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Molecular analysis of Mycobacterium tuberculosis infection of human macrophages : the role of Protein… Poirier, Valérie 2015

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Molecular Analysis of Mycobacterium tuberculosis Infection of Human Macrophages:  The Role of Protein Tyrosine Phosphatase A  by Valérie Poirier  B.Sc., Bishop’s University, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Experimental Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  February 2015  © Valérie Poirier, 2015 ii  Abstract  To survive in the human host macrophage, Mycobacterium tuberculosis, the etiological agent of Tuberculosis, infects human macrophages and inhibits two key cellular events required for the elimination of invading organisms; phagosome acidification and fusion with lysosomes. This is partially dependent on Protein tyrosine phosphatase A (PtpA) secreted into the cytosol of the macrophage. PtpA first disrupts phagosome acidification by binding to subunit H of the proton pump, and second, inhibits phagosome-lysosome fusion by dephosphorylating and inactivating the membrane fusion regulator, hVPS33B. The ability of M. tuberculosis to actively interfere with host trafficking events allows this pathogen to replicate and persist inside the macrophage and prevent antigen presentation required to initiate an adaptive immune response.   In this work, we explored the global macrophage’s response to infection emphasizing on PtpA’s role in this process. We analyzed the macrophage global proteomic responses and focused on the activity of signalling pathways by determining the phosphorylation status of host proteins upon infection with M. tuberculosis strains.    We found that PtpA affects the macrophage’s response by modulating various proteins involved in RNA metabolism, immunity and defence, and cellular respiration pathways. We further show that PtpA promotes M. tuberculosis survival by dephosphorylating the host kinase GSK3α on amino acid Y279 leading to inhibition of GSK3α and arrest of macrophage apoptosis. GSK3α has pro- and anti-apoptotic activities and dephosphorylation of Y279 inhibits its ability to initiate apoptosis. In this regard, activation of the host apoptosis executioner, caspase-3, is iii  blocked in M. tuberculosis-infected macrophages compared to cells infected with ΔptpA mutant strain.    Taken together, these findings reveal one of the long-sought effectors behind the inhibition of apoptosis of the host by virulent M. tuberculosis. Moreover, we are the first to simultaneously determine proteome-wide protein expression levels of human macrophages infected with M. tuberculosis and outline molecular signatures of the global and PtpA-dependent proteomic patterns of macrophages during infection.    We have now established that PtpA significantly contributes to successful infection and survival of M. tuberculosis inside the macrophage. Understanding the mechanism of action of PtpA during M. tuberculosis infection may lead to the development of novel anti-mycobacterial drugs.       iv  Preface Parts of this thesis have been published in the peer-reviewed journals listed below:  1. Poirier, V., and Av-Gay, Y. (2012). Mycobacterium tuberculosis modulators of macrophage     cellular events. Microbes Infect. 14, 1211-1219      In this work, I was responsible for writing a review article on the current knowledge of     Mycobacterium tuberculosis effectors for which a cognate substrate, a cellular partner or a     signalling pathway have been experimentally identified within the human host. Y. Av-Gay     participated in writing and editing the manuscript. This work is located in Sections 1.2.1 and      1.2.2.  2. Poirier, V., Bach, H., and Av-Gay, Y. (2014) Mycobacterium tuberculosis promotes anti     apoptotic activity of the macrophage by PtpA-dependent dephosphorylation of host GSK3α.       J Biol Chem. 289, 29376-29385      In this study, I designed and performed all experiments except those of work related to Figure     11, which was performed by H. Bach. I wrote the manuscript. Y. Av-Gay contributed to the     designing of the study and writing the manuscript. A version of this published work is located      in Sections 3.1 and 4.1.  3. A version of Chapter 3 and 4 (Sections 3.2 and 4.2) will be submitted for publication in the     near future. In these sections, I designed and performed all experiments. G. Av-Gay performed v      all statistical analyses. I wrote the manuscript except for Section 2.14, which was written     by G. Av-Gay. Y. Av-Gay participated in designing the study and writing the manuscript.   The work presented in this thesis was conducted in accordance with the University of British Columbia Research Policies and Procedures, the University Biosafety Committee, the Public Health Agency of Canada guidelines, and with approval from the University of British Columbia Office of Research Services. Biohazard Approval Certificate: B10-0112. Work performed with radioactive materials was approved by the University of British Columbia Advisory Committee on Radioisotopes and Radiation Hazards. Radioisotope License: MEDI-3375-18.     vi  Table of Contents Abstract ........................................................................................................................................... ii Preface............................................................................................................................................ iv Table of Contents ........................................................................................................................... vi List of Tables ...................................................................................................................................x List of Figures ................................................................................................................................ xi List of Abbreviations ................................................................................................................... xiii Acknowledgements ........................................................................................................................xx Dedication ................................................................................................................................... xxii Chapter 1: Introduction ................................................................................................................... 1 1.1 Tuberculosis ...................................................................................................................... 1 1.1.1 Epidemiology of Human Tuberculosis ...................................................................... 1 1.1.2 Origin of Human Tuberculosis .................................................................................. 5 1.1.3 Mycobacterium tuberculosis ...................................................................................... 6 1.2 Phagocytosis ..................................................................................................................... 8 1.2.1 Phagosome Formation and Maturation ...................................................................... 8 1.2.2 Inhibition of Phagosome Maturation by M. tuberculosis ........................................ 11 1.2.2.1 Biological Effectors Mediating M. tuberculosis Survival in Macrophages ... 13 1.2.2.1.1 LAM ........................................................................................................ 14 1.2.2.1.2 SapM ....................................................................................................... 16 1.2.3 Inhibition of Antigen Processing and Presentation ................................................. 18 1.2.3.1 19 kDa Lipoprotein ........................................................................................ 18 1.3 PtpA ................................................................................................................................ 20 vii  1.3.1 Overview ................................................................................................................. 20 1.3.2 VPS33B and the HOPS Complex ............................................................................ 21 1.3.3 Subunit H of the V-ATPase Pump .......................................................................... 23 1.4 Host Responses to M. tuberculosis Infection ................................................................. 24 1.4.1 Apoptosis Versus Necrosis ...................................................................................... 24 1.4.2 Apoptosis During Infection with Virulent Versus Avirulent Mycobacterium Strains ................................................................................................................................. 27 1.5 Techniques for Identifying Intracellular Substrates for M. tuberculosis Effectors During Infection .......................................................................................................................... 30 1.5.1 Phosphoproteome Analysis ..................................................................................... 30 1.5.2 iTRAQ ..................................................................................................................... 31 1.6 Aims of the Study ........................................................................................................... 34 Chapter 2: Methods and Materials ................................................................................................ 36 2.1 Reagents and Chemicals ................................................................................................. 36 2.1.1 Commercial Reagents .............................................................................................. 36 2.1.2 Antibodies ................................................................................................................ 36 2.2 Bacteria ........................................................................................................................... 37 2.2.1 Strain Maintenance .................................................................................................. 37 2.2.2 Bacterial Preparation for Infection .......................................................................... 37 2.2.3 In Vivo Bacterial Uptake by Macrophage ................................................................ 37 2.3 Cell Culture ..................................................................................................................... 38 2.3.1 Tissue Culture Maintenance and Differentiation ..................................................... 38 2.3.2 Macrophage Infection .............................................................................................. 38 viii  2.3.3 Macrophage Cellular Extraction for Western Blot Analysis ................................... 39 2.3.4 Macrophage Cellular Extraction for iTRAQ Analysis ............................................ 39 2.4 DNA Manipulation ......................................................................................................... 40 2.4.1 RNA Extraction and cDNA Synthesis ..................................................................... 40 2.4.2 DNA Cloning ........................................................................................................... 40 2.4.3 Quantitative Polymerase Chain Reaction (qPCR) ................................................... 41 2.5 Protein Expression and Purification ............................................................................... 43 2.5.1 Expression of Recombinant Proteins in E. coli ....................................................... 43 2.5.2 Expression of Recombinant Proteins in M. smegmatis ........................................... 44 2.6 Phosphoproteome Analysis by KinetworksTM Phospho-Site Screen (KPSS) Assay...... 44 2.7 In Vitro Phosphatase Activity Assay .............................................................................. 45 2.8 In Vivo and In Vitro Western Blot Analysis ................................................................... 45 2.9 Determination of PtpA and GSK3α Dissociation Constant ............................................ 47 2.10 In Vitro Protein Kinase Assay ........................................................................................ 47 2.11 Radiometric Protein Kinase Assay ................................................................................. 48 2.12 Isobaric Tags for Related and Absolute Quantitation (iTRAQ) and LC-MS/MS  Analyses .......................................................................................................................... 48 2.13 Data Analysis and Interpretation .................................................................................... 49 2.14 Statistical Analysis .......................................................................................................... 49 Chapter 3: Results ......................................................................................................................... 51 3.1 M. tuberculosis Promotes Anti-Apoptotic Activity of the Macrophage by PtpA-Dependent Dephosphorylation of Host GSK3α ............................................................. 51 3.1.1 Introduction ............................................................................................................. 51 ix  3.1.2 PtpA Interferes with Multiple Host Macrophage Signalling Pathways .................. 52 3.1.3 KinetworksTM Analysis ............................................................................................ 53 3.1.4 M. tuberculosis PtpA Dephosphorylates the Human Kinase GSK3α ..................... 57 3.1.5 GSK3α Transcription Levels are not Decreased by PtpA ....................................... 57 3.1.6 PtpA Dephosphorylates GSK3α Under In Vivo Growth Conditions ....................... 59 3.1.7 PtpA Interferes with Host Macrophages Apoptosis Early During Infection ........... 63 3.1.8 M. tuberculosis Blocks Proteolytic Cleavage of Inactive Caspase-3 into Active Caspase-3 ................................................................................................................. 65 3.2 Global Proteomic Analyses of Macrophage Response to M. tuberculosis Infection ..... 67 3.2.1 Introduction ............................................................................................................. 67 3.2.2 Modulation of Host Protein Expression in Response to M. tuberculosis ................ 68 3.2.3 Identification of Macrophage Proteins Modulated by M. tuberculosis Infection .... 71 3.2.4 Validation of Proteomics Expression Levels ........................................................... 74 3.2.5 PtpA Effect on Macrophage Protein Expression Levels ......................................... 77 Chapter 4: Discussion ................................................................................................................... 80 4.1 M. tuberculosis PtpA Inhibits Host Apoptosis Early During Infection .......................... 80 4.2 M. tuberculosis Causes Concerted Up-Regulation and Random Down-Regulation of Host Macrophage Proteins .............................................................................................. 85 4.2.1 Host Proteins Targeted by M. tuberculosis ............................................................. 86 4.2.2 Host Proteins Targeted by M. tuberculosis PtpA .................................................... 92 4.3 Future Directions ............................................................................................................ 97 References ....................................................................................................................................101  x  List of Tables Table 1. Oligonucleotides Used for DNA Cloning ....................................................................... 41 Table 2. Oligonucleotides Used for Quantitative PCR ................................................................. 42 Table 3. Identification of Proteins Dephosphorylated by the Addition of Recombinant PtpA to a Phosphorylated THP-1 Cellular Extract ....................................................................................... 53 Table 4. Phosphoproteome Analysis of Host Signalling Proteins Affected by PtpA ................... 56 Table 5. M. tuberculosis-Infected Macrophage Proteins Modulated One Standard Deviation Above or Below the Mean ............................................................................................................ 72 Table 6. ΔptpA M. tuberculosis-Infected Macrophage Proteins Modulated 1.8 Standard Deviations Above or Below the Mean .......................................................................................... 79               xi  List of Figures Figure 1. Diagnosis and Notification of Multidrug-Resistant TB (2012) ....................................... 3 Figure 2. Schematic Representation of the Mycobacterial Cell Wall ............................................. 7 Figure 3. Receptor and Signalling Interactions During Phagocytosis of Microbes ........................ 9 Figure 4. Stages of Phagosome Maturation Exemplified by E. coli Infection ............................. 11 Figure 5. Blockage of Phagosome Maturation by M. tuberculosis............................................... 13 Figure 6. Inhibition of Phagosome Maturation by M. tuberculosis Effectors .............................. 16 Figure 7. Subunit Arrangement of the HOPS Complex ............................................................... 22 Figure 8. Schematic Representation of V-ATPase Exclusion and Inhibition of Phagosome Acidification by PtpA ................................................................................................................... 24 Figure 9. Modes of Macrophage Cell Death Caused by Virulent and Avirulent Mycobacteria .. 29 Figure 10. Sample Processing and Analysis Workflow for Global Proteome Profiling with Quantification Using Mass Tag Labeling of Peptides .................................................................. 33 Figure 11. Phosphoproteomic Analysis of Host Macrophage Proteins Dephosphorylated by PtpA Revealed by 2D Gel Electrophoresis ............................................................................................ 52 Figure 12. Global Phosphoproteome Analysis of THP-1 Cells Infected with M. tuberculosis and ΔptpA M. tuberculosis................................................................................................................... 54 Figure 13. Transcriptional Levels of GSK3α Post-Infection ........................................................ 58 Figure 14. Western Blot Analyses of PtpA Dephosphorylation of GSK3α In Vivo ..................... 59 Figure 15. Western Blot Analyses of PtpA Dephosphorylation of GSK3α In Vitro .................... 60 Figure 16. Kinase Assays of PtpA Dephosphorylation of GSK3α In Vitro ................................. 61 Figure 17. PtpA Interacts with GSK3α In Vitro ........................................................................... 63 Figure 18. PtpA Reduces Trancriptional Levels of Caspase-3 Early in Infection ........................ 65 xii  Figure 19. M. tuberculosis Blocks Activation of Caspase-3 In Vivo ............................................ 66 Figure 20. Boxplots of Host Proteins Obtained from Infected Versus Uninfected Macrophages 69 Figure 21. M. tuberculosis-Modulation of Host Protein Expression ............................................ 70 Figure 22. Global Expression Level Scatterplot of Normalized Infected Versus Uninfected Log-Ratios ............................................................................................................................................ 73 Figure 23. Western Blot Analyses of M. tuberculosis-Dependent Host Protein Expression    Levels ............................................................................................................................................ 75 Figure 24. Transcriptional Levels of Five M. tuberculosis-Modulated Proteins .......................... 76 Figure 25. Boxplots of Host Proteins Obtained from Infected Versus Uninfected Macrophages 77 Figure 26. Global Expression Level Scatterplot of Normalized M. tuberculosis Versus ΔptpA M. tuberculosis Log-Ratios ................................................................................................................ 