MraZ Transcriptionally Controls the Critical Level of FtsL Required for Focusing Z-Rings and Kickstarting Septation in Bacillus subtilis

ABSTRACT

The bacterial division and cell wall (dcw) cluster is a highly conserved region of the genome which encodes several essential cell division factors, including the central divisome protein FtsZ. Understanding the regulation of this region is key to our overall understanding of the division process. mraZ is found at the 5′ end of the dcw cluster, and previous studies have described MraZ as a sequence-specific DNA binding protein. In this article, we investigate MraZ to elucidate its role in Bacillus subtilis. Through our investigation, we demonstrate that increased levels of MraZ result in lethal filamentation due to repression of its own operon (mraZ-mraW-ftsL-pbpB). We observed rescue of filamentation upon decoupling ftsL expression, but not other genes in the operon, from MraZ control. Our data suggest that regulation of the mra operon may be an alternative way for cells to quickly arrest cytokinesis, potentially during entry into the stationary phase and in the event of DNA replication arrest. Furthermore, through time-lapse microscopy, we were able to identify that overexpression of mraZ or depletion of FtsL results in decondensation of the FtsZ ring (Z-ring). Using fluorescent d-amino acid labeling, we also observed that coordinated peptidoglycan insertion at the division site is dysregulated in the absence of FtsL. Thus, we reveal that the precise role of FtsL is in Z-ring maturation and focusing septal peptidoglycan synthesis.

IMPORTANCE MraZ is a highly conserved protein found in a diverse range of bacteria, including genome-reduced Mycoplasma. We investigated the role of MraZ in Bacillus subtilis and found that overproduction of MraZ is toxic due to cell division inhibition. Upon further analysis, we observed that MraZ is a repressor of its own operon, which includes genes that encode the essential cell division factors FtsL and PBP2B. We noted that decoupling of ftsL alone was sufficient to abolish MraZ-mediated cell division inhibition. Using time-lapse microscopy, we showed that under conditions where the FtsL level is depleted, the cell division machinery is unable to initiate cytokinesis. Thus, our results pinpoint that the precise role of FtsL is in concentrating septal cell wall synthesis to facilitate cell division.

INTRODUCTION

Bacterial cell division is a highly orchestrated process that typically results in the creation of two identical daughter cells through binary fission (1–3). Many species of bacteria encode a conserved gene neighborhood known as the division cell wall (dcw) cluster (4, 5). In general, the peptidoglycan biosynthesis genes are found toward the 5′ end, and genes encoding important cell division factors, including ftsZ, are found at the 3′ end (5–7). Gram-positive bacteria have an additional conserved region, the ylm operon, downstream of ftsZ, which includes genes for additional cell division factors such as sepF and divIVA (8, 9).

At the very 5′ end of the dcw cluster is a gene encoding a DNA binding protein, MraZ (previously known as yllB), which is conserved in diverse lineages. In a range of species, including Escherichia coli and many Firmicutes, mraZ is found within a short operon consisting of itself, mraW (rsmH; yllC), ftsL (yllD) and pbpB (pbp2B) (10). In genome-reduced Mycoplasma species, the dcw cluster consists of mraZ and mraW followed by ftsA and ftsZ alone (11).

Previous work in E. coli and Mycoplasma showed that overexpression of mraZ results in a change in transcriptional regulation of its own operon, causing lethal filamentation in E. coli and cell enlargement in Mycoplasma (11–13). In E. coli, this phenotype could be resolved by coexpression of the gene immediately downstream of mraZ, mraW, a putative 16S rRNA (and possibly DNA) methyltransferase (12, 14). Work in Burkholderia cenocepacia has shown that P_mra is the sole transcription start site of the dcw cluster, and MraZ can bind to the promoter sequence and presumably act as a transcriptional regulator (15). Recent investigation of Neisseriaceae family organisms revealed that deletion of mraZ among other factors may have allowed for the evolution of alternative growth modes (16). A recent report about Staphylococcus aureus proposes a role for MraZ in virulence regulation (17).

In this report, we show that overexpression of mraZ is toxic in Bacillus subtilis due to cell division inhibition similar to what has been reported in other organisms thus far. Through fluorescence microscopy, we show that MraZ is a DNA-associated protein. Using transcriptional reporter assays and transcriptome sequencing (RNA-Seq) analysis, we show that MraZ functions as a transcriptional repressor of the mra operon and thus an inhibitor of cell division. Additionally, we demonstrate that MraZ-mediated lethal cell division inhibition is driven primarily by the reduction in the levels of FtsL, a critical divisome component that is turned over rapidly (18). Finally, we provide evidence that coalescence of FtsZ protofilaments during maturation of FtsZ ring (Z-ring) assembly is impaired upon MraZ overproduction. More specifically, FtsL depletion also leads to similar impairment of Z-ring assembly and even promotes disassembly of what appears to be mature Z-rings. However, depletion of FtsL did not have any observable effect on Z-rings that were already constricting. Using fluorescent d-amino acid labeling, we observed that under conditions when FtsL is depleted, peptidoglycan insertion does not occur in a focused manner, which in turn results in impairment of septation. Thus, our results illustrate that the main role of FtsL is in promoting Z-ring maturation, focusing Z-ring treadmilling, and providing the initial impetus required for septum formation in the form of organized peptidoglycan synthesis.

(A preprint of this article was posted on bioRxiv [19]).

RESULTS

Overproduction of MraZ is lethal to B. subtilis and is dependent on DNA binding.

