Abstract
Bacterial growth and division require insertion of new peptidoglycan (PG) into the existing cell wall by PG synthase enzymes. Emerging evidence suggests that many PG synthases require activation to function, however it is unclear how activation of division-specific PG synthases occurs. The FtsZ cytoskeleton has been implicated as a regulator of PG synthesis during division, but the mechanisms through which it acts are unknown. Here we show that FzlA, an essential regulator of constriction in Caulobacter crescentus, links FtsZ to PG synthesis to promote division. We find that hyperactive mutants of the PG synthases FtsW and FtsI specifically render fzlA, but not other division genes, non-essential. However, FzlA is still required to maintain proper constriction rate and efficiency in a hyperactive PG synthase background. Intriguingly, loss of fzlA in the presence of hyperactivated FtsWI causes cells to rotate about the division plane during constriction and sensitizes cells to cell wall-specific antibiotics. We demonstrate that FzlA-dependent signaling to division-specific PG synthesis is conserved in another α-proteobacterium, Agrobacterium tumefaciens. These data establish that FzlA links FtsZ to cell wall remodeling, serving both to activate and spatially orient PG synthesis during division. Overall, our findings support the paradigm that activation of SEDS-PBP PG synthases is a broadly conserved requirement for bacterial morphogenesis.
Bacterial division is driven by the insertion of new cell wall material at midcell in a tightly regulated manner, allowing for determination of cell shape and maintenance of envelope integrity1,2. The cell wall is made of peptidoglycan (PG), a meshwork consisting of glycan strands crosslinked by peptide stems3,4. PG synthesis requires the coordination of glycan polymerization and peptide crosslinking by either coupled monofunctional glycosyltransferases (GTases) and transpeptidases (TPases), or bifunctional enzymes that contain both activities, with these proteins being more generally referred to as PG synthases1.
Monofunctional PG synthase pairs have been implicated as the primary synthetic enzymes of the elongation (elongasome) and division (divisome) machineries. A paradigm has been proposed whereby a shape, elongation, division, and sporulation (SEDS) family GTase is functionally coupled to a penicillin binding protein (PBP) TPase, which together facilitate cell wall synthesis5–7. Through characterization of the elongation-specific PG synthases RodA and PBP2 in Escherichia coli, it has been postulated that SEDS-PBP enzymes require activation to function5. Specifically, mutations in RodA or PBP2 that increase GTase activity in vitro and PG synthesis in cells render other components of the elongasome non-essential, arguing that their normal function is to activate the RodA-PBP2 complex5. Intriguingly, analogous mutations in the division-specific SEDS-PBP enzymes, FtsW and FtsI, allow cells to constrict faster than normal8, suggesting that these mutations promote formation of an activated PG synthase complex5,9. However, it is unclear precisely how SEDS-PBP activation normally occurs during division.
Recent studies have established that the conserved cytoskeletal protein FtsZ10,11, which recruits the division machinery to a ring-like structure at midcell12–14, is coupled to PG synthesis activation during division. In multiple organisms, the C-terminal linker domain of FtsZ was found to be required for regulating cell wall integrity15–17 and shape, as well as PG chemistry16,18. Moreover, in E. coli and Bacillus subtilis, FtsZ dynamics were demonstrated to drive PG synthase dynamics in both organisms, as well as division site shape in E. coli and constriction rate in B. subtilis19,20. Collectively these data indicate that, at least in some organisms, FtsZ acts as a “dynamic scaffold” or “dynamic activator” of PG synthesis likely impinging on FtsWI. However, the signaling pathway connecting these two endpoints remains unresolved.
We previously demonstrated that an essential FtsZ-binding protein, FzlA21, is required for division and regulates the rate of constriction in the α-proteobacterium Caulobacter crescentus22. Mutations in FzlA with diminished affinity for FtsZ were found to have slower constriction rates and altered cell pole shape, indicative of reduced PG synthetic activity during division22. We therefore postulated that FzlA facilitates a link between FtsZ and PG synthesis by serving as an upstream activator of PG synthases and, here, set out to test this hypothesis.
Results
fzlA lies upstream of ftsWI in a PG synthesis pathway
We reasoned that if FzlA impacts constriction through PG synthases, it likely acts on the division-specific SEDS family GTase FtsW and/or the monofunctional PBP TPase FtsI. To assess if FzlA activates FtsWI, we leveraged fast-constricting strains containing hyperactive mutant variants of FtsI and/or FtsW termed ftsW**I*8,9 and ftsW*9. ftsW**I* bears the mutations F145L and A246T in FtsW and I45V in FtsI, whereas ftsW* contains only the FtsW A246T mutation9. These mutations are thought to stabilize an activated form of the FtsWI complex5,9, leading to increased rates of cell constriction8 via unrestrained PG synthesis.