78  xiii  List of Abbreviations 2D   Two-Dimensional ABCB10  ATP-Binding Cassette Sub-Family B Member 1  ALPHAScreen Amplified Luminescent Proximity Homogeneous Assay Screen AraLAM   Arabinofuranosyl-Terminated LAM  ATP   Adenosine-5’-Triphosphate γ32P-ATP  [γ32P]-Adenosine-5’-Triphosphate  ATP5I   ATP Synthase Subunit ε BCG        Bacillus Calmette-Guérin BCL-2   B-Cell Lymphomal Leukaemia-2 BIRC6   IAP Repeat-Containing Protein-6 Brg1   Brahma-Related Gene-1 Ca2+   Calcium  CaM    Calmodulin   CaMKII   Ca2+/CaM Kinase II  cDNA   Complementary DNA CIITA   Class II, Major Histocompatibility Complex, Transactivator CIRBP   Cold-Inducible RNA-Binding Protein  CO   Complement (ΔptpA::ptpA M. tuberculosis) CO2   Carbon Dioxide COX7B  Cytochrome c Oxidase Subunit 7B CR3   Complement Receptor 3 CTSG   Cathepsin G xiv  DNA   Deoxyribonucleic Acid DOCK10  Dedicator of Cytokinesis Protein-10  DTT        Dithiothreitol ECL   Enhanced Chemiluminescence EDTA   Ethylenediaminetetraacetic Acid EEA1   Early Endosome Antigen-1  FACS       Fluorescence-Activated Cell Sorting FBS   Fetal Bovine Serum FcγR   Fc Gamma Receptor GAPDH  Glyceraldehyde 3-Phosphate Dehydrogenase GEF   Guanine Nucleotide Exchange Factor GSK3α    Glycogen Synthase Kinase-3 Alpha GSK3β    Glycogen Synthase Kinase-3 Beta GST    Glutathione S-Transferase GTP   Guanosine-5’-Triphosphate H2AFZ  Histone H2A  HAT    Histone Acetyl Transferase  HCl   Hydrogen Chloride HDAC   Histone Deacetylase  HEPES  4-(2-Hydroxyethyl)-1-Piperazineethanesulfonic Acid  His   Histidine HOPS       Homotypic Vacuole Fusion and Protein Sorting HPLC   High-Performance Liquid Chromatography xv  HRP   Horseradish Peroxidase  hVPS   Human Vacuolar Protein Sorting hVPS34  Human Vacuolar Protein Sorting Protein 34  hVPS33B  Human Vacuolar Protein Sorting Protein 33B ICAM1  Intercellular Adhesion Molecule-1 IFNγ   Interferon-Gamma IgG   Immunoglobulin G IL1β   Interleukin-1β IPTG      Isopropyl-β-D-Thio-Galactoside iTRAQ  Isobaric Tags for Related and Absolute Quantitation Kd   Dissociation Constant kDa   KiloDalton KO   Knockout (ΔptpA M. tuberculosis) LAM    Lipoarabinomannan  LAMP   Lysosomal Associated Membrane Protein LB   Luria-Bertani LC   Liquid Chromatography LLO   Listeriolysin O LpqH   19 kDa Lipoprotein  ManLAM   Mannosylated LAM  MAPK   Mitogen-Activated Protein Kinase  M. bovis  Mycobacterium bovis M. bovis BCG  Mycobacterium bovis Bacillus Calmette-Guérin xvi  MDR        Multidrug-Resistant MEK1/2  Mitogen-Activated Protein Kinase Kinase-1/2 MgCl2   Magnesium Chloride MHC-I   Major Histocompatibility Complex Class I MHC-II   Major Histocompatibility Complex Class II  MIF   Migration Inhibitory Factor M. kansasii  Mycobacterium kansasii   MMTS  S-Methyl Methanethiosulfonate  MnCl2   Manganese Chloride MOI        Multiplicity of Infection MOWSE  Molecular Weight Search mRNA   Messenger RNA MS/MS  Mass Spectrometry M. smegmatis  Mycobacterium smegmatis MTBC   M. tuberculosis Complex M. tuberculosis Mycobacterium tuberculosis MW   Molecular Weight NaCl   Sodium Chloride Na2HPO4  Disodium Hydrogen Phosphate Na3VO4  Sodium Orthovanadate NI   Uninfected Ni-NTA   Nickel-Nitriloacetic Acid  NSF   N-Ethylmaleimide-Sensitive Factor xvii  OADC        Oleic Acid Dextrose Catalase Complex PAMP   Pathogen-Associated Molecular Pattern PBS        Phosphate Buffered Saline PCD   Programmed Cell Death PCR   Polymerase Chain Reaction PH   Pleckstrin Homology  Pi   Inorganic Phosphate pI   Isoelectric Point PI   Phosphatidylinositol  PI3K   Phosphatidylinositide 3-Kinase PI3P    Phosphatidylinositol 3-Phosphate  PiLAM   Phosphoinositol-Capped LAM   PlcA   Phospholipase A PlcB   Phospholipase B PMA        Phorbol-12-Myristate-13-Acetate PMSF        Phenylmethanesulfonyl Fluoride pNP    p-Nitrophenol pNPP    p-Nitrophenyl Phosphate  PRPF31  U4/U6 Small Nuclear Ribonucleoprotein Prp31 PRR   Pattern Recognition Receptor PtpA        Protein tyrosine phosphatase A qPCR   Quantitative Polymerase Chain Reaction R   Gnu xviii  RCF   Relative Centrifugal Force RILP   Rab-Interacting Lysosomal Protein RNA   Ribonucleic Acid RPMI   Roswell Park Memorial Institute  SapM    Secreted Acid Phosphatase of M. tuberculosis   SARS2   Seryl-tRNA Synthetase-2  SD   Standard Deviation SDS   Sodium Dodecyl Sulphate SDS-PAGE  Sodium Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis Ser        Serine SNAP   Soluble NSF Attachment Protein SNARE  Soluble N-Ethylmaleimide-Sensitive Factor Attachment Protein Receptor  SNRPD1  Small Nuclear Ribonucleoprotein Sm D1  SOD2   Superoxide Dismutase-2 TB        Tuberculosis TCEP   Tris(2-Carboxyethyl)phosphine TEAB   Tetraethylammonium Bromide TGN   Trans-Golgi Network  Thr        Threonine TLR2   Toll-Like Receptor 2  TNF-α   Tumor Necrosis Factor-Alpha Tyr         Tyrosine V-ATPase  Vacuolar H+-ATPase xix  VPS         Vacuolar Protein Sorting VPS11   Vacuolar Protein Sorting 11 VPS16   Vacuolar Protein Sorting 16 VPS18   Vacuolar Protein Sorting 18 VPS33B  Vacuolar Protein Sorting 33B VPS34   Vacuolar Protein Sorting 34 VPS39   Vacuolar Protein Sorting 39 VPS41   Vacuolar Protein Sorting 41 WHO   World Health Organization WT         Wild-Type (M. tuberculosis) XDR        Extensively Drug-Resistant Y   Tyrosine xx  Acknowledgements  The first person that I wish to thank is my supervisor Dr. Yossef Av-Gay. The work presented in this thesis would not be possible had it not been from his strong and constant support to persevere through the hard work of my graduate studies. Dr. Av-Gay accepted me into his laboratory in 2008 as a Master's student, and throughout the entirety of my graduate studies Dr. Av-Gay presented me with numerous opportunities to learn and experience the various aspects of professional scientific research. These opportunities allowed me to meet leaders within the scientific community and benefit from participating in several international conferences. Dr. Av-Gay has inspired me to become a better scientist, to continue my work in this field and has made me more sensitive to a disease that ravages developing countries. I offer Dr. Av-Gay my enduring gratitude for his welcoming heart, expertise and for broadening my vision of science. He is a selfless human being and displayed nothing but patience and mentorship during my graduate studies.   I thank my committee members, Dr. Zakaria Hmama and Dr. Steven Pelech, for their support. They have provided me with helpful comments and questions over the years that helped me to question myself and my work on a fundamental level that is critical to scientific research.   I also thank all the past and present members of the Av-Gay laboratory, with special thanks to Mary Ko for all of her technical assistance, and Melissa Richard-Greenblatt for her enduring friendship over the years and her ability to create a lively environment in the laboratory. xxi   I would like to acknowledge financial support from Canadian Institute of Health Research (CIHR) and the University of British Columbia, and also, the British Columbia Center for Disease Control for providing us access to a BCL-3 laboratory.    Special thanks are owed to my loving family who helped me get through the emotional ups and downs of being so far away from home during my Ph.D studies.    Finally, I thank my husband Stefan Szary for supporting me morally and financially throughout my years of education. His support gave me the luxury of choosing my path. But most importantly, I deeply thank him for his unconditional care, love and advice that helped me become the woman I am today.   Dedication           xxii  Dedication “The Great White Plague, tuberculosis, is never done with its  insidious work; for numberless years it has been silently, continuously  and unrelentingly pursuing its destructive course, never remitting its  deadly work, always reaching out for new victims, and today it is  the most universal scourge of the human race.”    Otis, Edward O. The Great White Plague: Tuberculosis. New York: Thomas Y. Crowell & CO, 1909. Print, p. 2.   1  Chapter 1: Introduction 1.1 Tuberculosis 1.1.1 Epidemiology of Human Tuberculosis  Among all infectious diseases, Tuberculosis (TB) is second to HIV/AIDS as the greatest killer worldwide (WHO Report, 2014). An estimated one-third of the world’s population carries latent TB, implying that people have been exposed to M. tuberculosis, but show no signs of illness and cannot yet transmit the disease. TB is mostly prevalent in developing countries where over 95% of TB-related deaths occur. In 2013, an estimated 9 million people fell ill with TB and 1.5 million people died of the disease (WHO Report, 2014). Although the number of people falling ill due to TB is declining annually, TB is still the leading killer of people living with HIV representing one quarter of all deaths (WHO Report, 2014).    By the beginning of the 20th century, before the discovery of antibiotics,  TB was the principal cause of death of human beings and was crowned with the names; “Great Killer” and “Captain of all Deaths” (1). The work of Albert Calmette and Camille Guérin led, in 1921, to the introduction of the first TB vaccine, known as the BCG vaccine, which relies on the administration of live attenuated M. bovis bacilli, the causative agent of bovine TB. The use of the BCG vaccine then became the prevalent strategy to combat human TB in the world (1). Although the vaccine has proven effective against severe forms of childhood TB and miliary TB (an extrapulmonary form of TB characterized by wide dissemination) (2), its efficacy varies widely against pulmonary TB, the most prevalent form of the disease in adults (3).  2   The introduction of first-line anti-mycobacterial drugs (antibiotics such as streptomycin, and chemotherapy such as isoniazid, ethambutol and pyrazinamide) in the late 1940s revolutionized TB treatment, and caused astonishing decreases in mortality rates worldwide (4). However, resistance to these drugs was observed soon after their introduction (5) and by the end of the 1960s, the antibiotic rifampicin had to be added to the anti-mycobacterial drug cocktail. This combination of drugs appeared so promising that the “TB plague” was widely considered resolved (6). In 1969, the medic chief of the National Institute of Health declared that “it was time to close the books about infectious diseases” (4), which led to a decline in funding and interest in TB control programs (7). This false sense of security and ensuing lack of precautionary measures culminated in the reemergence of TB in the 1980s at the cost of many more millions of lives.    Besides the lack of rigor from health authorities, the reemergence of TB has been attributed to several factors such as; the HIV/AIDS pandemic, the aging of the world population, and the increase in world travel, including immigration, from countries of high TB prevalence to developed nations. Social inequalities that generate poverty, malnutrition and unsanitary living conditions are also major contributing factors (8). With the arrival of HIV, improperly administered two-drug therapies were not sufficient to fight TB. For the first time, the world was facing both mycobacterial resistance to antibiotics and a pathogen of unprecedented strength.    Multidrug-resistant TB (MDR-TB), primarily caused by the inappropriate use of antibiotics, is a strain of M. tuberculosis that does not respond to the two most effective first-line anti-TB drugs, isoniazid and rifampicin (WHO). MDR-TB is treatable and curable but requires 3  extensive chemotherapy with second-line drugs such as aminoglycosides, fluoroquinolones and thioamides. The emergence of MDR-TB is of great concern since second-line drugs are difficult to procure to developing countries, are more expensive, and more toxic than first-line drugs. Treatment can also take up to two years with severe adverse effects in patients (9). The MDR-TB strain is found in all countries surveyed (Figure 1).   World Health Organization. Global Tuberculosis Control 2012. Reprinted with permission from World Health Organization.  Figure 1. Diagnosis and Notification of Multidrug-Resistant TB (2012) Worldwide distribution of MDR-TB with the highest incidence rate in South Africa, the Russian Federation, India, and eastern European and central Asian countries with over 1001 cases identified (10).   4   Extensively drug-resistant TB (XDR-TB) is resistant to the two most powerful first-line anti-TB drugs (isoniazid and rifampicin), flouoroquinolones and at least one of the second-line anti-TB drugs (amikacin, kanamycin or capreomycin) (WHO). XDR-TB developed from MDR-TB as a result of the misuse or mismanagement of second-line drugs. Because of the resistance of XDR-TB to both first- and second-line anti-TB drugs, options for treatment are limited and the likelihood of success is much smaller than in patients with drug-susceptible TB or even with MDR-TB (WHO).    The combination of co-infection with HIV and the emergence of multidrug-resistant strains due to patient non-compliance pose a significant challenge to treating TB and increasing the survival rate. Despite intensive efforts to reduce the incidence of TB, no new anti-mycobacterial drugs had been deployed since the mid 1960s (11). However, in 2005, screening efforts led to the discovery of a novel molecule with anti-mycobacterial properties (12). This compound, now commercially known as bedaquiline, targets the M. tuberculosis proton pump Adenosine Triphosphate (ATP) Synthase and is reported to treat MDR-TB when administered in conjunction with second-line drugs (12). Due to the promising outcome of its use, bedaquiline was granted accelerated approval by the United States Food and Drug Administration in 2012 to treat MDR-TB (WHO). However, information regarding the safety of this new drug remains limited (WHO) and research focused on understanding the mechanism of action of M. tuberculosis should be a top priority in order to open new avenues for pharmaceutical interventions to treat TB.  5  1.1.2 Origin of Human Tuberculosis  In 1882, Dr. Robert Koch proved that the acid-fast bacillus M. tuberculosis causes TB (4). TB is caused by the highly virulent human pathogen Mycobacterium tuberculosis (M. tuberculosis). There is evidence of TB occurring in Ancient Egypt and the pre-Columbian societies of Central and South America as it has been identified in Egyptian and Peruvian mummies, a testament to the enduring threat it poses to every human society (13,14).     M. tuberculosis is a member of the Mycobacterium tuberculosis complex (MTBC). This complex includes the closely related species M. bovis, M. microti, M. africanum and M. canettii (15), which share 99.9% similarity at the nucleotide level in their genomes and identical 16S rRNA (16,17). Due to their extreme homogeneity, it is believed that the members of the MTBC are the clonal progeny of a single successful ancestor of the tubercle bacilli from East Africa approximately 3 million years ago (18). This indicates that our hominid ancestors may well have suffered from TB. It is surmised that clonal expansion took place only recently, following an evolutionary bottleneck that occurred 20,000 to 35,000 years ago, which gave rise to what we now know as the members of MTBC (17,19,20).      Despite their unusual high degree of conservation in housekeeping genes, and based on the assumption that they are all derived from a common ancestor, mycobacteria differ widely in phenotypic characteristics, including host preferences and pathogenicity (18). M. tuberculosis, M. africanum and M. canettii are exclusive human pathogens, whereas M. microti is a rodent pathogen, and M. bovis is a pathogen infecting a variety of hosts including cattle and man (19). These differences arise from genomic variations stemming from successive loss of DNA that 6  took place before the expansion of the M. tuberculosis clone (18,21), conferring the strains to a more or less pathogenic state and different host specificity (22). Furthermore, genomic studies show that M. tuberculosis is the direct descendant of tubercle bacilli that existed before clonal expansion (19) indicating that this common ancestor was already a human pathogen.   1.1.3 Mycobacterium tuberculosis  M. tuberculosis is a slow growing, aerobic facultative intracellular bacterium of the Actinomycetales order and Corynebacterineae suborder. It is an airborne pathogen that preferentially colonizes the respiratory system. In infected alveolar macrophages, M. tuberculosis normally goes into a dormant state as long as the immune system of the host predominates (23).   The genome of M. tuberculosis has a high G + C content of 65.6%, relative to E. coli (GC content ~ 50%), and contains approximately 4,000 genes. A genomic feature differentiating M. tuberculosis from other bacteria is its large amount of genes coding for fatty acid metabolic enzymes (about 250 genes in M. tuberculosis compared to 50 genes in E. coli, which has a similar genome size) (24). This unique characteristic of the tubercle bacillus confers it a complex cell envelope that is distinguished from the cell wall of Gram-positive and Gram-negative bacteria. Indeed, beyond the peptidoglycan, its envelope contains an additional layer exceptionally rich in unusual lipids, glycolipids and polysaccharides such as mycolic acids, mycocerosic acid, lipoarabinomannan, arabinogalactan and phenolthiocerol (Figure 2) (25,26). This thick lipid-rich coat contributes to host inflammatory reactions and forms a permeability barrier preventing entry of environmental solutes (27). This confers M. tuberculosis with high 7  intrinsic resistance to antibiotics and host antimicrobial mechanisms (28). Besides this highly hydrophobic cell wall, M. tuberculosis has gained resistance to antibiotics with mechanisms such as hydrolytic or drug-modifying enzymes and drug-efflux systems (25). Taken together, these unique features of M. tuberculosis contribute to “successful” infection and survival within the extremely hostile environment of the host macrophage.     Smith. Yale J Biol Med. 2011. 84 (4): 361-369. Reprinted with permission from Elsevier.  Figure 2. Schematic Representation of the Mycobacterial Cell Wall This illustration shows the cell wall of M. tuberculosis comprising of several layers of diverse cell wall lipid types ranging from simple fatty acids to complex long-chain lipids such as mycolic acids. The outer layers surround the traditional phospholipid bilayer of the cell.   8  1.2 Phagocytosis 1.2.1 Phagosome Formation and Maturation M. tuberculosis infects the human lung and is internalized by circulating alveolar macrophages; the cells that provide the first line of defence against mycobacterial invasion (29). Macrophages, along with neutrophils and dentritic cells, are professional phagocytes. They phagocytose large particles, including microorganisms, destroy them and, if required, illicit an immune response by promoting antigen presentation that activates an adaptive immune response (30).   Phagocytosis occurs via the simultaneous recognition of pathogen-associated molecules by pattern recognition receptors or by opsonins; host factors, such as antibodies or complement proteins, that attach to the pathogen membrane allowing it to be recognized by phagocytic receptors (Figure 3) (30,31). The best characterized phagocytic receptors involved in the phagocytosis of M. tuberculosis are the Fc gamma receptor (FcγR), the mannose receptor and the complement receptor 3 (CR3).  9   Underhill et al. Annu. Rev. Immunol. 2002. 20: 825-52. Reprinted with permission from Annual Reviews.  Figure 3. Receptor and Signalling Interactions During Phagocytosis of Microbes Phagocytosis of the invader is initiated by the simultaneous binding of several pattern-associated molecules and opsonins to multiple host receptors. This binding causes a signalling cascade leading to phagocytosis and various other responses.   The FcγR binds to the Fc region of immunoglobulin G (32). It plays an important role in inducing phagocytosis and acting as a link between cellular and humoral branches of the immune system (33). The mannose receptor, expressed on the surface of tissue macrophages (34), mediates endocytosis and phagocytosis of microorganisms that express mannose-rich cell walls (35) such as M. tuberculosis. Lastly, the CR3 is a versatile receptor capable of recognizing diverse ligands (36). For instance, the CR3 binds to endogenous ligands such as extracellular matrix and coagulation proteins, and to glycoconjugates of pathogens such as M. tuberculosis lipopolysaccharides. Interactions with these ligands contribute to the host recognition of these 10  microorganisms and their clearance (37-39). These receptors work synergistically to augment their activity and trigger phagocytosis of the pathogen (40).   Upon phagocytosis, M. tuberculosis is trapped within an organelle called a phagosome. Phagosomes are derived from the plasma membrane and must acquire microbicidal features, characteristic of innate immunity, to kill and digest the engulfed microbe (41). This process is known as phagosome maturation and includes fusion and fission events with compartments of the endocytic pathway, i.e. the route undertaken by vesicles derived from the plasma membrane to deliver their contents to lysosomes for degradation (30). As illustrated in Figure 4, phagosomes containing foreign particles, including invading microorganisms, interact with the endosomal pathway allowing for the exchange of endocytic solute materials and membrane components between the phagosomes and the endosomes. This results in the acidification of the phagosomes through the acquisition of vacuolar H+-ATPase pumps (V-ATPase) and proteolytic enzymes (lysosomal hydrolases) (42). These fusion events modify the function of the phagosome to reflect the content of the lysosome (42). Moreover, they initiate the production of reactive oxygen intermediates and the processing of antigens for degradation of the organism (43).  11   Poirier and Av-Gay. Microbes and Infection. 2012. 14 (13): 1211-1219. Reprinted with permission from Elsevier.  Figure 4. Stages of Phagosome Maturation Exemplified by E. coli Infection Upon phagocytosis, the phagosome undergoes a series of fusion and fission events with vesicles of the endocytic pathway culminating in the formation of the phagolysosome. During this maturation, the phagosome acquires antimicrobial properties such as an acidic pH and a high concentration of hydrolytic enzymes. These events lead to the digestion of the invader and presentation of antigens on the surface of the phagocyte to initiate an adaptive immune response.   1.2.2 Inhibition of Phagosome Maturation by M. tuberculosis   Interference with host cellular pathways is a common trait of bacterial pathogenesis. For instance, once internalized, Listeria monocytogenes blocks the fusion of the phagosome with lysosomes and escapes the phagosome by secreting exotoxins LLO, PlcA and PlcB (44). Salmonella enterica, by contrast, activates acid tolerance genes allowing it to adapt to the low phagosomal pH (45). Lastly, Legionella pneumophila redirects phagosome maturation by altering maturation of the endocytic vesicle in which it resides. Rather than fusing with early 12  endosomes, late endosomes and lysosomes, phagosomes containing L. pneumophila fuse with membranes derived from the endoplasmic reticulum creating a niche suited for bacterial replication (46).   M. tuberculosis is also able to circumvent the macrophage killing machinery by inhibiting phagosome maturation and preventing the process of phagosome fusion with lysosomes (Figure 5). Armstrong and Hart (47) were the first to report on phagosome maturation arrest in macrophages infected with M. tuberculosis. They showed by electron microscopy that approximately 70% of phagosomes containing M. tuberculosis did not fuse with lysosomes (47). The blockage of phagosome maturation allows M. tuberculosis to avoid proteolytic degradation and antigen presentation required to initiate an adaptive immune response (47). Phagosomes containing M. tuberculosis are therefore characterized by the absence of several phagosomal markers: the V-ATPase pump (43), hydrolases such as cathepsin D (48,49), the small GTPase RAB7 (50,51) and lysosomal markers such as Lysosomal Associated Membrane Protein (LAMP) and Rab-Interacting Lysosomal Protein (RILP) needed for phagosome-lysosome fusion (52,53). The absence of phagosomal markers in M. tuberculosis infected macrophages indicates that active interference with macrophage trafficking events occurs within the majority of phagosomes containing M. tuberculosis.   13   Poirier and Av-Gay. Microbes and Infection. 2012. 14 (13): 1211-1219. Reprinted with permission from Elsevier.  Figure 5. Blockage of Phagosome Maturation by M. tuberculosis The fusion of the early endosome with the late endosome is inhibited by M. tuberculosis. The resulting phagosome lacks several markers required for phagolysosome formation, has a relatively neutral pH and contains few hydrolytic enzymes required for the pathogen degradation. M. tuberculosis remains intact in the phagosome.   1.2.2.1 Biological Effectors Mediating M. tuberculosis Survival in Macrophages  As a result of phagosome maturation arrest by M. tuberculosis, infected macrophages undergo impaired antigen processing and presentation (54), reduced responsiveness to interferon-gamma (IFNγ) (55), reduced production of cytokines, reactive oxygen and nitrogen intermediates (56), and suppression of host cell apoptosis (57). Attenuation of the host cells is attributed to an array of M. tuberculosis effectors composed of proteins, glycopeptides and lipids described below.   