Abstract
Rod shaped bacteria grow by adding material into their cell wall via the action of two spatially distinct enzymatic systems: The Rod system moves around the cell circumference, while the class A penicillin-binding proteins (aPBPs) are unorganized. To understand how the combined action of these two systems defines bacterial dimensions, we examined how each system affects the growth and width of Bacillus subtilis, as well as the mechanical anisotropy and orientation of material within their sacculi. We find that rod diameter is not determined by MreB, rather it depends on the balance between the systems: The Rod system reduces diameter, while aPBPs increase it. RodA/PBP2A can both thin or widen cells, depending on its levels relative to MreBCD. Increased Rod system activity correlates with an increased density of directional MreB filaments, and a greater fraction of directionally moving PBP2A molecules. This increased circumferential synthesis increases the amount of oriented material within the sacculi, increasing their mechanical anisotropy and reinforcing rod shape. Together, these experiments explain how the combined action of the two main cell wall synthetic systems build rods of different widths, a model that appears generalizable: Escherichia coli containing Rod system mutants show the same relationship between the density of directionally moving MreB filaments and cell width.
Introduction
While the length of Bacillus subtilis rods increases as a function of their growth rate 1, their width remains constant across different growth conditions 2. How bacteria define and maintain these rod shapes with such precision is not understood, but it must involve mechanisms controlling both the rate and location of glycan insertion into the peptidoglycan (PG) sacculus, the enveloping heteropolymer meshwork that holds cells in shape 3. In order to understand how bacteria grow in defined shapes, we need to understand not only where these enzymes act, but how their activity affects the geometry and arrangement of material within the sacculus.
The PG for cell elongation is synthesized by two families of penicillin-binding proteins (PBPs). The class A penicillin-binding proteins (aPBPs) both polymerize and cross-link glycans. The class B penicillin-binding proteins (bPBPs) cross-link 4,5 the glycans polymerized by the glycosyltransferase RodA 6. bPBPs and RodA are components of the “Rod complex”, a group of proteins essential for rod shape (Figure 1A). In B. subtilis, this contains RodA, the class B transpeptidase PBP2A, MreC, MreD, RodZ, and filaments of MreB, an actin homolog 7,8. MreB polymerizes together with other MreB homologs into short, highly curved filaments on the membrane 9,10. To maximize their interaction with the membrane these curved filaments deform to the membrane, orienting along the direction of highest inward membrane curvature, pointing around the rod width 11. Oriented by their association with MreB filaments, the Rod complex moves directionally around the cell circumference, driven by the PG synthetic activity of RodA/PBP2A 12-15. This radial motion of independent filament-enzyme complexes are believed to insert PG strands around the rod width 16.
aPBPs also affect rod shape, as B. subtilis cells lacking them are thinner 17. Single molecule and biochemical studies indicate aPBPs and the Rod system are both spatially and functionally distinct 6,14: While Rod complexes move around the cell width, aPBPs have never been seen to move directionally. Rather, they diffuse within the membrane, occasionally pausing and remaining immobile for a few seconds 14. Furthermore, the synthetic activity of either aPBPs or the Rod complex can be inhibited without affecting the activity or motions of the other 6,14. Given that both synthetic systems are required for growth, but that Rod complex activity is spatially organized while aPBP activity is not (Figure 1A), it is not clear how these two PG synthetic machineries work together to create a rod-shaped sacculus of a given width.
Current models of rod width have focused on MreB filaments, attributing the altered widths of Escherichia coli MreB mutants to possible changes in MreB filament curvature, twist, angle, or localization to negative Gaussian curvature 18-24. Not only do these models lack the effects of aPBP mediated synthesis, they are: A) theoretical, as these proposed changes in filament curvature or twist have not been structurally validated, or B) difficult to reconcile in B. subtilis, which has no detectable negative Gaussian curvature 11 or skew in filament angle relative to the cell body 11-13,25.
Rather than focusing on MreB alone, we sought to develop a more thorough understanding of how the synthesis of both organized and unorganized PG affects the width and growth of B. subtilis, as well as their effects on the organization and mechanics of cell wall material. We find that aPBP and Rod complex-mediated PG synthesis have opposing effects on rod width, and that cell diameter depends on their balance. The rate cells add surface area is largely unaffected by the level of any one system, unless both become limiting. As MreBCD expression increases and rods thin, we observe a greater density of directionally moving MreB filaments and a greater fraction of directionally moving enzymes. Increased Rod complex activity creates a greater proportion of oriented material within the sacculus, causing the rod to stretch less across its width and more along its length in response to internal turgor. Finally, we show that the different widths of E. coli Rod mutants also show correlation between the density of directionally moving MreB filaments and cell width, giving a simple, generalizable model that may explain the role of the Rod system in cell width.
Results
The Rod and aPBP systems have opposing effects on cell diameter
The width of rod-shaped bacteria has been attributed to properties encoded within MreB filaments 19,21,24,26. We reasoned that if width were defined by MreB, then the mreBCD genes from larger bacteria should produce cells with larger diameters. To test this, we replaced the B. subtilis mreBCD operon with the mreBCD operon from Bacillus megaterium, a nearly 2-fold wider bacterium. Surprisingly, these cells grew as rods only slightly wider than wild type (WT) B. subtilis, and by further overexpressing B. megaterium mreBCD cells became even thinner (36 nm) (Figure 1B, S1A), suggesting that MreB filaments themselves do not encode a specific rod width.
We next examined how width changed as we independently controlled the levels of the two main B. subtilis PG synthetic systems. We created strains where expression of PBP1 (the major aPBP, encoded by ponA) was under IPTG control. As PG synthesis by the Rod system depends upon MreB 14, we created strains where the native B. subtilis mreBCD operon was under xylose control. We grew these strains at different induction levels, then measured their steady-state widths using fluorescence microscopy. For a subset of inductions, we measured the relative protein abundance using proteomic mass spectrometry, normalizing to the levels in WT cells grown in CH media (Supplemental File 1).
Varying mreBCD inductions revealed the Rod system has a thinning effect on rod shape. In the lowest inductions supporting growth, rods were ∼2-fold wider than WT, with some cells losing rod shape. As we increased mreBCD induction, cells became thinner, reaching WT width when MreB abundance recapitulated WT levels (Figure 1C, Figure S1B). Inductions above this point resulted in cells becoming 33 nm thinner than WT, and thinner still (58 nm) in mreBCD merodiploids. As above, these results demonstrate MreB filaments do not define a given rod diameter, rather as previously hypothesized 11,27, they indicate the Rod system reduces cell diameter.
Different ponA inductions revealed that aPBPs have a widening effect: With no IPTG, cells contained 0.25 the amount of WT PBP1, and were ∼23% thinner than WT, similar to ΔponA strains 17 (Figure 1D, Figure S1C). As we increased ponA induction cells became wider, reaching WT widths when PBP1 abundance recapitulated WT levels. Inductions above this point caused cells to become increasingly wider, and by expressing ponA under stronger promoters or in merodiploids, we could produce rods almost twice WT diameter.
