Transposon mutagenesis and sequencing in Cglu

The MB001 strain of Cglu was chosen for this analysis. It is a commonly used derivative of ATCC 13032 that has been engineered to remove three defective prophage elements [14]. We used the Tn5-based transposome system for transposon mutagenesis to yield a library of approximately 200,000 mutant colonies. Sequencing the transposon-chromosome junctions in the library yielded 10,419,920 reads that mapped to 200,940 unique insertions sites throughout the genome (Fig 2A). This high-density insertion map allowed the global identification of essential genes in Cglu. Such genes were identified by plotting the number of unique insertions per annotated gene relative to gene length (Fig 2B). The analysis revealed a bimodal distribution with the majority of genes displaying an increasing number of insertions proportional to their length and a small subset of genes that harbored very few insertions regardless of their size (Figs 2B and S1). Genes likely to be essential for growth in rich medium were defined as those harboring one or fewer unique insertions per 127 bp. With this cutoff, the percentage of essential genes was constant across a broad-range of gene lengths except for those shorter than 100bp (S1 Fig). For these smaller genes, an essentiality designation likely reflects technical factors of gene size and transposon density rather than biological function. We therefore excluded genes shorter than 100 bp from the final essential gene list along with tRNAs, rRNAs, and insertion elements. With these exclusions (131 in all, S1 Table), a total of 322 essential genes were identified in Cglu (S1 Table), which corresponds to roughly 11% of the total gene content, a percentage that is in-line with similar analyses of other bacteria, including Mtb [15–18]. Of the 490 total essential genes in Mtb, 445 have homologs in Cglu. However, only roughly half (231/445) of these are essential in Cglu (S2 Table). Thus, consistent with the conditional dispensability of the arabinan component of the AG and the MA layer of the envelope [6–9], Cglu is more tolerant to mutations and therefore should broadly enable studies aimed at defining the biological function of essential Mtb genes.

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Fig 2. Overview of the Tn-seq analysis.

(A) Location of all transposon insertions mapping to the Cglu MB001 genome. The cyan lines represent the location of each gene on the genome. The black bars represent the location of each transposon insertion with the height of each bar representing the detection frequency of each insertion. The numbers are the genome coordinates in base pairs. (B) The number of unique transposon insertions mapped to every gene of MB001 plotted as a function of gene length. For non-essential genes (black dots), the number of unique transposon insertions mapped linearly correlates with gene length. Genes that fall below the blue dashed line (red dots) have two-standard-deviations fewer unique transposon insertions than the mean value and therefore are designated as essential genes. (C) View of the Tn-Seq data for the highly conserved dcw locus encoding many essential genes. (D) The ftsW gene is shown as an example of Domain Essential (DE) gene. DE genes are genes that tolerate transposon insertions in some portion of their the 3’ ends but are depleted of insertions at the 5’ end. (E) Pie chart depicting the breakdown of essential, non-essential and DE genes categorized by the Tn-Seq analysis. Numbers in parenthesis correspond to the number of genes in the specified category.

https://doi.org/10.1371/journal.pgen.1008284.g002

Support for the validity of our essential gene designations came from the fact that many genes expected to play vital roles in bacterial physiology were identified as such. For example, widely conserved genes in the division-cell wall (dcw) cluster with known essential roles in PG precursor biosynthesis (murC, murD, murE, murF and mraY) or cell division (divIC, ftsI, ftsQ and ftsZ) were found in to be depleted of transposon insertions in the profiling data (Fig 2C). Surprisingly, the Tn-Seq data indicated that ftsW, which encodes a cell division PG polymerase [19] essential for the growth of most bacteria, had a high number of insertions mapping within the reading frame, suggesting that it may not be essential in Cglu. Upon closer inspection, all of the insertions were found to be located in the 3’ end of the gene (Fig 2D). Notably, FtsW in Cglu and other Corynebacterineae possesses an extended C-terminal sequence absent from bacteria such as E. coli and B. subtilis (S2 Fig). In mycobacteria, this positively charged tail was previously shown to bind FtsZ and proposed to regulate its assembly into the Z-ring [20,21], the key cytoskeletal structure required for cell division [22]. Our results suggest that the FtsW-FtsZ interaction may not be essential for the viability of Cglu. Based upon the example of ftsW, we computationally searched for genes in which only 5’ end of the reading frame was essential. This analysis led to the identification of 17 “domain essential” genes (S1 Table) bringing the total essential gene content of Cglu to 339 (Fig 2E).

Identification of ethambutol responsive loci

Ethambutol (EMB) inhibits the function of arabinosyl transferases responsible for the polymerization of arabinan chain of AG [12,13] as well as lipoarabinomannan (LAM), an arabinan-containing glycolipid found in the envelope of the Corynebacterineae [23,24]. Recent work has found that EMB treatment inhibits polar growth in Cglu and mycobacteria and causes defects in daughter cell separation [25,26]. We therefore reasoned that new factors involved in growth or division might be identified as loci causing EMB hypersensitivity when inactivated. To this end, the Cglu transposon library was propagated in the presence of a sublethal concentration of EMB (0.3 μg/ml) and then analyzed by transposon sequencing. Loci associated with sensitivity to ethambutol (ste genes) were then identified as those displaying a significant reduction in mapped insertions following EMB treatment relative to an untreated control.

A total of 49 ste loci were identified in the Tn-Seq analysis (Figs 3A and S3A and S3 Table). As an indication that the screen worked as expected, several genes encoding enzymes in the AG biogenesis pathway were identified as hits along with other loci encoding factors with predicted roles in envelope biogenesis, including two loci implicated in daughter cell separation: cgp_1735 and ftsE-ftsX. The cgp_1735 gene, which we will refer to as ripC, is homologous to a gene of the same name in mycobacteria and encodes an exported protein with an N-terminal coiled-coil domain and a C-terminal PG endopeptidase domain that cuts crosslinks in the cell wall matrix (Fig 3A and 3B) [27]. The ftsE and ftsX genes encode an ABC-transporter-like complex (FtsEX) (Fig 3A and 3B) that was originally implicated in the control of cell wall hydrolase activity in E. coli [28] and Streptococcus pneumoniae [29]. FtsEX was subsequently found to be involved in the activation of PG hydrolase activity in a number of organisms, including mycobacteria [30], and was recently suggested to function in complex with RipC to promote cell separation in Cglu [31].

