Heterologous Production of 1-Tuberculosinyladenosine in Mycobacterium kansasii Models Pathoevolution towards the Transcellular Lifestyle of Mycobacterium tuberculosis

We introduced the M. tuberculosis-specific gene pair Rv3377c-Rv3378c into the M. kansasii genome within an integrative plasmid containing hygromycin resistance to produce M. kansasii::Rv3377-78c (see Materials and Methods). As a control for subsequent experiments, an integrative “empty vector” (EV) was employed. After labeling with [¹⁴C]adenosine and lipid extraction using chloroform and methanol, radio-thin-layer chromatography (radio-TLC) was used to detect adenosine-linked lipids in M. kansasii::Rv3377-78c clones for comparison to M. tuberculosis strain H37Rv (Fig. 1a). Conventional molybdenum-based sprays followed by charring broadly detected all lipids as a loading control, suggesting a lack of broad lipid changes detectable at the TLC level after gene transfer (Fig. 1b). Whereas uncomplemented bacterial extracts showed material at the origin and one weak band by radio-TLC, Rv3377c-Rv3378c complementation generated at least five additional lipid species. Three of these novel lipids comigrated with compounds from M. tuberculosis strain H37Rv. Both results strongly suggested the successful genetic transfer of M. tuberculosis-associated adenosine-linked lipids to M. kansasii. Phosphomolybdic acid reagent (PMA) (5%) staining showed that similar amounts of total lipids were spotted for each M. kansasii sample (Fig. 1b).

High-performance liquid chromatography-mass spectrometry (HPLC-MS) was used to chemically identify the compounds produced by M. kansasii::Rv3377-78c, in which the expected retention of 1-TbAd (22.7 min) and N⁶-TbAd (5.8 min) were known (Fig. 1c to g). Whereas the M. kansasii::EV control did not release compounds that comigrated with either 1-TbAd or N⁶-TbAd, M. kansasii::Rv3377-78c produced high-intensity (6.7 × 10⁶ to 7.0 × 10⁶ counts) signals (m/z 540.354) matching the expected retention time and mass (m/z 540.3544) of the proton adducts of 1-TbAd. The extractions were performed at a range of pHs (4.5 to 7.4) since both the Dimroth reaction that generates N⁶-TbAd and the capture of lysosomotropic agents are sensitive to pH, as previously explained (18, 22, 23). Similar to results with M. tuberculosis in which 9% of the TbAd pool was released (22), we observed stronger (∼10-fold) signals in the pellet compared to the supernatant. However, there was no clear impact of altering preextraction pH for 2 h to the release nor relative abundance of 1-TbAd and N⁶-TbAd, and thus such effects, if they exist, did not occur under the tested conditions (Fig. 1c to g). Similar to patterns observed from M. tuberculosis, more 1-TbAd than N⁶-TbAd was recovered from M. kansasii::Rv3377-78c (Fig. 1f and g). In four tested cultures of M. kansasii::Rv3377-78c, 1-TbAd represented 0.125% ± 0.04% of the total lipid mass (versus 0.76% in M. tuberculosis) (data not shown).

To characterize any overt phenotypic effects of 1-TbAd production on M. kansasii, we assessed its influence on in vitro characteristics of the bacterial culture. M. kansasii::Rv3377-78c grew similarly to WT M. kansasii and M. kansasii::EV in Middlebrook 7H9 broth (Fig. 2a) and on 7H10 agar (Fig. 2b). Carotenoid pigments become integrated into bacterial cell membranes, maintain membrane fluidity, and provide support against external stressors (24). Since the production of 1-TbAd requires the same intermediate GGPP as that of the yellow pigment that M. kansasii produces when exposed to light, we then tested M. kansasii::Rv3377-78c’s ability to turn yellow after light exposure in order to rule out the possibility that a pigment-related phenomenon might affect our outcomes (Fig. 2b). M. kansasii::Rv3377-78c retained photochromogenic abilities by turning yellow after exposure to light at room temperature within the same time frame as M. kansasii::EV.

Congo red is an amphiphilic dye that binds to the mycobacterial cell membrane. When grown on Congo red-containing 7H10 plates, different mycobacteria retain the dye distinctly, and this feature has been associated with differences in the interactions of bacterial cells within the colony (25). In this study, M. tuberculosis absorbed the dye and became red, while M. kansasii colonies remained white on the plate. Since 1-TbAd is found on the cell surface and has an amphipathic character (17), we tested the effect of its production on intracolony bacterial interactions of M. kansasii::Rv3377-78c. Visual inspection of colonies (Fig. 2c) and absorbance at 488 nm (Fig. 2d) showed no difference in retention of Congo red between M. kansasii::EV and M. kansasii::Rv3377-78c.

