Stress-Induced Reorganization of the Mycobacterial Membrane Domain

In our previous study, we showed that the IMD was no longer detectable by sucrose density gradient in late stationary phase (150 h) (14), but little is known about its behavior during the growth phase transition. To begin dissecting the initial response of the IMD to growth-inhibiting conditions, we monitored the localization of this domain in the early stationary growth phase by using the previously established IMD reporter strain described above. In this strain, endogenous copies of the essential IMD-associated proteins galactosyltransferase GlfT2 and mannosyltransferase Ppm1 have been replaced with the fluorescent fusion proteins HA-mCherry-GlfT2 and Ppm1-mNeonGreen-cMyc, respectively (13). As previously observed, at the 18-h time point in the growth curve, when the cells are in logarithmic growth, the IMD markers were enriched at the growth poles. However, when the cells entered into stationary phase (38 h; optical density at 600 nm [OD₆₀₀] of 3.45), the intense polar fluorescence of the IMD was diminished (Fig. 1A). This intensity change in IMD-associated proteins implied either that (i) the IMD was disassembled and proteins became randomly associated with the conventional plasma membrane (i.e., PM-CW) or formed aggregates within the cytoplasm or (ii) the IMD was spatially reorganized in the absence of polar growth. To distinguish between these two possibilities, we fractionated the cell lysate from a stationary-phase culture (48-h time point) along a sucrose density gradient (Fig. 1B). We found PimB′, an IMD marker, predominantly in the IMD fractions (4 to 6), with a small amount of this peripheral membrane protein released to the cytosol (fraction 1) as well as localized to the PM-CW (with MptA as a marker; fractions 8 to 12). Importantly, in addition to PimB′, both HA-mCherry-GlfT2 and Ppm1-mNeonGreen-cMyc continued to be detectable in the IMD fractions, despite the lack of active elongation of the cells. This implies that the IMD is maintained but spatially rearranged during the log-phase–stationary-phase transition.

In stationary phase, mycobacteria undergo cell division without cell elongation (15), creating a population of cells shorter in cell length (Fig. 1C). During this stationary-phase cell division, cells no longer elongate by adding new cell envelope components at their cell poles but continue to produce cell envelope components for septum synthesis. PG is a key component of the mycobacterial cell envelope and is essential for elongation at the cell poles as well as septum synthesis. A key step in PG synthesis is mediated by MurG, a glycosyltransferase that synthesizes lipid II from its precursor, lipid I. MurG is one of the proteins associated with the IMD proteome (13) and has been found enriched in the subpolar region of M. smegmatis cells (12). Therefore, we speculated that the spatial rearrangement of the IMD correlates with the location of PG synthesis. To address this question, we visualized de novo PG synthesis by using metabolic labeling. We incubated another reporter M. smegmatis strain expressing HA-mCherry-GlfT2 with alkyne-functionalized d-Ala-d-Ala dipeptide (alkDADA) (16) for 15 min and detected the probe by using copper-catalyzed azide-alkyne cycloaddition of a fluorescent azide label. In correlation with active cell envelope elongation, log-phase cells incorporated quantitatively more alkDADA into their cell wall than did stationary-phase cells (Fig. 1D). Furthermore, alkDADA was predominantly incorporated into cell poles, slightly distal to the apparent subpolar location of the IMD marker (see Fig. S1 in the supplemental material). In contrast, the fluorescent signal of de novo PG synthesis was decreased in stationary phase (Fig. 1D) and was localized more prominently in the septa (Fig. S1). Thus, the decrease in polar enrichment of the IMD during stationary phase correlated with the spatially rearranged nonpolar PG biosynthesis.