These results demonstrate that the aPBPs and Rod system have opposing effects: The circumferentially moving Rod complex reduces cell diameter, while the spatially unorganized aPBPs increase diameter. We hypothesized that a balanced expression of each system could produce WT diameter rods. We combined the IPTG-inducible ponA and xylose-inducible mreBCD alleles into one “dual-inducible” strain. We found six different pairs of xylose and IPTG concentrations that produced rods of WT diameter (Figure 1E, Figure S1D), even though individually, each induction resulted in perturbed diameters in the singly-inducible parental strains (Figure 1C-D). Relative quantitation of protein levels revealed that, in the induction pairs at or beneath WT levels, these cells contained reduced, but relatively balanced amounts of PBP1 relative to MreB. To determine this balance in native conditions, we measured the widths and protein levels of WT cells grown in different media that result in different growth rates (Figure 1F, Figure S1E). Together, this data suggested width depends on the balance between the aPBP and the Rod systems. By plotting the ratio of the [Fold change PBP1] / [Fold change MreB] for all conditions in the data sets against their width (Figure 1G), we found that B. subtilis maintains its diameter within ∼5% of WT only when this ratio was within a range of 0.8 to 1.5; outside of this range, cell diameter diverged. WT cells in different media showed PBP1/MreB ratios ranging from 0.9 to 1.5. Likewise, the induction pairs yielding WT widths in the dual-inducible strain were within 0.8 to 1.0, only when their levels did not exceed WT. In contrast, the single-inducible ponA or mreBCD strains were within 5% of WT width only when the ratio was within this range; outside of it, width rapidly diverged. Together, these results indicate that, at the level of PG insertion, cell width is affected by the levels of the two opposing synthetic systems that insert material into the sacculus (Figure 1H).
RodA can function outside of the Rod system to widen cells, but only when PBP2A is also in excess
Given that RodA acts within the Rod complex with PBP2A (encoded by pbpA) to synthesize PG 14, titrations of rodA and pbpA should show the same “thinning effect” as mreBCD. However, rodA overexpression can restore WT width to ΔaPBP cells 6, a widening activity similar to ponA. To investigate this discrepancy, we made two sets of strains where rodA or pbpA (as the only elongation-specific bPBP in the cell) were under the control of increasingly strong inducible promoters. As before 28,29, low rodA or pbpA yielded wide cells; and as induction increased, cells gradually thinned to WT width, suggesting both are required for functional Rod complexes. However, while we expected excess RodA to create thinner cells, once rodA induction exceeded the level required for normal width, it behaved like excess ponA, fattening cells far beyond WT diameter 6 (Figure 2A). In contrast, pbpA inductions beyond this point had a negligible effect on diameter.
Given that RodA and PBP2A must interact to synthesize PG 30, we tested if the widening caused by RodA overexpression required excess PBP2A. We created two strains that had rodA under xylose control; one contained native pbpA, and the other had pbpA under IPTG control (both as the only elongation-specific bPBP in the cell). As before, when pbpA was under native control, depletion or overexpression of rodA caused diameter to increase (Figure 2B). In contrast, when pbpA was held at the lowest induction required for WT width and rodA was simultaneously overexpressed, cells remained at WT widths. However, these RodA-overproducing cells could be made increasingly wider by increasing pbpA induction (Figure 2B), demonstrating excess RodA requires excess PBP2A to increase width.
These results suggested RodA/PBP2A widens cells when their levels exceed the rest of the Rod complex. If this hypothesis is correct, wide cells caused by RodA/PBP2A overexpression should become thinner if MreBCD is increased, perhaps recruiting the diffusive excess RodA/PBP2A into circumferentially-moving, “thinning” Rod complexes. Indeed, while strong induction of rodA made wide cells, simultaneous overexpression of a second copy of mreBCD reduced cell diameter (Figure 2C), a narrowing far greater than what we observed when overexpressing mreBCD in otherwise WT strains (Figure 1C). Thus, the aPBP-like widening activity of RodA/PBP2A likely occurs once they exceed some level of MreBCD, possibly reflecting a saturation of binding sites within Rod complexes.
Growth rates are maintained across a wide range of enzyme levels, unless both systems become limiting
We next examined how each PG synthetic system affected the rate of cell growth in our dual-induction strain using two assays: 1) single-cell measurements of the rate of surface area addition 31 under agar pads and 2) population measures of growth in a shaking plate reader. First, we examined the growth of the dual-inducible strain at the mreBCD and ponA inductions that produced WT widths (Figure 1E). At the lowest induction pairs, growth was greatly reduced in both assays, and increased with each increasing induction pair (Figure 2D), up to the induction pair that produced WT PBP1 and MreB protein levels. Thus, growth can be reduced if both PG synthesis systems become limiting.
Next, we assayed growth as we titrated either mreBCD or ponA while holding the other constant. Similar to E. coli MreB studies 22,32,33, both assays showed no difference in growth across a wide range of mreBCD inductions, except at the lowest induction where cells frequently lost rod shape. Similarly, both assays saw no difference in growth across our ponA induction range. Thus, even though these cells have different geometries, they add new surface area at the same rate. We next examined how extremely low levels of aPBPs affected growth. Similar to previous observations 17,34,35, both ΔponA and ΔaPBP (ΔpbpF, ΔpbpG, ΔpbpD, ΔponA::kan) strains showed a marked reduction in bulk growth, a defect rescued by overexpression of RodA 6,36. However, the single cell measurements revealed a surprise: Both ΔponA and ΔaPBP cells showed no defect in their single-cell growth rate, adding surface area at the same rate as WT, as did RodA overexpressing ΔaPBP cells (Figure 2D). Given ΔaPBP cells display an increased rate of lysis that can be suppressed by RodA overexpression 6,36, it appears the population growth rate defect of aPBP-deficient cells arises not from a reduction in growth rate, but from an increased frequency of death.
Increased MreBCD increases the density of directionally moving MreB filaments and the fraction of directionally moving synthetic enzymes
To gain a mechanistic link between the level of the PG synthetic systems to cell width, we sought to develop microscopic measures of their activity. While the fraction of stationary aPBPs would be difficult to quantify, Rod complexes exhibit a more quantifiable phenotype: As their motion is driven by PG synthesis, the cellular activity of the Rod system can be measured by quantitating the number of directionally moving MreB filaments 37. We developed an analysis method that, using total internal reflection fluorescence microscopy (TIRFM) data, determines the density of MreB filaments moving directionally around the cell. Filaments undergoing directed motion are detected by taking advantage of the temporal correlation occurring between adjacent pixels across the cell width as objects move through them. These objects are then counted over a given time and then normalized to the total cellular surface area (Figure 3A, S3A-F). To evaluate the robustness of this approach to filament number, velocity, angle, and length, we simulated TIRFM time-lapses of MreB filaments in cells (Figure S3G-K). This revealed this approach was more accurate than counting filaments in the same data with particle tracking. Furthermore, when applied to real TIRFM movies of MreB filaments, this approach yielded numbers comparable to tracking MreB filaments inside cells imaged with TIRF - structured illumination microscopy (TIRF-SIM) (Figure S3L). With this analysis tool, we examined how the density of directionally moving MreB filaments related to rod width as we titrated the expression of an inducible mreB-sw-msfGFP, mreCD operon expressed as the only source at the native locus. This analysis revealed a well fit, apparently linear correlation between directional filament density and width (Figure 3B). Thus, the density of directionally moving MreB filaments reduces cell width, possibly by promoting directional PG synthesis.
To this end, we next examined how MreBCD levels affected PBP2A motion. Multiple studies have noted PBP2A molecules exist as a mixed population; some diffuse within the membrane, while others move directionally 12-14. We titrated mreBCD induction as we tracked the motions of PBP2A molecules (expressed at the native locus as a HaloTag fusion, sparsely labeled with JF-549 38). At low mreBCD, only a small fraction of PBP2A moved directionally. As we increased mreBCD induction, an increasing fraction of PBP2A molecules moved directionally (Figure 3C, Movie SM1), demonstrating that mreBCD limits the amount of directional PBP2A.
