Abstract
Increasing experimental evidence supports the idea that Mycobacterium tuberculosis has evolved strategies to survive within lysosomes of activated macrophages. To further our knowledge of M. tuberculosis response to the hostile lysosomal environment, we profiled the global transcriptional activity of M. tuberculosis when exposed to the lysosomal soluble fraction (SF) prepared from activated macrophages. Transcriptome sequencing (RNA-seq) analysis was performed using various incubation conditions, ranging from noninhibitory to cidal based on the mycobacterial replication or killing profile. Under inhibitory conditions that led to the absence of apparent mycobacterial replication, M. tuberculosis expressed a unique transcriptome with modulation of genes involved in general stress response, metabolic reprogramming, respiration, oxidative stress, dormancy response, and virulence. The transcription pattern also indicates characteristic cell wall remodeling with the possible outcomes of increased infectivity, intrinsic resistance to antibiotics, and subversion of the host immune system. Among the lysosome-specific responses, we identified the glgE-mediated 1,4 α-glucan synthesis pathway and a defined group of VapBC toxin/anti-toxin systems, both of which represent toxicity mechanisms that potentially can be exploited for killing intracellular mycobacteria. A meta-analysis including previously reported transcriptomic studies in macrophage infection and in vitro stress models was conducted to identify overlapping and nonoverlapping pathways. Finally, the Tap efflux pump-encoding gene Rv1258c was selected for validation. An M. tuberculosis ΔRv1258c mutant was constructed and displayed increased susceptibility to killing by lysosomal SF and the antimicrobial peptide LL-37, as well as attenuated survival in primary murine macrophages and human macrophage cell line THP-1.
INTRODUCTION
Mycobacterium tuberculosis infects a third of the world's population and causes death to millions of infected individuals annually. While 90% of the infected population is able to prevent progression into active disease, incomplete sterilization of the infecting bacilli, typically within granulomatous lesions formed in the lungs, leads to latent tuberculosis (TB), the asymptomatic form of the disease. It is a longstanding paradigm that these lesions provide a niche environment that induces TB latency, where the bacterium is believed to enter a state of bacteriostasis or very slow replication with low energetic and metabolic activities and retains the ability to resume growth under permissive conditions, leading to disease reactivation (1).
Macrophages represent a large proportion of the cell populations that are present in a TB lung granuloma (2, 3). Their phagocytic abilities are responsible for eliminating most intracellular microbes, and as such macrophages are important players in host innate immunity (4, 5). However, upon phagocytosis, internalized M. tuberculosis is able to survive and replicate within the phagosome by blocking its fusion with lysosomes according to a process that involves several mycobacterial lipid and protein factors (6). Following the onset of cell-mediated immunity, however, macrophage activation overrides phagosome maturation arrest and delivers M. tuberculosis into the lysosomal compartment (7, 8), characterized by an increased acidic environment and containing a plethora of bactericidal molecules, including hydrolytic enzymes, oxygenated lipids, fatty acids, reactive oxygen species and nitrogen intermediates, and antimicrobial peptides. However, killing of mycobacteria in activated macrophages appears to be a protracted event, as evidenced by the detection of low numbers of viable bacilli 7 days postinfection (7, 8). With an increasing number of mycobacterial factors reported to be specifically implicated in M. tuberculosis survival within activated macrophages (9), it seems that this pathogen has evolved strategies to adapt and survive within this hostile compartment, thereby challenging the idea that the lysosomal compartment is a dead end for M. tuberculosis. Specific M. tuberculosis responses to the lysosomal environment could therefore be exploited to identify novel targets and develop novel anti-TB drugs. However, there is limited knowledge on the behavior and physiology of M. tuberculosis in the lysosomal compartment.
Transcriptional profiling of M. tuberculosis from infected macrophages of human or mouse origin has been the typical approach to decipher the behavior of intramacrophage M. tuberculosis (10–15). A comparative study between resting and gamma interferon (IFN-γ)-activated macrophages identified a specific subset of mycobacterial genes that were distinctly modulated in activated macrophages, thereby supporting a lysosome-specific transcriptional reprogramming in M. tuberculosis with the potential to adapt to the inhospitable lysosomal microenvironment (11). While these studies have captured dynamic global transcriptional changes in M. tuberculosis during macrophage infection, contradictory observations were also reported, likely due to underlying experimental differences between these macrophage infection models, for instance, the macrophage type and M. tuberculosis strains employed and/or the time postinfection at which the transcriptome was assessed. The unsynchronized infection process throughout the macrophage population could generate a transcriptional profile representative of a combination of M. tuberculosis gene responses to multiple microenvironments encountered during macrophage infection which prevent the dissection of responses pertaining to each of the subcellular environmental niches encountered by M. tuberculosis during its intramacrophage life. To address these limitations, gene expression studies have been conducted in defined in vitro culture settings that feature one particular stress or growth condition possibly encountered by M. tuberculosis during macrophage infection, including hypoxia (16), nitric oxide (17, 18), iron limitation (19), acidic pH (20), gradual oxygen depletion (21, 22), nutrient starvation (23, 24), antibiotic pressure (25), and stationary phase (26). These studies have allowed the identification of M. tuberculosis genes that respond specifically to a particular environmental cue or growth condition.
Our work aims to study the transcriptional response of M. tuberculosis to the lysosomal content using RNA sequencing (RNA-seq). M. tuberculosis was exposed to the lysosomal soluble fraction (SF) prepared from activated macrophages. Previous work has shown that the lysosomal SF possesses mycobactericidal activity in a dose- and time-dependent manner (27). Here, upon exposure to SF conditions that led to an absence of apparent mycobacterial replication, we report a unique transcriptional signature as part of M. tuberculosis adaptive response to the hostile lysosomal environment.
MATERIALS AND METHODS
Ethics statement.
All of the animal experiments were carried out under the guidelines of the National Advisory Committee for Laboratory Animal Research (NACLAR) in the AAALAC-accredited NUS animal facilities (http://nus.edu.sg/iacuc/). NUS has obtained a license (VR008) from the governing body Agri-Food & Veterinary Authority of Singapore (AVA) to operate an Animal Research Facility. The animal experiments described in this work were approved by the IACUC from the National University of Singapore under protocol number R2014-00723.
Bacterial strains and growth conditions.
The parental strain of M. tuberculosis CDC1551, its derived mutant, and complemented strains were grown in Middlebrook 7H9 medium (Difco) supplemented with 10% ADS [50 g bovine fraction V albumin, 20 g d-(+)-glucose, 8.1 g sodium chloride per liter], 0.05% Tween 80, and 0.5% glycerol or on Middlebrook 7H11 agar containing oleic acid-albumin-dextrose-catalase (OADC; Becton Dickinson) and 0.5% glycerol. When appropriate, hygromycin and kanamycin were added at 80 and 20 μg/ml, respectively. Hygromycin was purchased from Roche. Kanamycin, streptomycin, and carbonyl cyanide m-chlorophenyl hydrazone (CCCP) were purchased from Sigma. For CFU enumeration, serial dilutions were performed in the Middlebrook 7H9 medium and plated on Middlebrook 7H11 agar. Plates were incubated at 37°C for 3 to 4 weeks.
Determination of the MIC of streptomycin.
Mid-log-phase mycobacterial cultures were grown in 7H9 medium and diluted to an optical density at 600 nm (OD600) of 0.02. The diluted bacterial suspension (200 μl) was added to 2-fold serially diluted streptomycin (5 μl) in a flat-bottom 96-well plate and incubated for 5 days. The OD600 of the cultures were measured using a Bio-Rad iMark microplate absorbance reader at 600 nm. The values were plotted against the log concentrations of streptomycin, and a sigmoidal dose-response curve was fitted to the plot. The MIC corresponded to the concentration which inhibits 100% of visible bacterial growth based on the OD600.
Extraction of lysosomal SF.
The lysosomal soluble fractions (SF) were extracted from activated bone marrow-derived macrophages (BMMOs) as previously described (27). T75 flasks of confluent BMMOs were incubated for 2 h at 37°C and 5% CO2 with 5 ml of 40 mg/ml iron-dextran (40 kDa) mixed with 2× Opti-MEM (Gibco) at a 1:1 ratio. The monolayers were rinsed twice in 10 ml of warmed sterile phosphate-buffered saline (PBS) to remove the excess Fe-dextran and chased overnight in culture medium. The cells were scraped in 5 ml of homogenization buffer (HB) (250 mM sucrose, 0.5 mM EGTA, 0.1% gelatin, and 20 mM Tris, pH 7.0), centrifuged at 1,500 rpm at 4°C for 10 min, and lysed by passing through a tuberculin syringe. The lysate was subjected to low-speed centrifugation at 1,000 rpm at 4°C for 10 min to remove debris, nuclei, and intact cells. The supernatant was applied to a MiniMACS column (Miltenyi Biotech) placed on a magnetic stand to retain the iron-loaded lysosomes. After two washes with HB, the column was removed from the magnetic stand, and the bound iron-loaded lysosomes were eluted twice with 500 μl of HB. The lysosomes were spun down at 12,000 rpm for 30 min and stored as a dry pellet at −20°C until use. To prepare the lysosomal SF, each pellet was resuspended in 200 μl of SF buffer (1% Tween 20, 20 mM sodium acetate, pH 5.5). The lysates from eight pellets (2.5 × 108 cells) were pooled and applied to two MidiMACS (Miltenyi Biotech) columns to remove iron. The flowthrough was collected and centrifuged at 100,000 rpm at 4°C for 50 min. The supernatant corresponding to the SF was collected and the total protein content was estimated using the bicinchoninic acid (BCA) protein assay kit (Thermo-Scientific Pierce). SF was stored at −80°C until use.
Bactericidal assays.
Bactericidal assays on M. tuberculosis strains were performed with SF and synthetic human cathelicidin (LL-37; Peptide Institute, Japan). LL-37 was reconstituted in 0.01% acetic acid for storage in −80°C until use. When required, 1 μg/ml CCCP was added to the medium. Mid-log-phase mycobacterial cultures of 5 × 105 CFU/ml were treated with the indicated concentrations of the bactericidal agents for the indicated periods of time. The number of surviving bacteria was enumerated by plating appropriate dilutions of the mixture on 7H11 agar and incubating at 37°C for 3 weeks.
RNA isolation and qualitative real-time PCR.