If fzlA lies upstream of ftsWI in a PG synthesis pathway, then the hyperactive variants ftsW**I* and/or ftsW* may bypass the essentiality of fzlA. Accordingly, we found that fzlA could be readily deleted in either the ftsW**I* or ftsW* strain backgrounds (Fig. 1A-B, Fig. S1A). This is a particularly striking finding given that depletion of FzlA in a WT background completely inhibits division and induces cell filamentation and death21. Interestingly, a number of ftsW**I*/ftsW* ΔfzlA cells appeared to be “S”-shaped with the direction of curvature in future daughter cells facing opposite directions, as opposed to the characteristic “C”-shape of pre-divisional WT and ftsW**I* Caulobacter cells (Fig. 1A, asterisk, discussed further below). Assessment of fitness revealed that strains lacking fzlA displayed a slight reduction in viability by spot dilution, compared to the corresponding hyperactive PG synthase mutant strains (Fig. 1C), whereas growth rate was unaffected (Fig. 1D). In addition, ftsW**I*/ftsW* ΔfzlA cells displayed an increase in length (Fig. 1E), indicative of a residual division defect. Because ftsW* ΔfzlA cells are longer than ftsW**I* ΔfzlA cells, we conclude that ftsW**I* suppresses loss of fzlA better than the single mutant.
We also observed that ftsW**I* suppresses length, width, and fitness defects associated with slowly constricting fzlA point mutants fzlANH2 and fzlANH3 (Fig. S2, Fig. S1B), further indicating that hyperactivated ftsWI are dominant to, and likely downstream of fzlA. To determine the contribution of the FtsZ-FzlA interaction to activation of FtsWI, we assessed cell morphology, fitness, and cell length of ftsW**I* strains containing FzlA mutants with decreasing affinity for FtsZ22 (Fig. S3, Fig. S1C; FzlA > FzlANH2 = FzlANH3 > FzlANH1; FzlANB2, FzlANB1 = no binding). We found that decreased affinity of FzlA towards FtsZ correlated with an increase in cell length (Fig. S3E), indicating that high-affinity binding to FtsZ is required for FzlA to signal to FtsWI.
FzlA plays a specific and unique role in activating FtsWI
To assess the specificity of the fzlA-ftsWI genetic interaction and potentially identify additional components of this pathway, we performed comparative transposon sequencing (Tn-Seq) on WT and ftsW**I* strains. Surprisingly, fzlA was the only essential gene to become non-essential in the ftsW**I* background, with few insertions in WT but plentiful insertions in ftsW**I* cells (Fig. 1F,G, Supplementary Table 1). All other known essential division genes, e.g. ftsZ (Fig. 1G), had few transposon insertions in either background. These data indicate that fzlA is specific and unique in its essential role upstream of ftsWI. We suspect that other essential division proteins participate in this pathway as well, but that they play additional essential functions in divisome assembly or activity.
FzlA contributes to efficient division in a hyperactive PG synthase background
Given that cells lacking fzlA in the hyperactive PG synthase backgrounds were elongated, we assessed constriction rate and division efficiency in these strains in more detail. Specifically, we performed time-lapse microscopy on ftsW**I* and ftsW* cells ± fzlA and tracked division in cells using MicrobeJ22,23 (Fig. 2A, Supplementary Video 1). Consistent with previous findings8, ftsW**I* and ftsW* cells constrict more quickly than WT (Fig. 2B). Intriguingly, the hyperactive PG synthase strains lacking fzlA constricted significantly more slowly than the corresponding strain with fzlA present, with constriction rates cut nearly in half (Fig. 2B). This suggests that hyperactivated FtsWI are not sufficient for efficient division and underscores the importance of FzlA in dictating constriction rate. As with cell length and fitness, ftsW**I* acted as a better suppressor to fzlA deletion, allowing for a faster constriction rate than did ftsW* (Fig. 2B).
