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Role of Mycobacterium tuberculosis protein tyrosine phosphatase A in the pathogenesis of tuberculosis Wong, Dennis Dick-Hang 2012

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ROLE OF MYCOBACTERIUM TUBERCULOSIS PROTEIN TYROSINE PHOSPHATASE A IN THE PATHOGENESIS OF TUBERCULOSIS by Dennis Dick-Hang Wong B.Sc., The University of British Columbia, 2007  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in  The Faculty of Graduate Studies  (Experimental Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  May 2012  © Dennis Dick-Hang Wong, 2012  ABSTRACT One of the main mechanisms by which the etiological agent of tuberculosis (TB), Mycobacterium tuberculosis (Mtb), survives in the host macrophage is by its capacity to arrest phagosome acidification and fusion with lysosomes. This Mtb feature is associated with phagosomal exclusion of the Vacuolar H+-ATPase (VATPase)  proton  pump,  which  normally  drives  luminal  acidification  of  membranous organelles. Although this phenomenon has been known for 20 years, the mechanism by which Mtb blocks phagosome acidification remains obscure. Research on Mtb pathophysiology shows that a wide array of Mtb lipid and protein molecules contribute to maintaining the mycobacterial phagosome in an immature state. We have previously found that Mtb protein-tyrosine Phosphatase A (PtpA) is required for Mtb infection of human macrophages. PtpA is secreted into the macrophage cytosol to inactivate the human VPS33B, a component of the Class C VPS Complex that regulates endosomal membrane fusion. VPS33B dephosphorylation by PtpA results in the inhibition of phagosome-lysosome fusion. In this work, we demonstrated that, in addition to its phosphatase activity, PtpA is also capable of binding to subunit H of the macrophage V-ATPase complex, indicating that PtpA can directly disrupt phagosome acidification. Indeed, we found that a Mtb strain expressing a VATPase-binding defective mutant PtpA protein failed to inhibit phagosome acidification, and expression of wild-type PtpA protein in E. coli-infected macrophages is sufficient to block acidification. Furthermore, we showed that the Class C VPS complex associates with V-ATPase during phagosome maturation,  	
    ii  identifying a novel role for V-ATPase in coordinating endocytic membrane fusion. PtpA interaction with host V-ATPase is required for the previously reported dephosphorylation of VPS33B and subsequent exclusion of V-ATPase from the phagosome during Mtb infection. Taken together, these findings reveal, for the first time, the long-sought mechanism behind the lack of acidification in the mycobacterial phagosome. Interestingly, we found that PtpA is also a substrate for the newly identified Mtb protein tyrosine kinase PtkA, which is encoded within a shared operon with PtpA, indicating a regulatory control of PtpA during Mtb infection. Understanding the pathophysiological importance of PtpA in Mtb infection  might  contribute  to  the  development  of  novel  antitubercular  therapeutics.  	
    iii  PREFACE Parts of this thesis have been published in the peer-reviewed journals listed below in chronological order: 1. Bach, H., Papavinasasundaram, K. G., Wong, D., Hmama, Z., & Av-Gay, Y. (2008) Mycobacterium tuberculosis virulence is mediated by PtpA dephosphorylation of human vacuolar protein sorting 33B. Cell Host Microbe 3(5): 316-322. In this study, I was responsible for performing several experiments, contributing to the development of work shown in Figure 2 (A, B, C, and E) and Figure 4 (A and C) and opening a lead into the work described in this thesis. H. Bach was responsible for the experimental design, performing experiments and writing the manuscript. K. G. Papavinasasundaram constructed the ΔptpA deletion Mtb mutant. Y. Av-Gay designed the study and wrote the manuscript. This published work is located in Section 1.3.2. 2. Bach, H., Wong, D., & Av-Gay, Y. (2009) Mycobacterium tuberculosis PtkA is a novel protein tyrosine kinase whose substrate is PtpA. Biochem J 420(2): 155-160. In this study, I was responsible for performing most of the experiments, contributing to the development of work shown in Figure 1, Figure 2 (A, B and D), Table 1 and Table 2. H. Bach was responsible for the experimental design and writing the manuscript. Y. Av-Gay wrote the manuscript. This published work is located in Section 3.2. 3. Chao, J., Wong, D., Zheng, X., Poirier, V., Bach, H., Hmama, Z., & AvGay,  Y.  (2010)  Protein  kinase  and  phosphatase  signaling  in  Mycobacterium tuberculosis physiology and pathogenesis. Biochim Biophys Acta 1804(3): 620-627. In this review article, I was responsible for writing part of the manuscript, contributing to the sections titled “3. Protein tyrosine phosphorylation”, “4. 	
    iv  Protein tyrosine phosphatase A (PtpA), “5. Protein tyrosine phosphatase B (PtpB)”, “6. Inhibitors of Mtb protein tyrosine phosphatases” and “7. Protein tyrosine kinase A (PtkA)”. J. Chao and X. Zheng contributed to the session titled “2. PknG”, Figure 1 and Table 1. V. Poirier contributed to the session titled “4. Protein tyrosine phosphatase A (PtpA)” and Figure 2. Y. Av-Gay wrote the “Introduction” and “Concluding Remarks”. J. Chao edited and finalized the manuscript. This published work is located in Sections 1.3.2 and 4.3. 4. Mascarello, A., Chiaradia, L. D., Vernal, J., Villarino, A., Guido, R. V., Perizzolo, P., Poirier, V., Wong, D., Martins, P. G., Nunes, R. J., Yunes, R. A., Andricopulo, A. D., Av-Gay, Y., & Terenzi, H. (2010) Inhibition of Mycobacterium tuberculosis tyrosine phosphatase PtpA by synthetic chalcones: kinetics, molecular modeling, toxicity and effect on growth. Bioorg Med Chem 18(11): 3783-3789. In this study, I was responsible for performing the in vivo analysis of the efficacy of a family of synthetic chalcones as a novel drug against Mtb in the THP-1 macrophage infection model. My work contributed to Figure 4 and Section 2.4 “Inhibitors of MPtpA reduced mycobacterial survival in macrophages”. Y. Av-Gay designed the study and participated in writing the manuscript. This published work is located in Section 4.3. 5. Wong, D., Bach, H., Sun, J., Hmama, Z., Av-Gay, Y. (2011) Mycobacterium tuberculosis protein tyrosine phosphatase (PtpA) excludes host vacuolar-H+-ATPase to inhibit phagosome acidification. Proc Natl Acad Sci U S A 108(48): 19371-6. In this study, I designed and performed all experiments. J. Sun and Z. Hmama assisted in work related to Supplemental Figure S7A and S8. I wrote the manuscript. Hmama, Z. helped with editing of the manuscript. Bach, H. was involved in experimental design. Y. Av-Gay participated in  	
    v  designing the study and writing the manuscript. This published work is located in Section 3.1. 6. Wong, D. and Av-Gay, Y. Mycobacterium tuberculosis Phosphatase Interference with Human Signaling Pathways. Manuscript in Preparation. In this work, I was responsible for writing a review article on the current knowledge of phosphatases in Mycobacterium tuberculosis. Y. Av-Gay assisted in writing and editing the manuscript. This work is located in Section 1.3.2. The work in this dissertation is conducted with approval from University of British Columbia Office of Research and in accordance with the University of British Columbia Polices and Procedures, Biosafety Practices and Public Health Agency of Canada Guidelines. Biohazard Approval Certificate: B10-0112. Work with radioactive materials is approval by University of British Columbia Committee on Radioisotopes and Radiation Hazards. Radioisotope License: MEDI-3175-13.  	
    vi  TABLE OF CONTENTS ABSTRACT ...........................................................................................................ii PREFACE .............................................................................................................iv TABLE OF CONTENTS ......................................................................................vii LIST OF TABLES .................................................................................................xi LIST OF FIGURES ..............................................................................................xii LIST OF ABBREVIATIONS ...............................................................................xiv ACKNOWLEDGEMENTS ................................................................................xviii CHAPTER 1: INTRODUCTION .............................................................................1 1.1 Tuberculosis .......................................................................................................... 1 1.1.1 Disease epidemiology ...................................................................................... 1 1.1.2 Origin of human Tuberculosis .......................................................................... 2 1.1.3 Mycobacterium tuberculosis ............................................................................ 3 1.1.4 Infection cycle .................................................................................................. 5 1.2 Phagocytosis ......................................................................................................... 8 1.2.1 Phagosome formation ...................................................................................... 8 1.2.2 Phagosome maturation .................................................................................. 10 +  1.2.2.1 Vacuolar H -ATPase ............................................................................................ 14 1.2.2.2 Class C VPS ......................................................................................................... 17  1.3 Mtb arrest of phagosome maturation ............................................................... 19 1.3.1 Overview ........................................................................................................ 19 1.3.2 Mtb phosphatase interference with host signaling pathways ......................... 22 1.3.2.1 PtpB ...................................................................................................................... 24 1.3.2.2 SapM .................................................................................................................... 26 1.3.2.3 PtpA ...................................................................................................................... 28  1.4 Aims of the study ................................................................................................ 32  CHAPTER 2: MATERIALS AND METHODS......................................................34 2.1 Reagents and chemicals .................................................................................... 34 2.1.1 Commercial reagents ..................................................................................... 34 2.1.2 Antibodies ...................................................................................................... 35  	
    vii  2.2 Bacteria ................................................................................................................ 35 2.2.1 Strain maintenance and generation ............................................................... 35 2.2.2 Bacterial preparation for infection .................................................................. 36 2.2.3 Mycobacterial lysis ......................................................................................... 37 2.2.4 Transformation of mycobacteria .................................................................... 37 2.3 Cell culture .......................................................................................................... 37 2.3.1 Maintenance of tissue culture and differentiation ........................................... 37 2.3.2 Single transfection of differentiated THP-1 .................................................... 38 2.3.3 Double transfection of differentiated THP-1 ................................................... 38 2.4 Vectors and DNA manipulation ......................................................................... 38 2.4.1 RNA isolation and cDNA synthesis ................................................................ 38 2.4.2 Cloning ........................................................................................................... 39 2.4.3 Site-directed mutagenesis ............................................................................. 41 2.5 Protein expression and purification .................................................................. 42 2.5.1 Expression of recombinant proteins in E. coli ................................................ 42 2.5.2 Expression of recombinant proteins in M. smegmatis ................................... 43 2.6 In vitro pull down assay ..................................................................................... 44 2.7 ALPHAScreen protein-protein interaction assay ............................................. 44 2.8 Mycobacterial “Split-Trp” protein-protein interaction assay .......................... 45 2.9 Infection ............................................................................................................... 46 2.9.1 Infection of THP-1 .......................................................................................... 46 2.9.2 Phagocytosis assay ....................................................................................... 46 2.10 Immunoprecipitation ........................................................................................ 46 2.11 Measurement of phagosomal pH .................................................................... 47 2.11.1 Overview ...................................................................................................... 47 2.11.2 Phagosomal pH of Mtb phagosomes ........................................................... 48 2.11.3 Phagosomal pH of E. coli phagosomes ....................................................... 49 2.12 Digital confocal microscopy ............................................................................ 49 2.12.1 Intracellular staining ..................................................................................... 49 2.12.2 Microscopy ................................................................................................... 50 2.13 In vitro phosphatase activity assay ................................................................ 50 2.14 In vitro PtpA secretion analysis ...................................................................... 50 2.15 FACS analysis of antibody specificity ............................................................ 51 2.16 In vitro kinase assay ......................................................................................... 52  	
    viii  2.17 Phospho-amino acid analysis ......................................................................... 52  CHAPTER 3: RESULTS......................................................................................54 3.1 Mtb PtpA excludes host Vacuolar-H+-ATPase to inhibit phagosome acidification ............................................................................................................... 54 3.1.1 Introduction .................................................................................................... 54 3.1.2 Mtb PtpA binds subunit H of human V-ATPase ............................................. 54 3.1.3 PtpA binds to amino acid 220-402 of subunit H ............................................. 55 3.1.4 The C-terminal alpha helix of PtpA binds subunit H ...................................... 55 3.1.5 PtpA binds V-ATPase subunit H in vivo ......................................................... 58 3.1.6 PtpA binding to subunit H is required for Mtb intracellular survival ................ 59 3.1.7 PtpA inhibits phagosome acidification ........................................................... 60 3.1.8 The V-ATPase machinery recruits the Class C VPS complex ....................... 63 3.1.9 Mtb PtpA blocks the interaction between Class C VPS and V-ATPase complexes ............................................................................................................... 64 3.1.10 VPS33B remains phosphorylated in THP-1 infected with binding-defective PtpA ........................................................................................................................ 66 3.1.11 PtpA binding to subunit H participates in the exclusion of the V-ATPase from the Mtb phagosome ................................................................................................ 68 3.2 PtpA is a substrate for the novel protein-tyrosine kinase Mtb PtkA .............. 73 3.2.1 Introduction .................................................................................................... 73 3.2.2 PtpA interacts with the protein encoded by Rv2232 ...................................... 75 3.2.3 ORF Rv2232 encodes an autophosphorylated protein-tyrosine kinase ......... 76 3.2.4 Sequence-function analysis of PtkA .............................................................. 78 3.2.4.1 Mutation of lysine residues ................................................................................... 78 3.2.4.2 Mutation of tyrosine residues ................................................................................ 79 3.2.4.3 Mutation of D85 .................................................................................................... 80  3.2.5 PtpA is a substrate of PtkA ............................................................................ 80  CHAPTER 4: DISCUSSION ................................................................................85 4.1 PtpA exclusion of the V-ATPase from phagosomal membrane ..................... 85 4.2 PtpA is a substrate for the protein-tyrosine kinase PtkA ............................... 91 4.3 Inhibitors against PtpA as novel antituberculosis drug ................................. 95 4.4 Future directions ................................................................................................. 98  REFERENCES ..................................................................................................102 	
    ix  APPENDIX A: SUPPLEMENTAL FIGURES FOR SECTION 3.1 .....................124  	
    x  LIST OF TABLES 	
   Table 1. Oligonucleotides used for DNA cloning in this study .............................41 Table 2. Oligonucleotides used for SDM in this study .........................................42 Table 3. Phagosomal pH of Mtb strains in THP-1 ...............................................62 Table 4. Autophosphorylation kinetic values of parental and mutated PtkA ........80 Table 5. Kd values of PtpA interacting with parental and mutated PtkA proteins.83  	
    xi  LIST OF FIGURES  Figure 1. Estimated TB incidence rates (2010). ....................................................2 Figure 2. Phylogeny of selected mycobacterial species. .......................................4 Figure 3. Mtb infection cycle.. ................................................................................6 Figure 4. Receptor and signaling interactions during phagocytosis of microbes .10 Figure 5. Stages of phagosome maturation. ........................................................11 Figure 6. Vacuolar H+-ATPase.. ..........................................................................17 Figure 7. Working model for Class C VPS function in yeast ................................19 Figure 8. Model of Mtb inhibition of mycobacterial phagosome maturation. ........22 Figure 9. Scheme of PtpA catalytic activity. .........................................................30 Figure 10. PtpA interacts with V-ATPase subunit H in vitro .................................56 Figure 11. Computer-generated model of PtpA complexed with the V-ATPase subunit H.. ............................................................................................................57 Figure 12. PtpA and V-ATPase subunit H interacts in vivo .................................59 Figure 13. Mtb PtpA inhibits phagosome acidification.. .......................................61 Figure 14. Co-immunoprecipitation of V-ATPase subunits with Class C VPS in THP-1...................................................................................................................64 Figure 15. Mtb PtpA disrupts the interaction between the Class C VPS and VATPase complexes during infection ....................................................................65 Figure 16. Western blot analysis of VPS33B phosphorylation in vivo. ................67 Figure 17. Confocal microscopy of THP-1 infected with E. coli or indicated Mtb strains.. ................................................................................................................70 Figure 18. Confocal microscopy of infected THP-1 macrophage doubly transfected with GFP-subunit H and DsRed2-VPS33B.. .....................................72 Figure 19. Rv2232 interacts with PtpA in vitro .....................................................75 Figure 20. Rv2232 autophosphorylation in vitro ..................................................76 Figure 21. Phospho-amino acid analysis of PtkA. ...............................................77 Figure 22. Mutational studies of PtkA catalytic mechanism .................................79 Figure 23. PtkA phosphorylates PtpA in vitro.. ....................................................81 Figure 24. PtkA phosphorylates PtpA in dose- and time-dependent manner ......81 	
    xii  Figure 25. PtkA phosphorylates PtpA on tyrosine residues .................................84 Figure 26. A model for the specific exclusion of V-ATPase and the inhibition of mycobacterial phagosome acidification by PtpA. ................................................88 Figure 27. Mtb survival in infected THP-1 treated with chalcone inhibitors.. .......96 Figure 28. In vitro analysis of subunit H and PtpA interaction. ..........................124 Figure 29. PtpAL146A is a phosphatase-active mutant defective in binding subunit H. .......................................................................................................................126 Figure 30. Western blot analysis of PtpA expression in complemented ΔptpA strain and in vitro PtpA secretion using α-PtpA antibodies ................................127 Figure 31. Calibration of THP-1 phagosomal pH during Mtb infection with FACS. ...........................................................................................................................128 Figure 32. Measurement of phagosomal pH in transfected THP-1 infected with E. coli......................................................................................................................130 Figure 33. Analysis of antibodies specificity.. ....................................................132 Figure 34. Immunostaining control experiment with Mtb-infected THP-1. .........134 Figure 35. Immunostaining control experiment with E. coli-infected THP-1. .....135  	
    xiii  LIST OF ABBREVIATIONS Ace  Acetamide  ADP  Adenosine diphosphate  Arf  ADP ribosylation factor  Arg  Arginine  Asp  Aspartate  BCG  Bacillus Calmette-Guérin  CFP  Culture filtrate proteins  CFU  Colony forming units  CORVET  Class C core vacuole/endosome tethering complex  CR3  Complement receptor 3  Cys  Cysteine  DMEM  Dulbecco's modified Eagle medium  DMSO  Dimethyl sulfoxide  DTT  Dithiothreitol  EDTA  Ethylenediaminetetraacetic acid  EEA1  Early endosome antigen 1  FACS  Fluorescence activated cell sorting  	
    xiv  FCS  Fetal calf serum  FcγRs  Fcγ Receptors  FITC  Fluorescein isothiocyanate  GFP  Green fluorescent protein  GST  Glutathione s-transferase  HBSS  Hank's buffered salt solution  HEPES  4-(2-hydroxyethyl)-1-piperazineethanesulphonic acid  His  Histidine  HIV  Human immunodeficiency virus  HOPS  Homotypic vacuole fusion and protein sorting  IPTG  Isopropyl-β-D-thio-galactoside  ITAM  Immunoreceptor tyrosine-based activation motifs  LAMP  Lysosome-associated membrane proteins  LPS  Lipopolysaccharide  MDR  Multi drug resistant  mAGP  Mycolyl-arabinogalactan-peptidoglycan  MES  2-(N-morpholino)ethanesulfonic acid  	
    xv  MOI  Multiplicity of infection  Mtb  Mycobacterium tuberculosis  NOX2  NADPH oxidase  NK  Natural killer  OADC  Oleic acid albumin dextrose complex  PBS  Phosphate buffered saline  Phox  Phagocyte oxidase  PFA  Paraformaldehyde  PI3P  Phosphatidylinositol 3-phosphate  PtkA  Protein-tyrosine kinase A  PMA  Phorbol-12-myristate-13-acetate  PMSF  Phenylmethanesulfonyl fluoride  PtpA  Protein-tyrosine phosphatase A  RILP  Rab7 interacting lysosomal protein  ROS  Reactive oxygen species  RNS  Reactive nitrogen species  SE  Succinimidyl ester  	
    xvi  Ser  Serine  SCID  Severe combined immunodeficient  SDM  Site-directed mutagenesis  SNARE  Soluble N-ethylmaleimide-sensitive factor (NSF) attachment protein receptor  TB  Tuberculosis  Thr  Threonine  TNF  Tumor necrosis factor  Trp  Tryptophan  Tyr  Tyrosine  VPS  Vacuolar protein sorting  WT  Wild-type	
  	
    XDR  Extreme drug resistant  	
    xvii  ACKNOWLEDGEMENTS The work of this thesis would not be possible without strong support from my supervisor Dr. Yossef Av-Gay. I started in Dr. Av-Gay’s lab as an undergraduate science co-op student. It was during this time when I was inspired by Dr. Av-Gay to undertake the challenge of graduate studies and Tuberculosis research. Throughout my graduate program, Dr. Av-Gay gave me numerous opportunities to experience and learn the various aspects of scientific research, preparing me to be an independent researcher in the future. I especially want to thank Dr. AvGay for his selfless guidance and patience during my first year as a PhD student, helping me to persevere through the hardships I encountered while I was adapting to my new role in the research laboratory. My committee members, Dr. Lisa Craig, Dr. Zakaria Hmama and Dr. Steven Pelech have provided me with constant support over the years. Thank you for all your helpful comments and discussions that greatly improved the quality of my work. I would also like to thank all the past and present members of the Av-Gay lab with special thanks to Mary Ko for all the technical assistance. I owe gratitude to Dr. Horacio Bach for mentoring me, sharing his knowledge in research and teaching me the basic research techniques when I first started in Dr. Av-Gay’s lab. I have to thank Joseph Chao, Jim Sun, and Xingji Zheng for their friendships over the years and for creating a lively and joyful environment in the lab. I would like to acknowledge financial support from Canadian Institute of Health Research (CIHR), Michael Smith Foundation For Health Research, and the University of British Columbia. I must thank the support of my family, especially my parents, for their continued support and sacrifice to give me the opportunity to pursue my dream. Finally, I would like to thank Eileen Zhou for her unconditional care, support and advice that help me become what I am today.  	