The ability to survive in mildly acidic environments is a key feature of mycobacteria, both environmental and pathogenic (26, 27). It was recently shown that 1-TbAd production confers a growth advantage over a pH range (5.0 to 5.4) comparable to that of an activated phagolysosome, which is not tolerated by most bacteria (22). As 1-TbAd can be shed extracellularly to deacidify the phagosomal environment as seen in M. tuberculosis, we hypothesized that the production of 1-TbAd by M. kansasii::Rv3377-78c may modulate the pH of the media (22). As expected, M. kansasii::Rv3377-78c was able to grow in lower pH than M. kansasii::EV in 7H9 culture medium (see Fig. S1 in the supplemental material). However, at 8 and 17 days postinoculation, both M. kansasii::EV and M. kansasii::Rv3377-78c slightly increased the pH of the media where there was bacterial growth, and to a similar extent (Fig. 3). Therefore, although we observed enhanced growth with M. kansasii::Rv3377-78c compared to M. kansasii::EV, we could not demonstrate a causative role for 1-TbAd raising the extracellular pH of the culture media under these conditions.

Prior work with M. tuberculosis estimated that 1-TbAd might naturally accumulate to micromolar concentrations in phagosomes, and 5 to 20 μM 1-TbAd alters lysosomal pH and morphology in human macrophages (22). The proposed lysosomotropic mechanism requires that 1-TbAd access a low-pH compartment where the uncharged conjugate base binds protons to raise pH and regenerate a concentration gradient that promotes further entry of uncharged conjugate base to the acidic compartment. Whereas this mechanism can relieve pH stress on the bacterium, the major alternative, which is not exclusive of lysosomotropism, is that 1-TbAd directly signals for bacterial growth and division. To distinguish these mechanisms, we “chemically complemented” WT M. kansasii with synthetic 1-TbAd and N⁶-TbAd added externally in media. In this experiment, TbAd (already carrying a proton) should not alter pH, but it would contact bacteria in high concentrations. As expected, the addition of synthetic 1-TbAd (pK_a ∼ 8.5) and N⁶-TbAd (pK_a ∼ 3.8) did not alter the pH of the 7H9 media (Fig. S2a). Next, we inoculated M. kansasii into pH-adjusted 7H9 broth (pH 4.0, 4.8, 5.0, 5.2, 5.4, and 6.7) containing 1, 5, 10, or 20 μM TbAd and monitored growth over 16 days (Fig. 4 and Fig. S2b). With increasing doses of 1-TbAd or its isomer N⁶-TbAd, we did not observe any promotion of M. kansasii growth in normal or acidic 7H9 broth (Fig. 4 and Fig. S1b). In fact, 1-TbAd partially inhibited growth at 20 μM, the highest dose tested, at normal pH. These data demonstrated that the protection from low pH in 7H9 culture media afforded by Rv3377c-Rv3378c complementation in M. kansasii is cell intrinsic, promotes growth only at low pH, and does not occur with direct exposure to protonated TbAd.

We aimed to identify whether 1-TbAd production alters/maintains the pH of the bacterial cytosol when grown in acidic media. M. kansasii::EV or M. kansasii::Rv3377-78c was stained with carboxyfluorescein diacetate succinimidyl ester (CFSE) to measure their intracellular pH while monitoring their growth in different pH-adjusted 7H9 broth (pH 4.0, 4.8, 5.0, 5.2, 5.4, 6.0, 6.7, and 7.2) overnight (Fig. 5). Lower initial pH of the media was associated with lowering intracellular pH over the experimental time frame, as expected. Importantly, we observed the expected growth advantage with 1-TbAd production at lower pH (5.0), but there was no apparent intracellular pH difference between M. kansasii strains at this or any pH. Thus, 1-TbAd production does not aid M. kansasii growth at low pH by maintaining the pH of the bulk intracellular milieu, suggesting that the growth advantage is provided by countering the effect of low pH in a specific region of the cell.

