MreB shares a common prokaryotic ancestor with actin and is present in almost all rod-shaped bacteria. MreB proteins have been implicated in a range of important cell processes, including cell morphogenesis, chromosome segregation and cell polarity. The mreB gene frequently lies at the beginning of a cluster of genes, immediately upstream of the conserved mreC and mreD genes. RNA analysis showed that in Bacillus subtilis mreB is co-transcribed with mreC and that these genes form part of an operon under the control of a promoter(s) upstream of mreB. Construction of an in-frame deletion of mreB and its complementation by mreB+ only, in trans, established that the gene is important for maintenance of cell width and cell viability under normal growth conditions, independent of polar effects on downstream genes. Remarkably, virtually normal growth was restored to the mreB null mutant in the presence of high concentrations of magnesium, especially when high concentrations of the osmoprotectant, sucrose were also present. Under these conditions, cells could be maintained in the complete absence of an mreB gene, with almost normal morphology. No detectable effect on chromosome segregation was evident in the mutant, nor was there an effect on the topology of nascent peptidoglycan insertion. A GFP–MreB fusion was used to look at the localization of MreB in live cells. The pattern of localization was similar to that previously described, but no tight linkage to nucleoid positioning was evident. Propagation of the mreB null mutant in the absence of magnesium and sucrose led to a progressive increase in cell width, culminating in cell lysis. Cell division was also perturbed but this effect may be secondary to the disturbance in cell width. These results suggest that the major role of MreB in B. subtilis lies in the control of cell diameter.
Most bacterial cells have a characteristic morphology. In rod-shaped bacteria, such as Escherichia coli and Bacillus subtilis, a number of genes involved in cell shape have been identified. Most of these are associated with cell wall synthesis or assembly. The mreB family of genes have recently come to the fore through the discovery that they encode actin homologues (van den Ent et al., 2001; Jones et al., 2001). The proteins undergo dynamic assembly/disassembly into filamentous structures, which have now been observed in a range of different bacteria (Carballido-López and Errington, 2003a; Shih et al., 2003; Figge et al., 2004; Gitai et al., 2004). The filaments or cables typically follow a helical path running around the cell periphery, and are probably closely apposed to the inner surface of the cytoplasmic membrane.
The location of mreB immediately upstream of mreC and mreD poses problems for dissecting its specific function in B. subtilis. Here we describe the construction of a non-polar in-frame deletion of mreB, and its use to investigate the specific functions of this gene in B. subtilis. We also describe the construction of a functional gfp–mreB fusion and its use in characterizing the localization of MreB in live cells under various conditions. The results provide strong support for a role in control of cell width but appear to exclude an important role in chromosome segregation.
mreB is co-transcribed with mreC in an operon
Previous results have suggested that mreB is essential and required for shape control (Jones et al., 2001) as well as for chromosome segregation (Soufo and Graumann, 2003). However, all of the previous experiments were complicated by possible polar effects on downstream gene expression. Previous work from several laboratories has suggested that the downstream genes mreC and mreD are both essential (Levin et al., 1992; Lee and Stewart, 2003; M. Leaver and J. Errington, unpublished). An RNase protection assay was used to examine whether mreB is transcribed as part of an operon with mreC. An RNase protection probe (‘mreBC’) was designed to produce a 360 nucleotide protected fragment when added to RNA from the wild-type cells, if mreB and mreC are co-transcribed, or to produce protected fragments of ∼200 nucleotides and ∼145 nucleotides if they are initiated from different promoters (Fig. 1A). The RNase protection using the mreBC probe produced a single fragment of 360 nucleotides from the wild-type cells, showing that mreB is co-transcribed with mreC.