To ensure that changes in constriction rate were not due to global differences in PG synthesis, we determined elongation rates across strains (Fig. 2C), which enabled calculation of the ratio of constriction to elongation rate (Fig. 2D). We saw the same trend as for constriction rate itself, with ftsWI** and ftsW* mutant strains having higher ratios of constriction to elongation and loss of fzlA giving lower ratios (Fig. 2D). Interestingly, elongation rate was inversely correlated with constriction rate in all mutant strains (Fig. 2C, Fig. 2D), perhaps reflecting competition between the elongasome and divisome for PG precursor substrate24. Altogether, these data support the conclusion that alterations to the ftsZ-fzlA-ftsWI pathway specifically affect constriction, with FzlA increasing the constriction rate in both WT and hyperactive PG synthase mutant backgrounds.
While tracking division in ftsW**I* ΔfzlA and ftsW* ΔfzlA cells to measure constriction, we noticed that some cells initiated constriction at one location, then aborted division at that location before successfully dividing at a second (or third or fourth) site (Fig. 2E, Supplementary Video 2). We quantified the frequency of such constriction failure events and found that 16.6-19.5% of the hyperactive PG synthase cells lacking fzlA aborted division at one site before successfully dividing at another, compared to a 0-0.3% failure rate for WT or hyperactive PG synthase cells with fzlA present (Fig. 2F). These data further demonstrate that ftsW**I* are not sufficient for efficient division, and that fzlA is required to ensure division processivity and efficiency.
fzlA is required for maintenance of proper cell shape
As mentioned earlier, deletion of fzlA in the hyperactive PG synthase backgrounds impacted global cell morphology, with many pre-divisional cells appearing “S-shaped”. In order to more carefully assess this phenotype, we imaged cells by scanning electron microscopy (SEM). We saw a relatively high frequency of S-shaped ftsW**I* ΔfzlA cells, whereas most WT or ftsW**I* cells displayed the typical “C-shaped” morphology characteristic of Caulobacter (Fig. 3A). We quantified the frequency of S-shaped cells in a population of dividing cells by phase contrast microscopy to assess penetrance of this morphological phenotype. We extracted outlines of individual cells and performed principal component analysis using Celltool to isolate variance in cell shape to features referred to as shape modes22,25. Shape mode 3 captured the variation due to degree of S-versus C-shape and we set a cutoff such that cells with a standard deviation |sd| > 1 from the mean for this shape mode are considered S-shaped (Fig. 3B,C). Means and medians were similar for degree of S-shape across populations, with no significant difference for means, and a statistically significant but numerically small difference for medians. However, there was an obvious and significant difference in variance in degree of S-shape across populations (Fig. 3B), corresponding with a large difference in the number of cells found to be S-shaped in different strains. Over a quarter (26.9%) of dividing ftsW**I* ΔfzlA cells displayed an S-shaped morphology, compared to 2.4% of WT and 1.1% of ftsW**I* cells that are S-shaped (Fig. 3D).
To shed light on the origin of S-shape, we next asked at what point during growth do ftsW**I* ΔfzlA cells begin to adopt this morphology. Using time-lapse microscopy, we observed that ftsW**I* ΔfzlA cells were C-shaped at the beginning of the cell cycle and began to twist or rotate about the division plane after constriction initiated. S-shape only became apparent in the latter part of constriction, when daughters had rotated ~180° relative to each other (Fig. 3E, Supplementary Video 3). This finding suggests that the fzlA-ftsWI pathway determines geometry of PG insertion at the site of division in a manner that influences global cell morphology, normally constraining cells in their characteristic C-shape as constriction progresses. These results also indicate that our quantification method for S-shape likely underestimates the number of twisted ftsW**I* ΔfzlA cells, since S-shape is not obvious by phase contrast until the end of constriction, and we quantified cell shape at all stages of the constriction process.
Changes in division site shape and formation of S-shaped cells have been previously linked to aberrant localization of FtsZ and the elongation factor, MreB, respectively19,26. To determine if cell twisting might be facilitated by mislocalized division or elongation machineries, we visualized FtsZ and MreB localization in ftsW**I* ΔfzlA cells using inducible fluorescent fusions of these proteins. However, mNG-FtsZ and Venus-MreB localization in ftsW**I* ΔfzlA cells was comparable to ftsW**I* cells (Fig. S4, S5). Additionally, we visualized the localization of PG synthesis using the fluorescent D-amino acid HADA27, in order to assess whether cell twisting might be induced by mislocalized PG synthesis in spite of properly localized FtsZ and MreB. However, we did not detect any gross changes in HADA localization (Fig. S6). Together, these findings suggest that cell twisting is likely induced by a finer scale alteration of PG synthesis at the division site due to disruption of the fzlA-ftsWI pathway.
