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Characterisation of substrate preference in staphylococcus aureus siderophore biosynthesis Kolesnikov, Maxim 2018

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  CHARACTERISATION OF SUBSTRATE PREFERENCE IN STAPHYLOCOCCUS AUREUS SIDEROPHORE BIOSYNTHESIS  by  Maxim Kolesnikov   B.Sc., The University of British Columbia, 2015  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Microbiology and Immunology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  September 2018  © Maxim Kolesnikov, 2018 ii   The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, a thesis/dissertation entitled:  CHARACTERISATION OF SUBSTRATE PREFERENCE IN STAPHYLOCOCCUS AUREUS SIDEROPHORE BIOSYNTHESIS  submitted by Maxim Kolesnikov  in partial fulfillment of the requirements for the degree of Master of Science in The Faculty of Graduate and Postdoctoral Studies   Examining Committee: Dr. Michael Murphy, Microbiology and Immunology Supervisor  Dr. Lindsay Eltis, Microbiology and Immunology Supervisory Committee Member  Dr. Lawrence McIntosh, Biochemistry Additional Examiner     Additional Supervisory Committee Members: Dr. Pauline Johnson, Microbiology and Immunology Supervisory Committee Member    iii   ABSTRACT Staphylococcus aureus is a common opportunistic pathogen and commensal resident of a majority proportion of the adult population. Emerging drug-resistant and hypervirulent strains such as methicillin-resistant S. aureus (MRSA) have reduced the efficacy of existing treatment options. Iron acquisition from the host is required for the establishment of infection. S. aureus possesses several mechanisms for iron acquisition, including the ferric-iron binding siderophores staphyloferrin A (SA) and staphyloferrin B (SB).    To explore the substrate preference of the SA and SB biosynthetic enzymes, crystal structures of biosynthetic enzymes were solved, and alternative substrates were tested. Crystal structures of the synthetases SfaD and SbnF were solved and compared to homologs from other species to define structural determinants of substrate preference. An analogue of SA, substituting D-lysine for D-ornithine during synthesis was produced in vitro and characterized using liquid chromatography and mass spectrometry. Furthermore, S. aureus was shown to be able to use this SA analogue for iron acquisition. Analogues of intermediates in the SB biosynthesis pathways were produced in vitro. The biosynthesis of a functional S. aureus siderophore analogue provided insights into the structures and substrate specificities of siderophore synthesis proteins. The modified siderophores may be of use to deliver antimicrobials into the cell or as a diagnostic for S. aureus infection.    iv   LAY SUMMARY Staphylococcus aureus is a common bacterial pathogen that causes severe diseases across the globe. Iron is an important metal that is needed by bacteria for growth. To infect the human body, S. aureus must steal iron from the host. To acquire iron S. aureus secretes small molecules called siderophores into the environment to find and bind iron. These are eventually reabsorbed by the bacterium. This work characterized the machinery that builds these siderophores in S. aureus and examined how it can be manipulated to produce modified products, including a new siderophore. Making modifications to these siderophores could be used to attach antibiotics as a new therapy or label to create diagnostic tools. If developed, this work may help design new compounds to fight S. aureus infections.     v   PREFACE  The work in this thesis involved contributions by several fellow scientists from the laboratories of Dr. Michael Murphy and Dr. Lindsay Eltis. Development and optimization of the HPLC detection assay and siderophore utilization bioassay was a collaborative effort between Dr. Jason Grigg, Mariko Ikehata and myself. Adam Crowe and Eugene Kuatsjah of the Eltis lab provided assistance with MS analysis of siderophore samples.    vi   TABLE OF CONTENTS ABSTRACT…………………………………………………………………………………….. iii LAY SUMMARY……………………………………………………………………….………. iv  PREFACE…………………………………………………………………………….………….. v TABLE OF CONTENTS………………………………………………………………………. vi LIST OF TABLES……………………………………………………………………………… ix LIST OF FIGURES……………………………………………………………………………... x LIST OF ABBREVIATIONS…………………………………………………………………. xii ACKNOWLEDGEMENTS…………………………………………………………………... xiv Chapter 1 Introduction…………………………………………………………………………. 1   1.1: Staphylococcus aureus pathogenesis………………………………………………… 1  1.2: Iron acquisition by Staphylococcus aureus………………………………………….. 3  1.2.1: Iron uptake strategies of S. aureus……………………………………………… 3   1.2.2: Siderophore-mediated iron acquisition…………………………………………. 4  1.2.3: Transport of SA and SB………………………………………………………… 7  1.2.4: Transport of xenosiderophores………………………………………………….. 8   1.2.5: Siderophore synthesis…………………………………………………………… 9   1.2.6: SA and SB synthesis pathways………………………………………………... 10  1.3: Applications of modified siderophores……………………………………………... 12   1.4: Objectives…………………………………………………………………………... 14  Chapter 2 Methods…………………………………………………………………………….. 15  2.1: Bacterial strains and growth conditions…………………………………………….. 15  2.2: Site-directed mutagenesis…………………………………………………………... 15 vii    2.3: Protein expression and purification………………………………………………… 16   2.4: NIS synthetase pyrophosphate release assay……………………………………….. 17   2.5: CO2 release assay of SbnH…………………………………………………………. 18  2.6: Synthesis of SB intermediates, SA and D-Lys-SA…………………………………. 19  2.7: SbnF structure determination……………………………………………………….. 20  2.8: SfaD structure determination……………………………………………………….. 23  2.9: HPLC analysis of SA and alternative SfaB products………………………………..25  2.10: Mass Spectrometry of SA and D-Lys-SA…………………………………………. 25  2.11: Bioassay of SA and D-Lys-SA samples…………………………………………… 25  Chapter 3 Results………………………………………………………………………………. 27  3.1: Characterization of the structure and substrate preference of SA pathway NIS synthetases………………………………………………………………………….. 27 3.1.1: Structure of SfaD………………………………………………………………. 27 3.1.2: Active site mutants of SfaD…………………………………………………… 29 3.1.3: Substrate preference of SfaD…………………………………………………...31 3.1.4: Substrate preference of SfaB…………………………………………………... 33 3.1.5: Validation of D-Lys-SA……………………………………………………….. 35  3.2: Characterization of the structure and substrate preference of SB pathway enzymes. 37  3.2.1: Structure of SbnF……………………………………………………………… 37  3.2.2: Active site mutants of SbnF…………………………………………………… 40  3.2.3: Substrate preference of SbnE………………………………………………….. 41  3.2.4: Substrate preference of SbnH………………………………………………….. 43  3.2.5: Substrate preference of SbnF………………………………………………….. 45 viii   Chapter 4: Discussion………………………………………………………………………….. 48  4.1: Substrate preference of SA pathway enzymes……………………………………… 48  4.1.1: Substrate preference of SfaD…………………………………………………...48   4.1.2: Production of a SA analogue…………………………………………………... 53  4.2: The SB pathway…………………………………………………………………….. 55  4.2.1: Substrate preference of SB pathway enzymes………………………………… 55   4.3: Conclusions…………………………………………………………………………. 59   4.4: Future directions……………………………………………………………………. 60  BIBLIOGRAPHY……………………………………………………………………………… 62  ix   LIST OF TABLES  Table 2.1: Bacterial strains used in this study……………………………………………………15  Table 2.2: List of plasmids used in this study…………………………………………………… 16 Table 2.3: List of primers used in this study…………………………………………………….. 16 Table 2.4: Data collection and refinement statistics for SbnF…………………………………... 22  Table 2.5: Data collection and refinement statistics for SfaD…………………………………... 24 Table 3.1: Structural similarity of SfaD to NIS synthetases…………………………………….. 28 Table 3.2: Specific activity of SfaD mutants……………………………………………………. 31  Table 3.3: Specific activity of SfaD……………………………………………………………... 33 Table 3.4: Specific activity of SbnE…………………………………………………………….. 43 Table 3.5: Specific activity of SbnF…………………………………………………………….. 48   x   LIST OF FIGURES  Figure 1.1: Schematic of siderophores, highlighting different types of iron-coordinating groups..5 Figure 1.2: SA (A) and SB (B) coordinate ferric iron in a hexadentate manner…………………. 7 Figure 1.3: Crystal structures of HtsA (A) and SirA (B) bound to SA and SB, respectively, show positively-charged substrate-binding pockets…………………………………………….. 8  Figure 1.4: Schematic of a generic condensation reaction catalysed by NIS synthetases………. 10  Figure 1.5: Biosynthetic pathway of SA……………………………………………………….... 11  Figure 1.6: Biosynthetic pathway of SB………………………………………………………… 12 Figure 3.1: Structure of SfaD……………………………………………………………………. 28 Figure 3.2: Conserved ATP binding site of SfaD compared to AcsD…………………………... 29  Figure 3.3: Pyrophosphate assay of SfaD and its active site mutants…………………………… 30 Figure 3.4: Activity of SfaD with D-ornithine and its analogues……………………………….. 32  Figure 3.5: Activity of SfaD with citrate and its analogues…………………………………….. 32  Figure 3.6: Detection of SA by HPLC…………………………………………………………... 34 Figure 3.7: D-Lys-SA detection by HPLC………………………………………………………. 35 Figure 3.8: Predicted structure and iron-coordination of D-Lys-SA……………………………. 36 Figure 3.9: MS analysis of SA and D-Lys-SA reaction mixtures……………………………….. 36  Figure 3.10: Bioassay analysis of D-Lys-SA samples…………………………………………... 37 Figure 3.11: Structure of SbnF…………………………………………………………………...38 Figure 3.12: The structure of SbnF contains two citrates bound in the predicted active site…… 39 Figure 3.13: Citrate bound in a predicted substrate-binding site in SbnF………………………. 40 Figure 3.14: Pyrophosphate assay of SbnF active site mutants…………………………………. 41  Figure 3.15: Activity of SbnE with the substrate citrate and analogues………………………… 42  xi   Figure 3.16: Activity of SbnE with the substrate L-DAP and analogues……………………….. 42 Figure 3.17: Possible products of SbnE citrate substitutions…………………………………….44 Figure 3.18: Activity of SbnH with citryl-L-2,3-diaminopropionate and analogues……………. 45 Figure 3.19: Possible substrates generated for SbnF from reactions with SbnE and SbnH…….. 46 Figure 3.20: Activity of SbnF with the substrate citryl-diaminoethane and analogues…………. 47 Figure 4.1: Progression of citric acid substitutions through the SA synthesis pathway………… 50  Figure 4.2: Progression of D-ornithine substitutes through the SA pathway…………………….51 Figure 4.3: Functional nucleophile substitutions in SfaD………………………………………..52 Figure 4.4 Comparison of D-ornithine and putrescine structures……………………………….. 52  Figure 4.5: Functional citrate substitutions in SfaD…………………………………………….. 