4-3. Cell wall structure of E. coli and B. subtilis...Cell wall structure 117 approximately 10 times thicker than in the E. coli cell wall, and therefore the B. subtilis cell wall

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Escherichia coli and Bacillus subtilis: The Frontiers of Molecular Microbiology Revisited, 2012: 115-148

ISBN: 978-81-308-0492-7 Editors: Yoshito Sadaie and Kouji Matsumoto

4-3. Cell wall structure of E. coli and B. subtilis

Junichi Sekiguchi* and Hiroki Yamamoto# Faculty of Textile Science and Technology, Shinshu University, 3-15-1 Tokida, Ueda 386-8567, Japan

Abstract. The structure of peptidoglycans of Escherichia coli and Bacillus subtilis is similar except for a few minor modifications, but murein (cell wall) structures are extremely different because the major cell wall constituents, anionic polymers, are not attached to peptidoglycans of E. coli but are attached to those of B. subtilis. Thickness of the cell walls in B. subtilis and the presence of an outer membrane in E. coli are other important differences in the cell wall. In E. coli, murein hydrolases have several functions such as being associated with peptidoglycan biosynthesis, cell separation, modification and recycling/turnover of the cell wall components. In B. subtilis, further various functions include association with cell lysis, cell motility, prophage-induced autolysis, competence, conjugation, lateral peptidoglycan expansion/modification, and sporulation and germination. Enzymatic properties and functions of peptidoglycan hydrolases and cell wall recycling enzymes are reviewed. Moreover, functions and biosynthesis of anionic polymers (teichoic acid, teichuronic acid, and lipoteichoic acid) are fully described.

Introduction Major differences between gram-negative and gram-positive bacteria depend on cell wall structure. Gram-negative bacteria including *Correspondence/Reprint request: Dr. Junichi Sekiguchi, Faculty of Textile Science and Technology, Shinshu University, 3-15-1 Tokida, Ueda 386-8567, Japan. E-mail: [email protected] #E-mail: [email protected]

Junichi Sekiguchi & Hiroki Yamamoto 116

Escherichia coli possess an inner (cytoplasmic) membrane and outer membrane, and the space (called the periplasmic space) between the membranes contains one to two layers of peptidoglycan. In contrast, gram-positive bacteria including Bacillus subtilis do not possess an outer membrane, but contain thick peptidoglycans (10-20 layers) modified by extensive anionic polymers [1]. The structure of peptidoglycans is similar in E. coli and B. subtilis, with a few exceptions [1-4], and is classified as A1γ [5]. In contrast with E. coli, B. subtilis produces spores in which one of the major components, the cortex, has a structure similar to peptidoglycan and plays a role in resistance to various stresses and in germination [1, 4]. The anionic polymers attached to vegetative peptidoglycans in B. subtilis are teichoic acid and teichuronic acid, but the latter is only produced under phosphate-limiting conditions [1, 4]. Lipoteichoic acid is a high glycerol-phosphate-containing material that is anchored into the membrane by one end. Anionic polymers make up 35% to 60% of the entire dry weight of the vegetative cell wall in B. subtilis [1]. In this chapter, cell wall structures including peptidoglycans and anionic polymers as well as properties and functions of peptidoglycan hydrolases are addressed, but peptidoglycan biosynthesis and cell shape associated with high-molecular weight penicillin-binding proteins (PBPs) and actin homologues are not reviewed. 1. Cell wall structure of E. coli and B. subtilis

(1) Peptidoglycan structure of E. coli and B. subtilis during the growth phase In peptidoglycans, the glycan strands are made of disaccharide repeats, N-acetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc), which are linked by β-1,4-glycosidic bonds (Fig. 1) [1-4, 6]. MurNAc is attached to a lactyl group of which the carboxy terminus is linked to a peptide stem consisting of L-alanine (L-Ala; position 1), D-glutamic acid (D-Glu; position 2), meso-diaminopimelic acid (A2pm; position 3), D-alanine (D-Ala; position 4) and D-alanine (position 5 in the case of neither cross-linkage nor peptide processing). Between D-Glu (γ-carboxyl group) and A2pm (amino group at L-configuration), a γ-glutamyl bond is formed (the normal peptide bond consists of a covalent linkage between an α-amino group and α-carboxyl group). The carboxyl group of D-Ala (position 4) in a peptidoglycan strand is cross-linked to the amino group (D-configuration) of the A2pm residue (position 3) in the neighboring strand by a D,D-peptide bond, to form a strong net structure that is resistant to turgor pressure. It is estimated that the turgor pressure is 5 atm (ca. 500 kPa) in E. coli. In B. subtilis, the cell wall is

Cell wall structure 117

approximately 10 times thicker than in the E. coli cell wall, and therefore the B. subtilis cell wall can withstand a turgor pressure of approximately 24 atm (ca. 2,431) [4, 7]. The cross-linkage index of the E. coli peptidoglycan is approximately 50% of the peptide stem, and the glycan chain length varies widely, with 20-30 disaccharide units [8, 9]. In contrast, the cross-linkage index of the B. subtilis peptidoglycan is between 29% and 33%, and the chain length varies between species, with approximately 100-200 disaccharide units [1, 10]. In E. coli, a cross-linkage between two A2pm via an L,D-peptide bond has been found (Fig. 1). The A2pm residues are often modified by covalent bonds with Lys and Arg in lipoprotein [3, 6]. The compound 1,6-anhydromuramic acid is present at the end of the peptidoglycan strands [3, 6, 11].