Mycobacterial cultures were incubated with RNAprotect bacterial reagent (Qiagen) for RNA stabilization. The pelleted bacteria were then resuspended in 100 μl Tris-EDTA (TE) containing 20 μg/ml lysozyme and incubated at room temperature for 20 min. RNA extraction was then performed using an RNeasy minikit (Qiagen) according to the manufacturer's instructions. Contaminating genomic DNA from the eluted total RNA was removed using the Turbo DNA-free kit according to the manufacturer's protocol. The RNA concentrations and purity were measured using a NanoDrop 1000 spectrophotometer (Thermo Scientific). Reverse transcription was performed on 10 ng bacterial RNA using the iScript cDNA synthesis kit (Bio-Rad). Real-time PCR was performed in a 96-well plate with each well containing 2 μl cDNA mix, 0.5 μl forward (F) and reverse (R) primers (0.5 μM final), and 25 μl SYBR green supermix with ROX (Bio-Rad) to a final volume of 50 μl. The list of primers is presented in Table S7 in the supplemental material. Samples were run in triplicate. Real-time PCR amplification was conducted with the ABI Prism 7500 sequence detector (Applied Biosystems) over 40 cycles and with an annealing temperature of 61°C. The expression of each target gene was based on relative quantification (RQ) using the comparative critical threshold (CT) value method. Relative quantification of a specific gene was evaluated in each reaction by normalization to the CT value obtained for the endogenous control gene, sigA. For validation of transcriptome sequencing (RNA-seq) data, fold changes (RQ values) were derived with reference to expression levels from M. tuberculosis incubated with SF buffer for 48 h. For validation of Rv1258c overexpression, the fold change was derived with reference to expression from WT M. tuberculosis.
RNA-seq library preparation.
Total DNA-free RNA sample was depleted of bacteria rRNA with Ambion's MICROBExpress kit (AM1905) per the manufacturer's instructions. Bacterial rRNA-depleted sample was processed using the TruSeq RNA sample preparation (v2) per the manufacturer's instructions. Library preparation entailed fragmentation, 1st- and 2nd-strand cDNA synthesis, end repair, A tailing, and ligation of adapters with multiplex indexes according to the manufacturer's instructions. Samples were enriched with 15 PCR cycles followed by Agencourt AMPure XP magnetic bead (Beckman Coulter, Brea, CA, USA) clean up according to the manufacturer's instructions. The quality of cDNA libraries was checked with Agilent DNA1000 chips (2100 Bioanalyzer; Agilent Technologies, Santa Clara, CA, USA). Next-generation sequencing was performed using an Illumina HiSeq 2000 flow cell with two 76-bp end runs. PhiX was used as a control.
RNA-seq data analysis.
RNA-seq data analysis was performed on the CLC Genomics platform. Sequence reads were aligned to the Mycobacterium tuberculosis CDC1551 parental reference genome (GenBank accession number NC_002755). The reads per kilobase per million (RPKM) value for each gene was generated. Differential gene expression analysis using the R edgeR package was performed for the following groups of data sets: incubation for 24 h or 48 h at 0, 10, or 20 μg/ml SF. The exact-test function was applied to determine the association of the differences in expression read counts within each group, and corresponding P values were adjusted using the default Benjamini & Hochberg procedure. Their adjusted P values, in −log10 scale on the y axis and fold changes in log2 scale on the x axis, were plotted as a volcano plot. Differential gene expression was determined by a false discovery rate (FDR) of <0.01. Genes with read counts of less than 5 from both SF-treated and nontreated groups were also eliminated. Further functional annotation clustering analysis was performed using Database for Annotation, Visualization and Integrated Discovery (DAVID), version 6.7 (28, 29), and TB Database (http://www.tbdb.org/).
Meta-analysis with in vitro and ex vivo models of M. tuberculosis.
Microarray-based transcriptome studies of M. tuberculosis in in vitro and ex vivo models of M. tuberculosis were selected for comparative analysis with the M. tuberculosis transcriptome generated in this study. For short-term primary murine macrophage (BMMO) (10, 11) and human macrophage (THP-1) infection studies (13), genes that were differentially expressed at 24 h or 48 h after infection were considered. For temporal studies based on BMMO (30), genes that exhibited significant temporal trends were considered. For analysis with in vitro models of M. tuberculosis persistence, differential M. tuberculosis transcriptomes generated from gradual hypoxic (21), defined hypoxic (22), nutrient starvation (24) and drug-tolerant persister (25) models were considered.
Construction of ΔRv1258c mutant and Ox-Rv1258c complemented strains.
The ΔRv1258c mutant strain was generated in the M. tuberculosis CDC1551 background by allelic exchange using the suicide plasmid backbone pYUB854, as previously described (31). Briefly, primers with relevant restriction enzyme sites (see Table S7 in the supplemental material) were designed to amplify 5′ and 3′ PCR fragments (∼1 kb) flanking the Rv1258c open reading frame (ORF) from genomic DNA of the M. tuberculosis CDC1551 parental strain. The fragments were cloned into the pYUB854 plasmid at its corresponding multiple cloning sites (MCS) flanking the hygromycin resistance gene, hyg. A PacI-restricted fragment containing the selection genes lacZ and sacB was obtained from pGOAL17 (32) and cloned into the pYUB854-PCR5′-3′ construct to obtain the final delivery vector, pYUB854-Rv1258c. The overexpressing complemented strain Ox-Rv1258c was constructed by introducing Rv1258c, under the strong constitutive mycobacterial hsp60 promoter, into the ΔRv1258c mutant strain using a promoter-less integrative plasmid, pMV306 (33). The mycobacterial hsp60 promoter was excised from pMV262 and cloned into MCS of pMV306 vector. The ORF of Rv1258c was amplified from M. tuberculosis CDC1551 parental genomic DNA using primers indicated in Table S7 and subsequently were inserted downstream of the hsp60 promoter to obtain the final delivery vector, pMV306-Rv1258c. The UV-irradiated plasmid solutions (1 μg) were electroporated into the respective M. tuberculosis strains as described previously (32). To identify the ΔRv1258c mutant, hygromycin-resistant white colonies were selected. Deletion at the Rv1258c locus was verified by PCR using primers listed in Table S7 and Southern blot analysis. To identify the Ox-Rv1258c strain, kanamycin-resistant colonies were selected. Quantitative reverse transcription-PCR (qRT-PCR) was used to detect increased transcriptional activity of Rv1258c.
Southern blot analysis.
Chromosomal DNA (1 μg) prepared from each M. tuberculosis strain was digested with EcoRI and XmaI for 4 h and subjected to 0.8% agarose gel electrophoresis. The agarose gel containing the digested DNA was chemically treated and transferred onto a nitrocellulose membrane (Millipore) according to Roche's digoxigenin (DIG) application manual. The membrane was UV fixed for 1 min and equilibrated with 10 ml preheated DIG Easy Hyb solution (Roche) at 65°C for 20 min, with gentle agitation. A DIG-labeled probe was amplified using the PCR DIG probe synthesis kit (Roche) according to the manufacturer's instructions and primers as listed in Table S7. For hybridization, about 5 to 25 ng/ml heat-denatured DIG-labeled DNA probe in DIG Easy Hyb solution was incubated with the membrane overnight at 65°C. Detection was performed using alkaline phosphatase (AP)-conjugated anti-DIG antibody (Roche) at a dilution of 1:5,000. The membrane was developed using nitroblue tetrazolium–5-bromo-4-chloro-3-indolylphosphate (NBT-BCIP)-AP substrate (Chemicon).
Macrophage survival assays.
Bone marrow cells were flushed from femurs of 6- to 8-week-old BALB/c mice, seeded onto petri dishes (4 femurs per dish; Greiner), and differentiated into macrophages over 6 days in BMMO complete medium supplemented with 10 ng/ml recombinant mouse macrophage colony-stimulating factor (rM-CSF; R&D Systems). Differentiated macrophages were recovered by dislodging them in cold 1× PBS containing 1 mM EDTA (pH 7.4) and washed once in 1× PBS. To prepare activated macrophages, the complete medium was supplemented with 10% horse serum (Gibco) and macrophages were activated with 100 U/ml recombinant mouse IFN-γ (Chemicon) and 50 ng/ml of tumor necrosis factor (TNF) for 48 h. Primary macrophages consistently represented 70 to 80% of the total cell population harvested, as determined by flow cytometry using a panmacrophage marker, anti-F4/80 antibody (eBioscience). A human THP-1 monocytoid cell line (ATCC TIB-202; ATCC, MD, USA) was maintained at 37°C and 5% CO2 in HEPES buffered RPMI 1640 (Sigma-Aldrich, St. Louis, MO, USA) medium with 10% heat-inactivated fetal bovine serum (FBS), 2 mM l-glutamine, 10 mM HEPES, 1 mM sodium pyruvate, 4,500 mg/liter glucose, and 1,500 mg/liter sodium bicarbonate (pH 7). When needed, cells were expanded into 75-cm2 flasks and were activated with retinoic acid (RA; 1 μM) and vitamin D3 (VD; 1 μM) for 3 days as described previously (34). For survival assays, BMMO monolayers (5 × 104 cells/well) or RAVD-activated THP-1 cells (2.5 × 104 cells/well) in 24-well tissue culture plates (Nunc) were incubated with mycobacteria at multiplicities of infection (MOI) of 2 and 5, respectively, for 45 min in their respective incomplete culture media (culture media without penicillin-streptomycin and FBS). Infected cells were washed twice with 1× PBS, and the respective complete culture medium without penicillin-streptomycin was added to each well. At the indicated time points, cells were washed with 1× PBS and lysed with 0.1% Triton X-100 (Sigma-Aldrich) to release the intracellular bacteria. The cell lysates were serially diluted in 7H9 medium and plated on 7H11 agar. The number of CFU was enumerated after incubation at 37°C for 16 days.
Statistical analysis.
Statistical significance was assessed by the Student t test, and two-tailed P values of less than 0.05 were considered statistically significant.
Accession number.
The data discussed in this publication have been deposited in NCBI's Gene Expression Omnibus (GEO) and are accessible through GEO series accession number GSE68337.
RESULTS AND DISCUSSION
The LivE model.
The lysosomal in vitro exposure (LivE) model consists of the direct exposure of M. tuberculosis to the soluble fraction of lysosomes (SF) purified from activated murine bone marrow-derived macrophages (BMMO) based on a previously described protocol (27). The mycobactericidal activity of SF preparations was determined by incubating mid-log-phase M. tuberculosis cultures with a range of SF total protein concentrations for 24 and 48 h. As previously reported (27), the mycobactericidal activity of SF was found to be both time and concentration dependent (Fig. 1A). M. tuberculosis remained viable and unaffected in its growth rate after 24 h of incubation within the range of SF concentrations tested, as evidenced by CFU values being comparable to those of the positive control (buffer only) at 24 h. In contrast, a significant and dose-dependent reduction in viable CFU was observed after 48 h of coincubation.
FIG 1.