53 Figure 4.6: Citric acid substitute progression through the SB pathway………………………… 56  Figure 4.7: Key catalytic residues in SfaD and SbnF…………………………………………… 58        xii   LIST OF ABBREVIATIONS  AMP    Adenosine monophosphate  ATP    Adenosine triphosphate CA-MRSA   Community-associated MRSA Can    Collagen adhesin C-DAE   Citryl-diaminoethane ClfA    Clumping factor A EHEC    Uropathogenic Escherichia coli FnBPA   Fibronectin-binding protein A Fur    Ferric uptake regulator HPLC    High-pressure liquid chromatography IPTG    Isopropyl β-D-thiogalactopyranoside Isd    Iron-surface-determinant Kd    Dissociation constant  LB    Luria-Bertani broth  L-DAP    L-2,3-diaminopropionic acid MALDI-MS   Matrix-assisted laser desorption/ionization mass spectroscopy  MESG    2-amino-6-mercapto-7-methylpurine ribonucleoside MRSA    Methicillin-resistant Staphylococcus aureus MS    Mass spectrometry  NEAT    Near iron transporter  NIS    NRPS-independent siderophore NRPS    Nonribosomal peptide synthetases xiii   PDB    Protein data bank PLP    Pyridoxal 5’-phosphate PNP    purine nucleoside phosphorylase PPi    Pyrophosphate  PVL    Panton-Valentine Leucocidin  SA    Staphyloferrin A SB    Staphyloferrin B  SCCmec   Staphylococcal cassette chromosome mec SD    Standard deviation SeMet    Selenomethionine  TMS    Tris minimal succinate TSST    Toxic shock syndrome toxin  WT    Wild-type   xiv   ACKNOWLEDGEMENTS   I would like to express my deepest gratitude to Dr. Michael Murphy for taking a chance on an undergraduate student with no prior experience and providing invaluable guidance, support and advice over the years I have been fortunate enough to be a member of his lab.   My deepest thanks go to my committee members Dr. Lindsay Eltis and Dr. Pauline Johnson for all the feedback on my project and their tolerance of all my foibles. Additionally, I would like to thank Dr. Johnson for giving me the chance to collaborate on a fascinating project that is not a part of this thesis.    Particular thanks go to the people who contributed directly to the work included in this thesis: Dr. Jason Grigg, Angelé Arrieta, Mariko Ikehata, Adam Crowe and Eugene Kuatsjah. Thank you all for making this project possible.   I would like to acknowledge all the past and present members of the Murphy lab who have been an inexhaustible well of support, great ideas, and help. Special thanks to Angelé, thank you for being the keystone that keeps us all going. To Angela, you were the best road-trip buddy and confidant I could have ever wished for. To Emily, for always making me laugh more than I should probably have. To Anson, for all those endless MTG nights. And finally, to all of the undergraduates that have worked, are working and will work to keep all of the small, thankless, things in order so that we may go on about our work without a worry in our minds.   Thanks to my close friends for making my life more colourful than I could ever have made it myself.  To Kyle, you are probably the best friend and roommate that has ever existed. To Chloe, thank you for being the catalyst of joy and inspiration in my life. I would not be the person I am without you. To my mother, спасибо тебе за то что ты есть и за всё что ты сделала. 1   Chapter 1: Introduction  1.1  Staphylococcus aureus pathogenesis  Staphylococcus aureus is a gram-positive opportunistic pathogen, frequently found as a human commensal in the general population [1]. S. aureus colonizes nearly a third of the adult population, residing commonly on the skin in locations such as the nasal cavity, underarms, groin, and forearm [2, 3]. The predominant site of a persistent S. aureus population is in the nasal cavity, with researchers linking nasal colonization to medical complications as early as the 1930s [4, 5]. Although often asymptomatic, S. aureus is currently known to cause a wide variety of infectious diseases including bacteremia, toxic shock syndrome, infective endocarditis as well as multiple skin and soft tissue infections [6]. Moreover, it is one of the five most commonly acquired nosocomial infections, imposing an annual economic cost on society that has been estimated to be as large as 13.8 billion dollars in the United States alone [7, 8].   Treatment of S. aureus has been exacerbated over the decades by the emergence of drug-resistant strains [9]. Early clinical use of penicillin, although effective, was limited by the rapid spread of penicillin-resistance in S. aureus, first identified in the 1940s [10, 11]. This was eventually followed by methicillin and vancomycin resistance in 1961 and 2002, respectively [12, 13]. Vancomycin resistance, in particular, has raised a great degree of concern as vancomycin treatment is currently one of the last therapeutic options available for methicillin-resistant Staphylococcus aureus (MRSA) infections [14]. Vaccine development has been unsuccessful with several trials failing at the phase II stage [15-17].  The global threat of S. aureus is most commonly associated with MRSA, the emergence of which was first described in 1961 shortly after the introduction of methicillin [11]. Globally, MRSA often represents more than half of infective S. aureus isolates, spread by existing clones  2   or horizontal transfer of the staphylococcal cassette chromosome mec (SCCmec) which confers resistance to methicillin and by extension most β-lactam antibiotics [18, 19]. Resistance to β-lactams, in conjunction with increased resistance to other antibiotics, has been exacerbated by changes in MRSA epidemiology in the 1990s which lead to infections in communities of healthy individuals with no prior hospitalization such as athletes and enlisted personnel [20, 21]. This is in contrast to MRSA infections initially being relegated to hospitals [22]. Such strains with increased virulence have now been reclassified as community-associated MRSA (CA-MRSA) and their emergence has expanded the reach of S. aureus infections and underscored the need for new treatment options [23].  The relative success of S. aureus in causing severe infections can be attributed to its wide range of virulence factors that provide mechanisms for evading the immune response from the host as well as factors for nutrient acquisition [24]. The presence and specificity of virulence factors vary between strains as many virulence genes are found on mobile genetic elements [25]. In general, S. aureus gains access to sterile tissues via open wounds and uses adhesin proteins such as clumping factor A (ClfA), collagen adhesin (Can), and fibronectin-binding protein A (FnBPA) to anchor itself in the host [26-29]. Secreted virulence factors such as toxins are regulated by the global virulence regulator Agr, for example, alpha-toxin and Panton-Valentine Leucocidin (PVL) which lyse red blood cells, leukocytes and damage surrounding tissue [30-34]. Lysis of the surrounding tissue provides access to essential nutrients, in particular, transition metals, such as iron. Additional toxins, such as the toxic shock syndrome toxin (TSST), which stimulates the release of IL-1, IL-2, TNF-α, and other cytokines, act to evade and overwhelm the immune system [32, 35]. S. aureus resistance to oxidative stress is provided by the production of the enzymes catalase and superoxide dismutase, which counteract hydrogen peroxide and  3   superoxide produced by leukocyte activity [36-42]. These factors contribute to the high persistence of S. aureus, with studies showing its ability to survive and proliferate within host macrophages and epithelial cells [43-45].   1.2 Iron acquisition by Staphylococcus aureus 1.2.1: Iron uptake strategies of S. aureus   Transition metals are essential nutrients to nearly all forms of life due to their use as enzymatic cofactors in key intracellular processes [46]. Iron, in particular, is a crucial component of numerous systems such as in respiration where iron-bound haem is incorporated into cytochromes for electron transport [47]. A strategy commonly used by humans and other vertebrates to prevent infection is the restriction of these nutrients from pathogens. The various mechanisms which sequester Fe, Mn, Zn, and other essential nutrients are collectively termed nutritional immunity and form a defence against microbial proliferation [48-51]. Within the human host, over 90% of iron is located intracellularly [52, 53]. The most abundant carrier of iron is the haemoprotein haemoglobin found within erythrocytes [53, 54]. Iron located in serum is bound by the proteins transferrin and lactoferrin, while additional mechanisms provide regulated circulation and recycling of iron within the host [51, 55-57]. S. aureus has evolved several distinct and redundant iron-uptake systems to overcome these limitations.    When presented with an iron-limited environment, S. aureus alters its protein expression profile via the ferric uptake regulator (Fur) [58, 59]. In the presence of iron, iron-bound Fur binds to the Fur box, a short segment of DNA with a defined consensus sequence, and represses downstream genes [58, 59]. When iron is limiting, apo-Fur dissociates from DNA, allowing gene  4   expression [58, 59]. The Fur regulon contains several gene loci encoding for iron and heme uptake systems.   As the most abundant source of iron in the human body, haem is a target of the S. aureus iron-surface-determinant (Isd) system [60-65]. This system contains several cell-wall anchored receptors as well as a membrane-bound transporter and intracellular proteins. Haemoglobin and haemoglobin-haptoglobin complexes are bound by receptors containing NEAr iron Transporter (NEAT) domains, which extract haem and transfer it via other proteins to the IsdEF ABC transporter [62, 66-68]. Intracellular haem is subsequently degraded, providing iron to the bacterium [62]. The acquisition of haem from haemoglobin is a crucial virulence factor that is utilized in concert with haemolytic toxins secreted by S. aureus during infection [69]. During colonization and stages of infection where haem is not available, such as survival within macrophage phagolysosomes, iron may be acquired instead via the use of siderophores, small molecular weight ferric-iron binding molecules secreted by many bacterial species [69].     1.2.2: Siderophore-mediated iron acquisition   Siderophores are a diverse family of ferric-iron chelators, produced and secreted into the environment by numerous microbial species [70-72]. The affinity of these small (100-1000 Da) molecules for iron has been estimated to be in the range of 1012-1052 M [71, 73]. To date, over 500 individual siderophores have been characterized in bacteria, fungi and certain plants [70-72, 74]. Consequently, siderophore-mediated iron acquisition is commonly a key virulence factor for many human pathogens, including Mycobacterium tuberculosis and Pseudomonas aeruginosa [75]. Siderophores are generally classified into three types based on the chemical groups that coordinate the Fe3+ atom: catecholates, hydroxamates, and carboxylates (Figure 1.1), additionally  5   mixed-type siderophores containing more than one type of chemical group have been characterized [76-78]. S. aureus produces two chemically distinct hydroxamate type siderophores: staphyloferrin A (SA) and staphyloferrin B (SB) [69].    Figure 1.1 Schematic of siderophores, highlighting different types of iron-coordinating groups.  Enterobactin (A), produced by Escherichia coli utilizes catecholate functional groups. Ferrioxamine (B) and pyochelin (C) represent examples of siderophores containing hydroxamate and mixed-type coordinating groups, respectively. Iron-coordinating groups are labelled in red.   SA is a carboxylate-type siderophore that is comprised of two citrate molecules joined by a D-ornithine backbone (Figure 1.2A) and is produced by a variety of staphylococci, including S. aureus and S. epidermidis [79, 80]. In S. aureus, SA biosynthesis and export is managed by products of the sfaABCD operon which is Fur regulated via an intergenic Fur box [81-83]. Assembly of SA has been shown to require a functioning TCA cycle as a source of citrate and is  6   therefore linked to environmental factors that may disrupt TCA cycle function in S. aureus, such as high glucose concentrations [82, 84]. As glucose is the second most abundant metabolite in human serum, the role of SA during infection is limited to solid tissue. Recent studies have identified a role for this siderophore in specific environments such as in abscesses or epithelial cells [85]. Also, the full role of SA during infection is yet to be determined.  