    xviii  I am also thankful to all those who helped me during this entire work.  	
    xix  CHAPTER 1: INTRODUCTION 1.1 Tuberculosis 1.1.1 Disease epidemiology Tuberculosis (TB) remains the leading cause of morbidity and mortality due to a single infectious agent (Figure 1). Despite intensive efforts to reduce the global spread of TB disease, one-third of the world’s population is believed to be exposed to Mtb, which causes nearly two million deaths annually.  WHO  estimated that one billion new infections would occur by 2020 if no urgent and efficient measures to control TB disease are taken [1]. Indeed, the emergence of multi-drug resistant (MDR) and extensively-drug resistant (XDR) TB are impacting upon the efficiency of the current treatment. MDR-TB is defined as a form of TB that is resistant to at least isoniazid and rifampin, which are two of the four front line antibiotics. On the other hand, XDR-TB is defined as resistance to second-line drugs, including fluoroquinolone, and at least one of the three injectable TB drugs, i.e. amikacin, kanamycin, and capreomycin [2]. While treatment for MDR-TB requires up to two years of multi-drug regimen, XDR-TB is virtually untreatable. In fact, in a recent outbreak of XDR-TB in Kwazulu-Natal in South Africa, 52 of the 53 patients died within an average of 25 days after being diagnosed [3]. Furthermore, the association with the Human Immunodeficiency Virus (HIV) has led to the explosive spread of TB in developing countries [4]. Patients who are co-infected with both HIV and TB have an up to 800 times greater risk of developing active TB disease symptoms and becoming infectious compared to people who are not infected with HIV [4]. To complicate this  	
    1  problem, despite promising trials for several novel anti-TB therapeutics, no new TB drug was introduced into the clinic in the last 40 years. Therefore, there is an urgent need to understand the physiology and the mechanism of TB pathogenesis in order to identify new targets for the development of novel therapeutics.  World Health Organization. Global Tuberculosis Control 2011. Figure 1. Estimated TB incidence rates (2010). This illustration shows the worldwide distribution of TB with the highest incidence rate in sub-Saharan Africa at 350 cases per 100,000 population [5]. Reprinted with permission from World Health Organization.  1.1.2 Origin of human Tuberculosis TB has plagued humankind throughout its recorded history. The etiological agent of TB, Mycobacterium tuberculosis (Mtb), was discovered by Robert Koch in  	
    2  1882. Recent genetic studies showed that Mtb might have evolved from an early progenitor, which might have infected early hominids in East Africa around 3 million years ago [6]. Comparison of the genome sequences of various species of the Mtb complex indicates that all members, including Mtb, Mycobacterium africanum, Mycobacterium canettii and Mycobacterium bovis, had a common African ancestor approximately 35,000 - 15,000 years ago [7, 8]. However, despite popular belief, Mtb did not arise from a bovine TB strain as evident from comparative genomics analysis and molecular phylogeny (Figure 2). In fact, M. bovis has undergone numerous deletions in its genome relative to Mtb, resulting in a smaller chromosome (~66 kbp truncation) as compared to Mtb [9]. As it is unlikely that the deletions could be repaired by recombination in clonal organisms, this observation rules out the possibility that Mtb might have evolved from M. bovis. Furthermore, the most recent common ancestor of both Mtb and M. bovis was more similar to Mtb than M. bovis in chromosomal content [10], and this common ancestor is believed to have already been a human pathogen. 1.1.3 Mycobacterium tuberculosis Mtb is a rod-shaped, slow growing, high G+C content (65.6%), aerobic facultative intracellular pathogen. It belongs to the order Actinomycetales and suborder Corynebacterineae together with other related microogranisms such as nocardiae, rhodococci and corynebacteria. Mtb is specially classified as an acido-alcohol resistant bacteria in the division of Actinobacteria due its cell envelope having a very different chemical nature, as compared to Gram-positive and Gram-negative bacteria. The cell envelope of Mtb is composed mainly of a 	
    3  mycolyl-arabinogalactan-peptidoglycan (mAGP) complex [11-15], which forms the inner part of the cell wall. Other additional lipids such as phthiocerol dimycocerosates,  glycopeptidolipids,  menaquinones,  and  glycosylated  phenolpthiocerols intercalate in the mycolic acid layer and form the outer region of the cell wall [16-18]. It is this thick lipid-rich coat and capsule that result in the distinctive property of acid fastness and make the Mtb cell wall 10 – 100 times more impermeable than that of the notably impermeable bacillus Pseudomonas aeruginosa [19]. The low permeability of the cell wall confers upon Mtb an intrinsic resistance to host antimicrobial mechanisms and antibiotics.  Smith et al. Nature Reviews Microbiology. 2009. 7: 537-544 Figure 2. Phylogeny of selected mycobacterial species. This illustration shows the evolutionary relationship between Mtb complex species. The deletion of chromosomal region RD9 differentiates Mtb from other the animal specific 	
    4  strains. Reprinted with permission from Macmillan Publishers Ltd, Nature Reviews Microbiology. 	
   1.1.4 Infection cycle Infection by Mtb occurs by aerosol route, and the infectious dose is estimated to be 5 to 10 bacteria. Once inhaled, Mtb is phagocytosed by the alveolar macrophages, inducing a localized inflammatory response. The infected macrophages  release  Tumour  Necrosis  Factor  (TNF)-alpha  and  other  inflammatory chemokines that drive the recruitment and activation of neutrophils, natural killer (NK) T cells, CD4+ T cells and CD8+ T cells [20, 21]. The recruited immune cells, in turn, initiate the inflammatory cascade of cytokines, intensifying cellular recruitment and remodeling of the infection site. It is this host response and recruitment of immune cells as an attempt to contain the infection that leads to the formation of the granuloma, a pathologic feature characteristic of TB (summarized in Figure 3) [22] . The granuloma, with extensive vascularization, consists of the core of infected alveolar macrophage, surrounded by other macrophages, monocytes, neutrophils, giant cells, foamy macrophages, and epithelioid macrophages [23, 24]. The lymphocytes, which are recruited to mediate the adaptive immune response, localize to the periphery of the granuloma associated with a fibrous capsule of collagen and other extracellular matrix components [23]. At this stage, generally 2-3 weeks post infection, the infected human host does not show apparent signs of disease or transmit the infection to others, and the mycobacterial infection is considered to be contained [23]. However, the granuloma can reactivate when the host becomes immune	
    5  compromised, which is usually the consequence of aging, malnutrition or coinfection with HIV. The disease progression in the granuloma is marked by diminished vascular structure and a more prominent fibrous sheath, creating a hypoxic and necrotic center [24]. Ultimately, the granuloma caseates and ruptures into a structureless mass of cell debris, releasing thousands of infectious bacilli into the lung airways [24]. This contributes to the development of a persistent cough, which further aid the aerosol spreading of TB. Figure 3 (next page). Mtb infection cycle. The illustration describes the life cycle of Mtb. Upon inhalation, Mtb is phagocytosed by alveolar macrophages. Host immune responses lead to granuloma formation as an attempt to control infection. However, Mtb reactivates when host immunity is compromised, and the granuloma caseates and ruptures, releasing infectious bacilli that contribute to a productive cough and spread of TB. Reprinted with permission from Nature Publishing Group; Nature Reviews Immunology.  	
    6  Russell et al. Nat Immunol. 2009. 10: 943-948  	
    7  1.1.5 Mycobacteria resistance to host killing As an intracellular pathogen, Mtb is capable of adapting to various environmental changes that it encounters within the host macrophage throughout the course of infection. Once Mtb is phagocytosed by the alveolar macrophage, the host cell attacks the invading pathogen with a wide range of antimicrobial mechanisms, including nutrient deprivation, hypoxia, reactive oxygen and nitrogen species (ROS and RNS), acidity and hydrolytic enzymes. A major mycobacterial component that allows Mtb to survive and persist within such a hostile host environment is its thick, waxy, lipid-rich cell wall, which, as mentioned above, is composed of high mycolic acid content. The various complex lipids within the mycobacterial cell envelope not only scavenge harmful reactive radicals from the extracellular environment but also contribute to intrinsic resistance against acidic or alkali conditions and antibiotics [25]. In addition to this protective capsule, successful TB pathogenesis is dependent on several other Mtb intracellular survival strategies. Numerous studies in the past have shown that Mtb is capable of neutralizing ROS and RNS [26], disrupting host signaling pathways [27-29], inhibiting apoptosis to suppress the adaptive immune response [30, 31], and, most importantly, arresting phagosome fusion with lysosomes [32-34]. A summary of some of these mechanisms, with the focus on the arrest of phagosome maturation, will be further discussed in Section.1.3. 1.2 Phagocytosis 1.2.1 Phagosome formation Phagocytosis is a major component of the innate immune defense against 	
    8  infectious agents such as bacteria and viruses. The interaction of the microorganism with the phagocyte can occur through recognition of pathogenassociated molecules, such as surface carbohydrates, peptidoglycans or lipoproteins, by pattern recognition receptors [35, 36]. Alternatively, foreign invading particles can be coated by opsonins such as immunoglobulin G (IgG) and components of the complement cascade that attach to the pathogen surface [37-39]. The opsonins are recognized by the phagocytes’ surface receptors such as Fcγ receptors (FcγRs) and complement receptor 3 (CR3) [37, 39]. Engagement of the host phagocyte receptors induces a complex network of signaling events (summarized in Figure 4). One such event is the phosphorylation of immunoreceptor tyrosine-based activation motifs (ITAMs) on FcγRs by Src-family kinases [40]. ITAM phosphorylation recruits and activates the tyrosine kinase SYK, which in turn phosphorylates various substrates [41]. These signaling pathways trigger cytoskeletal rearrangement, mediated by Rho GTPases Rac1 and Cdc42, and alterations in membrane trafficking, regulated by Rab and ADP Ribosylation Factor (Arf) GTPases, allowing the phagocyte to extend pseudopod around the foreign particle [42, 43]. Phosphoinositides such as  phosphatidylinositol-4,5-bisphosphate  and  phosphatidylinositol-3,4,5-  trisphosphate also play a role in mediating the engulfment of the foreign particle [44]. Accumulation of phosphoinositides at the site of particle engagement initiates actin assembly and drives pseudopod formation, ultimately leading to the encapsulation of the foreign particle into a vacuole termed as phagosome.  	
    9  	
   Underhill et al. Annu. Rev. Immunol. 2002. 20:825-52  Figure 4. Receptor and signaling interactions during phagocytosis of microbes. Phagocytosis is initiated through recognition of the invading foreign particles or microbes by multiple receptors. Receptor engagement elicits a network of signal transduction pathways with the phagocyte, leading to various responses. Reprinted with permission from Annual Reviews: Annual Reviews in Immunology.  1.2.2 Phagosome maturation Phagosome maturation starts immediately after the nascent compartment pinches off from the surface plasma membrane. Through progressive interactions with components in the endocytic pathway, the phagosome remodels its membrane and luminal contents [45]. It is unclear whether these interactions involve complete fusions with the incoming membranes or “kiss and run” 	
    10  transient passage. It is also unclear whether the maturation process occurs in a linear manner where the phagosome interacts with the early endosomes, late endosomes and lysosomes sequentially or a more complex network of membrane transfers occurs simultaneously. However, there is evidence that upon formation of the nascent phagosome, tubular structures from lysosomes located in the perinuclear region extend along the microtubules to fuse with and deliver crucial contents to the phagosome at an early stage [46]. This indicates that phagosome maturation might occur through a more complex process involving multi-directional interactions with various membrane compartments.  Figure 5. Stages of phagosome maturation. Upon phagocytosis, the nascent phagosome interacts sequentially with components of the endocytic pathway to acquire antimicrobial properties, including an acidic pH caused by the proton pumping V-ATPase and hydrolytic enzymes such as Cathepsins. Progressive interaction with early endosome, late endosome and lysosome ultimately leads to the formation of the phagolysosome where the invading microbe is eliminated, and antigens are presented on the surface of the phagocyte to initiate adaptive immune response. 	
    11  Nevertheless, the phagosome maturation process can be characterized by several stages that culminate in the creation of a highly acidic, oxidative and degradative compartment (summarized in Figure 5). Upon detaching from the plasma membrane, the nascent phagosome retains properties similar to the early endosome, having a mildly acidic environment (pH 6.0) with poor hydrolytic activity characterized by the presence of the inactive Procathepsin D protease [47]. The small GTPase Rab5 plays a key role during this early phase of phagosome maturation [48]. Rab5 associates with phagosome immediately after phagocytosis and facilitates the recruitment of Rab5 effector proteins such as VPS34 [49]. VPS34 is a class III phosphatidylinositol-3-kinase that generates phosphatidylinositol-3-phosphate (PI3P) on the early phagosomal membrane. PI3P is responsible for recruiting and anchoring effector proteins, including the early endosome antigen 1 (EEA1), to the cytosolic face of the phagosome through FYVE and PX domains [50, 51]. EEA1, which also interacts directly with Rab5, participates in tethering the phagosome with early endosomes [52]. Furthermore, EEA1 binds to the SNARE (soluble N-ethylmaleimide-sensitive factor (NSF) attachment protein receptor) protein syntaxin13 to drive phagosome fusion with early endosomes [53]. PI3P is also required for the recruitment and assembly of the NADPH oxidase complex (NOX2), which interacts with membrane phosphoinositides via the PX domains of its p40phox and p47phox subunits, on the phagosomal membrane [54]. NOX2 generates superoxide in the phagosomal lumen through a redox chain by transferring electrons from cytosolic NADPH to phagosomal oxygen [54]. The superoxide is in turn converted into  	
    12  several other powerful microbicidal ROS and RNS, such as hydrogen peroxide, hydroxyl radicals and peroxynitrite when combined with nitric oxide radicals [55]. Further enrichment of PI3P and association of Rab5 downstream effector proteins in the phagosomal membrane are followed by the recruitment of the Rab7 GTPase, a characteristic marker of the late phagosome [56, 57]. The exchange of Rab5 for Rab7 is mediated by the Class C VPS HOPS complex, a key regulator of membrane tethering and fusion in the endocytic pathway [58, 59]. The interaction with both Rab5 and Rab7 indicates that the Class C VPS HOPS complex is crucial for the transition from early to late phagosome [59]. The composition and function of the Class C VPS complex will be discussed further in Section 1.2.4. A key downstream effector molecule recruited by Rab7 is the Rabinteracting lysosomal protein (RILP), which drives the centripetal movement of late phagosomes to perinuclear lysosomes by linking the membrane to the dynein–dynactin motor complex [60]. Fusion of the late phagosome with lysosomes is then driven by the Class C VPS HOPS complex, which tethers the apposed membranes and mediates the assembly of SNARE complex, completing the membrane coalescence event [61, 62]. At this stage, the phagosome has an acidic luminal pH (5.5 – 6.0) due to the acquisition of the proton-pumping vacuolar H+-ATPase (V-ATPase), whose structure and function will be further discussed in Section 1.2.3. The late phagosome is also enriched in proteases and lysosomal-associated membrane proteins (LAMPs), which are either imported from the Golgi complex or acquired by fusion with late endosomes [45]. 	
    13  The phagosome maturation process ultimately leads to the formation of the phagolysosome, a terminal microbicidal organelle loaded with an arsenal of antimicrobial effectors that completely degrade the invading microorganisms. Further accumulation of the V-ATPases creates a profoundly acidic lumen (pH 4.5 – 5.0), the hallmark of phagosome maturation. Small antimicrobial peptides such as defensins and cathelicidins [63, 64], which directly disrupt membrane integrity through the formation of multimeric ion channels on bacterial pathogens, are delivered to the phagolysosome. The phagolysosome also has an elevated content  of  acid-activated  carboxypeptidase,  hydrolytic  phospholipase  A2,  enzymes,  including  α-hexosaminidase,  lysosomal  β-glucuronidase,  lysozyme and cathepsins, that target protein, lipid and carbohydrate components of ingested particle [65, 66]. These acid-activated hydrolases mediate the complete destruction of the invading foreign particles, whose constituents are then presented as peptide antigens by MHC molecules on the phagocytes’ surface to activate adaptive immunity. 1.2.2.1 Vacuolar H+-ATPase The ubiquitous eukaryotic V-ATPase is composed of a membrane bound sector (V0) and a cytosolic stalk and catalytic head (V1) [67, 68]. It is structurally and mechanistically related to the ATP synthase (F-ATPase), which is also a multisubunit protein complex that operates through a rotary mechanism to synthesize ATP [69]. However, unlike the ATP synthase, the V-ATPase hydrolyzes ATP to drive active transport of protons across membranes. The integral V0 domain, which has a molecular mass of 260 kDa, consists of five different subunits (a, d, 	
    14  c, c’ and c’’) that are responsible for proton translocation while the cytosolic domain contains eight different subunits (A, B, C, D, E, F, G, H) that are mainly involved in harnessing the energy from the hydrolysis of ATP (Figure 6). The complex is present on the membranous organelles such as lysosomes, endoplasmic reticulum and the Golgi apparatus. ATP hydrolysis is catalyzed by subunit A and B, which form the heterohexameric structure of the catalytic head [70, 71]. The energy from ATP hydrolysis drives the rotation of the entire rotary assembly consisting of the cytoplasmic central stalk (subunit D and F), subunit d and the integral membrane proteolipid ring (c, c’, c’’) [72]. Proton transport is dependent on a crucial glutamate residue in each of the proteolipid subunits that undergoes reversible protonation during the rotation [73]. The glutamate residue is protonated as it engages the cytoplasmic hemi-channel in subunit a. Rotation of the proteolipid ring then exposes the protonated residue to the lumenal hemichannel, where an arginine residue forces the deprotonation of the glutamate residue leading to proton translocation [74]. Subunit E, C, G and H form the peripheral stator that maintains the orientation of the V-ATPase subunits during the rotational catalysis [75]. Interestingly, subunit H is the only subunit of VATPase of which an analogous subunit is not found in F-ATPase, distinguishing the V-ATPase complex from the F-ATPase complex. A deletion mutant of the subunit H gene in yeast showed that it is essential for the activity but not the assembly of the enzyme complex, suggesting that subunit H is a regulator of the V-ATPase [76]. This finding was further supported by the observation that when V1 and V0 domains of the V-ATPase complex dissociate, subunit H remains  	
    15  bound to cytosolic V1 domain and inhibits the unproductive ATPase activity of the dissembled V1 domain [77]. In addition to the elucidation of the proton transport mechanism, recent studies showed that the V-ATPase complex is capable of interacting with other proteins to regulate their activities [78-80]. In particular, the small GTPases ADP-ribosylation factor nucleotide site opener (ARNO) and Arf6, which are endocytosis regulators involved in carrier-coat vesicle formation, have been found to be recruited directly by V-ATPase to endosomes, in a pH dependent manner, to initiate vesicle formation. These observations reveal that V-ATPase is a multi-functional enzyme complex that modulates a network of cellular processes. Furthermore, studies of Drosophila neurons and mammalian renal medulla cells have shown interaction between subunits of V-ATPase and SNARE proteins, reinforcing the notion that V-ATPase might act as a scaffolding structure to modulate membrane trafficking and fusion [81, 82]. Accumulation of the V-ATPase complex and acidification of the phagosome lumen is therefore not only crucial for the inhibition of bacterial survival and activation of acid-activated hydrolytic enzymes, but also proper trafficking of the phagosome within the endocytic pathway.  	