Figure 1. Peptidoglycan structure of E. coli and B. subtilis and bond specificity of various peptidoglycan hydrolases. D-Glu and m-A2pm-(NH2) are linked by a γ-glutamyl bond. In E. coli, m-A2pm-NH2 and m-A2pm cross-linkage as well as D-Ala-m-A2pm cross-linkages are found. The figure is modified from a review article [20]. GlcNAc, N-acetylglucosamine; MurNAc, N-acetylmuramic acid; L-Ala, L-alanine; D-Glu, D-glutamic acid; m-A2pm, meso-diaminopimelic acid; m-A2pm-NH2, amidated m-A2pm; D-Ala, D-alanine; MurNAc1,6Anh, 1,6-anhydro-N-acetylmuramic acid. Numbers in the figure indicate the following: 1, endo-β-N-acetylglucosaminidase; 2, endo-β-N-acetylmuramidase; 2’, lytic transglycosylase; 3, endo-N-acetylmuramoyl-L-alanine amidase; 4, LD-endopeptidase; 5, DL-endopeptidase; 6, DD-endopeptidase; 7, DD-carboxypeptidase; and 8, LD-carboxypeptidase.


In B. subtilis, several modifications in peptidoglycans have also been observed. Muropeptide analyses (after muramidase hydrolysis) have indicated that at least one free carboxyl group of A2pm is amidated, approximately 17% of muropeptides are not acetylated on the glucosamine moiety (causing a higher resistance to muramidase), anhydromuropeptides are found at the reducing terminus, and glycine is found at position 5 in the stem peptide [1]. Activation of a cell wall modification pathway (O-acetylation of peptidoglycan and D-alanylation of teichoic acid) confers lysozyme resistance [12]. Research on the three-dimensional structure of the gram-negative sacculus has indicated that a single layer of glycans lie parallel to the cell surface, roughly perpendicular to the long axis of the cell, encircling the cell in a disorganized hoop-like fashion [13]. (2) Peptidoglycan structure of B. subtilis during the sporulation phase B. subtilis produces spores (endospore) under starvation conditions (Figs. 2, 3). Spores consist of the core (cytoplasmic space), cytoplasmic membrane, primordial cell wall, cortex, and coats (inner and outer layers) from the center. Therefore, cell wall components are located at the primordial cell wall and cortex. It is believed that the primordial cell wall is similar to the vegetative cell wall, and it contains peptidoglycan. However, it is unknown whether the primordial cell wall contains anionic polymers. The cortex has a thick layer that consists of peptidoglycans with several modifications. Fifty percent of the muramic acid residues are present as a muramic acid δ-lactam and the cross-linkage index of the cortex is only 2.9% (Fig. 2) [1]. Approximately 23% of the stem peptides in the mature cortex are present as single L-Ala, and 26% have a tetrapeptide chain, which may be cross-linked [1]. Anionic polymers have not been reported in the cortex. The muramic acid δ-lactam is synthesized by modification of peptidoglycan [1, 4]. MurNAc peptide is first digested by N-acetylmuramoyl-L-alanine amidase (CwlD), followed by deacetylation of MurNAc (PdaA) and lactam formation by an unknown protein [14-18]. 2. Peptidoglycan hydrolases of E. coli and B. subtilis Many redundant peptidoglycan hydrolases are found in E. coli and B. subtilis, and therefore the physiological properties of peptidoglycan hydrolases are often ambiguous. However, recent research on multiple mutations has led to a better understanding of the functions of peptidoglycan hydrolases.


Figure 2. Spore peptidoglycan (cortex) structure of B. subtilis. δ indicates the delta-lactam structure, which is a unique structure of the cortex. Numbers are the same as those shown in the legend to Fig. 1.

Figure 3. Life cycle and peptidoglycan hydrolases of B. subtilis. Typical peptidoglycan and cortex hydrolases are shown in the figure. Gray arrows indicate the direction from the sigma factors to gene products. The figure is modified from a review article [22].

(1) Peptidoglycan hydrolases in E. coli The structure of peptidoglycan in E. coli is almost identical to that of vegetative peptidoglycan in B. subtilis except for some minor modifications