14  1.2.2.1.1 LAM   Lipoarabinomannan (LAM) is a major lipoglycan component of the M. tuberculosis cell wall (Figure 2) contributing to bacterial survival in the phagosomal milieu. LAM includes three types of lipoglycans classified according to the presence and structure of the capping motif: mannosylated LAM (ManLAM), phosphoinositol-capped LAM (PiLAM) and arabinofuranosyl-terminated LAM (AraLAM) (58). ManLAM is found in pathogenic mycobacterial species (e.g. M. tuberculosis, M. leprae and M. bovis) whereas PiLAM and AraLAM are mainly found on non-pathogenic mycobacterial species (59).    ManLAM is considered to be a key contributor to host-mycobacteria relationships by possessing immunomodulatory activities. Its main role is the inhibition of calcium (Ca2+) concentration rise in macrophages upon infection, leading to interference with phagosome maturation (60). Ca2+ serves as an essential cell signalling molecule, especially for the phagosome maturation process (60). ManLAM specifically blocks the Ca2+ rise and interferes with a sorting pathway that delivers lysosomal hydrolases and V-ATPases from the trans-Golgi network (TGN) to the phagosome (61). ManLAM also inhibits the Ca2+/calmodulin PI3K hVPS34 (human Vacuolar Protein Sorting protein 34) pathways and blocks the syntaxin 6-dependent delivery of cargo from the TGN to the early endosome (62).   Upon phagocytosis of a pathogen, intracellular Ca2+ concentration normally rises leading to the recruitment of the early endosomal marker, the small GTPase RAB5, to the phagosomal membrane. RAB5 facilitates fusion events between early and late organelles of the endocytic pathway (63). RAB5, along with calmodulin (CaM) and its effector Ca2+/CaM kinase II 15  (CaMKII), recruit type III PI3K VPS34 to the phagosomal membrane (64). PI3K VPS34 is essential for the production of the lipid regulator phosphatidylinositol 3-phosphate (PI3P) on the phagosomal membrane (65). PI3P serves as a membrane tag that signals phagosomes to mature down the phagolysosome biogenesis pathway (62) and affects localization and function of proteins containing PI3P-binding domains (FYVE zinc finger, PH and PX). Proteins containing these domains are involved in membrane trafficking, endosomal protein sorting, multisubunit enzyme assembly at the membrane and fusion (66). One protein containing the FYVE domain is Early Endosome Antigen-1 (EEA1), which is a tethering molecule and a RAB5 effector essential for fusion of early and late endosomes (62). Its FYVE domain binds to PI3P causing membrane fusion by interacting with a family of proteins termed SNARE (Soluble N-ethylmaleimide-sensitive factor Attachment protein REceptor) composed of syntaxin 6, syntaxin 13 and additional elements such as the priming factor NSF and alpha-SNAP (67). SNARE proteins drive fusion of membranes (67) and interaction between PI3P and SNAREs cause early endosomes to fuse with late endosomes allowing for the delivery of the endosomal cargo between them (68).    Phagocytosis of M. tuberculosis results in a decrease in Ca2+ level in the macrophage by an unknown mechanism (60) and is linked to the mycobacterial inhibition of the Ca2+ effector protein CaMKII (69). This inhibition hinders the recruitment of PI3K VPS34 to the phagosomal membrane, the production of PI3P via the PI3K dependent pathway and the subsequent recruitment of EEA1, which drives fusion of early and late endosomes (Figure 6) (61,64). Although studies have shown that ManLAM is behind the blockage of intracellular Ca2+ rise leading to phagosome maturation arrest, it is still not known how ManLAM prevents intracellular Ca2+ levels from rising.  16   Philips. Cell Microbiol. 2008. 10 (12): 2408-15. Reprinted with permission from John Wiley and Sons.  Figure 6. Inhibition of Phagosome Maturation by M. tuberculosis Effectors Mycobacterial effectors (depicted in red) impair different pathways by diminishing or inhibiting activity of host proteins (shown in blue) required for phagosome maturation. The cell wall glycolipid ManLAM and the PI3P phosphatase SapM interfere with the recruitment of EEA1 to and the production of the phagosomal membrane protein PI3P at the mycobacterial phagosome. The secretion of PtpA dephosphorylates VPS33B impairing membrane fusion between the phagosome and the lysosome.   1.2.2.1.2 SapM  Prior to the discovery of SapM in M. tuberculosis, acid phosphatases were thought to play a role in microbial pathogenicity (70,71). Saleh et al. (72) isolated a 28-kDa protein from M. tuberculosis culture filtrate and characterized it as acid phosphatase SapM, for secreted acid phosphatase of M. tuberculosis. SapM functions in an acidic-to-neutral pH range (70); the optimum pH for enzymatic hydrolysis of the artificial protein-tyrosine phosphatase substrate pNPP (p-nitrophenyl phosphate) was observed at pH 6.5-7.5 (72) by quantitation of released pNP (p-nitrophenol) (73) and Pi (74).  17   SapM was the first mycobacterial acid phosphatase to be identified and it appears to be the only one secreted by M. tuberculosis. Vergne and colleagues observed that SapM takes an active part in preventing M. tuberculosis phagosome maturation. They found that the exclusion of the membrane tag PI3P from phagosomes was only partly explained by the decrease in Ca2+ levels (75) and that SapM hydrolyzes the phagosomal membrane protein PI3P to PI (Figure 6) (75). They concluded that SapM dephosphorylation of PI3P prevents the recruitment of EEA1 to the phagosomal membrane and the fusion of phagosomes with late endosomes (75). Vergne et al. (75) also tested whether intracellular Ca2+ levels can restore PI3P levels on phagosomes containing live mycobacteria. They found that despite the Ca2+ rise, a transient spike in PI3P levels occurs in only 10% of all phagosomes observed. This indicated that reduced Ca2+ levels alone are not sufficient to explain the reduction of PI3P levels on mycobacterial phagosomes and reinforces the role of SapM in phagosome maturation arrest (75). To accomplish complete arrest, M. tuberculosis maintains a PI3P-free environment during its long-term stay in infected macrophages (60,75). Thus, in addition to ManLAM, a second mycobacterial macromolecule, SapM, is required for complete inhibition of PI3P accumulation at the phagosomal membrane, impeding EEA1 recruitment and fusion with late endosomes (75).   The role of SapM in infection remains a conundrum as it is found to be secreted into the lumen of the phagosome but the mechanism by which it gains access to the cytoplasmic face of the phagosome where it hydrolyzes PI3P is unknown (75). It is predicted that SapM contains an export signal allowing it to be exported to the cytosol. Nevertheless, SapM was experimentally proven to directly interfere with host cellular pathways by hydrolyzing host PI3P into PI, resulting in phagosome maturation arrest (75,76). 18  1.2.3 Inhibition of Antigen Processing and Presentation  As a result of phagosome maturation arrest, M. tuberculosis is able to alter antigen processing and presentation and evade the host adaptive immune response. Major histocompatibility complex class II (MHC-II) molecules loaded with bacterial antigenic peptides, the products of bacterial digestion in phagolysosomes, traffic to the cell surface and present specific M. tuberculosis antigens to helper T cells, which initiate an adaptive immune response. Once the antigens are presented to helper T cells, the proinflammatory cytokine IFNɣ is secreted to activate infected macrophages by inducing a microbicidal response through the generation of reactive oxygen and nitrogen species, and by further amplifying antigen processing and presentation (77). IFNɣ plays a crucial role in activating an immune response and controlling macrophage infection by increasing the expression of MHC-II molecules and accessory molecules required for antigen presentation (78). Interestingly, upon binding to Toll-like receptor 2 (TLR2) located on macrophages, M. tuberculosis triggers a potent proinflammatory immune response early during infection. In later stages of infection, the proinflammatory response results in inhibition of antigen processing and presentation. It has been shown that this inhibition of the proinflammatory response is required to combat the infection (79).            1.2.3.1 19 kDa Lipoprotein  The M. tuberculosis cell wall 19 kDa lipoprotein (LpqH) is identified as a pathogen-associated molecular pattern (PAMP) protein (79,80). The function of LpqH in M. tuberculosis grown in vitro and its biological activity are not yet identified, but it is proposed to have a role as an antigenic modulator of host immune processes (79). LpqH is shown to bind to the macrophage TLR2 and selectively inhibit IFNγ induction of the CIITA gene, the MHC-II gene 19  and accessory protein genes required for antigen presentation (81,82). CIITA, the MHC-II transactivator, regulates the expression of genes encoding MHC-II molecules and other genes involved in antigen processing and presentation by promoting the binding of transcription factors to the MHC-II promoter (82). Upon binding to TLR2, LpqH elicits a proinflammatory immune response in the host. This immune response helps combat infection by inducing expression of cytokines, bacterial killing and apoptosis (83). Interestingly, LpqH seems to have a pleiotropic effect on the immune system, i.e. its presence results in the activation of an immune response contributing to host defence, but also results in the inhibition of a subset of IFNγ responsive genes leading to persistence of the infection (84). However, over a prolonged period of time some of these proinflammatory innate immune responses are downregulated to limit damage to surrounding tissues (82).   The mechanism by which LpqH inhibits expression of MHC-II and antigen processing and presentation remains unclear. It is known, however, that the inhibition is dependent on TLR2 and Mitogen-Activated Protein Kinase (MAPK) signalling (p38 and/or ERK) (85), and that the expression of CIITA is suppressed (81,85). Chromatin remodeling has been suggested to be responsible for the inhibition of a subset of IFNγ-induced genes by M. tuberculosis, since CIITA expression is dependent on changes in chromatin structure (82,85).   Eukaryotic DNA transcription is controlled by chromatin structure wrapped around histones. Unstructured histone tails can be targeted for post-translational modifications (such as phosphorylation, methylation and acetylation) that modulate gene transcription. Histone acetylation is a transcription activator and it was observed that MHC2TA, the gene encoding 20  CIITA, is silenced by deacetylation of histone in response to LpqH (86). Pennini and colleagues (85) showed that CIITA promoter activity is inhibited by LpqH by a mechanism that involves inhibition of chromatin remodeling in a TLR2- and MAPK-dependent manner. They also showed that IFNγ-induced acetylation of histones H3 and H4 was inhibited by LpqH. This indicates that the inhibition of chromatin remodeling and histone acetylation at the CIITA promoter accounts for the inhibition of CIITA mRNA expression (85). Two models were developed by the same group in order to explain the inhibition of chromatin remodeling by LpqH. One model involves the reduced expression of histone acetyl transferase (HAT), known to catalyze acetylation of histones H3 and H4, and/or an increased expression of histone deacetylase (HDAC) that suppresses transcription through the deacetylation of histones. LpqH is proposed to inhibit the expression/activity of HAT required at the CIITA promoter or increase the expression/activity of HDAC that regulates the CIITA promoter (85). The second model involves inhibiting the recruitment of certain transcription factors (e.g. Brahma-related gene-1 - Brg1) or the activation of CIITA repressors (85). Regardless of the mechanism, LpqH inhibits CIITA expression via TLR2 and MAPK signalling resulting in deacetylation of histones, a loss of chromatin remodeling and a decrease in subsequent expression of MHC-II and accessory molecules (85). Additional studies are required to confirm the exact mechanism of inhibition.  1.3 PtpA 1.3.1 Overview   Upon macrophage infection, M. tuberculosis actively expresses a low molecular weight phosphatase, named Protein-tyrosine phosphatase A (PtpA), which was shown to play an essential role in M. tuberculosis pathogenicity (87). PtpA is secreted into the macrophage’s 21  cytosol (87) where it interferes with and disrupts key components of the macrophage endocytic pathway. PtpA blocks phagosome acidification and maturation via interaction with V-ATPase subunit H and dephosphorylation of VPS33B, respectively (Figure 6) (76,88).   1.3.2 VPS33B and the HOPS Complex  Vacuolar Protein Sorting 33B (VPS33B), a substrate of PtpA within human macrophages, is a key cytoplasmic, peripheral membrane protein ubiquitously expressed in eukaryotic cells (76). VPS33B is recruited from the cytosol and, along with three other vacuolar sorting proteins (VPS11, VPS16 and VPS18), forms what is known as the Class C complex (89). The Class C complex binds two additional proteins (VPS39 and VPS41) forming the HOPS (HOmotypic fusion and vacuole Protein Sorting) complex, a large multimeric tethering factor essential for vesicle fusion (Figure 7) (90). The HOPS complex is localized to the phagosomal membrane where it is required for phagosomal trafficking to the lysosome (91). It is needed during the tethering and docking stages of vesicle fusion between phagosomes and lysosomes (92) where it plays a role in regulating the assembly of SNAREs, and cytosolic or membrane-bound proteins  required in several membrane fusion steps in secretory and endocytic pathways (93-95), through interaction with endolysosomal Rab GTPases (96,97). Specifically, the HOPS complex associates with GTP-RAB7 through the interaction of VPS39 and VPS41 to promote tethering, SNARE-mediated membrane fusion of the phagolysosome and degradation of the pathogen (98). These interactions permit the exchange of cytosolic contents and subsequent degradation of the invader (99).   22   Solinger and Spang. FEBS J. 2013. 280 (12): 2743-57. Reprinted with permission from John Wiley and Sons.  Figure 7. Subunit Arrangement of the HOPS Complex The organization of the HOPS complex according to Brocker et al. (100). The HOPS complex regulates membrane fusion events at lysosomes through interaction with RAB7 and SNAREs.    Recently, studies from our laboratory showed that M. tuberculosis PtpA interferes with the process of phagosome-lysosome fusion by dephosphorylating VPS33B (76). VPS33B dephosphorylation blocks the assembly of the HOPS complex and prevents its interaction with RAB7 and SNAREs. To support this finding, phagosomes harboring a M. tuberculosis mutant (where PtpA was deleted; ΔptpA) were shown to undergo increased fusion with lysosomes and increased transfer of lysosomal contents when compared to phagosomes containing the parental strain, H37Rv M. tuberculosis (76). Therefore, dephosphorylation of VPS33B inhibits fusion between the phagosome and the lysosome allowing M. tuberculosis to survive and grow within the enclosed and protected environment of the phagosome (76).  23  1.3.3 Subunit H of the V-ATPase Pump  The V-ATPase pump is key to the cell’s ability to combat infections. V-ATPase controls phagosome acidification by transporting protons across phagosomal membranes (101). During phagosome maturation, lysosomes extend along microtubules to fuse with and deliver the V-ATPase to the phagosome where the proton pump recruits the HOPS complex to allow for phagolysosome fusion (102). This fusion generally results in a reduction in phagosomal pH from 6.5 to approximately 4.5, and in the activation of hydrolytic enzymes required to eliminate pathogens (103).     Studies from our laboratory showed that M. tuberculosis PtpA binds to subunit H of the V-ATPase pump, which brings PtpA in close proximity to VPS33B (Figure 8) (88). PtpA then dephosphorylates VPS33B and prevents the formation of the HOPS complex required for proper phagolysosome fusion and pathogen elimination (88). Since PtpA is responsible for disrupting the interaction between V-ATPase and the HOPS complex, dephosphorylation of VPS33B by PtpA occurs as a consequence of PtpA binding to subunit H (88). Hence, the inhibition of the phagolysosome fusion and phagosome acidification is dependent on both PtpA phosphatase activity (dephosphorylation of VPS33B) and its ability to bind to subunit H of V-ATPase (88).   24   Wong et al. Trends in Microbiology. 2013. 21 (2): 100-109. Reprinted with permission from Elsevier.  Figure 8. Schematic Representation of V-ATPase Exclusion and Inhibition of Phagosome Acidification by PtpA During phagosome maturation, the V-ATPase pump recruits the HOPS complex for fusion of the phagosome and lysosome membranes. PtpA, secreted into the cytosol, binds to subunit H of the V-ATPase pump and disrupts the interaction between the pump and the HOPS complex. The binding to subunit H brings PtpA in proximity to VPS33B and dephosphorylates it. This results in the shutting down of the membrane fusion machinery and a phagosome with a pH too high to activate hydrolytic enzymes for M. tuberculosis degradation.   1.4 Host Responses to M. tuberculosis Infection 1.4.1 Apoptosis Versus Necrosis  Macrophages infected with M. tuberculosis can undergo two types of cell death; programmed cell death (PCD) or necrosis. Apoptosis and autophagy are examples of PCD and are phenomena of widespread biological importance in which unwanted cells or damaged cellular contents are neatly removed without causing damage to tissues or initiating an inflammatory response (104,105). PCD is also an important innate defence mechanism against intracellular pathogens that prevents the spreading of infections (106,107). On the one hand, 25  apoptosis is regulated by a variety of cellular signalling pathways that culminate in the activation of a set of enzymes named caspases (108). Caspases are cysteine proteases that play key biological roles in the apoptotic, necrotic and inflammatory response pathways (109). Caspases are usually activated by the release of apoptotic mediators from the mitochondria or by ligation of a death receptor (108). Moreover, apoptosis is characterized by structural changes in the nucleus, such as nuclear condensation and fragmentation, chromosomal DNA cleavage, and packaging of deceased cells into apoptotic bodies whose specific morphological features are recognized by surrounding phagocytes (105,110). All of these morphological changes are achieved without the breakdown of the plasma membrane to avoid triggering an inflammatory response and damage to surrounding tissues (110). On the other hand, autophagy functions primarily as an intracellular recycling mechanism (111), which includes the sequestration of long-lived proteins and large cytoplasmic organelles into vesicular compartments followed by their delivery to lysosomes for degradation (112). The degraded metabolites are then reused by the cell for energy needs or macromolecule synthesis (111). Autophagy is triggered by various signals such as nutrient deprivation, damaged organelles, accumulated misfolded proteins, and oxidative stress (113). Autophagy can also provide cytoprotective functions to the host cell as it can be triggered by infection (113). The autophagic response then contributes to the elimination of invading pathogens (114). Several years ago, a link between autophagy activation and M. tuberculosis infection was revealed (115). The induction of autophagy led to a decrease in M. tuberculosis survival rates in infected macrophages, which indicated that the fusion of autophagosomes containing M. tuberculosis with lysosomes for degradation overcame the phagosome maturation block imposed by M. tuberculosis (115). Therefore, autophagy also acts as an important defence mechanism against pathogens (107). 26   Contrary to PCD, necrosis has traditionally been regarded as unorganized and disruptive cell death sharing more similarities with a train wreck than with surgical culling. Necrosis is the end result of a bioenergetic catastrophe initiated by toxic insults, ischemia or physical damage (110). Morphologically, necrosis is different from PCD and is characterized by rapid permeabilization of the membrane causing general swelling of the cell, vacuolation of the cytoplasm, loss of cell membrane integrity, and induction of inflammation due to the release of potentially toxic intracellular contents and proinflammatory molecules in the extracellular space from lysed cells (116). Unlike apoptosis and autophagy, necrosis provides detrimental effects to the surrounding cells and tissues (117).    The ability of M. tuberculosis to manipulate and control the mode of death of infected host cells plays an essential role in its survival and transmission. On the one hand, apoptosis of M. tuberculosis-infected macrophages is directly associated with mycobacterial killing (118) and with enhanced stimulation of T cell responses from antigen presentation (119). Necrosis, on the other hand, has also been observed in M. tuberculosis infection and serves to propagate infection. Indeed, necrosis is an essential part of the M. tuberculosis granuloma. The granuloma is a pathologic feature characteristic of TB and forms in later stages of infection when the immune system cannot eliminate foreign particles (77). It is characterized by the accumulation of immune cells such as infected alveolar macrophages surrounded by monocytes, lymphocytes, giant cells, neutrophils, and foamy macrophages (120,121). Lymphocytes localize to the periphery of the granuloma and associate with collagen fibers and other extracellular matrix components forming a capsule around the cells (120). At this stage, the infected human host does not show signs of disease or transmit the infection to others, and the granuloma remains an organized structure as 27  long as the immune system prevails over the pathogen (120). However, when the immune system fails to contain the infection (e.g. when the host becomes immunocompromised), M. tuberculosis reactivates and the granuloma is marked by a caseous necrotic center (121). This leads to the rupture of the granuloma into a mass of formless cell debris, releasing viable infectious bacilli into the airways and spreading of TB (121). Thus, necrosis of the granuloma represents an important mode of spreading and transmission to surrounding phagocytes for M. tuberculosis (122).    A fine balance exists between the two types of cell death during M. tuberculosis infection; one allows the host cell to have complete control of cell death leading to apoptosis and elimination of the pathogen, the other allows M. tuberculosis to manipulate host cell death signalling pathways leading to necrosis, transmission of the bacteria and subsequent progression of the disease (123).   