Increased MreBCD:PBP1 correlates with an increased amount of oriented material and structural anisotropy of the cell wall
To gain more insight into how an increased density of directional rod complexes can reduce rod width, we used polarization microscopy to understand how increased circumferential synthesis affected the organization of material within the cell wall. Polarization microscopy reports on both the angle and extent of orientation within optically anisotropic (or birefringent) materials 39, and has been used to assay the orientation of various materials, including plant cell walls 40-42 and mitotic spindles 43. Polarization microscopy revealed purified WT sacculi show birefringence, indicating that some fraction of the material within them is oriented. Focused at their surface, sacculi showed a predominant slow axis oriented along the rod length (Figure 4A, Movie SM2). Given that amino acids have a higher refractive index compared to sugars, this suggests (in agreement with previous models 44) the peptide crosslinks are predominantly oriented along the rod length, and the glycans are oriented around the circumference.
We then examined how the balance between the Rod and aPBP systems affected the relative amount of oriented material in the wall. We grew our dual-induction strain under three different induction conditions to create varied width cells: wide (high ponA, low mreBCD), normal (ponA and mreBCD induced at WT levels), and skinny (low ponA, high mreBCD) (Figure 4B), purified their sacculi, and quantified their total retardance with polarization microscopy (Figure 4C, Movie SM3). Retardance is the differential optical path length for light polarized parallel and perpendicular to the axis of molecular alignment; alternatively, it is defined as birefringence (Δn) times the physical path length through an anisotropic material. Wide cells (high ponA, low mreBCD) had the lowest retardance, skinny cells (low ponA, high mreBCD) had the highest retardance, and normal cells were in between (Figure 4D). Because retardance depends on both the thickness of material (path length) and the degree of its orientation (Δn), we normalized the retardance of each sample to the mean thickness of the cell walls of each induction condition, obtained using transmission electron microscopy (Figure S4A). This revealed that the skinny cell walls had more highly ordered material (Δn) per path length (nm) of cell wall thickness, and that wide cell walls had the least (Figure 4E). Thus, in agreement with recent atomic force microscopy studies showing orientated glycans in E. coli require MreB 16, these experiments demonstrate that as Rod system activity increases, so does the amount of oriented material in the wall.
The sacculi of E. coli and B. subtilis are mechanically anisotropic, stretching more along their length than across their width 44-46. To test how the ratio of oriented to unoriented PG synthesis affected this property, we grew our dual-inducer strain at the three mreBCD:ponA inductions above, labeled their walls with Alexa-488-D-amino carboxamides 4, and assayed the dimensions of their cell walls before and after hyperosmotic shocks (Figure 4F-G). This revealed that increased Rod system activity correlated with an increased mechanical anisotropy of the sacculus: As we increased the expression of MreBCD relative to PBP1A, rods shrank less across their width, and more along their length (Figure 4H, S4B-C). Thus, the Rod system acts to reinforce rod shape against internal turgor by promoting oriented PG synthesis around the rod.
E. coli Rod mutants also show a correlation between cell width and the density of directionally moving filaments
Previous studies have examined how MreB and PBP2 mutations affect the shape of E. coli, hypothesizing that their abnormal widths arise from changes in the curvature, twist, or angle of MreB filaments. Our observations in B. subtilis suggested an alternative explanation: Abnormal width might arise from simply changing the amount of Rod system activity. We tested this by measuring the density of directional GFP-MreB filaments in these same mutant E. coli strains. As a benchmark, we first assayed the width of E.coli as we titrated the expression of mreB-sw-msfGFP using CRISPRi against msfGFP 47,28. As in B. subtilis, this yielded an inverse relationship between directional MreB filament density and cell width (Figure 5A). Examining each group of mutants showed the same result: 1) An identical trend was observed for the mutations hypothesized to change filament twist 19, 2) as well as in the mutations designed to change filament curvature 24. And 3) notably, the same correlation between directional filament density and cell width was observed in strains where E. coli mrdA (PBP2) was replaced with mrdA genes from other species 22. TIRF-SIM imaging of these MreB mutants revealed some insight into these effects (Movie SM4): Some of the wide mutants showed either A) longer but fewer filaments, or B) a large fraction of immobile filaments. Conversely, some thinner mutants appeared to have more, but shorter filaments. Finally 4), while Colavin et al. hypothesized that RodZ reduces width by changing MreB filament curvature 24, we found that increased RodZ induction increased MreB filament density as cells thinned (Figure 5B, Movie SM5). This suggests that the decrease in cell width most likely occurs from to RodZ’s ability to nucleate MreB filaments 48 rather than changing their curvature. Thus, the density of directionally moving Rod complexes correlates with cell width in both B. subtilis and E. coli across multiple genetic perturbations.
Discussion
The shape of bacteria is defined by their cell walls; these experiments demonstrate that the two systems that insert PG into it have opposing roles on its shape. Due to the intrinsic orienting of MreB filaments around the rod width 11, the Rod system inserts circumferentially oriented material around the rod circumference, reducing its diameter. As the number of MreB filaments increases, so does the fraction of directional enzymes and the amount of oriented material in the wall. In contrast, the aPBPs do not move circumferentially, inserting material that isotropically enlarges the sacculus. Our data indicates the macroscopic shape of the sacculus arises from the nature of the material inserted into it: The more it is oriented around the rod circumference by the Rod complex, the less the rod stretches across its width, and the more it stretches along its length (Figure 5C).
If the balance between the two PG synthetic systems is perturbed, the shape of the sacculus becomes altered, though its rate of expansion remains constant. As both systems utilize the same pool of lipid II 49, the flux through each may depend on their relative levels; if one is reduced, the flux through the other may increase. This would explain why disrupting the Rod system causes cells to swell 27,29,50-52: In the absence of Rod-mediated thinning, aPBPs add more material uniformly over the cell surface. Likewise, in the absence of aPBP-mediated widening, increased flux through the Rod system would explain why cells become extremely thin 17,53. However, if both systems are equivalently reduced, cells grow with normal widths, but at slower rates; as long as the activities are balanced - identical shape arises from the balanced levels of enzymes, but growth is reduced due to their combined activity becoming limiting. This would explain why ponA mutations rescue mreB deletions 54; equally crippling both systems may rebalance the activities such that the cell retains normal shape and viability.
Implications for the role of MreB in rod width determination
Given that 1) mreBCD from the wide bacterium B. megaterium creates close to normal diameter B. subtilis rods, and 2) B. subtilis diameter depends on mreBCD levels, we find it unlikely that any property of MreB filaments defines a given cell diameter. Rather, MreB appears to be one component of a rod-thinning system, working in opposition to aPBP-mediated widening. Indeed, in vitro studies have revealed MreB filaments are extremely curved (> 200 nm), allowing them to deform to, and orient around the width of bacteria of any larger width 9,11. While it remains possible that given MreB mutations or interactions with RodZ could indeed alter filament curvature or twist 19,21,24,26, these experiments suggest a more parsimonious explanation for their effect on Rod activity: These mutations alter MreB’s polymerization dynamics or cellular distribution. Reducing the amount or number of active MreB polymers would reduce Rod activity, causing cells to widen. Likewise, mutations altering MreB’s polymer length distribution or tendency for filaments to bundle would cause the distribution of Rod activity to become non-uniform, causing some parts of the cell to thin while others would widen, as previously observed for certain MreB mutants 12,27.
Additionally, our experiments reinforce observations that aPBPs and RodA (when in excess to MreBCD) serve reparative, anti-lytic roles: aPBP-deficient cells grow at the same rate as WT, yet have an increased frequency of death, lysing as thin rods without losing shape 6,36. Likewise, aPBP-mediated synthesis increases upon endopeptidase overexpression in E. coli 55. Given the active state of aPBPs correlates with single aPBP molecules displaying periods of transient immobility 14,56, their synthesis may be localized to small (<50 nm) regions. Combined, these observations support a model where aPBPs synthesize material to fill gaps in the PG meshwork 36,14. Gaps could arise via mechanical damage, hydrolases 55, or between the imperfectly oriented strands built by the Rod complex 11. If this model is correct, the different spatial activities of these two systems might allow the sacculus to maintain integrity at any Rod/aPBP ratio: the fewer the Rod complexes, the larger the gaps filled by aPBPs.