To determine whether global shape regulation depends on the FtsZ-FzlA interaction, we assessed S-shaped cell frequency in strains containing mutants of FzlA displaying decreasing affinities towards FtsZ. We observed that affinity of the mutant FzlA for FtsZ was inversely correlated with the frequency of S-shaped cells (Fig. S7), verifying that the FtsZ-FzlA interaction is important for maintaining proper morphology.
The ftsZ-fzlA-ftsWI pathway contributes to resistance to PBP-targeting antibiotics
Because FzlA is important for regulation of PG synthesis in the context of determining constriction rate and cell shape, we hypothesized that it might also contribute to resistance to cell wall-targeting antibiotics. To test this, we challenged cells with antibiotics targeting PG synthetic processes and assessed resulting cell fitness. ftsW**I*, as has been previously shown9, displayed sensitivity to cephalexin (Fig. 4A), which inhibits FtsI and other penicillin-binding proteins in Caulobacter 28,29. Interestingly, deletion of fzlA in the ftsW**I* background exacerbated sensitivity to cephalexin (Fig. 4A). We found a similar trend upon treatment with mecillinam, which targets the elongation-specific PG synthase PBP230,31, whereby the minimum inhibitory concentration (MIC) for ftsW**I* cells was decreased compared to WT, with deletion of fzlA further lowering the MIC (Fig. 4B). We also treated cells with the β-lactam ampicillin32 and with the cell wall targeting antibiotics vancomycin, which blocks transpeptidation by a distinct mechanism from β-lactams33, and fosfomycin, which inhibits cell wall synthesis by blocking PG precursor availability34 (Fig. S8). Neither hyperactivation of ftsWI nor loss of fzlA yielded a change in MIC in the presence of any of these antibiotics (Fig. S8). fzlA therefore supports robust cell wall synthesis in the presence of certain PG-targeting drugs, perhaps by compensating for inactivation of specific PBPs. To determine if the interaction between FtsZ and FzlA is important for maintaining cell wall integrity, we assessed sensitivity to cephalexin using the panel of fzlA mutants which display varying affinities towards FtsZ (Fig. S9). We found that mutants with decreased FzlA affinity towards FtsZ in fact became more sensitive to cephalexin (Fig. S9), demonstrating that the entire ftsZ-fzlA-ftsWI pathway is required for promoting cell wall integrity during antibiotic treatment.
Since ftsW**I* cells are more sensitive to perturbation of other PG synthetic activities even when fzlA is present, we asked if any normally non-essential division genes become more important for fitness in an ftsW**I* background, as they might help bolster resistance to assaults on PG synthesis. Examination of the ftsW**I* Tn-Seq data indicated that pbpX (encoding a bifunctional PG synthase that localizes to midcell)35,36, and to a lesser extent ftsX (encoding a cell separation factor)37 and dipM (encoding an envelope maintenance/cell separation factor)37–40, had fewer transposon insertions in an ftsW**I* background than in WT (Fig. 1F, S10). Because ftsW**I* cells have misregulated division site PG synthase activity, we suspect that FtsEX, DipM, and PbpX become important for ensuring robust PG synthesis during constriction and, later, efficient cell separation. Surprisingly, the normally non-essential nhaA locus, coding for a putative sodium-proton antiporter41, was also predicted by Tn-Seq to become essential in ftsW**I* cells (Fig. 1F, Fig. S10B). Disruption of nhaA in the presence of sucrose has been shown to arrest division41, suggesting nhaA may be important for division under certain conditions. It is unclear why it also becomes important upon PG synthesis mis-regulation, but its role in osmoregulation may contribute to its apparent synthetic lethality with ftsW**I*.