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[1-4, 6, 9]. Therefore, each linkage of peptidoglycan is assumed to be hydrolyzed by peptidoglycan hydrolases of E. coli as found in B. subtilis (Fig. 1; Tables 1 and 2) [19-22]. However, a limited number of enzyme types have been reported in E. coli (Table 1). In E. coli, PBP2 and RodA are required for the synthesis of glycan strands during elongation, and periplasmic amidases, which aid in cell separation, are minor players, cleaving only one-sixth of the peptidoglycan that is turned over by lytic transglycosylases [23]. (1-1) N-acetylmuramoyl-L-alanine amidases E. coli has two families consisting of Amidase 3 and Amidase 2 domains. The former family contains AmiA, AmiB, and AmiC of which enzymes cleave the septum to separate daughter cells after cell division [25, 26, 54]. The latter family contains the lipoprotein AmiD and the cytoplasmic protein AmpD, and they are associated with peptidoglycan turnover [28, 30, 31, 54]. AmiD hydrolyzes muropeptides with anhMurNAc-L-Ala and MurNAc-L-Ala linkages, and AmpD hydrolyzes the anhMurNAc-L-Ala linkage [28, 30, 31]. The compound anhMurNAc is formed from peptidoglycans by lytic transglycosylase, and the structure terminates the glycan linkage in peptidoglycans [6, 71]. GlcNAc-anhMurNAc-peptides are transported from the periplasmic space to the cytoplasm by AmpG permease and then digested to GlcNAc-anhMurNAc and stem peptides by AmpD [30, 31] for peptidoglycan turnover [6, 54]. E. coli genome encodes four factors with the LytM domain, which shows high sequence similarity with lysostaphin (protein_M23 family) [72]. Two of the factors, EnvC and NlpD, have major functions in cell separation and are located at septa with AmiB and AmiC, but not AmiA [72]. EnvC and NplD stimulate enzymatic activities of AmiA, AmiB, and/or AmiC, and they are not enzymes but spatiotemporal regulators for cell separation [72]. (1-2) Endopeptidases Endopeptidases include LD-endopeptidase which cleaves the L-Ala-D-Glu linkage, DL-endopeptidase which cleaves the D-Glu-meso-A2pm linkage, DD-endopeptidase which cleaves the D-Ala-meso-A2pm cross-linkage (meso-A2pm contains D- and L-configurations at chiral carbons in its molecule), and LD-endopeptidase which cleaves the meso-A2pm-D-Ala linkage (Fig. 1). In E. coli, only DD-endopeptidases have been reported and they consist of four families (peptidase_S13 containing PBP4, peptidase_S11 containing PBP7 and PBP8, peptidase_M74 containing MepA, and AmpH) (Table 1). The genes dacB encoding PBP4 and pbpG encoding PBP7 and PBP8 (proteolytic


degradation product from PBP7) are associated with cell shape and septum cleavage [32-34, 37-44]. MepA is a penicillin-insensitive endopeptidase [45, 46]. PBP4 contains additional DD-carboxypeptidase activity [32, 40]. AmpH, a paralog of β–lactamase AmpC, is a bifunctional enzyme exhibiting DD-endopeptidase and DD-carboxypeptidase activities [47]. Mutations of the ampC and/or ampH genes in E. coli lacking PBPs 1a and 5 produce morphologically aberrant cells [48]. Two types of carboxypeptidases (DD-carboxypeptidase and LD-carboxypeptidase) have been reported in E. coli. DD-carboxypeptidase cleaves the D-Ala-D-Ala linkage from the terminal peptide stem and LD-carboxypeptidase cleaves the meso-A2pm-D-Ala linkage from the terminal peptide after DD-carboxypeptidase digestion (Fig. 1). PBP5, PBP6, and PBP6B encoded by dacA, dacC, and dacD, respectively, belong to the DD-carboxypeptidases, and the former two PBPs are associated with cell shape [34, 36-39, 49]. PBP6B is not essential for cell growth [52]. LdcA is a cytoplasmic LD-carboxypeptidase and plays a role in cell morphology, lysis, and murein recycling [53, 54]. PBP5 is localized in the cylindrical envelope as well as the division site, and recent research has shown that the cellular localization of PBP5 is determined predominantly by sites of active peptidoglycan synthesis, which provide pentapeptide substrate that is accessed most efficiently by the membrane-bound form of PBP5 [49]. β-N-Acetylglucosaminidase and lytic transglycosylase are involved in digestion of the glycan part of peptidoglycan in E. coli. NagZ is a glucosaminidase, which plays a role in cell wall recycling [55, 56]. For recycling of peptidoglycan amino sugars, GlcNAc-anhMurNAc derived from GlcNAc-anhMurNAc-peptides is digested by NagZ to produce GlcNAc and anhMurNAc, and these monosaccharides are further digested and recycled through two pathways to produce UDP-GlcNAc [54]. (1-3) Lytic transglycosylases Lytic transglycosylases have seven members in E. coli and are classified into six families (SLY, MltA, MltB, MltC, MltD, and MltF) (Table 1). SltY (Slt70), MltA, and MltB (Slt35) belong to the SLY, MltA, and MltB families, respectively, and they are associated with cell wall recycling [6, 57-65]. Six members are anchored to the outer membrane (MltA-MltF) but SltY (Slt70) is a soluble periplasmic enzyme, and GlcNAc-anhMurNAc-tetrapeptide is released from the end of the glycan strand by most lytic transglycosidases, except for MltE [54, 68]. MltE is only an endolytic enzyme [68]. Triple mutants lacking Slt70, MltA, and MltB grow normally but show a dramatically reduced rate of murein turnover [58]. A decrease in murein turnover products reduces the induction of β-lactamases [58]. MltC and MltE are in the same


