The major and minor wall teichoic acids prevent the sidewall localization of vegetative dl-endopeptidase LytF in Bacillus subtilis

Authors


*E-mail jsekigu@shinshu-u.ac.jp; Tel. (+81) 268 21 5344; Fax (+81) 268 21 5345.

Summary

Cell separation in Bacillus subtilis depends on specific activities of dl-endopeptidases CwlS, LytF and LytE. Immunofluorescence microscopy (IFM) indicated that the localization of LytF depended on its N-terminal LysM domain. In addition, we revealed that the LysM domain efficiently binds to peptidoglycan (PG) prepared by chemically removing wall teichoic acids (WTAs) from the B. subtilis cell wall. Moreover, increasing amounts of the LysM domain bound to TagB- or TagO-depleted cell walls. These results strongly suggested that the LysM domain specifically binds to PG, and that the binding may be prevented by WTAs. IFM with TagB-, TagF- or TagO-reduced cells indicated that LytF−6xFLAG was observed not only at cell separation site and poles but also as a helical pattern along the sidewall. Moreover, we found that LytF was localizable on the whole cell surface in TagB-, TagF- or TagO-depleted cells. These results strongly suggest that WTAs inhibit the sidewall localization of LytF. Furthermore, the helical LytF localization was observed on the lateral cell surface in MreB-depleted cells, suggesting that cell wall modification by WTAs along the sidewall might be governed by an actin-like cytoskeleton homologue, MreB.

Introduction

The bacterial cell wall (CW) is mainly composed of mesh-like peptidoglycan (PG) and covalently linked anionic polymers such as wall teichoic acids (WTAs). PG is built from long-glycan strands cross-linked by peptide side-chains (Warth and Strominger, 1971; Foster and Popham, 2002). It was reported that CW assembly in Bacillus subtilis occurred in both the cylindrical part of the wall and the septum (Mobley et al., 1984; Clarke-Sturman et al., 1989; Merad et al., 1989). Moreover, recent experiments involving fluorescent vancomycin (Van-FL) suggested that PG synthesis of the septum depends on divisome, whereas that of the sidewall occurs in a helical pattern governed by an actin-like homologue, Mbl (Daniel and Errington, 2003). Mbl forms a dynamic helical filament with other actin-like homologues, MreB and MreBH, beneath the cytoplasmic membrane along the sidewall (Jones et al., 2001; Carballido-López and Errington, 2003; Carballido-López et al., 2006). On the other hand, another approach involving Van-FL and ramoplanin, an antibiotic that specifically binds to the reducing end of the nascent glycan chain and lipid II, labelled with a fluorophore, BodipyFl, revealed that the sidewall PG synthesis is governed in an Mbl-independent manner, as the helical pattern along the sidewall was observed even in an mbl null mutant (Tiyanont et al., 2006).

Anionic polymers, which are 35–60% of the vegetative CW, are mainly composed of major and minor WTAs under non-phosphate-limiting conditions (Foster and Popham, 2002). The major WTA-biosynthetic enzymes are encoded by tagA, tagB, tagD, tagE, tagF, tagO, gtaB and tagP (yvyH) (Foster and Popham, 2002; Lazarevic et al., 2002). A recent report has demonstrated that six genes, the exceptions being tagE and gtaB, are essential in B. subtilis (Kobayashi et al., 2003). Moreover, an ABC transporter encoded by two essential genes, tagG and tagH, is required for WTAs translocation and linkage to PG (Lazarevic and Karamata, 1995). Biosynthesis pathway of major WTA was essential but recently, D'Elia et al. (2006) reported that tagO, whose product catalyses the first step in the WTA biosynthesis pathway, is dispensable for cell viability, and that a tagO null mutant shows slow growth, aberrant morphology and septation, and non-uniform PG thickness. In addition, they defined that tagB, tagD and tagF are essential in the presence of tagO, but not in its absence. TagB and TagD are involved in the linkage unit synthesis of major and minor WTAs, and TagF is required for chain polymerization of the major WTA (Foster and Popham, 2002; Lazarevic et al., 2002). It remains to be resolved why only tagO is not essential and why the essential nature of other tag genes can be suppressed by the deletion in tagO. On the other hand, GtaB, a UDP-glucose pyrophosphorylase involved in the glucosylation of the major WTA, is not essential (Soldo et al., 1993; Varón et al., 1993). In addition, the biosynthesis pathway of minor WTA is not essential (Lazarevic et al., 2002; Freymond et al., 2006). The ggaA and ggaB genes are required for the biosynthesis of the galactosamine-containing minor WTA.

The cell separation event following septation is the final step of cell division in bacteria (Errington and Daniel, 2002). B. subtilis produces, at least, three dl-endopeptidases, CwlS (YojL) (Fukushima et al., 2006), LytE (CwlF) (Ishikawa et al., 1998; Margot et al., 1998) and LytF (CwlE) (Margot et al., 1999; Ohnishi et al., 1999), during vegetative growth. A triple mutant lacking these enzymes exhibited aggregated microfibre formation, indicating a cell separation defect (Fukushima et al., 2006). Among these vegetative dl-endopeptidases, LytF plays a major role in cell separation especially after the middle vegetative growth phase (Ohnishi et al., 1999; Yamamoto et al., 2003). The lytF gene is transcribed by EσD RNA polymerase, and a lytF mutant shows a long-chained cell morphology (Ohnishi et al., 1999) similar to that of a sigD mutant (Helmann et al., 1988). On the other hand, the lytE gene is transcribed by EσA and EσH RNA polymerases, and a lytE mutant shows a slightly chained cell morphology, especially in the early vegetative growth phase (Ishikawa et al., 1998; Ohnishi et al., 1999). Carballido-López et al. (2006) have reported that a LytE–GFP fusion is localized not only at cell separation sites and poles but also along the sidewall under slight overexpression conditions. The former localization appears to be a septum-dependent manner, and the latter one in an MreBH-dependent helical manner. MreBH is one of the actin-like homologues and plays an important role in cell morphogenesis by interacting with the C-terminal dl-endopeptidase domain of LytE (Carballido-López et al., 2006). In addition, lytE and mreBH mutants show similar CW-related defects under low-Mg2+ conditions (Carballido-López et al., 2006). Moreover, Bisicchia et al. (2007) reported that the essential YycFG two-component system positively regulates the expression of two vegetative dl-endopeptidase genes, lytE and cwlO (yvcE). They revealed that a lytE cwlO double mutant strain is not viable and that cells depleted of CwlO and lacking LytE exhibit loss of lateral CW synthesis and cell elongation. Based on these findings, it is thought that LytE plays at least two roles; one is in cell separation at the septum, and the other in CW turnover along the sidewall. Recently, it was reported that the cwlS gene is expressed by EσH RNA polymerase during the late vegetative and stationary phases (Britton et al., 2002), and that CwlS is the third vegetative dl-endopeptidase in B. subtilis (Fukushima et al., 2006). Subcellular localization analysis involving immunofluorescence microscopy (IFM) revealed that LytE, LytF and CwlS are potentially localized at cell separation sites and both poles (Yamamoto et al., 2003; Fukushima et al., 2006). Moreover, IFM and Western blot analysis revealed that the enzymes were degraded by CW-bound and extracellular proteases WprA and Epr, respectively, during the vegetative growth phase. The N-terminal domains of CwlS, LytE and LytF include four, three and five tandem repeats of the LysM motif, respectively, which appears to be a general PG-binding module (Bateman and Bycroft, 2000; Buist et al., 2008), separated by serine-rich regions. Thus we presumed that the LysM domains of CwlS, LytE and LytF play an important role in their specific localization at cell separation sites.