    16  Forgac et al. Nat. Rev. Mol. Cell. Biol. 2007. 8:917-929  Figure 6. Vacuolar H+-ATPase. This illustration shows a model of the overall structure of the S. cerevisiae V-ATPase complex, comprising of the V1 cytosolic domain and V0 membrane integral domain. The Arg-735 residue in subunit a is the key residue involved in proton translocation. Reprinted with permission from Nature Publishing Group: Nature Reviews Molecular Cell Biology.  1.2.2.2 Class C VPS The Class C Vacuolar Protein Sorting (VPS) complex comprises four subunits, VPS11, 16, 18 and 33, that were first identified in classical genetics screens in Saccharomyces cerevisiae [83, 84]. Mutations in each of the four genes resulted in a failure of vacuolar lysosome formation in mutant yeast. This observation, together with biochemical studies, led to the conclusion that the Class C VPS complexes play a key role in the endocytic trafficking pathway, especially at the endosome-to-lysosome stage of transport where it functions together with Rab  	
    17  GTPases to tether membrane compartments and to assemble trans-SNARE complexes for membrane fusion [62, 85, 86]. In particular, Class C VPS complex was found to bind to the vacuolar SNARE syntaxin7 to regulate its activity [87]. More recent studies in S. cerevisiae further indicate that the Class C VPS complex acts as a core protein assembly that can interact with other VPS accessory subunits to participate at different stages of membrane transport. Specifically, the Class C VPS complex can interact with VPS8 and VPS3 to form the CORVET (class C core/vacuole endosome tethering) complex on Rab5positive early endosomes to mediate fusion with late endosomes and with VPS41 and VPS39 to form the HOPS (homotypic fusion and protein sorting) complex that, together with Rab7, regulates lysosomal transport [88, 89] (Figure 7). Although the CORVET complex has not yet been identified in mammalian cells, it likely exists and has the same functions given the high similarity in the transport machinery between yeast and mammals. Peplowska et al. [88] showed that the HOPS and CORVET complexes could reversibly interconvert and also identified two intermediates depending on which subunits had been exchanged (Figure 7). Two models for the conversion have been postulated [88]. The first model assumes that the intermediates can assemble de novo and bind to different Rab GTPases on endocytic membranes. The second model links the observed order of intermediates to the Rab GTPase cycle, displacing and recruiting Rab GEFs (guanine exchange factors) and Rab effectors as necessary. In fact, VPS39, the GEF that interacts with and activates Rab7, can also bind to Rab5, indicating that the conversion of CORVET to HOPS is dependent on and coincides with the  	
    18  exchange of Rab5 for Rab7 [59, 88]. The phagosome maturation process is therefore largely dependent on the dynamics of the interaction between Rab5 and Rab7 GTPases and the Class C VPS complex, and the recruitment of their respective downstream effector proteins to direct the membrane trafficking and fusion along the endocytic pathway.  Nickerson et al. Curr Opin Cell Biol. 2009. 21: 543-551  Figure 7. Working model for Class C VPS function in yeast. Class C VPS complex serves as the core for the CORVET and HOPS complexes through reversible association with CORVET-specific subunits (VPS3 and VPS8) and HOPS-specific subunits (VPS39 and VPS41). CORVET and HOPS regulate membrane fusion events at late endosomes and lysosomes, respectively, through interaction with Rab GTPases. Hybrid intermediate complexes containing both HOPS and CORVET subunits are known to exist. Reprinted with permission from Elsevier Inc.: Current Opinion in Cell Biology. 1.3 Mtb arrest of phagosome maturation 1.3.1 Overview The ability of Mtb to persist and replicate within the host macrophage after phagocytosis is central to TB pathogenesis [90]. As mentioned above, intracellular survival of Mtb results from a combination of factors including a unique cell wall structure, which physically shields the bacterium from  	
    19  bactericidal and hydrolytic enzymes [91], and secretion of enzymes to combat host ROS and RNS [92, 93]. Although these factors contribute to Mtb persistence within the macrophage, one recurring and highly important feature of this pathogen is inhibition of the normal phagosome maturation process, thereby abrogating physical fusion of phagosome with lysosomes and ultimately protecting the bacterium from a bactericidal environment (summarized in Figure 8). Mtb arrests phagosome maturation at an early stage, resulting in a mycobacterial phagosome that possesses similar properties of the early endosome, characterized by membrane acquisition of the early endosome marker Rab5 [94]. However, the key Rab5 effectors, EEA1 and VPS34, are excluded from the mycobacterial phagosome [27]. This phenotype is due to Mtb’s capability of interfering with the macrophage Ca2+ fluxes during phagosome maturation [95]. The block in Ca2+ fluxes, mediated by Mtb cell wall glycolipid ManLAM (mannose-capped lipoarabinomannan), results in the inhibition of the Ca2+/Calmodulin protein kinase CaMKII signaling cascade, which is required for the recruitment of VPS34 to the phagosomal membrane [28]. The absence of VPS34 impairs the generation of PI3P in the phagosomal membrane, resulting in the failure to recruit EEA1 and the inhibition of the phagosome maturation process. Another major characteristic of the mycobacterial phagosome is the lack of acidification, restricting the compartment to a pH of 6.4 where Mtb could survive and persist. It is a common theme among intracellular pathogens that 	
    20  enter the host cells through the phagocytic or endocytic pathway to possess mechanisms to counter the acidic environment within the phagosome. For instance, Salmonella enterica maintains the pH homeostasis by upregulation of its acid tolerance response genes [96]. Yersinia pseudotuberculosis blocks phagosome acidification by direct inhibition of V-ATPase activity [97]. Legionella pneumophila secretes SidK into the host phagocyte to inhibit vacuole acidification through interaction with V-ATPase subunit A [98]. In the case of Mtb, it is long accepted that the lack of phagosome acidification is mainly due to exclusion of the V-ATPase from the phagosomal membrane [99]. Recent studies have also demonstrated that Mtb might be capable of disrupting the Rab7 GTPase mediated membrane trafficking pathway. Sun et al. [100, 101] showed that nucleoside diphosphate kinase (Ndk) is secreted by M. bovis BCG into the cytosol of infected cells to act as a GTPase-activating protein (GAP) that catalyzes the conversion of the active Rab7-GTP to the inactive Rab7-GDP form. The lack of active Rab7 on the phagosomal membrane results in the failure to recruit RILP, thus halting phagosome fusion with lysosomes.  	
    21  Philips. Cell Microbiol. 2008. 10:2408-15  Figure 8. Model of Mtb inhibition of mycobacterial phagosome maturation. Mtb proteins or lipids are colored red, and host proteins or lipids are illustrated in blue. SapM, a PI3P phosphatase, and the cell wall glycolipid ManLAM together interfere with PI3P production on the phagosomal membrane, thereby leading to the exclusion of EEA1 from the mycobacterial phagosome. PtpA shuts down the membrane fusion machinery through dephosphorylation of VPS33B in the Class C VPS complex. A Rab7 GTPase-activating protein (GAP) also disrupts Rab7 activation, impairing phagosome trafficking to the lysosome. Reprinted with permission from Blackwell Publishing Ltd: Cellular Microbiology.  1.3.2 Mtb phosphatase interference with host signaling pathways In addition to the aforementioned mycobacterial proteins and lipids, Mtb utilizes a wide repertoire of signal transduction systems to sense and respond to the hostile host environment and modulate the host signaling pathways. These  	
    22  systems include eleven “two-component” systems, eleven eukaryotic-like proteinserine/threonine  kinases  (STPKs)  (PknA-PknL),  two  protein-tyrosine  phosphatases (PTPs) (PtpA and PtpB), and the newly identified protein-tyrosine kinase (PTK) (PtkA) [32, 102-105]. The role of mycobacterial kinases in Mtb pathophysiology, particularly the regulation of cell wall biogenesis and dormancy, has been widely studied, spurring research efforts in the identification of compounds targeting these signal transduction components. In recent years, as a better understanding of the cross-talk between host and mycobacterial signaling pathways is gained, there was an immense interest in targeting this host-pathogen interface for novel antituberculosis drugs. Among the proteins secreted by Mtb into the host cytosol, three phosphatases, PtpA, PtpB and SapM, are essential for Mtb survival in human macrophages and in animal models of infection [32, 34, 106, 107]. While SapM was found to be a lipid phosphatase that inhibits the proper generation of phosphatidylinositol 3-phosphate (PI3P) that is essential for phagosome recruitment of EEA1 [34], PtpA was shown by us to target the human VPS33B in order to inhibit phagosome-lysosome fusion [32]. Although the host cognate substrate of PtpB remains to be elusive, this phosphatase was shown to contribute to the establishment of latent infections. Thus,  identifying  potential  inhibitors  that  specifically  target  these  Mtb  phosphatases is the main focus of many laboratories with the ultimate goal to generate new TB drugs [106, 108-113].  	
    23  1.3.2.1 PtpB PtpB, along with PtpA, was first identified from the genome sequence of Mtb H37Rv through its homology to known eukaryotic PTPs. Specifically, PtpB possesses the typical C(X)5R(S/T) signature motif in the active site phosphate binding loop (P-loop) of eukaryotic PTPs. Koul et al. [104] showed that Mtb PtpB is secreted into culture media and its phosphatase activity is specific for phosphotyrosine substrates. Additionally, crystallographic analyses revealed that PtpB possesses the distinct features of dual phosphotyrosine binding sites and a two-helix lid structure that covers and protects the active site of the enzyme in an oxidative environment [114]. From these observations, it has been postulated that Mtb secretes PtpB into either the phagosome or host cytosol during infection to interact with host proteins and disrupt macrophage signaling pathways. However, a recent study, which re-examined the sequence homology and enzymatic activity of PtpB, indicates that PtpB, within its C(X)5R(S/T) signature motif, contains the conserved AGK and DRT motifs (CFAGKDRT) found in phosphatase  and  tensin  homolog  (PTEN)  and  myotubularin  (MTM)  phosphoinositide lipid phosphatase, respectively [115]. With an active site region that also resembles the eukaryotic dual-specificity phosphatase that can dephosphorylate both phosphotyrosine and phosphoserine/threonine, PtpB was found to possess the unique property of triple-specificity for phosphoinositides, phosphotyrosine and phosphoserine/phosphothreonine [115]. Based upon these results, it is reasonable to suggest that PtpB might also disrupt host phosphoinositide metabolism and its associated signaling pathways, which are  	
    24  known to have a key role in phagosome maturation. However, thus far, the cognate substrates of PtpB remain elusive. Despite the lack of information on PtpB targets within the host macrophage, there is compelling evidence that PtpB is essential for TB pathogenesis [106, 107].  Indeed, ptpB deletion in Mtb H37Rv strain led to  decreased bacterial survival in IFN-γ activated murine J774A.1 macrophage-like cells and a significant reduction of Mtb persistence in the organs of infected guinea pigs 6 weeks post-infection as compared to infection with the parental strain [107]. This late replication defect of the ΔptpB mutant coincides with the development of efficient T lymphocyte-mediated responses, which supports a role for PtpB in Mtb’s evasion of adaptive immune response. A more recent study examining the effect of PtpB on IFN-γ-mediated signaling pathways showed that expression of PtpB in activated RAW264.7 murine macrophage-like cells resulted in reduced phosphorylation of the ERK1/2 and p38 mitogen-activated protein kinases (MAPK) [116]. Suppression of ERK1/2 and p38 activities led to decreased production of IL-6 [116], which has been implicated in the induction of innate and adaptive immune response during Mtb infection. Furthermore, PtpB expression resulted in the inhibition of macrophage apoptosis in response to IFNγ activation as evident from increased Akt phosphorylation and decreased caspase-3 activation [116]. In fact, it has been previously observed that mycobacterial infection can induce the activation of ERK1/2 and p38 MAPK during entry into macrophages; however, this activation of MAPK signaling cascades is only observed with attenuated or non-pathogenic strains whereas 	
    25  pathogenic species quickly diminish the host kinase activities upon infection [117]. Although there is no evidence suggesting that ERK1/2 and p38 are direct substrates of PtpB, these observations, taken together, indicate that PtpB participates in disrupting host signal transductions to subvert host immune response to Mtb infection and to promote Mtb intracellular survival. It is interesting to note that Mtb antigens such as ESAT-6, Heparin binding hemagglutinin (HBHA) and various cell wall glycolipids and glycoproteins are known to elicit the MAPK and PI3K/Akt signaling pathways [118-121]. It is therefore likely that PtpB plays a key role as a damper balancing host immune detection with evasion during Mtb infection. The relevance of PtpB to Mtb intracellular survival was further confirmed by a recent study showing that specific inhibition of PtpB impairs intracellular survival of Mtb within murine macrophages [106]. Multiple studies have already identified compounds that specifically target and inhibit PtpB; however, PtpB’s exact mechanism of action within the host macrophage remains to be elucidated. 1.3.2.2 SapM SapM (secreted acid phosphatase of Mtb) was first identified from fractionated culture filtrate of the Mtb H37Rv strain, which was known to contain phosphatase activity, as a non-specific acid phosphatase [122]. Interestingly though, SapM does not exhibit significant homology with any prokaryotic acid phosphatase. In addition, SapM lacks any conserved sequence motifs of protein-serine/threonine phosphatases, protein-tyrosine phosphatases, metallophosphoesterases, and histidine phosphatases. The absence of any identifiable motifs posed questions 	
    26  on SapM function. However, SapM does possess two histidine residues that are highly conserved in fungal acid phosphatases. In fact, three homologous sequences were identified in SapM, belonging to known fungal acid phosphatases from Penicillium chrysogenum, Aspergillus nigerficuum and Kluyveromyces lactis, could be found [122-125]. Furthermore, the presence of a typical N-terminal signal peptide indicates that SapM is secreted by Mtb through the general secretion system (Sec) pathway [122]. The secretory nature of SapM, therefore, indicates that it might function to interact with host molecules during infection. When SapM was tested on various phospho-substrates, it was initially found that it does not have activity on phospholipids such as phosphatidylcholine, phosphatidylethanolamine, and phosphatidic acid [122]. Rather, it exhibits a notable activity on phosphoenolpyruvate, glycerophosphate, GTP, NADPH, phosphotyrosine, and trehalose-6-phosphate [122]. Subsequent studies by Vergne et al. [34] showed that PI3P, a membrane trafficking regulatory lipid essential for phagosomal acquisition of lysosomal constituents, is retained on phagosomes harboring dead mycobacteria but is continuously eliminated from phagosomes with live bacilli. While host macrophage PI3P phosphatases such as MTMs were not responsible for the absence of PI3P from the mycobacterial phagosome, SapM was found to possess PI3P phosphatase activity and to be responsible for the removal of PI3P [34]. This indicated that SapM, together with the cell wall glycolipid ManLAM, mediates the arrest of phagosome maturation during Mtb infection through disrupting the recruitment of PI3P effector proteins 	
    27  such as EEA1. Indeed, addition of SapM to an in vitro assay inhibited phagosome fusion with late endosomes [34]. A more recent study of SapM in M. bovis BCG demonstrated that deletion of sapM results in a better vaccine strain that improved survival of mice challenged with Mtb [126]. The ΔsapM BCG vaccine was more effective than the parental vaccine in inducing recruitment and activation of CD11c+ MHC-IIint CD40int dendritic cells (DCs) to the draining lymph nodes. Reduced activation of DCs in animal vaccinated with wild type BCG could be the result of SapM inhibition of phagosome maturation, thereby preventing the subsequent antigen presentation and the initiation of adaptive immunity. 1.3.2.3 PtpA As mentioned before, PtpA was identified from the genome sequence of Mtb H37Rv through its homology to known eukaryotic PTPs. The gene encoding PtpA is located within an apparent operon together with ptkA and Rv2235 upstream and downstream, respectively. PtpA is classified as a low molecular weight PTP (LMW-PTP) based upon its sequence and structural similarity with the family of 18-kDa enzymes involved in cell growth regulation in eukaryotes [127]. It contains the conserved C(X)5R(S/T) signature motif typical of functional PTPs, where the Cys-11 residue serves as the nucleophile, attacking the phosphoryl group on the Tyr residue of the protein substrate. The active site Arg17 residue forms bidentate hydrogen bonds with the phosphoryl group in the substrate through its guanidinium group, stabilizing the transition state during catalysis. The invariant Asp-126 residue is critical for release of substrate and regeneration of active enzyme (Figure 9). Despite overall topology similarity with 	
    28  the eukaroytic LMW-PTPs, significant deviations regarding the amino acids involved can be found in the crevice leading to the active site region. Different physiological function may have arisen from the different substrate specificities [127]. Interestingly, the presence of an additional cysteine (Cys-16) residue in the active site may form an intramolecular disulfide bridge with Cys-11. The formation of the disulfide bond was proposed to protect the nucleophilic Cys-11 from excessive oxidation, conferring rapid sensing and responding to the redox status of the environment [128]. On the other hand, a more recent study showed that PtpA, when treated with reactive nitrogen species (RNS) in vitro, is nitrosylated on the solvent-exposed Cys-53 residue, leading to a 50% reduction in phosphatase activity. On the other hand, both Cys-11 and Cys-16, which have lower solvent-accessibility, are unaffected by the treatment [129]. While the existence of the active site disulfide bridge in PtpA has not been observed directly, Cys-53 nitrosylation indicates that PtpA could be susceptible to the oxidative environment in the host macrophage. This would then raise questions on how PtpA could maintain its activity in the macrophage to disrupt host signaling pathways. It is tempting to speculate that another reduction system to protect PtpA from oxidative or nitrosative stress might exist. The exact mechanism of redox regulation of PtpA and its role in Mtb pathogenesis remains to be elucidated.  	
    29  Bach et al. Cell Host Microbe. 2008. 3: 316-22  Figure 9. Scheme of PtpA catalytic activity. PtpA cysteine at position 11 attacks nucleophilically the phosphorylated tyrosine substrate. PtpA aspartate at position 126 protonates the phenolic oxygen and a cysteinyl-phosphate intermediate is formed facilitating the cleavage of the P-O bond. The transient intermediate is hydrolyzed by an activated water molecule. Reprinted with permission from Elsevier Inc.: Cell Host & Microbe. The fact that PtpA is secreted from Mtb and the absence of a gene encoding a typical protein tyrosine kinase (PTK) in the Mtb genome sequence indicates that PtpA is involved in dephosphorylation of host tyrosine phosphorylated protein and thus may be involved in the modulation of host signaling pathways during infection. Indeed, initial studies in our laboratory demonstrated that PtpA is released by Mtb H37Rv into culture media and that PtpA expression is up regulated upon Mtb entry into host macrophages [104, 105].  Furthermore, PtpA export into the host macrophage cytosol can be  detected directly with immuno-electron microscopy and western blot analysis of the cytosolic fractions of infected macrophages. More importantly the phenotype observed after expression of neutralizing single-chain antibodies against PtpA within macrophages clearly demonstrate the presence of PtpA within the cytosol of the infected macrophage [32].  Yet, the exact mechanism by which PtpA  translocates into the host macrophage cytosol remains to be identified. The lack of an amino-terminal signal sequence rules out the SecA1 and twin-arginine 	
    30  translocation (Tat) pathway for the secretion of PtpA. While the SecA2 or ESX/Type VII export systems are possible candidates responsible for PtpA secretion, there is also evidence indicating that bacterial proteins less than 70 kDa in molecular size can cross the phagosomal membrane [130]. This observation might be explained with the recent discovery that the ESX-1 secretion system with its substrate ESAT-6 can perturb host cell membrane to facilitate the translocation of mycobacterial proteins into the macrophage cytosol [131]. Further study is required to investigate whether PtpA is a direct substrate for the ESX/Type VII secretion system. Within Mtb-infected macrophage, PtpA was found to disrupt key components of the endocytic pathway in order to arrest phagosome maturation [32]. Initial studies showed that expression of PtpA in RAW 264.7 murine macrophage-like cells results in a decrease of phagocytic activity and an increase of F-actin nucleation on phagosomes in vivo and in vitro. The authors of this study postulated that F-actin polymerization might physically block direct contact between donor and acceptor membranes, thereby inhibiting phagosome– lysosome fusion. However, in a separate study, PtpA was shown to be dispensable for Mtb growth in a mouse infection model [132]. Nevertheless, using substrate trapping and radioactive γ32P-ATP	
   labeling approaches, the human Vacuolar Protein Sorting 33B (VPS33B), a subunit of the Class C VPS complex, was identified as the cognate substrate of PtpA. As the Class C VPS is a key regulator of membrane trafficking, tethering and the assembly of transSNAREs complexes in the endocytic pathway [133], PtpA dephosphorylation of 	
    31  VPS33B inactivates the membrane fusion mechanism, leading to inhibition of phagosome-lysosome fusion [32]. In support of this, deletion of ptpA in Mtb H37Rv impaired the intracellular survival of the bacteria with host human THP-1 macrophage-like cells, and phagosomes harboring the ΔptpA strain showed increased fusion with the lysosomes and accumulation of lysosomal markers as compared to the parental strain. PtpA is, therefore, essential for the pathogenesis of TB. 1.4 Aims of the study Although the cognate substrate of PtpA within human macrophages has been identified, the complete mechanism of action of PtpA in relation to phagosome maturation arrest has not been characterized. In particular, from the substrate trapping experiment performed to identify VPS33B, we observed that recombinant wild type PtpA protein, which should not have substrate-trapping capability, was able to pull down another human protein with an apparent MW of 50 KDa from THP-1 macrophage-like cell lysate. MALDI-TOF mass spectrometry identified this protein as the subunit H of the V-ATPase complex, indicating that PtpA might be capable of directly disrupting phagosome acidification, an essential arm of the phagosome maturation process. The significance of this discovery stems from the lack of a mechanistic explanation for the long established paradigm of Mtb pathogenesis: the absence of acidification in the mycobacterial phagosome. The overall objective of this thesis is therefore to characterize the complete mechanism by which PtpA contributes to Mtb persistence within the macrophage. The working hypothesis for this project is 	
    32  that Mtb PtpA interferes with multiple host macrophage pathways that are essential for phagosome maturation in order to promote Mtb intracellular survival. To investigate this hypothesis, our initial goal was to investigate whether the subunit H of V-ATPase is a bona fide catalytic or binding substrate of PtpA. When the interaction between PtpA and V-ATPase subunit H was established to be genuine, our aim was to identify the downstream effects of this host-pathogen protein-protein interaction on the various aspects of the phagosome maturation process. In the second part of the project, we performed biochemical studies on PtkA, encoded by an open reading frame (Rv2232) upstream of ptpA within the PtpA operon, and investigated its functional relationship with PtpA. Overall, the study has provided significant insight into Mtb evasion of host macrophage killing. Specifically, we discovered the long-sought mechanism behind the lack of acidification in mycobacterial phagosome and identified a novel pathway required for successful phagosome maturation. The complete elucidation of human signaling pathways disrupted by PtpA will perhaps lead to new therapeutics to combat the global spread of TB.  	