To investigate the role of mraZ in B. subtilis, we constructed an IPTG (isopropyl-β-d-thiogalactopyranoside)-inducible copy of mraZ at an ectopic locus. We grew cultures of wild-type (WT) B. subtilis and cells containing inducible mraZ and plated serial dilutions on lysogeny broth (LB) agar with and without 1 mM IPTG. We found that when grown on IPTG, cells containing inducible mraZ (mraZ⁺) were unable to grow at any dilution, in contrast to the WT control (Fig. 1A). This is similar to what has previously been shown in E. coli (12). Additionally, we investigated whether overexpression of mraZ resulted in a growth defect in liquid medium. After the addition of 1 mM IPTG (3 h), cells overproducing MraZ display a drop in cell density at an optical density of 600 nm (OD₆₀₀), indicating cell lysis (Fig. 1B). CFU/mL enumeration revealed an approximately 1,000-fold drop in viable cell count between WT (3.8 × 10⁶) and mraZ⁺ (1.9 × 10³) 6 h after the addition of IPTG. To study the cause of lethality, we observed mraZ⁺ cells under the microscope. When grown in the presence of 1 mM IPTG for 2 h, mraZ⁺ cells are extremely filamentous (14.6 μm ± 6.9 μm), in contrast to the WT control (3.4 μm ± 1 μm) (Fig. 1CD and Fig. S1B to D in the supplemental material). This phenotype is indicative of cell division arrest in rod-shaped organisms. Eventually, the mraZ-overexpressing cells go on to lyse, explaining the lethal phenotype observed on solid and liquid media.

FIG 1.

- Overproduction of MraZ is lethal to B. subtilis and is dependent on DNA binding. (A) Spot assay of WT (PY79) B. subtilis and cultures containing inducible mraZ⁺ (MW189), mraZ^R15A (MW256), or mraZ^R86A (MW350), serially diluted and spotted onto agar in the absence and presence of 1 mM IPTG. The cartoon on the right depicts the control of mraZ/mutants (mraZ*) under native promoter or IPTG-inducible promoter. (B) Growth curve of WT mraZ⁺, mraZ^R15A, and mraZ^R86A in the presence of 1 mM IPTG. Readings were taken every hour at OD₆₀₀. (C) Fluorescence micrographs of WT B. subtilis (i and ii), mraZ⁺ (iii and iv), mraZ^R15A (v and vi), and mraZ^R86A (vii and viii) in the presence and absence of 1 mM IPTG. Cells were induced with IPTG for 2 h prior to imaging. Cell membrane was stained with SynaptoRed (FM4-64). Scale bar = 1 μm. (D) Quantification of cell length from microscopy in panel C. n = 100 cells; ****, P < 0.0001.

MraZ belongs to the AbrB and SpoVT family of transcription factors and contains two highly conserved DXXXR motifs (Fig. S1A) (20–22). Previous work by Eraso et al. (12) had shown that a single point mutation of the first motif from arginine to alanine (R15A) was sufficient to prevent lethality in E. coli. We sought to identify whether this was the case for B. subtilis MraZ and generated point mutations in both DXXXR motifs—R15A and R86A. As described earlier, we serially diluted and plated cells expressing both the R15A and R86A variants on LB agar with and without 1 mM IPTG. When grown in the presence of IPTG, neither mutant caused lethality, and cells grew like the WT B. subtilis control (Fig. 1A). This was also seen in liquid culture (Fig. 1B). When imaged under the microscope, cells expressing the R15A and R86A mutants were observed to be phenotypically WT-like in length (3.3 μm ± 1.28 μm and 3.5 μm ± 0.9 μm, respectively), in contrast to the filamentation observed in the unmutated mraZ overexpression strain (Fig. 1C and D). Thus, the ability of MraZ to cause filamentation, and thereby induce lethality, relies on the presence of both of these motifs.

MraZ associates with the chromosome through DXXXR motifs.

To monitor whether MraZ localizes to the chromosome, we constructed a C-terminal fusion of MraZ to the green florescent protein (GFP). When overexpressed on solid medium, mraZ-gfp results in the formation of translucent colonies (Fig. 2A); however, there is no significant growth defect in liquid medium (Fig. 2C). This phenotype is in contrast to the overproduction of untagged MraZ, which is toxic on both solid and liquid media. Overproduction of MraZ-GFP results in cells that are slightly shorter (10.4 μm ± 6.7 μm) than those produced by overproduction of untagged MraZ (15.7 μm ± 6.9 μm) but are nonetheless filamentous in comparison to the WT control (3.7 μm ± 1.1 μm), (Fig. 2B and D). This suggests that MraZ-GFP may only be partially functional. GFP signal can be seen as multiple nucleoid-associated foci in the absence of inducer or exclusively at the chromosome, overlapping the DNA-specific DAPI stain in the presence of inducer (Fig. 2B) (as noted previously [23]). The coating of entire nucleoid suggests that MraZ has the potential to play a larger role in nucleoid organization and/or as a transcriptional factor with control of many genes spread throughout the genome.

FIG 2.

MraZ associates with the chromosome through DXXXR motifs. (A) Spot assay of cultures containing WT (PY79) B. subtilis, mraZ⁺ (MW189), mraZ-gfp (MW295), mraZ^R15A-gfp (MW296), and mraZ^R86A-gfp (MW351) serially diluted and spotted onto plates with and without 1 mM IPTG. (B) Micrographs of cells expressing mraZ-gfp, mraZ^R15A-gfp, and mraZ^R86A-gfp in the absence of IPTG (top), the presence of IPTG (middle), or the presence of IPTG and chloramphenicol (bottom). Cells were induced with 1 mM IPTG for 2 h prior to imaging without or with 6.25 μg/mL chloramphenicol for the last 60 min. Membrane is visualized with SynaptoRed, and DNA is stained with DAPI. Scale bar = 1 μm. (C) Growth curve of WT, mraZ⁺, mraZ-gfp, mraZ^R15A-gfp, and mraZ^R86A-gfp cultures following the addition of 1 mM IPTG; readings were taken every hour at OD₆₀₀. (D) Quantification of cell length in panel B. n = 100 cells, ****, P < 0.0001. (E) The stability of MraZ-GFP and its mutants was confirmed through immunoblotting. Cell lysates from induced and uninduced cultures were probed with anti-GFP and anti-Sigma A (loading control) antibodies.