Nutrient deprivation is a well-established model to study the stress response of mycobacteria, with early work dating back to the 1930s (17, 18). It is also relevant to M. tuberculosis pathogenesis, as tissue infected with M. tuberculosis in mice or human patients expresses genes involved in adaptation to nutrient limitation (19). Interestingly, M. smegmatis exposed to mild nutrient starvation in phosphate-buffered saline with Tween 80 (PBST) continues to divide without cell separation, resulting in a filamentous morphology with multiple nucleoids and multiple septa (20). This response to PBST suggests that septum biosynthesis continues even under limited nutrient availability conditions. These observations led us to question how the IMD responds to nutrient deprivation in M. smegmatis. We grew cells expressing HA-mCherry-GlfT2 and Ppm1-mNeonGreen-cMyc to logarithmic phase in normal Middlebrook 7H9 medium and then replaced the medium with PBST. As expected, the cells no longer grew upon nutrient depletion (Fig. S2A). Consistent with the previous report that PBST-treated cells do not undergo cell separation (20), the average cell length was maintained relatively comparable to that of actively growing cells over 70 h of starvation (Fig. 2A; Fig. S2B). After 20 h of starvation, the IMD foci were no longer enriched at the poles but were reorganized along the cell sidewall (Fig. 2A). We took the 48-h time point and quantified the polar IMD enrichment as the ratio of mean fluorescence intensities between the polar cap and sidewall region. As shown in Fig. 2B and Fig. S3, the polar enrichment of the IMD was significantly reduced over time during the PBST starvation, and this delocalization was already evident at the earliest (6 h) time point (Fig. S3). The IMD remains biochemically isolatable in a sucrose gradient even after 36 h under starvation conditions (Fig. 2C), as evidenced by the enrichment of HA-mCherry-GlfT2 and endogenous PimB′ in fractions 4 to 6. We note that there was a minor and more significant relocation of HA-mCherry-GlfT2 and Ppm1-mNeonGreen-cMyc to the denser part of the sucrose gradient, suggesting that some proteins may not stably associate with the IMD under this stress condition. Finally, in starved cells alkDADA incorporation appeared to decrease, and it labeled almost exclusively the septa, with some cells having multiple septa at various fluorescence intensities (Fig. 2D; Fig. S1). Taken together, these data suggest that the IMD and its associated enzymes are spatially rearranged and maintained under two different nongrowing conditions, stationary phase and starvation, perhaps to support nonpolar cell envelope biosynthetic activities, such as septal PG synthesis.

As mentioned above, MurG is an IMD-associated glycosyltransferase involved in PG biosynthesis (13). Consistent with the association of this critical enzyme with the IMD, our results indicated that IMD enrichment at the cell poles correlates with polar PG biosynthesis. These observations implied that active PG synthesis at the poles could be a prerequisite for the polar enrichment of the IMD. However, it is also possible that IMD enrichment at the pole is triggered by different metabolic cues regardless of polar PG biosynthesis activity. To distinguish these possibilities, we tested if blocking PG biosynthesis triggered the delocalization of the IMD from the polar region. We inhibited PG biosynthesis chemically by using d-cycloserine (DCS), an inhibitor of d-alanine:d-alanine ligase and alanine racemase (21), which are involved in producing d-alanyl–d-alanine, a key substrate for the synthesis of the pentapeptide precursor. When the IMD reporter cell line was treated with 40 µg/ml DCS, the cells immediately ceased growth (Fig. S4A), and the IMD foci became delocalized from the polar region within 6 h of DCS treatment (Fig. 3A and B), similar to the responses found under stationary phase and starvation conditions. We examined the presence of the IMD in a sucrose gradient and found that it was maintained after DCS-induced inhibition of PG biosynthesis (Fig. 3C). These data indicated that perturbation of PG synthesis triggers the delocalization of the IMD from the pole.

To confirm that the rearrangement of the IMD is not due to an off-target effect of DCS, we inhibited PG biosynthesis genetically by using a diaminopimelate (DAP) auxotroph. This mutant cannot synthesize DAP, an essential amino acid for formation of the PG pentapeptide and therefore requires an exogenous supply of DAP. We transformed this strain with a vector to express the previously established IMD marker (mTurquoise-GlfT2-FLAG) and tracked the subcellular localization of the IMD during logarithmic growth in the presence of an exogenous supply of DAP or under DAP depletion. As expected, the IMD marker was associated with the IMD fractions on the sucrose gradient (Fig. 4A, fractions 4 to 6) and enriched at the poles when the cells were actively elongating (Fig. 4B and C). However, when the medium was depleted of DAP, the cells ceased to grow at 8 h (Fig. S4B). Furthermore, at approximately the same time as the cells stopped growing, the IMD marker relocalized along the cell sidewall (Fig. 4B and C). Importantly, the IMD fractions remained isolatable by density gradient fractionation (Fig. 4D, fractions 4 to 6). These observations recapitulate the results obtained using DCS, suggesting that reduced PG biosynthesis is one critical factor that leads to delocalization of the IMD from the pole. Importantly, the IMD is spatially rearranged but maintained as a biochemically isolatable membrane domain.