    33  CHAPTER 2: MATERIALS AND METHODS 2.1 Reagents and chemicals 2.1.1 Commercial reagents DMEM, Fetal calf serum (FCS) and HBSS were purchased from Gibco Laboratories (Burlington, Ontario, Canada). Other culture reagents were purchased from Sigma-Aldrich (St. Louis, MO). Protease inhibitor mixture, PMSF, trypsin-EDTA, and glutathione-agarose beads were purchased from SigmaAldrich. Protein A-agarose beads were from Bio-Rad laboratories (Hercules, CA). Nickel-Nitriloacetic acid (Ni-NTA) polyhistidine-Tag purification resin was purchased from Qiagen (Mississauga, ON, Canada). pHrodo Succinimidyl Ester (SE), Alexa-Fluor 350 Carobxylic Acid SE and Alexa-Fluor 488 Carobxylic Acid SE were purchased from Invitrogen (Carlsbad, CA).  Aldehyde/sulfate latex  beads (diameter, 4 µm) were obtained from Interfacial Dynamics (Portland, OR). [γ-32P]-adenosine 5`-triphosphate (ATP) and [γ-32P]-guanosine 5`-triphosphate (GTP) were purchased from Perkin Elmer (Boston, MA). Restriction enzymes, Pfu DNA polymerase, and T4 DNA ligase were purchased from Fermentas (Burlington, ON, Canada) or New England Biolabs (Ipswich, MA). Luria-Bertani (LB) broth and LB agar were from Fisher Scientific (Pittsburgh, PA). Oleic acid, dextrose and catalase complex (OADC) and 7H9 and 7H10 agar culture media were from Difco Laboratories, (Detroit, MI). Kanamycin was purchased from Gibco Laboratories. Hygromycin was purchased from Roche. MinElute PCR purification and plasmid purification Mini and Midi Kits were from Qiagen.  	
    34  2.1.2 Antibodies Rabbit polyclonal anti-VPS33B and rabbit polyclonal anti-subunit E of V-ATPase (ATP6V1E1) IgG were purchased from Proteintech Group (Chicago, IL). Rabbit polyclonal anti-subunit H of V-ATPase (ATP6V1H) IgG and rabbit polyclonal antisubunit B of V-ATPase (ATP6V1B) IgG were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Mouse monoclonal Anti-His-tag antibody and rabbit polyclonal anti-VPS18 IgG were purchased from Applied Biological Materials (Vancouver, British Columbia, Canada). Mouse polyclonal anti-subunit H of V-ATPase IgG was purchased from Sigma-Aldrich, 4G10 Antiphoshotyrosine antibody was purchased from Millipore (Billerica, MA). Rabbit anti-PtpA was described previously [134]. Secondary horseradish peroxidase (HRP) conjugated goat anti-rabbit antibodies was purchased from Sigma-Aldrich. Secondary horseradish peroxidase (HRP) conjugated goat anti-mouse antibodies was purchased from Cedarlane Labs. Texas Red conjugated goat anti-rabbit IgG, Alexa Fluor 488-conjugated goat anti-rabbit IgG and Alexa Fluor 488-conjugated goat anti-mouse IgG were purchased from Invitrogen. FITC-conjugated antirabbit and anti-mouse IgG were purchased from Sigma Aldrich. Specificity of the commercial antibodies was tested with FACS analysis, Western blot analysis and immunostaining control experiments. 2.2 Bacteria 2.2.1 Strain maintenance and generation Mtb H37Rv and derivative strains were grown in Middlebrook 7H9 broth (BD Diagnostic Systems, Mississauga, ON, Canada) supplemented with 10% (v/v) 	
    35  OADC (oleic acid, albumin and dextrose solution; BD Diagnostic Systems) and 0.05% (v/v) Tween-80 (Sigma-Aldrich) at 37°C standing or rolling in incubator. Red fluorescent Mtb H37Rv was generated by transforming the wild type Mtb H37Rv strain with pSMT3 vector encoding the DsRed protein under the control of the Hsp60 promoter (pSMT3-DsRed). For complementation of ΔptpA, DNA sequences encoding wild type PtpA or mutant PtpA under the control of the native promoter, or the hps60 promoter, were cloned into the pKP201 integrative vector [135]. The ΔptpA strain was co-transformed with the resulting plasmids and pBSint, a non-replicating plasmid that provides integrase in trans but is subsequently lost from the cells, thereby reducing the chances of integrasemediated excision of the complementing DNA [136]. Transformants were selected on 7H11 medium with 50 µg/mL hygromycin. 2.2.2 Bacterial preparation for infection Bacteria in mid-log phase were harvested by centrifugation at 12,000 rpm. In indicated experiments, bacteria were stained by pHrodo SE (Invitrogen), AlexaFluor 488 carboxylic acid SE (Invitrogen) or Alexa-Fluor 350 carboxylic acid SE (Invitrogen) at a final concentration of 10 µg/mL at 37°C for 1 hour. After staining, the bacteria were washed three times with 7H9 with 0.05% Tween-80. Prior to infection of differentiated THP-1 macrophage-like cells, the bacteria are also opsonized with human serum for 30 minutes at 37oC, washed three times with 7H9 with 0.05% Tween-80 and passed through 25 gauge needles several times to prevent bacteria clumping.  	
    36  2.2.3 Mycobacterial lysis Bacterial cell pellets were resuspended in 500 µL of 50 mM Tris pH 7.4, 100 mM NaCl, 5 mM EDTA, 1% Triton X-100 and 1 mM PMSF, then mixed with 80 mg of 0.1 mm silica beads (Biospec) and lysed by bead-beating at 30 seconds intervals for 3 times. Subsequently, soluble lysates were obtained by centrifugation at 12,000 rpm for 15 minutes. 2.2.4 Transformation of mycobacteria Competent mycobacteria were prepared by harvesting mid-log phase bacteria and washing three times with 10% glycerol. 200 µL of competent cells were then mixed with plasmid DNA in a 0.2 cm electroporation cuvette (Bio-Rad, Hercules, CA). Cells were electroporated with a 2.5V pulse, and allowed to recover in 7H9 supplemented with 10% OADC and 0.05% Tween-80 in the absence of antibiotics for 2 hours (M. smegmatis) or overnight (Mtb) at 37oC. Transformants were selected on 7H10-OADC agar supplemented with appropriate antibiotics. 2.3 Cell culture 2.3.1 Maintenance of tissue culture and differentiation The human monocytic leukemia THP-1 cell line (TIB-202, American Type Culture Collection, Manassas, VA) was maintained in 125 cm2 tissue culture flasks (Corning Inc., Corning, NY) in RPMI 1640 medium supplemented with 10% FBS, and 1% L-Glutamine. THP-1 cells were seeded onto 12-well (1.0 x 106 /well) or 24-well (5.0 x 105 /well) tissue culture plates and differentiated into macrophagelike cells with 40 ng/mL 12-phorbol 13-myristate acetate (PMA) in complete RPMI media. 	
    37  2.3.2 Single transfection of differentiated THP-1 THP-1 cells were transfected with the mammalian pEGFP vector harboring the GFP-tagged PtpA constructs (pEGFP-N1-GFP) using the Nucleofector Cell Line Nucleofector Kit V (Lonza) according to manufacturer’s protocol. Transfected macrophages were seeded in 12 well or 96 well plates and differentiated with PMA as described above. Expression of GFP-tagged proteins was measured 12 hours after transfection by spectrofluorometry (Emission 535 nm). 2.3.3 Double transfection of differentiated THP-1 THP-1 cells were seeded onto coverslips in 24 well plates at a density of 1 million cells per well, and differentiated with PMA at 40 ng/mL overnight. Transfection was carried out by magnetofection using PolyMAG reagent (Chemicell, Berlin, Germany). In brief, 1.5 µg of each DNA construct (pEGFP-N1-H and pDsRed2N1-VPS33B) was mixed with 1 µL of PolyMAG reagent in a volume of 200 µL serum free and supplement free RPMI 1640 (Sigma). This mixture was incubated for 15 minutes at room temperature and then added to each well in 300 µL of complete RPMI media. Thereafter, plates were placed on a magnetic field and incubated for 30 minutes at 37oC. The magnet was then removed and cells were incubated overnight for expression of the fluorescent proteins. 2.4 Vectors and DNA manipulation 2.4.1 RNA isolation and cDNA synthesis Total RNA was prepared from lysates of THP-1-derived macrophages (1 x 107 cells) using Qiagen RNeasy Mini kit according to the manufacturer's instructions. Synthesis of cDNA was performed using Revert Aid First strand cDNA synthesis 	
    38  kit (Fermentas). For each cDNA synthesis 5 µg of total RNA and 0.5 µg oligo(dT) primers were used. 2.4.2 Cloning The gene encoding Mtb PtpA was PCR-amplified from Mtb H37Rv genomic DNA using the PtpA-F1 primer with an EcoRI site, and PtpA-R1 primer with an XhoI site. The amplified gene was inserted into pGEX-4T3 vector (GE Healthcare) to generate the pGEX-4T3-PtpA plasmid for GST-tagged PtpA fusion protein expression. For cloning into the pALACE vector to create pALACE-PtpA for Histagged fusion protein expression construct, the gene encoding Mtb PtpA was amplified from Mtb H37Rv genomic DNA using the PtpA-F2 primer, containing an AflII site, and PtpA-R2 primer, containing a ClaI site. For constructing the ΔptpA complementation plasmid, the ptpA gene was amplified from pGEX-4T3-PtpA using PtpA-F3 primer, containing an EcoRI site, and PtpA-R2 primer, containing a ClaI site. The amplicon was inserted downstream of the Mtb hsp60 promoter in the pMV261 mycobacterial vector, generating a pMV261-PtpA plasmid. The entire promoter-gene construct was then excised using XbaI and ClaI restriction enzymes and inserted into the pKP201 mycobacterial complementation vector, generating pKP201-hPtpA plasmid. To construct the PtpA-GFP mammalian transfection construct, ptpA was amplified from pGEX-4T3-PtpA using PtpA-F4 primer, containing an XhoI site, and PtpA-R4 primer, containing an EcoRI site. The amplified gene was inserted into pEGFP-N1 vector (Clontech), generating pEGFP-N1-PtpA plasmid. The gene encoding subunit B (ATP6V1B1) of the human V-ATPase was amplified from THP-1 cDNA using B-F1, containing a XhoI 	
    39  site, and B-R1, containing a HindIII site. The amplicon was inserted into pRSETb his-tagged fusion protein expression vector, generating pRSET-b-ATP6V1B1 plasmid. The genes encoding subunit H (ATP6V1H) of the human V-ATPase was amplified from cDNA prepared from THP-1 using H-F1, containing a XhoI site, and H-R1, containing a HindIII site. The amplicon was inserted into pRSET-b, a his-tagged fusion protein expression vector (Invitrogen), generating pRSET-bATP6V1H plasmid. Human VPS33B his-tagged fusion protein expression plasmid (pBO1-VPS33B) was purchased from Genecopoeia Inc. (Rockville, MD). To construct VPS33B-DsRed2 mammalian transfection construct, VPS33B gene was amplified from pBO1-VPS33B using 33-F1 primer, containing a HindIII site and 33-R1 primer, containing a EcoRI site. The amplicon was inserted into pDsRed2-N1 (Clontech), generating the pDsRed2-N1-VPS33B plasmid. To construct Subunit H-GFP mammalian transfection construct, the gene encoding subunit H (ATP6V1H) was amplified from pRSET-b-ATP6V1H using H-F2 primer, containing a XhoI site, and H-R2 primer, containing an EcoRI site. The amplified gene was inserted into pEGFP-N1 vector, generating pEGFP-N1-H plasmid. Mtb ptkA (Rv2232) was amplified from its genomic DNA using the PtkA-F1 primer, containing an AflII site, and PtkA-R1 primer, containing a ClaI site. The amplified gene was inserted into pET151DTOPO to generate pET151DTOPOPtkA His-tagged PtkA fusion protein expression construct. To clone ptkA into the pALACE vector, the ptkA gene was excised from pET151DTOPO-PtkA using the AflII and ClaI restriction sites, and the insert was ligated with the pALACE vector digested with the same enzymes, generating pALACE-PtkA.  	
    All plasmid  40  constructs were subsequently verified by sequencing at Eurofins MWG Operon (Ebersberg, Germany). The oligonucleotides used for DNA cloning in this study are described in Table 1. Table 1. Oligonucleotides used for DNA cloning in this study Oligonucleotides PtpA-F1 PtpA-R1 PtpA-F2 PtpA-R2 PtpA-F3 PtpA-F4 PtpA-R4 B-F1 B-R1 H-F1 H-R1 33-F1 33-R1 H-F2 H-R2 PtkA-F1 PtkA-R1 a  Sequence (5’ è 3’)a ATATATGAATTCCGTGTCTGATCCGCTG ATATATCTCGAGTCAACTCGGTCCGTTC ATATATCTTAAGGTGTCTGATCCGCTG ATATATATCGATTCAACTCGGTCCGTTC ATATATGAATTCGTGTCTGATCCGCTG ATATCGCTCGAGATGTCTGATCCGCTG ATATATGAATTCGACTCGGTCCGTTCCGCGC ATATATCTCGAGAATGGCCATGGAG ATATATAAGCTTCTAGAGCGCAGTGTC ATATATCTCGAGAATGACCAAAATGG ATATATAAGCTTTTAGCTTCGGGCGGC ATAGTGAAGCTTATGGCTTTTCCCCATC ATATATGAATTCGGGCTTTCACCTCACTC ATATCGCTCGAGATGACCAAAATGGATATC ATATATGAATTCGGCTTCGGGCGGCAGCGGTC CACCCTTAAGGTGTCTTCGCCTCGTGAAC ATATATATCGATTCAGACCACCTAGCGCCT  The recognition sequence for the restriction site is underlined.  2.4.3 Site-directed mutagenesis Site-directed mutagenesis (SDM) was carried out according to the QuikChange II-E site-directed mutagenesis kit (Stratagene, La Jolla, CA) protocol. Pfu DNA polymerase (Fermentas) was used in the SDM reactions. Oligonucleotides were designed with 15 bases flanking both sides of the mutated codons (Table 2). . The products obtained after PCR amplification were digested with DpnI restriction enzymes and chemically transformed into E. coli strain DH5α. The positive clones were grown in LB media containing appropriate antibiotics, and the plasmids were extracted using Qiagen Mini Prep Plasmid Extraction Kit. Point  	
    41  mutations were verified by sequencing at Eurofins MWG Operon (Ebersberg, Germany). Table 2. Oligonucleotides used for SDM in this study Oligonucleotides PtkA-D85A-F PtkA-D85A-R PtkA-K184M-F PtkA-K184M-R PtkA-K217M-F PtkA-K217M-R PtkA-K270M-F PtkA-K270M-R PtkA-Y146A-F PtkA-Y146A-R PtkA-Y150A-F PtkA-Y150A-R PtkA-Y262A-F PtpA-Y262A-R PtpA-Y67A-F PtpA-Y67A-R PtpA-Y128A-F PtpA-Y128A-R PtpA-Y129A-F PtpA-Y129A-R PtpA-Y128-129A-F PtpA-Y128-129A-R PtpA-L146A-F PtpA-L146A-R a  Sequence (5’ è 3’)a CAGCTGGTGATCTTCGCTCTGGACGGCACGCTG CAGCGTGCCGTCCAGAGCGAAGATCACCAGCTG GCCGTCGCCACCTCCATGGCAGAGCCGACCGCA TGCGGTCGGCTCTGCCATGGAGGTGGCGACGGC GGCTCGCGAGGCAGCATGGTCGACGTGCTGGCC GGCCAGCACGTCGACCATGCTGCCTCGCGAGCC GGCATCTTTATCGACATGACCTCCACCACCGTC GACGGTGGTGGAGGTCATGTCGATAAAGATGCC GAGGCGATCGTAGCCGCCCGGGCCGACTACAGC GCTGTAGTCGGCCCGGGCGGCTACGATCGCCTC GCCTACCGGGCCGACGCCAGCGCCCGCGGTTGG CCAACCGCGGGCGCTGGCGTCGGCCCGGTAGGC GTGGTCGGCTGGGGCGCCGGGCGCGCCGACTTT AAAGTCGGCGCGCCCGGCGCCCCAGCCGACCAC TTGCGAGCCCACGGCGCCCCTACCGACCACCGG CCGGTGGTCGGTAGGGGCGCCGTGGGCTCGCAA GATGTCGAGGATCCCGCCTATGGCGATCACTCC GGAGTGATCGCCATAGGCGGGATCCTCGACATC GTCGAGGATCCCTACGCCGGCGATCACTCCGAC GTCGGAGTGATCGCCGGCGATGGGATCCTCGAC GATGTCGAGGATCCCGCCGCCGGCGATCACTCCGAC GTCGGAGTGATCGCCGGCGGCGGGATCCTCGACATC CATCGAATCCGCCGCGCCCGGCCTGCACGAC GTCGTGCAGGCCGGGCGCGGCGGATTCGATG  The substituted nucleotide is highlighted in bold font.  2.5 Protein expression and purification 2.5.1 Expression of recombinant proteins in E. coli GST-tagged PtpA was expressed in E. coli strain BL21 transformed with pGEX4T3-PtpA plasmid. An overnight starter culture of the transformed BL21 was grown and inoculated into 1 L of LB media with 100 µg/mL ampicillin. The culture was then grown to an OD600 of 0.6 at 37°C and expression was induced with 0.4 mM IPTG at 19°C overnight. The bacteria were harvested by centrifugation at  	
    42  5,000 rpm, resuspended in PBS containing 1 mM DTT, 0.1 mM PMSF and lysed by sonication. The soluble fraction of the bacterial lysates was obtained by centrifugation at 13,000 rpm and GST-tagged proteins were purified from the soluble fraction by affinity chromatography on glutathione-agarose resin (SigmaAlrich). To express and purify histidine-tagged recombinant protein, pRSET-bATP6V1H, pBO1-VPS33B, pRSET-b-ATP6V1B1 and pET151D-TOPO-PtkA plasmid were transformed into E. coli strain BL21. The transformed BL-21 was used to grow overnight starter cultures, which were then inoculated into 1 L of LB media supplemented with appropriate antibiotics. The cultures were then grown to an OD600 of 0.6 at 37°C, and expression was induced with 0.4 mM IPTG at 19°C or room temperature overnight. The bacteria were harvested by centrifugation  at  5,000  rpm,  resuspended  in  lysis  buffer  (50  mM  Na2HPO4/NaH2PO4 pH = 7.4, 500 mM NaCl, 10 mM imidazole and 1 mM PMSF) and lysed by sonication. The soluble fraction of the bacterial lysates was obtained by centrifugation at 13,000 rpm and GST-tagged proteins were purified from the soluble fraction by affinity chromatography on Ni-NTA polyhistidine-tag purification resin (Qiagen). 2.5.2 Expression of recombinant proteins in M. smegmatis Competent M. smegmatis cells were transformed with pALACE-PtpA or pALACE-PtkA plasmid by electroporation and plated at 37°C on 7H10 plates containing 1% dextrose and 50 µg/mL hygromycin. Starter cultures of the transformed bacteria were grown over two nights in 7H9 media containing 0.05% 	
    43  Tween-80, 50 µg/mL hygromycin and supplemented with 1% dextrose. The starter cultures were inoculated into 1 L of 7H9 media containing 0.05% Tween80, 50 µg/mL hygromycin and supplemented with 1% dextrose. The cultures were then grown at 37°C to an OD600 of 1.0, and the cells were harvested by centrifugation at 5000 rpm for 30 minutes. The cell pellets were resuspended in 7H9 media supplemented with 0.05% Tween-80, 50 µg/mL hygromycin. Acetamide was added to a final concentration of 0.2% to induce protein expression at room temperature for 24 hours. The bacteria were harvested by centrifugation at 5000 rpm for 30 minutes, resuspended in lysis buffer (50 mM Na2HPO4/NaH2PO4 pH = 7.4, 500 mM NaCl, 10 mM imidazole and 1 mM PMSF) and lysed by sonication. The soluble fraction of the bacterial lysates was obtained by centrifugation at 13,000 rpm and histidine-tagged proteins were purified by affinity chromatography on Ni-NTA polyhistidine-tag purification resin (Qiagen). 2.6 In vitro pull down assay In vitro pull down assays were carried out by incubation of THP-1 lysate with recombinant wild-type his-tagged PtpA proteins at 4oC overnight. The mixture was purified using Ni-NTA resin (Qiagen) and resolved in SDS-PAGE. Captured proteins were identified with MALDI-TOF mass spectrometry and analyzed with western blotting to confirm identity. 2.7 ALPHAScreen protein-protein interaction assay ALPHAScreen assays performed using the Histidine (Nickel Chelate) Detection Kit (Perkin Elmer) according to the manufacturer’s protocol. Specifically, purified 	
    44  GST-tagged recombinant proteins were first biotinylated using the EZ-link Biotinylation Kit (Pierce). The biotinylated recombinant proteins were serial diluted in white opaque 384-well microplates (PerkinElmer), and purified histagged recombinant proteins were added to same wells. Nickel-chelating acceptor beads were added to the proteins and incubated for 30 minutes at 25oC. Streptavidin donor beads were added to the wells and the reactions were incubated for 30 minutes at 25oC. The reaction kinetics were then monitored on the Fusion-α-HT Multimode Microplate Reader (PerkinElmer) for luminescence signal generated from protein-protein interaction (counts per second (cps)). 2.8 Mycobacterial “Split-Trp” protein-protein interaction assay In vivo interaction was detected using a modified “Split-Trp” assay first developed by O’Hare et al. [137, 138]. Interaction between two proteins of interest, which are individually fused to the split protein fragments (Ntrp and Ctrp), would reconstitute the activity of the N-(5’-phosphoribosyl)-anthranilate isomerase enzyme in tryptophan synthesis; thereby, enabling the auxtrophic ΔhisA M. smegmatis to grow on tryptophan deficient media. Briefly, the DNA encoding PtpA and subunit H were cloned into pJC10 and pJC11 vectors, generating translational fusion constructs with the Ntrp and Ctrp fragments of the N-(5’phosphoribosyl)-anthranilate isomerase under the control of the acetamidase promoter respectively [138]. The resulting plasmids were transformed into the ΔhisA M. smegmatis strain. Positive transformants were grown in LB broth supplemented with 0.05% Tween-80, 50 µg/mL hygromycin and 30 µg/mL  	
    45  apramycin to OD600 of 1.0 and spotted onto 7H9 agar medium with 1% glucose, 60 µg/mL histidine and 0.02% acetamide to induce protein expression. 2.9 Infection 2.9.1 Infection of THP-1 Infection of THP-1 macrophage-like cells was performed using human serumopsonized Mtb or E. coli DH5α at multiplicity of infection (MOI) of 10:1 or 1:1. For Mtb intracellular survival studies, infected macrophages were harvested at defined time points, lysed with 0.025% SDS, serially diluted, and plated on 7H10 agar medium supplemented with OADC and appropriate antibiotics. The plates were incubated at 37oC for 2 weeks to allow for growth of visible colonies. 2.9.2 Phagocytosis assay THP-1 macrophage-like cells were transfected and seeded as mentioned above (Section 2.3.2 and 2.3.3). E. coli was labeled with 50 ug/mL Alexa Fluor 350 carboxylic acid SE (Invitrogen) at 37oC for 1 hour. The bacteria were then washed with PBS and opsonized with human serum. The transfected cells were infected with labeled E. coli at a MOI of 10:1. Phagocytosis was synchronized as described previously [139]. Non-internalized bacteria were washed away after incubation at 37oC for 15 minutes, and phagocytosis was measured by spectrofluorometry. 2.10 Immunoprecipitation THP-1 cells were infected and lysed as above (Section 2.9.1). Lysates were centrifuged and soluble fractions were used in co-immunoprecipitation assays.  	