1.4.2 Apoptosis During Infection with Virulent Versus Avirulent Mycobacterium Strains  Alveolar macrophages clear bacteria from the lungs and thus are well equipped to phagocytose and kill ingested bacteria. Several microbes have developed strategies to prevent infected macrophages from eliminating pathogens via the conventional phagolysosome route. M. tuberculosis is an example of a pathogen that possesses a fundamental virulence strategy to block phagosome maturation allowing it to survive and replicate inside the macrophage phagosome (29). In such instances, macrophages turn to an additional level of defence to kill the pathogen, and thus resort to apoptosis as a disposal route (124,125). Despite efforts deployed by the macrophage to eliminate the invader, some pathogens have found ways to also block 28  apoptosis and evade this second layer of host defence. One pathogen that has developed such an anti-apoptotic tactic is the human blood parasite Leishmania donovani (126).    M. tuberculosis has been shown to interfere with apoptosis in macrophages both in vitro and in vivo, and this appears to be directly related to its virulence capacity (29,125,127,128). Virulent strains of mycobacteria, such as M. tuberculosis H37Rv, M. tuberculosis Erdman and M. bovis, inhibit apoptosis as a virulence mechanism whereas avirulent or attenuated strains, such as M. kansasii, the vaccine strain M. bovis BCG, and the laboratory strain H37Ra, activate apoptosis (29,57,129). The ability of virulent mycobacteria to block apoptosis is of great importance for the pathogen as death of the host cell removes its supportive growth environment. Thus, mycobacterial virulence factors that modulate the apoptotic response of macrophages promote host cell survival and allow pathogen replication (29).   This model, stipulating that virulent mycobacteria inhibit apoptosis whereas attenuated or avirulent mycobacteria stimulate apoptosis, is controversial due to conflicting results obtained in several studies. Some studies have reported that the deletion of the virulent M. tuberculosis anti-apoptotic genes, nuoG, secA2 and ndk, conferred a pro-apoptotic phenotype to the pathogen (130-132). Conversely, it has been reported that infection of macrophages with pathogenic M. tuberculosis in which the nuoG and secA2 anti-apoptotic genes had been reintroduced, stimulated macrophage death by apoptosis to an extent greater than even avirulent strains (133-135). This discrepancy can be partly attributed to bacterial load, the type of host cells, periods of infection and the stimulatory milieu used in the different studies.  29   Moreover, recent work provides evidence that pathogenic M. tuberculosis does indeed have the ability to evade the host immune response by blocking apoptosis and favouring necrosis (Figure 9) (136-139). According to these studies, virulence is a determining factor in the balance between apoptotic versus necrotic cell death; virulent mycobacteria inhibit apoptosis of macrophages early in infection, but stimulate necrosis at later time points where the host cell lyses resulting in the spreading of the infection. Moreover, Butler and colleagues observed that different strains of pathogenic M. tuberculosis (GC1237 and H37Rv) induced equivalent levels of cell death, but caused apoptosis and necrosis in cells to a different degree, indicating that the balance between apoptotic and necrotic cell deaths could be strain-specific (140).    Porcelli and Jacobs. Nat Immunol. 2008. 9 (10): 1101-1102. Reprinted with permission from Nature Publishing Group.  Figure 9. Modes of Macrophage Cell Death Caused by Virulent and Avirulent Mycobacteria Avirulent mycobacteria induce apoptosis of the host. The phagocyte sequesters the bacteria within apoptotic bodies that are taken up by surrounding macrophages. Apoptosis clears the microbial content of the phagocyte and prevents the release of toxic intracellular contents avoiding tissue inflammation. In contrast, virulent mycobacteria possess virulence factors that cause the host cell to undergo necrosis; a process that permeabilizes the phagocyte’s membrane, allowing the bacteria to escape and spread to surrounding phagocytes.  30   A great degree of uncertainty remains in elucidating the relationship between virulence of mycobacteria and control of host cell death. It is clear, however, that macrophage cell death results from a complex meshing of pro- and anti-apoptotic stimuli partly shaped by the virulence of the invader. Other virulence-independent or anti-apoptotic mechanisms such as the binding of ManLAM and LpqH to TLR2 (141,142), the activities of NuoG (130), SecA2 (131) and Ndk (132), and the up-regulation of host anti-apoptotic proteins such as the BCL-2 family members (143), may also directly stimulate necrosis and determine the mode of death undertaken by the macrophage. Nonetheless, the detailed relationship between the control of cell death pathways and virulence of mycobacteria has yet to be characterized and it is necessary to understand how M. tuberculosis achieves intracellular growth while modulating host cell death to its advantage.  1.5 Techniques for Identifying Intracellular Substrates for M. tuberculosis Effectors During Infection  1.5.1 Phosphoproteome Analysis  Signal transduction mediated by protein kinase catalyzed phosphorylation is a key post-translational modification needed to analyze cellular phosphorylation events occurring in macrophages infected with different strains of M. tuberculosis. The experimental approach chosen to analyze signal transduction in this instance is a refined proteomics method, termed phosphoproteome analysis.   Phosphoproteome analysis is a large-scale Western assay that utilizes phosphosite-specific antibodies against a predefined set of key signalling proteins to identify modification events and quantify global protein kinase phosphotransferase activity in cells (144). One notable 31  advantage of phosphoproteome analysis is its high customizability with respect to phosphorylation events involved in specific cellular processes characterized for a particular species. The technique is usually protein phosphosite-specific and targets defined signalling proteins identified as taking active roles in major eukaryotic signalling networks (145). The assay can also be applied to protein kinases mediating cellular phosphorylation events of the cell population of interest under the desired treatment conditions. For this project, we monitored and compared the phosphorylation status of a predefined set of signalling protein kinases from cellular extracts of uninfected human macrophage-like cells and of macrophage-like cells infected with M. tuberculosis H37Rv and ΔptpA M. tuberculosis. These modification events were verified using biochemical assays such as Western blotting.  1.5.2 iTRAQ  To understand the mechanism by which macrophages fight M. tuberculosis and to analyze the extent of proteomic reprogramming macrophages undergo during infection, a multiplexed peptide quantitation methodology, termed Isobaric Tags for Related and Absolute Quantitation (iTRAQ), was employed. iTRAQ is a quantitative mass spectrometry (MS)-based proteomics approach used to perform a global proteomic analysis of cellular proteins (146), in conjunction with tandem mass spectrometry (MS/MS), to identify the resultant peptides (147). iTRAQ precisely quantifies and identifies cellular proteins for quantitative protein analysis by placing isobaric mass labels on peptides in a peptide mixture (Figure 10) (147).    To better understand the evasive nature of M. tuberculosis and its molecular manifest on the macrophage response to infection, we conducted a global quantitative proteomic profiling of 32  infected macrophages. By examining four independent controlled infection experiments, we identified hundreds of proteins whose expression is modulated upon infection in all replicates. We showed that the macrophage response to M. tuberculosis infection mainly includes simultaneous and concerted up-regulation of proteins while the down-regulated response appears unorganized and random. The contribution of the M. tuberculosis secreted virulence factor PtpA was also assessed.   33   Figure 10. Sample Processing and Analysis Workflow for Global Proteome Profiling with Quantification Using Mass Tag Labeling of Peptides Proteins derived from each sample were labeled with distinct iTRAQ labels (113-116). Identification and quantification took place in the MS/MS scan by comparing the peak intensities of the iTRAQ reporter ions. An iTRAQ 4-plex chemical labeling is presented in this figure, although an 8-plex was also used in the experiments. Author’s own work. 34  1.6 Aims of the Study  Data assembled from studies of M. tuberculosis infection of human macrophages indicates that PtpA’s role in infection extends beyond acting on its two known substrates; VPS33B and subunit H of the V-ATPase pump (76,88). In fact, recombinant PtpA was found to dephosphorylate multiple substrates in macrophage cell extracts (148). These findings suggested that M. tuberculosis PtpA actively affects multiple host macrophage signalling pathways.    We explored M. tuberculosis-host interactions using two approaches: a kinome analysis to identify host protein kinases modulated by PtpA and a proteomic assay to analyze global and PtpA-dependent proteomic changes during M. tuberculosis infection.     The first part of this thesis was assigned to the study of the kinome response of the human macrophage-like cell line, THP-1, upon M. tuberculosis and ΔptpA M. tuberculosis infection. Performing a phosphoproteome analysis, the phosphorylation status of a set of predefined host protein kinases involved in key signalling pathways was identified. The modulation effect of PtpA on the host protein kinase, GSK3α, was assessed by performing various biochemical assays and the impact of this PtpA-dependent modulation on the host in M. tuberculosis infection was identified.    The second part focused on analyses of the global proteomic response of THP-1 cells to M. tuberculosis infection. Using the multiplex iTRAQ proteomic strategy, the proteome-wide protein relative expression levels of human macrophages infected with different strains of M. tuberculosis (H37Rv, ΔptpA and ΔptpA::ptpA) were simultaneously determined. We 35  assembled data from four independent iTRAQ experiments and identified proteins that were consistently modulated by M. tuberculosis. We validated these findings by biochemical assays. We also examined the specific contribution of PtpA to the infection process. These experiments provide insight into the global and PtpA-dependent effects of M. tuberculosis from a proteomic view of infected human macrophages.  36  Chapter 2: Methods and Materials 2.1 Reagents and Chemicals 2.1.1 Commercial Reagents  Endotoxin-free RPMI 1640 medium, phorbol-12-myristate-13-acetate (PMA), protease inhibitor mixture, phosphatase inhibitor cocktail, PMSF and glutathione-agarose beads were purchased from Sigma-Aldrich (St. Louis, MO). FBS and kanamycin were purchased from Gibco Laboratories (Burlington, ON), L-glutamine, penicillin and streptomycin from StemCell Technologies (Vancouver, BC), gentamicin from Invitrogen (Burlington, ON), and hygromycin from Roche Diagnostics (Laval, QC). Ni-NTA polyhistidine-tag purification resin was purchased from Qiagen (Mississauga, ON), ATP from PerkinElmer (Boston, MA), and LB broth from Thermo Fisher Scientific (Waltham, MA). OADC was purchased from Difco Laboratories (Detroit, MI) and 7H9 culture media from Becton, Dickinson and Company (Sparks, MD).   2.1.2 Antibodies  Affinity purified rabbit polyclonal anti-phosphoGSK3α (pY279) antibody was purchased from Invitrogen, rabbit polyclonal anti-GSK3α from Abcam (Toronto, ON), rabbit polyclonal anti-caspase-3 IgG from Cell Signalling Technology (Danvers, MA), mouse monoclonal anti-ICAM1 from eBioscience (San Diego, CA), and rabbit polyclonal anti-CTSG and rabbit polyclonal anti-IL1β from Antibodies-online.com (Atlanta, GA). Secondary HRP-conjugated goat anti-rabbit antibody was purchased from Sigma-Aldrich, Alexa Fluor 680-conjugated goat anti-rabbit IgG from Invitrogen, and Alexa Fluor 780-conjugated goat anti-mouse from Mandel Scientific (Guelph, ON). The specificity of the commercial antibodies was tested with Western blot analysis and immunostaining control experiments. 37  2.2 Bacteria 2.2.1 Strain Maintenance  M. tuberculosis H37Rv and its derivative strains were grown in Middlebrook 7H9 broth supplemented with 10% (v/v) OADC and 0.05% (v/v) Tween 80 (Sigma-Aldrich) at 37 °C standing in the incubator. The PtpA-null mutant and the PtpA complemented strains were generated in our laboratory by allelic replacement and were isolated by a sequential two-step selection protocol as previously described (149).  2.2.2 Bacterial Preparation for Infection Bacteria in mid-log phase were harvested by 5 min centrifugation at 2,602 relative centrifugal force (RCF) and were subsequently washed three times with 7H9 with 0.05% Tween 80. For infection of differentiated THP-1 macrophage-like cells, bacteria were opsonized with human serum for 30 min at 37 °C.  2.2.3 In Vivo Bacterial Uptake by Macrophage   To ascertain that infection with the different M. tuberculosis strains investigated resulted in comparable rates of infection of macrophages, THP-1 cells (1.0 x 106) were seeded on six-well plates and differentiated as explained in Section 2.3.1. Bacteria were washed three times with 7H9 broth with 0.05% Tween 80, labeled with 100 μg/ml Alexa Fluor 488 carboxylic acid SS (Invitrogen), and opsonized with 10% human serum. THP-1 cells were infected with the labeled bacteria at a multiplicity of infection (MOI) of 10:1 for 2 h at 37 °C in a humidified atmosphere of 5% CO2. Noninternalized bacteria were washed away, and cells were incubated for another 2 h at 37 °C in a humidified atmosphere of 5% CO2. THP-1 cells were then washed 38  with PBS, scraped off the plate, and fixed with 2.5% paraformaldehyde as described by Wong and colleagues (88). Uptake was measured with FACS in a FACSCalibur Flow Cytometer (BD Bioscience) and data was analyzed using the FlowJo 8.7 software. The mean fluorescence intensities (the geometric mean from the distribution in FACS histograms) of Alexa Fluor 488 (FL1) were used to calculate bacterial uptake. The mean fluorescence intensities of uptake of wild-type M. tuberculosis were used as a reference.  2.3 Cell Culture 2.3.1 Tissue Culture Maintenance and Differentiation The human monocytic leukemia cell line THP-1 (TIB-202; American Type Culture Collection, Manassas, VA) was cultured in 125 cm2 tissue culture flasks (Corning Inc., Corning, NY) in RMPI 1640 medium supplemented with 10% FBS, 1% L-glutamine, 1% penicillin and 1% streptomycin. Cells were seeded in 10 cm (diameter) tissue culture dishes at a density of  7.0 x 106 cells/dish and differentiated into a macrophage-like cell line with 20 ng/ml PMA in RPMI 1640 medium supplemented with 10% FBS and 1% L-glutamine (incomplete RPMI) at 37 °C in a humidified atmosphere of 5% CO2 for 18 h.  2.3.2 Macrophage Infection  Bacterial cells were washed with Middlebrook 7H9 broth supplemented with 0.05% (v/v) Tween 80 (Sigma-Aldrich). Infection of THP-1 macrophage-like cells was performed using human serum-opsonized M. tuberculosis at a MOI of 10:1 in RPMI 1640 medium. After 3 h of incubation at 37 °C and 5% CO2, cells were washed with RPMI 1640 medium to remove non-39  internalized bacteria and re-incubated at 37 °C and 5% CO2 in incomplete RPMI containing 100 μg/ml gentamicin for a predetermined number of hours.   2.3.3 Macrophage Cellular Extraction for Western Blot Analysis At defined time points after infection, infected THP-1 macrophage-like cells were washed twice with cold PBS and cellular extracts were harvested in lysis buffer (20 mM Tris-HCl, 5 mM EDTA, 1% Triton X-100, 1 mM DTT, 1 mM PMSF, 1 mM phosphatase inhibitor cocktail (Sigma-Aldrich), pH 7.2) by drawing the solution in and out of a blunt syringe 15-20 times. The cellular extracts were centrifuged for 10 min at 20,199 RCF and passed through a 0.22 μm filter column (Millipore Corporation, Billerica, MA).  2.3.4 Macrophage Cellular Extraction for iTRAQ Analysis   Four hours post-infection, M. tuberculosis-, ΔptpA M. tuberculosis- and ΔptpA::ptpA M. tuberculosis-infected THP-1 cells were washed twice with RPMI 1640 medium and scraped in 3 ml of the medium. Cells were centrifuged for 10 min at 180 RCF, resuspended in 490 μl iTRAQ lysis buffer (25 mM ammonium bicarbonate, 5 mM sodium fluoride, 5 mM sodium orthovanadate, 10 mM sodium β-glycerophosphate pentahydrate, pH 7-7.5) and transferred to screw-cap tubes containing 0.5 g of 0.5 mm glass beads. Ten μl of 10% sodium dodecyl sulphate (SDS) were added to the tubes and cellular extracts were bead-beated (BioSpec Products Inc., Bartlesville, OK) for 2 min (30 sec on/30 sec off) at a speed of 5.0. The lysates were then centrifuged for 20 min at 20,199 RCF and the supernatants were passed through a 0.22 μm filter column.  40  2.4 DNA Manipulation 2.4.1 RNA Extraction and cDNA Synthesis Total RNA was extracted from M. tuberculosis-infected THP-1-derived macrophages (7.0 x 106 cells) at defined time points using the RNAspin Mini Kit according to the manufacturer’s instructions (GE Healthcare, Burnaby, BC). RNA was reversed transcribed to cDNA using the EasyScript cDNA Synthesis Kit following the manufacturer’s protocol (ABM, Richmond, BC). For each cDNA synthesis, 1 µg of total RNA, measured by an Epoch Microplate Spectrophotometer (BioTek, Winooski, VT), and 0.5 µM oligo(dT) oligonucleotide primers were used.   2.4.2 DNA Cloning The plasmid pANT7-GSK3α was purchased from DNASU Plasmid Repository (Tempe, AZ). Because this vector was not created for bacterial inducible expression, GSK3α was PCR-amplified from the plasmid DNA using the GSK3α F primer with a BamH1 site and GSK3α R primer with an EcoR1 site. The amplified gene was digested and inserted into the pGEX-6P-3 vector (GE Healthcare) to generate the pGEX-6P-3-GSK3α plasmid for GST-tagged GSK3α fusion protein expression.  The human GSK3α His-tagged fusion protein expression plasmid, pBO1-GSK3α, was purchased from GeneCopoeia Inc. (Rockville, MD). The gene encoding RAB7 was PCR-amplified from cDNA from prepared THP-1 cells using the RAB7 F primer with a BamH1 site and RAB7 R primer with an XhoI site. The amplified gene was inserted into the pET22b vector 41  (Millipore Corporation) to generate the pET22b-RAB7 plasmid for His-tagged RAB7 fusion protein expression.   The M. tuberculosis ptpA gene was previously PCR-amplified from M. tuberculosis H37Rv genomic DNA using the PtpA F primer with an EcoR1 site and the PtpA R primer with an Xho1 site. The amplified gene was inserted into the pGEX-6P-3 vector to generate the pGEX-6P-3-PtpA plasmid for GST-tagged PtpA fusion protein expression.   All plasmid constructs were verified by sequencing at Eurofins MWG Operon (Ebersberg, Germany). The oligonucleotides used for DNA cloning in this study are described in Table 1.  Oligonucleotides Sequence (5’ → 3’)* GSK3α F TATATAGGATCCATGAGCGGCGGCGGGCCTTCG GSK3α R TATATAGAATTCGGAGGAGTTAGTGAGGGTAGG PtpA F ATATATGAATTCCGTGTCTGATCCGCTG PtpA R ATATATCTCGAGTCAACTCGGTCCGTTC RAB7 F TATATAGGATCCATGACCTCTAGGAAGAAAGTG RAB7 R TATATACTCGAGTCAGCAACTGCAGCTTTC *The recognition sequence for the restriction site is underlined.  Table 1. Oligonucleotides Used for DNA Cloning  2.4.3 Quantitative Polymerase Chain Reaction (qPCR) Total RNA from lysates of M. tuberculosis-infected THP-1-derived macrophages was extracted at defined time points and reversed transcribed as previously described (Section 2.4.1). Primers specific for the mRNA of the genes of interest were designed using Primer-BLAST (National Center for Biotechnology Information, Bethesda, MD) (Table 2). Control PCR 42  amplifications for the expression of gene-specific mRNAs were performed on cDNA templates from uninfected PMA-differentiated THP-1 cells to confirm the specificity of the designed primers. Each qPCR reaction contained 2 X EvaGreen qPCR Mastermix (ABM), 15 ng cDNA and 1 μM of each primer, and was analyzed in quantification mode on a DNA Engine Opticon instrument (Bio-Rad Laboratories, Mississauga, ON). The following cycling conditions were used: 95 °C for 10 min, 40 cycles of 95 °C for 15 sec, 52 °C for 15 sec, and 60 °C for 30 sec with data collection during each cycle. Mock reactions (no reverse transcriptase) were also included with each experiment to confirm the absence of genomic DNA contamination. Ct values were converted to copy numbers using standard curves. Results were analyzed using GraphPad Prism 5.0 software. All values of gene-specific mRNA were internally normalized to cDNA expression levels of housekeeping gene GAPDH.  Oligonucleotides Sequence (5’ → 3’) Caspase-3 F TGAGGCGGTTGTAGAAGAGTTT Caspase-3 R GCTCGCTAACTCCTCACGG CTSG F ATGACGAGGAAGCGATGGTG CTSG R TGGCAGGAAGTACTGGAGTAGA GAPDH F GAAGGTGAAGGTCGGAGTC GAPDH R GAGGGATCTCGCTCCTGGAAGA GSK3α F GCTCACCCCTGGACAAAGGTGTT GSK3α R CGCACAGGCCTCTAGTGGGGA ICAM1 F GACCATCTACAGCTTTCCGGC ICAM1 R CATTCAGCGTCACCTTGGCT IL1β F AGATGAAGTGCTCCTTCCAGG IL1β R CATGGCCACAACAACTGACG MIF F CCGGACAGGGTCTACATCAACT MIF R TGGGCGGGCCTAGAACA SOD2 F GTGGAGAACCCAAAGGGGAG SOD2 R GAAACCAAGCCAACCCCAAC  Table 2. Oligonucleotides Used for Quantitative PCR  43  2.5 Protein Expression and Purification 2.5.1 Expression of Recombinant Proteins in E. coli To express and purify His-tagged recombinant protein, pBO1-GSK3α and pET22b-RAB7 plasmids were transformed into chemically competent BL21 E. coli cells. The transformed cells were used to grow starter cultures overnight at 37 °C, and were then inoculated into 2 L of LB media (Thermo Fisher Scientific) supplemented with 100 μg/ml ampicillin (Bio Basic Inc., Toronto, ON). The cultures were then grown to an OD600 of 0.6 at 37 °C, and expression was induced with 0.4 mM IPTG (Bio Basic Inc.) at room temperature overnight. The bacteria were harvested by centrifugation at 4,066 RCF, resuspended in lysis buffer (50 mM Na2HPO4, 290 mM NaCl, 10 mM imidazole, pH 8.0, and 1 mM PMSF) and lysed by sonication. The soluble fraction of the bacterial lysates was obtained by centrifugation at 20,199 RCF and His-tagged proteins were purified from the soluble fraction by affinity chromatography on Ni-NTA polyhistidine-tag purification resin.   GST-tagged PtpA and GSK3α were expressed in BL21 E. coli cells transformed with pGEX-6P-3-PtpA and pGEX-6P-3-GSK3α plasmids. A starter culture of the transformed BL21 was grown overnight at 37 °C and inoculated into 2 L of LB media with 100 μg/ml ampicillin. The culture was grown at 37 °C to an OD600 of 0.6 and expression was induced with 0.4 mM IPTG at room temperature overnight. The bacteria were harvested by centrifugation at 4,066 RCF, resuspended in lysis buffer (50 mM Na2HPO4, 290 mM NaCl, 10 mM imidazole, pH 8.0, and 1 mM PMSF) and lysed by sonication. The soluble fraction of the bacterial lysates was obtained by centrifugation at 20,199 RCF and GST-tagged proteins were purified from the soluble fraction by affinity chromatography on glutathione-agarose resin (Sigma-Alrich). 