A ROLE FOR PROTEIN KINASE G IN FOLATE METABOLISM AND INTRACELLULAR
SURVIVAL IN MYCOBACTERIA
by
KERSTIN ANDREA WOLFF
Submitted in partial fulfillment of the requirements
for the degree of Doctor of Philosophy
Dissertation Adviser: Dr. Liem Nguyen
Department of Molecular Biology and Microbiology
CASE WESTERN RESERVE UNIVERSITY
January 2012
2
Table of Contents
List of Tables 8
List of Figures 9
Acknowledgements 12
List of Abbreviations 13
Abstract 16
Chapter 1: Introduction 18
1.A. Tuberculosis and Other Mycobacterial Infections 19
1.A.1. Active and Latent M. tuberculosis Infections 20
1.A.2. Mycobacteria and Intrinsic Drug Resistance 23
1.A.3. MDR and XDR TB 27
1.B. Targeting Resistance Mechanisms to Potentiate Antibiotics 28
1.C. Folate Metabolism and Antifolate Antibiotics 33
1.C.1. Folate Antagonism in Chemotherapies and Antifolate
Resistance in Mycobacterial Infections 37
1.C.2. Potentiation of Antifolates in Mycobacteria 39
1.D. Biofilms 42
1.E. Protein kinase G (PknG) 44
1.F. MutT3 50
1.G. RplM 54
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Chapter 2: Materials and Methods 57
2.A. Bacterial strains, chemicals, media and growth conditions 58
2.B. Cloning, construction of plasmids and construction of strains 58
2.B.1. Targeted deletion of pknGMsm and mutT3Msm and
T11A T11E replacement of rplMMsm with rplMMsm or rplMMsm
in M. smegmatis by modified recombineering method 59
2.B.2. Targeted deletion of mutT3Mtb and replacement of
T11A T11E rplMMtb with rplMMtb or rplMMtb in M. tuberculosis
by specialized transduction 62
2.B.3. Deletion of ntpA in E. coli by one-step phage λ-Red-based
recombineering method 64
2.B.4. Cloning for in trans expression of proteins 65
2.C. RT-PCR 77
2.D. SDS-PAGE, Coomassie Brilliant Blue staining and Western Blot 77
2.E. Antibiotic Susceptibility Testing 79
2.F. Cellular aggregation, Hydrophobicity Index and Congo red
binding assay 80
2.G. Zeta Potential 80
2.H. Biofilm Assays 80
2.I. Kinase Assays 81
2.J. Radioactive nucleotide binding assays 82
2.K. Macrophage infection, lysosomal trafficking and microcopy 82
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Chapter 3: Protein Kinase G is Required for Intrinsic Antibiotic
Tolerance in Mycobacteria 84
3.A. Abstract 85
3.B. Introduction 86
3.C. Results 88
3.C.1. Identification of MAR4, an M. smegmatis pknG transposon
mutant displaying multidrug sensitivity 88
3.C.2. Targeted deletion of pknG and complementation
experiments confirm the MDR function of PknG in
pathogenic and non-pathogenic mycobacteria 89
3.C.3. Loss of pknG leads to altered cell surface properties 97
3.D. Discussion 101
3.E. Acknowledgements 103
Chapter 4: Modulation of Folate Metabolism by Protein Kinase G
Determines Biofilm Growth and Subcellular Localization of
Pathogenic Mycobacteria in Macrophages 104
4.A. Abstract 105
4.B. Introduction 106
4.C. Results 109
4.C.1. PknG is required for survival of folate starvation 109
4.C.2. Chemical inhibition of PknG activity potentiates classical
antifolate antibiotics 110
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4.C.3. PknG is required for biofilm growth 114
4.C.4. PknG is involved in regulation of lysosomal delivery in
macrophages via modulation of mycobacterial folate
metabolism 119
4.D. Discussion 123
4.E. Acknowledgements 126
Chapter 5: Protein Kinase G Regulates Folate Metabolism
Through Phosphorylation of Ribosomal Protein RplM 127
5.A. Abstract 128
5.B. Introduction 129
5.C. Results 132
5.C.1. PknG does not mediate regulation of folate metabolism
through phosphorylation of GarA 132
5.C.2. mutT3 is required for survival of folate starvation and
biofilm growth 133
5.C.3. Preliminary characterization of MutT3 as a Nudix
Hydrolase 137
5.C.4. RplM, a novel substrate for PknG, co-purifies with MutT3 144
5.C.5. Phosphorylation of RplM by PknG on conserved
threonine 11 mediates regulation of folate metabolism 153
5.D. Discussion 158
5.E. Acknowledgements 163
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Chapter 6: Discussion and Future Directions 164
6.A. Future Experiments 165
6.A.1. Hierarchy of PknG, RplM and MutT3 165
6.A.2. RplM point mutant alleles in M. tuberculosis and effect on
lysosomal trafficking 167
6.A.3. Subcellular localization of RplM/MutT3-containing
complexes 168
6.A.4. Biochemical characterization of MutT3 enzymatic
function 171
6.A.5. MutT3 deletion effect on lysosomal trafficking 173
6.A.6. Intracellular folate levels 174
6.B. Discussion 175
6.B.1. Folate significance during colonization of host
macrophages 175
6.B.1.a. Folate and cellular dormancy 176
6.B.1.b. Immune evasion and cell wall maintenance 178
6.B.2. Regulation of folate metabolism 182
6.B.3. Implications for TB and MDR or XDR TB 187
Bibliography 191
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List of Tables
Table 1-1. Tuberculosis drugs 26
Table 2-1. Primers used in this study 67
Table 2-2. Plasmids and phasmids used in this study 70
Table 3-1. Susceptibility of M. smegmatis strains to antibiotics 91
Table 3-2. Surface properties of M. smegmatis strains 99
Table 5-1. MutT3 co-purified proteins identified by LC/MS/MS 148
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List of Figures
Figure 1-1. A simple plateau model for persistent infection in
mice 22
Figure 1-2. Schematic structure of mycobacterial cell wall 24
Figure 1-3. Concept of drug potentiation by targeting resistance 30
Figure 1-4. Folate metabolism and antagonism in bacteria 34
Figure 1-5. Protein kinase G is involved in lysosomal delivery to
colonized phagosomes. 46
Figure 1-6. Chemical inhibition of PknG accelerates phagosome-
lysosome fusion 47
Figure 1-7. PknG-dependent trafficking of pathogenic
mycobacteria 49
Figure 1-8. Scheme of the 7,8 Dihydroneopterin Triphosphate
Pyrophosphatase (DNHTPase) Catalytic Reaction and
Effect of the ntpA Deletion on E. coli's Folate
Concentration 53
Figure 1-9. 3D-model of the E. coli 50S subunit showing the
position of each phosphorylated ribosomal protein. 56
Figure 3-1. Identification of MAR4, a M. smegmatis pknG
transposon mutant 90
Figure 3-2. Construction and complementation of the ΔpknGMsm
strain 95
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Figure 3-3. Antibiotic susceptibilities of M. tuberculosis strains 96
Figure 3-4. Loss of PknG does not affect growth rate in liquid,
shaking medium 98
Figure 4-1. Requirement of PknGMtb activity for survival of folate
starvation and exploitation for antifolate potentiation. 111
Figure 4-2. Requirement of PknGMsm activity for survival of folate
starvation. 113
Figure 4-3. PknGMtb is required for folate-dependent biofilm
growth in M. tuberculosis. 115
Figure 4-4. PknGMsm is required for folate-dependent biofilm
growth in M. smegmatis. 117
Figure 4-5. PknG is involved in lysosomal delivery to infected
phagosomes through folate levels. 120
Figure 5-1. PknG does not mediate control over folate
metabolism through phosphorylation of GarA. 134
Figure 5-2. Folate de novo biosynthesis pathway. 136
Figure 5-3. mutT3Mtb is required for survival of folate starvation
and biofilm growth. 138
Figure 5-4. mutT3Msm is required for survival of folate starvation
and biofilm growth. 140
10
Figure 5-5. ntpA, a dihydroneopterin-triphosphate
pyrophosphatase, is required for survival of folate
starvation. 143
Figure 5-6. MutT3 binds nucleotides in vitro. 145
Figure 5-7. RplM, a novel substrate of PknG, co-purifies with
MutT3. 149
Figure 5-8. MutT3 co-purified with RplM and is not regulated
translationally. 151
Figure 5-9. RplM is phosphorylated on threonine 11 by PknG in
vitro. 154
Figure 5-10. Phosphorylation of RplMMsm on threonine 11 is
required for survival of folate starvation and biofilm
growth. 156
Figure 5-11. Model of folate biosynthesis regulation via PknG,
RplM and MutT3. 161
11
Acknowledgements
Thank you to my adviser, Dr. Liem Nguyen, for taking a risk with me as his
first graduate student, even though I was the only student in my year that couldn’t
apply for additional funding, for opening the world of genetics to me and for being a
mentor in more than just science.
Hoa Nguyen, who has been with me on the roller coaster that has been this
project from the start, and who has become one of my closest friends, thank you.
Marissa Sherman, who has only recently joined our group but has quickly
become an essential part of my day (and the greatest cat sitter ever!), thank you.
Past and present members of our lab have helped me in many ways, thank you Joe Timpona, Sam Ogwang, Rich Cartabuke, Ajay Singh, and Soumya Gogula.
My committee members have been invaluable in steering me along my
graduate journey; thank you for your advice, your patience and your foresight, Dr.
Arne Rietsch, Dr. W. Henry Boom, Dr. Jonathan Karn, and Dr. Patrick Viollier.
I owe more to my parents than I could express, for letting me cross an ocean, for spending their money on American university fees, and for supporting me in every way. Thank you a million times.
My parents-in-law and their family, thank you for welcoming me with open arms and making me feel at home in the U.S., with or without my gall bladder.
Finally to my wife, Niki Smith, without whom I am nothing, and who, conveniently also solves every Adobe® Illustrator® problem I create. Thank you for
always bearing with me. I love you.
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List of Abbreviations
AIDS Acquired immune-deficiency syndrome
BCG Bacillus Calmette-Guérrin
BMDMs bone marrow derived macrophages
bp base pair
5-CHO-H4PteGlu folinic acid
5-CH3-H4PteGlu 5-methyl-tetrahydrofolate
CFU colony forming unit
dGTP deoxy-guanosine-5-triphosphate
DHFR dihydrofolate reductase
DHPS dihydropteroate synthethase
DOTS daily observed treatment schedule
EMB ethambutol
ETH ethionamide
FA folinic acid
FPLC fast protein liquid chromatography
GAIT interferon-γ-activated inhibitor of translation
GDH glutamate dehydrogenase
GDP guanosine 5’-disphosphate
GDP-manp guanosine 5’-diphosphate-mannose-1-phosphate
GFP green fluorescent protein
13
GLU glutamate
GTP guoanosine 5’-triphosphate
H2Pte dihyrdopteroate
H2PteGlu dihydrofolate
H4PteGlu tetrahydrofolate
Hcy homocysteine
HIV human immunodeficiency virus
HPLC high performance liquid chromatography
INH isoniazid
KEGG Kyoto Encyclopedia of Genes and Genomes
KGD alpha-ketoglutarate decarboxylase
LAM lipoarabinomannan
LB Luria-Bertani
LTBI latent tuberculosis infection
MAC Mycobacterium avium complex
MDR TB multiple drug resistant tuberculosis
mRNA messenger RNA
MS/MS tandem mass spectrometry
MTB Mycobacterium tuberculosis
MTHFS 5,10-methenyl-tetrahydrofolate synthase
MutT3 mutator protein T3
NTM Non-tuberculous mycobacterial infection
14
NtpA (dihydro)neopterin triphosphate pyrophosphohydrolase
OD# optical density at # nanometers
ODH 2-oxoglutarate dehydrogenase pABA para-aminobenzoic acid
PAS p-aminosalicylic acid pGp guanosine 5’,10’-bisphosphate
PI(3)P phosphoinositol-3-phosphate
PknG protein kinase G ppGpp guanosine 5’,10’-bispyrophosphate
RIF rifampicin
RplM ribosomal protein L13
SAM S-adenosine methionine
SHMT serine hydroxymethyltransferase
SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis
SSDB Sequence Similarity Database
(RT)-PCR (reverse transcription)-polymerase chain reaction
TB tuberculosis
TBS Tris-buffered saline
WHO World Health Organization
XDR TB extensively drug resistant tuberculosis
15
A Role for Protein Kinase G in Folate Metabolism and Intracellular
Survival in Mycobacteria
Abstract
by
KERSTIN ANDREA WOLFF
The eukaryotic-like serine/threonine protein kinase G (PknG) of
Mycobacterium tuberculosis has been shown to be important for survival of the bacillus in host macrophages, presumably by preventing lysosomal delivery. It was therefore suggested that PknG serves as a virulence factor, although the underlying mechanism has remained uncharacterized.
Here we present evidence that PknG regulates de novo folate biosynthesis that is required for static growth conditions including those found in surface biofilms and in host macrophages. Deletion of pknG resulted in defects in folate synthesis, growth under static conditions, as well as resistance of mycobacterial
16
species to classical antifolates and other antibiotics. We have identified a novel substrate for PknG, the ribosomal protein RplM of the large ribosomal subunit. RplM is phosphorylated by PknG at a unique, mycobacterially conserved residue, threonine 11, and co-purified with MutT3, a putative Nudix enzyme that exhibits a dihydroneopterin-triphosphate pyrophosphatase activity, catalyzing the first committed step of de novo folate biosynthesis. We demonstrate that inhibition of
PknG has the potential to be exploited for potentiating the anti-mycobacterial activity of classical antifolate drugs and we present evidence that the previously demonstrated role of PknG in blocking phagolysosomal synthesis in infected host macrophages is mediated through its function in mycobacterial folate metabolism.
Our work suggests that folate antagonism could be used to sensitize pathogenic mycobacteria to available chemotherapeutic reagents, as well as the innate bactericidal activity of host phagocytic cells.
17
Chapter 1.
Introduction
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1.A. Tuberculosis and Other Mycobacterial Infections
Mycobacterial infections are one of the leading causes of death in humans
through disease world-wide 1, encompassing infections such as tuberculosis (TB),
leprosy, Buruli ulcers, and opportunistic non-tuberculosis mycobacterial (NTM)
infections in immune-compromised individuals, especially patients with acquired
immune deficiency syndrome (AIDS) 2. These diseases are caused by Mycobacterium
tuberculosis (MTB), Mycobacterium leprae, Mycobacterium ulcerans, and environmental mycobacteria, such as Mycobacterium avium complex (MAC),
Mycobacterium kansasii, Mycobacterium chelonae and Mycobacterium fortuitum 2; 3, respectively. The World Health Organization (WHO) has estimated that one third of
the world’s population is currently infected with M. tuberculosis, the causative agent
of TB 1. Only about ten percent of those infected will develop active disease, unless
they are also afflicted with immune-compromising conditions such as AIDS that
highly increase the likelihood of active disease development 1. The lung typically is the affected organ, but the brain, kidneys, spine, and other body parts can also be infected. Mycobacterial loads in the affected organs and whether intracellular M. tuberculosis are dormant or actively growing depend on containment that must be successfully mediated by the host immune system (see section 1.A.1) 4; 5. This
process is severely impaired in immune-compromised individuals, and as a result there is greater opportunity for M. tuberculosis to transition from latency to active
TB in AIDS patients 4. Geographical areas with the highest incidence rates of TB are
located in sub-Saharan Africa and Southeast Asia, coinciding with human
19
immunodeficiency virus (HIV) hot spots, where up to 75 percent of HIV-infected
patients also are infected with M. tuberculosis 1. Accordingly, twenty-four percent of
all TB associated deaths occur in people co-infected with HIV 1.
1.A.1. Active and Latent TB Infections
Infections with TB are classified as either active or latent (LTBI) infections that differ significantly in infectiousness, symptoms and appropriate treatment regimens 6.
The vast majority of M. tuberculosis infections are LTBI cases, in which the
infected individual does not feel sick and does not exhibit any symptoms, since they do not develop active TB disease 6. Latent infection is diagnosed by either a positive
reaction to the tuberculin skin test or the QuantiFERON® blood test in combination with a negative chest X-ray. Patients with LTBI cannot spread M. tuberculosis to
others 6; 7. The daily observed treatment schedule (DOTS) for latent, drug susceptible TB infection consists of six to nine months of daily oral INH doses 8.
Latency in the bacillus is associated with reduced metabolic activity and slow or absent cell division and growth 4; 6; 11; 12.
About ten percent of latently infected persons will develop active TB at some time in their lives, with sharply increased likelihood of disease development by up to ten percent per year in immune-compromised patients, such as individuals co- infected with HIV 1; 6; 9. Fifty percent of individuals who develop active TB do so
within the first two years after initial infection 6. Symptoms of active TB disease
include weight loss, loss of appetite, fever and night sweats, fatigue and chills,
20
persistent coughing, hemoptysis, and chest pain 6. Active TB disease is diagnosed by chest X-ray and culturing of M. tuberculosis from patient sputum samples, along with positive tuberculin skin or QuantiFERON® blood tests. Patients with active TB
disease are infectious and spread M. tuberculosis by aerosolization when coughing 6.
The WHO estimates that a person with active TB who does not seek treatment will infect between ten and twelve people per year 1.
M. tuberculosis is internalized to phagosomes by host macrophages (see also section 1.E.), and like all intracellular pathogens must be able to survive and persist within infected host cells 10. The course of TB infections typically occurs in distinct phases 4; 5. The first phase following initial infection is a burst of active replication of
M. tuberculosis in the infected host tissue, followed by a phase of immune-mediated containment, in which a balance established between the host immune system and the pathogen leads to a constant, low bacterial load (Fig. 1-1) 4; 5. During this stage,
M. tuberculosis transitions from active growth to a dormant form, which is
metabolically distinct 4; 6; 11; 12. In this persistent state, M. tuberculosis becomes
highly tolerant to antibiotics compared to actively dividing cells 13; 14; 15; 16.
While the process by which M. tuberculosis establishes successful persistence is poorly understood, it has been shown that both host and mycobacterial factors are involved in phagosome maturation of infected host macrophages 17; 18; 19. More recently, mycobacterial protein phosphatases and kinases have been identified to perform key roles in establishing the survival niche in mature, but non-lysosomally fused phagosomes 19; 20; 21. One such kinase is the mycobacterial eukaryotic-like
21
Figure 1-1. A simple plateau model for persistent infection in mice. Low-dose infection of C57BL/6 mice with M. tuberculosis results in initial active replication in the lung (1st month), followed by immune-mediated containment, and a persistence phase that is characterized by a constant bacterial load. This provides a convenient model to study the molecular basis of mycobacterial and host responses to persistent infection. Parallel studies with human tissues are required to determine the extent to which persistence in mice resembles latent tuberculosis. CFU, colony- forming units.
Reproduced with permission from 4.
22
serine/threonine protein kinase G (PknG), which was suggested to support survival
of pathogenic mycobacteria in the host macrophage by preventing phagolysosomal fusion and therefore acidification of the mycobacterial compartment 20 (see also section 1.E.). Finally, in patients who develop active tuberculosis disease, immune-
mediated containment fails, often due to other factors weakening the immune
response, e.g. HIV infection, other diseases or aggressive immune-suppressing
therapy. As a consequence, the bacilli transition from the latent state to active
growth and expand in the affected tissue, causing symptoms and making spread to
other individuals possible 5. Interestingly, in yet another case of phenotypic
association between antibiotic tolerance and virulence 22; 23; 24, the transition from active division to the persistent state can be triggered by exposing M. tuberculosis to antibiotics 25.
1.A.2. Mycobacteria and Intrinsic Drug Resistance
Although mycobacteria are classified as Gram-positive bacteria, which do not have an outer membrane on top of their peptidoglycan layer, the mycobacterial cell wall is extremely thick and multi-layered, with varied hydrophobicity among the layers, and thereby poses an effective obstacle for the entry of most chemical compounds (Fig. 1-2) 3; 23; 26; 27. The peptidoglycan network is covered by an arabinogalactan layer, and both of these components are hydrophilic and likely limiting for the penetration of hydrophobic compounds (Fig. 1-2) 3. A layer of long chain fatty acids called mycolic acids of about seventy to ninety carbon units in length linked to acyl lipids overlays these two layers and forms a waxy, non-fluid
23
Figure 1-2. Schematic structure of mycobacterial cell wall. (a) Schematic
structure of mycobacterial cell wall depicted according to Minnikin's model 28.
Intercalation of hydrophilic arabinogalactan and hydrophobic mycolate containing layers creates an extremely impermeable envelope for antibiotic penetration. Small molecules and nutrients are transported through porin channels that are deposited through these layers. Modified from 29.
Reproduced with permission from 23.
24
barrier that is very efficient at restricting transport of both hydrophobic and
hydrophilic molecules (Fig. 1-2) 26; 30. As a result, mycobacteria are highly
problematic pathogens resistant to most of the typically used antibiotics and other
chemotherapeutic agents, even to extents greater than seen in Gram-negative
bacteria, simply by virtue of their complex cell envelope 3; 23. For example, the cell
wall permeability for hydrophilic compounds measured in Mycobacterium chelonae
was roughly three orders of magnitude lower than that of the Escherichia coli outer
membrane, and ten times lower than the permeability measured for the outer
membrane of Pseudomonas aeruginosa 31.
While permeability and slow influx across the mycobacterial cell wall is a necessary factor for the high intrinsic resistance of mycobacteria to most common antibiotics, it is not sufficient and usually must be supported by dedicated resistance mechanisms operating in the cytosol 3. Apart from several drug degradation and inactivation processes that occur inside mycobacteria, they also possess over 30 drug efflux transporters that have been shown to be involved in resistance to aminoglycosides, chloramphenicol, fluoroquinolones, isoniazid, linezolid, rifampicin, tetracycline and other toxic compounds 32; 33; 34; 35.
Cumulatively, the extremely high level of intrinsic resistance to most
antimicrobial drug classes exhibited by M. tuberculosis has left us with a severely
limited arsenal of useful anti-tuberculosis drugs (Table 1-1) 36. Complicating matters further, these antimycobacterial drugs must be used in regimens consisting of a minimum combination of four drugs to be effective for treatment of active
25
Table 1-1. Tuberculosis drugs.
Antibiotics Year of discovery
First-line antibiotics Isoniazid 1952 Rifampin 1966 Pyrazinamide 1952
Ethambutol 1961
Rifabutina 1980
Rifapentinea 1965
Second-line antibiotics Cycloserine 1952 Ethionamide 1956 P-aminosalicylic acid 1946
Streptomycin 1944 Amikacin 1972
Kanamycina 1957
Capreomycin 1963
Levofloxacina (Levaquin, DR3355, Daiichi) 1986
Moxifloxacina (Avelox, BAY 12-8039) 1996
Gatifloxacina (Tequin, AM-1155) 1992
Antibiotics Year of discovery
aNot approved by the U.S. Food and Drug Administration (FDA) for TB treatment 11.
Reproduced with permission from 36.
26
tuberculosis disease 36, making TB and NTM infections one of the most difficult
human disease groups that are caused by bacteria to treat at the current time.
1.A.3. MDR and XDR TB
The five first-line drugs used to treat M. tuberculosis infections, isoniazid
(INH), rifampicin (RIF), ethambutol (EMB), pyrazinamide, and streptomycin were
all discovered more than sixty years ago (Table 1-1) 23; 36. They have been used
regularly since then and therefore are encountered by M. tuberculosis strains
constantly 23; 36. Furthermore, the current standard regimen (DOTS) for TB is
comprised of six to nine months of daily antibiotic treatment with a combination of
four out of these five drugs 23; 36. These lengthy regimens often lead to poor patient
adherence and incomplete courses of treatment, resulting in frequent exposure of M.
tuberculosis to sub-optimal doses of drugs 23; 36. This provides ample opportunity for the bacillus to acquire additional resistance by amassing sequential mutations in drug-target encoding genes 23; 36; 37. Due to the rapid mutational frequency
characteristic of bacteria, we now face the increasingly wide spread of multiple drug
resistant (MDR) and extensively drug resistant (XDR) M. tuberculosis strains 23; 36; 37.
MDR TB strains exhibit acquired resistance to at least the two most potent first-line
drugs (RIF and INH) 1; 36; 37; 38. XDR TB strains are resistant to any fluoroquinolones and to at least one of the three injectable second-line drugs (capreomycin, kanamycin, and amikacin) (Table 1-1) in addition to RIF and INH resistance 1; 36; 37; 38.
Infections with such strains require even more prolonged and aggressive treatment
courses consisting of combinations of numerous second-line drugs that often exhibit
27
toxic side effects and are expensive to administer 36; 39. An example can be found in the South-African province KwaZulu-Natal, where extraordinarily high rates of
HIV/AIDS and MDR TB coincide. In this region poor patient compliance to treatments requiring between 18 and 24 months of hospitalization for drug administration has led to an outbreak that accounts for one sixth of the worldwide
XDR TB cases 40. Roughly eighty percent of these active XDR TB patients are also
infected with HIV 40. During the 2006 outbreak, the mean patient survival time from sputum collection amounted to only 16 days 40. Per month, about 30 new cases of active XDR TB are diagnosed in KwaZulu-Natal alone 40.
As the spread of these infections is diminishing our already limited arsenal of effective antibiotics even further, the situation is becoming more and more problematic, and we desperately require new, alternative drugs 37; 41; 42; 43. Indeed,
some XDR M. tuberculosis strains have become virtually untreatable with the
currently available anti-TB medicines 41; 42; 43.
1.B. Targeting Resistance Mechanisms to Potentiate Antibiotics
The current prevalence of drug-resistant mycobacterial strains has created a dire need for alternative TB therapies to use in patients infected with MDR or XDR
TB. However, development of completely novel anti-tubercular drugs is both time- intensive and costly; roughly twelve to fifteen years and US $500 million are required to facilitate development of a new drug from the laboratory stage to its distribution on the market 44. Despite massive investments of effort and capital into
28
this approach, only a few compounds have proceeded into pre-clinical or clinical
stages, and none have been newly approved for treatment.
An alternative approach to this pathway that may help relieve the current crisis is presented by the concept of “targeting resistance” 45. This drug potentiation
approach makes use of our knowledge of molecular resistance mechanisms in order
to (re)sensitize pathogenic bacteria to already available drugs 45. This is achieved by coadministration of existing drugs and inhibitors suppressing resistance mechanisms to these drugs, thereby allowing ineffective drugs to (re)gain their antimicrobial activity (Fig. 1-3) 45; 46; 47. Because we are currently using only a limited number of drugs due to existing resistance in M. tuberculosis, potentiation by targeting resistance may become an important trend in the new era of drug development for TB. Drug potentiation could be used to regain and extend current
TB drug efficacy, as well as to make available for use other approved drugs that are currently inactive against the bacillus 45. The extension of lifespan and utility of
valuable approved antibiotics of known pharmacology, toxicology, and treatment
schedule represents a unique advantage of the drug potentiation approach 45.
Recent findings indicate that several available drug classes encompassing
both TB and non-TB antibiotics are promising candidates for potentiation by
targeting resistance mechanisms 45. Efforts are underway exploring this approach using β-lactams, ethionamide, and classical antifolates 45.