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Fig 3. Identification of sensitive to ethambutol (ste) mutants by Tn-seq.

(A) Shown are transposon insertion profiles for selected regions of the chromosome after growth in medium without and with EMB as indicated. The height of each line in the profile represents the number of sequencing reads corresponding to a transposon insertion at the indicated genome position. The ratio listed below each gene corresponds to the ratio of the total number of sequencing reads for all insertions in a given gene following growth in medium with EMB relative to the no drug control. P-value reports the statistical significance of the difference in Tn-Seq profiles for each condition based on the Mann-Whitney U test. Profiles highlighted in yellow correspond to genes that showed a statistically significant reduction in transposon reads following EMB treatment (ratio <0.33 and p-val <0.05). (B) Schematic showing the envelope localization and membrane topology of factors implicated in cell separation. The TMHMM algorithm predicts that both SteA and SteB have an N-in/C-out topology. (C) Overnight cultures of MB001 (WT) and its indicated deletion derivatives were normalized to an OD₆₀₀ of 0.5, serially diluted, and spotted (5 μl) onto BHI agar medium with and without 1 μg/ml EMB as indicated. Plates were incubated for 24 hours at 30°C and photographed. Note that mutants with cell separation defects that form aggregates were vortexed for 3 seconds to resuspend the cells before OD₆₀₀ measurements were taken for normalization.

https://doi.org/10.1371/journal.pgen.1008284.g003

To validate the screening results, deletion alleles for several of the identified ste genes were constructed and the corresponding mutants were tested for EMB hypersensitivity. All of the deletion mutants constructed were found to display some degree of sensitivity to EMB (Figs 3C, S3B), indicating that the screen was faithfully identifying loci responsive to EMB. The morphology of each mutant was also assessed by fluorescence microscopy following staining with the membrane dye FM4-64. The number of septa per cell was also quantified following growth in the presence of a fluorescent D-amino acid (FDAA), which labels septa and other sites of new cell wall synthesis [32]. Indicative of their expected cell separation defect [26,27,31], mutants deleted for ripC or the ftsEX locus yielded a population of cells that were longer than average relative to wild-type and possessed one or more unresolved septum (Fig 4A and 4B). As previously observed [26], cellular compartments that were ‘trapped’ in the middle of chaining cells failed to elongate properly, presumably because their cell poles remained attached to the neighboring cells. Among the other ste mutants constructed, only deletions in steA and steB were found to cause a cell separation defect that resembled cells inactivated for ripC or ftsEX (Fig 4A and 4B).

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Fig 4. Cell separation defect of ste mutants.

(A) Overnight cultures of MB001 (WT) and its indicated derivatives were diluted 1:1000 in BHI and grown at 30°C. When the OD₆₀₀ of the cultures reached 0.2, cells were stained with FM 4–64 (1.5 μg/ml) for 2 minutes before they were spotted on agarose pads for visualization by fluorescence microscopy. Scar bar: 2 μm. (B) Assessment of cell separation defects by quantification of septa/cell. Overnight cultures of the strains from (A) were diluted 1:1000 in BHI and grown at 30°C. When the OD₆₀₀ reached 0.2, cells were gently pelleted, resuspended and diluted 1:10 in BHI supplemented with the fluorescent D-amino acid HADA (1 μM). Following growth at 30°C for 3 hours, labeled cells were loaded on a CellASIC ONIX microfluidic plate, flushed with BHI and imaged by fluorescence microscopy. The number of septa per cell was tallied using the Cell Counter plugin in FIJI (n > 300 for each strain).

https://doi.org/10.1371/journal.pgen.1008284.g004

The steA and steB genes are located in an apparent operon that is highly conserved within the Corynebacteriales order (S4A Fig). SteA is a predicted to be a bitopic transmembrane protein with an N-terminal cytoplasmic domain and a small C-terminal domain at the membrane surface (Fig 3B). SteB is also a single pass transmembrane protein, but it is predicted to have a small N-terminal tail in the cytoplasm and a soluble C-terminal domain exposed to the envelope (Fig 3B). As expected from the Tn-Seq data where steA displayed a greater depletion of transposon insertions following EMB treatment than steB (Fig 3A), a ΔsteA mutant was found to have a more severe EMB-sensitivity phenotype than a ΔsteB mutant. The double ΔsteAB mutant was as sensitive to EMB as a single ΔsteA mutant, indicating that steA is epistatic to steB (S5 Fig). Importantly, the phenotypes of all steA/B deletion mutants could be complemented upon ectopic expression of the missing gene(s), indicating that the phenotypes of the deletions were not due to adverse effects on the expression of nearby genes (S5 Fig). Given the importance of SteA and SteB for cell division and their conservation among the Corynebacterineae, we chose to investigate their functions further. Studies of the other ste loci will be pursued as part of a separate line of investigation.

Genetic evidence that SteA and SteB function in the RipC cell separation pathway

In addition to RipC, Cglu encodes a second PG hydrolase called RipA that plays a role in cell separation [27]. Unlike a ΔripC mutant, cells inactivated for RipA alone were not found to display a separation defect [27]. However, deletion of ripA in a ΔripC background was found to exacerbate the cell separation defect displayed by cells in which only RipC was inactivated [27]. We re-created these mutants in the MB001 background and confirmed these findings (S6 Fig). Furthermore, we found that although deletion of ripA alone did not change the EMB-sensitivity of MB001 cells, it enhanced the EMB sensitivity of ΔripC cells (Fig 5). These results suggest that RipA and RipC contribute to distinct cell separation pathways. To determine whether SteA and SteB contribute to one of these pathways or form yet another mode of cell separation, we combined the ste deletions with those of ripC or ripA and assessed their EMB sensitivity. Deletion of ripA was found to enhance the EMB sensitivity of cells deleted for steA or steB, but ΔsteA ΔripC and ΔsteB ΔripC mutants displayed the same EMB sensitivity as the single ste deletion mutants (Fig 5). We therefore conclude that SteA and SteB are likely functioning as part of the RipC cell separation pathway. Further support for this possibility is presented below.

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Fig 5. EMB sensitivity of double mutant strains.

