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Role of aspartate aminotransferases AspB and AspC in the WhiB7-controlled intrinsic drug resistance system.. Ng, Carol Ka Lo 2012

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ROLE OF ASPARTATE AMINOTRANSFERASES AspB AND AspC IN THE WhiB7-CONTROLLED INTRINSIC DRUG RESISTANCE SYSTEM OF MYCOBACTERIA  by  Carol Ka Lo Ng  BSc (Hon), University of British Columbia 2005  A THESIS SUMBITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE  in  THE FACULTY OF GRADUATE STUDIES (Microbiology and Immunology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  October 2012  © Carol K L Ng, 2012  Abstract Intrinsic resistance of the intracellular pathogen Mycobacterium tuberculosis is one of the main reasons that the disease tuberculosis is difficult to treat and why it remains as one of the world’s most prevalent and dangerous infectious diseases. The intrinsic resistance regulator WhiB7 controls a regulon that contains many genes predicted to have physiological functions including aspartate aminotransferases, aspB and aspC. Multi-drug susceptibility was observed in an aspC mutant and an aspB constitutive expression strain. The expression of aspC was positively regulated by WhiB7 while expression of aspB downregulated whiB7 expression. The fitness of Mycobacterium smegmatis was affected negatively by oxaloacetate and positively by α-ketoglutarate, substrates of aspartate aminotransferase, which then altered the growth inhibition by antibiotics. Recombinant AspB and AspC both catalyze measurable transamination of aspartate and α-ketoglutarate. AspC plays an important role in mycobacteria physiology as deletion of this gene caused many growth deficiencies. Furthermore, the physiological role of AspC extended beyond amino acid intermediary metabolism to redox homeostasis and oxidative stress detoxification. These results revealed a link between intrinsic antibiotic resistance and metabolism mediated through AspB and AspC. Since antibiotic resistance in mycobacterium is a complex function of its physiology, it is important to screen for tuberculosis drugs under growth conditions that resemble those found in vivo.  ii  Preface Dr. Gaye Sweet performed the catalase activity assay experiments. Experimental design and analysis of the data were a collaborative effort between Dr. Sweet and Carol Ng. Steady-state kinetic determination of AspB was done in collaboration with Antonio Ruzzini in the group of Dr. Lindsay E. Eltis (University of British Columbia). Purified protein was supplied by Carol Ng. The work in this thesis is conducted with approval from University of British Columbia Office of Research and in accordance with the University of British Columbia Policies and Procedures, Biosafety Practices and Public Health Agency of Canada Guidelines. The protocols in this work were approved by the UBC Biosafety Committee. The certificate number is B07-0169.  iii  Table of contents  Abstract ................................................................................................................................ ii Preface ................................................................................................................................. iii Table of contents.................................................................................................................. iv List of tables ........................................................................................................................ vii List of figures .......................................................................................................................viii List of abbreviations .............................................................................................................. x Acknowledgments ................................................................................................................xii 1. Introduction ...................................................................................................................... 1 1. 1. Mycobacterium tuberculosis and antibiotic resistance ................................................................... 1 1. 2. The WhiB family of proteins ............................................................................................................ 2 1. 3. Oxidative stress and tuberculosis pathogenesis .............................................................................. 3 1. 4. Physiological changes induced by antibiotics .................................................................................. 5 1. 5. Mycobacterial amino acid aminotransferases ................................................................................. 7 1. 6. Project rationale .............................................................................................................................. 8  2. Materials and methods ...................................................................................................... 9 2. 1. Cloning and reagents ..................................................................................................................... 13 2. 2. Growth conditions ......................................................................................................................... 13 2. 3. Constitutive expression strain construction .................................................................................. 13 2. 4. Mutant construction ...................................................................................................................... 14 2. 5. Construction of double mutant ΔaspBΔaspC ................................................................................ 14 2. 6. Drug susceptibility testing.............................................................................................................. 15 2. 7. Growth kinetics .............................................................................................................................. 15 2. 8. Antibiotic kill curve ........................................................................................................................ 15 2. 9. RNA isolation.................................................................................................................................. 16 2. 10. Quantitative real-time PCR .......................................................................................................... 16 iv  2. 11. NAD-NADH pool isolation and NAD cycling assay ....................................................................... 16 2. 12. Hydrogen peroxide and menadione disc assay ........................................................................... 17 2. 13. Preparation of cell lysate ............................................................................................................. 17 2. 14. Catalase assay .............................................................................................................................. 18 2. 15. Phylogenic analysis of AspBMtb and AspCMtb ................................................................................ 18 2. 16. Cloning of aspB and aspC in pGEX4T-1 ........................................................................................ 18 2. 17. Purification of GST-tagged AspBMtb and AspCMtb ......................................................................... 18 2. 18. Thrombin cleavage of GST-tagged AspBMtb and AspCMtb ............................................................. 19 2. 19. Cloning of aspB and aspC in pET expression vectors ................................................................... 19 2. 20. Expression of recombinant His-tagged AspBMtb and AspCMtb ...................................................... 19 2. 21. Purification of recombinant His-tagged AspBMtb and AspCMtb ..................................................... 20 2. 22. Removal of His tag from AspBMtb ................................................................................................. 20 2. 23. Cloning and expression of AspCMtb without fusion tags .............................................................. 21 2. 24. Cloning and expression of AspCMtb in M. smegmatis ................................................................... 21 2. 25. Aspartate aminotransferase assay............................................................................................... 21  3. Results............................................................................................................................. 23 3. 1. Expression of whiB7 upregulates aspC .......................................................................................... 23 3. 2. Expression of whiB7 and aspC is upregulated by tetracycline....................................................... 24 3. 3. Optimal whiB7 induction by tetracycline requires aspC ................................................................ 26 3. 4. Constitutive expression of aspB downregulates whiB7 and aspC in tetracycline-induced cultures ............................................................................................................................................................... 26 3. 5. The ∆aspC mutant exhibits growth deficiencies ............................................................................ 28 3. 6. The ΔaspC is able to catalyze glutamate but not aspartate .......................................................... 30 3. 7. α-ketoglutarate complements ΔaspC mutant’s growth deficiencies ............................................ 30 3. 8. Passaging the ΔaspC mutant in media containing α-ketoglutarate suppresses the growth rate defect but does not shorten lag phase .................................................................................................. 32 3. 9. The ΔaspB ΔaspC double mutant is not an amino acid auxotroph ............................................... 32 3. 10. Opacification of 7H10 by ΔaspC mutant ...................................................................................... 34 v  3. 11. The ΔwhiB7 mutant, ΔaspC mutant, and aspB constitutive expression strains are antibiotic sensitive ................................................................................................................................................. 35 3. 12. Complementation of antibiotic susceptibility of the ΔaspC mutant ........................................... 35 3. 13. Effects of antibiotics on the growth kinetics of the ΔaspC mutant ............................................. 38 3. 14. Differential killing of the ΔaspC mutant by clarithromycin ......................................................... 42 3. 15. Oxaloacetate and α-ketoglutarate modulate the rate of recovery from stationary phase growth and adaptation to bacteriostatic drugs ................................................................................................. 43 3. 16. Deletion of aspC causes a shift in intracellular redox homeostasis............................................. 46 3. 17. Increased oxidative stress in the ΔaspC mutant .......................................................................... 46 3. 18. Phylogenic analysis corroborates aspB and aspC as putative aminotransferases ...................... 50 3. 19. Complementation of aspartate auxotrophy in E. coli with AspBMtb and AspCMtb ........................ 56 3. 20. Amino acid auxotroph DL39 does not exhibit general growth deficiencies and does not show increased antibiotic susceptibility ......................................................................................................... 58 3. 21. Expression and purification of AspBMtb and AspCMtb ................................................................... 61 3. 22. Aspartate aminotransferase activity of AspBMtb and AspCMtb ...................................................... 73  4. Discussion ....................................................................................................................... 75 4. 1. aspB and aspC expression is linked to whiB7 expression .............................................................. 75 4. 2. Intermediary metabolism of mycobacteria is linked to intrinsic antibiotic resistance ................. 78 4. 3. Role of AspB and AspC in mycobacteria physiology ...................................................................... 82 4. 4. Implicated roles for AspC in redox homeostasis and oxidative stress detoxification ................... 84 4. 5. Concluding remarks ....................................................................................................................... 87  References .......................................................................................................................... 88  vi  List of tables Table 1. Plasmids and strains used in this study. ........................................................................................ 10 Table 2. Primers used in this study ............................................................................................................. 12 Table 3. Growth properties of ΔaspB and ΔaspC mutants in cultures containing aspartate and glutamate as carbon and/or nitrogen sources. ............................................................................................................ 31 Table 4. Antibiotic susceptibility profiles of the ΔwhiB7 mutant, hsp-aspB expression, and the ΔaspC mutant strains of M. smegmatis. ................................................................................................................ 36 Table 5. Effect of antibiotics and putative aspartate transaminase substrates on time of recovery from stationary phase growth arrest................................................................................................................... 45 Table 6. Functions of anchor residues in family I aminotransferases. ....................................................... 55 Table 7. Amino acid auxotrophic strain DL39 does not show changes in antibiotic susceptibility. ........... 59 Table 8. Expression of AspCMtb in E. coli and M. smegmatis ....................................................................... 72 Table 9. Specific activities of purified aspartate aminotransferases. ......................................................... 74 Table 10. Comparison of steady state kinetic parameters determined for AspCMtb and AspCEco. ......... 74  vii  List of figures Figure 1. Detoxification of reactive oxygen species...................................................................................... 4 Figure 2. Oxidative damage via Fenton’s reaction occurs in 3 steps ............................................................ 6 Figure 3. Reaction catalyzed by aspartate aminotransferases ................................................................... 23 Figure 4. Expression of aspC was upregulated by whiB7 ............................................................................ 25 Figure 5. Expression responses of whiB7, aspB, and aspC to tetracycline ................................................. 25 Figure 6. Interactive regulation of aspB, whiB7 and aspC .......................................................................... 27 Figure 7. Growth kinetics of M. smegmatis wild-type, ∆aspB, ∆aspC, hsp-aspBMtb, hsp-aspCMtb, and ∆aspC complemented ............................................................................................................................................ 29 Figure 8. Growth of the ΔaspC mutant was modulated by addition of aspartate aminotransferase substrates.................................................................................................................................................... 31 Figure 9. Allowing the ΔaspC mutant to adapt to growth in media containing α-ketoglutarate did not suppress the extended lag phase growth deficiency.................................................................................. 33 Figure 10. Growth kinetics of double mutant ΔaspBΔaspC ........................................................................ 33 Figure 11. Opacification of 7H10 by the ΔaspC mutant.............................................................................. 34 Figure 12. Complementation of antibiotic susceptibility of the ΔaspC mutant by constitutive expression of aspCMtb or addition of α-ketoglutarate ................................................................................................... 37 Figure 13. The time required for wild-type mc2155 to adapt and grow in media containing antibiotics is dependent on antibiotic concentration ...................................................................................................... 39 Figure 14. The difference between wild-type and the ΔaspC mutant growth kinetics in media containing antibiotic ..................................................................................................................................................... 40 Figure 15. Differential killing of the ΔaspC mutant by clarithromycin ....................................................... 42 Figure 16. Effect of α-ketoglutarate and oxaloacetate supplementation on growth kinetics and antibiotic inhibition of growth in wild-type M. smegmatis ........................................................................................ 44 Figure 17. The ΔaspC mutant has an increased redox state ....................................................................... 47  viii  Figure 18. Sensitivity of mutants ΔaspB and ΔaspC to oxidative stress compounds hydrogen peroxide and menadione ........................................................................................................................................... 48 Figure 19. Higher oxidative stress in the ΔaspC mutant reflected in catalase activity ............................... 49 Figure 20. Phylogenetic tree showing amino acid homology of various aminotransferases ..................... 52 Figure 21. Neighbor-joining phylogenic tree of aspartate aminotransferases from different organisms . 54 Figure 22. Vector maps of pUC19::hsp-aspB and pUC19::hsp-aspC........................................................... 57 Figure 23. Amino acid auxotroph DL39 has a lower cell density at stationary phase ................................ 59 Figure 24. Oxaloacetate inhibited growth of E. coli at 40mM and 80mM ................................................. 60 Figure 25. Expression vector maps for GST-tagged recombinant protein expression ............................... 62 Figure 26. Purification of AspBMtb and AspCMtb using GST affinity chromatography .................................. 63 Figure 27. Removal of GST-tag from recombinant protein GST-AspBMtb.................................................... 64 Figure 28. GST-AspCMtb contain many putative secondary thrombin cleavage sites ................................. 66 Figure 29. Thrombin cleavage of GST-AspCMtb ............................................................................................ 67 Figure 30. His-tagged AspBMtb and AspCMtb expression vectors.................................................................. 68 Figure 31. Purified AspBMtb and His-tag removal ........................................................................................ 69 Figure 32. Purification of C-terminal His-tagged AspCMtb ........................................................................... 70 Figure 33. Schematic representation of the proposed antibiotic induced regulatory network of whiB7, aspB, and aspC ............................................................................................................................................ 77  ix  List of abbreviations Abbreviation ADC AES AspAT AT BCA cDNA cfu CLR DMSO DNA DTT Eco EDTA GST HEPES his hsp / hsp60 INH IPTG LB MDR MIC mRNA Msm Mtb MTT MWCO  Expanded albumin dextrose catalase allelic exchange substrate aspartate aminotransferase aminotransferase bicinchoninic acid complementary deoxyribonucleic acid colony forming unit clarithromycin dimethyl sulfoxide deoxyribonucleic acid dithiothreitol Escherichia coli ethylenediaminetetraacetic acid glutathione s-transferase 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid hexa-histidine heat shock promoter isoniazid isopropyl β-D-1-thiogalactopyranoside Luria-Bertani multidrug resistant minimum inhibitory concentration messenger ribonucleic acid Mycobacterium smegmatis Mycobacterium tuberculosis [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide; thiazolyl blue molecular weight cut off  NAD+ NADH NADPH OAA OADC PAGE PB PBS PCR PEG PMSF qRT-PCR RNI  nicotinamide adenine dinucleotide (oxidized form) nicotinamide adenine dinucleotide (reduced form) nicotinamide adenine dinucleotide phosphate oxaloacetate oleic acid albumin dextrose catalase polyacrylamide gel electrophoresis Proskauer Beck phosphate buffered saline polymerase chain reaction polyethylene glycol phenylmethanesulfonylfluoride quantitative real-time PCR reactive nitrogen species x  ROS SDS SPT TB TET WT XDR YT αKG  reactive oxygen species sodium dodecyl sulfate spectinomycin tuberculosis tetracycline wild-type extensively drug resistant yeast extract tryptone alpha ketoglutarate  xi  Acknowledgments I wish to offer my deepest gratitude to Dr. Charles Thompson for giving me this opportunity to do a graduate project under his supervision, and whose guidance and support enabled me to explore and expand my abilities. I would like to thank my committee members, Dr. William Mohn and Dr. Lindsay Eltis for their counsel and support. I wish to acknowledge the generosity of the Hallam, Eltis, Mohn, and Murphy labs for the use of their equipment. Special thanks to Dr. Santiago Ramón-García who first started me out on this project and for continual enthusiasm and input into this research. I would like to also express my sincere gratitude towards Dr. Gaye Sweet for invaluable guidance throughout this program and for critical reading of this thesis. It was an honour and pleasure to work alongside my lab mates in the Thompson lab both past and present, especially Leah Lim and Jan Burian, their friendship and lively discussions was what created such a positive work environment. Above all, I owe a great debt of gratitude to my family and friends whose infinite support is from where I drew the courage to pursue my passion in research. Profound thanks go to my parents for their unfailing love and support throughout the pursuit of this degree. Words cannot express how much their support has meant throughout my life, especially now. The research project was funded in part by a Graduate Entrance Scholarship from the University of British Columbia. Additional funding was provided by CIHR and Lung Association grants awarded to Dr. Charles Thompson.  xii  1. Introduction 1. 