29
Active Resistance ANTIBIOTIC Mechanism
Resistant Bacterium
ANTIBIOTIC
Susceptible Bacterium
POTENTIATOR Inactive Resistance Mechanism
Figure 1-3. Concept of drug potentiation by targeting resistance. An active
resistance mechanism allows survival of bacterial pathogens in the face of
antibiotics. A potentiator that inhibits the resistance mechanism would (re)sensitize
the bacteria to the antibiotic, thus enhancing antibacterial activity.
Reproduced with permission from 45.
30
β-lactams are the most widely used group of antibiotics today. They make up
a broad class of drugs including penicillin and penicillin derivatives, cephalosporins,
monobactams and carbapenems, that targets bacterial cell wall synthesis at the peptidoglycan layer 48; 49. Currently, β-lactams are the only successful clinical
example of potentiation through targeting resistance, but inhibitors of β-lactamases
have prolonged the life of β-lactams for more than thirty years and are commonly co-administered with β-lactams in clinical use 50. As mycobacterial susceptibility to
β-lactams in the absence of β-lactamase activity, in the case of M. tuberculosis this
activity is exhibited by BlaC, is well within clinically achievable levels 51; 52, effective chemical inactivation of β-lactamases should similarly increase β-lactam sensitivity in these bacteria. Indeed, recent in vitro studies have shown three FDA-approved inhibitors to effectively inhibit purified BlaC protein 53; 54. Further in vitro data show
that a combination of meropenem/clavulanate kills both aerobically and
anaerobically grown M. tuberculosis with great efficiency 55, is effective against
thirteen tested XDR M. tuberculosis strains 55, and significantly reduces
mycobacterial counts in mouse peritoneal macrophages infected with M. tuberculosis upon treatment within clinically achievable dose ranges 56; 57 . Another study demonstrated imipenem to double the survival rate of infected mice 58 and one human study using imipenem alone in MDR TB patients with poor predicted
outcomes achieved a 70% cure rate as well 58. Treatment of TB patients with an amoxicillin/clavulanate combination significantly reduced M. tuberculosis counts with early bactericidal activity comparable to patients treated with the frontline
31
drug INH in an independent study 59, and perhaps most significantly, an advanced
XDR TB patient was treated with meropenem/clavulanate in conjunction with other
drugs after all previous other treatments had failed and has since fully recovered 60.
Ethionamide (ETH) is one of the most commonly used drugs in the treatment of MDR TB, but ETH usage is complicated by its low therapeutic index 61; 62. This
index signifies a narrow concentration range in which ETH is therapeutically
effective yet safe to use without eliciting toxic side effects in the patient 61; 62; 63. The
minimal dose required for inhibition of M. tuberculosis growth causes adverse side
effects including hepatitis and gastrointestinal conditions 64. ETH is a structural thioamide analogue of INH that must be metabolically activated by EthA to form adducts in reaction with nicotinamide adenine dinucleotide (NAD) that target synthesis of mycolic acids, the major mycobacterial cell wall component, by inhibiting InhA 65; 66; 67; 68; 69; 70; 71; 72. EthA is transcriptionally repressed by EthR,
leading to lowered activation and efficacy of ETH, and therefore, targeting EthR by inhibitors to achieve derepression of EthA transcription should sensitize mycobacteria and potentiate the antimycobacterial activity of ETH 64; 70; 73; 74.
Hydrophobic ester compounds can bind the DNA-binding domain of EthR and prevent binding of EthR to the ethA operator 73; 74; 75; 76. When tested as potentiators in combination with ETH, these esters were shown to significantly increase the effects of ETH in in vitro cultures 73; 74; 75; 76. An intelligently designed ester based on a lead compound identified in a screen for EthR inhibitors, BDM31343, allows for twentyfold lower ETH doses to inhibit growth of M. tuberculosis in vitro and
32
threefold lower ETH doses to significantly reduce bacterial loads in lungs of infected
mice 64; 74. Potentiation of ETH by hydrophobic ester inhibitors of EthR could therefore not only re-sensitize XDR TB strains to ETH, but also allow for elimination or reduction of toxic side effects in MDR TB patients due to lower required doses 45.
Evidence obtained from numerous studies performed in vitro, in animals, and
in humans, has clearly illustrated that β-lactams potentiated by β-lactamase
inhibitors and ETH potentiated by EthR inhibitors could be used as an effective
addition to the treatment of drug resistant TB. This powerfully demonstrates the
potential value of potentiation by targeting resistance as a tool for relieving the need
for alternative options in TB treatment 45.
1.C. Folate Metabolism and Antifolate Antibiotics
One group of antibiotics that seems promising for potentiation in mycobacterial infections is the antifolates, which target folate biosynthesis and metabolism 45. The term folate is used to refer to a number of B vitamins of similar
but distinct chemistry, all of which are absolutely essential for the existence of
cellular organisms in all kingdoms of life 45. Most naturally occurring folates are
derivatives of tetrahydrofolate (H4PteGlu, Fig. 1-4), a reduced folate compound, but
the commonly used nutritional supplement vitamin B9 or folic acid is chemically
synthesized 45. While all folates consist of three molecular components, namely a
two-ring pteridine nucleus derived from guanosine-5’-triphosphate (GTP), a para-
aminobenzoic acid (pABA) group, and one or more glutamate residues attached via
33
Figure 1-4. Folate metabolism and antagonism in bacteria. (A) Simplified interconversions of folate derivatives in de novo folate synthesis and one-carbon metabolic network. DHFS, dihydrofolate synthase; Gly, glycine; Met, methionine; MT, methionine synthase; Pte, pteroate; Ser, serine. (B) Chemical structure of monoglutamylated tetrahydrofolate and its derivatives carrying C1 groups at various levels of oxidation attached to N-5 or/and N-10.
Reproduced with permission from 45.
34
amide linkages, they are varied in the nature of the one-carbon group attached to
45 the N-5 and/or N-10 positions of H4PteGlu (Fig. 1-4B) .
Folates are critically important for all cellular activities because H4PteGlu derivatives are required as co-enzymes in one-carbon unit transfer reactions (Fig. 1-
4A) 45. These reactions are required for the biosynthesis of purines, thymidines, glycine, panthotenate and methionine. Additionally, in bacteria, folates also act as co-enzymes for synthesis of formyl-methionyl-tRNA, critical for initiation of translation 77; 78; 79. Therefore, folate molecules are essential for the synthesis of the building blocks of macromolecules such as nucleic acids and proteins. Consequently, folate deficiency interferes with almost all processes in actively dividing cells and ultimately leads to cell death 45. Yet another effect of folate starvation manifests itself in interference with recycling of both homocysteine (Hcy, Fig. 1-4) and S- adenosine methionine (SAM) 45. This elevates cellular Hcy concentrations, a condition called homocysteinemia, and reduces cellular methylation activities 45. As a result, folates are of particular importance during periods of rapid cell division and growth 77; 78.
Folate metabolism is regarded as bipartite, with a clear division between de novo biosynthesis (upstream) and utilization (downstream) of folates (Fig. 1- 4A).
De novo folate biosynthesis is a process comprised of several steps: (i) synthesis and modification of the pteridine group from guanosine-5’-triphosphate, (ii) synthesis of pABA from chorismate, (iii) condensation of pteridine and pABA to form dihydropteroate (H2Pte), and (iv) glutamylation which adds one or more glutamate
35
45 groups to form dihydrofolate (H2PteGlu) that is reduced to form H4PteGlu . This de novo biosynthesis pathway is present in most bacteria and plants, but absent in many other species, including mammals 45. The downstream utilization of folate, also called one-carbon metabolism, involves co-enzyme action of the various active forms of H4PteGlu in distinct reactions donating or accepting one-carbon units for
the formation of purines, thymidines, glycine, panthotenate, methionine, and
formyl-methionyl tRNA, as mentioned above (Fig. 1-4) 45; 77; 78; 79.
It is not surprising that compounds as essential to the vast majority of cellular processes as the folates were soon recognized as promising candidates to target in chemotherapeutic treatments of many conditions and diseases and were one of the first antibiotics invented 45. Folate antagonism has been exploited
successfully for therapies of cancers, malaria, psoriasis, rheumatoid arthritis, graft- versus-host disease, and bacterial infections 80; 81; 82. Folate antagonists, called antifolates, were initially used extensively to treat a variety of infectious diseases, but their use has been scaled back since the 1960s 80; 83. This is due in part to emergence of resistant strains, to the cytotoxicity of antifolates that can cause side effects in the patient, and most importantly due to the introduction of new drugs more effective against certain pathogens and diseases 80; 83. Other infectious
diseases, for example urinary tract infections, pneumonia caused by Pneumocystis
jiroveci, and shigellosis, are still commonly treated with a combination therapy
consisting of trimethoprim and sulfonamide antifolates that makes use of the
synergy between the two drug classes 83; 84; 85. Furthermore, the same combination
36
regimen is also used effectively as a prophylactic treatment for recurring and drug-
resistant infections 83; 84; 85.
While there are a multitude of well-studied proteins contributing to the
processes of folate metabolism, currently used folate antagonists seem to
exclusively target biosynthesis and reduction of folate 86; 87; 88. Whereas trimethoprim and folate analogs such as methotrexate inhibit the reduction step through inhibition of dihydrofolate reductases (DHFR), sulfonamides and sulfone drugs are pABA analogs that outcompete pABA in the condensation with the pteridin group, catalyzed by dihydropteroate synthase (DHPS) (Fig. 1-4) 86; 87; 88. A factor that makes antifolates a particularly attractive group of antibiotics is the absence of these targeted enzymes acting in de novo folate biosynthesis in humans and other mammals that must rely on their diet for folate intake, making antifolates a group of drugs that will specifically affect the bacterial pathogens, but not infected host cells 86.
1.C.1. Folate Antagonism in Chemotherapies and Antifolate Resistance in
Mycobacterial Infections
Since both steps in de novo folate biosynthesis and downstream folate utilization have been targeted by successful antimicrobial and antineoplastic
antifolate drugs since the 1940s, acquired resistance to antifolates in pathogenic
bacteria is by now wide-spread 45. This resistance is typically acquired by amassing
mutations that alter either expression levels or protein structures of the targeted
enzymes 80; 83. DHFR can become resistant to trimethoprim through point mutations
37
of active-site residues, thus altering its affinity for trimethoprim 89; 90. While clinical
resistant strains frequently show a variety of mutations, mutations of residues
crucial for trimethoprim affinity are highly conserved among the isolates 89. For
example, a point mutation in the DHFR gene that changes a conserved Isoleucine
residue (Isoleucine 94 in M. tuberculosis DHFR) to Leucine confers 50-fold higher
trimethoprim resistance in Streptococcus pneumoniae 89. This same mutation is commonly found in DHFR from mammalian, parasitic and bacterial resistant isolates
89; 90. Likewise, single point mutations at the Serine 53 or Proline 55 residues within
DHPS are found in resistant isolates of M. leprae and render the enzyme resistant to
sulfonamides and sulfone drugs 91; 92. The two residues are located in the drug
binding region of M. tuberculosis DHPS and are highly conserved throughout bacteria and protozoa 91; 92. Since only these two enzymes are currently being
targeted, combined mutations in DHFR and DHPS encoding genes confer resistance
to all available antifolates 86; 87; 88. Most studies of trimethoprim and sulfonamide resistance thus far use bacteria distantly related to M. tuberculosis. Due to recent increases in the use of antifolates as prophylactic treatments in HIV patients 93, many of whom are co-infected with M. tuberculosis, the bacillus is bound to encounter antifolates regularly and therefore a better understanding of antifolate resistance mechanisms in M. tuberculosis is urgently needed 94.
Despite this lack of intricate knowledge concerning resistance mechanisms in
mycobacteria, antifolate drugs have been used in the treatment of mycobacterial
infections. For example, PAS (p-aminosalicylic acid) is currently used as a second-
38
line drug for TB 95 and the sulfone drug Dapsone has long been effective in
monodrug regimens for the treatment of leprosy 96. More recently, a study
suggested that the frontline TB-drug INH, which is not a classical antifolate
antibiotic, may also target folate metabolism through DHFR inhibition by its adducts
97. With the current desperate need for alternative treatment options of MDR and
XDR TB infections, interest in the potential use of combination regimens of classical antifolates against M. tuberculosis has been renewed 98; 99; 100. As a result, several
recent in vitro studies and a case report suggest antifolate combinations such as
those of co-trimoxazole (trimethoprim plus sulfamethoxazole) could be effective
against TB 98; 99; 100. M. tuberculosis clinical strains isolated from TB patients were
shown to be widely susceptible to clinically achievable concentrations of co-
trimoxazole 98, and even to sulfamethoxazole alone 99. However, since exposure of M.
tuberculosis to antifolates is likely to rise due to the World Health Organization’s call
for widespread use of co-trimoxazole in the prophylactic treatment of HIV-AIDS
patients to prevent opportunistic infections, selection for resistant strains may soon
occur and prematurely render this promising class of drugs useless 82; 93. It would
therefore be prudent to seek ways to potentiate classical antifolate antibiotics by
targeting resistance mechanisms, similar to the approach used for β-lactams, so that
M. tuberculosis strains that acquire antifolate resistance could be re-sensitized and
remain treatable by this group of drugs 45.
1.C.2. Potentiation of Antifolates in Mycobacteria
39
A method for boosting antifolate efficacy by utilizing combinations of drugs
that target individual steps in folate biosynthesis is already in place (see section
1.C.1). Trimethoprim is commonly coadministered with sulfonamides, for example
sulfamethoxazole in the co-trimoxazole combination, in order to achieve a
synergistic effect and increase efficacy 83 (Fig. 1-4). However, in many cases
including that of M. tuberculosis, the synergistic effect of trimethoprim on
sulfonamides remains questionable and has not been proven conclusively 98; 99; 101.
More problematically, bacterial strains have already acquired resistance mutations, making them untreatable with both trimethoprim and sulfonamides 86; 87; 88.
Therefore, novel potentiation approaches targeting resistance mechanisms not rooted in point mutations of DHFR and DHPS might be more effective in both potentiating available antifolates and preventing additional emergence of resistant strains.
A recent study aimed at targeting intrinsic antifolate resistance in mycobacteria has the potential to reveal valuable targets for such resistance- targeted potentiation approaches 102. To systematically identify novel antifolate
resistance determinants, a genetic screen was first employed using a saturated
transposon-insertion library of M. smegmatis 102. The mutants are tested for
increases in susceptibility to a number of classical folate antagonists to identify
genes that encode for antifolate tolerance contributing proteins 102. Hits are confirmed by chemical complementation using folate derivatives of both the de novo synthesis and the one-carbon interconversion network 102. This chemogenomic
40
profiling approach allows for identification of novel determinants previously
unknown to function in mycobacterial intrinsic antifolate resistance and identified fifty such components of the mycobacterial antifolate resistome (unpublished data)
102. One of these identified genes was later confirmed to encode for a protein that acts as a 5,10-methenyl-tetrahydrofolate synthase (MTHFS) homolog, and disruption of this gene resulted in up to 64-fold lower resistance to various of the tested trimethoprim/sulphonamide combinations 102. A series of genetic knockout
and complementation studies, including the human MTHFS allele, indicated that
MTHFS activity is required for mycobacterial intrinsic antifolate resistance, despite
there being no function for MTHFS in the typically targeted pathways of folate de
novo biosynthesis or folate reduction 102. The role of MTHFS in intrinsic antifolate
resistance was confirmed in M. tuberculosis and Escherichia coli 102; 103, suggesting
that this determinant functions ubiquitously among bacteria. Pharmaceutical
inactivation of MTHFS activity should therefore result in sensitization of M.
tuberculosis to classical antifolates, potentially including current anti-TB drugs that
target folate metabolism (PAS, INH, etc.). Potentiation of the anti-TB efficacy of
classical antifolates may also allow for reduction of effective therapeutic doses,
thereby minimizing their cytotoxicity that has been a clinical problem.
Considering these promising reports concerning the potential of making
classical antifolate antibiotics a useful, readily available alternative to the aggressive
and often not effective treatments required in MDR and XDR TB infections 36; 39, fundamental studies of molecular mechanisms conferring both acquired and
41
intrinsic antifolate resistance in M. tuberculosis and related mycobacteria need to be conducted 45. The knowledge obtained from these studies will be essential for
strategic implementations of antifolate use for TB, and will reveal valid targets for
the resistance-targeted potentiation of classical antifolates 45.
1.D. Biofilms
Biofilms are surface-bound communities of microbial cells that can consist of
one or multiple bacterial or fungal species and occur both in natural environments,
such as lake water, and in clinical settings, for example on catheters in patients or on
the surface of implants 104; 105; 106; 107. Bacteria growing in biofilms display enhanced
tolerance for antibiotics, a phenomenon that can be ascribed to the fact that biofilm
growth is metabolically distinct from planktonic growth, resembling static-phase or
persistent growth stages 104; 105; 106; 107. In accordance with the idea of separate metabolic requirements, there is evidence that the stringent response, a stress response to amino acid starvation that is triggered by exposure to antifolates, affects biofilm formation 108; 109.
Biofilm development is typically modeled in four distinct phases, consisting of (a)
attachment of planktonically growing cells to a surface, (b) secretion of polysaccharides,
protein and extracellular DNA resulting from autolysis that then form an extracellular
matrix in which cells aggregate, (c) biofilm maturation, and (d) biofilm dispersal 104.
While the most extensively studied and best understood bacterial biofilm
models pertain to Escherichia coli, Vibrio cholerae, Bacillus subtilis, Pseudomonas
aeruginosa and Staphylococcus epidermis 104; 110, mycobacteria, including M.
42
tuberculosis, have been shown to form biofilms both in vivo and in vitro 111; 112; 113; 114.
Mycobacteria differ from most other biofilm-forming bacterial species in that they
do not externalize polysaccharides to form an extracellular matrix, but rather
aggregate through short-chain and free mycolic acids of their waxy cell envelope 112;
113. In other aspects however, mycobacterial biofilms behave just like those of other
pathogenic bacteria and have been shown to play a role in the pathogenesis of
Mycobacterium chelonae and Mycobacterium ulcerans 114; 115; 116; 117. Mycobacterial
biofilms also exhibit higher antibiotic tolerance than planktonic mycobacteria,
which is consistent with the idea that biofilm growth may represent a late static
growth stage similar to persistence, as M. tuberculosis in its persistent state during
latent infection also exhibits higher levels of resistance than actively growing M.
tuberculosis 22. Conversely, exposure to antibiotics is one of the factors inducing the
transition from active growth to persistence 22, and indeed a recent study in M. tuberculosis has revealed a correlation of biofilm growth and the development of persisters with increased antibiotic tolerance that provide an important reservoir of cells that can repopulate colonized sites upon removal of drug treatment 108; 113; 118.
The details of mycobacterial biofilms are not well understood, but it has been
shown that alterations in cell wall characteristics and composition, as well as in
sliding motility, result in defects in biofilm formation in both M. tuberculosis and M.
smegmatis 113; 119; 120. Furthermore it has been demonstrated that M. tuberculosis
requires OdhA, the E1 subunit of the 2-oxoglutarate dehydrogenase (ODH) complex,
also known as the α-ketoglutarate dehydrogenase (KGD) complex, an enzyme
43
complex of the citric acid cycle, for biofilm formation 113. Interestingly, the enzymatic activity of ODH is controlled by the phosphorylation state of OdhI and its homolog GarA in Corynebacterium glutamicum and M. tuberculosis, respectively 121;
122; 123. OdhI/GarA can be phosphorylated by the eukaryotic-like serine/threonine
kinases PknG and PknB in both M. tuberculosis and in the related actinobacterium
Corynebacterium glutamicum 121; 122; 124. Deletion of Rv2454c and Rv2455c, the two
genes encoding for the subunits of ODH complex, leads to defects in biofilm
formation, which could be bypassed by exogenous supplementation of CO2 113.
1.E. Protein kinase G (PknG)
Protein kinases can be sub-divided into 3 groups defined by the residue to which the phosphate group is transferred. Histidine kinases typically are part of two-component signaling systems, where they first autophosphorylate on a histidine residue upon reception of a stimulus, and then proceed to transfer the phosphate group onto the aspartic acid residue of a response regulator 125; 126. These two-component systems are mainly found in bacteria and eukaryotes outside the
animal kingdom 125; 126; 127. Tyrosine kinases have evolved separately and use
distinct mechanisms of phosphorylation in bacteria and eukaryotes 128. Third,
serine/threonine kinases exist both in eukaryotes 129 and in bacteria, where they are
called eukaryote-like or eukaryotic-type serine/threonine kinases 130. While their importance in signaling has been extensively studied for many years in eukaryotes
131, our understanding of their roles in bacteria has only recently begun to expand.
Eukaryotic-type serine/threonine kinases have been found to play roles in
44
pathogenicity, stress responses and developmental processes in various bacteria 130;
132.
M. tuberculosis contains 11 putative eukaryotic-type serine/threonine
protein kinase-encoding genes in its genome 133; 134. For many of these kinases either function or substrates remain unknown 10, but it was soon found that kinase
inhibitors prevented uptake by and infection of murine macrophages by
Mycobacterium leprae, indicating crucial roles in pathogenicity 135.
While the vast majority of mycobacterial serine/threonine kinases exhibit typical receptor-kinase structures with a trans-membrane domain and an intracellular kinase domain, two kinases, PknG and PknK, appear to be soluble rather than membrane-associated 132; 133; 134.
PknG is conserved throughout the mycobacterial genus, including M. leprae,
which indicates that it plays an important role in bacterial survival in the host cell,
since M. leprae is dependent on its host and therefore has reduced its genome to a
minimum set of genes that are absolutely required for intracellular survival 136. It
has been shown that PknG plays a critical role in M. tuberculosis’ ability to establish
the phagolysosomal fusion block inside host macrophages, although its mechanism
of action and targets involved in this process remain unknown 20; 137. Both genetic deletion of pknG and chemical inhibition of PknG by a specific inhibitor 20; 138 in M.
bovis BCG resulted in increased fusion of lysosomes to phagosomes containing
mycobacteria, correlating with decreased survival of internalized mycobacteria (Fig.
1-5, 1-6) 20; 23. It was therefore suggested that PknG may act as a virulence factor
45
Figure 1-5. Protein kinase G is involved in lysosomal delivery to colonized phagosomes. Macrophages were infected with M. bovis BCG or M. bovis BCG/ΔpknG
expressing green fluorescent protein (GFP) for 1 hour, followed by a 2-hour chase.
Cells were processed as described in supplementary materials online and stained
with antibody to LAMP followed by Alexa Fluor 568 (LAMP-568). Scale bar, 10 μm.
Reproduced with permission from 20.
46
Figure 1-6. Chemical inhibition of PknG accelerates phagosome-lysosome fusion. Macrophages were infected with M. bovis BCG for 1 hour, followed by a 2- hour chase, in the absence (left) or presence (right) of AX20017 (20 μM), fixed, permeabilized, and stained with antibody to LAMP followed by Alexa Fluor 568.
Scale bar, 10 μm.
Reproduced with permission from 20. 47
that is secreted into the host cytosol and actively modifies host molecules to
establish the phagolysosomal fusion block 20; 23; 139. It has also been suggested that
this virulence function may have been acquired in pathogenic mycobacteria that
regulate PknG expression translationally to levels dramatically elevated over those
seen in non-pathogenic mycobacterial species, and that virulence function depends
upon expression levels 140.
Additionally, recent findings confirmed that, as had been shown for
Corynebacterium 121; 123, PknG is involved in the tricarboxylic acid cycle as well as glutamine utilization in Mycobacterium, where it phosphorylates GarA on threonine
21, directly adjacent to threonine 22, which is phosphorylated by PknB 122; 124; 141.
The two phosphorylation events are mutually exclusive 121; 122; 124; 141. GarA in its unphosphorylated state inhibits action of ODH complex and, depending on substrate concentration, either inhibits or activates glutamate dehydrogenase (GDH) 122. ODH converts 2-oxoglutarate to succinate, generating CO2, which has been shown to play a role in biofilm formation in mycobacteria 113. GDH converts 2-oxoglutarate to glutamate. Glutamine and its derived molecules, such as polyglutamines, may serve as components that are required for the integrity of the mycobacterial cell wall, thereby helping to pose an effective barrier to diffusion of antibiotics of diverse size and polarity, and are also important for static-phase growth 122; 132; 141.
PknG can autophosphorylate on four N-terminally located threonine residues, threonines 23, 32, 63 and 64, which are distinct from the typically
48
Figure 1-7. PknG-dependent trafficking of pathogenic mycobacteria. (a) Model of PknG-mediated phagosome-lysosome fusion blockage by pathogenic mycobacteria. PknG (purple) is secreted by pathogenic mycobacteria into the phagosomal lumen and cytosol of macrophages to block the fusion of phagosomes to lysosomes. Deletion of the pknG gene leads to rapid lysosomal transfer and destruction of the resulting mutant.
Reproduced with permission from 23.
49
autophosphorylated residues located in a conserved activation loop in other
mycobacterial serine/threonine kinases 142. Consistent with this observation,
autophosphorylation of PknG is not required for activation of its kinase activity,
setting it further apart from the trans-membrane domain containing kinases in
mycobacteria 142. Strikingly, autophosphorylation is essential for establishing the phagolysosomal fusion block in infected host macrophages 142.
Cumulatively, while no exclusive substrate for PknG has been identified, it is evident that PknG is required for the ability of M. tuberculosis to prevent lysosomal fusion to the colonized phagosome 20 (Fig. 1-7) 23, a function that is dependent on
autophosphorylation of PknG 142 and potentially based on expression levels 140.
Furthermore, PknG may influence survival inside the host through regulation of glutamine and glutamate levels, which are crucial for static-phase growth in the persistent state 121; 122; 124; 141.
1.F. MutT3
MutT motif hydrolases or Nudix hydrolases are pyrophosphatases that catalyze the hydrolysis of a nucleoside diphosphate linked to a variable moiety x 143;
144. They are characterized by the presence of about 10 conserved amino acids within a 20 amino acid sequence and occur throughout all classes of life 143; 144.
Substrates however are very diverse in nature and include nucleoside triphosphates, dinucleoside polyphosphates, nucleotide sugars, and co-enzymes 143; 144.
Furthermore, substrate specificity typically is determined by a preference for
certain substrates, but most members of the Nudix family are promiscuous and can
50
hydrolyze a variety of substrates 143; 144. It follows that these enzymes function in a number of metabolic pathways as diverse as the Nudix substrates 143; 144. For
example, the prototype Nudix hydrolase, E. coli MutT is mainly involved in hydrolysis of 8-oxoguanine, a potentially mutagenic form of deoxy-guanosine-5’- triphosphate (dGTP), and therefore helps to prevent mutations in newly synthesized
DNA 143; 144; 145. Most Nudix hydrolases seem to be involved in the clearance of
potentially deleterious or hazardous molecules or in balancing metabolites and metabolic intermediates, preventing their build-up and acting as “housekeeping” enzymes 143; 144; 146. Recently, however, it has been shown that some Nudix hydrolases serve as enzymes in intermediate steps of biosynthetic pathways 146; 147;
148 and as regulators of cellular growth in response to specific environmental stresses 149.