While many different activities affect the shape of the sacculus, such as its cleavage by hydrolases or rigidification by wall teichoic acids 11, the first process defining its geometry is the spatial coordination controlling where glycans are inserted into it. While these experiments give a coarse-grained description into how each synthetic system affects cell shape, a fine scale, mechanistic understanding remains to be determined - How does aPBP activity make cells wider? How does increased Rod activity make cells thinner? Understanding the physical mechanisms causing these changes will require not only a better understanding of the molecular architecture of the sacculus, but also an investigation into how the enzymes downstream of glycan insertion affect the shape and mechanics of sacculi as they subsequently modify, remodel, and break down the nascently-polymerized glycan architecture.
Author Contributions
B. subtilis strains were cloned by MD, YS, and MK. All width and bulk growth measurements of B. subtilis were done by MD, and E. coli widths by YS. E. coli CRISPRi strains were cloned by AV, who was supervised by SVT. Single cell growth rates were done by YS and MK. TIRFM and tracking of PBP2A was done by MK. All TIRFM of MreB was done by YS. TIRF-SIM of MreB was done by YS, EG, and JR. Purified sacculi and proteomic mass spec sample preps were done by MD. YS wrote the code for analysis of single cell growth rate, filament density, and simulations of data. Polarization microscopy and analysis was conducted by JR, RO, and EG. SW did the FDAA synthesis, osmotic shocks, and TEM. The paper was written by EG, MK, MD, and YS.
Supplemental Material
Supplemental figures
Supplemental Movie Legends
Movie S1 – Example single molecule movies of PBP2A-HaloTag (labeled with JF5491) at different mreBCD induction levels in bMK385 (amyE::Pxyl-mreBCD::erm, ΔmreBCD, HaloTag-11aa-pbpA::cat). Xylose concentrations are indicated in each panel. Frames are 300 msec apart.
Movie S2 – Z-stack of LC-PolScope images of sacculi purified from wild type Py79. (Left) Z-stack of the retardance of each plane. (Right) Color map of birefringence at each plane, with color wheel at upper left giving orientation reference. Z-steps were taken in 100 nm increments.
Movie S3 – Z-stack color map of birefringence of sacculi purified from strain bMD620 (amyE::Pxyl-mreBCD::erm, ΔmreBCD, yhdG::Pspank-ponA::cat, ΔponA::kan), where cells were induced to “Wide” (0.5 mM xylose, 0.1 mM IPTG) and “Normal” widths (5 mM xylose, 0.025 mM IPTG). Color wheel at upper left indicates orientation direction. Z-steps were taken in 100 nm increments.
Movie S4 – TIRF-SIM movies of different mreB-SW-msfGFP mutants. Cells were grown in LB at 37°C, then placed under an LB agar and imaged at 37°C. Frames are 1 sec apart. Scale bars are 1 um.
Movie S5 – TIRF-SIM movies of strain KC717(mreB::msfGFP-mreB, ProdZ<>(frt araC PBAD)). Cells were grown for 5 hours in LB with the indicated arabinose concentrations, were placed under an LB agar pad and imaged at 37°C. Frames are 1 sec apart. Scale bars are 1 um.
Materials and Methods
Media and culture conditions
For all experiments, unless otherwise noted, B. subtilis and B. megaterium were grown in casein hydrolysate (CH) medium (where indicated, xylose and/or isopropyl thiogalactoside (IPTG) was added), and E. coli strains were grown in lysogeny broth (LB) medium (where indicated, arabinose or anhydrotetracycline (ATc) was added), at 37°C with rotation. When pre-cultures reached exponential phase the next day, after overnight growth at 25°C, they were further diluted into fresh growth medium (and where indicated, with the specified concentrations of inducer) and were grown for at least 3 hours at 37°C with rotation, to an OD600 of ∼0.3 to 0.7.
Sacculi purification
20 mL cultures were grown in 250 mL baffled flasks at 37°C with vigorous shaking, to an OD600 of ∼0.5, were harvested and were centrifuged at 5,000 × g for 5 min at 4°C. Cell pellets were resuspended in 1 mL of ice-cold phosphate buffered saline (PBS), were centrifuged at 6,000 × g for 30 sec, were resuspended in 500 μL of PBS, and were killed by boiling in a water bath for 10 min. Cells were centrifuged at 6,000 × g for 2 min, were resuspended in 500 μL of PBS containing 5% sodium dodecyl sulfate (SDS), and were boiled in a water bath for 25 min; this was repeated once, except boiling was for 15 min. To remove the SDS, the samples were centrifuged at 6,000 × g for 2 min, and were resuspended in 500 μL of PBS; this was repeated 5 times. The cells were centrifuged at 6,000 × g for 2 min, were resuspended in 1 mL of 50 mM Tris-HCl, pH 7.5, containing 10 mM NaCl, and 2 mg of pronase from Streptomyces griseus (MilliporeSigma, MA), and were incubated at 60°C for 90 min with gentle shaking. To remove the pronase, the samples were boiled twice in PBS/5% SDS, followed by 6 rounds of washes in PBS, exactly as described in the steps above that precede the pronase treatment. The samples were centrifuged at 6,000 × g for 2 min, were resuspended in 1 mL of 25 mM Tris-HCl, pH 8.5, containing 0.5 M NaCl, 5 mM MgCl2, and 100 U of salt-active nuclease (SAN) (ArticZymes, Norway), and were incubated at 4°C overnight with gentle mixing. To remove the SAN, the samples were boiled twice in PBS/5% SDS, followed by 6 rounds of washes in PBS, exactly as described in the steps above that precede the pronase treatment. The sacculi were centrifuged at 6,000 × g for 2 min, the supernatant was removed, and the pellets were stored at −80°C.
Polarization microscopy
Purified sacculi were resuspended in PBS and placed on ethanol-cleaned No. 1.5 coverslips under a cleaned glass slide. Polarization images were acquired on an inverted LC-PolScope mounted on a Nikon Ti-E equipped with a 60x/1.4NA Plan Apo oil immersion objective and oil condenser with matching NA, and a Hamamatsu Photonics Flash4 camera. Z-stacks in 100 nm steps were taken of each sample using 50 msec acquisitions of 546/12 nm light. All image acquisition, processing, and display, including colored display and line maps were prepared using the OpenPolScope Hardware Kit and plugins for ImageJ/Micro-Manager from OpenPolScope.org.
Calculation of retardance
Z-stacks of the computed total retardance for each slice were exported from OpenPolScope. These stacks were cropped at one frame above and beneath the focal planes of the top and the bottom of the cells, then projected into a single plane using ImageJ. To avoid getting high retardance values from the edge effects arising from the sides of the cell or the septa, we selected the retardance at the middle of the cell, using the average of line scans (5 pixels long and 3 wide) that were drawn down the center of cells, taking care to avoid edges and septa.