44  2.5.2 Expression of Recombinant Proteins in M. smegmatis The competent cells M. smegmatis were used for transformation of the pALACE-PtpA plasmid. Transformed bacteria were used to grow starter cultures over two nights in 7H9 media containing 0.05% Tween 80, 50 μg/ml hygromycin and supplemented with 1% ADS (0.5% bovine serum albumin-fraction V, 0.2% dextrose, 0.85% saline) enrichment. The starter cultures were inoculated into 2 L of 7H9 media containing 0.05% Tween 80, 50 μg/ml hygromycin and supplemented with 1% ADS and grown at 37 °C to an OD600 of 1.0. Cells were harvested by centrifugation at 4,066 RCF for 30 min and resuspended in 7H9 media supplemented with 0.05% Tween 80, and 50 μg/ml hygromycin. Expression of the protein was induced with 0.2% acetamide at room temperature for 24 h. The bacteria were harvested by centrifugation at 4,066 RCF for 30 min, resuspended in lysis buffer (50 mM Na2HPO4, 290 mM NaCl, 10 mM imidazole, pH 8.0, and 1 mM PMSF) and lysed by sonication. The soluble fraction of the bacterial lysates was obtained by centrifugation at 20,199 RCF and His-tagged proteins were purified by affinity chromatography on Ni-NTA polyhistidine-tag purification resin.  2.6 Phosphoproteome Analysis by KinetworksTM Phospho-Site Screen (KPSS) Assay  THP-1 cells were infected with wild-type M. tuberculosis H37Rv or the H37Rv strain in which the ptpA gene was deleted (ΔptpA M. tuberculosis) as previously described (Section 2.3.2) and cellular extracts were harvested 18 h post-infection (Section 2.3.3). The macrophage lysates were prepared for phosphoproteome analysis (KPSS 1.3) according to the manufacturer’s instructions (Kinexus Bioinformatics Corporation, Vancouver, BC). Data was analyzed according to statistical confidence provided by experience in analyzing over ten thousand 45  screens. According to Kinexus, the significance levels of change are over 25% variability in intensity.  2.7 In Vitro Phosphatase Activity Assay To assess for phosphatase activity of PtpA in vitro, the artificial chromogenic substrate pNPP was used. The reaction contained 100 mM Tris-HCl pH 8.0, 5 μM MgCl2, 2 μM MnCl2, 1 μM DTT and 30 mM pNPP with 5 μM of purified recombinant PtpA. The reaction was incubated at 37 °C for 1 h. The absorbance (A450) was measured after 1 h using a Bio-Rad Model 680 microplate reader.  2.8 In Vivo and In Vitro Western Blot Analysis  Western blot analyses of cell extracts from infected THP-1 cells harvested 18 and 48 h post-infection (Sections 2.3.2 and 2.3.3) were performed as follow: 50 μg of THP-1 cell extracts were resolved by SDS-PAGE and transferred onto a nitrocellulose membrane (Bio-Rad Laboratories). The blots were probed with affinity purified rabbit polyclonal anti-phosphoGSK3α (pY279) IgG, anti-GSK3α or anti-caspase-3 antibody (final IgG dilution for all antibodies, 1:1,000) and incubated overnight at 4 °C. For detection of phosphorylated GSK3α (Y279) and non-phosphorylated GSK3α, HRP-conjugated goat anti-rabbit (final IgG dilution, 1:3,500) antibody was used as the secondary detection reagent and the blot was developed by enhanced chemiluminescence (ECL) (Thermo Fisher Scientific). For detection of caspase-3, Alexa Fluor 680 goat anti-rabbit antibody was used as the secondary detection reagent (final IgG dilution, 1:10,000) and detection was done using an Odyssey Infrared CLx Imager (LI-COR Biosciences, Lincoln, NE).  46  For iTRAQ analyses, lysates of infected THP-1 cells were harvested 4 h post-infection (Sections 2.3.2 and 2.3.4). Briefly, 100 μg of cell extracts were resolved by SDS-PAGE and transferred onto a nitrocellulose membrane. The blots were probed with affinity purified mouse monoclonal anti-ICAM1 antibody (final IgG dilution, 1:250), affinity purified rabbit polyclonal anti-IL1β IgG (final IgG dilution, 1:1,000), or affinity purified rabbit polyclonal anti-CTSG antibody (final IgG dilution, 1:500) and incubated overnight at 4 °C. For detection of ICAM1, Alexa Fluor 780 goat anti-mouse antibody was used as the secondary detection reagent (final IgG dilution, 1:5,000). For IL1β and CTSG, Alexa Fluor 680 goat anti-rabbit antibody was used (final IgG dilution, 1:10,000). An Odyssey Infrared CLx Imager (LI-COR Biosciences) was employed for chemiluminescence detection. To ensure identical protein loading of the different samples, Ponçeau staining of the blots was performed.  In other experiments, 1 to 4 μM of recombinant GSK3α were incubated with 0.15 mM ATP for 1 h at 37 °C. Then, a fixed concentration of recombinant PtpA (0.04 μM) was added to different GSK3α concentrations and incubation was continued for 45 min at 37 °C. Samples were then resolved by SDS-PAGE and transferred onto a nitrocellulose membrane. The blot was probed with rabbit anti-phosphoGSK3α (pY279) IgG and HRP-conjugated goat anti-rabbit antibody was used as the secondary detection reagent as described above. The blots were developed by ECL. Blots were stripped and reprobed with anti-GSK3α antibody to confirm that total GSK3α protein levels were identical in each reaction samples.   47  2.9 Determination of PtpA and GSK3α Dissociation Constant  The interaction between PtpA and GSK3α was measured using a Fusion-α-HT Multimode Microplate Reader (PerkinElmer) and the ALPHAScreen Histidine (Nickel Chelate) Detection Kit (PerkinElmer). Purified GST-tagged recombinant PtpA was biotinylated using the EZ-Link Biotinylation Kit (Thermo Scientific) and diluted in the assay buffer (25 mM HEPES, 100 mM NaCl, and 0.1% Tween 20, pH 7.4) in 384-well microplates (PerkinElmer). Purified His-tagged recombinant GSK3α was added to wells containing PtpA. Nickel chelating acceptor beads were further added to the proteins and the microplate was incubated for 30 min at room temperature. Streptavidin donor beads were then added to the reactions and incubation was continued for 1 h at room temperature. Kinetics of the reactions was monitored in the ALPHAScreen apparatus by luminescence signals generated from protein-protein interactions (counts per second (cps)). Dissociation constants were determined using GraphPad 5.0 software.  2.10 In Vitro Protein Kinase Assay  Three separate subsets of reactions containing 2, 3 and 4 μM recombinant GSK3α were autophosphorylated in a kinase buffer (50 mM Tris-HCl, 5 mM MgCl2, 2 mM MnCl2, 1 mM DTT, pH 7.5) containing 10 μCi γ32P-ATP (PerkinElmer) for 30 min at 37 °C. After this incubation period, 0.04 μM of PtpA was added to the second subset of reactions and 0.04 μM of PtpA and 1.5 mM or 5 μM of the phosphatase inhibitors, sodium orthovanadate (Na3VO4) or BVT 948, to the third. The first subset of reactions served as controls. Incubation of all three subsets was continued for 15 min at 37 °C. At the end of the incubation period, reactions were stopped with the addition of SDS sample loading buffer and heated at 95 °C for 8 min. The samples were resolved by 12% SDS-PAGE. The gel was silver-stained, dried and exposed to a 48  screen overnight. The 32P-radioactively labeled protein bands were detected by a PhosphorImager SI apparatus (GE Healthcare). Bands corresponding to phosphorylated GSK3α were cut, submerged in scintillation fluid (Beckman Coulter Inc.) and analyzed by scintillation counting using a Beckman Coulter LS 6500 (Beckman Coulter, Mississauga, ON).  2.11 Radiometric Protein Kinase Assay The kinase assay was performed as previously described (Section 2.10) until the end of the second incubation period. Reactions were spotted onto phosphocellulose paper (GE Healthcare), dried and washed thoroughly with 1% phosphoric acid six times for 10 min. Radioactivity levels were measured by submerging the phosphocellulose papers in scintillation fluid and analyzed by scintillation counting.  2.12 Isobaric Tags for Related and Absolute Quantitation (iTRAQ) and LC-MS/MS  Analyses THP-1 cells were infected with M. tuberculosis, ΔptpA M. tuberculosis and the complemented strain (ΔptpA::ptpA), and cell extracts were prepared as described above (Sections 2.3.2 and 2.3.4, respectively). Protein concentrations in cell extracts were determined using a bicinchoninic acid assay (Sigma-Aldrich). Protein samples (200 μg from each test) were sent to the University of Victoria Genome British Columbia Proteomics Centre for iTRAQ analyses. In brief, 85 µg of each sample were precipitated overnight in acetone at 4 C, then resolubilized in 0.5 M Tetraethylammonium Bromide (TEAB), 0.2% SDS. Proteins were then reduced with TCEP and alkylated with MMTS. Proteins were digested in solution with trypsin (Promega, Madison, WI) at 37 °C overnight and labeled with the appropriate iTRAQ label (113-119, 121), 49  an isobaric stable isotope tag, at room temperature for 1 h. iTRAQ labeled peptides were then combined and separated by strong cation exchange HPLC. HPLC fractions containing peptides were concentrated by speed-vac, then analyzed and quantified by LC-MS/MS. The length of the reverse gradient used was 2 h per HPLC fraction. Samples were analyzed by reversed phase nanoflow (300 nl/min) HPLC with nano-electrospray ionization using a LTQ-Orbitrap mass spectrometer (LTQ-Orbitrap Velos, Thermo Fisher Scientific) operated in positive ion mode.  2.13 Data Analysis and Interpretation  All data was analyzed using Proteome Discoverer 1.4 (Thermo Fisher Scientific) and the search engine MASCOT v2.4 (Matrix Science, Boston, MA). Raw data files were searched against the Uniprot-SwissProt database with allspecies filter and mammalian species only.   2.14 Statistical Analysis   Raw data files from each of the four experimental trials were collected and analyzed using R (gnu). All four trials included M. tuberculosis/uninfected ratios for detected proteins. Additionally, the first two trials included ΔptpA M. tuberculosis/uninfected and ΔptpA::ptpA M. tuberculosis/uninfected ratios. Approximately two thousand proteins were detected in each trial, of which only 845 were detected in all trials. All data was log-transformed and each trial was normalized individually. Trials were normalized before elimination of contaminants and missing data.    For detection of anomaly, protein ratios of uninfected and M. tuberculosis-infected macrophages were compared using two methods. The first involved isolating all proteins for 50  which the expression ratio in at least three of four assays was one standard deviation above/below the mean expression ratio. For normalized data, the mean for each assay is zero and the standard deviation is one. For instance, to isolate up-regulated proteins, all proteins with a normalized log-ratio above one in at least three of the four assays were kept. The second method involved taking the mean ratio for each protein over all four assays and isolating those proteins for which this mean was outside of the ±1 standard deviation interval about the mean.   A method was implemented for the initial two iTRAQ assays which included all three treatment conditions, i.e. infection with M. tuberculosis, ΔptpA M. tuberculosis, and ΔptpA::ptpA M. tuberculosis. This was inspired by the apparent co-linearity between each of these covariates. Proteins for which the infected/uninfected ratio was 1.8 standard deviations above the mean for all three possible infected/uninfected ratios were considered consistently up-regulated proteins for that sample, and therefore not PtpA-specific. The same was done to isolate proteins consistently down-regulated for each sample. These generally up- or down-regulated samples had proteins ranked based on their mahalanobis distances (150) from the mean of the given sample where the three variables used in calculation of the correlation matrix were the three infected treatment conditions. Furthermore, the remaining proteins were analyzed in accordance to their M. tuberculosis/ΔptpA M. tuberculosis ratios, organized into up- and down-regulated lists based on the 1.8 standard deviation criterion, and ranked based on their distance from the mean.    51  Chapter 3: Results 3.1 M. tuberculosis Promotes Anti-Apoptotic Activity of the Macrophage by PtpA-Dependent Dephosphorylation of Host GSK3α 3.1.1 Introduction  M. tuberculosis pathogenicity partly relies on the inhibition of phagosome acidification and maturation (76,88). These processes are blocked, in part, by the low molecular weight protein-tyrosine phosphatase, PtpA, secreted by M. tuberculosis into the macrophage cytosol (76). We showed that PtpA dephosphorylates the macrophage protein-serine/threonine kinase GSK3α on the tyrosine residue 279. GSK3α is a multifunctional protein kinase that acts as a master regulator of numerous signalling pathways including the insulin response, glycogen regulation, cell survival and apoptosis (151,152). GSK3α’s activity is positively regulated by the phosphorylation of the tyrosine residue located in the activation loop, Y279, and this phosphorylation is essential for the full activity of the enzyme (153). Studies have shown that apoptotic stimuli increase GSK3α’s activity by phosphorylation of  Y279 (154), providing evidence for a role for tyrosine phosphorylation in apoptosis. Considering this, we demonstrated that dephosphorylation of GSK3α on amino acid Y279 leads to modulation of the enzyme’s anti-apoptotic activity, promoting pathogen survival early during infection. This was exemplified by the inhibition of the proteolytic cleavage and activation of the apoptosis executioner, caspase-3, in M. tuberculosis-infected macrophages. These findings revealed that PtpA significantly contributes to inhibiting host cell apoptosis at early stages of infection.  52  3.1.2 PtpA Interferes with Multiple Host Macrophage Signalling Pathways   Protein phosphatases play key roles in signal transduction in a number of pathways (155). To decipher the multifaceted activity of PtpA on macrophage signalling pathways, we previously performed an in vitro dephosphorylation assay in which recombinant PtpA was incubated with macrophage cell extracts. As seen in Figure 11 and Table 3, this resulted in the dephosphorylation of multiple host proteins including human VPS33B (hVPS33B), a host substrate previously identified in our laboratory (76).    Figure 11. Phosphoproteomic Analysis of Host Macrophage Proteins Dephosphorylated by PtpA Revealed by 2D Gel Electrophoresis A cellular extract of differentiated THP-1 cells was incubated with γ32P-ATP in a kinase buffer for 30 min at 30 °C. Recombinant PtpA was added to one of the samples for 15 min and mixtures were electrophoresed onto 2D SDS-Polyacrylamide gels using isoelectric focusing in a 4-7 pH gradient for the first dimension and 10% SDS-PAGE for the second dimension. The radiolabeled spot profile was obtained by exposing overnight to a PhosphorImager screen. Sixteen spots demonstrating reduced phosphorylation upon addition of PtpA were identified by mass spectrometry (Table 3).      kDa 53  Spot No. Protein identification* MOWSE Score^ MW  (pI) Coverage (%) 1 Xin B (CAF25191) Phosphoinositide 3-Kinase Class-3 (NP002638) 5.8e+16 3.17e+15 122.1 (5.2) 101.5 (6.4) 15 12 2 Rabaptin (NP004694) Rabaptin-5 (AAC70781) Rabaptin-4 (3832516) 3.25e+07 3.25e+07 1.63e+07 99.3 (4.9) 95.6 (4.9) 95.5 (4.9) 21 19 19  VPS39 (AAH15817) 1.61e+07 90.3 (6.6) 19 3 Exocyst Complex Component Sec6 (O60645) cGMP-Dependent Protein Kinase-1 (Q13976) Gamma Adducin (Q9UEY8) 1.31e+02 1.21e+02 1.56e+02 86.8 (5.8) 76.3 (5.7) 79.1 (5.9) 12 11 7 4 Unknown  (AAH04303) 3.1e+103 52.5 (5.9) 64  Hypothetical Protein  8.36e+97 52.4 (5.5) 51 5 MHC Class I Antigen Cw*7 (P10321) 5.31e+01 40.7 (5.6) 7  MHC Class I Antigen Cw*1 (P30499) 2.36e+01 40.9 (5.5) 6 6 Not Determined    7 GTPase, IMAP Family Member-7 (Q8NHV1)  1.06e+16 34.5 (6.1) 63 8 Annexin A13 (P27216) 1.19e+31 35.5 (5.5) 60 9 N-Myc Interactor-STAT Interactor (Q13287) Cdc42 Effector Protein-4 (Q9H3Q1) 3.64e+43 3.7e+42 35.1 (5.2) 37.9 (5.1) 57 41 10 Not Determined    11 Vacuolar Protein Sorting 33B (AAF91174) 6.7e+52 70.6 (6.3) 64 12 Ras-Related Protein Rab-7L1 (O14966) 2.15e+69 23.1 (6.7) 70 13 Ras-Related Protein Rab-28 (Rab-26) (P51157) 6.5e+48 24.9 (5.7) 44 14 Syntaxin 18 (Q9P2W9) MHC Class I Antigen Cw*3 (70076)  1.34e+04 38.7 (5.4) 40.7 (6.0) 21 15 15 Arfaptin-1 (P53367) 1.86e+01 41.7 (6.2) 10 16 MHC Class I Antigen Cw*17 (Q95604) 1.63e+01 41.2 (6.3) 19 * Accession numbers are shown in parenthesis, ^ Score based on peptide frequency. Table 3. Identification of Proteins Dephosphorylated by the Addition of Recombinant PtpA to a Phosphorylated THP-1 Cellular Extract A cellular extract of differentiated THP-1 cells was incubated with recombinant PtpA. The phosphorylation status of several THP-1 cell proteins was found to be modulated by PtpA. These proteins were identified by mass spectrometry. The numbers seen in Table 3 correspond to spots shown in Figure 11. MOWSE score or Molecule Weight Search allows calculation of the probability of matching N peaks by random chance and is based on the frequency of a peptide molecular weight being found in a protein of a given molecular weight range (156). Accession numbers are shown in parenthesis.   3.1.3 KinetworksTM Analysis To identify other macrophage signalling proteins that might be affected by M. tuberculosis PtpA, we conducted a specific large-scale proteomics analysis of signalling networks, termed KinetworksTM analysis, which tracks the kinome and phosphoproteome (145). This multi-immunoblotting method uses an array of phospho-specific antibodies against defined 54  human signalling proteins and networks (145). To test PtpA’s effect on signal transduction pathways, we monitored and compared the phosphorylation status of a predefined set of signalling proteins from uninfected macrophages, macrophages infected with M. tuberculosis and macrophages infected with ΔptpA M. tuberculosis. We chose to investigate events occurring 18 h post-infection. Based on the selective phosphoproteome analysis, as seen in Figure 12, we observed that PtpA affects the phosphorylation pattern of a series of host signalling proteins.    Figure 12. Global Phosphoproteome Analysis of THP-1 Cells Infected with M. tuberculosis and ΔptpA M. tuberculosis Simultaneous detection of selected host proteins and their activation status using a multiple immunoblotting technique. Accurate intensity values for each protein are the accumulated signal obtained over a given scan time for each blot. These are shown as numerical values in Table 4. The three gels represent (A) uninfected THP-1 cells, (B) THP-1 cells infected with M. tuberculosis, and (C) THP-1 cells infected with ΔptpA M. tuberculosis. Each lane was probed with one or more antibodies. The highlighted proteins are host signalling proteins showing a phosphorylation change greater than 25% (red: upregulation, green: downregulation). Antibodies against the phosphorylated proteins were as follows: lane 1, molecular size standard; lane 2, NR1 (S896); lane 3, PKR1 (T451); lane 4, STAT5A (Y694); lane 5, PKCα (S657), Src (Y418); lane 6, JNK (T183+Y185), RSK1/3 (T359+S363/T356+S360); lane 7, MEK3/6 (S189/S207), PKCα/β2 (T638+T641); lane 8, ERK1 (T202+Y204), ERK2 (T185+Y187), S6Kα p70 (T389), S6Kα p85 (T389); lane 9, PKCε (S729), SMAD1/5/9 (S463+S363/S463+S465/S465+S467); lane 10, STAT3 (S727); lane 11, JUN (S73); lane 12, RAF1 (S259), STAT1α (Y701), STAT1β (Y701); lane 13, PKBα (AKT1) (T308), PKCδ (T507); lane 14, PKBα (AKT1) (S473); lane 15, GSK3α (S21), GSK3β (S9), MSK1 (S376); lane 16, α-Adducin (S726), γ-Adducin (S693), CDK1/2 (Y15), Src (Y529); lane 17, GSK3α (Y279), GSK3β (Y216); lane 18, p38α MAPK (T180+Y182), RB (S780); lane 19, NPM (S4), MEK1/2 (S218+S222); and lane 20, CREB1 (S133), RB (S807+S811).  55  As illustrated in Figure 12, cellular extracts of uninfected and M. tuberculosis-infected macrophages were subjected to simultaneous screens. Changes in phosphorylation were measured based on the intensity of 38 predefined phosphoproteins (Table 4) shown as bands in each gel. The phosphorylation levels of the three different treatments were compared in terms of the relative fold change in phosphorylation. The fold change was calculated by comparing the accumulated signal of proteins obtained over a given scan time (normalized counts per minute) from uninfected macrophages and macrophages treated with M. tuberculosis, to the accumulated signal of proteins from macrophages treated with ΔptpA M. tuberculosis (control; accumulated signal set as 1) (Table 4). Due to the high sensitivity of the assay in determining the phosphorylation state of phosphoproteins, a change in phosphorylation greater than 25% between treated cells is considered significant according to the supplier of the screening kit (Kinexus Bioinformatics Corporation). Kinexus Bioinformatics Corporation has performed over ten thousand screens and thorough analysis of all data determined this value. A change in phosphorylation of less than this percentage may be due to experimental and/or biological variation.         56  Protein Full Name Abbreviation Epitopes Control ΔptpA M. tuberculosis Fold Change Uninfected        M. tuberculosis Adducin Alpha (ADD1)     Adducin Gamma (ADD3) B23 (Nucleophosmin, Numatrin, Nucleolar  Protein NO38) Cyclin-Dependent Protein Kinase-1/2 cAMP Response Element Binding Protein-1  Extracellular Regulated Protein Kinase-1  (p44 MAP Kinase)  Extracellular Regulated Protein Kinase-2  (p42 MAP Kinase)  Glycogen Synthase Kinase-3 Alpha  Glycogen Synthase Kinase-3 Alpha  Glycogen Synthase Kinase-3 Beta  Glycogen Synthase Kinase-3 Beta  Jun N-Terminus Protein Kinase  (Stress-Activated Protein Kinase (SAPK))-1/2/3  Jun Proto-Oncogene-Encoded AP1 Transcription  Factor S73 MAPK/ERK Protein Kinase-1/2 (MKK1/2)  MAP Kinase Protein Kinase-3/6 (MKK3/6)  MAP Kinase Protein Kinase-6 (MKK6)  Mitogen- and Stress-Activated Protein Kinase-1  N-Methyl-D-Aspartate (NMDA) Glutamate  Receptor-1 Subunit Zeta  Mitogen-Activated Protein Kinase p38 Alpha  Protein Kinase B Alpha (Akt1)  Protein Kinase B Alpha (Akt1)  Protein Kinase C Alpha  Protein Kinase C Alpha/Beta-2  Protein Kinase C Delta  Protein Kinase C Epsilon  Double-Stranded RNA-Dependent Protein Kinase  Raf 1Proto-Oncogene-Encoded Protein Kinase  Retinoblastoma-Associated Protein  Retinoblastoma-Associated Protein  Ribosomal S6 Protein Kinase-1/3   p85 Ribosomal Protein S6 Kinase-2  p70 Ribosomal Protein S6 Kinase Alpha  SMA- and Mothers Against Decapentaplegic  Homologs-1/5/9   Src Proto-Oncogene-Encoded Protein Kinase  Src Proto-Oncogene-Encoded Protein Kinase  Signal Transducer and Activator of Transcription-1  Alpha Signal Transducer and Activator of Transcription-1  Beta Signal Transducer and Activator of Transcription-3  Signal Transducer and Activator of Transcription-5  α-Adducin γ-Adducin B23 [NPM]  CDK1/2 CREB1 ERK1  ERK2  GSK3α GSK3α GSK3β GSK3β JNK  JUN  MEK1/2  MEK3/6  MEK6  MSK1 NR1  p38α MAPK PKBα [AKT1] PKBα [AKT1] PKCα PKCα/β2 PKCδ PKCε PKR1 RAF1 RB RB RSK1/3  S6Kα p85 S6Kα p70 SMAD1/5/9   Src Src STAT1α  STAT1β  STAT3 STAT5 S726 S693 S4  Y15 S133 T202 + Y204  T185 + Y187  S21 Y279 S9 Y216 T183 + Y185  S73  S217 + S221 S189/S207 S207 S376 S896  T180 + Y182 T308 S473 S657 T638/T641 T507 S729 T451 S259 S780 S807 + S811 T359 + S363/ T356   + S360 T412 T389 S463 + S465/S463     + S465/S465            + S467 Y418 Y529 Y701  Y701  S727 Y694 0 0 0  1 1 0  0  0 1 0 1 1  0  0 0 0 1 1  0 0 0 1 1 1 0 1 1 1 0 0  1 1 0   0 1 1  1  1 0     2.00 0.70      0.65  0.93 ND       0.57 0.85     1.50 0.94 0.81  1.31 0.88 0.45    1.16 1.03     0.89 0.56  1.04  0.79     ND 0.84      0.33  0.61 1.15       0.67 0.90     1.25 0.62 0.51  1.54 0.47 0.78    0.88 0.85     1.57 0.86  0.90  0.86 ND, not determined. Table 4. Phosphoproteome Analysis of Host Signalling Proteins Affected by PtpA A total of 38 phospho-specific antibodies targeting key host signalling proteins were used. The trace quantity of each protein band was measured by the area under its intensity profile curve and corrected for the individual scan times (recorded time before saturation occurs). Values for the control samples were set to 1 or 0. A value of 0 indicates that no immunoreactive signal was detected for this protein. An immunoreactive signal was detected for only 17 proteins. Values for uninfected THP-1 cells and cells infected with M. tuberculosis show the fold change relative to their respective control samples (ΔptpA M. tuberculosis).   