MreB is essential
To investigate the function of mreB but avoid possible polar effects on mreC or mreD an in-frame deletion of mreB was constructed, lacking all but the first and last 42 bases of the mreB coding region. The deletion was introduced into B. subtilis strain (2056) carrying a xylose-inducible copy of the mreBCD cluster (Pxyl-mreBCD) at the amyE locus. The resultant strain (3722) was xylose-dependent. When DNA from this strain (3722) was transformed into a wild-type recipient strain (168) few colonies were obtained and all of these grew poorly. Analysis of their genotype revealed that they had all been co-transformed with the Pxyl-mreBCD construct at the amyE locus (the mreB and amyE loci are distant from each other and should be unlinked by transformation), indicating that mreB is essential.
If the in-frame deletion had no polar effect on downstream genes then complementation with mreB alone should generate cells with wild-type growth and morphology. The deletion of mreB was transformed into recipient strains containing ectopic copies of either Pxyl-gfp-mreB (3723) or PspacHY-mreB (3724). In both cases, transformants were readily obtained and the transformants had retained the inducible mreB-only constructions at amyE. Therefore, both PspacHY-mreB and Pxyl-gfp-mreB complemented the ΔmreB mutation. With the Pxyl-gfp-mreB construct, normal colonial growth required the addition of about 0.5% xylose (Fig. 2A). With the very strong and leaky PspacHY promoter, mreB function was obtained in the absence of inducer. The Pxyl-gfp-mreB ΔmreB construct allowed the phenotype arising specifically from depletion of MreB to be characterized. Depletion of the GFP–MreB, by the removal of xylose, caused cells to lose control over their cell width, becoming progressively swollen over several generations and eventually lysing (Fig. 2C).
MreB mutants can be stabilized in SMM medium
The cell lysis that occurred during depletion of MreB complicated interpretation of the phenotype. Previous observations with a disruption of mreC, suggested that addition of an osmotic stabilizer might avoid the problem of lysis at least partially (Lee and Stewart, 2003). A mixture of sucrose, MgCl2 and maleic acid (SMM) that had previously been used to stabilize protoplasts (Chang and Cohen, 1979), was added to the growth medium (PAB) and did indeed result in the amelioration of the ΔmreB phenotype. Surprisingly, in the SMM-supplemented medium the mreB-depleted cells maintained an almost wild-type rod shape (Fig. 2E; compare with 2C).
On the basis of the above result it seemed possible that an mreB null mutant could be constructed and propagated in the presence of SMM. The ΔmreB mutation was transformed into B. subtilis wild-type strain (168), as above, but this time in the presence of SMM. Now many transformants were obtained, and the major class of transformants were SMM-dependent (Fig. 3A and B) and did not contain an ectopic copy of mreB at amyE. The absence of mreB from this strain was confirmed by polymerase chain reaction (PCR) (not shown) and Western blotting (Fig. 3D). An RNase protection assay was used, as described earlier, to examine whether transcription through to mreC is affected by the deletion of mreB. The RNase protection using the mreBC probe produced a single fragment of about 220 nucleotides with the ΔmreB RNA (Fig. 1B). This showed that in the ΔmreB strain read-through into mreC occurs. Taken together with the fact that analogous in-frame deletions of mreC and mreD both exhibit a different, spheroidal morphology in SMM (M. Leaver and J. Errington, unpublished), the ΔmreB mutation does not appear to impair the function of the downstream genes. The lack of a 360 nucleotide protected fragment in the mutant further confirmed that the mreB gene has been deleted from these cells.
MreB mutants require increased levels of Mg2+ for cell viability
The above experiments established that the ΔmreB strain is devoid of the MreB protein and viable in the presence of SMM. Initially it was thought that the sucrose-containing medium acted as an osmotic protectant, thereby preventing cell lysis. Surprisingly, further investigation revealed that stabilization of the ΔmreB mutant only required the Mg2+ component of SMM. Growth of the ΔmreB strain in PAB with no additions resulted in cell lysis (Fig. 3G), in the presence of 0.3 M sucrose the cells became swollen (Fig. 3I) whereas in the presence of 25 mM MgCl2 the cells maintained a rod shape (Fig. 3H) and the growth rate was indistinguishable from wild type (Fig. 3E). Addition of 2.5 mM MgCl2 or MgSO4 was sufficient to restore wild-type growth rate. Other divalent cations were also examined for their ability to rescue the ΔmreB mutant (not shown). Concentrations of up to 1 mM Mn2+ did not stimulate growth and precipitation prevented higher concentrations from being tested. Precipitation was also a problem with high calcium concentrations. However, 1 mM Ca2+ partially restored growth, although at a substantially lower rate than observed with 1 mM Mg2+.