Overnight cultures of MB001(WT) and its indicated derivatives were normalized to an OD₆₀₀ of 0.5, serially diluted 10-fold, and spotted (5 μl) onto agar medium with or without 0.4 μg/ml EMB as indicated. Plates were incubated for 36 hours at 30°C and photographed. Note that a lower concentration of EMB was used in this experiment relative to that shown in Fig 3C in order to better discern the severity of the EMB hypersensitivity phenotype for each strain.

https://doi.org/10.1371/journal.pgen.1008284.g005

Mutants defective for SteA, SteB, RipC, and FtsEX are delayed for V-snapping

The multiple septa observed in the ΔsteA, ΔsteB, ΔripC, and ΔftsEX mutants could either be due to a defect in cell separation or to septal closure being slowed such that new ones form before the old septa are completed. Like other bacteria, cell division in Cglu is mediated by a cytokinetic ring apparatus organized and controlled by polymers of the tubulin-like protein FtsZ [33]. We monitored the lifetime of the Z-ring constriction process as a proxy for the rate of septal closure. A fluorescent monomeric superfolding variant of GFP (msfGFP) fused to FtsZ (msfGFP-FtsZ) was produced from an ectopic site on the chromosome in the wild-type and mutant strains, and the lifetime of the Z-ring from formation to resolution was monitored by time-lapse microscopy. In the same cells, the progress of septum completion was monitored with a fluorescent trehalose reporter, 6-TMR-Tre [34]. This labeled molecule is incorporated into the non-covalently attached trehalose glycolipids that form the outer leaflet of the mycomembrane. Previous work has shown that 6-TMR-Tre infiltrates the newly completed septum through perforations in the peripheral peptidoglycan just before the daughter cells abruptly separate via a mechanical fracturing event called V-snapping [26]. The time delay between 6-TMR-Tre infiltration and V-snapping therefore provides a useful kinetic parameter with which to characterize the efficiency of the cell separation process.

In wild-type cells and all of the mutants assayed, the characteristic lifetime of the Z-ring constriction process remained unchanged (Fig 6A and 6B). However, while wild-type and the ΔripA mutant showed the expected short lag between septal infiltration of trehalose glycolipid and V-snapping (7.6 and 4.8 min, respectively), the time between these two events was significantly extended in the ΔsteA, ΔsteB, ΔripC, and ΔftsEX mutants (between 31.7 and 45 min) (Fig 6C and 6D). We therefore conclude that the cell division defect in these mutants is the same and related to the cell separation process, not slowed septal closure.

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Fig 6. Assessment of the cell division phenotype of ste mutants.

(A) Shown are representative time-lapse fluorescence images used to follow Z-ring dynamics in MB001(WT) cells and its indicated derivatives. Yellow arrowheads denote Z-ring formation, and they transition to a red arrowhead at the time of observed resolution (note the expansion of the fluorescent band). These strains expressed msfGFP-ftsZ from the integrated pHCL106 plasmid as well as native ftsZ. Overnight cultures were diluted 1:1000 in BHI and grown at 30°C. When the OD₆₀₀ reached 0.1, cells were loaded into a CellAsic microfluic plate and imaged by phase-contrast and fluorescence microscopy at 4-minute intervals. (B) Quantification of the Z-ring lifetime for each strain. The life-time of the Z-ring was defined as the time elapsed between Z-ring formation and its resolution, which was observed as the transition from a constricted band to an expanded band of lower fluorescence intensity. Z-ring lifetimes were manually measured for each strain and plotted as the cumulative percentage of cells with a lifetime less than or equal to a given time on the X-axis. The lifetime of thirty Z-rings was quantified for each strain. (C) The same cells in (A) were also monitored for the infiltration of trehalose glycolipid by exposing the cells with BHI supplemented with 100 μM 6-TMR-Tre for two minutes every 1 hour during imaging [34]. Representatives time lapse fluorescence images taken at 4 min intervals are shown for each strain. Yellow arrowheads denote septal infiltration of trehalose glycolipid and red arrows denote the V-snapping event. Note that the septa of ΔftsEX, ΔripC, ΔsteA and ΔsteB mutants eventually undergo V-snapping, but they are not shown here due the extreme delay. (D) V-snap delay was defined as the time elapsed between infiltration of trehalose glycolipid at the septum and V-snapping. It was quantified manually for each strain shown in (C) and plotted as cumulative percentage of cells with a V-snap delay less than or equal to a given time on the X-axis. Fifty infiltration events were quantified for each strain. For (C) and (D), solid lines represent best-fit curves of each dataset with the Hill equation: cumulative percentage = , in which τ characterizes the lifetime of Z-ring or the V-snap delay.

https://doi.org/10.1371/journal.pgen.1008284.g006

SteA and SteB are recruited to the division site

To investigate whether SteA and SteB are likely to be playing a direct role in the cell separation process, we assessed their subcellular localization. Fusions with mScarlet-I (mScar) [35] were constructed and produced from the attB1 site in the chromosome. Both mScar-SteA and mScar-SteB fusions restored the normal cell length distribution to cells lacking the native protein (S7A–S7C Fig). The mScar-SteA fusion was also found to reduce the number of observed septa per cell back to normal (Fig 7A). However, there was a notable difference in the ability of the fusions to rescue the EMB-sensitivity of mutant cells. The mScar-SteB fusion came close to fully complementing the EMB^S phenotype of ΔsteB mutants (S7D Fig). By contrast, mScar-SteA only weakly rescued the EMB-sensitivity of ΔsteA cells (Fig 7A), indicating that there is not a perfect correspondence between cell separation and EMB resistance. We infer from this result that SteA may have a second function unrelated to cell separation and that the fusion may be defective for this activity (See Discussion).

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Fig 7. SteA and SteB localize to the division site.