In particular, the Ndx8 Nudix hydrolase of Thermus thermophilus has been shown to be involved in regulation of guanosine 3',5'-bispyrophosphate (ppGpp) levels independently of the RelA/SpoT synthesis and degradation cycles that tightly regulate this bacterial alarmone 149. ppGpp is synthesized by RelA in response to amino acid starvation and initiates a growth arrest to ensure cell survival until sufficient amino acid levels are encountered or re-established, a process called the stringent response 150; 151. Once the triggers for this response are removed, SpoT
degrades ppGpp to guanosine-5-diphosphate (GDP) and the bacterial cell can re-
enter the growth phase 151; 152. Ndx8 on the other hand degrades ppGpp to 3’,5’-
bisphosphate (pGp), and may act in a relay system with SpoT, thereby helping to
51
control the transition from growth arrest to active growth after amino acid
starvation conditions have been removed 149. Interestingly, one specific trigger of amino acid starvation that induces the ppGpp-mediated stringent response is the action of classical antifolate antibiotics 153.
Three Nudix enzymes from E. coli (NtpA), Lactococcus lactis (YlgG) and
Arabidopsis (AtNUDT1) have been shown to catalyze the removal of pyrophosphate
from dihydroneopterin-triphosphate (DHN-3P) to create dihydroneopterin (DHN)
in an Mg2+-dependent manner 147; 148. DHN-3P is one of the earliest intermediates in folate de novo biosynthesis, and its conversion to DHN, which is eventually turned into folate, constitutes the first committed step in this process (Fig. 1-8) 86; 147; 148.
For over fifty years, this reaction had previously been thought to occur by
spontaneous dephosphorylation 148; 154.
While no homologous DHN-3Pase enzyme catalyzing this reaction has been so far identified in M. tuberculosis, SSDB (Sequence Similarity Database) gene cluster search at the Kyoto Encyclopedia of Genes and Genomes (KEGG:
http://ssdb.genome.jp/ssdb-bin/ssdb_gclust?org_gene=mtu:Rv0410c&type=g; Date
of access: September 02, 2011) suggested the putative Nudix enzyme (MutT3) as a
likely candidate DHN-3Pase. The amino acid sequences of MutT3 from M. smegmatis (MutT3Msm) and M. tuberculosis (MutT3Mtb) show 78% homology
(http://tuberculist.epfl.ch/blast_output/Rv0413.fasta.out; Date of access:
September 02, 2011), and no significant homology with the prototypic E. coli MutT3,
52
Figure 1-8. Scheme of the 7,8 Dihydroneopterin Triphosphate
Pyrophosphatase (DNHTPase) Catalytic Reaction and Effect of the ntpA
Deletion on E. coli's Folate Concentration. (Left) Reaction catalyzed by the
product of the gene ntpA, a DHNTPase. (Right) Folic acid concentration of parental
cells and cells lacking DHNTPase activity. Each bar represents the mean of four separate cultures, two of which were grown in M9 minimal medium and two in
Luria Broth. Cells were harvested and assayed microbiologically using Lactobacillus casei (ATTC 7469) 155. Bar A: parent cells containing plasmid without insert
(MG1655 harboring pTrc 99A); bar B: parent cells harboring plasmid with ntpA
(MG1655-pTrc 99A-NtpA); bar C: ntpA deletion mutant harboring plasmid without
insert (MG1655ntpA− − pTrc 99A); bar D: ntpA deletion mutant harboring plasmid with ntpA insert (MG1655ntpA− − pTrc 99A-NtpA).
Reproduced with permission from 147. 53
but in a 33 amino acid region, 8 residues that define the Nudix motif 143; 144 are
perfectly conserved 156. Mycobacterial MutT3 does not appear to function as either a mutator protein that hydrolyzes 8-oxo-guanine and dGTP, like MutT1 and MutT4, or as an 8-oxo-guanosine triphosphatase like MutT1 and MutT2 156. Strikingly, mutT3 is
located three genes upstream of pknG, with highly conserved synteny of the locus
across the mycobacterial genus and even some related actinobacteria. Proximity of
gene location often indicates similar function or involvement in the same pathways,
suggesting that MutT3 and PknG may act in the same pathway or regulate each
other.
1.G. RplM
The ribosomal protein RplM, also known as L13, is encoded by rplM, which is
part of a single transcriptional unit together with rpsI, a gene encoding ribosomal
protein RpsI or S9 157; 158; 159. RplM is an essential protein and constitutes a component of the large ribosomal subunit 157, but can also act in sub-ribosomal particles 160 or outside the ribosomal context 161. On the large ribosomal subunit, the
C-terminus of RplM is located near the factor binding site between ribosomal
proteins L3 and L6, towards the back of the 50S subunit, while the N-terminal end
interacts with ribosomal proteins L20 and L21 (Fig. 1-9) 162; 163. Interestingly, RplM
has been found to directly interact with Obg, an essential GTP-binding protein that
is required for regulation of the σB-mediated stress response in B. subtilis 164; 165.
Obg co-fractionated with ribosomal particles and stress response regulators, indicating a role for ribosomal function mediated by RplM interaction in induction
54
of the σB-mediated stress response in B. subtilis 165. While interactions of mycobacterial Obg with RplM have not been assayed, Obg does associate with fully assembled ribosomes as well as both ribosomal subunits in M. tuberculosis 166. RplM
has also been detected in active sub-ribosomal particles that had bound both 23S
and 5S rRNA in Thermus aquaticus 160. In a non-ribosomal setting, RplM has been
demonstrated to be involved in anti-termination together with ribosomal proteins
of the small subunit, S4, and large subunit, L3 and L4, in E. coli 161. As part of anti-
terminating transcription complexes, RplM was shown to be responsible for an 11-
fold increase in terminator read-through in in vitro transcription reactions 161.
Comprehensive analysis of phosphorylated proteins in ribosomes of E. coli
identified RplM to be phosphorylated on 5 residues, tyrosines 16, 44 and 53 and
threonines 45 and 50 167. The protein kinases responsible for this phosphorylation remain unknown, as does function of phosphorylation of these residues. However, almost all of the phosphorylated residues are located on the solvent interface of RplM, suggesting that they may be important for interactions with ribosome-associated proteins (Fig1-9) 167.
55
Figure 1-9. 3D-model of the E. coli 50S subunit showing the position of each phosphorylated ribosomal protein. The phosphorylated ribosomal proteins in the
50S subunit are represented by green color, while the phosphorylated residues mapped by mass spectrometry and unphosphorylated proteins are colored red and pink, respectively. Coordinates of the E. coli 50S subunit were obtained from the
Protein Data Bank (Acc. # 2AW4). (A) Depicting the 50S subunit from the solvent side, (B) gives a view of the large subunit from the 30S interface, and (C) is the side view of the large subunit showing the phosphorylated proteins around the SRL.
Specific regions, peptidyl transferase center (PTC), central protuberance (CP), sarcin-ricin loop (SRL), L1 and L7/L12 stalks and the 50S ribosomal proteins were labeled in the model generated by PyMol software (DeLano Scientific LLC). Red arrow indicates position of L13 on the 50S subunit.
Reproduced with permission from 167. 56
Chapter 2.
Materials and Methods
57
2.A. Bacterial strains, chemicals, media and growth conditions.
Depending on the experiments, M. smegmatis strains were grown using 7H9
(Difco), biofilm 112 liquid medium, 7H10 (Difco), NE 168, or Luria-Bertani (LB) agar medium supplemented with 0.5% Tween 80 at 37˚C. M. tuberculosis strains were grown using 7H9 (Difco), Sauton’s medium without Tween 80 113, or 7H10 (Difco)
agar medium supplemented with 0.05% Tween 80 and OADC (BD) at 37˚C. All E. coli strains were grown in LB medium or on LB agar medium at 37˚C. Kanamycin was used at 50µg/ml; hygromycin at 75µg/ml for mycobacteria and at 100µg/ml for E. coli; and ampicillin was used at 100µg/ml for E. coli.
Media and supplements were obtained from Sigma (St. Louis, MI) unless otherwise indicated, AX20017 was synthesized by GLSynthesis Inc (Worcester, MA), chelerythrine was obtained from Sigma (St. Louis, MI), [γ-32P]-ATP was from
PerkinElmer (Waltham, MA), and Guanosine 5’-triphosphate-[8-3H] ([8-3H]-GTP)
was from MP Biomedicals (Solon, OH).
2.B. Cloning, construction of vectors and construction of strains.
All primers used in this study and their sequences are listed in table 2-1.
Primers were synthesized by Fisher Scientific (Pittsburgh, PA) or Eurofins MWG
Operon (Huntsville, Alabama). All PCRs were performed using the Expand Long
Template PCR kit (Roche Molecular Biochemicals). Sequences were verified by
cloning PCR products into pGEM-T Easy vector (Promega, Madison, WI) and
sequencing (Biotic Solutions, New York, NY or ACGT Inc., Wheeling, IL). Plasmids
used in this study are listed in table 2-3. Restriction endonucleases and T4 ligase
58
were obtained from New England Biolabs (Beverly, MA). Preparation of competent cells and transformation were performed as previously described 169.
2.B.1. Targeted deletion of pknGMsm and mutT3Msm and replacement of rplMMsm
T11A T11E with rplMMsm or rplMMsm in M. smegmatis by modified recombineering
method.
Targeted gene deletion or replacement in M. smegmatis was performed by
recombineering method 170 with modifications 171.
The temperature-sensitive plasmid pVN701B, which expresses
mycobacteriophage Che9c recombination proteins 170, kanamycin resistance genes
and the counter-selectable sacB marker gene 172; 173, was constructed by removing
the SpeI-XbaI fragment of pJV53 170 and inserting it into the XbaI site of pPR27 172.
pknGMsm flanking regions were amplified using primers MS-PknG-del1 and
MS-PknG-del2 (Table 2-1) for the 5’-pknG-flanking region (551bp) and primers MS-
MutT3del3 and MS-MutT3del4 (Table 2-1) for the 3’-pknG-flanking region (528bp), respectively. pVN740 was constructed by cloning the 5’-pknGMsm-flanking region
directionally into pYUB854 adjacent to the hygromycin-resistance gene using
SpeI/HindIII sites and cloning the 3’-pknGMsm-flanking region directionally into pYUB854 adjacent to the hygromycin-resistance gene using XbaI/KpnI, respectively.
Wild-type M. smegmatis mc2155 carrying plasmid pVN701B were induced to
express the recombineering system from the vector and transformed with the linear
pknG deletion cassette fragment removed from pVN740 by digestion with SpeI/KpnI.
Selection for replacement of chromosomal DNA with the Ωhyg-containing
59
deletion/replacement cassette by homologous recombination was performed on
7H10 agar medium containing hygromycin and kanamycin. Removal of pVN701B
was performed by shifting cultures of ∆pknGMsm to 39ºC in 7H9 medium containing
only hygromycin and confirming loss of kanamycin resistance as well as gain of
ability to grow on 7H10 agar medium supplemented with 1% (wt/vol) sucrose.
pknGMsm-Ωhyg junctions were amplified using primers P1 + P2 (Table 2-1) for the 5’-
pknGMsm-Ωhyg junction and primers P3 + P4 (Table 2-1) for the 3’-pknGMsm-Ωhyg junction, respectively.
mutT3Msm flanking regions were amplified using primers MS-MutT3del1 and
MS-MutT3del2 (Table 2-1) for the 5’-mutT3-flanking region (606bp) and primers
MS-MutT3del3 and MS-MutT3del4 (Table 2-1) for the 3’-mutT3-flanking region
(500bp), respectively. pVN755 was constructed by cloning the 5’-mutT3Msm-flanking region or 5’-PrplM-flanking region directionally into pYUB854 adjacent to the hygromycin-resistance gene using SpeI/HindIII sites and cloning the 3’-mutT3Msm- flanking region directionally into pYUB854 adjacent to the hygromycin-resistance gene using XbaI/KpnI, respectively. Wild-type M. smegmatis mc2155 carrying
plasmid pVN701B were induced to express the recombineering system from the
vector and transformed with the linear pknG deletion, mutT3 deletion or rplM
replacement cassette fragment removed from pVN755 by digestion with SpeI/KpnI.
Selection for replacement of chromosomal DNA with the Ωhyg-containing
deletion/replacement cassette by homologous recombination was performed on
7H10 agar medium containing hygromycin and kanamycin. Removal of pVN701B
60
was performed by shifting cultures of ∆mutT3Msm to 39ºC in 7H9 medium containing
only hygromycin and confirming loss of kanamycin resistance as well as gain of
ability to grow on 7H10 agar medium supplemented with 1% (wt/vol) sucrose.
mutT3Msm-Ωhyg junctions were amplified using primers MMsm1 + P2 (Table 2-1) for the 5’-mutT3Msm-Ωhyg junction and primers P3 + MMsm2 (Table 2-1) for the 3’- mutT3Msm-Ωhyg junction, respectively.
The 872bp-region of the chromosome upstream of the rplMMsm
promotor area was amplified using primers MS-RplMrep1 and MS-RplMrep2 (Table
T11A T11E 3-1), and mutant alleles (715bp) of rplMMsm encoding RplMMsm or RplMMsm
were generated by a two-stage PCR procedure. Primers MS-RplMT11Arev and MS-
RplMT11Afwd (Table 2-1) were used together with primers MS-RplMrep3 and MS-
RplMrep4 (Table 2-1), respectively, and PCR products let anneal. Primers MS-
RplMrep3 and MS-RplMrep4 (Table 2-1) were then added to amplify the mutant
T11A T11E rplM or rplM alleles after the native promotor region (PrplM). pVN895 and pVN896 were constructed by cloning the 5’-PrplM-flanking region directionally into pYUB854 adjacent to the hygromycin-resistance gene using SpeI/HindIII sites and
T11A T11E cloning the PrplM- rplM region or the PrplM- rplM region directionally into pYUB854 adjacent to the hygromycin-resistance gene using XbaI/BspHI (pVN895 and pVN896). Wild-type M. smegmatis mc2155 carrying plasmid pVN701B were
induced to express the recombineering system from the vector and transformed
with the linear rplM replacement cassette fragment removed from pVN895 or
pVN896 by digestion with SpeI/BspH1. Selection for replacement of chromosomal
61
DNA with the Ωhyg-containing deletion/replacement cassette by homologous
recombination was performed on 7H10 agar medium containing hygromycin and
kanamycin. Removal of pVN701B was performed by shifting cultures of
mc2155/rplMT11A and mc2155/rplMT11E to 39ºC in 7H9 medium containing only
hygromycin and confirming loss of kanamycin resistance as well as gain of ability to
grow on 7H10 agar medium supplemented with 1% (wt/vol) sucrose. Insertion of
Ωhyg was confirmed by PCR using primers RMsm1 and RMsm2 (Table 2-1) to observe according molecular weight shift. Primer MS-RplMrep3 (Table 2-1) was used to confirm sequence had changed to mutant allele.
2.B.2. Targeted deletion of mutT3Mtb and replacement of rplMMtb with
T11A T11E rplMMtb or rplMMtb in M. tuberculosis by specialized transduction.
Targeted gene deletion or replacement in M. tuberculosis H37Rv was performed by specialized transduction 174.
mutT3Mtb flanking regions were amplified using primers TB-MutT3del1 and
TB-MutT3del2 (Table 2-1) for the 5’-mutT3Mtb-flanking region (502bp) and primers
TB-MutT3del3 and TB-MutT3del4 (Table 2-1) for the 3’-mutT3Mtb-flanking region
(570bp), respectively. pVN755 was constructed by cloning the 5’-mutT3Mtb-flanking directionally into pYUB854 adjacent to the hygromycin-resistance gene using
SpeI/HindIII sites and cloning the 3’-mutT3Mtb-flanking region directionally into pYUB854 adjacent to the hygromycin-resistance gene using XbaI/KpnI, respectively.
After PacI-l digestion to linearize the recombinant cosmids, they were ligated to shuttle phasmid phAE87 digested with PacI, and the ligation packaged with the
62
GIGAPackIII GOLD system (Stratagene). Phages containing the recombinant
phasmid ph∆mutT3Mtb were used to transduce E. coli NM554 and transductants were selected on hygromycin. Phasmid DNA was extracted from pooled transductants and used to transform M. smegmatis mc2155 by electroporation.
Plaques obtained at 30ºC were isolated and used to transduce M. tuberculosis H37Rv and CDC1551. Transductants were selected on hygromycin and gene deletion/replacement confirmed by PCR using primers MMtb1 + MMtb2 (Table 2-1) to observe according molecular weight shift. Strains were designated ∆mutT3Mtb.
The region of the chromosome upstream of the rplMMtb promotor area
(716bp) was amplified using primers TB-RplMrep1 and TB-RplMrep2 (Table 2-1),
and mutant alleles of rplMMtb (676bp) in which the phosphorylated residue threonine 11 (T11) was replaced by an alanine (T11A) or a glutamate residue
(T11E) were generated by a two-stage PCR procedure. Primers TB-RplMT11Arev and
TB-RplMT11Afwd (Table 2-1) were used together with primers TB-RplMrep3 and TB-
RplMrep4 (Table 2-1), respectively, and PCR products let anneal. Primers TB-
RplMrep3 and TB-RplMrep4 (Table 2-1) were then added to amplify the mutant
T11A T11E rplM or rplM alleles after the native promotor region (PrplM). pV897 and pVN898 were constructed by cloning the 5’-PrplM-flanking region directionally into pYUB854 adjacent to the hygromycin-resistance gene using SpeI/XhoI sites and
T11A T11E cloning the PrplM-rplM region or the PrplM-rplM region directionally into pYUB854 adjacent to the hygromycin-resistance gene using XbaI/KpnI, respectively.
After PacI-l digestion to linearize the recombinant cosmids, they were ligated to
63
shuttle phasmid phAE87 digested with PacI, and the ligation was packaged with the
GIGAPackIII GOLD system (Stratagene). Phages containing recombinant phasmids
T11A T11E phrplMMtb or phrplMMtb were used to transduce E. coli NM554 and transductants were selected on hygromycin. Phasmid DNA was extracted from pooled transductants and used to transform M. smegmatis mc2155 by electroporation. Plaques obtained at 30ºC were isolated and used to transduce M. tuberculosis H37Rv and CDC1551. Transductants were selected on hygromycin and gene deletion/replacement confirmed by PCR using primers RMsm1 + RMsm2 (Table
2-1) to observe according molecular weight shift. Primer TB-RplMrep3 was used to
confirm sequence had changed to the mutant allele. Strains were designated
H37Rv/rplMT11A, H37Rv/rplMT11E, CDC1551/rplMT11A and CDC1551/rplMT11E.
2.B.3. Deletion of ntpA in E. coli by one-step phage λ-Red-based recombineering method.
The chromosomal gene encoding the E. coli mutT3 homolog (ntpA) was deleted using a one-step phage λ-Red-based recombineering method as previously described 175; 176. The KanR cassette was amplified using primers NtpA-Kan1 and
Ntpa-Kan2, which carried extensions homologous to the flanking DNA regions of
ntpA (Table 2-1). PCR products were gel-purified and directly electroporated into E.
coli TB10 cells (MG1655, nadA::Tn10 λcI857 Δ(cro-bioA)) 177 that had been heat- induced to express the λ-Red system 175; 176. Kanamycin-resistant transformants were selected and correct replacement of ntpA by KanR verified by PCR using primers annealing adjacent to homologous sequences, NtpA1 + NtpA2 (Table 2-1).
64
The obtained ∆ntpA locus was transferred to the wild type E .coli strain MG1655 by
P1 phage-mediated transduction as previously described 177.
2.B.4. Cloning for in trans expression of proteins.
pVN578 and pVN579 were constructed as previously described. 20
pknGMtb was amplified using primers TB-PknG-1 and TB-PknG-2 (Table 2-1)
and cloned directionally into pET15b using NdeI/XhoI to fuse the N-terminus to a
6xHis-tag, creating pVN792.
garAMsm was amplified using primers MS-GarA-1 + MS-GarA-2 (Table 2-1)
and cloned directionally into pMV361 using EcoRI/HindIII to be expressed from
Phsp60, creating plasmid pVN821.
mutT3Msm was amplified using primers MS-MutT3-1 + MS-MutT3-2 (Table 2-
1) and cloned directionally into pMV361 using EcoRI/HindIII to be expressed from
Phsp60, creating plasmid pVN753. For addition of a C-terminal 6xHis-tag, mutT3Msm
was amplified using primers MS-MutT3-1 + MS-MutT3-2-6xHis (Table 2-1) and cloned directionally into pVN747 or pET11c using NdeI/HindIII or NdeI/BamHI to be expressed from PSOD or PT7, creating pVN823 and pVN835, respectively. For expression from its native promotor PmutT3, mutT3Msm was amplified using primers
MS-pro-MutT3 + MS-MutT3-2-6xHis (Table 2-1) and cloned directionally into pCV125 using NdeI/HindIII, creating pVN866.
mutT3Mtb was amplified using primers TB-MutT3-1 + TB-MutT3-2 (Table 2-
1) and cloned directionally into pMV361 using EcoRI/HindIII to be expressed from
Phsp60, creating plasmid pVN771. For expression from its native promotor PmutT3,
65
mutT3Mtb and its upstream promotor region was amplified using primers TB-pro-
MutT3 + TB-MutT3-2 (Table 2-1) and cloned directionally into pVN839 using
NdeI/HindIII, creating pVN840.
ntpA was amplified using primers NtpA-1 + NtpA-2 (Table 2-1) and cloned
directionally into pMV361 using EcoRI/HindIII to be expressed from Phsp60, creating plasmid pVN773.
ylgG sequence was optimized for codon usage in mycobacteria, synthesized by GenScript (Piscataway, NJ) with added restriction sites, and cloned directionally into pMV361 using EcoRI/HindIII to be expressed from Phsp60, creating plasmid pVN776.
rplMMsm was amplified using primers MS-RplM-1 and MS-RplM-2 (Table 2-1) and cloned directionally into pET15b using NdeI/BamHI to be expressed from T7, creating pVN844.
rplMMtb was amplified using primers TB-RplM-1 and TB-RplM-2 and cloned
directionally into pET15b using NdeI/BamHI to be expressed from T7, creating
pVN885. Mutant alleles of rplMMtb in which the residues threonine 11 (T11),
threonine 12 (T12) and/or serine 14 (S14) were replaced by an alanine (T11A,
T12A, S14A) were generated by a two-stage PCR procedure. Primers TB-RplM3T-fwd,
TB-RplMT11A-fwd, TB-RplMT12A-fwd or TB-RplMS14A-fwd (Table 2-1) were used together with primer TB-RplM-2 (Table 2-1) and PCR products let anneal with primers TB-RplM3T-1, TB-RplMT11A-1, TB-RplMT12A-1 or TB-RplMS14A-1 (Table 2-1),
respectively. Primers TB-RplM-1 and TB-RplM-2 (Table 2-1) were then added to
66
Table 2-1. Primers used in this study.
Primer Sequence
MS-PknG-1 gaattccatatgacttcacccgagaacc
MS-PknG-2 aagcttccatgaaagcacggtcgacgtg
P1 atcggcagcaacctgttctcgttc
P2 ctgcacgacttcgaggtgttcga
P3 gcgattcagcctggtatgatcagc
P4 ccgtcttctacagtcgtcgtagg
PknG-del1 actagtaccaggtggacatcgtcgtcaaga
PknG-del2 aagcttgatcagggttctcgggtgaagtca
PknG-del3 tctagactggtcgacctggccaacagc
PknG-del4 ggtacccaatgggaattgacgggtcgatcc
RT-PknG1 gccaccgacatctacaccgt
RT-PknG2 ggtgtgcgccaccagcag
TB-PknG-1 CATatggccaaagcgtcagagaccgaa
TB-PknG-2 CTCGAGttagaacgtgctggtgggccgga
MMsm1 gatctgcgtctgcatcatcgacga
MMsm2 acctgatccagctgcgtgacaag
MMtb1 gcctctgacagatatgacgacc
MMtb2 ggcaaggctgtatctgtgcacc
P2 ctgcacgacttcgaggtgttcga
P3 ccgattcagcctggtatgatcagc
MS-MutT3del1 ACTAGTgcaactggatcagcgcctcgtc
MS-MutT3del2 AAGCTTccgttctccgacatcacccagc
MS-MutT3del3 TCTAGAtgctcaaccggcagcgctgatc
MS-MutT3del4 GGTACCcccgtcaggaactcgaagcgc
67
TB-MutT3del1 ACTAGTcccagcaattgcaccaacgattcg
TB-MutT3del2 AAGCTTcgtatgcctaacaccatctgtcgg
TB-MutT3del3 TCTAGAcggatcagttcgctgctgtaagc
TB-MutT3del4 GGTACCggagtggacatcatccagttgcg
MS-MutT3-1 CATATGTCTAGAgcaactggatcagcgcctcgtc
MS-MutT3-2 AAGCTTtcagcgctgccggttgagcaac
MS-MutT3-2-6xHis GGATCCAAGCTTTCAATGATGATGATGATGGTGGCTGCTGCCGCT GCCGCGCGGCACCAGgcgctgccggttgagcaacgg
MS-pro-MutT3 CATATGcgatggcctgctcgtagcgct
TB-MutT3-1 GAATTCCATATGccgacagatggtgttaggcatacg
TB-pro-MutT3 TCTAGAcaccgacagcgtggtgtacagc
TB-MutT3-2 AAGCTTgcttacagcagcgaactgatccg
NtpA-1 GAATTCCATAtgaaggataaagtgtataagcgtcc
NtpA-2 GGATCCAAGCTTtcaggcagcgttaattacaaactgt
NtpA-Kan1 tcgatcttagtggtcatctacgcacaagatacgaaacgggtccggggatccgtcgacc
NtpA-Kan2 cgcctgccggttgctccaggacttagtgagcgccgccgcgtgtaggctggagctgcttcg
MS-GarA-1 GAATTCCATATGgatgtgacggtagagaccacatcg
MS-GarA-2 AAGCTTGGATCCccatgggtcaggcgttcgagc
RMsm1 ccgaaaatctcgcacacaagccaac
RMsm2 cgctctcggtcaccacctcg
MS-RplMrep1 ACTAGTcagtgcgtcagtagtagggagc
MS-RplMrep2 AAGCTTcgtcctattcatatgccgtttctgg
MS-RplMrep3 TCTAGAgtgcgtaccggttccctctatg
MS-RplMrep4 TCATGAgtttcggtcacatcggtcactgg
TB-RplMrep1 ACTAGTccatgaagacctcctcagggagg
TB-RplMrep2 CTCGAGcggaactagcgtgcccaagct
TB-RplMrep3 TCTAGAgcgcaacagacccctgtgtggt
68
TB-RplMrep4 GGTACCggctggggtggtttcggtcat
MS-RplMT11Arev taccacgaacgcgtggcgtca
MS-RplMT11Afwd tgacgccacgcgttcgtggta
MS-RplMT11Erev gtaccacgaacgcgtctcgtca
MS-RplMT11Efwd tgacgagacgcgttcgtggtac
TB-RplMT11Arev cacgatcgcgtggcgtcaccc
TB-RplMT11Afwd gggtgacgccacgcgatcgtg
TB-RplMT11Erev cacgatcgcgtctcgtcaccc
TB-RplMT11Efwd gggtgacgagacgcgatcgtg
MS-RplM-1 GAATCCCATATGgtgcctacttacacgccgaagg
MS-RplM-2 AAGCTTGGATCCgtttcggtcacatcggtcactgg
TB-RplM-1 CATATGagcgctgtgcccacgtacgc
TB-RplM-2 GGATCCggtggtttcggtcattgcgcca
TB-RplM3T-fwd acgccgcgcgagcgtggta
TB-RplM3T-1 taccacgctcgcgcggcgtcacccgccttgggcgcgtacgtgggcacagcgctCATATG
TB-RplMT11A-fwd acgccacgcgatcgtggta
TB-RplMT11A-1 taccacgatcgcgtggcgtcacccgccttgggcgcgtacgtgggcacagcgctCATAG
TB-RplMT12A-fwd acaccgcgcgatcgtggta
TB-RplMT12A-1 taccacgatcgcgcggtgtcacccgccttgggcgcgtacgtgggcacagcgctCATAG
TB-RplMS14A-fwd acaccacgcgagcgtggta
TB-RplMS14A-1 taccacgctcgcgtggtgtcacccgccttgggcgcgtacgtgggcacagcgctCATATG
Capital letters represent restriction sites or tags added to genetic sequence (lower case).