The fzlA-ftsW genetic interaction is conserved in diverse α-proteobacteria
FzlA homologs are encoded in nearly all sequenced α-proteobacterial genomes, but not outside this group. To assess the conservation of FzlA’s role in regulating PG synthesis, we sought to characterize the genetic interaction between fzlA and PG synthases in another α-proteobacterium, Agrobacterium tumefaciens. A. tumefaciens and Caulobacter display disparate growth patterns driven by distinct machineries during elongation, with A. tumefaciens exhibiting polar elongation and Caulobacter elongating primarily at midcell and through dispersed growth 27,42–44. However, the components of the division machinery, including FtsZ, FzlA, and FtsWI, are largely conserved. To test if the genetic interaction between fzlA and ftsW is conserved, we made an IPTG-dependent FzlA depletion construct in a WT background or in a background with a single hyperactivating mutation in A. tumefaciens ftsW (F137L, the equivalent of Caulobacter FtsW F145L) at the ftsW locus. Depletion of FzlA in a WT background resulted in reduced viability, FtsW depletion in A. tumefaciens18. Importantly, we found that the decrease in viability division arrest, and ectopic pole formation at midcell (Fig. 5, Fig. S11), reminiscent of and morphology defects associated with depletion of FzlA were rescued by ftsW F137L (Fig. 5, Fig. S11). These data indicate that FzlA’s essential role in regulating division-specific PG synthesis is conserved in another α-proteobacterium and further highlight the importance of FzlA as a key regulator of constriction and cell morphology.
Discussion
Here we have described a conserved PG synthesis activation pathway in which FtsZ and FzlA signal through FtsWI to regulate wall synthesis during division in α-proteobacteria (Fig. 6, left panel). Specifically, the FtsZ-FzlA-FtsWI pathway determines geometry of cell wall insertion at the site of division, sets the constriction rate, and promotes cell wall integrity (Fig. 6, left panel). FtsW**I* can still receive input from FzlA which, in combination with their intrinsic hyperactivity, leads to shorter, faster-constricting cells with sensitivity to cell wall antibiotics (Fig. 6, middle panel). In the absence of fzlA, ftsW**I* cells lose critical regulation of PG synthesis, leading to twisting during division, slower constriction, and increased sensitivity to cell wall antibiotics. We establish FzlA as a key intermediary in signaling from FtsZ to FtsWI and demonstrate that this division-specific SEDS-PBP pair require activation for normal division. Notably, our observations indicate that FtsWI activity is regulated in multiple ways, likely including input into their catalytic rates and modulation of the fine-scale geometry by which they insert new PG for constriction.
Our findings provide the foundation for further mechanistic investigation into this pathway and raise a number of questions. For one, the nature and timing of the activation signal(s) are still unknown: is there a signal always emanating from FtsZ-FzlA that induces constriction as soon as FtsW arrives, or is constriction triggered by a discrete cellular event, such as clearance of the chromosomal termini or the arrival of a sparkplug factor that jumpstarts FtsWI activityŒ Additionally, it is unclear why hyperactivation of ftsWI together or ftsW alone can rescue loss of fzlA – are TPase and GTase activities impacted similarly, or does one predominate in regulating constriction rateŒ A recent described here in fact hyperactivates FtsW. Finally, we have no evidence that FzlA and study demonstrated that FtsW is activated by FtsI7, so it is possible that the ftsI* mutation FtsWI directly interact, and suspect that other intermediary factor(s) transduce the activation signal from FzlA to FtsWI.
Our model advances the idea that regulation of SEDS-PBP pairs for growth and division is conserved at numerous levels. The finding that FzlA governs division-specific PG synthesis in both Caulobacter and A. tumefaciens argues that α-proteobacteria use FzlA as a conserved and dedicated FtsWI activator. FzlA is absent outside of this clade, however, so we propose that other divisome components serve as FtsWI activators in other organisms. More broadly, our findings expand the paradigm for PG synthesis by SEDS-PBP PG synthase pairs in bacteria and provide evidence that the requirement for PG synthase activation is conserved. Elongation is facilitated by the coordination of the SEDS family GTase RodA and the monofunctional TPase PBP2, orthologs to FtsW and FtsI, respectively5. The proposed model for elongation activation, as described for E. coli, holds that these PG synthases are activated by another protein, MreC, forming an activated complex that in turn regulates assembly and directional motion of the polymerizing scaffold MreB5. In this system, hyperactivating mutations in RodA or PBP2 allow for bypass of the activator, MreC, similar to our finding that FtsW**I* can bypass the activator FzlA. Our data provide experimental support for the proposal that the requirement for activation of the SEDS-PBP pair of PG synthases is generally conserved for elongation and division.
