iron metabolism in mammals occurs at the systemic and cellular levels. For a comprehensive review of iron metabolism in humans see references (35, 36). Briefly, ingested iron is absorbed in the duodenum where it is reduced to ferrous iron in the brush border of enterocytes. Once internalized, ferrous iron can be used by the cell, stored, or exported through ferroportin (37). Exported iron is promptly bound by the plasma glycoprotein, transferrin, to be transported around the body to target cells for use by metalloenzymes, for heme biosynthesis, or stored intracellularly in ferritin. Tight regulation of iron metabolism is advantageous for humans   6 as it can prevent damage associated with iron overload and, conversely, anemia due to iron deficiency. Also, it serves as a method to protect against microbial pathogens by rendering iron scarcely available as a type of nutritional immunity, as mentioned previously. Due to the importance of iron, the study of nutritional immunity has been strongly focused on iron-withholding strategies. However, the host can also restrict access to other nutrient transition metals, namely copper, manganese, and zinc (see (31, 38, 39) for reviews). The majority of host iron is found intracellularly as heme bound to hemoglobin within erythrocytes (35). The high-affinity iron binding capabilities of transferrin and lactoferrin, which is abundant in mucosal secretions, function to maintain extremely low extracellular iron concentrations. Iron distribution in the body is regulated to ensure the amount of extracellular iron is kept to a minimum, limiting the amount available to invading pathogens. Furthermore, the body can strengthen iron-withholding defenses when infection is sensed by eliciting a hypoferremic response. Inflammation triggers the release of the peptide hormone, hepcidin, from the liver which works to decrease iron export into the blood (40). Iron can further be sequestered at infectious foci by lactoferrin that is released in neutrophil granules (41). Host iron-withholding strategies make an important contribution to the outcome of host-pathogen interactions. In response to this type of evolutionary arms race for iron, microbes have evolved mechanisms to circumnavigate host-inflicted iron starvation and access host iron sources.  1.2.2 S. aureus iron-sparing response S. aureus has proven to be capable of infecting a variety of host niches, including skin, soft tissue, respiratory, bone, joint, and endovascular tissue (4). Despite the differences in these tissue tropisms, one key environmental condition shared amongst them is iron limitation. Iron   7 deprivation triggers gene expression and metabolic changes to elevate iron scavenging. In S. aureus, the iron-binding protein Fur (ferric uptake regulator) mediates this response through derepression of several iron acquisition systems and by modulating the expression of different virulence factors (42, 43). Fur also coordinates a metabolic rearrangement, termed the iron-sparing response, to decrease the iron demands of the cell by reducing expression of non-essential iron-containing pathways (44–46). Concomitant up-regulation of glycolytic and fermentative pathways allows for suppression of the TCA cycle, which relies on many iron-containing enzymes (44). These iron-sparing metabolic changes also favor the production of acidic metabolites that decrease local pH in the area of infection. The acidic microenvironment in turn favors S. aureus, as it improves the efficacy of iron chelation from host iron-binding proteins, like transferrin, by siderophores (46).  1.3 Iron and heme uptake by S. aureus Successful iron uptake from the human host is integral to infection and pathogenesis of S. aureus and most other microbial pathogens (47, 48). Iron starvation is a major sensory cue to many pathogens upon entering the host that can trigger transformation from a colonizing to invasive phenotype. Some iron uptake systems are highly conserved between different species of staphylococci, while others are constrained to specific lineages and make up some of the defining characteristics that differentiate commensal from highly invasive species of staphylococci. The success of S. aureus as a major colonizer throughout the human body has been attributed, at least in part, to the breadth of iron uptake systems it possesses. By comparison to other coagulase positive staphylococcal species, S. aureus stands out in the number of systems it possesses dedicated to heme and iron uptake (Table 1-1). S. aureus has the capacity to exploit   8 a variety of iron sources within the host, including transferrin and lactoferrin, free inorganic iron, heme, and hemoproteins to satisfy nutritional needs. Numerous studies have demonstrated the importance of iron acquisition to S. aureus pathogenesis (50–52). Outlined in the following sections are the major systems employed by S. aureus to achieve iron uptake (for comprehensive reviews see (47, 53, 54)).  Table 1-1 Distribution of iron transport related genes identified in eight staphylococcal speciesa.  Product Gene names Sa Sp Sd Si Se Ss Sh Sc Staphyloferrin B production sbnABCDEFGHI + + + + – – – – Staphyloferrin B ABC transporter sirABC + + + + – – – – Staphyloferrin A production sfaABCD + + + + + + + + Staphyloferrin A ABC transporter htsABC + + + + + + + + Transcriptional repressor of iron uptake fur + + + + + + + + Catechol-type siderophore uptake  sstABCD + – – – + + + + Hydroxomate-type siderophore uptake fhuCBG + + + + – + + + Lipoprotein receptor for hydroxymate-type siderophores fhuD + – – – – + + + Iron-responsive surface determinant (heme uptake) isdABCDEFG, srtB + – – – – – – – Iron-manganese ABC transporter mntABC (sitABC)b + + –c + + + – + Heme-regulated ABC transporter (heme detoxification) hrtAB + + + + + + + + Ferrous iron transporter fepABC + – + + – – + + Ferrous iron uptake homolog feoABC + + + + + – – + Sa, S. aureus Mu50; Sp, S. pseudintermedius ED99; Sd, S. delphini 8086; Si, S. intermedius NCTC11048; Se, S. epidermidus RP62A; Ss, S. saprophyticus ATCC15305; Sh, S. haemolyticus JCSC1435; Sc, S. carnosus TM300. a adapted from reference (55) b sitABC in S. epidermidus c sitA is a pseudogene in S. delphini    1.3.1 S. aureus heme acquisition Heme represents over 75% of the total iron pool in mammals, with the majority found in erythrocytes bound to hemoglobin (56). S. aureus is a hemolytic pathogen, capable of producing   9 hemolysins under iron restricted conditions, as in human blood (24). The ability to lyse erythrocytes and liberate hemoglobin into the extracellular milieu can be a fruitful source of iron for the pathogen as each erythrocyte contains ~280 million molecules of hemoglobin, with four molecules of heme bound to a hemoglobin heterotetramer (~ 1 billion iron atoms/cell) (57).  Heme uptake in S. aureus is primarily achieved using the iron-responsive surface determinant (Isd) system. S. aureus and S. lugdunensis are the only two staphylococcal species known to harbor the isd locus and it is considered to be an adaptation to an invasive lifestyle (58). This nine-gene system, isdABCDEFGHI, allows for high affinity hemoglobin binding at the cell surface, heme extraction, and relay into the cell cytoplasm (Figure 1-1) (59, 60). Briefly, cell-wall anchored proteins IsdB and IsdH can bind hemoglobin and hemoglobin-haptoglobin complexes, respectively, from which heme is extracted and then transported through the cell wall by IsdA and IsdC (61, 62). It is proposed that the architectural arrangement of these four proteins facilitates a funneling effect to unidirectionally move heme through the cell wall to the bacterial membrane (63). IsdC functions as the central conduit to transfer heme to an ABC transporter consisting of substrate-binding lipoprotein, IsdE, associated membrane permease, IsdF, and unidentified ATPase for internalization of heme (64, 65). IsdD is predicted to be an associated membrane protein but its function has not yet been defined. Cytoplasmic heme is either incorporated into bacterial heme-containing proteins or degraded by the heme-degrading enzymes, IsdG or IsdI, to liberate nutrient iron (66). IsdG and IsdI are differentially regulated, non-canonical heme monooxygenases that cleave the porphyrin ring to release iron and generate biliverdin-like products, known as staphylobillins (67). The Isd pathway is important for S. aureus pathogenesis as evidenced by attenuated staphylococcal virulence and/or burden in single deletion mutants of isdA, isdB, isdC, isdG, isdH, or isdI in murine models of infection (52, 68–  10 70). The Isd system is also required for abscess formation and thus can contribute to persistence and dissemination within the host (71). Heme is the most abundant iron reservoir in the host and, perhaps unsurprising, purportedly the preferred iron source for S. aureus (51).   Figure 1-1 Schematic of heme uptake and degradation by the Isd system in S. aureus. IsdB and IsdH bind extracellular hemoglobin (Hb) and hemoglobin-haptoglobin complex (Hb-Hp) at the cell surface. Heme (Hm) is relayed through the cell wall by IsdA and IsdC and internalized by IsdEF. IsdD is of unknown function. IsdI and IsdG degrade internalized heme using the reductase IruO to liberate iron and produce staphylobillins.     11 1.3.2 S. aureus non-heme iron uptake systems A second iron uptake strategy employed by S. aureus during infection is the biosynthesis and secretion of siderophores (53). Siderophore production is a common method used by microbes to overcome iron limitation and access extracellular host iron. These low molecular weight molecules can chelate ferric iron from host iron-binding proteins like lactoferrin and transferrin and courier it back to the bacterial cell (72, 73). Siderophores are broadly classified according to the functional groups used in iron coordination. These include catecholate, hydroxamate and a-hydroxycarboxylate types, as well as “mixed-types” ( see (74) for a review). Staphyloferrin A (SA) and staphyloferrin B (SB) are two ferric iron-chelating siderophores produced by S. aureus. Both staphyloferrins are a-hydroxycarboxylate-type and coordinate iron, at least in part, through citrate-derived moieties (75). Enzymes for SA and SB synthesis and export are encoded by the sfaABCD and sbnABCDEFGHI loci, respectively (50, 75). Iron-bound SA and SB are selectively imported by dedicated surface transporters, HtsABC and SirABC, respectively, in conjunction with the FhuC ATPase (73, 76). Iron release from internalized ferric-SA complexes requires iron-regulated expression of NtrA, a nitroreductase (77). The iron release mechanisms for both ferric-SA and ferric-SB have not been defined. Expression and utilization of both staphyloferrins is iron-regulated and under the control of Fur (50, 76). SA and SB have both been implicated in pathogenesis of S. aureus but SB appears to play a greater role in severe disease phenotypes and promotion of staphylococcal virulence in abscess and endocarditis models of infection (50, 78). S. aureus can obtain iron by acquiring exogenous siderophores produced by other organisms, referred to as xenosiderophores. S. aureus harbors the fhu (ferric hydroxamate uptake) system for hydroxamate-type siderophores and sstABCD (staphylococcal siderophore transporter) for uptake   12 of catechol-siderophores though S. aureus does not synthesize either of these siderophore types (73, 79). The hydroxamate siderophore, deferoxamine, requires expression of iruO for iron release in vivo (77), but otherwise, the mechanism(s) for iron utilization from these siderophores remains largely unknown. The ability to thieve and utilize both foreign catecholate and hydroxamate-type siderophores as iron sources may provide S. aureus with an advantage for growth in polymicrobial populations. Less studied in S. aureus are conserved free iron uptake systems. S. aureus possesses homologs of the fepABC (Fe-dependent peroxidase) system, but the exact function of the system has not been defined. S. aureus fepABC and its Escherichia coli homolog, efeOUB, have been connected to both ferrous and ferric iron uptake, as well as heme-iron extraction (80–82). sitABC (staphylococcal iron transporter) is used for ferric iron uptake in certain staphylococcal species. However, the homologus sitABC locus in S. aureus is named mntABC (manganese transporter) and has been characterized primarily for manganese import. This importer is predicted to be in the Fur regulon, which suggests it could have a role in S. aureus iron uptake as well (83, 84). Dedicated ferrous iron uptake systems in S. aureus are poorly understood. The feoABC (ferrous iron importer) is a ferrous iron importer found in many bacteria that is well-studied in Gram-negative bacteria, but is poorly characterized in Gram-positive pathogens. It is upregulated under iron-limited conditions in a Fur-dependent manner in S. aureus (85, 86). The contribution of free iron uptake to S. aureus pathogenesis has been sparsely characterized, mainly due to the overshadowing effects of the rigorously studied heme and siderophore-based iron acquisition and the low abundance of bioavailable inorganic iron. However, further study of free iron uptake system could elucidate how S. aureus accesses iron in anaerobic and acidic host niches, where ferrous iron predominates and siderophores are rendered ineffective (48).   13 1.4 Staphyloferrin biosynthesis in S. aureus Siderophore production is a virulence determinant and is required for optimal survival and proliferation of S. aureus. As aforementioned, S. aureus synthesizes two endogenous siderophores, SA and SB, that promote growth in iron-restricted environments. Staphyloferrin biosynthesis is achieved through Non-ribosomal peptide synthetase-Independent Siderophore (NIS) biosynthesis (75, 87). NIS synthesis systems involve intermediates and enzymes that are freely dissociable from each other during assembly. Primary metabolites generally serve as the source of building blocks to generate siderophores and can be chemically modified prior to incorporation into the siderophore. In addition to the synthetases required for assembly of the siderophore, NIS biosynthetic gene clusters generally encode enzymes that synthesize the necessary precursors to be assembled by the NIS synthetases (88). While functionally similar, SA and SB are chemically distinct and are produced by separate biosynthetic pathways.  1.4.1 Staphyloferrin A SA biosynthesis and efflux are mediated by the proteins encoded in the sfaABCD locus (75, 76). Staphyloferrin A biosynthetic genes are found broadly across both coagulase-positive and coagulase-negative staphylococci. Staphyloferrin A biosynthesis requires two synthetases, SfaD and SfaB, to condense two molecules of citrate with a central D-ornithine molecule in a stepwise manner to yield a functional SA molecule (75). SfaC is a putative amino acid racemase that catalyzes formation of D-ornithine from L-ornithine before to incorporation into the siderophore (75, 76).  Lastly, SfaA is a membrane transport protein responsible for efflux of apo-SA (89). Notably, the SA locus does not contain a dedicated citrate synthase and is consequently reliant on the TCA cycle citrate synthase, CitZ, for precursor citrate molecules (90). Thus, production of   14 SA relies heavily on central metabolism and is effected under conditions of extreme iron-deprivation as the TCA cycle is down-regulated as a part of the iron-sparing response (44–46). Additionally, the presence of glucose in growth conditions can hinder SA biosynthesis as carbon catabolite repression stringently limits the TCA cycle in favor of glycolysis (91).  1.4.2 Staphyloferrin B SB biosynthetic genes, sbnABCDEFGHI, encode not only the biosynthetic machinery for assembly and efflux of SB, but also the enzymes necessary to generate precursors from metabolites in central metabolism (Figure 1-2) (50, 87, 92). SbnC, SbnE, and SbnF are the three synthetases and SbnH is a decarboxylase that together are required for SB assembly from one molecule of each α-ketoglutarate (α-KG) and citrate, and two molecules of L-2,3-diaminopropionate (L-Dap) (87). There is some ambiguity in the literature as to the reported final structure of SB with the linear form most often depicted. However, chemical synthesis of SB confirms that the correct structure of SB contains a cyclic hemiaminal α-KG moiety and is referred to as the hemiaminal form (Figure 1-2B) (93, 94). Linear SB presumably spontaneously converts to hemiaminal SB, which represents its biological form (93, 94). The precursor molecules essential for SB biosynthesis are synthesized by SbnA, SbnB, and SbnG. α-KG and L-Dap are provided by the sequential enzymatic activities of SbnA and SbnB (92, 95). SbnA first catalyzes the formation of a novel amino acid intermediate, N-(1-amino-1-carboxyl-2-ethyl)-glutamic acid (ACEGA) from O-phospho-L-serine (OPS) and L-glutamate. SbnB subsequently oxidatively hydrolyzes ACEGA using NAD+ to produce L-Dap and α-KG precursors (92). OPS substrate is postulated to be shunted from the serine biosynthesis pathway stemming from 3-phosphoglycerate produced in glycolysis (90, 95). SbnG is a dedicated citrate synthase that   15 produces citrate from oxaloacetate and acetyl-CoA (90, 96, 97). As such, production of SB occurs regardless of TCA cycle activity (90). Efflux of assembled apo-SB into the extracellular milieu occurs via SbnD and an additional, as yet unidentified exporter (89). Lastly, SbnI, the ninth gene product, is a heme-responsive transcriptional regulator for SB production (98). A model has been proposed by which SbnI senses heme as a means to reduce SB synthesis in favor of heme acquisition (98). This model provides a mechanism for the previous findings that S. aureus demonstrates a heme-iron preference in vitro (51).      16   Figure 1-2 Physical map of the sbn genetic locus and SB biosynthetic pathway. (A) The SB biosynthetic locus consists of nine genes that encode enzymes required for synthesis of precursor molecules (orange), assembly of the siderophore (green), and efflux (blue). sbnI (red) encodes a heme-responsive regulator of SB biosynthesis. (B) Schematic of SB precursor molecule synthesis and use by synthetases, SbnE, SbnF, and SbnC, and decarboxylase, SbnH, to yield linear SB which interconverts to the biological form, hemiaminal SB.    17 SB is believed to be the primary siderophore expressed during invasive infection, while SA is speculated to serve more of a house-keeping role in iron acquisition conducive to commensalistic colonization (48). Observations that support this notion include the local metabolite concentrations found on skin compared to serum. Skin has relatively lower glucose and higher iron concentrations that may therefore allow TCA cycle activity and support SA production (99–103). Conversely, the concentration of glucose found in human serum is sufficient to suppress SA synthesis in vitro (90). As SB biosynthesis can occur autonomously from TCA cycle activity, SB production is unencumbered by serum glucose concentrations. Some of the most strongly up-regulated genes in the iron-restricted host include sbnA-I and the genes encoding the SB surface receptor and cognate ABC transporter, sirABC (24, 46, 104). Furthermore, presence of the SB biosynthetic locus is largely restricted to the more virulent, coagulase-positive species of staphylococci potentiating that SB production is a lineage specific adaptation to support a more invasive lifestyle.  1.5 S. aureus iron homeostasis and regulation 1.5.1 Intracellular iron metabolism and management S. aureus uses extreme measures to obtain iron from the host, but acquisition must be carefully coordinated with storage and efflux to ensure iron homeostasis. In many bacteria, including S. aureus, iron metabolism is regulated by Fur (105). Fur is a homodimeric metalloprotein with Fe2+ affinity tuned to monitor the labile iron pool of the cell (reviewed in (106)). Fur primarily functions as a transcriptional repressor of iron-responsive genes. In iron-replete conditions, Fur binds iron as a co-repressor and can interact with a consensus sequence, designated the Fur box, found within the promoter region of target genes (107). Fur-binding to   18 DNA blocks RNA polymerase and resultantly prevents transcription. Conversely, under conditions of iron deprivation, iron dissociates from Fur relieving transcriptional repression and allowing for synthesis of iron homeostatic systems. In addition, staphylococcal Fur influences the expression of proteins involved in biofilm formation, oxidative stress, and a subset of secreted virulence factors (42, 46, 108, 109). Bacterial pathogens can store iron in one of three types of proteins, ferritin, bacterioferritin, and Dps proteins. S. aureus possesses ferritin and Dps homologs, FtnA and MrgA, respectively (110). FtnA has been implicated as an iron storage molecule; however, studies of MrgA suggest that it likely functions as a DNA-binding protein for oxidative stress resistance rather than as an iron storage protein (110, 111). An alternative strategy to storage to alleviate iron toxicity is the export of iron-containing compounds or metabolites. A mechanism of heme detoxification by S. aureus is discussed in Section 1.6.3. S. aureus likely encounters a gradient of iron concentrations during infection. It is currently not well understood how S. aureus coordinates iron uptake and maintains homeostasis in accordance with iron availability in different host niches beyond the regulatory control of Fur.  1.5.2 S. aureus iron management in response to oxidative stress Reactive oxygen species (ROS) produced endogenously as a by-product of aerobic bacterial growth or exogenously by microbial competitors can cause oxidative stress to bacteria (29, 112). Also, eukaryotic hosts can deliberately produce ROS in phagocytes to attack engulfed bacteria. This oxidative burst response is an important part of the innate immune response against S. aureus (113). ROS have multiple toxic effects, including oxidation of cysteine and methionine residues damaging proteins and disruption of iron-sulfur clusters. However, the most damaging   19 effects are attributed to reaction of hydrogen peroxide with ferrous iron to form hydroxyl radicals. This primarily effects DNA due to the association of iron with nucleic acids (114). Such Fenton chemistry-induced damage can have deleterious effects on the growth of S. aureus (115), and elevated intracellular iron levels have been demonstrated to predispose S. aureus to killing by monocytes and macrophages (116, 117). The efficient sensing of redox stress and subsequent induction of resistance mechanisms have been considered crucial for pathogens that have to combat oxidative immune defenses (116). To cope with redox stress, many bacteria, especially aerobes and pathogens, have evolved sophisticated oxidative stress response systems orchestrated by transcription factors that sense specific ROS. The main strategy employed to defend against oxidative damage is production of enzymes to degrade ROS. In S. aureus these include superoxide dismutases (SodA and SodM), catalase (KatA), alkyl hydroperoxide reductase (AhpCF), thiol-dependent peroxiredoxin homolog bacterioferritin comigratory protein (Bcp), and thioredoxin reductase (TrxB) (110, 118–120). S. aureus uses PerR (peroxide response regulator), a member of the Fur family of metal-dependent regulators, for the control of the oxidative stress response (109, 110). PerR has primarily been characterized as a manganese-dependent repressor of the PerR regulon. However, PerR has also recently been described to use Fe2+ to sense very low levels of hydrogen peroxide (121). Upon sensing iron and/or peroxides, PerR regulon repression is relieved and allows for expression of antioxidant genes (KatA, AhpCF, Bcp, and TrxB) and iron storage proteins including FtnA and MrgA. Upregulation of FtnA and MrgA may have a protective role for DNA by sequestering intracellular iron and effectively limiting free iron in the cell (122, 123). As well, PerR is a metal-dependent repressor of the metalloregulators Fur and PerR, itself (110). PerR-dependent control is important for pathogenicity in murine skin abscess model of infection (110).   20 Another response to peroxide stress is increased expression of Mn2+ importers, mntABC and mntH, controlled by the manganese homeostasis regulator, MntR (83). As there can be metal co-factor plasticity amongst mononuclear iron-requiring enzymes, some enzymes can replace iron with manganese. This change in cofactors may allow microbes to maintain enzymatic activity and prevent iron-induced damage under redox stress (124). Additionally, several manganese complexes can catalyze the disproportionation of hydrogen peroxide (125). The interplay between iron homeostasis, PerR regulation, and the oxidative stress response allows S. aureus to coordinate the intracellular availability of free iron with the level of antioxidant proteins present in the cell (111). Since iron can exacerbate oxidative stress, and also serves as a co-factor for some ROS degrading enzymes (eg. KatA requires heme-iron for catalase activity), there is significant regulatory cross-talk between iron homeostasis and oxidative-stress resistance networks. As PerR is repressed by elevated manganese concentrations, there is a regulatory link between MntR control of manganese uptake and expression of the PerR regulon and resistance to redox stress (83). Together, Fur, PerR, and MntR form an integrated network controlling oxidative stress resistance as well as iron and manganese homeostasis.  1.6 S. aureus heme homeostasis and regulation 1.6.1 Heme biosynthesis Heme is advantageous to the pathogen as it is a cofactor for aerobic respiration and catalase to resist oxidative immune defenses (126, 127). During infection, most bacterial pathogens either acquire host heme or synthesize heme endogenously. The contribution of the heme uptake Isd system is well established in S. aureus pathogenesis. However, much less is known about the role of heme biosynthesis in infection (128). Heme biosynthesis is vital in murine infection   21 models, where inactivation of the S. aureus heme biosynthetic pathway results in a less invasive phenotype that is defective in colonizing the murine heart and liver (127). Similarly, in a murine model of osteomyelitis, a S. aureus strain deficient in heme synthesis has reduced colonization and bone destruction (129, 130). Together the data suggests that heme biosynthesis is required for full pathogenesis and that heme acquisition alone is insufficient for organ colonization (128). Heme biosynthesis in S. aureus follows a recently described “noncanonical” or “transitional” pathway (Figure 1-3) (131–133). In the last few years, the terminal steps of heme synthesis in Gram positive phyla, Firmicutes and Actinobacteria, was determined to diverge from the classical heme synthesis pathway (131). As with most bacteria, heme biosynthesis in S. aureus begins with charged glutamyl-tRNAGlu to form the precursor d-amino-levulinic acid (ALA) by glutamyl tRNA reductase (GtrR), which is subsequently transformed to uroporphyrinogen III in three enzymatic steps by PbgS, HmbS, and UroS (formerly HemB, HemC, and HemD) (134). UroD (formerly HemE) carries out a decarboxylation reaction to yield coproporhyrinogen III and is the first committed step of the classical heme biosynthetic pathway (135). Until recently, the terminal steps of the classical heme pathway were considered universally conserved for all heme synthesizing organisms. It is after the synthesis of coproporphyrinogen III that the non-canonical pathway used by Gram positive bacteria diverges from the classical pathway. The last step in the classical pathway involves insertion of ferrous iron into the protoporphyrin IX ring by protoporphyrin ferrochelatase (PpfC, formerly HemH) to form protoheme IX, more commonly called heme (136). Conversely, in the non-canonical pathway, the antepenultimate step is insertion of the iron into coproporphyrin III to yield coproheme III by coproporphyrin ferrochelatase (CpfC) (133). Coproheme decarboxylase (ChdC, formerly HemQ), an enzyme found in members of the Firmicutes and Actinobacteria, catalyzes the final step in non-canonical   22 heme biosynthesis by the oxidative decarboxylation of coproheme into heme (132, 133, 137, 138). ChdC has also been demonstrated to have low peroxidase activity, but in the presence of peroxide the bound heme is susceptible to degradation (139, 140). ChdC may play a dual role as a decarboxylase in heme synthesis and a regulatory protein in heme homeostasis, catalyzing the degradation of bound heme when it is not required for cellular metabolism (131).    Figure 1-3 Bacterial heme biosynthesis. The heme synthesis pathway of most bacteria begins with charged glutamyl-tRNAGlu to form the universal precursor ALA and coproporphyrinigen III is formed through a series of conserved enzymatic steps. The classical pathway (blue) forms heme through the protoporphyrinogen IX intermediate; most organisms including Gram negative bacteria and eukaryotes use this pathway. The noncanonical pathway (green), performed by most Gram positive bacteria, produces heme through the coproporphyrin III intermediate. Shown for eash step is the enzyme name followed by the common protein annotation in bold. Obtained with permission from reference (128). Copyright 2016 Elsevier Publishers.   