We tagged mraZ^R15A and mraZ^R86A to gfp to generate C-terminal GFP fusions to identify whether mutation of the DXXXR binding motif prevents MraZ from colocalizing with the DNA. In both cases, GFP signal can be seen diffused in the cytoplasm, and the cells are WT-like in length (R15A: 3.4 μm ± 1.1 μm, R86A: 3.4 μm ± 0.9 μm) (Fig. 2B and D). The diffused localization of the DXXXR mutants became more obvious upon chloramphenicol treatment (Fig. 2B; 24) or in the stationary phase, where the nucleoids are more condensed (Fig. S2). We confirmed the stable production of tagged proteins via western blotting (Fig. 2E), indicating that the diffuse signal is a result of MraZ-GFP mislocalization and that MraZ binding to the chromosome is dependent on the presence of both DXXXR DNA binding motifs.

MraZ represses expression of the mra operon through the MBRs.

The promoter of the mra operon contains multiple MraZ binding repeats (MBRs) in a diverse range of species, including Mycoplasma species, E. coli, and B. subtilis (11). In B. subtilis, this repeat consists of three GTGG[A/T]G motifs separated by a 4-nucleotide spacer (11; Fig. 3A). We were able to observe similar patterns in 7 other species that belong to the Firmicutes phylum and generated a sequence logo of the consensus sequence showing conservation of the GTGG repeat in the upstream region of the mra operon (Fig. S3A). Following a BLAST search of the PY79 genome, only one double or triple repeat was found within the promoter region of mraZ. In addition, we conducted a search of the PY79 genome through Pattern Locator (25) and were only able to identify a triple repeat in the upstream region of the mra operon, confirming what was found through BLAST. Probing for only double repeats in Pattern Locator resulted in 13 hits (Table S2A). Besides MBRs upstream of mraZ, only two hits landed in an intergenic region—one partially overlapping the open reading frames of genes mraY and murD (part of the dcw cluster) and the other overlapping yaaL and the intergenic region upstream of bofA. A search using a GTGGAG or GTGGTG single repeat resulted in 464 and 332 hits, respectively. However, whether MraZ binds at these sites remains to be investigated.

FIG 3.

MraZ represses expression of the mra operon through the MraZ binding repeats (MBRs). (A) Cartoon representation of the mra promoter region and operon showing the binding repeats in yellow and the nucleotide sequence of the region below. Graphical representation of the transcriptional fusion of P_mra to gfp (MW385) and of the mra promoter in which the MBRs were mutated (MBR*; MW525) or switched with the −35/–10 region of gpsB (ΔMBR; MW478). (B) Immunoblot of cultures containing a transcriptional fusion of P_mra to gfp (MW368, lanes 1 and 2) with either mraZ⁺ (MW385, lanes 3 and 4), mraZ^R15A (MW389, lanes 5 and 6), or mraZ^R86A (MW429, lanes 7 and 8) in the presence and absence of 1 mM IPTG. Whole-cell lysates were probed with anti-GFP and anti-Sigma A (loading control) antibodies. (C) Immunoblot of cultures containing GFP-based transcriptional reporter under the control of WT mra (MBR; MW385, lanes 1 and 2), a mutant promoter either lacking the MBRs (ΔMBR; MW478, lanes 3 and 4) or with the MBRs mutated (MBR*; MW525, lane 5 and 6), with and without mraZ overexpression. Whole-cell lysates were probed with anti-GFP and anti-SigA antibodies. (D) Volcano plot of RNA-Seq analysis from overexpression of mraZ (MW189) with 1 mM IPTG. Points in red are genes within the mra operon. (E) Table showing the log fold change of RNA levels for the mra operon genes in mraZ⁺ (MW189), mraZ^R15A+ (MW256), and ΔmraZ (MW192) compared to the WT (PY79) control. (F) Venn diagram showing genes differentially regulated as identified through RNA-Seq when mraZ (MW189) and mraZ^R15A (MW256) were overexpressed or mraZ was deleted (MW192) relative to the WT (PY79). The total number of differentially regulated genes is indicated in the parenthesis.

To investigate the regulation of the mra operon in B. subtilis, we constructed a GFP-based transcriptional reporter of the mraZ promoter that includes all three MBR repeats (P_mra-gfp) (Fig. 3A). This construct was introduced in the WT and mraZ⁺ backgrounds, and the cell lysates were western blotted and probed with anti-GFP and anti-SigA antibodies (Fig. 3B). We observed a single band corresponding to the size of GFP from cultures containing P_mra-gfp, indicating the native level of expression from the transcriptional reporter (Fig. 3B, lanes 1 and 2). In the mraZ⁺ background we could detect GFP in the minus inducer control; however, in the plus inducer condition GFP was below the detectable range (Fig. 3B, lanes 3 and 4). This indicates that overproduction of MraZ leads to strong repression of the mra promoter. We next tested whether the overproduction of either of the DXXXR mutants (R15A or R86A) disrupted the ability to repress the promoter of mraZ. Our results indicate that neither R15A nor R86A is able to repress P_mra-gfp (Fig. 3B, lanes 5 to 8). Thus, MraZ-mediated repression of the mra promoter is dependent on its ability to bind DNA through both DXXXR motifs.

We were interested in determining whether the MraZ-dependent repression of the mra promoter was dependent or independent of the MBRs. To this end we constructed a variation of the fusion in which the −35/–10 region of P_mra was switched with that of gpsB—a constitutively expressed cell division gene that is not part of the dcw cluster (Fig. 3A; P_mra(ΔMBR)). Additionally, we mutated each of the MBRs (P_mra(MBR*)). We introduced each of these fusions into WT or mraZ⁺ strain backgrounds. In these backgrounds we then overexpressed mraZ and western blotted the resulting cultures. We found that overexpression of mraZ did not result in repression of P_mra in the absence of intact MBRs (Fig. 3C). Therefore, these results confirm that the MBRs are a requirement for MraZ-mediated repression of the mra operon.