    46  Briefly, rabbit anti-VPS33B or rabbit anti-subunit H IgG were added to 2 mg or 4 mg of soluble lysate (final IgG dilution, 1:100) and the mixture was incubated at room temperature for one hour. Affi-Gel Protein-A Agarose Resin (Bio-Rad) was blocked with 0.5% BSA, washed with PBS, then added to the mixture and incubated on a shaker at room temperature for one hour. The resin was then washed with PBS, and SDS sample buffer was added to release IgG and bound material. The resulting samples were resolved on SDS-PAGE and transferred onto a nitrocellulose membrane for Western blot analysis with the indicated primary antibodies. Protein A-HRP was used as the secondary detection reagent. For Western blot analysis with anti-phosphotyrosine 4G10, recombinant VPS33B protein was phosphorylated in kinase buffer as previously described [32] with 0.5 µM ATP and assayed as a control along with the immunoprecipitated samples. 2.11 Measurement of phagosomal pH 2.11.1 Overview Assessment of the phagosomal pH of E. coli and Mtb phagosomes was performed according to a previously published procedure with modifications [140, 141]. Before infection, bacteria were labeled with the pH-sensitive pHrodo probe (Invitrogen), which emits red fluorescence in acidic environment, and pHinsensitive probes Alexa Fluor 350 carboxylic acid SE (Blue) or Alexa Fluor 488 carboxylic acid SE (Green) (Invitrogen). E. coli Phagosomal pH was measured with spectrofluorometry in the Fusion-Alpha HT microplate reader (Perkin Elmer) and Mtb phagosomal pH was measured with a FACSCalibur flow cytometer (BD Bioscience). The fluorescence ratio values of the pH-sensitive and pH-insensitive 	
    47  probes were used to determine phagosomal pH according to a calibration curve. pH calibration was performed by incubating Mtb- or E. coli-infected THP-1 macrophage-like cells in 10 mM phosphate-citrate buffer of pre-determined pH (5.0 - 8.0) containing 145 mM KCl, 1 mM MgCl2, 0.5 mM CaCl2 and 10 mM glucose, together with 20 µM nigericin and 4 µM monensin, which were added as ionophores in order to equilibrate intracellular and intraphagosomal pH with extracelluar pH. As a positive control, concanamycin (50 nM) was used to inhibit macrophage V-ATPase proton transport. 2.11.2 Phagosomal pH of Mtb phagosomes Mtb strains were first labeled with 20 µM pHrodo SE (Invitrogen) at 37oC for one hour. The bacteria were washed with 7H9 supplemented with 0.05% Tween-80 three times before being labeled with 25 µg/mL Alexa Fluor 488 carboxylic acid SE (Invitrogen). The bacteria were washed and opsonized with human serum then used to infect THP-1-derived macrophages at a MOI of 10:1 for 2 hours at 37oC. Non-internalized bacteria were washed away, and cells were reincubated at 37oC for another 2 hours. Cells were then washed, scraped off the plate and fixed with 2.5% paraformaldehyde. pHrodo is stable after fixation with paraformaldehyde [142]. Fixed cells were then analyzed by flow cytometry, and the data was processed with the FlowJo 8.7 software. Color compensation was performed to prevent signal overlapping. Mean fluorescence intensities of pHrodo and Alexa Fluor 488 were used to calculate pHagosomal pH.  	
    48  2.11.3 Phagosomal pH of E. coli phagosomes Transfection of THP-1, E. coli labeling with pHrodo SE and Alexa Fluor 350 carboxylic acid SE and THP-1 infection were performed as described before. For pH calibration, the infected cells were incubated in 10 mM phosphate-citrate buffer with pre-determined pH (5.0 - 8.0) after non-internalized bacteria were washed away. pHrodo fluorescence (Emission 620 nm) and Alexa Fluor 350 fluorescence (Emission 460 nm) were measured simultaneously in the Fusion-α HT microplate reader (Perkin Elmer). 2.12 Digital confocal microscopy 2.12.1 Intracellular staining Immunofluorescence staining was performed on cells adhered to glass coverslips. Cell fixation was performed using 2.5% paraformaldehyde in PBS for 20 minutes at 37°C. Subsequently, the cells were washed with PBS and incubated in PBS containing 0.2% saponin and 1% normal goat serum (permeabilization buffer) for 15 minutes at room temperature. Specific primary antibody (10 µg/mL) in permeabilization buffer was added to the coverslips and incubated for 45 minutes at room temperature. Then, the cells were washed three times with PBS and incubated with suitable secondary FITC-, Alexa-Fluor 488- or Texas Redconjugated antibody for 45 minutes. Cells were then washed extensively with PBS. Double immunostaining was performed sequentially with rabbit antiVPS33B at 10 µg/mL and mouse anti-subunit H at 10 µg/mL in permeabilization buffer. Texas Red-conjugated goat anti-rabbit IgG (Invitrogen) and Alexa-Fluor  	
    49  488-conjugated goat anti-mouse IgG (Invitrogen), both of which are highly crossadsorbed to minimize cross-reactivity, were used at a dilution of 1:1000. 2.12.2 Microscopy Fixed adherent cells on cover slips were mounted on microscope slides in FluorSaveTM (Calbiochem-Novabiochem, La Jolla, CA) to prevent photobleaching. Digital confocal microscopy using Axioplan II epifluorescence microscope (Carl Zeiss Inc., Thornwood, NY) equipped with 63x/1.4 Plan-Apochromatobjective (Carl Zeiss Inc) was used to analyze the slides. Images were captured using Retiga EX CCD digital camera (QImaging, BC, Canada) operated by Northern Eclipse software (Empix Imaging Inc. Ontario, Canada). 2.13 In vitro phosphatase activity assay para-nitrophenyl phosphate (pNPP) was used as an artificial chromogenic substrate to assay for PtpA phosphatase activity in vitro. The phosphatase reaction contained 50 mM Tris-HCl pH 7.4, 5 mM MgCl2, 2 mM MnCl2, 1 mM DTT and 50 mM pNPP together with 20 µg of purified recombinant PtpA proteins. The reaction was incubated at 37oC, and the absorbance A450 was measured at 10 minutes interval on Bio-rad Model 680 microplate reader. 2.14 In vitro PtpA secretion analysis The DNA encoding His-tagged PtpA under the control of the hsp60 promoter was cloned into the pMV261 vector. The resulting plasmid was introduced into the wild type H37Rv Mtb strain, and the transformed strain was grown to an OD600 = 0.8 in 50 mL of protein-free Sauton’s medium (containing (per L) 4 g of L-  	
    50  asparagine, 0.5 g K2HPO4, 0.5 g MgSO4, 50 mg ferric ammonium citrate, 2 g citric acid, 1 mg ZnSO4 and 60 mL glycerol) supplemented with 0.05% Tween-80. To test whether covalent labeling with fluorescent dyes affect secretion of proteins by Mtb, pHrodo and Alexa-Fluor 488 labeling was performed according to the manufacturer’s protocol. The unlabeled and labeled bacteria were then reinoculated into 50 mL of fresh Sauton’s medium supplemented with 0.05% Tween-80 and grown for 48 hours at 37oC. The culture filtrate proteins were then harvested and precipitated with 10% TCA and 0.1% SDS as described before [143]. The samples were resolved on 10% SDS-PAGE and PtpA secretion into the culture supernatant was analyzed with Western blotting using rabbit anti-PtpA and mouse anti-His-tag antibodies. 2.15 FACS analysis of antibody specificity H37Rv Mtb harboring a plasmid construct for the expression of DsRed (pSMTDsRed) were grown to O.D.600 = 0.8 in 7H9 medium supplemented with 10% OADC and 0.05% Tween-80. The bacteria were washed three times with 7H9 supplemented with 0.05% Tween-80 (7H9T), and 1.0 x 108 was resuspended in 100 µL 7H9T. The indicated primary antibodies were added to the bacteria at a final concentration of 5 µg/mL and incubated at 37oC with shaking for one hour. The bacteria were then washed three times with 7H9T and incubated with either Alexa-Fluor conjugated goat anti-rabbit IgG (Invitrogen) or Alexa-Fluor 488 goat anti-mouse IgG (Invitrogen) secondary antibodies at a dilution of 1:1000 for one hour at 37oC with shaking. The bacteria were then washed with 7H9T and fixed with 2.5% paraformaldehyde for 30 minutes at 37oC. The fixed bacteria were 	
    51  subjected to analysis with FACSCalibur for antibody binding. Expression of DsRed in Mtb allowed specific gating to distinguish the bacterial population from background noise signal. Color compensation was performed to prevent signal overlapping. 2.16 In vitro kinase assay The in vitro kinase reactions contained 25 mM Tris–HCl, pH 7.2, 5 mM MgCl2, 2 mM MnCl2, 1 mM DTT, 10–300 ng of recombinant PtkA kinase, and 0–500nM of substrates. The reactions were started by addition of 10 µCi of γ-[32P]ATP (Perkin-Elmer) or 10 µCi of γ-[32P]GTP followed by 5–30 minutes incubation at room temperature (25°C). At the end of the incubation period, reactions were stopped by the addition of SDS-sample loading buffer and heated at 95 °C for 5 minutes. Samples were resolved by SDS–PAGE on 8% polyacrylamide gels and the gels were silver-stained, and dried. The  32  P-radioactively labeled protein  bands were detected using a PhosphorImager SI (Molecular Dynamics). Bands corresponding to the phosphorylated proteins were cut out and analyzed by scintillation counting (Beckman Coulter LS 6500). Kinase inhibitors were added to the kinase reaction buffer at the following concentrations: staurosporine (100 nM), tartaric acid (10 mM), wortmannin (1 nM), methyl piperazine (100 nM), and phenyl piperazine (100 nM). 2.17 Phospho-amino acid analysis Recombinant PtkA was incubated with γ-[32P]-ATP in the kinase buffer described above (Section 2.16). The sample was hydrolyzed with 6 N HCl at 110oC for 1 hour and separated on a cellulose TLC plate by ascending chromatography [144]. 	
    52  For PtpA phosphorylation by PtkA, the reaction was stopped with sample buffer, loaded into SDS-PAGE 12% containing 8 M urea, and subjected to electrophoresis. The gel was electroblotted onto an Immobilon PVDF membrane [145]. Phosphorylated proteins bound to the membrane were detected by autoradiography using a PhosphorImager apparatus. The  32  P-labelled protein  band corresponding to the migration of PtpA was excised from the membrane and hydrolyzed following the same procedure as described above. After hydrolysis, samples were separated on ascending TLC chromatography [144]. After migration, radioactive amino acids were detected by autoradiography. Phosphoserine,  phosphothreonine  and  phosphotyrosine  standards  were  separated on cellulose TLC plate in parallel and visualized by staining with ninhydrin.  	
    53  CHAPTER 3: RESULTS 3.1 Mtb PtpA excludes host Vacuolar-H+-ATPase to inhibit phagosome acidification 3.1.1 Introduction Mtb pathogenicity partly depends on its ability to inhibit phagosome acidification and maturation processes after engulfment by macrophages. We showed that secreted PtpA, a key protein tyrosine phosphatase needed for Mtb pathogenicity within host macrophages, binds to subunit H of the host V-ATPase machinery, a multi-subunit protein complex in the phagosome membrane that drives luminal acidification. Furthermore, we showed for the first time that macrophage Class C VPS complex, a key regulator of endosomal membrane fusion, associates with V-ATPase during phagosome maturation, indicating a novel role for V-ATPase in coordinating membrane trafficking and phagosome-lysosome fusion. PtpA interaction  with  V-ATPase  is  required  for  the  previously  reported  dephosphorylation of VPS33B and subsequent exclusion of V-ATPase from the phagosome during Mtb infection. These findings showed that inhibition of phagosome acidification in the mycobacterial phagosome is directly attributed to PtpA, revealing the mechanism behind the long-established paradigm of acidification block by Mtb. 3.1.2 Mtb PtpA binds subunit H of human V-ATPase Earlier, we used a substrate trapping assay with a mutant of PtpA to pull down the catalytic substrate of PtpA, VPS33B, from THP-1 cell lysate [32]. Interestingly, when we used wild type recombinant PtpA as bait, we were able to pull down  	
    54  another 50 kDa macrophage protein [32]. We identified this protein by MALDITOF mass spectrometry to be subunit H of human V-ATPase (Figure 28A, Appendix A) and verified its identity by Western analysis (Figure 10A). In vitro kinase assay and phosphoamino acid analysis showed that subunit H is phosphorylated on threonine and therefore could not serve as a catalytic substrate for PtpA, which is a tyrosine phosphatase (Figure 28B and 28C, Appendix A). In vitro protein-protein interaction analysis of PtpA and subunit H demonstrated direct binding of mycobacterial PtpA to host V-ATPase subunit H (Figure 10B and 10C). A hyperbolic curve fitting the 1:1 Langmuir Binding Model and a dissociation constant (Kd) of 1.1 x 10-7 M indicate a relatively high binding affinity. 3.1.3 PtpA binds to amino acid 220-402 of subunit H We determined the PtpA binding site on subunit H by constructing overlapping polypeptide fragments of subunit H. As shown in Figure 10D, only fragments that included the linker region (amino acid 220-402) that connects the N-terminal and C-terminal domains of subunit H were able to bind to PtpA with high affinity. This is consistent with previous predictions that the linker region of yeast subunit H is involved in protein-protein interactions [146]. 3.1.4 The C-terminal alpha helix of PtpA binds subunit H Subunit H was reported to interact with di-leucine based motifs on the Human Immunodeficiency Virus (HIV) Nef protein [78]. Therefore, we performed sitedirected mutagenesis to investigate whether di-leucine motifs or other leucine 	
    55  residues on PtpA participate in binding subunit H (Figure 29A, Appendix A). One mutation in the C-terminal alpha helix (PtpAL146A) abolished PtpA binding to subunit H (Figure 10E). These findings, together with the results presented above, are consistent with the predicted interaction of PtpA bound with subunit H using the protein-docking algorithm, 3D-Garden (Figure 11A and 11B) [147]. Interestingly, the PtpAL146A mutant retained its phosphatase activity with kinetics similar to the wild type protein (Figure 29B, Appendix A). This result is in line with our observation that subunit H is not a catalytic substrate but rather a binding partner of PtpA.  Figure 10. PtpA interacts with V-ATPase subunit H in vitro. (A) In vitro pulldown using recombinant His-tagged Mtb PtpA captured human V-ATPase 	
    56  subunit H from THP-1 macrophage lysate. The eluates were analyzed on Western blots using rabbit anti-subunit H and anti-PtpA antibodies. (B) Recombinant PtpA was subjected to ALPHAScreen Assay with increasing concentration of V-ATPase subunit H. GST served as negative control. Curve fitting yielded a dissociation constant (Kd) of 1.1 x 10-7 M. (C) The reciprocal experiment with increasing PtpA concentration yielded similar results. His-tagged Rv0323c protein was used as negative control. (D) Mapping of the PtpA binding site on V-ATPase subunit H. Overlapping truncated recombinant subunit H protein fragments were incubated with recombinant PtpA proteins in ALPHAScreen assay. Amino acid 220-402 of subunit H is the minimal region required for binding PtpA. (E) Site-directed mutagenesis generated a recombinant mutant PtpA protein (PtpAL146A) that failed to interact with subunit H. 	
   	
    Figure 11. Computer-generated model of PtpA complexed with the VATPase subunit H. Based upon 3D-Garden macromolecular docking analysis [147], the crystal structure of Mtb PtpA (PDB accession number 1U2P) and S. cerevisiae V-ATPase subunit H (PDB accession number 1HO8) were oriented into the predicted binding position. (A) Surface representation of PtpA and subunit H as separated molecules. The C-terminal alpha helix of PtpA is colored in magenta. L146A mutation on PtpA likely alters the C-terminal alpha helix, leading to the defective binding. (B) Computer prediction of PtpA binding on subunit H with the C-terminal alpha helix of PtpA docking on to the cleft region between the N-terminal and C-terminal domain of subunit H. 	
    	
    57  3.1.5 PtpA binds V-ATPase subunit H in vivo We applied immunoprecipitation with specific antibodies to investigate the in vivo interaction between PtpA and subunit H during macrophage infection. Antibodies against subunit H were able to pull down PtpA from lysates of THP-1 cells infected with H37Rv Mtb expressing wild type PtpA, confirming the physiological relevance of PtpA-subunit H interaction during Mtb infection (Figure 12A). To further monitor PtpA interaction with subunit H, we used a modified “Split Trp” mycobacterial assay for detecting protein-protein interactions [137, 138]. In this system, a tryptophan auxotrophic strain of M. smegmatis grows in the absence of tryptophan only if the tested proteins interact with each other. As shown in Figure 12B, co-expression of subunit H and PtpA fusion proteins restored mycobacterial growth on 7H9 agar plates, confirming protein-protein interaction within a surrogate host. Co-expression of subunit H with the bindingdefective PtpAL146A did not lead to mycobacterial growth, indicating that this phosphatase-active PtpA mutant lost its ability to bind subunit H. The phosphatase-defective PtpAD126A retained its ability to bind to subunit H, further indicating the interaction is independent of the catalytic activity of PtpA (Figure 12B).  	