69
Table 2-2. Plasmids and phasmids used in this study.
Plasmid Relevant Features References / Company
pCV125 KanR, mycobacterial integrative vector carrying 178 lacZ pET11c AmpR, replicative E. coli vector (OripMB1) carrying Novagen a built-in PT7 promotor followed by a lac operator and a multiple cloning site. Carries lacI.
pET15b AmpR, replicative E. coli vector (OripMB1) carrying Novagen a built-in PT7 promotor followed by a lac operator, an N-terminal 6xHis-tag sequence, a thrombin site and a multiple cloning site. Carries lacI.
pJV53 KanR, mycobacterial replicative vector expressing 179 the Che9c 60-61 genes under control of an acetamide-inducible promotor
pPknGK181M KanR, pSD5 -derived, expressing M. tuberculosis 20 pknGK181M provided by Jean Pieters
pPR27 GenR, mycobacterial vector, replicated from a 173
temperature sensitive Ori and expresses a
sucrose counter selectable marker sacB
pMV361 KanR, mycobacterial integrative vector carrying a 180 built-in heat shock promotor (Phsp60) for translational fusion
pVN578 KanR, pMV361-derived, expressing M. tuberculosis This study 171 pknG from Phsp60
pVN579 KanR, pMV361-derived, expressing M. smegmatis This study 171 pknG from Phsp60
pVN701B KanR GenR, pJV53-derived vector replicating from This study 171 a temperature-sensitive OriTs and expressing a sucrose counter selectable marker sacB as well as mycobacteriophage Che9c recombination proteins
pVN740 HygR, pYUB854 -derived vector with M. This study 171 smegmatis pknG flanking-regions cloned orientationally flanking the hygromycin cassette
70
pVN747 HygR, pMS2–derived replicative vector carrying a 181 built-in PSOD promotor
pVN753 KanR, pMV361-derived, expressing M. smegmatis This study mutT3 from Phsp60
pVN755 HygR, pYUB854-derived vector with M. smegmatis This study mutT3 flanking-regions cloned orientationally flanking the hygromycin cassette
pVN771 KanR, pMV361-derived, expressing M. tuberculosis This study mutT3 from Phsp60
pVN773 KanR, pMV361-derived, expressing E. coli ntpA This study from Phsp60
pVN776 KanR, pMV361-derived, expressing L. lactis ylgG This study from Phsp60
pVN791 HygR, pYUB854-derived vector with M. This study tuberculosis mutT3 flanking-regions cloned orientationally flanking the hygromycin cassette
pVN792 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. tuberculosis pknG from PT7
pVN798 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged E.coli ntpA from PT7
pVN821 KanR, pMV361-derived, expressing M. smegmatis This study garA from Phsp60
pVN823 HygR, pVN747-derived, expressing c-terminally This study 6xHis-tagged M. smegmatis mutT3 from PSOD
pVN835 AmpR, pET11c-derived, expressing c-terminally This study 6xHis-tagged M. smegmatis mutT3 from PT7
pVN839 KanR, integrative mycobacterial vector carrying This study OriC, KanR, and integration sites from pMV361 and PSOD and multiple cloning sites from pVN747
pVN840 KanR, pVN839-derived, expressing M. tuberculosis This study mutT3 from it native promotor PmutT3
71
pVN844 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. smegmatis rplM from PT7
pVN866 KanR, pCV125-derived, expressing c-terminally This study 6xHis-tagged M. smegmatis mutT3 from its native promotor PmutT3
pVN885 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. tuberculosis rplM from PT7
pVN889 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. tuberculosis rplMT11A,T12A,S14A from PT7
pVN890 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. tuberculosis rplMT11A from PT7
pVN891 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. tuberculosis rplMT12A from PT7
pVN892 AmpR, pET15b-derived, expressing n-terminally This study 6x-His-tagged M. tuberculosis rplMS14A from PT7
pVN895 HygR, pYUB854-derived vector with M. smegmatis This study rplMT11A-containing flanking-regions cloned orientationally flanking the hygromycin cassette
pVN896 HygR, pYUB854-derived vector with M. smegmatis This study rplMT11E-containing flanking-regions cloned orientationally flanking the hygromycin cassette
pVN897 HygR, pYUB854-derived vector with M. This study tuberculosis rplMT11A-containing flanking-regions cloned orientationally flanking the hygromycin cassette
pVN898 HygR, pYUB854-derived vector with M. This study tuberculosis rplMT11E-containing flanking-regions cloned orientationally flanking the hygromycin cassette
pYUB854 HygR, E. coli replicative vector carrying a 174 hygromycin cassette and lambda cos sites
72
phAE87 TM4-derived mycobacterial shuttle phasmid, 182 conditionally replicative at 30°C
ph∆mutT3Mtb HygR, phAE87 –derived phasmid carrying PacI- This study digested mutT3Mtb-deletion cassette of pVN791
phrplMMtbT11A HygR, phAE87 –derived phasmid carrying PacI- This study digested rplMMtbT11A-replacement cassette of pVN987 phrplMMtbT11E HygR, phAE87 –derived phasmid carrying PacI- This study digested rplMMtbT11E-replacement cassette of pVN989 HygR, hygromycin resistance. KanR, kanamycin resistance. GenR, gentamycin resistance. AmpR, ampicyclin resistance
73
T11A,T12A,S14A T11A T12A S14A amplify the mutant rplMMtb , rplMMtb , rplMMtb or rplM alleles and products cloned directionally into pET15b using NdeI/BamHI to be expressed from
T7, creating pVN889, pVN890, pVN891 and pVN892, respectively.
For expression and purification of PknGMtb-6xHis, BL21 transformed with
pVN792 were grown at 37ºC until OD600 had reached 0.65 and induced with 0.1mM
IPTG at 22ºC, 200rpm for 16 hours, as previously described 20. Soluble fraction of cell lysates was prepared in TBS buffer containing protease inhibitor cocktail by sonication (15 cycles of 10 seconds on ice with chilling intervals) and diluted 1:1 with wash buffer (50mM sodium phosphate, 300mM NaCl, and 45mM imidazole), loaded onto a cobalt metal affinity spin column (Pierce) pre-equilibrated with the same buffer. The column was washed 7 times with 2 column volumes of wash buffer and bound protein was eluted 4 times with 1 column volume of elution buffer (50 mM sodium phosphate, 300 mM NaCl, and 250 mM imidazole). Eluted fractions were pooled and stored at -80ºC with 50% glycerol.
For expression and purification of MutT3Msm-6xHis from E.coli, BL21 transformed with pVN835 were grown at 37ºC until OD600 had reached 0.5 and
induced with 0.5mM IPTG at 30ºC, 250rpm for 3 hours. Soluble fraction of cell
lysates were prepared in lysis buffer (50mM sodium phosphate, 300mM NaCl, and
10mM imidazole) containing protease inhibitor cocktail by sonication (15 cycles of
10 seconds on ice with chilling intervals), loaded onto a nickel metal affinity spin
column (Qiagen) preequilibrated with the same buffer. The column was washed 6
times with 2 column volumes of wash buffer (50mM sodium phosphate, 300mM
74
NaCl, and 20mM imidazole) and bound protein was eluted 4 times with 1 column
volume of elution buffer (50 mM sodium phosphate, 300 mM NaCl, and 500 mM
imidazole). Eluted fractions were pooled and stored at -80ºC with 10% glycerol.
For purification of MutT3Msm-6xHis from M. smegmatis, 7H9 medium
2 containing hygromycin was inoculated with 5 OD600 units/L of mc 155 transformed
with pVN823 and cultures grown at 37ºC, 240rpm for 72 hours. Cell pellets were
harvested at 4ºC, 4000rpm for 15 minutes and washed twice with TBS buffer
containing a protease inhibitor cocktail (Roche Molecular Biochemicals). After
resuspension in 1/10 culture volume of TBS buffer + protease inhibitors, cells were
disrupted by sonication (25 times for 10 seconds on ice with 1 minute cooling
intervals), and lysate spun for 20 minutes at 10,000rpm, 4ºC. Supernatant was
filtered at 4ºC through 0.22µm filters (Denville) and diluted 1:1 with wash buffer
(50mM sodium phosphate, 300mM NaCl, and 20mM imidazole), loaded onto a
cobalt metal affinity spin column (Pierce) preequilibrated with the same buffer. The
column was washed 10 times with 2 column volumes of wash buffer and bound
protein was eluted 3 times with 1 column volume of elution buffer (50 mM sodium
phosphate, 300 mM NaCl, and 150 mM imidazole). Eluted fractions were pooled and
exchanged into 20mM Tris-HCl buffer, pH8.0 using PD-10 desalting columns (GE
Healthcare) before removing remaining contaminants using a strong anion
exchange column (HiTrap Q FF, GE Healthcare) and a 0-1M NaCl gradient. Fractions
were analyzed for presence of MutT3Msm-6xHis by SDS-PAGE stained with
Coomassie Brilliant Blue (see Methods section 3.C.7.) and fractions containing
75
protein (20mM Tris-HCL, 100-150mM NaCl) were pooled and concentrated using 9
kDa molecular weight cut-off spin concentrators (Pierce). Obtained purified fraction
was stored at -80ºC with 10% glycerol.
For expression and purification of RplMMsm-6xHis, RplMMtb-6xHis,
T11A,T12A,S14A T11A T12A S14A RplMMtb , RplMMtb RplMMtb , and RplMMtb , BL21 was transformed with pVN844, pVN885, pVN890, pVN891, and pVN892, respectively,
and grown at 37ºC until OD600 had reached 1. These cultures served as a 1/100 seed
for LB medium supplemented with ampicillin and the cultures were then grown at
37ºC, 240rpm overnight. Soluble fraction of cell lysates were prepared in lysis
buffer (50mM sodium phosphate, 300mM NaCl, and 10mM imidazole, pH9.0)
containing protease inhibitor cocktail by sonication (15 cycles of 10 seconds on ice
with chilling intervals), loaded onto a nickel metal affinity spin column (Qiagen) pre-
equilibrated with the same buffer. The column was washed 10 times with 2 column
volumes of wash buffer (50mM sodium phosphate, 300mM NaCl, and 20mM
imidazole, pH9.0) and bound protein was eluted 3 times with 1 column volume of
elution buffer (50 mM sodium phosphate, 300 mM NaCl, and 500 mM imidazole,
pH9.0). Eluted fractions were pooled, protein exchanged into thrombin buffer
(20mM Tris-HCl, pH8.4, 0.15M NaCl, 2.5mM CaCl2) using PD-10 desalting columns
(GE Healthcare) and 6xHis tag was removed by thrombin cleavage (Novagen) at a
5:8 ratio of recombinant protein (mg) : thrombin (U) at 4ºC for 20 hours. Cleavage
reactions were concentrated using 9 kDa molecular weight cut-off spin
concentrators (Pierce), and the cleaved tag and other contaminants were removed
76
by fast protein liquid chromatography (FPLC) using a HiLoad 16/60 Superdex 75
prep grade column (GE Healthcare). Fraction containing purified, untagged protein
was concentrated using 9 kDa molecular weight cut-off spin concentrators (Pierce) and stored at -80ºC with 10% glycerol.
2.C. RT-PCR.
Primers (Table 2-1) were synthesized by Fisher Scientific (Pittsburgh, PA).
RNA from M. smegmatis strains was isolated according to manufacturers protocol using the RNeasy minikit (Qiagen) and reverse transcribed as previously published
140 using the Omniscript RT kit (Qiagen). RT-PCRs were performed using the Expand
Long Template PCR kit (Roche Molecular Biochemicals). cDNA of pknG was
amplified using primers RTpknG1 and RTpknG2 (Table 2-1), which anneal to
conserved sequences present in both pknGMsm and pknGMtb.
2.D. SDS-PAGE, Coomassie Blue staining and Western blots.
For preparation of whole cell extracts, mycobacterial cells were washed
three times in Tris-HCl-buffered saline (TBS) containing protease inhibitors (Roche
Molecular Biochemicals) and disrupted by glass bead beating using a Fastprep 24
(MP Biomedicals, Solon, OH) for 6 cycles of 40 seconds at frequency 60s-1 with chilling intervals. Glass beads, unbroken cells and insoluble material were removed by centrifugation (20,000rpm, 20 min) and the total cell lysate treated with SDS- sample buffer and heated at 95ºC for 10 minutes.
SDS-PAGE was performed using the Biorad Protean III system. Lysates, protein purification fractions or kinase assay samples were heated for 10 min at
77
95°C in SDS-sample buffer, centrifuged briefly and loaded on 10 or 15 %
polyacrylamide SDS-PAGE gels and resolved at 20mA per gel.
Gels used for kinase assay or Western blot were blotted by semi-dry transfer
(BioRad, Hercules, CA) onto PVDF membranes for 4 hours at 0.8mA/cm2 of membrane.
Coomassie Brilliant Blue G-250 (Fisher Bioreagents, Pittsburgh, PA) staining of SDS-PAGE gels or PVDF membranes was done for 30 minutes at room temperature.
PknG Western Blot was performed using 35µg total protein of M. smegmatis strains (except ∆pknGMsm/pVN578 and ∆pknGMsm/pVN579, where 10 µg total protein
were loaded) or M. bovis BCG per lane. Membranes were incubated with 1/5000
20 polyclonal antibody raised against PknGMtb in 5% milk in TBS supplemented with
1% Tween 80 (TBS-T) (Sigma) for 3 hours at room temperature. Secondary
antibody was horseradish-peroxidase conjugated goat anti-rabbit IgG antibody
(Pierce) in 5% milk in TBS-T. Detection was done using Pierce ECL Western blotting
kit and signals were recorded on film.
His-tag Western blot was performed using 20µg of mycobacterial cell extract
or 0.1µg of purified 6x-His-tagged protein per lane. Membranes were incubated with
1/1000 penta-His antibody (Qiagen) in 3% BSA (Sigma) in TBS supplemented with
1% Tween 80 (TBS-T) (Sigma) overnight at 4ºC. Secondary antibody was
horseradish-peroxidase conjugated goat anti-mouse IgG antibody (Pierce) in 5%
milk in TBS-T. Detection was done using Pierce ECL Western blotting kit and signals
78
were recorded on film.
2.E. Antibiotic susceptibility testing.
Antibiotic susceptibility of M. smegmatis strains was estimated using a diffusion assay with antibiotic-containing filter paper discs (BD Diagnostic Systems) as previously described 168. Minimal antibiotic inhibitory concentrations (MIC) were
® determined using E-test (AB Biodisk) strips on NE agar medium with 0.04 OD600
units of cells and read out after 48-72 hour growth at 37°C as described previously
168.
Antibiotic susceptibility of E. coli strains was estimated using an adapted
168 filter paper disc assay where cultures were grown to OD600 = 1, NE medium replaced by LB agar medium and plates read out after 24 hours.
In M. tuberculosis strains, erythromycin susceptibility was assessed by
streaking 10µl of cultures at OD600 = 1 onto 7H10-OADC medium with or without
erythromycin (100mg/L). Growth was assessed after 4 week incubation at 37°C.
Sulfachloropyridazine susceptibility on solid medium was assessed either by streaking 10µl of cultures at OD600 =1 or by replicating (96-well layout) or spotting
2µl volumes of serial dilutions of cultures of defined OD600 onto 7H10-OADC
medium containing titrations of sulfachloropyridazine and folinic acid. Growth was compared after 4 week incubation at 37°C. Sulfachloropyridazine susceptibility in
liquid medium was assessed by daily monitoring of growth (OD600) in 7H9-OADC
medium containing increasing amounts of antibiotic (0-100µg/ml) over a 7 day
incubation period at 37°C and 200rpm (n=3).
79
2.F. Cellular aggregation, Hydrophobicity Index and Congo red binding assays.
Cellular aggregation assays were performed as previously described 183.
Hydrophobicity assays were performed as previously described 183. Briefly,
M. smegmatis strains were grown to late exponential phase in 7H9 medium and single cell suspensions were prepared and washed with PBS. OD600 was adjusted to
1.0 and mixed with hexadecane (Sigma). After 15 minute incubations at room temperature allowing for phase separation, the aqueous phase was harvested and
OD600 measurements used to assay percentage of cells transitioning from hydrophilic to hydrophobic phase.
Congo Red binding assay was performed as previously described 183. Briefly,
M. smegmatis strains were grown in the presence of 20µg/ml Congo red for 72 hours in 7H9 medium, washed until supernatant was completely colorless, cell resuspended in acetone to extract bound Congo Red and OD at 488nm monitored.
Cells were then pelleted and dried and Congo red binding expressed as OD488/g dry cell mass.
2.G. Zeta Potential.
Cell surface charge of M. smegmatis strains was measured by zeta potential
183 determination as previously described . Briefly, 1 OD600 units of PBS-washed single cell suspensions were used for measurements in a ZetaMaster zetameter (Malvern Instruments) and expressed in mV.
2.H. Biofilm assays.
Biofilm assays were done as described 112; 113. Briefly, strains were grown in
80
7H9 liquid medium supplemented with glucose (M. smegmatis) or OADC (M.
tuberculosis) and appropriate antibiotics until OD600 had reached 0.5-0.8 (M. smegmatis) or >1 (M. tuberculosis). Growing cultures (10 µl for M. smegmatis, 100 µl for M. tuberculosis) were used to inoculate 60x15mm polystyrene petri plates containing 10 ml of biofilm medium (M. smegmatis) 112 or Sauton’s medium without
Tween 20 113. Depending on assay, medium was supplemented with antifolates (0-
50 µM) (Sigma), kinase inhibitors, folic acid (0-1mM) (Schircks Laboratories) or
folinic acid (0-1mM) (Schircks Laboratories). Standing cultures were incubated at
30ºC and observed for 7 days (M. smegmatis) or at 37ºC under humidified
conditions and wrapped in parafilm and observed for 6 weeks (M. tuberculosis). For
quantitation of biofilm growth, the liquid medium and any planktonic cells were
removed with a syringe connected to a sterile needle from beneath the biofilm layer and the remaining biofilm biomass harvested. Viable bacteria counts were determined by colony forming unit (CFU) assay or total protein per plate determined by Bradford method (Bio-Rad Laboratories, CA).
2.I. Kinase assays.
In 20µl volumes, 0.5µg of purified PknGMtb-6xHis (depending on assay pre- treated for 15 minutes at 37°C with 5mM AX20017) were incubated with 1.5µg of purified substrate (RplM or MutT3 forms, with or without 6x-His-tag, purified from
E. coli or M. smegmatis) in 10mM HEPES, 20µM DTT, 40µM MnCl2 with 10µCi of [γ-
32P]-ATP (PerkinElmer) for 30 minutes at 37°C. Reactions were stopped by addition
of SDS-sample buffer and boiling at 95°C for 10 minutes and resolved on 15%
81
polyacrylamide SDS-PAGE gels (see section 2.E.). Gels were transferred by semi-dry
blot (BioRad) onto PVDF membranes (BioRad) and stained with Coomassie Brilliant
Blue R-250 (Fisher) before exposing to autoradiography imaging screens for 6 days.
Screen were read on a Storm 820 PhosphoImager (GE Healthcare) and analyzed with
ImageQuant software (GE Healthcare).
2.J. Radioactive nucleotide binding assays.
Radioactive nucleotide binding assays were adapted from previously
184; 185 described protocols . Briefly, in triplicates, purified MutT3Msm-6xHis was incubated with [8-3H]-GTP (MP Biomedicals) in the presence or absence of
unlabeled GTP (Sigma) in buffer (50mM Tris-HCl, pH 7.4, 100mM NH4Cl, 10mM
MgCl2, 1mM dithiothreitol) in 200µl total volumes for 20 minutes at 37°C before stopping reactions by addition of 3ml chilled stopping solution (50mM Tris-HCl, pH
7.4, 10mM NH4Cl, 10mM MgCl2). Reactions were immediately filtered through
0.45um pore size nitrocellulose membrane discs (25mm) and membranes washed with three 3ml volumes of chilled stopping solution before soaking in scintillation fluid (PerkinElmer). Released radioactive signal that had been retained on membranes was read by a Beckman LS6500 liquid scintillation counter.
3 For titration of MutT3Msm-6xHis, [8- H]-GTP was kept constant at 6.4µCi
(2.5µM). For titration of labeled GTP, MutT3Msm-6xHis was kept constant at 10 or
20µg. For outcompetition of [8-3H]-GTP by unlabeled GTP, [8-3H]-GTP and
MutT3Msm-6xHis were kept constant at 6.4µCi (2.5µM) and 20µg, respectively.
2.K. Macrophage infection, lysosomal trafficking and microscopy.
82
Macrophage infections and lysosomal trafficking assays were adapted from previously published methods 20; 186. Bone marrow–derived macrophages were generated by incubating bone marrow monocytes from C57BL/6 mice (provided by
Kurt Lu) for 7 days in high-glucose-Dulbecco’s Modified Eagle Medium containing
25% L-929 conditioned medium, 1% penicillin, 1% streptomycin, 4.5g/L glucose
(HyClone) supplemented with 4 mM L-glutamine (Sigma), 15% heat-inactivated fetal calf serum (Sigma), and 0.02mg/L macrophage-stimulating growth factor
(Sigma) at 37°C and 10% CO2. Non-activated macrophages were seeded on MatTek
(Ashland, MA) glass bottom 14mm microwell dishes with 1.5 coverglass and let
adhere for 2 hours (37°C, 10%CO2) prior to infection with M. bovis BCG or ∆pknGBCG
expressing GFP (provided by Jean Pieters) 20; 186 (MOI 625) for 1 hour (37°C,
10%CO2). After removal of non-phagocytosed bacteria, infected macrophages were
subjected to a 16-hour chase with 30µM sulfachloropyridazine (Sigma) and/or 2mM
folinic acid (Schircks Laboratories). After the chase period, lysosomal compartments
were labelled for 30 minutes (37°C, 10%CO2) with 1µM neutral red (Invitrogen), an acidic organelle–selective cell-permeant fluorescent probe, and fixed with 4% formaldehyde (Fisher) for 20 minutes before staining with DAPI (Invitrogen) for 5 minutes. Slides were mounted in ProLong Gold antifade reagent (Invitrogen). Assays were analysed on a Zeiss LSM510 confocal microscope using LSM510 REL3.5
software provided by the manufacturer. 100 events were counted for each condition
from triplicate slides in 2 independent experiments.
83
Chapter 3.
Protein Kinase G is Required for Intrinsic Antibiotic Tolerance in
Mycobacteria
Kerstin A. Wolff,† Hoa T. Nguyen,† Richard Cartabuke, Ajay Singh, Sam Ogwang,
and Liem Nguyen
Department of Molecular Biology and Microbiology, School of Medicine, Case Western
Reserve University, Cleveland, Ohio 44106
† These authors contributed equally to this research.
84
3.A. Abstract.
The eukaryote-like serine/threonine protein kinase G (PknG) of M.
tuberculosis has previously been reported to be involved in survival of the bacillus in infected host macrophages, contributing to its virulence and pathogenesis. We
identified pknG in a transposon mutagenesis based screen for genome wide
determinants of multiple drug resistance in mycobacteria, and demonstrate that
PknG is required for intrinsic multiple drug tolerance (MDR) in both pathogenic and
non-pathogenic mycobacteria. Deletion of pknG leads to altered cell surface
properties and increased susceptibility to a wide variety of antibiotics in both M.
smegmatis and M. tuberculosis, indicating that PknG is a genetic factor underlying
the commonly observed phenotypic association between antibiotic resistance and
virulence.
85
3.B. Introduction.
Traditionally, host persistence and antibiotic resistance have been regarded
as separate phenomena, but phenotypic association of the two is commonly
observed both in vivo and in vitro 22; 23; 24. For example, during persistence in the
host, M. tuberculosis exhibits reduced antibiotic susceptibility when compared to
actively dividing bacteria 13; 14; 15; 16. Conversely, the transition from active growth to persistence can be induced by supplementation of antibiotics 25.
Like for all intracellular pathogens, it is crucial for M. tuberculosis to be able to survive and persist within infected host cells 10. While we do not have a complete understanding of persistence in the host, which is established by colonizing the early phagosome and arresting its maturation, several factors from both host and M. tuberculosis have been shown to play crucial roles in this process 4; 5; 17; 18; 19. Among
the mycobacterial factors are glycolipids and the RD1 specialized secretion system
17; 18; 19. In addition, mycobacterial protein phosphatases and kinases have recently been identified to perform key roles in establishing the survival niche within host phagosomes 19; 20; 21. One such kinase is the mycobacterial eukaryotic-like
serine/threonine protein kinase G (PknG), which was suggested to support survival
of pathogenic mycobacteria in the host macrophage by preventing phagolysosomal
fusion and therefore acidification of the mycobacterial compartment 20. Thus, PknG has been thought to function as a virulence factor of M. tuberculosis 20; 187. However,
genes encoding PknG are ubiquitously conserved throughout the Mycobacterium genus, including non-pathogenic mycobacteria such as M. smegmatis 122; 140.
86
Sequence identity between the M. smegmatis and M. tuberculosis homologs of PknG
is ~80%, and the gene locus is also highly conserved in both bacteria 140.
Cumulatively, these facts suggest that PknG performs a physiologically relevant function in all mycobacterial species besides the previously proposed virulence function in the pathogenic species.
M. tuberculosis displays a high level of intrinsic antibiotic tolerance to most available antibiotics, making treatment of M. tuberculosis infections difficult and
creating a requirement for long treatment schedules with a limited arsenal of useful
drugs 23. This intrinsic tolerance of antibiotics is mediated by a variety of factors including several drug degradation and inactivation processes that occur inside mycobacterial cells. Additionally, over 30 drug efflux transporters that have been shown to be involved in resistance to aminoglycosides, chloramphenicol, fluoroquinolones, isoniazid, linezolid, rifampicin, tetracycline and other toxic compounds also contribute to intrinsic drug resistance 32; 33; 34; 35. Finally, the extremely thick and multi-layered mycobacterial cell wall, which is composed of multiple layers of varied hydrophobicity and thereby poses an effective obstacle for the entry of most chemical compounds 3; 23; 26; 27.