Mycobactericidal activity of a lysosomal soluble fraction (SF) prepared from activated primary murine macrophages. Mid-log-phase in vitro cultures of M. tuberculosis CDC1551 strain were coincubated for 24 or 48 h with a lysosomal SF prepared from activated primary murine macrophages at the indicated concentrations or with SF buffer only. (A) The treated bacteria were then plated on 7H11 agar and enumerated for viable CFU after 16 days of incubation at 37°C. The dotted line represents the initial inoculum. (B) Results are expressed as a percentage of viable CFU obtained with buffer only at their respective times of incubation. Data shown are the means ± standard deviations (SD) from triplicates.
Based on the growth profiles observed, we defined the following LivE conditions, ranging from noninhibitory to cidal upon SF exposure. Noninhibitory conditions consist of exposing M. tuberculosis to 10 to 30 μg/ml SF for 24 h, which led to growth comparable to 24 h of incubation with buffer only (24 h control) (Fig. 1). The subinhibitory condition was achieved by exposing M. tuberculosis to 10 μg/ml SF for 48 h and was characterized by a significant decrease in cell viability compared to the 48-h buffer control (Fig. 1) but a greater number of viable CFU compared to the 24-h control (Fig. 1A). The inhibitory condition was obtained upon incubation of M. tuberculosis in the presence of 20 μg/ml SF for 48 h, which resulted in a concentration of viable bacteria that was comparable to the inoculum concentration and significantly lower than that obtained with the 48-h untreated control (Fig. 1A and B). This suggested that incubation with 20 μg/ml SF for 48 h led to an apparent replication arrest, which can be the result of (i) a true arrest in replication where mycobacteria cease dividing but do not die, as described for other stress conditions, such as hypoxia (35) or starvation (36), or (ii) equal killing and replication rates that cancel each other out. Finally, the cidal condition was observed when M. tuberculosis was incubated with 30 μg/ml SF for 48 h, which resulted in a drastic reduction in viable CFU compared to the 48-h control (Fig. 1B).
RNA sequencing of M. tuberculosis in the LivE model.
To investigate the transcriptome profile of M. tuberculosis upon exposure to SF, M. tuberculosis was exposed to noninhibitory (10 μg/ml SF, 24 h), subinhibitory (10 μg/ml SF, 48 h), and inhibitory (20 μg/ml SF, 48 h) conditions, with buffer only (0 μg/ml SF 24 h and 48 h) as the reference control. Illumina sequencing was performed on biological triplicates of cDNA libraries prepared from mRNA extracted from the SF-treated M. tuberculosis cultures. High-quality paired-end sequence reads were generated for each sample and were aligned with the M. tuberculosis CDC1551 parental reference genome, revealing coverage of more than 264 for all samples and indicating a high accuracy in the sequences generated. More than 89% of the sequence tags were mapped to the annotated CDS in the sense orientation. Differential expression analysis was performed with the R edgeR package (see Materials and Methods). We observed that the number of differentially expressed M. tuberculosis genes increased with increasing growth-inhibitory SF conditions, with more genes being induced than repressed, as illustrated in the volcano plots (Fig. 2). In addition, the majority of the genes found to be modulated under noninhibitory conditions were further modulated under the subinhibitory and inhibitory conditions.
FIG 2.
Volcano plots of M. tuberculosis genes in the LivE model. Transcriptomes of M. tuberculosis exhibited differential expression under noninhibitory (10 μg/ml SF, 24 h) (A), subinhibitory (10 μg/ml SF, 48 h) (B), and inhibitory (20 μg/ml SF, 48 h) (C) conditions. A total of 4,293 M. tuberculosis CDC1551 genes were annotated.
The inhibitory LivE condition (iLivE) of 20 μg/ml for 48 h was then selected for further analysis, where the apparent replication appears to resemble the nonreplicative state described for mycobacteria exposed to other environmental stresses, such as hypoxia (35) or nutrient starvation (36) (Fig. 1). The iLivE M. tuberculosis genes were short-listed based on an FDR of <0.01 and disregarding genes with expression read counts of <5 (see Table S1 in the supplemental material). Gene function annotation was performed using DAVID and TBDB databases. The distribution of iLivE M. tuberculosis genes into different functional categories showed that a significant number of genes were involved in cell wall remodeling and substrate transport, intermediary metabolism and respiration, lipid metabolism, information pathways, regulatory proteins, and virulence, detoxification, and adaptation (Table 1). In total, there were 264 upregulated (Table 2) and 106 downregulated (Table 3) iLivE M. tuberculosis genes.
TABLE 1.
Functional categories representing the iLivE M. tuberculosis transcriptome
| Functional categorya | No. of genesb |
|
|---|---|---|
| Induced | Repressed | |
| Cell wall and cell processes | 27 | 15 |
| Information pathways | 17 | 2 |
| Insertion sequences and phages | 10 | |
| Intermediary metabolism and respiration | 41 | 20 |
| Metabolism | 13 | 13 |
| PE/PPE | 5 | 7 |
| Pseudogenesc | 8 | |
| Regulatory proteins | 15 | 2 |
| Virulence, detoxification, and adaptation | 29 | 11 |
| Unknown | 8 | 5 |
| Conserved hypotheticals | 91 | 31 |
| Total | 264 | 106 |
Functional annotations were obtained from TB Database (http://www.tbdb.org).
The number of induced and repressed genes is given for each gene category.
Genes that are not annotated in the M. tuberculosis H37Rv background.
TABLE 2.
Genes induced in iLivE M. tuberculosis
| Functional categorya | Designation for strain: |
Gene | Description | Fold change | SD | |
|---|---|---|---|---|---|---|
| CDC1551b | H37Rv | |||||
| Cell wall and cell processes | MT0201 | Rv0191 | Sugar transporter family protein | 1.93 | 0.2 | |
| MT0409 | Rv0399c | lpqK | Putative lipoprotein | 2.37 | 0.5 | |
| MT0623 | Rv0593 | mce2E (lprL) | Mce family protein | 1.81 | 0.2 | |
| MT0700 | Rv0671 | lpqP | Lipoprotein | 2.09 | 0.3 | |
| MT0961 | Rv0934 | pstS1 (phoS1) | Periplasmic phosphate-binding lipoprotein | 1.90 | 0.8 | |
| MT1013 | Rv0985c | mscL | Large-conductance ion mechanosensitive channel | 1.95 | 0.2 | |
| MT1297 | Rv1258c | tap | Putative Tap-like membrane efflux pump | 5.94 | 0.3 | |
| MT1503 | Rv1456c | Antibiotic transport membrane ABC transporter | 1.82 | 0.5 | ||
| MT1519 | Rv1473 | Probable macrolide transport ATP-binding protein ABC transporter | 1.89 | 0.4 | ||
| MT1642 | Rv1607 | chaA | Cation/proton antiporter | 1.97 | 0.3 | |
| MT1926 | Rv1877 | Drug transporter | 1.94 | 0.4 | ||
| MT1973 | Rv1922 | Peptidase, putative | 1.88 | 0.8 | ||
| MT1997 | Rv1946c | lppG | Possible lipoprotein | 1.84 | 0.9 | |
| MT2016 | Rv1964 | yrbE3A | Membrane protein | 2.05 | 1.3 | |
| MT2031 | Rv1979c | Amino acid permease | 2.03 | 0.5 | ||
| MT2040 | Rv1986 | Putative amino acid transporter | 2.08 | 0.9 | ||
| MT2097 | Rv2037c | Conserved transmembrane protein | 1.84 | 0.5 | ||
| MT2100 | Rv2040c | Sugar ABC transporter, permease protein | 1.87 | 0.7 | ||
| MT2101 | Rv2041c | Sugar ABC transporter, sugar-binding protein | 1.83 | 1.2 | ||
| MT2334 | Rv2273 | Probable conserved transmembrane protein | 2.00 | 1.2 | ||
| MT2339 | Rv2281 | pitB | Putative phosphate permease | 3.59 | 1.5 | |
| MT2469 | Rv2398c | cysW | Sulfate transport membrane protein ABC transporter | 1.93 | 0.6 | |
| MT2471 | Rv2400c | sbp | Sulfate ABC transporter substrate-binding protein | 1.85 | 0.4 | |
| MT2598 | Rv2522c | Peptidase, M20/M25/M40 family | 1.80 | 0.2 | ||
| MT2901 | Rv2835c | ugpA | Probable Sn-glycerol-3-phosphate transport integral membrane protein ABC transporter | 2.22 | 0.8 | |
| MT3080 | Rv3000 | Possible conserved transmembrane protein | 3.11 | 1.4 | ||
| MT3951 | Rv3843c | Probable conserved transmembrane protein | 1.87 | 0.4 | ||
| Information pathways | MT0691 | Rv0662c | DNA-binding protein, CopG family, transcriptional repressor | 2.08 | 1.0 | |
| MT0699 | Rv0670 | end | AP endonuclease, family 2 | 2.01 | 0.5 | |
| MT1338 | Rv1299 | prfA | Peptide chain release factor 1 | 2.01 | 0.3 | |
| MT1463 | Rv1420 | Probable excinuclease ABC (subunit C nuclease) | 1.77 | 0.3 | ||
| MT1589 | Rv1537 | dinX | DNA polymerase IV DinX | 2.09 | 0.5 | |
| MT2042 | Rv1988 | ermMT | rRNA adenine N-6-methyltransferase, putative | 2.62 | 0.4 | |
| MT2247 | Rv2191 | DNA polymerase III, epsilon subunit, putative | 1.90 | 0.4 | ||
| MT2488 | Rv2415c | comE operon protein 1, putative | 2.50 | 0.7 | ||
| MT2535 | Rv2460c | clpP2 | ATP-dependent Clp protease proteolytic subunit 2 | 2.12 | 0.5 | |
| MT2536 | Rv2461c | clpP1 | Probable ATP-dependent CLP protease proteolytic subunit 1 | 2.18 | 0.8 | |
| MT2741 | Rv2667 | clpC2 | ATP-dependent protease ATP-binding subunit | 2.10 | 0.8 | |
| MT2902 | Rv2836c | dinF | DNA damage-inducible protein F, putative | 1.92 | 0.6 | |