Although addition of 25 mM Mg2+ produced an almost normal rod shape, the cells were wider than wild-type cells. However, combination of sucrose and MgCl2 gave an almost wild-type morphology (Fig. 3J) so the SMM medium was subsequently used to study the effects resulting from the loss of mreB in the absence of morphological change.
Morphological effects of the ΔmreB mutation
The process of cell lysis was investigated using time-lapse microscopy. Cultures were grown in PAB containing SMM and then diluted into PAB (no added magnesium). Cells were then incubated for 30 min at 37°C before the cell samples were transferred onto a thin film of 1.2% agarose (containing PAB medium but no magnesium) for microscopy. A typical time-lapse is shown in Fig. 4 (Supplementary material,Fig.S1). Initially the ΔmreB cells elongated rapidly but then they became progressively wider (arrowhead A). This widening appeared to occur at or near new division sites, whereas the old poles retained a constant width. It appeared that as the ΔmreB cells grew and came into contact with each other they twisted and some cells burst (arrowheads B and D). Lysis of one cell often appeared to be followed by the spreading of lysis to adjacent cells in the chain. The cytoplasmic contents of these adjacent cells gradually faded away without swelling (sequence of cells beginning at arrowhead C and ending with arrowhead E), suggesting that the chains of cells were not separated off from each other by completed division septa.
In contrast, wild-type cells grew and divided normally under these conditions (Supplementary material, Fig.S2). The addition of 25 mM magnesium to the slide also resulted in normal growth of the ΔmreB strain (Supplementary material, Fig.S3); the cells maintained a rod shape and no lysis was observed. The addition of only sucrose to the growth medium resulted in a retarded growth rate with the cells twisting and swelling (Supplementary material, Fig.S4).
MreB is not required for cylindrical cell wall synthesis
We previously reported that in B. subtilis Mbl but not MreB is required for cylindrical cell wall synthesis (Daniel and Errington, 2003). Fluorescently labelled vancomycin (Van-FL) is thought to stain sites that are involved in nascent PG synthesis (Daniel and Errington, 2003). Growth of the ΔmreB strain in the presence of SMM allowed the sites of newly incorporated PG to be visualized in the absence of significant morphological defects. As shown in Fig. 5A the ΔmreB mutant cells showed patches of Van-FL staining along the length of the cell cylinder and at likely division sites, indistinguishable from the patterns obtained with wild-type cells (Fig. 5C).
No significant chromosome segregation defect in the complete absence of MreB
Soufo and Graumann (2003) have reported that MreB is required for proper chromosome segregation and that following depletion of MreB large numbers of anucleate cells are produced. Unfortunately, these experiments were complicated by both possible polar effects on the downstream genes by using a depletion strain and the extreme morphological defects observed for mreB mutant cells, which could potentially impact on chromosome segregation or maintenance indirectly. Availability of a complete null mutant of mreB and the ability to grow cells in the presence of SMM allowed us to assess the role of mreB in chromosome segregation. ΔmreB cultures were grown in several different media supplemented with SMM and samples were taken during the exponential phase of growth. The samples were then fixed with glutaldehyde and the DNA was stained with 4,6-diamidino-2-phenylindole (DAPI). The number of anucleate cells produced by the ΔmreB mutant in a series of experiments was always <0.1% (n = 1000), which was indistinguishable from wild-type cultures. Images of mutant and wild-type cells from a typical experiment are shown in Fig. 6. No significant impairment of chromosome maintenance was evident.