57  Out of the 38 signalling proteins tested, an accumulated signal was detected for 17 macrophage phosphoproteins. Among these, several displayed significant changes in phosphorylation between macrophages infected with M. tuberculosis and the control ΔptpA mutant. These include Protein Kinase C Alpha (PKCα; fold change of 1.25 compared to the control), Double-Stranded RNA-Dependent Protein Kinase (PKR1; 1.54), Protein Kinase C Alpha/Beta-2 (PKCα/β2; 0.62), Raf 1 Proto-Oncogene-Encoded Protein Kinase (RAF1; 0.47), Protein Kinase C Delta (PKCδ; 0.51), Mitogen- and Stress-Activated Protein Kinase-1 (MSK1; 0.67), Src Proto-Oncogene-Encoded Protein Kinase (Src; 1.57), Glycogen Synthase Kinase-3 Alpha (GSK3α; 0.33), and Glycogen Synthase Kinase-3 Beta (GSK3β; 0.61). From this list, Src kinase was the only protein previously shown to be associated with M. tuberculosis infection (157).  3.1.4 M. tuberculosis PtpA Dephosphorylates the Human Kinase GSK3α Among all host signalling proteins whose phosphorylation status was affected by PtpA (Figure 12, Table 4), we identified human Glycogen Synthase Kinase-3 (GSK3) as another potential substrate for mycobacterial PtpA. We chose to investigate GSK3α for further analysis, because GSK3α Y279 showed a 67% decrease in phosphorylation in macrophages infected with M. tuberculosis compared to macrophages infected with the ΔptpA mutant strain. GSK3α was identified as the protein with the greatest level of dephosphorylation by M. tuberculosis.   3.1.5 GSK3α Transcription Levels are not Decreased by PtpA  To rule out the possibility that the GSK3α dephosphorylation observed in the KinetworksTM analysis (Figure 12, Table 4) was caused by reduced levels of expression due to 58  PtpA’s effect on GSK3α transcription levels, we examined levels of GSK3α transcripts by qPCR. RNA from uninfected THP-1 cells and from THP-1 cells infected with M. tuberculosis and ΔptpA M. tuberculosis was harvested 18 h post-infection corresponding to the time point at which lysates were harvested for the KinetworksTM analysis. As seen in Figure 13, qPCR profiling revealed a general modest increase in GSK3α transcript levels in cells infected with both M. tuberculosis and the ΔptpA mutant without significant difference between the two. The difference in GSK3α transcript levels between M. tuberculosis- and ΔptpA M. tuberculosis-infected cells may be attributed to GSK3α‘s response to its inhibition via PtpA’s catalyzed dephosphorylation. Nonetheless, the difference is not significant and we concluded that PtpA does not affect GSK3α expression levels and that the dephosphorylation observed in the phosphoproteome analysis is due to a bona fide dephosphorylation of GSK3α by PtpA.                    Figure 13. Transcriptional Levels of GSK3α Post-Infection Quantitative PCR analysis comparing mRNA levels of GSK3α from different infection conditions. RNA from uninfected and treated THP-1 cells (treated with M. tuberculosis and ΔptpA M. tuberculosis) was extracted (A) 4 h and (B) 18 h after infection and reversed transcribed. Data observed show the expression levels of GSK3α. Transcript abundance was determined relative to housekeeping gene GAPDH. Data shown are the means ± standard deviation of three independent experiments. The difference in GSK3α transcript levels between M. tuberculosis- and ΔptpA M. tuberculosis-infected cells was not significant (p-value of 0.8868 for A and 0.5193 for B). **, p < 0.001. Significant differences were compared by Student’s t test. 59  3.1.6 PtpA Dephosphorylates GSK3α Under In Vivo Growth Conditions   To further examine GSK3α dephosphorylation by PtpA, we conducted Western blot analyses in which we tested cell extracts from macrophages infected with M. tuberculosis and the ΔptpA mutant (Figure 14). The phosphorylation level was monitored using the same anti-phosphoGSK3α (pY279) antibody used in the KinetworksTM analysis (Figure 12). As seen in Figure 14, GSK3α phosphorylation levels were found to be higher in extracts obtained from macrophages infected with the ΔptpA mutant compared to macrophages extracts obtained from infection by the parental M. tuberculosis strain, confirming our KinetworksTM analysis screening. We used an anti-GSK3α antibody to confirm that total protein levels of GSK3α was not reduced in the lysates from macrophages infected with M. tuberculosis with functional PtpA.   Figure 14. Western Blot Analyses of PtpA Dephosphorylation of GSK3α In Vivo THP-1 cells were infected with M. tuberculosis or ΔptpA M. tuberculosis. Cellular extracts were harvested 18 h post-infection and 50 μg were used for Western blotting in which the anti-phosphoGSK3α (pY279) antibody was utilized. The bottom panel represents the membrane probed with anti-GSK3α. The molecular mass of GSK3α is 50.981 kDa.   To determine whether GSK3α is a direct substrate of PtpA, we used two separate approaches: biochemical assays to monitor catalysis and a protein-protein interaction analysis to determine interaction between the two proteins. Western blot analysis of recombinant GSK3α to which PtpA was added was performed and we showed that Y279 is dephosphorylated by PtpA in vitro (Figure 15A). To confirm the dephosphorylating effect of PtpA on GSK3α, two Western blot analyses were performed in which the protein-tyrosine phosphatase inhibitors sodium 60  orthovanadate (Na3VO4) (Figure 15B) and BVT 948 (Figure 15C) were added to a GSK3α reaction containing PtpA. Although these are non-specific protein-tyrosine phosphatase inhibitors, their inhibitory effect on PtpA is noticeable bringing the GSK3α Y279 phosphorylation level closer to its basal level.      Figure 15. Western Blot Analyses of PtpA Dephosphorylation of GSK3α In Vitro (A) Different concentrations of GSK3α (1, 2, 3 and 4 μM) with and without PtpA (0.04 μM) were incubated and developed by enhanced chemiluminescence. The anti-phosphoGSK3α (pY279) antibody was utilized. The bottom panel represents the Ponçeau stained membrane showing equal loading of samples. The molecular mass of GSK3α is 50.981 kDa. (B, C) A fixed concentration of GSK3α (3 μM) with and without PtpA (0.04 μM) to which the tyrosine phosphatase inhibitor Na3VO4 (1.5 mM) or BVT 948 (5 μM) was added was incubated and developed by enhanced chemiluminescence. The anti-phosphoGSK3α (pY279) antibody was utilized. The bottom panels represent the Ponçeau stained membranes showing equal loading of the recombinant GSK3α samples.   A more sensitive radioactive assay monitoring GSK3α kinase activity revealed that its autophosphorylation levels were significantly reduced in the presence of recombinant PtpA (Figure 16A). This phenomenon was improved upon addition of the protein-tyrosine phosphatase inhibitor Na3VO4 (Figure 16B) and completely reversed by the addition of BVT 948 (Figure 61  16C). The extent of γ32P-ATP incorporation into GSK3α confirms that GSK3α is a self-phosphorylating autokinase dephosphorylated by PtpA.     Figure 16. Kinase Assays of PtpA Dephosphorylation of GSK3α In Vitro (A) Dephosphorylation of GSK3α by PtpA was tested in an in vitro kinase assay. GSK3α (1 and 2 μM) was autophosphorylated in a kinase buffer containing 10 μCi γ32P-ATP with or without PtpA (0.04 μM) and was resolved onto a 12% SDS gel and exposed to a PhosphorImager screen for radiolabeled band localization. Bands corresponding to phosphorylated GSK3α were cut from the gel, and the radioactive incorporation was measured by a scintillation counter. This graph represents the radioactivity of the dried gel. Results are expressed as ± standard deviation of three independent experiments. The p-value of 2 μM GSK3α with PtpA is 0.0214. (B, C) The inhibiting effect of PtpA on GSK3α’s activity was tested by radiometric analysis. GSK3α (2, 3 and 4 μM) was autophosphorylated in a reaction containing kinase buffer and 10 μCi γ32P-ATP. PtpA (0.04 μM) and Na3VO4 (1.5 mM) or PtpA (0.04 μM) and BVT 948 (5 μM) were added to the sample mixtures and were spotted onto phosphocellulose paper. Radioactivity levels were measured by a scintillation apparatus. *, p < 0.05; **, p < 0.001; ***, p < 0.0001. Significant differences were compared by Student’s t test.  62  To assess whether PtpA binds to GSK3α, ALPHAScreen (Amplified Luminescent Proximity Homogeneous Assay), which is used to monitor protein-protein interactions, was performed. Recombinant GSK3α was immobilized to beads by its His-tag while GST-PtpA was immobilized by biotinylation according to the manufacturer’s protocol (PerkinElmer). The results show direct and dose-dependent interaction between PtpA and GSK3α (Figure 17). A hyperbolic curve fitting the 1:1 Langmuir binding model and a dissociation constant (Kd) of 4.0 × 10-9 M indicate a high level of affinity between the two proteins. PtpA binding to its known host substrate, hVPS33B, has a similar strength with an assessed Kd value of 2.1 x 10-9 M (76).     63   Figure 17. PtpA Interacts with GSK3α In Vitro Protein-protein interaction between GSK3α and PtpA was determined using ALPHAScreen technology. (A) The Kd was calculated using biotinylated PtpA and increasing concentration of His-tagged GSK3α. His-tagged RAB7 protein served as the negative control. Curve fitting yielded a Kd of 4.0 x 10-9 M. (B) The reciprocal experiment with increasing concentration of PtpA yielded a Kd of 3.1 x 10-9 M. GST was used as negative control.  3.1.7 PtpA Interferes with Host Macrophages Apoptosis Early During Infection GSK3α plays a key role in the control of cell fate in both pro- and anti-apoptotic manners (158,159). Previous studies have shown that phosphorylation of GSK3α Y279 is essential for the full activity of the enzyme (153) and that apoptotic stimuli increase its tyrosine phosphorylation activity (154). Although a low level of phosphorylation can be achieved by autophosphorylation, the protein kinases Mitogen-Activated Protein Kinase Kinase-1/2 (MEK1/2) are the most likely 64  protein kinases to phosphorylate GSK3α Y279 (Dr. Steven Pelech, personal communication). We observed that modulation of GSK3α’s tyrosine phosphorylation activity correlates with an interference with apoptosis of the macrophage, the programmed self-destruction process considered to be a major defence mechanism against M. tuberculosis.  To investigate whether dephosphorylation of GSK3α Y279 by PtpA affected apoptosis, measurements of transcriptional and translational expression levels of the apoptotic executioner, caspase-3, were taken (160). Comparative qPCR levels of caspase-3 were performed on cellular extracts from either uninfected macrophages or infected with M. tuberculosis, the ΔptpA mutant or with the complemented ΔptpA mutant (ΔptpA::ptpA) where the original ptpA gene was reintroduced via a replicative plasmid into the mutant strain. As seen in Figure 18, we observed a significant difference in the expression levels of caspase-3 between cells infected with M. tuberculosis and those infected with the ΔptpA mutant. Eighteen hours post-infection, caspase-3 transcription levels were two-fold higher in the ΔptpA mutant-infected macrophages compared to wild-type M. tuberculosis-infected macrophages (Figure 18A) indicating that suppression of  caspase-3 expression by M. tuberculosis is PtpA-dependent early during infection. Caspase-3 expression levels were even lower in macrophages infected with M. tuberculosis than in uninfected macrophages. Interestingly, the inhibition of caspase-3 expression was overturned between 18 and 48 h post-infection where the caspase-3 transcript in M. tuberculosis-infected macrophages increased, surpassing its levels in the ΔptpA mutant-infected macrophages (Figure 18B). This turn of events indicated that PtpA interference with the apoptotic pathway is transient and macrophages are capable of initiating apoptosis regardless of the presence of PtpA. 65   Figure 18. PtpA Reduces Trancriptional Levels of Caspase-3 Early in Infection Quantitative PCR analysis comparing mRNA levels of caspase-3 from different infection conditions. RNA from uninfected and treated THP-1 cells (treated with M. tuberculosis, ΔptpA M. tuberculosis and ΔptpA::ptpA M. tuberculosis) was extracted (A) 18 h and (B) 48 h after infection and reversed transcribed. Data observed show the expression levels of caspase-3. Transcript abundance was determined relative to housekeeping gene GAPDH. Data shown are the means ± standard deviation of three independent experiments. The difference in transcript levels in cells infected with M. tuberculosis and ΔptpA M. tuberculosis 18 and 48 h post-infection were significant (p-value of 0.0179 for A and 0.0097 for B). *, p < 0.05; **, p < 0.001. Significant differences were compared by Student’s t test.  3.1.8 M. tuberculosis Blocks Proteolytic Cleavage of Inactive Caspase-3 into Active Caspase-3  To test whether PtpA’s dephosphorylation of GSK3α resulted in modulation of caspase-3 activity, we monitored caspase-3 proteolytic degradation using the anti-caspase-3 antibody. As seen in Figure 19, M. tuberculosis infection inhibited the cleavage of inactive caspase-3 (31.6 kDa) into its active forms (17/19 kDa). Cellular extracts from uninfected cells and from cells infected with the ΔptpA mutant showed both active and inactive caspase-3 whereas those from M. tuberculosis-infected macrophages showed only inactive caspase-3. Macrophages infected 66  with the ΔptpA::ptpA strain showed similar activation of caspase-3 to macrophages infected with the ΔptpA mutant strain, indicating that the observed effect of the complemented strain is not optimal and in agreement with other complementation phenotypes we have observed (76,161). Complementation of the ΔptpA mutant strain was done using epichromosomal plasmid to reintroduce the ptpA gene to the genome. As observed in Figure 19, the ΔptpA::ptpA strain does not exhibit a wild-type phenotype, but rather a ΔptpA phenoptype. Since we know little about ptpA regulation of gene expression, it is possible that the plasmid did not get expressed from this plasmid. Moreover, the last step of specialized transduction, which was used to generate the original ΔptpA mutant (76), requires alteration of the entire locus within M. tuberculosis genome which may have resulted in deletion of regulatory elements needed for full PtpA expression.                             Figure 19. M. tuberculosis Blocks Activation of Caspase-3 In Vivo THP-1 cells were uninfected (NI), infected with M. tuberculosis (WT), the ΔptpA mutant (KO) or the complement ΔptpA mutant (CO) and cellular extracts were harvested 48 h post-infection. A total of 50 μg of cellular extract were used for Western blotting in which the anti-caspase-3 antibody was utilized. The molecular mass of inactive caspase-3 is 31.608 kDa and 17/19 kDa for active caspase-3. The cellular extract of RAW 264.7 cells treated with 5 μM staurosporine for 5 h was used as the positive control (+ve). The bottom panel represents the Ponçeau stained membrane showing equal loading of samples.  67  3.2 Global Proteomic Analyses of Macrophage Response to M. tuberculosis Infection 3.2.1 Introduction M. tuberculosis possesses multiple strategies to subvert recognition by the immune response. To better understand the evasive nature of the tubercle bacilli and its molecular manifest on the macrophage response to infection, we compared the proteome, i.e. the global protein expression profile, of macrophages infected with M. tuberculosis H37Rv to that of uninfected macrophages. In addition, due to its role in pathogenesis (76,88,148), we further extended our study to identifying the contribution of PtpA to the macrophage response by comparing the proteome of macrophages infected with the mutant ΔptpA M. tuberculosis to that of macrophages infected with the parental M. tuberculosis strain.    In total, four independent proteomic experiments were performed and 9,784 host proteins were detected. Using a number of statistical methods (Section 2.14), we identified 27 proteins whose expression ratios lie outside of a 90% confidence interval about the mean. The majority of these proteins belong to functional classes as diverse as chromatin remodeling and transcription regulation, protein synthesis, immunity and defence, and cell signalling. We verified some of our findings by selecting five proteins whose expression showed significant modulation by infection for quantitative PCR for transcriptional analysis and Western blot for validation of protein expression. In addition, the contribution of the M. tuberculosis secreted virulence factor PtpA was assessed and showed a pronounced PtpA-dependent modulation of 11 proteins involved in RNA metabolism, immunity and defence, and cellular respiration.  68  3.2.2 Modulation of Host Protein Expression in Response to M. tuberculosis   Four independent sets of THP-1 cells infected with M. tuberculosis and the corresponding controls were analyzed by iTRAQ resulting in the detection of 42,007 peptides from cellular extracts of macrophages and in the identification of 9,784 proteins. After removing the repeats, 4,868 distinct proteins were identified. To determine the general effects of M. tuberculosis on host protein expression levels, assessment of expression ratios of identified proteins comparing uninfected to infected macrophages was performed using two-sided t tests for each sample‘s log-ratios. Using this approach we identified a total of 845 proteins in all four independent experiments.  Disparity in values obtained for the expression ratios was observed between experiments (Figure 20A); in three of the four infection experiments, infected versus uninfected log-ratios displayed a mean and a 95% confidence interval below zero. Log-ratios in the fourth experiment displayed a mean and 95% confidence interval above zero. Thus, we normalized the data so that each sample has a standard deviation of approximately one and a mean of zero (Figure 20B). In this manner, the relative modulation of protein expression could be measured in relation to all other proteins in the same experiment, across samples and between independent experiments.   69   Figure 20. Boxplots of Host Proteins Obtained from Infected Versus Uninfected Macrophages Log-ratios measured 4 h post-infection. (A) Log-ratios for four trials of experiments where ratios are between M. tuberculosis-infected and uninfected macrophages. (B) Normalized log-ratios for four trials of experiments where ratios are between M. tuberculosis-infected and uninfected macrophages. Each sample is normalized to have a mean and a standard deviation of approximately zero and one, respectively.    To study the effect of M. tuberculosis on the macrophage proteome, three different strains were used for the infection experiments; the wild-type H37Rv strain (WT), the gene knockout strain ΔptpA M. tuberculosis (KO), and the corresponding complemented strain (CO). We identified proteins that are either consistently up- or down-regulated for each of the strains and determined their expression levels by means of the ratio between infected over uninfected macrophages. The ratios were plotted against one another for each individual sample. We observed that up-regulated proteins (Figures 21A and 21C) show a strong linear relationship between each ratio indicating that upon independent M. tuberculosis infection, similar proteins become up-regulated at similar rates. Alternatively, there appears to be a lack of linearity of the macrophage response in terms of down-regulated protein levels (Figures 21B and 21D). 70    Figure 21. M. tuberculosis-Modulation of Host Protein Expression Proteins for which the infected to uninfected expression ratios are above and below the mean 4 h post-infection. (A) The M. tuberculosis/ΔptpA::ptpA M. tuberculosis log-ratio is plotted and shows a strong linear relationship between these expression ratios for up-regulated proteins. (B) The M. tuberculosis/ΔptpA::ptpA M. tuberculosis log-ratio is plotted and shows little to no linear relationship between these expression ratios for down-regulated proteins. (C) The M. tuberculosis/ΔptpA M. tuberculosis log-ratio is plotted and shows a strong linear relationship between these expression ratios for up-regulated proteins. (D) The M. tuberculosis/ΔptpA M. tuberculosis log-ratio is plotted and shows little to no linear relationship between these expression ratios for down-regulated proteins. Proteins for which one of these ratios was not present in the sample were not included. 71  3.2.3 Identification of Macrophage Proteins Modulated by M. tuberculosis Infection  To identify host proteins modulated by M. tuberculosis infection, we plotted the average infected over uninfected expression level ratios of the 845 distinct proteins. We then determined the statistically relevant proteins by selecting those whose average expression level ratios fell outside of ±1 standard deviation from the global mean for at least three of the four experiments (Table 5). We further shortlisted proteins by taking the mean ratio for each protein over all four trials and excluded those whose ratios were within 1.8 standard deviations of the mean (Table 5). Using these two methods, we identified 27 unique biomarker proteins to be modulated in M. tuberculosis infection (Figure 22, Table 5).        