(A) Functionality of SteA variants. Constructs encoding the indicated mScar-SteA variants under P_steA control were integrated in the genome of the ΔsteA mutant. To quantify the number of septa per cell, the indicated strains were grown in BHI that was supplemented with NADA to label PG for five hours before imaging by phase-contrast, and for fluorescence (green and red channels). SteA_N indicates the N-terminal portion of SteA covering residue 3–347. TM_A indicates the C-terminal portion of SteA (residue 331–393) containing the transmembrane domain. Constructs encoding the indicated SteA variants are as follows: pHCL171: mScar-SteA; pHCL172: mScar-SteA_N; pHCL173: mScar-SteA_N-TM₁; pHCL174: mScar-SteA_N-TM₂; pHCL175: TM_A; and pHCL57: SteA. Septa per cells were scored using the Cell Counter plugin in the FIJI software [59] and the percentage of cells with zero, one or more than one septum were calculated and plotted. (B-C) Phase-contrast and fluorescence micrographs of Cglu cells producing mScar-SteA (B) or mScar-SteB (C), as the sole copy of the corresponding gene, from integrated plasmids pHCL171 or pHCL170, respectively. (D) Correlation of cell length-dependent mid-cell accumulation of mScar-SteA with septal PG synthesis, the infiltration of the trehalose glycolipid, and Z-ring formation at the population level. Overnight cultures of the appropriate strains were diluted 1:1000 in BHI and grown at 30°C. When the OD₆₀₀ reached 0.2–0.3, cells were processed (when necessary) and spotted on an agarose pad for phase-contrast and fluorescence microscopy. Cells were grouped according to their cell lengths into bins of 0.3 μm increments and the percentage of cells with septal signal were calculated for each bin. With the exception of mScar-SteA (see below), solid lines represent the mean value while the shaded areas represent standard error of the mean. To follow the timing of Z-ring assembly, exponentially growing MB001(attB1HCL106) cells expressig msfGFP-FtsZ were used (n = 2, 535 cells and 734 cells, respectively). To visualize mid-cell localization of SteA, HL2(attB1HCL171) cells were used (n = 157 and 879 cells, respectively). Black filled spheres represent mean values and error bars represent standard error of the mean. Septal PG labeling and trehalose-glycolipid infiltration were followed in both strains. For septal PG labeling, exponentially growing cells were pulse-labeled with 1 μM NADA for 5 min before imaging. The plotted curve represents the average of two highly similar curves (low standard error) obtained from the two strains MB001(attB1HCL106) (n = 734) and HL2(attB1HCL171) (n = 879). For trehalose-glycolipid labeling, cells were grown in the BHI medium supplemented with 100 μM 6-TMR-Tre for 5 min and washed twice with BHI by gentle pelleting and resuspension before imaging. As with the septal PG labelling, the curve represents the average of the results from the two strains: MB001(attB1HCL106) (n = 535) and HL2(attB1HCL171) (n = 330). (E) Time-lapse microscopy showing coincidental mid-cell accumulation of mScar-SteA and commencement of septal PG construction. Unlabeled HL2(attB1HCL171) cells in the exponential growth phase were loaded in a CELLASIC ONIX microfluidic device. Five minutes before the start of time-lapse microscopy, the cells were exposed to medium supplemented with 1 μM NADA to label sites of active peptidoglycan synthesis (see Methods for details). Images were taken at 8 minutes intervals. Yellow arrows indicate concomitant appearance of NADA and mScar-SteA signals at the cell septum. Cyan arrow indicates dissolution of the septal mScar-SteA signal upon V-snapping.

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Cells expressing mScar-SteA or mScar-SteB fusions displayed one of two localization patterns. In recently separated cells that remained tethered to their siblings, the mScar signal decorated the cell periphery, consistent with a membrane localization (Fig 7B and 7C). In cells that were presumably further along in the division cycle, cells displayed tight band of fluorescence at the prospective division site with minimal signal observed along the cell periphery (Fig 7B and 7C). Thus, SteA and SteB are recruited to the cytokinetic ring and are likely to be directly participating in the division process.

We next investigated the timing of SteA recruitment to the division site relative to other measurable cell division events: Z-ring formation assessed using msfGFP-FtsZ, septal PG construction assessed by labeling of the septum with the FDAA called NADA [32], and the infiltration of labeled trehalose-glycolipids into cell septum. The analysis was carried out using population level measurements of individually labeled cells to determine the cell length (a proxy for age) at which 50% of the population displayed label localized at the division site. As expected, Z-ring formation, septal PG synthesis, and infiltration of trehalose-glycolipids at the cell septum occurred in apparent chronological order with characteristic cell lengths of septal labeling of 1.76, 2.75, and 3.49 μm, respectively (Fig 7D). The characteristic cell length of mScar-SteA recruitment to the division site was 2.64 μm, which coincides with the inception of septal PG assembly (Fig 7D). To further investigate the relative timing of these two events, we simultaneously monitored nascent PG synthesis and mScar-SteA localization at the single-cell level in a time-lapsed experiment. Cells producing mScar-SteA were imaged within a microfluidic device, which allowed constant infusion with media containing NADA. Consistent with the population level analysis, in every one of the more than thirty cells that were manually monitored in time-lapse, the mScar-SteA and NADA signals appeared at the division site simultaneously (Fig 7E). Furthermore, the mScar-SteA signal was observed to disperse from the cell septum immediately following V-snapping (Fig 7E), indicating that SteA does not linger at the new cell poles following division.

Septal localization of SteA requires its cytoplasmic and transmembrane domains

Next, we investigated the domain requirements for SteA localization and function. A series of mScar-SteA variants were constructed in which different parts of the protein were either deleted or substituted with another domain (Fig 7A). Even though all of the protein fusions accumulated to comparable levels in Cglu (S8 Fig), only the full-length mScar-SteA protein was capable of rescuing the the cell separation phenotype of the ΔsteA mutant or partially rescuing EMB hypersensitivity (Fig 7A). In accordance with the functionality of the fusions, only mScar-SteA showed specific enrichment at the septum (S9 Fig). The N-terminal domain of SteA (mScar-SteA_N) displayed diffuse cytoplasmic signal, whereas mScar fused to the C-terminal transmembrane domain of SteA (mScar-TM_A) labeled the peripheral membrane as well as septal membranes (S9 Fig). To test whether the native SteA transmembrane domain plays a role in SteA function or localization beyond anchoring the N-terminal domain to the membrane, it was replaced by unrelated transmembrane domains from two putative tail-anchored membrane proteins of Streptomycetes coelicolor: SecE (TM₁) and PkaB (TM₂) [36]. These constructs, mScar-SteA_N-TM₁ and mScar-SteA_N-TM_2, displayed a peripheral localization signal consistent with membrane recruitment, but they were not functional and did not promote specific septal labeling (Figs 7A and S9). We conclude that septal localization of SteA is critical for proper cell separation and that both the N-terminal and transmembrane domains of the proteins are required for its function.