| MT2904 | Rv2838c | rbfA | Ribosome-binding factor A | 2.02 | 0.5 | |
| MT2905 | Rv2839c | infB | Translation initiation factor IF-2 | 2.01 | 0.6 | |
| MT2942 | Rv2874 | dipZ | Cytochrome c biogenesis protein | 2.19 | 0.5 | |
| MT3686 | Rv3580c | cysS | Cysteinyl-tRNA synthetase 1 | 2.62 | 0.2 | |
| MT3942 | Rv3834c | SERYL-tRNA synthetase | 1.84 | 0.4 | ||
| Insertion sequences and phages | MT0850 | Rv0829 | IS1605′, transposase, truncation | 2.08 | 2.4 | |
| MT0873 | Rv0850 | IS1606′, transposase | 2.02 | 1.2 | ||
| MT0948 | Rv0921 | IS1535, resolvase | 2.13 | 1.0 | ||
| MT2069 | Rv2013 | IS1607, transposase | 2.67 | 0.8 | ||
| MT2070 | Rv2014 | IS1607, transposase | 2.11 | 0.2 | ||
| MT2497 | Rv2424c | IS1558, transposase | 2.48 | 1.1 | ||
| MT2732 | Rv2655c | Possible PhiRv2 prophage protein | 2.03 | 0.1 | ||
| MT2735 | Rv2646 | Integrase | 1.94 | 1.0 | ||
| MT2953 | Rv2885c | IS1539, transposase | 1.80 | 0.7 | ||
| MT3573.3 | Bacteriophage protein | 1.85 | 0.2 | |||
| Intermediary metabolism and respiration | MT0098 | Rv0089 | Putative methyltransferase | 3.50 | 1.4 | |
| MT0207 | Rv0197 | lpqS | Molybdopterin oxidoreductase | 1.80 | 0.7 | |
| MT0337 | Rv0322 | udgA | UDP-glucose 6-dehydrogenase | 1.91 | 0.7 | |
| MT0511 | Rv0492c | Oxidoreductase, GMC family | 1.94 | 0.3 | ||
| MT0560 | Rv0536 | galE3 | NAD-dependent epimerase/dehydratase family protein | 1.80 | 0.4 | |
| MT0777 | Rv0753c | mmsA | Methylmalonate-semialdehyde dehydrogenase | 2.24 | 1.2 | |
| MT0888 | Rv0865 | mog | Probable molybdopterin biosynthesis protein | 1.81 | 0.1 | |
| MT0916 | Rv0892 | Monooxygenase, flavin-binding family | 1.84 | 0.6 | ||
| MT1128 | Rv1096 | Polysaccharide deacetylase, putative | 1.83 | 0.3 | ||
| MT1295 | Rv1256c | cyp130 | Probable cytochrome P450 | 1.79 | 0.5 | |
| MT1339 | Rv1300 | papM (hemK) | N-methylase | 1.91 | 0.3 | |
| MT1368 | Rv1326c | glgB | 1,4-α-Glucan branching enzyme | 1.89 | 0.5 | |
| MT1369 | Rv1327c | glgE | Glucanase | 1.88 | 0.4 | |
| MT1424 | Rv1380 | pyrB | Probable aspartate carbamoyltransferase | 1.82 | 0.6 | |
| MT1511 | Rv1464 | csd | Cysteine desulfurase | 1.89 | 0.4 | |
| MT1512 | Rv1465 | Nitrogen fixation protein NifU-related protein | 1.77 | 0.3 | ||
| MT1636 | Rv1600 | hisC1 | Probable histidinol-phosphate aminotransferase | 1.77 | 0.3 | |
| MT1658 | Rv1622c | cydB | Membrane cytochrome D ubiquinol oxidase subunit II | 1.85 | 0.3 | |
| MT1659 | Rv1623c | cydA | Membrane cytochrome D ubiquinol oxidase subunit I | 1.98 | 0.3 | |
| MT1667 | Rv1631 | coaE | Probable dephospho-CoA kinase | 1.80 | 0.5 | |
| MT1690 | Rv1652 | argC | Probable N-acetyl-gamma-glutamyl-phosphate reductase | 1.80 | 0.9 | |
| MT1767 | Rv1726 | Oxidoreductase, FAD binding | 1.91 | 0.7 | ||
| MT1902 | Rv1854c | ndh-2 | NADH dehydrogenase | 1.76 | 0.6 | |
| MT1987 | Rv1937 | Ferredoxin reductase, electron transfer component, putative | 1.78 | 0.7 | ||
| MT2103 | Rv2043c | pncA | Pyrazinamidase/nicotinamidas | 1.90 | 0.5 | |
| MT2274 | Rv2217 | lipB | Lipoate biosynthesis protein B | 1.91 | 0.7 | |
| MT2336 | Rv2276 | cyp121 | P450 heme-thiolate protein | 2.25 | 1.0 | |
| MT2511 | Rv2436 | rbsK | Ribokinase | 1.84 | 0.9 | |
| MT2572 | Rv2497c | bkdA (pdhA) | Probable branched-chain keto acid dehydrogenase E1 component, alpha subunit | 1.78 | 0.4 | |
| MT2797 | Rv2725c | hflX | GTP-binding protein | 1.99 | 0.3 | |
| MT2965 | Rv2897c | Mg chelatase | 4.77 | 0.7 | ||
| MT2967 | Rv2899c | fdhD | Formate dehydrogenase accessory protein | 1.99 | 0.4 | |
| MT2968 | Rv2900c | fdhF | Possible formate dehydrogenase H | 2.09 | 0.2 | |
| MT3065 | Rv2987c | leuD | 3-Isopropylmalate dehydratase small subunit | 1.86 | 0.2 | |
| MT3066 | Rv2988c | leuC | 3-Isopropylmalate dehydratase large subunit | 2.10 | 0.4 | |
| MT3224 | Rv3137 | Inositol monophosphatase family protein | 1.95 | 0.7 | ||
| MT3283 | Rv3192 | Putative luciferase | 2.52 | 1.2 | ||
| MT3514 | Rv3406 | Putative dioxygenase | 2.46 | 0.7 | ||
| MT3687 | Rv3581c | ispF | Probable 2C-methyl-d-erythritol 2,4-cyclodiphosphate synthase | 1.82 | 0.2 | |
| MT3813 | Rv3710 | leuA | 2-Isopropylmalate synthase | 1.77 | 0.4 | |
| MT3950 | Rv3842c | glpQ1 | Glycerophosphoryl diester phosphodiesterase | 2.43 | 0.5 | |
| Metabolism | MT0590 | Rv0564c | gpsA (gpdA1) | Probable glycerol-3-phosphate dehydrogenase | 2.06 | 0.2 |
| MT0776 | Rv0752c | fadE9 | Acyl-CoA dehydrogenase | 2.43 | 1.1 | |
| MT1162 | Rv1130 | prpD | Possible methylcitrate dehydratase | 3.62 | 0.3 | |
| MT1163 | Rv1131 | gltA1 (prpC) | Probable methylcitrate synthase | 3.70 | 0.3 | |
| MT1518 | Rv1472 | echA12 | Possible enoyl-CoA hydratase | 1.80 | 0.6 | |
| MT1983 | Rv1933c | fadE18 | Acyl-CoA dehydrogenase, putative | 1.91 | 0.4 | |
| MT1984 | Rv1934c | fadE17 | Acyl-CoA dehydrogenase | 3.09 | 1.5 | |
| MT2243 | Rv2188c | pimB | Mannosyltransferase | 1.96 | 0.2 | |
| MT2599 | Rv2523c | acpS | Holo-[acyl-carrier protein] synthase | 1.90 | 0.3 | |
| MT2730 | Rv2953 | Enoyl reductase, may be involved in phenolpthiocerol and phthiocerol dimycocerosate (dim) biosynthesis | 2.18 | 1.4 | ||
| MT3081 | Rv3001c | ilvC | Ketol-acid reductoisomerase | 1.98 | 0.3 | |
| MT3082 | Rv3002c | ilvH (ilvN) | Acetolactate synthase, small unit | 2.31 | 0.4 | |
| MT3083 | Rv3003c | ilvB | Acetolactate synthase, large unit | 2.37 | 0.5 | |
| PE/PPE | MT0369 | Rv0354c | ppe7 | PPE family protein | 1.86 | 0.4 |
| MT0778 | Rv0754 | PE_PGRS11 | 2.41 | 0.6 | ||
| MT2505 | Rv2430c | ppe41 | PPE41 | 1.77 | 0.9 | |
| MT3637 | Rv3533c | PPE62 | 2.10 | 0.0 | ||
| MT3701 | Rv3595c | pe_pgrs59 | PE/PGRS protein | 1.85 | 0.2 | |
| Regulatory proteins | MT0222 | Rv0212c | nadR | AsnC family transcriptional regulator | 4.02 | 0.6 |
| MT0368 | Rv0353 | hspR | Probable heat shock protein transcriptional repressor | 2.22 | 0.5 | |
| MT0481 | Rv0465c | Transcriptional regulator | 2.24 | 0.9 | ||
| MT0514 | Rv0494 | Transcriptional regulator, GntR family | 1.97 | 0.1 | ||
| MT0605 | Rv0576 | Transcriptional regulator, ArsR family | 2.57 | 0.1 | ||
| MT0849 | Rv0827 | kmtR | Transcriptional regulator, ArsR family | 1.93 | 0.7 | |
| MT1161 | Rv1129c | Probable transcriptional regulator protein | 4.38 | 0.8 | ||
| MT1440 | Rv1395 | Transcriptional regulator, AraC family | 1.86 | 0.1 | ||
| MT1520 | Rv1473A | Possible transcriptional regulatory protein | 2.22 | 1.1 | ||
| MT1960 | Rv1909c | furA | Ferric uptake regulation protein | 3.86 | 1.3 | |
| MT2039 | Rv1985c | Transcription regulator | 2.60 | 0.1 | ||
| MT2073 | Rv2017 | Transcriptional regulatory protein | 2.75 | 1.9 | ||
| MT2386 | Rv2324 | Transcriptional regulator, AsnC family | 1.89 | 1.1 | ||
| MT2980 | Rv2912c | Probable transcriptional regulatory protein, TetR family | 2.19 | 0.4 | ||
| MT3290.1 | Rv3197A | whiB7 | Probable transcriptional regulatory protein, WhiB-like | 3.56 | 0.6 | |
| Virulence, detoxification and adaptation | MT0134 | Rv0126 | treS | Trehalose synthase, alpha-amylase family protein | 1.92 | 0.4 |
| MT0254 | Rv0240 | vapC24 | Possible toxin | 2.04 | 0.5 | |
| MT0265 | Rv0251c | hsp | Heat shock protein, HSP20 family | 3.06 | 2.4 | |
| MT0289 | Rv0277c | vapC25 | Possible toxin | 2.25 | 0.6 | |
| MT0365 | Rv0350 | dnaK | Probable chaperone protein | 3.00 | 1.0 | |
| MT0366 | Rv0351 | grpE | Probable GrpE protein | 3.36 | 1.5 | |
| MT0367 | Rv0352 | dnaJ1 | Chaperone protein | 2.70 | 1.0 | |
| MT0397 | Rv0384c | clpB | Endopeptidase ATP binding protein chain B, heat shock protein F84.1 | 1.77 | 0.4 | |
| MT0456 | Rv0440 | groEL2 | 60-kDa chaperonin 2 | 2.78 | 1.3 | |
| MT0574 | Rv0549c | vapC3 | Possible toxin | 1.85 | 0.6 | |
| MT0575 | Rv0550c | vapB3 | Possible antitoxin | 2.23 | 0.6 | |
| MT0618 | Rv0589 | mce2A | Mce family protein | 1.76 | 0.2 | |
| MT0621 | Rv0591 | mce2C | Mce family protein | 2.56 | 0.1 | |
| MT0685 | Rv0656c | vapC6 | Possible toxin | 2.65 | 0.3 | |
| MT0693 | Rv0665 | vapC8 | Possible toxin | 2.02 | 2.7 | |
| MT1296 | Rv1257c | Probable oxidoreductase | 2.27 | 0.2 | ||
| MT1959 | Rv1908c | katG | Catalase-peroxidase | 2.34 | 0.2 | |
| MT1961 | Rv1910c | Hypothetical exported protein | 2.09 | 0.1 | ||
| MT2004 | Rv1955 | higB | Possible toxin | 2.72 | 0.9 | |
| MT2005 | Rv1956 | higA | Possible antitoxin | 2.05 | 0.6 | |
| MT2018 | Rv1966 | mce3A | Mce family protein | 2.05 | 0.4 | |
| MT2489 | Rv2416c | eis | Enhanced intracellular survival protein | 3.73 | 0.6 | |
| MT2503 | Rv2428 | ahpC | Alkyl hydroperoxide reductase C protein | 2.19 | 0.6 | |
| MT2504 | Rv2429 | ahpD | Alkyl hydroperoxide reductase D protein | 2.22 | 1.1 | |
| MT2941 | NA | Prevent-host-death family protein | 3.00 | 3.8 | ||
| MT3526 | Rv3417c | groEL1 | 60-kDa chaperonin 1 | 3.18 | 2.3 | |
| MT3527 | Rv3418c | groES | 10-kDa chaperonin | 2.79 | 1.6 | |
| MT3771 | Rv3670 | ephE | Epoxide hydrolase | 2.11 | 0.8 | |
| MT3949** | Rv3841 | bfrB | Bacterioferritin | 2.21 | 0.7 | |
Conserved hypothetical, unknown, and pseudogenes are listed in Table S1 in the supplemental material.