    58  Figure 12. PtpA and V-ATPase subunit H interacts in vivo. (A) PtpA coimmunoprecipitated with subunit H from THP-1 infected with Mtb expressing PtpA. Immunoprecipitation (IP) was performed with either rabbit anti-subunit H (α-H) antibodies (Ab) or rabbit IgG as negative control. (B) PtpA and subunit H interacted in the “Split-Trp” protein fragment complementation assay to facilitate Ntrp and Ctrp reassembly required for Trp biosynthesis, thus enabling growth of the M. smegmatis Trp- strain coexpressing Ntrp-PtpA and subunit H-Ctrp under acetamide (ACE) induction (Middle panel). Ntrp-PtpAL146A failed to interact with subunit H-Ctrp to restore M. smegmatis growth. Ntrp-ESAT6 and CFP10-Ctrp were used as a positive control. The negative control consisted of Ntrp and Ctrp fragments alone. Top panel shows that all transformed strains are capable of growing on Trp-supplemented media. No growth could be observed in the absence of acetamide induction and exogenous Trp (Bottom panel) (C) Loss of PtpA binding to V-ATPase subunit H impairs Mtb survival within host macrophages. V-ATPase binding-defective PtpAL146A expressing Mtb strain showed a 2-log reduction in bacterial load compared to the wild type strain at 6 days post-infection.  3.1.6 PtpA binding to subunit H is required for Mtb intracellular survival To test whether PtpA binding to subunit H is required for Mtb pathogenicity, we investigated the ability of the binding-defective PtpAL146A Mtb strain to survive in  	
    59  THP-1 macrophage-like cells. As shown in Figure 12C, a ΔptpA strain complemented with a construct encoding PtpAL146A was attenuated within the macrophage in a manner similar to that of the ΔptpA knockout strain, while the parental and the complement strains were able to establish a stable infection after 3 days. Expression and stability of the wild type and mutant PtpA proteins in these strains were confirmed with Western blot analysis (Figure 30A, Appendix A). At 6 days post-infection, the binding defective strain showed a 2-log reduction in CFU compared to the wild type strain. The PtpAD126A strain also exhibited a similar intracellular survival phenotype as the ΔptpA strain. While the parental and complemented strains, entered into growth phase within the macrophage, the ΔptpA and the binding-defective strains were continually cleared, establishing the importance of PtpA interaction with the V-ATPase machinery for Mtb survival within macrophages. 3.1.7 PtpA inhibits phagosome acidification The in vivo PtpA-subunit H interaction and the impaired intracellular survival of the binding-defective PtpAL146A strain indicated that PtpA interferes with the phagosome acidification process. To examine this hypothesis, we analyzed the pH of Mtb-containing phagosomes by FACS. Parental and mutant strains were dually labeled with the pH-sensitive pHrodo fluorescent dye and the pHinsensitive Alexa Fluor 488 prior to infection of THP-1. Covalent labeling of the bacteria with fluorescent dyes does not affect PtpA secretion (Figure 30B, Appendix A). Overlaid FACS histograms showed a clear increase in the mean fluorescence intensity of pHrodo in phagosomes harboring ΔptpA, ΔptpA 	
    60  complemented  with  the  phosphatase-inactive  ptpAD126A,  and  ΔptpA  complemented with the binding-defective ptpAL146A (corresponding to pH 5.8, 5.6, and 5.55 respectively), as compared to phagosomes containing parental Mtb and ΔptpA complemented with wild type ptpA (corresponding to pH 6.7 and 6.4, respectively) (Figure 13A and 13B, Figure 31, Appendix A and Table 3). These results indicated a direct functional role for PtpA in the inhibition of Mtb phagosome acidification, which is dependent on both the binding ability to subunit H and phosphatase activity of PtpA.  Figure 13. Mtb PtpA inhibits phagosome acidification. (A) The indicated Mtb strains were labeled with the pH-sensitive fluorescent dye (pHrodo) and used to infect THP-1 macrophage-like cells. Phagosomal pH of the infected macrophages was measured with FACS. Wild type Mtb maintained a 	
    61  phagosomal pH of around 6.7 while phagosomes of both ΔptpA and PtpAL146A strains were acidified to around pH 5.5. * = p < 0.05; ** = p < 0.01, significant difference compared with H37Rv by Student’s t test. (B) Overlaid FACS histograms of the pHrodo fluorescence intensity of the wild type H37Rv and ΔptpA phagosomes show a shift in phagosomal pH in the presence of PtpA. (C) THP-1 macrophage-like cells were transfected with GFP-tagged constructs of wild type and mutant PtpA proteins and infected with pHrodo-labeled E. coli. Phagosomal pH was measured with spectrofluorometry during the course of infection. Wild type PtpA inhibits E. coli phagosome acidification (pH 6.5) while the binding-defective PtpAL146A failed to block acidification (pH 5.6). Concanamycin (CMA) was added as a positive control for inhibition of phagosome acidification. Untransfected THP-1 and THP-1 transfected with pEGFP vector served as negative control. (D) Digital confocal microscopy of the infected macrophages in (C) confirms the inhibition of phagosome acidification by Mtb PtpA. Green: Expression of GFP or GFP-tagged PtpA constructs. Red: pHrodo fluorescence in acidified phagosomes. Blue: Alexa Fluor 350 labeled E. coli.  WT ΔptpA Complement D126A L146A  pHrodo MFI  AF 488 MFI  Fluorescence Ratio (pHrodo/AF488)  pH  Standard Error  p value  42.23 47.53 44.20 50.87 49.17  15.40 14.17 15.15 14.47 13.90  2.74 3.36 2.92 3.52 3.54  6.73 5.78 6.42 5.58 5.55  0.38 0.04 0.01 0.09 0.09  NA 0.012 NS 0.0067 0.0063  Table 3. Phagosomal pH of Mtb strains in THP-1 	
   AF488: Alexa Fluor 488; MFI: Mean Fluorescence Intensity; N/A: Not Applicable; NS: Not Significant.  To  examine  PtpA’s  ability  to  modulate  phagosome  acidification  independently of other Mtb proteins, we transfected THP-1 macrophage-like cells with plasmids expressing GFP-fused parental or mutated forms of PtpA. All constructs were expressed at similar levels, and the survival and phagocytic ability of the macrophages were not impaired by the transfection (Figure 32A and 	
    62  32B, Appendix A). Transfected macrophages were infected with E. coli labeled with pHrodo and Alexa Fluor 350, and the phagosomal pH was assessed by spectrofluorometry (Figure 32C, Appendix A). Expression of wild type PtpA within THP-1 macrophage-like cells inhibited acidification of E. coli containing phagosomes, whereas both catalytically inactive PtpAD126A and binding mutant PtpAL146A were unable to block phagosome acidification, showing a marked decrease in phagosomal pH similar to untransfected and empty vector control macrophages (Figure 13C). Digital confocal microscopy of the infected macrophages confirmed these findings (Figure 13D and Figure 32D, Appendix A). These results established that PtpA alone is able to prevent effectively phagosome acidification in the macrophage. This inhibition depends on both PtpA phosphatase activity and its ability to bind to host V-ATPase machinery. 3.1.8 The V-ATPase machinery recruits the Class C VPS complex The interaction of PtpA with both subunit H and the previously described VPS33B [32] indicated that V-ATPase and Class C VPS complexes might be in close proximity or even interacting directly during phagosome maturation and endosome-lysosome fusion. To investigate this hypothesis, we conducted immunoprecipitation  experiments  in  THP-1  macrophage-like  cells  using  antibodies against VPS33B as bait and probed for subunits H, B and E of the VATPase complex and VPS18 of the Class C VPS complex by Western blot. In uninfected THP-1, faint bands corresponding to subunits of the V-ATPase complex were captured with VPS33B and VPS18 (Figure 14). This indicates that V-ATPase interacts transiently with the Class C VPS to regulate normal 	
    63  endosome-lysosome fusion in resting cells. Furthermore, this interaction is upregulated upon phagocytosis of E. coli as demonstrated by the prominent bands of V-ATPase subunits captured 2 hours post-infection (Figure 14), further supporting that the Class C VPS complex interaction with V-ATPase plays a key role in phagosome maturation and fusion with lysosomes.  Figure 14. Co-immunoprecipitation of V-ATPase subunits with Class C VPS in THP-1. V-ATPase subunits (H, B and E) can be captured with VPS33B and VPS18 from uninfected THP-1 macrophage-like cells when immunoprecipitation was performed using 4 mg of soluble lysate. The interaction was not detected when 2 mg of lysate were used. Prominent bands corresponding to V-ATPase subunits can be pulled down from 2 mg of lysate from E. coli-infected THP-1 macrophage-like cells.  3.1.9 Mtb PtpA blocks the interaction between Class C VPS and V-ATPase complexes When THP-1 macrophage-like cells were infected with Mtb H37Rv, V-ATPase subunit H, B or E were not captured using antibodies against VPS33B (Figure 15A). Mtb inhibition of V-ATPase and Class C VPS interaction was maintained 24 hours post-infection (Figure 15B). However, the V-ATPase subunits coimmunoprecipitated with VPS33B and VPS18 in THP-1 macrophage-like cells 	
    64  infected with ΔptpA Mtb. Complementation of ΔptpA mutant with wild type ptpA restored the ability of Mtb to inhibit Class C VPS interaction with the V-ATPase complex. Furthermore, complementation with ptpAD126A but not ptpAL146A restored Mtb inhibition of Class C VPS and V-ATPase interaction (Figure 15A). These results indicated that Mtb infection disrupted the V-ATPase and Class C VPS association, that PtpA is responsible for blocking this interaction, and that PtpA binding to subunit H is specifically necessary for this disruption. Figure 15 (next page). Mtb PtpA disrupts the interaction between the Class C VPS and V-ATPase complexes during infection. Immunoprecipitation (IP) was performed with anti-VPS33B (α-VPS33B) using 2 mg of soluble lysate from THP-1 macrophage-like cells infected with the indicated strain for 2 hours and followed by Western analysis with the indicated antibody. Rabbit IgG was used as negative control. (A) The Class C VPS (VPS33B and VPS18) interacts with the V-ATPase complexes (Subunit B, H, and E) from THP-1 infected with E. coli, ΔptpA and PtpAL146A. The parental H37Rv, PtpAD126A and the complemented strain disrupted the host proteins interaction. (B) Mtb disruption of V-ATPase interaction with Class C VPS can be observed 24 hours post-infection. Immunoprecipitation (IP) was performed with either rabbit IgG (IgG) or antiVPS33B (α-VPS33B) using 2 mg of soluble lysate from THP-1 infected for 24 hours. For the ΔptpA and ΔptpA complemented with PtpAL146A strains, V-ATPase interaction with Class C VPS remains upregulated 24 hours post-infection, suggesting continued delivery of lysosomal contents and killing of the bacteria in the phagosomes.  	
    65  3.1.10 VPS33B remains phosphorylated in THP-1 infected with bindingdefective PtpA VPS33B phosphorylation is required for phagosome-lysosome fusion and its dephosphorylation by Mtb PtpA blocks this process [32]. Since the Class C VPS  	
    66  complex is recruited to the V-ATPase, we asked whether PtpA binding to VATPase subunit H affects the phosphorylation status of VPS33B during Mtb infection. To address this question, we immunoprecipitated VPS33B and analyzed its phosphorylation with the anti-phosphotyrosine antibody 4G10. When E. coli was used to infect macrophages, VPS33B was phosphorylated (Figure 16). However, as expected, infection with Mtb inhibited VPS33B phosphorylation. Consistent with these findings, ΔptpA strain failed to reduce VPS33B phosphorylation, and this was reversed by complementation with wild type ptpA (COMP). Interestingly, VPS33B remained phosphorylated in macrophages infected with ΔptpA complemented with either the phosphatase-defective ptpAD126A, or the binding-defective ptpAL146A, despite being catalytically active (Figure 16). This indicates that the binding of PtpA to subunit H of V-ATPase is a prerequisite for VPS33B dephosphorylation. This is consistent with our findings that the loss of PtpA interaction with subunit H abolished its functions within the host cell.  	
   Figure 16. Western blot analysis of VPS33B phosphorylation in vivo. VPS33B remained phosphorylated in macrophages infected with the PtpAL146A expressing strain. Top panel was probed with anti-phosphotyrosine antibody 4G10, and bottom panel is probed with anti-VPS33B to ensure equal loading.  	
    67  3.1.11 PtpA binding to subunit H participates in the exclusion of the VATPase from the Mtb phagosome Disruption of the interaction between Class C VPS and V-ATPase by PtpA suggests that PtpA directly impairs V-ATPase trafficking to the Mtb phagosome. To verify this hypothesis, we used digital confocal microscopy to monitor Class C VPS and V-ATPase localization within infected macrophages, represented by VPS33B  and  subunit  H,  respectively.  In  control  experiments,  where  macrophages were infected with E. coli, both VPS33B and subunit H co-localized to the phagosome (Figure 17A). In contrast, phagosomes containing Mtb were decorated with VPS33B but excluded subunit H, indicating a lack of V-ATPase recruitment. To directly assign a role to PtpA, we examined cells that were infected with the ΔptpA strain and found co-localization of VPS33B and subunit H on the mycobacterial phagosomal membrane. These findings were confirmed with the observation that macrophages infected with ΔptpA complemented with wild type ptpA led to a phenotype similar to that observed with wild type strain (Figure 17B). Additionally, co-recruitment of subunit H and VPS33B to bacillicontaining phagosomes was observed for ΔptpA strain complemented with either the catalytically inactive ptpAD126A or the binding-defective ptpAL146A. Therefore, PtpA binding to V-ATPase subunit H is required for blocking V-ATPase trafficking to the phagosome, which ultimately leads to the inhibition of Mtb phagosome acidification. Although we have shown that VPS33B and subunit H antibodies are highly specific (Figure 33, Figure 34 and Figure 35, Appendix A), to overcome 	
    68  any potential cross-reactivity, we also performed double transfection of THP-1 with GFP and DsRed2 fused constructs of subunit H and VPS33B respectively, allowing for direct visualization of these proteins and their localization within the macrophage. Co-localization of GFP-subunit H and DsRed2-VPS33B in uninfected macrophages provides further support that V-ATPase and Class C VPS functions in the same pathway to regulate normal endosome-lysosome fusion. When transfected macrophages were infected with the same Mtb strains, similar patterns of subunit H and VPS33B localization was observed as compared to the immunofluorescent stained macrophages (Figure 18). This result further supports that PtpA binding to subunit H is needed for the exclusion of V-ATPase from the phagosome.  	
    69  Figure 17. Confocal microscopy of THP-1 infected with E. coli or indicated Mtb strains. (A) Localization of VPS33B and V-ATPase subunit H was detected with immunofluorescence staining. V-ATPase was excluded from the Mtb phagosome in macrophages infected with the wild type strain while the ΔptpA phagosome acquired V-ATPase. The phagosomes of the binding-defective 	
    70  PtpAL146A expressing strain also failed to exclude host V-ATPase. (B) Quantification of the confocal data shown in (A). Values are the mean ± SD of phagosome colocalization with V-ATPase subunit H in 50-80 cells from three independent experiments. *** = p < 0.001, significant difference compared with H37Rv by Student’s t test.  	
    71  	
   Figure 18. Confocal microscopy of infected THP-1 macrophage doubly transfected with GFP-subunit H and DsRed2-VPS33B. (A) The transfected macrophages were infected with E. coli or indicated Mtb strains, and the localization of subunit H and VPS33B was directly visualized. V-ATPase was 	
    72  excluded from the Mtb phagosome in macrophages infected with the wild type strain while the ΔptpA phagosome acquired V-ATPase. The phagosomes of the binding-defective PtpAL146A expressing strain also failed to exclude host VATPase. (B) Quantification of the confocal data shown in (A). *** = p < 0.001, significant difference compared with H37Rv by Student’s t test. 	
   3.2 PtpA is a substrate for the novel protein-tyrosine kinase Mtb PtkA 3.2.1 Introduction Protein tyrosine phosphorylation has long been recognized to play a key role in the regulation of numerous fundamental cellular processes in eukaryotes [148] and various bacterial species [149-151]. Thus far, bacterial phosphotyrosine signaling has been shown to be involved in cell division, antibiotic production, capsule synthesis, and host infection [150-155]. The first indication of bacterial protein-tyrosine kinase activity was demonstrated in Escherichia coli, in which the presence of phosphotyrosine in partial hydrolysates of proteins was shown [156]. Since then, a large number of protein-tyrosine kinases and phosphatases were identified in various bacterial species. These include two protein kinases from Streptococcus pneumoniae that autophosphorylate on tyrosine residues and are involved in the regulation of capsular polysaccharide production [150, 152]. Pathogenic Yersiniae contain an extrachromosomal virulence plasmid that encodes a PTP named YopH, which is injected into the host through a Type III secretion system encoded by the same plasmid [157]. Once inside the host cells, YopH modulates host-signaling pathways by dephosphorylating p130Cas to inhibit phagocytosis of the bacteria [157].  	
    73  In Mtb, the presence of tyrosine phosphorylation activity has been predicted since the identification of a 55 kDa protein that was recognized by the 4G10 anti-phosphotyrosine antibody in cell extracts from virulent but not avirulent Mtb  strains  [158].  Furthermore,  Mtb  possesses  two  protein-tyrosine  phosphatases, PtpA and PtpB, further implying the existence of phosphotyrosine signaling activity in Mtb [104, 105]. Previous studies have shown that genes encoding kinases are often located in the proximity or in the same operon with the genes encoding their substrates [159]. Studies on bacterial PTKs and PTPs also showed that their corresponding genes are frequently clustered in an operon such that the expression of the PTKs and PTPs can be coordinated and regulated in concert [e.g. 160]. Within the Mtb genome, ptpA is located in an apparent operon with Rv2232 and Rv2235 upstream and downstream, respectively. The Rv2232 gene encodes a protein annotated as a member of the Haloacid Dehalogenase likehydrolases (HAD) super-family, which catalyze the hydrolysis of various molecules. Interestingly though, with more than 3000 sequenced proteins, this super-family  comprises  enzymes,  such  as  phosphatases,  ATPases,  phosphonatase (P-C bond hydrolysis), and sugar phosphomutases [161, 162] that are specialized in phosphoryl transfer. In this study, we show that the Rv2232 encoded protein is a novel protein-tyrosine kinase that phosphorylates PtpA.  	
    74  3.2.2 PtpA interacts with the protein encoded by Rv2232 As described earlier, genes in the proximity of Mtb kinases and phosphatases are proposed to act as substrates or be under the control of these regulatory proteins. We used AlphaScreen technology to determine and measure the interaction between PtpA and Rv2232. As shown in Figure 19, the protein encoded by Rv2232 binds to PtpA (Kd = 3.0 µM). Interestingly, in the presence of ATP, a stronger affinity was measured as reflected by a Kd of 1.3 µM. GTP was also able to serve as a phosphate donor with a Kd of 1.8 µM. These results suggest that: (i) the presence of the phosphate donor increases the interaction affinity of the protein encoded by Rv2232 for PtpA, (ii) GTP serves as an alternative phosphate donor, and (iii) ATP is the preferred substrate based on the lower Kd.  Figure 19. Rv2232 interacts with PtpA in vitro. Protein–protein interaction between Rv2232 and PtpA was measured using AlphaScreen technology. The 	
    75  dissociation constant (Kd) was calculated using biotinylated Rv2232 and His6tagged PtpA in the presence or absence of 50 µM ATP (A) or GTP (B). GST was used as negative control. In the absence of ATP or GTP, Rv2232 interacts with PtpA in vitro with a Kd of 3.0 µM. Addition of either ATP (A) or GTP (B) results in a stronger interaction between Rv2232 and PtpA with a Kd of 1.3 µM (ATP) and 1.8 µM (GTP). GST negative control shows a Kd of 2.9 x 10 7 µM, indicating absence of significant protein-protein interaction.  3.2.3 ORF Rv2232 encodes an autophosphorylated protein-tyrosine kinase Since ptpA is clustered with Rv2232, and rationalizing that as a result of such proximity, the Rv2232-encoded protein might serve as a substrate for the tyrosine phosphatase PtpA, the phosphorylation status of Rv2232 was investigated. Indeed, in vitro phosphorylation assays using purified recombinant protein expressed from the Rv2232 gene revealed that this protein possesses ATP concentration- and time-dependent autophosphorylation activity (Figure 20).  Figure 20. Rv2232 autophosphorylation in vitro. Recombinant PtkA is autophosphorylated in an ATP dose- (A) and time- (B) dependent manner. The 	
    76  upper panels in (A) and (B) show phosphorylation visualized by autoradiography, and the lower panels represent the silver-stained gel. Molecular size markers (MM) were shown. To determine the identity of the phosphorylated residues, phospho-amino acid analysis of the [32P]-phosphorylated protein was performed under acidic conditions. As shown in Figure 21, this protein was found to be phosphorylated on tyrosine residues. These data provided support for the protein encoded by Rv2232 to be the first protein tyrosine auto-kinase identified in Mtb, and thus we named it as PtkA.  Figure 21. Phospho-amino acid analysis of PtkA. PtkA is phosphorylated on tyrosine residues as revealed by phospho-amino acid analysis. The migration of hydrolysed protein was detected by TLC against phospho-amino acids standards 	
    77  (Retention Factor (Rf): phosphor-serine (p-Ser)=0.22; phosphor-threonine (pThr)=0.25; phosphor-tyrosine (p-Tyr)=0.30; PtkA=0.29).. 3.2.4 Sequence-function analysis of PtkA Bacterial protein-tyrosine kinase signatures, termed Walker A and B motifs [163], and the glycine-rich loop GXGXXGXV motif [164], are not present in PtkA. As we were unable to find any conserved kinase signature from in silico analysis of PtkA, we constructed a series of mutants in order to identify key functional residues in the protein. We mutated all three lysine residues with alanine residues in Rv2232 rationalizing that they might be required for ATP binding. Furthermore, all three tyrosine residues present in Rv2232 were mutated to determine potential phosphorylation sites. Lastly, the first aspartate (D85) in the DXD motif, which is conserved and essential for the catalytic activity in the HAD super-family members, was mutated to determine its role in the catalytic mechanism of Rv2232 [165]. All of the mutated proteins were individually tested for their ability to undergo autophosphorylation by means of radiolabeled γ-[32P]ATP incorporation in an in vitro kinase assay (Table 4). 3.2.4.1 Mutation of lysine residues Binding of ATP in an enzymatic autophosphorylation reaction depends on lysine residues [166]. Thus, we constructed PtkA mutants K184M, K217M, and K270M and found all of them unable to bind ATP in a dose-dependent manner (Figure 22). Moreover, a concomitant increase of 2 to 14-fold in the KM values revealed a decrease in the affinity of these mutants for ATP as detailed in Table 1.  	