SB is also a carboxylate type siderophore, and is produced by a smaller number of staphylococcal species than SA [86]. SB is synthesized form L-2,3-diaminopropionic acid (L-DAP), citrate and α-ketoglutarate (Figure 1.2B) [87]. In S. aureus, SB is assembled and exported by products of the sbn operon [87]. Synthesis of SB is supported by additional enzymes for biosynthesis of the precursors citrate (SbnG), L-DAP and α-ketoglutarate (SbnA, SbnB and SbnI) [88-92]. Gene deletion mutants of S. aureus unable to produce SB show a significantly decreased ability to survive iron-restricted conditions and decreased virulence in mouse infection models [93]. These conclusions have been supported by gene expression data showing upregulation of the sbn operon in serum [58, 94].     7    Figure 1.2 SA (A) and SB (B) coordinate ferric iron in a hexadentate manner. Groups that participate in iron binding are coloured in red.   1.2.3: Transport of SA and SB   Ferric-iron bound SA and SB are taken up from the environment via the ABC transporters HtsABC and SirABC, respectively [83, 95]. Both receptors are specific for their respective siderophore with dissociation constants (Kd) in the sub-nanomolar range [96, 97]. However, both transporters require the function of an ATPase named fhuC, as neither operon encodes a dedicated ATPase to power transport across the membrane [98]. Crystal structures of HtsA in complex with SA and SirA in complex with SB show both siderophores bound in positively-charged pockets that undergo conformational changes upon binding (Figure 1.3) [96, 97].   8    Figure 1.3 Crystal structures of HtsA (A) and SirA (B) bound to SA and SB, respectively, show positively-charged substrate-binding pockets.  1.2.4: Transport of xenosiderophores  In addition to dedicated transport of SA and SB, S. aureus encodes a nonspecific receptor system targeting hydroxamate type siderophores produced by other bacteria for transport. This is mediated by the receptors FhuD1 and FhuD2, as well as the permease FhuBG [99, 100]. This complex is presumed to have a lower affinity for ligands, allowing it to transport a broad spectrum of xenosiderophores available in the immediate environment of the bacterium. Currently, four siderophores have been identified as transported via this system: aerobactin, coprogen, ferrichrome A and desferrioxamine B [86]. Similarly, the SstABCD system is able to transport catecholate-type siderophores such as bacillibactin and ferrated catecholamines [101]. The presence of these systems provides an advantage in complex communities and may play a role in infections with several bacterial pathogens present.       9   1.2.5: Siderophore synthesis  Despite the great diversity in their structures, most siderophores are assembled by one of two distinct methods: nonribosomal peptide synthetases (NRPS) and the NRPS-independent siderophore (NIS) synthetase pathway [102]. Most hydroxamate, catecholate, and mixed-type siderophores are assembled from amino and/or aryl acids by NRPSs, large multifunctional enzyme complexes [102, 103]. Homologous complexes synthesize peptide antibiotics in other species [104]. NRPSs consist of a single or multiple polypeptide chains that form an assembly line with different catalytic sites for siderophore production [70, 102]. Generally, an NRPS consists of an adenylation domain, peptidyl carrier protein and a condensation domain [70, 102]. Synthesis of the siderophore in these active sites is defined by the covalent attachment of intermediates to the individual domains [70, 102]. Several examples of this pathway have been characterized and the mechanisms involved described [70, 102].   The lesser studied NIS pathway synthesizes the majority of carboxylate type siderophores [102, 105, 106]. These include aerobactin and achromobactin found in various bacterial genera. This pathway consists primarily of so-called NIS-synthetases with other enzyme classes occasionally involved in siderophore assembly [102, 107]. Unlike NRPS-mediated synthesis, all NIS synthetases are free enzymes without covalent retention of intermediates.   Most of the NIS synthetases discovered to-date cluster together in a single superfamily based on sequence and structure and can be divided into categories based on their substrate specificities. Type A synthetases utilize citric acid, type B α-ketoglutarate, and type C monoamide/monoester derivatives of a citric acid molecule, often the product of a previous synthesis step [102]. A mechanism for the NIS family was proposed based on the aerobactin pathway and validated with the structure of the type A synthetase AcsD from Pectobacterium  10   chrysanthemi [105-107]. The active site in AcsD consists of an ATP-binding site that is conserved across the known NIS synthetase family, as well as the catalytic arginine R305 and ATP-coordinating histidine H444, shown to play a key role in catalysis [107]. All three classes catalyse a Mg2+-dependent condensation of a carboxylic acid with either an amine or alcohol. The carboxylate is initially adenylated by a single ATP molecule, followed by displacement of the AMP group by the nucleophilic amine or alcohol (Figure 1.4 B) [107]. Catalytic residues and the Mg2+ ion act to correctly position substrates in the active site and polarize bonds to facilitate nucleophilic attack [107].    Figure 1.4 Schematic of a generic condensation reaction catalysed by NIS synthetases.  (A) Initial adenylation of a carboxylic acid displaces a pyrophosphate group and is followed (B) by a nucleophilic attack by either an amine or alcohol. R-COO is a citric acid, α-ketoglutarate or citrate-derived molecule.   1.2.6: SA and SB synthesis pathways  S. aureus synthesizes SA and SB via two distinct NIS pathways [81, 87]. SA is assembled by products of the Fur-regulated sfa gene locus, which encodes the NIS synthetases  11   SfaD, SfaB, the putative ornithine racemase SbnC, and a membrane-embedded efflux protein, SfaA [81]. As described previously, SA consists of two citrate molecules bridged by a D-ornithine backbone. Citrate is likely diverted from the TCA cycle, as SA production shuts down when the TCA cycle is inactive, and SbnC likely provides D-ornithine via conversion from L-ornithine pools present in the cell [81, 82]. SA is synthesized by the sequential reactions of SfaD and SfaB (Figure 1.5) [81]. Both enzymes group by protein sequence with type-B NIS synthetases but are defined as type A due to their utilization of citrate [81]. SfaD activates citrate by adenylation to form a citrate-adenylate intermediate, which reacts with the δ-amine of D-ornithine. The remaining α-amine present on the intermediate acts as the nucleophile on the citrate-adenylate intermediate produced by SfaB. SA can be synthesized in vitro using SfaD, SfaB and the necessary substrates [81].    Figure 1.5 Biosynthetic pathway of SA. SA is synthesized by the sequential reactions of SfaD and SfaB from two molecules of citric acid and one molecule of D-Ornithine.    12   SB is synthesized in four steps by three NIS synthetases and a pyridoxal 5’-phosphate (PLP)-dependent decarboxylase encoded on the sbn locus (Figure 1.6) [87]. PLP-dependent enzymes exist as a Schiff base which is displaced by the substrate [108]. Synthesis begins with the type A synthetase SbnE condensing L-DAP with citrate via the β-amine, producing the first intermediate [87]. This is followed by PLP-dependent decarboxylation by SbnH of the L-DAP originating carboxyl group. Addition of another L-DAP molecule by the type C SbnF and α-ketoglutarate by the type B SbnC results in the final SB molecule. In vitro synthesis of SB has been shown using these four enzymes and the necessary substrates [87].    Figure 1.6 Biosynthetic pathway of SB. SB is synthesized by the NIS synthetases SbnE, SbnF, and SbnC as well as the decarboxylase SbnH from two molecules of L-DAP and one molecule each of citric acid and α-ketoglutarate.   1.3 Applications of modified siderophores    Modified siderophores have been proposed as a tool for both scientific and clinical purposes with extensive research into siderophore conjugation to other effector molecules [109]. Several naturally occurring examples exist such as analogues of hydroxamate type siderophores called sideromycins, produced by several species from the Streptomyces and Actinomyces genera [110, 111]. These molecules consist of siderophores or siderophore mimics conjugated to  13   antimicrobial molecules, such as albomycin where the siderophore is coupled to a nucleoside analogue via a peptide linker [109]. Efforts to produce siderophore conjugates for possible “trojan horse” applications began in earnest in the 1980s and have resulted in multiple proof-of-concept studies with in vitro activity against target species [109]. In particular, catechol conjugates to β-lactam antibiotics, such as between pyoverdine from Pseudomonas aeruginosa and ampicillin, were shown to be effective [112]. Similarly, siderophores conjugated to cationized BSA to increase binding to immune cells were used to successfully vaccinate and prevent uropathogenic Escherichia coli (EHEC) infection in mice [113]. However, high rates of resistance have been reported for these approaches and although there is currently a conjugate siderophore undergoing clinical trials, no compounds have yet made it to the clinic [109, 114].   Aside from bactericidal applications, other uses for conjugated siderophores have been proposed. A vibrioferrin conjugate to a modified fluorophore was shown to selectively target several vibrios including V. parahaemolyticus, V. cholerae and V. vulnificus in a complex bacterial community containing S. aureus and E. coli [115]. These results suggest the possibility for applications in the scientific community for rapid analysis of bacterial diversity.   Synthesis of siderophore conjugates has been limited in some cases by the difficulty of purifying or synthesizing the siderophore. No protocol exists for high-yield purification of SA or SB and while both have been chemically synthesized, yields remain a significant challenge [116, 117]. While recent progress has been made with synthesizing some modified siderophores in vitro, very little work has been done with carboxylate type siderophores and enzymes of the NIS pathway.      