**, DosR-dependent genes.
TABLE 3.
Genes repressed in iLivE M. tuberculosis
| Functional categorya | Designation in strain: |
Gene | Description | Fold change | SD | |
|---|---|---|---|---|---|---|
| CDC1551b | H37Rv | |||||
| Cell wall and cell processes | MT0046 | Rv0040c | mtc28 | Secreted proline-rich protein | 0.53 | 0.3 |
| MT0182 | Rv0173 | mce1E (lprK) | Mce family protein | 0.48 | 0.2 | |
| MT0356 | Rv0341 | iniB | Isoniazid-inducible gene protein | 0.47 | 0.2 | |
| MT0911 | Rv0888 | Probable exported protein | 0.54 | 0.1 | ||
| MT1235 | Rv1197 | esxK | ESAT-6-like protein | 0.34 | 0.1 | |
| MT1236 | Rv1198 | esxL | ESAT-6-like protein | 0.42 | 0.1 | |
| MT1729 | Rv1690 | lprJ | Probable lipoprotein | 0.43 | 0.1 | |
| MT1779 | Rv1737c | narK2 | Nitrite extrusion protein, MFS | 0.32 | 0.1 | |
| MT1932 | Rv1884c | rpfC | Probable resuscitation-promoting factor | 0.55 | 0.3 | |
| MT2411 | Rv2346c | esxQ | ESAT-6-like protein | 0.55 | 0.2 | |
| MT2412 | Rv2347c | esxP | ESAT-6-like protein | 0.53 | 0.2 | |
| MT2420 | Rv2346c | esxO | ESAT-6-like protein | 0.47 | 0.2 | |
| MT2458 | Rv2389c | rpfD | Resuscitation-promoting factor | 0.51 | 0.2 | |
| MT3988 | Rv3874 | esxB (cfp10) | 10-kDa ESAT-6-like protein | 0.56 | 0.2 | |
| MT3989 | Rv3875 | esxA (esat-6) | ESAT-6-like protein | 0.52 | 0.2 | |
| Information pathways | MT2669 | Rv2592c | ruvB | Holliday junction ATP-dependent DNA helicase | 0.45 | 0.3 |
| MT3347 | Rv3249c | Transcriptional regulator, TetR family | 0.41 | 0.3 | ||
| Intermediary metabolism and respiration | MT0037 | Rv0032 | Aminotransferase | 0.47 | 0.1 | |
| MT0266 | Rv0252 | nirB | Probable nitrite reductase [NAD(P)H], large subunit | 0.54 | 0.2 | |
| MT0738 | Rv0711 | atsA | Possible arylsulfatase | 0.51 | 0.3 | |
| MT1449 | Rv1405c | Methyltransferase | 0.43 | 0.2 | ||
| MT1603 | Rv1552 | frdA | Probable fumarate reductase | 0.51 | 0.0 | |
| MT1604 | Rv1553 | frdB | Fumarate reductase, iron-sulfur subunit | 0.42 | 0.0 | |
| MT1606 | Rv1555 | frdD | Fumarate reductase membrane anchor subunit | 0.42 | 0.1 | |
| MT1778** | Rv1736c | narX | Probable nitrate reductase | 0.51 | 0.9 | |
| MT1904 | Rv1856c | Possible oxidoreductase | 0.54 | 0.2 | ||
| MT2063** | Rv2007c | fdxA | Ferredoxin | 0.47 | 0.6 | |
| MT2088** | Rv2029c | pfkB | 6-Phosphofructokinase | 0.36 | 0.5 | |
| MT2401 | Rv2338c | moeW | Possible molybdopterin biosynthesis protein | 0.53 | 0.2 | |
| MT3194 | Rv3111 | moaC | Molybdenum cofactor biosynthesis protein C 2 | 0.34 | 0.2 | |
| MT3423 | Rv3322c | Possible methyltransferase | 0.37 | 0.1 | ||
| MT3424 | Rv3323c | moaDE | Molybdopterin cofactor biosynthesis protein D/E | 0.37 | 0.2 | |
| MT3426 | NA | moaB3 | Probable pterin-4-alpha-carbinolamine dehydratase | 0.28 | 0.2 | |
| MT3427 | NA | moaA3 | Molybdenum cofactor biosynthesis protein A 3 | 0.27 | 0.2 | |
| MT3849 | Rv3741c | Possible oxidoreductase | 0.57 | 0.3 | ||
| MT3850 | Rv3742c | Possible oxidoreductase | 0.52 | 0.3 | ||
| MT3969 | Rv3854c | ethA | Monooxygenase | 0.53 | 0.3 | |
| Metabolism | MT0038 | Rv0033 | acpA (acpP) | Acyl carrier protein | 0.48 | 0.3 |
| MT0175 | Rv0166 | fadD5 | Probable fatty-acid-CoA ligase | 0.55 | 0.1 | |
| MT0258 | Rv1467c | fadE15 | Acyl-CoA dehydrogenase | 0.45 | 0.2 | |
| MT1702 | Rv1662 | pks8 | Probable polyketide synthase | 0.55 | 0.2 | |
| MT2559 | Rv2485c | lipQ | Carboxylesterase family protein | 0.50 | 0.3 | |
| MT2667 | Rv2590 | fadD9 | Fatty acid-CoA ligase | 0.50 | 0.3 | |
| MT3216** | Rv3130c | tgs1 | Triacylglycerol synthase | 0.24 | 0.1 | |
| MT3326 | Rv3229c | desA3 | Linoleoyl-CoA desaturase, putative | 0.55 | 0.2 | |
| MT3348 | Rv3250c | rubB | Rubredoxin | 0.41 | 0.3 | |
| MT3349 | Rv3251c | rubA | Rubredoxin | 0.47 | 0.3 | |
| MT3350 | Rv3252c | alkB | Transmembrane alkane 1-monooxygenase | 0.44 | 0.3 | |
| MT3591 | Rv3847c | lipF | Probable esterase/lipase | 0.53 | 0.2 | |
| MT3933 | Rv3825c | pks2 | Mycocerosic acid synthase | 0.51 | 0.2 | |
| PE/PPE | MT1233 | Rv1195 | pe13 | PE family protein PE13 | 0.29 | 0.1 |
| MT1234 | Rv1196 | ppe18 | PPE family protein PPE18 | 0.33 | 0.0 | |
| MT1745 | Rv1705c | ppe22 | PPE family protein PPE22 | 0.49 | 0.2 | |
| MT1746 | Rv1706c | ppe23 | PPE family protein PPE23 | 0.47 | 0.1 | |
| MT2166 | Rv2107 | pe22 | PE family protein PE22 | 0.42 | 0.1 | |
| MT3427.1 | Rv3347 | ppe55 | PPE family protein PPE55 | 0.27 | 0.2 | |
| MT3854 | Rv3746c | pe34 | PE family protein | 0.42 | 0.2 | |
| Regulatory proteins | MT3230 | Rv3143 | Probable response regulator | 0.53 | 0.2 | |
| MT3870 | Rv3765c | tcrX | Probable two-component transcriptional regulatory protein | 0.51 | 0.2 | |
| Virulence, detoxification, and adaptation | MT0052 | Rv0046c | 1-l-myo-inositol-1-phosphate synthase | 0.49 | 0.3 | |
| MT0176 | Rv0167 | yrbE1A | Conserved integral membrane protein | 0.54 | 0.2 | |
| MT0178 | Rv0169 | mce1A | Mce family protein | 0.54 | 0.1 | |
| MT0179 | Rv0170 | mce1B | Mce family protein | 0.54 | 0.1 | |
| MT0180 | Rv0171 | mce1C | Mce family protein | 0.52 | 0.2 | |
| MT0181 | Rv0172 | mce1D | Mce family protein | 0.50 | 0.2 | |
| MT0183 | Rv0174 | mce1F | Mce family protein | 0.46 | 0.2 | |
| MT2087 | NA | Universal stress protein family protein | 0.35 | 1.0 | ||
| MT2090** | Rv2031c | hspX | Alpha crystallin, 14-kDa antigen | 0.25 | 0.0 | |
| MT2698** | Rv2623 | TB31.7 | Universal stress protein family | 0.43 | 0.5 | |
| MT3598 | Rv3494c | mce4F | Mce family protein | 0.50 | 0.1 | |
Conserved hypothetical and unknown genes are listed in Table S1 in the supplemental material.
**, DosR-dependent genes.
Validation of RNA-seq data.