23 Regulation of heme biosynthesis in S. aureus was recently defined (141). GtrA generates the heme precursor ALA, the first committed step in heme biosynthesis. It is also a critical regulator of S. aureus heme biosynthesis. GtrR is regulated post-transcriptionally by heme abundance and the integral membrane protein HemX (141). When heme levels are proficient in cells, GtrR is kept at low levels but increases in response to heme deficiency. HemX post-transcriptionally regulates GtrR but the specific mechanism by which HemX alters GtrR abundance remains unidentified (141). HemX is annotated as a member of the cytochrome c assembly protein family supporting a role in heme-binding and trafficking at the membrane. Futhermore, a HemX mutant strain has increased heme synthesis and activates the heme stress response (HssRS), supporting a role in heme homeostasis and suggesting that regulation of heme is essential to avoid self-induced heme toxicity (141). Heme-bound GtrR is proposed to be susceptible to degradation by cellular proteases, unlike apo-GtrR (142, 143). These findings are consistent with the mechanism of heme synthesis regulation described in model Gram negative organisms where GtrR protein levels are regulated by heme and proteolysis (142).  Other studies have suggested further roles of S. aureus ferrochelatase, CpfC, in maintenance of heme homeostasis. CpfC has a regulatory iron-binding site that upon iron binding, results in substrate inhibition (144). This finding has implications for how iron availability may influence heme biosynthesis. Iron abundance may inhibit CpfC as a way to avoid accumulation of Fe-coproporphyrin III or possibly when the heme requirements of the cell are satisfied, iron is made available for other processes (144). S. aureus depends on many heme-requiring proteins but is coincidentally sensitive to heme toxicity so intracellular heme levels must be closely monitored. Regulation of heme biosynthesis is one mechanism to maintain heme homeostasis. Many   24 bacterial pathogens encode complete heme biosynthetic pathways, but the contribution of heme biosynthesis to pathogenesis remains largely understudied.  1.6.2 Intracellular heme metabolism and management During infection of host heme- and hemoglobin-rich niches, pathogens can experience heme toxicity. Free heme is bacteriocidal towards S. aureus at low-micromolar concentrations (66, 126). S. aureus heme homeostasis is ensured through the concerted efforts of three mechanisms: Fur repression of heme uptake when excess iron is available, degradation of heme by heme oxygenases, and lastly a heme-sensing system that controls efflux of intracellular heme. S. aureus encodes a two-component system, the heme-sensor system (HssRS), to sense heme and activate a response to alleviate heme toxicity (145). HssS is a transmembrane histidine kinase that senses excess heme or cellular stress mediated by excess heme exposure. The exact ligand of HssS is unknown but excess exogenous or endogenous heme leads to activation. When activated, HssS phosphorylates HssR, its cognate response regulator, which binds a direct repeat in the promoter region of the heme-regulated transporter (hrtAB) (146). HrtA is a putative ATPase and with the permease, HrtB, comprises a proposed heme efflux pump (46, 145). Deletion of hssR results in intracellular accumulation of heme and hypervirulence demonstrating the important function HrtAB has in maintain heme homeostasis (146). Currently, it is unknown how heme is trafficked to the exporter. Internalized heme does not likely exist as a free molecule in the cytoplasm, but rather as a complex with heme-binding proteins or chaperones to facilitate intracellular trafficking. Not only does free heme have low solubility, heme has toxic effects due to the reactive nature of heme-iron and the ability to damage DNA through heme-generated ROS (126). ROS can also initiate non-enzymatic modifications and degradation of heme itself (46).   25 Whether heme is synthesized de novo or host-acquired, the molecular mechanisms responsible for intracellular heme trafficking in S. aureus have not been evaluated in detail. Sensing or regulatory pathways that connect heme synthesis, heme availability, hemoprotein abundance, or heme stress activation have not been fully defined. The role of heme-trafficking in maintenance of heme and iron homeostasis requires further research.  1.6.3 Heme degradation An alternative to reduce cellular heme levels by efflux or decreased heme uptake is heme degradation. Intracellular heme concentrations in S. aureus can, at least in part, be managed by the activity of heme-degrading proteins, IsdI and IsdG (147, 148). IsdI and IsdG are two paralogous, non-canonical heme oxygenases that degrade heme to form novel chromophores, staphylobilins, with the release of iron and formaldehyde (67, 149). Referred to as the IsdG-family heme oxygenases, IsdI and IsdG are structurally and mechanistically distinct from canonical heme oxygenases (149–151). IruO (iron utilization oxidoreductase) has been identified in S. aureus as the in vivo source of reductant for IsdI and IsdG heme degradation by transfer of electrons from NADH (152). In vitro, heme degradation facilitated by IruO occurs more rapidly with IsdI than IsdG raising the possibility that IsdG utilizes an as yet unidentified reductase (48, 152). This role could be fulfilled by NtrA, a nitroreductase that has been implicated in heme-iron utilization in vivo (77). Both IsdI and IsdG are Fur-regulated; however, IsdG is post-translationally regulated by heme (68). IsdG has a unique flexible loop that, through an unknown mechanism, targets IsdG for degradation in the absence of heme (153). Differential regulation of IsdI and IsdG has been proposed to allow for fine-tuning heme homeostasis in the cell (68).   26 Fur regulation of heme oxygenases, IsdI and IsdG, was recently discovered to be required for fine-tuning heme homeostasis in S. aureus (154). Though investigators initially sought to investigate the fate of heme degradation products (staphylobillins), they observed that dysregulation of IsdI and IsdG led to increased expression of genes linked to oxygen-independent energy production (154). This increased reliance on fermentation for energy production was hypothesized to be due to insufficient heme to populate cytochromes for aerobic respiration. Unregulated heme oxygenase activity was demonstrated to be detrimental for the cell and Fur regulation ensures that the cell has sufficient quantities of heme (154).   1.7 Spatiotemporal expression of iron uptake systems by S. aureus in the host The spatial organization of pathogenic and commensal microbes over the landscape of a single human is driven by host landmarks and specific host-pathogen interactions. The term microbiogeography was coined to describe the processes that generate the spatial patterns of microbial distribution to the scale of a single infection (155). Factors that impact microbiogeography include host receptors for microbial attachment, physiochemical and nutritional gradients (eg. pH and oxygen), access to nutrients, and the immune system. The distribution of microbes is not random throughout the body, rather, it is influenced by the microenvironments that can vary in how well they can suppress or support microbial growth. Microbial spatial patterns across the human body have recently inspired significant research and interest (156–158). S. aureus is known for its ability to infect diverse tissue types and one critical host factor that likely influences microbiogeography is access to iron. Host environments are generally low in iron and the accessible biological iron pools vary between tissue tropisms. Not well understood is how S. aureus coordinates and adapts its iron uptake strategy based on the site   27 of infection and how iron uptake changes over the course of infection. The possibility that additional iron regulatory mechanisms exist is substantiated by observations of differential expression of iron acquisition systems or conversely, variation in the necessity of certain iron uptake systems for colonization of different organs. Evidence for adaptation of iron uptake strategies employed to suit different anatomic sites has been reported in the development of S. aureus pneumonia in a murine model of infection. Establishment and pathogenesis in the model were not dependent on hemoglobin binding or heme uptake systems, which suggested that colonization of distinct host niches involves the utilization of different iron sources (159). Recent study of the spatiotemporal expression of FhuD, a lipoprotein required for uptake of iron-hydroxamate siderophores, demonstrated that fhuD2 is expressed in multiple organs in vivo and that expression is spatially and temporally controlled, increasing at later stages of infection in some organs (160). Though monitoring in vivo bacterial gene expression over the course of infection presents with inherent challenges, these studies can provide insight into how certain iron sources are preferred at different infectious foci over the course of infection. It is tempting to hypothesize that S. aureus has mechanisms to coordinate spatiotemporal expression of iron-uptake systems according to the site of infection. Together, there is a close relationship between nutrient supply and gene expression and mounting evidence that host sensing plays a key role in S. aureus adaptation of iron uptake and survival strategies used to advance pathogenesis and infection. Identification of the molecular mechanisms that control nutrient source preference during infection could allow drugs to be designed to disrupt essential iron acquisition pathways.    28 1.7.1 S. aureus iron source preference S. aureus demonstrates heme-iron preference in vitro (51). This preference may seem logical as heme represents the most abundant iron pool in the vertebrate host or heme may be the form of iron most needed by S. aureus. Mechanistic insight into how this nutritional preference is controlled has been provided by study of SbnI, the ninth gene product encoded by the sbn locus for SB biosynthesis (98). All proteins encoded by the sbn gene cluster play roles in either biosynthesis or export of SB, except for the SbnI. Though conserved in staphylococcal species harboring an sbn locus, SbnI has no characterized enzymatic role in SB production and is not required for in vitro SB synthesis (87). SbnI is a heme-responsive transcriptional regulator required for full expression of sbnD-H and thus controls SB biosynthesis (98). SbnI binds to a DNA fragment within the sbnC coding region; however, SbnI can bind heme and heme-bound SbnI does not bind DNA (98). A model by which SbnI is required for transcription of parts of the SB biosynthetic operon and senses intracellular heme to reduce SB synthesis in favour of heme acquisition is proposed (Figure 1-4). SbnI is reported as the first example in S. aureus of fine-tuning iron-regulated genes beyond Fur and of a DNA-binding regulatory protein that senses heme to control gene expression for siderophore synthesis (98).    29   Figure 1-4 Schematic of SbnI regulation of SB biosynthesis and SB-mediated iron acquisition. In the absence of heme, SbnI is a positive regulator of SB biosynthesis. SB is exported into the extracellular mileau by SbnD where upon binding ferric iron it is recognized by SirA for import of the Fe-SB complex. Receptor-bound Fe-SB is translocated across the cell membrane through a transmembrane permease, SirBC, in conjuction with the FhuC ATPase. Iron is released from SB by an unknown mechanism. Shown in the grey box: SbnI can bind heme as a negative regulatory molecule. Heme binding by SbnI abrogates interaction with DNA and thus SB biosynthesis.  1.8 Objectives Heme acquisition by the Isd system, heme biosynthesis, and siderophore-mediated iron uptake are required for full S. aureus pathogenesis in diverse host environments. For therapeutics to be rationally developed to target iron acquisition systems in S. aureus, we must study and understand the environmental conditions and types of infections under which certain iron pathways are essential. Presently, studies have primarily focused on in vivo characterization of   30 the necessity of iron-acquisition systems in various models of infection. However, study of the spatiotemporal regulatory mechanisms required for adaptation of iron uptake systems, beyond Fur, have been limited. The recent characterization of SbnI provides a mechanism for how S. aureus demonstrates iron source preference, but the structural determinants that allow this protein to carry out its heme-dependent regulatory role remain unknown. I hypothesize that the structural features of SbnI are important for its direct or indirect regulatory role in SB biosynthesis. SbnI lacks obvious sequence similarity to any known transcription factors or heme-binding proteins. Therefore, X-ray crystallography was used to gain insight into the function of SbnI. Structural analysis of a C-terminal truncated construct revealed that SbnI shares low sequence identity but high structural homology with an archeael free serine kinase. A combination of UV-visible spectroscopy, 31P NMR, HPLC, kinetic analysis, and in vivo growth experiments were used to determine that SbnI produces OPS using ATP and L-serine. Furthermore, SbnI-generated OPS is the in vitro and in vivo source of OPS for SbnA in SB biosynthesis by S. aureus. Determination of SbnI as a free serine kinase and its contribution to the functional modularity of the sbn locus will be discussed in Chapter 3. SbnI transcriptional regulation of SB biosynthesis is negatively regulated by heme. The goals of this study were to determine the full-length structure of SbnI, how it interacts with heme, and if heme-binding alters L-serine kinase activity. As the C-terminal truncated construct of SbnI was defective in heme-binding, the structure of full-length SbnI homolog from Staphylococcus pseudintermedius was solved using X-ray crystallography to gain insight into the structural determinants for SbnI heme-binding. Site-directed mutagenesis, UV-visible spectroscopy, and molecular docking experiments were used to examine this interaction. Presented in Chapter 4 is   31 an analysis of the dual function of SbnI in SB biosynthesis, as a transcriptional regulator of the sbn locus and enzyme for biosynthesis of SB precursor molecule OPS. A model of heme-binding by SbnI and determination that SbnI acquires heme from IsdI are also presented. These results build on the model of SbnI as a heme dependent regulator of SB biosynthesis and provides a mechanism for how S. aureus can adapt to different iron-restricted niches in the host. Although great strides have been made in recent years understanding the terminal steps of heme biosynthesis in Gram positive bacteria, there remains an open question as to how synthesized heme is trafficked and delivered to heme-requiring proteins in the cell. The aim of this study was to determine the structure of the S. aureus terminal enzyme in heme biosynthesis, ChdC, and to determine whether it is capable of heme-transfer to cytosolic heme-binding proteins. Based on UV-visible spectroscopy and kinetic analysis of heme transfer reactions, a model of how ChdC, IsdI, IsdG, and SbnI participate in a heme trafficking network in S. aureus is presented in Chapter 5.    32 Chapter 2: Methods  2.1 Cloning, expression, and protein purification for biochemical assays and structure determination E. coli strains and plasmids used for protein expression are summarized in Table 2-1.  Table 2-1 E. coli strains and plasmids used in this study.   E. coli strains and plasmids Description Source or reference Strains   E. coli   BL21 (λDE3) F− ompT gal dcm lon hsdSB (rB− mB+)λ (DE3 [lacI lacUV5-T7 gene 1 ind1 sam7 nin5]) Novagen Plasmids   pET28a-sbnI IPTG-inducible expression vector containing sbnI; KmR This study pET28a-sbnI1-240 IPTG-inducible expression vector containing sbnI1-240; KmR This study pET28a-sbnI-H3A IPTG-inducible expression vector containing sbnIH3A; KmR This study pET28a-sbnI-E20A IPTG-inducible expression vector containing sbnIE20A; KmR This study pET28a-sbnI-D58A IPTG-inducible expression vector containing sbnID58A; KmR This study pET28a-sbnI-C155A IPTG-inducible expression vector containing sbnIC155A; KmR This study pET28a-sbnI-C168A IPTG-inducible expression vector containing sbnIC168A; KmR This study pET28a-sbnI-C244A IPTG-inducible expression vector containing sbnIC244A; KmR This study pET28a-sbnI-H3A/C244A IPTG-inducible expression vector containing sbnIH3A/C244A; KmR This study pET28a-spsbnI IPTG-inducible expression vector containing spsbnI; KmR This study pET28a-chdC IPTG-inducible expression vector containing chdC; KmR This study pET15b-isdI IPTG-inducible expression vector containing isdI; AmpR (161) pET15b-isdG IPTG-inducible expression vector containing isdG; AmpR (161)   33 E. coli strains and plasmids Description Source or reference pET52b-isdI IPTG-inducible expression vector containing isdI; AmpR, N-terminal Strep-tag This study pET28a-iruO IPTG-inducible expression vector containing iruO; KmR (152) pET28a-sbnA IPTG-inducible expression vector containing sbnA; KmR (92) a IPTG (isopropyl β-D-thiogalactopyranoside)  b KmR, and AmpR designate resistance to kanamycin and ampicillin respectively.  2.1.1 Cloning, expression, and purification of SbnI and SbnI variants Constructs with an N-terminal His6 tag and thrombin cleavage site were generated in pET28a vectors for recombinant expression of S. aureus full-length SbnI (residues 1-254) and C-terminal truncated construct SbnI1-240 (residues 1-240), with the first codon mutated from the native TTG to a common start codon, ATG. The S. aureus SbnI nucleotide sequence can be accessed in the GenBank database under accession code NC_009641.1 (90178-90942) (gene locus NWMN_RS00380) and the amino acid sequence can be accessed through NCBI Protein Database under NCBI accession WP_001015549.1. Briefly, a megaprimer-based whole-plasmid synthesis PCR cloning protocol was used to clone constructs amplified from chromosomal DNA from S. aureus strain Newman (162).  S. aureus SbnI variants SbnI H3A, SbnI E20A, SbnI D58A, SbnI C155A, SbnI C168A, and SbnI C244A were produced using a single primer mutagenesis method (163). Mutagenesis primers used in this study are summarized in Table 2-2. A double site-directed mutant, SbnI H3A/C244A, was generated by performing a subsequent round of mutagenesis on the pET28a-sbnI-H3A plasmid with the sbnI C244A mutagenesis primer. All clones were introduced into E. coli BL21 (λDE3) and confirmed by DNA sequencing.    34 Table 2-2 Primers used for site-directed mutagenesis of S. aureus sbnI active site and putative heme-binding site.  Primer name Primer sequence sbnI H3A 5‘ /5phospho/atg aat gcc att cat gaa cat tta aaa ttg g 3’ sbnI E20A 5’ /5phospho/att gat ctt cac gcc aca ttc gaa cct tta ag 3’ sbnI D58A 5’ /5phospho/tat atg gtt ata gcc ggt gtg cat cgg tat aca ag 3’ sbnI C155A 5’ /5phospho/gca agt tat agt ggt gcc tgt tct gta gag aga att gc 3’ sbnI C168A 5’ /5phospho/ggt aca tat cct gcc ctt tct caa caa gat g 3’ sbnI C244A 5’ /5phospho/gcc aat atg aga gcc tat act gaa aaa gta tac ttg g 3’  a Bolded oligonucleotides are mutated from the S. aureus strain Newman sequence b 5phospho – primers were synthesized 5’ phosphorylated  Recombinant full-length SbnI, SbnI1-240, SbnI H3A, SbnI E20A, SbnI D58A, SbnI C155A, SbnI C168A, SbnI C244A and SbnI H3A/C244A constructs were overexpressed in E. coli BL21 (λDE3) cells. Cultures were grown in 2x yeast extract tryptone (2xYT) media supplemented with 25 µg/mL kanamycin at 30°C to an OD600 of 0.7-0.9. Cultures were then induced with 0.5 mM isopropyl β-D-thiogalactopyranoside (IPTG) and grown for an additional 18 h at 20°C. Cells were pelleted by centrifugation at 4400 x g for 7 min at 4 °C and resuspended in buffer containing 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, 2 mM tris(2-carboxyethyl)phosphine (TCEP), and 10 mM imidazole on ice. Approximately 5 mg of DNase was added to cell suspension prior to lysis at 4°C using an EmulsiFlex-C5 homogenizer (Avestin). Insoluble material was removed by centrifugation at 39,000 x g for 1 h and recombinant protein was purified from soluble lysate using a HisTrap nickel affinity column (GE Healthcare) by elution with an imidazole gradient. Protein was dialyzed against 50 mM HEPES (pH 7.4), 100 mM NaCl, 5% (v/v) glycerol, and 2 mM TCEP and then cleaved with thrombin at a 1:500 ratio by weight of His6 protein to remove the His6 tag over 18 hr at 4°C. Subsequently, recombinant protein was dialysed into 50 mM HEPES (pH 7.4), 5% (v/v) glycerol, and 2 mM TCEP and further purified by anion exchange chromatography using a Source 15Q column (GE   35 Healthcare). Purified protein was obtained by elution with a NaCl gradient and further dialyzed into 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, 2 mM GSH. The sample was concentrated to approximately 20 mg/mL, flash frozen, and stored at -80°C. Selenomethionine-incorporated SbnI1-240 was produced by methods previously described (164) and purified as described above for native SbnI1-240.  2.1.2 Cloning, expression, and purification of S. pseudintermedius SbnI Full-length S. pseudintermedius SbnI homolog, termed SpSbnI for this study, (residues 1-254) nucleotide sequence can be accessed in the GenBank database under GenBankAccession NC_017568.1 (2180050-2180814) (gene locus SPSE_RS10030) and the amino acid sequence can be accessed through NCBI Protein Database under NCBI Accession WP_015728696.1. Briefly, SpSbnI construct was cloned from S. pseudintermedius strain ED99 chromosomal DNA using a megaprimer-based whole-plasmid synthesis PCR cloning strategy (162). The clone was introduced into Escherichia coli BL21 (λDE3) and confirmed by DNA sequencing. Recombinant SpSbnI constructs was overexpressed in E. coli BL21 (λDE3) cells. Cultures were grown in 2xYT media supplemented with 25 µg/mL kanamycin at 30°C to an OD600 of 0.7-0.9. Cultures were then induced with 0.5 mM IPTG and grown for an additional 18 h at 20°C. Cells were pelleted by centrifugation at 4400 x g for 7 min at 4 °C and resuspended in buffer containing 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, 2 mM tris(2-carboxyethyl)phosphine (TCEP), and 10 mM imidazole on ice. 5 mg of DNase was added to cell suspension prior to lysis at 4°C using an EmulsiFlex-C5 homogenizer (Avestin). Insoluble material was removed by centrifugation at 39,000 x g for 1 h and recombinant protein was purified from soluble lysate using a HisTrap nickel affinity column (GE Healthcare) by elution   36 with an imidazole gradient. Protein was dialyzed against 50 mM HEPES (pH 7.4), 100 mM NaCl, 5% (v/v) glycerol, and 2 mM TCEP and then cleaved with thrombin at a 1:500 ratio by weight of His6 protein to remove the His6 tag over 18 hr at 4°C. Subsequently, recombinant protein was dialysed into 50 mM HEPES (pH 7.4), 100 mM NaCl, 5% (v/v) glycerol, and 2 mM TCEP and further purified by anion exchange chromatography using a Source 15Q column (GE Healthcare). Purified protein was obtained by elution with a NaCl gradient and further dialyzed into 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, 2 mM glutathione (GSH). The sample was concentrated to ~20 mg/mL, flash frozen, and stored at -80°C.   2.1.3 Cloning, expression, and purification of ChdC A construct with an N-terminal His6 tag and thrombin cleavage site was generated in pET28a vector for recombinant expression of S. aureus strain Newman full-length ChdC, residues 1-250, using a megaprimer-based whole-plasmid synthesis PCR cloning strategy (162). All clones were introduced into Escherichia coli BL21 (λDE3) and confirmed by DNA sequencing. The S. aureus ChdC nucleotide sequence can be accessed in the GenBank database under GenBankAccession AP009351.1 (633718-634470) (gene locus NWMN_0550) and the amino acid sequence can be accessed through NCBI Protein Database under NCBI Accession BAF66822.1.   Recombinant ChdC was overexpressed in E. coli BL21 (λDE3) cells. Cultures were grown in 2xYT media supplemented with 25 µg/mL kanamycin at 30°C to an OD600 of 0.7-0.9. Cultures were then induced with 0.5 mM IPTG and grown for an additional 18 h at 20°C. Cells were pelleted by centrifugation at 4400 x g for 7 min at 4 °C and resuspended in buffer containing 50 mM Tris (pH 7.5), 300 mM NaCl, and 20 mM imidazole on ice. 5 mg of DNase was added to   37 cell suspension prior to lysis at 4°C using an EmulsiFlex-C5 homogenizer (Avestin). Insoluble material was removed by centrifugation at 39,000 x g for 1 h and recombinant protein was purified from soluble lysate using a HisTrap nickel affinity column (GE Healthcare) by elution with an imidazole gradient. Protein was dialyzed against 50 mM Tris (pH 7.5) and then cleaved with thrombin at a 1:500 ratio by weight of His6 protein to remove the His6 tag over 18 hr at 4°C. Subsequently, recombinant protein was further purified by anion exchange chromatography using a Source 15Q column (GE Healthcare). Purified protein was obtained by elution with a NaCl gradient and further dialyzed into 50 mM Tris (pH 7.5), 100 mM NaCl. The sample was concentrated to ~ 20 mg/mL, flash frozen, and stored at -80°C.   2.1.4 Cloning, expression, and purification of IsdI An isdI construct with N-terminal Strep-tag was generated in pET52b using megaprimer-based whole-plasmid synthesis PCR cloning strategy (162). Template DNA for isdI was subcloned from previously made construct in pET15b containing S. aureus strain Newman DNA sequence (147). The clone was introduced into E. coli BL21 (λDE3) and confirmed by DNA sequencing.  IsdI containing strep tag was expressed in E. coli BL21 (λDE3). Cultures were grown in 2xYT media supplemented with 100 µg/mL ampicillin at 30°C to an OD600 of 0.7-0.9. Cultures were then induced with 0.5 mM IPTG and grown for an additional 18 h at 25°C. Cells were pelleted by centrifugation at 4400 x g for 7 min at 4 °C and resuspended in buffer containing 100 mM Tris (pH 8.0), 150 mM NaCl, and 1 mM ethylenediaminetetraacetic acid (EDTA) on ice. 5 mg of DNase was added to cell suspension prior to lysis at 4°C using an EmulsiFlex-C5 homogenizer (Avestin). Insoluble material was removed by centrifugation at 39,000 x g for 1 h.   38 Recombinant protein was purified from soluble lysate using gravity flow 13 mL column containing Strep-tactin Superflow high capacity resin (IBA Life Sciences). Bound protein was eluted using elution buffer containing 100 mM Tris (pH 8.0), 150 mM NaCl, 1 mM EDTA, and 2.5 mM desthiobiotin. Protein was dialyzed against 50 mM HEPES (pH 7.4), 300 mM NaCl, and 5% (v/v) glycerol. The sample was concentrated to ~20 mg/mL, flash frozen, and stored at -80°C. IsdI containing His6 tag was expressed in E. coli BL21 (λDE3) cells from the plasmid pET15b, purified by His-tag affinity chromatography, and digested with the tobacco etch virus protease to remove His6 tag as previously described (147). Protein was dialyzed against 50 mM HEPES (pH 7.4), 300 mM NaCl, and 5% (v/v) glycerol. The sample was concentrated to ~20 mg/mL, flash frozen, and stored at -80°C.  2.1.5 Expression and purification of SbnA His6-tagged SbnA was expressed in E. coli BL21 (λDE3) cells from the plasmid pET28a, purified by His-tag affinity chromatography, and digested with thrombin to remove the His6 tag. The protein was further purified by anion exchange chromatography using the previously published method for improved SbnA solubility (92). SbnA was dialyzed into 50 mM Tris pH 8, 100 mM NaCl, and 2 mM TCEP, concentrated to ~20 mg/mL and stored at -80°C.  2.1.6 Expression and purification of IruO IruO containing His6 tag was expressed in E. coli BL21 (λDE3) cells from the plasmid pET28a, purified by His-tag affinity chromatography, and digested with thrombin to remove His6 tag and further purified by anion exchange chromatography as previously described (152).   39 Protein was dialyzed against 50 mM HEPES (pH 7.4), 300 mM NaCl, and 5% (v/v) glycerol. The sample was concentrated to ~20 mg/mL, flash frozen, and stored at -80°C.  2.1.7 Expression and purification of IsdG IsdG containing His6 tag was expressed in E. coli BL21 (λDE3) cells from the plasmid pET15b, purified by His-tag affinity chromatography, and digested with the tobacco etch virus protease to remove His6 tag as previously described (147). Protein was dialyzed against 50 mM HEPES (pH 7.4), 300 mM NaCl, and 5% (v/v) glycerol. The sample was concentrated to ~20 mg/mL, flash frozen, and stored at -80°C.  2.2 Crystallization, data collection, and structure determination 2.2.1 SbnI1-240 structure determination Selenomethionine-labeled SbnI1-240 crystals were grown by sitting drop vapor diffusion at 4°C in 2 uL drops with a 1:1 mixture of ~20 mg/mL SbnI1-240 in 50 mM HEPES (pH 7.4), 100 mM NaCl, 5% (v/v) glycerol, and 2 mM TCEP with reservoir solution containing 0.18 M HEPES (pH 7.5) and 20% (w/v) PEG 8000. Crystals were briefly soaked in reservoir buffer supplemented with 30% (v/v) glycerol for cryoprotection and flash frozen in liquid nitrogen. A single wavelength anomalous diffraction dataset was collected at the Canadian Light Source (CLS) on beamline 08B1-1 (165). The data was processed and scaled using XDS (166, 167). Crystals were of space group P31 with one molecule in the asymmetric unit. Five selenomethionine sites were identified for phasing to build a preliminary model using AutoSol (initial figure of merit of 0.35) and Autobuild (187 of 240 residues built) programs in Phenix (168). Manual building was done using Coot (169) and refinement was performed with   40 phenix.refine using translation libration screw (TLS) parameters with three TLS groups (170). The refined structure contains Met1-Ala240, one glycerol, and 12 water molecules. Data collection and refinement statistics are summarized in Table 2-3. Structure figures were generated in PyMOL (The PyMOL Molecular Graphics System, Version 1.8 Schrödinger, LLC).   Table 2-3 X-ray diffraction data collection and refinement statistics for SbnI1-240.  Data Collectiona Resolution Range (Å) 42 – 2.50 (2.59 – 2.50) Space group P31 Unit cell dimensions a, b, c (Å) 55.1, 55.1, 92.7 Unique reflections 10,883 (1,564) Completeness (%) 99.9 (100) Redundancy 2.9 (2.9) Average I/σI 15.8 (2.2) Rmerge 0.051 (1.007) CC1/2 0.998 (0.651) Wilson B-factor (Å2) 56.4 Anisotropy 0.501 Refinement Rwork (Rfree) 0.219 (0.259) Number of water molecules 12 RMSD bond length (Å) 0.003 Average B-values (Å2) 95.0 Ramachandran plot (%) Most favored regions 97.5 Disallowed regions 0.4 PDB ID 5UJE aData collection values in parentheses represent the data for the highest resolution shell.  