1. Mycobacterium tuberculosis and antibiotic resistance The acid-fast bacterium M. tuberculosis is the causative agent of tuberculosis. Tuberculosis remains one of the top infectious causes of mortality, with 8.8 million incident cases and 1.4 million deaths worldwide in 2010 [1]. The current treatment for tuberculosis is a combination of four antibiotics administered over a course of 6 months. Noncompliance or abandonment of treatment is the major impediment to effective therapy due to the long and sometimes unpleasant regimen required to cure tuberculosis. This further contributes to the emergence of drug resistant mutants. The development of drug-resistance is a growing problem for tuberculosis control. Multidrug-resistant TB (MDR-TB) and extensively drug- resistant TB (XDR-TB) are forms of tuberculosis that are even more difficult and expensive to treat because they fail to respond to standard first- and second-line therapy [2,3]. There are an estimated 290,000 cases of MDR-TB in 2010 making antibiotic resistant tuberculosis a global health concern [1]. Acquired drug resistance to the two most powerful front-line antibiotics used to treat tuberculosis, isoniazid and rifampicin, is on the rise. In 1994, mono resistance to isoniazid and rifampicin was found at a rate of 4.1% and 0.2% respectively [4]. In a 2012 report, the frequency of these resistances had increased to 9.3% and 0.3% [5]. To further hinder treatment, mycobacteria have high intrinsic resistance to most naturally occurring antibiotics, including many commonly-used broad-spectrum drugs. In contrast to acquired drug resistance, which occurs via random mutation or horizontal gene transfer, intrinsic resistance is the natural resistance of the bacterial species and is typically chromosomally encoded. The intrinsic resistance of mycobacteria has conventionally been attributed to the mycolic acid-containing cell envelope as it may serve as a barrier to antibiotic entry by diffusion [6]. In addition to decreased permeability of the bacterial envelope, the M. tuberculosis chromosome contains intrinsic resistance genes that encode efflux pumps [7] [8], β-lactamases [9] [10], and ribosome methyltransferases [11] [12]. Clearly, M. tuberculosis has evolved many mechanisms to counteract the actions of antibiotics. Mutations conferring antibiotic resistance in M. tuberculosis are often found associated with a cost which reduces the fitness of the bacterium [13] [14]. Furthermore, physiological state and growth rate of the bacteria can also influence antibiotic susceptibility [15,16]. Intrinsic resistance mechanisms of M. tuberculosis would appear to have strong links to the general physiology of the bacterium.  1  1. 2. The WhiB family of proteins WhiB homologues are highly conserved throughout the Actinomycetales order with most species containing multiple orthologs [17]. WhiB proteins are small (87 to 130 amino acid residues), putative transcription factors with conserved structural characteristics such as a glycine-rich helix-turn-helix motif that could interact with RNA polymerase. WhiB proteins also possess four near-invariant cysteine residues arranged as C-X19-36-C-X-X-C-X5-7-C which may act as ligands for a metal cofactor and are predicted to have a role in sensing redox change [18]. WhiB proteins have been implicated in a variety of functions from sporulation in Streptomyces coelicolor [17] to oxidative stress in Corynebacterium glutamicum [19]. The M. tuberculosis genome has seven annotated whiB genes implicated in a variety of physiological processes including cell division, pathogenesis, starvation and detection of stress conditions [20]. All M. tuberculosis WhiB proteins, except for WhiB2, have disulfide reductase activity [20]. While these assays typically are based on reduction of insulin, WhiB1 can catalyze reduction of at least one M. tuberculosis protein [20]. WhiB1 acts in a thioredoxin-like manner to reduce GlgB, a mycobacterial enzyme important in carbohydrate metabolism [21]. WhiB2 regulates expression of genes that are involved in cell division [22]. WhiB3 controls fatty acid metabolism [23]. The seven M. tuberculosis whiB genes were differentially expressed during different growth phases and in response to environmental stresses such as pH, nutrient starvation, antibiotic assault, and oxidative stress [24,25] [26]. Experimental evidence has shown that both WhiB1 and WhiB3 coordinate redox-sensitive Fe-S clusters that are sensitive to oxygen and nitric oxide [27] [28]. In general, WhiB proteins are used by M. tuberculosis to coordinate specific expression of proteins that the bacterium requires for survival under unfavourable conditions. The transcriptional regulator WhiB7 has a central role in regulation of intrinsic multidrug resistance [29]. WhiB7 controls expression of antibiotic resistance genes including tap (Rv1258c), an efflux transporter, erm, a methyltransferase (Rv1988), and eis (Rv2416c), an aminoglycoside acetyltransferase [29]. The efflux pump Tap confers tetracycline resistance by extruding the antibiotic from the cell [30,31]. ermMT encodes a methyltransferase which causes methylation of 23S rRNA leading to resistance against macrolides [11]. The gene eis, initially named for enhanced intracellular survival in macrophages [32], confers kanamycin resistance by acetylating the antibiotic as well as host signalling molecules resulting in suppression of host immune responses [33] [34]. Deletion of whiB7 in M. tuberculosis, M. smegmatis, or M. bovis BCG renders the strain sensitive to a broad spectrum of antibiotics such as macrolides, rifampicin, streptomycin, tetracycline and chloramphenicol [29]. The expression of whiB7 is induced by antibiotics, fatty acids, redox-modifying compounds, and immediately after entry to macrophages [29] [35,36]. The M. smegmatis whiB7 mutant is deficient in mycothiol, the major low-molecular-weight thiol produced by mycobacteria, linking whiB7 to redox homeostasis [35]. These results suggest that WhiB7 links bacterial physiology and redox homeostasis to intrinsic antibiotic resistance.  2  1. 3. Oxidative stress and tuberculosis pathogenesis The success of M. tuberculosis as an intracellular pathogen depends on its survival within host macrophages. The macrophage creates a highly oxidative environment by producing reactive oxygen species (ROS) [37] [38] in response to infection. To survive the exposure to exogenous, host-derived oxidative stress, M. tuberculosis must employ strategies to sense and respond to changes in its microenvironment. WhiB proteins have been implicated in this process. WhiB1, WhiB2, and WhiB3 transcription activators differentially bound to DNA sequences in promoters depending on its redox state [27] [39] [40]. Similarly, the activity of WhiB7 was strongly induced by a more reduced cytoplasmic environment [35]. These sensors allow M. tuberculosis to detect oxidative stress and make the appropriate response, such as expression of detoxifying factors and modulating cellular metabolism, to promote survival inside the host. In order to survive within macrophages, M. tuberculosis must withstand the toxicity of ROS produced by phagocyte NADPH oxidase (NOX2/gp91phox) [41]. Activated macrophages generate superoxide molecules and other ROS by transporting electrons across the plasma membrane from cytosolic NADPH to molecular oxygen [42]. The thick, mycolic acid-containing cell wall of M. tuberculosis limits diffusion of toxic molecules but the bacteria also contain an arsenal of mechanisms to detoxify ROIs before they can cause damage to the bacteria. To counteract the deleterious effects of ROS such as superoxides and hydroxyl radicals, M. tuberculosis requires superoxide dismutase (SOD) and catalase to convert the ROS to water and molecular oxygen. The M. tuberculosis genome contains two SODs: sodA and sodC. The function of catalase in M. tuberculosis is performed by a single enzyme, KatG, which functions in two other capacities: peroxidase and peroxynitritase [43] [44] [45]. AhpCD, another oxidative stress detoxifier, encode alkyl hydroperoxidases to neutralize the alkyl hydroperoxides generated by the reaction between lipids and superoxides and is required for survival in macrophages [46,47,48,49]. SodC is localized to the mycobacterial cell envelope [50] and is essential to resistance against the initial oxidative burst generated by IFNγ-activated macrophages [51]. sodA and katG are required for growth of M. tuberculosis in vitro [52] and appear to be important during mouse infections. SodA and KatG are exported by the SecA2 virulence secretion system; M. tuberculosis ΔsecA2 is unable to grow in nonactivated macrophages and showed reduced growth in mice due to the bacteria being defective in SodA and KatG export [53,54]. KatG confers protection against both hydrogen peroxide [55] [56] and peroxynitrite [45]. The M. tuberculosis katG mutant was attenuated in wild-type mice but virulent in mice lacking NOX2 [41]. While more research is required for better understanding of the mechanisms of ROS detoxification in M. tuberculosis, their detoxification genes are needed for virulence.  3  Figure 1. Detoxification of reactive oxygen species  Figure 1. Detoxification of reactive oxygen species. Organisms that grow aerobically and pathogens that encounter immune cells in the host are routinely exposed to oxidative stress in the form of reactive oxygen species (ROS) such as oxygen radicals, hydrogen peroxide, and hydroxyl radicals. Detoxification enzymes, SodA and SodC, convert oxygen radicals to hydrogen peroxide. Hydrogen peroxide can react to form hydroxyl radicals through Fenton’s reaction, requiring the catalase activity of KatG to degrade hydrogen peroxide to water and molecular oxygen.  4  1. 4. Physiological changes induced by antibiotics Antibiotics and their antibacterial activities are rigorously studied for their potential use as therapeutics. Traditionally it is thought that antibiotics cause growth inhibition by a singular cellular target within systems such as DNA replication or repair, protein synthesis, or cell wall turnover. However, global expression studies have revealed that changes in expression profiles induced by subinhibitory concentrations of antibiotics can be diverse and some are within genes not related to the target function [57] [58] [59] [60] [61] [62]. In mycobacteria, inactivation of one of a number of genes with roles in physiology and metabolism leads to a multidrug sensitive phenotype; mspA, a porin protein [63], fbpA, a cell wall biosynthesis gene [6], pknG, a serine/threonine kinase related to glutamine/glutamate metabolism [64,65] and asnB, an asparagine synthetase [66]. The fact that these genes with physiological roles in transport, metabolism, or cell envelope functions are activated in response to antibiotic exposure implicates physiology of the bacterium to play a role in antibiotic resistance. The redox-responsive transcription regulator WhiB7 is a central regulator of intrinsic resistance. In addition to induction of expression by antibiotics, the activation of whiB7 is triggered by small molecules that are known to perturb respiration, redox balance and transmembrane ion flux [35]. Furthermore, the expression of whiB7 is highly dependent on the reducing potential in the cytoplasm and on the main mycobacterial thiol antioxidant, mycothiol [35], placing this transcription regulator at the junction between redox homeostasis and antibiotic resistance. Indeed, antibiotics have been shown to cause both oxidative [67] [68] and reductive stress [69]. A large body of work by the Collins lab indicates that bactericidal antibiotics instigate the production of hydroxyl radicals produced from hydrogen peroxide via Fenton’s reaction and cause bacterial cell death via oxidative damage [67] [70] [71]. Many bacteria produce catalases and peroxidases to scavenge hydrogen peroxide and protect themselves from reactive oxygen species. E. coli and many other bacteria rely on the action of alkylhydroperoxide reductase (Ahp), a NAD(P)H peroxidase, for the detoxification of hydrogen peroxide [72] [73]. The action of Ahp, like other antioxidation systems, involves the transfer of electrons between pools of NAD(P)H, FADH, and thiol redox compounds (Burian, J, 2012, in review). Mycothiol and other thiol-reducing agents require reducing power in order to eliminate the oxidative stress. Therefore, the cytoplasmic redox potential of mycobacteria is kept reduced in order to maintain a pool of thiols in preparation for any oxidoreductive stress brought on by the host immune system [46]. During infection, the cytoplasmic redox potential of M. tuberculosis is highly reduced. This redox imbalance is likely generated through metabolic pathways such as hypoxia, inhibition of respiration by nitric oxide, and β-oxidation of lipids [74], pathways which are upregulated in M. tuberculosis during infection. These processes all generate a high NADH/NAD+ ratio making the regeneration of NAD+ important for redox homeostasis and survival of the bacilli [75] [76]. A role for redox imbalance has also been implicated in the mechanism of antibiotic action. In contrast to the traditional thought that antibiotics act on a single target, a proposed mechanism by Kohanski, Collins and coworkers points to an 5  additional underlying mechanism in which bactericidal antibiotics cause oxidative stress and bacterial death is brought on by the toxic accumulation of reactive oxygen species [67,71,77]. While hydrogen peroxide cannot directly oxidize DNA, it can react very rapidly with transition metals. Hydroxyl radicals are generated through reaction of hydrogen peroxide with ferrous iron via Fenton’s reaction (Figure 2 reaction 1). Hydroxyl radicals are powerful oxidants and react with the base or sugar residues of DNA, resulting in base modification and DNA breakage (Figure 1 reaction 2). The ferric iron can be reduced back to ferrous state by a reductant and the reaction cycles to generate more hydroxyl radicals (Figure 1 reaction 3)[78] [79] [80]. The tricarboxylic acid (TCA) cycle is a source of the reductant NADH and TCA cycle mutants were less susceptible to bactericidal drugs [67]. The results which led to this proposed pathway were derived from E. coli experiments but similar results have been observed with Staphylococcus [67], Pseudomonas [80], Listeria [81], Leishmania [82], and Acinetobacter [83] suggesting that this phenomenon may extend to all bacteria (and protozoans) and perhaps even Mycobacterium. The poorly understood but significant role of redox homeostasis and antioxidation in M. tuberculosis resistance could prove to be a good target candidate for tuberculosis therapy and deserves additional study.  Figure 2. Oxidative damage via Fenton’s reaction occurs in 3 steps  Figure 2. Oxidative damage via Fenton’s reaction occurs in 3 steps. Hydroxyl radicals are formed through reactions with hydrogen peroxide and free iron. Radicals cause DNA damage and mutagenesis. Iron II is regenerated by a reducing agent (NADH).  6  1. 5. Mycobacterial amino acid aminotransferases Bacteria encode enzymes for the synthesis of all twenty amino acids required for protein synthesis [84]. Pyridoxal 5′-phosphate-dependent transaminase performs the final biosynthetic step in these pathways, converting keto acid precursors into α-amino acids. In the hopes of identifying inhibitors of aminotransferases for target-based tuberculosis drug development, the structures of several aminotransferases of M. tuberculosis have been solved, including phosphoserine aminotransferase (SerC, Rv0884c) [85], histidinol phosphate aminotransferase (HisC2, Rv3772) [86] and branched-chain amino acid aminotransferase (IlvE, Rv2210c) [87]. There are two putative aspartate aminotransferases annotated in M. tuberculosis genome, aspB (Rv3565) and aspC (Rv0337c). The transamination reaction of aspartate serves as a link between amino acid and carbon metabolism. Aspartate aminotransferases form aspartate from oxaloacetate, an intermediate of the TCA cycle, via a transamination reaction with glutamate. Aspartate can then act as an intermediate for the biosynthesis of other amino acids such as asparagine, threonine, isoleucine, methionine, and lysine [88]. Furthermore, aspartate is required in central physiological pathways such as pyrimidine ribonucleotide, purine nucleotide and nicotinamide dinucleotide biosynthesis [88]. Aspartate aminotransferases catalyze the reversible reaction of transamination between a dicarboxylic α-amino acid and the corresponding α-keto-acids by a ping-pong bi-bi mechanism with pyridoxal 5’-phosphate (PLP) as a cofactor [89]. The M. tuberculosis genome contains a large conserved gene cluster involved in cholesterol catabolism as a carbon source during infection [90]. The expression of genes for growth of M. tuberculosis on lipids such as cholesterol is controlled by two TetR- type regulators, KstR (Rv3574) and KstR2 (Rv3557c) [91,92]. aspB belongs to a regulon controlled by the transcriptional repressor KstR2 [92]. Deletion of KstR2 in M. smegmatis de-repressed genes in the regulon including aspB, resulting in increased expression [92]. aspB is located on the M. tuberculosis genome within an operon that includes fatty acid CoA ligase and acyl-CoA dehydrogenases [92]. KstR2 control the expression of genes required for utilizing cholesterol as a carbon source during infection is implicated [91]. The aspC gene does not appear to be part of any operon. aspC was identified as being required for growth in vitro [93].  7  1. 6. Project rationale Expression of both aspB (Rv3565) and aspC (Rv0337c) genes were dependent on WhiB7 in a microarray experiment (unpublished data, Thompson Lab). When whiB7 was expressed, both by antibiotic induction and from a plasmid promoter, the expression of aspB was downregulated. Conversely, the expression of aspC was upregulated along with whiB7 expression. As putative aspartate aminotransferases, AspB and AspC have predicted functions in amino acid intermediary metabolism. That aspB and aspC were in the WhiB7 regulon strongly suggested that they could contribute to antibiotic resistance in Mycobacteria. WhiB7 responds to changes to cell metabolism by sensing and responding to changes in cytoplasmic redox potential. Based on the observation that WhiB7 responds to redox stress and controls expression of metabolic genes such as aspB and aspC, we hypothesized that mycobacterial intermediary metabolism plays a role in the WhiB7-controlled antibiotic resistance system mediated through AspB and AspC.  8  2. Materials and methods Table 1. Plasmids and strains used in this study. Plasmid/Strain  pMV361::aspB  Relevant features Single-copy, L5 integration-proficient mycobacterial expression vector containing kanamycin resistance, integrase gene, and the hsp60 promoter upstream of a multiple cloning site pMV361 expressing aspB (Rv3565) from hsp60  pMV361::aspC  pMV361 expressing aspC (Rv0337c) from hsp60  pMV361::whiB7Mtb  pMV361 expressing whiB7 (Rv3197a) from hsp60  pMV361::whiB7Msm p004S-AESaspB  pMV361 expressing whiB7 (MSMEG_1953) from hsp60 plasmid containing AES for aspBMsm deletion  pYUB854-AESaspC  plasmid containing AES for aspCMsm deletion  pJV53  acetamide inducible expression of the Che9c recombination genes  [95]  pYUB870 pGEX4T-1::aspB pGEX4T-1::aspC pET19b::aspB pET30b::aspC pET19b::aspC  for transient expression of γδ –resolvase pGEX4T-1 (GE) cloned with aspB for GST-tagged AspB expression pGEX4T-1 (GE) cloned with aspC for GST-tagged AspC expression pET19b (Novagen) cloned with aspB for His-tagged AspB expression pET30b (Novagen) cloned with aspC for His-tagged AspC expression pET19b (Novagen) cloned with aspC for His-tagged AspB expression  [96]  pET30b::aspC1 pGro7, pTf-2, pG-KJE8, pTf16  pET30b (Novagen) cloned with aspC for tag free AspC expression  pMV361  expression of chaperone proteins GroEL, GroES, GrpE, Tf, DnaJK  pYUB1062::aspC  acetamide inducible protein expression vector expression in mycobacteria pYUB1062 cloned with aspC expression in mc24517  pUC19::hsp-aspBMtb  expression of aspB from hsp60 promoter in E. coli  pUC19::hsp-aspCMtb  expression of aspC from hsp60 promoter in E. coli  E. coli TOP 10  general cloning  pYUB1062  E. coli Rosetta  TM  2  protein expression  E. coli BL21 (DE3)  protein expression. λDE3 expresses T7 RNA polymerase  2  mycobacteria strain for T7-based expression  2  wild-type M. smegmatis  2  ∆whiB7 mutant  2  mc 155 ∆aspB  ∆aspB mutant  mc2155 ∆aspC  ∆aspC mutant  mc 155 mc 155 ∆whiB7  [94]  Takara Bio [97]  protein expression. pRARE2 expresses tRNAs for rare codons  E. coli BL21 mc 4517  References  [97]  9  mc2155 ∆aspB ∆aspC  ∆aspB ∆aspC double mutant  mc2155 pMV361  wild-type vector control  mc2155 pMV361::aspB  constitutive aspBMtb expression  mc2155 pMV361::aspC  constitutive aspCMtb expression  mc2155 pMV361::whiB7Mtb  constitutive whiB7Mtb expression  mc2155 pMV361::whiB7Msm  constitutive whiB7Msm expression  ΔaspC pMV361::aspCMtb  mutant ΔaspC with constitutive expression of aspCMtb  DL39  E. coli mutant in aspC, tyrB, and ilvE; auxotrophic for Val, Leu, Ile, Tyr, Phe, Asp  MG1655  wild-type laboratory strain of E. coli K-12  DL39 pUC19  DL39 vector control  DL39 pUC19::hspaspBMtb  DL39 that constitutively expresses aspBMtb  DL39 pUC19::hspaspCMtb  DL39 that constitutively expresses aspCMtb  [98]  Table 1. Plasmids and strains used in this study.  10  Table 2. Primers used in this study Primer  Sequence (restriction site)  Purpose  BOO-13  TTTCGAATTCGTGACGGATCGTGTC (EcoRI)  aspBMtb forward primer  BOO-14  GCAAAGCTTCTATTGGCTCGGCAG (HindIII)  aspBMtb reverse primer  BOO-15  AGGGAATTCGTGGACAACGATGGC (EcoRI)  aspCMtb forward primer  BOO-16  TAGAAGCTTCTATTGCCGGTAACTG (HindIII)  aspCMtb reverse primer  BOO-30  CCATGAATTCATGACTGCTCCGACCACGG (EcoRI)  whiB7Msm forward primer  BOO-31  AAATAAGCTTGATCAGGCGGCGGC (HindIII)  whiB7Msm reverse primer  BOO-32  GATATAGAATTCGTGTCGGTACTGACAGTCCCC (EcoRI)  whiB7Mtb forward primer  BOO-33  TAGAAAGCTTCTATGCAACAGCATCCTTGC-3' (HindIII)  whiB7Mtb reverse primer  BOO-34  TTTTTTTTCCATAAATTGGCGTTGTTGTTGCTGCGGG (PflMI)  AES for Msmeg_6017 (aspBMsm)  BOO-42  TTTTTTTTCCATTTCTTGGGCGGTCGTTCATCTCACCC (PflMI)  AES for Msmeg_6017 (aspBMsm)  BOO-36  TTTTTTTTCCATAGATTGGCGTGCGAGGCAACTCCTATG(PflMI)  AES for Msmeg_6017 (aspBMsm)  BOO-43  TTTTTTTTCCATCTTTTGGCCAACGACCGCTTCCTGTC (PflMI)  AES for Msmeg_6017 (aspBMsm)  BOO-61  TTTTTTCTTAAGAAGCCCGCAGCCACAGAGG (AflII)  AES for Msmeg_0688 (aspCMsm)  BOO-62  TTTTTTTCTAGATCGTGGACTGGGTGAACGTG (XbaI)  AES for Msmeg_0688 (aspCMsm)  BOO-63 BOO-64 BOO-103 BOO-105 BOO-119 BOO-120  TTTTTTAGATCTCCCGAGGTGTACGACATCCAC (BglII) TTTTTTACTAGTGCCGTGGTCGCAACCTTTC (SpeI) GACCCGGCGCCAAGAAGACC CCGCGAAGGTCAGGCACGTC CCGTCGACCCCACGCGCAAG GACGTTCGGCGACGCGGTCC  AES for Msmeg_0688 (aspCMsm) AES for Msmeg_0688 (aspCMsm) forward primer upstream of aspC reverse primer downstream of aspC forward primer upstream of aspB reverse primer downstream of aspB  BOO-92  ACGTTCTCGGTGGTGCTGCG  reverse primer for hyg  BOO-104  CACCGATCCGGAGGAACTGGC  Hyg down (fw)  prAB49  CCAAAAACCATCTGCTGGAG  mysAMsm forward primer for qRT-PCR  prAB50  AGGTTGCCTTCCTGGATGAG  prAB47a  TCCATTGCGATGACTGCTC  prAB48  GTTCTCGGCGAACCACAG  prAspB_fw GGGTCTGCTGACGCACTAC  mysAMsm reverse primer for qRT-PCR whiB7Msm forward primer for qRTPCR whiB7Msm reverse primer for qRTPCR aspBMsm forward primer for qRT-PCR  prAspB_rv  CAGCGAGTTGTCGGTGTACT  aspBMsm reverse primer for qRT-PCR  prAspC_fw ATCCCGGCACCCGACTACCC  aspCMsm forward primer for qRT-PCR  prAspC_rv  CCACGATCGCCTTGGTGCGT  aspCMsm reverse primer for qRT-PCR  BOO-78  TTTTGGATCCGT GAC GGA TCG TGT CGC (BamHI)  aspBMtb forward primer  BOO-79  TTTTGAATTCCTA TTG GCT CGG CAGC (EcoRi)  aspBMtb reverse primer  BOO-80  TTTTGGATCCGTG GAC AAC GAT GGC ACCATTG (BamHI)  aspCMtb forward primer 11  BOO-81  TTTTGAATTCCTA TTG CCG GTA ACT GACCAGGAAG (EcoRI)  aspCMtb reverse primer  BOO-106  TTTTCATATG GT GAC GGA TCG TGT CGC (NdeI)  aspBMtb forward primer  BOO-107  TTTTGGATCC CTA TTG GCT CGG CAGC (BamHI)  aspBMtb reverse primer  BOO-108  TTTTCATATG GTG GAC AAC GAT GGC ACCATTG (NdeI)  aspCMtb forward primer  BOO-109  TTTTGGATCC CTA TTG CCG GTA ACT GACCAGGAAG (BamHI) TTTT GGTACC TTG CCG GTA ACT GACCAGGAAG (KpnI) mutated stop codon  aspCMtb reverse primer  BOO-110  aspCMtb reverse primer  Table 2. Primers used in this study  12  2. 