To validate the gene expression changes observed by RNA-seq analysis of the iLivE M. tuberculosis transcriptome, we selected a number of genes that were either highly modulated or of functional relevance and performed qRT-PCR on M. tuberculosis exposed to the same iLivE conditions (20 μg/ml for 48 h). The icl gene was used as a negative control, given its nonsignificant regulation under iLivE (fold change, 1.24; P value of 0.16). A comparable trend in fold changes obtained with both qRT-PCR and RNA-seq was observed for all selected genes (Fig. 3), thereby validating the iLivE M. tuberculosis transcriptome profile generated by RNA sequencing.
FIG 3.
Validation by quantitative real-time PCR analysis of selected iLivE M. tuberculosis genes. Exponential M. tuberculosis culture was coincubated with SF under inhibitory conditions (20 μg/ml SF, 48 h) or with buffer only (control). Total RNA was extracted and real-time PCR was performed using specific primers listed in Table S7 in the supplemental material. For each gene, the average from technical triplicates was calculated and expressed as fold change compared to the gene expression level measured in the control. Fold changes (black bar) were compared to those obtained by RNA-seq (open bar). Data shown are the means ± standard deviations (SD) from triplicates. *, DosR-dependent genes.
iLivE M. tuberculosis expresses a unique transcriptome. (i) General stress responses.
The hostile nature of the lysosomal content undoubtedly imposes stress on M. tuberculosis. Under iLivE conditions, we observed the induction of several markers of general stress response, which includes a number of chaperone protein-encoding genes (groEL1, groEL2, groES, grpE, dnaJ1, dnaK, and hspR) that are involved in the folding and translocation of polypeptides and DNA repair (dinF and dinX) (37) (Table 2). These genes have also been reported to be upregulated during BMMO infections (10, 11, 30) and in lungs from TB patients (38), indicative of a stressful environment during infection partly contributed by the lysosomal contents. Furthermore, upregulation of the Clp proteases, particularly clpP1, clpP2, and clpC2, suggests prevalent protein degradation in iLivE M. tuberculosis. This observation may indicate a homeostatic response to prevent toxic accumulation of misfolded and aggregated proteins generated under stressed conditions (39). ClpP1 and ClpP2 have been reported previously to play an important role in M. tuberculosis pathogenesis and represent potential drug targets (40). More recently, ClpP1 has been used in a novel target mechanism-based whole-cell screening assay and was used to successfully identify bortezomib as a new lead compound for tuberculosis therapy (41).
(ii) Metabolic reprogramming.
Metabolic adaptations to host fatty acids and cholesterol by intracellular M. tuberculosis have been reported in transcriptomics studies from macrophage and mouse infections (11, 13, 15, 30, 42). Upregulation of prpC and prpD was observed in iLivE M. tuberculosis (Table 2). These genes encode key enzymes of the methylcitrate cycle and help M. tuberculosis detoxify propionyl-coenzyme A (CoA), a product from fatty acid catabolism during intracellular survival (43). However, induction of isocitrate lyase (icl) was not observed in iLivE M. tuberculosis (Fig. 2C), a key bifunctional enzyme that is induced simultaneously with prpC and prpD in intracellular M. tuberculosis (30) to utilize fatty acids via the glyoxylate shunt and methylcitrate cycle (44). The sole induction of icl has been reported in the presence of palmitic acid (11), suggesting that icl expression is directly modulated by the presence of fatty acids, which may explain its lack of induction in the fatty acid-free iLivE model. Reinforcing the notion that fatty acids are the preferred carbon source of intracellular M. tuberculosis, we also found a number of iLivE M. tuberculosis genes predicted with enzymatic functions involved in biochemical activation and β-oxidation of fatty acids. These include acyl-CoA dehydrogenase (fadE9, fadE15, fadE17, and fadE18), fatty acid-CoA ligase (fadD5 and fadD9), enoyl-CoA hydratase (echA12), and lipases (lipF and lipQ) (Table 2). Most of these genes have been reported previously to be modulated under various in vitro and ex/in vivo conditions (10, 11, 21, 24, 30) with the exception of fadE17 and fadE18, which seem to be specifically upregulated in iLivE M. tuberculosis. Thus, it appears that the lysosomal content represents an environmental signal for the bacterium to upregulate genes involved in fatty acid beta-oxidation, perhaps in anticipation of the next round of infection upon lysis of the host cell.
(iii) Cell wall remodeling.
With a lipid-rich cell wall envelope, mycobacterial cell wall remodeling is also tightly associated with its lipid metabolism (45) and can be a possible adaptive mechanism of M. tuberculosis when coping with a constantly changing host environment (46). In line with the apparent nonreplicative state of iLivE M. tuberculosis, we detected a downregulation of pks2 and desA3 (Table 3), which are essential genes for the biosynthesis of main cell wall components of mycobacteria (47). On the contrary, pks2 expression was found to be induced upon phagosome acidification (12, 20), an environmental cue that is not represented in the LivE model, where pH is maintained at 6.8. Interestingly, desA3 was previously proposed to be involved in regulating the membrane fluidity necessary for physiological function (48). Repressed expression of desA3 in iLivE M. tuberculosis suggests reduced cell membrane fluidity, leading to limited barrier permeability that could limit drugs from gaining access to their bacterial targets and consequently conferring phenotypic drug resistance.
Furthermore, consistent with observations in the short-term macrophage infection model (10, 11), the mce1 operon (yrbE1A and mce1A-F) was downregulated in iLivE M. tuberculosis (Table 3). Deletion of this operon has been associated with (i) accumulation of free mycolic acids in the mycobacterial cell wall (49), (ii) a hypervirulent infection profile in mice with an impaired ability to trigger a proinflammatory response (50), and (iii) in vitro phenotypic drug tolerance (51). Therefore, mce1 downregulation in iLivE M. tuberculosis may lead to changes in its cell wall mycolic acid composition, which could contribute to altered drug susceptibility of M. tuberculosis.
Finally, in iLivE M. tuberculosis we measured the upregulation of pstS1, which encodes a mycobacterial cell wall adhesin that has been demonstrated to promote phagocytosis of mycobacteria via binding to the mannose receptor (52). This implies an increased ability of bacilli to infect neighboring host cells when released from apoptotic macrophages.
Downregulation of dosR-dependent genes.
Induction of the transcriptional factor DosR, involved in activation of the dormancy program in M. tuberculosis (53), has been associated with hypoxic conditions within TB granulomas (54) and reactive nitrogen intermediates produced by activated macrophages (55). The Dos regulon, which comprises ∼49 genes regulated by the DosR-S/T two-component system, has been described to respond to a variety of signals and stresses, including low-oxygen tension, S-nitrosoglutathione (GSNO), ethanol, and carbon monoxide (56). As expected, dosR expression was minimally modulated in the iLivE model (fold change, −1.3; P value of 0.25), since this model does not incorporate any of the above-mentioned stimuli. However, and interestingly, we observed significant modulation of 17 dosR-dependent genes in iLivE M. tuberculosis, among which 15 were downregulated (Table 3). The majority of these repressed dosR-dependent genes were also found downregulated in the long-term BMMO infection model (at day 8) (Table 3), and this was attributed to the sudden loss of cue(s) driving the DosR response at the later stage during macrophage infection (30), where the bacterium presumably has transited to a persistent state. Consistent with this, transient and early induction of dosR-dependent genes during the first 4 to 8 h, followed by a gradual decline to baseline within 24 h, was also reported in a defined hypoxia model (22).
Thus, these observations suggest that the gene expression signature of iLivE M. tuberculosis partially overlaps that profiled at the late stage of macrophage infection, which further supports the idea that mycobacteria at this stage of infection are exposed to a lysosomal environment. The data also suggest that genes previously identified as part of the Dos regulon also are regulated independently of DosR.
ESAT-6 and PE-PPE family of proteins.
Genes encoding a cluster of ESAT-6-like proteins (esat-6, cfp10, esxQ, esxP, esxK, and esxQ) were notably downregulated in iLivE M. tuberculosis (Table 3). Consistent with this, the expression of esxQ, esxP, esxK, and esxQ genes was also found downregulated in resting and activated macrophages (10). ESAT-6 and CFP-10 have been identified as both virulence factors and protective antigens (57, 58). In contrast, the PE/PPE genes that were modulated in iLivE M. tuberculosis exhibited various expression trends in previous transcriptomics studies of macrophage (10, 11), mouse (42) infection models, and under in vitro stresses (21, 24) (see Table S1 in the supplemental material). Our analysis singled out PPE41 based on its consistent induced profile observed in M. tuberculosis from infected BMMO (11) and human macrophages (13), mouse lungs (42), and iLivE M. tuberculosis (see Table S1). The PE25/PPE41 protein complex was shown to induce dendritic cell activation and drive Th2-biased immune responses (59), whereas Th1-biased immune responses have long been known to be protective against tuberculosis (60). Downregulation of ESAT-6-like proteins and upregulation of PPE41 in iLivE M. tuberculosis indicates that the lysosomal environment contributes to subversion of the host immune system by M. tuberculosis toward nonprotective immune responses.
Fighting oxidative stresses.
Exposure to inhibitory SF conditions upregulated several M. tuberculosis genes (furA, katG, ahpC, and ahpD) (Table 2) that are known to be instrumental in combating oxidative stresses mediated by reactive oxygen species (ROS) and reactive nitrogen intermediates (RNI) (61–63), which are abundantly produced in an activated macrophage. While furA and katG have been reported to be upregulated under in vitro oxidative stresses (11), their regulation under nonoxidative conditions, such as nutrient starvation (24), gradual hypoxia (21), static growth (26), and iLivE conditions (see Table S1 in the supplemental material), suggests that these genes respond to multiple environmental stimuli. Alternatively, intrinsic ROS production by the mycobacterial cell itself could also be responsible for inducing these genes. This hypothesis is supported by the upregulation of Rv1464 and Rv1465 ORFs (Table 2) from the SUF operon that encodes the alternative mycobacterial iron-sulfur cluster machinery (64). Furthermore, high production of ROS was detected in nonreplicating nutrient-starved mycobacteria and was attributed to cytochrome P450 (CYP)-based metabolism of ketone bodies generated from triacylglycerol (TAG) stores during nutrient starvation (our unpublished observations). Coincidentally, cyp121 and cyp130 were found to be upregulated in iLivE M. tuberculosis (Table 2). Thus, intracellular production of ROS through CYP activity upon lysosomal exposure represents an interesting possibility which remains to be further investigated.
Respiratory status.