2.2.2 SpSbnI structure determination SpSbnI crystals were grown by sitting drop at 4°C in a 1:1 mixture of ~20 mg/mL SpSbnI in 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, and 2 mM GSH with reservoir solution containing 4 M sodium formate. Crystals were flash frozen in liquid nitrogen.   41 Diffraction data was collected at the CLS on beamline 08B1-1 (165). SpSbnI crystallized in the space group P21212 with two molecules in the asymmetric unit. The structure was solved using molecular replacement with SbnI1-240 coordinates as a search model in PhaserMR from Phenix (168). The structure was manually edited using Coot (169) and refinement was performed with phenix.refine using TLS refinement using nine TLS groups. The refined structure has all 254 residues modeled for each protomer, three formates, and 119 water molecules. All residues were modeled but there was poor side-chain electron density for residues 94-142, 153-173, and 220-229 and poor main chain electron density for residues 103-105.  SpSbnI co-crystals were obtained using a protein solution consisting of ~11 mg/mL SpSbnI incubated with 10 mM ADP, 20 mM O-phospho-L-serine (OPS), and 10 mM MgCl2 in 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, and 2 mM GSH. Crystals were grown by sitting drop at 4°C in a 1:1 mixture of the aforementioned protein solution with reservoir solution containing 0.16 M calcium acetate, 0.1 M imidazole (pH 8), and 8% (w/v) PEG 8000. Crystals were briefly soaked in reservoir buffer supplemented with 30% (v/v) ethylene glycol for cryoprotection and flash frozen in liquid nitrogen. Diffraction data was collected at the Stanford Synchrotron Radiation Lightsource (SSRL) on beamline 9-2 and data was processed and integrated by AUTOXDS (171, 172). SpSbnI co-crystal crystallized in the space group P21212 with two molecules in the asymmetric unit. The structure was solved using molecular replacement with SpSbnI coordinates as a search model in PhaserMR from Phenix (168). The structure was manually edited using Coot (169) and refinement was performed with phenix.refine using TLS refinement using nine TLS groups. The refined structure has all 254 residues modeled for each protomer and 187 water molecules. Though the crystallization solution contained both products, ADP and OPS, only clear electron density was seen for the   42 diphosphate of ADP and a modeled magnesium ion for protomer A and the diphosphate, sugar, and magnesium ion for protomer B.  Data collection and refinement statistics for both structures are summarized in Table 2-4. Structure figures were generated in PyMOL (The PyMOL Molecular Graphics System, Version 1.8 Schrödinger, LLC). Domain analysis was done using the Dali server for comparison of the protein structure against structures in the PDB (173).   Table 2-4 X-ray diffraction data collection and refinement statistics for SpSbnI structures.   SpSbnI  SpSbnI + ADP Data Collectiona Resolution Range (Å) 45 - 2.10 (2.15 – 2.10) 37.8 – 1.9 (1.97 – 1.9) Space group P21212 P21212 Unit cell dimensions a, b, c (Å) 78.8, 111.4, 67.7 101.6, 73.1, 88.4 α, β, γ (°) 90, 90, 90 90, 90, 90 Unique reflections 35,448 (2,538) 52,399 (5081) Completeness (%) 99.8 (98.8) 99.7 (98.7) Redundancy 9.0 (8.4) 6.6 (6.0) Average I/σI 16.9 (2.1) 17.7 (2.1) Rmerge 0.087 (1.061) 0.046 (0.867) CC1/2 0.999 (0.809) 1.000 (0.890) Wilson B-factor (Å2) 39.3 37.5 Anisotropy 0.41 0.09 Refinement Rwork (Rfree) 0.217 (0.259) 0.207 (0.242) Number of water molecules 119 187 RMSD bond length (Å) 0.004 0.006 Average B-values (Å2) 67.0 59.4 Ramachandran plot (%) Most favored regions 98.0 96.4 Disallowed regions 0 0 PDB ID 5UJD  a Data collection values in parentheses represent the data for the highest resolution shell.    43 2.2.3 ChdC structure determination ChdC crystals were grown by sitting drop vapor diffusion at room temperature in a mixture containing 1 µL ~20 mg/mL ChdC in 50 mM Tris (pH 7.5), 100 mM NaCl with 1 µL reservoir solution containing 0.2 M potassium thiocyanate, 12% (w/v) PEG 3350, and 0.2 µL 0.2 M hexamine cobalt (III) chloride. Crystals were directly flash frozen in liquid nitrogen. Diffraction data was collected at the SSRL beamline 9-2. Data was processed and integrated using AUTOXDS (171, 172). ChdC crystallized in the space group P21212 with ten molecules in the asymmetric unit. The ChdC structure from Geobacillus stearothermophilus (PDB ID: 1T0T) was used to generate a homology model using SWISS-model for coordinates as a search model for molecular replacement using PhaserMR from Phenix (168). The structure was manually edited using Coot (169) and refinement was performed with phenix.refine. The refined structure has between 242-246 residues, out of 250, modeled for each protomer. Residues in the loop (residues 112-120) had poor electron density that varied modestly between protomers. The ChdC structure had 15 hexamine cobalt(III) molecules modelled at crystallographic symmetry axes and the inside edge of domain I and a total of 439 water molecules modelled. Anomalous signal was detected where the hexamine cobalt (III) chloride molecules were modeled. Data collection and refinement statistics are summarized in Table 2-5.  ChdC and heme co-crystals were obtained using a protein solution consisting of ~20 mg/mL ChdC incubated with equimolar heme in 50 mM Tris (pH 7.5), 100 mM NaCl buffer solution. Crystals were grown by sitting drop at room temperature in a 1:1 mixture of the aforementioned protein solution with reservoir solution containing 0.1 M MES (pH 6) and 8% (w/v) PEG 6000, with a final pH 6. Crystals were briefly soaked in reservoir buffer supplemented with 20% (v/v) glycerol for cryoprotection and flash frozen in liquid nitrogen. Diffraction data was collected at   44 the SSRL beamline 9-2. Data was processed to 6.5 Å and integrated using Mosflm (174) and scaled using Aimless (175, 176). The ChdC pentamer structure previously solved from S. aureus was used as a search model for molecular replacement using PhaserMR from Phenix (168). ChdC with heme crystallized in the space group I23 with five molecules in the asymmetric unit as a homopentamer. Due to the low resolution of the dataset, no atomic detail could be discerned.  Table 2-5 X-ray diffraction data collection and refinement statistics for ChdC structures.   ChdC ChdC + heme Data Collectiona  Resolution Range (Å) 38.4 – 2.52 (2.61 – 2.52) 55.8 – 6.52 (6.75 – 6.52) Space group P21212 I23 Unit cell dimensions  a, b, c (Å) 123.4, 138.2, 156.8 208.8 Unique reflections 91,292 (8844) 3075 (305) Completeness (%) 99.1 (95.7) 99.9 (99.3) Redundancy 5.1 (4.8) 17.6 (19.0) Average I/σI 12.8 (1.7) 14.5 (3.8) Rmerge 0.080 (0.684) 0.109 (0.859) CC1/2 0.991 (0.702) 0.997 (0.924) Wilson B-factor (Å2) 44.9 401.4 Anisotropy 0.22 1.30 Refinement  Rwork (Rfree) 0.22 (0.27) 0.21 (0.45) Number of water molecules 439 0 RMSD bond length (Å) 0.007 0.015 Average B-values (Å2) 51.2 447.8 Ramachandran plot (%)  Most favored regions 97 90.9 Disallowed regions 0.3 0.9 aData collection values in parentheses represent the data for the highest resolution shell.    45 2.3 Bioinformatic analysis 2.3.1 Genomic neighborhood analysis of SbnI and homologs Protein homology between genomic regions carrying SbnI orthologs was plotted using a custom Biopython script bio.links.py based on output from BLASTP 2.2.2.28 (e-value ≤ 1.00e-40) (https://github.com/minevskiy/bioinformatics). Species used for comparison were found by BLAST search of SbnA or SbnI amino acid sequence and STRING analysis of SbnI. Orthologous genes are indicated in the same color. A multiple sequence alignment of SbnI orthologs in the genomic neighborhood analysis was generated using T-COFFEE Expresso and visualized using Jalview (177–179). A second multiple sequence alignment of staphylococcal homologs was also generated using T-COFFEE Expresso and visualized using Jalview (177–179). Staphylococcal species for comparison were found by a BLAST search of S. aureus SbnI protein sequence against the Staphylococcus group (taxid:90964).    2.3.2 SbnI1-240 structure and conservation analysis Domain analysis was performed using the Dali server for comparison of the protein structure against structures in the PDB (173). Sequence conservation was mapped onto the SbnI1-240 structure using ConSurf (180). The multiple sequence alignment used for the analysis was generated using default ConSurf parameters and the S. aureus SbnI amino acid sequence as the search sequence. Superimposition with SerK (PDB ID: 5X0E) was performed using the align function in PyMOL (181).     46 2.3.3 SpSbnI conservation and molecular surface electrostatics analyses Conservation pattern of SpSbnI monomer and dimer generated using ConSurf (180). The multiple sequence alignment used for the analysis was generated using default ConSurf parameters and the S. pseudintermedius SbnI amino acid sequence as the search sequence. 17 sequences were used. Electrostatic potential molecular surface map of SpSbnI was generated using default settings in the APBS plug-in, an interface to the Adaptive Poisson-Boltzmann Solver (APBS) (182). The results were visualized in PyMOL. The electrostatic potential was set to ±5 kT/e so a blue color indicates regions of positive potential (> +5 kT/e) and red represents negative potential (< -5kT/e) values. Superimposition of SpSbnI with SbnI1-240 or SerK (PDB ID: 5X0E) was performed using the align function in PyMOL (181).  2.3.4 Generation of model of full-length S. aureus SbnI and heme-binding A model of dimeric SbnI was generated using SWISS-MODEL (183). The SpSbnI dimeric crystal structure was used as a template structure. Molecular docking was used to model where heme binds to SpSbnI dimer and model of SbnI dimer. The heme ligand and SpSbnI and SbnI receptor PDB files were converted to PDBQT format using AutoDock Tools (184). The docking parameters were set with default values and the size of the grid box was set as 56 Å × 58 Å × 42 Å to encompass the SpSbnI or SbnI protomer. The molecular docking screening was performed by AutoDock Vina (185).  2.3.5 ChdC structural superimposition and analysis A structure similarity search against structures in the PBD was conducted using the Dali server (173). Sequence conservation was mapped onto the ChdC structure using ConSurf (180).   47 The multiple sequence alignment used for the analysis was generated using default ConSurf parameters and the S. aureus ChdC amino acid sequence as the search sequence. Superimposition of S. aureus ChdC with G. stearothermophilus Mn(III) coproheme bound ChdC (PDB ID: 5T2K) were performed using align function in PyMOL (181). A multiple sequence alignment of ChdC sequences from species with structures available in the PDB was generated using Clustal Omega (186).   2.4 Heme reconstitution of proteins Purified apo-SbnI, IsdI, IsdG, or ChdC was incubated for 20-30 min at 4°C with 1.2 M equivalents of heme solution. Heme was prepared fresh by dissolving in 0.1 M NaOH and adjusting the pH to 8.5-9. Excess and non-specifically bound heme was removed by gel filtration chromatography on a Sephadex G-25 column (1 cm x 4 cm). The concentration of holo-protein was determined by quantifying heme using pyridine hemochrome assay using ε418 extinction coefficient of 191.5 mM-1 cm-1 as previously described (187). For SbnI, a bicinchoninic acid (BCA) assay was used for protein quantification (188) to calculate heme binding stoichiometry.   2.5 Determination of SbnI oligomeric state in solution Samples of SbnI, SbnI with equimolar heme, and SbnI1-240 were analyzed by dynamic light scattering (DLS) using a DynaPro Plate Reader (Wyatt Technologies). Protein was diluted to 0.5 mg/mL with 50 mM HEPES (pH 7.4), 300 mM NaCl, 5% (v/v) glycerol, and 2 mM GSH and results were generated based on averaging five, 5 second acquisitions. Data were collected at room temperature. Values reported are an average of data collected.    48 2.6 UV-visible spectroscopic analysis of SbnI OPS production using SbnA UV-visible spectra were collected using Varian Cary 50 UV-visible spectrophotometer. SbnA-PLP spectrum was recorded at a concentration of 15 µM in 50 mM HEPES pH 7.4, 100 mM NaCl, 5% (v/v) glycerol. The spectrum of SbnA aminoacrylate aldimide complex was recorded immediately after addition of 30 µM OPS (92). The spectral shift observed when SbnA binds OPS was used to evaluate OPS production by SbnI. The spectrum of 15 µM SbnA in 50 mM HEPES pH 7.4, 100 mM NaCl, 5% (v/v) glycerol, 20 mM MgCl2, 25 mM L-serine, and 5 mM ATP was recorded before and after the addition of 15 µM SbnI, SbnI E20A, SbnI D58A, or SbnI1-240. Phosphate donor specificity was also examined using 5 mM ADP in place of ATP.   2.7 Heme-binding by SbnI and variants UV-visible spectra were recorded in a conventional spectrophotometer (Cary50) with the optical path length of 1 cm in a quartz cuvette. All spectra were measured at room temperature and reactions were carried out in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol. To examine heme binding by SbnI variants, 5 µM of SbnI, SpSbnI, SbnI1-240, SbnI H3A, SbnI C155A, SbnI C168A, SbnI C244A or SbnI H3A/C244A protein was mixed with 5 µM heme and spectra were immediately recorded. Further readings were taken as indicated and samples for the 1 or 2 h reading were incubated on ice and kept in the dark.   2.8 UV-visible spectroscopic analysis of oxidized, reduced, and CO bound forms of SbnI and SpSbnI bound to heme Reduced holo-SbnI and holo-SpSbnI was prepared by adding 2-3 mg of sodium hydrosulfite (dithionite) to a 1 mL of 5 µM protein. The UV-visible spectrum was measured immediately.   49 Investigation of heme-bound SbnI and SpSbnI interaction with carbon monoxide was carried out by bubbling 950 µL of buffer in quartz cuvette with CO for 5 min, holo-SbnI or holo-SpSbnI was added to obtain a final concentration of 5 µM holo-protein, and 2-3 mg of dithionite was added to the cuvette. The headspace of the cuvette was exchanged with CO and the cuvette was subsequently sealed, inverted to mix, and UV-visible spectra were recorded. All spectra were measured at room temperature and reactions were carried out in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol.  2.9 UV-visible spectroscopic analysis of heme transfer reactions All spectra were measured at room temperature and reactions were carried out in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol.  2.9.1 Heme transfer from IsdI to SbnI and variants Spectral analysis of IsdI heme transfer to SbnI, SbnI H3A, SbnI C155A, SbnI C168A, SbnI C244A and SbnI H3A/C244A constructs was conducted by first measuring the absorbance of 5 µM holo-IsdI. Subsequently, 5 µM of apo-SbnI variant was added to the cuvette, mixed and spectra were recorded immediately.    2.9.2 Heme transfer from IsdI to SbnI in the presence of IruO IsdI heme transfer to SbnI in the presence of IruO was performed with IsdI reconstituted with 0.5 molar equivalents of heme. Reactions were assessed by first measuring the spectrum of 5 µM holo-IsdI then adding a mixture of 5 µM SbnI, 5 µM IruO, and 100 µM NADPH (final concentration) and immediately measuring the spectrum every minute for 5 or 10 minutes.   50 Appropriate controls were carried out excluding SbnI, IruO, or NADPH. A control reaction using SbnI bound to equimolar heme and then adding IruO and NADPH was also measured.  2.9.3 Heme transfer from ChdC to IsdI, IsdG, and SbnI All spectra were measured at room temperature and reactions were carried out in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol. Spectral analysis of ChdC heme transfer to SbnI, IsdI, or IsdG constructs was conducted by first measuring the absorbance of 5 µM holo-ChdC. Subsequently, 5 µM of SbnI, IsdI, or IsdG was added to the cuvette, mixed and spectra were recorded immediately. IsdI heme transfer to ChdC was similarly conducted by first measuring the absorbance of 5 µM holo-IsdI followed by addition of 5 µM ChdC and spectra were recorded immediately.   2.9.4 Heme transfer from IsdG to SbnI All spectra were measured at room temperature and reactions were carried out in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol. IsdG heme transfer to SbnI was conducted by first measuring the absorbance of 5 µM holo-IsdG followed by addition of 5 µM SbnI and spectra were recorded immediately.  2.10 IsdI heme transfer to SbnI pulldown assay Strep-tagged IsdI (150 µL of 50 µM) was incubated with 45 µM heme prior to binding to Strep-tactin Superflow high capacity resin (streptactin beads) (50 µL suspended volume). Samples were washed with 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol to remove any unbound protein. Holo-IsdI bound to strep beads was subsequently incubated with   51 excess apo-SbnI, SbnI H3A, SbnI C244A, SbnI H3A/C244A, SbnI C168A, SbnI C155A, SbnI1-240, or SpSbnI (150 µl of 100 µM) or buffer as negative control for 5 minutes. Samples were centrifuged and supernatant (referred to as flowthrough) was removed and stored on ice. Samples were washed again with 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol prior to IsdI elution form the streptactin beads using 50 mM HEPES (pH 7.4), 100 mM NaCl, 5% (v/v) glycerol, 5 mM desthiobiotin. Streptactin beads were spun down and the supernatant (referred to as eluent) was removed and stored on ice. All samples were run on SDS-polyacrylamide gel and UV-visible absorption spectra were recorded. Quantification of IsdI heme transfer to SbnI variant was based on the percent of heme transferred from IsdI. The percentage was calculated based on the amount of holo-IsdI eluted when incubated with buffer compared to incubation with SbnI variant. No heme transfer is equal to the A412/A280 of holo-IsdI incubated with buffer. The amount of holo-IsdI was calculated based on the A412 (wavelength at which holo-IsdI absorbs maximally) to A280 ratio of the eluent containing IsdI after incubation with the SbnI variant. Statistical analyses were conducted using one-way ANOVA.   2.11 Stopped-flow kinetic analysis of enzyme heme off-rate All reactions were performed in 50 mM HEPES (pH 7.4), 300 mM NaCl, and 5% (v/v) glycerol at 20°C. The rates of dissociation of heme from SbnI, IsdI, IsdG, and ChdC were measured by single-wavelength stopped-flow spectroscopy with apomyoglobin as a heme scavenger (189). Apomyoglobin was prepared from myoglobin (Sigma-Aldrich) (190). Heme dissociation reactions were carried out with 2.5 µM holo-SbnI, holo-IsdI, holo-IsdG, or holo-ChdC (reconstituted with heme as described in Section 2.4) in one syringe and 12.5 µM, 25 µM, or 50 µM apomyoglobin in the second syringe. Reactions were monitored over time by recording   52 the absorbance at 408 nm, the maximal absorbance for holomyoglobin. 1000 time points logarithmically distributed over the time frame were acquired using Pro-Data SX software. The change in absorbance was plotted versus time and fit by a triple-exponential equation for SbnI and ChdC and a double-exponential equation for IsdI and IsdG to determine the first-order rate constants for heme dissociation. Off-rates were calculated from five independent reactions that were averaged.   2.12 Stopped-flow kinetic analysis of heme transfer reactions All reactions were performed in 50 mM HEPES (pH 7.4), 300 mM NaCl, and 5% (v/v) glycerol at 20°C.  2.12.1 IsdI heme transfer to SbnI IsdI heme transfer reactions to SbnI were carried out with 2.5 µM holo-IsdI in one syringe and 12.5 µM, 25 µM, 50 µM, or 100 µM apo-SbnI. The wavelength of maximal absorbance change was determined to be 426 nm based on a difference absorption spectrum between holo-IsdI and holo-SbnI. Reactions were monitored for 120 s at 426 nm. 1000 time points logarithmically distributed over the time frame were acquired using Pro-Data SX software. The change in absorbance was plotted versus time and fit by a triple exponential curve to determine the rate of heme transfer from IsdI to SbnI.  2.12.2 IsdG heme transfer to SbnI IsdG heme transfer reactions to SbnI were carried out with 2.5 µM holo-IsdG in one syringe and 12.5 µM, 25 µM, 50 µM apo-SbnI. The wavelength of maximal absorbance change was   53 determined to be 430 nm based on a difference absorption spectrum between holo-IsdG and holo-SbnI. Reactions were monitored for 300 s at 430 nm. 1000 time points logarithmically distributed over the time frame were acquired using Pro-Data SX software. The change in absorbance was plotted versus time and fit by a triple exponential curve to determine the rate of heme transfer from IsdG to SbnI.  2.12.3 ChdC heme transfer to SbnI, IsdI, and IsdG ChdC heme transfer reactions to SbnI, IsdI, and IsdG were carried out with 2.5 µM holo-ChdC in one syringe and 12.5 µM, 25 µM, 50 µM of apo-SbnI, apo-IsdI, or apo-IsdG. ChdC heme transfer reactions were monitored for 180 s at 430 nm for apo-SbnI, 240 s at 416 nm for apo-IsdI, and 240 s at 417 nm for apo-IsdG. 1000 time points logarithmically distributed over the time frame were acquired using Pro-Data SX software. The change in absorbance was plotted versus time and fit by a triple exponential curve to determine the rates of heme transfer from ChdC to SbnI and ChdC to IsdG. Data for ChdC heme transfer to IsdI was fit by a double exponential curve to determine the rate of heme transfer.  2.13 Fluorescence quenching of SbnI Heme binding by SbnI was measured by intrinsic tryptophan fluorescence quenching by heme. Fluorescence-detected heme titrations into SbnI were completed using 250 nM samples of SbnI in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol. Heme was added to the buffered protein solution in 50-250 nM increments and allowed to reach equilibrium before readings were measured. The titrations covered a heme concentration range of 50-1000 nM. Fluorescence emission spectra were acquired for a 290 nm excitation using a Cary Eclipse   54 fluorescence spectrophotometer (Agilent Technologies). Emission spectra were acquired in the 305 to 400 nm range with a step size of 1 nm and slit widths of 10 nm. The dissociation constant (KD) was calculated from the decrease in the area under the fluorescence curve across 305 to 400 nm as a function of increasing heme concentration. The data was fit by an equation for nonlinear regression one-site binding kinetics using GraphPad Prism 7.0a.  2.14 HPLC analysis of SbnI kinase activity Kinase activity of SbnI was detected using HPLC to examine production of ADP from ATP. The reaction mixture was composed of 50 mM HEPES (pH 7.4), 100 mM NaCl, 2.5% glycerol, 50 mM L-serine, 0.25 mM ATP or ADP, 10 mM MgCl2, and 5 µM of SbnI, SbnI E20A, or SbnI D58A. The reaction was carried out for 1 hour at room temperature (22ºC). The protein was removed by centrifugation using a 3K Nanosep® column. The filtrate was 0.2 µm filtered and analyzed by HPLC using a Waters 2695 Separations HPLC module (Milford, MA) equipped with a Waters 2996 photodiode array detector and a Luna 3 µm PFP(2) 50 x 4.6 mm LC column (Phenomenex) using a linear gradient of 0 to 15% methanol in 0.1 M ammonium acetate, pH 4.5 over 10 minutes at 1 mL min-1. Analytes were detected by the absorbance at 258 nm.  2.15 31P NMR spectra of SbnI kinase reaction The reaction mixture contained 50 mM HEPES (pH 7.4), 100 mM NaCl, 2.5% (v/v) glycerol, 10 mM MgCl2, 48 mM L-serine, 5 mM ATP, and 5% D2O. NMR spectra were collected at 25 °C using a broadband frequency probe with Z-magnetic field gradient in a Bruker Avance III 500 MHz spectrometer. One-dimensional 31P spectra were recorded at different time points before and after addition of 4.8 µM SbnI until reaction completion. The spectra were referenced to   55 2,2,6,6-tetramethylpiperidine (TMP), which was set to 0 ppm. The spectra were processed and using TopSpin™ (Bruker). The chemical shifts of ATP (191) and OPS were assigned using reference spectra (Figure 2-1).    Figure 2-1 31P NMR spectra of ATP and OPS standards.  The buffer used contained 50 mM HEPES pH 7.4, 100 mM NaCl, 10 mM MgCl2, and 2.5% (v/v) glycerol.  2.16 Steady-state kinetic analysis of SbnI serine kinase activity 2.16.1 Pyruvate kinase/lactate dehydrogenase coupled assay ATP-dependent serine kinase activity of SbnI was measured using a pyruvate kinase/lactate dehydrogenase (PK/LDH) coupled assay. The assay is based on a reaction in which the regeneration of hydrolyzed ATP is coupled to the oxidation of NADH (192). The rate of NADH absorbance decrease at 340 nm (A340 nm = 6220 M-1 cm-1) is proportional to the rate of ATP conversion to ADP by SbnI-kinase activity. Coupled reactions contained 50 mM HEPES pH 7.4, 100 mM NaCl, 2.5% (v/v) glycerol, 10 mM MgCl2, 2 mM phosphoenolpyruvate, 1/50 of the final reaction mixture volume of PK/LDH enzyme (from rabbit muscle, Sigma-Aldrich, cat. P-0294), 5 mM ATP, and 100 mM L-serine. The mixture was incubated for 5 min to remove any   56 contaminating ADP. Continuous measurement at 340 nm was recorded for 2 minutes prior to addition of 0.5 µM SbnI, SbnI1-240, SbnI E20A, or SbnI D58A enzyme to start the reaction. The assay was run for 10 minutes. To determine kinetic parameters, the initial velocities of SbnI, heme-bound SbnI, and SbnI1-240 kinase reactions in the presence of varying concentrations of ATP with 100 mM L-serine and in the presence of varying concentrations of L-serine with 5 mM ATP were recorded. Phosphate donor specificity was investigated with 10 mM ADP and phosphate acceptor specificity was examined with 50 mM L-threonine, and 50 mM His-Ser dipeptide. In addition, alternative substrates, 50 mM L-Dap and 50 mM a-KG were tested as phosphate acceptors. All data was collected on a Varian Cary 50 UV-visible spectrophotometer at room temperature (22ºC) and a total of three replicates were collected for each reaction condition.  2.16.2 SbnA and phosphate release coupled assay SbnI kinase activity was also measured using an assay with SbnA. In a reaction mixture, SbnI-dependent OPS production was measured using SbnA and L-glutamate to release inorganic phosphate and ACEGA (95). A coupled enzymatic assay was used to detect inorganic phosphate release from OPS as previously described (193). The reaction mixture contained 50 mM HEPES pH 7.4, 100 mM NaCl, 2.5% (v/v) glycerol, 10 mM MgCl2, 10 mM ATP, 100 mM L-serine, 0.2 U purine nucleoside phosphorylase, 400 µM 2-amino-6-mercapto-7-methylpurine riboside, 5 µM SbnA, and 2 mM L-glutamate. The mixture was incubated for 10 min to remove any contaminating inorganic phosphate and establish a baseline. Continuous measurement at 360 nm was recorded for 2 minutes prior to addition of 0.5 µM SbnI or SbnI1-240 enzyme to start the reaction. The assay was run for 15 minutes. To determine kinetic parameters, the initial   57 velocities of SbnI kinase reactions in the presence of varying concentrations of ATP with 100 mM L-serine and in the presence of varying concentrations of L-serine with 10 mM ATP were recorded. All data was collected on a Varian Cary 50 UV-visible spectrophotometer at room temperature (22ºC) and a total of three replicates were collected for each reaction condition. The concentration of inorganic phosphate release from OPS was determined using the extinction coefficient A360 nm = 11000 M-1cm-1 (193).  2.16.3 Software for kinetic analysis Data were fit by nonlinear regression using a Michaelis-Menten model in GraphPad Prism version 7.0a.  2.17 S. aureus bacterial strains  Experiments were performed with a derivative of S. aureus USA300 LAC cured of the 27-kb plasmid encoding macrolide resistance (194). The plasmid-cured USA300 LAC is referred to as USA300 throughout. Transposon insertion mutants JE2 serC::ΦNΣ; EmR (SAUSA300_1669) and  JE2 sbnI::ΦNΣ; EmR (SAUSA300_0126) were obtained from the Nebraska Transposon Mutant Library (NTML) containing the resistance cassette ermB, which confers resistance to erythromycin (25). Transposons were transduced to USA300 background strain using phage 80α and confirmed by PCR using published methods (195). USA300 serC::ΦNΣ and  USA300 sbnI::ΦNΣ transposon insertion mutants are referred to as serC and sbnI throughout (Table 2-4). S. aureus RN6390 sirA mutant and RN6390 htsABC mutant strains were provided by Dr. David Heinrichs, University of Western Ontario (76).    58 Table 2-6 S. aureus strains used in this study.  S. aureus strains Description Source or reference USA300 USA300 LAC cured of antibiotic resistance plasmid (194) JE2 serC::ΦNΣ JE2 serC::ΦNΣ; EmR (SAUSA300_1669) (25) JE2 sbnI::ΦNΣ JE2 sbnI::ΦNΣ; EmR (SAUSA300_0126) (25) serC USA300 serC::ΦNΣ; EmR (SAUSA300_1669) This study sbnI USA300 sbnI::ΦNΣ; EmR (SAUSA300_0126) This study sirA RN6390ΔsirA::KmR; SB transport-deficient mutant (104) htsABC RN6390ΔhtsABC:TcR; SA transport-deficient mutant (76) aEmR, KmR, and TcR designate resistance to erythromycin, kanamycin, and tetracycline respectively.  