1. Cloning and reagents E. coli TOP 10 (Invitrogen) was used for general cloning procedures. Media components were purchased from Difco. All DNA primers were synthesized by Integrated DNA Technologies, Inc. All PCRs were performed using Dynazyme EXT polymerase (New England Biolabs) according to the manufacturer's protocol with buffer sometimes supplemented with 5% (v/v) DMSO. Restriction enzymes, purchased from New England Biolabs and Rapid DNA Dephos & Ligation Kit from Roche Applied Bioscience, were used in all molecular cloning protocols following standard recombinant DNA techniques. Plasmids pET19b and pET30b were purchased from Novagen. Plasmid DNA pGEX4T-1 was purchased from GE Healthcare. Fine chemicals were acquired from Sigma unless otherwise indicated. Strains, plasmids, and oligonucleotides used are summarized in Tables 1 and 2.  2. 2. Growth conditions The bacterial strains used are listed in Table 1. E. coli strains were grown in LB medium at 37°C with shaking. Antibiotics (ampicillin 100 μg/ml, kanamycin 300 μg/ml, chloramphenicol 34 μg/ml) were added to the medium when necessary. M. smegmatis mc2155 strains and its derived mutants were routinely grown in Middlebrook 7H9 broth (Difco) supplemented with 0.05% tyloxapol, 0.2% glycerol, and 10% albumin-dextrose-catalase (ADC; Difco) or on Middlebrook 7H10 agar (Difco) supplemented with 0.5% glycerol, and 10% oleic acid-albumin-dextrose-catalase (OADC; Difco). Cultures were aerated using an orbital shaker at 200 rpm or culture roller drum at level 5. For experiments investigating nutritional growth requirements of M. smegmatis, Proskauer Beck (PB) medium was used (36.7 mM KH2PO4, 37.8 mM NH4Cl, 2.4 mM MgSO4•7H2O, 11.6 mM magnesium citrate, 2% v/v glycerol). 0.05% tyloxapol (v/v) was added to 7H9 and PB medium to promote dispersed growth except in the case with drug susceptibility testing using alamar blue or MTT assay in microwells. Minimal media 56/2 (Glucose, 0.3%; KH2PO4, 0.5 g/L; Na2HPO4, 0. 93 g/L; (NH4)2SO4, 0.2 g/L; tri-sodium citrate 0.14 g/L; MgSO4.7H2O, 20 mg/L; FeSO4.7H2O, 2.5 mg/L; Ca(NO3)2.4H2O, 0.7 mg/L; amino acids 50 μg/ml; 1.5% agar, pH 7.6) was used as the minimal medium for E. coli nutritional requirement studies[99].  2. 3. Constitutive expression strain construction Strains that constitutively express M. smegmatis whiB7 (MSMEG_1953), M. tuberculosis whiB7 (Rv3197a), aspB (Rv3565) and aspC (Rv0337c) in M. smegmatis were generated by cloning into the integrative vector pMV361 that drives expression from a heat-shock promoter (hsp60). The genes were amplified from M. tuberculosis H37Rv or M. smegmatis mc2 155 genomic DNA (for primers refer to Table 2) with engineered EcoR1 and HindIII restriction sites (whiB7Msm, BOO-30 and BOO-31; whiB7Mtb, BOO-32 13  and BOO-33; aspB, BOO-13 and BOO-14; aspC, BOO-15 and BOO16). The PCR fragments were digested and cloned into pMV361 digested with EcoR1 and HindIII. The cloned vectors were transformed into mc2 155 to create hsp promoter-driven constitutive expression strains (hsp-whiB7Mtb, hsp-aspBMtb and hspaspCMtb).  2. 4. Mutant construction Knockout strains of aspBMsm (MSMEG_6017) and aspCMsm (MSMEG_0688) in M. smegmatis mc2155 were generated by allele exchange using recombineering technique [95]. The allelic exchange substrate (AES) for aspCMsm deletion was prepared by amplifying the upstream and downstream flanking regions and cloning them on either side of the hygromycin-resistant gene in the shuttle vector pYUB854. The upstream region was amplified from genomic DNA using primers BOO-61 and BOO-62 and cloned between AflII and XbaI sites in pYUB854. The downstream region was amplified with primers BOO-63 and BOO-64 and cloned between BglII and SpeI. The linear aspCMsm AES was removed from pYUB854 by AflII and SpeI digestion and electroporated into mc2155 expressing Che9c recombination genes from pJV53. Construction of the mc2155 ΔaspBMsm mutant was carried out following a similar strategy. The aspBMsm flanking regions were amplified from genomic DNA using primers BOO-34 and BOO-42 for the upstream region and BOO-36 and BOO-43 for the downstream region. A PflMI restriction enzyme site was engineered at the 5’ end of each primer. The amplicons were digested with PflMI and cloned into PflMI-digested p0004s vector arms. P0004s is a delivery vector that contains lambda phage cos sites, a hyg resistance marker, and a sacB cassette (T. Hsu and W. R. Jacobs, Jr., unpublished data). The linearized aspBMsm AES was generated with BamHI digestion and electroporated into mc2155 expressing recombineering functions from pJV53. For deletion of both aspB and aspC, colonies resistant to hygromycin and kanamycin were screened for precise allele replacements by PCR using primers specific to the hygromycin cassette (BOO-92 and BOO-104) and primers outside the flanking sequences (BOO119 and BOO-120 for aspB and BOO-103 and BOO-105 for aspC).  2. 5. Construction of double mutant ΔaspBΔaspC The double mutant was constructed by first removing the hygromycin resistance gene from the ΔaspB single deletion mutant. The specialized res-hyg-res gene cassette employed to construct the aspB knockout mutant contains the specific DNA binding sites (res) for a site-specific γδ –resolvase, the product of the tnpR gene of E. coli transposon Tn1000, cloned under the expression control of mycobacterial promoter hsp60. Excision of the hygromycin resistance cassette was achieved by transient expression of the γδ –resolvase from the pYUB870 plasmid, with subsequent loss of the sacB14  expressing pYUB870 plasmid via negative selection via plating onto medium containing sucrose [96]. The second mutation, ΔaspC, was introduced into the unmarked ΔaspB mutant using the original hygromycin selection as in the construction of the single deletion mutant.  2. 6. Drug susceptibility testing MTT and resazurin assays were used to determine whether expression of aspBMtb or aspCMtb or deletion of aspBMsm or aspCMsm affected the drug susceptibility of M. smegmatis. Stationary phase cultures were diluted to an A600 nm of 0.005 and added to microtiter plates containing 2-fold dilutions of antibiotics and incubated at 37°C for 48 hours. At 48 hours MTT reagent was added. Cultures were incubated at 37°C for 2 hours before addition of solubilising solution (10% SDS). For E. coli, cultures were seeded into the wells at OD600nm =0.001 and incubated at 37°C for 6 hours. MTT reagent was added followed by incubation for 30 min at 37°C prior to addition of solubilisation solution. The absorbance at 570nm was measured to quantify the amount of purple formazan precipitate formed. The minimal inhibitory concentration (MIC) was set as the lowest concentration of antibiotic to give a reduction of more than 90% of OD570nm reading of the untreated control MIC. For resazurin assay, the initial setup was essentially the same but the cultures were incubated for 24 hours before the addition of the resazurin reagent followed by an additional incubation of 24 hours at 37°C. A change from blue to pink indicated growth of bacteria, and the MIC was defined as the last concentration at which resazurin remained blue.  2. 7. Growth kinetics M. smegmatis strains were seeded into the microtitre wells of honeycomb plates with antibiotics (for susceptibility testing) or intermediary metabolites (for nutritional growth requirement determination) and incubated in a Bioscreen C kinetic growth reader at 37 °C with constant shaking at maximum amplitude. Growth was monitored by the machine by measuring OD600nm.  2. 8. Antibiotic kill curve Time-kill curves were performed in shaken flask cultures in 7H9 media at 37°C. Wild-type and mutant ΔaspC was inoculated to give a starting OD600nm of 0.001 and grown to early exponential phase (OD600nm = 0.5) at which point a bactericidal concentration of clarithromycin was added at 15 μg/ml (about 30 times MIC). Surviving bacteria were counted by cfu plating every 2 hours onto 7H10 (10% OADC). Colonies were counted after 48 h incubation at 37°C.  15  2. 9. RNA isolation Total mRNA from M. smegmatis was isolated by a standard technique [100]. Briefly, M. smegmatis was grown to early exponential phase (OD600nm = 0.4-0.6). Some samples were treated with tetracycline at 0.8 μg/ml (equivalent to four times the MIC for wild-type) for 1 hour at 37°C. Guanidine lysis buffer (5 M guanidine thiocyanate, 17 mM sodium lauroyl sarcosinate, 28.5 mM tri-sodium citrate, 0.5% (v/v) Tween 80, 0.7% (v/v) 2-mercaptoethanol) was added to the culture at a ratio of 4 to 1. Bacterial cells were then pelleted by centrifugation (4000 rpm for 15min at 4°C) and disrupted in 1 ml of TRIzol reagent (Life Technologies) with 0.1 mm silica beads in a FastPrep-24 bead-beater (MP Biomedicals). Cell debris was removed by centrifugation at full speed in a microcentrifuge for 10 min and the lysate was extracted with an equal volume of chloroform/isoamyl alcohol 24:1. Total nucleic acid was precipitated using 30 mM sodium acetate in isopropyl alcohol. Nucleic acid samples were then treated for 30 min at 37 °C with Turbo DNase (Ambion) and final RNA clean-up was done using RNA spin mini columns (GE Healthcare) according to the manufacturer's instructions.  2. 10. Quantitative real-time PCR cDNA was synthesized from 250 ng of total mRNA using qScript cDNA Synthesis kit (Quanta Biosciences) as per manufacturer’s instructions. Each 25-μl reaction mixture included cDNA from 25 ng of RNA, 1 μM of each primer pair, and Perfecta SYBR Green SuperMix (Quanta Biosciences). The reactions were monitored in a CFX96 real-time System (Bio-Rad Laboratories Inc.). Thermocycling parameters used were 95 °C for 3 min, and 35 cycles of 95°C for 30 s, 55°C for 30 s, 72°C for 30 s. Amplification specificity was assessed by melting curve analyses. Oligonucleotides for amplification of M. smegmatis whiB7, aspB and aspC are listed in Table 2. The degrees of expression change were normalized to an internal control, mysA, using primers prAB49 and prAB50 and the significance of differential gene expression in each sample, relative to the non-treated wild-type control was calculated using the 2-ΔΔCt method. Analysis was performed on triplicate biological samples that were assayed in duplicate. Genomic DNA contamination was measured by real-time PCR of RNA not treated with reverse transcriptase and found to be insignificant.  2. 11. NAD-NADH pool isolation and NAD cycling assay NAD+ and NADH levels were measured by a sensitive cycling assay [101,102,103]. M. smegmatis strains were grown to early exponential phase (OD600nm = 0.2-0.4). Some cells were treated with tetracycline (0.8 μg/ml) for 1 hour before extraction of NAD+ and NADH. Cells were pelleted by centrifugation and split in two for extraction with 0.2 M HCl (for NAD+ determination) or 0.2 M NaOH (for NADH 16  determination). The samples were incubated at 50°C for 10 min and then cooled on ice. The cell extracts were neutralized by adding an equal volume of 0.1 M NaOH (for NAD+ determination) or 0.1 M HCl (for NADH determination). Cell debris was removed via centrifugation and the supernatant used in the cycling assay immediately. In a microtiter plate, 10 μl of extract was added to 180 μl of reagent solution consisting of 110 mM Bicine buffer, pH 8.0, 11% (v/v) ethanol, 4.4 mM EDTA, pH 8.0, 0.47 mM MTT, and 3.7 mM phenazine ethosulfate. The assay was initiated by the addition of 10 μl alcohol dehydrogenase (1 mg/ml in 0.1 M Bicine buffer, pH 8.0) and the absorbance at 570nm was read at intervals of 20 seconds. The concentrations of NAD+ and NADH in the extracts were extrapolated from a standard curve constructed using 0.01-0.05 mM solutions of NAD+ or NADH. The change in absorbance (∆A570nm/min) reflects the concentration (mM) of NAD+ or NADH in extracts. The final concentration of NAD+ and NADH was expressed as micromolar per gram of dry weight of cells (μM/g). All determinations were based on triplicate biological experiments.  2. 12. Hydrogen peroxide and menadione disc assay Sensitivity to hydrogen peroxide was assessed by disc diffusion. Twenty ml of 7H9 agar [0.2% (v/v) glycerol and 1.5% (w/v) agar] were poured into standard 100 mm diameter petri plates. Plates were overlaid with 10 ml of 7H9 top agar [(0.2% (v/v) glycerol and 0.5% (w/v) agar] inoculated with stationary phase culture to 105 cfu/ml. Five μl of 30% hydrogen peroxide solution or 50 mM menadione was applied to a blank assay disc (6 mm diameter, BD) placed in the centre of the plate. Zones of inhibition were measured after incubation at 37°C for 48 hours.  2. 13. Preparation of cell lysate The catalase activity in whole cell lysate was determined in untreated and hydrogen peroxide treated cultures. Cultures were grown to exponential phase (OD600 = 0.5) and subjected to 3.6 mM and 20 mM hydrogen peroxide treatment or PBS control for 1 hour at 37°C. Cells were harvested by centrifugation (5000 rpm for 25 min at 4°C) and the cell pellet was washed three times in dH2O and resuspended in 0.1 mM phenylmethylsulfonyl fluoride, 50 mM triethylamine (pH 7.8), concentrating the cells 200-fold. The cells were lysed with 0.1 mm silica beads in a FastPrep-24 bead-beater (MP Biomedicals). The glass beads and cell wall were pelleted by centrifugation at 10,000 X g for 30 min at 4°C. The supernatant was transferred to a fresh tube and its protein concentration was determined using Pierce Micro BCA Protein Assay Kit according to manufacturer’s instructions.  17  2. 14. Catalase assay The determination of catalase activity was based on an assay previously described by Beers and Sizer [104]. Briefly, 10 μL of the lysate was added to 890 μL of 100 mM potassium phosphate, pH7.0. 100 μL of 100 mM H2O2 was added and the decrease in absorbance at 250 nm was monitored for 2 min using a Varian Cary 5000 spectrophotometer. The linear part of the curve was used to quantitate the rate of H2O2 removal using molar extinction coefficient ε250=16.69 M-1cm-1. One unit of catalase was defined as the amount of lysate that catalysed the decomposition of 1 μmol H2O2 per min at 25°C. The catalase activity was expressed as specific activity, units per mg total protein.  2. 15. Phylogenic analysis of AspBMtb and AspCMtb Protein sequences were obtained from Uniprot (www.uniprot.org) and aligned using Clustal X2, followed by manual adjustments [105]. Phylogenetic trees were constructed using the Neighbor-joining algorithm in Mega 5 [106].  2. 16. Cloning of aspB and aspC in pGEX4T-1 The genes aspB and aspC from M. tuberculosis H37Rv were cloned from genomic DNA using primers BOO-78 and BOO-79 (aspB) and BOO-80 and BOO-81 (aspC). The PCR fragments were ligated into pGEX4T-1 (GE Healthcare) pre-digested with BamH1 and EcoR1 for expression of recombinant fusion proteins with an N-terminal Glutathione S-transferase (GST) tag. The recombinant fusion proteins contained an engineered thrombin cleavage site for the removal of the tag after affinity purification. The expression vectors were transformed into BL21 E. coli; transformants with ampicillin resistance were selected. A single colony was inoculated into selective media and grown overnight with shaking at 37°C. The overnight culture was diluted 200-fold into 2xYT media containing ampicillin (100 μg/ml) and grown at 37°C until OD600nm 0.4 was reached. Expression was induced with 0.25 mM IPTG for 17 hours at room temperature. Cells were harvested by centrifugation (4000 rpm, 15 min) and stored at -80°C.  2. 17. Purification of GST-tagged AspBMtb and AspCMtb Purification of GST-tagged AspB and AspC was based on the method described by Frangioni and Neel [107]. Cells were resuspended in STE buffer (10mM Tris, pH 8.0, 150 mM NaCl, 1mM EDTA) with 1.5% sarkosyl and 2% triton. Cells were lysed by bead beating in a FastPrep-24 (MP Biomedicals) for 3 rounds of 30 seconds at level 5. The lysate was centrifuged for 40 min at 10 000 g to remove unbroken cells and 18  inclusion bodies. The supernatant was then loaded onto a Glutathione Sepharose 4B column (GE Healthcare) and the column was washed with STE buffer. Purified GST-AspBMtb and GST-AspCMtb were eluted from the column using 10 mM reduced glutathione in STE buffer (Figure 24). De-salting and buffer exchange into 10 mM Tris, pH8.0 was performed using centrifugal filter units (Amicon) with a 30 kDa MWCO. The purity of the purified recombinant fusion protein was assessed using SDS-PAGE.  2. 18. Thrombin cleavage of GST-tagged AspBMtb and AspCMtb Removal of the affinity tag was performed at room temperature for 10 min at a 100:1 molar ratio of the protein with human α-thrombin (Haematologic Technologies Inc.). At the end of the incubation, thrombin was inactivated using 1 mM PMSF followed by a second round of Glutathione Sepharose affinity chromatography.  2. 19. Cloning of aspB and aspC in pET expression vectors DNA fragments encoding the genes aspB Rv3565 (BOO-106 and BOO-107) and aspC Rv0337c (BOO-108 and BOO-109) were cloned from M. tuberculosis genomic DNA, using primers with engineered NdeI and BamH1 recognition sequences. The PCR products were directionally cloned into pET19b for heterologous expression of recombinant AspB and AspC with an N-terminal hexa-histidine (His) fusion tag. Additionally the aspC gene was amplified without the stop codon using primers BOO-108 and BOO110 and was directionally cloned between NdeI and KpnI sites of pET30b to attach a His tag at the Cterminus of the recombinant protein. Both pET19b and pET30b included an engineered enterokinase cleavage site between the His-tag and recombinant protein.  2. 20. Expression of recombinant His-tagged AspBMtb and AspCMtb The expression vectors were electroporated into BL21 (DE3) or Rosetta™ 2 (Novagen®) E. coli expression strains. The transformants were plated onto LB agar plates containing ampicillin (100 μg/ml), kanamycin (30 μg/ml) and chloramphenicol (34 μg/ml) whenever necessary. For expression with chaperone proteins, pET30b::aspC was co-transformed with one of the following: pGro7, pTf-2, pG-KJE8, or pTf16 (Takara Bio Inc.) and plated on kanamycin and chloramphenicol selection plates. Starter cultures were made from a single colony inoculated into LB broth containing antibiotics for plasmid maintenance and grown overnight at 37°C with shaking. The starter culture was diluted 200-fold into 1 litre 2xYT media, grown at 37°C with shaking until an OD600nm of ~0.4, and induced using 0.1 mM 19  IPTG. To induce the expression of chaperone proteins, L-arabinose (1 mg/ml) and/or tetracycline (2 ng/ml) were used. Induction proceeded at room temperature overnight (~17 h). The cells were harvested by centrifugation at 4000 g for 15 min and frozen at -80°C until further use.  2. 21. Purification of recombinant His-tagged AspBMtb and AspCMtb The cell pellets were thawed on ice and resuspended in lysis buffer containing 50 mM NaH2PO4 pH 7.8, 300 mM NaCl, 0.06mM pyridoxal 5′-phosphate hydrate, 2% Triton and 1 mM DTT. The cells were lysed by five 30 second cycles of sonication on ice. The lysate was clarified by centrifugation at 10 000 g for 40 min. The recombinant protein was purified using a Ni-NTA Agarose (Qiagen) column according to manufacturer’s protocol. Briefly, the clarified supernatant was loaded onto the column which was then washed with 10 column volumes of wash buffer (50 mM NaH2PO4 pH 8.0, 300 mM NaCl, 100 mM imidazole). The recombinant protein was eluted using 2 column volumes of elution buffer (75 mM HEPES pH 7.8, 150 mM NaCl and 250 mM imidazole). The recombinant His-tagged AspB protein was observed to have a molecular weight of ~45 kDa and the recombinant His-tagged AspC protein had an observed molecular weight of ~51 kDa, consistent with sizes predicted by their nucleotide sequences. The eluted recombinant proteins were dialyzed in 2 l of 50 mM HEPES pH 7.8 overnight at 4°C. The volumes of the purified recombinant proteins were reduced using PEG 8000. The His tag from both recombinant proteins was removed using the engineered enterokinase site. The purity of the recombinant proteins was examined by SDS PAGE.  2. 22. Removal of His tag from AspBMtb The His tag was removed by incubating a 6000:1 molar ratio of AspB with enterokinase (NEB Inc.) for 10 h at room temperature. The reaction was stopped by the addition of 1.5 mM PMSF followed by the removal of the enterokinase with a Benzamidine Sepharose (Sigma) column. Centrifugal filter units (Amicon) with a 30 kDa MWCO were used for concentration and buffer exchange into 10 mM HEPES buffer pH 7.8. The removal of the His tag was monitored via SDS-PAGE and it was determined that >95% of the fusion tags had been cleaved. Purified AspB aliquots were frozen with liquid nitrogen and stored at -80°C until use.  20  2. 23. Cloning and expression of AspCMtb without fusion tags The aspC gene fragment was cut out from pET19b::aspC using NdeI and BamH1 and cloned into pET30b creating pET30b::aspC1 for expression of AspC without any fusion tags. The plasmid was transformed into Rosetta 2™ E. coli and kanamycin-resistant colonies were selected. Expression of AspC was analogous to steps described in section “Expression of recombinant His-tagged AspB and AspC” (Section 2. 21) except AspC expression was induced with 1 mM IPTG for 3 h at 37°C or 0.05 mM IPTG overnight at 15°C. The expression and solubility of AspC was monitored using SDS PAGE.  2. 24. Cloning and expression of AspCMtb in M. smegmatis The aspC gene fragment was removed from pET30b::aspC using NdeI and KpnI and cloned into pYUB1062 [97] creating pYUB1062::aspC. The expression vector for C-terminus His tagged AspC fusion protein was electroporated into mc2 4517 M. smegmatis expression strain [97] and plated onto 7H10 hygromycin selection plates. A starter culture was prepared by inoculating a single colony into 10 ml 7H9 containing hygromycin B and the culture was grown at 37°C for 24 h. 1 ml of the primary culture was inoculated into 50 ml of the same 7H9 media and was grown at 37°C to OD600nm 0.6-0.8. Subsequently expression of AspC was induced with 0.2% (w/v) acetamide and expression proceeded at 37°C for 22 h. After 22 h, the cells were harvested by centrifugation (4000 rpm for 15min at 4°C), resuspended in lysis buffer II (50 mM NaH2PO4 pH 7.8, 300 mM NaCl) and lysed with 0.1 mm silica beads in a FastPrep-24 bead-beater (MP Biomedicals). Expression and solubility of AspC was observed using SDS PAGE.  2. 