SteB promotes robust septal localization of SteA

To determine whether any of the other factors required for cell separation are required for the recruitment of SteA to the division site, we visualized the localization of mScar-SteA in mutants inactivated for SteB, RipC, or FtsEX. All of these mutant strains contain multiple septa resulting from their cell separation defect. In each case mScar-SteA labeled these septa (Fig 8A). However, we noticed that the intensity of mScar-SteA septal bands in the ΔsteB mutant was consistently lower when compared to the other two mutants. The lower septal mScar-SteA signal in ΔsteB cells was not due to a reduction in the overall mScar-SteA protein level (S10 Fig). Rather, we measured a statistically significant increase in the non-septal mScar-SteA signal in the ΔsteB mutant (2.74 ± 0.17, 23 cells) compared to those of ΔftsEX (1.88 ± 0.09, 18 cells) or ΔripC cells (2.19 ± 0.14, 24 cells) (Fig 8B). As a result, the ratio of septal mScar-SteA signal to background in the ΔsteB mutant (1.07 ± 0.37) was significantly reduced compared to mutants inactivated for ftsEX (2.20 ± 0.72) or ripC (1.85 ± 0.72) (Fig 8C). These results indicate that SteB is required for optimal recruitment of SteA to the division site. The observation that SteA retains some ability to localize in the absence of SteB may explain why the ΔsteB mutant exhibits a milder EMB sensitivity phenotype than cells lacking SteA.

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Fig 8. SteB is required for robust recruitment of SteA to the division site.

(A) Representative images of the indicated Cglu mutants producing mScar-SteA under P_steA from genome integrated pHCL171. Overnight cell cultures were diluted 1:1000 in BHI and grown at 30°C. When the OD₆₀₀ reached 0.2–0.3, cell cultures were diluted 10-fold and loaded in a CELLASIC ONIX microfluidic device for phase-contrast and fluorescence microscopy. Scale bar: 2 μm. (B) Reduced accumulation of mScar-SteA signal at the division site in cells lacking SteB. Representative examples of mScar-SteA fluorescence profiles in the indicated strains using cells in (A) marked by yellow arrows. Cell detection and fluorescence quantification was performed using Oufti [60]. To quantify the background signal and the septal signal, the fluorescence profiles were fitted with multi-exponential Gaussians plus background (black line) using Matlab. The height of each Gaussian peak was taken as the value of septal signals. (C) Bar graphs comparing the septal mScar-SteA signal to background ratio of the indicated strains (31 septa from at least 18 cells from each strain were used for measurements). P-values (Tukey-Kramer test) report the statistical significance between the difference of the means using Prism software.

https://doi.org/10.1371/journal.pgen.1008284.g008

SteA and SteB form a complex that interacts with RipC

The requirement of SteB for robust recruitment of SteA to the division site suggested that the two proteins might interact at the septum. To investigate their potential interaction further, we employed a two hybrid assay we recently developed called POLAR based on PopZ-linked apical recruitment in E. coli. This method takes advantage of the polar organizing protein called PopZ from Caulobacter cresentus [37–39] and its ability to form polar foci in E. coli. Bait proteins are fused to the H3H4 peptide of PopZ and sfGFP. When they are produced in E. coli cells that also make full-length PopZ, the bait fusions are recruited to the cell pole by the H3H4 peptide-PopZ interaction (Fig 9A). Prey proteins are fused to mScar and their interaction with the bait is assessed based on whether or not they are also recruited to the pole (Fig 9A). The assay is thus similar to previous cytological interaction detection systems based on DivIVA fusions [40,41], but it benefits form the more robust and specific polar accumulation of PopZ in E. coli relative to DivIVA. An in-depth description of the POLAR method and the types of protein-protein interactions it can detect will be presented in a separate report.

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Fig 9. SteA and SteB form a complex.

(A) Schematic depicting the expected localization patterns of a prey depending on whether or not it interacts with the bait in the POLAR two hybrid assay. (B) Top row: representative fluorescence images of TB28 E. coli cells expressing the indicated the bait and prey proteins. The indicated baits were expressed using the following medium-copy plasmids: pHCL149 (TM control) and pHCL202 (SteA). The TM control harbors residues 2–55 of Pseudomonas aeruginosa PAO1 PonB covering its transmembrane domain that has a N-in/C-out topology. The preys were expressed from genome-integrated plasmids (pHCL147: ^ssDsbA and pHCL194: SteB). Cell cultures were grown from a single colony in LB supplemented with antibiotics at 37°C for two hours. To induce the bait and prey proteins, cells were collected by centrifugation and resuspended in M9 supplemented with 0.2% arabinose and 100 uM IPTG and grown for two hours at 37°C. Cells were spotted on an agarose pad for imaging. Bottom row: demographs showing fluorescence distribution of the corresponding prey and bait fusions at the population level. Single-cell fluorescence quantification was performed using Oufti [60]. Cells were oriented using a custom-written MATLAB script such that the cell pole with the higher bait fluorescence was located on the right of the demograph. At least 250 cells were used to generate each demograph.

https://doi.org/10.1371/journal.pgen.1008284.g009

To assess the SteA-SteB interaction, H3H4-sfGFP-SteA was used as the bait. The fusion successfully resulted in the recruitment of SteB-mScar to the cell pole whereas a control membrane-anchored bait protein H3H4-sfGFP-TM did not (Fig 9B). Additionally, a control periplasmic mScar protein was also not recruited to the pole by H3H4-sfGFP-SteA (Fig 9B). In addition to visual inspection of individual cells, the POLAR results can also be viewed at a population level using a demograph in which quantified fluorescence traces of cells are stacked according to cell length and visualized as a heat map. Such population level results also support a specific interaction between the H3H4-sfGFP-SteA bait and SteB-mScar (Fig 9B). Given that E. coli is only distantly related to Cglu and unlikely to encode interaction partners for Cglu proteins, we conclude from the POLAR results that SteA and SteB are likely to interact directly to form a complex.