family and are located in the outer membrane (Table 1) [66, 67]. The function of MltD is probably wall recycling [69]. Recently, MltF was reported as an outer membrane-bound periplasmic protein and its C-terminal is a catalytic domain and the N-terminal appears to modulate the lytic behavior of the C-terminal domain [70]. PBP1B (ponA) that is associated with peptidoglycan biosynthesis and expresses bifunctional transglycosylation and transpeptidation reactions is essential for cell growth [6, 73]. Since PBP1B, MipA and MltA form a complex [73], MltA may be a member of murein biosynthetic proteins and digest preexisting (old) peptidoglycans during cell growth [6]. (2) Peptidoglycan hydrolases in B. subtilis B. subtilis produces a complement set of enzymes capable of hydrolyzing the shape-maintaining and stress-bearing peptidoglycan layer of its own cell wall (Fig. 3). Some of these peptidoglycan hydrolases can trigger cell lysis, and therefore can be called autolysins or suicide enzymes [1, 4]. Autolysins have been implicated in several important cellular processes, such as cell wall turnover, cell separation, competence, and flagellation (motility), in addition to cell lysis, and they act as pacemaker and space maker enzymes for wall growth [20-22]. Therefore, fine-tuning of autolysin activity through efficient and strict regulation is essential for bacterial survival. Only the essential two-component system YycFG (WalRK) in B. subtilis regulates autolysins and anionic polymer formation in the cell wall, and double mutations of the autolysins, LytE and CwlO (yvcE), show synthetic lethality [74, 75]. Since these enzymes are located on the side wall, both peptidoglycan synthesis and degradation of old peptidoglycan on the side wall are required for cell growth [1]. One third of the peptidoglycan hydrolases are associated with sporulation and germination (Table 2). These are unique functions in spore-forming bacteria. B. subtilis has several prophages and transposon-like elements, and autolysin genes are found in these regions [21, 22]. Although carboxypeptidases acting on the carboxylic end of the peptide stem of peptidoglycan are not autolysins, they are required for peptidoglycan recycling and turnover [76]. Since the expression phase, localization, and amount of cell wall hydrolases are different, even in similar domain-containing enzymes, these variations reflect many cellular functions in vivo. (2-1) N-Acetylmuramoyl-L-alanine amidase N-Acetylmuramoyl-L-alanine amidases belong to the largest group of cell wall hydrolases, and one of two vegetative major autolysins, LytC (CwlB), belongs to this group (Table 2). The amidase genes are classified into four


families: XlyA, LytC, SpoIIP, and AmiE. Three out of five members of the XlyA family are associated with prophages. Prophage PBSX has two amidases, XlyA and XlyB. The gene xlyA forms a host cell lysis system with the upstream genes, xhlA and xhlB (holin) [81, 82]. The gene blyA encoding amidase in prophage SPβ is also a member of a host cell lysis system [83]. The gene cwlA is the first autolysin cloned in B. subtilis [77, 78], and it is localized in a skin element, which is removed during the late stage of sporulation. CwlH is physiologically the only functional amidase in this family during the cell cycle, and the gene is transcribed by EσE during sporulation [79]. Mother cell lysis at the end of sporulation is carried out by CwlH and the other amidases, CwlC and LytC [79, 80, 88]. The LytC family consists of LytC, CwlC, CwlD, YrvJ and YqiI, and the former three enzymes are reported to be functional during the cell cycle. LytC is the major autolysin [84-86], and the lytC gene is part of a three-gene operon, lytABC [87]. It is transcribed by EσA and EσD, and the latter is the main RNA polymerase for its transcription [87]. The transcription of lytC is regulated by the degSU two-component system and pleiotropic sin mutation through sigma D expression [93, 94]. The σD–dependent autolysins, LytC, LytD, and LytF, may be related to morphological and functional heterogeneity for vegetative B. subtilis populations [140]. LytB is a modifier of LytC, and LytB has high amino acid sequence similarity to SpoIID [85, 90, 91]. The function of LytC is varied, and LytC affects not only cell lysis during the vegetative growth phase, but also cell wall turnover, motility, antibiotic-induced cell lysis, and mother cell lysis [86, 87, 89, 92, 95]. The gene cwlC is transcribed by EσK, and its product is a mother cell-specific lytic enzyme as well as CwlH [79, 96, 97]. CwlD is a protein related to spore cortex synthesis, and its disruption completely lacks muramic-delta-lactam in the cortex [15, 16]. The cortex containing muramic-delta-lactam is a good substrate for germination-specific cortex-lytic enzymes, and therefore the disrupted spores lack the late stage of germination followed by outgrowth [14]. However, in vitro activity of CwlD has not been proven. PdaA is a polysaccharide deacetylase and its disruption lacks muramic-delta-lactam in the cortex [17]. Since PdaA is only active for the substrate –[GlcNAc-MurNAc]n- and deacetylates the acetyl group of MurNAc, it is proposed that CwlD amidase acts before PdaA to synthesize muramic-delta-lactam [17, 18]. During the early sporulation phase, the developing spore (forespore) is wholly engulfed by the adjacent mother cell. A prerequisite for engulfment is the removal of peptidoglycan from the septum that separates the forespore from the mother cell, and the process depends on the autolysins SpoIID and SpoIIP and the membrane-bound protein SpoIIM. SpoIIP localizes to the