25. Aspartate aminotransferase assay The aspartate aminotransferase activity of purified recombinant AspB and AspC was assayed by coupling the aspartate aminotransferase reaction with malate dehydrogenase, and measuring the oxidation of NADH spectrophotometrically at 340 nm [108] [109]. Briefly, the final substrate mixture contained 200 mM HEPES buffer, pH 7.5; 10 mM aspartate; 10 mM α-ketoglutarate; 0.1 mM NADH, 2 μM pyridoxal 5′phosphate; and 0.5 μg/ml malate dehydrogenase. Reaction rates at 25°C were determined by following the decrease in absorbance at 340 nm with a Varian Cary 5000 spectrophotometer. Calculations were based on extinction coefficient for NADH (ε340 = 6220 M-1cm-1). One unit of AAT activity equals 1 μmol of product generated per minute at 25°C. Steady-state kinetic parameters, Km values were determined for aspartate and α-ketoglutarate in the assay system described above. Initial reactions rates were measured by varying the α-ketoglutarate 21  concentration while aspartate concentration was held constant at 10mM and vice versa as aspartate concentration was varied as α-ketoglutarate was fixed at 1.5 mM. The concentration of the enzyme used in each assay was 130 nM. Curves were fit using the equation for a substituted-enzyme mechanism in LEONORA.  22  3. Results 3. 1. Expression of whiB7 upregulates aspC The transcription factor encoded by whiB7 is required for the induction of genes that encode conventional resistance determinants, including an antibiotic modifying enzyme (eis [110,111]), a target modifying enzyme (erm37[12,112]), and antibiotic transporter (tap [113]) [29]. Intriguingly, Morris et al. reported that some primary metabolic genes were also under whiB7 control. Further analyses of the microarray data suggested that expression of two homologous genes annotated as aspartate transaminases (aspB, Rv3565; aspC, Rv0337c) were affected by whiB7 expression. Comparing expression levels between whiB7 overexpressing M. tuberculosis and wild-type, aspB was downregulated three-fold (ratio of 0.34) and aspC was upregulated ratio of 3.64) (unpublished data, Thompson lab). Members of this family of proteins catalyze a reversible transamination reaction involving interconversions of α-keto acids and their partnered amino acids [114], thereby serving as central hubs of metabolism. Although these enzymes may catalyze reactions involving multiple amino acid/ α-keto acid partners, they are often called glutamate-oxaloacetate transaminases (Figure 3). Figure 3. Reaction catalyzed by aspartate aminotransferases  Figure 3. Reaction catalyzed by aspartate aminotransferases. The genes aspB and aspC are both annotated to be aspartate aminotransferases. Aspartate aminotransferases catalyze a pyridoxal5’phosphate-dependent (PLP-dependent) reversible reaction that transfers an amino group from glutamate to α-ketoglutarate forming aspartate and oxaloacetate [115].  Initial experiments were designed to explore the interdependencies of aspB, aspC, and whiB7 expression and how this might relate to metabolism and intrinsic drug resistance. whiB7Mtb (Rv3197a) and whiB7Msm (MSMEG_1953) were cloned into pMV361 to provide constitutive expression in M. smegmatis. Since whiB7 orthologs in M. tuberculosis and M. smegmatis share 75% amino acid identity, M. smegmatis was adopted as a convenient surrogate system to study the regulation of gene expression of whiB7, aspB, and aspC. mysA (MSMEG_2758) encoding the primary sigma factor σA, was used as an internal control for normalization of aspB, aspC, and whiB7 expression levels. Rates of aspB and aspC transcription were 23  measured by qRT-PCR in wild-type when whiB7Mtb or whiB7Msm were constitutively expressed from the heat-shock protein (hsp) promoter [94,116,117]. Relative to the wild-type strain transformed with the empty vector, constitutive transcription of whiB7Mtb and whiB7Msm resulted in a 7.5-fold increase in aspC expression (Figure 4). Under these conditions aspB expression was not significantly changed by whiB7Mtb. This result suggests that aspC, but not aspB, expression may be controlled by transcription of whiB7.  3. 2. Expression of whiB7 and aspC is upregulated by tetracycline In M. tuberculosis (and Streptomyces lividans), whiB7 is induced by various antibiotics including tetracycline [29]. To validate the use of M. smegmatis as a surrogate model system, cultures were treated with tetracycline at 0.8 μg/ml (about four-times the MIC of tetracycline in the wild-type strain) for 1 hour and the expression of whiB7Msm was measured using qRT-PCR. This treatment induced expression of whiB7Msm over 1000-fold (Figure 5A). The expression of aspC in the same tetracyclinetreated RNA samples was induced 4-fold (Figure 5B). To more clearly understand the dependence of aspB and aspC expression on whiB7 and antibiotics, transcription was measured in both wild-type and ∆whiB7 after tetracycline treatment. The expression of aspC in untreated ∆whiB7 cultures was slightly (about 50%) depressed relative to the wild-type strain (Figure 5B). While aspC was induced more than four-fold in wild-type by tetracycline, this did not occur in the tetracycline-treated ∆whiB7 strain (Figure 5B). These observations demonstrated whiB7dependent upregulation of aspC in response to tetracycline. Parallel experiments showed that expression of aspB was not significantly affected by tetracycline treatment or whiB7 deletion (Figure 5B)  24  expression of aspB/aspC (Log2 fold change compared to WT)  Figure 4. Expression of aspC was upregulated by whiB7  16  aspB expression aspC expression  8 4 2 1 0.5  hsp-whiB7Msm  hsp-whiB7Mtb  Figure 4. Expression of aspC was upregulated by whiB7. Expression of aspB and aspC in M. smegmatis that constitutively expresses whiB7Mtb or whiB7Msm from hsp60 promoter was measured by qRT-PCR. Constitutive expression of whiB7Mtb increased the expression of aspC seven-fold compared to wild-type. The expression of aspB was not significantly affected. Mean data from 3 biological experiments are reported.  Figure 5. Expression responses of whiB7, aspB, and aspC to tetracycline  B  2048 1024 512 256 128 64 32 16 8 4 2 1 WT + tetracycline  aspB + tetracycline  aspC + tetracycline  expression of aspB/aspC (Log2 fold change compared to WT)  whiB7 expression (Log2 fold change compared to WT)  A  8  aspB expression aspC expression  4 2 1 0.5  whiB7 untreated  wild-type + tetracycline  whiB7 + tetracycline  Figure 5 Expression responses of whiB7, aspB, and aspC to tetracycline. (A) Wild-type and mutants ∆aspB and ∆aspC were treated with 0.8 μg/ml tetracycline for 1 h and whiB7 expression levels compared to untreated wild-type. whiB7 was strongly upregulated in response to tetracycline and the deletion of aspB did not affect this response. The expression of whiB7 in response to tetracycline treatment was significantly reduced in ∆aspC. (B) Expression of aspC in wild-type and ∆whiB7. Expression of aspC increased in response to tetracycline treatment. Deletion of whiB7 downregulated aspC in untreated samples and abrogated upregulation of aspC in response to tetracycline. Mean data from 3 biological experiments are reported. 25  3. 3. Optimal whiB7 induction by tetracycline requires aspC Cultures of mutant strains ∆aspB and ∆aspC were tetracycline treated and relative whiB7 expression levels were determined using qRT-PCR to assess whether those genes affected whiB7 expression in response to tetracycline. As shown in Figure 5A, the ∆aspB mutant strain had the same whiB7 tetracycline induced upregulation as the wild-type while the whiB7 expression in the ∆aspC mutant was significantly lower (340-fold versus over 1000-fold in wild-type). This suggested that whiB7 upregulation in response to tetracycline is enhanced by aspC.  3. 4. Constitutive expression of aspB downregulates whiB7 and aspC in tetracycline-induced cultures The M. tuberculosis aspBMtb and aspCMtb genes were cloned and expressed in M. smegmatis from an integrative vector with constitutive transcription from hsp promoter (pMV361:: aspBMtb and pMV361::aspCMtb) generating strains that constitutively expressed aspBMtb and aspCMtb (hsp-aspBMtb and hsp-aspCMtb). To determine whether constitutive aspB and aspC expression affected expression of whiB7 in response to antibiotics, whiB7 transcripts were measured in cultures constitutively expressing these genes with and without tetracycline induction. In untreated wild-type cultures, constitutive expression of aspB (hsp-aspBMtb) reduced expression of whiB7 about 95% and the strong tetracycline induced upregulation of whiB7 expression was eliminated (Figure 6A). Measurement of aspC in these samples revealed a similar trend; upregulation of aspC in response to tetracycline was not observed in the hspaspBMtb strain (Figure 6B). In contrast, expression of aspCMtb increased whiB7 expression by 3.8-fold in untreated cultures but did not significantly alter whiB7 upregulation in tetracycline treated cultures. (Figure 6A).  26  Figure 6. Interactive regulation of aspB, whiB7 and aspC  B aspC expression (Log2 fold change compared to WT)  whiB7 expression (Log2 fold change compared to WT)  A  1024 256 64 16 4 1 0.25 0.0625  wild-type  tetracycline  -  +  hsp-aspCMtb  -  +  hsp-aspBMtb  -  +  8 4 2 1 0.5  tetracycline  hsp-aspBMtb  wild-type  hsp-aspBMtb  -  +  +  Figure 6. Interactive regulation of aspB, whiB7 and aspC. (A) whiB7 expression in aspBMtb and aspCMtb constitutive expression strains. Constitutive expression of aspBMtb downregulated whiB7 expression in untreated and tetracycline treated cultures. Constitutive expression of aspCMtb increased levels of transcription in untreated cultures. (B) Expression of aspC in the hsp-aspBMtb strain. Constitutive expression of aspBMtb also prevented tetracycline-induced upregulation of aspC. Mean data from 3 biological experiments are reported.  27  3. 5. The ∆aspC mutant exhibits growth deficiencies aspB and aspC are both annotated as aspartate aminotransferases, enzymes that are central to intermediary metabolism. To test the hypothesis that aspB and/or aspC are needed for growth, knockout mutant strains of aspBMsm (MSMEG_6017) and aspCMsm (MSMEG_0688) were constructed using a mycobacterial recombineering system [95]. Growth kinetics of mutant and constitutive-expression strains were analyzed in either nutrient rich (7H9 supplemented with ADC; Figure 7A) or minimal (Proskauer Beck (PB) medium; Figure 7B). Strains ∆aspB, hsp-aspBMtb and hsp-aspCMtb grew at the same rate as wild-type M. smegmatis in both media. However, in both complete 7H9 media and minimal media PB ∆aspC exhibited growth deficiencies, including a prolonged lag phase, decreased rate of exponential growth, and decreased cell density at stationary phase. For example, in PB medium, deletion of aspC increased the doubling time approximately twofold (from 6-6.5 hrs to 12-14 hrs); rolling cultures of wild-type M. smegmatis achieved a stationary phase OD600nm of 5.0-5.5 while ∆aspC only reached OD600nm of 3.0-3.4. The lag phase of the ∆aspC mutant was generally 6 hours (in 7H9 rich medium) and 12 hours (in PB minimal medium) longer than that of wildtype (Figures 7A and 7B). Finally, ∆aspC also grew as smaller colonies on both 7H10 and PB agar plates. Transformation of ∆aspC with an integrative construct that constitutively expressed aspCMtb (pMV361::aspCMtb) fully restored growth defects on both media (results with PB cultures are shown in Figure 7C).  28  Figure 7. Growth kinetics of M. smegmatis wild-type, ∆aspB, ∆aspC, hsp-aspBMtb, hsp-aspCMtb, and ∆aspC complemented  B  A 2.0  1.0  mc2155 aspB aspC hsp-aspB Mtb hsp-aspC Mtb  2.0  OD 600  1.5  OD600  2.5  mc2155 aspB aspC hsp-aspB Mtb hsp-aspC Mtb  1.5  1.0 0.5  0.5  0.0  0.0 0  10  20  30  40 time (h)  50  60  70  80  0  20  40  60 80 time (h)  100  120  140  C 2.5 mc 2155 aspC  2.0  OD600  aspC pMV361::aspC Mtb 1.5  1.0  0.5  0.0 0  10  20  30  40  50  60 70 time (h)  80  90 100 110 120  Figure 7. Growth kinetics of M. smegmatis wild-type (black), ∆aspB (red), ∆aspC (orange), hsp-aspBMtb (green), hsp-aspCMtb (blue), and ∆aspC complemented (purple). Strains were grown in nutrient-rich 7H9 with ADC (A) and PB minimal media (B) (C). Growth curves are representative of quadruplicate wells. Bars indicate standard error.  29  3. 6. The ΔaspC is able to catalyze glutamate but not aspartate Since the use of amino acids to provide carboxylic acids or ammonia as intermediary metabolites require aminotransferases, the ability of aspB and aspC mutants to utilize aspartate and glutamate as carbon and nitrogen sources were tested. Wild-type and ∆aspB and ∆aspC mutants were inoculated into PB salts (without NH4Cl) containing glycerol (1% v/v) and 40mM glutamate or aspartate to test the ability of the strains to use these amino acids as nitrogen sources. In other cultures, glycerol was omitted in order to determine whether these amino acids could serve as sources of both carbon and nitrogen. In all these media, the aspB mutation had no growth phenotype; its growth kinetics were similar to wild-type. Growth of the ∆aspC mutant was not impaired (relative to wild-type) when cultures were grown with glutamate as nitrogen source; growth was slightly impaired in media with glutamate as the source of both carbon and nitrogen (Table 3). In contrast, the aspC mutant strain grew poorly with aspartate as its nitrogen source and could not grow using aspartate as both carbon and nitrogen sources.  3. 7. α-ketoglutarate complements ΔaspC mutant’s growth deficiencies To further investigate whether the growth deficiencies observed in ∆aspC mutant were due to a lack of one of its putative substrates, growth rates were determined in PB minimal medium supplemented with aspartate, glutamate, oxaloacetate or α-ketoglutarate. Oxaloacetate did not significantly affect the growth of the ∆aspC mutant at 20mM and 40mM (data not shown). The presence of α-ketoglutarate increased the growth rate of ∆aspC in a dose-dependent manner and allowed it to grow at a rate similar to that of wild-type at 60 mM α-ketoglutarate (Figure 8A). The prolonged lag phase of ∆aspC was not shortened by α-ketoglutarate (Figure 8A). Aspartate and glutamate did not improve the growth of ∆aspC and did not affect the growth of wild-type cultures. However, at higher concentrations of aspartate and glutamate growth of ∆aspC mutant was further delayed, showing that the mutant may have defects in catabolising these amino acids (Figure 8B).  30  Table 3. Growth properties of ΔaspB and ΔaspC mutants in cultures containing aspartate and glutamate as carbon and/or nitrogen sources. Growth was tested on minimal Proskauer Beck salts 1 supplemented with the indicated carbon and nitrogen sources at 37°C. Addition to PB salts  Strain mc2155  ΔaspB  ΔaspC  1% glycerol + 40 mM L-glutamate  +++  +++  +++  1% glycerol + 40 mM L-aspartate  +++  +++  +  40 mM L-glutamate  ++  ++  +  40 mM L-aspartate  ++  ++  -  Table 3. Growth properties of ΔaspB and ΔaspC mutants in cultures containing aspartate and glutamate as carbon and/or nitrogen sources.  Growth represented by symbols (in order of decreasing growth) +++, ++, +, - . 1  36.7 mM monopotassium phosphate, 2.4 mM magnesium sulfate, 0.4 mM sodium citrate, pH 7.4  Figure 8. Growth of the ΔaspC mutant was modulated by addition of aspartate aminotransferase substrates  B  A WT +40mM Asp  2.5  WT  2.4  aspC  WT (80 mM Glu)  aspC (20 mM -KG)  2.0  WT (40 mM Asp)  2.0  aspC (60mM -KG)  aspC  OD600  OD600  aspC (80 mM Glu)  1.6  1.5  1.0  aspC (40 mM Asp)  1.2 0.8  0.5 0.4 0.0 0  20  40  60  80 time (h)  100  120  140  0.0 0  20  40  60 time (h)  80  100  120  Figure 8. Growth of the ΔaspC mutant was modulated by addition of aspartate aminotransferase substrates. (A) Effect of α-ketoglutarate on growth kinetics of ∆aspC in PB media. The supplementation of α-ketoglutarate at 60mM into the medium restored the doubling rate to wild-type levels. (B) Addition of glutamate and aspartate at high concentrations (80 mM and 40 mM respectively) inhibits growth of ΔaspC mutant but wild-type growth in unaffected. Growth curves are representative of quadruplicate wells. Error bars indicate standard error. OD600, OD at 600 nm.  31  3. 8. Passaging the ΔaspC mutant in media containing α-ketoglutarate suppresses the growth rate defect but does not shorten lag phase The growth deficiencies of mutant ΔaspC could be partially complemented by the inclusion of αketoglutarate in the growth medium. When α-ketoglutarate was supplemented in the medium, both the reduced growth rate and lowered cell density at stationary phase of the ΔaspC mutant was restored to wild-type levels. However, the longer lag phase remained (Figure 8A). Since the prolonged lag phase could be the time required for ΔaspC mutant to make the necessary metabolic adaptations for growth on α-ketoglutarate, pre-cultures of wild-type, mutant ΔaspC and complemented strain ΔaspC pMV361::aspC were grown in 7H9 medium. The second pre-culture of the ΔaspC mutant was inoculated into the same media that also contained 40 mM α-ketoglutarate. The stationary phase cultures were washed and re-inoculated in PB (with 0.05% tyloxapol) medium with and without 40 mM αketoglutarate, and seeded into microwells in quadruplicates for growth analysis using the Bioscreen C kinetic growth reader. The mutant ΔaspC grown in PB media without α-ketoglutarate still exhibited growth deficiencies (longer lag, reduced growth rate, and lowered cell density at stationary phase) compared to wild-type (Figure 9). Addition of α-ketoglutarate to the medium suppressed only the growth rate and cell density at stationary phase but not the prolonged lag regardless of whether or not the pre-culture contained α-ketoglutarate. This result shows that the prolonged lag phase of the ΔaspC mutant is not due to a nutritional deficiency or toxicity that can be overcome with passaging in αketoglutarate-containing media.  3. 9. The ΔaspB ΔaspC double mutant is not an amino acid auxotroph The ΔaspBΔaspC double mutant was able to grow on a standard media for testing auxotrophy M9 minimal media (Figure 10A). In 7H9 liquid media, growth kinetic analyses revealed that the double mutant ΔaspBΔaspC had growth deficiencies similar to the single mutant ΔaspC (Figure 10B). The double mutant had a prolonged lag phase as with the single mutant ΔaspC, but the double mutant replicated faster (doubling rates of 5.7 hours and 4.7 hours for ΔaspC and ΔaspBΔaspC respectively). It is likely the growth deficiencies in both strains were due to the deletion of aspC but the second deletion in aspB may relieve slightly the toxicity caused by the single ΔaspC mutant. Understanding the physiological roles of aspB and aspC individually would contribute to understanding the effect of deletions of both aspB and aspC to M. smegmatis physiology.  32  Figure 9. Allowing the ΔaspC mutant to adapt to growth in media containing α-ketoglutarate did not suppress the extended lag phase growth deficiency  2.5  mc 2 155 aspC pMV361::aspC  2.0  OD600  aspC aspC (+40 mM KG)  1.5  aspC* (+40 mM KG)  1.0 0.5 0.0 0  20  40  60  80  100  120  time (h)  Figure 9. Allowing the ΔaspC mutant to adapt to growth in media containing α-ketoglutarate did not suppress the extended lag phase growth deficiency. Growth of wild-type mc2 155, mutant ΔaspC, and complemented strain ΔaspC pMV361::aspC in PB media. Constitutive expression of aspCMtb complemented the mutant’s growth deficiencies. Addition of α-ketoglutarate suppressed only the reduced doubling rate and lower cell density at stationary but not the prolonged lag phase. Passaging the mutant in α-ketoglutarate (ΔaspC*) did not shorten the lag phase of the mutant.  Figure 10. Growth kinetics of double mutant ΔaspBΔaspC  B  A 2.0  2.0  mc 2155  mc 2155 aspB  aspB aspBaspC  1.0  aspC aspBaspC  1.0  0.5  0.5  0.0  1.5  aspC  OD600nm  OD600nm  1.5  0.0 24  29  29  34  34  Time (h)  39  39  24  29  29  34  34  39  39  Time (h)  Figure 10. Growth kinetics of double mutant ΔaspBΔaspC. (A) in M9 minimal salt media and (B) in rich media 7H9 (10% ADC). The double mutant was not auxotrophic and displayed growth defects similar to mutant ΔaspC.  33  3. 10. Opacification of 7H10 by ΔaspC mutant The ΔaspC mutant generated a white, opaque, diffuse zone surrounding the colonies (opacification) when grown on Middlebrook 7H10 (with 10% OADC) (Figure 11A). The white effect can be mimicked by reducing the pH of the media.  Figure 11. Opacification of 7H10 by the ΔaspC mutant  Figure 11. Opacification of 7H10 by the ΔaspC mutant. Growth of mutant ΔaspC on 7H10 with 10% OADC yields a white opaque colour.  34  3. 11. The ΔwhiB7 mutant, ΔaspC mutant, and aspB constitutive expression strains are antibiotic sensitive Deletion of whiB7 in M. tuberculosis or M. smegmatis made them sensitive to an assortment of antibiotics including rifampicin and erythromycin [29,35]. To test whether the expression of asp genes affect antibiotic sensitivity, strains in which they were constitutively expressed (hsp-aspBMtb, hsp-aspCMtb) or inactivated (∆aspB, and ∆aspC) were subjected to antibiotic susceptibility profiling using resazurin or MTT assays. MICs were determined for fourteen antibiotics representing different structures and targets. Their MICs were compared to wild-type with no plasmid or transformed with an empty pMV361 vector. While the drug sensitivities of ∆aspB and hsp-aspCMtb strains were identical to wild-type, ∆aspC and hspaspBMtb strains were more sensitive to many antibiotics (Table 4). ∆whiB7 and hsp-aspBMtb were 4 to 16fold more sensitive than wild-type to drugs targeting the ribosome, including chloramphenicol, clarithromycin, clindamycin, roxithromycin, spectinomycin and tetracycline. The ΔaspC mutant was also more susceptible to drugs targeting protein synthesis including clindamycin, spectinomycin, clarithromycin and roxithromycin. While both hsp-aspBMtb and ∆aspC sensitivity profiles affected activities of the same set of ribosome targeting antibiotics, they exhibited a different susceptibility profile for drugs targeting RNA polymerase (rifampicin) or affecting cell envelope synthesis (isoniazid, clofazimine, and possibly triclosan). Transformation of the mutant ΔaspC with a plasmid encoding aspCMtb complemented the antibiotic sensitivity (Figure 12). Furthermore, the increased antibiotic susceptibility of mutant ΔaspC was also studied using growth kinetic analysis (Figure 14).  3. 12. Complementation of antibiotic susceptibility of the ΔaspC mutant The antibiotic susceptibility of M. smegmatis ΔaspC was partially complemented (tetracycline, clarithromycin, and spectinomycin were tested) by constitutive expression of the M. tuberculosis aspC gene as determined by MTT assay (Figure 12). Similarly, addition of α-ketoglutarate to the growth medium increased the growth rate of the mutant and suppressed the antibiotic susceptibility of ΔaspC (Figure 12). It is interesting to note that addition of α-ketoglutarate caused the strain to become more resistant to isoniazid (32-fold more resistant that wild-type (Figure 12).  