In this study, we have demonstrated that the N-terminal CW-binding domain of LytF is required for its specific localization at cell separation site and poles, and that the LysM motif in the domain is involved in the specific binding to naked PG not modified by WTAs. Moreover, the binding to the sidewall was mainly inhibited by anionic polymers, major and minor WTAs, in the vegetative CW.

Results

Localization of LytF depends on the N-terminal CW-binding domain

In this research, we used a LytF−6xFLAG fusion strain to obtain brighter foci than LytF−3xFLAG (Yamamoto et al., 2003) and also confirmed that the LytF−6xFLAG protein retains a cell separation activity as well as LytF (data not shown). We previously reported that the LytF−3xFLAG fusion protein is potentially localized at cell separation sites and cell poles after cell division in a wprA epr double mutant (Yamamoto et al., 2003). To determine whether this specific localization depends on the N-terminal CW-binding domain (CWBE) or the C-terminal catalytic one (CTDE) of LytF, we constructed two strains, YM1047 and YM1051, carrying cwbE– and ctdE−6xflag fusion genes at the lytF locus respectively. Then we carried out IFM to detect the fusion proteins. IFM of the CWBE−6xFLAG expressing cells clearly indicated that the fusion protein was localized at cell separation sites and poles (Fig. 1D–F). The localization pattern was very similar to that of LytF−6xFLAG (Fig. 1A–C). In addition, Western blot analysis indicated that LytF– and CWBE−6xFLAGs were detected in the CW fraction (Fig. 2). On the other hand, we could not observe any CTDE−6xFLAG foci on the cell surface (Fig. 1G–I). As supporting this result, Fig. 2 showed that CTDE−6xFLAG was secreted in the culture medium but not localized on the CW. These results strongly suggest that the specific localization of LytF−6xFLAG depends on the N-terminal CWBE domain including five direct repeats of the LysM motif. Moreover, to examine the septum localization of LytF in PBP 2B-depleted cells, we constructed a pbpB-conditional mutant, HY1054. For this purpose, we put the pbpB gene downstream of an isopropyl-β-d-thiogalactopyranoside (IPTG)-inducible promoter, Pspac. When IPTG was removed, cells began to elongate and form filaments, and the septal localization of LytF was absent (Fig. S1). The result suggests that septal PG synthesized by PBP 2B is required for the specific localization of LytF at cell separation sites, consistent with the LytE–GFP localization, as reported by Carballido-López et al. (2006).

Figure 1.

Localization of the LytF–, CWBE– and CTDE−6xFLAG fusion proteins.
A–C. Phase contrast (A) and LytF−6xFLAG localization (B) images, and an overlay image (C) of (A) and (B) (strain MH1022).
D–F. Phase contrast (D) and CWBE−6xFLAG localization (E) images, and an overlay image (F) of (D) and (E) (strain YM1047).
G–I. Phase contrast (G) and CTDE−6xFLAG localization (H) images, and an overlay image (I) of (G) and (H) (strain YM1051).
The OD600 value at the sampling time was 1.7 (late exponential phase). The exposure times were 0.1 s for phase contrast (A, D and G) and 0.1 s (gain 2) for Cy3 (B, E and H). In the case of (H), the background of the image is raised in the image processing in order to detect weak signals. Scale bars, 10 μm.

Figure 2.

Western blot analysis of LytF–, CWBE– and CTDE−6xFLAG fusions. Cell surface proteins (lane C) and culture supernatant proteins (lane S) were prepared and subjected to Western blot analysis as described in Experimental procedures. The molecular masses of the protein standards (Bio-Rad) are indicated on the left. B. subtilis MH1022 (LytF−6xFLAG; 55 kDa), YM1047 (CWBE−6xFLAG; 43 kDa) and YM1051 (CTDE−6xFLAG; 19 kDa) were cultured in LB medium at 37°C and were harvested at the late exponential phase (OD600, 1.8). Proteins from the cell surface (lane C; equivalent to 5 OD600 cells per lane) and from the culture supernatant (lane S; equivalent to 5 OD600 cells per lane) were applied on a 12% polyacrylamide gel. Asterisks indicate the degradative products of LytF−6xFLAG.

GST−2xLysM protein specifically binds to PG in vitro

To further examine the CW-binding ability of the CWBE and CTDE domains, we constructed two expression plasmids, pGEX−2xLysM, carrying a glutathione-S-transferase (gst)–2xlysM fusion gene, and pGEXEtCTD, carrying a gst–ctdE (catalytic domain of LytF) fusion gene. We were able to purify the GST−2xLysM and GST–tECTD fusion proteins in a soluble fraction of Escherichia coli BL21 (pGEX−2xLysM or pGEXEtCTD) (data not shown). Although we also tried to purify the GST−1xLysM, −3xLysM, −4xLysM and −5xLysM fusion proteins, we could not obtain them as soluble proteins because of their high insolubility (data not shown). Therefore the GST−2xLysM protein was used for the CW-binding assay in vitro. CW was prepared from B. subtilis 168 cells at the transition stage (OD600 ∼ 2.0) in LB medium, and PG was prepared by chemically removing WTAs from CW as described under Experimental procedures. We found that the intact GST protein (data not shown) and the GST–tECTD fusion protein (Fig. 3B) bound to neither CW nor PG under the assay conditions. The latter result clearly indicated that the C-terminal catalytic domain of LytF does not have the CW-binding activity, supporting the IFM observation of the CTDE−6xFLAG localization (Fig. 1H and I). On the other hand, in the case of the GST−2xLysM fusion protein, small amount of the protein is able to bind to vegetative CW (Fig. 3A, lane P for CW). Moreover, large amount of the protein binds to PG (Fig. 3A, lane P for PG). This suggests that PG prepared by treatment with 10% trichloroacetic acid (TCA) is a better substrate for the binding of the LysM domain than CW. Because the substances removed from CW on the TCA treatment are mainly anionic polymers such as WTAs (Pollack and Neuhaus, 1994), we inferred the PG binding of the LysM domain might be prevented by WTAs.