septal membrane by interacting with SpoIIM and SpoIID [98, 99]. SpoIIP is a bifunctional enzyme that expresses amidase and DD-endopeptidase activities, and SpoIID is a new family of lytic transglycosylase [99]. (2-2) Endopeptidases Peptidoglycan has a peptide stem containing L-Ala-D-Glu (LD-), D-Glu-meso-A2pm (DL-), and cross-linking D-Ala-meso-A2pm (DD-) linkages (Fig. 1), and these cleaving endopeptidases have been found in B. subtilis. LD-endopeptidase has two families containing CwlK and LytH. The cwlK gene is expressed during the vegetative growth phase [102], but the lytH gene is under the control of mother cell-specific σK, and it is required for the production of single L-Ala side chains in the spore cortex [19]. DL-endopeptidase has two families (family I and family II). Family I consists of only one candidate, YqgT, and family II consists of six enzymes (Table 2). LytF (CwlE) and CwlS are cell-separation enzymes, and these genes are transcribed by EσD and EσH, respectively [103, 104, 109]. LytE (CwlF) has two functions (cell separation and side wall synthesis), and it is transcribed by EσA and EσH [103, 107, 108]. CwlO is an unstable enzyme, and its degradation products are often found in supernatant [110]. However, during the early exponential growth phase, CwlO is localized on the sidewall (unpublished data). The functions of LytF, CwlS and LytE are correlated with localization of proteins on the cell surface, because LytF and CwlS are located at the septum, LytE is at the septum and sidewall [106], and CwlO is at the sidewall (unpublished results). Although LytE and CwlO are synthetic lethal enzymes, which express DL-endopeptidase activity [141], their N-terminal (non-catalytic) domains are different; LytE has three repeats of the cell wall-binding LysM motif, and CwlO has a function-unknown domain [109, 110]. One of the actin homologs, MreBH, is localized as a helical form at the side wall and affects LytE localization [142]. The N-terminal domains of LytF and CwlS also contain five and four LysM repeats, respectively [106, 109]. The cwlT gene is localized in the integrative-conjugative element (ICEBs1) region, and the C-terminal of CwlT exhibits DL-endopeptidase activity [113]. Since the N-terminal region of CwlT exhibits N-acetylmuramidase activity, CwlT is able to digest both glycan and peptide sides and it is a powerful two-domain autolysin [113]. The physiological function of CwlT remains unclear, but it may play a role in partial hydrolysis at a conjugation site. PgdS (YwtD) is involved in γ-polyglutamic acid degradation [111], and γ-polyglutamic acid is a homopolymer of D- or L-glutamic acid produced by B. subtilis var. natto [112]. Since this structure consists of a γ-glutamyl bond, it is possible to digest the D-Glu-meso-A2pm


linkage of peptidoglycan, but PgdS is not able to hydrolyze peptidoglycan (unpublished result). B. subtilis produces a proteinaceous DL-endopeptidase inhibitor IseA, which inhibits peptidoglycan hydrolytic activities of LytF, LytE, CwlS and CwlO [143]. DD-endopeptidases that digest the cross-linked DD-bond in B. subtilis are the C-terminal domain of CwlP (CwlP-C) [114], the bifunctional protein SpoIIP [98, 99] and the sporulation protein SpoIIQ [144, 145]. These amino acid sequences exhibit high amino acid sequence similarity with lysostaphin (Gly-Gly endopeptidase of Staphylococcus aureus). However, CwlP-C has no activity to hydrolyze S. aureus and S. thermophilus cell walls and peptidoglycans [114]. (2-3) N-Acetylglucosaminidases N-Acetylglucosaminidase has four families: LytD (endo-N-acetylglucosaminidase), LytG (exo-N-acetylglucosaminidase), glycohydrolase family 18 (endo-N-acetylglucosaminidase) and NagZ (exo-N-acetylglucosaminidase) (Table 2). LytD (CwlG) is the major vegetative autolysin with LytC (CwlB), and it is under the control of σD [115, 116]. Double mutants deficient in LytC and LytD exhibit greatly impaired motility on a swarm plate, whereas single mutants are motile, and antibiotic-induced cell lysis and cell wall turnover are also impaired by the double mutations [89, 92]. LytG is the exo-glucosaminidase responsible for peptidoglycan structural determination during vegetative growth, and it is involved in cell division, lysis and motility [117]. YdhD, YaaH, YvbX, and YkvQ belong to glycohydrolase family 18 and exhibit high amino acid sequence similarity with the N-acetylglucosaminidase SleL, which is a cortical fragment-lytic enzyme of B. cereus [118]. The ydhD and yaaH genes are transcribed by EσE

and associated with sporulation and germination [119, 120]. NagZ (YbbD) is a β-N-acetylglucosaminidase and is involved in muropeptide rescue and recovery collaborating with N-acetylmuramoyl-L-alanine amidase AmiE (YbbE) [101]. The nagZ and amiE genes make an operon with murQ (ybbI), murR (ybbH) and murP (ybbF) [101]. (2-4) Lytic transglycosylases and endo-N-acetylmuramidases Lytic transglycosylase has three families (GSLE, SLT and SpoIID), and endo-N-acetylmuramidase has only one family, SLT (Table 2). GSLE is a germination-specific lytic enzyme, and the family contains SleB, CwlJ and function-unknown YkvT. Previously, SleB was reported to be a cortex-hydrolyzing amidase [122, 123], but muropeptide analysis suggests that SleB