Transmission electron microscopy of cell wall thickness
Overnight cultures were diluted into fresh medium and grown to an OD600 of ∼0.3 to 0.5. Cells were pelleted by centrifugation at 5,000 x g and fixed by resuspending in 100 mM MOPS pH 7 containing 2% paraformaldayde, 2.5% gluteraldehyde, and 1% DMSO overnight at 4°C. Cells were centrifuged at 5,000 x g and were washed 3 times with 100 mM MOPS pH 7. The pellet was stained with 2% osmium tetroxide in 100 mM MOPS for 1 hr, washed twice with deionized water, and stained overnight with 2% uranyl acetate. The pellet was washed twice with deionized water and dehydrated by washing once with 50% ethanol, once with 70% ethanol, once with 95% ethanol, and then three times with 100% (v/v) ethanol. Samples were prepared for resin infiltration by washing once with 50% ethanol, 50% propylene oxide (v/v) and then once with 100% propylene oxide. All wash steps were for 5 min. Infiltration of resin was achieved by incubation with 50% Embed 812 (EMS, PA)/50% propylene oxide for 1 hr, followed by incubation with 67% Embed 812/33% propylene oxide for 1hr, and incubation with 80% Embed 812/20% propylene oxide for 1 hr. Samples were then incubated twice with Embed 812 for 1 hr, followed by an overnight incubation in molds. The molds were baked at 65°C for 18 hr before sectioning.
Serial ultrathin sections (80 nm) were cut with a Diatome diamond knife (EMS, PA) on a Leica Ultracut UCT (Leica Microsystems, Germany) and collected on 200-mesh thin-bar formvar carbon grids. Sections were imaged on an FEI Tecnai transmission electron microscope.
Cell wall thickness measurements were performed using a custom-built MATLAB (Mathworks, MA) script. Image intensity profiles extracted from lines were drawn perpendicular to a user-input line defining the middle of the cell wall. The distance between the two lowest points below a threshold within 40 nm of the middle of the cell wall was measured as the cell wall thickness at ∼30 points in each cell.
Measurements of cell diameter
Cultures grown to an OD600 of ∼0.3 to 0.7 were stained with 0.5 μg/mL FM 5-95 (Thermo Fisher, MA) for 1 min, and were concentrated by centrifugation at 6,000 x g for 30 sec. The cell pellet was resuspended in ∼1/20 volume of growth medium, and 3 μL was applied to ethanol-cleaned No. 1.5 coverslips under a 3% agarose pad containing growth medium. Fluorescent cells were imaged with the top surface of the agarose pad exposed to air, in a chamber heated to 37°C. Epifluorescence microscopy was performed using a Nikon Eclipse Ti equipped with a Nikon Plan Apo λ 100×/1.4NA objective and an Andor camera. Cell contours and dimensions were calculated using the Morphometrics software package 2.
Measurements of single-cell growth rate
Cultures grown to an OD600 below 0.3 were concentrated by centrifugation at 6,000 x g for 30 sec. The cell pellet was resuspended in growth medium, and applied to No. 1.5 glass-bottomed dishes (MatTek Corp., MA). All cells were imaged under a 2% agarose pad containing growth medium, with the top surface exposed to air, in a chamber heated to 37°C. Phase-contrast microscopy was performed using a Nikon Eclipse Ti equipped with a Nikon Plan Apo λ 100×/1.4NA objective and an Andor camera. We used a custom-built package in MATLAB to perform segmentation on phase-contrast time-lapse movies, then calculated the growth rate of the surface area of single B. subtilis chains.
Measurements of bulk growth rate
For cell culture measurements of growth rate, overnight cultures were diluted in fresh medium and grown for ∼3 hr at 37°C to an OD600 of ∼0.3 to 0.7. The cultures were diluted back to a calculated OD600 of 0.07 in 100-well microtiter plates, and their growth rates were measured in a Bioscreen-C Automated Growth Curve Analysis System (Growth Curves USA, NJ) plate reader, at 37°C with continuous shaking. Growth rates were calculated from OD600 measurements that were recorded every 5 min for at least 6 hr.
TIRF microscopy of Halo-PBP2A
Cultures grown to an OD600 below 0.3 were labelled with 10 nM JF549 3, and were concentrated by centrifugation at 6,000 x g for 30 sec. The cell pellet was resuspended in growth medium and applied to ethanol-cleaned No. 1.5 coverslips. All cells were imaged under a 2% agarose pad containing growth medium with the top surface exposed to air, in a chamber heated to 37°C. TIRFM and phase-contrast microscopy were performed using a Nikon Eclipse Ti equipped with a Nikon Plan Apo λ 100×/1.45NA objective and a Hamamatsu ORCA-Flash4.0 V2 sCMOS camera. Fluorescence time-lapse images were collected by continuous acquisition with 300 msec exposures.
Analysis of the density of MreB
Phase images of bacteria were segmented using Morphometrics 4, and the width and length of each cell was calculated. Next, the fluorescence time-lapses were analyzed based on the segmentation mask of the phase image (Figure S3C). Filament counting was performed in several steps (Figure S3). First, kymographs were generated for each row of pixels along the midline of the cell. Next, the kymographs for each row were placed side by side, converting the TIRF time lapse data into a single 2D image (Figure S3D). To identify filaments in the kymograph, closed contours were generated in the 2D image (Figure S3E). We only selected contours within a given size range (0.04 μm2 to 0.17 μm2). For these contours, we calculated the total intensity (the sum of the intensities of the pixels in the contour), the centroid, the velocity (calculated from the slope of the major axis line of the contour) (Figure S3F), and time (from the centroid). Next, to identify cases where the same MreB filament appears in multiple sequential kymographs, each object in a given kymograph is linked to a corresponding object in the next and previous kymographs based on the above properties of the object (see Figure S3F for details). Finally, the counting is verified manually by numbering each filament on the 2D image (Figure 3A). To test the performance of the filament counting, we analyzed simulated data with different filament density, velocity, and orientation settings (Figure S3). All of the image analyses were performed using MATLAB.
Simulation of directionally moving MreB
The Image Correlation Spectroscopy 5 MATLAB package was used for the simulation of MreB moving around the cell. The following parameters were set for the MreB simulations: velocity, orientation, filament number and filament length. The default velocity setting is 30 nm/sec and the default orientation is 0, which means MreB moves perpendicular to the central axis. The default filament length is set to 250 nm and each MreB monomer is assumed to be 5 nm. The cell width is set to 1 um and the cell length is set to 4 μm. The pixel size is 65 nm and the time interval is 1 sec, which is the same as the TIRF imaging obtained with our Nikon Eclipse Ti equipped with a Hamamatsu ORCA-Flash4.0 V2 sCMOS camera. The particles are randomly distributed on the surface of the cell. Each simulation data point was repeated 5 times for the counting analysis. To compare the correlated motion approach against particle tracking, we counted the number of tracks observed after tracking the simulated data using the Linear Motion LAP tracker in FIJI 6 with TrackMate v3.8 7. The threshold for spot size was 0.195 μm and the intensity threshold was 10 counts. The search radius is 0.085 μm, the link radius is 0.085 μm, and the gap size is 1. All traces longer than 6 frames that had moved more than 250 nm were considered directional motions.
TIRF-SIM imaging of E. coli strains
Cells were prepared as described in “Media and culture conditions”. Cells were placed under an LB agarose pad, on a cleaned No.1.5 coverslip, and imaged at 37°C. Live-cell SIM data were acquired as described previously 8 on a Zeiss Axio Observer.Z1 inverted microscope outfitted for structured illumination. An Olympus 100×/1.49NA objective was used instead of the Zeiss 1.45NA objective because the slightly larger NA of the Olympus objective gives higher tolerance for placing the excitation beams inside the TIRF annulus. Data was acquired at 1 sec frame rates, with 20 msec exposures from a 488 nm laser for each rotation. TIRF-SIM images were reconstructed as described previously8.
TIRF-SIM imaging of B. subtilis strains
Cells were prepared as described in “Media and culture conditions”. Cells were placed under a CH agarose pad in a No. 1.5 glass-bottomed dish (MatTek Corp., MA) for imaging. Images were collected on a DeltaVision OMX SR Blaze system in TIRF mode, using an Edge 5.5 sCMOS camera (PCO AG, Germany) and a 60x objective. 75 msec exposures from a 488 nm diode laser were used for each rotation. Spherical aberration was minimized using immersion oil matching. Raw images were reconstructed using SoftWoRx (GE Healthcare, MA) software.