While the triple repeat is found only within P_mra, GTGG[A/T]G is a common sequence within the genome, and we sought to identify whether MraZ transcriptionally regulates any other genes in B. subtilis through RNA-Seq analysis. We were particularly interested, as MraZ-GFP appears to bind promiscuously throughout the chromosome (Fig. 2B). RNA was extracted from B. subtilis WT and cells overproducing MraZ and MraZ^R15A and, additionally, from cells lacking mraZ. Pooled RNA samples of each biological triplicate were then submitted for RNA-Seq and were sequenced twice as technical replicates. We identified 766 differentially regulated genes when mraZ was overexpressed (Fig. 3F). Through comparison of the data set obtained when the inactive MraZ^R15A was overproduced, we were able to eliminate some of these nonspecific changes in gene expression (Fig. 3F). We then compared the remaining 682 differentially regulated genes against those that were up- or downregulated in the ΔmraZ background. This resulted in a set of 38 genes; however, on further analysis, 34 of those 38 genes were found to be upregulated when mraZ is overexpressed as well as when mraZ is deleted, making it unlikely that these genes are part of the MraZ regulon (File S1). Three of the remaining four differentially regulated genes were part of the mra operon (mraW, ftsL, and pbpB) and are downregulated when mraZ is overexpressed and upregulated when mraZ is deleted (Fig. 3D and E). Additionally, we observed that mraZ overexpression results in a decrease in the mRNA level of pksJ, which encodes a polyketide synthetase, and in the ΔmraZ strain level of pksJ mRNA is increased (File S1). However, whether MraZ directly regulates this gene and the physiological reasons for the change in expression under these conditions are yet to be elucidated. Overall, based on our analysis, it appears that MraZ primarily acts as an autoregulatory transcriptional repressor of the four-gene operon (mraZ-mraW-ftsL-pbpB).

Repression of ftsL by MraZ drives cell division arrest.

Two of the four genes in the mra operon (ftsL and pbpB) are known to be essential in B. subtilis (10, 26), and we sought to identify if either of these genes was responsible for the filamentation we observed when MraZ was overproduced. To that end, we uncoupled the expression of mraW, ftsL, and pbpB from MraZ control by placing each of these genes individually under an IPTG-inducible promoter. We then expressed mraZ with each of the other genes in the operon to determine if there was any resolution to the filamentation we previously observed. We first confirmed that mra repression was occurring at all IPTG concentrations tested to identify the lowest concentration that resulted in filamentation (Fig. S1B to E). Based on our results, we chose 25 μM IPTG for this set of experiments.

When mraW is uncoupled from MraZ repression, cells are still filamentous (11.6 μm ±5.8 μm), and exhibited similar cell length compared to overexpression of mraZ alone (10.8 μm ±6.0 μm) (Fig. 4A and B). This result is in contrast to what was previously observed in E. coli, where coexpression of mraZ and mraW results in a rescue of the lethal phenotype (12). When pbpB is overexpressed with mraZ, the cells are again filamentous and similar in length to mraZ overexpression (12.0 μm ± 6.0 μm). This result suggests that although pbpB is an essential gene, immediate cell division arrest may not be due to repression of its expression, as PBP2B appears to be stable (27). To further confirm the expression of mraW and pbpB in our system, we tagged both genes to gfp. While MraW-GFP displayed diffused localization and GFP-PBP2B exhibited division site localization (28), neither of them was able to restore normal cell division (Fig. S3B to D). In contrast, when we uncoupled expression of ftsL from MraZ control, we observed cells that were much shorter (5.7 μm ± 2.1 μm) and closer to wild type in length (3.9 μm ± 1.0 μm). Thus, it appears that the repression of ftsL is the likely cause of immediate cell division arrest in MraZ-overproducing cells, as FtsL is rapidly turned over (18, 27, 29–32).

FIG 4.

Repression of ftsL by MraZ drives cell division arrest. (A) Fluorescence micrographs of WT (PY79) B. subtilis (i) and cells expressing mraZ⁺ (MW293) (ii), mraW⁺ (MW379) (iii), ftsL⁺ (MW330) (v), or pbpB⁺ (BAH1) (vii) and of cells coexpressing mraZ⁺/mraW⁺ (MW387) (iv), mraZ⁺/ftsL⁺ (MW333) (vi), and mraZ⁺/pbpB⁺ (BAH4) (viii) in the presence of 25 μM IPTG. Cultures were induced for 2 h prior to imaging. Membrane is visualized with SynaptoRed; Scale bar = 1 μm. (B) Quantification of cell length from microscopy in panel B. n = 100; ****, P < 0.0001, **, P = 0.06, *, P = 0.0264. (C) Time-lapse microscopy of cultures containing mraZ⁺ harboring ftsZ-gfp (MW205) in the absence or presence of IPTG. Images were taken every 15 min for 2 h. Yellow arrows indicate constriction-efficient FtsZ rings (or sites where they used to be), and white arrows follow newly assembled FtsZ rings. Scale bar = 1 μm.

Depletion of FtsL results in decondensation of the FtsZ ring.

Knowing that decoupling of ftsL expression relieves filamentation when MraZ is overproduced, we wanted to study the immediate effects of FtsL depletion on divisome assembly. To investigate this, we utilized a strain containing ftsZ-gfp (33) to track FtsZ dynamics when MraZ is in excess. Using time-lapse microscopy, we monitored the localization pattern of FtsZ-GFP at 15-min intervals in the mraZ⁺ strain in the absence and presence of IPTG. In the absence of IPTG, cells progress through the division cycle normally (Fig. 4C; Movie S1). However, after the addition of IPTG, cells can be observed dividing regularly, with the Z-ring forming and constricting initially. After 60 min, Z-rings that have formed but have not constricted begin to decondense. By 75 min after induction, multiple Z-rings can be seen immediately adjacent to one another (Fig. 4C; Movie S2), suggesting a likely impairment of the formation of functional Z-rings that are capable of undergoing constriction.