Figure 3.

In vitro cell wall (CW)-binding assays with the GST−2xLysM protein. In vitro CW-binding assays were examined in 60 μl of PBST buffer [80 mM NaH2PO4, 20 mM Na2HPO4, 100 mM NaCl (pH 7.5) and 1% Tween 20] containing 20 μg of the purified protein, and CW or peptidoglycan (PG) corresponding to 75 μg of PG. After 15 min incubation on ice, the reaction mixture was centrifuged to separate the supernatant, as the non-binding fraction, and the pellet, as the CW- or PG-binding fraction. Lane M, size marker (Bio-Rad, each 1 μg); lane S, non-binding fraction; lane P, binding fraction. The ratios of the GST−2xLysM bands in lanes S and P calculated with a Lane and Spot Analyser (ATTO) are shown under each lane.
A and B. CW- and PG-binding assays with GST−2xLysM (A) and GST–tECTD (B).
C. GST−2xLysM-binding assay to CWs prepared from TagB-depleted and TagO-depleted cells. B. subtilis HY1055 (Pspac-tagB) and MH1023 (Pspac-tagO) were first cultured in LB medium with 0.8 and 0.4 mM IPTG, respectively, at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with IPTG (0.8 mM, TagB+ 0.4 mM, TagO+) or without IPTG (TagB- and TagO-). After incubation for 3 h, cells were harvested. CW amount in the binding assay was normalized in PG amount (75 μg in 60 μl of each reaction mixture) as described in Experimental procedures.
D. GST−2xLysM-binding assay to CW and PG prepared from a tagO null mutant strain. B. subtilis YM1052 (tagO::kan) was cultured in LB medium supplemented with 25 mM MgCl2. Cells were grown to an OD600 of 1.6 and then harvested. CW and PG were prepared as described in Experimental procedures. Each reaction mixture (60 μl) included CW or PG corresponding to 75 μg of PG.

CW-binding assay of the GST−2xLysM protein on CWs prepared from WTA mutants

Wall teichoic acids of B. subtilis consist of major and minor forms, which differ in the repeating unit; the major WTA is a polymer of glycerol phosphate, whereas the minor form is made from N-acetylgalactosamine and glucose phosphate (Foster and Popham, 2002; Lazarevic et al., 2002). Both TagB and TagO are required for the linkage unit biosynthesis of major and minor WTAs (Foster and Popham, 2002; Lazarevic et al., 2002). Indeed, it has been reported that depletion of TagO caused a significant decrease in the CW phosphate content (Soldo et al., 2002; D'Elia et al., 2006). As a binding assay has revealed that the PG-binding ability of the LysM domain appears to be prevented by anionic polymers among CW components, we examined the binding of the GST−2xLysM fusion protein to CWs prepared from TagB- and TagO-depleted cells (Fig. 3C). In vitro binding assay to CW prepared from TagB-depleted cells revealed a considerably increased amount of GST−2xLysM binding to CW (Fig. 3C, lane P for TagB-). This appeared to exhibit that the ratio of unmodified PG is increased in the TagB-depleted CW, as the PG amounts of the TagB+ and TagB- CWs are normalized in the assay conditions. In addition, a similar binding assay for CW from TagO-depleted cells indicated that TagO depletion gave rise to an increased amount of GST−2xLysM binding to CW (Fig. 3C, lane P for TagO-). These findings strongly suggested that the LysM motif in the CWBE domain specifically recognizes and binds to PG, and that the binding is inhibited by CW modification with anionic polymers such as WTAs. Moreover, we found that considerable amount of GST−2xLysM bound to CW prepared from a tagO null mutant (Fig. 3D, lane P for ΔtagO CW) as compared with the wild-type CW (Fig. 3A, lane P for CW). Furthermore, a large amount of GST−2xLysM binding was observed in the ΔtagO PG prepared from the ΔtagO CW (Fig. 3D, lane P for ΔtagO PG). These results appeared to suggest that CW modification might not completely lack in the mutant, and that the unknown CW modification may be removable by 10% TCA treatment.

Helical localization of LytF on the lateral cell surface in WTA mutants affecting the linkage unit biosynthesis

To confirm the results of the CW-binding assay in vitro, we observed the LytF localization pattern in a tagB-conditional mutant. As the tagB gene is essential and the product is involved in the linkage unit biosynthesis of major and minor WTAs (Soldo et al., 2002; D'Elia et al., 2006), here we used an IPTG-inducible conditional mutant, HY1058. When IPTG was reduced to 0.1 mM, the LytF−6xFLAG was seen not only at cell separation sites and poles but also on the lateral cell surface (83.7% of 92 cells) (Fig. 4C and D). Interestingly, the latter signals formed a helical pattern in most cells (Fig. 4D and G, and Movie S1). Judging from the results of the in vitro CW-binding assay, we thought that the helical LytF cables along the sidewalls might correspond to the regions of naked PG not modified by WTAs. Moreover, TagB-depleted cells showed an aberrant morphology, and LytF was localized on almost the whole cell surface (100% of 50 cells) (without IPTG, Fig. 4E and F). In addition to the results for the tagB-conditional mutant, a similar helix pattern and whole cell surface localization were seen in TagO-reduced (81.7% of 82 cells) (0.08 mM IPTG, Fig. 4J and K) and TagO-depleted (100% of 58 cells) (without IPTG, Fig. 4L and M) cells respectively. TagO is also required for the first step of the linkage unit biosynthesis of major and minor WTAs (Soldo et al., 2002), but D'Elia et al. (2006) reported that the tagO gene was dispensable, and that a tagO null mutant showed slow-growing phenotype and aberrant cell morphology. Thus we examined the LytF localization in a tagO null mutant (Fig. 4N and O). The result indicated that LytF is localizable on the whole cell surface as well as in the case of TagO-depleted cells. These observations strongly suggest that WTAs inhibit the LytF localization on the lateral cell surface.