14   1.4  Objectives   The emergence of S. aureus as a hypervirulent and multi-drug resistant pathogen has underlined the need for novel treatment methods. Acquisition of iron from the host plays an important role during infection and can be hijacked by siderophore conjugates as shown in several other bacterial species. Additionally, new ideas for alternative uses of conjugated products present an opportunity to provide tools to the field of S. aureus research. The relative lack of research into carboxylate type siderophore conjugation with useful molecules and low yields from chemical synthesis provide a rationale for determining a method for modified siderophore production in vitro. For the first part of my research, I characterized the substrate preferences of the NIS synthetases SfaD, SbnE, and SbnF as well as the decarboxylase SbnH. This was followed by the production of a modified SA, detected and validated by a combination of HPLC and MS. Utilization of the modified siderophore by S. aureus for iron-acquisition was demonstrated by bioassay. Structures of the NIS synthetases SfaD and SbnF provided insight into the similarity of these proteins to other members of the family and were used to generate several active site mutants to confirm catalytic residues. This study has provided insights into the structure and substrate preferences of the NIS synthetase family of enzymes and created an initial protocol for the production of a modified SA.        15   Chapter 2: Methods  2.1  Bacterial strains and growth conditions  DNA was propagated using Escherichia coli strain DH5α. Proteins were expressed using E. coli strain BL-21 (DE3). Strains were grown in Luria-Bertani broth (LB). For selection and plasmid maintenance all media was supplemented with 25 μg ml-1 of kanamycin.   Table 2.1: Bacterial strains used in this study  Bacteria Strain name Description or genotype Source or reference Escherichia coli BL-21 F– ompT gal dcm lon hsdSB(rB–mB–) λ(DE3 [lacI lacUV5-T7 gene  ind1 sam7 nin5]) [malB+]K-12(λS) Novagen Escherichia coli DH5α F– endA1 glnV44 thi-1 recA1 relA1 gyrA96 deoR nupG purB 20 φ80dlacZΔM15 Δ(lacZYAargF) U169, hsdR17(rK–mK+), λ– Life Technologies Staphylococcus aureus JE-2 USA300, SCCmecIV , pvl+ University of Nebraska Medical Center  2.2 Site-directed mutagenesis   SfaD and SbnF were predicted to contain several conserved catalytic residues. To validate these predictions, whole plasmid site-directed mutagenesis was used to generate point mutations in the target amino acid sequence. SfaD R351A, H496N, SbnF R292A, and H426N were generated using the same method. Using existing protein expression constructs on the pET28A vector, complementary mutagenesis primers were designed and purchased from Integrated DNA Technologies with a 5’ phosphate tag. Amplification reactions contained Phusion polymerase (NEB), Ampligase (Epicentre) as well as sufficient nucleotides to amplify and ligate mutant plasmid. Amplicons were subsequently digested with DpnI for 3 hours at 37°C  16   to remove methylated or hemi-methylated DNA. Ten μl of digestion product was transformed into heatshocked E. coli DH5α cells. Mutations were confirmed by sequencing (Genewiz Inc).  Table 2.2: List of plasmids used in this study  Plasmid name Description Antibiotic Resistance Source or references pET28a Expression vectors carrying an N terminal His-tag/thrombin/T7 tag Kanamycin in E. coli EMD Biosciences pET28a-SbnE pET28a derivative for SbnE expression Kanamycin in E. coli [87] pET28a-SbnH pET28a derivative for SbnH expression Kanamycin in E. coli [87] pET28a-SbnF pET28a derivative for SbnF expression Kanamycin in E. coli [87] pET28a-SbnC pET28a derivative for SbnC expression Kanamycin in E. coli [87]    Table 2.3: List of primers used in this study  Description Names of Primer(s) Primer sequence (5’-3’) SfaD R351A R351A 5’mutagenesis CATATTACAGGTGAGATTGCTACATTA TCAGAACAAACG SfaD H496N H496N 5’mutagenesis GCATTGCATTAGAGGCTAACTTGCAAAA TGCTATTGC SbnF R292A R292A 5’mutagenesis GAACACTT CAACGAAAGCAGTGTTGGCG CCTCATAC SbnF H426N H426N 5’mutagenesis GGTATTGCCTTTGAATCGAATGCACAAAAT ATGATGCC  2.3 Protein expression and purification   Proteins were purified using the same protocol unless otherwise noted. One liter of LB supplemented with 25 μg/mL kanamycin was inoculated with 5 mL of an overnight culture of BL-21 cells with the desired protein encoded within a pET-28a vector. The culture was grown at 37°C for 4-6 h until an OD600 of 0.6-1. The culture was then induced by addition of 0.3 mM isopropyl β-D-thiogalactopyranoside (IPTG) and incubated for 16-20 h at 22°C with constant  17   shaking at 200 rpm. Cell pellets were collected by centrifugation at 4000 rpm for 7 minutes, resuspended in 40 mM Tris (pH 8.0), 100 mM NaCl and lysed at 10,000-13,000 psi using an EmulsiFlex-C5 homogenizer (Avestin). Supernatants containing protein were isolated by centrifugation at 18,000 rmp for 1 h. Collected supernatant was loaded on a 5 mL HisTrap HP column (GE Healthcare) and eluted with a linear imidazole gradient (0-500 mM). Collected protein was dialyzed overnight into 40mM Tris (pH 8.0) at 4°C. Dialysed fractions were further purified by anion exchange chromatography using a Source 15Q column (GE Healthcare) and eluted with a NaCl gradient (0-500 mM). Collected fractions were dialyzed overnight into 40mM Tris, pH 8.0, containing 100 mM NaCl at 4°. All proteins were concentrated to >10mg/mL, flash frozen and stored at -80°C. Selenomethionine-incorporated (SeMet) SfaD was expressed by methods previously described using a defined minimal media with supplemented selenomethionine [118]. Purification was done using the same procedure as all other proteins used in this study.   2.4  NIS synthetase pyrophosphate release assay  NIS synthetases catalyse a condensation reaction utilizing one molecule of adenosine triphosphate (ATP) and releasing one molecule each of adenosine monophosphate (AMP) and pyrophosphate (PPi). The S. aureus NIS synthetases SfaD, SbnE, and SbnF conform to this mechanism and were assayed for turnover with wild-type (WT) and alternative substrates using the EnzCheck pyrophosphate assay kit (Sigma). In the presence of inorganic phosphate, the substrate 2-amino-6-mercapto-7-methylpurine ribonucleoside (MESG) is converted to ribose 1-phosphate and 2-amino-6mercapto-7-methylpurine by purine nucleoside phosphorylase (PNP). This results in a shift of the absorbance maximum from 330 nm to 360 nm. Phosphate is  18   generated by inorganic pyrophosphatase which converts PPi to phosphate. The measured absorbance gain at 360 nm represents turnover of the enzyme.   UV-visible spectra were measured using a Varian Cary 50 UV-visible spectrophotometer. A 200 μL reaction contained 1 mM ATP, 1 mM of any additional enzyme substrate, 200 μM MESG, 10 μL 20X reaction buffer (1.0 M Tris-HCl, 20 mM MgCl2, pH 7.5, 2 mM sodium azide), 1 U PNP, 0.03 U inorganic pyrophosphatase and 10 μM NIS synthetase. Reactions were started by addition of the NIS synthetase and followed for 5-10 minutes.   Absorption data were plotted as standard deviation (SD) + error. Initial substrate concentrations were kept consistent unless derived from a mixed isomer stock, in which case all calculated rates defined as apparent. To determine reaction rates, the slope of a linear portion of the progress curve was measured for a period of one to three minutes starting at 30 seconds after the addition of enzyme. The period measured depended on the enzyme studied: SfaD 3 min., SbnE 2 min., SbnF 1 min. All rates calculated for SbnF are defined as apparent due to substrate concentration being derived from an estimate.  2.5  CO2 release assay of SbnH  SbnH catalyses the second step in the SB pathway and is a decarboxylase to produce CO2 and citryl-diaminoethane. The rate of CO2 formation by SbnH in the presence of either its WT substrate or modified products produced by SbnE was measured via a coupled enzymatic assay with phosphoenolpyruvate carboxylase (catalyses the addition of bicarbonate to phosphoenolpyruvate to form oxaloacetate) and malic dehydrogenase (catalyses the NADH-dependent reduction of oxaloacetate to form malate) [119].   Oxidation of NADH by malic dehydrogenase was measured at 340 nm for three minutes. The reactions were performed in 75 mM Tris-HCl, 25 mM NaCl, 8 mM MgSO4, pH 8.0 (Buffer  19   A).  To remove residual CO2 from the reactions, Buffer A (200 mL) was purged with N2 for 1 hour and then placed in an anaerobic glove box maintained under a positive N2 atmosphere to equilibrate for 16 hours. The powdered forms of the substrates, malic dehydrogenase, and phosphoenolpyruvate carboxylase were placed into the glove box, allowed to equilibrate overnight and were dissolved in Buffer A. SbnH was also introduced into the glove box where all the following steps were undertaken at 25 °C. Reactions were performed in a total volume of 500 μL and monitored at 340 nm for three minutes using a NanoDrop 2000c UV-visible spectrophotometer (ThermoFisher). Each reaction contained 0.2 mM NADH, 500 μM phosphoenolpyruvate, 4.1 units/mL of malate dehydrogenase, 1.0 units/mL of phosphoenolpyruvate carboxylase and an estimated 200 μM SbnH substrate.  The reactions were initiated with the addition of 10 μM SbnH. All reactions were run in triplicate.   For data analysis, SbnH enzyme activity was defined as an absorbance change after three minutes that was greater than the negative control.  2.6  Synthesis of SB intermediates, SA and D-Lys-SA  As SbnH and SbnF, the second and third enzymes in SB synthesis, respectively, use pathway intermediates which are not commercially available, a one-pot reaction mixture was adapted from in vitro synthesis of SA and SB, as described by other groups [81, 87]. This method was also used to generate SA and D-Lys-SA used for HPLC and MS analysis.  To generate substrate for SbnH, 1 mL reactions containing 1 mM citrate, 1 mM L-DAP, 3 mM ATP, 1mM MgCl2 and 10 μM SbnE were set up in water and incubated overnight at room temperature. Protein was filtered out by centrifugation using nanosep protein centrifugal filters (PALL Laboratory) at 10,000 g for 10 minutes. The resulting filtrate was collected and used for subsequent assays. Alternative products were generated by replacing either citrate or L-DAP in  20   the reaction mixture. SbnF product generation followed the same procedure but was supplemented with 10 μM SbnH.   Production of SA and testing of alternative SfaB products were set up as 1 mL reactions containing 2 mM citrate, 1 mM D-ornithine, 3 mM ATP, 1 mM MgCl2 and 10 μM of SfaD. For alternative SfaB products, D-ornithine or citrate was substituted out. The reactions were incubated overnight, and protein was removed by filtration. SfaB to 10 μM was added to continue the reaction and removed following 6-8 hours of incubation. The reaction products were processed with the same procedure as described above.   To estimate the concentration of the final product in the reaction mixtures, samples were analysed by HPLC. Reaction mixtures were analysed using an Agilent 1260 Infinity HPLC equipped with a Waters Xbridge 3.5 μm Amide column (2.1 x 100 mm). Compounds were mixed 50:50 with acetonitrile prior to injection, run in 1.25 mM ammonium acetate (pH 5.1 by glacial acetic acid) and eluted with acetonitrile. Peaks at 210 nm corresponding to ATP and AMP were identified by comparison with standards and a decrease of the ATP peak was measured. The product was estimated to be at 90% of precursor concentration.  