    78  3.2.4.2 Mutation of tyrosine residues We have shown earlier that PtkA is phosphorylated on tyrosine residues (Figure 21). To precisely map the tyrosine phosphorylation site, Y146A, Y150A and Y262A mutants were constructed. Only the Y262A mutant failed to undergo autophosphorylation, while the Y146A and Y150A mutants incorporated  32  P in a  dose-dependent manner (Figure 22). These results indicate that Y262 is the autophosphorylated tyrosine in the PtkA backbone. Interestingly, the affinity of these proteins for ATP binding and the enzymatic efficiency of  32  P incorporation  were not affected as reflected by similar values measured for KM and Kcat/KM respectively (Table 4).  Figure 22. Mutational studies of PtkA catalytic mechanism. Autophosphorylation activity of a series of purified recombinant PtkA point mutant 	
    79  proteins was assessed by incubation with increasing concentrations of γ-[32P] ATP. The reactions were resolved on silver-stained SDS-PAGE, and phosphorylation was visualized with autoradiography. Molecular size markers (MM) were shown. 3.2.4.3 Mutation of D85 The mutation D85A, which represents the first aspartate in the conserved DXD motif within the HAD super-family was found to be essential for PtkA autophosphorylation activity (Figure 22). This finding is supported by a considerable decrease in the enzymatic efficiency according to the measured Kcat/KM for this mutant, as well as by the significant decrease in the ATP-binding affinity (Table 4). Table 4. Autophosphorylation kinetic values of parental and mutated PtkA PtkA Wild type Y146A Y150A  Vmax (pmolŸ min-1Ÿ mg-1) 1331 1877 1967  Km (nM) 27 54 68  Kcat (s-1) 9.73 x 10-4 1.40 x 10-4 1.47 x 10-3  Kcat/Km (M-1Ÿ s-1) 3.60 x 104 2.59 x 104 2.16 x 104  Y262A D85A K184M K217M K270M  855 533 9 x 10-5 117 302  59 118 40 89 379  6.38 x 10-4 3.98 x 10-4 6.79 x 10-11 8.71 x 10-5 2.26 x 10-4  1.08 x 104 3.40 x 103 1.68 x 10-3 9.76 x 102 5.97 x 102  3.2.5 PtpA is a substrate of PtkA To test whether PtkA can serve as a substrate for the tyrosine phosphatase PtpA, PtpA was supplemented in the autophosphorylation reaction. Interestingly, we found the opposite of the hypothesized relationship. PtkA autophosphorylation did not decrease upon addition of PtpA. Moreover, PtpA acted as a substrate of PtkA. As shown in Figure 23, PtkA phosphorylates PtpA (lane 1) and the generic 	
    80  tyrosine kinase substrate myelin basic protein (MBP) (lane 2). Moreover, PtkA phosphorylated PtpA in an ATP dose- and time-dependent manner (Figure 24A and 24B), demonstrating that the proitein-tyrosine phosphatase PtpA is a cognate substrate of PtkA.  Figure 23. PtkA phosphorylates PtpA in vitro. PtkA was incubated with PtpA in an in vitro kinase reaction including γ-[32P]-ATP. Myelin basic protein (MBP) was used as an artificial substrate. The results show that PtpA is a cognate substrate for PtkA. Molecular size markers (MM) were shown.  Figure 24. PtkA phosphorylates PtpA in dose- and time-dependent manner. (A) Increasing concentrations of PtpA were incubated with PtkA in an in vitro 	
    81  kinase reaction with γ-[32P]-ATP, subjected to silver-stained SDS-PAGE and visualized with autoradiography. An increase in PtpA phosphorylation signal can be observed as PtpA concentration increases, indicating a genuine PtkA kinase activity on PtpA. (B) PtkA and PtpA were incubated in an in vitro kinase reaction over the indicated time periods. Increase in PtpA phosphorylation over time can be observed. The lower panel in (B) represents the silver-stained SDS-PAGE, showing equal loading of PtpA. Molecular size markers (MM) were shown. To determine the amino acid residues affecting the interaction between PtkA and PtpA, we performed protein-protein interaction studies with a series of PtkA mutants in an AlphaScreen assay. The results listed in Table 5 show that K184M, Y146A, and Y150A PtkA mutants interacted with PtpA as reflected with Kd values ranging between 0.85 - 1.5 µM. This range is similar to the Kd of 1.27 µM obtained for the wild-type protein. However, K217M, K270M, Y262A and D85A mutants showed Kd values between 3- to 6-fold higher than that obtained with native PtkA (Kd = 1.27 µM) (Table 5). These results support the finding that Y262A is the phosphorylated residue (Kd = 6.6 µM), as a decrease of 5-fold in the interaction was measured with respect to the wild type protein (Kd = 1.27 µM). In addition, K217M and K270M, with Kd of 4.98 and 7.08 µM, respectively, appear to be the two residues involved in ATP binding as shown in Table 4. The measured Kd of D85A mutant (3.32 µM) also indicated that this residue is involved in the reaction as an interaction with PtpA of about 3-fold weaker than the wild type protein (Kd = 1.27 µM) was measured. We cannot accurately determine if K184 is an essential amino acid for the catalytic activity of the enzyme, as its autophosphorylation was not observed (Figure 22). Nevertheless, an interaction similar to the wild-type protein was measured (Kd = 1.47 µM).  	
    82  Table 5. Kd values of PtpA interacting with parental and mutated PtkA proteins PtkA  PtpA  Kd (µM)  Wild type Y146A Y150A  Wild type Wild type Wild type  1.27 0.97 0.85  Y262A D85A K184M  Wild type Wild type Wild type  6.60 3.32 1.47  K217M K270M Wild type  Wild type Wild type Y128A/Y129A  4.98 7.08 1.24  3.2.6 PtkA can phosphorylate both Y128 and Y129 residues of PtpA Phospho-amino acid analysis of PtkA-phosphorylated PtpA showed that PtpA, like PtkA, is phosphorylated on tyrosine residues (Figure 25A). To determine the specific tyrosine phosphorylated residue, we replaced the three tyrosine residues in PtpA with alanine, generating Y67A, Y128A, and Y129A variant forms of the PtpA protein. All three PtpA tyrosine variants were able to incorporate 32P (Figure 25B, lanes 2-5). The proximity of both Y128 and Y129 residues indicated that upon mutation of one of these residues, the phosphorylation would undergo on the adjacent residue. Therefore, we constructed a double mutant, where both Y128 and Y129 were replaced with alanine. PtkA failed to phosphorylate this double mutant (Figure 25B, lane 1), indicating that both residues exist in the phosphorylated form. The binding affinity between PtkA and the PtpA Y128-129A double mutant was not affected by the residue replacement, as a Kd of 1.24 µM is very similar to the Kd of 1.27 µM obtained between PtkA and the parental PtpA.  	
    83  Figure 25. PtkA phosphorylates PtpA on tyrosine residues. (A) Phosphoamino acid analysis of PtpA phosphorylated by PtkA in an in vitro kinase reaction with γ-[32P]-ATP. Retention factor (Rf): phosphoserine = 0.49; phosphothreonine = 0.55; phosphotyrosine = 0.59; PtpA = 0.61). (B) Site directed mutagenesis of PtpA was performed to mutate candidate tyrosine phosphorylation sites to alanine. The recombinant mutant proteins were purified and subjected to in vitro kinase reaction with PtkA in the presence of γ-[32P]-ATP. The reactions were resolved on silver-stained SDS-PAGE and visualized with autoradiography. PtkA failed to phosphorylate PtpAY128A-Y129A, pointing to Y128 and Y129 as the residues phosphorylated by PtkA.  	
    84  CHAPTER 4: DISCUSSION 4.1 PtpA exclusion of the V-ATPase from phagosomal membrane More than a decade ago, Sturgill-Koszycki et al. demonstrated that macrophages fail to acidify phagosomes containing Mycobacteria because these phagosomes did not accumulate the V-ATPase responsible for phagosomal acidification [99]. This suggested that Mtb actively inhibits the fusion of its phagosome with vesicles harboring the V-ATPase complex [99]. Despite the lack of a mechanistic explanation, the exclusion of the V-ATPase complex during Mtb infection was a long-established paradigm. We showed that the lack of phagosome acidification is directly attributed to Mtb secreted protein PtpA, which specifically inhibits VATPase trafficking to the mycobacterial phagosome during phagosome maturation. During Mtb infection, PtpA positions itself within a cleft linker region between the N-terminal and C-terminal domains of the V-ATPase subunit H, which was predicted to serve as a platform for the interaction with other proteins [146]. In fact, the V-ATPase subunit H and the HIV Nef protein were previously shown to interact through a specific di-leucine motif [78]. We were able to identify a catalytically active Leucine mutant of PtpA (Leu146Ala), which lost its ability to bind to subunit H but retained its phosphatase activity. Due to its location within the core region of PtpA, this mutation most likely altered the conformation of the entire C-terminal alpha helix, leading to the defective binding. These and our binding sites mapping results together with a computer-generated model based upon a protein-docking algorithm (3D-Garden) confirmed the cleft region of 	
    85  subunit H as the binding site for the C-terminal alpha helix of PtpA (Figure 11A and 11B). The observed binding to the V-ATPase subunit H provided the first evidence that PtpA has a major role in interfering with the Mtb phagosome acidification process. Indeed, we showed that the loss of PtpA interaction with the host V-ATPase machinery resulted in the failure to block phagosome acidification and impaired Mtb survival within the host macrophage. The interaction with V-ATPase subunit H and dephosphorylation of VPS33B are both required for PtpA inhibition of macrophage phagosome-lysosome fusion and phagosome acidification. In fact, the loss of either its phosphatase activity or binding to subunit H renders PtpA non-functional within the host cell. We have shown earlier that Mtb PtpA dephosphorylates the host macrophage protein VPS33B, a key regulator of membrane fusion, leading to inhibition of phagosome-lysosome fusion [32]. The identification of V-ATPase subunit H as an additional key partner to the process of PtpA-mediated phagosome maturation arrest suggested a possible link between Class C VPS and V-ATPase. During phagosome maturation, lysosomal V-ATPase is directly recruited to the phagosomal membrane for luminal acidification [46]. The Class C VPS complex has been implicated in this process as a membrane tethering factor. Although studies of Drosophila neurons and mammalian renal medulla cells have shown interaction between subunits of V-ATPase and SNARE proteins [81, 82], our studies of PtpA in Mtb demonstrate for the first time a direct interaction between the Class C VPS and V-ATPase. We further showed that this interaction could be detected, albeit at a weaker level, in non-infected 	
    86  macrophages, indicating that association of Class C VPS and V-ATPase is not limited to phagosome maturation only, but occurs transiently in resting cell to regulate normal endosome-lysosome fusion. However, this association is likely upregulated during phagocytosis in order to deliver lysosomal contents to the phagosome. This provides compelling evidence that V-ATPase is a key player in the membrane fusion machinery, particularly in cooperating with the Class C VPS complex to tether the fusing phagosomal or endosomal membrane with the lysosome (Figure 6). Disruption by Mtb PtpA further exemplifies the importance of this interaction in host defense mechanism. The exact mechanism by which PtpA is secreted across the phagosomal membrane remains unclear. However, using electron microscopy, neutralizing antibodies and Western blot analysis we have previously shown that PtpA is present in the host cytosol milieu [32, 167]. There is evidence suggesting that bacterial proteins less than 70 kDa in molecular size can cross the phagosomal membrane [130]. This observation might be related to the more recent discovery that the ESX-1 secretion system with its substrate ESAT-6 can activate the inflammasome and perturb host cell membrane to facilitate the translocation of mycobacterial proteins into the macrophage cytosol [131, 168]. Interestingly, a recent study showed that Mtb deletion mutant in secA2, which encodes for a secondary general secretion pathway that does not require the typical N-terminal signal peptide, failed to inhibit phagosome acidification [169]. This similar phenotype to the ΔptpA strain suggests that the SecA2 protein secretion system  	
    87  might be responsible for the secretion of PtpA. Further investigation is needed to elucidate the mechanistic details of PtpA secretion.  Figure 26. A model for the specific exclusion of V-ATPase and the inhibition of mycobacterial phagosome acidification by PtpA. During infection, the V-ATPase recruits the Class C VPS complex, possibly associated with VPS39 and VPS41 as the HOPS complex, aiding the tethering of the fusing phagosomal and lysosomal membranes. PtpA, secreted into the host cytosol by Mtb, binds to subunit H of the V-ATPase complex, disrupting the interaction between the two protein complexes and localizing itself near its catalytic substrate VPS33B. PtpA then dephosphorylates and inactivates VPS33B, thereby shutting down the membrane fusion machinery. Binding to subunit H therefore allows PtpA to specifically inhibit V-ATPase trafficking to the mycobacterial phagosome. With these results, in our current model, we suggest a two-step process for PtpA exclusion of V-ATPase and inactivation of VPS33B. During Mtb infection, PtpA binding to subunit H may first disrupt initial membrane tethering and localize PtpA to the proximity of its catalytic substrate VPS33B (Figure 26). Subsequent dephosphorylation of VPS33B would then inactivate the entire membrane fusion machinery and its downstream effectors, preventing delivery of V-ATPase to the  	
    88  mycobacterial phagosome. This would explain the failure of the phosphatasedefective mutant PtpAD126A, which retains the capability of binding to V-ATPase and disrupting V-ATPase and Class C VPS complexes association, to prevent VATPase trafficking to the Mtb phagosome. Our observation that VPS33B remains phosphorylated in both the phosphatase-defective and binding-defective strain supports such a two-step process, indicating that PtpA binding to subunit H is a prerequisite for VPS33B dephosphorylation. This is also supported by our confocal data where both the phosphatase-defective PtpAD126A and bindingdefective PtpAL146A strains failed to inhibit V-ATPase trafficking to Mtb phagosomes. Yet, the mycobacterial phagosome is not an isolated compartment; rather, it interacts extensively with the early endosomes to acquire nutrients, such as iron-bound transferrin, to allow Mtb survival within the host cell [170]. In fact, the Mtb cell wall glycolipid, phosphatidylinositol mannoside (PIM), is capable of stimulating homotypic fusion of early endosomes in an ATP-, cytosol-, and Nethylmaleimide sensitive factor-dependent manner [171]. Recent advances in the study of membrane trafficking in S. cerevisiae show that vesicle fusion in the endocytic pathway depends on two membrane tethering protein complexes, CORVET (class C core vacuole/endosome tethering) and HOPS (homotypic fusion and protein sorting) [88, 172]. Class C VPS complex serves as the core of both CORVET and HOPS complexes through reversible association with CORVET-specific (VPS3 and VPS8) and HOPS-specific (VPS39 and VPS41) accessory subunits, which mediate early to late endosome fusion events and 	
    89  fusion with lysosomes, respectively [reviewed in 173]. Although the CORVET complex has not yet been identified in mammalian cells, it likely exists and has the same functions given the high similarity in the transport machinery between yeast and mammals. On the other hand, the mammalian HOPS complex has been identified and is known to play the same role as the S. cerevisiae homologs in mediating the conversion of Rab5 to Rab7 and tethering the fusing membranes. Our results indicate that V-ATPase may specifically interact with the HOPS complex during phagosome-lysosome fusion (Figure 26). Therefore, by binding to the V-ATPase, PtpA could specifically localize to the phagosomelysosome interface while CORVET-mediated early phagosome-endosome fusion process will remain intact. This specific localization mechanism would then allow PtpA to distinguish HOPS from CORVET, thereby specifically excluding VATPase to inhibit mycobacterial phagosome acidification. In fact, a link between the V-ATPase and HOPS complexes was suggested by a previous study demonstrating that HOPS specific subunit VPS41 failed to function in yeast which had mutations in the V-ATPase complex [174]. This indicates that in macrophages, the HOPS-specific subunits, VPS39 and VPS41, might be direct effectors of the V-ATPase complex during phagosome-lysosome fusion (Figure 26), and that the CORVET complex might be recruited to the phagocytic pathway through a different mechanism. As the HOPS-specific subunits are known to interact with activated Rab7 during endosome maturation [58], it is likely that Rab7 is the upstream activator of V-ATPase association with the Class C VPS complex. Further experiments will be needed to fully characterize these host  	
    90  cellular pathways in the context of mycobacterial infection. Nonetheless, our study clearly demonstrated that the absence of V-ATPase and the lack of mycobacterial phagosome acidification are directly attributed to the Mtb protein tyrosine phosphatase PtpA. 4.2 PtpA is a substrate for the protein-tyrosine kinase PtkA As protein phosphorylation plays a fundamental role in a wide range of cellular processes, it is not surprising that a large number of structurally distinct protein kinases have evolved [175, 176]. The HAD enzyme super-family comprises enzymes with hydrolytic activities, but kinase activities were not reported as of yet. We provided evidence that while Mtb PtkA was annotated as a HAD superfamily member it exhibited an authentic protein-tyrosine kinase activity and its substrate was PtpA. Although earlier evidence indicated that Mtb possesses protein-tyrosine phosphorylation activity [158], post-genomic bioinformatics analysis failed to identify a corresponding protein-tyrosine kinase. Extensive in silico analysis revealed that PtkA does not possess any signature or pattern related to tyrosine kinases. For instance, the Walker A and B motifs [163], which are present in bacterial autophosphorylating tyrosine kinases, are absent in PtkA. Our findings are in line with recent data describing novel protein-tyrosine kinases and exemplified by the MasK protein from Myxococcus xanthus [177] and WaaP from Pseudomonas aeruginosa [178], which do not contain conserved pattern homologous to classical tyrosine kinases [163] but have been reported as selfphosphorylated tyrosine kinases. Therefore, we conclude that protein tyrosine 	
    91  kinase activity does not necessarily require the canonical protein-tyrosine kinase motifs. We carried out several independent experiments to demonstrate that PtkA possesses tyrosine kinase activity. These include the incorporation of  32  P on  tyrosine residues, detection of the radioactive phosphotyrosine residues in a TLC autoradiogram, and mutational analysis of PtkA. These experiments showed that the amino acid Y262 is the autophosphorylated tyrosine residue on PtkA. This finding indicates that Y146 and Y150 are not involved in the catalytic activity of PtkA. In an attempt to determine other residues involved in the catalytic activity of PtkA, we carried out mutational analyses on all three lysine residues in this protein. Mutations of these residues resulted in no incorporation of 32P, indicating that these residues could play a role in the enzymatic mechanism. Since individual mutations of each of the three lysines reduced the enzymatic activity to nearly undetectable levels, we, at this stage are not able to determine which one of them is providing the hydrogen bonds to both the phosphate and the nucleophilic [179]. Interestingly, we found that although the mutant K184M was not phosphorylated, it binds PtpA similarly as the wild-type protein. The inability of all three lysine mutants to incorporate  32  P is in agreement with previous  observations that two lysines are required for ATP binding in tyrosine kinase Ptk of Acinetobacter johnsonii [163]. The core mechanism of HAD proteins is the transfer of a phosphoryl group from a specific phosphate ester to an active site aspartate, and then to a water  	
    92  molecule. The nucleophilic mechanism in HAD super-family members is driven by the first aspartate in the DXD motif. To demonstrate that this residue is involved in the catalytic activity of PtkA, D85 residue was replaced by alanine. The resulting mutant protein did not show any kinase activity indicating that D85 residue is indeed involved in the catalytic mechanism of the enzyme. These findings further supports the essentiality of the DXD motif for the catalytic activity of the HAD super-family members [161] and the inclusion of PtkA in this superfamily. Our findings that PtpA is the cognate substrate of PtkA are in agreement with the observations that PtpA is secreted and acts on host proteins. Therefore, it seems unlikely that its substrate would be present within the bacilli. Although preliminary reports have shown that PtkA is upregulated in the SCID mouse model of Mtb infection [180] and in murine macrophages [181], the role of PtpA phosphorylation by PtkA still needs to be defined. Mutational analyses of PtpA showed that PtkA could phosphorylate two adjacent tyrosine residues of PtpA. This result is in concordance with the findings that in human low-molecularweight protein tyrosine phosphatase B, two adjacent tyrosine residues are phosphorylated by a protein-tyrosine kinase [182]. Thus, we hypothesize that phosphorylation of PtpA is needed to retain its activity within the host macrophages. Alternatively, phosphorylation might be needed for the PtpA secretion process from Mtb, through the phagosome membrane to the host cytosol. Nevertheless, due to the discrepancy between the molecular mass of PtpA, (about 18 kDa) and the reported tyrosine phosphorylated 55 kDa protein in  	