Figure 4.

Figure 4.

Localization of LytF−6xFLAG in several WTA mutants. Localization patterns of LytF−6xFLAG in tagB (A–G), tagO (H–O), tagF (P–U) and ggaAB (V and W) mutants were observed as follows. For culturing conditional mutants of tagB (HY1058; A–G), tagO (MH1031; H–M) and tagF (HY1059; P–U), the strains were first cultured in LB medium with the addition of IPTG (0.8 mM for HY1058, and 0.4 mM for MH1031 and HY1059) at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with or without IPTG. After incubation for 3 h, cells were harvested and fixed. Three sections at different levels in the z-axis after deconvolution were taken in a typical TagB-reduced cell (G).
N and O. Phase contrast (N) and LytF−6xFLAG localization (O) images of a tagO null mutant strain (HY1060). After the strain was cultured in LB medium with 25 mM MgCl2 at 37°C to an OD600 of 1.8, cells were harvested and fixed.
V and W. Phase contrast (V) and LytF−6xFLAG localization (W) images of a ggaAB double null mutant strain (MH1036). Cells were cultured in LB medium at 37°C to an OD600 of 1.8. The exposure times were 0.1 s for phase-contrast images and 0.1 s (gain 2) for Cy3 images.
Scale bars, 10 μm.

Figure 4.

Figure 4.

Localization of LytF−6xFLAG in several WTA mutants. Localization patterns of LytF−6xFLAG in tagB (A–G), tagO (H–O), tagF (P–U) and ggaAB (V and W) mutants were observed as follows. For culturing conditional mutants of tagB (HY1058; A–G), tagO (MH1031; H–M) and tagF (HY1059; P–U), the strains were first cultured in LB medium with the addition of IPTG (0.8 mM for HY1058, and 0.4 mM for MH1031 and HY1059) at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with or without IPTG. After incubation for 3 h, cells were harvested and fixed. Three sections at different levels in the z-axis after deconvolution were taken in a typical TagB-reduced cell (G).
N and O. Phase contrast (N) and LytF−6xFLAG localization (O) images of a tagO null mutant strain (HY1060). After the strain was cultured in LB medium with 25 mM MgCl2 at 37°C to an OD600 of 1.8, cells were harvested and fixed.
V and W. Phase contrast (V) and LytF−6xFLAG localization (W) images of a ggaAB double null mutant strain (MH1036). Cells were cultured in LB medium at 37°C to an OD600 of 1.8. The exposure times were 0.1 s for phase-contrast images and 0.1 s (gain 2) for Cy3 images.
Scale bars, 10 μm.

LytF localization in WTA mutants affecting the main chain polymerization

In the previous section, we revealed that the linkage unit biosynthesis enzymes affect the sidewall localization of LytF. Next, we examined two main chain polymerization enzymes, TagF (Fig. 4P–U) for major WTA and GgaAB (Fig. 4V and W) for minor one. As the tagF gene is essential and the product is involved in the main chain polymerization of major WTA (Pooley et al., 1992), here we used an IPTG-inducible conditional mutant, HY1059. We observed the helical pattern of LytF−6xFLAG in TagF-reduced cells (61.8% of 110 cells) (Fig. 4R and S) and the whole cell surface localization in TagF-depleted cells (100% of 60 cells) (Fig. 4T and U). Moreover, we performed IFM with a double null mutant (MH1036) of ggaA and ggaB, which are involved in minor WTA synthesis (Freymond et al., 2006). As a result, we observed a weak helical pattern of LytF−6xFLAG along the sidewalls (46% of 50 cells) in the mutant strain (Fig. 4V and W). This helical pattern was very similar to those observed in the TagB-, TagO- or TagF-reduced cells (Fig. 4D, K and S). However, whole cell surface localization of LytF was not observed in the ggaAB mutant (Fig. 4W), as compared with in the TagF-depleted cells (Fig. 4U). Taken together, these results strongly suggest that LytF is localizable in a helical manner on the cylindrical part of major WTA-reduced cells and minor WTA-lacking ones, and that major and minor WTAs are principal hindering components of LytF on the lateral cell surface.

WTA modification along the sidewalls is governed by an actin-like homologue, MreB

It is now clear that LytF forms a helical pattern along the sidewalls on the major WTA-reduced or minor WTA-lacking cell surface. Recent reports revealed that PG synthesis on the lateral cell surface occurs in a helical manner in B. subtilis (Daniel and Errington, 2003; Tiyanont et al., 2006). They supposed that the helical PG synthesis along the sidewalls might be governed by one of the actin-like homologues. Thus we examined whether or not actin-like homologues are involved in the modification of major and minor WTAs. For this purpose, we constructed three strains: one is an mreBH null mutant (MH1042), and the other two conditional mutants of mbl (HY1067) and mreB (HY1071). In the case of the mreB-conditional mutant, an in-frame deletion (ΔmreB) was introduced at the mreB locus as described previously (Formstone and Errington, 2005). To observe the patterns of localization of LytF−6xFLAG in these mutants, we carried out IFM observation (Fig. 5). Figure 5A–D showed that the mreBH null mutation did not affect the LytF localization, suggesting that MreBH is not involved in the WTA modification on the lateral CW. Moreover, Fig. 5E–H indicated that no significant difference of the LytF localization was observed in an mbl-conditional mutant strain with or without 12 mM xylose. Furthermore, we examined the effect of MreB depletion on the LytF localization. Interestingly, Fig. 5K–N clearly showed that LytF−6xFLAG was localizable in a helical manner on the lateral cell surface in the MreB-reduced (80.6% of 72 cells) and MreB-depleted cells (84.7% of 59 cells), this being very like the helical pattern observed in WTA-reduced cells (Fig. 4D, K and S). These results appeared to suggest that MreB depletion might affect the CW modification by WTAs. Moreover, the helical localization was seen in an mreB null mutant grown without Mg2+ (Fig. 5S and T) and in the presence of 2.5 mM MgCl2 (Fig. 5Q and R), but not in the presence of 25 mM MgCl2 (Fig. 5O and P). Formstone and Errington (2005) have reported that an in-frame mreB null mutant restored normal growth and morphology with the addition of 25 mM MgCl2. Our results appeared to indicate that teichoic acid modification along the sidewall is also restored in the mreB null mutant by high Mg2+ supplementation. Taken together, these results strongly suggest that the CW modification along the sidewall might be governed by MreB.

Figure 5.

Figure 5.