2.7  SbnF structure determination   Initial SbnF crystals were grown by setting up the JCSG Core Suites crystallization screen (QIAGEN) using the sitting-drop vapor diffusion method at 4 °C in 2-μl drops with a 1:1 mixture of ~16.7 mg/mL SbnF and reservoir condition. Hits were identified based on visual examination under a microscope and selected for further optimization. Crystals grown for structure determination were grown with reservoir conditions of 0.18 M tri-ammonium citrate and 20% w/v PEG 3350 at 4 °C. For cryoprotection crystals were soaked for ~30 seconds in reservoir buffer supplemented with 30% (v/v) glycerol. Crystals were then flash-frozen in liquid  21   nitrogen and stored until data collection. Crystals were initially screened on a Rigaku MicroMax 007-HF generator, VariMax HR optics, and Saturn CCD 944+ detector. Diffraction data were collected at the Stanford Synchrotron Radiation Lightsource beamline 7-1. Diffraction data were processed and scaled using XDS [120]. Crystals grew in the space group P212121 with two molecules in the asymmetric unit (Table 2.4). Phase determination by molecular replacement was conducted using the NIS synthetase AsbB from Bacillus anthracis (PDB entry code 3TO3), using the AutoSol program in Phenix [121]. Model building and refinement were performed using the Phenix suite and further visualized using COOT [121, 122]. Structure figures were made using Chimera [123].       22   Table 2.4: Data collection and refinement statistics for SbnF Data collection  Space group P212121 Cell dimensions      a, b, c (Å) 71.72, 137.96, 150.00     α, β, γ ()  90, 90, 90 Resolution range (Å) 37.48 - 1.80 (1.86 - 1.80) * R-merge 0.050 (0.335) I / σI 13.5 (3.1) CC1/2 0.996   Refinement  Resolution (Å) 1.8 No. reflections 136049 R-work 0.1653 (0.2304) R-free 0.1946 (0.2818) No. atoms      Macromolecules 9572     Ligand 39    Water 1313 Average B-factor (Å2) 28.50     Protein 27.20     Ligand/ion 43.70     Water 37.90 R.m.s deviations      Bond lengths (Å) 0.017     Bond angles () 1.52 Ramachandran favored (%) 97 Ramachandran outliers (%)  0.17 *Values in parentheses are for the highest-resolution shell.     23   2.8 SfaD structure determination   SfaD crystal conditions were identified and optimized in the same manner as for SbnF. Crystals were grown with a reservoir condition of 0.1 M MES (pH 5.0), 5% w/v PEG 6000 with a final pH of 6.0 at 4 °C. For cryoprotection crystals were soaked for ~30 seconds in reservoir buffer supplemented with 25% (v/v) glycerol. Data were collected at the Stanford Synchrotron Radiation Lightsource beamline 7-1. Diffraction data were processed and scaled using XDS. Selenomethionine (SeMet) crystals of SfaD were grown in the same reservoir conditions as native crystals. Data for SeMet crystals were collected at the Canadian Light Source beamline 08B1-1. Diffraction data were processed and scaled using XDS. Phasing of the SeMet dataset was done by PhaserMR and an initial structure was built by Autobuild from the phenix suite [121]. The structure was refined as for SbnF    24   Table 2.5: Data collection and refinement statistics for SfaD Data collection  Space group C2221 Cell dimensions      a, b, c (Å) 77.74, 116.81, 165.45     α, β, γ ()  90, 90, 90 Resolution range (Å) 38.9 – 2.0 (2.1 - 2.0)* R-merge 0.0228 (0.125) I / σI 15.03 (5.12) CC1/2 0.999 (0.936)   Refinement  Resolution (Å) 2.0 No. reflections 101622 R-work 0.1771 (0.2096) R-free 0.2207 (0.2506)  No. atoms 5583     Macromolecules 5108    Water 475 Average B-factor (Å2) 30.60     Protein 29.97     Water 39.16 R.m.s deviations      Bond lengths (Å) 0.008     Bond angles () 1.01 Ramachandran favored (%) 97 Ramachandran outliers (%)  0 *Values in parentheses are for the highest-resolution shell.     25   2.9  HPLC analysis of SA and alternative SfaB products   As direct measurement of SfaB activity was not possible due to background signals from protein aggregation, reaction mixtures were set up with all successful substitute substrates identified for SfaD as described above and analysed by HPLC to detect final products. Reaction mixtures were analysed using an Agilent 1260 Infinity HPLC equipped with a Waters Xbridge 3.5 μm Amide column (2.1 x 100 mm). Compounds were mixed 50:50 with acetonitrile prior to injection, run in 1.25 mM ammonium acetate (pH 5.1 by glacial acetic acid) and eluted with acetonitrile.   Detection of iron-bound SA and alternative products of SfaB was done by continuous monitoring at 340 nm during the sample run. Additional validation of the detected peaks was carried out by supplementation of the sample by 0.8 mM FeCl3 before injection and comparison of peak height to un-supplemented samples.  2.10  Mass spectrometry of SA and D-Lys-SA  SA and D-Lys-SA reactions prepared and processed as described above were filtered using a 0.2 μm syringe filter and diluted to an estimated concentration of 100 μM with acetonitrile-water (3:97 vol/vol) supplemented with 0.1% formic acid. Diluted sample was directly injected into an Agilent 6550 time of flight mass spectrophotometer. Analysis of the sample was performed with electrospray ionization in negative ion mode with a scanning range of 200-500 or 200-600 m/z. The peaks corresponding to SA and D-Lys-SA were determined based on the characteristic [M-H+] -479 and 493 m/z fragments.  2.11  Bioassay of SA and D-Lys-SA samples  To evaluate whether D-Lys-SA could be used by S. aureus for iron acquisition, a bioassay on iron-chelated media previously described was used [83]. In brief, S. aureus JE2 was  26   seeded into Tris Minimal Succinate (TMS) square agar plates [124]. D-Lys-SA samples were prepared by HPLC purification by collecting the 340 nm absorption peak corresponding to D-Lys-SA. The collected peak sample was concentrated and applied to sterile paper disks placed on the seeded agar plates. A control sample using SA collected and processed in an identical manner and a negative control sample of filtered, sterile H2O were applied to filter discs as well. Agar plates containing samples were incubated at 37°C for 24-48 h and the growth radius was recorded.         27   Chapter 3: Results  3.1 Characterization of the structure and substrate preference of SA pathway NIS synthetases  3.1.1: Structure of SfaD  SA is synthesized by the sequential reaction of the NIS synthetases SfaD and SfaB from two molecules of citrate and one molecule of D-ornithine (Figure 1.5). SfaD is the first enzyme in the pathway, catalysing a condensation reaction between one molecule each of D-ornithine and citrate, utilizing one molecule of ATP in the process. Although the use of citrate defines SfaD as a type A NIS synthetase, its protein sequence is most similar to type B synthetases. Of the available structures in the protein data bank (PDB), the closest homolog based on protein sequence is the type C NIS synthetase AsbB from Bacillus anthracis (26% identity). The closest type A homolog is IucA from Klebsiella pneumoniae (24% identity).   To determine whether the structure of SfaD represented a novel type of NIS synthetase, the crystal structure of SeMet labelled SfaD was solved by anomalous dispersion and used to phase a native dataset of SfaD to a final resolution of 2 Å. The structure represents a nearly complete polypeptide, with 4 residues missing at the N-terminus, 2 at the C-terminus and no discernable density for three flexible loop regions of 13, 4 and 10 residues in length (Figure 3.1). The structure does not have substrates or other large molecules bound. Visually, the structure of SfaD conforms to the architecture of previously described NIS synthetases and comprises of three α-helix rich domains in a “cupped hand” topology. However, comparison of SfaD with the type A NIS synthetase IucA from K. pneumoniae and type A synthetase AcsD from Pectobacterium chrysanthemi shows limited structural similarity with RMSD values, based on alignment of the Cα atoms, of 3.0 Å and 3.4 Å, respectively (Table 3.1). A structural similarity  28   search of structures in the PDB, identified the type C synthetase IucC from K. pneumoniae as the most similar with an RMSD value of 3.0 Å (Table 3.1). Superimposition with an ATP-bound structure of AcsD shows that, although the overall structures are dissimilar the ATP-binding pocket is relatively well conserved (Figure 3.2). Repeated attempts to grow crystals in the presence of the native substrates, as well as to soak compounds into existing crystals, resulted in several crystals and datasets which neither improved resolution nor contained density for substrates or missing residues.  Table 3.1: Structural similarity of SfaD to NIS synthetases      NIS synthetase RMSD (Å) Z-Score Number of residues aligned PDB entry IucC 3.0 36.5 506 6CN7 AcsD 3.4 36.2 539 2W02 IucA 3.0 35.6 514 5JM8 AsbB 3.2 30.5 541 3TO3 DfoC 3.0 29.8 535 5O7O AlcC 3.1 29.2 538 2XO2 Superimposition of SfaD with structures deposited in the PDB using the DALI server, ranked by Z-score [125]. Hits with a Z-Score lower than 2 are considered spurious.     Figure 3.1 Structure of SfaD.  (A) The structure of SfaD conforms to the fold of previously described NIS synthetase structures but lacks density for several flexible loop regions. (B) A close-up view of the active site region  29   denoted by the red box in (A). The red line connects endpoints of a 13 amino acid segment absent from the structure.    Figure 3.2 Conserved ATP binding site of SfaD compared to AcsD.  Superposition of SfaD (cyan) to an ATP-bound AcsD (tan) structure shows a conserved ATP binding site. Conserved residues within hydrogen bonding distance are displayed as sticks. Atoms are coloured by heteroatom, and ATP carbon atoms are coloured purple.   3.1.2: Active site mutants of SfaD.  As the structure of SfaD did not provide details regarding enzyme function due to missing residues and lack of bound substrates, a biochemical approach was taken to define the catalytically important residues. The function of key residues in homologous NIS synthetases has been defined by other groups and is best characterized in the type A synthethase AcsD form the achromobactin synthesis pathways in P. chrysanthemi. Sequence alignment of SfaD with AcsD and other type A synthetases identified a conserved predicted catalytic arginine and an ATP-coordinating histidine at positions 351 and 496 respectively. The R351A and H496N site- 30   directed mutants were generated by PCR, transformed into E. coli and confirmed by sequencing. Purified protein mass was confirmed by MALDI-MS. Catalysis of ATP by SfaD and other NIS synthetases produces pyrophosphate (PPi) as a by-product of the reaction. As the SfaD substrates are commercially available, the reaction can be monitored spectroscopically at 360 nm by coupling PPi release to conversion of a fluorescent nucleotide (Section 2.4). Using a standard reaction containing defined concentrations of substrates, the specific activity of SfaD and active site mutants was determined (Section 2.4). Lack of enzyme turnover was defined as a specific activity at or below the negative control rate. Neither the R351A nor the H496N mutant showed detectable activity (Table 3.2 and Figure 3.2). Histidine 208 is another conserved residue in NIS synthetases and in the AcsD structure this residue interacts with the central carboxyl group of citric acid. A H208A variant was made and was found to retain wild-type activity (Table 3.1).   Figure 3.3 Pyrophosphate assay of SfaD and its active site mutants.  Activity of active site SfaD mutants was measured by a coupled pyrophosphate release assay monitored at 360 nm for 10 minutes. No absorbance change was detected for either the R351A (blue) or H496N (green) mutant. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).    