72                                     Up-Regulated Proteins                                                                                  Down-Regulated Proteins Proteins Num. of Peptides Avg. Normalized Inf./Uninf. Ratio p-value  SD Involved in Infection (Ref.) Proteins  Num. of Peptides Avg. Normalized Inf./Uninf. Ratio     p-value    SD Involved in Infection (Ref.) GPNMB 2 1.26 0.1836 1.4664 (162) DOCK10 2 -1.77 0.1307 1.7113  SNRPD1 HMGN2 2 3 2.05 1.16 0.0299 0.2011 1.0531 1.4271  (163) CTSG ADRBK1 6 5 -1.91 -1.33 0.3969 0.0535 3.8686 0.8619 (164) (165) ICAM1 6 2.16 0.1018 1.8507 (166,167) RANBP2 8 -1.40 0.0519 0.8951 (168) H2AFZ ATP5D 4 2 1.56 1.12 0.0355 0.0091 0.8525 0.3700  (169) ITPR1 CEP170 4 3 -1.10 -1.53 0.2812 0.0367 1.6859 0.8469 (170) RPL23A 4 1.52 0.0019 0.2920  ARHGEF2 5 -1.19 0.0903 0.9619 (171) HIST1H4 FAU IL25 DPY30 GRPEL1 FIS1 CHTOP IL1β SOD2 8 1 2 2 2 1 2 4 7 1.54 1.33 0.53 0.88 1.17 1.18 1.39 2.24 1.92 0.0809 0.0059 0.4318 0.0967 0.0166 0.0444 0.0649 0.2461 0.0776 1.1920 0.3786 1.1594 0.7349 0.4796 0.7091 0.9750 3.1216 1.4506 (85)  (172-174)   (175)  (176,177) (176,178) NCAPG BIRC6 WDR3 MIF 2 2 2 3    -1.06 -2.28 -1.31 -1.97    0.1538    0.0804    0.2572    0.0804   1.1115   1.7579   1.8737   2.5254      (179)      (180)  Table 5. M. tuberculosis-Infected Macrophage Proteins Modulated One Standard Deviation Above or Below the Mean This list was obtained by eliminating all proteins for which less than three sample ratios were within one standard deviation of the mean. Proteins in bold are proteins for which the average standardized M. tuberculosis/uninfected ratio (over all samples) was above or below 1.8 standard deviations of the mean. For each protein, two-tailed p-values were calculated using a t test over the four trials. p-value < 0.05 indicates significance at the 95% level. p-value < 0.1 indicates significance at the 90% level.73   Figure 22. Global Expression Level Scatterplot of Normalized Infected Versus Uninfected Log-Ratios Includes 845 proteins detected in all four trials of the experiments where highlighted proteins are those with mean expression ratios above or below a one standard deviation confidence interval about the mean. Blue dots correspond to up-regulated proteins, and green dots correspond to down-regulated, as per Table 5. The extreme values marked by a circle were determined to be contaminants (P35527, P04264, P13645).  Sixteen of the identified proteins were previously reported to be involved in the macrophage response to microbial infections. Up-regulated proteins include the innate immune proteins Superoxide Dismutase-2 (SOD2) (176,178), Intercellular Adhesion Molecule-1 (ICAM1) (166,167), and Interleukin-1β (IL1β) (176,177). Among known down-regulated proteins, we recognized Cathepsin G (CTSG) (164), Migration Inhibitory Factor (MIF) (180), and IAP Repeat-Containing Protein-6 (BIRC6) (179).  74  Eleven proteins were never previously shown to be associated with or affected by infection. These include the up-regulated RNA metabolism protein Small Nuclear Ribonucleoprotein Sm D1 (SNRPD1) and the chromatin remodeling and transcription protein Histone H2A (H2AFZ), and the down-regulated signalling protein Dedicator of Cytokinesis Protein-10 (DOCK10).  3.2.4 Validation of Proteomics Expression Levels  To experimentally validate our findings, we selected five representative proteins, previously reported to be modulated in microbial infections, from those listed in Table 5 and compared our iTRAQ findings to those obtained from either Western blot analysis (for protein levels) or quantitative PCR (qPCR for transcription levels).    Western analysis was conducted for ICAM1, IL1β and CTSG. Based on the iTRAQ results (Table 5), protein levels of ICAM1 and IL1β were expected to be up-regulated in infected cells and those of CTSG, to be down-regulated. As anticipated, levels of ICAM1 and IL1β in M. tuberculosis-infected cells were up-regulated (Figure 23A-B) correlating with and confirming the modulations observed in our proteomic analyses. However, as seen in Figure 23C, in contrast to our iTRAQ findings, CTSG levels remained unchanged when comparing infected to uninfected macrophages.  75    Figure 23. Western Blot Analyses of M. tuberculosis-Dependent Host Protein Expression    Levels THP-1 cells were infected with M. tuberculosis and cellular extracts were harvested 4 h post-infection. A total of 100 μg of cellular extract were employed for Western blotting in which (A) anti-ICAM1, (B) anti-IL1β, and (C) anti-CTSG antibodies were utilized. The molecular mass of ICAM1 is 57.825 kDa, of IL1β is 30.748 kDa, and of CTSG is 28.837 kDa. The bottom panels represent the Ponçeau stained membrane showing equal loading of samples.   The mRNA levels of ICAM1, IL1β, SOD2, CTSG, and MIF were quantified by qPCR. The average mRNA fold change was assessed 2, 4 and 18 h post-infection and was compared to the relative change in protein levels observed in the corresponding iTRAQ experiments. As seen in Figure 24, qPCR analysis revealed up-regulation in mRNA levels of ICAM1, IL1β and SOD2 in cells infected with M. tuberculosis 4 h post-infection (Figure 24A-C). The increase in mRNA abundance correlates with our proteomic data (Table 5). Transcription levels of CTSG and MIF in infected cells were unchanged contradicting the proteomic results that detected down-regulation 4 h post-infection (Figure 24D-E, Table 5). Levels of CTSG mRNA remained 76  unchanged in infected macrophages and increased 18 h post-infection (Figure 24D) while mRNA levels of MIF showed no significant change in any of the time points monitored (Figure 24E).    Figure 24. Transcriptional Levels of Five M. tuberculosis-Modulated Proteins Quantitative PCR analysis comparing mRNA levels of (A) ICAM1, (B) IL1β, (C) SOD2, (D) CTSG and (E) MIF from uninfected and infected cells. RNA from uninfected and M. tuberculosis-infected THP-1 cells was extracted 2, 4 and 18 h after infection and reversed transcribed. Data observed show the expression levels of all genes. Transcript abundance was determined relative to the housekeeping gene GAPDH. Data shown are the means ± standard deviation of three independent experiments.   77  3.2.5 PtpA Effect on Macrophage Protein Expression Levels  To assess the contribution of PtpA to the macrophage’s response to infection, we compared the macrophage proteome in response to infection with WT or ΔptpA KO strains 4 h post-infection and normalized the data as described above to give each sample a mean of zero and a standard deviation of approximately one (Figure 25A-B).     Figure 25. Boxplots of Host Proteins Obtained from Infected Versus Uninfected Macrophages Log-ratios measured 4 h post-infection. (A) Log-ratios for two trials of experiments where ratios are between M. tuberculosis- and ΔptpA M. tuberculosis-infected macrophages. (B) Normalized log-ratios for two trials of experiments where ratios are between M. tuberculosis- and ΔptpA M. tuberculosis-infected macrophages. Each sample is normalized to have a mean and a standard deviation of approximately zero and one respectively.   In total, we identified 1,281 proteins with 11 macrophage proteins whose expression ratios fell outside of a ±1.8 standard deviation interval for macrophages infected with the KO strain (Figure 26, Table 6). Among these PtpA-dependent proteins, three have previously been reported to be involved in the infection process; the up-regulated stress response protein Cold-78  Inducible RNA-Binding Protein (CIRBP) (181), and the down-regulated cell death regulator Glycogen Synthase Kinase-3 Alpha (GSK3α) (182), and transporter ATP-Binding Cassette Sub-Family B Member 1 (ABCB10) (183). GSK3α is the only PtpA-dependent protein that has been identified before to be involved in mycobacterial infections (148,184-186). Among the eight other proteins, the most modulated ones include the down-regulated RNA metabolism protein splicing factor U4/U6 Small Nuclear Ribonucleoprotein Prp31 (PRPF31), and protein synthesis protein tRNA synthetase Seryl-tRNA Synthetase-2 (SARS2).   Figure 26. Global Expression Level Scatterplot of Normalized M. tuberculosis Versus ΔptpA M. tuberculosis Log-Ratios Includes 1,281 proteins detected in two trials of the experiments. Highlighted proteins are those with mean M. tuberculosis/ΔptpA M. tuberculosis ratios above or below a 1.8 standard deviation confidence interval about the mean. Blue dots correspond to up-regulated proteins, and green dots correspond to down-regulated proteins, as per Table 6.79                                         Up-Regulated Proteins                                                                                     Down-Regulated Proteins Proteins Num. of Peptides Avg. Normalized WT/KO Ratio p-value SD Involved in Infection (Ref.) Proteins  Num. of Peptides Avg. Normalized WT/KO Ratio p-value SD Involved in  Infection (Ref.) RPL27 CIRBP  PURB  2 2 1 1.86 2.29 1.98 0.2470 0.2589 0.3189 1.0763 1.3966 1.5345  (181) GSK3α  PRPF31 SARS2 1 2 3 -2.28 -3.14 -3.18 0.5163 0.5464 0.5523 3.4015 5.1394 5.3055 (148,182,184-186)       COX7B 1 -2.10 0.6820 5.4357        ATP5I  BYSL1 ABCB10 GID8 2 2 2 3 -2.32 -2.07 -2.35 -2.07 0.4905 0.1992 0.4462 0.5259 3.1823 0.9451 2.8042 3.1700   (183)  Table 6. ΔptpA M. tuberculosis-Infected Macrophage Proteins Modulated 1.8 Standard Deviations Above or Below the Mean The list was obtained using two methods: (i) eliminating all proteins for which the M. tuberculosis/ΔptpA M. tuberculosis sample ratios were within 1.8 standard deviations of the mean for both samples, and (ii) eliminating all proteins for which the mean M. tuberculosis/ΔptpA M. tuberculosis ratio was within the ±1.8 standard deviation interval about the mean. For each protein, two-tailed p-values were calculated using a t test over the two trials. p-value < 0.05 indicates significance at the 95% level. p-value < 0.1 indicates significance at the 90% level.       80  Chapter 4: Discussion 4.1 M. tuberculosis PtpA Inhibits Host Apoptosis Early During Infection  M. tuberculosis pathogenicity relies upon its ability to sense changes in the environment and respond to host defence assaults. One of the ways to do so is by actively interfering with macrophage physiological pathways (187). One specific strategy utilizes the secreted phosphatase, PtpA,  to block both phagosome maturation and acidification (76,88); two key processes required for digestion of invading microorganisms and initiation of an adaptive immune response (47).   Earlier studies from our laboratory showed that several regulatory phosphoproteins in diverse signalling pathways in macrophages changed significantly in their phosphorylation status upon mycobacterial infection (186). Following the rationale that some of these changes are dependent on M. tuberculosis’ PtpA, we comparatively monitored PtpA’s contribution to the phosphorylation status of key human signal transduction proteins during M. tuberculosis infection. We have now shown that PtpA modulates global phosphorylation patterns of macrophage proteins and that these modulations can impact the fate of the host cell.    The original phosphoproteome analysis that we performed compared the effect of infecting macrophages with live or dead Mycobacterium bovis BCG on phosphoprotein levels (186). Notably, Glycogen Synthase Kinase-3 Beta (GSK3β) was amongst the most phosphorylated proteins upon M. bovis BCG infection (186). Interestingly, the phosphorylation pattern of the M. bovis BCG and the M. tuberculosis phosphoproteome analyses (Figure 12, Table 4) showed some contradicting results exemplified by GSK3α and GSK3β. In the M. bovis 81  BCG phosphoproteome analysis, GSK3α and GSK3β were hyperphosphorylated on Y279/216 in cells infected with live M. bovis BCG with a fold change of 1.29 and 1.57, respectively, compared to the uninfected control cells (186). In the M. tuberculosis phosphoproteome analysis, GSK3α and GSK3β showed reduced phosphorylation in infected cells compared to uninfected cells and had a fold change of 0.51 and 0.66, respectively (Table 4). This discrepancy can be attributed to the genotypic differences of the two strains as M. bovis BCG is an avirulent vaccine strain (188). It is also well documented that macrophages respond differently to M. bovis BCG than they do to M. tuberculosis (189). PtpA is expressed in M. bovis BCG (87), but its function is still under investigation. Potential reduction or absence of secreted PtpA in this vaccine strain could explain the hyperphosphorylation occurring in macrophages harbouring this non-virulent strain.    It is worth noting that in comparison to uninfected cells, the relative fold change of the GSK3 isoforms in macrophages infected with ΔptpA M. tuberculosis resembled that of the isoforms in macrophages infected with live M. bovis BCG. In fact, an increase in phosphorylation for GSK3α (fold change of 1.54) and for GSK3β (1.07) is observed in macrophages infected with ΔptpA M. tuberculosis when compared to the phosphorylation status of these isoforms in uninfected macrophages (Table 4). This data is similar to the results obtained from our previous phosphoproteome analysis when comparing the phosphorylation status of the isoforms in M. bovis BCG-infected cells versus uninfected cells (186). Moreover, PKCα/β2, PKCδ, and RAF1 also had a similar relative fold change in ΔptpA M. tuberculosis- and M. bovis BCG-infected macrophages when compared to uninfected macrophages (Table 4) 82  (186). It appears that the attenuation caused by the ΔptpA mutation (76) makes the mutant strain behave more like the avirulent M. bovis BCG strain.   The observed dephosphorylation of GSK3α and GSK3β provided the first evidence that PtpA may be involved in apoptosis; an effective defence strategy employed by the immune system to limit the growth of intracellular pathogens. More than two decades ago, apoptosis was first recognized to be manipulated by intracellular pathogens to their own advantage (190,191). Apoptosis, as a host defence mechanism, plays such a crucial role in the destruction of microorganisms. Throughout evolution, pathogens have acquired apoptosis-modulating genes (29). GSK3α and GSK3β play an essential role in the regulation of the apoptotic pathway. Studies have shown that phosphorylation of GSK3 Y279/216 is critical for the full activation of these kinases (153) and the induction of apoptosis (154,192,193). We have shown that infection of macrophages with an attenuated mycobacterial strain increases phosphorylation of GSK3 Y279/216 (186), indicating that macrophages are primed for apoptosis. On the one hand, results from both phosphoproteome analyses indicate that the apoptotic pathway is turned on in macrophages infected with the attenuated strains, i.e. in M. bovis BCG and ΔptpA M. tuberculosis, via phosphorylation of GSK3α and GSK3β Y279/216. On the other hand, we have demonstrated that the virulent strain M. tuberculosis H37Rv inactivates GSK3α and GSK3β by dephosphorylation of Y279/216, which promotes survival of the host cell (Table 4).    Several in vitro studies revealed that the rate of apoptosis is increased in macrophages infected with mycobacteria (194,195). However, virulent strains of mycobacteria were shown to minimize apoptosis of macrophages in comparison to avirulent or attenuated strains (29,57), 83  reinforcing the idea that M. tuberculosis has developed strategies to block apoptosis to promote its survival within the host cell. The ability of M. tuberculosis to block apoptosis is of great importance for the pathogen as death of the host cell restricts its supportive growth environment (29). In agreement with this, we demonstrated that GSK3α dephosphorylation of Y279 could be interpreted as an anti-apoptotic feature of this pathogen.   To confirm the role of PtpA in inhibiting macrophage apoptosis, we measured transcriptional levels of caspase-3, a protease that plays a critical role in the execution-phase of apoptosis of the host, and showed that mRNA levels of caspase-3 were significantly reduced in M. tuberculosis-infected cells (Figure 18). This indicates that M. tuberculosis blocks early expression of caspase-3 to prevent apoptosis. Moreover, we showed that a significant difference in caspase-3 transcript levels existed between cells infected with M. tuberculosis and those infected with ΔptpA M. tuberculosis (Figure 18A) confirming that inhibition of caspase-3 gene expression is, in part, dependent on PtpA. The increased caspase-3 gene expression in the ΔptpA mutant may be a result of the mutant’s attenuated phenotype.    Dephosphorylation of host GSK3α by PtpA leads to prevention of host cell apoptosis during early stages of infection. In fact, PtpA's anti-apoptotic role lessens at later stages of infection (Figure 18B), but does not necessarily signify resumption of host macrophages apoptosis. Indeed, we found that proteolytic cleavage of caspase-3 in M. tuberculosis-infected macrophages is blocked 48 h post-infection despite PtpA’s non-engagement. On the one hand, as shown in Figure 19, inactive caspase-3 is expressed in all four treatments, but is only cleaved to active caspase-3 in two treatments: in uninfected cells and in cells infected with ΔptpA 84  M. tuberculosis. On the other hand, macrophages infected with M. tuberculosis and the complemented mutant strain show no cleavage and limited cleavage of caspase-3, respectively.     The inhibition of apoptosis by virulent strains of mycobacteria is a long-established paradigm. Virulent M. tuberculosis secretes bacterial proteins that inhibit apoptosis of the macrophage to promote immortality of the host cell (29). The identification of these proteins is essential to understand the pathogenesis of TB and to understand how these effectors manipulate the host defence mechanism. To date, only a few M. tuberculosis virulence factors have been reported to inhibit apoptosis. We present evidence that the inhibition of apoptosis of the host macrophage early in infection is directly attributed to M. tuberculosis secreted protein PtpA, which inhibits the cleaving of inactive pro-caspase-3 into active caspase-3. Other yet to be identified M. tuberculosis proteins may also be involved in this process. This provides novel insight into M. tuberculosis’ pathogenicity within macrophages and better mechanistic understanding of how it is able to circumvent the macrophage’s killing machinery.    In addition to its role in apoptosis and phagosome maturation, we have shown that PtpA is a versatile virulence factor that influences several signalling pathways to facilitate and maximize replication and intracellular survival. As illustrated in Figure 12 and Table 4, PtpA modulates the phosphorylation status of several signalling proteins such as RAF1, PKCδ, GSK3β, PKCα/β2, MSK1, PKCα, PKR1, and Src (148). Moreover, in vitro phosphorylation analysis of macrophage proteins by recombinant PtpA resulted in the identification of multiple host signalling substrates including hVPS33B, a cognate PtpA substrate we had identified before 85  (Figure 11, Table 3) (76,148). Future experiments will reveal the multifaceted characteristics of PtpA mechanisms enabling M. tuberculosis’ invasion and survival inside the macrophage.  4.2 M. tuberculosis Causes Concerted Up-Regulation and Random Down-Regulation of Host Macrophage Proteins In recent years, several large-scale expression studies investigating global transcriptional responses of macrophages exposed to M. tuberculosis have been performed (157,176,196-208). The emergence of these studies has undeniably contributed to a better understanding of the extent of transcriptomic reprogramming of the macrophage during mycobacterial infection. However, a broader systems-biology approach would provide a better understanding of TB specific host-pathogen interactions and insights into the macrophage biology. Furthermore, understanding the mechanisms by which M. tuberculosis manipulates the host response could provide fertile grounds for novel interventions by means of chemotherapy or vaccine development, especially with the targeting of the most consistent protein changes in infected macrophages. Proteomics provides us with a higher level of understanding of biological functions than transcriptomic analysis as it takes into account protein levels, post-translational modifications and stability, as well as protein subcellular localization.  In this study, we provided insight into the macrophage’s global response to M. tuberculosis infection using a quantitative proteomic approach, and show how the specific virulence factor, PtpA, affects the macrophage proteome. Our first finding indicated that the macrophage acts mainly by modulating protein expression of a variety of specific “responder proteins” that are activated by all strains examined in the study in a concerted up-regulated 86  manner. Contrary to this, down-regulation of protein levels seems to be random and varies from strain to strain and between experiments. Retrospectively, it seems more logical from an energetic point of view to induce a well defined set of proteins required to cope with infection rather than shutting down specific set of housekeeping or resting state homeostasis proteins which will in turn affect the macrophage’s general ability to sustain infection.   To the best of our knowledge this is the first study to simultaneously determine proteome-wide protein relative expression levels of human macrophage-like cells infected with M. tuberculosis, and outline molecular signatures of the global and PtpA-dependent proteomic pattern of macrophages during M. tuberculosis infection.  4.2.1 Host Proteins Targeted by M. tuberculosis  Using the iTRAQ methodology, we identified 845 distinct host proteins whose expression level was modified by M. tuberculosis infection. Among these, 27 were statistically validated and were shown to form the “responder” family of proteins. The immune response proteins (ICAM1, IL25, IL1β, SOD2) were found to be up-regulated. We also found proteins involved in chromatin remodeling and transcription regulation (HMGN2, H2AFZ, HIST1H4, DPY30, CHTOP) to be up-regulated as well as proteins involved in protein synthesis (RPL23A, FAU). This pattern indicates that upon infection, the host turns on the transcriptional and translational machineries to synthesize proteins to fight the pathogen. This is supported by the lack of down-regulation of proteins involved in transcription and translation as the majority of down-regulated proteins belong to the cell signalling (DOCK10, ADRBK1, ITPR1, ARHGEF2) and, surprisingly, immunity and defence (CTSG, MIF, BIRC6) functional classes. The 87  unexpected down-regulation of proteins involved in specific immunity may be a direct outcome of virulence factors secreted by M. tuberculosis that target these host proteins.   Of the 27 modulated proteins, we analyzed proteins with the highest variation index. These include BIRC6 (relative fold change of -2.28 (179)) which has previously been associated with bacterial or viral infections and SNRPD1 (2.05) and DOCK10 (-1.77), which had not been previously linked to infection (Table 5).   