Here we demonstrate a role for PknG in mycobacterial multiple drug
resistance, which appears to be conserved in both non-pathogenic and pathogenic
mycobacteria, thereby making PknG a direct link between antibiotic susceptibility
and intracellular persistence.
87
3.C. Results.
3.C.1. Identification of MAR4, a M. smegmatis pknG transposon mutant
displaying multidrug sensitivity.
A library of more than 5,000 transposon mutants was generated from wild
type M. smegmatis mc2155 179 by Himar1-mediated mutagenesis 168, and deposited in 96 well-plates. The library was subjected to exhaustive screens to identify
mutants sensitive to one or more antibiotics. This screening process has identified
genome-wide drug resistance determinants in mycobacteria, and a subgroup of the
obtained drug sensitive mutants encompassing those that displayed a decreased
Multiple Antibiotic Resistance phenotype was designated as the MARs. One of the
MAR mutants, MAR4, exhibited increased sensitivity to many currently clinically
important antibiotics when compared to wild-type M. smegmatis (Table 3-1). These
antibiotics are of diverse chemical structures and function, and include drugs
targeting cell wall synthesis (imipenem, vancomycin, and ethambutol), protein
synthesis (erythromycin), folate synthesis (sulfachloropyridazine), and
transcription (rifampicin) (Table 3-1). Of these, two (rifampicin and ethambutol)
are frontline antibiotics that are part of the standard regimen to treat tuberculosis.
E-test® drug sensitivity testing (AB Biodisk, Sweden) was performed 168 to obtain
representative minimal inhibitory concentrations (MICs) that determined that
MAR4 was 8-, 12-, 15-, 4-, and >8-fold more susceptible to erythromycin,
vancomycin, imipenem, ethambutol, and rifampin, respectively, than wild-type M.
smegmatis mc2155 (Table 3-1).
88
An arbitrary PCR method 168 first localized the transposon insertion in MAR4 to msmeg_0786, which encodes for a homolog of protein kinase G (PknGMsm), with
140 almost 80% identity to the M. tuberculosis homolog PknGMtb (Fig. 3-1). The
disruption of pknG in M. smegmatis MAR4 was further confirmed by PCR amplification using primers flanking the putative open reading frame (MS-PknG1 and MS-PknG2, Table 2-1 and arrows in Fig. 3-1A). The mutant gene generated a larger fragment corresponding to the inserted 1825 base pairs of the Himar1- transposon, which showed decreased mobility on an agarose gel (Fig. 3-1B).
Sequencing of the junction region from this PCR product identified the insertion site
at the dinucleotide TA1015-1016, introducing a stop codon after the triplet encoding
residue Ile338 (Fig. 3-1A, bottom). To confirm loss of the pknG gene product, Western
20 blots were done using a polyclonal antibody raised against PknGMtb . The antibody
recognized a protein band of ~ 78 kDa corresponding to the molecular weight of
2 PknGMsm in cell extract of wild type M. smegmatis mc 155 but failed to do so in
protein extract of MAR4, confirming the deletion of the pknG gene and loss of its
gene product (Fig. 3-1C).
3.C.2. Targeted deletion of pknG and complementation experiments confirm the MDR function of PknG in pathogenic and non-pathogenic mycobacteria.
Since secondary mutations or polar effects to downstream genes can cause false attribution of the drug susceptibility phenotype, a targeted pknG mutant (the
∆pknGMsm strain) was constructed by a modified version of the previously published recombineering approach 170 to independently assess if the multi-drug sensitive
89
Figure 3-1. Identification of MAR4, a M. smegmatis pknG transposon mutant.
(A) Synteny of the pknG loci in M. smegmatis and M. tuberculosis genomes and the
Himar1 transposon insertion in MAR4. Homologous genes are uniformly illustrated.
The nucleotide sequence at the pknG-Himar1 junction shows truncation of PknGMsm
338 at Ile . (B) PCR amplification using primers flanking pknGMsm (Table 2-1, arrows in panel A) from genomic DNA of wild-type M. smegmatis mc2155 and MAR4. (C)
Western blot assay using PknGMtb antibody that recognizes PknG in cell extracts of M.
bovis BCG and wild-type M. smegmatis mc2155 but not in MAR4 extract.
Reproduced with permission from 171. 90
Table 3-1. Susceptibility of M. smegmatis strains to antibiotics.
MIC (mg/liter)a Strain EM VA IP EB RI mc2155 32 48 6 0.25 >32 MAR4 4 4 0.38 0.064 4 mc2155/pknGK181M 12 16 1 0.064 4
ΔpknGMsm strain 3 4 0.38 0.064 4
K181M ΔpknGMsm/pknG strain 3 4 0.38 ND ND
ΔpknGMsm/pknGMtb strain 32 48 6 0.25 >32
ΔpknGMsm/pknGMsm strain 32 48 6 0.25 >32 aAbbreviations: EM, erythromycin; VA, vancomycin; IP, imipenem; EB, ethambutol; RI, rifampin; ND, not determined.
Reproduced with permission from 171.
91
phenotype of MAR4 was due to pknG disruption. Vector pVN701B expressing the mycobacteriophage Che9c recombination proteins gp60 and gp61 (which encode for homologs of RecE and RecT, respectively) from pJV53 170 and replicating from a
173 temperature-sensitive origin of replication (Orits) from pPR27 was constructed
(Fig. 3-2A). Chromosomal DNA sequences encompassing 551bp and 528bp of the 5’
and 3’-flanking regions, respectively, adjacent to pknGMsm were cloned into vector pYUB854 174 to flank the hygromycin-resistance cassette Ωhyg. The resulting pknG
deletion substrate (5’-pknG-flanking-region_ Ωhyg_3’-pknG-flanking-region) was
excised by SpeI/KpnI digestion (Fig. 3-2B). Homologous recombination promoted
by the induced mycobacteriophage system replaced pknGMsm with Ωhyg on the
chromosome, leading to hygromycin resistance in the resulting mutants (Fig. 3-
2B,D). Besides being resistant to both hygromycin and kanamycin (KanR carried by pVN701B), these transformants showed reduced growth on medium supplemented with sucrose (Fig. 3-2D, left panel), indicating the presence of pVN701B. Shifting cultures to 39°C in the absence of kanamycin removed pVN701B to prevent further undesired random recombination events (Fig. 3-2D).
Homologous recombination replacing chromosomal pknG with the hygromycin cassette in ∆pknGMsm mutant candidates was tested by PCR and sequencing using primers annealing to chromosomal regions adjacent to the homologous sequences of the pknGMsm flanking regions: PCR A (primers P1 and P2)
recovered the 5’-ΔpknG-Himar1 transition; PCR B (primers P3 and P4) recovered
the Himar1-3’-ΔpknG transition (Table 2-1 and Fig. 3-2B). All tested hygromycin
92
resistant transformants showed correct homologous recombination leading to
deletion of pknGMsm (Fig. 3-2C).
For complementation, plasmids expressing the M. smegmatis (pVN579) or M. tuberculosis (pVN578) pknG gene from the heat shock promotor (Table 2-2) 20 were
transformed into ΔpknGMsm, generating strains ΔpknGMsm/pknGMsm and
ΔpknGMsm/pknGMtb.
Next, reverse transcription PCR (Fig. 3-2E) was used to analyze the
transcription of pknG in all generated M. smegmatis strains. RNA was isolated and
reverse transcribed, followed by amplification of pknG cDNA using primers
RTpknG1 and RTpknG2 (Table 2-1) that recognized both M. smegmatis and M.
tuberculosis pknG sequences and thus allowed for monitoring of transcription of
either pknG gene into mRNA. Whereas transcripts of pknG were readily detected in
RNA samples isolated from wild type M. smegmatis mc2155, transcripts of pknG in
ΔpknGMsm were not detectable. Transformation of pVN578 or pVN579 restored transcripts of pknG (Fig. 3-2E).
Similarly, at the protein level, PknG was present in protein extracts of wild
type M. smegmatis and the complemented strains ΔpknGMsm/pknGMsm and
ΔpknGMsm/pknGMtb, but absent in ΔpknGMsm (Fig. 3-2F). Notably, compared to M. bovis
BCG (far left, Fig. 3-2F), a dramatic reduction of PknG expression was observed in wild type M. smegmatis, in agreement with previously published observations 140.
The reduced signal is likely caused by lower PknGMsm production in M. smegmatis
rather than by reduced specificity of the antibody (generated using PknGMtb), as in
93
trans expression of PknGMsm or PknGMtb from the same vector yielded comparable
signals on Western blots (lanes 4 and 5, Fig. 3-2F). The low expression of PknGMsm in
M. smegmatis was completely abolished in the ΔpknGMsm mutant (lane 3, Fig. 3-2F),
just as in the transposon mutant, MAR4 (Fig. 3-1C). Strains ΔpknGMsm/pknGMsm and
ΔpknGMsm/pknGMtb overproduced PknGMsm and PknGMtb, respectively (Fig. 3-2F).
Cumulatively, these results confirmed targeted deletion and complementation of pknG.
The generated strains allowed for analysis of PknGMsm function in antibiotic resistance. As before, antibiotic susceptibility was tested using E-test® diffusion
strips (AB Biodisk, Sweden). The antibiotic susceptibility level of ΔpknGMsm was identical to MAR4 for all tested antibiotics (Table 3-1). Importantly, in trans
expression of pknGMsm from vector pVN579 completely restored wild type levels of
antibiotic resistance to ΔpknGMsm (Table 3-1), confirming that the observed
phenotype is a direct function of pknGMsm expression. Expression of pknGMtb also
rescued ΔpknGMsm, suggesting that PknG provides a physiological, multiple drug tolerance function to not only non-pathogenic, but also to pathogenic mycobacteria
(Table 3-1).
Next, multiple drug susceptibility was assayed in M. tuberculosis H37Rv,
20 where phenotypes for ΔpknGMtb are similar to those observed in M. smegmatis, with increased susceptibility to the representative drugs erythromycin (Fig. 3-3A) and sulfachloropyridazine (Fig. 3-3B). Again, expression of pknGMtb restored wild-
94
Figure 3-2. Construction and complementation of the ΔpknGMsm strain. (A)
Construction of pVN701B. (B) The pknGMsm deletion cassette and pknG locus in wild-
2 type M. smegmatis mc 155 and its derived ΔpknGMsm strain. (C) PCR confirmation of
pknG deletion in eight random mutant candidates. (D) Removal of pVN701B by a
temperature shift resulted in kanamycin sensitivity and sucrose resistance of the
ΔpknGMsm strain. (E) Reverse transcription (RT)-PCR detection of pknG transcription
in M. smegmatis strains. (F) Western blot detection of PknG in M. smegmatis strains.
Reproduced with permission from 171. 95
Figure 3-3. Antibiotic susceptibilities of M. tuberculosis strains. (A) Growing
cultures (10 μl; optical density at 600 nm, 1.0) of wild-type M. tuberculosis H37Rv,
its derived ΔpknGMtb mutant, and the complemented ΔpknGMtb/pknGMtb (pVN578)
strain were plated on solid 7H10-OADC medium 19 in the absence (−) or presence
(+) of erythromycin (100 mg/liter) and incubated at 37°C for 4 weeks. (B) M. tuberculosis H37Rv (circles), the ΔpknGMtb strain (triangles), and the
ΔpknGMtb/pknGMtb strain (squares) were grown at 37°C for 7 days with shaking (200 rpm) in liquid 7H9-OADC 168 supplemented with increasing concentrations of
sulfachloropyridazine. Growth was recorded as optical density at 600 nm 19
normalized to the absorbance of control (no antibiotic).
Reproduced with permission from 171. 96
type levels of antibiotic resistance to the deletion mutant (Fig. 3-3). These results
confirm that PknG indeed performs similar functions in multiple drug tolerance of
pathogenic as well as non-pathogenic mycobacterial species.
Finally, to test whether the kinase activity of PknG is required for its function
K181M 181 20 in antibiotic tolerance, a kinase-dead form of PknGMtb, PknG (Lys → Met) , was introduced into wild type M. smegmatis (plasmid pPknGK181M, Table 2-2).
Antibiotic susceptibility assays showed that overproduction of this dominant negative PknGK181M 20 form resulted in reduction of antibiotic resistance of wild type
M. smegmatis, similar to the effect caused by pknG deletion (Table 3-1). This result strongly supports the role of PknG phosphorylation activity in intrinsic multiple drug resistance of mycobacteria.
3.C.3. Loss of pknG leads to altered cell surface properties.
Proliferation of wild-type M. smegmatis and ΔpknGMsm in vitro was monitored by optical density measurements at a wavelength of 600nm. While no differences in growth rates in either 7H9 or biofilm liquid medium 112 could be observed between wild type and ΔpknGMsm mutant in non-static growth conditions
(Fig. 3-4A), increased cell-cell aggregation compared to wild type was observed for the mutant strain. Quantification of cellular aggregation showed an increase by 53%
(Table 3-2) in ΔpknGMsm compared to wild type. In trans expression of PknGMsm
rescued the aggregation phenotype of the mutant.
97
Figure 3-4. Loss of PknG does not affect growth rate in liquid, shaking medium.
2 In vitro growth of wild type M. smegmatis mc 155 (squares) and ΔpknGMsm (circles) in 7H9 (filled symbols) and biofilm liquid medium 112 (open symbols). Saturated
cultures growing in 7H9 medium were used to inoculate fresh medium to initial
optical density at 600nm (OD600) of 0.2 and incubated at 37ºC with shaking
(250rpm). At indicated time intervals, growth of the cultures was monitored by measuring OD600.
98
Table 3-2. Surface properties of M. smegmatis strains.
Strain
2 Surface Property mc 155 ΔpknGMsm ΔpknGMsm/pknGMsm
Cellular 18.8 +/- 3.6 72.0 +/- 2.5 24.7 +/- 1.3 Aggregationˆ
27.3 +/- 2.41 13.63 +/- 5.34 29.79 +/- 2.96 Hydrophobicity Index˜
Congo Red 224 +/- 45 119 +/- 16 257 +/- 48 Binding˚
Zeta Potential¯ -20.47 +/- 1.26 -27.13 +/- 1.57 -23.67 +/- 0.71
ˆ Percentage of aggregated cells ˜Arbitrary units, expressed as the percentage of cells recovered in the organic phase. ˚10−4*A488 ¯ Expressed in mV
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To assess whether this was due to alterations in hydrophobicity of the cell
wall, the hydrophobicity index of the strains was measured by hexadecane partition
procedure 183. Single cell suspensions adjusted to units of 1 OD were mixed with
various volumes of hexadecane and the hydrophilic phase recovered and assessed
for reduction in optical density. The results indicated that the surface of ΔpknGMsm is
more hydrophilic than that of wild type, and ΔpknGMsm/pknGMsm was rescued completely (Table 3-2). Further confirmation of altered hydrophobicity was obtained by Congo Red binding 183. Congo Red is a hydrophobic diazo-dye that binds
lipids, lipoproteins and other macromolecules found in the mycobacterial cell wall
and has been used as an indicator of phenotypic drug susceptibility 188. Consistent
with the observed decrease in hydrophobicity index of ΔpknGMsm, ΔpknGMsm bound only half as much Congo red as wild type or complemented cells (Table 3-2).
To assess whether the loss of PknG also results in charge alterations on the cell surface, zeta potential measurements were performed on single cell
183 suspensions as previously described using a Malvern ZetaMaster . ΔpknGMsm
showed an increase in negative cell surface charge over wild type by 34.8%, and in
trans expression of PknGMsm restored wild type charge to ΔpknGMsm (Table 3-2).
100
3.D. Discussion.
The presented work constitutes a comprehensive set of evidence suggesting
that PknG plays a role in intrinsic mycobacterial resistance to multiple antibiotics of diverse structures, targets and functions. Both transposon-mediated (MAR4) and targeted mutagenesis (ΔpknGMsm and ΔpknGMtb) resulted in mutant strains with increased multiple antibiotic susceptibility (Table 2-3) in both non-pathogenic and pathogenic mycobacterial species. The non-specific sensitivity of MAR4, ΔpknGMsm
and ΔpknGMtb to antibiotics with diverse chemical structures and mechanisms of action (Table 2-3, Fig. 3-3) suggests that PknG controls either (i) the activities of dedicated multiple drug resistance mechanisms such as efflux pumps, (ii) the integrity of the mycobacterial cell envelope through PknG involvement in essential cellular functions, or (iii) the physiology and fitness of mycobacteria. The results obtained in this study support the latter hypothesis, as strain ΔpknGMsm showed altered cell wall characteristics (Table 2-4). Loss of PknGMsm resulted in a cell wall both more hydrophilic and more negatively charged than that of wild type M.
smegmatis (Table 2-4). Reduced cell wall hydrophobicity has previously been
reported to be important for multiple antibiotic tolerance in mycobacteria 168; 189.
Additionally, recent findings that PknG is involved in the tricarboxylic acid cycle as well as glutamine utilization in Corynebacterium and Mycobacterium also favor this possibility 121; 122; 124. Glutamine and its derived molecules, such as polyglutamines, may serve as components that are required for the integrity of the mycobacterial cell wall, thereby helping to pose an effective barrier to diffusion of antibiotics of
101
diverse size and polarity. Furthermore, the likelihood that PknG does not directly
control dedicated MDR systems is supported by the fact that increased expression of
neither PknGMtb nor PknGMsm in the ΔpknGMsm strain (lanes 4 and 5 in Fig. 3-2F)
leads to an increase in intrinsic resistance to any of the tested antibiotics compared
to wild-type M. smegmatis (Table 2-3).
Expression analyses revealed that the amount of PknG protein detectable in
the cytoplasm of M. smegmatis is dramatically reduced in comparison to those found
in M. tuberculosis or M. bovis BCG (Fig. 3-2F). These results are in agreement with a
recently published observation that PknG expression is translationally regulated in
mycobacteria 140, suggesting the existence of regulatory mechanisms to control
expression levels of PknG in mycobacterial species. A dramatic increase of PknGMtb
expression may have provided pathogenic mycobacteria with an additional amount of the protein required for its new function in virulence 20. This possibility was also suggested in a recent study that addressed differential expression of PknG in various mycobacterial species in relation to its virulence function 140.
Together with previous reports 20; 124; 140, our data suggest that signaling
circuitries surrounding PknG may serve as a phenogenetic network that controls
persistence in the infected host and intrinsic antibiotic resistance to multiple drugs
in mycobacteria 20.
102
3.F. Acknowledgements
We thank Jean Pieters, Julia van Kessel and Graham F. Hatfull for providing
materials; Abram Stavitsky and Kien Nguyen for critical reading of the manuscript.
Support was provided by start-up funds from the School of Medicine, a STERIS
Infectious Diseases Research Grant and a CFAR Developmental Award from Case
Center For AIDS Research. Sam Ogwang is a recipient of a Forgarty AIDS
International Training and Research Program (AITRP) Fellowship.
103
Chapter 4.
Modulation of Folate Metabolism by Protein Kinase G Determines
Biofilm Growth and Subcellular Localization of Pathogenic
Mycobacteria in Macrophages
Kerstin A. Wolff, Hoa T. Nguyen and Liem Nguyen
Department of Molecular Biology and Microbiology, School of Medicine, Case Western
Reserve University, Cleveland, Ohio 44106
104
4.A. Abstract
The eukaryotic-like serine/threonine protein kinase G (PknG) of M. tuberculosis has been shown to be involved in preventing lysosomal fusion to
phagosomes containing the bacilli in infected host macrophages, thereby creating an
intracellular niche. It was therefore suggested that PknG serves as a virulence factor
that directly modifies host signaling cascade proteins. Other reports however have
shown involvement of PknG, which is highly conserved in Actinobacteria, in
glutamate metabolism, cell surface properties, and intrinsic multiple antibiotic
tolerance in both non-pathogenic and pathogenic mycobacteria.
Here we present evidence that PknG regulates mycobacterial folate
metabolism in late stage growth conditions, and is required for survival of folate
starvation and biofilm growth. We also demonstrate that inhibition of PknG has the
potential to be exploited to potentiate classical antifolate drugs that target folate
biosynthesis in other steps for use against M. tuberculosis. Lastly, we present
evidence suggesting that the previously demonstrated role of PknG in inhibition of
phagolysosomal fusion is mediated through regulation of mycobacterial folate levels
rather than modification of host signaling cascades.
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4.B. Introduction
Upon infection of host macrophages, M. tuberculosis colonizes the host
phagosome by preventing acidification of this compartment 10; 12; 190 and modulation
of other antimicrobial mechanisms of the macrophage 191. Acidification normally
occurs via fusion of lysosomal compartments to the bacteria-containing phagosomes,
and colonization of the phagosome is achieved by arresting its maturation and
preventing lysosomal delivery 10; 12; 190; 192. The underlying mechanisms of this arrest
have not yet been fully elucidated, but both host and mycobacterial factors play
crucial roles in the process 4; 5; 17; 18; 19. One such factor is the soluble eukaryote-like
serine/threonine kinase PknG of M. tuberculosis that has been shown to play a
critical role in M. tuberculosis’ ability to establish the phagolysosomal fusion block,
though its mechanism of action and targets remain unknown 20; 137. In mouse
macrophage infections, both genetic deletion of pknG and chemical inhibition of
PknG by a specific inhibitor, AX20017 20; 138, in M. bovis BCG resulted in increased
fusion of lysosomes to phagosomes containing mycobacteria, and this correlated
with decreased survival of internalized mycobacteria (Fig. 1-5, 1-6) 20; 23. It has therefore been proposed that PknG is actively secreted into the macrophage cytosol where it acts as a virulence factor and modifies eukaryotic host substrates that are active in signalling cascades necessary for establishment of lysosomal trafficking to the colonized phagosome 20.
Recently it has been shown that PknG is also involved in bacterial glutamate
metabolism 121; 122; 124, maintenance of mycobacterial cell surface properties 171, as
106
well as intrinsic multiple drug resistance 171, determining PknG as a genetic factor underlying the often observed phenotypic association of virulence and antibiotic resistance. One example of this association is biofilm growth, which has been suggested to represent a growth condition similar to static growth during persistence of mycobacteria in the host 104; 105; 106; 107; 112; 113. Bacteria growing in biofilms, like persistent mycobacteria during latent infection, display enhanced tolerance for antibiotics, a phenomenon that is likely due to the metabolic differences between biofilm growth and planktonic growth 104; 105; 106; 107.
Mycobacteria, including M. tuberculosis, have been shown to form biofilms both in vivo and in vitro 111; 112; 113; 114, and biofilms of Mycobacterium chelonae and
Mycobacterium ulcerans have been shown to play a role in pathogenesis 114; 115; 116;
117. Consistent with the idea that biofilm growth represents a late static growth stage similar to host persistence, mycobacterial biofilms also exhibit higher antibiotic tolerance than planktonic mycobacteria, similar to persistent M. tuberculosis during latent infection 22, and biofilm of M. tuberculosis has been linked
to the formation of drug-tolerant persisters 113.
Here we present further evidence suggesting that the role of PknG in
phagolysosome biosynthesis is mediated through the physiology of the bacillus
rather than a role in host signaling control. We show that PknG regulates folate
metabolism in the bacterium, specifically in late stage growth conditions that mirror
latent infection and persistence inside the host macrophage. We demonstrate that
regulation of folate metabolism by PknG is required for mycobacterial survival of
107
folate starvation induced by classical antifolate antibiotics and for biofilm growth.
We also present evidence that inhibition of PknG phosphorylation has the potential
to be exploited for boosting the antimycobacterial activity of classical antifolates.
Lastly, we show that the previously demonstrated inhibition of phagosome- lysosome fusion in infected macrophages by PknG 20 is mediated through regulation of bacterial folate levels, rather than through active modification of eukaryotic host proteins in signaling cascades.
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4.C. Results
4.C.1. PknG is required for survival of folate starvation.
We had previously identified pknG, the gene encoding for the eukaryotic-like serine/threonine protein kinase G in mycobacteria, in a transposon mutagenesis screen for genome-wide drug resistance determinants in mycobacteria. We had confirmed the requirement for PknG kinase activity for intrinsic tolerance of M. tuberculosis to many clinically important antibiotics. This observation suggested a physiological function of PknG based on modification of mycobacterial substrates rather than a virulence function based on phosphorylation of host signaling factors
171. The identified drug classes included antifolates 171, a group of antibiotics that encompasses both sulfadrugs and trimethoprim and inhibits de novo folate biosynthesis. In a later, independent transposon mutagenesis screen that looked for antifolate resistance determinants in mycobacteria 102 pknG was identified again, further indicating that PknG plays a role in susceptibility of mycobacterial species to
this group of antibiotics. The previously observed multi-drug tolerance function
thus seemed to be a consequence of a defect in folate metabolism 171.
171 As previously shown , deletion of pknGMtb led to increased susceptibility to the sulfadrug sulfachloropyridazine when compared to wild type M. tuberculosis
H37Rv (Fig. 4-1A, upper row). To exclude polar effects on downstream genes, we
expressed pknGMtb in trans from plasmid pVN578 (Table 2-2) in the ∆pknGMtb mutant strain (strain ∆pknGMtb/pknGMtb) and could completely restore wild-type levels of
sulfachloropyridazine resistance (Fig. 4-1A, upper row). Notably, supplementation
109
of exogenous folinic acid, a storage form of folate, bypassed the requirement for
PknG (Fig. 4-1A, bottom row), indicating that the function of PknG in antifolate
resistance is directly related to de novo folate biosynthesis or folate metabolism.
Similar results were observed in M. smegmatis (Fig. 4-2A), indicating that this role of
PknG is conserved in both pathogenic and non-pathogenic mycobacteria.
4.C.2. Chemical inhibition of PknG activity potentiates classical antifolate
antibiotics.