31   Table 3.2 Specific activity of SfaD mutantsa   SfaD mutant Specific activity (nmole min-1 μM-1) Physiological reaction 560 ± 40 Negative control (no D-ornithine) 60 ± 40 R351A N.D. H496N N.D. H208A 550 ± 40 aThe specific activity was determined using a coupled pyrophosphate release assay. The reactions were initiated by adding 10 μM SfaD. All measurements were performed in triplicate. N.D. (Not Detected) defined as activity below negative control sample.   3.1.3: Substrate preference of SfaD  To characterize its substrate preference, the specific activities of SfaD with analogues of the native substrates D-ornithine and citric acid were determined using the PPi release assay described previously. Analogues were selected based on commercial availability and presence of the required amine and carboxyl groups necessary for the condensation reaction to occur as described (Figure 1.4). Successful catalysis by SfaD was defined as a significant specific activity above the negative control. SfaD turnover in reactions with a single substituted component was detected with 6 of out 7 D-ornithine and 3 out 8 citric acid substitutions tested (Table 3.3). Additionally, SfaD is able to utilize either enantiomer of ornithine and lysine (Figure 3.4 A and B). Replacement of citrate with alternate compounds showed turnover with DL-isocitric acid, hydroxycitrate and tricarballylic acid (Figure 3.5 B and D). The specific activities measured for DL-isocitric acid, diaminobutyric, diaminopropionic acids and 5-hydroxylysine are apparent due to the mixed isomer stocks used (Figure 3.4 C).  32    Figure 3.4 Activity of SfaD with D-ornithine and its analogues.  (A) Turnover as measured by absorbance at 360 nm for SfaD physiological reaction (red), L-ornithine substitution (blue) and control reaction without D-ornithine present (black). (B) Absorbance of physiological reaction (red), negative control (black), DL-5 hydroxylysine (green), L-Lysine (orange) and D-Lysine (purple). (C) Absorbance of physiological reaction (red), negative control (black), DL-diaminobutyric acid (yellow), DL-diaminopropionic acid (blue) and putreschine (purple). Reactions were measured continuously for 10 minutes. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).    Figure 3.5 Activity of SfaD with citrate and its analogues.  (A) SfaD turnover as measured by absorbance at 360 nm with WT substrates (red), glutaric acid (blue), 3OH3CH3 glutaric acid (blue) and a negative control (black). (B) Turnover of physiological reaction (red), negative control (black), tricarballylic acid (blue) and diglycolic  33   acid (brown). (C) SfaD turnover with WT components (red), negative control (black), DL-malic acid (orange) and malonic acid (green) with separate negative controls for either (yellow and cyan). (C) Turnover of WT SfaD reaction (red), negative control (black), hydroxycitrate (green) and DL-isocitric acid (brown). Reactions were monitored for 10 minutes. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).  Table 3.3 Specific activity of SfaDa   D-ornithine substitute Specific activity (nmole min-1 μM-1) Citrate substitute Specific activity (nmole min-1 μM-1) L-ornithine 420 ± 40 Glutaric acid N.D. D-lysine 610 ± 40 3-hydroxy-3-methylglutaric acid N.D. L-lysine 170 ± 40 DL-malic acida N.D. DL-2,3-Diaminopropionic acida 580 ± 40 Malonic acid N.D. DL-2,4-Diaminobutyric acida 620 ± 40 Diglycolic acid N.D. DL-5-Hydroxylysinea 480 ± 40 DL-Isocitric acidb 350 ± 40 Putrescine N.D. Hydroxycitrate 380 ± 40   Tricarballylic acid 550 ± 40 Physiological reaction (D-ornithine + citrate) 560 ± 40 Negative control (- D-ornithine) 60 ± 40  aAll measurements were performed in triplicates. N.D. (Not Detected) is defined as activity below negative control sample. bReported specific activity is apparent.    3.1.4: Substrate preference of SfaB  The synthesis of SA is catalysed by the sequential reactions of SfaD and SfaB. SfaB is a type A synthetase that catalyses the condensation of citrate with the δ-citryl-D-ornithine product of SfaD (Figure 1.5). The identification of alternative substrates for SfaD allowed testing of both substrates of SfaB. Several attempts were made to adapt the previously described PPi release assay for use with SfaB. However, high background signals due to aggregation of protein interfered with measurements. Instead, reaction mixtures containing SfaD and substrates were set up and incubated overnight. These reactions were filtered to remove SfaD, SfaB was added and  34   incubated for 6-8 hours. Following the second incubation, SfaB was removed and the sample was analysed by HPLC using an amide column. Control reactions were set up without SfaB to distinguish the final product from the SfaD-generated intermediate. Further controls included a physiological reaction mixture to generate SA. Iron-bound SA can be detected by HPLC by absorption at 340 nm and is distinct from intermediate signal (Figure 3.6). As no iron was supplemented into the reaction mixture, synthesized SA is believed to be a mix of apo and holo-SA. Analysis of reactions with citrate and each of the identified SfaD substrate analogues showed a single peak present for the reaction with D-lysine (Figure 3.7A). No peak was observed in the intermediate control (no SfaB present). Henceforth this product is referred to as D-Lys-SA. As the WT product of SfaB is a functional siderophore, approximately equimolar FeCl3 was added to the reaction mixture prior to HPLC analysis to evaluate whether the product was capable of binding ferric iron. The peak in the HPLC trace of FeCl3-supplemented sample eluted at the same time point but with a greater absorption at 340 nm than sample without supplementation (Figure 3.7B).   Figure 3.6 Detection of SA by HPLC.  35   HPLC trace of wild type synthesis reaction run through an amide column and monitored at 340 nm. SA tR 17.4 min. Reaction control tR 16.8 min.  Figure 3.7 D-Lys-SA detection by HPLC. D-Lys-SA (tR = 17.4 min) was detected via HPLC as a single peak at 340 nm by analysing the reaction mixture using an amide column (A). D-Lys-SA peak signal was significantly increased by the addition of FeCl3 to the reaction mixture prior to analysis (tR = 17.5 min) (B).   3.1.5: Validation of D-Lys-SA  The modified product detected by HPLC, D-Lys-SA, represents an extension of the D-ornithine backbone by one methylene group and would increase the molecular weight from 480.123 Da to 494.137 (Figure 3.8). To confirm the identity of this molecule, reaction mixtures were tested by negative ion-mode MS. MS analysis confirmed the presence of a species with a weight of 493.157 that was absent in reactions lacking SfaB (Figure 3.9B). Analysis of reaction mixtures of citrate with D-ornithine to produce SA showed a peak corresponding to a weight of 479.120 that was absent in reactions without SfaB (Figure 3.9A).    36    Figure 3.8 Predicted structure and iron-coordination of D-Lys-SA.     Figure 3.9 MS analysis of SA and D-Lys-SA reaction mixtures. MS analysis of SA reaction mixtures showed an m/z peak at 479.120 (A) under negative-ion mode. The same conditions showed an m/z peak at 493.157 (B) for the D-Lys-SA reaction mixture.   To evaluate whether S. aureus was able to use D-Lys-SA for iron acquisition, the product was purified by HPLC and concentrated. A bioassay on iron-limiting media containing S. aureus showed halos of growth for both SA and D-Lys-SA with no growth for an H2O control (Figure 3.10).    37    Figure 3.10 Bioassay analysis of D-Lys-SA samples.  Addition of 10 μl to a filter disk (white circle) located on solid, iron-chelated, agar media of either purified D-Lys-SA (A), SA (B) or H2O (C) produced halos of S. aureus growth for the D-Lys-SA and SA samples. Extent of the growth halo has been emphasized with a red circle for clarity.   3.2  Characterization of the structure and substrate preference of SB pathway enzymes  3.2.1: Structure of SbnF  SB is synthesized by a four-enzyme pathway encoded on the sbn locus (Figure 1.6). SbnF, is a type-C synthetase that catalyses the condensation of one molecule of L-DAP and citryl-diaminoethane (Figure 1.6). For insights into enzymes of the SB pathway the crystal structure of SbnF was solved by molecular replacement with the NIS synthetase AsbB (36% amino acid identity) and refined to a resolution of 1.8 Å. The structure consists of two molecules in the asymmetric unit and each protein is missing three residues in a single flexible loop region (Figure 3.11). SbnF retains the “cupped hand” domain architecture of NIS synthetases and SfaD. Comparing the structure directly to AsbB (used for molecular replacement) by superposition of the Cα atoms generates an RMSD value of 1.8 Å (546 residues aligned).  Alignment with the structure of AcsD gives an RMSD value of 3.2 Å (517 residues aligned). Interestingly, alignment with SfaD (RMSD 2.4 Å) (500 residues aligned) shows a high degree of similarity between both enzymes. Two molecules of citrate, present in the crystallization conditions, are bound within the  38   protein (Figure 3.12). Alignment with an ATP-bound structure of AcsD (PDB entry 2W02) identifies the placement of one of the citrates in the triphosphate region of the ATP binding pocket. The other citrate molecule is bound in a putative citryl-diaminoethane (C-DAE) binding site (PDB entry 2X3J) (Figure 3.13). Attempts to crystallize the other NIS synthetases in the pathway (SbnE and SbnC) or obtain substrate-bound structures of SbnF were unsuccessful.   Figure 3.11 Structure of SbnF. SbnF is found as a dimer in the asymmetric unit. The fold and domain organization are similar to other NIS synthetases.     39    Figure 3.12 The structure of SbnF contains two citrates bound in the predicted active site.  Residues within hydrogen-bonding distance have been displayed as sticks. One citrate is located in a histidine-coordinated pocket, whereas the second citrate appears to mimic a triphosphate in the ATP binding site.    40    Figure 3.13 Citrate bound in a predicted substrate-binding site in SbnF. A superposition of the structure of SbnF (green) and that of AcsD bound to substrate (tan, PDB entry 2X3J). The citryl-diaminoethane bound to AcsD (blue) and citrate bound to SbnF (grey) and residues within hydrogen bond distance are drawn as sticks. Oxygen and nitrogen atoms are coloured red and blue respectively.  3.2.2: Active site mutants of SbnF  SbnF shares a higher degree of sequence and structural similarity to other characterized NIS synthetases than SfaD. Sequence alignment of SbnF with other synthetases showed the presence of catalytic arginine and histidine residues at positions 292 and 426, respectively. To confirm their role in catalysis, site-directed mutants R292A and H426N were generated by PCR, transformed into E. coli and confirmed by sequencing. Mass of expressed protein was confirmed by MALDI-MS. To test the activity of both mutants, the specific activity of WT protein was first defined using the previously described PPi release assay. Turnover was defined as specific activity above the negative control. WT substrate had a specific activity of 360 ± 40 nmole min-1  41   μM-1 and a negative control without L-DAP present gave a calculated specific activity of 200 ± 40 nmole min-1 μM-1. No activity was detected for either mutant (Figure 3.14). Activity of the control reaction (no L-DAP) can be attributed to SbnF turnover with citrate only.    