Localization of LytF−6xFLAG in mreBH, mbl and mreB mutants.
A–D. Localization of LytF−6xFLAG in the wild type (MH1022; A and B) and an mreBH null mutant (MH1042; C and D) strains. The strains were cultured in LB medium at 37°C to an OD600 of 1.8.
E–N. Localization of LytF−6xFLAG in an mbl-conditional (HY1067; Pxyl-mbl) (E–H) and an mreB-conditional (HY1071; Pxyl-mreB) (I–N) mutant strains. The strains were first cultured in LB medium with xylose (12 mM for HY1067 and 2 mM for HY1071) at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with or without xylose. When cells reached an OD600 of 1.8, cells were harvested and fixed.
O–T. Localization of LytF−6xFLAG in an mreB null mutant strain (HY1075) grown with 25 mM (O and P), 2.5 mM (Q and R) and without (S and T) MgCl2. The strain was first cultured in LB medium supplemented with 25 mM MgCl2 at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with or without MgCl2. After incubation for 3 h, cells were harvested and fixed. The exposure times were 0.1 s for phase-contrast images and 0.1 s (gain 2) for Cy3 images.
Scale bars, 10 μm.

Figure 5.

Figure 5.

Localization of LytF−6xFLAG in mreBH, mbl and mreB mutants.
A–D. Localization of LytF−6xFLAG in the wild type (MH1022; A and B) and an mreBH null mutant (MH1042; C and D) strains. The strains were cultured in LB medium at 37°C to an OD600 of 1.8.
E–N. Localization of LytF−6xFLAG in an mbl-conditional (HY1067; Pxyl-mbl) (E–H) and an mreB-conditional (HY1071; Pxyl-mreB) (I–N) mutant strains. The strains were first cultured in LB medium with xylose (12 mM for HY1067 and 2 mM for HY1071) at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with or without xylose. When cells reached an OD600 of 1.8, cells were harvested and fixed.
O–T. Localization of LytF−6xFLAG in an mreB null mutant strain (HY1075) grown with 25 mM (O and P), 2.5 mM (Q and R) and without (S and T) MgCl2. The strain was first cultured in LB medium supplemented with 25 mM MgCl2 at 37°C to an OD600 of 0.5. Cells were harvested and inoculated at an OD600 of 0.03 into fresh LB medium with or without MgCl2. After incubation for 3 h, cells were harvested and fixed. The exposure times were 0.1 s for phase-contrast images and 0.1 s (gain 2) for Cy3 images.
Scale bars, 10 μm.

Discussion

In this report, we revealed that the N-terminal putative CW-binding domain of the vegetative dl-endopeptidase, LytF, plays an important role in the specific localization to cell separation sites and poles (Figs 1 and 2). In addition, a depletion experiment on an essential protein, PBP 2B, for septum formation demonstrated that septum biosynthesis is required for the LytF localization, suggesting that the N-terminal CW-binding domain of LytF binds to septal PG synthesized through the transpeptidase activity of PBP 2B (Fig. S1). The CW-binding domain of LytF consists of five direct repeats of the LysM motif separated by serine-rich regions. The LysM motif has been reported to be a general PG-binding module (Bateman and Bycroft, 2000; Buist et al., 2008), and is conserved in some cell separation enzymes in bacteria, e.g. MurA of Listeria monocytogenes (Carroll et al., 2003), AcmA of Lactococcus lactis (Steen et al., 2003; 2005) and Sle1 of Staphylococcus aureus (Kajimura et al., 2005). Interestingly, it has been reported that a sensor protein having the LysM motif is also required for the recognition of symbiotic bacteria in plants (Madsen et al., 2003; Radutoiu et al., 2003). Thus, the LysM motif must be one of the targeting domains required for the septum localization of cell separation enzymes in bacteria. Indeed in B. subtilis, all three vegetative dl-endopeptidases, LytE, LytF and CwlS, which are associated with cell separation, retain the LysM repeats in the N-terminal region (Yamamoto et al., 2003; Fukushima et al., 2006). In addition, we revealed that a GST−2xLysM fusion protein binds to PG prepared from vegetative CW in vitro, and that the binding is prevented by anionic polymers such as WTAs (Fig. 3A). As it is thought that the LysM motif specifically binds to PG (Bateman and Bycroft, 2000; Steen et al., 2003; 2005; Buist et al., 2008), our results appear to be quite reasonable. These observations have also been supported by the results of a similar binding assay involving CWs prepared from TagB- and TagO-depleted cells (Fig. 3C). TagB and TagO are required for the linkage unit biosynthesis of major and minor WTAs (Foster and Popham, 2002; Lazarevic et al., 2002; Soldo et al., 2002; D'Elia et al., 2006). Moreover, IFM for LytF−6xFLAG in either TagB-, TagO- or TagF-reduced cells showed that the fusion protein bound to lateral CW in a helical manner in addition to cell separation sites and poles (Fig. 4D, G, K and S, and Movie S1). Supporting our results, it has been reported that AcmA of L. lactis was localized at cell separation sites and that its localization was hindered by CW constituents (Steen et al., 2003; 2005). They have supposed that lipoteichoic acid is a candidate for hindering component. On the other hand, it is well known that WTAs such as teichoic and teichuronic acids are major components for CW modification in B. subtilis (Foster and Popham, 2002; Lazarevic et al., 2002). In addition, as Soldo et al. (1999) reported that the tua operon, which is involved in the teichuronic acid biosynthesis under phosphate-limiting conditions, was not transcribed during vegetative growth in LB medium, we inferred that the WTAs in the B. subtilis CW are the principal candidates for the inhibiting components of the LytF localization. Indeed, as compared with the wild-type CW, an increased amount of the GST−2xLysM protein bound to PG (Fig. 3A), which was chemically prepared from the wild-type CW by treatment with 10% TCA, and CWs prepared from either TagB- or TagO-depleted cells in which both major and minor WTAs would be reduced (Fig. 3C). Moreover, the binding assays involving CW and PG prepared from a tagO null mutant strain appeared to suggest that unknown CW modification might still remain in the mutant cells (Fig. 3D). This unknown CW modification may suppress the lethality of the tagO null mutation.