Particle tracking of JF549-Halo-PBP2A
Particle tracking was performed using the software package FIJI 6 and the TrackMate v3.8 7 plugin. For calculation of particle velocity, the scaling exponent α, and track orientations relative to the midline of the cell, only tracks persisting for 7 frames or longer were used. Particle velocity for each track was calculated from nonlinear least squares fitting using the equation MSD(t) = 4Dt + (vt)2, where MSD is mean squared displacement, t is time interval, D is the diffusion coefficient, and v is speed. The maximum time interval used was 80% of the track length. To filter for directionally moving tracks, we discarded those with a velocity lower than 0.01 nm/sec. Tracks were also excluded if the R2 for log MSD versus log t was less than 0.95, indicating a poor ability to fit the MSD curve.
Osmotic shock experiments
Overnight, exponentially growing cultures (as described in “Media and culture conditions”) were diluted into fresh CH medium, grown at 37°C to an OD600 of 0.1 to 0.2, then stained by growing in 100 μM Alexa Fluor 488-D-Lysine-NH2 for 1 hr. Without washing, cells were then loaded into a CellASIC microfluidic flow cell (MilliporeSigma, MA) pre-conditioned with media and washed in the chamber via channels 6 and 5. Media in channel 6 was replaced with 5 M NaCl, and the flow cell was resealed and imaged immediately. After collecting images across the whole chip pre-shock, 5 M NaCl was flowed into the chip via channel 6 and imaged immediately.
Alexa Fluor 488-D-Lysine-NH2
Alexa Fluor 488-D-Lysine-NH2 was synthesized as in Lebar et al., 2014. Briefly, Boc-D-Lys(Cbz)-OH (Bachem, Switzerland) was reacted with carbonyldiimidazole (CDI) (MilliporeSigma, MA) in dimethylformamide (DMF) for 1.5 hr, then aqueous ammonia was added and stirred for 6 hr to form the carboxamide Boc-D-Lys(Cbz)-NH2. The Cbz protecting group was removed by catalytic hydrogenation (20% Pd(OH)2/C) in methanol. The product, Boc-D-Lys-NH2, was reacted with CDI in DMF for 1.5 hr, then Alexa Fluor 488 carboxylic acid in DMF was added and reacted for 6 hr to yield Boc-D-Lys(Alexa Fluor 488)-NH2. The Boc protecting group was removed by stirring in neat trifluoroacetic acid (TFA) for 30 min. The reaction was stopped by dropwise addition of TFA solution to ice-cold ether. The precipitate was then HPLC-purified to obtain Alexa Fluor 488-D-Lysine-NH2.
CRISPRi titration of MreB expression
We used complementarity-based CRISPR knockdown to titrate the MreB expression level in E. coli. The degree of MreB-SW-msfGFP repression is controlled by introducing mismatches between the guide RNA and the target DNA 9. The repression strength can be tuned by modulating spacer complementarity to msfGFP using different numbers of mismatches. To repress msfGFP using CRISPR knockdown, we placed the dCas9 cassette under the control of a Ptet promoter and different plasmids to target msfGFP. We induced dCas9 at a constant high level with 100 ng/ml of ATc and changed the degree of guide RNA complementarity with different plasmids. For pcrRNA plasmid we use four different guide RNAs with 10, 11, 14, and 20 bp of complementarity. For pAV20 plasmid we use four different guide RNAs with 5, 10, 14, and 20 bp of complementarity. The cells were grown and imaged in LB containing 50 ug/ml kanamycin.
Protein extraction and labelling
Cell cultures were grown to an OD600 of ∼0.4 to 0.6 (cell amounts were normalized by harvesting the equivalent of 3 mL of culture at an OD600 of 0.5). Cells were centrifuged at 6,000 × g for 30 sec, washed once in 1 mL of ice-cold 20 mM Tris-HCl, pH 7.5, and 10 mM EDTA (TE), were resuspended in 100 μL of TE, and were killed by boiling in a water bath for 10 min. All samples were frozen at −80°C overnight (or up to 1 week). Thawed samples were digested with 50 μg of lysozyme (Thermo Fisher, MA) in the presence of 1 mM PMSF, at 37°C for 15 min.
Protein extraction was achieved utilizing a Covaris S220 ultrasonicator (Covaris, MA), under denaturing conditions upon the addition of urea-based Protein Extraction Buffer DF (Covaris, MA), followed by ice-cold methanol/chloroform precipitation. Proteins were digested with trypsin (Promega, WI). Each resulting peptide mixture was labeled with one of a set of up to eleven isotopic tandem mass tags (TMTs) (Thermo Fisher, MA).
Peptide fractionation and mass spectrometry
Equal amounts of each TMT-labelled sample were combined and fractionated by electrostatic repulsion-hydrophobic interaction chromatography, on an Agilent (Santa Clara, CA) 1200 HPLC system using a PolyWAX LP 200 x 2.1 mm, 5 μm, 300Â column (PolyLC, MD). Peptides were separated across a 70 min gradient from 0% of “buffer A” (90% acetonitrile, 0.1% acetic acid) to 75% of “buffer B” (30% acetonitrile, 0.1% formic acid), with 20 fractions collected over time. Each fraction was dried in a SpeedVac (Eppendorf, Germany) and resuspended in 0.1% formic acid before injection to a mass spectrometer.
LC-MS/MS was performed on a Thermo Orbitrap Elite (Thermo Fisher, MA) mass spectrometer equipped with a Waters nanoACQUITY HPLC pump (Waters Corp., MA). Peptides were separated on a 150 μm inner diameter microcapillary trapping column packed with ∼3 cm of C18 Reprosil 5 μm, 100 Å resin (Dr. Maisch GmbH, Germany), followed by an analytical column packed with ∼20 cm of Reprosil 1.8 μm, 200 Å resin. Separation was achieved by applying a gradient of 5–27% acetonitrile in 0.1% formic acid, over 90 min at 200 nl/min. Electrospray ionization was achieved by applying a voltage of 2 kV using a home-made electrode junction at the end of the microcapillary column and sprayed from fused-silica PicoTips (New Objective, MA). The Orbitrap instrument was operated in data-dependent mode for the mass spectrometry methods. The mass spectrometry survey scan was performed in the Orbitrap in the range of 410 –1,800 m/z at a resolution of 12 × 104, followed by the selection of the twenty most intense ions for HCD-MS2 fragmentation using a precursor isolation width window of 2 m/z, AGC setting of 50,000, and a maximum ion accumulation of 200 msec. Singly-charged ion species were not subjected to HCD fragmentation. Normalized collision energy was set to 37 V and an activation time of 1 msec. Ions in a 10 ppm m/z window around ions selected for MS2 were excluded from further selection for fragmentation for 60 sec.
Mass spectrometry analysis: Raw data were submitted for analysis in Proteome Discoverer 2.1.0.81 (Thermo Scientific) software. Assignment of MS/MS spectra was performed using the Sequest HT algorithm by searching the data against a protein sequence database that including all entries from B. subtilis (UniProt proteome ID UP000018540) and other known contaminants such as human keratins and common lab contaminants. Sequest HT searches were performed using a 15 ppm precursor ion tolerance and required each peptide’s N- and C-termini to adhere with trypsin protease specificity, while allowing up to two missed cleavages. TMT tags on peptide N-termini and lysine residues (+229.162932 Da) were set as static modifications while methionine oxidation (+15.99492 Da) was set as a variable modification. An MS2 spectra assignment false discovery rate (FDR) of 1% on protein level was achieved by applying the target-decoy database search. Filtering was performed using Percolator (64-bit version 10). For quantification, a 0.02 m/z window centered on the theoretical m/z value of each of the TMT reporter ions and the intensity of the signal closest to the theoretical m/z value was recorded. Reporter ion intensities were exported in a results file of the Proteome Discoverer 2.1 search engine in Microsoft Excel format. The total signal intensity across all peptides quantified was summed for each TMT channel, and all intensity values were adjusted to account for potentially uneven TMT-labelling and/or sample handling variance for each labelled channel. For our final relative protein quantitation analysis, all contaminants from the database search were removed from the results, and only the remaining B. subtilis proteins were used to re-normalize all protein abundances.