1-TbAd is hypothesized to promote virulence of M. tuberculosis by countering phagosome acidification, enhancing survival of the pathogen in cellulo (19, 28, 29). We therefore wished to test the virulence of M. kansasii::Rv3377-78c. In C57BL/6 mice, M. tuberculosis expands in a logarithmic scale within the early course of infection. In sharp contrast, M. kansasii remains at its initial levels of infection, suggesting that it is a good model for acquisition-of-virulence studies within the mouse (3). We hypothesized that the pulmonary bacterial load of M. kansasii in mice as a measure of virulence would be enhanced with 1-TbAd production. During pilot experiments, we infected mice with M. kansasii::EV and M. kansasii::Rv3377-78c through aerosolization, and infections with either strain resulted in a pulmonary burden within 1 log unit of the initial infection up to day 42, with no clear differences between both groups with the sample sizes used (Fig. S3).

Despite repeated attempts to standardize the inoculum, we consistently noted a higher number of 1-TbAd-producing M. kansasii::Rv3377-78c 1 day after aerosol infection compared with M. kansasii::EV (not shown). To test whether Rv3377c-Rv3378c was altering the dose administered or instead enhancing establishment or early growth, mice were aerosolized with M. kansasii::EV or M. kansasii::Rv3377-78c and their lungs were collected and homogenized shortly (4 h) after infection and compared to bacterial counts 24 h after infection (Fig. 6a and b). Equivalent 4-h CFU counts were observed for M. kansasii::EV and M. kansasii::Rv3377-78c; only the latter multiplied successfully 24 h later (Fig. 6a). M. kansasii::Rv3377-78c showed a 1.5-fold increase in numbers from 4 to 24 h, while M. kansasii::EV numbers remained equal (Fig. 6b). The experiment was performed three times, once at a low dose of 100 CFU/lung and twice at a higher dose of 750 CFU/lung, with M. kansasii::Rv3377-78c consistently attaining larger numbers by 24 h postinfection in all three instances (Fig. S4).

We hypothesized that 1-TbAd was enhancing proliferation of M. kansasii in the lungs by promoting survival in resident macrophages. First, using bone marrow-derived macrophages (BMDMs), we infected cells in culture with M. kansasii::EV or M. kansasii::Rv3377-78c and collected cell lysates at 4 and 24 h postinfection: the infections proceeded similarly with both bacteria unlike what we had observed in mouse lungs in vivo (Fig. S5). Alveolar macrophages are resident lung macrophages that phagocytose infectious agents entering the lower airways (30). We assessed whether fitness would be altered in an ex vivo infection of murine alveolar macrophages. In two independent experiments, M. kansasii::Rv3377-78c increased in numbers by CFU counts from 4 to 24 h postinfection, while M. kansasii::EV numbers were largely unchanged and lower than the M. kansasii::Rv3377-78c numbers after 24 h (Fig. 6c and d). M. kansasii::Rv3377-78c exhibited a 2.0-fold increase compared to the 1.1-fold increase seen for M. kansasii::EV (Fig. 6e). This is consistent with our observations in murine lungs and with the conclusion that 1-TbAd provides an advantage to M. kansasii during the early stages of pulmonary infection by specifically promoting survival or growth in resident alveolar macrophages.

We validated a translaryngeal infection model wherein WT M. kansasii was directly introduced into the upper respiratory tract (Fig. S6 and Materials and Methods) (31). Mice were monitored for 42 days; bacteria persisted at the same log CFU as the initial infection, and the mice did not become overtly sick (Fig. S6). To test the effect of 1-TbAd in a high-dose competitive infection, wherein we expected high numbers of bacteria to allow us to see subtle changes in bacterial burden, we used the translaryngeal infection model to generate a mixed infection with 3 × 10⁶ CFU of a 1:1 WT M. kansasii and M. kansasii::Rv3377-78c (Fig. 7a). Lung homogenates plated on 7H10 plates with and without 50 μg/ml hygromycin revealed a statistically significant decrease in the ratio of M. kansasii::Rv3377-78c to WT M. kansasii over time, with an initial decrease in the proportion of M. kansasii::Rv3377-78c in the bacterial population from 0.48 (week 0) to 0.27 (week 1), stabilizing at the latter proportion over time (Fig. 7a). These findings show no beneficial effect of 1-TbAd production for M. kansasii survival in vivo.