35  Table 4. Antibiotic susceptibility profiles of the ΔwhiB7 mutant, hsp-aspB expression, and the ΔaspC mutant strains of M. smegmatis. The MIC of the strains including wild-type towards the antibiotics was determined using the MTT cell viability assay described in Section 2 .6. fold susceptibility relative to wild-type target  compound  ethambutol isoniazid triclosan capreomycin chloramphenicol clarithromycin clindamycin protein synthesis fusidic acid roxithromycin spectinomycin streptomycin tetracycline RNAP rifampicin other clofazimine cell wall  ∆whiB7  hsp-aspBMtb  ∆aspC  --4 -4 8 8 2 8 16 -4 ---  --4 -4 8 4 -8 4 -4 ---  -2 ---16 2-4 2-4 16 2 --8 2  Table 4. Antibiotic susceptibility profiles of the ΔwhiB7 mutant, hsp-aspB expression, and the ΔaspC mutant strains of M. smegmatis.  36  fold susceptibility of  aspC relative to wild-type  Figure 12. Complementation of antibiotic susceptibility of the ΔaspC mutant by constitutive expression of aspCMtb or addition of α-ketoglutarate  16.00  aspC aspC::hsp-aspC  4.00  aspC + -ketoglutarate  1.00 0.25 0.06 0.02  TET  CLR  SPT  INH  Figure 12. Suppression of antibiotic susceptibility of the ΔaspC mutant by constitutive expression of aspCMtb or addition of α-ketoglutarate. Mutant ΔaspC, complemented strain ΔaspC pMV361::aspCMtb, and wild-type mc2 155 were grown in the presence of 2-fold dilutions of tetracycline, clarithromycin, spectinomycin, and isoniazid in PB medium. After 2 days of growth at 37°C, the MIC was determined using MTT cell viability assay. The constitutive expression of aspCMtb or presence of α-ketoglutarate (50 mM) in the media partially suppressed the antibiotic sensitivity of the mutant.  37  3. 13. Effects of antibiotics on the growth kinetics of the ΔaspC mutant The kinetic growth response of wild-type M. smegmatis and the ∆aspC mutant to nine of the antibiotics used in the antibiotic susceptibility profiling were also assessed with growth curves. Nine antibiotics (clarithromycin, tetracycline, chloramphenicol, clindamycin, clofazimine, fusidic acid, rifampicin, roxithromycin, and spectinomycin) were tested at five concentrations each, together with control growth curves of wild-type and the ∆aspC mutant (92 growth curves in total). Treatment of cultures with antibiotics, generally affected growth both by prolonging the lag phase and slightly increasing the doubling time. With both phenomena, the time for the strain to achieve a certain cell density increased. The growth curve data was plotted to show the effect of antibiotic concentration and the time to reach half of the maximum OD600nm – as illustrated in Figure 13. Figure 13A shows growth curves of antibiotic-treated wild-type mc2155 and Figure 13B shows an alternate presentation of the same data, to show the effect of antibiotic concentration on growth delay. With this type of graph, the slope is proportional to the growth inhibition effect. The steeper the slope, the greater the increases in lag phase with higher antibiotic concentration. The growth curves of wild-type mc2155 and the ∆aspC mutant in the presence of nine different antibiotics were analyzed using this graphical representation (Figure 14). With clarithromycin, roxithromycin, rifampicin, and clindamycin, the curves representing the ∆aspC mutant have steeper slope and lay above and to the left of the wild-type. This shows that the growth kinetic inhibition (i.e. length of lag phase, doubling rate) was greater in ∆aspC mutant than in the wild-type and that the mutant experiences orders of magnitude of more growth inhibition than wildtype. With fusidic acid, spectinomycin, and clofazimine the curves representing the ∆aspC mutant has a similar slope compared to wild-type but the mutant curve still lie above the wild-type curve. This demonstrated that while the growth inhibition of ∆aspC mutant was more than wild-type, the effect of increasing concentrations was similar. When there was no change in susceptibility in the ∆aspC mutant compared to wild-type, the curves overlap, as is the case with tetracycline and chloramphenicol. The growth curve data correlated with the results of the MTT assay for MIC determination (Table 4). When the mutant growth curve had a steeper slope than the wild-type curve (clarithromycin, roxithromycin, rifampicin), the fold susceptibility was higher (eight- to sixteen-fold more susceptible as compared to wild-type). A slight increase in susceptibility (two- to four-fold more susceptible as compared to wild-type) corresponded to the mutant curve having a similar slope to the wild-type curve but with the mutant curve lying above the wild-type curve (clindamycin, fusidic acid, and spectinomycin). The growth curve analysis once again showed that deletion of aspC caused the strain to be more susceptible to a number of antibiotics. In addition, this analysis revealed that the susceptibility of the ∆aspC mutant increase was due the increased growth lag, akin to the effect of oxaloacetate and antibiotics on the growth of wild-type (Table 5 and Figure 16).  38  Figure 13. The time required for wild-type mc2155 to adapt and grow in media containing antibiotics is dependent on antibiotic concentration  A  B 150  2.0  untreated antibiotic 2 g/ml  125  antibiotic 1 g/ml  100  antibiotic 0.5 g/ml  1.0 x  x  x  x  x  0.5  0.0  antibiotic 0.25 g/ml  time (h)  OD600  1.5  75 50 25  0  15 30 45 60 75 90 104 119 133  time (h)  0 0.0  0.5  1.0  1.5  2.0  antibiotic conc  Figure 13. The time required for wild-type mc2155 to adapt and grow in media containing antibiotics is dependent on antibiotic concentration. (A) The time for recovery from stationary phase was increased by inoculation into various concentrations (0.25 – 2.0 μg/ml) of antibiotics. The growth at OD600nm was monitored for 6 days. The time spent in lag phase was proportional to the concentration of the antibiotic. The point at which half-maximum OD600nm (about 0.83) is marked by a red ‘x’ on each growth curve (B) The data from growth curves is represented as time to reach half-maximum OD600nm versus antibiotic concentration.  39  Figure 14. The difference between wild-type and the ΔaspC mutant growth kinetics in media containing antibiotic  aspC is 8-16 fold more susceptible  125  WT  WT aspC  125  NG  WT aspC  150  aspC  100  75 50  time (h)  125  100  time (h)  time (h)  Rifampicin  Roxithromycin  Clarithromycin 150  75 50  100 75 50  25  25  25 0  0  0 0.0  0.2  0.4  0.6  0.8  0  1.0  2  concentration ( g/ml)  4  6  0  8  5  10  15  20  25  30  concentration ( g/ml)  concentration (g/ml)  aspC is 2-4 fold more susceptible Clindamycin  Fusidic acid WT aspC  150  time (h)  time (h)  75 50  100  WT aspC  125  125 100  Spectinomycin  150  100 75 50  0  0  0  2  4  6  8  aspC  60 40 20  25  25  WT  80  time (h)  NG  0 0  10  concentration ( g/ml)  20  30  40  50  60  0  10  concentration (g/ml)  20  30  40  50  60  concentration (g/ml)  aspC is not more susceptible Clofazimine WT  NG  aspC  150  WT aspC  150  125  NG  75  100 75  100 75  50  50  50  25  25  25  0 0  2  4  6  8  10  12  concentration ( g/ml)  14  16  0 0.0  aspC  125  time (h)  100  WT  150  125  time (h)  time (h)  Chloramphenicol  Tetracycline  NG  0 0.5  1.0  1.5  concentration (g/ml)  2.0  0  4  8  12  16  20  24  28  32  concentration (g/ml)  s 40  Figure 14. The difference between wild-type and the ΔaspC mutant growth kinetics in media containing antibiotics. Stationary phase cultures of wild-type and mutant ΔaspC were inoculated into media containing various concentration of nine antibiotics (clarithromycin, tetracycline, chloramphenicol, clindamycin, clofazimine, fusidic acid, rifampicin, roxithromycin, and spectinomycin). Growth of the bacteria was monitored at OD600nm for six days. The time for each cell growth to reach half-maximum OD600nm of its respective strain was plotted against antibiotic concentrations. No growth is indicated on the second segment of y-axis marked ‘NG’. The difference between susceptibility of mutant ΔaspC and wild-type towards an antibiotic determined by MTT MIC determination is indicated by border colors around each plot. Orange (8 to 16 fold more susceptible), brown (2 to 4 fold more susceptible), grey (not more susceptible)  41  3. 14. Differential killing of the ΔaspC mutant by clarithromycin Time-kill curve experiment was performed to evaluate the susceptibility of mutant ΔaspC compared to wild-type mc2155. The mutant ΔaspC was previously observed to have increased susceptibility to clarithromycin using both the MTT cell viability assay (Table 4) and growth curve analyses (Figure 14). Here the ∆aspC strain exhibited a slower rate of death compared to wild-type (Figure 15). After 24 hours, wild-type mc2155 had decreased 5 log10 cfu/mL while mutant ΔaspC only had a decrease of 3 log10 cfu/mL suggesting that mutant ΔaspC is more resistant to killing by clarithromycin.  Figure 15. Differential killing of the ΔaspC mutant by clarithromycin  10 1 1  wild-type aspC  10 1 0  cfu/ml  10 9 10 8 10 7 10 6 10 5 10 4 0  2  4  6  8  10  24  time (h) Figure 15. Differential killing of the ΔaspC mutant by clarithromycin. Logarithmic phase cultures were treated with clarithromycin (15μg/ml). Samples were taken every 2 hours for cfu plating. The ∆aspC maintained higher cell viability compared to wild-type at every time point. Data shown represents results from 2 biological experiments.  42  3. 15. Oxaloacetate and α-ketoglutarate modulate the rate of recovery from stationary phase growth and adaptation to bacteriostatic drugs In addition to the classical lag phase observed after inoculation of stationary phase cultures into fresh medium, M. smegmatis also displayed a lag phase in response to bacteriostatic antibiotic treatments. When inoculated from antibiotic-free cultures into media containing low concentrations of bacteriostatic antibiotics (tetracycline, spectinomycin, or clarithromycin), onset of growth of M. smegmatis was delayed (Figure 16A). In both cases, α-ketoglutarate in the medium consistently decreased the delay caused by antibiotics (Figure 16B-D). In contrast, oxaloacetate consistently prolonged the growth delay of the bacteria due to the presence of antibiotics (Figure 16B-D). Data are summarized in Table 5 and explained in more detail below. Wild-type cultures growth in PB medium supplemented with 40 mM α-ketoglutarate consistently entered exponential phase 4 to 5 hours earlier than in medium without α-ketoglutarate (three independent experiments are shown in Figure 16B-D; Table 5). Conversely, growth in medium supplemented with 40 mM oxaloacetate delayed exponential phase by 24 to 27 hours (Figure 16B-D; Table 5). Based on these observations, we hypothesized that the drug sensitivity of ∆aspC might be related to changes in intracellular concentrations of oxaloacetate and α-ketoglutarate. To test this hypothesis, we analyzed the effects of oxaloacetate and α-ketoglutarate on the growth inhibition activity of antibiotics. We used growth curve analysis to observe the growth inhibition of antibiotics alone and in combination with oxaloacetate or α-ketoglutarate. Because ∆aspC exhibited sensitivities to antibiotics of diverse classes we also chose to use drugs representing different ribosomal targets in order to determine whether the effects of oxaloacetate and α-ketoglutarate are also similarly nonspecific with respect to the class of antibiotic. Individually tetracycline (0.125 μg/ml), clarithromycin (0.25 μg/ml) and spectinomycin (4 μg/ml) delayed the growth of wild-type from 36 hours to 51, 63, and 64 hours respectively (Table 5, Figure 16A). When oxaloacetate was added to the medium at 40 mM the growth of M. smegmatis was delayed further by 64, 96, and 118 hours for tetracycline, clarithromycin, and spectinomycin respectively as compared to media without oxaloacetate. Conversely, αketoglutarate promoted growth with and without antibiotic treatment. When α-ketoglutarate was added to the medium, the growth delay of antibiotics was decreased to 44, 36, and 53 hours for tetracycline, clarithromycin and spectinomycin respectively. In summary, oxaloacetate potentiated the growth inhibition actions of antibiotics whereas α-ketoglutarate increased the fitness of the bacterium, antagonizing the inhibition of antibiotics.  43  Figure 16. Effect of α-ketoglutarate and oxaloacetate supplementation on growth kinetics and antibiotic inhibition of growth in wild-type M. smegmatis  A  B  2.5  no antibiotic  2.5  no antibiotic/supplement  2.0  oxaloacetate TET  1.5  TET+ -ketoglutarate TET+ oxaloacetate  TET  2.0  -ketoglutarate  SPT  OD600  OD600  CLR  1.5 1.0 0.5 0.0 20  1.0 0.5  40  60  80 time (h)  100  120  0.0 20  140  40  60  80 time (h)  100  120  140  D  C 2.5  2.5  no antibiotic/supplement  no antibiotic/supplement -ketoglutarate  -ketoglutarate  2.0  2.0  oxaloacetate  oxaloacetate CLR  1.5  SPT+ -ketoglutarate SPT+ oxaloacetate  1.0 0.5 0.0 20  OD600  OD600  SPT  CLR+ -ketoglutarate CLR+ oxaloacetate  1.5 1.0 0.5  40  60  80 time (h)  100  120  140  0.0 20  40  60  80 time (h)  100  120  140  Figure 16. Effect of α-ketoglutarate and oxaloacetate supplementation on growth kinetics and antibiotic inhibition of growth in wild-type M. smegmatis. (A) Growth of wild-type mc2155 in PB medium with ¼fold MICs of tetracycline (0.125 μg/ml, TET, yellow), spectinomycin (4.0 μg/ml, SPT, purple), or clarithromycin (0.25 μg/ml, CLR, orange). (B-D) α-ketoglutarate and oxaloacetate were added into PB at 40 mM. Oxaloacetate (tan) delayed growth of M. smegmatis and potentiated the effects of tetracycline (B), spectinomycin (C), and clarithromycin (D) (blue). The onset of growth was accelerated by the addition of α-ketoglutarate (grey) and the growth delay due to tetracycline, spectinomycin, and clarithromycin was diminished with α-ketoglutarate (green). Growth curves are representative of quadruplicate wells. Error bars indicate standard error. OD600, OD at 600 nm. Data are summarized in Table 5.  44  Table 5. Effect of antibiotics and putative aspartate transaminase substrates on time of recovery from stationary phase growth arrest. The table summarizes the time required to establish exponential growth defined at OD600nm = 0.5 in PB media supplemented with oxaloacetate and α-ketoglutarate at 40mM and/or ¼-fold MIC sub-inhibitory concentrations of tetracycline (0.125 μg/ml), clarithromycin (0.25 μg/ml) or spectinomycin (4 μg/ml). Time to reach OD600nm = 0.5 (hours) Antibiotic (μg/ml)  Proskauer Beck  PB + a-ketoglutarate  PB + oxaloacetate  no treatment  36  31  58  tetracycline (0.125)  51  44  64  clarithromycin (0.25)  63  36  96  spectinomycin (4)  64  53  118  Table 5. Effect of antibiotics and putative aspartate transaminase substrates on time of recovery from stationary phase growth arrest  45  3. 16. Deletion of aspC causes a shift in intracellular redox homeostasis By analogy to other bacteria, and metabolic pathways predicted by genome sequences, aspartate is likely to be a precursor for the nicotinamide adenine dinucleotide (NAD+) biosynthetic pathway in mycobacteria [118]. WhiB7 also plays a role in redox homeostasis and influences expression of aspB and aspC. Based on this information, we speculated that asp genes might have roles in maintaining intracellular redox balance. Since NADH and NAD+ levels and ratios are important indicators of redox, we measured the levels of reduced (NADH) and oxidized (NAD+) nicotinamide adenine dinucleotide; these were quantified in wild-type, ∆whiB7, ∆aspB, ∆aspC mutants, hsp-aspBMtb, and hsp-aspCMtb overexpression strains in early exponential phase (Figure 17A). A trend observed in mutant ∆aspB, hspaspBMtb and hsp-aspCMtb was a decrease in the pools of NAD+ (63%, 54%, and 58% of wild-type levels respectively) and no significant changes in NADH. The overall effect on the NADH:NAD+ ratios of ∆aspB, hsp-aspBMtb and hsp-aspCMtb was minor. The ∆aspC mutant was notably different in that its pool of NADH and NAD+ increased. This reflected the fact that while its NAD+ levels were unchanged, NADH levels were elevated 2.5-fold relative to wild-type (Figure 17A). The higher NADH:NAD+ ratio indicates a more reduced intracellular redox state (Figure 17B). In conclusion, deletion of aspC resulted in an increased pool of NADH and NAD+ and a major shift in intracellular redox suggesting that it may be linked to redox homeostasis.  3. 17. Increased oxidative stress in the ΔaspC mutant The ΔaspC mutant was more sensitive to hydrogen peroxide and to ROS inducer menadione, as determined via a disc assay. Stronger growth inhibition of the ΔaspC mutant compared to wild-type was indicated by larger zones of inhibitions (Figure 18). It should be noted that the observed sensitivity may be a by-product of the slower outgrowth of the ΔaspC mutant which would allow more time for the diffusion of compound from the disc into the medium. Results showing sensitivity of the ΔaspC mutant to hydrogen peroxide and menadione led to the hypothesis that aspC may be linked to oxidative stress response in mycobacteria. Catalase is one of the pathways for detoxification of oxidative stress. In mycobacteria, catalase (KatG) is expressed in response to hydrogen peroxide [119] [120] by relieving the repression of katG by the oxidative stress response regulator OxyS [121]. The amount of catalase typically reflects the level of oxidative stress in the bacterium. The catalase activities of exponential phase cultures were determined by directly measuring the decrease of hydrogen peroxide by catalase in lysates. The catalase activity measured in the lysate of the ΔaspC mutant was determined to be about four times that of wild-type (Figure 19A). Deletion or constitutive expression of aspB did not significantly affect catalase activity (Figure 19A). Treatment with hydrogen peroxide (20 mM) for 1 hour resulted in no change in wild-type catalase activity whereas the ΔaspC mutant had a decreased catalase activity (about 36% decrease) (Figure 19B). 46  Figure 17. The ΔaspC mutant has an increased redox state  A  B NAD+  ***   M/g dry wt  8  *  *  *  6 4 2 0  WT  hiB w  7  pB  as  pC  as  1.5  NADH  ratio NADH: NAD+  10  1.0  0.5  0.0 C Mtb B Mtb -asp -a s p hsp hsp  **  WT  7 hiB w  pB a s  pC B Mtb aspC Mtb as -asp hsp hsp  Figure 17. The ΔaspC mutant has an altered redox state. (A) Determination of intracellular levels of NAD+ and NADH in wild-type, ∆whiB7, ∆aspB, ∆aspC mutants, hsp-aspBMtb, and hsp-aspCMtb overexpression strains. Mutant ∆aspB, hsp-aspBMtb and hsp-aspCMtb had decreased concentrations of oxidized NAD+. (B) The redox ratio (NADH:NAD+) was calculated from NAD+ and NADH levels from A. The ∆aspC mutant had more than double the concentration of NADH compared to wild-type causing the redox potential of the cytoplasm to become more reduced as indicated by the NADH:NAD+ ratio. Results from triplicate measurements of 3 biological experiments. Values are the means ± standard error. * (P ≤ 0.05), ** (P ≤ 0.01), *** (P ≤ 0.001).  47  Figure 18. Sensitivity of mutants ΔaspB and ΔaspC to oxidative stress compounds hydrogen peroxide and menadione  A  B 40  diameter of ZOI (mm)  diameter of ZOI (mm)  80 70 60 50 40  mc2 155  ΔaspB  ΔaspC  ΔaspC pMV361::aspC  30  20  10  mc2 155  ΔaspB  ΔaspC  ΔaspC pMV361::aspC  Figure 18. Sensitivity of mutants ΔaspB and ΔaspC to oxidative stress compounds hydrogen peroxide and menadione. The diameter of the zone of inhibition around the disc containing (A) 5 μl of 30% hydrogen peroxide and (B) 5 μl of 50 mM menadione solution was measured after incubation at 37°C for 48 h. The ΔaspC mutant was more sensitive to both hydrogen peroxide and menadione as indicated by the larger zones of inhibition compared to wild-type. The sensitivity was suppressed in the complemented strain. Mutant ΔaspB did not exhibit any sensitivity to either hydrogen peroxide or menadione.  48  Figure 19. Higher oxidative stress in the ΔaspC mutant reflected in catalase activity  A  B catalase activity (x 1000 units mg -1)  catalase activity (x 1000 units mg -1)  *** 40  30  20  10  0  mc2 155  ΔaspB  hsp-aspB  ΔaspC  40  30  20  ns  10  Strain H 2 O2 (mM)  0  mc2 155 0  mc2 155 20  Δ aspC 0  Δ aspC 20  Figure 19. Higher oxidative stress in the ΔaspC mutant reflected in catalase activity. (A) Catalase activity measured in total lysate of wild-type, ∆aspB, ∆aspC mutants, and hsp-aspBMtb, strains. The catalase activity found in ∆aspC was four times that of wild-type. (B) Catalase activity in response in hydrogen peroxide treatment (20 mM H2O2 for 1 hour). The catalase activity decreased after hydrogen peroxide exposure in the ∆aspC mutant but was not affected in wild-type. ns (not significant), *** (P≤ 0.0001).  49  3. 