The pronounced down-regulation of the apoptosis inhibitor BIRC6 (relative fold change of -2.28) (209) may demonstrate the extent of stress the macrophage is exposed to during infection. Indeed, one of M. tuberculosis’ fundamental virulence strategies is to block phagosome maturation (29). Thus, the macrophage cannot eliminate the pathogen by the conventional phagosome-lysosome fusion route and an additional level of defence is applied; that is, the macrophage turns to apoptosis as a disposal route (124,125). As such, the down-regulation of the apoptosis inhibitor BIRC6 may represent a host-mediated event reflecting the macrophage’s attempt at eliminating M. tuberculosis.  The SNRPD1 protein is a component of the spliceosome; a complex molecular machine involved in the splicing of all cellular pre-mRNAs (210). Previous studies have shown that high concentrations of SNRPD1 result in the production of regulatory T cells (Tregs) (211), which actively suppress the immune system by inhibiting or down-regulating the induction, proliferation, differentiation and effector functions of immune cells such as T cells, B cells, and dentritic cells (212,213). The immunosuppressive activity of Tregs is suggested to be down-88  regulated during immune responses to infectious microorganisms to facilitate their elimination. However, some pathogens, such as M. tuberculosis (214), manipulate Tregs to suppress the host immune response and prevent systemic inflammatory immune responses (215). A M. tuberculosis-induced SNRPD1 up-regulation (relative fold change of 2.05) could represent another tactic employed by the pathogen to reduce the host inflammatory response that contributes to its survival.   The guanine nucleotide exchange factor DOCK10 (relative fold change of -1.77) is also down-regulated. DOCK10 is an activator of Rho GTPases and thus promotes diverse functions dependent on Rho-family proteins (216). Studies have shown that interleukin-4 (IL4) induces expression of DOCK10 in B cells (217). Down-regulation of DOCK10 may be a result of the pathogen targeting DOCK10 as IL4 expression was shown to be induced during M. tuberculosis infection (218). A consequence of a reduced DOCK10 expression could be impaired signalling in B cells and therefore, impaired proliferation and differentiation of B cells, and production of immunoglobulin. Although humoral response to TB is not well understood and established as a protective response, down-regulation of DOCK10 could reduce levels of M. tuberculosis-specific antibodies circulating in the blood and impair the ability of the adaptive immunity to respond to infection.  To validate the iTRAQ data, we investigated the correlation between the transcriptional and translational levels of five proteins whose modulation has previously been reported in mycobacterial infections. We selected SOD2 (relative fold change of 1.92) (176,178), ICAM1 89  (2.16) (167), IL1β (2.24) (176,177), CTSG (-1.91) (164), and MIF (-1.97) (180) for further analysis (Table 5).    The proteomic data of the five modulated proteins was confirmed at the transcriptional level by performing gene expression profiling using qPCR. We also verified the protein expression levels of ICAM1, IL1β and CTSG by Western blotting. As observed in the iTRAQ data (Table 5), mRNA levels of ICAM1, IL1β and SOD2 were up-regulated 4 h post-infection (Figure 24A-C). Their up-regulated expression observed between uninfected and M. tuberculosis-infected cells is persuasive.   ICAM1 exhibited a four-fold increase 4 h after infection (Figure 24A). ICAM1 is a glycoprotein expressed at the surface of immune cells. It plays a role in the migration of macrophages to the site of infection and in the presentation of antigens to T cells (167). Its expression has been reported to be significantly enhanced on macrophages harbouring M. tuberculosis (219). Western blot analysis of ICAM1 showed upregulated expression levels in M. tuberculosis-infected cells (Figure 23A), which is consistent with our iTRAQ results (Table 5) and reports in the literature.   The cytokine IL1β displayed a 10-fold increase in mRNA levels between uninfected and infected cells 4 h post-infection (Figure 24B). IL1β is produced by activated macrophages and acts as an important mediator of the inflammatory response through its involvement in several cellular activities such as T cell proliferation, B cell maturation and proliferation, cell differentiation, and apoptosis (177). IL1β was previously observed in the supernatants of 90  macrophages stimulated with M. tuberculosis. It is believed to have effector function in the cellular response to M. tuberculosis infection and play a role in the resistance of macrophages (220). Western blot analysis of IL1β showed a pronounced increase in expression levels in M. tuberculosis-infected macrophages (Figure 23B). Protein and mRNA expression levels of IL1β were up-regulated across both analytical platforms validating the iTRAQ results (Table 5) and reinforcing the affirmation that IL1β is an important player in the fight against M. tuberculosis.  The mRNA levels of the antioxidant SOD2 increased 12-fold 4 h post-infection (Figure 24C). This up-regulation is justified by the sudden oxidative bursts generated by the macrophage upon phagocytosis of M. tuberculosis. Expression of SOD2 must be increased to defend the host from its own toxic assaults from superoxide anions (O2•-) and reactive oxygen species (ROS) (221-223). In such instances, SOD2 scavenges O2•- offering the host cytotoxic protection. Up-regulation of SOD2 expression is, therefore, a crucial defence mechanism employed by the macrophage, but also a tool used by M. tuberculosis to protect itself from O2•- killing since the protection of the host from O2•- and ROS converts the macrophage into a sanctuary for M. tuberculosis to grow.   A lack of association exists between gene and protein expression levels of CTSG and MIF. As seen in Figure 24D-E, mRNA levels of CTSG and MIF from M. tuberculosis-infected cells show minimal up-regulation for most time points observed. This up-regulation does not correlate with their respective down-regulated protein levels identified by iTRAQ 4 h post-infection (Table 5) and results cannot be directly compared. Moreover, results from Western blot 91  analysis show no change in protein expression levels of CTSG, which seemingly contradict the iTRAQ data (Figure 23C). Transcription levels of CTSG, which encodes a serine protease that possesses antibacterial activities in the killing and digestion of engulfed pathogens, were shown to be down-regulated in macrophages upon M. tuberculosis infection (164), which is in agreement with our iTRAQ results. However, our Western blot and qPCR data does not compare with our iTRAQ results or the literature as expression levels of CTSG are unchanged (Figure 23C) or slightly above basal levels (Figure 24D). Moreover, a 2.5-fold increase in mRNA expression levels is even observed in infected cells 18 h post-infection (Figure 24D). These findings may indicate that M. tuberculosis is only marginally capable of inhibiting CTSG expression to basal levels early during infection.   Lastly, MIF is a proinflammatory cytokine involved in the innate immune response to pathogens. It has been shown to be secreted by and up-regulated in macrophages upon exposure to M. tuberculosis (224). MIF-deficient macrophages have revealed impaired mycobacterial killing (180). Our qPCR data did not correlate with the proteomic results as a slight up-regulation is noticed in infected cells at all time points observed (Figure 24E).  The interpretation of transcriptomic data can be variable due to the high frequency of false positives (225). An additional feature is that the level of transcription of a gene may not reflect its level of expression. Indeed, mRNA transcripts may not always be translated into proteins, may be degraded (226) or, through alternative splicing, may give rise to several different proteins (227). It is also possible that, in the case of CTSG, the protease is more stable within the macrophage by means of possessing a lower degradation rate or higher half-life during 92  infection. By contrast, MIF mRNA may be regulated post-translationally by events such as protein-targeted degradation. Post-translational modifications are strategies commonly employed by pathogens to modulate key host signalling pathways to their advantage. This would play a pivotal role in the pathogenesis of the infection due to MIF’s important functions in immunity and explain why down-regulated protein levels were observed in the iTRAQ analyses, but not in the qPCR assay.  4.2.2 Host Proteins Targeted by M. tuberculosis PtpA   The proteomic response of macrophages infected with the PtpA mutant strain resembled that of macrophages infected with the parental or complement strains (Figure 21A-B). As observed in Figure 21C, a strong linear relationship exists between each of the ratios indicating that upon M. tuberculosis and ΔptpA M. tuberculosis infection similar proteins become up-regulated at comparable rates. However, little concordance is observed in the rates of protein down-regulation, and, as for the other M. tuberculosis strains, infection with the PtpA mutant caused proteins to be down-regulated at seemingly random rates (Figure 21D). By examining two independent controlled infections, we were able to identify modulation of expression of 11 PtpA-dependent host proteins with statistical confidence (Table 6). Surprisingly, the majority of the PtpA-dependent proteins were down-regulated and included members of the immunity and defence (GSK3α, ABCB10) and cellular respiration (COX7B, ATP5I) functional classes. Protein synthesis and respiration require production of ATP by the host. The down-regulation of proteins involved in the electron transport chain leading to a reduction in ATP synthesis seems counterintuitive as reducing ATP production could be detrimental to the host and the pathogen. However, targeting the energy generator of the host may be a strategy employed by 93  M. tuberculosis to weaken the host. Indeed, reducing the production of ATP to levels that would affect protein synthesis of the host and impair its ability to counterattack while still serving as a niche for the pathogen may represent a tactic. Thus, we propose another role for PtpA, which is to facilitate interference with the energy generator of the host, and thereby secure the infiltration and survival of M. tuberculosis.  Among the PtpA-dependent proteins with the greatest modulation, we identified CIRBP (relative fold change of 2.29) (181), GSK3α (-2.28) (148,182), and ABCB10 (-2.35) (183), which have previously been shown to be involved in the infection process, and PRPF31 (-3.14), and SARS2 (-3.18) (Table 6), which have not yet been reported to be associated with infection.    CIRBP is a nuclear protein that responds to stresses such as mild hypothermia (228), exposure to UV irradiation (229), and hypoxia (230) by up-regulation. Recently, CIRBP was described as a new proinflammatory mediator of shock as it is up-regulated in macrophages during septic shock, induces inflammatory responses by stimulating the release of TNF-α, and causes tissue injury (181). The significant up-regulation (relative fold change of 2.29) observed in our iTRAQ results may reflect the stress level the macrophage endures and the corresponding measures it takes to eliminate the pathogen.    GSK3α is a protein-serine/threonine kinase that acts as a regulatory switch for numerous signalling pathways including cell survival and apoptosis (151). In a recent study, we showed that PtpA directly dephosphorylates human GSK3α promoting M. tuberculosis survival by blocking apoptosis of the host early in infection (148). This finding was exemplified by the 94  impaired activation of the apoptosis executioner, caspase-3, in M. tuberculosis-infected macrophages compared to ΔptpA mutant-infected cells.    We showed in our previous study that mRNA levels of GSK3α are not affected by PtpA (148). Here we show that GSK3α protein levels are down-regulated by PtpA (relative fold change of -2.28). As observed for CTSG and MIF, GSK3α stability may contribute to the discrepancy between mRNA and protein levels. Regardless, we now show that PtpA contributes to both the protein levels and the phosphorylation status of GSK3α, which may reflect a common feedback mechanism.    The transmembrane protein ABCB10 is located in the inner mitochondrial membrane and is involved in intracellular trafficking and compartmentalization of peptides (183,216). Previous work done with the Saccharomyces cerevisiae ABCB10 homolog identified the protein as a mitochondrial peptide transporter (214). Degradation products of mitochondrial inner membrane proteins are exported via this transporter, released from the mitochondria (210,211), and presented on the cell surface by major histocompatibility class I (MHC-I) molecules (215). PtpA-dependent down-regulation of ABCB10 (relative fold change of -2.35) supports a putative physiological role in antigen processing (183).   PRPF31 is a protein involved in the formation of the spliceosome and in the splicing of pre-mRNAs (231). Studies have shown that depletion of PRPF31 inhibits spliceosome formation and pre-mRNA splicing (231). Modulation of PRPF31 has never been identified the context of bacterial infection. However, infected macrophages must synthesize large amounts of proteins 95  for self-defence. Therefore, the proper functioning of PRPF31 for RNA processing is deemed crucial for normal macrophage functions. Our iTRAQ analysis indicated a significant down-regulation of PRPF31 (relative fold change of -3.14). We therefore hypothesize that the high demand for spliced mRNA for protein synthesis in infected macrophages becomes a target for M. tuberculosis. The down-regulation of PRPF31 caused by PtpA could signify that splicing of pre-mRNAs is severely compromised and the host defence tactics, markedly weakened.  The aminoacyl-tRNA synthetase SARS2 is a component of the mitochondrial translational apparatus (232). This enzyme catalyzes the ligation of the serine amino acid to mitochondrial tRNA acceptors. Previous studies have shown that mutation in SARS2 greatly impacts acylation of tRNAs and the absence of acylated tRNASer leads to the degradation of nonacylated transcripts (233). The pronounced PtpA-dependent down-regulation of SARS2 (relative fold change of -3.18) observed in our iTRAQ analysis may result in decreased serine aminoacylation of mRNA transcripts and adversely affect mitochondrial translation systems. As such, an impaired mitochondrial protein synthesis apparatus would eventually lead to a defective energy supply provided by the mitochondria and impaired antigen processing. To support this claim, we have identified the expression levels of two mitochondrial proteins, COX7B and ATP5I, involved in the generation of ATP, to be significantly down-regulated in M. tuberculosis-infected macrophages (relative fold change of -2.10 and -2.32 respectively, Table 6). Moreover, the mitochondrial ABCB10 transporter potentially involved in antigen processing was also downregulated by PtpA (-2.35, Table 6). Thus, we suggest that to promote its survival, M. tuberculosis releases PtpA, which targets the mitochondrial translational machinery of the host and impairs its energy generator and weakens its self-defence strategies. 96   Previous studies from our lab analyzed the macrophage’s phosphoproteome response to mycobacterial infection. We investigated the phosphorylation status of key host macrophage kinases during infection with live or heat killed M. bovis BCG (186) or M. tuberculosis or ΔptpA M. tuberculosis (Figure 12) (148). Overall, we noted changes in host signalling pathways caused by mycobacterial infection including apoptosis (up-regulation of c-JUN, SAPK and GSK3β), cytoskeletal arrangement (up-regulation of α-Adducin), Ca2+ signalling (up-regulation of NR1) and macrophage activation (down-regulation of PKCε) (186). We also showed that M. tuberculosis PtpA dephosphorylates an abundance of macrophage proteins (Figures 11 and 12, Tables 3 and 4) (76,148). These events may contribute to downstream effects resulting in the observed protein expression changes we describe in this report. Future experiments will reveal the macrophage’s multiphasic response that enables the symbiotic nature of M. tuberculosis infection.   M. tuberculosis is a host-adapted human pathogen and a deeper biological understanding of the host-pathogen interactome is necessary to get a general idea of what artillery is deployed by the host against M. tuberculosis, and to draw a more detailed picture of how M. tuberculosis manipulates the host response to its advantage. Despite overlooking many notable proteins in the process due to the high-throughput nature of our screening, we expand existing knowledge of the global response of macrophages to M. tuberculosis infection and show the PtpA-specific effect of M. tuberculosis on the organization of the macrophage proteome. We also present novel host targets never observed in M. tuberculosis infection or any other pathogenic infections. These findings provide novel insight into the global proteomic organization (pathways turned on or off) of the macrophage when challenged by M. tuberculosis and into the macrophage protein content. 97  They also help in the understanding of the complex and unique interplay between M. tuberculosis and the host macrophage, and of the manipulative nature of M. tuberculosis to circumvent the macrophage’s killing machinery.   4.3 Future Directions  The M. tuberculosis virulence factor PtpA is known for its inhibitory action on phagosome maturation. Its export into the host cytoplasm indicates that secretion of PtpA is essential to the sabotage of the antimicrobial and defence mechanisms of the macrophage, allowing M. tuberculosis to establish growth within the human macrophage. In this thesis, we presented a new role for PtpA as an anti-apoptotic effector and described PtpA’s influence on the host macrophage proteomic response upon infection. In particular, we demonstrated that PtpA can disrupt multiple host pathways, such as the cellular respiration pathway, to promote M. tuberculosis’ intracellular survival. Based on our work thus far, the interference of PtpA with numerous signalling pathways can translate into several research directions.   The PtpA-dependent inhibition of caspase-3 proteolytic cleavage partially explains the mechanism behind the inhibition of apoptosis during macrophage infection with M. tuberculosis. As of now, we can only speculate that inhibition of caspase-3 cleavage and activation is due to GSK3α dephosphorylation by PtpA. Future work involving PtpA will aim at determining the exact signal transduction pathway engaged following dephosphorylation of GSK3α by PtpA and leading to the inhibition of caspase-3 activation. The pathway affected by PtpA will be established to confirm that inhibition of caspase-3 cleavage and activation in M. tuberculosis infection is truly caused by dephosphorylation of GSK3α Y279. To achieve this, several 98  experiments can be performed such as an immunoprecipitation of GSK3α substrates by GSK3α. Using antibodies against GSK3α, cellular extracts of uninfected, M. tuberculosis- and ΔptpA M. tuberculosis-infected cells will be immunoprecipitated, resolved on SDS-PAGE and proteins interacting with GSK3α will be identified by mass spectrometry. In the case that no stable complexes of GSK3α and interacting proteins form, a 2D SDS-PAGE assay will be carried out. In this instance, recombinant GSK3α will or will not be incubated with cellular extracts of uninfected and infected cells. Lysates will be incubated with anti-GSK3α antibodies. The immunocomplexes will be purified, resolved by 2D SDS-PAGE and phosphorylated proteins will be identified by mass spectrometry. Moreover, an antibody microarray using cellular extracts of uninfected, M. tuberculosis- and ΔptpA M. tuberculosis-infected cells and anti-GSK3α can be performed to yield insights into the differential binding of macrophage proteins to GSK3α and examine how these interactions are affected by PtpA. Lastly, the use of a specific and potent GSK3α inhibitor is another way to determine if GSK3α’s inhibition results in reduced caspase-3 cleavage and activation. The chemical compound lithium chloride, which selectively inhibits GSK3α, will be used.   In this work, we also showed that, in M. tuberculosis infection, PtpA dephosphorylates GSK3β (Figure 12, Table 4). GSK3α and GSK3β are two isoforms encoded by distinct genes. They share 98% homology in their kinase domain and perform similar functions in several settings (234). Despite their resemblance, GSK3α and GSK3β interact with proteins of their own and depending on the cell type, distinct functions of the isoforms have been demonstrated in cell survival (235-237). It would be interesting to examine whether dephosphorylation of GSK3β by PtpA also leads to inhibition of caspase-3 activation and if so, to what extent. One way to 99  determine the outcome of  GSK3β dephosphorylation will be by obtaining macrophages from GSK3α knockout mice, infect them with M. tuberculosis and the ΔptpA mutant (as described in Section 2.3.2), harvest the lysates (Section 2.3.3), and perform a Western blot analysis using anti-caspase-3 (Section 2.8). Knowing whether the inhibition occurs via GSK3α, GSK3β or both would allow us to look at the effects of this dephosphorylation on GSK3α- and GSK3β-specific pathways and understand the complex connections among all of the molecules involved in these pathways, as well as to understand how these work in M. tuberculosis infection.   Finally, we demonstrated both the global and PtpA-specific effect of M. tuberculosis on the macrophage proteomic response. On the one hand, we found that M. tuberculosis causes a general up-regulation of housekeeping proteins, such as those involved in transcriptional and translational regulation, as well as proteins involved in immunity and defence. On the other hand, we discovered that PtpA forces a down-regulation of proteins involved in cellular respiration, and, surprisingly, in immunity and defence. Based on these results, the number of new research avenues that can be pursued is vast.   One appealing area of research would be to investigate the global phosphoproteomic response of the host macrophage upon M. tuberculosis infection. A quantitative proteomic analysis approach, such as iTRAQ, would determine the phosphorylation status of thousands of proteins upon M. tuberculosis infection. Hypothetical knot research of modulated kinases would indicate downstream host signalling cascades affected by PtpA and the precise protein composition change of human macrophages infected with M. tuberculosis. Knowledge of the pathways affected would provide new information for comparative studies concerning 100  pathological states. Revealing downstream host signalling cascades affected by M. tuberculosis and, more specifically by PtpA, could have a major impact on drug discovery and development to prevent and better manage this notorious disease.  101  References 1. Ducati, R. G., Ruffino-Netto, A., Basso, L. A., and Santos, D. S. 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