C-C chemokine receptor 2 (CCR2) is an essential component for defense in the airways; Ccr2^−/− mice lose the ability to recruit non-tissue-resident immune cells and succumb to M. tuberculosis infection (32). We used these mice to test whether the short-term alveolar macrophage phenotype could be recapitulated in a longer-term in vivo setting without the interference of recruited immune cells in WT mice that might explain the lack of a phenotype in the previous experiment. We aerosol infected Ccr2^−/− mice with a mixed 1:1 bacterial suspension of WT M. kansasii and M. kansasii::Rv3377-78c (Fig. 7b). Interestingly, although WT M. kansasii exhibited a slight increase in numbers over time, M. kansasii::Rv3377-78c steadily decreased within the same period. A statistically significant decrease in the M. kansasii::Rv3377-78c to WT M. kansasii ratio over time was noted, with a steady, nonstabilizing decrease in the proportion of M. kansasii::Rv3377-78c in the bacterial population (Fig. 7b). With these unexpected results, to be sure that our method of identifying M. kansasii::Rv3377-78c (hygromycin resistance) in the mixed infection was valid, we compared hygromycin resistance of M. kansasii::EV (Fig. S7a) and M. kansasii::Rv3377-78c (Fig. S7b) after 4 weeks in vivo from separate aerosol infections. Hygromycin resistance declined by up to 20% initial levels (Fig. S7b); this is clearly less than the 40 to 70% decline observed in the proportion of M. kansasii::Rv3377-78c during mixed infections (Fig. 7). Neither strain appeared more fit in the mouse in the separate infection (Fig. S7c). Thus, 1-TbAd production clearly did not enhance M. kansasii survival in any of these in vivo infection models but in contrast may hinder fitness in the long term.

M. kansasii ATCC 12478 and M. tuberculosis H37Rv were grown in Middlebrook 7H9 broth (BD Difco, MD, USA) as previously described (3). To test the ability of M. kansasii::Rv3377-78c to produce yellow pigment, fully formed colonies were additionally exposed to white light and incubated at room temperature for 7 days. Where indicated and to ensure single-cell suspensions, liquid bacterial cultures were declumped by slowly passaging the cell suspensions through 22-gauge (G) needles five times, 25-G needles five times, and 26-G needles three times, followed by low-speed centrifugation at 50 × g for 5 min with passage through a 5-μm filter. To generate M. kansasii::Rv3377-78c, a 2.4-kb PCR fragment spanning Rv3377c-Rv3378c was generated using primers BamHI-Rv3377c-Rv3378c-F and HindIII-Rv3377c-Rv3378c-R (see Table S1 in the supplemental material) using high-fidelity Phusion DNA polymerase (New England Biolabs). The fragment was subsequently digested with BamHI and HindIII (all restriction enzymes from New England Biolabs) and cloned into the episomal plasmid pMV261 with the constitutive mycobacterial hsp60 promoter and a selective apramycin resistance marker. The hsp60-Rv3377c-Rv3378c fragment was shuttled into the integrative vector pMV306 containing a hygromycin resistance cassette using XbaI and HindIII. All ligations were done using T4 DNA ligase (Fermentas). The resulting plasmid pMV306::Hsp60-Rv3377c-Rv3378c was verified by Sanger sequencing (Genome Québec) to ensure the absence of frameshift or point mutations during the cloning process. An unaltered version of pMV306 with a hygromycin resistance cassette was used to create the empty vector (EV) control strain M. kansasii::EV. Following electroporation, M. kansasii::EV and M. kansasii::Rv3377-78c were grown in the presence of 100 μg/ml hygromycin (Wisent).

M. kansasii::EV, M. kansasii::Rv3377-78c, and M. tuberculosis were grown to mid-log phase and subsequently incubated with 0.25 μCi/ml radiolabeled [8-¹⁴C]adenosine (American Radiolabeled Chemicals) for 14 days. Polar lipid fractions were extracted using CHCl₃−CH₃OH−0.3%NaCl (vol/vol/vol) (22, 37). Extracted lipids were spotted on a TLC Silica Gel 60 (Millipore Sigma) with CHCl₃−CH₃OH−H₂O 10:5:1 (vol/vol/vol) used as the mobile phase solvent. The radiolabeled signature was developed using Storm 840 PhosphorImager (GE Healthcare) to visualize adenosine-linked lipids in each lane. [8-¹⁴C]adenosine 1:100 was used as a no-lipid staining control. The plate was stained with 5% phosphomolybdic acid reagent (PMA) (Sigma) and heated briefly using an industrial blow dryer to visualize the total amounts of lipids loaded in each lane.