Helical localization of GFP–MreB
We previously reported that a GFP fusion to the N-terminus of MreB was non-functional and did not localize (Jones et al., 2001). However, by using a different length linker between the GFP and MreB a functional gfp–mreB fusion was generated (strain 2565). The fusion, controlled by a xylose-inducible promoter (Pxyl), was integrated into the chromosome at the mreB locus. The fusion was judged to be functional as the cells were viable in the presence of xylose and the fusion was the only copy of mreB present in the cell. The GFP–MreB fusion showed a helical pattern of localization (Fig. 7A) with one or two discrete structures per cell, similar to the configuration described previously on the basis of immunofluorescence microscopy (Jones et al., 2001). Analysis of 3D reconstructions suggested that the structures observed generally corresponded to 1–1.25 helical turns around the periphery of the cell. The number of such structures tended to increase in parallel with cell length. Unfortunately no consistently identifiable structure could be found that was associated with any particular cell length, as reported by Defeu Soufo and Graumann (2004). In contrast to their report, we also observed helical structures during stationary phase. The appearance of the GFP–MreB fusion differed from that previously described for GFP–Mbl in the presence of spaces between discrete helical elements and in the general absence of fluorescence in the polar regions of the cells (Fig. 7B; arrows). A strain (2566) containing a gfp–mreB construct at an ectopic locus (amyE), as well as the wild-type chromosomal copy of mreB, was also created and this showed a similar helical pattern of localization (data not shown).
MreB helical filaments form independently of the nucleoid
Chromosome segregation is thought to be an active process (Sharpe and Errington, 1999) and it has been suggested that mreB may play an important role in this process (Soufo and Graumann, 2003). Consistent with this idea, it has been reported that the localization of MreB is influenced by the nucleoid (Defeu Soufo and Graumann, 2004). To investigate a possible relationship between MreB and the nucleoid during the cell cycle, we looked at the localization of MreB under conditions in which chromosome replication is uncoupled from cell growth. In the presence of the replication elongation inhibitor HPUra (Brown, 1972), cells elongate at about the normal rate but chromosomes fail to replicate and segregate and division is delayed. Cells expressing gfp–mreB (strains 2565 and 2566) were treated with HPUra and, after a sufficient time for several doublings, were viewed microscopically to determine the localization of GFP–MreB. Figure 7D shows a typical chain of filamentous cells formed following HPUra treatment. The regions that lacked DNA still contained helix-like structures (Fig. 7D; arrowheads), showing that the placement of such structures is not dependent on the nucleoid. Although the GFP–MreB signal tended to be brighter around the nucleoid, this might reflect the site where de novo synthesis of MreB is.
As an alternative way to look for a possible link between MreB localization and the nucleoid, cells depleted for DnaA, a key initiator protein for DNA replication, were analysed. Depletion of DnaA also uncouples cell growth and chromosome segregation except that the block in DNA replication is at the initiation stage. An ectopic copy of gfp–mreB was introduced into cells bearing a conditional (IPTG-dependent) copy of dnaA (strain 3702). In the absence of inducer MreB localization again occurred independently of the nucleoid, along the length of the cell (Fig. 7C). Furthermore, the arrowhead in Fig. 7C points to a typical anucleate cell formed under these conditions, which clearly contained MreB structures. The above results established that MreB structures are not obligatorily associated with nucleoids.
Construction of an in-frame deletion of mreB
The key technical advance reported here is the construction of a complete deletion of mreB that does not affect expression of the downstream genes in the mre operon. Expression of an ectopic copy of mreB alone was sufficient to complement this in-frame deletion, showing that the deletion has no significant polar effect on mreC or mreD. Repression of the ectopic mreB copy resulted in increased cell width and ultimately cell lysis (Fig. 2C).
Surprisingly, an increased level of Mg2+ dramatically ameliorated the phenotype of the ΔmreB mutant enabling an mreB null mutant to be constructed. The discovery that an mreB null could be grown in the presence of SMM enabled the roles of mreB in B. subtilis to be studied in the absence of significant morphological defects.