We next wanted to assess the ability of the SteAB complex to interact with RipC. However, the assay was complicated by the RipC-mScar localization pattern in cells producing the control bait fusion H3H4-msfGFP-TM, which it is not expected to interact with. The bait displayed a primarily unipolar localization signal as expected, but RipC-mScar localized to both poles in a manner uncorrelated to the pattern of the control bait (Fig 10A and 10B). Even though cells in the demograph were oriented such that the pole with higher bait accumulation was oriented to the right, RipC-mScar intensity was distributed roughly equally between the two poles (Fig 10A). This bipolar localization of RipC-mScar in the E. coli periplasm likely reflects some level of aggregation due to the predicted intrinsically disordered and coiled coil domains present in the N-terminal portion of RipC. Accordingly, a RipC truncation mutant lacking the C-terminal NlpC/p60 catalytic domain (^ssDsbA-mScar-RipC_N) also showed bipolar localization when it was secreted to the E. coli periplasm (S11A Fig). Nevertheless, when H3H4-msfGFP-FtsEX was used as the bait, the mScar-RipC signal became biased to the pole with the highest bait signal. In this case the correlation index (CI) between the FtsEX bait and the RipC prey localization pattern was 0.96 ± 0.04, much higher that the 0.14 ± 0.12 index value between RipC and the control bait (Fig 10B). Given that FtsEX and RipC are known to interact robustly in mycobacteria, we conclude that the POLAR assay can accurately detect RipC interactions despite its tendency to generate a bipolar signal when fused to mScar. We therefore assessed whether or not RipC interacts with SteAB using H3H4-msfGFP-SteA as a bait co-expressed with untagged SteB. As with FtsEX, the intensity of the polar RipC-mScar signal was highly correlated with the H3H4-msfGFP-SteA bait signal (CI = 0.64 ± 0.07), albeit to a slightly lesser degree than FtsEX (Fig 10B). A similar correlation of polar signals (CI = 0.67 ± 0.12) was obtained when a prey harboring only the N-terminal domain of RipC was used instead of the full-length protein (S11A and S11B Fig). This result indicates that RipC interacts with the SteAB complex primarily if not entirely through its N-terminal domain. To determine which component(s) of the SteAB complex directly contacts RipC, we used the N-terminal domain of RipC fused to H3H4-msfGFP via a TM domain (H3H4-msfGFP-TM-RipC_N) as the bait. It recruited SteB-mScar but not mScar-SteA prey to the cell poles (Fig 10C), suggesting that SteB directly binds RipC via its N-terminal coiled-coil rich domain. This interaction requires the C-terminal part of SteB as a SteB variant lacking the extracellular domain (SteB_N-mScar) failed to be recruited to the cell pole by the RipC_N bait (Fig 10C). Notably, SteB_N-mScar retains its ability to interact with SteA (S11C Fig). Using the POLAR assay, we were unable to detect a direct interaction between FtsEX and SteA or SteB (S11D Fig). Overall, the interaction data suggest that SteAB forms part of the FtsEX-RipC complex at the septum in Cglu to promote septal PG remodeling and daughter cell separation via V-snapping (Fig 10D).

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Fig 10. RipC interacts with both the SteAB and FtsEX complexes.

(A & C) Top row: representative fluorescence images of TB28 E. coli cells expressing the indicated the bait and prey proteins. The indicated baits were expressed using the following plasmids: pHCL149 (TM control), pHCL204 (SteAB), pHCL205 (FtsEX), pHCL213 (RipC_N). The preys were expressed from genome-integrated plasmids (pHCL221: RipC, pHCL214: SteA, pHCL194: SteB and pHCL196: SteB_N). Cells were grown and imaged as in Fig 9B. (B) Fluorescence correlation between the bait and the prey at the cell pole. To quantify the relative distribution of the the bait and prey between the poles, we first calculated fractional difference of the bait signal (D_bait) and the prey signal (D_prey) between the poles. The correlation between the bait and the prey signal distribution was represented by the correlation index (CI), which is the ratio of D_prey to D_bait. CI value of 1 indicates complete positive correlation, 0 indicates no correlation while -1 indicates complete negative correlation. A least 215 cells were used in each calculation (n = 2 for the FtsEX experiment and n = 3 for the control and the SteAB experiments). (D) Summary of the protein-protein interaction network of the RipC cell separation pathway.

https://doi.org/10.1371/journal.pgen.1008284.g010

Media, bacterial strains, and plasmids

All Cglu strains used are derivatives of MB001 [14]. Unless mentioned otherwise, strains were grown in Brain-Heart Infusion Medium (BHI) that was supplemented with 15 μg/mL kanamycin (Kan), when necessary. Plasmids were maintained in either DH5a(λpir) or S17-1(λpir). All E. coli strains used in the reported POLAR two-hybrid experiments are derivatives of TB28 [51] and are grown in LB (1% tryptone, 0.5% yeast extract, 0.5% NaCl). Whenever necessary, antibiotics were used at 25 (Kan), 15 (chloramphenicol; Cm), 50 (ampicillin; Amp) or 5 (tetracycline; Tet) μg/mL. Growth conditions for microscopy experiments are described in the figure legends and the appropriate Methods section. Detailed information about plasmid constructions can be found in S1 Text. All strains are listed in S4 Table and all plasmids are listed in S5 Table.

Gene deletion in Cglu

We used the temperature-sensitive plasmid pCRD206 to perform allelic replacement in Cglu essential as described previously [52] with some modifications. Briefly, sequences corresponding to approximately 700 bp upstream and downstream of the desired deletion were inserted into pCRD206. The resulting plasmid was transformed into the appropriate recipient strain. Transformants were selected and propagated on BHI-Kan agarose plates at 25°C. A few colonies were then purified on BHI-Kan agarose and grown at 37°C for 36 hours. One or two of the resulting Kan^R colonies were then grown in BHI liquid medium lacking antibiotic at 25°C overnight. The lower temperature allows replication from the plasmid origin, which selects for a second recombination to remove the plasmid from the chromosome. An aliquot of the overnight culture (10 μL) was then spread on a BHI agarose plate supplemented with 10% sucrose, which further selects against sacB in the vector, and the plate was then incubated overnight at 30°C. The resulting colonies were replica patched onto BHI and BHI-Kan plates to identify Kan^S colonies lacking plasmid. Deletion alleles in Kan^S isolates were finally confirmed by colony PCR.