Potentiation of currently available TB drugs by targeting resistance mechanisms to
these agents has recently gained attention because of the rapid spread of MDR and
XDR TB spread that has limited the current options for TB chemotherapy 44. We sought to explore whether the kinase activity of PknG mediates its function in antifolate resistance, and as a result, whether chemical inhibition of PknG activity could be exploited to potentiate the antimycobacterial activity of classical antifolates. For this purpose, the PknG-specific kinase inhibitor AX20017 20 that
specifically inhibits PknG activity (Fig. 4-1B) was titrated along with increasing
concentrations of sulfachloropyridazine (Fig. 4-1C). While inhibition of PknG in the
absence of antibiotics did not affect growth of wild-type M. tuberculosis H37Rv,
addition of AX20017 increased the antimycobacterial activity of
sulfachloropyridazine in a concentration-dependent manner (Fig. 4-1C). These data
indicate that not only the kinase activity of PknG is required for intrinsic
mycobacterial tolerance to antifolates, but also that this mechanism of resistance
110
Figure 4-1. Requirement of PknGMtb activity for survival of folate starvation and exploitation for antifolate potentiation. (A) Sulfachloropyridazine susceptibility of M. tuberculosis strains. Growing cultures (10μl; optical density at
600 nm, 1.0) of wild-type M. tuberculosis H37Rv, its derived ΔpknGMtb mutant, and
111
the complemented ΔpknGMtb/pknGMtb (pVN578) strain were plated on solid 7H10-
OADC medium 19 containing increasing amounts (0-5µM) of sulfachloropyridazine
(SL) and either 0 or 1mM folinic acid (FA) and incubated at 37°C for 4 weeks. (B) In vitro kinase assay using radioactive [γ-32P]-ATP to assess autophosphorylation activity of purified PknGMtb-6xHis (0.5µg). 15 minute pre-incubation with 5mM
AX20017 markedly reduced inhibited kinase activity of PknG. (C) Potentiation of sulfachloropyridazine activity by chemical inhibition of PknG. Serial dilution of growing H37Rv culture (2μl volumes) were spotted in triplicate onto solid 7H10-
OADC medium containing increasing amounts of sulfachloropyridazine (SL) and
AX20017 (0-2mM). Growth was recorded after 3 weeks of incubation at 37°C.
112
Figure 4-2. Requirement of PknGMsm activity for survival of folate starvation. (A)
Sulfachloropyridazine susceptibility of M. smegmatis strains. Growing cultures (50µl;
2 optical density at 600 nm, 0.6) of wild-type M. smegmatis mc 155, its derived ΔpknGMsm
mutant, and the complemented ΔpknGMsm/pknGMsm (pVN579) and ΔpknGMsm/pknGMtb
(pVN578) strains were plated on solid NE medium containing 0 or 1mM folic acid (FA),
paper discs containing 0.25µg of sulfachloropyridazine (SL) applied and incubated at
37°C for 4 days. (B) Potentiation of sulfachloropyridazine activity by chemical inhibition
of PknG. Growing cultures (50µl; optical density at 600 nm, 0.6) of wild-type M.
smegmatis mc2155 were plated onto solid NE medium and paper discs containing 0.25µg
sulfachloropyridazine (SL) and 0.1 µg kinase inhibitor chelerythrine (Che) and incubated
at 37°C for 4 days.
113
can be targeted by chemical inhibition in order to potentiate classical antifolate
antibiotics against mycobacteria.
4.C.3. PknG is required for biofilm growth.
We observed a greater increase in susceptibility of ∆pknGMtb to antifolates and other antibiotics in late-stage growth than in log-phase 171, suggesting importance of PknG involvement in folate regulation in late-stage or latent infection- like conditions. Since biofilm growth has been shown to be correlated with the ability to form persisters displaying increased antibiotic tolerance in M. tuberculosis
113, we sought to explore whether biofilm behavior might be altered upon loss of
pknGMtb.
Biofilm assays using a liquid-air interface model 112; 113 indeed showed that the ∆pknGMtb strain showed a delay in formation of mature biofilms compared to wild-type M. tuberculosis H37Rv (Fig. 4-3A, left), corresponding with decreased biofilm biomass (Fig. 4-3A, right). In trans expression of pknGMtb rescued the mutant
phenotype (Fig. 4-3A). Strikingly, supplementation of exogenous folinic acid in the
medium reestablished normal biofilm growth in the ∆pknGMtb strain (Fig. 4-3A).
Similar results were obtained in M. smegmatis (Fig. 4-4A). These results suggest that
PknG plays a role in biofilm formation through regulation of folate biosynthesis.
To assay whether kinase activity of PknG was required, AX20017 was added
to biofilm cultures of both wild type and ∆pknGMtb/pknGMtb strains, which resulted in
immature biofilms closely resembling that of the ∆pknGMtb strain (Fig. 4-3B).
Likewise, inhibition of PknGMsm by the Pkcα inhibitor chelerythrine yielded similar
114
115
Figure 4-3. PknGMtb is required for folate-dependent biofilm growth in M. tuberculosis. (A) Biofilm growth of wild type M. tuberculosis H37Rv, its derived
ΔpknGMtb mutant, and the complemented ΔpknGMtb/pknGMtb (pVN578) strain was
assayed in the presence or absence of 1mM folinic acid (FA). Biofilm was pictured at
6 weeks (left). Surface biofilm growth was quantified at week 6 by measuring total
protein per biofilm plate (right). Supplementation of folinic acid bypassed the
requirement for PknG. (B) Surface biofilm growth of M. tuberculosis H37Rv strains
was assayed in the absence or presence of 1mM AX20017 (PknG inhibitor). Growth
was recorded at week 6 after inoculation. (C) Effect of folate starvation on biofilm formation. Biofilm growth of wild type M. tuberculosis H37Rv was assayed in the presence of increasing amounts of the antifolate drug sulfachloropyridazine (SL) and either 0 or 1mM folinic acid. Growth was quantified at week 6 after inoculation by measuring total protein per biofilm plate.
116
Figure 4-4. PknGMsm is required for folate-dependent biofilm growth in M. smegmatis. (A) Biofilm growth of wild type M. smegmatis mc2155 and its derived
ΔpknGMsm mutant was assayed in the presence or absence of 1mM folic acid (FA).
Biofilm was pictured after 1 week (left). Surface biofilm was quantified after 1 week of growth by measuring total protein per biofilm plate (right). Supplementation of folate bypassed the requirement for PknGMsm. (B) Surface biofilm growth of M. smegmatis mc2155 was assayed in the absence or presence of 1mM chelerythrine
117
(Che). Growth was recorded at week 1 after inoculation (left) and quantified by colony forming unit (CFU) measurement of surface biofilm.
118
results in an M. smegmatis background (Fig. 4-4B). PknG kinase activity is therefore
required for normal biofilm growth.
To further clarify whether folate availability mediates normal biofilm
formation, biofilm assays were conducted in the presence of sulfachloropyridazine.
Folate starvation in wild-type M. tuberculosis H37Rv led to decreased biofilm
formation in a dose-dependent manner, and could be overcome by supplementation of exogenous folinic acid (Fig. 4-3C).
Cumulatively, these data support a model in which biosynthesis and availability of folate are crucial in late-stage growth such as in the biofilm setting. In this condition, PknG kinase activity is required to establish this required folate level by an unknown mechanism.
4.C.4. PknG is involved in regulation of lysosomal delivery in macrophages via modulation of mycobacterial folate metabolism.
In light of our data and the implied importance of mycobacterial substrates of PknG, we sought to determine whether an alternate explanation to the proposed virulence role of PknG in phagolysosomal fusion 20 might be the requirement for mycobacterial folate biosynthesis in late-stage growth of M. tuberculosis infections.
Failure to produce sufficient amounts of folate would have led to decreased survival, which would also account for increased processing and clearance by infected macrophages upon loss of PknG function, as observed with heat-killed bacteria 20.
The increase in phagolysosomal fusion we observed in bone marrow derived
murine macrophages infected with ∆pknGBCG strain was comparable to previously
119
120
Figure 4-5. PknG is involved in lysosomal delivery to infected phagosomes through folate levels. Bone marrow derived murine macrophages were infected with GFP-expressing M. bovis BCG or its derived ΔpknGBCG mutant (green) and
chased with either 2mM folinic acid (FA) or 30µM sulfachloropyridazine (SL) before
staining with DAPI (blue) and Neutral Red (red). Fixed macrophages were pictured
at 170x magnification (top) and colocalization quantified (bottom). Folinic acid
reestablished failure of macrophages to deliver lysosomes to phagosomes in cells
infected with ΔpknGBCG and in cells infected with BCG that was folate-starved by classical antifolate antibiotics.
121
published observations 20 (Fig. 4-5). Supplementation of exogenous folinic acid in the medium reestablished the inability of infected macrophages to deliver lysosomes to phagosomes containing mycobacteria (Fig. 4-5), bypassing the requirement for PknG. Furthermore, treatment of infected macrophages with sulfachloropyridazine, which targets the bacteria, but not the host cells, led to increased lysosomal delivery, similar to that observed in the absence of PknG (Fig.
4-5).
These results indicate that latent growth conditions such as those of biofilm or macrophage infections require folate biosynthesis that is involved in lysosomal trafficking to phagosomes containing pathogenic mycobacteria, and that PknG is necessary for this process. It also indicated that substrates for PknG might be prokaryotic in nature.
122
4.D. Discussion
In this report we present evidence suggesting a novel function of PknG in the
physiology and metabolism of mycobacterial species. We demonstrate that PknG
regulates mycobacterial folate metabolism, which is of particular importance in static growth conditions. We also show that this regulation of folate is required for mycobacterial persistence to classical antifolate antibiotics, biofilm growth, and, most importantly, for successful prevention of lysosomal delivery of pathogenic mycobacteria after internalization by host macrophages 20. It was previously
suggested that PknG may be actively secreted into the host cell, where it interferes
with host signalling cascades, possibly involving Pkcα 127, thereby mediating the
control over phagolysosomal fusion and aiding the survival of pathogenic
mycobacteria in macrophages 20. Our results, on the other hand, indicate a mycobacterial function for PknG instead that is in agreement with previous reports proposing roles for PknG in glutamate metabolism, multiple antibiotic tolerance and cell wall maintenance 121; 122; 124; 171.
While the intracellular bacilli that persist during latent TB infections are generally regarded to be non-replicating and growing very slowly 193; 194; 195, recent reports have shown that they are metabolically active 196 and should therefore require folate for the operation of metabolically essential one-carbon transfer
reactions 45; 77; 78; 79. Our results indicating that modulation of mycobacterial folate
metabolism by PknG is most important in conditions of static growth, are in agreement with the observations that pknG is upregulated during latent infection in
123
mice 196, and that is plays a role in stationary phase metabolism 196; 197. Dependence on folate levels in an intracellular setting would also account for the presence of pknG in the obligate intracellular pathogen M. leprae, which has reduced its genome to the bare minimum. It may also be the reason for the generally higher expression levels of PknG in pathogenic mycobacterial species 140. The involvement of PknG in
multiple antibiotic resistance and cell wall integrity of both non-pathogenic and
pathogenic bacteria 157 is mediated through bacterial folate levels. This may partially account for the association of stationary phase growth, latency, and biofilm
growth with higher levels of antibiotic resistance to a variety of drugs 22; 13; 14; 15; 16.
Non-pathogenic mycobacteria do not normally infect eukaryotic hosts. The
conservation of pknG in such typically environmental, saprophytic species may be accounted for by a requirement to frequently form biofilms, for which PknG activity is also required.
We also present evidence that, chemical inhibition of PknG leads to potentiation of classical antifolate antibiotics against M. tuberculosis. Therefore,
PknG provides a potential target for development of potentiators that boost activity of classical antifolates 45. Considering the role of PknG in regulation of folate metabolism that is required for preventing lysosomal delivery of pathogenic mycobacteria, co-administration of PknG inhibitors with antifolates might not only result in potentiation of antimycobacterial activity of TB drugs, but also promote
phagosome-lysosome fusion, leading to improved killing and clearance of M.
tuberculosis by host phagocytic cells. It is therefore critical to further study the
124
pathways that PknG is involved in, as well as to identify its targets and molecular mechanism of action.
125
4.E. Acknowledgements.
We thank Jean Pieters (BCG/GFP, ∆pknGBCG, ∆pknGMtb), Julia van Kessel and
Graham F. Hatfull (pJV53), Bryan Doreian and Kurt Lu (mouse bone-marrow derived macrophages). Support was provided by an NIH R01 grant (AI 087903), start-up funds from the School of Medicine, a STERIS Infectious Diseases Research
Grant, and a CFAR Developmental Award from Case Center For AIDS Research.
126
Chapter 5.
Protein Kinase G Regulates Folate Metabolism in Mycobacteria
Through Phosphorylation of Ribosomal Protein RplM
Kerstin A. Wolff and Liem Nguyen
Department of Molecular Biology and Microbiology, School of Medicine, Case Western
Reserve University, Cleveland, Ohio 44106
127
5.A. Abstract
We have previously demonstrated that the eukaryotic-like serine/threonine protein kinase G (PknG) of M. tuberculosis modulates folate metabolism, cell wall
integrity, intrinsic multiple drug resistance, biofilm formation, and lysosomal
trafficking to phagosomes, allowing for persistence of pathogenic mycobacteria in
host macrophages. This modulation of folate metabolism is mediated through its
kinase activity.
In this study, we sought to determine the mycobacterial substrate(s) and
pathway(s) targeted by PknG. We identify a novel substrate for PknG, the RplM
protein of the large ribosomal subunit. RplM is phosphorylated by PknG on
threonine 11 that is conserved in mycobacterial species. RplM co-purifies with
MutT3, a putative Nudix hydrolase identified as an effector of PknG function in
folate-dependent biofilm growth. We hypothesize that MutT3 functions as a
dihydroneopterin-triphosphate pyrophosphatase that catalyzes the first committed
step of folate de novo biosynthesis.
128
5.B. Introduction
In previous chapters, we demonstrated that the prokaryotic-like
serine/threonine kinase G (PknG) is required for antibiotic tolerance in both M.
tuberculosis and non-pathogenic mycobacteria 171, where it also regulates folate
metabolism that is required for static growth resembling M. tuberculosis persistence inside the host macrophage. PknG-mediated regulation of folate metabolism is required for mycobacterial survival of folate starvation triggered by classical antifolate antibiotics, as well as for biofilm growth and the successful prevention of
phagolysosome biosynthesis in infected macrophages 20. These results suggest that
the substrate(s) of PknG is likely a mycobacterial protein, rather than host proteins
acting in signaling cascades 20; 139. We therefore sought to elucidate the bacterial pathways and substrates that are affected by PknG, and, more specifically, how PknG modulates folate metabolism in mycobacteria.
In this chapter, we identify RplM as a novel substrate for PknG. RplM, also known as L13, is an essential ribosomal protein and is a component of the large ribosomal subunit 157. It has also been shown to act in sub-ribosomal particles 160
and outside the ribosomal context 161. RplM has been found to directly interact with
Obg, an essential GTP-binding protein that is required for induction and regulation
of the σB-mediated stress response in B. subtilis 164; 165. RplM has also been detected
in active sub-ribosomal particles that bind both 23S and 5S rRNA in Thermus
aquaticus 160. In a non-ribosomal setting, RplM has been demonstrated to be
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involved in anti-termination together with ribosomal proteins of the small and large subunits in E. coli 161. RplM is phosphorylated on several residues in E. coli, although
the involved kinases remain unknown 167. These phosphorylated residues are located on
the solvent interface of RplM, suggesting that they may be important for interactions with
ribosome-associated proteins 167. We show here that mycobacterial RplM is
phosphorylated by PknG on threonine 11, which is exclusively conserved
throughout the Mycobacterium genus.
RplM is co-purified with MutT3, a putative Nudix hydrolase and an effector of
PknG. MutT motif hydrolases, also known as Nudix hydrolases, catalyze the
hydrolysis of a nucleoside diphoshate linked to a variable moiety x, removing
pyrophosphate 143; 144. Their manifold substrates include nucleoside triphosphates,
dinucleoside polyphosphates, nucleotide sugars and co-enzymes 143; 144. Most Nudix hydrolases seem to be involved in the clearance of potentially deleterious or hazardous molecules, acting as “housekeeping” enzymes 143; 144; 146. Some Nudix
hydrolases however have been shown to act in metabolic pathways, such as ppGpp
regulation in response to amino acid starvation 149 and the catalysis of the removal
of pyrophosphate from dihydroneopterin-triphosphate in de novo folate biosynthesis 147; 148. We hypothesize that mycobacterial MutT3 functions as a dihydroneopterin-triphosphate pyrophosphatase (DHN-3Pase) catalyzing this first committed step in the pathway, in accordance with previous observations that this
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Nudix hydrolase does not appear to function as either a mutator protein or an 8-
oxo-guanosine triphosphatase 156.
We also present evidence that phosphorylation of threonine 11 on RplM and
the presence of MutT3 are required for antifolate resistance and biofilm formation.
These results indicate a signaling cascade involving PknG, RplM and MutT3 that is required for modulation of folate metabolism. This signaling cascade is important for static growth conditions including biofilm surfaces and the phagosomal niche within host phagocytic cells.
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5.C. Results
5.C.1. PknG does not mediate regulation of folate metabolism through phosphorylation of GarA.
Only one bacterial substrate has so far been identified for PknG. PknG of C.
glutamicum phosphorylates OdhI, which then inhibits 2-oxoglutarate
dehydrogenase (ODH) complex 121; 123. A later study confirmed that, along with
PknB, PknG can phosphorylate GarA, the mycobacterial homolog of OdhI, in vitro 122.
GarA, in its unphosphorylated state inhibits action of ODH complex, as is the case in
C. glutamicum, and, depending on substrate concentration, either inhibits or
activates glutamate dehydrogenase (GDH) 122.
To address regulation of folate biosynthesis by PknG through
phosphorylation of GarA, both limitations in CO2 and glutamate had to be addressed.
Since ODH complex is inhibited by unphosphorylated GarA 122, pknG deletion should lead to inhibition of ODH and therefore decreased levels of CO2. We therefore performed biofilm assays in the presence of 5% CO2 to determine whether the
defect in biofilm formation of ∆pknGMsm could be bypassed by exogenous
supplementation of CO2, as had been shown with an ODH mutant 113. While biofilm formation overall was accelerated, loss of PknG still led to a significant delay in the mutant (data not shown). It therefore appears that CO2 is important for biofilm formation, but not sufficient to compensate for the lack of PknG in mycobacteria.
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As GarA also acts as an inhibitor of GDH 122, lack of PknG should lead to decreased levels of glutamate. Since glutamate is a building block of folate, we quantified biofilm formation of ∆pknGMsm in the presence and absence of exogenous
2 folinic acid. While GarA overexpression (strain mc 155/garAMsm) indeed leads to a
biofilm formation defect similar to that resulting from loss of PknG, supplementation of
exogenous folate does not at all alleviate the biofilm defect of the GarA overexpressing
strain (Fig. 5-1). Therefore, phosphorylation of GarA by PknG is not how PknG controls folate biosynthesis.
5.C.2. mutT3 is required for survival of folate starvation and biofilm growth.
To examine mycobacterial proteins involved in folate biosynthesis that might be regulated by PknG, this pathway was investigated by SSDB (Sequence Similarity
Database) gene cluster search at Kyoto Encyclopedia of Genes and Genomes (KEGG: http://ssdb.genome.jp/ssdb-bin/ssdb_gclust?org_gene=mtu:Rv0410c&type=g). The
only step in folate biosynthesis in mycobacteria for which the catalytic enzyme
remains unknown is the conversion of dihydroneopterin-triphosphate (DHN-3P) to
dihydroneopterin (DHN) (Fig. 5-2). This conversion had been thought to occur in a
spontaneous chemical reaction until 2007, when a dedicated Nudix hydrolase was
identified as the catalytic enzyme in L. lactis and Arabidopsis 148. KEGG suggested that this reaction is catalyzed in mycobacteria by the putative Nudix enzyme MutT3.
MutT3 is encoded only three genes upstream of pknG by Rv0413 and msmeg_0790 in
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Figure 5-1. PknG does not mediate control over folate metabolism through phosphorylation of GarA. Biofilm growth of M. smegmatis strains. Surface biofilm
2 growth of M. smegmatis mc 155, its derived ΔpknGMsm mutant, and the
2 overexpressing mc 155/garAMsm strain was assayed in the absence or presence of
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1mM folinic acid (FA). Surface biofilm growth was recorded at week 1 after inoculation and quantified by measuring total protein per biofilm plate.
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Figure 5-2.Folate de novo biosynthesis pathway. De novo folate biosynthesis pathway in M. tuberculosis. Intermediates are shown in green boxes, classical antifolate antibiotics in orange ellipses, enzymes in black.
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M. tuberculosis and M. smegmatis respectively, and shares the same intergenic region in a locus conserved across the mycobacterial genus and some related actinobacteria (Fig. 5-3A and Fig. 5-4A). Furthermore, proximity of mutT3 and pknG indicates similar function or involvement in the same pathways.
Strain ∆mutT3Mtb showed increased susceptibility to classic antifolate
antibiotics when compared to wild-type M. tuberculosis H37Rv, similar to the phenotype observed in ∆pknGMtb, and in trans expression of mutT3Mtb from plasmid
pVN771 (Table 2-2) restored wild-type levels of resistance (Fig 5-3C). Similar to
deletion of pknGMtb, supplementation of exogenous folate in the form of folinic acid bypassed the requirement for mutT3Mtb (Fig 5-3C). Similar results were obtained in
M. smegmatis assays (Fig. 5-4C), indicating that MutT3 might be involved in folate biosynthesis and be regulated by PknG.
Biofilm assays also showed similar phenotypes for ∆mutT3Mtb and ∆pknGMtb, and again genetic or chemical complementation could be achieved by in trans expression of mutT3PknG or exogenous addition of folinic acid, respectively (Fig. 5-
3D). M. smegmatis experiments yielded similar results (Fig. 5-4D).
5.C.3. Preliminary characterization of MutT3 as a Nudix Hydrolase
Nudix hydrolases are pyrophosphatases that catalyze the hydrolysis of a
nucleoside diphoshate linked to a variety of moieties. While most members of this
family are involved in clearance of potentially deleterious metabolites from the cell
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138
Figure 5-3. mutT3Mtb is required for survival of folate starvation and biofilm growth. (A) The mutT3Mtb deletion cassette and mutT3 loci in wild-type M. tuberculosis H37RV and its derived ΔmutT3Mtb strain. (B) PCR confirmation of
mutT3Mtb deletion using primers MMtb1 + MMtb2 (Table 2-1) shows shift corresponding to replacement of mutT3Mtb with the hygromycin resistance cassette.
Sequencing also confirmed the replacement of mutT3Mtb gene by the hygromycin cassette (not shown) (C) Biofilm growth of M. tuberculosis strains. Biofilm growth of wild type M. tuberculosis H37Rv, its derived ΔpknGMtb and ΔmutT3Mtb mutants, and
the complemented ΔmutT3Mtb/mutT3Mtb and ΔmutT3Mtb/ylgG strains was assayed in
the presence of either 0 or 1mM folinic acid. Growth was quantified at week 6 after
inoculation by measuring total protein per biofilm plate. (D) Sulfachloropyridazine
susceptibility of M. tuberculosis strains. Serial dilutions of growing cultures of wild-
type M. tuberculosis H37Rv, its derived ΔpknGMtb and ΔmutT3Mtb mutants, and the complemented ΔmutT3Mtb/mutT3Mtb and ΔmutT3Mtb/ylgG strains were replicated
onto solid 7H10-OADC medium containing sulfachloropyridazine (SL) and either 0
or 1mM folinic acid (FA) and incubated at 37°C for 3 weeks.
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140
Figure 5-4. mutT3Msm is required for survival of folate starvation and biofilm growth. (A) The mutT3Msm deletion cassette and mutT3Msm loci in wild-type M.
2 smegmatis mc 155 and its derived ΔmutT3Msm strain. (B) PCR confirmation of mutT3Msm deletion using primers MMtsm1 + P2 and P3 + MMsm2 (Table 2-1) amplify the 5’- and 3’-transitions from genomic DNA to the inserted hygromycin cassette, respectively. (C) Biofilm growth of M. smegmatis strains. Surface biofilm growth of M.
2 smegmatis mc 155, its derived ΔpknGMsm and ΔmutT3Msm mutants, and the complemented ΔmutT3Msm/mutT3Msm, ΔmutT3Msm/mutT3Mtb, ΔmutT3Msm/ntpA and
ΔmutT3Mtb/ylgG strains was assayed in the absence or presence of 1mM folic acid
(FA). Growth was recorded at week 1 after inoculation. (D) Sulfachloropyridazine susceptibility of M. smegmatis strains. Growing cultures of wild-type M. smegmatis
2 mc 155, its derived ΔpknGMsm and ΔmutT3Msm mutants, and the complemented
ΔmutT3Msm/mutT3Msm, ΔmutT3Msm/mutT3Mtb, and ΔmutT3Mtb/ylgG strains were plated onto solid NE medium containing either 0 or 1mM folic acid (FA), paper discs containing 1mg of sulfachloropyridazine (SL) applied to the center and plates incubated at 37°C for 4 days.
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and prevent build-up of metabolic intermediates, some Nudix hydrolases serve as enzymes in intermediate steps of biosynthetic pathways 146; 147; 148. In particular,
Nudix enzymes of E. coli (ntpA), Lactococcus lactis (ylgG) and Arabidopsis have only recently been shown to catalyze the removal of pyrophosphate from dihydroneopterin-triphosphate (DHN-3P) 147; 148, one of the earliest intermediates in folate biosynthesis, to create dihydroneopterin (DHN), which is eventually turned into folate 86; 147; 148 (Fig. 1-8). This reaction had been thought to occur by spontaneous chemical dephosphorylation for over 50 years 148; 154.
To determine whether loss of DHN-3P pyrophosphatase activity yielded similar sensitivity profiles, as had been observed in mycobacterial backgrounds, we created strain ∆ntpA in E. coli MG1665 by one-step phage λ-Red -based
recombineering method 176; 177; 184 and performed a sulfamethoxazole sensitivity
assay (Fig. 5-5). Deletion of ntpA led to increased susceptibility to the sulfadrug (Fig.
5-5), indicating a general requirement of DHN-3Pase for bacterial antifolate
resistance.
If mutT3 indeed encodes a DHN-3Pase activity, deletion of mutT3 should be complemented by in trans expression of other DHN-3Pases. ∆mutT3Mtb regained wild-type levels of resistance to sulfachloropyridazine when L. lactis ylgG (pVN776) was expressed in trans (Fig. 5-3D). Similar results were obtained in M. smegmatis mc2155 by in trans expression of ntpA (pVN773) or ylgG (pVN776) (Fig. 5-4D).
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Fig. 5-5. ntpA, a dihydroneopterin-triphosphate pyrophosphatase, is required for antifolate resistance in E. coli. Sulfamethoxazole sensitivity of E.coli strains.
Growing culture of wild-type E. coli MG1665, its derived ∆ntpA or the complemented
∆ntpA/ntpA strains (50µl, OD600=1.0) were plated onto solid LA medium and paper discs containing 1mg sulfamethoxazole were applied. Chemical complementation was achieved by exogenous supplementation of 1mM folinic acid (FA). Plates were incubated at 37ºC overnight.
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Expression of DHN-3Pases also restored surface biofilm growth to normal levels in
strain ∆mutT3Mtb/ylgG (Fig. 5-3C), further strengthening the hypothesis that MutT3 performs DHN-3Pase activity in mycobacteria, and that folate levels underlie the biofilm and antifolate resistance phenotypes.
Next we tested whether MutT3 can bind GTP by radioactive nucleotide
184; 185 binding assay . A titration of purified MutT3Msm at saturating amounts of tritium-labeled GTP showed binding occurring in a MutT3-dependent manner (Fig.
5-6A). When the amount of tritiated GTP was titrated for defined amounts of purified MutT3Msm, binding was GTP-dependent and leveled out at plateaus (Fig. 5-
6B), indicating that saturation of binding sites was occurring. Finally, when purified
3 MutT3Msm was incubated with saturating levels of [8- H]-GTP and increasing
amounts of unlabeled GTP, cold GTP could outcompete labeled GTP (Fig. 5-6C).