While these data suggest that ongoing PG synthesis is a prerequisite for polar IMD localization, PG biosynthesis may not be the sole factor for the spatial rearrangement. The spatial change of the IMD in response to growth-phase transition and nutrient deprivation suggests a more global metabolic response. Indeed, when we added isoniazid (INH), an inhibitor of fatty acid biosynthesis, we observed a similar delocalization of the IMD within 5.5 h at two different concentrations of the drug (Fig. S5). These observations, generated using different chemical and genetic inhibition methods, suggest that the subcellular localization of the IMD is likely governed by the general metabolic status of the cell.

We have so far demonstrated that the IMD reorganizes in cells that are not growing. We next asked if the IMD restores the polar localization when cells are returned to a nutrient-replete condition. To monitor cell envelope elongation during recovery from starvation, we used an amine-reactive dye to stain the existing cell walls of starved cells (30 h in PBST) and placed them in fresh Middlebrook 7H9 medium. We determined the resumed envelope elongation by using fluorescence microscopy (Fig. 5A and B). As a control, logarithmically growing cells continued to elongate, with an average extension of 2.8 ± 0.88 µm (mean ± standard deviation) after 3 h of growth in fresh medium. In contrast, the starved cells added only 1.9 ± 0.81 µm to their total cell length during the same 3-h period under the nutrient-replete condition (Fig. 5B). The IMD in starved cells (in PBST) was less enriched in the polar region prior to recovery and became more enriched at 3 h post-nutrient repletion (Fig. 5A), suggesting that cell envelope elongation is slower or delayed in the initial recovery phase after starvation, when polar enrichment of the IMD is not fully restored.

To provide further insights, we used time-lapse microscopy and monitored the IMD in cells starved in PBST for 6 h and then allowed to recover in fresh Middlebrook 7H9 medium (Fig. 5C). In most cells (25 of 29), we observed a pattern of mild IMD polar enrichment and minimal cell growth during PBS starvation (Fig. 5C; Fig. S6). In these cells, we observed polar enrichment within ~4 h of recovery, followed by cell growth within ~6 h of recovery (Fig. 5C; Fig. S6). To determine the location of PG synthesis, we probed recovering cells with alkDADA (Fig. S7). Under this growth condition, the IMD started to become more enriched in the polar region at 4 h post-nutrient repletion (Fig. 5D; Fig. S7). Importantly, this is the same time point when PG metabolic labeling becomes more prominently enriched in the polar region. Structural illumination microscopy (SIM) revealed that the enrichment of the IMD and the enriched synthesis of PG did not colocalize, but the IMD was distinctively subpolar to the PG synthesis (Fig. 5E). Taken together, enrichment of the polar IMD is spatiotemporally correlated with the initiation of polar PG synthesis and cell elongation.

Markerless knock-in M. smegmatis strains expressing both HA-mCherry-GlfT2 and Ppm1-mNeonGreen-cMyc (SUMA187) or HA-mCherry-GlfT2 alone (SUMA86) were previously established (13). These strains were grown in Middlebrook 7H9 medium supplemented with 0.05% Tween 80, 11 mM glucose, and 14.5 mM NaCl at 30°C with shaking. Growth was monitored via the OD₆₀₀. For starvation experiments, M. smegmatis cultures were grown to logarithmic phase, spun down to remove the medium, and resuspended in PBS supplemented with 0.05% Tween 80. The DAP auxotroph (mc²1620) was transfected with the pMUM43 vector to express mTurquoise-GlfT2-FLAG (13). The DAP auxotroph was grown in Middlebrook 7H9 supplemented with Middlebrook ADC supplement, amino acids (l-lysine, l-methionine, and l-threonine at 40 mg/ml and dl-homoserine at 80 µg/ml), and DAP (200 µg/ml) as previously described (25).