Figure 3.14 Pyrophosphate assay of SbnF active site mutants. Activity of active site SfaD mutants was measured by a coupled pyrophosphate release assay monitored at 360 nm for 5 minutes. No turnover was detected for either the R292A (blue) or H426N (orange) mutant. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).  3.2.3: Substrate preference of SbnE   To characterize the substrate preference of enzymes in the SB pathway, activity of the first three enzymes (SbnE, SbnH and SbnF) with substrate analogues was measured. The same group of analogues used for SfaD characterization was used to provide a comparison to the SA pathway. SbnE, the first enzyme in SB synthesis and a type A NIS synthetase, utilizes one molecule each of L-DAP, citric acid and ATP (Figure 1.6). Specific activity with substrate analogues was measured as described using a PPi release assay. To validate detection of enzyme function, specific activity with known physiological substrates or control reactions lacking citrate or L-DAP were tested. Physiological substrates produced a specific rate of 650 ± 40 nmole min-1 μM-1. Control reactions produced rates of 40 ± 40 (no citrate) and 70 ± 40 nmole min-1 μM-1 (no L-DAP).  42   Testing of the citrate analogues showed turnover with tricarballylic acid, isocitric acid, and hydroxycitrate, mirroring the results recorded with SfaD (Table 3.4 and Figure 3.15B). In contrast with SfaD, no substitute displayed a specific activity higher than the natural substrate (Table 3.4). Furthermore, none of the L-DAP replacements displayed a specific activity higher than the negative control (Table 3.4 and Figure 3.16)   Figure 3.15 Activity of SbnE with the substrate citrate and analogues.  Testing of SbnE citrate substitutions showed no absorbance change for several tested compounds (A) when compared against a WT SbnE reaction (red). (B) Turnover was detected with DL-isocitric acid (green), hydroxycitrate (brown) and tricarballylic acid (blue). Reactions were monitored for 5 minutes. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).   Figure 3.16 Activity of SbnE with the substrate L-DAP and analogues. Reactions of SbnE in the presence of L-DAP replacements did not produce any absorbance increase. Reactions were monitored for 5 minutes. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).   43   Table 3.4 Specific activity of SbnEa   D-ornithine substitute Specific activity (nmole min-1 μM-1) Citrate substitute Specific activity (nmole min-1 μM-1) L-ornithine N.D. Glutaric acid N.D. D-lysine N.D. 3-Hydroxy-3-methylglutaric acid N.D. L-lysine N.D. DL-Malic acid N.D. DL-2,4-Diaminobutyric acid N.D. Malonic acid N.D. DL-5-Hydroxylysine N.D. Diglycolic acid N.D. Putrescine N.D. DL-Isocitric acid 230 ± 40   Hydroxycitrate 550 ± 40   Tricarballylic acid 480 ± 40 Physiological reaction (L-DAP + citrate) 650 ± 40 Negative control (no L-DAP) 70 ± 40  aAll measurements were performed in triplicate. N.D. is defined as activity below the negative control sample.   3.2.4: Substrate preference of SbnH   SbnH is the second enzyme in the SB synthesis pathway and decarboxylates the product of the first enzyme SbnE, releasing CO2 (Figure 1.6). The identification of several functional substrate analogues for SbnE provided an opportunity to study the substrate preference of SbnH. As the orientation of hydroxycitrate and isocitric acid in the active site of SbnE is unknown, several potential products are possible (Figure 3.17). To track turnover of SbnH, a CO2 release assay in a positive pressure nitrogen atmosphere glovebox was adapted from an existing protocol. This assay couples CO2 release to utilization of NADH by malic dehydrogenase, resulting in a decrease in absorbance at 340 nm. The products of the one-pot SbnE reactions were tested as substrates. SbnE was filtered out and substrates were allowed to equilibrate in the glovebox overnight. To confirm that turnover of enzyme could be detected, physiological substrates and products of control reaction lacking either the enzyme SbnE or the substrate L-DAP were tested. The absorption changes observed were not linear with time. Enzyme activity  44   was qualitatively assessed by the absorption change after 3 minutes. Physiological substrate produced an absorbance reading of -0.164 ± 0.012, control reactions produced readings of -0.058 ± 0.008 (No-SbnE) and -0.057 ± 0.009 (No L-DAP). The background signal seen in the control reaction mixtures lacking SbnE or L-DAP can potentially be attributed to residual CO2 present in the glovebox or assay buffers.   Using this metric, SbnH showed turnover with SbnE products derived from all three citrate analogues tested (tricarballylic acid, hydroxycitrate and isocitric acid substituted SbnE products) as well as the physiological substrate (Figure 3.18). Control reactions lacking either SbnE or the SbnE substrate L-DAP showed a significantly smaller decrease in absorbance (Figure 3.18).  Figure 3.17 Possible products of SbnE citrate substitutions.  (A) SbnE is capable of producing the WT product citryl-L2,3-diaminopropionic acid, a tricarballylic substitution product (D) and two possible products for isocitrate (B and C) and hydroxycitrate (E and F) substitutions.    45    Figure 3.18 Activity of SbnH with citryl-L2,3-diaminopropionate and analogues.    Activity of SbnH with products of SbnE reactions containing L-DAP and either citrate or the indicated alternative substrate. SbnH showed catalysis with all three modified SbnE products. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).   3.2.5: Substrate preference of SbnF  SbnF is a type C NIS synthetase catalysing condensation of a molecule of L-DAP to the product of SbnH, utilizing a molecule of ATP in the process. Due to the uncertain nature of hydroxylysine and isocitric acid placement in the SbnE active site, several different SbnH products are possible (Figure 3.19). Substrates derived from tricarballylate, hydroxycitrate and isocitrate instead of citrate were produced using SbnE and SbnH. Specific activity of SbnF was determined as described previously using a PPi release assay. Turnover was defined as specific activity above the negative control. WT substrate produced a specific activity of 360 ± 40 nmole min-1 μM-1 and a negative control without L-DAP present gave a calculated specific activity of 200 ± 40 nmole min-1 μM-1. Assay of SbnF activity showed turnover the hydroxycitrate and tricarballylate derived substrates (Table 3.5, Figure 3.20 A and B). The specific activity with the  46   isocitrate-derived substrate was not statistically significantly different from the control (Table 3.5 and Figure 3.20 C).    Figure 3.19 Possible substrates generated for SbnF from reactions with SbnE and SbnH. (A) SbnH is capable of producing the WT product citryl-diaminoethane, a tricarballylic substitution product (D) and two possible products for isocitrate (B and C) and hydroxycitrate (E and F) SbnE substitution products.     47    Figure 3.20 Activity of SbnF with the substrate citryl-diaminoethane and analogues. Enzyme turnover as measured by absorbance at 360 nm was detected with (A) hydroxycitrate (blue) and (B) tricarballylic acid (green). (C) Specific activity with isocitrate was not significant from control (black). Reactions were monitored for 5 minutes. All measurements performed in triplicate. Data plotted as standard deviation and error (thin dotted line).  Table 3.5 Specific activity of SbnFa  D-ornithine substitute Specific activity (nmole min-1 μM-1)bc Physiological reaction 160 ± 40 DL-Isocitric acid N.D. Hydroxycitrate 310 ± 40 Tricarballylic acid 210 ± 40  aAll measurements were performed in triplicate. N.D. is defined as activity below the negative control sample.  bReported parameters are apparent.  cValues normalised to negative control (no L-DAP).     48   Chapter 4: Discussion   4.1  Substrate preference of SA pathway enzymes  4.1.1: Substrate preference of SfaD  To date six NIS synthetases have been structurally characterized. Of those, two are type A NIS synthetases. SfaD is categorized as a type A synthetase due to it using citrate as a substrate, but it is more closely related to type B and C synthetases based on amino acid sequence identity [81]. To define the structural features of SfaD, a crystal structure of apo-SfaD was solved. Comparison of SfaD to other synthetases shows a similar domain architecture, but relatively high RMSD values when superimposed with other NIS synthetases indicates SfaD has structurally unique features. Conservation of the ATP binding site and key catalytic residues suggests that SfaD is conserved in the core active site region but is much more variable on the periphery. The significance for this variability is unknown. Comparison with type C synthetase SbnF showed that SfaD is structurally most similar to another NIS synthetase from S. aureus.  Unfortunately, the structural model of SfaD is missing residues 204 to 217, including several residues predicted to form the citrate-binding pocket of the active site. Attempts to crystallize SfaD and active site mutants of SfaD in complex with substrates to stabilize this loop were unsuccessful. The SfaD crystal structure leaves unanswered questions regarding the high degree of substrate promiscuity.  The general enzymatic mechanism of NIS synthetases has been described biochemically and supported by structural analysis of the active site [102, 107]. In addition, some of these synthetases were shown to turn over non-native substrates, with several reported cases of nucleophile substitution and one case of citrate substitution [126-128]. The high degree of  49   structural similarity within the family and conservation of key amino acid residues suggested that S. aureus homologs may also be able to accommodate a variety of substrate analogues.   Using a PPi release assay to measure specific activity, SfaD was found to process six analogues of D-ornithine and three analogues of citrate (Figures 4.1 and 4.2). All functional D-ornithine replacements retained a similar chemical structure with a terminal carboxyl group and a pair of amine groups (Figure 4.3). Previously, only the D isomer of ornithine was shown to give rise to SA [81]. Here both the D and L enantiomers of ornithine and lysine are substrates of SfaD, consistent with more stringent substrate selection resting with SfaB, the second SA synthetase. Additionally, the lack of any enzyme activity with putrescine, suggests that the carboxylate group is required for substrate recognition by SfaD, as the condensation reaction occurs at the δ-amine (Figure 4.4). Functional citrate substitutes retained the same overall length and carboxyl groups in the native substrate (Figure 4.5). No activity was detected with shorter substitutes such as malate and malonate. Similarly, removal of the central carboxyl group, such as in the case of glutaric acid was sufficient to prevent reaction. Successful analogues show the central hydroxyl group is not essential, as isocitrate or even hydroxycitrate retain activity (Figure 4.5). This degree of substrate promiscuity found in SfaD is thus far unique in NIS synthetases as previous examples of functional substitutions were limited to only a handful of compounds [107, 126-128].    50    Figure 4.1 Progression of citric acid substitutions through the SA synthesis pathway. No citrate substitute product was detected by HPLC, only the WT product citrate gives rise to WT SA.   51    Figure 4.2 Progression of D-ornithine substitutes through the SA pathway.  