There are prominent differences between the models for elongasome and divisome activation, however. The elongasome appears, in essence, to be a stripped down and when either RodA or PBP2 is hyperactivated through mutation, all elongasome version of the divisome45: the elongasome contains fewer proteins than the divisome45, components except MreB and the PG synthases are rendered dispensable5. This would suggest that for elongation, the cell needs an activated SEDS-PBP pair and a spatial regulator to orient their motion5. Conversely, hyperactive FtsWI in Caulobacter only allows for disruption of FzlA, with the rest of the divisome remaining essential. This may be because division is a more complex process than elongation, requiring invagination and fission of all layers of the envelope in coordination with DNA segregation and cell cycle progression. This complexity necessitates functions in addition to PG synthesis and remodeling provided by components of the divisome. PG synthesis during division likely requires more regulation, as well. Whereas PG synthesis during elongation comprises insertion of new PG in the same plane as old cell wall material, PG synthesis during constriction requires a lasting, directional change to shape the new cell poles. So while there are key similarities in the paradigm of PG synthase activation, regulation of division likely requires a more complicated network of inputs to manage the additional outputs and constraints discussed above. In summation, this work provides evidence that the requirement for SEDS-PBP activation is conserved across multiple modes of PG synthesis, which has broad implications for determining the speed of division, cell shape, and cell wall robustness.
Methods
Strains, growth conditions, and growth determination
Caulobacter crescentus strains were derived from the NA1000 WT strain. Unless otherwise indicated, Caulobacter colonies were isolated from solid 1.5% agar peptone yeast extract (PYE) plates grown at 30 °C and cells were grown in liquid culture in PYE shaking at 30 °C. Where indicated, Caulobacter cells were treated with 6 µg/ml of cephalexin. Antibiotic MIC analysis was performed using antibiotic test strips (Liofilchem), which include a concentration gradient of 0.016-256 mg/L for all antibiotics tested. Where indicated, cells were treated with 0.3% xylose to drive inducible gene expression. Spot dilutions for Caulobacter were performed by serially diluting cells at the indicated fraction (1/10 or 1/2), before plating. Growth rates were obtained by measuring optical density at 600 nm (OD600) values of cells every 30 minutes. Caulobacter cell synchrony was performed as previously described12. Briefly, log phase cells were washed with M2 salts (6.1 mM Na2HPO4, 3.9 mM KH2PO4, 9.3 mM NH4Cl)46, resuspended in 1:1 M2:Percoll, then centrifuged at 11,200 x g. The swarmer band was isolated, and cells were subsequently washed twice in M2, then resuspended in PYE.
A. tumefaciens were grown in A. tumefaciens glucose and (NH4)2SO4 (ATGN) minimal medium47, with 0.5% glucose at 28°C. E. coli strains were grown in LB medium at 37°C. IPTG was added at a concentration of 1 mM when necessary. For A. tumefaciens spot dilutions, cells were grown overnight in ATGN minimal medium in the presence of IPTG at 1 mM concentration, washed, then pre-depleted of IPTG for 16 hours where indicated. Cells were then serially diluted (ten-fold) and spotted on ATGN minimal medium with the presence or absence of IPTG. To make the ΔfzlA PlacfzlA strain (PBA199), first, a mini-Tn7 vector containing IPTG inducible fzlA, along with the pTNS3 helper plasmid, were introduced into DtetRA a-attTn7 cells (PBA44) via electroporation as previously described48. Deletion of fzlA (for PBA199) and allelic exchange of ftsW (for PBA232) were subsequently performed by transferring the corresponding suicide vector to A. tumefaciens via conjugation with E. coli S17.
Plasmids (Supplementary Table 2) and strains (Supplementary Table 3) used in this study can be found in the supplementary information. Plasmids were created using standard molecular cloning procedures including PCR, restriction digestion, and ligation. Mutagenesis of ftsW for A. tumefaciens was performed using a QuikChange Lightning Multi SiteDirected Mutagenesis Kit (Agilent Genomics), with primers designed using Agilent’s QuikChange Primer Design Program, as previously described22. pEG1345 was constructed using an NEBuilder HiFi DNA Assembly Cloning Kit (NEB).