18. Phylogenic analysis corroborates aspB and aspC as putative aminotransferases The function of AspB and AspC was analyzed using a phylogenetic approach based on amino acid sequence homology. Both AspB and AspC are annotated as aspartate aminotransferases. Aligning AspBMtb and AspCMtb sequences showed that they share 30% amino acid identity. Under the premise that aminotransferases with similar catalytic activities would exhibit sequence homology, AspB and AspC amino acid sequences were compared to five classes of aminotransferases that are closely related to aspartate aminotransferases functionally and phylogenetically. Figure 20 shows comparison of 87 sequences of enzymes that are biochemically characterized as aspartate aminotransferases: alanine aminotransferases (EC 2.6.1.2), tyrosine aminotransferases (EC2.6.1.5), kynurnine-oxoglutarate aminotransferases (EC2.6.1.7), histidinol-phosphate aminotransferases (EC 2.6.1.9), aromatic amino acid aminotransferases, and branched-chain amino acid aminotransferases. As an internal control M. tuberculosis IlvE was included as it is the only mycobacterial aminotransferase to be structurally and functionally characterized [87]. As predicted, the M. tuberculosis IlvE clustered with the branched chain amino acid aminotransferases of Pseudomonas, Staphylococcus and Haemophilus. AspB was found clustered with aspartate aminotransferases of S. solfataricus and T. thermophilus. AspC was found with other aspartate aminotransferases as well (Bacillus sp., G. stearothermophilus) but the proximity to alanine aminotransferases could suggest another function. To examine the phylogenetic relationship of mycobacterial AspB and AspC with aspartate aminotransferases from plants, animals, protozoa, eubacteria and archeabacteria, a phylogram was constructed using the Neighbor-joining method. Aspartate aminotransferases are classified into the aminotransferase family I which is again separated into subgroups on the basis on their amino acid sequence [122,123] [124]. Subgroup Ia generally contain aspartate aminotransferases from eubacteria, eukaryotes, animals and plants while subgroup Ib contain almost exclusively aspartate aminotransferases from prokaryotes, including protozoa, archaebacteria and eubacteria [125]. The amino acid sequence identities between subgroups Ia and Ib are only ~15% [125]. Figure 21 shows a phylogenetic tree of aspartate aminotransferases from different organisms. In the tree, AspB and AspC from M. tuberculosis clustered with other bacterial aspartate aminotransferases supporting potential structural and functional similarity to other bacterial aspartate aminotransferases. It is interesting that while AspB and AspC clustered with other Ib subgroup prokaryote-type aspartate aminotransferases, their sequences did not cluster with the main subgroup Ib branch: each occupies a separate periphery branch in the phylogeny tree. Extensive work has been done in the field of eukaryotic and prokaryotic aspartate aminotransferases and key catalytic residues important for their function have been defined. The conserved active residues important to the function of family I aminotransferases have been thoroughly studied using X-ray crystallographic studies and site-directed mutagenesis experiments. We performed sequence alignment 50  analysis between aspartate aminotransferase from E. coli (AspC) and cytosolic aspartate aminotransferase from Sus scrofa (pig) and found that AspB and AspC from M. tuberculosis and M. smegmatis also appeared to have most of the conserved active residues of family I aminotransferases (Table 6). In addition, the signature amino acid motifs surrounding the conserved residues were found in AspB and AspC. An interesting note was that AspC from M. tuberculosis and M. smegmatis had an arginine residue that aligns with Arg292, which is found in subgroup Ia aspartate aminotransferases. This residue recognizes the distal carboxyl groups of dicarboxylate substrates. Aspartate aminotransferases from subgroup Ib typically rely on a Lys109 to perform this function but this residue was not found in either AspB or AspC. Studies on the substitution of Arg292 have shown that aspartate aminotransferase activity was greatly decreased by the mutation while other pyridoxal 5’-phosphate-dependent catalytic activities such as β-decarboxylation and racemization of L-aspartate increased [126]. Based on phylogenic analysis, AspB and AspC possess sequence similarities to the family I within the protein superfamily of aminotransferases which includes aspartate aminotransferases. The arginine catalytic residue characteristic of subgroup Ia was found in AspC but not AspB. The absence of this important catalytic residue could have consequences for the enzymatic activity of AspB.  51  Figure 20. Phylogenetic tree showing amino acid homology of various aminotransferases sp|Q949X3|HIS8 ARATH Histidinol-phosphate aminotransferase chloroplastic OS Arabidopsis thaliana GN HPA1 PE 2 SV 1 sp|Q9PII2|HIS8 CAMJE Histidinol-phosphate aminotransferase OS Campylobacter jejuni GN hisC PE 1 SV 1 sp|P34037|HIS8 ZYMMO Histidinol-phosphate aminotransferase OS Zymomonas mobilis GN hisC PE 3 SV 3 sp|P17731|HIS8 BACSU Histidinol-phosphate aminotransferase OS Bacillus subtilis GN hisC PE 3 SV 3 sp|Q82AA5|HIS8 STRAW Histidinol-phosphate aminotransferase OS Streptomyces avermitilis GN hisC PE 3 SV 1 sp|P16246|HIS8 STRCO Histidinol-phosphate aminotransferase OS Streptomyces coelicolor GN hisC PE 3 SV 1  Histidinol-  sp|P06986|HIS8 ECOLI Histidinol-phosphate aminotransferase OS Escherichia coli (strain K12) GN hisC PE 1 SV 2 sp|P10369|HIS8 SALTY Histidinol-phosphate aminotransferase OS Salmonella typhimurium GN hisC PE 3 SV 2  phosphate AT  sp|Q8ZFX6|HIS8 YERPE Histidinol-phosphate aminotransferase OS Yersinia pestis GN hisC PE 3 SV 1 sp|Q8Z5J9|HIS8 SALTI Histidinol-phosphate aminotransferase OS Salmonella typhi GN hisC PE 3 SV 1 sp|A5UGY2|HIS8 HAEIG Histidinol-phosphate aminotransferase OS Haemophilus influenzae (strain PittGG) GN hisC PE 3 SV 1 sp|P0A678|HIS8 MYCTU Histidinol-phosphate aminotransferase OS Mycobacterium tuberculosis GN hisC PE 3 SV 1 sp|A4QFG6|HIS8 CORGB Histidinol-phosphate aminotransferase OS Corynebacterium glutamicum (strain R) GN hisC PE 3 SV 1 sp|P07172|HIS8 YEAST Histidinol-phosphate aminotransferase OS Saccharomyces cerevisiae (strain ATCC 204508 / S288c) GN HIS5 PE 1 SV 2 sp|A5I245|HIS8 CLOBH Histidinol-phosphate aminotransferase OS Clostridium botulinum (strain Hall / ATCC 3502 / NCTC 13319 / Type A) GN hisC PE 3 SV 1 sp|Q8BTY1|KAT1 MOUSE Kynurenine--oxoglutarate transaminase 1 OS Mus musculus GN Ccbl1 PE 2 SV 1  Kynurenine-  sp|Q08415|KAT1 RAT Kynurenine--oxoglutarate transaminase 1 mitochondrial OS Rattus norvegicus GN Ccbl1 PE 1 SV 1 sp|Q16773|KAT1 HUMAN Kynurenine--oxoglutarate transaminase 1 OS Homo sapiens GN CCBL1 PE 1 SV 1  oxoglutarate AT  sp|Q71RI9|KAT3 MOUSE Kynurenine--oxoglutarate transaminase 3 OS Mus musculus GN Ccbl2 PE 1 SV 1 sp|Q58FK9|KAT3 RAT Kynurenine--oxoglutarate transaminase 3 OS Rattus norvegicus GN Ccbl2 PE 2 SV 1 sp|Q6YP21|KAT3 HUMAN Kynurenine--oxoglutarate transaminase 3 OS Homo sapiens GN CCBL2 PE 1 SV 1 sp|O85746|TYRB KLEPN Tyrosine aminotransferase OS Klebsiella pneumoniae GN tyrB PE 1 SV 1  Aromatic  sp|P74861|TYRB SALTY Aromatic-amino-acid aminotransferase OS Salmonella typhimurium GN tyrB PE 3 SV 3  amino acid AT  sp|P04693|TYRB ECOLI Aromatic-amino-acid aminotransferase OS Escherichia coli (strain K12) GN tyrB PE 1 SV 1 sp|P72173|AAT PSEAE Aspartate aminotransferase OS Pseudomonas aeruginosa GN aspC PE 3 SV 2 sp|P44425|AAT HAEIN Aspartate aminotransferase OS Haemophilus influenzae (strain ATCC 51907 / DSM 11121 / KW20 / Rd) GN aspC PE 3 SV 1 sp|Q02636|ATTY RHIME Tyrosine aminotransferase OS Rhizobium meliloti GN tatA PE 3 SV 1 sp|P43336|PHHC PSEAE Aromatic-amino-acid aminotransferase OS Pseudomonas aeruginosa GN phhC PE 3 SV 2 sp|P58661|AAT SALTY Aspartate aminotransferase OS Salmonella typhimurium GN aspC PE 3 SV 1 sp|P00509|AAT ECOLI Aspartate aminotransferase OS Escherichia coli (strain K12) GN aspC PE 1 SV 1 sp|P05201|AATC MOUSE Aspartate aminotransferase cytoplasmic OS Mus musculus GN Got1 PE 1 SV 3 sp|P17174|AATC HUMAN Aspartate aminotransferase cytoplasmic OS Homo sapiens GN GOT1 PE 1 SV 3 sp|P00503|AATC PIG Aspartate aminotransferase cytoplasmic OS Sus scrofa GN GOT1 PE 1 SV 3 sp|P13221|AATC RAT Aspartate aminotransferase cytoplasmic OS Rattus norvegicus GN Got1 PE 1 SV 3 sp|P28011|AAT1 MEDSA Aspartate aminotransferase 1 OS Medicago sativa GN AAT-1 PE 2 SV 2 sp|P46644|AAT3 ARATH Aspartate aminotransferase chloroplastic OS Arabidopsis thaliana GN ASP3 PE 1 SV 1 sp|P00505|AATM HUMAN Aspartate aminotransferase mitochondrial OS Homo sapiens GN GOT2 PE 1 SV 3 sp|P23542|AATC YEAST Aspartate aminotransferase cytoplasmic OS Saccharomyces cerevisiae (strain ATCC 204508 / S288c) GN AAT2 PE 1 SV 3 sp|P05202|AATM MOUSE Aspartate aminotransferase mitochondrial OS Mus musculus GN Got2 PE 1 SV 1 sp|P00506|AATM PIG Aspartate aminotransferase mitochondrial OS Sus scrofa GN GOT2 PE 1 SV 2 sp|P00507|AATM RAT Aspartate aminotransferase mitochondrial OS Rattus norvegicus GN Got2 PE 1 SV 2 sp|O67781|AAT AQUAE Aspartate aminotransferase OS Aquifex aeolicus GN aspC PE 3 SV 1 tr|P96847|P96847 ASPB (ASPAT) Mycobacterium tuberculosis aspB P2 tr|A0R503|A0R503 Aminotransferase class I Mycobacterium smegmatis mc(2)155) GN MSMEG 6017 PE 3 SV 1  Aspartate AT  AspB  sp|Q9V0L2|AAT PYRAB Aspartate aminotransferase OS Pyrococcus abyssi (strain GE5 / Orsay) GN aspC PE 3 SV 1 sp|P14909|AAT SULSO Aspartate aminotransferase OS Sulfolobus solfataricus (strain ATCC 35092 / DSM 1617 / JCM 11322 / P2) GN aspC PE 1 SV 2 sp|Q9SIE1|PAT ARATH Bifunctional aspartate aminotransferase and glutamate/aspartate-prephenate aminotransferase OS Arabidopsis thaliana GN PAT PE 1 SV 2 sp|O33822|AAT THEAQ Aspartate aminotransferase OS Thermus aquaticus GN aspC PE 3 SV 1 sp|Q56232|AAT THET8 Aspartate aminotransferase OS Thermus thermophilus (strain HB8 / ATCC 27634 / DSM 579) GN aspC PE 1 SV 1 sp|P58350|AATB1 RHIME Aspartate aminotransferase B OS Rhizobium meliloti GN aatB PE 3 SV 1 sp|O86459|AAT RHILP Aspartate aminotransferase OS Rhizobium leguminosarum bv. phaseoli GN aspC PE 3 SV 1 sp|Q02635|AATA RHIME Aspartate aminotransferase A OS Rhizobium meliloti GN aatA PE 3 SV 1 sp|Q60013|AAT STRVG Aspartate aminotransferase OS Streptomyces virginiae GN aspC PE 3 SV 1 sp|Q55128|AAT SYNY3 Aspartate aminotransferase OS Synechocystis sp. (strain ATCC 27184 / PCC 6803 / N-1) GN aspC PE 3 SV 1 sp|P33447|ATTY TRYCR Tyrosine aminotransferase OS Trypanosoma cruzi PE 1 SV 2 sp|P17735|ATTY HUMAN Tyrosine aminotransferase OS Homo sapiens GN TAT PE 1 SV 1  Tyrosine AT  sp|P04694|ATTY RAT Tyrosine aminotransferase OS Rattus norvegicus GN Tat PE 1 SV 1 sp|Q8QZR1|ATTY MOUSE Tyrosine aminotransferase OS Mus musculus GN Tat PE 1 SV 1 sp|P53001|AAT1 BACSU Aspartate aminotransferase OS Bacillus subtilis GN aspB PE 3 SV 1 sp|P23034|AAT BACY2 Aspartate aminotransferase OS Bacillus sp. (strain YM-2) PE 1 SV 1 sp|Q59228|AAT GEOSE Aspartate aminotransferase OS Geobacillus stearothermophilus GN aspC PE 3 SV 1 sp|P63498|AAT MYCTU Probable aspartate aminotransferase OS Mycobacterium tuberculosis GN aspC PE 3 SV 1  AspC  tr|A0QQA8|A0QQA8 MYCS2 Aspartate aminotransferase OS Mycobacterium smegmatis (strain ATCC 700084 / mc(2)155) GN MSMEG 0688 PE 4 SV 1 sp|Q8QZR5|ALAT1 MOUSE Alanine aminotransferase 1 OS Mus musculus GN Gpt PE 2 SV 3 sp|P25409|ALAT1 RAT Alanine aminotransferase 1 OS Rattus norvegicus GN Gpt PE 1 SV 2 sp|P13191|ALAT1 PIG Alanine aminotransferase 1 (Fragment) OS Sus scrofa GN GPT PE 1 SV 1 sp|P52892|ALAT YEAST Probable alanine aminotransferase OS Saccharomyces cerevisiae (strain ATCC 204508 / S288c) GN ALT2 PE 1 SV 1  Alanine AT  sp|P24298|ALAT1 HUMAN Alanine aminotransferase 1 OS Homo sapiens GN GPT PE 1 SV 3 sp|Q8TD30|ALAT2 HUMAN Alanine aminotransferase 2 OS Homo sapiens GN GPT2 PE 1 SV 1 sp|Q8BGT5|ALAT2 MOUSE Alanine aminotransferase 2 OS Mus musculus GN Gpt2 PE 2 SV 1 tr|Q1CNM9|Q1CNM9 YERPN Branched chain amino acid aminotransferase OS Yersinia pestis bv. Antiqua (strain Nepal516) GN ilvE PE 3 SV 1 sp|P0A1A5|ILVE SALTY Branched-chain-amino-acid aminotransferase OS Salmonella typhimurium GN ilvE PE 1 SV 2 sp|P0A1A6|ILVE SALTI Branched-chain-amino-acid aminotransferase OS Salmonella typhi GN ilvE PE 3 SV 2 sp|P0AB80|ILVE ECOLI Branched-chain-amino-acid aminotransferase OS Escherichia coli (strain K12) GN ilvE PE 1 SV 2 sp|Q5HF24|DAAA STAAC D-alanine aminotransferase OS Staphylococcus aureus (strain COL) GN dat PE 3 SV 1 sp|O85046|DAAA LISMO D-alanine aminotransferase OS Listeria monocytogenes GN dat PE 3 SV 2 sp|O86428|ILVE PSEAE Branched-chain-amino-acid aminotransferase OS Pseudomonas aeruginosa GN ilvE PE 1 SV 2 sp|A0R066|ILVE MYCS2 Branched-chain-amino-acid aminotransferase OS Mycobacterium smegmatis (strain ATCC 700084 / mc(2)155) GN ilvE PE 1 SV 1 sp|O32954|ILVE MYCLE Probable branched-chain-amino-acid aminotransferase OS Mycobacterium leprae GN ilvE PE 3 SV 1 sp|Q10399|ILVE MYCTU Branched-chain-amino-acid aminotransferase OS Mycobacterium tuberculosis GN ilvE PE 1 SV 1  52  tr|A0QQA8|A0QQA8 MYCS2 Aspartate aminotransferase OS Mycobacterium smegmatis (strain ATCC 700084 / mc(2)155) GN MSMEG 0688 PE 4 SV 1 sp|Q8QZR5|ALAT1 MOUSE Alanine aminotransferase 1 OS Mus musculus GN Gpt PE 2 SV 3 sp|P25409|ALAT1 RAT Alanine aminotransferase 1 OS Rattus norvegicus GN Gpt PE 1 SV 2 sp|P13191|ALAT1 PIG Alanine aminotransferase 1 (Fragment) OS Sus scrofa GN GPT PE 1 SV 1 sp|P52892|ALAT YEAST Probable alanine aminotransferase OS Saccharomyces cerevisiae (strain ATCC 204508 / S288c) GN ALT2 PE 1 SV 1 sp|P24298|ALAT1 HUMAN Alanine aminotransferase 1 OS Homo sapiens GN GPT PE 1 SV 3 sp|Q8TD30|ALAT2 HUMAN Alanine aminotransferase 2 OS Homo sapiens GN GPT2 PE 1 SV 1 sp|Q8BGT5|ALAT2 MOUSE Alanine aminotransferase 2 OS Mus musculus GN Gpt2 PE 2 SV 1 tr|Q1CNM9|Q1CNM9 YERPN Branched chain amino acid aminotransferase OS Yersinia pestis bv. Antiqua (strain Nepal516) GN ilvE PE 3 SV 1 sp|P0A1A5|ILVE SALTY Branched-chain-amino-acid aminotransferase OS Salmonella typhimurium GN ilvE PE 1 SV 2 sp|P0A1A6|ILVE SALTI Branched-chain-amino-acid aminotransferase OS Salmonella typhi GN ilvE PE 3 SV 2 sp|P0AB80|ILVE ECOLI Branched-chain-amino-acid aminotransferase OS Escherichia coli (strain K12) GN ilvE PE 1 SV 2 sp|Q5HF24|DAAA STAAC D-alanine aminotransferase OS Staphylococcus aureus (strain COL) GN dat PE 3 SV 1 sp|O85046|DAAA LISMO D-alanine aminotransferase OS Listeria monocytogenes GN dat PE 3 SV 2 sp|O86428|ILVE PSEAE Branched-chain-amino-acid aminotransferase OS Pseudomonas aeruginosa GN ilvE PE 1 SV 2 sp|A0R066|ILVE MYCS2 Branched-chain-amino-acid aminotransferase OS Mycobacterium smegmatis (strain ATCC 700084 / mc(2)155) GN ilvE PE 1 SV 1  IlvE  sp|O32954|ILVE MYCLE Probable branched-chain-amino-acid aminotransferase OS Mycobacterium leprae GN ilvE PE 3 SV 1 sp|Q10399|ILVE MYCTU Branched-chain-amino-acid aminotransferase OS Mycobacterium tuberculosis GN ilvE PE 1 SV 1  Branched  sp|Q93Y32|BCAT1 ARATH Branched-chain-amino-acid aminotransferase 1 mitochondrial OS Arabidopsis thaliana GN BCAT1 PE 1 SV 2 sp|P54689|ILVE HAEIN Branched-chain-amino-acid aminotransferase OS Haemophilus influenzae (strain ATCC 51907 / DSM 11121 / KW20 / Rd) GN ilvE PE 3 SV 1  chain AT  sp|P99138|ILVE STAAN Probable branched-chain-amino-acid aminotransferase OS Staphylococcus aureus (strain N315) GN ilvE PE 1 SV 1 sp|Q5HIC1|ILVE STAAC Probable branched-chain-amino-acid aminotransferase OS Staphylococcus aureus (strain COL) GN ilvE PE 3 SV 1 sp|P39576|ILVE2 BACSU Branched-chain-amino-acid aminotransferase 2 OS Bacillus subtilis GN ilvK PE 1 SV 5 sp|O31461|ILVE1 BACSU Branched-chain-amino-acid transaminase 1 OS Bacillus subtilis GN ilvE PE 1 SV 1 sp|O07597|DAAA BACSU D-alanine aminotransferase OS Bacillus subtilis GN dat PE 3 SV 1 sp|P47176|BCA2 YEAST Branched-chain-amino-acid aminotransferase cytosolic OS Saccharomyces cerevisiae (strain ATCC 204508 / S288c) GN BAT2 PE 1 SV 1 sp|P38891|BCA1 YEAST Branched-chain-amino-acid aminotransferase mitochondrial OS Saccharomyces cerevisiae (strain ATCC 204508 / S288c) GN BAT1 PE 1 SV 1 sp|P54687|BCAT1 HUMAN Branched-chain-amino-acid aminotransferase cytosolic OS Homo sapiens GN BCAT1 PE 1 SV 3 sp|O15382|BCAT2 HUMAN Branched-chain-amino-acid aminotransferase mitochondrial OS Homo sapiens GN BCAT2 PE 1 SV 2 sp|O35854|BCAT2 RAT Branched-chain-amino-acid aminotransferase mitochondrial OS Rattus norvegicus GN Bcat2 PE 1 SV 1  Figure 20. Phylogenetic tree showing amino acid homology of various aminotransferases: Both AspB and AspC clustered with other aspartate aminotransferase sequences. The phylogenetic tree was constructed with full-length amino acid sequences using MEGA 5.0. , based on Figure found in [127]. As a control the sequences of the branched chain aminotransferase (IlvE) of M.tuberculosis and M.smegmatis were included and both clustered with other branched chain aminotransferase sequences showing that amino acid sequences of functionally similar proteins cluster together in this tree.  53  Figure 21. Neighbor-joining phylogenic tree of aspartate aminotransferases from different organisms AATc AAQ02891 A.aegypti  Animal  AAT cy P00504 Chicken  cytoplasmicAspATs  AAT cy NM012571 Rat AAT cy P33097 Bovine AAT cy P00503 Pig AATm AAQ02892 A.aegypti  Animal  AAT m P00506 Pig  mitochondrialAspATs  AAT m P12344 Bovine AAT m NM013177 Rat AAT m AAH00525 Human  Plant mitochondrial  AAT m P46643 A.thaliana AAT m AAA98603 Glycine.max  AspATs  AAT m BAA04993 P.miliaceum AAT p 1908424A M.sativa AAT cy CAA43779 M.sativa  Plant plastidic,  AAT cy AAC50015 Glycine.max  cytoplasmic &  AAT cy P46645 A.thaliana  chloroplastic AspATs  AAT ch P46644 A.thaliana AAT CBA04235 N.meningitidis AAT AAC23265 H.influenzae AAT P00509 E.coli AAT AAD56399 A.hydrophila AAT AAF94452 V.cholerae AAT CAA04961 Moraxella AAT CAA48188 S.cerevisiae AAT CAA64186 B.circulans AATB3 AY040867 Bsubtilis AAT1 NP894860 P.marinus AAT ABA23301 A.variabilis AAT3 BAA10796 Synechocystis AAT MSMEG 6017aspB AAT Rv3565 M.tuberculosis  AspB  AAT YP 879876 M.avium AAT ro04781 R.jostii AAT NP 342388 S.solfataricus AAT NP390118 B.subtilis168 AAT P23034 Bacillus.sp  Bacteria AspATs  AAT NP683147 T.elongatus AAT3 BAA10261 Synechocystis AAT BAB86290 P.lapideum AAT Q60013 S.virginiae AAT CAB77419 S.coelicolor AAT EFD67395 S.lividans AAT JC4537 T.aquaticus AAT X99521 T.aquaticus AAT YP703251 R.jostii AAT1 AAA71965 S.meliloti AAT2 L05064 R.meliloti AAT2 NP894611 P.marinus AATos BAB86539 O.sativa AATat Q9SIE1 A.thaliana AATpp Q5F4K8 P.pinaster AAT2 BAA10205 Synechocystis AAT ro05465 R.jostii AAT MSMEG 0688aspC AAT Rv0337c M.tuberculosis  AspC  0.2  Figure 21. Neighbor-joining phylogenic tree of aspartate aminotransferases from different organisms: AspB and AspC from M.tuberculosis and M.smegmatis were found to have strong sequence homology with other bacterial aspartate aminotransferases. The phylogenetic tree was based on figure in [128] and constructed with full-length amino acid sequences using MEGA 5.0.  54  Table 6. Functions of anchor residues in family I aminotransferases. Based on [127] Residue in hierarchical level of invariance  Alpha division of PLP-dependent  residue functional role  AspBMtb  AspCMtb AspBMsm AspCMsm  K-258  K-234  K-265  K-237  K-260  D-202  D-232  D-205  D-227  G-177  G-207  G-180  G-202  R-364  R-403  R-364  R-404  Y-64  Y-92  Y-67  Y-87  G-102  NF  G-105  NF  N-174  N-204  N-177  N-199  P-175  P-205  P-178  P-200  Y-205  Y-235  Y-208  Y-230  R-242  R-273  R-245  R-268  G-244  G-275  G-247  G-270  NF  R-299  NF  R-300  NF  NF  NF  NF  D-222  proteins Aminotransferase  G-197  superfamily R-386  Y-70 Family I aminotransferases  G-110 N-194 P-195 Y-225 R-266 G-268  Subgroup Iα  R-292  Subgroup Iβ  Y-109  forms Schiff base with PLP forms salt bridge by H-bonding with the pyridine N-1 of PLP accommodates the turn at the interdomain interface forms salt bridge by H-bonding with α-COO- of substrate H-bond to phosphate OP2 of PLP and stabilizes transition state unknown role H bonds to hydroxyl O-3' of PLP regulating electron distribution has cis conformation within span of interdomain interface H bonds to hydroxyl O-3' of PLP regulating electron distribution H bonds with phosphates OP2 and OP4 of PLP unknown role recognizes the distal carboxyl groups of dicarboxylate substrates recognizes the distal carboxyl groups of dicarboxylate substrates  Table 6. Functions of anchor residues in family I aminotransferases.  NF: residue not found  55  3. 19. Complementation of aspartate auxotrophy in E. coli with AspBMtb and AspCMtb The homologies of AspB and AspC suggested their functions as transaminases but did not clarify their substrates. Furthermore, since the M. smegmatis strain with mutations in both aspB and aspC had no amino acid auxotrophy (Figure 10), the metabolic functions of these genes were unclear. E. coli was used to explore the metabolic function of AspBMtb and AspCMtb. In E. coli (and by analogy Mycobacterium), aspartate auxotrophy is dependent on several alleles, including the aspC ortholog. In order to observe auxotrophy for aspartate in an E. coli aspC mutant, the genes ilvE and tyrB, encoding a branched-chain aminotransferase and aromatic amino acid aminotransferase respectively, also need to be mutated [129]. Aminotransferases have discrete specificities but there is also cross-specificities and activities. Aromatic amino acid aminotransferase TyrB and branched-chain amino acid aminotransferase IlvE can complement the mutation in the other gene [130]. Aspartate aminotransferase AspC can also catalyze the transamination of the keto-acids phenylpyruvate and 4-hydroxyphenylpyruvate to form phenylalanine and tyrosine respectively, albeit at a lower rate with lower affinity for the substrates [129,131]. We obtained the auxotroph DL39 with mutations in aspC, ilvE, and tyrB auxotrophy for leucine, isoleucine, valine, tyrosine, phenylalanine and aspartate [98] from the E. coli Genetic Stock Center. We hypothesized that extra-chromosomal expression of aspBMtb and aspCMtb would be able to complement the activity of the mutated aspC in DL39 if AspB and AspC were indeed able to catalyze the aspartate aminotransferase reaction. The genes aspBMtb and aspCMtb along with the hsp60 promoter was cloned from pMV361-aspB and pMV361-aspC respectively using restriction sites XbaI and HindIII and ligated into pUC19 (Figure 22). The mycobacterial hsp60 promoter is active in E. coli as it is orthologous to the GroEL protein of E. coli [132]. The constructs pUC19::hsp-aspBMtb and pUC19::hsp-aspCMtb were transformed into DL39 and plated onto LB plates with ampicillin selection. The transformants were grown onto 56/2 minimal media agar plates [99] with and without amino acids (leucine, isoleucine, valine, tyrosine, phenylalanine, and aspartate at 0.3 mM). Both DL39 pUC19::aspBMtb and DL39 pUC19::aspCMtb exhibited the same auxotrophy as DL39, requiring supplementation of all the six amino acids for growth. The hsp60 promoter drives constitutive expression of aspBMtb and aspCMtb although higher levels of expression might be required to complement the auxotrophy. AspB and AspC may have lower catalytic efficiency and lower substrate affinity than E. coli AspC for the transamination of aspartate thus requiring higher expression levels to compensate for the mutation. Also, cloning the indigenous aspCEco gene in the same manner to act as a positive control would help indicate whether the hsp60 promoter is driving sufficient levels of expression.  56  Figure 22. Vector maps of pUC19::hsp-aspB and pUC19::hsp-aspC  Figure 22. Vector maps of pUC19::hsp-aspB and pUC19::hsp-aspC. Constructs were transformed into the amino acid auxotroph DL39.  57  3. 20. Amino acid auxotroph DL39 does not exhibit general growth deficiencies and does not show increased antibiotic susceptibility The deletion of aspC in M. smegmatis caused many growth deficiencies in the strain (Figure 7). To determine whether this phenomenon can be extended to other bacteria, the growth kinetics of the amino acid auxotroph DL39, which has also has a mutation in its aspC gene, was studied. The growth curve of DL39 was very similar to that of its wild-type, MG1655. The two strains entered into exponential growth at similar time and the doubling rates during exponential growth were similar. The growth of DL39 decreased more quickly than MG1655 growth resulting in a lower cell density at stationary phase; the M. smegmatis mutant ΔaspC also displayed a decreased cell density in stationary phase (Figure 23). The M. smegmatis ΔaspC mutant also showed increased antibiotic susceptibility. To test whether deletion of aspC would also cause increased antibiotic susceptibility in E. coli, the MICs of amino acid auxotroph DL39 against tetracycline, clarithromycin, and spectinomycin were determined using MTT assay and compared with the MICs of those antibiotics in wild-type MG1655. There was no significant difference in antibiotic susceptibility between wild-type MG1655 and amino acid auxotroph DL39 (Table 7). Additionally, oxaloacetate was able to delay growth of M. smegmatis at high concentrations (40-80 mM) leading us to test whether oxaloacetate can also be toxic to E. coli. At 40 mM oxaloacetate, growth of E. coli MG1655 was inhibited by approximately 45% while at 80 mM growth was further depressed to 22% of untreated control. There was no significant difference in susceptibility towards oxaloacetate between wild-type MG1655 and amino acid auxotroph DL39 (Figure 24).  