Among B. subtilis WTAs, the major WTA is a very important component of the B. subtilis CW as its deficiency affects cell morphology (D'Elia et al., 2006), whereas the minor WTA is not (Estrela et al., 1991; Freymond et al., 2006). However, it has been unclear how and where CW modification occurs. Our IFM observation results for some WTA-related mutants provided us with several important clues relating to the mode of CW modification with major and minor WTAs. In TagB- or TagO-reduced cells, LytF is localized not only at cell separation sites and poles but also in a helical manner along the sidewall (Fig. 4D, G and K, and Movie S1). Moreover, whole cell surface localization of LytF was observed in TagB- or TagO-depleted cells in which the linkage unit biosynthesis of major and minor WTAs would be abolished (Fig. 4F and M). Furthermore, LytF cables were readily observed in TagF-reduced cells in which the main chain polymerization of major WTA would be affected (Fig. 4S). In addition, helical LytF localization was observed in a double null mutant of ggaA and ggaB in which the main chain synthesis of minor WTA would be abolished (Fig. 4W), but whole cell surface localization seen in TagB-, TagO- and TagF-depleted cells (Fig. 4F, M and U) was not observed in the ggaAB null mutant ones (Fig. 4W). These results suggest that major WTA is a main hindering component for the LytF localization on the cylindrical part of the rod-shaped cell. Based on the findings of the in vitro CW-binding assay, it was thought that CW modification by WTAs might be poor in the LytF-binding regions. Thus there is a possibility that WTA modification might be reduced at the septum rather than in the sidewall. On the other hand, Formstone et al. (2008) reported that WTA synthesis enzymes localized not only along the sidewall but also to the cell division sites, suggesting that WTA modification might not be reduced at the septum. To answer this discrepancy, the exact nature of septum localization of LytF is currently under study.

In this study, we found that helical LytF localization was observed in several WTAs mutants (Fig. 4D, G, K and S, and Movie S1). These results appeared to suggest, at least, two possibilities. One is that WTA modification and nascent PG incorporation are simultaneously occurred in a helical manner along the sidewall, and the other is that PG synthesis occurs in a helical manner but WTA modification does not. As supporting the former possibility, Formstone et al. (2008) revealed that teichoic acid synthetic enzymes (TagB/F/G/H/O) form a large multienzyme complex and localize in a helical pattern.

Carballido-López et al. (2006) demonstrated that MreBH appears to form a filamentous and helical structure complex with other actin-like homologues, MreB and Mbl, just beneath the cytoplasmic membrane. In addition to this complex formation by three MreBs, MreBH controls the lateral cell surface localization of dl-endopeptidase LytE by means of a physiological interaction, and this interaction is especially required for survival at low Mg2+ concentrations (Carballido-López et al., 2006). The authors inferred that MreBH and LytE might play roles in the helical pattern insertion of both PG-synthesizing and PG-hydrolysing activities. Thus, we examined whether or not the CW modification by WTAs might be governed by helical scaffolds, e.g. an actin-like homologue, Mbl, MreB or MreBH, just beneath the cytoplasmic membrane. Our results suggested that MreBH and Mbl did not affect the LytF localization (Fig. 5D and H), suggesting that these two actin-like homologues might not be involved in the WTA modification. On the other hand, our IFM observations indicated that the helical LytF localization was observed only on the cell surface in both MreB-reduced and MreB-depleted cells (Fig. 5L and N). This helical pattern was very like that observed in the WTAs mutants (Fig. 4D, G, K and S), strongly suggesting that an actin-like filament, MreB, might govern CW modification by major and minor WTAs. However, whole cell surface localization of LytF was not observed in MreB-depleted cells (Fig. 5N) as compared with in TagB-, TagO- or TagF-depleted cells (Fig. 4F, M and U). We presume that regular and helical CW modification by WTAs may be abolished, but the irregular modification continued in MreB-depleted cells because the substrates of WTAs were supplied. On the other hand, they were not supplied in TagB-, TagO- or TagF-depleted cells. Thus, whole cell surface localization of LytF may be observed (Fig. 4F, M and U). Moreover, as helical LytF cable appeared to be seen in the nascent PG region, helical PG synthesis might occur in MreB-depleted cells as well as in the WTAs mutants. Formstone and Errington (2005) revealed that an mreB null mutant restores the normal growth and cell morphology in the presence of high concentrations of Mg2+, and that MreB is not required for cylindrical PG synthesis and chromosome segregation in the presence of SMM (sucrose, maleic acid and MgCl2). Thus we examined whether the helical LytF localization changes in the presence of high concentrations of Mg2+. The result clearly indicated that the helical LytF localization in an mreB null mutant is lost in the presence of the 25 mM MgCl2 (Fig. 5P), suggesting that CW modification was restored in the mreB null mutant under high Mg2+ concentrations. This result appeared to support a very recent finding that an mreB disruption did not affect the helical localization of Tag proteins with the addition of sucrose, maleic acid and Mg2+ (Formstone et al., 2008). Taken together, these results suggest that CW modification by WTAs along the sidewall might be governed by an actin-like cytoskeleton homologue, MreB, in a helical manner. It appears to be quite reasonable if CW modification is carried out in a helical manner as well as PG synthesis and PG hydrolysis. Further experiments are needed to demonstrate whether or not CW modification occurs in a helical manner, and to determine what mechanism and factors lie between a bacterial actin-like homologue, MreB, and CW modification by WTAs.

Experimental procedures

Bacterial strains and plasmids

The strains of B. subtilis and E. coli, and plasmids used in this study are listed in Table 1. B. subtilis WEC, a double mutant strain of wprA and epr without any antibiotic resistance genes, was used as the parent strain throughout this study.

Table 1.  Bacterial strains and plasmids used in this study.
Strain and plasmidRelevant genotypeSourcea or reference
  • a. 

    Sources shown before and after the arrows indicate donor DNA and recipient cells on transformation respectively.

  • BGSC, Bacillus Genetic Stock Center, Ohio State University.