Absence of a chromosome segregation defect in the ΔmreB mutant
It has recently been reported that MreB has a major role in chromosome segregation and that upon depletion of MreB the number of anucleate cells rises dramatically to ∼25% of the cell population (Soufo and Graumann, 2003). The MreB depletion strategy used by Soufo and Graumann (2003) was complicated by two factors. First, the severe morphological defects that arise when MreB is depleted. Second, likely polar effects on downstream gene expression resulting from depletion of the first gene of the mre operon. By growing the ΔmreB mutant under conditions in which the rod shape was maintained, the possible effect on chromosomal segregation could be reassessed. As shown in Fig. 6, the frequency of anucleate cell formation under these conditions was insignificant and the efficiency of segregation was indistinguishable from that of the wild type. Thus, MreB probably does not have a significant role in chromosome segregation in B. subtilis. However, it remains a possibility that one of the other actin homologues, mbl or mreBH (Carballido-López and Errington, 2003b), helps to support chromosome segregation in this organism.
We also investigated if the chromosomes exerted any influence on the localization of MreB. Localization of a fully functional GFP–MreB fusion protein (Fig. 7A) was similar to that observed previously by immunofluorescence (Jones et al., 2001) and to results reported elsewhere (Defeu Soufo and Graumann, 2004). When chromosome replication was uncoupled from cell extension, MreB localized almost normally over areas with or without the nucleoid. Strikingly MreB structures could be found in anucleate cells (arrowhead Fig. 7C). It seems likely that effects on chromosome segregation in cells affected for mreB function are a secondary consequence of the effects on cell morphology.
Possible role of MreB in control of cell diameter
The loss of MreB from B. subtilis causes cells to become progressively wider over several generations. This was clearly seen when the ΔmreB mutant was grown in the absence of Mg2+ (Fig. 4; Supplementary material, Fig.S1). In this case the cells became wider and new poles were often misshapen, perhaps because the division plane is less precisely orientated when the cell has an increased diameter. Following prolonged growth the cells swelled and eventually burst. This appeared to be stimulated by contact with other cells, perhaps due to torsional stresses. Interestingly, when lysis occurred it often appeared to spread to adjacent cells in the chain, again suggesting a possible effect on cell division in the ΔmreB mutant. Alternatively, this could be an indirect consequence of the increased cell diameter. Loss of MreB may lead to a change in resistance of the cell wall to osmotic or mechanical stress, due to the absence or incorrect assembly of a cell wall component. At present it remains unclear precisely how MreB exerts control over cell diameter.
Amelioration of the lethal phenotype of mreB by Mg 2+
The mechanism by which Mg2+ was able to restore an almost wild-type cell morphology and growth rate to the ΔmreB mutant is currently unknown. Mg2+ has many important cellular roles and there are several general areas related to the cell wall that it might influence, including: the level of cross-linking in the cell wall, peptidoglycan structure and conformation, anionic polymer structure and conformation, and the stabilization or activity of proteins or enzyme complexes involved in cell wall synthesis. Consistent with one or more roles in this area, peptidoglycan precursors have been reported to build up in cells limited for Mg2+ (Garrett, 1969). This raises the formal possibility, which we cannot yet exclude, that mreB has a role in Mg2+ uptake. However, mreB is not the first mutant to be reported as being rescued by high Mg2+ concentrations. mreD is a membrane-bound protein currently thought to be required for peptidoglycan synthesis in the cylindrical part of the cell (Burdett, 1979; M. Leaver and J. Errington, unpubl. data). A temperature-sensitive mreD mutant (rodB1 mutation) (Rogers et al., 1968; Varley and Stewart, 1992) displaying a reduced growth rate and spheroidal morphology, was rescued by high Mg2+ concentration (Rogers et al., 1976; Rogers and Thurman, 1978). This mutant was shown not to be significantly affected in Mg2+ uptake. More recently, cells lacking penicillin-binding protein 1 (ponA), a protein which localizes at the septum and forms part of the cell division machinery with a role in synthesis of septal PG (Scheffers and Errington, 2004; Scheffers et al., 2004), have also been shown to require increased concentrations of divalent cations for growth (Murray et al., 1998). Mendelson and Favre (1987) reported that macrofibre twist development in B. subtilis was also influenced by the concentration of Mg2+. The 3D organization of macrofibres is thought to reflect some feature of the mechanism of peptidoglycan incorporation into the wall.