is a lytic transglycosylase [124]. The sleB-inactivated mutant is unable to complete germination mediated by L-alanine as a germinant [122]. CwlJ is a paralog of SleB, and cwlJ disruption leads to slow germination [125], while cwlJ sleB-deficient spores do not degrade their cortex and are extremely inefficient in giving rise to viable cells. The gene cwlJ is under the control of σE in the mother cell compartment, and sleB is under the control of σG in the forespore compartment [124-127]. The SLT family consists of CwlQ, and the SpoIID family consists of SpoIID (Table 2) [98, 99]. Endo-N-acetylmuramidase in B. subtilis also belongs to the SLT family (Table 2) [113, 114, 128]. CwlP-M (medium region of CwlP), CwlT-N (N-terminal region of CwlT) and CwlQ are orthologs of E. coli soluble lytic transglycosylases, SltY (Slt70), and MltA [6], and also a new type of lysozyme (goose lysozyme) [129]. It is difficult to deduce the enzymatic activity by their sequences, because their amino acid sequences are very similar [113, 114, 128]. CwlP and CwlT are located in the phage SPβ and the conjugative element ICEBs1 regions, respectively. Since they are two-domain autolysins, they are more effective than other autolysins for digesting peptidoglycan [113, 114]. CwlQ is a small protein containing the bifunctional autolysin domain. CwlQ exhibits both endo-N-acetylmuramidase and lytic transglycosylase activities, but its physiological function is unknown [128]. YocH has a typical cell wall binding domain (2 repeats of LysM) in its N-terminal. Recently, it was reported that YocH expresses muranolytic activity, but the bond specificity is unknown [130]. Peptidoglycan hydrolase genes, yocH, cwlO and lytE, are positively controlled by the essential two-component system, YycFG (WalRK) [74, 75]. In contrast, the DL-endopeptidase inhibitor gene iseA (yoeB), peptidoglycan deacetylase gene pdaC (yjeA), and teichoic biosynthetic genes, tagAB and tagDEF, are negatively controlled by YycFG (WalRK) [74, 75]. These results indicate that efficient peptidoglycan hydrolysis is stimulated and anti-peptidoglycan degradation is repressed by YycFG. (2-5) Carboxypeptidases In both gram-positive and gram-negative bacteria, PBPs are responsible for the polymerization and cross-linking of peptidoglycan and PBPs have been divided into three classes based on sequence similarities [1, 137]. Class A high-molecular weight PBPs (PBP-As) are bifunctional enzymes with transglycosylase and transpeptidase activity. Class B high-molecular weight PBPs (PBP-Bs) have a C-terminal transpeptidase domain associated with other function-unknown domain(s). PBP-As are required for peptidoglycan biosynthesis and PBP-Bs are required for cell septation and


maintenance of cell shape. Class C is a group of low-molecular weight PBPs (PBP-Cs), which affect peptidoglycan maturation [1]. Carboxypeptidase digests a stem peptide of peptidoglycan from its carboxy terminal. In B. subtilis, a carboxypeptidase group consists of VanY, PBP-C3, PBP-C1, and PBP-C2 families (Table 2). PBP-C3 consists of PBP5 (dacA), PBP5* (dacB), and PBPI (dacF) [131-135], and PBP-C1 consists of PBP4a (dacC) [136, 137]. PBP-C2 consists of PBP4* (pbpE) and PBPX (pbpX) [134, 138]. Since PBP-As and PBP-Bs are not cell wall hydrolases and the other chapter addresses this content, PBP-Cs are only listed in Table 2. The gene dacA is expressed during the vegetative growth phase, and mutants occupy almost 82% of pentapeptide side chains in total muropeptides [131, 132]. The genes dacB and dacF are transcribed by EσE and EσF, respectively [133-135]. The cortex of dacB dacF double mutants is highly cross-linked, and this modification results in failure to achieve normal spore core hydration and in a decrease in spore heat resistance [134]. The gene dacC is transcribed by EσH, but the function is unknown [136]. The gene pbpE is transcribed during the stationary phase and its expression is greatly increased under high salt stress conditions. Mutants are salt-sensitive and show increased sensitivity to cell envelope active antibiotics and cell wall defects [139]. 3. Teichoic acids of the B. subtilis cell wall

(3-1) Teichoic acids in gram-positive bacteria Teichoic acids (TAs), being anionic polymers, are either covalently coupled to peptidoglycan (WTA) or anchored to membrane lipids (LTA) [1, 146, 147]. According to previous studies, WTA biosynthesis appears to occur on the intracellular side of the cytoplasmic membrane [1, 146, 148] (Fig. 4). In contrast, LTA polymerization is performed on the extracellular surface of the membrane [149-152] (Fig. 5). The gram-positive model organism B. subtilis 168 produces two types of WTA under non-phosphate-limiting conditions. They consist of a common disaccharide linkage unit followed by glycerol-phosphate (GroP) repeats or glucosyl-N-acetylgalactosamine-1-phosphate (GlcGalNAcP) repeats [148]. On the other hand, under phosphate-limiting conditions, teichuronic acids are predominant in the B. subtilis cell wall [153]. In B. subtilis W23 and S. aureus, the main chains of both WTAs are structurally identical and consist of ribitol-phosphate (RboP) repeats [154]. LTA is also an anionic polymer linked to the cytoplasmic membrane via a glycolipid anchor. The main chain consists of a poly(GroP) polymer in most gram-positive bacteria [155].