Plasmid integration at the attB1 site in Cglu

For complementation studies, genes were integrated at the attB1 site on the chromosome using the pK-PIM vector [53]. The plasmid was introduced into the recipient by conjugation.

Conjugation into Cglu

Plasmids were transferred into Cglu using the E. coli strain S17-1(λpir). Conjugation was performed according to [54] with some modifications. Briefly, overnight cultures of recipient strains grown in LB supplemented with 4% (w/v) glucose (LB-Glu4) were diluted 1:5 in the same medium and incubated for two hours at 30°C. After adjusting the density to an OD₆₀₀ of 2.0, each culture was incubated in a 48.5°C water bath for 9 minutes. Overnight cultures of the donor strain were diluted 1:100 in LB supplemented with the appropriate antibiotics and grown at 37°C until OD₆₀₀ reached 1.0. When the recipient cultures cooled to room temperature, the donor and recipient cultures were mixed at 1:3 ratio, gently pelleted to collect cells, which were then resuspended into 150–200 μL LB-Glu4 medium. The cell suspension was pipetted onto a 0.4 μm sterile filter placed on a LB-Glu4 agarose plate and incubated. Following overnight incubation at 30°C, cells were collected from the filter and resuspended into LB and plated on BHI-Kan agar plates that were supplemented with 30 μg/mL nalidixic acid to select against E. coli donors. Plasmid integration was confirmed by colony PCR using a primer mixture consisting of ATTB1g-UP, ATTB1g-UN, ATTB2g-UP and ATTB2g-UN [53]. Using WT gDNA as the template, the PCR reaction generates two bands of 464 bp and 700 bp in size, which represent uninterrupted attB1 and attB2 sites respectively. Therefore, specific disappearance of the 464-bp band was used as a diagnostic for plasmid integration at the attB1 site.

Preparation of electrocompetent Cglu cells

Cells of Cglu were grown to stationary phase, and 10 mL of this culture was diluted into 1 L BHIS (BHI + 91 g/L sorbitol) that was supplemented with 25 g glycine, 0.4 g isoniazid, and 0.1% Tween 80 and incubated shaking at 18°C. The culture was chilled on ice for 1 hour when the OD₆₀₀ reached 0.5 (typically in 16–18 hours). Cells were then pelleted at 4000 x g for 20 min, washed once with 500 ml chilled 10% glycerol, and then three additional times with 100 ml chilled 10% glycerol. The cell density was adjusted to an OD₆₀₀ of 20 before use for electroporation.

Tn5 transposon insertion library construction and sequencing

The Tn5 transposome was purchased from Epicentre (now discontinued) or prepared in-house using Tn5 transposase purified as described previously [55]. For the in-house preparation, the transposon was amplified by polymerase chain reaction (PCR) using 5’-monophosphorylated primers ME Plus 9–3’ primer (CTGTCTCTTATACACATCTCAACCATCA) and ME Plus 9–5’ primer (CTGTCTCTTATACACATCTCAACCCTGA) using the <KAN-2> transposon supplied by Epicentre as the template. The transposon was purified using the QIAquick PCR purification kit (Qiagen) and eluted with TE buffer (10 mM Tris pH 8.0 and 1 mM EDTA). The Tn5 transposome was reconstituted by vortexing a 1:1 mixture of 1 μM purified transposase [55] with the purified transposon (200 ng/μL) and used immediately after a 30-minute incubation at room temperature. Consistent with another report [56], we found that the Tn5 transposase prepared according to Picelli et al. 2014 loses activity quickly following purification.

To generate mutant libraries, transformation was carried out by electroporating 1 μL of commercial or homemade Tn5 transposome with 100 μl of freshly prepared MB001 electrocompetent cells. Each transformation reaction typically generated between two to five thousand transformants. We pooled transformants from 50 electroporations performed over 4 days to generate our library consisting of approximately 230,000 total transformants.

DNA libraries for Tn-Seq analysis were prepared by a modified version of a published protocol [15]. Genomic DNA was extracted from strains using the Wizard Genomic DNA Purification Kit (Promega) and further purified using Genomic DNA Clean & Concentrator (Zymo). Genomic DNA was fragmented using a Qsonica Q800RS Sonicator for 12 minutes (using a 15 second on and 15 second off pulse cycle) at 20% amplitude. Fragmented DNA was purified with 1.8× volume Agentcourt AMPure XP beads (Beckman Coulter, Inc.) and eluded into 30 μl water. Purified fragmented DNA was then treated with terminal deoxynucleotidyl transferase (TdT; Promega) in a 20 μl reaction with 1 μL 9.5mM dCTP/0.5mM ddCTP, 4 μl 5× TdT reaction buffer and 0.5 μl rTdT at 37°C for 1h, then at 75°C for 20min. TdT-treated DNA was purified with Performa DTR Gel Filtration Cartridge (EdgeBio). Purified, TdT-treated DNA was used as a template in a PCR reaction to amplify the transposon junctions using the Easy-A Hi-Fi Cloning System (Agilent Technologies). The primers used were:

PolyG-1st-1 5’-GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCTGGGGGGGGGGGGGGGG-3’ and Tn5-1st-1 5’-ACCTGCAGGCATGCAAGCTTCAGGG-3’.

A second nested PCR was then performed to further amplify the transposon junctions and append the sequencing barcode. The primers used were generic NEBNext Multiplex Oligos for Illumina (NEB) and:

Tn5-2nd-1 5’-AATGATACGGCGACCACCGAGATCTACACTCTTTTCAGGGTTGAGATGTGTATAAGAGA-3’.

The final product was run on a 2% agarose gel, and fragments ranging from 200–500 bp were gel purified using QIAquick Gel Extraction Kit (Qiagen). Libraries were sequenced at the Tufts University Core Facility on a HiSeq 2500 (Illumina) or a Miseq (Illumina) on a 1× 50 single end run. All sequencing data generated in this study are deposited in Sequence Read Archive under accession PRJNA548135.