The ability of Mg2+ to rescue mutants affected in two different aspects of peptidoglycan synthesis (i.e. mreD and ponA) as well as mreB, suggests that it might act relatively indirectly, for example by stiffening the cell envelope, so as to counteract the deleterious effects on peptidoglycan synthesis or assembly. In this case, MreB might have a function in assembly or maturation of the peptidoglycan. However, the pattern of Van-FL staining in the mreB mutant in the presence of Mg2+, was normal (Fig. 4).
The other major cell wall polymers involved in cell shape determination are the anionic polymers, teichoic acid and teichuronic acid (Rogers et al., 1968; 1971; Boylan and Mendelson, 1969; Boylan et al., 1972; Pollack and Neuhaus, 1994). Under Mg2+ limitation the amount of teichoic acid present in the wall increases (Ellwood, 1970). This supports the idea that anionic polymers are required for the scavenging and uptake of cations from the environment. Little is known about the localization and mechanism of insertion of the teichoic acids into the cell wall and it remains a possibility that MreB is required for the coordination of this process. In this case, increased levels of Mg2+ may help to stabilize cells with randomly incorporated teichoic acid.
One final facet of wall function in which MreB could act would be in the organization of cell wall turnover. This could involve the coordination of the activity of the cell wall autolysins, perhaps by spatially organizing their export or localization.
In conclusion, in the ΔmreB mutant cylindrical extension of the cell wall occurs, probably, via the helical mode of insertion dependent on Mbl (Daniel and Errington, 2003), but the diameter of the cell gradually increases. Eventually, catastrophic width increase occurs, followed by lysis. In principle, it seems that MreB might act continuously to restrain the diameter of the elongating cell. Alternatively it could act discontinuously, e.g. at the division site, to reset the correct diameter each time the cell divides. A possible defect in cell division has been noted above and it may be relevant that in C. crescentus, at least, MreB is seen to switch from a helical localization to become concentrated in a region that will become the future plane of cell division. Whatever the mechanism, the ability of B. subtilis to regulate its cell width is essential to its survival and the role of mreB in this process remains an important question for future work.
Bacterial strains and plasmids
Bacillus subtilis strains and plasmids used in this study are listed in Table 1.
Table 1. B. subtilis strains and plasmids used in this study.
Bacillus subtilis strains were grown in either Difco antibiotic medium 3 (PAB), casein hydrolysate (CH) medium (Sterlini and Mandelstam, 1969) or S medium (Sharpe et al., 1998) supplemented, where required, with xylose (0.5%) and/or IPTG (0.5 mM). Where SMM was added the above media were made at 2× concentration and diluted 50:50 with a 2× SMM solution (1 M sucrose, 33.7 mM maleic acid, 40 mM MgCl2, pH 7.0). E. coli strains were grown at 37°C in 2× YT (Sambrook et al., 1989), supplemented with ampicillin (100 µg ml−1) as required.
DNA manipulations and E. coli DH5α transformations were carried out using standard methods (Sambrook et al., 1989). B. subtilis strains were transformed according to the method of Anagnostopoulos and Spizizen (1961) as modified by Jenkinson (1983). Selection for B. subtilis transformants was carried out on nutrient agar (Oxoid), supplemented with antibiotics, as required, with: kanamycin (5 µg ml−1) chloramphenicol (5 µg ml−1), erythromycin (1 µg ml−1), lincomycin (25 mg ml−1) and/or spectinomycin (50 µg ml−1), with xylose (0.5%), IPTG (0.5 mM) and SMM (0.5×), as necessary.