2.18 S. aureus bacterial growth curve to assess serine auxotrophy  Bacterial growth curves to test serine auxotrophy were performed in a chemically-defined medium with 0.4% (w/v) glucose (CDMG) as previously described (90), with or without L-serine. Briefly, colonies of wildtype USA300 or serC transposon insertion mutant were inoculated from tryptic soy agar into 2 mL CDMG overnight at 37ºC. Cells were normalized to an OD600 of 0.1 and washed twice with CDMG lacking L-serine (CDMG-L-Ser) and 5 µL of the resuspension was used to inoculate 200 µL aliquots of CDMG or CDMG-L-Ser in 96 well plates. Cultures were grown in a TECAN plate reader for 24 hours at 37ºC with a 10 second shake every 10 min, and the OD600 was measured every 30 min. Data are representative of three independent experiments, and error bars signify standard error of the mean.  2.19 Disc diffusion bioassays to detect siderophore production Concentrated spent culture supernatants were prepared from 10 mL cultures of S. aureus USA300, serC transposon insertion mutant, and sbnI transposon insertion mutant grown for 16 hrs in Chelex-100 treated Tris minimal succinate (TMS), as previously described (79), in a flask   59 to volume ratio of 10:1 at 37ºC with shaking at 200 rpm without antibiotic selection. Growth was assessed via OD600 and culture densities normalized. Bacterial cells were pelleted by centrifugation and culture supernatants were filter sterilized and lyophilized overnight. Dried material was resuspended in 0.5 mL sterile ddH2O. To assess growth promotion of concentrated culture supernatants, S. aureus strain RN6390 sirA mutant (growth of this mutant is dependent on SA in supernatant) or htsABC mutant (growth of this mutant is dependent on SB in supernatant) derivatives, as previously described (76), were seeded into TMS agar containing 10 µM ethylenediamine-N, N’-bis(2-hydroxyphenylacetic acid) (EDDHA) to 2 x 105 cells/mL. 10 µL of concentrated supernatant was applied to sterile paper discs placed on TMS agar containing the seeded reporter strains, and growth radii about the discs were measured after 24 hr incubation at 37ºC. The reported growth radius has the disc radius (3 mm) subtracted. Statistical analyses were conducted using 2-way ANOVA.   60 Chapter 3: SbnI is a free serine kinase that generates O-phospho-L-serine for staphyloferrin B biosynthesis in S. aureus 3.1  Introduction Primary metabolites generally serve as the source of building blocks to generate siderophores and can be first chemically modified prior to incorporation into the siderophore. In addition to the synthetases required for assembly of the siderophore, NIS biosynthetic gene clusters generally encode enzymes that synthesize the necessary precursors to be assembled by the NIS synthetases (88). The SB biosynthetic operon consists of nine genes, sbnABCDEFGHI. The enzymatic capacity required for precursor synthesis, production, and export of SB can be completed by the enzymes encoded by sbnA-H using the four precursors: acetyl-CoA, oxaloacetate, O-phospho-L-serine (OPS), and L-glutamate (50, 87, 92).  These can be produced by S. aureus from two of the four most abundantly available nutrients in human serum, glucose and L-glutamine (102). L-glutamate can be generated from L-glutamine by glutamate synthase (gltBC). Both acetyl-CoA and oxaloacetate can be generated directly from glycolysis. Lastly, OPS is an intermediate in the serine biosynthesis pathway stemming from 3-phosphoglycerate produced in glycolysis and is predicted to be the source of OPS for SB biosynthesis (Figure 3-1) (90, 95).     61  Figure 3-1 Metabolic pathways for the production of L-Dap and a-KG from glucose or L-serine in S. aureus.  Highlighted in light green is the contribution of SbnI in this pathway, which feeds into the previously characterized SB biosynthetic pathway (light blue). 3P stands for 3-phospho.  SbnI, the ninth gene product, is a heme-responsive transcriptional regulator for SB production (98). SbnI is required for full expression of sbnD-H and thus controls SB-mediated iron acquisition. Based on primary sequence analysis, homology was not detected between SbnI and any characterized transcription factors or heme-binding proteins (98), but it is annotated as containing an N-terminal ParB-like domain. ParB is an essential component of the chromosome segregation system in bacteria (196). However, the conserved N-terminal ParB-like domain in SbnI is not responsible for interaction with DNA in ParB (197), leaving the role of the ParB-like domain in SbnI unknown. The aim of this study was to gain insight into how SbnI functions using X-ray crystallography. The SbnI structure revealed striking structural homology to a recently characterized free serine kinase, SerK, from the archaea T. kodakarensis (198). SerK can   62 phosphorylate free L-serine using ADP to generate OPS for cysteine biosynthesis (199). Herein, we demonstrate that SbnI is also a free serine kinase that uses ATP to phosphorylate L-serine to yield OPS and ADP (Figure 3-1). The structure of SbnI, supported by site-directed mutagenesis, suggests that it follows a similar open-close reaction mechanism as proposed for SerK. Additionally, SbnI-generated OPS can be used by SbnA in vitro and serves as the in vivo source of OPS for SB production. To our knowledge, this is the first example of a bacterial free serine kinase and the first described free serine kinase that is ATP dependent. This function earns SbnI an enzymatic role in the SB biosynthetic pathway in addition to its heme-dependent transcriptional regulatory function.  3.2 Results 3.2.1 Structure determination of SbnI Full length (254 amino acid) SbnI was not amendable for X-ray crystallography due to its propensity to precipitate. To improve solubility and stability in solution, several expression constructs were made containing varying N- and C-terminal truncations based on predicted secondary structure and disorder identified using PSIPRED and DISOPRED, respectively (200, 201). One construct containing a 14-amino acid C-terminal truncation, SbnI1-240, had improved stability in solution and produced well-diffracting crystals. The structure of selenomethionine-labelled SbnI1-240 was determined to 2.5 Å resolution using single wavelength anomalous dispersion in space group P31with one molecule in the asymmetric unit (Figure 3-2A). Data collection and refinement statistics are summarized in Table 2-3. All 240 residues were modeled with 98% of residues in the most favored regions of the Ramachandran plot.     63  Figure 3-2 Structure of SbnI1-240. The overall fold of SbnI1-240 is shown as a cartoon and divided into two domains with N- and C-termini and alpha helices labelled. Domain I shown in teal contains the conserved, core ParB/Srx domain (a2 and a3) and domain II is colored raspberry. (B) Structural superimposition of SbnI1-240 (raspberry), and the “open” conformation of SerK (yellow, PDB ID: 5X0B). (C) Superimposition with the “closed” conformation of SerK (blue, PDB 5X0E) in cartoon putty representation, where the size of the tube depends on the B-factor in each structure. (D) Structural superimposition of SbnI1-240 (raspberry), and SerK (yellow, PDB ID: 5X0E). Selected active site residues, AMP, and OPS are drawn as sticks. Mg2+ is drawn as a green sphere, O, N, and P atoms colored red, blue, and orange, respectively. AMP and OPS carbons are colored cyan.  SbnI1-240 is comprised of two domains, domain I and domain II. Domain I consists of residues M1-Q83 and I205-A240 and domain II is composed of residues Y84-N204. Domain I includes a conserved core ParB/Srx fold, corresponding to the annotated ParB-like domain based on primary sequence. This fold has been described in a functionally diverse ParB/Srx superfamily of proteins. Members are found in varied biological contexts and thus far are described to bind a nucleotide for kinase, ATPase, or DNase activity (199, 202). To our knowledge, no ParB/Srx   64 family member has been found to be directly involved in siderophore biosynthesis. The ParB/Srx core domain is comprised of a 4-strand mixed β-sheet and two α-helices, a2 and a3 (Figure 3-2A) and contains an absolutely conserved GXXR motif, GVHR59-62 in SbnI.  Domain II is composed of a mixed a/b fold with a central 4-stranded antiparallel b-sheet surrounded by 5 a-helices. A pair of antiparallel b-strands abut the main sheet and serve as a linker to Domain I.  B-factor analysis reveals that domain II has a relatively high average B-factor of 109 Å2, compared to 75 Å2 in domain I, suggesting domain II has more disorder in the crystal and the domains are connected by a flexible linker.  To gain functional insight into SbnI, a search of the SbnI1-240 structure against structures in the PDB was performed with the Dali server. Five proteins all belonging to the functionally diverse superfamily of ParB/Srx proteins were identified (Z score < 3.9). The most striking observation was the high structural similarity SbnI shared with the top search result, SerK (PDB ID: 5X0B). Superimposition of SbnI1-240 with SerK (PDB ID: 5X0B) using PDBeFOLD has a root mean square distance (RMSD) of 2.0 Å for 194 Cα despite sharing only 19% amino acid sequence identity across the aligned residues (Figure 3-2B). The other structures also share modest sequence identity (12-23%) and include sulfiredoxin (Srx) from Homo sapiens (PDB ID: 2RII, RMSD of 2.7 Å across 84 residues), chromosome partitioning protein (ParB) from Sulfolobus solfactaricus (PDB ID: 5K5D, RMSD of 2.5 Å across 71 residues), oncogenic suppressor (Osa) from Shigella flexneri (PDB ID: 4OVB, RMSD of 3.7 Å across 83 residues), and chromosome segregation protein (Spo0J) from Thermus thermophilus (PDB ID: 1VZ0, RMSD of 5.4 Å across 76 residues) (198, 202–205).  SerK is a free serine kinase from T. kodakarensis that uses ADP to phosphorylate L-serine to generate OPS for cysteine biosynthesis. Of the proteins annotated in the ParB/Srx family, SerK is   65 the only identified kinase though Osa and Srx both possess ATPase activity (202, 206). Overall, the structures of SerK and SbnI are very similar. The SerK domain II has a high average B-factor and superimposition of SbnI1-240 with the SerK structure in a “closed” conformation (PDB ID: 5X0E) suggests how SbnI may possess similar domain flexibility in solution (Figure 3-2C). Additionally, structural superimposition and multiple sequence alignments revealed that several residues important for substrate and product binding identified in the SerK crystal structure (PDB ID: 5X0E) are conserved in SbnI. Moreover, the active site architecture is highly conserved between the two proteins (Figure 3-2D, Figure 3-3). Of the active site residues, SerK Glu30 was identified as a catalytically essential residue and Asp69 is required for magnesium ion binding; site-directed mutagenesis of either of these residues abolished SerK kinase activity (198). The homologous residues in SbnI are Glu20 and Asp58.    66  Figure 3-3 Sequence alignment of SbnI with SerK and SbnI homolog proteins generated using T-Coffee Expresso.  The colored triangles represent the proposed catalytic residue (red), residues recognizing adenosine of AMP, ADP, or ATP (blue), the residue interacting with the magnesium ion (black), the residues that interact with the phosphate group of the OPS, AMP, ADP, or ATP (green), and the residues that interact with the Ser moiety of OPS (orange), as proposed by Nagataa et al. (198). Abbreviations (gene): TK, SerK from T. kodakarensis (TK0378); Sa, SbnI from S. aureus (SAUSA300_0126); Sp, SbnI homolog from S. pseudintermedius (UH47_00825); Bbr, SbnI homolog from Brevibacillus brevis (BGP74_RS01830); Pl, SbnI homolog from Paenibacillus larvae (B7C51_RS23590); Bba, SbnI homolog from Bacillus badius (A3781_RS10685); Mm, SbnI homolog from Marininema mesophilum (BLV90_RS00780); Mn, SbnI homolog from Methylobacterium nodulans (MNOD_RS32840); Rs, SbnI homolog from Ralstonia solanacearum (BC350_RS18860); and Sd, SbnI homolog from Shewanella denitrificans (SDEN_RS03055).    67 The genomic context of sbnI was analysed to compare with the genomic neighborhoods of homologs. All staphylococcal sbnI homologs are part of the nine gene SB biosynthetic cluster. More distant homologs co-occur with sbnA and sbnB homologs, either alone or in combination with different putative siderophore biosynthetic enzymes (Figure 3-4). More distant SbnI homologs are shorter and alignments suggest that they have an abbreviated domain II (Figure 3-3). Interestingly, a putative sbnI ortholog was identified upstream of the sbnA-H gene locus in Ralstonia solanacearum, which was previously thought to lack a SbnI homolog but still produce SB (207). A multiple sequence alignment of SerK and SbnI homologs used in the genomic neighborhood analysis reveals that certain key residues important for catalysis, substrate, and product binding identified in SerK are fully conserved (Figure 3-3). Notably, these include the SerK catalytic residue Glu30, magnesium ion binding residue Asp69, residues implicated in interacting with the β phosphate of ADP or phosphate group of OPS, His72 and Arg73, and residues that interact with the serine moiety of OPS, Trp102 and Thr223 (Figure 3-3). More variability is seen with the SerK residues interacting with the adenosine group, raising the possibility that SbnI and other homologs may use a different phosphate donor or binding-mode. Overall, the genomic neighborhood and sequence analyses suggest that free serine kinases are found in diverse species belonging to Firmicute and Proteobacteria phyla.    68   Figure 3-4 Illustration of gene neighborhoods containing SbnI homologs from diverse species from Firmicute and Proteobacteria phyla.  Each predicted gene is represented by an arrow showing the direction of transcription. The S. aureus sbn locus is labelled by gene name and homologous genes are indicated in the same color. Grey links connect protein homologs with e-value≤1e-40. This figure highlights that SbnI homologs appear in the same genomic context as SbnA and SbnB homologs in these bacterial genomes. The bottom scale shows the length of depicted genomic regions in nucleotide base pairs. Region coordinates used for each species are as follows (GenBank:nucleotide region): Staphylococcus aureus USA300 FPR3757 (CP000255.1:134324-145881), Staphylococcus pseudintermedius E104 (LAWU01000001.1:151604-163186), Brevibacillus badius NBRC 110488 (NZ_BDFB01000004.1:355053-396579), Paenibacillus larvae SAG 10367 (NZ_CP020557.1:4429364-4439754), Bacillus badius DSM 5610 (NZ_LVTO01000018.1:5060-15111), Marininema mesophilum DSM 45610 (NZ_FNNQ01000001.1:200002-206967), Methylobacterium nodulans ORS 2060(NC_011894.1:7032971-7044380), Ralstonia solacearum CQPS-1 (NZ_CP016915.1:169574-186973), and Shewanella denitrificans OS217 (NC_007954.1:663307-674451).    69 Consurf analysis of SbnI1-240 was used to map conserved regions to the molecular surface (Figure 3-5). Highly conserved residues including those that form the kinase active site delineate a groove between domain I and II (Figure 3-5AB). Structural alignment with the structure of the SerK ternary product complex (PDB ID: 5X0E) revealed that among SbnI homologs, the putative active site is highly conserved, while the remainder of the protein surface is variable (Figure 3-5C).   Figure 3-5 Conservation of surface residues of SbnI1-240 generated using ConSurf. (A, B) Ribbon and surface representations of SbnI1-240 have AMP and OPS modelled based on a structural alignment with the SerK ternary product complex (PDB ID: 5X0E). (C) Surface representation of SbnI1-240 after 180° rotation about the x-axis. Conserved amino acids are colored bordeaux, residues of average conservation are white, and variable amino acids are turquoise.  3.2.2  SbnI is a dimer in solution The oligomeric state of SbnI was analyzed using dynamic light scattering (DLS). Since SbnI contains 7 Cys residues, the analysis was conducted in the presence of GSH as a reductant. The calculated molecular weight based on amino acid sequence of full-length SbnI is 30 kDa and the measured molecular weight by DLS was 61 ± 6 kDa with an average of 24 ± 7% polydispersity, implying it predominantly forms a dimer in solution (Figure 3-6). The molecular weight of SbnI1-240 by DLS was 28 ± 3 kDa with 34 ± 1% polydispersity (Figure 3-6). SbnI1-240 has a calculated weight of 28 kDa, implying it is primarily a monomer in solution. These data indicate   70 that the C-terminal 14 amino acids excluded from the SbnI1-240 construct are important for dimerization of the full-length protein.   Figure 3-6 Representative DLS results for analysis of SbnI and SbnI1-240 oligomerization state.  Percent mass-weighted size distribution of (A) SbnI and (C) SbnI1-240 with calculated molecular weight (Mw) and polydispersity (Pd) are shown. The correlation function data measured for (B) SbnI and (D) SbnI1-240.   3.2.3 SbnI is a serine kinase that uses L-serine and ATP to generate OPS OPS is a substrate for SbnA in SB biosynthesis (92, 95), lending support to our hypothesis that SbnI produces OPS for use by SbnA. To thus test if SbnI is capable of producing OPS, the   71 spectral changes that occur when SbnA binds OPS were used to assay SbnI activity. SbnA has a characteristic absorption maxima at 412 nm attributed to an internal Schiff base formed between its pyridoxal 5’-phosphate (PLP) cofactor and an active site lysine. Adding OPS to SbnA causes a rapid change in UV-visible spectra with the appearance of absorption peaks at 324 nm and 467 nm (Figure 3-7A), characteristic of the formation of an external aminoacrylate intermediate (92). This spectral change is specific to OPS and does not occur with O-acetyl-L-serine or L-serine (92). SbnA incubated with SbnI, L-serine, and ADP resulted in no change in the UV-visible spectrum of SbnA. However, SbnA incubated with SbnI, L-serine, and ATP led to a shift in the UV-visible spectra indicative of OPS production and reaction with SbnA-PLP to form the external aminoacrylate (Figure 3-7BC). No spectral change was observed when SbnI was omitted indicating that only the SbnI enzymatic product could react with SbnA-PLP. Therefore, we conclude that the reaction product is most likely OPS and SbnI activity is ATP-dependent.   Figure 3-7 Detection of O-phospho-L-serine produced by SbnI via reaction with SbnA.  (A) UV-visible absorption spectra of SbnA and SbnA in complex with OPS. (B) Shown are spectra of SbnA incubated with SbnI-generated OPS from L-serine and ATP but not (C) ADP. SbnI variants (D) E20A and (E) D58A are defective in producing OPS as indicated by no change in the SbnA absorption spectra. (F) SbnI1-240 is capable of producing OPS but at a slower rate compared to full-length SbnI.    72 Phosphate acceptors, alternative to L-serine, were tested using a pyruvate kinase/lactate dehydrogenase (PK/LDH) assay for detection of ATP conversion to ADP. L-threonine, α-KG, and L-Dap were not phosphate acceptors (data not shown). Additionally, SbnI did not phosphorylate the serine residue in a His-Ser dipeptide (data not shown).  SbnI-mediated conversion of ATP to ADP and generation of OPS were monitored using HPLC and 31P NMR. Incubation of SbnI with ATP and excess L-serine led to turnover of ATP to ADP as detected by HPLC (Figure 3-8A) indicating SbnI has ATPase activity. Unlike SerK, no turnover of ADP to AMP was detected with ADP as a phosphate donor by HPLC (Figure 3-8A). ADP was not generated when L-serine was excluded, indicating that this activity requires the presence of L-serine.    Figure 3-8 HPLC analysis of the nucleotide reaction products.  HPLC trace of ATP, ADP, and AMP standards (black lines) with retention times 1.4, 1.5, and 2.9 min, respectively. (A) SbnI, L-serine, and ADP (green line) or ATP (blue line), control reactions of ATP and L-serine excluding SbnI (red line) and of SbnI and ATP excluding L-serine (grey line). (B) Reactions of variants SbnI1-240 (green line), SbnI D58A (red line), or SbnI E20A (blue line) with ATP and L-serine.  31P NMR was also used to monitor SbnI-mediated ATPase activity and generation of OPS from L-serine. The observed disappearance of the ATP g-phosphate 31P signal with the   73 concomitant appearance of a 31P signal of OPS demonstrates transfer of the ATP g-phosphate to L-serine to yield ADP and OPS (Figure 3-9). The chemical shift of SbnI-generated OPS was consistent with the 31P NMR spectrum measured for an OPS standard (Figure 2-1). Attempts to obtain a co-crystal structure of SbnI with identified substrates (or ATP analog, adenyl-imidodiphosphate) or products, both in the presence and absence of heme, have not been met with success.   Figure 3-9 A stack plot of 31P NMR spectra for a single reaction of SbnI mediated conversion of L-serine and ATP to OPS and ADP.  The reaction was initiated by the addition of SbnI with a final concentration of 4.8 µM. The initial concentrations of ATP and L-serine were 5 and 48 mM, respectively. The reaction buffer contained 50 mM HEPES pH 7.4, 100 mM NaCl, 10 mM MgCl2, and 2.5% (v/v) glycerol.   74 3.2.4 SbnI active site variants The role of Glu20 and Asp58 in SbnI kinase function were tested by site-directed mutagenesis. Two mutants each containing a single alanine substitution, SbnI E20A and SbnI D58A, were generated. Using the PK/LDH assay, we determined that the mutants were incapable of turning over ATP to ADP in the presence of L-serine. Additionally, no ADP could be detected by HPLC in reactions containing the SbnI mutants incubated with ATP and L-serine (Figure 3-8B); together, these data allow us to conclude that these mutants are catalytically inactive. These data also correlate with SbnA-PLP UV-visible absorption spectra that demonstrated that these SbnI variants also do not produce OPS (Figure 3-7DE). The importance of these residues in catalysis is consistent with the reaction mechanism presented for SerK in which substrate binding promotes conformational closure positioning the catalytic Glu30 (Glu20 in SbnI) close to the hydroxyl group of bound L-serine. Glu30 is a catalytic base that deprotonates the hydroxyl group of L-serine. The deprotonated hydroxyl can then attack the phosphorus atom of the ADP b-phosphate to yield OPS and AMP (198). Our results suggest SbnI uses a similar two-ligand binding sequential mechanism, but instead uses L-serine and ATP to yield ADP and OPS.  3.2.5 Kinetic analysis of SbnI kinase activity To obtain kinetic parameters for the SbnI kinase activity, SbnI enzymatic turnover was monitored using an established coupled assay for ADP using PK/LDH. The steady-state kinetic parameters of SbnI reaction with ATP and L-serine were determined and are summarized in Table 3-1. The saturating concentration of L-serine was beyond conditions permissive to the assays and thus Km could not be accurately determined. A second assay to measure SbnI enzymatic turnover employed SbnA-dependent turnover of OPS coupled to a phosphate release   75 detection assay. SbnA is the in vivo acceptor of OPS generated by SbnI most likely to facilitate synthesis of SB substrates L-Dap and a-KG, in concert with SbnB. Using excess concentrations of SbnA, the coupled assay demonstrated that SbnA could use SbnI-generated OPS and supplied L-glutamate to generate its products, N-(1-amino-1-carboxyl-2-ethyl)-glutamic acid and inorganic phosphate. The kinetic parameters of SbnI reaction with ATP and L-serine using the SbnA coupled assay were determined and are summarized in Table 3-1. The kinetic parameters measured using both methods agree with rates and Km values in the same order of magnitude. SbnI mutants E20A and D58A were catalytically insufficient to accurately measure enzyme rates. Compared to SerK, SbnI has a Km for ATP one order of magnitude lower than SerK has for ADP and a higher Km for L-serine by 2-orders of magnitude.   Table 3-1 Apparent steady-state kinetic parameters of SbnI and SbnI1-240.   Km (mM) kcat (min-1) kcat/Km (mM-1 min-1) ATP    SbnI 0.6 ± 0.1  3.9 ± 0.1  6.8 ± 1.0 SbnIa 0.2 ± 0.1 3.9 ± 0.1  17.3 ± 3.6  SbnI1-240 1.2 ± 0.3 2.1 ± 0.1  1.7 ± 0.7  ADP    SerKb 2.4 ± 0.5 12240 ± 720  5100 L-serine    SbnI 340 ± 40 14.3 ± 0.8  0.04 ± 0.01  SbnIa 150 ± 20 10.0 ± 0.6  0.07 ± 0.01 SbnI1-240 900 ± 450 2.1 ± 0.1 0.02 ± 0.01 SerKb 5.1 ± 0.5 13100 ± 300 2600 a Kinetics measured using SbnA coupled assay b Data from Makino et. al. (199)  The enzymatic activity of SbnI1-240 was also measured since the 14-amino acid C-terminal truncation does not exclude any regions in the SerK structures identified as required for substrate and product binding or catalysis. SbnI1-240 displayed decreased kinase activity compared to full-  76 length SbnI. These data correlate with the HPLC and spectral data with SbnA for SbnI1-240 having intermediate activity relative to full-length SbnI (Figure 3-7F, 3-8B). Plots of initial velocities used for determination of kinetic constants are included in the supporting information (Figure 3-10). Given that SbnI1-240 is a monomer in solution (see Section 3.3.2), we conclude that dimerization/oligomerization of SbnI is not essential for its kinase activity.    Figure 3-10 Plots of initial velocities used for determination of kinetic constants for SbnI and SbnI1-240.  Initial velocities of the L-serine kinase reactions of 0.5 µM SbnI or SbnI1-240 in the presence of varying concentrations of ATP with 100 mM L-serine (A, C, E) and in the presence of varying concentrations of L-serine with 10 mM ATP (B, D, F).    77 3.2.6 Physiological function of an ATP-dependent serine kinase activity in S. aureus The serC gene encodes the enzyme responsible for OPS synthesis from 3-phosphohydroxypyruvate in the S. aureus serine biosynthesis pathway. To our knowledge, prior to this work, SerC activity was the only identified metabolic source of OPS in S. aureus and was the assumed source of OPS for SB biosynthesis (90, 95). To test if other metabolic sources of OPS exist in S. aureus, wildtype USA300 and a serC transposon insertion mutant USA300 strain (serC) were grown in chemically defined media containing glucose with and without L-serine in a TECAN plate reader. In L-serine replete medium, both wildtype USA300 and the serC mutant grew, although the serC mutant entered stationary phase at a lower cell density and failed to reach biomass equivalent to that of the WT culture (Figure 3-11A). However, in stark contrast, in a growth medium lacking L-serine, wildtype USA300 grew while the serC mutant failed to grow altogether (Figure 3-11A). Thus, we conclude that the serC transposon mutant is a bona fide serine auxotroph; a serine auxotrophy phenotype was previously observed for an E. coli serC mutant strain (208). We also conclude that SerC activity is the sole source of OPS for serine biosynthesis; since the medium is iron-restricted, SbnI would be expressed and, if it contributed to OPS biosynthesis for use in the L-serine biosynthetic pathway, we would have not observed L-serine auxotrophy for the serC mutant (see Figure 3-1).   78   Figure 3-11 S. aureus serC mutant is a serine auxotroph but can produce SB. (A) Growth kinetics of S. aureus USA300 wildtype (black lines) and serC transposon insertion mutant (red lines) strains in chemically defined medium with glucose (CDMG) (solid lines) and CDMG without L-serine (CDMG – L-Ser) (dashed lines). (B) Agar plate disc diffusion bioassays were performed using culture supernatants prepared from S. aureus USA300 strains (wildtype and sbnI and serC transposon insertion mutants), as indicated on the x-axis, that were grown for 16 hrs in chelex-treated Tris minimal succinate (c-TMS) media. The black dots [labeled SA (DsirA)] are a measure for the presence of SA in culture supernatants and the grey dots [labeled SB (DhtsABC)] are a measure of SB in culture supernatants based on the growth radius around the disc. The disc radius (3 mm) is subtracted from the reported growth radius. Lines represent the standard deviation from the mean. *** p-value < 0.0002, **** p-value < 0.0001.  To test if SbnI could serve as a source of OPS for SB biosynthesis, the serC transposon insertion mutant was tested for its ability to make SB. A disc diffusion assay was used to detect SB in spent culture supernatant of serC and sbnI transposon insertion mutants, compared to the wildtype USA300 S. aureus strain. Briefly, spent culture supernatants were applied to sterile filter discs on iron-restricted agar seeded with either sirA or htsABC mutant S. aureus strains. The sirA mutant strain is defective for SB uptake so growth promotion around the disc in iron-restricted media is dependent on the presence of SA in the supplied culture supernatant. The htsABC mutant strain is unable to take up SA and thus growth around the disc is reliant on the presence of SB in supplied culture supernatant. Similar to wildtype S. aureus, the serC   79 transposon insertion mutant was capable of producing SB (Figure 3-11B), indicating the presence of another pathway for OPS production independent of SerC, almost assuredly via SbnI (Figure 3-1).  The sbnI mutant was impaired for SB production (Figure 3-11B), consistent with previously published results using the disc diffusion assay (98). Decreased SB production by the sbnI mutant is due to the necessity of SbnI for full expression of sbnDEFGH (98). A serC sbnI double mutant is expected to be SB deficient. However, as the sbnI mutation is pleiotropic on SB biosynthesis due to its requirement for transcription of sbn biosynthetic genes, assaying for SB production does not inform on a role in OPS production. Nonetheless, our results support the hypothesis that SbnI serves as a second metabolic route for OPS biosynthesis from L-serine in S. aureus. Production of OPS from L-serine and ATP is a previously unrecognized route for OPS synthesis in bacteria.  