58  Figure 23. Amino acid auxotroph DL39 has a lower cell density at stationary phase  1.6  MG1655  1.4  DL39  OD600  1.2 1.0 0.8 0.6 0.4 0.2 0.0 0  5  10  15  20  25  30  35  time (h) Figure 23. Amino acid auxotroph DL39 has a lower cell density at stationary phase. E. coli strains MG1655 and DL39 in quadruplicate wells were grown in a Bioscreen C kinetic growth reader at 37 °C with constant shaking at maximum amplitude. Growth was monitored by the Bioscreen by measuring OD600nm. The growth kinetics of E. coli MG1655 and amino acid auxotroph DL39 in LB show that the mutant has a similar doubling rate as compared to wild-type but enters into stationary phase earlier, resulting in a lower final cell.  Table 7. Amino acid auxotrophic strain DL39 does not show changes in antibiotic susceptibility. The MICs of wild-type (MG1655) and amino acid auxotroph DL39 for antibiotics tetracycline, clarithromycin, and spectinomycin were determined by MTT assay in LB medium. The MICs of wild-type (MG1655) and amino acid auxotroph DL39 for antibiotics tetracycline, clarithromycin, and spectinomycin were similar. MIC90 (μg/ml) MG1655  DL39  0.16  0.31  clarithromycin  16  16  spectinomycin  16  16  tetracycline  Table 7. Amino acid auxotrophic strain DL39 does not show changes in antibiotic susceptibility.  59  Figure 24. Oxaloacetate inhibited growth of E. coli at 40mM and 80mM  100  MG1655 DL39  % inhibition  80 60 40 20 0 20 mM  40 mM  80 mM  oxaloacetate Figure 24. Oxaloacetate inhibited growth of E. coli at 40mM and 80mM. MG1655 and DL39 were grown in LB in the presence of oxaloacetate. Percent inhibition compared to untreated control of the strains was determined using MTT viability assay. No difference in oxaloacetate susceptibility was observed between wild-type MG1655 and amino acid auxotroph DL39.  60  3. 21. Expression and purification of AspBMtb and AspCMtb To confirm AspB and AspC as aspartate aminotransferases, the activities of the purified enzymes would need to be determined experimentally. The asp genes were cloned into a pGEX-based vector (GE Lifesciences) for expression as recombinant GST-tagged proteins (Figure 25). Protein production in E. coli BL21 was verified after overnight induction with IPTG at room temperature. While the GST affinity purification system was chosen for properties of increasing protein solubility and purifying soluble GST fusion proteins, most of the GST-tagged AspB and AspC were insoluble under non-denaturing conditions. Inclusion of 1.5% sarkosyl in the lysis buffer increased solubility of the proteins and 2% triton increased binding to glutathione matrix [107]. Affinity purification on a glutathione sepharose column yielded purified GST-AspB and GST-AspC as observed by SDS PAGE and silver staining (Figure 26). The purified GST-tagged proteins were assayed for aspartate aminotransferase activity using the assay described in section 2.25 but no catalysis was observed suggesting that the GST moiety may be sterically hindering the activities of the Asp proteins due to its relatively large size. Thus, we proceeded to remove the GST-tag via cleavage of an engineered thrombin cleavage site between the tag and the Asp proteins. Thrombin specifically cleaved and released AspB from the GST tag. However, attempts at removing free GST using the same glutathione matrix left a significant amount of free GST in the flow-through with the cleaved AspB protein. Additional purification steps were required to separate the free GST from AspBMtb after protease cleavage (Figure 27).  61  Figure 25. Expression vector maps for GST-tagged recombinant protein expression  Figure 25. Expression vector maps for GST-tagged recombinant protein expression. The genes aspB and aspC from M. tuberculosis were cloned into pGEX4T-1 (GE) for expression of GST-tagged recombinant AspB and AspC in E. coli BL21.  62  Figure 26. Purification of AspBMtb and AspCMtb using GST affinity chromatography  Figure 26. Purification of AspBMtb and AspCMtb using GST affinity chromatography. GST-tagged AspB and AspC were expressed in BL21 E. coli and purified on a Glutathione Sepharose 4B column. (A) Purification of GST-AspB. Lane 1 whole cell lysate. Lane 2 unbound fraction. Lane 3 wash fraction. Lane 4 GST-AspB elution. (B) Purification of GST-AspC. Lane 5 whole cell lysate. Lane 6 unbound fraction. Lane 7 wash fraction. Lane 8 GST-AspC elution. M, molecular weight marker. The eluted tagged proteins were of the expected size: AspB approximately 67 kDa. and AspC approximately 73 kDa.  63  Figure 27. Removal of GST-tag from recombinant protein GST-AspBMtb  Figure 27. Removal of GST-tag from recombinant protein GST-AspBMtb. Lane 1 to 4 GST protein still remaining in solution along with cleaved AspB after clean-up with a glutathione sepharose column. Lane 5 uncut GST-AspB. M, molecular weight marker. Thrombin successfully removed the GST-tag from AspB but free GST was difficult to remove from solution.  64  Removal of the GST fusion tag from GST-AspC using the engineered thrombin cleavage site was unsuccessful. Thrombin has secondary recognition sites that cleaved within AspCMtb (Figure 28). The primary recognition site for thrombin is P4-P3-P-R/K● P1’-P2’ with the bond between R/K●P1’ being the scissile bond (as denoted by ●). At P4 and P3 are hydrophobic residues and residues at P1’ and P2’ are non-acidic [133]. The thrombin cut site between the GST and recombinant AspCMtb is LVPR●GS. However, thrombin also cuts at secondary sites such as P2-R/K●P1’ where P2 or P1’ is a glycine residue (Figure 28). This less stringent secondary site can be found throughout AspCMtb and this could allow for the many degradation products observed when GST-AspCMtb was cleaved with thrombin (Figure 29). Many conditions, including shortened incubation time, lower protease to protein ratio, and lower reaction temperature, were altered in attempt to reduce the affinity of thrombin for the secondary recognition sites but all were found to be ineffective (Figure 29). In addition, the AspC protein appeared unstable and seemed to degrade spontaneously. The GST tag expression system was effective at achieving expression of both AspB and AspC in soluble form. However, the relatively large size of the GST protein, 26 kDa compared to 41 kDa and 47 kDa for AspB and AspC respectively, made it necessary to remove the fusion tag as it hindered the catalytic activities of AspB and AspC. Moreover, the fusion tag removal system with thrombin was problematic as thrombin possesses too many secondary activities.  65  Figure 28. GST-AspCMtb contain many putative secondary thrombin cleavage sites  A Amino acid position P4  P3  P2  P1  P1’  P2’  L  V  P  R  G  S  hydrophobic  hydrophobic  G  R/K  G  hydrophobic  P  R  L/G  G/T/R/M/V P/G/V  R  P3’  P4’  reference [133] [133] [134]  S/A/G/T W/G/F/S  R  V/L/S/R  [135]  B MSPILGYWKIKGLVQPTRLLLEYLEEKYEEHLYERDEGDKWRNKKFELGLEFPNLPYYIDGDVKLTQSMAIIRYIADKHNM LGGCPKERAEISMLEGAVLDIRYGVSRIAYSKDFETLKVDFLSKLPEMLKMFEDRLCHKTYLNGDHVTHPDFMLYDALDV VLYMDPMCLDAFPKLVCFKKRIEAIPQIDKYLKSSKYIAWPLQGWQATFGGGDHPPKSDLVPRGSVDNDGTIVDVTTH QLPWHTASHQRQRAFAQSAKLQDVLYEIRGPVHQHAARLEAEGHRILKLNIGNPAPFGFEAPDVIMRDIIQALPYA QGYSDSQGILSARRAVVTRYELVPGFPRFDVDDVYLGNGVSELITMTLQALLDNGDQVLIPSPDYPLWTASTSLAGG TPVHYLCDETQGWQPDIADLESKITERTKALVVINPNNPTGAVYSCEILTQMVDLARKHQLLLLADEIYDKILYDDAK HISLASIAPDMLCLTFNGLSKAYRVAGYRAGWLAITGPKEHASSFIEGIGLLANMRLCPNVPAQHAIQVALGGHQSIE DLVLPGGRLLEQRDIAWTKLNEIPGVSCVKPAGALYAFPRLDPEVYDIDDDEQLVLDLLLSEKILVTQGTGFNWPAPD HLRLVTLPWSRDLAAAIERLGNFLVSYRQZ Figure 28. GST-AspCMtb contain many putative secondary thrombin cleavage sites. (A) thrombin recognition cleavage sites. Thrombin cleaves bond between P1 and P1’. While thrombin typically cuts the bond on the carboxyl end of an arginine residue, many other residues can be substituted in the recognition sequence. (B) Amino acid sequence of recombinant fusion protein GST-AspCMtb. The AspCMtb sequence is in bold print. The engineered thrombin cut site is highlighted in yellow while putative secondary cut sites are highlight in blue.  66  Figure 29. Thrombin cleavage of GST-AspCMtb  Figure 29. Thrombin cleavage of GST-AspCMtb. 50 μg of GST-AspCMtb was cleaved by varying amount of thrombin at 4°C for 18 hours: Lane 1 9000 ng. Lane 2 1800 ng. Lane 3 360 ng. Lane 4 72 ng Lane 5 15 ng. Lane 6 3 ng. Lane 7 0.6 ng. Lane 8 0.12 ng. Lane 9 no thrombin control. M, molecular weight marker. . Protein instability and secondary thrombin recognition sites cause non-specific degradation of GSTAspCMtb. Low temperature and high protein to protease ratio did not diminish non-specific cleavage.  67  Given the difficulties with the GST constructs, His affinity purification was chosen for purification of AspB and AspC as the His tag is a relatively small fusion tag. The aspB and aspC genes were cloned into pETbased vectors for expression (Figure 30) in Rosetta2 E. coli which possesses the plasmid pRARE2 (Novagen) allowing for the expression of tRNAs for rare codons as mycobacteria have a G/C-rich genome while the genome of E. coli is A/T-rich. His-AspB was expressed in E. coli, found to be soluble and then purified using affinity chromatography on a nickel column followed by removal of His tag via enterokinase cleavage (Figure 31). N-terminal His tagged AspC was expressed but was found in inclusion bodies so another vector for addition of a C-terminus His tagged onto AspC was cloned as well (Figure 30B). As with purification of GST-AspC, addition of sarkosyl in the lysis buffer increased solubility of HisAspC and allowed for purification on a nickel column (Figure 32). However, due to the low yield (100 μg per litre of culture) of His-AspC purification, other expression systems were explored.  Figure 30. His-tagged AspBMtb and AspCMtb expression vectors  Figure 30. His-tagged AspBMtb and AspCMtb expression vectors. aspBMtb was cloned into pET19b for expression of recombinant AspB with a N-terminus histidine tag. aspCMtb was cloned into pET30b for expression of recombinant AspC with a C-terminus histidine tag.  68  Figure 31. Purified AspBMtb and His-tag removal  Figure 31. Purified AspBMtb and His-tag removal. N-terminal His-tagged AspBMtb (~ 45 kDa) was expressed in E. coli Rosetta 2 (DE3) and purified to apparent purity using nickel column affinity chromatography. Tag removal using the engineered enterokinase cleavage site proceeded at room temperature for 2 h. Lane 1 Marker; Lane 2 purified His-AspB; Lane 3 cleavage negative control (no enterokinase added); Lane 4 protein to enterokinase ration at 500:1; Lane 5 1000: 1; Lane 6 2500:1; Lane 7 5000:1. Addition of too much protease resulted in a secondary cleavage product at approximately 31 kDa (Lanes 4, 5 and 6). The optimal ratio of protein to protease was determined to be 5000:1 (Lane 7)  69  Figure 32. Purification of C-terminal His-tagged AspCMtb  Figure 32. Purification of C-terminal His-tagged AspCMtb. His-tagged AspCMtb (~ 51 kDa) was expressed in E. coli Rosetta 2 (DE3) and purified to apparent purity using nickel column affinity chromatography. Lane 1 Marker; Lane 2 total expression in whole cells; Lane 3 insoluble fraction; Lane 4 cell free lysate; Lane 5 unbound fraction; Lane 6 blank; Lane 7 eluted His-AspCMtb. A large portion of recombinant AspC was found in the insoluble fraction (Lane 3) reducing the yield of the purification.  70  In order to obtain soluble AspC for purification many different systems were tested. Expression conditions such as low IPTG concentration induction, low temperatures, or heat shock did not increase protein solubility. Variation of the His tag placement or even expression without His tag also did not improve solubility. Co-expression of AspC with chaperone proteins such as GroEL-ES that can aid in protein folding and increase the recovery of soluble proteins did not improve the solubility of AspC. Even expression in M. smegmatis [97], a close relative of M. tuberculosis, generated mostly insoluble AspC. The systems tested for expression of soluble AspC are summarized in Table 8.  71  Table 8. Expression of AspCMtb in E. coli and M. smegmatis. A variety of systems were explored to generate enzymatically active M. tuberculosis AspC protein. One M. smegmatis and four E. coli and aspC expression plasmids were constructed and tested for expression under a variety of conditions. All constructs generated recombinant AspCMtb detectable as bands using SDS PAGE, only one (designated in bold print) was used for protein purification (in bold) that was subsequently used in biochemical analyses (aspartate aminotransferase activity using methods described in section 2.21 and 2.25). The results are found in Figure 32 and Table 9. Expression plasmid  Fusion tag  Expression strain  pGEX4T-1::aspC  N-terminal GST  BL21  pET19b::aspC  N-terminal His  Rosetta 2 (DE3)  pET30b::aspC  C-terminal His  Rosetta 2 (DE3)  pET30b::aspC  C-terminal His  BL21 (DE3)  Expression conditions 0.25 mM IPTG, 17 h, room temperature 42°C for 30 min then 0.5 mM IPTG, 17 h, room temperature 42°C for 30 min then 0.5 mM IPTG, 4 h, 37°C 42°C for 30 min then 0.2 mM IPTG, 17 h, room temperature 42°C for 30 min then 0.1 mM IPTG, 17 h, 10°C 1 mM IPTG, 5 h, 37°C 1 mM IPTG, 5 h, 37°C 1 mM IPTG, 5 h, 37°C 1 mM IPTG, 5 h, 37°C  pET30b::aspC  no fusion tag  Rosetta 2 (DE3)  pYUB1062::aspC  C-terminal His  mc2 4517  1 mM IPTG, 5 h, 37°C 0.5 mM IPTG, 3 h, 37°C 0.05 mM IPTG, 20 h, 15°C 0.05 mM IPTG, 22 h, 37°C  comments soluble with 1.5% sarkosyl and 2% triton tRNAs for rare codons supplied by pRARE2 tRNAs for rare codons supplied by pRARE2 tRNAs for rare codons supplied by pRARE2 tRNAs for rare codons supplied by pRARE2  dnaK-dnaJ-grpE and groES-groEL co-expression (pG-KJE8, Takara Bio Inc.) groES-groEL co-expression (pGro7, Takara Bio Inc.) groES-groEL-tig co-expression (pG-Tf2, Takara Bio Inc.) tig co-expression (pTf16, Takara Bio Inc.) tRNAs for rare codons supplied by pRARE2 tRNAs for rare codons supplied by pRARE2 Expression in M. smegmatis  Table 8. Expression of AspCMtb in E. coli and M. smegmatis  72  3. 22. Aspartate aminotransferase activity of AspBMtb and AspCMtb Purified recombinant his-tagged AspBMtb and AspCMtb expressed from E. coli each migrated as a single band on SDS-PAGE at their predicted theoretical masses (45 kDa and 51 kDa respectively) (Figures 31 and 32). The preparations were assayed for aspartate aminotransferase activity. The assay measures glutamate formation by transamination from aspartate to α-ketoglutarate. The specific activity measured for recombinant AspBMtb was measured at 2.62 x 10-2 units per milligram and recombinant AspCMtb has a specific activity for aspartate transamination at 5.34 x 10-2 units per milligram (Table 9). The specific activities of purified AspB and AspC were comparable to reported specific activities of other purified aspartate aminotransferases (Table 9) Steady-state kinetic parameters for AspBMtb were determined for the reaction of glutamate formation by transamination from aspartate to α-ketoglutarate (Table 10). The Michaelis constants, Km for aspartate and α-ketoglutarate determined for AspBMtb is comparable to that reported for E. coli aspartate aminotransferase AspCEco by Toney and Kirsch [136]. However, the catalytic constant, kcat, is over 500fold smaller than AspCEco making the efficiency of AspBMtb, as calculated by kcat/Km, a poor aspartate aminotransferase.  73  Table 9. Specific activities of purified aspartate aminotransferases. Enzyme  Organism  Specific activity (U/mg)  Reference  aspartate aminotransferase (from heart) AspC aspartate aminotransferase -2  Sus scrofa (pig)  620-690  [137]  E. coli Medcago sativa L.(alfalfa root) Bacillus sp. strain YM-2 M. tuberculosis M. tuberculosis  200-232 130-240  [138] [139] [140]  220  [141]  2.62 x 10-2 5.34 x 10-2  this study this study  aspartate aminotransferase AspB AspC  Table 9. Specific activities of purified aspartate aminotransferases.  Table 10. Comparison of steady state kinetic parameters determined for AspBMtb and AspCEco. Kinetic data for AspC from E. coli retrieved from Toney and Kirsch [136] AspB (Mtb) AspC (Eco) -1 -1 -1 Substrate kcat (s ) Km (mM) kcat/Km (M s ) kcat (s ) Km (mM) kcat/Km (M-1s-1) aspartate 1.07 ± 0.07 270 ± 10 1.83 87 000 0.29 ± 0.01 160 α -ketoglutarate 0.36 ± 0.03 800 ± 50 0.47 340 000 -1  Table 10. Comparison of steady state kinetic parameters determined for AspCMtb and AspCEco.  74  4. Discussion 4. 1. aspB and aspC expression is linked to whiB7 expression The putative aspartate aminotransferases aspB and aspC were both identified as genes in the WhiB7controlled regulon, suggesting that they might have roles in intermediary metabolism and intrinsic drug resistance. The transcription regulator WhiB7 is expressed in response to antibiotics as well as other stress conditions such as heat shock, iron starvation, and entry into stationary phase [24]. whiB7 is also implicated in M. tuberculosis virulence because it is upregulated within resting or activated macrophages [36]. A screen of hundreds of bioactive compounds revealed that the expression of whiB7 can be induced by compounds having diverse structures and targets and that the induction of whiB7expression is strongly affected by the reducing potential of the cytoplasm [35]. Here we further explore the link between metabolism and intrinsic drug resistance showing aspB and aspC have essential roles in both processes. In E. coli, aspartate is synthesized by aspCEco in which oxaloacetate receives an amino group from glutamate. The expression of aspCEco is reported to be constitutive [142] [143], based only on the observation that aspartate does not repress expression, unlike other transaminase paralogues [144]. The aspB and aspC genes were first linked to antibiotic resistance through microarray studies of the WhiB7-controlled transcriptome of M. tuberculosis. Given the homologies of M. tuberculosis and M. smegmatis aspB, aspC, and whiB7 genes (80%, 88%, and 71% amino acid identity respectively), we decided to study their regulation using qRT-PCR in the M. smegmatis model system. WhiB7 induced expression of aspC when whiB7 was either constitutively expressed from a heat shock promoter or highly induced by tetracycline (Figure 4 and 5B). However, there was no measurable change in aspBSMG in response to whiB7Mtb expression or tetracycline induction, suggesting that aspB is not antibiotic induced or under WhiB7’s control. The fact that aspCMsm expression responded to tetracycline as well as either whiB7Mtb (heat shock promoter expression) or whiB7Msm (heat shock promoter and tetracycline induced) demonstrated that whiB7 regulation of aspC is similar in both organisms and that M. smegmatis was a reliable surrogate model. The observations that expression of aspC was regulated by WhiB7 and aspC was upregulated by tetracycline pointed towards a role for AspC in the intrinsic resistance mechanisms of mycobacteria. whiB7 expression is autoregulated [29,35] and its activity as a transcriptional activator depends on other regulatory signals including the reducing potential of the cytoplasm [35]. One of these elements could be AspC (or dependent genes) because the activation (often over 1000-fold) of whiB7 in response to tetracycline was diminished (but not eliminated) in the ∆aspC mutant (Figure 5A). While aspC expression alone upregulated whiB7 slightly, the fact that there was no significant change in whiB7 expression in response to tetracycline in the hsp-aspCMtb strain provided evidence that aspC acts generally to amplify whiB7 expression in response to a primary metabolic inducer (Figure 6A). Results showed that while 75  expression of whiB7 is autoregulatory, full activation of the whiB7 promoter depends on WhiB7 [35] as well as AspC. It is likely that in response to antibiotics AspC expression acts to change the physiological state of the bacterium, which then promotes whiB7 expression. Thus whiB7 and aspC are components of the same autoregulatory loop. In contrast, constitutive expression of aspB strongly inhibited both whiB7 and aspC. Even under noninducing (no tetracycline) conditions, the expression of whiB7 was reduced by more than 90% (Figure 6A). After tetracycline treatment, whiB7 induction was reduced by 99.98% in the strain constitutively expressing aspBMtb (Figure 6A). AspB activity appears to generate a signal for downregulation of whiB7 that overrides the strong upregulatory signal generated by tetracycline. It would follow that if AspB expression prevented whiB7 expression, then aspC would also not be upregulated. Indeed, levels of aspC in ∆whiB7 (Figure 5B) and aspBMtb constitutive expression strains (Figure 6B) were very similar. aspB expression was unaffected by tetracycline treatment (Figure 5B), whiB7 or aspC expression; other aspects of aspB regulation remain to be elucidated. A model used for coordinated regulation of both aspB and aspC by whiB7 is presented schematically in Figure 33. Expression of aspB causes downregulation of whiB7 by suppressing signals generated by antibiotics that upregulate whiB7. The expression of aspC is dependent on whiB7. When whiB7 is upregulated in response to tetracycline, aspC expression is also upregulated in a whiB7-dependent manner; when aspB expression depresses the tetracycline-induced upregulation of whiB7, aspC upregulation is also abolished. The study on interrelation of whiB7, aspB, and aspC gene regulation showed that the regulation of expression of whiB7, a mycobacterial regulator of intrinsic resistance, is connected to expression of metabolic genes aspB and aspC.  76  Figure 33. Schematic representation of the proposed antibiotic induced regulatory network of whiB7, aspB, and aspC  Figure 33. Schematic representation of the proposed antibiotic induced regulatory network of whiB7, aspB, and aspC. Antibiotic treatment results in physiological stress which upregulates expression of whiB7. WhiB7 induces expression of the transcriptome which includes aspC. AspC acts to enhance the whiB7-inducing signals, which further amplifies whiB7 expression. AspB neutralizes/suppresses the signal, so that whiB7 and aspC are not upregulated when aspB is expressed.  77  4. 