D-Lysine substitution gives rise to a final product in the full SA pathway reaction.   52    Figure 4.3 Functional nucleophile substitutions in SfaD.  SfaD is capable of utilizing native D-ornithine (A), L-ornnithine (B), DL-diaminobutyric acid (C), DL-diaminopropionic acid (D), D and L enantiomers of lysine (E and F) as well as DL-5-hydroxylysine. All functional D-ornithine substitutions retain a terminal carboxylic acid group and a pair of amines.    Figure 4.4 Comparison of D-ornithine and putrescine structures. SfaD is not capable of catalysing a reaction using putrescine, which is a decarboxylated derivative of its WT substrate D-ornithine.  53    Figure 4.5 Functional citrate substitutions in SfaD.  SfaD is capable of catalysing reactions containing either the carboxylic acid citrate (A), tricarballylic acid (B), Hydroxycitrate (C) or Isocitrate (D). All functional citrate analogues retain the three carboxyl groups found on citrate.   4.1.2 Production of a SA analogue.   The second and final enzyme in SA synthesis pathway, SfaB, is a type C NIS synthetase that adds a second citric acid group to the intermediate produced by SfaD. Although the continuous assay was not suitable likely due to the relative instability of the protein in solution, analysis by HPLC of incubated reactions provided some evidence regarding the substrate preference of SfaB. Of the nine alternate SfaD products tested, only with D-lysine was a product formed detectable by HPLC (Figures 4.1 and 4.2). Selectivity for D isoform is consistent with enantiomer specificity in assembly of the final siderophore product [81]. Exclusion of the diaminobutyric and diaminopropionic acid derived products may be due to the amino group  54   being poorly positioned for condensation with a second citrate. The use of absorption at 340 nm in the HPLC analysis may have failed to detect products unable to bind iron sufficiently. However, for the analogue to be useful as a siderophore it must retain the ability to bind ferric iron. Overall these findings suggest that SfaB is the major substrate selection point in the SA pathway, as it successfully excluded all but one of the alternate SfaD products.  The detection of an SfaB product and the signal increase detected by HPLC upon addition of FeCl3 suggested that this molecule may bind ferric iron and act as a siderophore. Modified siderophores require uptake by native receptors to be useful tools in “trojan horse” and other applications. The ability of S. aureus to bind and take up modified siderophore would be an indication of the viability of these approaches in this species.   MS analysis of complete and control reaction mixtures detected the presence of peaks with masses within error of the predicted mass of both SA and D-Lys-SA. The predicted mass and structure of D-Lys-SA corresponds to the structure of corynebactin from Corynebacterium diphtheria [129]. However, the stereochemistry of corynebactin and D-Lys-SA are unknown and these siderophores may be stereoisomers. Furthermore, a bioassay confirmed that S. aureus was capable of utilizing the new siderophore to acquire iron from iron restricted media. It is not known whether S. aureus is capable of utilizing corynebactin for iron acquisition. The smaller size of the halo in the D-Lys-SA sample may be a consequence of a difference in concentration as we did not quantify the SA or D-Lys-SA purified by HPLC. The fact that S. aureus is capable of growth using a modified form of SA suggests that further modifications of SA and SB, potentially conjugated to other molecules, may be a viable approach.      55   4.2  The SB pathway  4.2.1: Substrate preference of SB pathway enzymes  The SB biosynthesis pathway in S. aureus is more complex than SA synthesis. Four enzymes are required to sequentially assemble SB: SbnE, SbnH, SbnF, and SbnC. Successful substitute selection requires processing by all four enzymes. An important difference with the sfa pathway is the presence of the PLP-dependent decarboxylase SbnH. To investigate whether the SB pathway could be manipulated to produce alternate products, the first three enzymes were assayed for their activity with substrate analogues. The first enzyme, SbnE, catalyses the condensation of citrate with L-DAP, similar to the condensation of citrate with D-ornithine catalysed by SfaD. SbnE and SfaD accepted the same citric acid replacements (Figure 4.6). However, in contrast to SfaD, SbnE, in the presence of citric acid, was found to not accept any replacements for L-DAP. As a result, a much higher degree of substrate selectivity is present at the initial stage of SB synthesis when compared to SA.   56    Figure 4.6 Citric acid substitute progression through the SB pathway. Two substitutes for citric acid were able to generate products that were used by all enzymes tested.    The SbnE product citryl-L2,3-diaminopropionate is decarboxylated by the PLP-dependent decarboxylase SbnH. Interestingly, SbnH decarboxylated the alternative SbnE  57   products; however, since the SbnE products were not characterized and more than one product could be formed, whether all or only some of these products are processed by SbnH was not determined.   The presence of one citrate molecule in the conserved ATP-binding site of SbnF and another in the putative C-DAE substrate site suggests that other carboxylic acids may interact with the active site of SbnF. This is observed in the PPi release assays where negative controls that contained citrate showed a change in absorbance, suggesting that SbnF is capable of turning over with citrate. Additionally, assaying for SbnF activity with alternative substrates produced by SbnH showed that tricarballylic acid and hydroxycitrate substitution products were utilized by SbnF. The exclusion of both forms of the isocitrate variant indicates that structural isomers are excluded in later stages of siderophore assembly, as both SbnE and SbnH utilized structural isomers of isocitrate. Attempts to obtain a native-substrate bound structure using WT and site-directed mutants of SbnF were unsuccessful.  Comparison of SfaD and SbnF to AcsD, a type A synthetase used for structural characterization of NIS synthetases, shows that catalytic residues are conserved (Figure 4.7). Active site mutants of SfaD R351A and SbnF R292A showed no discernable activity. Furthermore, no activity was detected for mutants of a predicted ATP-coordinating residue; SfaD H496N and SbnF H426N. The histidine residues can be clearly identified in the predicted ATP-biding pocket of the SfaD and SbnF structures. Additionally, the SfaD H208A mutant was generated. This residue was predicted to coordinate citrate based on structures of other synthetases but has not previously been described as essential. Phosphate assay results showed that this mutant indeed retained functionality with its native substrates. These findings confirm  58   that the S. aureus NIS synthetases SfaD and SbnF retain the same mechanism described in other NIS synthetases and show a high degree of structural conservation in the active site region.      59    Figure 4.7 Key catalytic residues in SfaD and SbnF. Superimposition of (A) SfaD (cyan) with AcsD (tan) and (B) SbnF (green) with AcsD (tan). Residues required for catalysis are drawn in sticks and are labelled.    The final enzyme in the SB pathway, SbnC, was not assayed due to difficulties with protein purification and stability, however, several observations regarding substrate preferences of enzymes in the SB pathway were made. The stringency of SbnE for L-DAP ensures that subsequent enzymes in the pathway are provided with the proper backbone for further assembly of the siderophore. Based on the exclusion of citrate substitution products by SfaB it would be possible for SbnC to act as the final control step in SB assembly.    4.3  Conclusions  The ability of S. aureus to scavenge iron from the human host has been repeatedly shown to play a crucial role in establishing an infection. The siderophores SA and SB form a key part of the iron-acquisition repertoire used by this pathogen. In my study, I was able to obtain structures of S. aureus NIS synthetases SfaD and SbnF. Comparison to existing NIS synthetase structures showed conservation of the core active site region with variability at the periphery. Furthermore, I made mutations of a catalytically essential arginine (SfaD R351A and SbnF R292A), a histidine that binds ATP (SfaD H496N and SbnF H426N), as well as a non-essential histidine (SfaD H208A), to confirm that both synthetases function in accordance with a previously described mechanism. Using enzymes from both SA and SB pathways I was able to show catalytic activity with a number of substrate analogues which provided insight into substrate specificities of the NIS synthetases SfaD, SbnE, SbnF and the decarboxylase SbnH. Finally, a modified SA was produced, detected and validated, showing that S. aureus is able to use modified SA for iron-acquisition.   60   4.4  Future Directions  The specificity of S. aureus NIS synthetases could be further characterized by defining the kinetic parameters with the physiological and alternate substrates identified in this study. To this end, I have collected preliminary enzyme rate data for the physiological and some alternate substrates.   Although progress has been made in determining the substrate binding sites of SfaD and SbnF, structures of both enzymes in complex with native and alternative substrates would shed light on why certain NIS synthetases are more promiscuous than others. Currently, I have generated crystals of site-directed mutants for both SbnF and SfaD in the presence of several substrates but have not been able to refine them to sufficient diffraction quality. Alternatively, computational docking methods could be attempted to model substrates into the apo-structures.   Similarly, a structure of SbnH in complex with substrate would reveal details about substrate preference that would complement the initial assay results obtained in this study. Crystals of inactive site-directed mutants have been generated, but solved structures did not possess discernable density for even a partial substrate molecule. If a crystallographic approach proves unsuccessful, an existing apo structure of SbnH may be used in docking trials to predict key coordinating residues which may be validated by site-directed mutants.   Finally, the aspect of generating a modified siderophore can be greatly expanded by testing additional substrates and making mutations on enzymes in both pathways to reduce substrate selectivity. The ability of S. aureus to use a modified SA for iron-acquisition presents an opportunity to further develop variants with altered backbone structure and work towards generating an easily modifiable siderophore. This could then be used in a semi-synthetic process  61   to produce conjugates with antibiotic or other functional groups and adapted for further studies of a clinical nature.     62   Bibliography  1. Lowy, F.D., Staphylococcus aureus infections. N Engl J Med, 1998. 339(8): p. 520-32. 2. Williams, R.E., Healthy carriage of Staphylococcus aureus: its prevalence and importance. 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