3.3 Discussion SbnI is a free L-serine kinase that makes OPS, which serves as a substrate for SbnA and is a precursor for SB biosynthesis. Insight into the kinase function of SbnI was gained based on homology identified by structural similarity with SerK, an enzyme in a biosynthetic pathway for cysteine in the thermophilic archaea T. kodakarensis (198). ATP is used as the phosphate donor in the reaction catalysed by SbnI, whereas SerK uses ADP. Interestingly, hyperthermophiles generally use ADP in place of ATP in key glycolytic enzymes as an adaptation to life at high temperatures, presumably because ADP is more stable than ATP (209). While SbnI is capable of phosphorylation of free L-serine, it was not capable of mediating phosphotransfer to serine residues within a His-Ser dipeptide. Bacterial Ser/Thr kinases such as S. aureus Stk1 (also named   80 PknB) (210) and cognate phosphatases function as molecular switches that have key roles in bacterial cell signaling as in eukaryotic systems (211). However, SbnI free L-serine kinase activity is functionally and enzymatically distinct from that of Ser/Thr kinases. SbnI1-240 retained partial catalytic efficiency compared to full-length SbnI. As SbnI1-240 is lacking 14 residues at the C-terminus and is monomeric, either the C-terminus or dimerization is required for full kinase function. The residues shown to be required for phosphotransfer in SerK (Glu30, Asp69) are conserved in SbnI and present in the truncated protein. Moreover, three of four residues interacting with the serine substrate in SerK are conserved (Glu30, Trp102, Thr223) and the fourth position is a conservative substitution of His225 with Phe203 in SbnI (Figure 3-2D). In contrast, interactions of SerK with ADP/AMP are poorly conserved. Of eight residues making key contacts, only three residues (Ser43, His72, Arg73) are conserved in SbnI and two of these interact with the phosphate groups. Attempts to obtain crystals with substrates or products bound to SbnI or SbnI1-240 have not been met with success and the binding mode of ATP to SbnI remains elusive. Structural analysis of SerK suggests that the conformational closure upon binding both substrates positions the catalytic glutamate (Glu30) to deprotonate L-serine to attack the terminal phosphate of ADP. With no published structure of substrate or product-free SerK, the apo-SbnI1-240 structure supports this proposed mechanism in that the unbound form is in an open conformation to expose the binding pocket. Measurement of SbnI enzyme kinetic parameters revealed that, compared to SerK, it has a relatively low selectivity for L-serine. The comparatively low kcat/Km could relate to the physiological role of SbnI in S. aureus. SerK supplies cysteine synthase with OPS to produce cysteine and may represent an ancient heterotrophic mechanism of amino acid metabolism (199). Interestingly, this cysteine synthase is   81 a distant SbnA homolog and a true OPS sulfhydrylase. Also, SerK is postulated to provide an advantage by enabling carbon from serine to be directed to glycolysis and gluconeogenesis by conversion to OPS (199). In vitro evidence suggests that S. aureus uses amino acids to support gluconeogenesis (212). However, serine is used to generate ATP and acetate rather than to facilitate gluconeogenesis as in T. kodakarensis (199, 212).  In contrast, SbnI kinase activity fulfils a distinct physiological role, providing substrate necessary for SB production. The comparatively low kcat/Km may allow SbnI to respond to a greater range of substrate concentrations, such that at high L-serine concentrations SbnI increases the rate of OPS production for SB production and possibly other metabolic processes. Note, the kinetics for SbnI were measured under non-saturating concentrations of serine and thus the derived parameters are apparent kinetic constants. Though SbnI has lower apparent catalytic efficiency at 25°C than SerK at 85°C (Table 3-1) (199), the activity of Srx, a related ParB/Srx family member, is similar to that of SbnI. The Srx ATP-dependent reduction of peroxidredoxin sulfinic acid in mammals and plants occurs with a catalytic efficiency (kcat/Km) in the range of 0.8 to 8.4 mM-1 min-1 comparable to SbnI (213, 214).  SbnI is a sufficient biological source of OPS for SB biosynthesis and contributes to the functional modularity of the sbn locus. A recognized characteristic of bacterial networks is a high degree of modularity and sparse connectivity between individual functional modules, where a functional module refers to a group of biological components that are spatially isolated or chemically specific and work together for a discrete biological function (215). The functional redundancy of enzymes encoded in the sbn locus for generation of precursor substrates, SbnG and SbnI, decreases dependence on central metabolism and contribute to the modularity of the sbn locus. SbnG is functionally redundant with the TCA cycle citrate synthase, CitZ (96).   82 Moreover, SbnG activity allows for SB biosynthesis to occur independent of TCA cycle activity (90). This functional independence is important because S. aureus elicits an iron-sparing response during infection resulting in down-regulation of the TCA cycle. We propose that the functional redundancy of SbnI with SerC for OPS production allows SB biosynthesis to occur independent of glycolysis as SerC substrate is funneled from 3-phosphoglycerate. Additionally, the serine biosynthetic pathway is regulated by negative feedback where SerA, metabolically upstream of SerC, is allosterically inhibited by L-serine and could limit the amount of SerC-derived OPS available to support SB synthesis when serine is abundant.  Thus, an alternative, SB-dedicated, OPS synthetic route via SbnI is advantageous. Together with SbnG, SbnI allows SB biosynthesis to occur autonomously from glycolysis and TCA cycle activity by generating precursor substrates dedicated to SB biosynthesis.  This observed metabolic redundancy may improve robustness and help buffer environmental perturbations S. aureus encounters during infection. Glucose is the preferred carbon source by S. aureus and available at concentrations to support growth in human blood (102). However, within staphylococcal abscesses, glucose is limiting and S. aureus likely survives through catabolism of secondary carbon sources, specifically lactate, peptides, and free amino acids (212, 216, 217). Resultant changes in central metabolism of S. aureus may not severely hinder SB biosynthesis due to the modularity of the sbn system. Despite variability in growth conditions, one constancy in S. aureus survival strategy is iron acquisition.  Serine is amongst the most abundant amino acids found in fluid obtained from S. aureus infected prosthetic joints and measured to be ~4 mM (26). SB biosynthetic genes including sbnI, but not SA biosynthetic genes, are up-regulated in this environment (26). Also, L-serine is one of the amino acids most rapidly consumed by S. aureus grown on amino acids in vitro (212).   83 Together, the combination of high L-serine in the extracellular milieu and the rapid consumption of L-serine may limit the activity of the endogenous serine biosynthetic pathway. Under these conditions OPS from serine biosynthesis is likely to be limiting to feed SB biosynthesis justifying the necessity of SbnI-generated OPS. Interestingly, SB production is restricted to more invasive coagulase-positive staphylococcal strains and the sbn locus may represent a lineage-specific innovation to support adaptation to a more invasive lifestyle. In contrast, SA biosynthetic genes are found across both coagulase-positive staphylococci and more commonly commensal coagulase-negative staphylococci. This newly determined enzymatic role of SbnI generates questions regarding how it carries out heme-dependent transcriptional regulation of the sbn locus. It remains unclear if SbnI kinase activity impacts its heme-dependent transcriptional regulation of the sbn locus. The structure of SbnI1-240 did not reveal a prototypical DNA-binding motif or heme binding site. The functional sites required for kinase activity could be structurally distinct from those involved in the transcriptional regulation and heme-binding. Full-length SbnI is a dimer yet SbnI1-240 is a monomer in the crystal structure and in solution.  The C-terminal 14 amino acids excluded from the SbnI1-240 construct are likely required for multimerization, which could be important for transcriptional regulatory function or heme-binding. Additionally, SbnI has a C-terminal extension of 34 amino acids compared to SerK and this C-terminal region is truncated in distant SbnI homologs not associated with a SB biosynthetic locus. This raises the possibility that kinase activity may be the core function of the non-SB-associated SbnI homologs. The C-terminal extension in SbnI could be a structural adaptation to facilitate multiple biological functions in SB precursor biosynthesis and sbn locus gene regulation. Heme-dependent SbnI regulatory activity may further enhance network robustness as a negative feedback loop from an iron acquisition   84 perspective. Characterization of SbnI heme-binding and effect on kinase activity remains the subject of future work.  OPS production by the kinase activity of SbnI represents a new biosynthetic path for production of this metabolite in bacteria. SbnI may serve as a metabolic adaptation to facilitate SB production when growing on non-preferred carbon sources. Furthermore, it demonstrates the metabolic flexibility S. aureus possesses to allow employment of iron uptake strategies in changing host environments.    85 Chapter 4: SbnI has two distinct roles in staphyloferrin B biosynthesis in S. aureus 4.1 Introduction Structural study of a C-terminal truncated construct of S. aureus SbnI (SbnI1-240) revealed homology to a free serine kinase and biochemical assays showed SbnI catalyzes phosphotransfer from ATP to serine to generate OPS (Chapter 3). OPS is a substrate of SbnA which, together with SbnB, produces L-Dap and a-KG, precursors for SB biosynthesis (92, 95). SbnI-generated OPS was demonstrated to be sufficient to support SB biosynthesis in vivo (Chapter 3). SbnI is characterized as a positive transcriptional regulator of the sbn locus that is required for in vivo SB synthesis (98). However, SbnI can also bind heme as a negative regulatory molecule to limit SB production, presumably by preventing SbnI interaction with DNA (98).  Herein we present insight into the dual function of SbnI in SB biosynthesis by studying the effect of heme-binding on L-serine kinase activity.  SbnI1-240 has decreased kinase activity and is monomeric, while full-length SbnI is dimeric. As crystals of full-length SbnI were not obtained, we crystallized the full-length SbnI homolog from Staphylococcus pseudintermedius (SpSbnI) with and without ADP, a product of the kinase reaction. SpSbnI formed a dimer through C-terminal domain swapping and a dimer of dimers through the formation of intermolecular disulfides. A co-crystal structure with ADP confirms the location of the L-serine kinase active site. Using site-directed mutagenesis, spectroscopic analysis, and molecular docking, we propose a model of SbnI heme-binding. Also, we demonstrated that SbnI can obtain heme from IsdI, consistent with a role as a heme-sensing protein, and provide a mechanism for how successful   86 Isd-mediated heme uptake could be sensed by SbnI, leading to decreased SB production as a way for S. aureus to control iron source preference.  4.2  Results 4.2.1 Heme binding by SbnI does not greatly hinder L-serine kinase activity In order to determine if heme binding to SbnI impacted serine kinase activity, steady state enzyme kinetics of heme-bound SbnI were measured using a pyruvate kinase/lactate dehydrogenase (PK/LDH) assay for detecting ADP (Figure 4-1). Heme-bound SbnI was purified by gel filtration to avoid confounding effects of excess heme in solution. The apparent steady-state kinetic parameters of SbnI:heme with ATP and L-serine are summarized in Table 4-1. Notably, the specificity constant (kcat/Km) measured for apo-SbnI differed by less than a factor of two from the value for heme-bound SbnI, which suggests that the presence of heme does not drastically alter L-serine kinase activity.    Figure 4-1 Plots of initial velocities versus substrate concentration used for determination of kinetic constants for SbnI bound to equimolar heme.  Initial velocities of the L-serine kinase reactions of 0.5 µM SbnI bound to equimolar heme in the presence of varying concentrations of (A) ATP with 100 mM L-serine present and in the presence of varying concentrations of (B) L-serine with 10 mM ATP present.   87 Table 4-1 Apparent steady-state kinetic parameters of the L-serine kinase reaction by heme-bound SbnI.   Km (mM) kcat (min-1) kcat/Km (mM-1 min-1) ATP    SbnIa 0.6 ± 0.1  3.9 ± 0.1  6.8 ± 1.0 SbnI+heme 0.75 ± 0.15 3.2 ± 0.2  4.3 ± 0.9  L-serine    SbnIa 340 ± 40 14.3 ± 0.8  0.04 ± 0.01  SbnI+heme 940 ± 30 28 ± 6 0.03 ± 0.01 a kinetic values obtained from Table 3-1 in Chapter 3.  4.2.2  Structure of SpSbnI and SpSbnI bound to ATP S. pseudintermedius is closely related to S. aureus and is a pathogen of companion animals such as dogs. SpSbnI shares 60% amino acid sequence identity with S. aureus SbnI. SpSbnI was crystallized and the structure was solved to 2.1 Å resolution using the S. aureus SbnI1-240 structure (PDB ID: 5UJE) for phasing by molecular replacement. Data collection and refinement statistics are summarized in Table 2-4. The coordinates and observed structure factor amplitudes have been deposited in the PDB under the accession code 5UJD. The protein crystallized with a dimer in the asymmetric unit. The structure of SpSbnI is very similar to the C-terminal truncated structure of S. aureus, SbnI1-240. SpSbnI protomers overlay with SbnI1-240 with a RMSD of 1.4 Å across 240 Cα atoms for protomer A and 1.9 Å across 240 Cα atoms for protomer B (181). The SpSbnI protomer consists of three domains. Domain I consists of residues M1-Q83 and I205-A240 and domain II contains Y84-N204 (Figure 4-2A). These domains are consistent with SbnI1-240 domain I and II. Domain I contains a conserved core a ParB/Srx fold that is consistent with the serine kinase active site architecture of SbnI1-240. Domain II has a similar overall fold as in SbnI1-240. Domain III encompasses the C-terminal 14 amino acids absent from the SbnI1-240 structure, M241-E254, and forms the dimer interface (Figure 4-2A). This interface involves C-  88 terminal domain swapping between the two protomers (Figure 4-2B). Domain III assembles into two mixed 3-strand β-sheets with one β-strand originating from the other protomer. The buried surface area at the SpSbnI dimer interface is 1650 Å2, which accounts for approximately 12% of the total solvent-accessible area of the structure (PDBePISA) (218). The interface involves 28 hydrogen bonds within residues 204-213 and 240-254.    Figure 4-2 Overall structure of SpSbnI.  (A) SpSbnI monomer colored by domain: N-terminal domain I (yellow), domain II (green), and C-terminanl dimerization domain III (purple). (B) SpSbnI crystallographic dimer formed by C-terminal domain swapping. (C) SpSbnI dimer of dimers linked by two pairs of intermolecular disulfides.   Despite inclusion of 2 mM GSH in the crystallization buffer to keep cysteine residues reduced, in the crystal structure domain II contains two pairs of intermolecular disulfide bonds each between Cys166 and Cys168 from a symmetry related molecule are observed to yield a dimer of dimers (Figure 4-2C, 4-3A). This second dimer interface has 200 Å2 of buried surface area. The residues involved in H-bonding are poorly conserved amongst SbnI homologs and include Gln126, Cys166, Cys168, and Ser171. Domain II in the SpSbnI structure also contains an intramolecular disulfide bond between residues Cys99 and Cys156 within each protomer (Figure 4-3B). Superimposition of SbnI1-240 and SpSbnI crystal structures suggests that S. aureus   89 can also oligomerize through formation of intermolecular disulfide bonds (Figure 4-3C). SbnI1-240 has a single cysteine, Cys168, located on the equivalent loop as the SpSbnI cysteines, Cys166 and Cys168. A superimposition of the SbnI1-240 and SpSbnI structures reveals that where SpSbnI forms an intramolecular bond between Cys99 and Cys156, SbnI1-240 has instead two adjacent cysteines, Cys155 and Cys156 (Figure 4-3D). Formation of a vicinal disulfide between sequence adjacent cysteines is very rare (219). More commonly, the cysteines are found in the reduced form and if they do form disulfides, the two bonds are almost always to different partners (219, 220). Whether SbnI Cys155 and Cys156 have a biological role requires further study.         90  Figure 4-3 SpSbnI disulfide bonds and conservation with S. aureus SbnI.  (A) Intermolecular and (B) intramolecular disulfide bonds of SpSbnI. Fo–Fc electron density omit maps are contoured at 3s.  SpSbnI residues are drawn as sticks with separate protomers colored teal or pink. Oxygen, nitrogen, and sulfur atoms colored red, blue, and yellow, respectively. (C) Superimposition of SbnI1-240 (blue, residue labels in bold font) and SpSbnI (teal and pink) homologous strands at the dimer of dimers interface seen in the SpSbnI crystal structure. (D) A superimposition of SbnI1-240 (blue, residue labels in bold font) and SpSbnI (teal) where SpSbnI forms an intramolecular bond between Cys99 and Cys156. Cysteine residues are represented as sticks with carbon colored either teal, blue, or light pink and sulfur atoms colored yellow.  A co-crystal structure of SpSbnI with ADP was solved to 1.9 Å resolution using the SpSbnI structure (PDB ID: 5UJD) for phasing. Data collection and refinement statistics are summarized in Table 2-4. As observed for apo-SpSbnI, the co-crystal structure was in space group P21212 and found in the dimer of dimer conformation. However, it differed from the apo-SpSbnI crystal   91 structure in that the molecules linked by disulfide bonds were within the asymmetric unit, and protomers related by crystallographic symmetry formed the dimer interface by C-terminal domain swapping (Figure 4-4AB). The protomers related by crystallographic symmetry are opposite of that observed in the apo-structure. The cell dimensions for ADP bound SpSbnI were a = 101.6 Å, b = 73.1 Å, c = 88.4 Å and differed from the cell dimensions observed for apo-SpSbnI, which were a = 78.8 Å, b = 111.4 Å, c = 67.7 Å. Though the crystallization solution contained both products, ADP and OPS, only clear electron density was seen for the diphosphate of ADP and a modeled magnesium ion for protomer A (Figure 4-4C) and the diphosphate, sugar, and magnesium ion for protomer B (Figure 4-4D). SpSbnI protomer A modelled with ADP and a magnesium ion based on electron density present in the crystal structure was superimposed with the SerK ternary complex with AMP and OPS (PDB ID: 5X0E). The coordinates of OPS from the SerK-bound structure were merged with SpSbnI-ADP coordinates to generate a model of with both products, ADP and OPS, bound to SbnI (Figure 4-5). The C-terminal domain partially encloses the L-serine kinase active site in SerK and this feature may be important for catalysis and partially explain the lower catalytic efficiency of SbnI1-240 (Chapter 3).        92  Figure 4-4 Structure of SpSbnI bound to ADP.  (A) SpSbnI crystallographic dimer formed by intermolecular disulfide bonds. Protomer A is colored light blue and protomer B is yellow (B) SpSbnI dimer of dimers formed by C-terminal domain swapping. (C, D) Fo–Fc electron density omit maps (contoured to 3s) of active sites for protomer A and B. SpSbnI residues are drawn as sticks with separate protomers colored light blue or yellow. ADP carbon is colored grey, water molecules are red and magnesium ions are green. Oxygen, nitrogen, and phosphorus atoms colored red, blue, and orange, respectively.   Figure 4-5 Active site of SpSbnI and ADP co-crystal structure with OPS modelled. Selected SpSbnI active site and nearby residues, ADP, and Mg2+ of protomer A. OPS was modeled based on superimposition with SerK (yellow, PDB ID: 5X0E). In light blue is protomer A and dark blue is protomer A related by crystallographic symmetry and forms a dimer by C-terminal domain swapping. Mg2+ is drawn as a green sphere, O, N, S, and P atoms colored red, blue, yellow, and orange, respectively. ADP and OPS carbons are colored grey.   93 4.2.3 Conservation and molecular surface electrostatics analyses of the SpSbnI structure ConSurf was used to identify conserved regions at the SpSbnI molecular surface (Figure 4-6A). The groove that separates domain I from domain III is highly conserved and partially formed from the adjacent protomer. As SpSbnI is a homodimer, there are two conserved grooves per dimer located on opposite faces. Conserved, solvent exposed residues in this groove include H19, E20, E23, R26, D58-R62, W89, G199-R202, G207-N211, R241-Y243, E246, Y249, and E252. These residues are highlighted in a multiple sequence alignment of staphylococcal SbnI homologs (Figure 4-7). Importantly, these include residues identified in Chapter 3 as being involved with substrate binding and catalysis in the serine kinase active site. However, the serine kinase active site only accounts for a small portion of the conserved surface area suggesting the groove may support another function. Highly conserved residues in domain III form the dimerization interface (Figure 4-6A). Absolutely conserved sequence motifs amongst staphylococcal SbnI homologs that form the C-terminal dimer interface include FNIXGRCLNL (residues 204-213) and RCYXEK(V/I)YL(V/I)E (residues 242-252); residue numbering from SpSbnI (Figure 4-7).  The APBS plugin in PyMOL were used to generate an electrostatic surface map for the SpSbnI structure (Figure 4-6B). The surface of SpSbnI has two regions of notable charged surface. The first is the outward facing end of domain II has a patch of negative charge. Conserved, negatively charged amino acids in this region include E105, E113, E117, and E129. The second surface is the between domain I and domain III and forms a region of positive charge, the same conserved area identified by ConSurf. Conserved, positively charged residues in this area include H19, R26, R40, H41, H61, R62, R202, R208, and R242.    94  Figure 4-6 Amino acid conservation and surface electrostatics of SpSbnI. (A) Conservation pattern of the SpSbnI protomer and dimer generated using ConSurf. The color-coding bar shows the coloring scheme; conserved amino acids are colored bordeaux, residues of average conservation are white, and variable amino acids are turquoise. (B) Electrostatic potential mapped on the SpSbnI protomer and dimer molecular surface; a blue color indicates regions of positive potential (> +5 kT/e) whereas red represents negative potential (< -5kT/e) values.     95  Figure 4-7 Multiple sequence alignment of staphylococcal SbnI homologs.  The sequence alignment was generated using T-Coffee Expresso. The dark red boxes represent regions identified in SpSbnI to be important for dimerization by C-terminal domain swapping, and the magenta box highlights the cysteine residues that form intermolecular disulfides for the dimer of dimers in the SpSbnI structure. The lime green boxes indicate regions identified in the conserved groove between domain I and domain III. The colored triangles represent the proposed heme-iron coordinating ligands (black), residues predicted to interact with heme (red), proposed catalytic residue for serine kinase activity (grey), residues predicted to recognise adenosine of ADP, or ATP (green), the residues that interact with the phosphate groups of the ADP or OPS in the model (blue), and the predicted residues that interact with the Ser moiety of OPS (magenta). All SbnI homologs used in the alignment are from species of the Staphylococcus genus (NCBI reference sequence): Sau, S. aureus (ABD21649); Sar, S. argentius (WP_001015552); Ssc, S. schweitzeri (WP_047560844); Seq, S. equorum (WP_069816870); Sag, S. agnetis (WP_060552121); Shy, S. hyicus (WP_039643510); Ssch, S. schleiferi (WP_050329672); Sarl, S. arlettae (WP_002510733); Sde, S. delphini (WP_096592770); Sin, S. intermedius (WP_086428370); Sps, S. pseudintermedius (WP_014614629); Slu, S. lutrae (WP_085237657).    96 4.2.4 Spectroscopic characterization of heme coordination structure of S. aureus SbnI and SpSbnI Heme binding was evaluated based on the UV-visible spectra of SbnI or SpSbnI incubated with equimolar heme. When SbnI was incubated with excess heme and purified by gel filtration, quantification of heme and total protein using the pyridine hemochrome (187) and BCA assays found SbnI monomer binds heme at a ratio of approximately 1:1. The ferric, ferrous and ferrous CO-bound spectra of heme-bound SbnI and SpSbnI were characterized using UV-visible spectroscopy to examine the heme-iron electronic state and coordination structure. SbnI-Fe(III) heme spectrum was characteristic of low-spin, 6-coordinate ferric heme coordinated by a thiolate with a Soret peak at 423 nm, a shoulder at 360 nm, and alpha and beta bands at 573 nm and 541 nm, respectively (Figure 4-8A). An intense, well-resolved shoulder at 355-365 nm is a hallmark for low-spin Fe(III) Cys (thiolate) ligation (221, 222). A Soret peak at 415-430 nm, and alpha and beta bands between 535-575 nm are characteristic of a low-spin, 6-coordinate electronic structure (222). Broad a/b bands are characteristic of His trans the cysteine (thiolate), while resolved a/b bands are more commonly observed for H2O or N-terminal amino group trans the cysteine (thiolate) (223). The SpSbnI Fe(III) complex has comparatively less defined spectral features. A prominent shoulder is 365 nm is present with a less intense Soret peak at 420 nm (Figure 4-8D). The SpSbnI heme complex had poorly defined bands in the a/b region with one broad band at 542 nm. The shoulder at 365 nm suggests thiolate coordination of the heme-iron, as observed in S. aureus SbnI-Fe(III) heme.     97  Figure 4-8 UV-visible spectra of heme-bound SbnI and heme-bound SpSbnI in the oxidized, reduced, and CO bound forms. Spectrum of 5 µM (A) S. aureus SbnI or (D) SpSbnI mixed with equimolar heme. Spectrum of 5 µM (B) SbnI or (E) SpSbnI incubated with equimolar heme immediately after reduction with dithionite. Spectrum of 5 µM (C) SbnI or (F) SpSbnI with equimolar heme after exposure to CO and subsequent reduction with dithionite. All reactions were carried out in 50 mM HEPES (pH 7.4), 100 mM NaCl, and 5% (v/v) glycerol.  Upon reducing the SbnI-Fe(III) heme complex with sodium hydrosulphite (dithionite) to Fe(II), the Soret and a/b bands shift to 425, 559, and 532 nm, respectively (Figure 4-8B). Similarly, reduction of the SpSbnI:heme complex resulted in formation of a sharp Soret peak at 425 nm and the appearance of defined a/b bands at 560 nm and 530 nm, respectively (Figure 4-8E). Such changes in the heme spectroscopic signature are reported for other proteins that bind Fe(III) heme 6-coordinate with thiolate ligation that undergo a redox-mediated ligand switch   98 upon reduction (223). These include the two bacterial gas sensors, EcDos of E. coli and CooA of Rhodospirullum rubrum (224, 225). Increased electron density via reduction of the heme iron is proposed to make 6-coordinate heme thiolates intrinsically disordered where the heme center can compensate by weakening the Fe-S interaction. The labile cysteine (thiolate) ligand is susceptible to replacement by a nearby residue if there is flexibility in the heme pocket (223, 226). The propensity of the cysteine to undergo ligand-switch upon reduction results in preservation of the low-spin 6-coordinate state when the heme iron is reduced to the ferrous form.  The spectral features that accompany this redox-mediated ligand switch are consistent with those observed for SbnI. The Soret peak for Fe(II) heme centers that are coordinated by two neutral donors is usually in the 420-430 nm range, accompanied by intense, well resolved, asymmetric visible region peaks (a > b) with the a band between 550-560 nm (223, 227). By contrast, the 5-coordinate, high-spin Fe(II) state with a sole cysteine ligand exhibits distinct spectral features with a Soret peak at 410 nm and a band in the visible region at 540 nm (223). Overall, the results are suggestive that upon heme iron reduction, SbnI loses cysteine ligation of the heme iron and it is replaced by a nearby neutral donor, like a histidine or methionine, to maintain the low-spin, 6-coordinate state of the Fe(II) heme.  Incubation of heme proteins with CO is commonly used to probe heme centers. When heme bound SbnI was exposed to CO and subsequently reduced with dithionite, a shift in the Soret peak to 421 nm, and alpha and beta bands to 570 nm and 540 nm, respectively, indicate a change occurred in the electronic environment of the heme and that CO was binding directly to the heme iron (Figure 4-8C). SpSbnI displayed similar spectral changes with a shift in the Soret peak to 422 nm and alpha and beta bands to 570 nm and 540 nm, respectively (Figure 4-8F). When a   99 cysteine ligand is replaced either by a redox-mediated ligand switch, or by CO directly, the CO-adducts typically display a Soret peak at 420 nm and approximately equivalent alpha and beta peaks around 560 and 540 nm (223). The spectra of SbnI and SpSbnI in the presence of CO displayed features that are characteristic of 6-coordinate, low-spin heme, similar to CO adducts of hemoglobin, myoglobin, and other proteins that maintain an axial His-Fe(II) ligation (228–230). Reaction of CO with the Fe(II) heme-thiolate of cytochrome P450 results in a Soret peak at 450 nm that is attributed to CO binding trans the cysteine (thiolate) (231, 232). Reaction of heme bound SbnI with CO did not result in a shift of the Soret peak to 450 nm suggesting in a loss of the cysteine (thiolate) ligand in SbnI in the CO-bound state, and suggesting that the putative His ligand is maintained.  4.2.5 SbnI heme affinity and heme off-rate Aggregation of recombinant SpSbnI prevented determination of heme binding affinity and binding kinetics. Full-length, recombinant S. aureus SbnI was sufficiently soluble for these studies. Intrinsic tryptophan fluorescence of SbnI was used to measure heme binding affinity as heme binding causes fluorescence quenching. Concentration-dependent quenching of fluorescence was observed for SbnI (250 nM) and a KD value of 125 ± 105 nM was determined (Figure 4-9AB). Attempts to measure heme-affinity by fluorescence quenching using lower protein concentration (50 nM) did not yield reproducible results likely due to instrument sensitivity and potentially the SbnI monomer-dimer equilibrium. Recently published heme-binding affinities for S. aureus IsdI and IsdG are 12.9 nM and 1.4 nM, respectively, and are 2-3 orders of magnitude lower than previously reported (233). These higher heme affinities of IsdI and IsdG were measured using intrinsic fluorescence quenching of 60-80 nM protein samples by   100 heme (233). Since KD values well below the protein concentration cannot be accurately measured by fluorescence quenching, our value for the SbnI heme affinity using 250 nM protein concentration likely represents and upper estimate of heme affinity rather than the actual KD.  