The respiratory status of M. tuberculosis is dependent on the microenvironment it encounters, such as the oxygen tension and availability of various carbon and nitrogen sources to act as terminal electron acceptors (65). Induction of bd-type terminal oxidase-encoding genes (cydA and cydB) and genes involved in nitrate respiration (narK2) were detected in iLivE M. tuberculosis (Table 2). This is consistent with a transitional respiratory state previously described for intracellular M. tuberculosis upon NO production following immune cell activation (65). In contrast, the fumarate reductase gene cluster frdABD was notably repressed in iLivE M. tuberculosis, which is consistent with the aerobic setup of the LivE model (Table 3). Induced expression of fumarate reductase was observed in activated BMMOs (11) in an NO-dependent manner and in hypoxic lung lesions from tuberculosis patients (38), and it was associated with anaerobic persistence (66). Interestingly, modulation of gene clusters encoding F0F1 ATP synthase (atpA-H) and NADH dehydrogenase 1 (nuoA-N), which are involved in aerobic respiration, was not observed in iLivE M. tuberculosis. Along with ribosomal proteins (rps), these regulons were also notably repressed during regulated slow growth (67) and in models of persistence where mycobacterial replication arrest is induced by reduced oxygen and/or nutrient availability (68, 69). As mentioned earlier, the true replicative/nonreplicative status of mycobacteria during SF exposure remains to be further characterized.
Meta-analysis with transcriptomes of intracellular M. tuberculosis from macrophage infection models.
The experimental setup of the LivE model was designed to (partially) mimic exposure of M. tuberculosis to the lysosomal content upon phagosome/autophagosome maturation during macrophage infection. Thus, we subjected the iLivE M. tuberculosis transcriptome profile to a comparative analysis with selected key transcriptome profiling studies of intracellular M. tuberculosis during infection in BMMO and human macrophages (THP-1). With the caveat that each study used different infection conditions, different macrophage types, and different M. tuberculosis strains, our analysis revealed that 193 out of 370 iLivE genes overlapped genes from these macrophage infection studies, while 177 did not overlap any of the studies considered (Fig. 4; see also Tables S2 and S3 in the supplemental material). As expected, the largest overlap was observed with the BMMO infection models (168 out of 193) (Fig. 4; see also Table S3), suggesting a partial recapitulation of intramacrophage lysosomal exposure in the LivE model. However, it is believed that in some of these BMMO infection models, mycobacteria reside primarily in a phagosomal environment due to phagosome maturation arrest. Therefore, it is possible that some of these overlapping genes are also modulated by stimuli present in the phagosome prior to lysosomal fusion. To further refine the lysosome-specific gene responses triggered under iLivE conditions, we excluded iLivE genes that were modulated during resting macrophage infection reported by Homolka et al. (10) and Schnappinger et al. (11), where mycobacteria are believed to reside mainly within phagosomes. Furthermore, since the process of infection is not synchronized, the bacilli retrieved from the infected macrophage population at each time point consist of a heterogeneous population of bacteria which have been exposed to various intracellular environments, ranging from early endosome to lysosomal compartment. Therefore, the mycobacterial transcriptional response measured is likely to be heterogeneous, reflecting the responses to various intracellular microenvironments. On the contrary, in the iLivE model, the mycobacterial transcriptional response is expected to be more homogeneous. Therefore, we postulated that fold changes measured in iLivE M. tuberculosis are likely to be greater in magnitude than those measured in activated macrophages. Thus, a more stringent cutoff value was arbitrarily implemented for each gene by either dividing (upregulation) or multiplying (downregulation) their expression fold change by 1.5, leading to selection of 41 iLivE M. tuberculosis genes (see Table S6). Among these 41 genes, most of the general stress response markers (groEL1, groES, and dnaJ1) were present, further supporting the increased level of stress experienced by bacilli exposed to the lysosomal environment. Interestingly, we found several antibiotic resistance-related genes, namely, eis (Rv2416c), ermMT (Rv1988), and tap (Rv1258c), with remarkably increased expression under iLivE conditions. Although the mechanisms by which they induce antibiotic resistance are different (70–73), expression of all of these genes is under the control of the transcriptional factor WhiB7 (70), one of the earliest and most highly induced transcriptional regulators in M. tuberculosis during BMMO infection (12), and in response to numerous stress conditions (70, 74, 75). Thus, these findings strongly support that the lysosomal content induces a WhiB7-mediated phenotypic drug resistance in M. tuberculosis.
FIG 4.
Venn diagram comparing transcriptional profiles of M. tuberculosis under iLivE conditions and during macrophage infections. THP-1, human macrophage model; iLivE, inhibitory condition of lysosomal in vitro exposure. The asterisk indicates that the gene is included if it is modulated in at least one of the three primary murine macrophage (BMMO) infection models.
In contrast, we observed a limited number of overlapping genes (36 out of 370) between THP-1 macrophage infection and the iLivE model (Fig. 4), which is likely attributable to inherent physiological differences between a human macrophage cell line and a primary mouse-derived macrophage (SF was prepared from activated BMMO). Nevertheless, genes involved in oxidative stresses (ahpC and ahpD) and fatty acid metabolism (prpC and prpD) were found in the 25 iLivE gene subset that were commonly regulated in both BMMO and THP-1 infection models (Fig. 4; see also Table S3 in the supplemental material), indicative of a comparable intramacrophage environment between human and mouse macrophages. From the subset of 11 iLivE genes that overlap THP-1 macrophages only (Fig. 4; see also Table S3), it is worth mentioning rpfD, the product of which has been implicated in resuscitating mycobacteria from dormancy (76).
Our meta-analysis also revealed a significant number of genes (177 iLivE M. tuberculosis genes) that did not overlap any of the macrophage infection models analyzed (see Table S2 in the supplemental material). These nonoverlapping genes could represent a subset of lysosome-inducible genes, which were previously not detected in macrophage models due to nonsynchronized infection conditions, where only a small percentage of mycobacteria harvested from the infected macrophages actually reside inside the lysosomal compartment at one time. Among these nonoverlapping genes, we found glgE, glgB, and treS to be significantly upregulated (see Table S2). These genes are involved in the α-1,4-glucan pathway implicated in detoxification of maltose-1-phosphate (M1P) to α-glucan (77). Interestingly, loss of glgE, which encodes a maltosyltransferase, was reported to impair the bacterium's replication ability in lungs and spleen from infected BALB/c mice (77). This defect was attributed to cell toxicity from the accumulation of the phosphosugar intermediate, thus revealing α-glucan synthesis as a potential target for antimicrobials (77). Upregulation of glgE in iLivE M. tuberculosis further indicates that targeting of GlgE and the α-glucan synthesis pathway represents a viable therapeutic approach to kill intraphagolysosomal mycobacteria.
Among the nonoverlapping iLivE gene subset, a number of mycobacterial toxin/antitoxin (TA) systems were also significantly induced. They include vapB3, vapC3, and vapC8 of the VapBC systems, which represents the largest family of bacterial TA systems (78). In addition, the HigBA1 TA system and three more VapC toxin-encoding genes, vapC24, vapC25, and vapC6, were upregulated (Table 2). Upregulation of TA systems in response to adverse conditions has been described in other bacterial species (79). Typically, bacterial TA systems are made of a toxin protein and a more labile antagonistic antitoxin, which can be a protein or noncoding RNA (80). Some of the mycobacterial VapC toxins have been shown to exert a bacteriostatic effect on mycobacterial growth through their RNase activity (81). While nutrient-starved M. tuberculosis cells and drug-tolerant M. tuberculosis persisters have been shown to express distinct sets of TA systems (25, 68), the VapBC TA systems could be exploited to induce toxicity in intramacrophage bacilli.
Finally, genes encoding cell wall-associated proteins, comprising transporters and lipoproteins, represent another prominent group of significantly induced nonoverlapping iLivE genes (see Table S2 in the supplemental material). The transporters were mainly associated with drug efflux (Rv0191, Rv1877, and Rv1456c) and uptake of sulfate, amino acids, and sugars (cysW, Rv1979c, and pitB), while the lipoproteins exhibited a plethora of predicted functions, including peptidoglycan cross-linking and remodeling (lpqK and Rv1922), degradation processes (Rv0671), host cell adhesion and invasion (Rv0593), and as solute binding proteins of ABC transport systems (sbp) (82). This could represent responses to (i) membrane stresses induced by membrane-perturbing agents (antimicrobial cationic peptides present in the lysosomal SF), (ii) intracellular toxic accumulation arising from SF-induced metabolic changes, and/or (iii) membrane remodeling for nutrient-scavenging activities or to facilitate escape into the cytosol.
Meta-analysis with M. tuberculosis transcriptomes from in vitro stress models.
Under in vitro conditions that are meant to reproduce some of the environmental cues encountered within a macrophage or a granuloma, exposure to gradual hypoxia (21), defined hypoxia (22), nutrient starvation (24), and antibiotics at incomplete sterilizing concentrations (25) induces M. tuberculosis persistence, characterized by nonreplication and phenotypic resistance to the TB drugs isoniazid and rifampin. The meta-analysis between iLivE M. tuberculosis transcriptome and M. tuberculosis transcriptomes profiled from these in vitro models indicates minimal overlap, which suggests that the physiological state of mycobacteria exposed to lysosomal SF significantly differs from that of hypoxic conditions, nutrient-starved conditions, or antibiotic-exposed mycobacteria (see Tables S4 and S5 in the supplemental material). Nevertheless, 115 iLivE genes overlapped at least one of these in vitro models of persistence, with only four genes (echA12, hsp, higA, and Rv2036) commonly regulated in all models considered, excluding the drug persisters model (Fig. 5; see also Table S5). The overlap between iLivE and at least one in vitro model here suggests that the same pathway is triggered by more than one environmental stimulus, as supported by a substantial number of regulatory proteins present in this gene subset (see Table S5), including multistress-induced transcriptional factor WhiB7. On the contrary, genes that are triggered by a specific environmental stimulus, such as acidic pH, induced aprABC locus (83), and hypoxia-driven dosR, were not found in the iLivE transcriptome due to the absence of these cues from the LivE model, highlighting a general limitation of in vitro models, where only one or a limited number of environmental stimuli are incorporated. On the other hand, a large number (255) of iLiveE genes did not overlap any of the in vitro models considered in this meta-analysis and include genes involved in virulence detoxification pathways, cell wall processes, and intermediary metabolism (see Table S4). It is plausible that these genes are modulated by lysosome-specific stimuli that are not represented in the other in vitro models. The iLivE model therefore allows investigation of M. tuberculosis responses specific to a relevant and defined intramacrophage microenvironment, where the identification and biochemical characterization of host molecules with antimycobactericidal activity can be undertaken (27).
FIG 5.
Venn diagram comparing transcriptional profiles of M. tuberculosis under iLivE conditions and various environmental stresses. EHR, defined hypoxic model; persisters, drug-tolerant persisters model; starvation, nutrient starvation model; NRP, gradual hypoxic model; iLivE, inhibitory condition of lysosomal in vitro exposure.
Validation of the LivE model.