Figure 4. Biosynthetic pathway of major wall teichoic acid in B. subtilis. Arrows indicate catalytic steps and ovals represent enzymes or transporters. Prenol-P-P indicates a lipid-linked precursor, undecaprenyl-pyrophosphate. GlcNAc, ManNAc, GroP, and Glc represent N-acetylglucosamine, N-acetylmannosamine, glycerol-phosphate, and glucose, respectively. After the major WTA precursor is synthesized on the inner side of the bacterial membrane, an ABC transporter (TagG and TagH) transports the precursor across the membrane. The biological roles of the anionic polymers WTA and LTA have been proposed as follows: cell shape determination [156-158], cell division [149, 159, 160], autolysis [106, 161-164], sporulation [159], immunogenicity and innate immune recognition [165, 166], pathogenicity [167-170], biofilm formation [171, 172], maintenance of cation homeostasis [147, 159, 173], protein secretion [174], and antibiotic resistance [175-178]. (3-2) Cell wall teichoic acids WTAs are anionic polymers, usually glycerol- or ribitol-phosphates, covalently linked to peptidoglycan of gram-positive bacteria. In B. subtilis 168, WTAs, which are 35% to 60% of the vegetative cell wall, are mainly composed of major and minor forms under non-phosphate-limiting conditions [1, 148]. These anionic polymers are linked to an N-acetylmuramyl residue in peptidoglycan via a linkage unit, which consists of


Figure 5. Biosynthetic pathway of lipoteichoic acid in B. subtilis. Arrows indicate catalytic steps and ovals represent enzymes or flippases (?). PtdGro and GroP indicate phosphatidylglycerol and glycerol-phosphate, respectively. After the synthesis of a diglucosyl-diacylglycerol (Glc2-DAG) glycolipid, an unknown flippase(s) (?) translocates the glycolipid anchor to the outer surface of the membrane. Four LtaS orthologs (LtaS, YfnI, YqgS, and YvgJ) add GroPs to the anchor to extend the poly(GroP) LTA polymer. YvgJ is an LTA primase producing GroP-Glc2-DAG, while LtaS, YfnI and YqgS are LTA synthases polymerizing the poly(GroP) main chain. two GroP residues linked to N-acetylmannosaminyl-N-acetylglucosamine phosphate (ManNAc-GlcNAcP). The major WTA has a chain length of approximately 45 to 60 repeated units of GroP. The hydroxyl group on C2 is often substituted by a glucosyl group or a D-alanine residue [1, 179]. The minor WTA, which is a polymer of glucosyl-N-acetylgalactosamine 1-phosphate, comprises 10% to 30% of total wall phosphates [148, 180]. Under phosphate-limiting conditions, B. subtilis 168 produces a teichuronic acid polymer containing a 1-3-linked glucuronic acid and N-acetylgalactosamine [148, 153]. On the other hand, it has been reported that B. subtilis W23 and S. aureus make poly(RboP) WTAs [1, 154, 181]. (3-3) Biosynthesis of wall teichoic acids In B. subtilis 168, the biosynthetic enzymes of the major WTA are encoded by the tag genes (tagABDEFO), gtaB, and mnaA [1, 148]. The


majority of WTA biosynthesis occurs on the inner surface of the cytoplasmic membrane and the pathway is thought to be as follows (Fig. 4). First, TagO couples with GlcNAc to a lipid-linked precursor, undecaprenyl-pyrophosphate, on the membrane surface [158]. The next step catalyzed by TagA is the addition of ManNAc, which is epimerized by MnaA [182]. After formation of the membrane-embedded GlcNAc-ManNAc disaccharide, TagB primase couples with the first GroP ester to the C4 position of the ManNAc sugar to form a linkage unit [182, 183]. TagF polymerase then adds GroPs, extending the poly(GroP) main chain [184, 185]. GroP is supplied as the activated precursor CDP-glycerol, the substrate for TagB and TagF, and is catalyzed by TagD [186]. Glucosylation of GroP subunits in the WTA main chain is performed by glucosyltransferase TagE [148, 187]. UDP-glucose, the substrate of TagE, is derived from glucose-1-phosphate by GtaB [188]. In addition to glucosylation of the major WTA, it is thought that products of the dltABCD operon are responsible for D-alanylation of the major WTA [189], but the modification mechanism has not been well characterized. The minor WTA polymer consisting of glucosyl-N-acetylgalactosamine 1-phosphate (GlcGalNAcP) is synthesized by GgaA and GgaB [148, 180]. The poly-(GlcGalNAcP) polymer is thought to be attached to peptidoglycan via a linkage unit, which is synthesized by TagO, TagA and TagB, as well as the major WTA [148]. The major and minor WTA precursors synthesized in the cytoplasm are translocated across the membrane by an ABC transporter comprising TagG and TagH [157]. Finally, a recent report has strongly suggested that TagT (YwtF), TagU (LytR) and TagV (YvhJ), which are involved in the widespread LytR-Cps2A-Psr (LCP) protein family, are required for attachment of WTAs to peptidoglycan [190]. Under phosphate-limiting conditions, products of the tuaABCDEFGH operon are responsible for biosynthesis of a teichuronic acid polymer consisting of a 1-3-linked glucuronic acid and N-acetylgalactosamine [153]. In the case of S. aureus, the biosynthetic pathway of poly(RboP) WTA has not been well characterized. However, very recently, the pathway has been resolved by a method based on in vitro reconstitution of the intracellular steps [154, 181]. The initial steps catalyzed by TarABDO (TagABDO) in the pathway are highly conserved in both B. subtilis 168 and S. aureus [181, 182]. The next step is catalyzed by a GroP primase TarF to transfer one unit of GroP from CDP-glycerol. TarI and TarJ supply CDP-ribitol, which is a substrate of the bifunctional ribitolphosphotransferase and polymerase TarL [154, 191]. Finally, TarL adds many RboP units to form the main poly(RboP) chain, and the WTA precursor is exported by an ABC transporter consisting of TarG and TarH to the cell surface [181, 192]. Interestingly, the poly(RboP) transporter TarGH expressed in B. subtilis can export a structurally different