M. smegmatis cells were grown in medium supplemented with 2 mM alkDADA for 15 min at 30°C, fixed in 2% formaldehyde, and probed with the Alexa Fluor 488 picolyl azide (Thermo Fisher) for 30 to 60 min at room temperature in the dark.

For chemical inhibition, cells were treated with 40 µg/ml DCS. For genetic inhibition, the DAP auxotroph cells were spun down and resuspended in medium lacking DAP.

M. smegmatis cells were treated with 5 mg/ml Alexa Fluor 488 NHS ester (succinimidyl ester; Thermo Fisher) for 1 min at room temperature in the dark, washed with PBST, returned to fresh Middlebrook 7H9, and incubated further at 30°C with shaking. After 3 h of growth, cells were fixed in 2% formaldehyde and washed with PBST.

To visualize fluorescence, a 2 μl aliquot of live or fixed M. smegmatis cells was spotted onto a 1% agar pad with water and covered with a glass coverslip. All static live cell images were taken under identical settings (100× objective and 175-ms exposure for phase-contrast microscopy, or 3-s exposure for fluorescence), using a Nikon Eclipse E600 microscope equipped with an ORCA-ER cooled charge-coupled-device camera (Hamamatsu) and Openlab software 5.5.2 (Improvision). Mean fluorescence intensity of alkDADA incorporation was calculated using Fiji (26), and the average fluorescence is indicated in the figures by a black bar. Polar enrichment was measured using a custom MatLab program that calculated mean fluorescence of the pixels at the pole (designated 10 px from the polar end), compared to the average sidewall pixel intensity (intensity of total cell minus the identified polar ends). Cell length was measured from phase-contrast images using Fiji. Statistical significance was determined via a t test. Time-lapse microscopy was performed as previously described (13, 27). Briefly, cells were loaded into a custom constant flow microfluidic device (11, 27) and observed by a DeltaVision PersonalDV wide-field fluorescence microscope in a controlled environmental chamber warmed to 37°C. SIM images were acquired via a Nikon Eclipse Ti N-SIM E microscope equipped with a Flash4.0 camera and a numerical aperture of 1.49. Images were taken at 100-ms exposures and reconstructed using NIS-Elements.

For density gradient fractionation, cells were pelleted, washed in 50 mM HEPES (pH 7.4) buffer, and resuspended in lysis buffer (25 mM HEPES [pH 7.4], 20% sucrose, 2 mM EGTA, and a protease inhibitor cocktail) at 1 g (wet weight) of pellet in 4 ml of lysis buffer. The cell suspension was then subjected to nitrogen cavitation at ~2,250 lb/in² for three times for 30 min each. The lysate was spun at 3,220 × g for 10 min at 4°C twice before it was loaded on a 20-to-50% sucrose gradient. Gradients were spun at 218,000 × g for 6 h at 4°C, fractionated into 1-ml fractions, and used for further biochemical analyses.

Protein samples were mixed with a reducing sample loading buffer, denatured on ice, and run in a 12% SDS-PAGE gel. Proteins were transferred to a polyvinylidene difluoride membrane (Bio-Rad) and blocked in 5% milk in PBST20 (PBS with 0.05% Tween 20). Membranes were incubated with primary antibodies (anti-HA [Sigma], anti-PimB′ [28], and anti-MptA [28]) at 1:2,000 dilutions, washed in PBST20, and incubated with horseradish peroxidase-conjugated secondary antibodies (GE Healthcare) at 1:2,000 dilutions. Blots were washed again in PBST20 and developed for chemiluminescence. Images were recorded using an ImageQuant LAS 4000mini (GE Healthcare). For Western blotting of sucrose gradients, an equal volume of each fraction was loaded into the gel. All experiments were repeated at least twice.