Light microscopy imaging and analysis
Images of log phase Caulobacter cells were obtained using either phase contrast microscopy, with cells grown on either 1% agarose PYE pads or 1% agarose dH2O pads, or, when indicated, fluorescence microscopy, with cells grown on 1% agarose dH2O pads. For fluorescence microscopy, mNG-FtsZ expression was induced for 1 hour with xylose then imaged through the GFP filter and venus-MreB expression was induced for 2 hours with xylose then imaged through the YFP filter. For determination of PG incorporation localization, cells were pulsed with 0.82 mM HADA for 5 minutes, washed twice with PBS, then visualized through the DAPI filter. For time-lapse imaging, as previously described22, synchronized cells were placed on 1% agarose PYE pads and imaged using phase contrast microscopy at room temperature (RT), with images being acquired at 5 minute intervals at 100x. Imaging of Caulobacter cells was performed using a Nikon Eclipse Ti inverted microscope with a Nikon Plan Fluor × 100 (numeric aperture 1.30) oil Ph3 objective and Photometrics CoolSNAP HQ2 cooled CCD (chargecoupled device) camera22. For A. tumefaciens phase contrast microscopy, exponentially growing cells were spotted on 1% agarose ATGN pads as previously described49, then imaged. For A. tumefaciens time-lapse microscopy, images were collected every ten minutes. Microscopy of A. tumefaciens cells was performed with an inverted Nikon Eclipse TiE with a QImaging Rolera em-c2 1K EMCCD camera and Nikon Elements Imaging Software.
For determination of dimensions of log phase cells, cell length and width were measured using MicrobeJ software, similar to as previously described22. Constriction rate and elongation rate were also determined using MicrobeJ23. Briefly, MicrobeJ software allowed for tracking of cells imaged by time-lapse microscopy throughout the division process, with automatic detection of constriction initiation and manual determination of cell separation. Cell length was found for cells at each time point, cell width was found at the site of constriction, and constriction time was calculated by multiplying the number of frames in which constriction was detected by 5 (since images were acquired every 5 minutes), allowing for calculation of constriction and elongation rates. Constriction failure rate was determined by counting the number of cells which initiated constriction at one division site, failed, then ultimately divided at a separate site. Prism was used for graphing and statistical analysis of calculated terms.
Cell shape analysis
For cell shape analysis, binary masks of phase contrast images of log phase cells were inputted into Celltool25, allowing for creation of cell contours, similar to as previously described22. Following alignment of cell contours (not allowing for reflection), a model of cell shape was created. The shape modes of interest were either plotted as histograms displaying the cell shape across two dimensions, or as single data points. R software was used to perform statistical analyses to compare population variances in shape modes across strains. Prism was used for graphing calculated terms.
Scanning electron microscopy sample preparation and imaging
For SEM, log phase cells were incubated on poly-lysine (1:10) coated glass cover slip for 15 minutes, then fixed for 1 hour using fixation buffer (1% glutaraldehyde, 0.02 M cacodylate, and 3 mM MgCl2). Cells were gradually dried by washing 3 times with wash buffer (3% sucrose, 0.02 M cacodylate, and 3 mM MgCl2), twice with dH2O, once each with 30%, 50%, 70%, 90%, 100% ethanol, once with 1:1 ethanol:hexamethyldisiloxane (HMDS), and once with HMDS at 5 minute intervals each, before desiccation overnight. Cover slips were mounted, then coated with a 15 nm gold palladium sputter coat. Samples were then imaged with a LEO/Zeiss Field-emission SEM.
Immunoblot analysis
Immunoblot analysis was performed similar to as previously described22, using a 1:5,000 – 1:6,666 dilution of α-FzlA primary antibody21, a 1:50,000 dilution of α-HU primary antibody50, and 1:10,000 of HRP-labeled α-rabbit secondary antibody (PerkinElmer) on nitrocellulose membranes. Chemiluminescent substrate (PerkinElmer) was added to facilitate protein visualization via an Amersham Imager 600 RGB gel and membrane imager (GE).
Transposon library preparation, sequencing, and analysis
Wild type Caulobacter crescentus NA1000 (EG2366) or ftsW**I* triple mutant (EG1557) cells were grown in a large culture (1 liter) to mid-log (0.4-0.6), washed of excess Mg2+ with 10% glycerol, and mutagenized with the Ez-Tn5 <Kan-2> transposome (Epicentre). Cells recovered by shaking at 30 °C for 90 minutes, then plated on kanamycin containing plates for 3 days at 30 °C in order to yield roughly 100-500 colonies per plate. Libraries were grown at 30 °C and comprised ~100,000-200,000 colonies each. Mutants were pooled into one library by scraping colonies from the surface of the agar and added into ~25-40 mL PYE. Pooled libraries were shaken to yield a homogenous slurry and sterile glycerol was added to 20%. Libraries were then frozen in liquid nitrogen and stored at −80 °C. Two libraries of each genetic background were prepared individually and compared as biological replicates.