The SbnI heme off-rate was determined based on the rate of heme transfer to the non-physiological heme acceptor apomyoglobin. Heme binding by apomyoglobin was followed by stopped-flow spectroscopy at 408 nm, the Soret maximum for holomyoglobin. The rate of heme transfer to apomyoglobin was independent of apomyoglobin concentration (data not shown). The transfer rate data was best fit by a triple exponential curve, as judged by the residual plot. The rate of heme dissociation from SbnI was measured to have an average k1 of 0.044 ± 0.003 s-1, k2 of 0.015 ± 0.001 s-1, and k3 of 0.003 ± 0.001 s-1. Each accounted for 31, 34, and 35% of the absorption change, respectively (Figure 4-9C).   Figure 4-9 SbnI heme affinity and heme off-rate.  (A) Representative emission spectra for 290 nm excitation for fluorescence-detected titration of heme into 250 nM of SbnI in 50 mM HEPES pH 7.4, 100 mM NaCl, 5% (v/v) glycerol. (B) Plotted is the area under the emission spectra curve against heme concentration. The vertical error bars represent the standard deviation of three technical replicates. (C) Measurement of heme release from SbnI and binding by apomyoglobin at a ratio of 1:10, SbnI:apomyoglobin. A plot of the change in absorbance at 408 nm, where holomyoglobin absorbs maximally, versus time was fit by a three-phase exponential function. A graph of the residuals is inset and displays a relatively random distribution.    101 4.2.6 Oligomeric state of heme-bound SbnI Apo-SbnI was previously shown to be dimeric and the SbnI1-240 C-terminal truncated construct to be monomeric under reducing conditions (Chapter 3). Here we investigated the effect of heme-binding on SbnI oligomerization using DLS. The calculated molecular weight of monomeric SbnI is 30 kDa, whereas apo-SbnI was measured to be 61 ± 6 kDa with an average of 24 ± 7% polydispersity using DLS (Chapter 3) and is presumed to form a dimer in solution. The average molecular weight of SbnI reconstituted with heme was determined as 43 ± 10 kDa with 29 ± 5% polydispersity, suggesting that heme may shift the monomer-dimer equilibrium toward its monomeric form (Figure 4-10). Alternatively, the heme-bound SbnI dimer may appear smaller than the apo-SbnI due to a structural change reducing the molecular radius of the dimer as all molecular weights were calculated using the theoretical hydrodynamic radius for a spherical protein. An estimate for SpSbnI oligomerization was not possible due to the propensity of the protein to aggregate and inability to obtain a consistent qualitative analysis by DLS.   Figure 4-10 Representative DLS results for analysis of heme-bound SbnI oligomerization state.  (A) Percent mass-weighted size distribution of heme-bound SbnI with calculated molecular weight (Mw) and polydispersity (Pd). (B) The correlation function data measured.    102 4.2.7 Site-directed mutagenesis to probe SbnI heme-binding mode Spectra of full-length dimeric SbnI bound to heme has a Soret peak at 423 nm, a shoulder at 360 nm, and well-resolved alpha and beta bands at 573 nm and 541 nm, respectively (Figure 4-11A). Comparatively, the C-terminal truncated monomeric construct, SbnI1-240, has altered heme binding as determined by a decrease in the Soret peak and alpha and beta bands compared to full-length SbnI spectra (Figure 4-11B), suggesting there is a change in the amino acid environment around the heme. The C-terminal 14 amino acids excluded from the SbnI1-240 construct may be important for interaction with heme or dimerization of the protein may be necessary for heme binding, or both. Within these 14 amino acids, a conserved cysteine was identified as a putative heme-iron coordinating residue. A site-directed mutant was made by replacing Cys244 with alanine (SbnI C244A). This variant was found to have altered heme binding based on the decrease in Soret peak and diminished alpha and beta bands in electronic spectra measured after incubation with heme (Figure 4-11C). Additionally, SbnI uniquely contains an N-terminal HXHXH motif (amino acids 3-7). A site-directed mutant of His3 to an alanine (SbnI H3A) was made. This variant was also found to have altered heme binding based on the decrease in Soret peak and diminished alpha and beta bands in electronic spectra measured after incubation with heme (Figure 4-11D). A SbnI double mutant of His3 and Cys244 both mutated to alanines (SbnI H3A/C244A) was generated and found to have additive effect on decreasing the Soret peak but did not completely abolish heme-binding (Figure 4-11E).  The crystal structures of SbnI1-240 and SpSbnI were examined for alternative surface-exposed cysteines that could be involved in heme binding. S. aureus SbnI Cys168 is predicted to participate in formation of the dimer of dimers by intermolecular disulfide bond formation (Figure 4-3C). A site-directed mutant of this residue, SbnI C168A, bound heme with UV-visible   103 spectra resembling wildtype SbnI (Figure 4-11G). Other surface exposed cysteines in S. aureus SbnI included Cys155 and Cys156. SpSbnI has a single Cys in this region, Cys156, but it was found to form an intramolecular disulfide bond with Cys99 in the structure (Figure 4-3D). Mutagenesis of SbnI to generate SbnI C155A had no change in the heme-bound spectra compared to wild-type SbnI indicating this residue is not involved in heme-binding (Figure 4-11F). S. aureus has two other Cys residues, Cys124 and Cys 209, that were not tested for a role in heme-binding. Cys124 is located in domain II and Cys204 is buried in the dimer interface formed by C-terminal domain swapping. Time-dependent SpSbnI heme-binding was also measured using UV-visible spectroscopy and it was found to have spectral features similar to SbnI but less hyperchromatic. SpSbnI has a Soret peak at 420 nm, a shoulder at 367 nm and broad alpha and beta bands at 570 nm and 542 nm, respectively (Figure 4-11H).   Figure 4-11 UV-visible spectroscopic analysis of heme binding by SbnI variants. (A-H) 5 µM of wild-type SbnI and the indicated variants were mixed with equimolar heme and spectra was recorded immediately (dotted line), at 10 minutes (grey), and after 1 hour (black).    104 4.2.8 Model of heme-bound SbnI Despite exhaustive attempts to obtain a crystal structure of heme-bound to SbnI, we were unable get a crystal structure of either S. aureus SbnI, SbnI1-240, or SpSbnI in complex with heme. SpSbnI co-crystallized with heme or Co(III)-protoporphyrin IX generated light red or pink diffraction-quality crystals, respectively. However, no iron or cobalt anomalous signal was detected nor was density for a porphyrin ring observed in electron density maps. Therefore, Autodock Vina was used to dock heme to the SpSbnI dimer to provide insight into initial heme binding by apo-SpSbnI. The area encompassing one SpSbnI protomer was selected for the search space and the top 19 solutions were recorded. The heme molecules were placed in a conserved groove formed between domain I of one protomer and domain III swapped from the adjacent protomer for 12 of these solutions and in the other 7, heme was placed in the kinase active site within domain I (Figure 4-12A). The solutions had calculated binding energies between -9.1 and -7.6 kcal/mol. The top four solutions had the heme placed within 3.9 Å of the top solution. The distance calculation is relative to the top solution where each atom in the top conformation is matched with the closest atom of the same element type in the other conformation, taking any symmetry of the heme molecule into account. The heme propionate groups are modelled to be within hydrogen bonding distance (3.0-3.2 Å) of amino acids from both protomers in the C-terminal domain swapped dimer. These include the side chains of Arg202, Asn204, and Arg208 of one protomer, and Thr245 and Glu246 of the adjacent protomer (Figure 4-12B), all of which are highly conserved amongst SbnI homologs from staphylococci. The only putative heme-iron ligand in the vicinity of the docked heme was Cys243, with the thiolate located 6.6 Å from the iron.    105 Mutagenesis of S. aureus SbnI His3 revealed it as a putative heme iron coordinating residue; however, is not conserved in SpSbnI. To investigate how this residue, and the surface exposed amino acid environment of SbnI, would impact heme placement, a model of S. aureus SbnI dimer was generated. The crystal structure of C-terminal truncated S. aureus SbnI (SbnI1-240) was deemed not suitable for the docking experiment as it had impaired heme-binding, was monomeric, and lacks the C-terminal Cys244 residue, a predicted heme-iron coordinating ligand. A homology model of dimeric SbnI was generated using SWISS-MODEL with the dimeric SpSbnI crystal structure as a template. A superimposition using PDBeFOLD of the SbnI model and SbnI1-240 crystal structure revealed a RMSD of 1.90 Å across 223 Cα atoms. Analysis of the dimeric S. aureus SbnI model using Molprobity found that 96.8% of modelled residues are in the favored regions of the Ramachandran plot, while 0.4% of residues are outliers (234). 95.4% of modelled residues are favored rotamers; however, 1.2% of bond angles are considered to have poor geometry (234).   Autodock Vina was used to dock heme to the SbnI dimeric model using an area encompassing one SbnI protomer for the search space. The resulting top 20 solutions were all within the conserved groove formed between domain I of one protomer and domain III swapped from the adjacent protomer SbnI dimer interface with calculated binding energies between -7.8 and -6.5 kcal/mol (Figure 4-12C). The amino acid environment surrounding the heme was similar to the SpSbnI top solution with the heme propionates within H-bonding distance of Arg202, Asn204, and Arg208. The Cys244 thiol was 7.8 Å and the epsilon-nitrogen of the imidazole ring of His3 was 17.1 Å from the modeled heme Fe (Figure 4-12D). Though the Cys243/Cys244 thiols in SpSbnI and SbnI are pointing away from the heme, upon heme binding   106 a modest conformational change could position the cysteine within coordinating distance of the heme iron.    Figure 4-12 Models of heme binding by SpSbnI and a full-length model of S. aureus SbnI dimer.  (A) 19 docking solutions were generated using as a search area that encompassed an SpSbnI protomer. (B) The top solution is displayed with amino acids predicted to form hydrogen bonds (3.0-3.2 Å), represented by dashed lines. Residues predicted to be involved in heme binding are drawn as sticks and labelled. Cys243 is positioned nearby the heme model with 6.6 Å between the heme iron and cysteine thiol. (C) Heme docking with the SbnI dimeric model placed the top 20 solutions all within the groove between domain I of one protomer and domain III of the adjacent protomer. (D) The amino acid environment surrounding the top solution places the heme within H-bonding distance of Arg202, Asn204, and Arg208. The Cys244 thiol was 7.8 Å and His3 was 15.9 Å from the modeled heme Fe. Carbon atoms are colored teal, maroon, light blue, or green and oxygen, nitrogen, and sulfur atoms are colored red, blue, and yellow, respectively. Heme carbon, oxygen, nitrogen, and iron atoms are colored grey, red, blue, and dark red, respectively.   107 The heme-docking models may represent the initial mode of heme binding into apo-SbnI. Additional conformational changes are likely necessary to accommodate heme binding by His3 and may require the unwinding of the N-terminal helix. Moreover, evidence of a shift in the monomer-dimer equilibrium upon heme binding by SbnI, as measured by DLS, also suggests structural changes occur upon heme-binding. Importantly, the top heme docking solutions positon the heme at a site that is structurally distinct from the kinase active site in the ParB/Srx core domain (Figure 4-13), consistent with our observations that kinase activity is largely independent of heme binding.    Figure 4-13 Superimposition of SpSbnI models bound to heme, ADP, and OPS.  (A) Superimposition of SpSbnI model with heme and SpSbnI modelled with ADP and OPS. Surface representation reveals that all three molecules are predicted to bind at adjacent, but structurally distinct sites. (B) Selected active site and nearby residues drawn as sticks and labelled.  Heme, ADP, and OPS carbon are colored grey and Mg2+ is represented by a green sphere. Oxygen, nitrogen, and sulfur atoms are colored red, blue, and yellow, respectively. Oxygen, nitrogen, sulfur, phosphorus, and heme iron atoms are colored red, blue, yellow, orange, and dark red, respectively.    108 4.2.9 IsdI can catalytically transfer heme to SbnI The Isd system is the primary means of heme uptake by S. aureus (61, 235). Once transported into the cell, heme is degraded by either IsdI or IsdG, two paralogous enzymes. As S. aureus demonstrates heme iron preference for heme in vitro (51) and little, if any, free heme should exist in the cytoplasm, we hypothesized that IsdI could serve as a heme source for SbnI. This hypothesis was tested by measuring the rate of heme transfer from IsdI to SbnI and compared to the heme off-rate from IsdI.  The rate of heme transfer from holo-IsdI to apo-SbnI was measured using stopped-flow spectroscopy. The heme transfer rates were determined by following the absorbance change at 426 nm, the wavelength of maximal difference in the visible region between holo-IsdI and holo-SbnI. Data were best fit by a triple exponential curve, as judged by the randomness of the residual plots. Heme transfer kinetic constants are k1 of 1.55 ± 0.08 s-1, k2 of 0.16 ± 0.01 s-1, and k3 of 0.05 ± 0.01 s-1, which account for 14%, 78%, and 8% of the absorption change, respectively (Figure 4-14A). IsdI and SbnI are both dimers that bind one heme in each protomer, so the overall rate of heme transfer reflected the rate of transfer of two heme molecules from one IsdI dimer to one or two SbnI dimers since SbnI was in excess. The observation of absorption changes with varying rates is consistent with the presence of multiple heme bound species; however, assignment of these rates to specific steps in heme transfer is not yet possible.  To determine whether the observed transfer from IsdI to SbnI was the result of an active transfer or heme release from IsdI followed by binding by SbnI, the heme transfer rates were compared to the rate of heme release from IsdI to apomyoglobin to measure the off-rate. The heme transfer rates were independent of the concentration of apomyoglobin. The transfer data were best fit by a double exponential curve, as judged by the randomness of the residual plot.   109 The rate constants of heme dissociation from IsdI were: kfast of 0.13 ± 0.01 s-1 and kslow of 0.04 ± 0.01 s-1, which account for 93% and 7% of the absorption change, respectively (Figure 4-14B). Importantly, the rate of IsdI heme transfer to SbnI was ~10 fold greater than the rate of passive heme dissociation from holo-IsdI indicative of the formation of an IsdI-SbnI complex to facilitate heme transfer.    Figure 4-14 Stopped-flow kinetic data of IsdI heme transfer to SbnI and measurement of the IsdI heme off-rate. (A) IsdI heme transfer to SbnI was measured. A plot of 426 nm, the wavelength of maximal absorbance change based on a difference absorption spectrum between holo-IsdI and holo-SbnI, vs time was fit by a triple exponential curve. (B) Heme release from IsdI was similarly measured using apomyoglobin. The change in absorbance was fit to a double exponential curve. The dashed line represents the represents the curve fit by the data, and the gray band represents the standard error of the average of five reactions. A graph of the residuals is inset and displays a relatively random distribution.  4.2.10 SbnI1-240, SbnI H3A, SbnI C244A, and SbnI H3A/C244A are deficient in accepting heme transferred from IsdI compared to wildtype SbnI IsdI heme transfer to apo-SbnI was measured using UV-visible spectroscopy. Apo-SbnI was added to holo-IsdI and within the time of mixing and reading the spectra, the Soret peak shifted   110 from 412 nm to 417 nm, and a shoulder around 360 nm, and alpha and beta bands at 573 and 541 nm, respectively, appeared (Figure 4-15A). Further incubation of the mixture resulted in the Soret peak shifting to 421 nm. The resulting spectra are characteristic of holo-SbnI (Figure 4-11A). The rapid spectral shift from that of holo-IsdI to that of holo-SbnI indicates that heme is transferred from IsdI to SbnI. Little to no spectral shift was observed upon addition of SbnI1-240, SbnI C244A, and SbnI H3A to IsdI suggesting these alterations impaired heme transfer (Figure 4-15BCDG). IsdI heme transfer to SbnI C155A and SbnI C168A resemble wildtype transfer (Figure 4-15EF), suggesting that His3 and Cys244 mutants have, in addition to altered heme bound spectra, a defect in accepting heme from IsdI.    Figure 4-15 IsdI heme transfer to SbnI and SbnI variants.  (A-G) The spectrum of 5 µM IsdI with equimolar heme was recorded (dashed line). 5 µM of SbnI or SbnI variant was added to the cuvette and spectra was recorded immediately (grey line) and after 1 hour (black line).    111  To assay heme transfer from IsdI to SbnI variants, a pull-down assay was employed to quantify the amount of heme that is transferred from IsdI to SbnI variants at equilibrium. The A280/A415 ratio of IsdI eluted after incubated with buffer was treated as a no transfer control. IsdI transferred a comparable amount of heme to SbnI C168A and SbnI C155A compared to wild-type SbnI (Figure 4-16). SbnI H3A, SbnI C244A, SbnI H3A/C244A and SbnI1-240 were impaired in accepting heme from IsdI (Figure 4-16). Spectra of the flow-through and eluent fractions collected from the pull-down assay is representative of the results from three replicates (Figure 4-17). Flow-through and eluent samples were analysed by SDS-PAGE. The flow-through contained protein with corresponding molecular weight to SbnI and the eluent contained protein with corresponding molecular weight to IsdI. The absence of SbnI in the eluent, as judged by SDS-PAGE, suggests against formation of a stable complex (data not shown).        112  Figure 4-16 Quantification of IsdI heme transfer to SbnI and SbnI variants.  Quantification of IsdI heme transfer to SbnI variants was based on the fraction of heme transferred from IsdI using a pulldown assay. Briefly strep-tagged holo-IsdI was bound to streptactin resin. SbnI variant or buffer (as no transfer control) was added, incubated for 1 minute, separated by centrifugation, and supernatant containing the SbnI variant was removed. The fraction of IsdI heme transferred was calculated based on the amount of holo-IsdI eluted from streptactin resin when incubated with buffer compared to incubation with the SbnI variant.  The amount of holo-IsdI was calculated based on the A412 (wavelength at which holo-IsdI absorbs maximally) to A280 ratio of the eluent containing IsdI. No heme transfer is equal to the A412/A280 of holo-IsdI incubated with buffer. Statistics are calculated based on multiple comparisons with wildtype SbnI. ** p-value < 0.0021, *** p-value < 0.0002.    113  Figure 4-17 IsdI heme transfer to SbnI and SbnI variants using the pull-down assay. UV-vivible spectra of the flow-through fraction (dashed line) and IsdI-strep eluent (black line) following pull-down assay to ascertain heme transfer from strep-tagged IsdI to SbnI variant.  The flow-through contains either (A) buffer control or (B-I) an SbnI variant that was incubated with heme-bound strep-tagged IsdI bound to strep resin and removed by centrifugation. Eluent contains strep-tagged IsdI eluted from streptactin beads.  4.2.11 IsdI heme transfer to SbnI outcompetes heme degradation In the presence of molecular oxygen and a reductant, IsdI functions as a heme-degrading enzyme (147, 150). IruO was identified as a flavin mononucleotide containing oxidoreductase that transfers electrons from NADPH to IsdI (152). In a bacterial cell, heme transfer from IsdI to   114 SbnI would compete with heme degradation. To test if IsdI preferentially degraded heme or transferred it to SbnI, IsdI heme transfer to SbnI was assayed in the presence of IruO and NADPH. IsdI heme degradation can be assessed by a decrease in the Soret peak at 412 nm. Instead, IsdI was found to transfer heme to SbnI in the presence of IruO and NADPH as indicated by the shift of the Soret peak from 412 nm to 424 nm and the appearance of spectra characteristic of SbnI bound to heme (Figure 4-18A). However, a gradual decrease in the Soret peak height suggested heme was still degraded with a half-life of approximately 17 minutes. The residual activity could be due to heme transfer equilibrium between SbnI and IsdI. Control reactions were monitored for IsdI degradation of heme in the presence of IruO and NADPH, IsdI heme transfer to SbnI in the presence of IruO only, no degradation of heme by IsdI in the presence of IruO only, and that no heme degradation by SbnI took place in the presence of IruO and NADPH (Figure 4-18BCDE).       115  Figure 4-18 IsdI heme transfer or degradation competition experiment between SbnI and IruO.  For these experiments, 5 µM IsdI was reconstituted with 0.5 molar equivalents of heme. (A) Spectra of IsdI before and after the addition of 5 µM SbnI, 5 µM IruO, and 100 µM NADPH. Spectra of positive controls (B) IsdI heme degradation in the presence of 5 µM IruO and 100 µM NADPH and (C) IsdI heme transfer to 5 µM SbnI in the presence of 5 µM IruO without NADPH. Negative controls show (D) no degradation of heme by IsdI in the presence of 5 µM IruO only, and (E) no heme degradation by 5 um SbnI previously reconstituted with equimolar heme in the presence of 5 µM IruO and 100 µM NADPH.   4.3 Discussion SbnI has two distinct roles in SB biosynthesis: first to serve as a heme-dependent regulator of the sbn locus and, second, to produce OPS, a precursor for SB production. Heme-binding has only a modest effect on enzymatic activity suggesting that these two functions are independent of each other and that the heme binding site is structurally distinct from the active site for serine kinase activity. These findings support a model where SbnI mediates the S. aureus iron source preference switch between heme and siderophore acquired iron by contributing precursor OPS to SB synthesis when heme-iron is not available and directly sensing heme to shut off SB synthesis when heme-iron is present. Adaptation of siderophore production to suit the host niche has been   116 proposed for the differential production of SA and SB, where siderophore production is dictated based on localized host metabolite concentrations of iron and glucose (90). Heme transfer from IsdI to SbnI offers a mechanism for S. aureus to control iron source preference based on the iron sources available in the host environment.  Site-directed mutagenesis and biochemical methods were used to probe the heme-binding site and heme-binding function of SbnI. Features of the UV-visible spectra of SbnI bound to Fe(III) heme were consistent with the spectral characteristics of hemoproteins that bind heme such that iron is hexacoordinate and low-spin with a cysteine (thiolate) ligand opposite a neutral donor, most commonly a histidine (223). Mutation of SbnI His3 and Cys244 resulted in altered heme-binding spectra and the ability of these variants to accept heme from IsdI was impaired suggesting that they may serve as the axial heme iron coordinating ligands. In contrast, mutagenesis of other conserved cysteine residues, Cys155 and Cys168, had no apparent effect on heme binding or transfer. To define the heme-iron coordinating ligands and elucidate the conformation of SbnI in the holo-form I sought to obtain a model of heme-bound SbnI based on a co-crystal structure but suitable crystals were not obtained. An alternative, albeit more limited, approach was taken to generate a model by computationaly docking heme into the apo-structure. Molecular docking predicted that SbnI and SpSbnI bind heme in a conserved groove between domain I and domain III swapped from the adjacent protomer. SbnI Cys244 and SpSbnI Cys243 are located within the groove that also contains conserved, positively charged amino acids that could function to stabilize the heme propionate groups, broadly supporting the model. However, it does not directly explain spectra that suggests SbnI binds heme iron hexacoordinated with Cys/His ligation. Alternatively, the model may represent an initial heme-binding site as heme   117 encounters apo-SbnI that undergoes a conformational change to form the binding pocket in the final heme-bound state.  The H3A and C244A mutations were additive in diminishing the Soret peak intensity similar to observations for other heme-binding proteins. For example, PhuS is a heme-trafficking protein in Pseudomonas aeruginosa that delivers heme to the heme oxygenase, HemO. Mutation of both His ligands to the heme iron did not eliminate heme-binding even though both His ligands are required for protein-protein interaction with HemO and subsequent heme transfer, highlighting the flexibility of the heme environment (236, 237). Similarly, S. aureus ChdC site-directed mutants of eight distinct residues in the substrate binding site all bound coproheme with >80% occupancy and had measured Kd values within an order of magnitude of wild-type (238). However, mutants that had a functional defect in catalytic competence were identified (238). Single amino acid substitutions to alanine may be insufficient to abrogate heme-binding for proteins with a high-affinity for heme such as SbnI. UV-visible absorption spectra suggest that SpSbnI also binds heme-iron in a low-spin, hexacoordinate manner with ligation by a cysteine (thiolate). The second residue participating in this interaction is unknown as SpSbnI does not have a homologous histidine at position 3. The lack of conservation argues against His3 serving as the heme iron coordinating ligand trans to the cysteine (thiolate) in the final heme bound form of S. aureus. His3 is only conserved amongst SbnI homologs from the closely related and recently distinguished staphylococcal species, Staphylococcus argentus (formerly S. aureus clonal complex 75) and Staphylococcus schweiteri (239, 240). This residue may have a role in heme transfer and another residue or a solvent molecule serves as the sixth ligand.   118 A minor portion of the top solutions from the docking experiment with SpSbnI placed the heme in the L-serine kinase active site. These are likely not physiological as the SbnI co-crystal structure with ADP confirms this region as the kinase active site and heme was not observed to have a large effect on SbnI L-serine kinase activity. These findings support a model where heme binds at a structurally distinct site from the kinase active site. The multiple sequence alignment of staphylococcal SbnI homologs illustrates that there are conserved residues in both the kinase and heme binding sites that are likely important for binding their respective ligand or substrate (Figure 4-7).  There is a clear relationship between redox status and iron homeostasis in S. aureus (109, 110, 116). S. aureus routinely encounters oxidative stress during infection and regulation of iron supply by the Fur regulon and oxidative stress resistance by the PerR regulon are important for S. aureus survival (110). As excess free iron can participate in damaging Fenton reactions, SB expressed during bacteremia may require additional layers of redox-sensitive control to limit iron uptake. The SbnI structure reveals the likely role of redox sensing in the function of SbnI based on the formation of the dimer of dimers via reversible disulfides. The formation of dimer of dimers could alter SbnI control of sbn gene expression, the enzymatic activity of SbnI to generate SB precursor molecule OPS, or both. Reversible disulfide bond formation and oligomeric state is a mechanism used by some bacterial transcription factors to sense oxidative stress (241, 242). The E. coli peroxide sensing global regulator, OxyR, regulates cell resistance to oxidative stress (243). Two cysteine residues in each OxyR monomer form an intramolecular disulfide bridge that induces tetramer formation and DNA-binding to activate transcription (241). Alternatively, AgrA in S. aureus senses oxidative stress using an intramolecular disulfide that leads to decreased affinity for DNA (242). Whether changes in the oligomeric state of SbnI, as a   119 result of changing redox conditions, effect its enzymatic or regulatory functions in vitro and in vivo is the subject of ongoing work.  SbnI is a heme sensing protein that can obtain heme from IsdI. IsdI binds heme with a KD of 12.9 ± 1.3 nM (233). The binding affinity is consistent with its function as a heme-degrading protein for maintenance of cytosolic heme homeostasis around 20-40 nM the measured concentration range for labile heme pools in Saccharomyces cerevisiae and eukaryotic HeLa cells (244, 245). Heme is physically transferred from IsdI to SbnI through a protein-protein interaction. SbnI was determined to have a high nanomolar heme binding affinity (KD = 125 ± 105 nM), which likely represents an upper estimate of heme affinity.  