Reads were mapped to the C. glutamicum MB001 genome (NCBI NC_022040.1). To identify EMB sensitive genes, we calculated the fold change in reads between the ethambutol conditions and the no drug conditions. We also performed a Mann-Whitney U test to determine whether differences in the insertion profiles were statistically significant. Genes in which insertions were at least 3-fold enriched or depleted and had a p-value lower than 0.05 by Mann-Whitney were defined as hits. Visual inspection of transposon insertion profiles was performed with the Sanger Artemis Genome Browser and Annotation tool.

POLAR two hybrid strains construction

Each strain harbors a replicative plasmid that expresses the bait and another genome-integrated plasmid that expresses the prey. We simultaneously introduced, by electroporation, both plasmids into the TB28 recipient that carried the helper plasmid pAH69 [57]. Transformants that integrated the prey plasmid at the HK022 phage attachment site and also harbored the bait plasmid were selected for by plating on LB agarose plates that contained Cm and Tet at 37°C, which prevented the replication of pAH69. Electrocompetent TB28/pAH69 cells were prepared by first diluting an overnight culture into LB supplemented with Amp and incubated shaking at 30°C. When OD₆₀₀ reached 0.15, the culture was shifted to 42°C for 30 minutes to induce integrase expression. The culture was then chilled on ice for 1 hour. Cells were then collected by centrifugation (4000 x g) for 20 min, washed once with 500 ml chilled 10% glycerol, and then three additional times with 100 ml chilled 10% glycerol.

Image acquisition and analysis

Growth conditions and staining procedures prior to microscopy are described in the figure legends. Prior to imaging, cells were immobilized on 2% agarose pads containing the appropriate growth medium, and covered with #1.5 coverslips [58]. Images were cropped and adjusted using FIJI software [59].

Microscopy: Images were obtained using a Nikon Ti inverted microscope that is fitted with a Nikon motorized stage with an OkoLab gas incubator with a slide insert attachment, an Andor Zyla 4.2 Plus sCMOS camera, Lumencore SpectraX LED Illumination, Plan Apo lambda 100x/1.45 Oil Ph3 DM objective lens, and Nikon Elements 4.30 acquisition software. Images in the green and red channels were taken using Chroma 49002 and 49008 filter cubes, respectively. The microscope was maintained at 30°C using a custom-made environmental control chamber.

Microfluidic time-lapsed experiment: Specific experimental setups are described in the figure legends. Cells were loaded into the CellASIC Onix B04 microfluidic plates (Millipore Sigma) that were attached to the microscope described above using using a multi-well insert. Each imaging chamber was flushed with the appropriate growth medium before loading the cells. During the course of the time-lapse, appropriate growth media were supplied to the cells using a constant flow rate of 60 psi. To avoid reduction of growth rate due to phototoxicity, we reduced the intensity of the excitation light using neutral density filters (at least ND8) in all our experiments.

Single-cell image analysis: Measurements of cell lengths and fluorescence signals at the single cell level were carried out using Oufti [60]. To identify cells with septal signals, the Oufti output was additionally analyzed using peakFinder, which is an add-on from Oufti, to identify the position of fluorescence peaks along the length of the cells. Custom-written Matlab code was used to further identify cells with a fluorescence peak that is located within 0.3–0.7 position within the normalized length of the cell. Plots of cummulative percentage of cells with a septal signal as a function of cell length were generated in MATLAB. The curves were fitted with the Hill coefficient using MATLAB as described in the figure legend.

Demograph generation: Using custom-written MATLAB code, cells were arranged from top to bottom according to their cell lengths. Additionally, in demographs of both the bait and prey, each cell was oriented such that the cell pole with the higher bait intensity was located on the right.

Correlation of bait and prey signal distribution between the cell poles: Only signals within 780 nm from the cell poles were used for the analysis. We first calculated the fractional difference in the bait signal between the left and right poles as designated in the demograph, using the equation , in which B_L and B_R represents the bait signals at the left and the right cell pole, respectively. The polar prey signals were analyzed similarly using the equation , in which P_L and P_R represents the bait signals at the left and the right cell pole, respectively. Finally, the correlation index, CI, was calculated using the equation (CI = 1: complete positive correlation, CI = 0: no correlation and CI = -1: complete negative correlation).

Fluorescent measurement of SteA or SteB protein levels

Overnight cultures of cells expressing mScar fusions to SteA, SteA variants, or SteB were diluted 1:1000 in fresh BHI and incubated shaking at 30°C until the optical density reached 0.5. An aliquot of cells (25 mL) was harvested by centrifugation and resuspended in 100 μL Lysis Buffer A (20 mM Tris 7.5, 10 mM EDTA, 1 mg/ml lysozyme, 1 mM PMSF, 10 μg/ml DNAseI and 100 μg/ml RNAseA). The cell suspensions were incubated at 37°C for 30 min before an equal volume of 2x SDS loading buffer was added. The samples were sonicated for 10 min to reduce the viscosity before they were resolved on a 4–20% Criterion TGX Precast Midi Protein Gel (Biorad). The gel was washed in H₂O for 10 min before it was imaged with the Cy3 and Cy5 channels on a Typhoon FLA Scanner (GE Healthcare). The image was scaled and cropped using FIJI [59].

Phylogenetic tree

For the phylogenetic tree showing the distribution of SteA, SteB and RecA homologs, the amino acid sequences of SteA (cgp_1603), SteB (cgp_1604) and RecA (cgp_2141) were used as a query in a BLASTp search against the NCBI “non redundant” (nr) database [61] with an e-value cutoff of 1e^-4 for each protein. A list of all the taxa for which significant BLAST results were found was then sorted. We used a complex and diverse set of 1773 bacterial taxa called “Representative Genomes” that is available on NCBI (ftp://ftp.ncbi.nlm.nih.gov/blast/db/, Representative_Genomes.00.tar.gz). The phylogenetic tree was constructed using PhyloT (http://phylot.biobyte.de/) and BLASTp results were plotted against the tree. The tree was visualized and annotated using iToL (http://itol.embl.de/) [62].

Genetic neighborhood analysis

A database of 1542 fully assembled representative bacterial genomes was created (https://www.ncbi.nlm.nih.gov/assembly). The same queries as for the phylogenetic tree were used in a tBLASTn against this database. The start and stop positions of the alignment were sorted for each genome and the distances between steA and steB or steA and recA were calculated for each organism.