Genomic DNA was extracted from one aliquot of each library using DNeasy Blood and Tissue Kit (Qiagen). Tn-Seq libraries were prepared for Illumina Next-Generation sequencing through sequential PCR amplifications using arbitrary hexamer primers and Tn5-specific primer facing outward for the first round, and indexing primer sets that include unique molecular identifier to filter artifacts arising from PCR duplicates for the second round. Libraries were then pooled and sequenced at the University of Massachusetts Amherst Genomics Core Facility on the NextSeq 550 (Illumina).
For analyses, reads were demultiplexed by index, then each sample Tn-Seq library was concatenated and clipped of the unique molecular identifier linker from the second PCR using Je51 and the following command:
Clipped reads were then mapped back to the Caulobacter NA1000 genome (NCBI Reference Sequence: NC_011916.1) using BWA52 and sorted using Samtools53:
Duplicate reads were removed using Je51 and indexed with Samtools53 using the following command:
5’ sites of inserted transposons from each library were converted into .wig files containing counts per position and viewed using Integrative Genomics Viewer54,55. Coverage and insertion frequency using a bedfile containing all open reading frames from NC_011916.1 with the outer 20% of each gene removed were determined using BEDTools56 and the following commands:
Comparison of transposon insertions was performed using the edgeR package in the Bioconductor suite57,58 using a quasi-likelihood F-test (glmQLFit) to determine the false discovery rate adjusted p-values reported here.
Author Information
PJL, CRM, AKD, PB, MH, GS-C, PC, RZ, and EDG planned the experiments, PJL, CRM, AKD, MH, GS-C, and RZ performed the experiments, PJL, CRM, AKD, MH, and EDG wrote the manuscript, and PJL, CRM, AKD, PB, MH, GS-C, PC, and EDG edited the manuscript.
Competing Interests
The authors declare no competing interests
Supplementary Video Legends
Supplementary Video 1:
Phase contrast time-lapse microscopy movies depicting division in WT or PG synthase hyperactive mutant cells with and without fzlA. As indicated, 5 minutes elapse between frames. Video playback is 10 frames per second.
Strain key (Caulobacter crescentus): WT (EG865), ftsW**I* (EG1557), ftsW**I* ΔfzlA (EG2170), ftsW* (EG1556), ftsW* ΔfzlA (EG2166).
Supplementary Video 2:
Phase contrast time-lapse microscopy movies depicting examples of constriction failure at the initial division site then initiation and completion at a subsequent site in ΔfzlA cells. As indicated, 5 minutes elapse between frames. Video playback is 10 frames per second.
Strain key (Caulobacter crescentus): ftsW**I* ΔfzlA (EG2170), ftsW* ΔfzlA (EG2166).
Supplementary Video 3:
Phase contrast microscopy movies depicting examples of cell twisting during division in multiple ftsW**I* ΔfzlA cells. As indicated, 5 minutes elapse between frames. Video playback is 10 frames per second. Strain key (Caulobacter crescentus): ftsW**I* ΔfzlA (EG2170).
Supplementary Tables
Supplementary Table 1:
Tn-Seq data and analysis for WT vs. ftsW**I* Caulobacter genes. Columns WT 1, WT 2, WI 1, and WI 2 contain the number of unique transposon insertions in each gene in each replicate of WT or ftsW**I* (WI) strain transposon insertion library. These values were used to determine the log2 fold-change (logFC), log counts per million reads (logCPM), PValue, false-discovery rate (FDR), and negative log10 of the FDR for each gene in WT versus ftsW**I*. Genes are ordered by significance (neglog(FDR)).
Supplementary Table 2:
List of plasmids used in this study.
Supplementary Table 3:
List of strains used in this study.
Acknowledgements
We would like to thank Josh Modell and Mike Laub for providing strains; the Manley lab, especially Ambroise Lambert, and members of the Xiao and Goley labs for helpful discussions; the Goley lab for feedback on the manuscript; Anant Bhargava and Adrien Ducret for help with time-lapse analysis; Mike Delannoy, Barbara Smith, and Selam Woldemeskel for developing the SEM protocol, and Mike and Barbara for training on SEM equipment; and Brandon King for providing support with experiments. This work funded in part by the National Institutes of Health, National Institute of General Medical Sciences through R01GM108640 (EDG and PJL), T32GM007445 (training grant support of PJL), R01GM111706 (PC and RZ), R25GM056901(training support of GS-C), and T32GM08515 (training grant support of RZ). PB and MH were supported by the National Science Foundation, IOS1557806.
References
Supplementary References
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