2. Intermediary metabolism of mycobacteria is linked to intrinsic antibiotic resistance Because expression of aspB and aspC were tightly linked to regulation of whiB7 and associated intrinsic resistance in mycobacteria, we reasoned that there might be a link between these aspartate aminotransferases and antibiotic resistance. Knock-out mutants of aspB and aspC were made using recombineering [95]. Both genes were non-essential, however aspC was required for optimal growth. This observation seemed consistent with results of transposon site hybridization (TraSH) screen of an insertion mutant library [93] as well as a high-density mutagenesis and deep sequencing study [52,93] to identify mutations that impair in vitro growth of M. tuberculosis. A deep sequencing transposon insertion mapping study recently revealed that certain portions of the coding sequence within the aspB coding sequence are required and function as non-protein-coding RNAs or cis regulatory elements [52]. The growth of the aspC transposon mutant was reduced compared to wild-type M. tuberculosis leading to the conclusion that aspC is essential for optimal growth. Observations that mutation of aspC affects growth of M. smegmatis, M. bovis BCG, and M. tuberculosis in culture suggest that the role of AspC is conserved across mycobacteria [93]. By analogy, it is also likely that aspC is required for optimal growth in vivo. However there is a complication with this analysis, it cannot be excluded that the preculture used for infection no longer contained aspC mutants and thus was not reported as required for survival during infection [145]. Our studies showed that intermediary metabolism of M. smegmatis was affected by the disruption of aspC; the ∆aspC mutant was impaired in using aspartate as nitrogen or carbon source (Table 3). Furthermore, the M. smegmatis ∆aspC mutant exhibited generalized growth defects such as slower growth rate and prolonged lag phase (Figure 7). Addition of α-ketoglutarate to the medium restored the growth rate to wild-type levels but the prolonged lag phase remained, suggesting that a different element caused that defect (Figure 8A). Passaging the ∆aspC mutant in α-ketoglutaratecontaining media to allow the strain to acclimatize did not shorten the lag, showing that the delay in growth was not due to the strain requiring time to activate alternative metabolic pathways (Figure 9). αketoglutarate was able to suppress other physiological deficiencies and consequently the antibiotic susceptibility of ΔaspC mutant. This demonstrates that AspC has roles in growth as well as antibiotic resistance (Figure 12), providing further evidence that mycobacterial intermediary metabolism is linked to intrinsic antibiotic resistance. Interestingly, the addition of oxaloacetate to the growth media produced a greater effect than αketoglutarate on the growth of wild-type. Oxaloacetate inhibited the growth of wild-type (Figure 16B-D). In essence, α-ketoglutarate seemed to increase bacterial fitness while oxaloacetate decreased fitness. The cause of the growth rate decrease in ∆aspC may be due to increased oxaloacetate in a model where AspC uses it as a substrate. α-ketoglutarate and oxaloacetate appeared to modulate the growth inhibitory effects of bacteriostatic antibiotic treatments (Table 5, Figure 16).  78  Oxaloacetate consistently potentiated the growth inhibition activity of tetracycline, clarithromycin and spectinomycin while α-ketoglutarate partially suppressed the inhibition by antibiotics. Similarily, growth of ΔaspC was multi-drug sensitive and addition of α-ketoglutarate to the medium suppressed this effect (Figure 12). It is hypothesized that increased oxaloacetate in ΔaspC leads to growth inhibition and antibiotic sensitivity. The fact that the antibiotic sensitivity profile of the ΔaspC mutant was similar to that of ∆whiB7 suggested that oxaloacetate metabolism may also be disrupted in the whiB7 mutant. Considering the transcriptional activation of aspC by WhiB7, WhiB7 may act to regulate oxaloacetate and α-ketoglutarate pools in the bacterium and AspC may be a primary determinant of intrinsic resistance. Oxaloacetate toxicity has been reported in Lactococcus fermentation causing reduced growth rates and lower biomass yields [146,147]. Although the reason for its toxicity was not explained, it was speculated that high oxaloacetate concentrations can competitively inhibit other essential metabolic enzymes, such as those in the TCA cycle. Transamination of oxaloacetate would relieve the stress [148]. Another method to detoxify oxaloacetate is to export it out of the cell via malate and citrate transporters [148,149]. These transporters have broad specificities and also transport other 2oxocarboxylates. The transport of 2-oxocarboyxlates such as oxaloacetate requires exchange with a partner (precursor/product exchange) and results in the generation of a membrane potential [149]. Addition of α-ketoglutarate may relieve the transmembrane stress generated by oxaloacetate (Figure 16B-D) either by acting as an exchange molecule having similar ionic properties, or by inducing expression of additional transporters. The opacification of 7H10 agar by the ΔaspC mutant suggested precipitation of components in the medium, perhaps involving the export of acidic metabolites. The precipitate might be the metabolite itself or other components of the medium7H10. Acidic metabolites such as pyruvate, malate, succinate, lactate, and fumarate have been detected in M. tuberculosis culture supernatant during growth [150]. Extracellular accumulation of secreted succinic acid and lactic acid was observed in anaerobic cultures of M. tuberculosis [74]. Under these conditions, decreased growth rates as well as accumulation of reduced cofactors such as NADH (resulting in a high NADH/NAD+ ratio), are probably due to the absence of oxygen as the terminal electron acceptor [74]. The secretion of succinic acid was due to reversal of the TCA cycle, allowing the bacterium to maintain sufficient membrane proton gradient for persistence, but not active growth [74]. The ΔaspC mutant exhibited shared features of M. tuberculosis physiology during hypoxic/anaerobic growth, including reduced growth rate, acid secretion, and a more reduced pool of NADH. It is plausible that deletion of aspC perturbed the metabolism of bacterium in such a way to cause the reversal of the TCA cycle. An atypical TCA cycle in the ΔaspC mutant could have implications in antibiotic resistance, as the activity of bactericidal antibiotics has been suggested to be linked to the TCA cycle [67]. Metabolic distress due to the deletion of aspC is not yet fully understood but is likely due to oxaloacetate toxicity (Figure 16B-D). Work thus far has shown that inactivation of aspC has widespread  79  consequences on the physiology of M. smegmatis and additional study is required to understand the mechanism. Similar to the ΔaspC mutant, constitutive expression of aspBMtb also increased susceptibility of M. smegmatis to antibiotics (Table 4), an effect that is likely mediated through whiB7. This is based on our studies demonstrating that whiB7 expression was strongly repressed by aspBMtb expression. In addition, the susceptibility profile of hsp-aspBMtb was similar that of ∆whiB7. We conclude that enzyme activity of AspB may create a signal which downregulates whiB7. Transcriptional activation of whiB7 promoted a reduced environment while oxidizing conditions decreased whiB7 transcription [35]. Drug susceptibility of the ΔaspC mutant was revealed by growth kinetics analyses that agreed with MTT-based MIC determination (Table 4). When the curve representing growth kinetics of the mutant was much steeper than the wild-type curve (clarithromycin, roxithromycin, rifampicin), the fold susceptibility of mutant ΔaspC was higher (eight to sixteen fold more susceptible as compared to wild-type). A slight increase in susceptibility (two to four compared to wild-type) was reflected in comparisons of mutant and wild-type curves. While these curves had similar slopes, the curve representing the mutant was positioned above the wild-type curve (clindamycin, fusidic acid, and spectinomycin) (Figure 22). The growth curve analysis once again showed that deletion of aspC caused the strain to be more susceptible to a number of antibiotics. In addition, this analysis revealed that the susceptibility increase was due to the ΔaspC mutant having an increased lag phase, akin to the effect of oxaloacetate and antibiotics on the growth of wild-type (Table 5 and Figure 16). Paradoxically, while deletion of aspC augmented inhibition of M. smegmatis growth by antibiotics, survival kinetics indicated this mutation reduced the bacteriocidal activity of clarithromycin (Figure 15). Clarithromycin, a macrolide, is generally considered to be bacteriostatic but the overwhelmingly high concentration used in our experiments was sufficient to reduce the cell viability of both wild-type and mutant. The difference in log reduction of cell numbers can be partly attributed to the fact that the growth of the ΔaspC mutant may have already plateaued when the antibiotic was added. Slow-growing or nongrowing cells are not very susceptible to many antimicrobial agents [151]. Indeed the ΔaspC mutant exhibits growth defects and doubles much more slowly than wild-type M. smegmatis, which could contribute to the reduced killing by clarithromycin and also promote more persisters that are tolerant to antibiotics. This resistance phenomenon may also be due to a subpopulation of persisters that are genetically identical (not mutants) but are less sensitive to killing by antibiotics because they are either not actively growing or metabolising very slowly [152] [153]. The observation that the ΔaspC mutant was simultaneously sensitive to the bacteriostatic effects of clarithromycin but resistant to the bacteriocidal killing by clarithromcyin at higher concentrations highlights that aspC has varied effects on the bacteria’s physiology. The hypothesized increased oxaloacetate in the ΔaspC mutant may potentiate the activity of bacteriostatic antibiotics while redox stress, reflected by the increased catalase activity may protect the bacteria from killing by bacteriocidal antibiotics. More experimentation using actively 80  growing cultures is required to determine whether the bactericidal and bacteriostatic effects of clarithromycin reflect independent physiological effects elicited by the aspC inactivation.  81  4. 3. Role of AspB and AspC in mycobacteria physiology Bioinformatic analysis of the sequences of aspB and aspC from M. tuberculosis and M. smegmatis suggested that both genes encode aspartate aminotransferases (Figures 20, 21, and Table 6). Both AspB and AspC from M. tuberculosis showed aspartate transaminase activity. The specific activities of AspB and AspC were much lower than specific activities reported for other aspartate aminotransferases (Table 9). Steady-state kinetic parameters for AspBMtb were determined for the reaction of glutamate formation by transamination from aspartate to α-ketoglutarate (Table 10) (experiments performed by Mr. Antonio Ruzzini). The Michaelis constants, Km for aspartate and α-ketoglutarate determined for AspBMtb were comparable to those reported for E. coli aspartate aminotransferase AspCEco by Toney and Kirsch [136]. However, the catalytic constant, kcat, is over 500-fold smaller than AspCEco making AspBMtb, a poor aspartate aminotransferase, as calculated by kcat/Km. The physiological role of AspB has yet to be fully characterized; perhaps AspB requires different conditions or substrates. Purified recombinant AspCMtb was also shown biochemically to possess aspartate aminotransferase activity. Over-expression of M. tuberculosis proteins in E. coli often results in the proteins aggregating to form insoluble inclusion bodies. It is estimated that only 30-50% of tuberculosis proteins can be expressed in a soluble form by E. coli [154]. We experienced similar problems with over-expression of AspB and AspC. AspB was expressed as a soluble recombinant protein, while AspC could be over-expressed, but was mostly present in inclusion bodies making purification of native protein difficult. Expression of AspC with GST protein tag (Figure 25) significantly improved solubility and allowed subsequent purification using affinity chromatography (Figure 26), but since the size of the GST tag is comparable to AspC (25 kDa and 47 kDa, respectively) its removal was required so that the activity of AspC would not be sterically hindered by the GST tag. Unfortunately, due to the presence of secondary protease cleavage sites and general instability of the protein, it was not possible to obtain purified, tag-free, AspC from this construct (Figures 28 and 29). Mycobacterial chaperones may be required to aid in the folding and soluble expression of M. tuberculosis proteins. Alternatively, codon usage might need to be optimized for expression of high GC content genes in E. coli, a more AT rich organism. M. smegmatis has been used successfully to produce soluble recombinant protein of M. tuberculosis proteins that were previously insoluble in E. coli [97,154,155]. Expression in Rosetta2™ E. coli and M. smegmatis [97] was attempted but recombinant AspC was still only found in inclusion bodies (Table 8). aspB maps within a cluster of cholesterol catabolic genes in the M. tuberculosis and M. smegmatis genomes that may be related to virulence of M. tuberculosis. aspB is within a putative operon controlled by a tetR-type transcriptional repressor, KstR2 [92]. aspB (MSMEG_6017) was expressed 100-fold more when kstR2 was deleted in M. smegmatis showing that KstR2 exerts negative regulation of aspB [92]. KstR2 controls the expression of genes in the cholesterol degradation pathway, a function that is important in M. tuberculosis virulence as cholesterol is a critical carbon source during chronic infection 82  [156,157]. Other genes in the operon, fadD3 (Rv3561), fadE32 (Rv3563), fadE33 (Rv3564), are essential for growth on cholesterol [158]. While aspB was not identified as an essential gene for growth on cholesterol, it is upregulated along with other genes in the operon during infection of both the resting and IFN-γ–activated macrophages [159], suggesting that AspB may be part of the metabolic adaptive pathway that M. tuberculosis requires for growth on cholesterol. Other genes in the KstR2 operon in addition to aspB are putative fatty acid CoA synthetase (fadD3) and acyl CoA dehydrogenases (fadE31, fadE32, and fadE33) with roles in β-oxidation. β-oxidative degradation of odd-chain-length fatty acids and cholesterol can yield molecules like propionyl CoA and acetyl CoA [160,161]. The degradation of branched chain amino acids (valine, leucine, and isoleucine) also produces propionate and requires actions of transaminases [162]. It has been speculated that AspB may act on a metabolite or intermediate of cholesterol degradation that mimics the structure of branched chain amino acids during β-oxidation-like degradation of a cholesterol degradation intermediate to produce propionyl CoA which can enter the methyl citrate cycle and be used for energy generation in [163].  83  4. 4. Implicated roles for AspC in redox homeostasis and oxidative stress detoxification Homologs AspB and AspC may both have aspartate aminotransferase activity but their roles in redox homeostasis, antioxidation, and antibiotic resistance differ. AspC acts in the WhiB7-directed pathway to modulate physiology, and may serve to counteract damage by oxidative stress. AspB, however, does not appear to do the same; deletion or constitutive expression strains of aspB did not show any significant changes in catalase activity or NAD+/H pools compared to wild-type. Moreover, AspB appears not to be the primary aspartate aminotransferase as deletion of the gene did not affect M. smegmatis growth whereas aspC was required for optimal aspartate catabolism and general fitness of the strain (Figure 7, Table 3). Deletion of both aspB and aspC did not cause M. smegmatis to become auxotrophic (Figure 10A). Amino acid transaminases have broad activities; aspartate auxotrophy in E. coli required deletion of three genes: aspC (aspartate aminotransferase), tyrB (aromatic amino acid aminotransferase), and ilvE (branched chain amino acid aminotransferase) [144]. Therefore, inactivation of additional transaminase genes may be needed to generate a M. smegmatis aspartate auxotroph. The ΔaspBΔaspC double mutant displayed many of the same growth defects of ΔaspC (Figures 7 and 10), further demonstrating that deletion of aspB does not significantly affect in vitro growth on glycerol. Antibiotics triggered changes in the cytoplasmic redox potential activated whiB7 expression. WhiB7 contributes to the maintenance of mycothiol pools (MSH and MSSM) in addition to regulating the reduction of oxidized mycothiol (MSSM) in response to antibiotics (erythromycin) [35]. Since aspC expression was positively regulated by WhiB7 and the deletion of aspC caused increased sensitivity to many antibiotics, we hypothesized that AspC may play a role in WhiB7-mediated maintenance of redox homeostasis. Quantification of NADH and NAD+ pools revealed that deletion of aspC resulted in a large increase in the pools of NADH and NAD+, as well as the NADH/NAD+ ratio (Figure 17). Inability to maintain redox homeostasis increases susceptibility to oxidative stress, as evidenced by mycothiol mutants exhibiting hypersensitivity to reactive oxygen compounds [164] [165] [166] [167] [168]. The ΔaspC mutant exhibited increased sensitivity to oxidative stress inducers like hydrogen peroxide and menadione (Figure 18). The growth deficiencies of the ΔaspC mutant may have contributed to this phenotype so additional experiments would need to be performed (such as a kinetic kill analysis) to confirm the sensitivity of the ΔaspC mutant to ROS inducers. We measured catalase activity in the ΔaspB and ΔaspC mutants to determine whether the sensitivity to oxidative stress inducers was due to changes in the detoxifying enzymes that maintain oxidative homeostasis. The catalase assay used in this study measured the disappearance of hydrogen peroxide and could reflect the activity of either catalase (KatG) or alkyl hydroperoxidase (AhpC). Paradoxically, while the ΔaspC mutant was apparently sensitive to peroxide, it had four-fold higher levels of catalase activity compared to wild-type indicating that the mutant requires the presence of more detoxifying enzymes to maintain oxidative homeostasis (Figure 19A). The increased catalase activity correlates with the ΔaspC mutant displaying increased sensitivity to 84  isoniazid, as KatG is responsible for activation of this prodrug [169] [170] [171]. It is interesting to note that while addition of α-ketoglutarate to the growth medium suppressed the drug susceptibility of the ΔaspC mutant to tetracycline, clarithromycin, and spectinomycin, the presence of α-ketoglutarate significantly increased the MIC of the strain to isoniazid making the strain 32-fold more resistant to isoniazid than wild-type (Figure 12). As previously noted, the activity of isoniazid, a prodrug, relies on a conversion catalyzed by KatG and the resistance to isoniazid in the presence of α-ketoglutarate represents a link between α-ketoglutarate and KatG and therefore oxidative stress detoxification. katG expression is induced in M. tuberculosis, M. bovis BCG, M. avium, and M. marinum in response to hydrogen peroxide [172] [173] [174] [175] [176]. With similar regulation by furA, one would expect katG would also be upregulated in M. smegmatis in response to hydrogen peroxide. However, the KatG response to hydrogen peroxide in M. smegmatis is still unclear. There has been evidence for [172] and against [176] the transcriptional upregulation of katG in M. smegmatis in response to hydrogen peroxide. Our measurement of catalase activity in wild-type cell lysates showed no change in catalase activity after hydrogen peroxide exposure (Figure 19B). It would seem that our results support lack of katG upregulation in response to hydrogen peroxide in M. smegmatis but it is plausible that there is post-transcriptional regulation of KatG. Nonetheless, the ΔaspC mutant, whose levels of catalase activity prior to hydrogen peroxide challenge are already elevated, responded to hydrogen peroxide with a decrease in catalase activity. The reason for this decrease in catalase activity detected after hydrogen peroxide exposure is unclear; the decrease in catalase activity may be due to activation of other detoxification enzymes such as AhpC. Alkyl hydroperoxidase is sufficient to protect M. tuberculosis from organic peroxides when katG is mutated [174]. Nonetheless, deletion of aspC results in a change in oxidative stress sensitivity and response of M. smegmatis to hydrogen peroxide oxidative stress. Paradoxically, the increased KatG activity could potentially increase oxygen radical formation; mycobacterial KatG possesses NADH oxidase activity in addition to catalase and peroxidase activities. KatG could oxidize NADH in an oxygen-dependent manner, forming either hydrogen peroxide or superoxide radicals [177]. The ΔaspC mutant concurrently has both increased catalase and NADH levels, supplying the substrates for radical formation via NADH oxidation, further adding to oxidative stress levels. The deletion of aspC may shift the metabolism of the bacterium to produce more ROS by-products during regular respiration. Aspartate is the precursor molecule in the mycobacterial NAD biosynthesis pathway. In the first step in this biosynthetic pathway aspartate is oxidized to α-iminosuccinate, producing hydrogen peroxide as a by-product. Deletion of aspC renders the strain unable to transaminate aspartate, leading to an accumulation of aspartate that may be fed into NAD+ biosynthesis reactions generating hydrogen peroxide and redox imbalance through increased NADH. Reducing NADH can drive the production of reactive oxygen from hydrogen peroxide through Fenton’s reaction, resulting in DNA damage (Figure 2) [178]. Bactericidal antibiotics have also been shown to cause cell 85  death via the generation of hydroxyl radicals [67]. AspC has a critical role in redox homeostasis and deletion of aspC causes an imbalance in redox homeostasis that increases susceptibility to antibioticgenerated oxidative stress.  86  4. 5. Concluding remarks Traditional understanding of antibiotic resistance focuses on the idea that antibiotics have one target and resistance is based on modification of that target or antibiotic. However, there is increasing evidence showing that physiology plays a role in determining intrinsic levels of antibiotic resistance. Intrinsic antibiotic resistance can be due to genes that are induced by antibiotics and when expressed, become functional antibiotic resistance determinants. In this study, we have presented evidence that AspB and AspC are aspartate aminotransferases that are involved in amino acid intermediary metabolism and WhiB7-controlled intrinsic resistance. Understanding of the microbial factors that affect intrinsic resistance in M. tuberculosis could be beneficial to the design of new regimens for the management of tuberculosis, contributing to the elimination of tuberculosis as a global health problem.  87  References  1. WHO (2011) Global tuberculosis control: WHO report 2011. WHO Press. 2. Gandhi NR, Nunn P, Dheda K, Schaaf HS, Zignol M, et al. (2010) Multidrug-resistant and extensively drug-resistant tuberculosis: a threat to global control of tuberculosis. 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