Strain construction
bMD277 containing amyE::Pxyl-mreBCD, minCD (B. megaterium)::erm was generated upon transformation of Py79 with a five-piece Gibson assembly reaction11, that contained the following PCR products. (1) A 1228 bp fragment containing sequence upstream of the amyE locus (amplified from Py79 genomic DNA using primers oMD191 and oMD108); (2) the 1673 bp erythromycin-resistance cassette loxP-erm-loxP (amplified from pWX467 [gift of D. Rudner] using primers oJM028 and oJM029); (3) a 1532 bp fragment containing the xylR gene, and the PxylA promoter with an optimized ribosomal binding sequence (amplified from pDR150 [gift of D. Rudner] using primers oMD73 and oMD226); (4) a 4106 bp fragment containing the mreB, mreC, mreD, minC and minD alleles of the B. megaterium mreB operon (amplified from strain QMB 1551 (ATCC 12872) genomic DNA using primers oMD227 and oMD228); and (5) a 1216 bp fragment containing the amyE terminator, and sequence downstream of the amyE locus (amplified from Py79 genomic DNA using primers oMD196 and oMD197).
bMD465 harboring amyE::Pxyl-mreBCD, minCD (B. megaterium)::erm, mreBCD, minC,D::spec (B. megaterium) was generated upon transformation of bMD277 with a four-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1275 bp fragment containing sequence upstream of the mreB gene (amplified from Py79 genomic DNA using primers oMD96 and oMD308); (2) a 4121 bp fragment containing the mreB, mreC, mreD, minC and minD alleles of the B. megaterium mreB operon (amplified from strain QMB 1551 (ATCC 12872) genomic DNA using primers oMD313 and oMD314); (3) the 1274 bp spectinomycin-resistance cassette loxP-spec-loxP (amplified from pWX466 [gift of D. Rudner] using primers oJM028 and oJM029); and (4) a 1152 bp fragment containing sequence downstream of the minD gene (amplified from Py79 genomic DNA using primers oMD300 and oMD315).
bMK355 harboring amyE::Pxyl-mreBCD::erm was built identical to bMD277, except that PCR product (4) was instead a 2493 bp fragment containing the mreB, mreC, and mreD alleles of the B. subtilis mreB operon (amplified from Py79 genomic DNA using primers oMD334 and oMK221).
bMD543 containing amyE::Pxyl-mreBCD::erm, ΔmreBCD, ΔminCD::cat was generated upon transformation of bMK355 with a three-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1305 bp fragment containing sequence upstream of the mreB gene (amplified from Py79 genomic DNA using primers oMD96 and oMD298); (2) the 1139 bp chloramphenicol-resistance cassette loxP-cat-loxP (amplified from pWX465 [gift of D. Rudner] using primers oJM028 and oJM029); and (3) a 1215 bp fragment containing sequence downstream of the minD gene (amplified from Py79 genomic DNA using primers oMD299 and oMD300).
bMD545 containing amyE::Pxyl-mreBCD::erm, ΔmreBCD, PmreB-minC,D::spec was generated upon transformation of bMD543 with a four-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1305 bp fragment containing sequence upstream of the mreB gene (amplified from Py79 genomic DNA using primers oMD96 and oMD379); (2) a 1582 bp fragment containing the minC and minD genes, and the minD terminator of the mreB operon (amplified from Py79 genomic DNA using primers oMD380 and oMD381); (3) the 1274 bp spectinomycin-resistance cassette loxP-spec-loxP (amplified from pWX466 using primers oJM028 and oJM029); and (4) a 1215 bp fragment containing sequence downstream of the minD gene (amplified from Py79 genomic DNA using primers oMD300 and oMD382).
bMD599 harboring ΔponA::kan was generated upon transformation of Py79 with a three-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1329 bp fragment containing sequence upstream of the ponA gene (amplified from Py79 genomic DNA using primers oMK001 and oMK002); (2) the 1577 bp kanamycin-resistance cassette loxP-kan-loxP (amplified from pWX470 [gift of D. Rudner] using primers oJM028 and oJM029); and (3) a 1313 bp fragment containing sequence downstream of the ponA gene (amplified from Py79 genomic DNA using primers oMK005 and oMK006).
bMK005 containing ΔponA::cat was built identical to bMD599, except that PCR product (2) was instead the 1139 bp chloramphenicol-resistance cassette loxP-cat-loxP (amplified from pWX465 using primers oJM028 and oJM029).
bMD554 containing yhdG::Phyperspank-ponA::cat was generated upon transformation of Py79 with a five-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1219 bp fragment containing sequence upstream of the yhdH gene (amplified from Py79 genomic DNA using primers oMD328 and oMD367), (2) the 1139 bp chloramphenicol-resistance cassette loxP-cat-loxP (amplified from pWX465 using primers oJM028 and oJM029); (3) a 1895 bp fragment containing the lacI gene, and the Phyperspank promoter with an optimized ribosomal binding sequence (amplified from pDR111 [gift of D. Rudner] using primers oMD232 and oMD234); (4) a 2799 bp fragment containing the ponA coding region (amplified from Py79 genomic DNA using primers oMD365 and oMK370); and (5) a 1252 bp fragment containing the yhdG terminator, and sequence downstream of the yhdG gene (amplified from Py79 genomic DNA using primers oMD371 and oMD372).
bMD586 harboring yhdG::Phyperspank-ponA::cat, ΔponA::kan was generated upon transformation of bMD554 with genomic DNA from bMD599.
bMD594 harboring yhdG::Pspank-ponA::cat was built identical to bMD554, except that PCR product (1) was instead a 1287 bp fragment made with primers oMD328 and oMD329; and PCR product (3) containing the Pspank promoter was instead amplified from pDR110 (gift of D. Rudner).
bMD598 containing yhdG::Pspank-ponA::cat, ΔponA::kan was generated upon transformation of bMD594 with genomic DNA from bMD599.
bMD619 containing amyE::Pxyl-mreBCD::erm, ΔmreBCD, PmreB-minCD::spec, yhdG::Pspank-ponA::cat was generated upon transformation of bMD545 with genomic DNA from bMD594.
bMD620 containing amyE::Pxyl-mreBCD::erm, ΔmreBCD, PmreB-minCD::spec, yhdG::Pspank-ponA::cat, ΔponA::kan was generated upon transformation of bMD619 with genomic DNA from bMD599.
bMD622 harboring amyE::Pxyl-mreBCD::erm, yhdG::Pspank-ponA::cat, ΔponA::kan was generated upon transformation of bMD598 with genomic DNA from bMK355.
bMD556 harboring yhdG::Phyperspank-rodA::cat was built identical to bMD554, except that PCR product (4) was instead a 1236 bp fragment containing the rodA coding region (amplified from Py79 genomic DNA using primers oMD364 and oMK369).