To further confirm that this effect was specifically due to a depletion of FtsL, we generated a strain that contains a CRISPR/CASi-based inducible-knockdown of ftsL (34). We ensured that filamentation occurs upon FtsL depletion in this strain background (Fig. S1F and G), as noted previously (34). We then introduced mNeonGreen-tagged ftsZ expressed from the native locus of ftsZ (35) to this FtsL depletion strain. Monitoring the localization dynamics of mNeonGreen-FtsZ upon FtsL depletion also revealed the impairment in Z-ring constriction as the assembled Z-rings appear to unravel (Fig. S3E and Movie S3). Of the 52 Z-rings monitored, 29 of them were immature (did not form a discrete Z-ring) and never reached maturity, and the remaining 23 appeared to be complete Z-rings but disassembled soon after and never matured (Fig. 5D). However, of the 20 Z-rings that displayed signs of constriction (diameter of the Z-ring shorter than the diameter of the short axis of the cell), all of them successfully completed septum formation (similar to recently proposed models [36, 37]). Given this evidence, we speculate that the main role of FtsL is in the formation of constriction-capable Z-ring assembly.

FIG 5.

Roles of MraZ and FtsL in controlling cell division. (A) Genes of the mra operon and the respective functions of their protein products. (B) FtsL is an intrinsically unstable protein that serves as a linchpin for the constriction-efficient divisome assembly. To achieve this, FtsL directly interacts with PBP2B (of the FtsW/PBP2B-SEDS/class B PBP complex), DivIC, DivIB, and perhaps also some FtsZ-associated proteins (ZAPs) either directly or indirectly. (C) Lethal phenotype of mraZ overexpression stems (at least initially) from the depletion of FtsL, which results in the lack of mature constriction-capable FtsZ (shown in green) ring formation. Additional supply of ftsL results in restoration of cell length that is similar to that of WT. (D) Depletion of FtsL (indicated by red arrows), depending on the stage of Z-ring assembly, leads to impairment of Z-ring maturation (top) promotes Z-ring disassembly (middle), or has no effect on Z-ring constriction (bottom).

Next, we used GFP-tagged DivIVA, a protein known to localize to concave intracellular features such as division sites and cell poles (38–40), as a proxy to study the status of septum construction in mraZ⁺ cells. For the WT, DivIVA-GFP is predominantly associated with division sites (Fig. S4A and B). However, in cells overexpressing mraZ, DivIVA-GFP remains mainly at the cell poles due to lack of cell division. Localization of DivIVA-GFP was minimal at nascent division sites as indicated by enriched membrane stain, suggesting that perhaps FtsL action (initiation of septal cell wall synthesis) is needed to effectively recruit DivIVA to nascent division sites. To further evaluate the precise role of FtsL in Z-ring constriction and initiation of peptidoglycan synthesis, we utilized fluorescein-d-lysine, a type of fluorescent d-amino acid (FDAA) used to monitor de novo peptidoglycan insertion activity (41). In the WT, FDAA labeling was present primarily at division sites and occasionally at poles, as reported previously (42–44). However, in mraZ⁺ cells (depletion of FtsL), FDAA labeling is patchy, and its incorporation at division sites appears diffused, in striking contrast to our WT control (Fig. S4C and D). This suggests that FtsZ-treadmilling-associated peptidoglycan insertion is occurring (44), but not in a focused way in the absence of FtsL, as Z-rings are unable to coalesce.

Additionally, we investigated the effect of deleting mraZ on cell division, as in this strain background expression of ftsL (and pbpB) is unregulated. Upon imaging cells lacking mraZ, we observed that, on average, these cells are smaller than the WT and appear very similar to cells overproducing FtsL (Fig. S3F and G), indicating hyperactivation of cytokinesis. This phenotype has previously been observed following deletion of the RasP protease, which is known to facilitate FtsL turnover (30). We did not observe a similar short-cell phenotype in the pbpB⁺ strain (Fig. 4AB). This set of data suggests that focusing Z-ring treadmilling and kickstarting the septal peptidoglycan synthesis may be the bottleneck of septum synthesis and that the level of FtsL acts as a critical timing device for septation, as suggested previously (30).

DISCUSSION

Cell division is a highly orchestrated and complex process, the timing of which must be tightly regulated. Numerous signals feed into the decision to divide, including, population density, nutrient availability, and the status of the chromosome. Previous reports have shown that MraZ is an important transcriptional regulator; indeed, in some species such as E. coli, MraZ regulates at least the first 9 genes in the dcw cluster and may be responsible for controlling transcription as far downstream as FtsZ (5, 45–47). Among the Gram-positive Actinobacteria phylum, work with Corynebacterium glutamicum has shown that mraZ mRNA is degraded by RNase III and that loss of rnc leads to accumulation of mraZ mRNA and, therefore, increased cellular levels of MraZ. This results in cell elongation through MraZ repression of ftsEX; however, whether MraZ in C. glutamicum is a transcriptional repressor of the mra operon is yet to be elucidated (48). Upon search of B. subtilis MraZ binding motifs in other organisms, we noted the high level of conservation in species within and outside the Firmicute phylum (Fig. S3A and Table S2B).

While the structure and function of MraZ and its multimeric crystal structure are fairly well characterized, at least in certain species (12, 49, 50), the role of its syntenous partner, MraW, remains unclear. MraW is predicted to be a 16S rRNA methyltransferase (N⁴ cytosine C1402). It has been reported that chloroplast MraW and 16S rRNA methylation may play a role in ribosome levels (51). Work with E. coli suggests that MraW may be able to regulate codon utilization (52) and possibly act as a transcriptional regulator through methylation of DNA (14). Additionally, results in S. aureus suggests that MraW may play a role in virulence (53). While E. coli MraZ and MraW appear to be regulated by each other, we were unable to identify a similar relationship in B. subtilis (12). Curiously, through our RNA-Seq analysis we were able to identify an upregulation of genes encoding ribosomal proteins when mraZ is deleted—conditions under which levels of MraW are likely increased (File S1). However, the biological relevance of this change remains to be elucidated.