Moreover, IsdI heme transfer to SbnI was found to occur in the presence of the in vivo reductase for IsdI heme degradation, IruO. Though IsdI is characterized as a heme-degrading enzyme, intact heme is required for a variety of cellular processes (147, 150). Thus, not all host-acquired heme is immediately degraded and IsdI may function as a heme-trafficking protein in addition to a heme-degrading enzyme. Heme acquisition provides an advantage during pathogenesis by enabling aerobic respiration and catalase activity. IsdI heme transfer to SbnI may function as a signal of active heme uptake by the Isd system leading to coordination of other iron uptake mechanisms, such as decreased SB production. As S. aureus is likely to encounter sudden, drastic changes in extracellular heme concentrations (eg. upon lysing red blood cells), it is possible that heme itself functions as an important environmental signal that is immediately sensed upon uptake to impart changes on gene transcription and facilitate niche adaptation. The ultimate fate of cytosolic heme is likely dependent on the intracellular and extracellular availability of iron and heme (66).  Based on interactions observed with the Isd system and IruO, SbnI may serve as a regulator of iron source preference in S. aureus through direct interaction with IsdI. Low-spin,   120 hexacoordinate heme iron with thiolate ligands are frequently components of signaling pathways as the labile nature of the Cys ligand render these heme centres well suited for small molecule sensing and transport (223). These results support the existence of a regulatory network between SbnI, IsdI, IruO, to optimize iron uptake strategies and creates a model for how intracellular heme homeostasis is maintained. The regulatory role of SbnI provides rationale for how S. aureus demonstrates heme iron preference as exogenous heme uptake would inhibit positive regulation of the sbn gene cluster, thereby limiting the production of SB. When infection is in a heme-rich environment, S. aureus may preferentially devote energy towards heme uptake rather than the metabolically taxing process of siderophore biosynthesis. Together, there is a close relationship between nutrient supply and gene expression and mounting evidence that host sensing plays a key role in S. aureus adaptation of iron uptake and survival strategies used to advance pathogenesis and infection.     121 Chapter 5: Role of the terminal heme biosynthetic enzyme, ChdC, in heme homeostasis and heme-trafficking   5.1  Introduction Heme is an integral nutritional requirement for bacteria that must be satisfied either by acquisition or synthesis. As a redox active molecule, heme is important for the function of many cellular proteins including as a cofactor for catalase to resist oxidative stress and by enzymes required for electron transport to enable cellular respiration (126, 127). Most bacterial pathogens encode a heme biosynthetic pathway but the contribution of heme biosynthesis to pathogenesis and the regulation of heme production is largely understudied (128). In S. aureus, a functional heme biosynthetic pathway is required for full pathogenesis and colonization of distinct organs in murine models of infection (127, 129). Additionally, a defect in the heme biosynthetic pathway is a defined auxotrophy observed in small colony variants (SCVs) (22). This phenotypic switch has implications for survival and persistence in the host. Unclear is if S. aureus relies on heme biosynthesis in specific spatiotemporal niches and whether there is variability in how endogenous and exogenously-acquired heme is partitioned to hemoproteins in the cell. However, a coordinated iron-regulatory response during infection likely exists to ensure heme homeostasis. Knowledge of prokaryotic heme biosynthesis has recently expanded to include a coproporphyrin-dependent heme biosynthetic pathway used by most Actinobacteria and Firmicutes (131, 132). S. aureus heme biosynthesis proceeds through this newly described non-canonical pathway that features ChdC, a coproheme decarboxylase, for catalysis of iron-coproporphyrin III (coproheme III) to heme (131, 137, 238). Prior to assignment in the heme biosynthetic pathway, ChdC (formerly HemQ) was thought to function as a chlorite dismutase   122 (139, 140), but with an obvious link to heme biosynthesis as a chdC mutant strain had a SCV phenotype under aerobic conditions (139). Additionally, ChdC was shown to degrade heme in the presence of minimal hydrogen peroxide, peracetic acid, or chlorite (139). The observation that heme bound to ChdC was redox sensitive led to the hypothesis that ChdC could sense iron availability or redox state (139). Thus, ChdC could play a dual role in maintaining S. aureus heme levels: first, as a decarboxylase in heme synthesis and, secondly, as a regulatory protein in heme homeostasis, catalyzing the degradation of bound heme when it is not utilized for cellular metabolism (131). Despite recent advances in defining the heme biosynthetic pathway in Gram positive bacteria, it is unclear how newly synthesized heme is trafficked and delivered to heme-requiring proteins in the cell. As free intracellular heme in the cell can be toxic, I hypothesized that ChdC can serve as a heme source for heme-utilizing proteins like SbnI and the paralogous heme-degrading proteins IsdI and IsdG. Previous research suggested holo-ChdC could supply heme-requiring proteins with heme and that apo-ChdC would allow heme biosynthesis to continue (139).   The aim of this study was to solve the structure of S. aureus ChdC and to determine whether it can participate in heme transfer reactions to other cytosolic heme-binding proteins, namely IsdI, IsdG, and SbnI. As described in Chapter 4, IsdI catalytically transfers heme to SbnI, here we extend these experiments to determine whether IsdG can transfer heme to SbnI. UV-visible spectroscopy and kinetic analysis of heme transfer reactions were used to further develop a model for intracellular heme trafficking in S. aureus and to understand how S. aureus may promote a coordinated response to the changing availability of nutritional iron sources during infection.    123 5.2  Results 5.2.1 Structure of S. aureus ChdC  The crystal structure of S. aureus ChdC (SaChdC) was solved to 2.5 Å with Rwork (Rfree) of 0.22 (0.27). The Geobacillus stearothermophilus ChdC structure (PDB ID: 1T0T, 58% sequence identity) was used for phasing by molecular replacement. S. aureus ChdC crystallized in space group P21212 with 10 molecules in the asymmetric unit organized as two homopentamers, each with 5-fold rotational symmetry (Figure 5-1A). The two homopentamers are related by non-crystallographic symmetry and the 5-fold axes are approximately perpendicular (Figure 5-1A). Modelled residues (97.0%) are in the most favored regions of the Ramachandran plot (168). Data collection and refinement statistics are summarized in Table 2-5. In all 10 protomers, a loop region between residues 113-119 had poor electron density and anywhere between 3 to 7 amino acids were not modelled. Modelled at crystallographic symmetry axes and the inside edge of domain I are 15 hexamine cobalt(III) molecules. Peaks in the anomalous electron density map for the cobalt ions was observed at these locations. The hexamine cobalt(III) molecules likely help stabilize the crystal contacts as its inclusion during optimization of crystal growth conditions altered crystal morphology and led to higher resolution diffracting crystals.     124   Figure 5-1 Structure of S. aureus ChdC. (A) S. aureus ChdC crystallographic dimer of pentamers represented as a cartoon with each protomer colored differently. The pentagon indicates 5-fold rotational symmetry. (B) The ChdC protomer is divided into three domains. N- and C-termini and long a-helices (>10 residues) are labeled.   Each ChdC protomer is made up of two ferredoxin-like a/b domains related by pseudo-2-fold symmetry with a central loop region that is directed outward from the pentamer. Domain I, the N-terminal ferredoxin-like a/b domain, consists of residues M1-V107 and I238-S250, domain II contains residues I108-H138 and forms the loop region, and domain III consists of residues S139-I235 and forms the C-terminal ferredoxin-like a/b domain (Figure 5-1B). Domain I is not known to have any catalytic function. Previous analysis of the loop region from diverse ChdC homologs was used to define the domain II (loop region) in SaChdC, discussed below (246). Domain III contains the putative coproheme decarboxylase active site and the absolutely conserved functional YP(M/F)X(K/R) motif for ChdCs (246), which is 145YPMNK149 in SaChdC. Interface analysis by PDBePISA (218) calculated an average of 1858 Å2 buried   125 interface area between protomers of the homopentamers with 40 predicted hydrogen bonds. Regions of the protein involved in hydrogen bonding at the interface include a-helix 2 (a2, residues 80-92) with the C-terminal residues 245-250 and between a-helix 3 (a3, residues 206-219) and N-terminal residues 2-9. The buried surface interface between asymmetric assembly of adjacent homopentamers observed in the crystal structure is 399 Å2 and is predicted by PDBePISA to not play a role in complex formation and rather is more likely a result of crystal packing. This analysis suggests that the functional unit in the SaChdC crystal is a homopentamer. Previous analysis of heme-bound SaChdC suggested that it is a hexamer in solution, as determined by analytical gel filtration (139). Conversely, apo-SaChdC eluted near the column’s void volume giving an estimated molecular mass of 1200 kDa, which was thought to be a multimer, nonglobular protein structure, or a highly disordered but soluble protein (139). ChdC structures solved to date include those from Geobacillus stearothermophilus, Listeria monocytogenes, Thermus thermophilus, and Thermopasma acidophilum (Table 5-1). Structural comparison of SaChdC with the structures of these ChdC homologs revealed that the overall fold is similar and they all appear to be a pentamer (Table 5-1). One exception identified was the structure of G. stearothermophilus PEG330-bound ChdC (GsChdC), which appears to form a higher-order complex as a 12-mer of pentamers, or 60-mer of protomers, to form a hollow sphere by crystallographic symmetry (PDB ID: 1T0T). Similar to the discrepancy between solution state and crystallographic analysis of oligomerization of S. aureus ChdC, L. monocytogenes appears to be hexameric in solution and pentameric in the crystal structures (247).       126 Table 5-1 ChdC structures, oligomerizations state, and area of protomer interfaces.  a, b, c, d from references (139), (238), (247), (138), respectively. Source PDB ID Amino acid sequence identity with SaChdC Superimposition apo-SaChdC (RMSD, across number of residues) Oligomerization Buried surface area between protomers (Å)  Ligands in active site  Staphylococcus aureus (apo-SaChdC) This work - - Higher-order complex (~1200 kDa) in solutiona, pentamer in crystal 1858 Å2 between protomers of pentamer  ligand-free Staphylococcus aureus This work - - Hexamer in solutiona, pentamer in crystal, 60-mer by crystallographic symmetry 1917 Å2 between protomers of pentamer,  163 Å2 between protomers of 60-mer Co-crystallized with heme Geobacillus stearothermophilus 1T0T 58% 0.8 Å, 237 Pentamer in crystal, 60-mer by crystallographic symmetry 1609 Å2 between protomers of pentamer,  731 Å2 between protomers of 60-mer PEG330 bound Geobacillus stearothermophilus 5T2K 58% 0.7 Å, 240 Pentamer in crystal 1726Å2 between protomers of pentamer  Mn(III) coproheme boundb Listereria monocytogenes 5LOQ 54% 0.7 Å, 240 Hexamer in solutionc Pentamer in crystal 1644 Å2 between protomers of pentamer  Fe(III) coproheme boundd Listereria monocytogenes 4WWS 54% 0.7 Å, 236 Hexamer in solutionc Pentamer in crystal 1584 Å2 between protomers of pentamer Ligand-free Thermus thermophilus 1VDH 44% 1.3 Å, 241 Pentamer in crystal 1728 Å2 between protomers of pentamer  Ligand-free Thermoplasma acidophilum 3DTZ 23% 2.4 Å, 216 Pentamer in crystal 1563 Å2 between protomers of pentamer 399 Å2 between dimer of pentamers Ligand-free   127 A co-crystal structure of G. stearothermophilus ChdC with a substrate analog, Mn(III)coproheme (PDB ID: 5T2K) and biochemical characterization of the ChdC reaction mechanism were recently published (238). ChdC coproheme decarboxylase catalyzes two sequential oxidative decarboxylations with hydrogen peroxide as the oxidant, coproheme III as substrate and cofactor, and heme and CO2 as the products. The proposed reaction mechanism involves use of an extended relay of propionate and sidechain mediated electron/proton transfers from the pair of propionate groups and to H2O2 bound to the iron, ultimately releasing CO2 and forming the vinyl groups of heme (238). This mechanism represents a novel route to convert propionate groups to vinyl groups. Superimposition of SaChdC with GsChdC: Mn(III)coproheme revealed that several important catalytic and substrate binding residues are structurally conserved in S. aureus ChdC (Figure 5-2). These include potentially mechanistically important residues Trp198 and Trp157, as well as Arg218, Arg131, Thr223, Gln185 which are involved in an H-bond network with propionate groups 2, 4, and 6 of the Mn(III)coproheme (238). Coproheme oxidations are suggested to occur through mediating amino acid residues, Tyr145 and Lys149, that are H-bonded to the reactive propionates 2 and 4, respectively, and play essential roles in the regioselective oxidation of these propionates (238). These two mediating residues are structurally conserved in SaChdC and likely support a similar enzymatic mechanism for coproheme decarboxylase activity (Figure 5-2B). Additionally, His172, the proximal coproheme Mn(III) ligand and Ile187 that forms part of the distal substrate binding pocket are structurally conserved (Figure 5-2B). In the Mn(III)-coproheme bound GsChdC crystal structure, the tetrapyrrole ring is slightly ruffled with the reactive propionates (2 and 4) slightly below the approximate porphyrin plane and pointing toward the protein core, away from the solvent (238).     128  Figure 5-2 Superimposition of apo and ligand-bound forms of G. stearothermophilus ChdC with S. aureus ChdC.  (A) Structural superimposition of SaChdC protomer in lime and Mn(III)coproheme bound GsChdC protomer in raspberry (PDB ID: 5T2K). (B) Close-up view of Mn(III)coproheme binding site of GsChdC (raspberry) and SaChdC (lime) highlights that the active site architecture is highly conserved and that residues are positioned similarly between the apo- and holo- structures. Selected active site residues and heme are drawn as sticks. Fe, O, and N atoms are colored purple, red, and blue, respectively. Heme carbons are colored grey.   ConSurf analysis of SaChdC was used to map conserved regions onto the surface of the structure (Figure 5-3). The C-terminal ferredoxin-like a/b domain (domain III), which contains the active site, is more highly conserved than the N-terminal ferredoxin-like a/b domain (domain I) ChdC (Figure 5-3A). Surface analysis of the SaChdC protomer reveals regions that interface with adjacent protomers to form the pentamer have areas of conservation and that surface exposed regions of domain I and the loop region of domain II are variable in sequence (Figure 5-3BC). Surface-exposed regions of conservation in the SaChd pentamer are concentrated around the active site in domain III (Figure 5-3D).       129  Figure 5-3 Conservation of SaChdC residues generated using ConSurf. (A) Ribbon representation of SaChdC protomer with domains labelled. (B) Surface representation of ChdC protomer and (C) after 180° rotation about the y axis. Blue circles denote areas of conservation that interface with adjacent protomers to form the pentamer. The black arrow points to the active site. (D) Surface representation of the SaChdC pentamer. Conserved amino acids are colored bordeaux, residues of average conservation are white, and variable amino acids are turquoise.  5.2.2 Structural comparison of ChdC homologs  The secondary structural elements of all ChdC crystal structures available in the PDB were compared with SaChdC (Figure 5-4A). Domain I and domain III of the ChdC structures are very similar and have an overall ferrodoxin-like fold. The loop region that constitutes domain II differs the most between structures (Figure 5-4A). In SaChdC, like GsChdC:Mn(III)coproheme, a portion of the loop (residues 111-121) could not be modelled suggesting that this region is dynamic. The dynamic loop in GsChdC:Mn(III)coproheme was postulated to accommodate substrate binding and product release as the closed conformer repositions residues within binding pocket that form contacts with propionates (238). Conversely, the domain II loop in apo-GsChdC is in a comparatively more open conformer (PDB ID: 1T0T). The domain II loop regions in both the apo- and Fe(III)coproheme bound L. monocytogenes ChdC (LmChdC) structures are oriented similarly in a more open, solvent exposed conformer (PDB IDs: 4WWS, 5LOQ) and resembles that of apo-GsChdC. It should be noted that domain II was only resolved for two of five   130 protomers in both LmChdC structures where crystal contacts with adjacent molecules stabilized the loop (138, 247). T. thermophilus ChdC (TtChdC) domain II loop has little defined secondary structure and folds down towards domain III in the crystal form (PDB ID: 1VDH). Lastly, T. acidophilum domain II loop has two a-helices and folds slightly towards domain III (PDB ID: 3DTZ). A sequence alignment of the ChdC homologs that are available in the PDB with SaChdC reveals there is sequence variability in domain II (Figure 5-4B). However, the functional YP(M/F)X(K/R) motif is conserved across all sequences (Figure 5-4B).     131  Figure 5-4 Structural overview of ChdC protomers.  (A) Cartoon representation of ChdC protomers. Domain I and III are oriented towards the top and bottom of the page, respectively, and domain II (loop region) is colored red. The top row of pentamers are ligand-free and the bottom row are substrate or substrate-analog bound, as indicated. (B) Sequence alignment of the five ChdC homologs investigated. The red box highlights domain II (loop region) and the green box denotes the conserved sequence motif for ChdCs. Red triangles indicate amino acids that form hydrogen bonds to generated the 60-mer observed in apo-GsChdC structure. SaChdC, S. aureus ChdC; GsChdC, G. stearothermophilus ChdC (PDB IDs: 1T0T, 5T2K); LmChdC, L. monocytogenes ChdC (PDB IDs: 5LOQ, 4WWS); TtChdC, T. thermophilus ChdC (PDB ID: 1VDH); TtChdC, T. acidophilum ChdC (PDB ID: 3DTZ).     132 5.2.3 GsChdC structure forms a 60-mer by crystallographic symmetry Further analysis of the crystal packing of the apo-GsChdC structure (PDB ID: 1T0T) revealed that the domain II loop is in contact with a protomer of an adjacent pentamer near the active site in domain III (Figure 5-5). These protein-protein contacts appear to facilitate the formation of a spherical 12-mer of pentamers (or 60-mer) with an open core that is 21 nm in diameter (10 nm inner diameter) (Figure 5-5A). Domain II loops of protomers from adjacent pentamers are related by 3-fold symmetry to form the protein-protein interfaces for 60-mer multimerization (Figure 5-5B). Residues involved in forming H-bonds at the 60-mer interface are between domain II and domain III and include Ser111, Asn112, Tyr113, Glu126, Arg129, Arg176, and Gln184 (Figure 5-5C), all of which are conserved in SaChdC and LmChdC except for Glu126 and Arg129 (Figure 5-4B). Interface analysis by PDBePISA predicts that these contacts plays an auxiliary role in formation of the 60-mer, as 713-752 Å2 buried interface area is between protomers of adjacent pentamers. By comparison, 1602-1626 Å2 buried interface area is between ChdC protomers that form the homopentamer. In the 60-mer, domain III of each protomer are oriented towards the inside of the sphere, while domain I of each protomer are facing outward from the spherical 60-mer. Superimposition of a GsChdC:Mn(III)coproheme protomer with the apo-GsChdC multimer reveals that formation of this higher-order complex encloses the active sites of GsChdC protomers (Figure 5-6AB). In this oligomerization state, the loop region of domain II is within close proximity Mn(III)coproheme binding site (Figure 5-6C).     133  Figure 5-5 Structure of GsChdC 60-mer and 60-mer multimerization interface. GsChdC 60-mer formed by crystallographic symmetry of 12 pentamers (PDB ID: 1T0T). Colored by pentamer. Higlighted by the red box is the region of the GsChdC protein oligomer in panel (B). (B) The loop region of domain II from adjacent protomers (colored red, magenta, and raspberry) from adjacent pentamers form 60-mer multimerization interfaces and are related by 3-fold symmetry. The domain II loop protrudes into the active site, near where a PEG330 (orange) is modelled in the GsChdC structure. (C) Selected residues predicted to form hydrogen bonds at 60-mer multimerization interface are drawn as sticks with the loop region of domain II colored red. GsChdC protomer carbon are colored red, and blue. These protomers belong to separate homopentamers. PEG330 carbons, O, and N atoms are colored orange, red, and blue, respectively.        134  Figure 5-6 GsChdC 60-mer multimerization encloses the active site. (A) Surface representation of Mn(III)coproheme bound GsChdC (PDB ID: 5T2K) protomer (raspberry) superimposed with apo-GsChdC (PDB ID: 1T0T) pentamer (dark blue). The black arrow points to the bound Mn(III)coproheme (grey). (B) Adjacent apo-GsChdC pentamer, constructed by crystallographic symmetry (green) covers the Mn(III)coproheme binding pocket. The red square highlights the region of the protein shown in panel (C). (C) The domain II loop of the adjacent protomer (colored red, rest of protomer is green) in the 60-mer oligomerization state interfaces near the ChdC active site, where Mn(III)coproheme substrate analog binds.  5.2.4 Low resolution crystal structure of SaChdC crystallized in the presence of heme Red crystals of SaChdC co-crystallized with heme were obtained and the structure was solved to 6.5 Å resolution. Data collection and refinement statistics are summarized in Table 2-5. The SaChdC pentamer was used for phasing by molecular replacement using Phaser-MR from Phenix (168). The top 10 solutions from molecular replacement were in space group I23 after searching for solutions in all space groups within the point group 23, supporting that the crystal was indeed in the space group I23. A solution with 5 molecules in the asymmetric unit gives an estimated Matthews coefficient of 2.70 Å3 Da-1 with an estimated solvent content of 54.4%.   135 Electron density for the donut shaped homopentamer could be distinguished as evidenced by a rendering of the 2Fo–Fc electron density map (contoured to 1s) with ChdC pentamer modelled (Figure 5-7A). After refinement with each protomer as a rigid body, the Rwork (Rfree) was 0.21 (0.45). Further analysis of crystal packing revealed that pentamers related by crystallographic symmetry form a higher order spherical multimer, consisting of 12 pentamers and yielding a 60-mer with an open core (Figure 5-7B). The radius of the sphere is 22 nm with an inner, hollow core of 10 nm. The buried surface interface between protomers of the same pentamer is 1917 Å2 and 163 Å2 between protomers of adjacent pentamers to form the 60-mer. Similar to GsChdC 60-mer, regions of SaChdC co-crystallized with heme that form the interfaces to construct the 60-mer are concentrated around the active site in domain III and domain II. Each domain III of the pentamers form the inner side of sphere and domain Is of the pentamers form the outer surface of the sphere. One caveat for the calculation of the buried surface area of the 60-mer interfaces is that residues were not modelled for part of the loop in domain II, the region that forms part of this interface in GsChdC, potentially underestimating the buried surface area. Another similarity with GsChdC was that both 60-mer crystals were in the same space group, I23, and had a similar unit cell edge length of 208.8 Å and 202.8 Å for SaChdC and apo-GsChdC (PDB ID: 1T0T), respectively. Due to low resolution of the data set, whether or not heme was indeed bound to the active site was not determined.   136  Figure 5-7 Structure of S. aureus ChdC co-crystallized with heme (A) 2Fo–Fc electron density map (contoured to 1s) for SaChdC homopentamer co-crystallized with heme. Colored by protomer. (B) SaChdC 60-mer constructed through crystallographic symmetry of 12 pentamers. Colored by pentamer.  5.2.5 Determination of ChdC:heme off-rate The S. aureus ChdC:heme off-rate was determined based on the rate of heme transfer to apomyoglobin. Heme binding by apomyoglobin was followed by visible stopped-flow spectroscopy at 408 nm, the Soret maximum for holomyoglobin. The transfer rate data was best fit by a triple exponential curve, as judged by the randomness of the residual plot and the rate of heme transfer to apomyoglobin was independent of apomyoglobin concentration. The rate of heme dissociation from S. aureus ChdC was measured to have an average k1 of 0.28 ± 0.01 s-1, k2 of 0.02 ± 0.01 s-1, and k3 of 0.01 ± 0.01 s-1. Each accounted for 23%, 49%, and 28% of the absorption change, respectively.     137 5.2.6 ChdC can actively transfer heme to SbnI and IsdG, but not IsdI Heme transfer from S. aureus ChdC to SbnI, IsdG, and IsdI was examined by UV-visible spectroscopy. ChdC bound to heme has a characteristic Soret peak at 403 nm. Upon addition of apo-SbnI the spectra shifted to that of holo-SbnI as evidenced by the a Soret peak shifted to 422 nm with a shoulder at 380 nm, and alpha and beta bands at 573 nm and 541 nm (Figure 5-8A). The rapid spectral shift from that of holo-ChdC to that of holo-SbnI suggests that heme is transferred from ChdC to SbnI. Similar results were observed when holo-ChdC was incubated with apo-IsdG. A shift in the Soret to 411 nm, characteristic of IsdG, suggested ChdC can transfer heme to IsdG as well (Figure 5-8B). Heme-bound IsdI has a Soret peak maxima at 412 nm, but only an intermediate spectral shift was observed upon addition of IsdI to holo-ChdC, to 407 nm, suggesting incomplete heme transfer (Figure 5-8C). To test whether the intermediate IsdI spectra represented an evenly mixed holo-ChdC/holo-IsdI population at equilibrium, the opposite reaction, IsdI-heme transfer to apo-ChdC, was measured and no spectral shift was observed (Figure 5-8D).    Figure 5-8 ChdC:heme transfer to SbnI, IsdG, and IsdI.  The spectrum of 5 µM ChdC with equimolar heme was recorded (dashed line) and subsequently 5 µM of (A) SbnI, (B) IsdG, or (C) IsdI was added to the cuvette and spectra was recorded immediately (black line). (D) Similarly, 5 µM IsdI with equimolar heme was recorded (dashed line) followed by addition of 5 µM ChdC and the spectrum was measured (black line).   138  To determine whether the heme transfer from ChdC to the various heme acceptors represented active transfer or a passive equilibrium with a free heme intermediate, the rates of heme transfer from holo-ChdC to SbnI, IsdG, and IsdI were measured using stopped-flow spectroscopy. The observed rates (kobs) were determined by the monitoring absorption changes in the Soret region.  The wavelengths of maximal change in absorption between holo-ChdC and holo-SbnI, holo-IsdG, or holo-IsdI, were at 430 nm, 417 nm, and 416 nm, respectively. Data were best fit by a double exponential curve for ChdC:heme transfer to IsdI and a triple exponential curve for reaction with SbnI and IsdG, as judged by the randomness of the residual plots. As the ChdC pentamer binds five molecules of heme and recipient proteins, SbnI, IsdG, and IsdI are each dimeric where each protomer binds one molecule of heme, the overall rate of heme transfer may reflect a complicated mechanism of heme transfer between oligomers. The observation of multiple rates (kobs) could be due the formation of intermediate species that transfer heme at different rates. However, assignment of these rates to heme transfer between specific species is not yet possible. The measured heme transfer rates and ChdC:heme off-rate are summarized in Table 5-2.   Table 5-2 Kinetic parameters of ChdC:heme off-rate and ChdC:heme transfer to SbnI, IsdG, and IsdI.    k1 (s-1)  k2 (s-1)  k3 (s-1)  ChdC:heme off-rate 0.28 ± 0.01 (23%) 0.02 ± 0.01 (49%) 0.01 ± 0.01 (28%) ChdC:heme ® SbnI 5.40 ± 0.04 (33%) 0.18 ± 0.01 (14%) 0.02 ± 0.01 (53%) ChdC:heme ® IsdG 4.47 ± 0.06 (24%) 0.08 ± 0.01 (14%) 0.01 ± 0.01 (62%) ChdC:heme ® IsdI 0.15 ± 0.01 (24%) 0.02 ± 0.01 (76%)      139 The rate of heme transfer from ChdC to SbnI, IsdI, and IsdG was compared to the rate of heme release (off-rate) of ChdC to discern if there was catalytic transfer of heme between the two proteins. The initial rate of ChdC:heme transfer to SbnI was an order of magnitude greater than the ChdC:heme off-rate. This result strongly suggests that heme bound by apo-SbnI is not first released into solution by ChdC, but instead is transferred from ChdC to apo-SbnI by a protein-protein interaction. Similarly, ChdC:heme transfer to IsdG was ~10 fold greater than the ChdC:heme off-rate indicating heme is catalytically transfered through a protein-protein interaction. However, heme transfer from ChdC to IsdI was within an order of magnitude of the ChdC:heme off-rate suggesting against a specific interaction between ChdC and IsdI. Rather, IsdI is more likely binding heme that dissociated from ChdC over the course of the experiment.   5.2.7 IsdG can transfer heme to SbnI To continue to build a model of intracellular heme transfer in S. aureus, heme transfer between IsdG and SbnI was measured. The equilibrium state of transfer from IsdG-heme to apo-SbnI was measured using UV-visible spectroscopy, and a shift in the spectrum was observed. Though the spectra did not completely resemble that of holo-SbnI, a red-shifted Soret and enhancement of a shoulder at ~360 nm was indicative that at least partial heme transfer may have occurred (Figure 5-9). Similar to study of SaChdC:heme transfer, the rate of heme transfer from holo-IsdG to apo-SbnI was measured using stopped-flow spectroscopy and compared to the IsdG heme off-rate. The IsdG heme off-rate data was best fit by a double exponential curve and using a biphasic model, k1 and k2 of IsdG heme transfer to SbnI were determined and summarized in Table 5-3. The rate of IsdG heme transfer to SbnI was ~5 fold greater than the measured IsdG heme off-rate. Rates are summarized in Table 5-2 and compared to IsdI heme off-rate and   140 transfer to SbnI. The measured IsdI and IsdG heme off-rates agree with previously published heme off-rates for these proteins (Table 5-3) (233).    