    93  Mtb [158], we do not rule out broader activity for PtkA depending on differential environmental signals. It is very likely that PtkA has multiple substrates within Mtb and controls a larger network of signal cascades. In fact, a recent study on Mtb STPKs phosphorylation motifs found that PtkA is phosphorylated on Ser/Thr residues in vivo when Mtb is exposed to peroxide stress, indicating another level of regulation for PtkA tyrosine kinase activity [183]. Since peroxide stress is a condition that Mtb encounters within the hostile host environment, PtkA activity might be controlled by a peroxide-sensing STPK during entry into the host macrophage. Therefore, it is tempting to speculate that this hypothetical signaling mechanism might allow Mtb to sense the phagosomal environment within the macrophage, thereby, activating or upregulating either PtpA’s activity or secretion to halt the phagosome maturation process. The exact relationship between Mtb STPK control of PtkA and PtkA phosphorylation of PtpA will require further investigation. Nevertheless, we have already demonstrated that PtpA and its secretion are essential to progression of Mtb infection [32]. This together with the large industrial know-how, and small molecule libraries of kinase inhibitors, raised the interesting notion that PtkA might yet be another attractive target for drug development against this notorious disease. To conclude, our data provide evidence that: (a) PtkA is a new tyrosine kinase belonging to the HAD super-family, (b) PtkA is able to use either ATP or GTP as phosphate donors, (c) PtkA neither possesses kinase signatures nor resembles a protein-tyrosine kinase, but is autophosphorylated in tyrosine residues, and (d) PtkA is able to autophosphorylate and transfer the incorporated  	
    94  phosphate group to the tyrosine phosphatase PtpA, although the role of PtpA phosphorylation in Mtb physiology remains to be determined.  4.3 Inhibitors against PtpA as novel antituberculosis drug The essentiality of PtpA in the intracellular survival and pathogenesis of Mtb in various infection models has spurred great interest in exploiting PtpA as a novel antitubercular drug target [111]. A major challenge for the development of Mtb PtpA inhibitors is to demonstrate efficacy in vivo due to the fact that PtpA is not essential for bacterial growth in in vitro. Therefore, the traditional high throughput screening approach for in vitro growth inhibition would not be effective for the discovery of PtpA inhibitors. Nevertheless, several programs have already developed selective inhibitors against PtpA. Platforms adopted from cancer research for the design of PTP inhibitors have been established, and compounds which can specifically inhibit PtpA enzymatic activity and reduce mycobacterial survival within host macrophages, have been successfully identified and characterized [106, 109, 110, 112, 113, 184]. Pioneering in the identification of PtpA inhibitory compounds, Manger et al. tested natural products such as stevastelins, roseophilins and prodigiosins, obtaining IC50 values between 8.8 µM and 28.7 µM [110]. However, as PtpA has a high sequence and structural similarity with human PTP, specificity is an issue with the identified compounds as they show inhibition against human PTPs including PTP1B and the dual-specificity phosphatase Cdc25A. More recently, using fragment-based discovery approach, where small chemical entities are 	
    95  screened for affinity and subsequently linked to generate specific inhibitors, a group of aryl difluoromethylphosphonic acid (DFMP) compounds were identified as potent inhibitors of PtpA with Ki value of 1.4 ± 0.3 µM [113]. This group of inhibitors, due to stringent design, was reported to have greater than 70-fold specificity for Mtb PtpA as compared to highly homologous human phosphatases. In another study, Chiaradia et al. [109] have recently characterized a family of synthetic chalcones, essential intermediate compounds in flavonoid biosynthesis in plants, as inhibitors of PtpA with IC50 values between 8.4 µM and 53.7 µM. When these compounds were analyzed for their efficacy in the THP-1 infection model, one inhibitor (named 4d), with minimal toxicity against the THP-1 macrophage-like cells, was able to reduce the bacillary load by 50% at 48 hours post infection and 77% at 96 hours post infection [111] (Figure 27). This was the first demonstration of the effectiveness of PtpA inhibitors as a potential antitubercular drug.  Figure 27. Mtb survival in infected THP-1 treated with chalcone inhibitors. Differentiated THP-1 macrophage-like cells were infected with the H37Rv Mtb 	
    96  strain and treated with 10 µM of the indicated inhibitor at 0 hr. A second and third dose were added at 24 hr and 48 hr, respectively. Survival of intracellular Mtb (CFU) was determined by plating on 7H10 agar. DMSO is used as a negative control (-). Data are expressed as mean CFU for triplicate wells with standard error. Our findings in this dissertation that PtpA binds to the host macrophage VATPase subunit H represents another avenue for targeting and inhibiting PtpA activity within the host cell. Since PtpA binding to the V-ATPase machinery precedes the dephosphorylation of VPS33B, targeting PtpA’s binding capability might prevent its localization to the phagosome-lysosome fusion interface and completely inactivate PtpA functions. The identification of the binding site for subunit H on PtpA might further aid the development of compounds that can specifically block the interaction with the host protein complex. It is interesting to note that the V-ATPase subunit H is also a binding target for HIV Nef, an accessory protein that is required for viral infectivity and pathogenicity [78]. HIV Nef plays a key role in downregulating the surface expression of CD4 and MHC-I on T cells by advancing the endocytosis and degradation of these cell surface proteins [185, 186]. Nef-dependent reduction of surface MHC-I protects HIV-infected primary T cells from recognition and killing by HIV-specific cytotoxic T lymphocytes whereas downregulation of CD4 prevents superinfection and allows optimal production of viral particles [187, 188]. It has been proposed that Nef, which binds to the cytoplasmic tail of CD4, drives internalization and degradation of CD4 by interacting with V-ATPase subunit H, which, in turn, binds to and recruits medium chain (µ2) of the adaptor protein complex 2 (AP-2) [78, 189]. AP-2 is responsible for the recruitment of clathrin  	
    97  proteins, driving the formation of clathrin-coated pit and the subsequent endocytosis of surface CD4. Therefore, the importance of the V-ATPase subunit H in both Mtb and HIV infections indicates that a single inhibitory compound that can prevent both PtpA and Nef recognition of subunit H might be capable of limiting both Mtb and HIV proliferation within the co-infected human host. However, toxicity could be an issue as in order for such an inhibitory compound to impede both PtpA and Nef binding, it would most likely have to target the host V-ATPase complex, which is essential for numerous human physiological processes. Nevertheless, continued advances in drug discovery approaches might allow the development of such an inhibitory compound with low toxicity profile. This would then be a potential novel therapeutic against the current twin epidemic of Mtb and HIV co-infection, “killing two birds with one stone”. 4.4 Future directions The work presented in this thesis described the role of PtpA in the arrest of phagosome maturation during Mtb entry into the host macrophage. PtpAmediated specific exclusion of the V-ATPase proton pump from the mycobacterial phagosome explains the mechanism behind the inhibition of phagosome acidification during macrophage infection with Mtb. PtpA is, therefore, is essential for the pathogenesis of TB, allowing Mtb to subvert macrophage antimicrobial mechanisms and persist within the human host. Furthermore, within Mtb, PtpA is a substrate for the protein tyrosine kinase PtkA, indicating a novel signal transduction pathway for the regulation of PtpA function. Based on the work thus far, the PtpA project has the potential to branch into several research 	
    98  directions, such as the global host macrophage phosphoproteome response to PtpA and the physiological relevance of PtkA phosphorylation of PtpA. To investigate the global phosphoproteome response of the host macrophages to PtpA during Mtb infection, a quantitative proteomic analysis approach based upon the iTRAQ (isobaric tag for relative and absolute quantitation) methodology can be performed to fully reveal downstream host signaling cascades affected by PtpA. From our previous study, we found that VPS33B possesses protein-tyrosine kinase activity, indicating that it might regulate multiple downstream signaling pathways that are involved in membrane trafficking and fusion [32]. Therefore, PtpA dephosphorylation of VPS33B suggests that PtpA can disrupt multiple host proteins in order to perturb phagosome maturation and to promote Mtb intracellular survival. Human macrophages will be infected with the parental H37Rv wild type (WT) and the ΔptpA Mtb strains, and the phosphorylation response of the infected macrophages to the different strains will be compared. The differential phosphorylation profiles of the host cells should then reveal the complete network of host signaling pathways that are modulated by PtpA, thereby, providing a complete overview of the physiological functions of PtpA in relation to Mtb pathogenicity. It will also be interesting to examine the effect of PtpA on the recruitment of proteins to the phagosome using the same iTRAQ approach. However, a hurdle of this analysis would be to isolate and obtain highly pure samples of Mtb-containing phagosomes from infected macrophages.  	
    99  To investigate the role of PtkA and its phosphorylation of PtpA in Mtb pathophysiology, we have constructed a ptkA deletion mutant in the H37Rv background. The in vitro growth characteristics and in vivo survival in various infection models, including THP-1 macrophage-like cells, can be examined. If PtkA is involved in regulating PtpA activity or secretion, we should observe phenotypes similar to those seen with the ΔptpA mutant strain. PtpA has also been demonstrated to be secreted into the culture media in vitro [105]. Therefore, to investigate whether PtkA controls secretion of PtpA, we can probe for the presence of PtpA in the culture filtrate proteins of the ΔptkA mutant strain as an initial study. Furthermore, as mentioned above, a previous study has shown that PtkA might be phosphorylated by Mtb STPK under peroxide stress conditions in vitro [183], indicating an upstream regulatory control of PtkA activity. It would be interesting to investigate whether PtkA can indeed be a substrate for one of the eleven STPKs in Mtb. Since the biological functions of Mtb STPKs, such as regulation of cell wall biosynthesis, dormancy and cell division, have been extensively studied [138, 190-196], identification of such a signal transduction pathway might provide further insight into the physiological role of PtkA within Mtb. It is possible that PtkA might, in addition to PtpA, regulate a larger network of Mtb proteins involved in host-pathogen interaction and adaptation to the hostile host environment. Lastly, within the ptpA operon, Rv2235 encodes a transmembrane protein annotated as conserved hypothetical protein. An effort to comprehensively identify all genes required for Mtb growth using the transposon site hybridization  	
    100  (TRaSH) technique predicted that Rv2235 is essential for Mtb growth [197]. However, the genomic location and transmembrane feature of Rv2235 indicates that this protein might also play a role in the regulation of PtpA secretion. It will be interesting to elucidate and characterize the function of the protein encoded by Rv2235.  	
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    Hunter, T. (1987) A thousand and one protein kinases. Cell 50, 823-829  120  177. Thomasson, B., Link, J., Stassinopoulos, A.G., Burke, N., Plamann, L., and Hartzell, P.L. (2002) The GTPase, MglA, interacts with a tyrosine kinase to control type-IV pili-mediated motility of Myxococcus xanthus. Mol Microbiol 46, 1399-1413 178. Zhao, X. and Lam, J.S. (2002) WaaP pf Pseudomonas aeruginosa is a novel eukaryotic type protein-tyrosine kinase as well as a sugar kinase essential for the biosynthesis of core lipopolysaccharide. J Biol Chem 277, 4722-4730 179. Morais, M.C., Zhang, W., Baker, A.S., Zhang, G., Dunaway-Mariano, D., and Allen, K.N. (2000) The crystal structure of bacillus cereus phosphonoacetaldehyde hydrolase: insight into catalysis of phosphorus bond cleavage and catalytic diversification within the HAD enzyme superfamily. Biochemistry 39, 10385-10396 180. Talaat, A.M., Lyons, R., Howard, S.T., and Johnston, S.A. (2004) The temporal expression profile of Mycobacterium tuberculosis infection in mice. Proc Natl Acad Sci U S A 101, 4602-4607 181. Srivastava, V., Rouanet, C., Srivastava, R., Ramalingam, B., Locht, C., and Srivastava, B.S. (2007) Macrophage-specific Mycobacterium tuberculosis genes: identification by green fluorescent protein and kanamycin resistance selection. Microbiology 153, 659-666 182. Tailor, P., Gilman, J., Williams, S., Couture, C., and Mustelin, T. (1997) Regulation of the low molecular weight phosphotyrosine phosphatase by phosphorylation at tyrosines 131 and 132. J Biol Chem 272, 5371-5374 183. Prisic, S., Dankwa, S., Schwartz, D., Chou, M.F., Locasale, J.W., Kang, C.M., Bemis, G., Church, G.M., Steen, H., and Husson, R.N. (2010) Extensive phosphorylation with overlapping specificity by Mycobacterium tuberculosis serine/threonine protein kinases. Proc Natl Acad Sci U S A 107, 7521-7526 184. Noren-Muller, A., Wilk, W., Saxena, K., Schwalbe, H., Kaiser, M., and Waldmann, H. (2008) Discovery of a new class of inhibitors of Mycobacterium tuberculosis protein tyrosine phosphatase B by biology-oriented synthesis. Angew Chem Int Ed Engl 47, 5973-5977 185. Benson, R.E., Sanfridson, A., Ottinger, J.S., Doyle, C., and Cullen, B.R. (1993) Downregulation of cell-surface CD4 expression by simian 	
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    123  APPENDIX A: SUPPLEMENTAL FIGURES FOR SECTION 3.1  	
    Figure 28. In vitro analysis of subunit H and PtpA interaction. (A) Recombinant PtpA pulled down V-ATPase subunit H from THP-1 cell lysate. Recombinant His-tagged PtpA proteins were used as a bait to pull down target proteins from THP-1 human macrophages lysates. Lane 1: THP-1 cell lysate alone. Lane 2: THP-1 cell lysate incubated with His-tagged wild type PtpA protein. PtpA pulled down V-ATPase subunit H (blue circle). MALDI-TOF mass spectrometry was used to analyze the captured proteins (1). (B) Phosphorylated subunit H is not a catalytic substrate of PtpA. (Top Panel) PhosphorImage of subunit H recombinant proteins phosphorylated with γ[32P]ATP and incubated with increasing concentration of PtpA recombinant proteins. Lane 1. Subunit H 	
    124  alone. Lane 2. PtpA alone. Lane 3. Subunit H and PtpA (1:1 molar ratio). Lane 4. Subunit H and PtpA (1:2 molar ratio). Lane 5. Subunit H and PtpA (1:3 molar ratio). (Bottom Panel) Silver stained SDS-PAGE of the corresponding gel. (C) Phosphoamino acid analysis shows subunit H is Thr-phosphorylated. Phosphoamino acid analysis was performed as described previously (1). Retention factor (Rf): Phospho-serine (P-Ser), 0.44; Phospho-threonine (P-Thr), 0.49; Phospho-tyrosine (P-Tyr), 0.55; Subunit H, 0.50.  	
    125  Figure 29. PtpAL146A is a phosphatase-active mutant defective in binding subunit H. (A) Site-directed mutagenesis of di-leucine motifs and leucine residues in PtpA. Highlighted amino acids are the residues mutated and tested. L146A mutation creates a catalytically active mutant PtpA that does not interact with subunit H of the V-ATPase. (B) PtpAL146A remains catalytically active with similar kinetics as the wild type enzyme despite point mutation on its C-terminal alpha helix. p-nitrophenyl phosphate (pNPP) is used as a chromogenic substrate to detect phosphatase activity of PtpA. The reactions were analyzed in a spectrophotometer for absorbance at 450 nm.  	
    126  Figure 30. Western blot analysis of PtpA expression in complemented ΔptpA strain and in vitro PtpA secretion using α-PtpA antibodies. (A) A weak band corresponding to PtpA expression is observed for the parental H37Rv as expected. ΔptpA strains complemented with wild type ptpA (COMP), ptpAD126A or ptpAL146A shows clear expression of PtpA proteins. Recombinant PtpA was used as a positive control. No expression was observed for the ΔptpA strain. (B) Covalent labeling of the bacteria with pHrodo and Alexa-Fluor 488 does not affect PtpA secretion as shown by the prominent bands corresponding to Histagged PtpA in the culture filtrate. Soluble lysates from the bacteria were analyzed to ensure expression of His-tagged PtpA. Recombinant PtpA was used as a positive marker control.  	
    127  Figure 31. Calibration of THP-1 phagosomal pH during Mtb infection with FACS. (A) Calibration curve of phagosomal pH in THP-1 macrophage-like cells infected with Mtb dual-labeled with pHrodo and Alexa Fluor 488. The fluorescence intensity ratios of the fluorescence probes were calculated and plotted for each pH. (B) Overlaid FACS histograms of the pHrodo fluorescence intensities of Mtb phagosomes at pre-calibrated pH (5.0 - 8.0). (C-F) Overlaid FACS histograms of pHrodo fluorescence intensities of THP-1 infected with the 	
    128  indicated Mtb strains. (C) Parental H37Rv. (D) ΔptpA complemented with the wild type ptpA gene. (E) ΔptpA complemented with the mutant ptpAD126A gene. (F) ΔptpA complemented with the mutant ptpAL146A gene.  	
    129  Figure 32. Measurement of phagosomal pH in transfected THP-1 infected with E. coli. (A) Expression of GFP-tagged wild type and mutant PtpA proteins were assessed by measurement of GFP fluorescence (Emission 535 nm) in transfected THP-1 macrophage-like cells. Similar levels of expression were found for the wild type and mutant proteins. Untransfected THP-1 serves as a negative control, and THP-1 transfected with the pEGFP vector was used as a positive control. (B) Phagocytosis of E. coli labeled with Alexa Fluor 350 by THP-1 was not affected by transfection and expression of GFP-tagged wild type and mutant PtpA proteins. There was no difference in the level of phagocytosis of THP-1 transfected cells with untransfected cells as shown by the similar levels of Alexa Fluor 350 fluorescence (Emission 460 nm). (C) Calibration of THP-1 phagosomal pH during E. coli infection with spectrofluorometry. The fluorescence ratios of pHrodo (Emission 620 nm) and Alexa Fluor 350 (Emission 460 nm) were plotted for each pH to generate the calibration curve of phagosomal pH in THP-1 	
    130  macrophage-like cells infected with E. coli dual-labeled with the pH-sensitive pHrodo and pH-insensitive Alexa Fluor 350 dyes. (D) Separate digital confocal microscopy fluorescent images of transfected THP-1 macrophage-like cells infected with dual-labeled E. coli. Green: Expression of GFP or GFP-tagged PtpA constructs. Red: pHrodo fluorescence in acidified phagosomes. Blue: Alexa Fluor 350 labeled E. coli. Localization of GFP-PtpA to the phagosomes can be observed.  	
    131  	
   Figure 33. Analysis of antibodies specificity. (A) FACS analysis of the ability of indicated antibodies to non-specifically recognize extracellular components of Mtb. None of the tested antibodies showed significant binding to the bacteria. Rabbit anti-LqpH (19 kDa lipoprotein that is known to be present on the Mtb cell 	
    132  wall) was used as a positive control. (B) Western blot analysis of antibodies specificity with soluble lysates from E. coli DH5α, Mtb H37Rv, THP-1, THP-1 infected with E. coli DH5α (THP-1α) and THP-1 infected with Mtb H37Rv (THP1Rv). 25 µg of each lysate was resolved on SDS-PAGE and probed with the indicated antibodies. The tested antibodies showed specific detection of the target proteins with minimal non-specific background signal.  	
    133  Figure 34. Immunostaining control experiment with Mtb-infected THP-1. Immunostaining control experiment was performed to examine potential crossreactivity among the secondary antibodies. Rabbit anti-VPS33B (Rα-VPS33B) and Mouse anti-subunit H (Mα-H) were used as primary antibodies. Texas Redconjugated goat-anti rabbit IgG and Alexa-Fluor 488-conjugated goat-anti mouse IgG were used as secondary antibodes. Fluorescent signal was only detected when the correct pair of primary and secondary antibodies was used. The secondary antibodies do not react with primary antibodies from another species, demonstrating the specificity of the secondary antibodies for double immunofluorescence staining.  	
    134  Figure 35. Immunostaining control experiment with E. coli-infected THP-1. Immunostaining control experiment was performed to examine potential crossreactivity among the secondary antibodies. Rabbit anti-VPS33B (Rα-VPS33B) and Mouse anti-subunit H (Mα-H) were used as primary antibodies. Texas Redconjugated goat-anti rabbit IgG and Alexa-Fluor 488-conjugated goat-anti mouse IgG were used as secondary antibodes. Fluorescent signal was only detected when the correct pair of primary and secondary antibodies was used. The secondary antibodies do not react with primary antibodies from another species, demonstrating the specificity of the secondary antibodies for double immunofluorescence staining.  	
    135  

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