In this report, we show that similar to previous studies conducted with other organisms, MraZ is a transcriptional regulator in B. subtilis (11–13, 16). Specifically, it is important in repressing the expression of two essential cell division genes, pbpB and ftsL, in addition to the nonessential mraW gene described above, which does not appear to have a direct role in cell division (10) (Fig. 5A). Although the essential nature of FtsL has been well characterized, the precise role of FtsL in the divisome complex remains unclear in B. subtilis. Studies of E. coli have elucidated that FtsL is involved in initiating membrane invagination and septal peptidoglycan synthesis that accompany Z-ring constriction under the direction of FtsN (54–57). However, FtsN is absent in B. subtilis (58). As shown in Fig. 5B, in B. subtilis, FtsL forms a complex with PBP2B (class B PBP; E. coli PBP3/FtsI homolog) and DivIC (FtsL paralog; E. coli FtsB homolog), and it is known that PBP2B also interacts with DivIB (E. coli FtsQ homolog) and FtsW (shape, elongation, division, and sporulation [SEDS] family glycosyltransferase) (59–65). It is noteworthy that although PBP2B is essential, its catalytic function is not (66). Thus, the essentiality of PBP2B comes from its scaffolding role. Based on our studies, we reveal that the FtsL level is integral for constriction-efficient FtsZ ring assembly (Fig. 5C). Upon MraZ overproduction (in the absence of enough FtsL) or when FtsL is specifically depleted using a CRISPR/CASi-based knockdown approach, we show that the Z-rings that have coalesced are unable to retain their structure and disassemble (Fig. 5D). This is reminiscent of what was noted when FtsZ-associated proteins (ZAPs) were absent (44). Similarly, defective Z-ring assembly has been noted when FtsW/PBP1 (homolog of B. subtilis PBP2B) levels are synthetically lowered in S. aureus (67). Perhaps the role of FtsL is to stabilize the FtsW/PBP2B complex and is conserved across multiple species. Absence of EzrA (one of the FtsZ membrane anchors) in a B. subtilis strain producing less FtsL is lethal, and overexpression of ftsL restores delayed FtsZ ring constriction in cells lacking ezrA (68). Thus, the FtsL complex may communicate with ZAPs and transduce that signal to concentrate the FtsZ treadmilling process (35, 36, 69, 70), similar to what has been reported in E. coli (54–56, 71), to kickstart cell division. However, how focusing of FtsZ treadmilling is triggered by the optimal level of FtsL in the absence of FtsN-like proteins remains to be investigated.

Among the divisome components, FtsL has emerged as a key factor to regulate cell division in a rapid manner (29, 30). FtsL is a transmembrane protein that is an essential part of the divisome, loss of which causes extreme filamentation in both E. coli and B. subtilis (31, 72). Previous studies have shown that FtsL is intrinsically unstable (73) and is rapidly turned over by the membrane protease RasP (18, 29, 30, 74). At elevated growth temperature, DivIB has been shown to protect FtsL from rapid degradation (18), and a strong complex between DivIC and FtsL has also been suggested to aid in the stabilization of FtsL (29, 59, 60). It was previously speculated that accumulation of FtsL at the divisome may be a rate-determining step (30). The results we report here further validate the previous findings that the level of FtsL is critical for cell division. This unstable nature of FtsL, combined with its essentiality, makes it an attractive control point to arrest cell division. In fact, during the DNA-damage response the known cell division inhibitor, YneA, appears to halt division by interacting with late-arriving divisomal proteins, including FtsL and PBP2B, in B. subtilis (75). Interestingly, YneA-mediated cell division inhibition could be rescued by ftsL overexpression (68). Similarly, another study discovered that during phage SP01 infection, a phage protein gp56 inhibits B. subtilis cell division by possibly directly interacting with FtsL (76). In this report, we show how in addition to posttranslational regulation of FtsL to regulate cell division, transcriptional regulation of the mra operon by MraZ could also be an efficient way to halt cell division due to the intrinsic instability of FtsL. Indeed, it has been proposed that the mra operon may be directly regulated by DNA replication initiation protein, DnaA, upon inhibition of DNA replication (32). In further support of this notion of transcriptional inhibition of cell division, it appears that during entry into stationary phase the transcriptional activity of mraZ significantly increases and that of the genes in the rest of the operon, mraW, ftsL, and pbpB, drops, presumably to eventually stop division (10) (Fig. S4E).

MATERIALS AND METHODS

Strain construction and general methods.

All B. subtilis strains utilized in this study are derivatives of PY79 (77) and were constructed via double recombination of circular plasmids into the chromosome. Information regarding strains and oligonucleotides/geneblocks can be found in Table S1. All plasmids were constructed using standard cloning protocols. Plasmids were transformed into the parental strain directly and integration confirmed via screening of the amyE locus for pDR111 (D. Rudner) based plasmids or the thrC locus for pDG1664 based plasmids. Primers oP4/oP5 were used for sequencing pDG1664-based plasmids, and oP209/oP210 were used for pDR111 plasmids. For generation of strains that require two separate genes of interest to be integrated into the amyE locus, a background containing a second synthetic amyE locus was used (AHB286: bkdB::TB917::amyE::catR; A. Camp).

(i) mraZ⁺. mraZ was amplified from the PY79 chromosome using primer pair oMW114/oMW115; the resulting fragment was then digested with HindIII and NheI and ligated into pDR111, an IPTG-inducible vector for integration at the amyE locus in B. subtilis. The resulting plasmid, pMW35, was then transformed into B. subtilis, producing MW189.