To support the relevance of our transcriptomics approach to study M. tuberculosis responses to the lysosomal environment, we selected one gene candidate, Rv1258c, which was found highly expressed under iLivE conditions and functionally relevant as a membrane Tap-like efflux pump (84–86). A previous study reported that M. tuberculosis transposon mutants of Rv1258c displayed reduced viability at 96 h postmacrophage infection (71). However, the authors did not confirm the observed attenuated phenotype by complementation study. Here, we constructed an M. tuberculosis ΔRv1258c mutant by homologous recombination and its complemented strain, Ox-Rv1258c, by introducing the Rv1258c ORF under the expression of the constitutive hsp60 promoter into the ΔRv1258c mutant (Fig. 6A). Deletion of Rv1258c was confirmed by Southern blotting (Fig. 6B) and did not affect the transcription of the downstream gene Rv1257c, as assessed by real-time PCR (data not shown). Furthermore, qRT-PCR analysis showed that expression of Rv1258c was restored in the complemented strain Ox-Rv1258c with a 22-fold increase in transcriptional activity compared to the parental expression, likely due to the use of the strong mycobacterial hsp60 promoter (Fig. 6C). The in vitro fitness of both the mutant and complemented strains was similar to that of the parental strain (Fig. 6D).
FIG 6.
Construction of M. tuberculosis ΔRv1258c mutant and its complemented strain, Ox-Rv1258c. (A) Schematic organization of CDC1551 M. tuberculosis Rv1257c-1258c locus in parental strain (WT). The mutant was obtained by introducing a hyg cassette into the Rv1258c ORF by double homologous recombination and complemented by reintegrating Rv1258c under the M. tuberculosis hsp60 promoter (Ox-Rv1258c) into its genome. Southern blot probe and restriction enzyme sites are indicated (E, EcoRI; X, XmaI; probe, double-headed arrow). (B) Southern blot analysis. M, DIG-labeled molecular ladder. (C) Transcriptional activity of Rv1258c in WT, ΔRv1258c, and Ox-Rv1258c strains as determined by real-time PCR analysis. Data are expressed as averages ± SD from triplicates. (D) In vitro growth kinetics of WT and Δrv1258c strains and its complemented strain in 7H9 medium.
To confirm the role of Rv1258c during macrophage infection that was previously reported (71), resting and activated (100 U/ml IFN-γ and 50 ng/ml TNF) BMMOs were infected with WT, ΔRv1258c, and complemented strains. A marked reduction in bacterial load of approximately one log was observed with the ΔRv1258c mutant compared to the WT at 2 to 3 days postinfection in both resting and activated BMMOs, followed by a restoration to parental levels at the later phase of infection (Fig. 7A and B). Parental infection profiles were observed with the complemented strain Ox-Rv1258c. The attenuation pattern observed in both resting and activated macrophages suggests that Rv1258c plays a role in M. tuberculosis survival regardless of the macrophage activation status. Furthermore, the transient nature of the attenuated phenotype supports that Rv1258c plays a critical role during the initial phase of infection. To further demonstrate the relevance of our observations, infection profiles of the WT, ΔRv1258c, and Ox-Rv1258c strains were determined in human monocyte-like THP-1 cells that were activated by retinoic acid and vitamin D3 prior to infection (34). Similar to the BMMO infection profile, an initial drop in viability was observed with the ΔRv1258c strain at day 1 postinfection, but the bacilli's replicative ability was quickly restored from day 2 onwards, possibly reflecting a differential environmental pressure between murine and human macrophages (Fig. 7C). The complemented strain restored partially parental levels of growth.
FIG 7.
M. tuberculosis ΔRv1258c strain survival profile in resting and activated macrophages. Resting (A) and activated (50 ng/ml of TNF and 100 U/ml of IFN-γ) (B) murine bone marrow-derived macrophages (BMMOs) were infected with M. tuberculosis strains at an MOI of 2. (C) Retinoic acid and vitamin D (RAVD)-activated THP-1 macrophages were infected at an MOI of 5. After 45 min of incubation, the infected cells were washed with PBS to remove extracellular bacteria (day 0 postinfection). At the indicated time points, the infected cells were lysed and viable CFU were recovered on 7H11 agar and enumerated. *, P < 0.05. Data shown are means ± SD from quadruplicates and are representative of two independent experiments.
To further examine the role of Rv1258c when M. tuberculosis encounters the lysosomal microenvironment, susceptibility to lysosomal SF killing was compared among WT, ΔRv1258c mutant, and complemented strains. Incubation with 37 μg/ml SF for 24 h drastically reduced the viability of the ΔRv1258c mutant to 3.3%, while the WT strain retained 17% viability (Fig. 8A). In contrast, Ox-Rv1258c exhibited greater resistance to SF killing than the WT, with 52% viability, correlating with the greater expression level of Rv1258c measured in the Ox-Rv1258c strain (Fig. 8A). Our results support a role for Rv1258c in M. tuberculosis survival during exposure to the lysosomal content. Previous literature proposed that the Tap efflux pump activity helps mycobacteria cope with the hostile environment by pumping out host-derived antimicrobial molecules (84–86). To test whether the Tap efflux pump activity is involved in the M. tuberculosis response to SF killing, the Ox-Rv1258c strain was incubated with SF in the presence of the proton motive force inhibitor CCCP. Since the Tap efflux pump requires energy to function, the presence of CCCP is expected to abrogate its activity. We observed that resistance to SF killing was comparable to that obtained in the absence of CCCP (Fig. 8B). In contrast, the enhanced resistance to streptomycin of Ox-Rv1258c was abrogated in the presence of CCCP, confirming the functionality of the Tap efflux pump in this M. tuberculosis strain (Table 4). Together, these data suggest that the mechanism(s) by which Rv1258c-encoded Tap is involved in M. tuberculosis resistance to lysosomal killing does not rely on its efflux pump activity. Mycobacterial efflux pumps have been described as part of the coupled biosynthesis/export machinery for mycobacterial cell wall components, and their respective mutants displayed defective growth in macrophage and mouse models (87). One could speculate that the absence of membrane-associated Tap in M. tuberculosis results in altered cell wall composition that may render the bacterium more permeable/vulnerable to antimicrobial compounds and compromise its cell wall-associated virulence, thereby impairing survival of this pathogen within its mammalian cell host.
FIG 8.
Susceptibility of ΔRv1258c and Ox-Rv1258c strains to SF and human antimicrobial peptides. (A) Mid-log-phase cultures of M. tuberculosis parental (WT), ΔRv1258c, and Ox-Rv1258c (complemented) strains were coincubated with 37 μg/ml SF for 24 h. (B) Mid-log-phase culture of the ΔRv1258c strain was coincubated with 37 μg/ml SF for 24 h and 1 μg/ml CCCP. (C) WT, ΔRv1258c, and Ox-Rv1258c strains were coincubated with LL-37 at 30 μg/ml for 72 h. The bacterial suspensions were then plated on 7H11, and CFU were determined after incubation at 37°C for 3 weeks. Data are expressed as the percentage of viable CFU when incubated in respective assay buffers (SF buffer and 0.1% acetic acid for LL-37). Data are the means ± SD from triplicates and are representative of two independent experiments. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.
TABLE 4.
Susceptibility of M. tuberculosis WT and Ox-Rv1258c strains to streptomycin
| Treatment | MICa (μg/ml) |
|
|---|---|---|
| WT | Ox-Rv1258c strain | |
| Streptomycin | 0.3 | 3.7 |
| Streptomycin + CCCP | 0.5 | 0.8 |
The MIC of streptomycin was assayed over a range of 2-fold dilutions of the compound and in the presence or absence of 1 μg/ml CCCP. Data shown are representative of two independent experiments.
Previous work suggested that ubiquitin-derived peptides isolated from murine lysosomes were involved in SF killing activity against M. tuberculosis (27). Other studies have pointed at the antimycobactericidal activity of other lysosomal small molecules, including LL-37, a multifunctional peptide belonging to the cathelicidin family and one of the most abundant antimicrobial molecules produced in various human host cells for M. tuberculosis, including lung epithelial cells, neutrophils, and macrophages (88, 89). To extend our observations to the human context, we assessed the susceptibility of WT, ΔRv1258c, and Ox-Rv1258c strains to LL-37. Similar to observations made with the murine lysosomal SF, the ΔRv1258c mutant exhibited greater susceptibility than its parental counterpart when incubated with 30 μg/ml of LL-37 for 72 h. In contrast, susceptibility of the Ox-Rv1258c strain was greater than that of the control bacteria (buffer only), suggesting a growth advantage conferred by Rv1258c overexpression in the presence of the antimicrobial peptide (Fig. 8C). Thus, these findings strongly suggest that (i) lysosomal killing of M. tuberculosis is likely mediated by several antimicrobial molecules and (ii) the mechanism by which Rv1258c is involved in M. tuberculosis resistance to lysosomal killing is not specific to one particular antimicrobial molecule. This notion fits well with the hypothesis that the absence of membrane-associated Tap renders the cell wall more vulnerable to antimicrobial peptide attack.
Conclusions.
The lysosome is the major digestive organelle, with a critical role at the end of the endocytic pathway in mammalian cells (90). Its lumen contains more than 50 acid hydrolases, including proteases, peptidases, phosphatases, nucleases, glycosidases, sulfatases, and lipases, and it is maintained at an acidic pH necessary for the optimal activity of these enzymes to degrade all types of macromolecules (90). Under conditions that induce autophagy, short-length peptides such as ubiquitin-derived peptides and human cathelicidin h-CAP-18/LL-37 have also been shown to accumulate in lysosomal and autophagosomal structures, respectively, in a macrophage (27, 83). In addition to nitro-oxidative, nutrient-limiting, and hypoxic stresses within the macrophage, the antimycobacterial activity of these peptides becomes efficacious on intramacrophage M. tuberculosis when its phagosome either fuses with lysosomes or colocalizes with autophagosomes, which is also designated for degradation through lysosomal fusion (90). It is undeniable that the lysosomal environment is one of the intracellular microenvironments encountered by M. tuberculosis during macrophage infection.
This work increases our knowledge of the possible adaptive strategies devised by M. tuberculosis to resist the hostile lysosomal microenvironment. It complements previous transcriptome studies with the common aim of deciphering the mechanisms involved in the survival of M. tuberculosis inside its host macrophage.
Supplementary Material
ACKNOWLEDGMENTS
We gratefully acknowledge the Novartis Institute for Tropical Diseases for their generosity in providing the M. tuberculosis CDC1551 parental strain for our study. We also thank the Genome Technology Biology team at GIS for their sequencing efforts and assistance in computational analysis and for providing their valuable high-throughput computing resources.
This work was supported by the Singapore-MIT Alliance Infectious Disease Interdisciplinary Research Group (SMART-ID-IRG).
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication. We have no conflicts of interest to declare.
Footnotes
Supplemental material for this article may be found at http://dx.doi.org/10.1128/IAI.00072-16.
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