M. kansasii::Rv3377-78c, M. kansasii::EV, and M. kansasii parent strain were grown in 30 ml of 7H9 media supplemented with albumin-dextrose-saline (5% bovine serum albumin fraction V, 2% anhydrous dextrose, and 0.87% sodium chloride) to late log phase. Bacterial cell pellet and supernatant were separated by centrifugation at 5,000 rpm for 5 min. The cell pellet was resuspended in 4 ml of phosphate-buffered saline (PBS) at pH 7.4 and distributed equally into four 2-ml screw-cap tubes. The cells were pelleted by centrifugation and further resuspended in 1 ml PBS at pH titrated to 7.4, 6.4, 5.5, and 4.5 with hydrochloric acid and incubated for 2 h at 37°C. At the end of 2 h, the cell pellet and the PBS supernatant were collected for lipid extraction. Ten volumes (3 ml) of chloroform/methanol (C/M) at a ratio of 1:2 were added to the cell pellet and supernatant for extraction for 1 h at room temperature. A second extraction under similar conditions was performed with 3 ml of C/M at a ratio of 1:1. The extracted fractions were pooled and dried under a stream of nitrogen gas. Lipids from the 1-ml PBS supernatant were extracted using acidified ethyl acetate by adding 3 μl of 6 N HCl and 1.4 ml of ethyl acetate and mixing for 30 min in an Orbitron shaker. The mixture was centrifuged at 2,000 rpm for 15 min to collect the upper organic phase and dried on glass under a stream of nitrogen gas at room temperature, and total lipids were weighed using an analytical balance. HPLC-MS separations were performed as described previously (17) using equal amount of lipid samples from different experimental conditions as determined by weight on a Mettler balance.

Bacterial cultures were grown on Congo red-containing 7H10 plates for 14 days, scraped into a 15-ml conical tube, washed with water until the supernatant became clear, and incubated with 2 ml dimethyl sulfoxide (DMSO) for 2 h (25, 38). Congo red was measured in the resultant supernatant at A₄₈₈. The values were normalized to the dry weight of the pellet to define the Congo red binding index.

For all pH experiments, liquid media were prepared as usual, and the pH was equilibrated to 4.0, 4.9, 5.0, 5.1, 5.2, 5.4, 6.0, 6.7, and 7.2 using 2 M HCl or NaOH. The optical density at 600 nm (OD₆₀₀) was adjusted to 0.34, and 222 μl of mid-log-phase declumped bacteria was added to 15 ml of freshly prepared, pH-adjusted 7H9 in 150-ml roller bottles (final OD₆₀₀ of 0.005). Triplicate cultures were made per condition (strain and pH) and incubated at 37°C, rolling in the dark. OD₆₀₀ was measured every 2 or 3 days using two 200-μl volumes of culture (technical duplicates) and a Tecan Infinite M200 Pro plate reader. At days 8 and 17, 1 ml was removed from each culture for centrifugation and recovery of supernatant, which was stored at 4°C until the extracellular pH was read using a micro pH combination electrode (AgCl) (Sigma-Aldrich), and Orion Star A111 meter (Thermo Scientific).

Synthetic 1-TbAd and N⁶-TbAd were produced as described previously (39). Bacterial cultures were grown to mid-log phase, declumped as described above, and then inoculated into 96-well plates in 200 μl of pH-adjusted 7H9 containing 1-TbAd, N⁶-TbAd, or vehicle (DMSO) control as indicated. The plates were incubated at 37°C in the dark, and OD₆₀₀ was measured every 1 to 3 days with a Tecan Infinite M200 Pro plate reader.

Bacterial cultures were grown to mid-log phase and declumped as previously described. A total of 5 × 10⁸ CFU was pelleted, the supernatant was completely removed, and the cells were resuspended in 0.3 ml PBS containing 100 μM carboxyfluorescein diacetate succinimidyl ester (CFDA-SE) (CellTrace CFSE; ThermoFisher) for 20 min at 37°C, shaking at 150 rpm in the dark. Bacteria were next diluted with 10 ml of 7H9 and incubated for 4 h at 37°C, rolling in the dark. A portion of bacteria was taken during this incubation for lysis in normal saline (0.9% NaCl) by beating with silica beads (MP Biomedicals, FastPrep-24) to extract free protein-CFSE conjugate to generate a fluorescence-to-pH standard curve. After 4 h, bacteria were washed and resuspended in normal saline to an OD₆₀₀ of 0.4. Twenty microliters of bacteria in saline was subcultured into 180 μl of freshly prepared, pH-adjusted 7H9 in 96-well plates (opaque-black for fluorescence [ThermoScientific Nunclon Delta Surface], translucent-colorless for absorbance [Falcon]). Lysed bacteria were plated similarly for the pH standard curve. Immediately, plates were placed in plate readers (Tecan Infinite M200 Pro) at 37°C, shaking and measuring fluorescence or OD₆₀₀ every 30 min. Fluorescence at 528 nm was measured from 490-nm excitation (pH sensitive), and 520-nm fluorescence was measured from 450-nm excitation (pH insensitive). To calculate pH, 7H9 background was subtracted from all data first (pH did not alter 7H9 fluorescence). Next, a standard curve of 490-nm excitation/450-nm excitation in relation to 7H9 pH was created from the CFSE-containing cell lysate. The 490/450 ratios calculated from the culture wells were applied to the standard curve to determine intracellular pH.