Construction of GFP fusions
A 400 base pair segment of the promoter-proximal part of mreB was amplified with primers mreB1 and mreB2 using the PCR and inserted into the XhoI and HindIII sites of plasmid pSG4902 (Wu and Errington, 2003), to produce plasmid pSG5451. The previously reported non-functional N-terminal GFP–MreB fusion used the BamHI site instead of the XhoI site (Jones et al., 2001). All primers used in this study are listed in Supplementary material (TableS1). Transformation of pSG5451 into B. subtilis strain 168 with selection for chloramphenicol resistance produced a xylose-dependent strain (2565), which carries a gfp fusion to mreB integrated into the chromosome at the mreB locus, as the only copy of the mreB in the cell and controlled by the Pxyl promoter. To generate a xylose-inducible copy of gfp–mreB at the amylase locus, full-length mreB was cloned using primers mreB1 and mreB3 into the XhoI and HindIII sites of pSG1729, which carries a selectable spectinomycin resistance marker (spc) (Lewis and Marston, 1999). The resultant plasmid (pSG5452) was transformed into B. subtilis strain 168 with selection for spectinomycin resistance, producing strain 2566.
To study the gfp–mreB fusion in a DnaA depletion background pSG5452 was transformed into strain PL10 with selection for spectinomycin resistance, creating strain 3702.
For fluorescence microscopy, cells from an overnight culture or from a culture grown to mid-exponential stage from a fresh overnight plate were diluted into fresh S medium containing 0.5% xylose and grown to late exponential phase/early stationary phase at 30°C. For live cell imaging, cells were mounted on microscope slides covered with a thin film of 1.2% agarose in water, essentially as described previously (Glaser et al., 1997). For time-lapse microscopy cells were mounted on microscope slides fitted with GeneFrames (ABgene) on a thin film of 1.2% agarose in PAB. When Nile red was used for staining the membranes, 5 µl of the culture was mixed with 1.2 µl of the Nile red (Molecular Probe) solution (12.5 µg ml−1) in an Eppendorf. When nucleoid staining was required a 6 µl sample of cell culture was mixed with 2 µl of DAPI (Sigma) solution (1 µg ml−1 in 50% glycerol) for 1 min before viewing. Image acquisition was performed as described previously (Lewis and Errington, 1997) using a Sony CoolSnap HQ cooled CCD camera (Roper Scientific) attached to a Zeiss Axiovert 200M microscope. The digital images were acquired and analysed with METAMORPH version 6 software. Fluorescent images were deconvolved within METAMORPH and assembled in Adobe PHOTOSHOP version 7.
Inhibition of DNA replication using HPUra
Cells were grown in S medium (Sharpe et al., 1998) and diluted back into the same medium to an OD600nm of 0.05 then grown to exponential phase. The DNA polymerase III inhibitor 6-(p-hydroxy-phenylazo)-uracil (Brown, 1972) was added to a final concentration of 200 µM to block DNA replication and the cells were incubated for several doubling times.
Depletion of MreB, DnaA or magnesium
To deplete MreB or DnaA the relevant cultures were grown in PAB, CH or S medium with 0.5% xylose (MreB) or 0.5 mM IPTG (DnaA) and diluted back to an OD600nm of 0.05 into medium with or without xylose or IPTG, respectively, and then incubated. To remove magnesium, cells were grown in PAB, supplemented with 25 mM magnesium to mid-exponential phase and then diluted back to an OD600nm of 0.05 into medium with differing levels of magnesium.