These results confirmed specific binding of GTP to MutT3 and strengthened the
hypothesis that mutT3 encodes a Nudix hydrolase.
5.C.4. RplM, a novel substrate of PknG, co-purifies with MutT3.
To investigate whether MutT3 was directly regulated by PknG phosphorylation, in vitro radioactive kinase assays were performed using purified
proteins. PknGMtb autophosphorylates, (Fig. 5-7A, lane 1), which could be inhibited
by addition of the specific inhibitor AX20017 (Fig. 5-AB, lane 2). While no substrate
phosphorylation was observed when using MutT3Msm-6xHis that had been recombinantly expressed in E. coli (Fig 5-AB, lane 9), a phosphorylated substrate
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145
Figure 5-6. MutT3 binds nucleotides in vitro. Radioactive nucleotide binding
assay using tritium-labeled GTP and purified MutT3Msm-6xHis. Radioactivity retained on membranes binding protein but not unbound nucleotides was measured in counts per minute (CPM) detected by liquid scintillation counting. Error bars represent standard deviations from triplicate experiments. (A) Saturating levels of tritium-labeled GTP (2.5µM) were incubated with 0 to 50µg purified MutT3. (B) 10 or 20µg MutT3 were incubated with 0 to 25µM tritium-labeled GTP. (C) 20µg MutT3 were incubated with 25µM labeled and 0 to 50µM unlabeled GTP.
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could be detected by autoradiograph when using MutT3Msm-6xHis purified from M. smegmatis (Fig. 5-7A, lane 7). This band however displayed a lower molecular weight compared to MutT3Msm-6xHis (Fig. 5-7A, lane 7 top panel vs. bottom panel).
When large amounts of the purified fraction were loaded, it could be detected by
Coomassie Blue stain (data not shown), suggesting that it was a peptide that co- purified with MutT3 from M. smegmatis lysate. Since no band was observed for
comparable fractions of M. smegmatis extract not expressing 6x-His-tagged
MutT3Msm, this co-purification seemed to occur in a specific, MutT3-dependent manner rather than through non-specific binding to the Nickel resin (Fig. 5-7A, lane
8). LC/MS/MS analysis of the MutT3Msm-6xHis fraction obtained from M. smegmatis identified 15 proteins associated with MutT3, 5 of which were ribosomal (Table 5-1).
The two proteins that were predicted to have a similar molecular weight as the detected PknG-phosphorylated band on the autoradiograph, RplM and SmpB, were selected for recombinant expression and purification in E. coli. While no phosphorylation was detectable for in vitro radioactive kinase assays using
SmpBMsm-6xHis (Fig. 5-7B, lane 4), RplMMsm-6xHis was phosphorylated by PknG (Fig.
5-7A, lane 6 and Fig. 5-7B, lane 2). Removal of the 6xHis-tag by thrombin cleavage
resulted in perfect alignment with the original band observed as co-purified with
MutT3Msm-6xHis (Fig. 5-7A, lane 4). Inhibition of PknG activity abolished RplM phosphorylation, indicating specific phosphorylation of RplMMsm by PknG (Fig. 5-7A,
lane 5). RplMMtb is likewise phosphorylated by PknGMtb (Fig. 5-9B, lane 3) and PknG
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Table 5-1. MutT3 co-purified proteins identified by LC/MS/MS.
Peptides N Name (95%) Cupin domain protein OS=Mycobacterium smegmatis (strain ATCC 1 700084 / mc(2)155) GN=MSMEG_5707 PE=4 SV=1 25 30S ribosomal protein S19 OS=Mycobacterium sp. (strain MCS) GN=rpsS 3 PE=3 SV=1 6 Pseudouridine synthase OS=Mycobacterium smegmatis (strain ATCC 4 700084 / mc(2)155) GN=rluB PE=3 SV=1 5 5 Putative uncharacterized protein OS=Plasmid pAL5000 PE=4 SV=1 4 Urease accessory protein UreE 1 OS=Mycobacterium smegmatis (strain 6 ATCC 700084 / mc(2)155) GN=MSMEG_1091 PE=4 SV=1 4 Pantothenate kinase OS=Mycobacterium smegmatis (strain ATCC 700084 7 / mc(2)155) GN=coaA PE=3 SV=1 3 Transcriptional regulator, TetR family protein OS=Mycobacterium smegmatis (strain ATCC 700084 / mc(2)155) GN=MSMEG_2153 PE=4 8 SV=1 4 50S ribosomal protein L27 OS=Mycobacterium smegmatis (strain ATCC 9 700084 / mc(2)155) GN=rpmA PE=1 SV=1 4 SsrA-binding protein OS=Mycobacterium smegmatis (strain ATCC 10 700084 / mc(2)155) GN=smpB PE=1 SV=1 3 11 Putative transposase OS=Mycobacterium smegmatis PE=4 SV=1 2 IS1549, transposase OS=Mycobacterium smegmatis (strain ATCC 700084 / 11 mc(2)155) GN=MSMEG_0074 PE=4 SV=1 2 Putative uncharacterized protein OS=Mycobacterium smegmatis (strain 12 ATCC 700084 / mc(2)155) GN=MSMEG_4029 PE=4 SV=1 2 50S ribosomal protein L16 OS=Mycobacterium smegmatis (strain ATCC 13 700084 / mc(2)155) GN=rplP PE=1 SV=1 1 50S ribosomal protein L13 OS=Mycobacterium smegmatis (strain ATCC 14 700084 / mc(2)155) GN=rplM PE=1 SV=1 1 50S ribosomal protein L21 OS=Mycobacterium smegmatis (strain ATCC 15 700084 / mc(2)155) GN=rplU PE=1 SV=1 1
Bolded proteins were of comparable size to observed band and were therefore recombinantly expressed in
E. coli and tested in hot in vitro kinase assays as substrates of PknG.
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149
Figure 5-7. RplM, a novel substrate of PknG, co-purifies with MutT3. (A) In vitro
kinase assay using radioactive [γ-32P]-ATP to assess phosphorylation by PknG. 0.5µg
purified PknGMtb-6xHis (with or without 15 minute pre-incubation with 5mM
AX20017) were incubated with 1.5µg of purified substrate candidate. Lane 1:
PknGMtb-6xHis, Lane 2: PknGMtb-6xHis + AX20017, Lane 3: RplMMsm-6xHis (purified
from E. coli (pVN844), Lane 4: PknGMtb-6xHis + RplMMsm (purified from E. coli
(pVN844)), Lane 5: PknGMtb-6xHis + RplMMsm (purified from E. coli (pVN844)) +
AX20017, Lane 6: PknGMtb-6xHis + RplMMsm-6xHis (purified from E. coli (pVN844)),
Lane 7: PknGMtb-6xHis + MutT3Msm-6xHis (purified from M. smegmatis (pVN823)),
Lane 8: PknGMtb-6xHis + control purification from M. smegmatis, Lane 9: PknGMtb-
6xHis + MutT3Msm-6xHis (purified from E. coli (pVN835)). (B) In vitro kinase assay
using radioactive [γ-32P]-ATP to assess phosphorylation by PknG. 0 or 0.5µg purified
PknGMtb-6xHis were incubated with 1.5µg of purified RplMMsm-6xHis or SmpBMsm-
6xHis. (C) Kinase assay using radioactive [γ-32P]-ATP and whole cell extract to assess specificity of RplM phosphorylation by PknG. Coomassie Blue stain (left) and autoradiograph (right) of hot kinase assay using 1mg whole cell extract of mc2155,
∆pknGMsm, ∆pknGMsm/pknGMtb or ∆mutT3Msm and 1.5µg RplM-6xHis followed by
6xHis-pulldown to enrich for 6xHis-MutT3.
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Figure 5-8. MutT3 co-purified with RplM and is not regulated translationally.
(A) Interaction model of RplMMsm and MutT3Msm-6xHis in vivo. RplM either directly interacts with MutT3 or their association is mediated by a third-party partner. (B)
Western blot assaying mutT3 expression in M. smegmatis strains. Total protein
2 extract of strains mc 155/mutT3Msm-6xHis and ∆pknGMsm/mutT3Msm-6xHis were
separated into soluble and insoluble fractions and probed with penta-His antibody
(Qiagen). No difference in expression levels was observed in the absence of PknG.
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inhibition also abolished phosphorylation (Fig. 5-8B, lane 4). RplM, the large
ribosomal subunit protein L13, is therefore phosphorylated by PknG in both
pathogenic and non-pathogenic mycobacterial species in vitro. These data show that
MutT3 associates with RplM in a complex involving other proteins, and that RplM is
a novel substrate of PknG in mycobacteria.
To investigate if MutT3 levels in the cell are regulated at a translational level,
2 Western blot assays were performed using strains mc 155/mutT3Msm-6xHis and
∆pknGMsm/mutT3Msm-6xHis in which 6xHis-tagged MutT3Msm is produced under the control of the native mutT3 promotor. No difference in protein levels was observed upon deletion of pknG, thereby excluding translational control exerted by PknG (Fig.
5-8B).
In order to test whether phosphorylation of RplM is PknG-specific, total cell
2 extract was prepared from M. smegmatis mc 155, the ∆pknGMsm, ∆mutT3Msm and
∆pknGMsm/pknGMtb strains and used in hot kinase assays with purified RplMMsm-
6xHis followed by 6xHis pull-down. A faint band at the correct molecular weight for
RplMMsm-6xHis was observed on autoradiographs with all extracts except the
∆pknGMsm mutant (Fig. 5-7C), indicating that native PknG at natural expression
levels can phosphorylate RplM. 6xHis-tagged RplM expressed in M. smegmatis from
the heat-shock promotor could not be detected, most likely due to too low
expression levels (data not shown). This result indicates that phosphorylation of
RplM is PknG-specific.
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5.C.5. Phosphorylation of RplM by PknG on threonine 11 mediates regulation of folate metabolism.
Of five potentially phosphorylated serine or threonine sites identified by
LC/MS/MS in the sequence of RplMMsm, three were conserved across the
mycobacterial genus (Fig. 5-9A, asterisks). A triple point mutant of RplMMtb in
which the three conserved residues threonine 11, threonine 12 and serine 14 were
167; 179; 180 mutated to alanines, which cannot accept phosphate groups . This RplMMsm
T11A,T12A,S14A could not be phosphorylated by PknGMtb, indicating that phosphorylation occurs on one or more of these residues (Fig. 5-9B, lane 5). Single
point mutants were therefore cloned and tested in hot kinase assays, showing
complete loss of phosphorylated signal upon mutation of threonine 11 to alanine
(Fig. 5-9B, lane 6), and partial loss upon mutation of threonine 12 (Fig. 5-9B, lane 7), likely due to steric interactions interfering with PknG access to the immediately adjacent threonine 11. Mutation of serine 14 did not affect phosphorylation levels
(Fig. 5-9B, lane 8). These data indicate that phosphorylation of RplM by PknG occurs specifically on the conserved threonine 11 residue.
To test whether phosphorylation of RplM mediates PknG regulation of folate metabolism and biofilm growth, the gene encoding RplM was replaced with alleles in which threonine 11 was changed to either alanine (T11A), which could not accept phosphate groups 167; 179; 180, or glutamate (T11E), mimicking constitutive phosphorylation 167; 179; 180. This was achieved by recombineering method in M.
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Figure 5-9. RplM is phosphorylated on threonine 11 by PknG in vitro. (A)
Sequence alignment of RplM homologs. (B) In vitro kinase assay using radioactive
32 [γ- P]-ATP to assess phosphorylation by PknG. 0.5µg purified PknGMtb-6xHis (with
or without 15 minute pre-incubation with 5mM AX20017) were incubated with
1.5µg of purified RplMMtb and its derived point mutants.
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smegmatis mc2155, generating strains mc2155/rplMT11A, mc2155/rplMT11E (Fig. 5-
10A, B). Replacement of rplM with the point mutant alleles was confirmed by sequencing and PCR (Fig. 5-10B).
2 M. smegmatis mc 155, its derived ∆pknGMsm mutant and strains
mc2155/rplMT11A and mc2155/rplMT11E were assayed biofilm growth assays (Fig. 5-
10C). Replacement with the non-phosphorylateable point mutant allele (strain
mc2155/rplMT11A) led to decreased surface biofilm formation, which mimicked the
defect observed in ∆pknGMsm. The requirement for native rplM could be bypassed by providing folinic acid in the medium (Fig. 5-10C). However, mc2155/rplMT11E also showed slighlty inhibited growth (Fig. 5-10C), indicating that while phosphorylation of RplM on threonine 11 is absolutely required for late stage growth, increased phosphorylation levels may be harmful for biofilm growth. These results strongly indicate that PknG mediates its regulation of folate control by balancing phosphorylation of RplM on threonine 11.
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156
Figure 5-10. Phosphorylation of RplMMsm on threonine 11 is required for
survival of folate starvation and biofilm growth. (A) The rplMMsm replacement
2 cassette and rplMMsm locus in wild-type M. smegmatis mc 155 and its derived
2 T11A 2 T11E mc 155/rplM and mc 155/rplM strains. (B) PCR confirmation of rplMMsm
replacement using primers RMsm1 + RMsm2 (Table 3-1) shows shift corresponding to
insertion of the hygromycin resistance cassette. (C) M. smegmatis biofilm growth.
2 Biofilm growth of wild type M. smegmatis mc 155, its derived ΔpknGMsm, mc2155/rplMT11A and mc2155/rplMT11E strains were assayed in the presence or absence of 1mM folinic acid (FA). Biofilm was pictured at 1 week (top). Surface biofilm growth was quantified at 1 week by measuring total protein per biofilm plate (bottom).
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5.D. Discussion
We have previously demonstrated that the eukaryotic-like serine/threonine
kinase G (PknG) of pathogenic and non-pathogenic mycobacterial species mediates antibiotic tolerance in both non-pathogenic and pathogenic mycobacteria 171. In this study, we found that PknG is involved in regulation of de novo folate biosynthesis, especially in late stage growth similar to the persistence of pathogenic mycobacteria inside the host macrophage. Furthermore, we have shown that PknG-dependent failure of host macrophages to deliver lysosomes to phagosomes 20 is mediated by
PknG through regulation of mycobacterial folate metabolism. These findings
question the proposed function of PknG as a virulence factor that modifies host
signaling networks in phagolysosome synthesis 20. We have identified a novel,
mycobacterial substrate of PknG, RplM, an essential protein that is a component of
the large ribosomal subunit 157. We showed that PknG phosphorylation of RplM
occurs on the mycobacterially conserved residue threonine 11 is important for a
continuation of the signaling cascade leading to MutT3. We provide preliminary
evidence that MutT3 functions as a nucleoside diphosphate hydrolase (Nudix)
enzyme. We hypothesize that MutT3 functions as a dihydroneopterin-triphosphate
pyrophosphatase (DHN-3Pase) that catalyzes the first committed step of folate de
novo biosynthesis. Our data suggested that this signaling axis is required for
mycobacterial resistance to antifolates and biofilm formation.
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Based on our data, we propose a model in which PknG, upon receiving
signals for a folate requirement such as seen in growth in macrophages or biofilms, phosphorylates RplM (Fig. 5-11), which forms a complex with MutT3. The phosphorylation event affects MutT3 activity, possibly by either promoting or preventing efficient assembly of the MutT3 and RplM involving complex (Fig. 5-11).
The nature of this complex remains unclear, but based on LC/MS/MS analysis of proteins co-purified with MutT3, this complex involves several ribosomal proteins
that may localize on the ribosome itself or on a sub-ribosomal complex (Model A, Fig.
5-11), or on an independent complex (Model B, Fig. 5-11) 160. MutT3 likely encodes a
Nudix pyrophosphohydrolase activity 143; 144 directly or indirectly affecting folate
metabolism that results in changes in folate availability. Since the absence of MutT3
can be complemented by in trans expression of YlgG (Fig. 5-3 and Fig. 5-4),
previously shown to catalyze dihydroneopterin-triphosphatase activity, and since deletion of DHN-3Pase (NtpA) in E. coli mimics phenotypes observed in mycobacteria, MutT3 likely acts as a DHN-3Pase in mycobacterial de novo folate biosynthesis 86; 147; 148.
Our previous results suggested that regulation of folate metabolism by PknG
is involved in lysosomal localization in infected macrophages (Chapter 4).
Knowledge of substrates and precise molecular mechanisms underlying this
observation may provide a basis for identification of inhibitors to this pathway. Such
inhibitors, along with inhibitors of PknG, may also be used to potentiate classical
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antifolates as well as to promote the innate mycobactericidal of host macrophages 45.
This signaling axis thus represents a novel target for development of antifolate potentiators that, when used together with antifolates, could also enhance the innate immunity of host macrophages against M. tuberculosis.
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Figure 5-11. Model of folate biosynthesis regulation via PknG, RplM and MutT3.
(Model A) PknG phosphorylates RplM, localized on the large subunit of the ribosome, on threonine 11. Phosphorylated RplM is released from the ribosome and associates 161
with a complex that includes MutT3. Interaction of RplM and this complex in turn
leads to alterations in de novo folate biosynthesis via MutT3 Nudix hydrolase
activity. (Model B) PknG phosphorylates RplM, localized on the large subunit of the ribosome, on threonine 11. Phosphorylated RplM signals to MutT3, also localized on
the large ribosomal subunit, which in turn leads to alterations in de novo folate
biosynthesis via MutT3 Nudix hydrolase activity.
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5.E. Acknowledgements.
We thank Jean Pieters, Julia van Kessel, Graham F. Hatfull, Magdalena
Taracila, Robert Bonomo, Bing Liu, Piet deBoer, Anne Walburger, Cathy Carlin, and
Hoa Nguyen for providing materials and expertise. Support was provided by an NIH
R01 grant (AI087903), start-up funds from the School of Medicine, a STERIS
Infectious Diseases Research Grant, and a CFAR Developmental Award from Case
Center For AIDS Research.
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Chapter 6.
Discussion and Future Directions
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6.A. Future Experiments
6.A.1. Hierarchy of PknG, RplM and MutT3
We have demonstrated that PknG, RplM, and MutT3 all are required for
survival of folate starvation and for biofilm formation, and thus likely act together in a folate metabolic pathway. While preliminary inferences can be made from our data supporting this hierarchy, more evidence will be needed to determine the
precise relationship among them. We proposed in our model (Fig. 5-11) that PknG
acts first, phosphorylating RplM, which then modulates MutT3 activity in some way.
In favor of our model, we demonstrated that the kinase activity of PknG itself
(Fig. 4-2B, Fig. 4-3B and Fig. 4-4B) and its phosphotransfer to threonine 11 of RplM
(Fig. 5-10 C, D) are specifically required for normal folate metabolism to occur in the
cell. However, a feedback loop in which phosphorylation of RplM as part of the
ribosome 157, or even as an anti-terminator 161, may regulate PknG levels and therefore PknG activity on RplM or other substrates, by way of translational or transcriptional control, respectively, cannot be excluded.
MutT3 likely acts downstream of PknG phosphorylating RplM, as it is not modified by PknG directly (Fig. 5-7A) but associates with RplM (Fig. 5-8A, Table 5-1).
MutT3 is not required for phosphorylation of RplM by PknG to occur in cellular extracts (Fig. 5-8B), further strengthening the hierarchy proposed in our model.
Likewise, MutT3 is not regulated transcriptionally or translationally by PknG (Fig. 5-
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8B), indicating that MutT3 activity is regulated by formation of a complex with RplM
rather that ribosomal or anti-termination activity of RplM.
To definitively determine the order of events and interactions of these three
proteins in the regulation of folate biosynthesis, we propose experiments in which
2 T11A 2 T11E strains ∆mutT3Msm, mc 155/rplM and mc 155/rplM will be tested in
antifolate susceptibility (see section 2.F.) and biofilm assays (see section 2.I.) alone or in combination with the specific PknG kinase inhibitor AX20017 20. If PknG kinase
action is the first event in the pathway as we propose, and if no feedback loop exists,
inhibition of PknG should not aggravate or otherwise alter the levels of
susceptibility to folate starvation or impairment of biofilm in comparison to
untreated strains. If however either MutT3 or RplM act before PknG and mediate
feedback control over PknG, PknG inhibition should lead to more severe phenotypes
2 T11A in ∆mutT3Msm or mc 155/rplM that mimics constitutively non-phosphorylated
RplM 181; 198; 199. On the other hand, strain mc2155/rplMT11E, which mimics constitutively phosphorylated RplM 181; 198; 199, should not exhibit any difference in phenotype regardless of treatment.
Additionally, we will assay total protein extracts of H37Rv and ∆mutT3Mtb for
expression levels of PknG to address either a feedback loop or a constitutive
mechanism in which MutT3 Nudix hydrolase activity impacts transcriptional or
translational regulation of pknG.
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6.A.2. RplM point mutant alleles in M. tuberculosis and effect on lysosomal trafficking
We have demonstrated that phosphorylation of RplM on threonine 11 by
PknG is required for survival of folate starvation and biofilm growth in M. smegmatis
(Fig 5-10 C, D). To confirm that this is also true for pathogenic mycobacteria, we are
T11A T11E currently working on generating strains H37Rv/rplM and ∆pknGMtb/rplM in
M. tuberculosis by specialized transduction 174 (see section 2.C.2.). Once these strains are obtained, they will be subjected to antifolate susceptibility (see section 2.E.) and biofilm assays (see section 2.H) to assess their phenotypes and whether they correspond to what we observed for M. smegmatis.
We propose that the phosphorylation event mediates folate metabolic control in latent growth conditions and that this in turn regulates the ability of M. tuberculosis to establish its intracellular localization 20 (see section 2.K. and Fig. 4-5).
T11A T11E Strains BCG/GFP/rplM and ∆pknGBCG/GFP/rplM constructed by specialized transduction (see section 2.C.2.) will be used in a lysosomal trafficking assay using murine bone-marrow derived macrophages (see section 2.K.) that would allow us to determine whether phosphorylation states of RplM are required for phagosomal localization of pathogenic mycobacteria independently of PknG. Strain
BCG/GFP/rplMT11A should exhibit increased lysosomal delivery in the presence of
T11E PknG, whereas ∆pknGBCG/GFP/rplM should remain localized in the phagosome in the absence of PknG. Furthermore, folate supplementation should rectify the
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T11A phagosomal localization defect of BCG/GFP/rplM , as seen with ∆pknGBCG/GFP,
T11E whereas sulfonamide treatment should send ∆pknGBCG/GFP/rplM to lysosomes.
6.A.3. Subcellular localization of RplM/MutT3-containing complexes
Since loss of PknG does not lead to alterations in MutT3 expression levels
(Fig. 5-7 D), phosphorylation of RplM does not likely affect MutT3 expression levels.
We failed to observe direct interaction between purified MutT3Msm and RplMMsm in vitro (data not shown). Taken together with the identification of a number of other co-purified proteins in MutT3 purification fractions by LC/MS/MS, several of which, like RplM, are ribosomal, it is likely that MutT3 localizes on a larger complex on which it comes into (in)direct contact with RplM, the phosphorylation status of which regulates MutT3 activity. The nature of this complex remains elusive and will have to be further investigated. The complex on which MutT3 performs its Nudix hydrolase activity could be ribosomal or sub-ribosomal in nature. We proposed two possible models (Fig. 5-11) that involve different complexes.
Model A proposes that the phosphorylation of threonine 11 on RplM leads to a specific release of phospho-RplM from the ribosome, followed by association with a complex involving MutT3 that regulates de novo folate biosynthesis (Fig. 5-11).
This model is based on the interferon-γ-activated inhibitor of translation (GAIT) complex found in eukaryotes, in which L13a, the human homolog of RplM, is phosphorylated and thereby released from the ribosome, making possible association with the GAIT system 200; 201; 202. The GAIT system proposes that L13a is
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not required for translation by the ribosome, and that the ribosome serves as a
depot for L13a until it is phosphorylated on a specific serine residue by death-
associated kinase-1 201; 202. Free phospho-L13a can then bind specifically to the
other components of the GAIT complex and repress translation of specific genes 200;
202. Similarly, RplM may be “stored” on the ribosome in mycobacteria until PknG
phosphorylates its threonine 11 in response to conditions requiring folate synthesis.
L13 is exclusively surface-associated without burrowing into the large ribosomal subunit in archaea 162; 203, and RplM of E. coli does not localize closely to the areas of the ribosome that bind message or are catalytically active (Fig. 1-9) 162; 163. It is likely
that RplM does not structurally support the ribosome, and can therefore be released
upon phosphorylation to perform extra-ribosomal functions, such as being an
essential component of the GAIT complex in eukaryots 200; 201; 202, acting as part of an anti-termination complex in bacteria 161, or as part of our proposed folate- synthesizing complex involving MutT3 (Fig 5-11).
Model B on the other hand proposes that both MutT3 and RplM localize on the large ribosomal subunit either as part of the ribosome or a sub-ribosomal complex (Fig 5-11). Some GTPases have been shown to co-fractionate with the ribosome where they regulate growth and stress responses 164; 165 and even show increased hydrolytic activity in the presence of ribosomal particles 204. This suggests that MutT3 may require ribosomal contact for performance of its activity, and this contact may only be possible upon phosphorylation of RplM, an event that may alter
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the ease with which MutT3 binds to the complex. Alternatively, MutT3 activity could
be activated by release from the complex, in which setting the phosphorylation
event of RplM may facilitate this release by altering affinities of neighboring
proteins or binding pockets that directly interact with MutT3.
To confirm whether MutT3 indeed localizes on sub-ribosomal particles, and
whether the ability to localize is determined by PknG phosphorylation, we propose
2 to perform subcellular fractionations of strains mc 155/pVN866, ∆pknGMsm/pVN866,
mc2155/rplMT11A/pVN866 and mc2155/rplMT11E/pVN866 that express 6x-His-
tagged MutT3Msm from the native MutT3 promotor. Ribosomal fractions will be prepared by standard protocols for mycobacterial samples using ultracentrifugation at 104,500 x g 205; 206 and assayed by Western blot for MutT3-6xHis (see section 2.E),
PknG (see section 2.E), and soluble as well as ribosomal controls. This assay will
allow us to determine whether MutT3 can associate with the ribosome or sub-
ribosomal complexes at all and whether association is promoted, decreased or
unaffected by phosphorylation of threonine 11 on RplM by PknG.
To test the hypothesis that RplM is released from the large ribosomal subunit,
2 2 subcellular fractionations of mc 155, mc 155 treated with AX20017, ∆pknGMsm, mc2155/rplMT11A and mc2155/rplMT11E will be prepared and ribosomal as well as
soluble fractions probed for presence of RplM by Western blot. This will allow us to
determine whether RplM can be released from the ribosome (wild-type), whether
PknG activity is required for the release (AX20017 treatment, ∆pknGMsm), and
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whether phosphorylation of threonine 11 is the essential release trigger
(mc2155/rplMT11A, mc2155/rplMT11E).
Should we find that RplM indeed is released from the ribosome to associate with MutT3 on a folate-synthesizing complex, we will seek to identify this complex by cross-linking experiments followed by pull-down of 6xHis-tagged MutT3 and submitting the pulled down materials to LC/MS/MS analysis.