(ii) mraZ^R15A/mraZ^R86A. mraZ was amplified with oMW114/oMW115 from a geneblock (Integrated DNA Technologies) that contained either the R15A (gMW4) or R86A (gMW5) mutation, before being digested and ligated into pDR111 as for MW189. The plasmids pMW49 (R15A) and pMW65 (R86A) were then transformed into B. subtilis PY79, producing strains MW256 and MW350, respectively.

(iii) mraZ-gfp. mraZ was amplified with oMW114/oMW172, the resulting product was then digested with HindIII and BamHI, and linker-gfp was amplified from geneblock gP1 with primers oMW173/oMW121; the fragment was then digested with BamHI and NheI before being ligated into pDR111. This generated plasmid pMW59, which was transformed into B. subtilis, generating strain MW295. Tagged point mutants (mraZ^R15A-gfp and mraZ^R86A-gfp) were constructed as described above with mraZ amplified from gMW4 or gMW5. The resulting plasmids pMW58 (R15A) and pMW66 (R86A) were then used to transform B. subtilis, generating strains MW296 and MW351, respectively.

(iv) P_mraZ-gfp. Promoter fusions of mraZ were constructed in the thrC integration plasmid pDG1664 (78). The promoter region of mraZ (500 bp upstream of the mraZ start codon) was amplified with oMW177/oMW178 from chromosomal DNA and gfp with oMW184/oMW185. The resulting fragments were digested with BamHI/XhoI and XhoI/EcoRI, respectively, and then ligated into pDG1664, producing plasmid pMW68. Another fusion that lacked the MraZ binding repeats was made by amplifying the region containing the sequence upstream of the MBRs with oMW177/oMW252. This was then digested with BamHI/XhoI and ligated with gfp that was amplified with two rounds of PCR with oMW253 (oMW254-R2)/oMW185 to add on the −35/–10 region from gpsB and digested with XhoI/EcoRI, which was then ligated into pDG1664, producing plasmid pMW81. To generate a version of the P_mra promoter where the MBRs were mutated (GTGGAGCGAAGTGGTGAATAGTGGTG were mutated to GatcAGCGAAcacGTGAATAGatcTG), we amplified gMW11, which contains alterations to all three binding repeats, while preserving the −10 region with oMW177/oMW304 and digested with BamHI/XhoI and ligated as before, creating plasmid pMW104. These plasmids were used to transform PY79, generating MW368, MW452, and MW523, respectively. pMW35 was then used to transform these strains, generating MW385, MW478, and MW525. Additionally, pMW49 and pMW65 were used to transform MW368, resulting in MW389 and MW429, respectively.

(v) ftsL⁺/mraW⁺/pbpB⁺. Inducible ftsL, mraW, and pbpB were constructed by amplifying the respective genes from PY79 chromosomal DNA with primer pairs oMW127/oMW128, oMW122/oMW123, and oMW275/oMW276; the resulting fragments were digested with SalI and NheI before being ligated into pDR111. This created plasmids pMW39 (ftsL⁺), pMW38 (mraW⁺), and pAH1 (pbpB⁺), which were then used to transform AHB286, which contains a second synthetic amyE locus, producing strains MW330, MW379, and BAH1, respectively. To construct strains that additionally have inducible mraZ, MW189 was transformed with pDAG32 (79) to switch the resistance cassette to catR, generating strain MW293. The chromosomal DNA from this strain was then used to transform MW330, MW379, and BAH1, resulting in strains MW387, MW333, and BAH4, respectively. pMW39 was additionally used to transform PY79 to produce MW207.

(vi) mraW-gfp. Inducible mraW-gfp was constructed by amplifying mraW with oMW122/oMW278 from PY79 DNA, and the resulting fragment was digested using SalI/NheI. gfp was amplified with the primer pair oP258/oP259, and the fragment was digested with NheI/SphI. The digested fragments were then ligated into pDR111, generating plasmid pAH5. This plasmid was then used to transform AHB286, generating strain BAH17. To construct a strain that has an additional inducible mraZ, DNA from MW293 was used to transform BAH17, creating strain BAH18.

(vii) gfp-pbpB. Genomic DNA from 1A1111 (28) from the Bacillus Genetic Stock Centre (BGSC), which contains a xylose-inducible copy of gfp-pbpB^1-825 at the native locus, was used to transform PY79, generating strain MW199. Additionally, the same genomic DNA was also used to transform MW189, producing strain MW195.

(viii) ΔmraZ. Chromosomal DNA from BGSC strain BKK15130 (80) was used to transform PY79 to generate MW192 (mraZ::kanR).

(ix) mraZ⁺ ftsZ-gfp. AD3007 (33), a strain which has an additional copy of ftsZ-gfp at the native locus, was transformed with pMW35, resulting in strain MW205.

(x) ftsL knockdown. To construct an ftsL knockdown strain, DNA from BEC15150 (BGSC [34)] was transformed into PY79, generating strain BAH16. To visualize the Z-ring when FtsL is depleted, bAB185 (35), a strain harboring a markerless copy of ftsZ fused to mNeonGreen at the native locus was transformed with genomic DNA of BEC15150, resulting in strain BAH9.

(xi) divIVA-gfp. pKR196 (38) was transformed into the second amyE strain AHB286, producing strain MW520. The chromosomal DNA from MW293 was then used to transform MW520, resulting in strain MW522.

Media and culture conditions.

Unless otherwise stated, overnight cultures of B. subtilis strains were grown at 22°C in lysogeny broth (LB) and diluted 1:10 in fresh LB before being grown at 37°C to the mid-log phase (OD₆₀₀, 0.5 to 0.8); cultures were then standardized to an OD₆₀₀ of 0.1 in fresh LB. Where induction of genes under IPTG required, unless otherwise stated, a final concentration of 1 mM IPTG was added when cultures were standardized, and the cultures were then grown for 2 h at 37°C. Where induction of genes under xylose control was required, a final concentration of 1% xylose was used unless otherwise stated.

Spot assays.