Construction of an in-frame deletion of mreB
Regions of DNA extending from just inside the borders of mreB to points upstream and downstream of mreB (∼4.5 kb each side) were PCR-amplified, using primers mreB4–7, from a strain (3427) containing a selectable kanamycin resistance determinant (neo) inserted just upstream of mreB (Ωneo3427). The products of these PCRs were then ligated together and reamplified with the outside end primers: mreB4 and mreB7. Transformation of the resulting PCR product into B. subtilis strain 2056 with selection for kanamycin resistance in the presence of xylose gave rise to a strain (3722) carrying an in-frame deletion of mreB (ΔmreB) and a xylose-inducible copy of the mreBCD cluster (Pxyl-mreBCD) at an ectopic locus (amyE). Chromosomal DNA of this strain was transformed into B. subtilis strain 2566 with selection for kanamycin resistance in the presence of xylose to replace the inducible mreBCD cluster with Pxyl-gfp-mreB generating strain 3723.
A copy of mreB was placed under PspacHY control at amyE by cloning a full-length copy of mreB into pPL82, which caries a cam selectable marker. Primers mreB8 and mreB9 were used to amplify mreB and this was inserted into the SphI and SpaI sites of pPL82 (Quisel et al., 2001). Transformation into B. subtilis strain 168 and selection for chloramphenicol resistance produced strain 3704. Chromosomal DNA from strain 3722 was then transformed into B. subtilis strain 3704 with selection for kanamycin resistance in the presence of IPTG, generating a strain (3724) with the in-frame deletion of mreB complemented by PspacHY-mreB.
Bacillus subtilis strain 168 was transformed with chromosomal DNA from strain 3722 with selection for kanamycin resistance in the presence of 0.5 M SMM. The transformants were screened for SMM dependence, AmyE+ phenotype and PCR was used to check for the loss of mreB giving strain 3725.
Culture samples (2 ml) were grown in the presence of SMM, as for microscopy, centrifuged and the cell pellets were frozen immediately in liquid nitrogen. Once sampling was complete, 200 µl of sample loading buffer (with proteinase inhibitor complete added; Roche) was added to the frozen pellet, and the samples were held on ice before sonication (two pulses of 10 µm amplitude for 12 s). The samples were then diluted 3:1 in 4× SDS-PAGE sample buffer, heated to 95°C for 5 min, and equal amounts of protein (adjusted according to the OD of the original culture) were separated by 12% SDS-PAGE. Proteins were subsequently transferred to a Hybond-P polyvinylidene difluoride membrane (Amersham Biosciences) by Western blotting. MreB was detected with anti-MreB antiserum 1:10 000 (Jones et al., 2001). Western blots were developed using an ECL detection kit (Amersham Biosciences).
Total RNA was isolated using a hot-phenol method (Furger et al., 2001). The total RNA pellets were resuspended in R-loop buffer (Ashe et al., 1995). For mreBC transcriptional analysis a 360 nucleotide region spanning mreB and mreC was PCR-amplified using primers BRNA1 and BRNA2. The DNA was then transcribed with SP6 RNA polymerase according to the manufacture's instructions (Promega) to make the ‘mreBC’ riboprobe. Full-length riboprobe was purified from a denaturing polyacrylamide gel in 300 µl of buffer (0.5 M ammonium acetate, 0.1% SDS, 1 mM EDTA) for 2 h at 37°C. Eluted RNA was ethanol-precipitated, resuspended in water and 100 cp of probe was hybridized with total RNA. RNase protection analysis was performed as described by Ashe et al. (1995).
Van-FL staining was carried out using vancomycin, BODIPY® FL conjugate (Molecular Probes) mixed with an equal amount of unlabelled vancomycin (final concentration 1 µg ml−1). Samples were then fixed and prepared as described previously (Daniel and Errington, 2003).
We thank Dr A. Furger for help and advice with the RNase protection. We thank members of our group for discussions and advice, notably M. Leaver for the gift of strain 3427, Dr H. Murray, Dr L. Hamoen and Dr R. Daniel for critical reading of the manuscript. This research was supported by grants from the Human Frontier Science Program and the UK Biotechnology and Biological Sciences Research Council.