6.A.4. Biochemical characterization of MutT3 enzymatic function
MutT3 contains the characteristic Nudix motif 156;143; 144, but does not appear
to function as a mutator protein or an 8-oxo-guanosine triphosphatase 156. The fact
that mutT3 is required for antifolate resistance and biofilm growth and that this
requirement could be bypassed by exogenous folate supplementation or expression of DHN-3Pases from other bacteria (Fig. 5-3 and Fig. 5-4) suggest that MutT3
performs the pyrophosphatase reaction converting dihydroneopterin-triphosphate
to dihydroneopterin-phosphate.
We have established a collaborative effort with the Amzel laboratory (Johns
Hopkins University, Baltimore, MD) that initially identified and characterized E. coli
NtpA 147. This collaborator will help us to characterize the DHN-3Pase activity of
MutT3 using their established methodology 147; 148; 207; 208; 135; 136. It is expected that several substrates will show moderate activity in this assay, as Nudix hydrolases tend to be promiscuous 143; 144, but a preference may be observed, as was the case
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for E. coli NtpA 147 and L. lactis YlgG 147. This study will further substantiate the role
of MutT3 in folate metabolism (Fig. 5-11).
We previously tested GTP and ATP hydrolysis by MutT3, followed by
LC/MS/MS analysis to detect breakdown products and did not observe significant
activity of MutT3 with these substrates (data not shown). It is possible that contact
with the ribosome may stimulate GTPase or DHNTP pyrophosphatase activity, as
has been observed for the E. coli GTPase YjeQ 204. Phosphorylation of RplM may lead to improved binding or other changes facilitating MutT3 function, and loss of MutT3 or PknG would therefore lead to similar phenotypes. This may be a regulation mechanism specific to mycobacteria, since PknG is not widely conserved throughout more distantly related bacteria such as E. coli 140.
If the previously proposed in vitro characterization fails to detect activity, we will repeat hydrolysis assays adding purified ribosomes to stimulate pyrophosphatase activity and monitor hydrolysis by LC/MS/MS with the help of Dr.
Guofang Zhang in the Nutrition Department (see also section 6.A.6). If ribosomal addition indeed leads to activity increases, we will try to modify the above- described colorimetric assay accordingly.
Furthermore, the Amzel laboratory will make efforts to obtain crystal structures of MutT3Msm and MutT3Mtb, which will allow for comparison and
superimposition of active site conformations and substrate binding pockets to E. coli
NtpA147 and other crystallized Nudix hydrolases to help elucidate possible
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substrates and mechanisms. Crystal structures could also help attempts to intelligently design inhibitors aimed at potentiation of classical antifolate antibiotics for treatment of MDR and XDR TB (see section 1.C.2.)
6.A.5. MutT3 deletion effects on lysosomal trafficking
We have demonstrated that mutT3Msm and mutT3Mtb are required for survival of folate starvation induced by classical antifolate antibiotics and biofilm growth (Fig 5-3 and Fig. 5-4). We further postulate that MutT3 is a Nudix hydrolase likely involved in folate de novo biosynthesis by acting as a DHN-3Pase (see Fig. 5-11 and summary in section 6.A.4), or otherwise mediates folate level control through
Nudix function with different substrates, e.g. by controlling growth via GTP or ppGpp level modulation (see section 1.F.), as part of an epigenetic network also involving PknG and RplM. We propose that MutT3 action mediates folate metabolic control in latent growth conditions and that this in turn regulates the lysosomal delivery to phagosomes containing M. tuberculosis in infected macrophages 20 (see section 2.L. and Fig. 4-5). We propose to generate strain ∆mutT3BCG/GFP by
specialized transduction (see section 2.C.2.) for use in a lysosomal trafficking assay
using murine bone-marrow derived macrophages (see section 2.L.) that would
allow us to determine whether mutT3BCG is involved in lysosomal trafficking and whether this is due to folate requirements of the bacterium in the macrophage. We predict that ∆mutT3BCG/GFP will exhibit increased lysosomal delivery comparable to
∆pknGBCG/GFP and that wild type characteristics will be restored when exogenous
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folate is provided. This would also allow us to determine whether inhibition of
MutT3 may be a valuable concept to consider for potentiation of classical antifolate antibiotics (see. Section 1.C.2.).
6.A.6. Intracellular folate levels
We have observed that supplementation of exogenous folate bypasses requirements for PknG, RplM phosphorylation and MutT3 for survival of folate starvation (Fig. 4-1, Fig. 4-2, Fig. 5-3, Fig. 5-4 and Fig. 5-10) and for biofilm formation (Fig. 4-3, Fig. 4-4, Fig. 5-3, Fig. 5-4 and Fig. 5-10), and the requirement for
PknG to prevent lysosomal delivery to phagosomes in infected macrophages (Fig. 4-
5). We have also shown that folate starvation leads to a similar increase in lysosomal delivery as observed for pknGBCG deletion (Fig. 4-5). We have therefore
proposed that PknG, RplM and MutT3 act in a pathway regulating intracellular folate
levels in static growth conditions in mycobacteria, likely by regulation of MutT3
DHN-3Pase activity in folate de novo biosynthesis. Loss of any of the three proposed
components of this regulatory pathway would therefore lead to deregulation and
reduction of folate levels in static growth conditions such as biofilm, and we propose
to confirm altered intracellular folate levels in strains ∆pknGMsm, ∆mutT3Msm, and
mc2155/rplMT11A.
Total folate levels will be quantified by a microbiological assay employing a
strain of L. casei that cannot synthesize folate de novo and that is therefore
dependent on exogenous supplementation of folate 209; 210.
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To analyze individual folate species levels inside the mycobacterial cell upon
loss of pknG or mutT3 or the ability to phosphorylate threonine 11 on RplM, the
above-listed strains will be grown in biofilm assays (see section 2.I.) and used for
extraction of intracellular folate species and analyis by LC/MS/MS.
6.B. Discussion
6.B.1. Significance of folate during mycobacterial colonization of host macrophages
Folates are essential co-enzymes in one-carbon unit transfer that, among other things, are involved in the biosynthesis of purines, thymidines, glycine, panthotenate and methionine, as well as bacterial formyl-methionyl-tRNA critical for initiation of transcription 77; 78; 79; 45 (see section 1.C.). Consequently, folate
molecules are essential for the synthesis of macromolecules such as nucleic acids
and proteins, and both deregulation of folate levels and folate starvation affect
almost all cell division 45. Yet another effect of folate starvation manifests itself in interference with recycling of both homocysteine (Hcy, Fig. 1-4) and S-adenosine methionine (SAM) 45. Folates are usually considered to be of particular importance during periods of rapid cell division and growth 77; 78 and a requirement for folate
biosynthesis during late-stage rather than exponential growth phases may appear
counter-intuitive at first. However, while intracellular mycobacteria in latent TB
infections are thought to be non-replicating and exhibit only very slow growth 193;
194; 195, they are metabolically active 196 and should therefore require folates for the
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essential one-carbon transfer reactions 45; 77; 78; 79. In fact, our results have indicated that modulation of mycobacterial folate biosynthesis and availability by PknG are of particular importance in such conditions of late stage or stationary growth. Support for this hypothesis is provided by observations that pknG is among the genes sharply upregulated during immune suppression and reactivation in mice infected with M. tuberculosis, and that pknG is likely required for stationary phase metabolism 196; 197. The requirement for folate biosynthesis and PknG regulation
thereof in the context of the macrophage may be rooted in (a) mycobacterial
dormancy and persister formation requiring specific folate forms, (b) the stringent
response, triggered by folate starvation, to facilitate survival in the face of anti-
bacterial immune mechanisms, (c) immune evasion mechanisms requiring
metabolic adjustments, or in (d) cell wall and cell surface adjustments involved in
adaptation to the phagosomal environment.
6.B.1.a. Folate and cell dormancy
Other than the requirement for folate forms in basic cell cycle mechanisms
such as DNA synthesis, DNA replication and protein synthesis that occur in any
metabolically active cell, specific folate forms appear to be important for the
establishment of dormant, non-replicating cellular states 211; 212; 213. In particular, 5-
CHO-H4PteGlu (see Fig. 1-4), the most stable naturally occurring form of reduced
folate, has been suggested to function as a folate storage form that is necessary for
induction and survival of dormancy in Neurospora and soy beans 211; 212; 213.
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Interestingly, lack of MTHFS in mycobacterial cells leads to elevation of
102 polyglutamylated forms of 5-CHO-H4PteGlu , and its homolog in E. coli, YgfA, is required for the formation of persisters that are non-replicating and display increased multiple antibiotic resistance 214, similar to what is thought to occur in TB
latency 15; 195. A role for MTHFS in intrinsic antifolate resistance was also confirmed in M. tuberculosis and Escherichia coli 102; 103, suggesting that this determinant functions ubiquitously among bacteria. Folate regulation by PknG may therefore
influence levels of particular folate forms that are required for M. tuberculosis to transition into a dormant, metabolically active, but not actively dividing, state that facilitates survival inside the host macrophage in chronic TB infections.
The current Wayne model of M. tuberculosis dormancy postulates that the cell progresses through two non-replicating stages, the first of which involves activation of glycine dehydrogenase 15; 194; 195. Since glycine biosynthesis requires folate co-enzymatic action, folate deficiency leads to glycine deficiency 77; 78; 79; and
the reaction catalysed by glycine dehydrogenase cannot occur, thus making the
transition from active growth to persistence ineffective or potentially even
impossible, and accounting for another reason that folate biosynthesis regulation by
PknG is necessary for successful colonization of macrophages by M. tuberculosis.
Another mechanism that has been shown to be important in MTB latency and
is triggered by antifolates 153 or otherwise caused folate deficiency is the stringent response, a stress response to amino acid starvation 108; 109. In this stress response,
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the alarmone ppGpp is synthesized by RelA/SpoT to trigger growth arrest and
ensure cell survival until sufficient amino acid levels are encountered or
reestablished 150; 151. In mycobacteria, RelA synthesis of ppGpp is activated by the presence of uncharged tRNAs 215, which are formed during amino acid starvation, as in the case of folate starvation leading to non-functional glycine and methionine synthesis 77; 78; 79. It has been shown that relA is 5-fold upregulated in nutritionally
starved M. tuberculosis cultures that mimic dormancy inside the macrophage 216.
Interestingly, the Ndx8 Nudix hydrolase of Thermus thermophilus has been shown to
be involved in regulation of guanosine 3',5'-bispyrophosphate (ppGpp) levels independently of the RelA/SpoT synthesis and degradation cycles, degrading ppGpp to 3’,5’-bisphosphate (pGp), and potentially acting in a relay system with SpoT 149.
Ndx8 may thereby provide an additional mechanism to control the transition from growth arrest to active growth after amino acid starvation conditions have been removed 149. While we have proposed a DHN-3Pase function for MutT3, we cannot exclude alternative substrates for MutT3, and rather than directly controlling folate metabolism, a response to altered folate biosynthesis and folate starvation may be controlled through the PknG-RplM-MutT3 cascade by MutT3 modulation of ppGpp levels in the mycobacterial cell, also providing important control over persistence in host macrophages.
6.B.1.b. Immune evasion and cell wall maintenance
While PknG is expressed in non-pathogenic mycobacterial species and in the
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related corynebacteria, expression levels are dramatically increased in pathogenic mycobacteria. There is evidence that PknG expression is translationally regulated
140, suggesting the existence of dedicated regulatory mechanisms to control
expression levels of PknG in mycobacterial species. It has been shown that PknG is
involved in the tricarboxylic acid cycle as well as glutamine utilization in
Corynebacterium glutamicum, which is used to produce and isolate large amounts of
glutamate industrially, and in vitro studies have shown that mycobacterial PknG can
also phosphorylate GarA, the glutamate regulator found to be its target in C.
glutamicum 121; 122; 124. However, glutamate levels in mycobacteria do not appear to be altered to significant amounts upon loss of pknG 141, and GarA does not play a role
in the folate mediated late stage growth functions we have observed (see Chapter 4).
It had therefore been suggested that elevated expression levels may have provided
the means for additional virulence functions of PknG in pathogenic mycobacteria 12;
20. This possibility was also proposed in a recent study that addressed differential
expression of PknG in various mycobacterial species in relation to its virulence
function 140. In light of our results, we also favor the hypothesis that GarA and
glutamate regulation do not represent the primary target of PknG in mycobacteria,
but we think it is probable that responses to folate requirements are a more likely
function for mycobacterial PknG. Glutamate may however have some impact, as it is
required for folate de novo biosynthesis (see Fig. 5-2).
Interestingly, it has been demonstrated that many bacteria that have adapted
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to survival inside host macrophages or other immune cells induce stress responses
upon host cell infection, thereby inducing degradation and repair or replacement of
bacterial proteins damaged by antimicrobial immune mechanisms 217; 218. Since
folate starvation induces the ppGpp-mediated stringent response 153, and RplM has
been shown to associate with Obg in B. subtilis 165 and M. tuberculosis 166 to mediate
the σB-mediated stress response in Bacillus subtilis 164; 165, the PknG-RplM-MutT3 cascade that we propose to regulate folate metabolism could contribute to intracellular survival and immune evasion by means of stress response mediation.
As folate availability is essential for synthesis of proteins and other molecules that impact cellular building blocks, it is not surprising that regulation of folate metabolism and biosynthesis by PknG is involved with multiple antibiotic resistance and cell wall integrity of both non-pathogenic and pathogenic bacteria 157.
Loss of PknG function results in altered cell surface properties, likely due to changes in cell wall composition 157. Stationary phase growth, TB latency and biofilm growth have all been shown to exhibit higher levels of antibiotic resistance to a variety of drugs 22; 13; 14; 15; 16, which is likely due to decreased permeability of these
compounds through a thick, intact, potentially specifically adapted cell envelope 168;
189. Glutamine and its derived molecules, such as polyglutamines, may serve as
components that are required for the integrity of the mycobacterial cell wall as well,
thereby helping to pose an effective barrier to diffusion of antibiotics of diverse size
and polarity. PknG-mediated inhibition of glutamate reactions thus may also
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contribute in ways not directly related to folate. It is possible that folate molecules
are simply necessary to maintain a normal cell wall, and therefore contribute to the
high intrinsic resistance to antibiotics found in mycobacteria, and likewise to lower
penetration rates of oxidative stress molecules generated in macrophages aiming to
degrade phagocytosed bacteria 217; 219. However, it has been shown that several
mycobacterial cell wall components actively modulate and suppress immune-
mediated mechanisms aimed at degradation of M. tuberculosis 217, and it is possible
that folates are required for synthesis of building blocks necessary to maintain the
needed amounts of these components as well as their delivery and proper
conformation on the cell surface.
More specifically, it has been demonstrated that one mechanism of immune
evasion in TB infection consists of early phagosomes containing M. tuberculosis
failing to recruit and activate hVPS34 220; 221, an effector kinase required for synthesis and accumulation of phosphatidylinositol-3-phosphate (PI(3)P) 222. PI(3)P signaling anchors later effectors that are required for phagosome maturation and
fusion to lysosomes in the phagosomal membrane 223; 224. M. tuberculosis actively modulates this failure to activate hPSV34 by shedding immunologically active lipoarabinomannan (LAM) and related lipoglycans, a lipid cell wall component that becomes distributed throughout the endocytic network of infected macrophages and prevents accumulation of Ca2+ in the host cytosol 225; 226. Lack of Ca2+
accumulation leads to failure to activate hVPS34 through calmodulin, and therefore
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leads to failure to generate PI(3)P and an arrest in maturation at an early phagosome state 222; 225. Synthesis of LAM and related lipoglycans require the building block mannose, a sugar that is converted to GDP-mannose-1-phosphate
(GDP-manp) 227. If mannose cannot be taken up from extracellular sources, as is likely the condition inside host macrophages, mycobacteria obtain mannose-6-
phosphate from the glycolytic pathway and then convert it in a series of reactions to
GDP-manp 227. Interestingly, enzymatic activity of glycolytic enzymes in humans,
mice and rats has been shown to be regulated by folate availability 228. Folate level
regulation by the proposed PknG-RplM-MutT3 cascade may therefore be necessary
for synthesis of LAM and other immunologically active lipoglycans through
regulation of glycolysis, thereby mediating immune evasion and maturation arrest
of mycobacterially infected phagosomes.
6.B.2. Regulation of folate metabolism
We have proposed a model in which PknG phosphorylates RplM, likely upon receiving cues regarding folate requirements in stationary phase growth like persistence in the macrophage or biofilm formation (Fig. 5-11). In our model, RplM co-localizes on a multi-protein complex with MutT3, and the phosphorylation event leads to a change in the shared environment of these two proteins and consequently leads to altered MutT3 activity, possibly by either promoting or preventing efficient binding of MutT3 to the complex (Fig. 5-11). The complex may be ribosomal or sub- ribosomal in nature, and we propose that MutT3 acts as a Nudix
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pyrophosphohydrolase 143; 144 to directly or indirectly affect folate metabolism and
lead to changes in folate availability. We believe the most likely function for MutT3
to be that of a DHN-3Pase in mycobacterial de novo folate biosynthesis 86; 147; 148, but we cannot exclude alternative substrates such as GTP or ppGpp.
While colonization of host macrophages represents a growth environment fairly specific to M. tuberculosis and other pathogenic mycobacteria, biofilm growth is not, and general folate regulation in late stage growth phases may be required in other bacteria as well. The investigated pathway involving phosphorylational control of a ribosomal protein and downstream regulation of pyrophosphohydrolase activity in the context of a multi-protein complex appears to function in both pathogenic and non-pathogenic mycobacterial species, and that this mode of control may not be a concept exclusive to the mycobacterial genus.
Beginning with PknG, which acts first in the hierarchy we have proposed, phosphorylational control of folate metabolism has to be considered. The eukaryotic-like serine/threonine kinases found in mycobacteria are involved in a variety of functions, including metabolic processes 132; 133; 134. PknG itself is
conserved throughout the entire mycobacterial genus, including M. leprae 140; 171, and a homolog has been identified in Corynebacterium glutamicum 121. It is
noteworthy that deletion of pknG in C. glutamicum leads to a set of phenotypes
distinct from those observed in mycobacteria; PknG seems to be primarily involved
in glutamate metabolic control in C. glutamicum 121 that remains mostly unaffected
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in mycobacteria 122; 141. Its main function in mycobacteria likely involves regulation of folate synthesis and metabolism, thereby affecting cell wall maintenance 159, survival of folate starvation, biofilm growth, multiple antibiotic tolerance 159 and, in
the case of M. tuberculosis, it affects inhibition of phagolysosomal fusion in infected host macrophages 20. Sequence identity of PknG from C. glutamicum and M.
tuberculosis is only 46%
(http://tuberculist.epfl.ch/quicksearch.php?gene+name=pkng&submit=Search),
while sequence identity within the mycobacterial genus amounts to at least 80% 140;
171. Therefore, substrate panels may not be identical between these two genera,
accounting for the observed phenotypic discrepancies. Also in favor of this theory is
the fact that the phosphorylated residue threonine 11 of RplM is conserved only
within the mycobacterial genus, and is not present in corynebacteria or in any bacteria lacking a PknG homolog (see Fig. 5-10). Regulation of folate metabolism by
PknG therefore appears to be a mechanism specific to mycobacteria, which may represent specific growth and infection modes in response to stimuli specific to mycobacterial molecules affected by folate biosynthesis, such as cell wall components exclusive to the mycobacterial cell envelope.
General phosphorylational control and involvement of ribosomal proteins or complexes in regulation of folate metabolic processes, on the other hand, are much more likely to occur in bacteria other than the mycobacterial genus. While further studies concerning the nature of the multi-protein complex on which RplM and
184
MutT3 appear to colocalize are required, it is likely that this complex is ribosomal or sub-ribosomal in its composition, although RplM has been shown to be active in non-ribosomal contexts as well 161. While dormant, non-culturable mycobacterial
cells are metabolically active 196, a decrease in ribosome numbers has been reported in M. smegmatis 229 that could free up ribosomal subunits, sub-ribosomal complexes
or single ribosomal protein components for regulation of other metabolic processes
such as folate biosynthesis.
Interestingly, phosphorylation of ribosomal proteins occurs in distinct profiles in vegetatively growing Dictyostelium discoideum, and also in response to starvation of these amoeba compared to actively growing, non-starved cells 230.
Among these phosphorylated ribosomal proteins in Dictyostelium was L13 230, a homolog of RplM 145. Phosphorylation of ribosomal components including RplM has also been identified in Escherichia coli, although no analysis of involved kinases and effects of phosphorylation has been performed 167. Phosphorylation of ribosomal
proteins similarly has been shown to occur in mammalian cells. It thus appears that
L13 is among proteins that can be phosphorylated both in isolated subunits or
mono-ribosomes as well as in poly-ribosomes, which markedly decrease
phosphorylation of other ribosomal proteins 231.
Cumulatively, it would appear that RplM phosphorylation is required in both actively dividing cells as well as dormant cells, though different residues may be phosphorylated by distinct kinases, depending on required modulation in the
185
different growth stages. Protein kinase activities have been shown to co-purify with
ribosomal fractions of Streptomyces collinus, an actinobacterium related to
mycobacteria 232. A number of ribosomal proteins were phosphorylated primarily
on serine and threonine residues, and this seemed to result in decreased
translational activity of the isolated ribosomes 232. By similar mechanisms, phosphorylation of RplM by PknG may lower the translational activity of ribosomes, which may become more accessible to MutT3, or alternatively free up complexes for association with and function in other metabolic processes such as folate metabolism.
Folate-dependent enzymes can act as translational regulators and modulate ribosomal activity, as is the case for thymidilate synthase, which can bind specific messenger RNAs (mRNA) directly, including its cognate mRNA as well as p53 mRNA, in eukaryotes 233. Binding of mRNA leads to translational repression of the message
233, demonstrating that folates influence basic cellular processes in multi-faceted
ways.
While ribosomal protein phosphorylation and ribosomal complex
involvement have not previously been described in the context of the de novo folate biosynthesis, non-ribosomal multi-enzyme complexes have been described to
associate with 5,6,7,8-tetrahydrofolate in microorganisms capable of folate
biosynthesis, and interestingly, intermediary enzymes of the pathway associate with
this same complex 234. Likewise, enzymes, whose activity is controlled by folate
186
biosynthesis via the requirement for folates as co-enzymes, have been reported to
occur in multi-protein complexes. Such is the case for example with the glycine
decarboxylase multi-enzyme system of plant mitochondria that is involved in
degradation of glycine 235. Glycine is a molecule that is essential for transition of intracellular M. tuberculosis into dormancy and therefore successful persistence of
TB 15; 194; 195, and glycine synthesis is also folate-dependent (see also section 6.B.1.).
Clearly, folate reactions required for both folate biosynthesis and
consumption can occur in large multi-protein complexes and regulate various
metabolic and other cellular pathways. Ribosomal involvement or binding of
ribosomal proteins to other protein complexes could therefore represent a widely
used strategy of tight control of folate levels inside the cell, which in turn modulate almost all cellular processes. Phosphorylation of ribosomal proteins occurs across all kingdoms of life and therefore may be a convenient control mechanism for such folate level modulation, although it is likely that the involved kinases, phosphorylated residues, and their resulting effects on folate levels will vary among organisms as well as across the growth or stress conditions that require folate adjustments.
6.B.3. Implications for TB and MDR or XDR TB
Due to the growing spread of MDR and XDR TB infections 23; 36; 37 that cannot be treated with the standard first-line TB drugs 1; 36; 37; 38, we face an urgent need for
alternative treatment options.
187
One such alternative approach is the concept of “targeting resistance” 45, which seeks to use current knowledge of molecular resistance mechanisms employed by bacteria in order to (re)sensitize pathogens to already available, approved drugs 45. This is achieved by coadministration of existing drugs and
inhibitors suppressing resistance mechanisms to these drugs, so that currently
ineffective drugs (re)gain antimicrobial activity (Fig. 1-3) 45; 46; 47. Classical antifolate
antibiotics attack folate de novo biosynthesis in only two steps (Fig. 1-4), and no
potentiators are available, although drugs targeting both steps are commonly
combined to achieve synergy and appear to be a promising group of drugs to use in
TB infection 98; 99; 100. However, resistance can readily be gained by point mutations
in one or both of the enzymes targeted 80; 83, and therefore identification of
potentiators that inhibit intrinsic antifolate tolerance mechanisms in steps not
involving these enzymes could present a valuable strategy for alternative
antimycobacterial treatment development 45; 102.
We have presented here evidence in favour of a regulatory pathway in mycobacteria involving the serine/threonine protein kinase PknG, the ribosomal protein RplM and the putative Nudix hydrolase MutT3 (Fig. 5-11), the disruption of which leads to increased susceptibility to classical antifolate antibiotics. Inhibitors targeting either of these three proteins or other steps that may occur in the proposed pathway should therefore act as potentiators of antifolate antimycobacterial activity, and we have shown here that, at least in vitro, chemical
188
inhibition of PknG can potentiate sulfadrug action against M. tuberculosis (Fig. 4-1).
Pharmaceutical inactivation of this regulatory pathway as well as other pathways regulating folate availability could possibly also potentiate the effects of current anti-TB drugs that target folate metabolism (PAS, INH, etc.) 45; 102. Potentiation of the anti-TB efficacy of classical antifolates may also allow for reduction of effective therapeutic doses, thereby minimizing the cytotoxicity that has been a clinical problem 45; 102.
Detailed studies concerning RplM modulation of MutT3 activity, the nature of the multi-protein complex involved, as well as MutT3 enzymatic activity and substrates should yield further targets for inhibition of folate level control and antifolate potentiator development. Such inhibitors could be obtained by rational design or small molecule screens 236; 237; 238, and would then have to be tested for
potentiation in in vitro mycobacterial cultures and in in vivo settings, progressing from macrophage experiments through mice studies and clinical trials 44; 45; 239.
Clinical trials may be significantly shortened in comparison to those involving entirely new drug classes, however, since potentiator targets in folate de novo
biosynthesis are not present in humans and these molecules should therefore not
affect patients, unless they exhibit off-target effects 86; 87; 88. Antifolates have been
extensively studied 80; 83, are used on a standard basis for human treatment of
various diseases 80; 81; 82, and already have been characterized for side effects 240; 241;
189
242, which might even be further reduced by potentiation-enabled lower antifolate
doses necessary to obtain sufficient antimycobacterial activity of these drugs 45.
We have demonstrated the requirement for PknG-mediated folate metabolism regulation in preventing lysosomal delivery to mycobacteria containing phagosomes (Fig. 4-5), which indicates that co-administration of inhibitors to PknG and other key players in this regulatory pathway together with antifolate antibiotics could also promote phagosome-lysosome fusion, leading to improved killing and clearance of M. tuberculosis by host immune system mechanisms. Targeting PknG, its substrate RplM, and downstream proteins involved in folate metabolic control such as MutT3 may therefore effectively be doubly valuable and facilitate both drug- and immune-mediated antimycobacterial mechanisms, providing a very promising alternative MDR and XDR TB treatment option.
190
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