Bacillus subtilis
168trpC2S.D. Ehrlich
MH1018trpC2ΔwprApMADWPRA→168
MH1019trpC2ΔeprΔwprApMADEPR→MH1018
MH1020trpC2 lytF::pCA6FLCE (lytF−6xflag cat)pCA6FLCE→168
YM1046trpC2 lytF::pCA6FLCWBE (cwbE−6xflag cat) (without the C-terminal catalytic domain of LytF)pCA6FLCWBE→168
YM1048trpC2Ω(lytF::ctdE−6xflag cat) (without the N-terminal 5xLysM domain of LytF)Supporting information
MH1022trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat)MH1020→MH1019
YM1047trpC2ΔeprΔwprA lytF::pCA6FLCWBE (cwbE−6xflag cat)YM1046→MH1019
YM1051trpC2ΔeprΔwprAΩ(lytF::ctdE−6xflag cat)YM1048→MH1019
MH1024trpC2Ω(ggaAB::spc)pBGABSp→168
HY1053trpC2 pbpB′::lacZ lacI bla ermC Pspac-pbpBpM4PBPB2→168
HY1055trpC2 tagB′::lacZ lacI bla ermC Pspac-tagBpM4TAGB→168
HY1056trpC2 tagF′::lacZ lacI bla ermC Pspac-tagFpM4TAGF→168
MH1023trpC2 tagO′::lacZ lacI bla ermC Pspac-tagOpM4TAGO→168
YM1052trpC2Ω(tagO::kan)pGtagOKm→168
MH1029trpC2Ω(thrC::Pxyl-mreB spc)pXTMreB→168
HY1064trpC2Ω(thrC::Pxyl-mbl spc)pXTMbl→168
HY1065trpC2Ω(thrC::Pxyl-mbl spc) Ω(mbl::kan)pBmblKm96→HY1064
MH1027trpC2Ω(mreBH::kan)pBmBH2941→168
HY1054trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) pbpB′::lacZ lacI bla ermC Pspac-pbpBHY1053→MH1022
HY1058trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) tagB′::lacZ lacI bla ermC Pspac-tagBHY1055→MH1022
HY1059trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) tagF′::lacZ lacI bla ermC Pspac-tagFHY1056→MH1022
MH1031trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) tagO′::lacZ lacI bla ermC Pspac-tagOMH1023→MH1022
HY1060trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) Ω(tagO::kan)YM1052→MH1022
MH1036trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) Ω(ggaAB::spc)MH1024→MH1022
MH1042trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) Ω(mreBH::kan)MH1027→MH1022
HY1066trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) Ω(thrC::Pxyl-mbl spc)HY1064→MH1022
HY1067trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) Ω(thrC::Pxyl-mbl spc) Ω(mbl::kan)HY1065→HY1066
HY1069trpC2ΔeprΔwprAΩ(thrC::Pxyl-mreB spc)HY1064→MH1019
HY1070trpC2ΔeprΔwprAΩ(thrC::Pxyl-mreB spc) ΔmreBpMADmreB→HY1069
HY1071trpC2ΔeprΔwprAΩ(thrC::Pxyl-mreB spc) ΔmreB lytF::pCA6FLCE (lytF−6xflag cat)MH1020→HY1070
HY1072trpC2Ωkan (kan is inserted between stop codon and terminator downstream of minD)pBminDKm→168
HY1073trpC2ΔmreBΩkanpMADmreB→HY1072
HY1075trpC2ΔeprΔwprA lytF::pCA6FLCE (lytF−6xflag cat) ΔmreBΩkanHY1073→MH1022
Escherichia coli
JM109recA1 supE44 endA1 hsdR17 gyrA96 relA1 thi-1Δ(lac-proAB)/F′[traD36 proAB+lacIqlacZΔM15]TaKaRa
C600supE44 hsdR17 thi-1 thr-1 leuB6 lacY1 tonA21Laboratory stock
BL21FompT hsdSB (rB mB) gal dcm′TaKaRa
Plasmids
pBluescriptII SK+blaTOYOBO
pBGABSpblaΔggaAB::spcThis study
pBmreBHfbblaΔmreBHThis study
pBmBH2961blaΔmreBH::kanThis study
pBmblfbblaΔmblThis study
pBmblKmblaΔmbl::kanThis study
pBminDKmbla kan (kan is inserted between stop codon and terminator downstream of minD)This study
pCA3xFLAGbla cat 3xflagYamamoto et al. (2003)
pCA3FLCEbla cat lytF−3xflagYamamoto et al. (2003)
pCA6xFLAGbla cat 6xflagThis study
pCA6FLCEbla cat lytF−6xflagThis study
pCA6FLCWBEbla cat cwbE (cell wall-binding domain of LytF)−6xflagThis study
pDG1727bla spcBGSC
pDG646bla ermCBGSC
pDG782bla kanBGSC
pDG783bla kanBGSC
pGEX-2Tbla gstGE Healthcare
pGEX−2xLysMbla gst−2xlysMThis study
pGEM-3Zf(+)bla lacZPromega
pGtagOKmblaΔtagO::kanThis study
pQECEtCTDbla 6xhis–ctdE (catalytic domain of LytF)Ohnishi et al. (1999)
pGEXEtCTDbla gst–ctdE (catalytic domain of LytF)This study
pMADbla ermC bgaBArnaud et al. (2004)
pMADWPRAbla ermC bgaBΔwprAThis study
pMADEPRbla ermC bgaBΔeprThis study
pMADmreBbla ermC bgaBΔmreBThis study
pMUTIN4lacZ lacI bla ermCVagner et al. (1998)
pM4PBPB2pMUTIN4::ΔpbpB (containing pbpB Shine–Dalgarno sequence)This study
pM4TAGBpMUTIN4::ΔtagB (containing tagB Shine–Dalgarno sequence)This study
pM4TAGFpMUTIN4::ΔtagF (containing tagF Shine–Dalgarno sequence)This study
pM4TAGOpMUTIN4::ΔtagO (containing tagO Shine–Dalgarno sequence)This study
pXTbla thrC::(Pxyl spc) ermCDerréet al. (2000)
pXTMblbla thrC::(Pxyl-mbl spc) ermCThis study
pXTMreBbla thrC::(Pxyl-mreB spc) ermCThis study

General methods

Bacillus subtilis strains were grown in Luria–Bertani (LB) medium (Sambrook et al., 1989) at 37°C unless otherwise noted. When necessary, chloramphenicol, kanamycin, spectinomycin and erythromycin were added to final concentrations of 5, 5, 100 and 0.3 μg ml−1 respectively. To culture conditional mutant strains of pbpB, tagB, tagF and tagO, IPTG was added to final concentrations of 0.4, 0.8, 0.4 and 0.4 respectively. To pre-culture xylose-inducible mbl and mreB mutants, 12 and 2 mM xylose, respectively, at final concentrations was added to LB medium. E. coli strains were cultured in LB medium at 37°C. If necessary, ampicillin was added to a final concentration of 100 μg ml−1.

DNA manipulations and E. coli transformation were performed by standard methods (Sambrook et al., 1989). B. subtilis transformation was performed by the conventional transformation procedure (Anagnostopoulos and Spizizen, 1961).