All spot assays were carried out similarly to those previously described (81). Briefly, strains were grown in liquid culture at 37°C with shaking until the mid-log phase (OD₆₀₀, 0.5 to 0.8) before being standardized to an OD₆₀₀ of 0.1. Standardized cultures were then serially diluted before 1 μL of each serial dilution was spotted onto either lysogeny agar (LA) or LA supplemented with 1 mM IPTG when required to induce expression of genes under IPTG control. Plates were incubated for approximately 14 h at 37°C before being observed for any growth defects.

Growth curves.

Strains to be analyzed by growth curve were grown to the mid-log phase (OD₆₀₀, 0.5 to 0.8) in liquid culture at 37°C with agitation and were then standardized to an OD₆₀₀ of 0.1 in LB. IPTG was added to a final concentration of 1 mM where required. An aliquot of 250 μL of each standardized culture was added in triplicate to a 96-well plate, which was then incubated at 37°C with agitation for 12 h in a Synergy H1 plate reader (BioTek). OD₆₀₀ readings were taken every hour, and growth curves were plotted using GraphPad Prism 9.

Microscopy.

Microscopy was carried out as previously described (82). Briefly, 500-μL aliquots of B. subtilis cultures were pelleted and then washed with 1× phosphate-buffered saline (PBS) via centrifugation. Pellets were then resuspended in 100 μL of PBS and were stained with SynaptoRed to stain the membrane at a final concentration of 1 μg/mL and with DAPI (4′,6-diamidino-2-phenylindole) to stain the DNA; a final concentration of 1 μg/mL. 5 μL of the stained cells was spotted onto the base of a glass-bottom dish (MatTek), and a 1% agarose pad made with PBS was placed gently on top. Imaging was carried out at room temperature using a DeltaVision Elite deconvolution fluorescence microscope with photos taken using a Photometrics CoolSnap HQ2 camera. All images were acquired by taking 17 Z-stacks at 200-nm intervals. Images were deconvolved though the SoftWorx imaging software provided by the microscope manufacturer. Analysis of cell lengths was done through ImageJ or SoftWorx. Statistical analysis such as one-way analysis of variance (ANOVA) and multiple-comparison or Student’s t test was carried out using GraphPad Prism 9.

Time-lapse microscopy.

As described previously, strains MW205 and BAH9 were grown to the mid-log phase (OD, 0.5 to 0.8) before 5 μL was aliquoted onto a MatTek dish and covered with a 1% agarose pad made with LB (82). Samples were allowed to adjust to the microscope chamber at 30°C for a period of 30 min before 10 μL of 10 mM IPTG (MW205) or 10 μL of 10% xylose (BAH9) was added to the agarose pad for the plus inducer conditions. Time-lapse imaging was immediately begun, and images were taken every 15 min for 2 h with 5 Z-stacks at 200-nm intervals. Image processing was carried out as described above.

Fluorescent d-amino acid labeling.

Fluorescein-d-lysine (FDL) was generated as previously described (41). An aliquot of 500 μL of B. subtilis cultures in mid-log phase were pelleted before being resuspended in 100 μL LB. A final concentration of 1 mM FDL was added, and the samples were incubated at 37°C with gentle shaking for 5 min. The reaction was then quenched with ethanol, and the cells were washed in 1× PBS before being prepared for imaging as described above.

Immunoblots.

Cultures were grown as previously described, and 1-mL aliquots standardized to an OD₆₀₀ of 1 were taken. Cells were pelleted and resuspended in 500 μL of protoplast buffer containing 0.5 M sucrose, 20 mM MgCl₂, 10 mM KH₂PO₄, and 0.1 mg/mL lysozyme. Samples were then incubated for 30 min at 37°C and prepared for SDS-PAGE. For analysis of P_mra activity across different growth phases, a 20-mL culture was standardized to an OD₆₀₀ of 0.1, and samples were taken every hour for 4 h. Each sample was standardized to an OD₆₀₀ of 1 in 1 mL and then prepared as previously described. Where required, MF339 cell lysate served as a control for free GFP (83). Following electrophoresis, samples were transferred to nitrocellulose membrane using the iBlot 2 transfer system (Thermo Fisher) and probed with antibodies against GFP (Kumaran Ramamurthi) and B. subtilis SigA (lab stock).

Bioinformatics.

Using the consensus sequence, blastn was used to find MBRs in B. subtilis by utilizing the PY79 whole-genome sequence (CP006881.1) as the query and the search string GTGGWGNNNNGTGGWGNNNNGTGGWG. Using algorithm parameters optimized for short nucleotide repeats with spaces (expect threshold 100,000, word size 7, match/mismatch cost 1,-1, gap cost existence 0, extension 2, no filters or masks). To generate a sequence logo (https://weblogo.berkeley.edu/; 84), sequences upstream of mraZ from the following species were used: B. subtilis, Bacillus cereus, Staphylococcus aureus, Staphylococcus epidermidis, Streptococcus pneumoniae, Enterococcus faecium, Lactococcus lactis, and Listeria monocytogenes. Visual examination of the sequences using the previously identified consensus sequence for B. subtilis mraZ binding site GTGG revealed the existence of ordered repeats containing the motif GTGGNNNNNAGTGGNGNNNNGTGG (11). The sequences can be found in Table S2B and the multiple-sequence alignment generated with the help of Clustal Omega (85) is shown in Fig. S3A. In addition, using the probe GTGGNNNNNNGTGG, Pattern Locator (25) was used to search the PY79 genome for potential MraZ binding repeats (Table S2A).

RNA-Seq.

Cultures were grown as previously described and treated with RNAprotect bacterium reagent (Qiagen) before RNA was extracted utilizing the RNeasy minikit (Qiagen). Samples were sent to the SeqCenter (Microbial Genome Sequencing Center, Pittsburgh, PA) for 12M paired-end sequencing and analysis. GraphPad Prism 9 was used to generate a volcano plot.

Data availability.

The data sets generated for this study are available in the Gene Expression Omnibus (GEO) repository (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE206572).