Male and female C57BL/6 and Ccr2^−/− mice (Jackson Laboratories) were used for experiments. Mice were approximately 6 to 16 weeks of age upon infection over all experiments; different groups were age and sex matched. All protocols were approved by independent ethics oversight at the Research Institute of the McGill University Health Centre (RI-MUHC) and followed the guidelines of the Canadian Council on Animal Care (CCAC). C57BL/6 mice were infected through aerosolization (ONARES, NJ, USA) of bacterial cultures at an OD₆₀₀ of 0.4 for 15 min as previously described (3). Alternatively, C57BL/6 mice were infected via translaryngeal intubation using 50 μl of a high-dose mixed bacterial suspension containing both WT M. kansasii and M. kansasii::Rv3377-78c at an OD₆₀₀ of 0.2. C57BL/6 Ccr2^−/− (C-C motif chemokine receptor 2 knockout) mice were infected through aerosolization of a low-dose mixed suspension at an OD₆₀₀ of 1.0. Mouse lungs were harvested at 4 or 24 h (early time points to measure short-term bacterial establishment) and 14, 21, 28, 42, or 56 days postinfection (later time points to measure long-term bacterial persistence) into 1 ml 7H9 and homogenized using an Omni Tissue Homogenizer TH (Omni International) at high speed for 45 s. Serial dilutions made in 7H9 liquid media from lung homogenates were plated on 7H10 plates containing polymyxin B, amphotericin B, nalixidic acid, trimethoprim, and azlocillin (PANTA) with or without 50 μg/ml hygromycin. CFU were counted 2 weeks after plating to determine bacterial burden.

Bone marrow was isolated from C57BL/6 murine tibiae and femora. Bone marrow-derived macrophage (BMDMs) were differentiated with recombinant macrophage colony-stimulating factor (M-CSF) (100 U/ml) (Peprotech) for a period of 7 days as previously described (40), after which they were lifted using 4 ml CellStripper solution (Corning) and seeded into the appropriate tissue culture plates. For alveolar macrophage (AM) isolation, the respiratory tract, including the trachea and lungs was isolated and repeatedly perfused with cold sterile PBS to collect the cells through bronchoalveolar lavage (BAL). Alveolar macrophages were enriched through adherence purification to tissue culture-treated 96-well plates over 24 h, at which point other cells were washed away. All mammalian cells were cultured in RPMI 1640 medium supplemented with nonessential amino acids, 10 mM HEPES, 10% fetal bovine serum (FBS) with or without penicillin-streptomycin (Wisent).

Macrophages were seeded into 96-well plates (100,000 cells/200 μl complete RPMI medium without antibiotics). Bacterial cultures were grown to an OD₆₀₀ of 0.2 to 0.5, clumps were removed to ensure single-cell suspensions and adjusted in complete RPMI medium (without antibiotics) to an OD₆₀₀ of 0.01. Macrophages were infected by replacing the medium with fresh medium containing bacterial suspension. After 4 h of infection, the wells were gently washed three times with PBS to remove extracellular bacteria and fresh complete RPMI medium (without antibiotics) was added to each well. At indicated time points, the plates were spun down at 2,000 × g for 5 min. Each well was subjected to PBS containing 1% Triton X-100 for 10 min at room temperature to induce macrophage lysis. Following serial dilution and plating, CFU were counted on 7H10 plates 2 weeks postplating to determine bacterial burden.

All calculations and statistical analyses were performed using Microsoft Excel or GraphPad Prism. Calculations included (i) the ratio of individual 24-h CFU values/mean 4-h CFU for murine lung and macrophage infection assays to determine bacterial proliferation (data point = 24-h CFU/mean 4-h CFU) and (ii) the proportion of M. kansasii::Rv33778c/total (values paired from individual mice) in competition assays to determine the comparative fitness of WT M. kansasii versus M. kansasii::Rv3377-78c (data point = CFU on 7H10 with hygromycin/CFU on 7H10).