Sample preparation for IFM observation

For IFM observation, cells from an overnight culture at 25°C in LB medium were 100-fold diluted in 5 ml of fresh LB medium. Then the cells were grown to the exponential phase at 37°C. A culture exhibiting an optical density at 600 nm (OD600) of 0.25 was centrifuged, and the cells were suspended in 5 ml of fresh LB medium. In the case of conditional mutants of pbpB, tagB, tagF and tagO, the cells were suspended in 5 ml of fresh LB medium with or without IPTG. For a xylose-inducible mutant of mbl (Pxyl-mbl) and mreB (Pxyl-mreB), 12 and 2 mM xylose, respectively, was added to the medium instead of IPTG. These cultures were allowed to grow until OD600 reached ∼1.8. Cells corresponding to 0.5 of an OD600 unit were harvested and fixed. Sample preparation for IFM observation was carried out as described previously (Yamamoto et al., 2003) with a minor modification, as follows. For the detection of anti-FLAG antibody with Cy3, a sheep anti-mouse IgG Cy3 conjugate antibody (Sigma) was used at 1:800 dilution.

Fluorescence microscopy

Fluorescence microscopy was performed as described previously (Yamamoto et al., 2003) with an Olympus BX61 microscope equipped with a BX-UCB control unit, a UPPlan Apo Fluorite phase-contrast objective (magnification, ×100; numerical aperture, 1.3), and standard filter sets for visualizing DAPI, FITC and rhodamine (for Cy3). The exposure times were 0.1 s for phase-contrast microscopy, 0.1 s (gain 2) for Cy3. Cells were photographed with a charge-coupled device camera (CoolSNAP HQ; Nippon Roper) driven by Metamorph software (version 4.6; Universal Imaging). For Cy3 imaging out of focus light was removed using the 2D Deconvolution utility of AutoDeblur software. All images were processed with Adobe Photoshop software. For z-axis imaging, fluorescence microscopy was performed with an AxioImager M1 microscope, a Plan-APOCHROMAT Fluorite differential interference objective (magnification, ×63; numerical aperture, 1.4), and standard filter sets for visualizing rhodamine (for Cy3). The exposure times were 0.1 s for phase-contrast microscopy, 0.1 s (gain 1) for Cy3. Cells were photographed with a charge-coupled device camera (AxioCam MRm; Carl Zeiss) driven by AxioVision software (version 4.6; Carl Zeiss). The 3D Deconvolution utility of AxioVision software was used for z-axis imaging. All images were processed with AxioVision and Adobe Photoshop software.

Preparation of CW, PG and cell surface proteins

Cell wall of the B. subtilis strain was prepared essentially as described previously (Fein and Rogers, 1976; Kuroda and Sekiguchi, 1990). Moreover, for preparation of purified PG, the CW was treated twice in 10% TCA at 37°C for 1 day to remove acid labile components such as WTAs and polysaccharide (DeHart et al., 1995). The amount of PG was calculated by measuring the OD540 value. One OD540 (ml−1) unit is equivalent to 6.45 mg ml−1 PG. For preparation of cell surface proteins, we used an extraction method involving high concentrations of LiCl described previously (Rashid et al., 1995). For concentration of proteins secreted in the culture medium, TCA precipitation (final concentration 2%) was used as described previously (Rashid et al., 1995).

Overexpression and purification of the GST−2xLysM and GST–tECTD fusion proteins

Escherichia coli BL21(pGX−2xLysM or pGXEtCTD) was cultured in 400 ml of LB medium containing 100 μg ml−1 ampicillin until an OD600 of approximately 1.5 at 37°C. Then IPTG was added to the culture to the final concentration of 1 mM, followed by further incubation for 0.5 h. The culture was then centrifuged, and the pellet was suspended in 10 ml of ice-cold PBST buffer [80 mM NaH2PO4, 20 mM Na2HPO4, 100 mM NaCl (pH 7.5) and 1% Tween 20]. After ultrasonication (Sonics and Materials) on ice, the suspension was centrifuged, and the supernatant was filtered through a 0.45-μm-pore-size membrane filter (Nalgene), followed by application to a GSTrap column (1 ml; GE Healthcare). The column was washed with 20 ml of ice-cold PBST buffer, and then the GST−2xLysM or GST–tECTD protein was eluted with 5 ml of elution buffer [50 mM Tris-HCl (pH 8.0), 10 mM reduced glutathione, 1% Tween 20]. The eluate was dialysed twice against 500 ml of PBST buffer at 4°C for more than 3 h.

SDS-PAGE and Western blot analysis

Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) of proteins was performed in 12% (w/v) polyacrylamide gels as described by Laemmli (1970). For sample preparation, an equal volume of 2× SDS-PAGE sample buffer (Laemmli, 1970) was added to a protein solution. After staining the gel, a Lane and Spot Analyser (Atto) was used to calculate the amount of protein in a band according to the manufacturer's instructions. Western blot analysis for the FLAG-fusion proteins was performed as described previously (Yamamoto et al., 2003).

In vitro binding assay to CW or PG

The CW-binding assay with the GST−2xLysM or GST–tECTD protein was examined in 60 μl of PBST buffer containing 20 μg of the purified protein, and CW or PG corresponding to 75 μg of PG. For the CW-binding assay to CWs prepared from TagB- or TagO-depleted cells, the CW amounts were normalized as to the PG amount as follows. A part of the TagB- or TagO-depleted CWs was treated twice in 10% TCA at 37°C for 1 day to remove WTAs, and then the PG amount was calculated by measuring the OD540 value. Finally, the TagB- or TagO-depleted CWs including 75 μg of PG was added to a 60 μl of the reaction mixture. After 15 min incubation on ice, the reaction mixture was centrifuged. Then the supernatant, as the non-binding fraction, was transferred to a new tube and an equal volume of 2× SDS-PAGE sample buffer (Laemmli, 1970) was added. The pellet, as the CW- or PG-binding fraction, was washed once with 60 μl of PBST buffer, and then 120 μl of 1× SDS-PAGE sample buffer was added to the pellet. After boiling for 5 min, samples were applied to an SDS-PAGE gel.

Acknowledgements

We would like to thank Dr T. Msadek and Dr M. Débarbouillé for kindly providing plasmids pXT and pMAD. We thank the members of our group for the helpful discussions and advice. We also thank Mr S. Ooiwa, Ms N. Hariyama and Mr T. Sakamoto for their technical assistance. This work was supported by Grants-in-Aid for Scientific Research (B) (19380047) and by the New Energy and Industrial Technology Department Organization (NEDO) to J.S., for Scientific Research (C) (19580085) to H.Y., and for the Global COE programmes (to J.S. and H.Y.) of the Ministry of Education, Culture, Sports, Science, and Technology of Japan.

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