Figure 5-9 IsdG heme transfer to SbnI.  The spectrum of 5 µM IsdG with equimolar heme was recorded (dashed line) and subsequently 5 µM of SbnI was added to the cuvette and the spectrum was recorded immediately (black line).  Table 5-3 Comparison of kinetic parameters of IsdG and IsdI heme off-rates and heme transfer to SbnI.   k1 (s-1)  k2 (s-1)  k3 (s-1)  IsdG heme off-rate 0.11 ± 0.01 (6%) 0.03 ± 0.01 (94%)   IsdG heme off-ratea 0.022 ± 0.002      IsdI heme off-rateb 0.13 ± 0.01 (93%) 0.04 ± 0.01 (7%)   IsdI heme off-ratea 0.092 ± 0.008      IsdG:heme ® SbnI 0.65 ± 0.03  (8%) 0.05 ± 0.01  (92%)   IsdI:heme ® SbnIb 1.55 ± 0.08 (14%) 0.16 ± 0.01 (78%) 0.05 ± 0.01 (8%) a Rates from reference (233). b Rates from Chapter 4.      141 5.3 Discussion SaChdC, amongst other ChdC homologs, has a pentameric structure with each protomer consisting of two ferredoxin-like a/b domains related by 2-fold pseudosymmetry. Interestingly, the same ferredoxin-like a/b domain structure in ChdC is shared by noncanonical IsdG-type heme oxygenases and heme-binding Dyp-type peroxidases, as noted previously (139, 248). Collectively, these enzymes belong to the functionally diverse CDE superfamily that use a versatile protein scaffold to carry out a variety of heme-requiring chemistries, where heme serves as either the cofactor, substrate, or product. Enzymes belonging to this superfamily are found in different oligomerization states, including dimer, trimer of dimers, homopentamer, and homohexamer (249). The different oligomerization states and subunit interactions, combined with distinct sets of conserved active site residues, likely influence the solvent and substrate accessibility of the heme-binding pocket, giving rise to the array of catalytic properties observed for these families belonging to the CDE superfamily (249).  SaChdC was found to crystallize in two different forms, as a pentamer and a spherical 60-mer (or 12-mer of pentamers) with a hollow center. One difference in the crystallization setup that may have led to the formation of the two oligomeric states was that the 60-mer resulted from SaChd that was co-crystallized with heme. It is tempting to hypothesize that the presence of heme contributed to the formation of the 60-mer. The high resolution crystal structure of GsChdC (1.75 Å, PDB ID: 1T0T) crystallized as a 60-mer where the protein interfaces involved in multimerization are near the active site, where heme would bind, in domain III and involve the loop in domain II. As the SaChdC 60-mer was of low resolution (6.5 Å), it is unknown if the heme is in fact bound to the active site and is directly involved in oligomerization. Previous reports have observed difference in ChdC oligomerization state depending on the presence of   142 heme (139, 247). Measurement by analytical gel filtration has shown that S. aureus apo-ChdC elutes near the column’s void volume giving an estimated molecular mass of 1200 kDa (predicted mass of 60-mer is 1763 kDa) and heme-bound SaChdC was hexameric would suggest that heme has the opposite effect on oligomerization (139). Conversely, study of LmChdC by size-exclusion chromatography that the majority of apo- and holo- protein was hexameric in solution but also observed peaks corresponding to higher oligomeric assemblies in both apo- and holo- samples, with a higher proportion of these larger oligomers in the latter (247). Other experimental factors that could have led to the two observed oligomeric states include differences in the crystallization solutions. The 60-mer was crystallized ~pH 6 and the pentamer was crystal was grown ~pH 7.4 (Section 2.2.3). The inclusion of hexamine cobalt (III) chloride as an additive to critical to improving apo-SaChdC crystal diffraction. These variations could contribute to the two multimerization states of SaChdC crystallized. The discovery that the apo-GsChdC structure formed a 60-mer has not been discussed in the literature despite intense bioinformatic analyses of this specific structure and other ChdC crystal structures (144, 238, 246, 249). The buried surface area between protomers of adjacent pentamers present in the high-resolution crystal structure (PDB ID: 1T0T) is an average of 731 Å2, with the 1917 Å2 of buried surface area between protomers of the same pentamer, and support that formation of the 60-mer is physiologically relevant. These interface areas are within the range observed for other bacterial higher-order multimers that form a sphere with a hollow core and are involved in iron homeostasis and storage. For example, ferritin forms a spherical nanocage made of 24 subunits that is 12 nm in diameter and has a hollow core. E. coli ferritin has an average buried surface area of 1109 Å2  between protomors of a subunit dimer and 420 Å2 between adjacent protomers related by 3-fold symmetry (PDB ID: 4XGS) (250). Encapsulin   143 nanocompartments are widely distributed in bacteria and archaea and form an icosahedral shell formed by oligomeric assembly of the protein (251, 252). These metabolic compartments have functionally diverse roles reflected by the function of the targeted encapsulin associated proteins that are sequestered (252). For example, Myxococcus xanthus encapsulin is 32 nm in diameter (26 nm inner diameter) and can sequester a number of proteins for nucleating iron deposition and act as an iron “mega-store”, supporting a hypothesis that its function involves regulated iron uptake to protect the bacteria from oxidative stress (253). M. xanthus forms a homo 180-mer where each protomer interfaces with three protomers with buried surface areas of 1587 Å2, 641 Å2, and 285 Å2 (PDB ID: 4PT2). Though formation of a ChdC 60-mer seen by crystallographic symmetry seems physiologically possible, it remains to be seen whether this higher ordered structure is biologically meaningful for ChdC function. The function of the loop in domain II, namely residues 112-121, has been mainly described in the context of ChdC coproheme decarboxylase activity (138, 238, 246, 247). The loop flexibility has been postulated to serve as a substrate access channel to accommodate substrate binding and product release (238). The observation that the loop region has poor electron density in crystal structures of apo- and substrate bound structures and molecular dynamic simulations of LmChdC have led to the hypothesis that the loop is dynamic to facilitate its function as a substrate access channel (138, 247, 254). It has been postulated that the closure of the loop is required to reposition residues within binding pocket that form contacts with propionates and appear to play a role in oxidative decarboxylation (238). Furthermore, a ligand-free ChdC would have the loop in a more open and solvent-exposed conformer with reference to apo-GsChdC structure (PDB ID: 1T0T) (238). Overlooked in this model is that the loop in apo-GsChdC is not solvent exposed and rather forms contacts with the adjacent protomer to form a 60-mer. Comparison of   144 the loop in domain II of ChdC structures available in the PDB and SaChdC presented here support that the position of the loop is not exclusively dependent on whether a substrate molecule is bound in the active site. Rather, evidence suggests that either the formation of a higher-order 60-mer or crystal contacts with adjacent molecules dictate the structure of the loop observed in the crystal structures (138, 247). The possibility that the loop in domain II has roles in both oligomerization and as a substrate access channel should not be excluded.  Very little is known about the fate of heme following biosynthesis in bacteria. Cytoplasmic heme levels are presumably kept extremely low due to the toxic and lipophilic nature of heme. Thus, it is unlikely that newly synthesized heme is released freely into the cell, suggesting ChdC, the terminal enzyme in the heme synthesis pathway, likely has a role in intracellular heme trafficking. Though this function is overshadowed by the newly assigned heme decarboxylase role, the potential for a role in heme supply motivated us to test the ability of S. aureus ChdC to transfer heme to apo- heme-requiring proteins, SbnI, IsdG, and IsdI. ChdC transferred heme to SbnI and IsdG but not IsdI at a rate an order of magnitude greater than the rate of passive dissociation of heme from ChdC (heme off-rate), suggesting the transfer is driven by an interaction between the donor and acceptor proteins. The exact mechanisms that govern heme transfer from between IsdI, IsdG, ChdC, and SbnI remain to be elucidated. It is possible that active transfer requires induced structural changes and potentially iron-ligand switches.  The observation that ChdC only transferred heme to IsdG but not IsdI supports the previous findings that these paralogous heme-degrading enzymes have differential roles in maintenance of heme homeostasis (68, 148, 255). IsdG homologs are found in bacteria that synthesize heme, but would not necessarily acquire it from the environment, including organisms like Thermus thermophilus, an extremophile found in thermal hot springs (256). Conservation of IsdG supports   145 that idea that it degrades excess heme that has been synthesized endogenously as a way to prevent heme-associated toxicity (256).  We have shown that ChdC can transfer heme directly to IsdG; however, the structural determinants that allow ChdC heme transfer to IsdG, but not IsdI, require further research. The differential roles of IsdG and IsdI may allow S. aureus to maintain intracellular heme homeostasis in response to a gradient of iron and heme concentrations encountered during infection. Based on in vitro heme transfer data, a model of intracellular heme transfer in S. aureus has been created (Figure 5-10). Overall, the model suggests that if heme is present, either from endogenous or exogenous sources, SbnI can accept it and impart negative regulation on the production of SB. The model also supports that under conditions that are conducive to heme biosynthesis, IsdG can serve as a safeguard to degrade excess heme through interaction with ChdC. Moreover, the model could benefit from in vitro and in vivo testing of the protein-protein interactions predicted to facilitate heme-trafficking in the cell. Alternatively, in silico methods could be used to probe the mechanism of heme transfer and potential protein-protein interactions. Future work to expand the model to include enzymes that use heme as a cofactor would help create a more complete picture of intracellular heme transfer.       146   Figure 5-10 Model of how ChdC, SbnI, IsdI, and IsdG participate in heme trafficking in S. aureus.  Arrows indicate the directionality of heme transfer between heme-binding proteins and the initial rate of heme transfer measured is indicated. In brackets is the heme off-rate measured for the above protein. This model is based exclusively on in vitro data.   Data suggestive that ChdC can form a 60-mer could be important for the role of this protein in heme homeostasis and supply. Many factors can play into the oligomeric state of a protein where different degrees of protein symmetry can have important biological and functional roles (257, 258). Ultimately, the role of ChdC 60-mer observed by crystallographic symmetry is unknown. One hypothesis is that oligomerization serves as a way to “store” enzymatic potential for when heme biosynthesis is needed as ChdC catalysis is relatively slow (kcat/KM = 1.8 x 10 M-1 s-1) (138). Formation of a 60-mer could improve stability and conserve space in the cell as larger protein complexes have decreased surface area to volume ratios which can be protective against denaturation and also decreases the amount of solvent required to hydrate proteins (257, 259).  An alternative hypothesis for this finding is that it could function as a heme storage complex, effectively storing 60 heme molecules in the active sites of the 60 protomers. The 60-mer structure I obtained was co-crystallized with heme, but it is unknown where the heme is bound in this crystal structure. Further optimization and a higher resolution data set of the crystal structure could discern if heme is indeed bound to the active site. Though the GsChdC 60-mer crystal   147 structure was not heme-bound, a PEG330 was modelled in the active site potentially stabilizing the formation of the higher oligomeric state (PDB ID: 1T0T). An attractive rationale for formation of the 60-mer is that it may allow for cooperative interaction between protomers and allosteric regulation, where formation of the complex changes affinity for substrates. Future studies to investigate ChdC protein dynamics in vitro and in vivo may answer questions regarding whether the 60-mer is a biologically relevant oligomeric state.    148 Chapter 6: Conclusion  Bioavailable iron is maintained at remarkably low concentrations in the mammalian host to effectively stifle microbial growth. However, iron depletion in host tissues can also serve as an environmental cue to alert S. aureus when it has entered a hostile host environment. In response, S. aureus switches to a more pathogenic phenotype by concomitantly upregulating the expression of iron-uptake systems and other virulence factors (42, 43). S. aureus growth and survival during infection is contingent upon exploiting the host iron pool. The multiplicity and functional redundancy of iron uptake systems S. aureus possesses is likely a reflection of the diverse host environments that S. aureus can infect, underlying its versatility as an opportunistic pathogen. Evidence that distinct iron uptake systems are more important in certain models of infections and colonization of specific organs suggests iron uptake is an important facet of host adaptation (50, 78, 127, 159). However, the spatiotemporal regulatory mechanisms by which S. aureus adapts to changing iron availability over the course of infection are ill-defined. Additionally, the intracellular network of heme-sensing and iron-binding proteins that are required for regulation and adaptation to the dynamic host environment, beyond Fur regulation, are largely unknown in S. aureus.  The work presented in this thesis contributes to our understanding of S. aureus adaptation of iron-uptake strategies to suit diverse host niches. Study of the structure and function of SbnI revealed that it has a bifunctional role in SB biosynthesis. OPS production from L-serine by SbnI provides a direct mechanism for adaptation of SB biosynthesis in environments where S. aureus catabolism must rely on amino acids, as in abscesses. Additionally, discovery of heme relay from IsdI to SbnI provides a mechanism by which S. aureus to senses heme to control iron-source   149 preference. Furthermore, study of heme transfer reactions between SbnI and proteins required for heme degradation and heme biosynthesis provide the framework for a model of hierarchical intracellular heme transfer in S. aureus.   6.1 SbnI provides an alternate route for OPS synthesis in S. aureus The role of SbnI in SB biosynthesis remained elusive as the enzymatic capacity required for precursor synthesis, production, and export of SB can be completed by the enzymes encoded by sbnA-H. Furthermore, the four precursors required for SB assembly, acetyl-CoA, oxaloacetate, OPS, and L-glutamate, can be generated by S. aureus from two of the four most abundantly available nutrients in human serum, glucose and L-glutamine (92, 102). Excluded from this model, however, is how S. aureus is able to produce SB in other host niches where nutrients like glucose and oxygen are limiting and resultantly alter central metabolism.  As a mechanism for persistence in host tissues S. aureus employs virulence factors to promote abscess formation that remodels its extracellular host environment (260). Abscess formation is a default, natural immune response to contain tissue trauma, but S. aureus can co-opt this response to enhance replication and dissemination. Within fibrin-walled abscess microenvironments, glucose and oxygen levels are depleted due to phagocytic activity (216, 217, 261). S. aureus can survive in this iron-restricted, glucose-limited, and hypoxic environment through catabolism of secondary carbon sources, including lactate, peptides, or amino acids, the major carbon sources available at the centre of abscesses (212).  As extracellular nutrient status changes subsequently drive alterations in S. aureus metabolism, the question of how S. aureus continues to support iron uptake strategies remains. Amino acid catabolic pathways and genes responsible for SB biosynthesis are upregulated in the   150 abscess environment (26, 262, 263). Additionally, a SB deficient mutant strain of S. aureus is attenuated in murine kidney abscess model (50). The unforeseen function of SbnI as a free serine kinase for OPS synthesis discussed in Chapter 3 offers a way to support SB biosynthesis in an environment where glucose is not available. SbnI serine kinase activity supports a model where SB can alternatively be made from strictly amino acid precursor molecules independent of glycolysis and serine biosynthesis. Overall, S. aureus appears to have evolved strategies to survive low iron stress and maintain SB production as infection progresses, enabling adaptation to changing extracellular environments. These findings further our understanding of how the metabolic flexibility of S. aureus can help tilt the host-pathogen axis in its favor.   6.2 Defining the function of SbnI is critical to understanding the spatiotemporal production of SB Many bacteria rely of siderophore systems to capture extracellular iron to fulfill nutritional iron needs. In S. aureus, iron deprivation results in derepression of staphyloferrin production through the global regulatory protein Fur. Examples are known in other bacteria of regulators of siderophore production, in addition to Fur, that are directly responsive to iron or an iron-siderophore complex (264–266). Regulation of siderophore iron acquisition pathways through heme-sensing is not common but was reported in the cyanobacterium Anabaena sp. PCC 7120, where the global iron-responsive regulator FurA is both heme and iron regulated (267). Iron binding by FurA promotes DNA interaction leading to transcriptional repression, typical of proteins belonging to the Fur family. However, heme binding by FurA relieves transcriptional repression and allows for expression of the regulon, including expression of the siderophore outer membrane transporter. SbnI shares similarities with FurA in that heme binding abrogates   151 interaction with DNA (267). However, unlike the global regulatory role of FurA, SbnI is thought to exclusively regulate sbn gene expression (98). SbnI establishes a key connection between staphyloferrin production and heme uptake and thus functions at the interface between two iron acquisition systems important for S. aureus pathogenesis.  Structural study of full-length SpSbnI aimed to add to our understanding of the function of SbnI as a heme-sensitive regulator of SB production. The results discussed in Chapter 4 disclosed that SpSbnI is dimeric under reducing conditions and provides rationale for the observed defects in oligomerization, L-serine kinase activity, and heme-binding observed in the S. aureus C-terminal truncated SbnI1-240. The results of this study suggest that SbnI has two functions in SB biosynthesis, transcriptional control of the sbn locus and L-serine kinase activity. Both activities are affected by heme-binding; however, heme altered transcriptional control to a greater extent. Thus, SbnI mediates the S. aureus iron source preference switch between heme and siderophore acquired iron by contributing precursor OPS to SB synthesis when heme-iron is not available and directly sensing heme to shut off SB synthesis when heme-iron is present. S. aureus transcriptome studies confirm expression of sbn locus in certain models of infection (24, 26, 262, 263) but it is difficult to discern the contribution of each function afforded by SbnI. The hypothesis that SbnI kinase activity and regulatory function are independent of each other requires further in vivo experiments. One approach to further understand the bifunctional role of SbnI would be to measure sbn gene transcription and SB production by S. aureus sbnI mutant strains complemented with an SbnI variants. SbnI variants tested could include mutants defective in either heme-binding or kinase-activity. To determine how available nutrient iron sources affect SbnI function, these strains could be grown in conditions where either ferric or heme iron   152 are given as the sole source of iron. In vivo study could also be used to validate SbnI as a critical determinant of adaptation of SB biosynthesis to glucose-deplete host environments.  A limitation of the results presented in Chapter 4 is the absence of an accurate structural model for SbnI or SpSbnI in complex with heme. As such, a complete mechanism for how heme binding alters SB production remains to be elucidated. SbnI bound to heme did not yield crystals using standard crystallization screens. Protein rigidity and stability favor crystal formation and heme-binding by SbnI may increase protein dynamics and flexibility, which would decrease the ordered protein-protein contacts that are prerequisite for crystallization (268). Heme-induced changes in the dynamics of tertiary or quaternary structure of SbnI may alter interaction with DNA or a protein partner thereby controlling transcription from the sbn locus. Alternative strategies such as small angle X-ray scattering or NMR spectroscopy can be employed to investigate quaternary structure and dynamics, respectively, of a SbnI in response to heme binding. However, the proneness of SbnI constructs to aggregate and molecular mass of dimeric SbnI render it poorly suitable for these techniques. Future efforts include investigation of SbnI homologs from different staphylococci or to leverage construct design to obtain protein samples suitable for a heme-bound crystal structure, NMR, or small angle X-ray scattering. This approach would reveal residues required for heme binding and associated conformational changes that allow SbnI to regulate transcription of sbn locus.  Interestingly no prototypical DNA-binding motif was identified in the SbnI structures. A direction that can be further explored is how SbnI directly imparts transcriptional regulatory control on the sbn locus and whether it is through specific interaction with a nucleic acid sequence (eg. dsDNA, ssDNA, or RNA), a protein partner, or if downstream intermediates in SB biosynthesis resulting from SbnI-generated OPS impact regulation. Convincing genetic evidence   153 that SbnI is required for expression of sbnD-H and that SB production is decreased during the initial stages of growth in sbnI deletion strain compared to wildtype S. aureus has been published (98). Nevertheless, how SbnI accomplishes this regulatory control remains unclear.  The observation that SpSbnI crystallized as a dimer of dimers through intermolecular disulfide bond formation suggests the redox status of the cell may impact SbnI oligomerization state and function. The role of reversible disulfide bond formation in SbnI and the effect of oxidative stress on SbnI-dependent regulation of sbn gene transcription and kinase activity offer new areas for exploration of function. As S. aureus encounters a gradient of iron concentrations during infection, it must also contend with oxidative stress imposed by the host as an immune defense (113). The detrimental effects of oxidative stress and excess iron may warrant redox-sensitive regulation of siderophore biosynthesis to avoid iron accumulation. Given the close relationship between oxidative stress and intracellular iron status in S. aureus (109, 110, 116), further studies to discern whether SbnI can integrate heme status and the redox state of the cell to affect SB biosynthesis will contribute to our understanding of the regulatory potential of SbnI.  6.3 ChdC participates in an intracellular heme transfer network  Due to the multifaceted lifestyle of S. aureus, critical to pathogenesis is an ability to quickly respond to the dynamic environment in the host. S. aureus possesses complex global regulatory networks to sense diverse signals and allow for a rapid response and fast metabolic adaptation (12). As heme can serve as a nutritional iron source, enzymatic co-factor, and a signalling molecule, we hypothesized sensing and regulatory pathways exist to connect heme synthesis with heme availability, hemoprotein abundance, and heme degradation. Moreover, how intracellular heme is transported from heme-biosynthetic enzymes to heme-requiring cellular   154 proteins is unclear. In Chapter 5, evidence for intracellular heme transfer reactions between ChdC, IsdI, IsdG, and SbnI were presented. While the experiments were carried out in vitro, they generate testable hypotheses for how the system may operate in vivo during S. aureus infection. A metabolomics approach could be taken to examine if S. aureus uses heme availability to regulate other metabolic decisions made over the course of infection (254). Mass spectrometry-based targeted metabolic profiling techniques have been used to compare the metabolic response of S. aureus to different antibiotics (269) and adaptation to glucose starvation (270). A similar method could be used to compare the metabolic changes that occur when S. aureus is grown on heme or non-heme iron sources. Alternatively, NMR methods to trace metabolic pathways and fluxes in response to different iron source availability could be used (271). NMR profiling of S. aureus metabolomic changes in response to iron-limitation with and without aeration has been previously explored (85). Data indicating that ChdC can form a 60-mer was discussed in Chapter 5. Changes in ChdC oligomerization state may have a regulatory role in relation to heme synthesis and heme-trafficking to supply hemoproteins with heme. Methods to test if the ChdC pentamer or 60-mer is the functional unit required for heme biosynthesis or transfer could be accomplished by mutating residues in the domain II loop region or designing a protein construct that has the loop entirely removed. Light-scattering techniques like DLS, size exclusion chromatography multi-angle light scattering or small-angle X-ray scattering could be used to measure solution-state oligomerization of apo- and holo-ChdC and ChdC variants. Furthermore, the catalytic and heme transfer competencies of these constructs could be tested.  Examples of oligomerization to form a nanocage as a means of controlling activity has been reported for encapsulin from M. xanthus. Encapsulin production is suggested to be regulated   155 post-translationally at the level of assembly to sequester iron and protect the cells from oxidative stress at different stages of growth (253). Structural analysis of the ChdC 60-mer revealed that formation of the higher order oligomer encloses the active sites, which could protect any bound heme from participating in reactions that generate ROS (126). Future studies to test if ChdC serves an antioxidant role in S. aureus as a heme storage complex could also be pursued. Ultimately, whether ChdC 60-mer assembly has a role in heme homeostasis and supply remains to be elucidated.   6.4 Concluding remarks Heme and iron are vital for bacterial growth and survival but are toxic at high concentrations. Thus, heme and iron homeostasis must be stringently regulated in bacteria as malfunctions result in growth defects and cell death (154). As such, many points of iron metabolism have been investigated as therapeutic targets for control of bacterial infections. These include targeting siderophore biosynthetic enzymes (272).  Recently, two antibiotics, baulamycins A and B, were reported to inhibit SB synthesis (273). Another promising approach is to exploit siderophore importers to bring in naturally occurring antimicrobials that mimic a siderophore, like albomycins, or rationally designed siderophore-drug conjugates (274–277). Heme uptake pathways have also been exploited in the same way to shuttle porphyrin-coupled antibiotics into the cell, such as the targeting the hemoglobin receptor of Porphyromonas gingivalis with deuteroporphyrin-metronidazole (278–280). Heme oxygenases are also potential targets as inhibiting their activity leads to increased intracellular heme concentrations. In silico drug design has aided in the development of molecules that can bind and inhibit the heme oxygenase, HemO,   156 from P. aeruginosa (281). These compounds inhibited the growth of P. aeruginosa when given hemoglobin as the sole source of iron (281). Heme-sensing proteins are proposed to be promising antibacterial targets because they are involved in the regulation of intracellular heme levels by influencing different processes (282). The role of heme-sensing by SbnI to direct this preference and contributions to SB precursor biosynthesis implicate it as an important element in mediating S. aureus iron and heme homeostasis. Yet, more research is required to fully understand SbnI function and the necessity of this heme-sensing protein during S. aureus infection. Iron is essential for microbial growth so targeting iron-related processes represents an interesting approach for antimicrobial drugs. However, effective clinical application of therapeutics targeting iron transport and metabolism requires a clear understanding of their functional roles and regulation in different tissues and infections.   157 References  1.  Jones, R. N. 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