bMD580 containing yhdG::Phyperspank-rodA::cat, ΔrodA::kan was generated upon transformation of bMD556 with a three-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1265 bp fragment containing sequence upstream of the rodA gene (amplified from Py79 genomic DNA using primers oMD388 and oMD389); (2) the 1577 bp kanamycin-resistance cassette loxP-kan-loxP (amplified from pWX470 using primers oJM028 and oJM029); and (3) a 1258 bp fragment containing sequence downstream of the rodA gene (amplified from Py79 genomic DNA using primers oMD386 and oMD387).
bMD592 harboring Pxyl-rodA::erm was generated upon transformation of Py79 with a four-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1265 bp fragment containing sequence upstream of the rodA gene (amplified from Py79 genomic DNA using primers oMD388 and oMD389); (2) the 1673 bp erythromycin-resistance cassette loxP-erm-loxP (amplified from pWX467 using primers oJM028 and oJM029); (3) a 1532 bp fragment containing the xylR gene, and the PxylA promoter with an optimized ribosomal binding sequence (amplified from pDR150 using primers oMD73 and oMD226); and (4) a 1271 bp fragment containing the rodA coding region (amplified from Py79 genomic DNA using primers oMD394 and oMD395).
bMD627 containing Pxyl-rodA::erm, ΔpbpH::spec was generated upon transformation of bMD592 with genomic DNA from bDR2487.
bMD563 containing yhdG::Phyperspank-pbpA::cat was built identical to bMD554, except that PCR product (4) was instead a 2204 bp fragment containing the pbpA coding region (amplified from Py79 genomic DNA using primers oMD316 and oMK368).
bMD573 harboring yhdG::Phyperspank-pbpA::cat, ΔpbpH::spec was generated upon transformation of bMD563 with genomic DNA from bRB776.
bMD574 harboring yhdG::Phyperspank-pbpA::cat, ΔpbpA::erm, ΔpbpH::spec was generated upon transformation of bMD573 with genomic DNA from bRB776.
bMD590 containing Pxyl-pbpA::erm was generated upon transformation of Py79 with a four-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1233 bp fragment containing sequence upstream of the pbpA gene (amplified from Py79 genomic DNA using primers oMD75 and oMD125); (2) the 1673 bp erythromycin-resistance cassette loxP-erm-loxP (amplified from pWX467 using primers oJM028 and oJM029); (3) a 1537 bp fragment containing the xylR gene, and the PxylA promoter containing the pbpA ribosomal binding sequence (amplified from pDR150 using primers oMD73 and oMD72); and (4) a 1230 bp fragment containing the pbpA coding region (amplified from Py79 genomic DNA using primers oMD393 and oMD68).
bMD597 harboring Pxyl-pbpA::erm, ΔpbpH::spec was generated upon transformation of bMD590 with genomic DNA from bDR2487.
bMD557 containing amyE::Pxyl-mreBCD::erm, yhdG::Phyperspank-rodA::cat was generated upon transformation of bMK355 with genomic DNA from bMD556.
bMD571 harboring amyE::Pxyl-mreBCD::erm, yhdG::Phyperspank-pbpA::cat was generated upon transformation of bMK355 with genomic DNA from bMD563.
bMD631 containing yhdG::Pspank-pbpA::phleo, ΔpbpH::spec, ΔpbpA::cat, Pxyl-rodA::erm was generated upon transformation of bRB773 with genomic DNA from bMD592.
bYS201 harboring HaloTag-11aa-pbpA::cat was generated upon transformation of Py79 with a four-piece Gibson assembly reaction, that contained the following PCR products. (1) A 1233 bp fragment containing sequence upstream of the pbpA gene (amplified from Py79 genomic DNA using primers oMD75 and oMD125); (2) the 1139 bp chloramphenicol-resistance cassette loxP-cat-loxP (amplified from pWX465 using primers oJM028 and oJM029); (3) a 962 bp fragment containing the native pbpA promoter, and B. subtilis-optimized coding sequence of the HaloTag protein (amplified from double-stranded DNA custom-ordered from DNA 2.0/ATUM [Newark, CA] using primers oYS598 and oYS599); and (4) a 1229 bp fragment containing the pbpA coding region (amplified from Py79 genomic DNA using primers oMD68 and oMD69).
bMK385 harboring amyE::Pxyl-mreBCD::erm, ΔmreBCD, PmreB-minCD::spec, HALO-11aa-pbpA::cat was generated upon transformation of bMD545 with genomic DNA from bYS201.
bYS19 containing mreB-SW-msfGFP was generated upon transformation of bMD88 12 with a three-piece Gibson assembly reaction that contained the following PCR products. (1) A 898 bp PCR fragment containing sequence upstream of the mreB gene (amplified from Py79 genomic DNA using primers oMD134 and oYS34); (2) a 774 bp fragment containing B. subtilis-optimized coding sequence of the msfGFP fluorescent protein (amplified from plasmid DNA custom-ordered from DNA 2.0/ATUM [Newark, CA] using primers oYS35 and oYS36); and (3) a 1629 bp fragment containing sequence downstream of the mreB gene (amplified from Py79 genomic DNA using primers oYS37 and oMD116). This strain is markerless, and selection was in the presence of 0.5% glucose for colonies that no longer required xylose for viability.
bYS977 harboring amyE::Pxyl-mreB-SW-msfGFP, mreCD::erm, was built identical to bMD277, except that PCR product (4) was instead a 3234 bp fragment containing mreB-SW-msfGFP, mreC, and mreD, amplified from bYS19 genomic DNA using primers oMD334 and oMK221).
bYS979 containing amyE::Pxyl-mreB-SW-msfGFP, mreCD::erm, ΔmreBCD, ΔminCD::cat was generated upon transformation of bYS977 with genomic DNA from bMD543.
bYS981 harboring amyE::Pxyl-mreB-SW-msfGFP, mreCD::erm, ΔmreBCD, PmreB-minC,D::spec was generated upon transformation of bYS979 with genomic DNA from bMD545.
AV88 containing 186::Ptet-dCas9, mreB-SW-msfGFP was obtained by transduction of strain LC69 with a P1 phage lysate obtained from strain NO50, and subsequent excision of the kanamycin resistance cassette by transient expression of a flippase. To make the plasmids expressing single-guide RNAs, the psgRNA plasmid was amplified three times with primers pairs V162/V165, V163/V166 and V164/V167. The three generated fragments were assembled together by Gibson assembly to obtain the pAV20 vector. To insert the anti-GFP single-guide RNAs into this vector, we annealed and phosphorylated the following pairs of oligonucleotides: V275/V283 for G5; V274/V282 for G10; V273/V281 for G14; and V272/V280 for G20. These were then inserted in pAV20 by restriction cloning with BsaI. As pAV20 carries two sgRNA insertion sites, a non-targeting oligonucleotide pair (V279/V287) was inserted into a second site during the same reaction.
Statistics
All P-values are reported in figures. P-values were calculated in GraphPad Prism using the Mann-Whitney test with a two-tailed P value. Means, SD, and N for all data points are reported in table S1.
Acknowledgements
We would like to thank Carl Wivagg, Ye Jin Eun, Leigh Harris, and Thomas Bernhardt for helpful advice and discussions, Georgia Squyres for reading of the manuscript and TIRF-SIM acquisition, and Luke Lavis for his generous gift of JF dyes. We thank Z. Gitai and K.C. Huang for providing strains. TIRF-SIM was performed at the Advanced Imaging Center at Janelia Research Campus, a facility jointly supported by the Gordon and Betty Moore Foundation and Howard Hughes Medical Institute. This work was funded by National Institutes of Health Grants R01GM114274 to RO, and DP2AI117923-01 to EG, as well as a Smith Family Award, Searle Scholar Fellowship, and support from the Volkswagen Foundation to EG and SVT. This work was performed in part at the Center for Nanoscale Systems at Harvard University, supported by NSF ECS-0335765.