substrate, poly(GroP) WTA, suggesting that the substrate specificity of the transporter may not depend on the WTA main chain polymer structure [192]. (3-4) Phenotypic characterization of WTA-deficient mutants It was originally thought that cell wall modification by the major WTA is essential for cell viability [193]. However, a recent report has revealed that tagO, whose product catalyzes the first step in the WTA biosynthesis pathway, is dispensable for cell viability, and that tagO null mutants show slow growth, aberrant morphology and septation, and non-uniform peptidoglycan thickness [194]. Moreover, this report showed that tagB, tagD, and tagF are essential in the presence of tagO but not in its absence. The apparent lethality is due to the accumulation of a toxic intermediate [194, 195]. Therefore, when tagO is deleted and no intermediates accumulate, cells are viable but show slow growth and shape malformations [194]. In addition to the toxic intermediate accumulation, it appears that the lethality with defects in the late steps of the WTA biosynthesis pathway is due to the sequestration of undecaprenol-linked WTA intermediates from the essential process of peptidoglycan synthesis [196]. This suggests that disorder of the WTA biosynthetic pathway affects peptidoglycan biosynthesis via undecaprenol recycling [196, 197]. In contrast, cells lacking the minor WTA show a rod shape and normal growth [106, 180]. In S. aureus, it has also been reported that the first-acting enzyme TagO (TarO) is dispensable for growth, whereas later-acting gene products, TagBD (TarBD) and TarFIJH, are essential for viability, and that this necessity of later-acting genes is suppressed in a tagO null genetic background [195]. In addition to the necessity of WTA for cell growth, pathogenicity critically depends on the expression of WTA in S. aureus [168, 198]. (3-5) Subcellular localization of WTA synthetic enzymes Peptidoglycan assembly occurs in both the cylindrical part and the septum of the B. subtilis cell wall [199-201]. Moreover, recent studies involving fluorescent vancomycin, which recognizes D-Ala-D-Ala residues of newly incorporated peptidoglycan precursor, have revealed that peptidoglycan synthesis at the septum depends on the divisome, whereas that along the sidewall occurs in a helical pattern governed by actin-like homologs, MreB and Mbl [202-205]. If peptidoglycan synthesis and WTA modification occur at the same position, attachment of the newly synthesized WTA precursor to peptidoglycan may also occur in a helical manner. To examine this hypothesis, subcellular localization patterns of the major WTA


synthetic enzymes have been demonstrated. Green fluorescent protein (GFP) fusions of TagB, TagF, TagG, TagH, and TagO have indicated helical localization patterns along the sidewall [206]. These observations suggest that WTA attachment to peptidoglycan may occur in a helix-like pattern on the cylindrical part of the cell. However, the helical localization of the WTA synthetic enzymes is not affected by disruption of any of the three MreB homologues [206]. (3-6) Lipoteichoic acids Lipoteichoic acids (LTAs) are anionic polymers anchoring to bacterial membranes in the cell envelopes of gram-positive bacteria [179]. LTAs of B. subtilis, S. aureus and L. monocytogenes consist of poly(GroP) linked via glycolipid anchors onto bacterial membranes [152, 179]. The glycolipid anchors are slightly different among these bacteria; diglucosyl-diacylglycerol (Glc2-DAG) is present in B. subtilis and S. aureus [207-209], and galactosyl-glucosyl-diacylglycerol (GalGlc-DAG) is present in L. monocytogenes [210, 211]. It is thought that the GroP backbone chain of LTA is substituted with D-alanine esters in B. subtilis and S. aureus [189, 212], and with both D-alanine esters and galactosyl residues in L. monocytogenes [210, 211, 213]. (3-7) Biosynthesis of lipoteichoic acids The initial step of LTA production is initiated by the synthesis of a glycolipid anchor consisting of a disaccharide linked to diacylglycerol (DAG) on the inner surface of the cytoplasmic membrane [150-152] (Fig. 5). The anchor in B. subtilis and S. aureus is composed of Glc2-DAG produced by the glycosyltransferases UgtP and YpfP, respectively [207, 208]. In the case of B. subtilis, two glucose moieties transferred from UDP-glucose are derived from glucose-1-phosphate by the UTP:α-glucose-1-phosphate uridyltransferase GtaB, which is also involved in the glucose modification pathway of WTA [188] (Figs. 4 and 5). The glycolipid anchor of L. monocytogenes is GalGlc-DAG, and thus two distinct glycosyltransferases, LafA and LafB, are required to produce Glc-DAG and GalGlc-DAG, respectively [211]. It appears that the glycolipid anchors are translocated to the outer surface of the membrane by a flippase LtaA in S. aureus [209]. In contrast, since no apparent homologue to LtaA is encoded on the B. subtilis genome, the mechanism of glycolipid anchor transport is still unclear. In contrast to glycolipid anchor formation, main chain polymerization in the LTA biosynthetic pathway has not been well characterized. Very recently, however, Gründling and Schneewind have reported the discovery of the key


enzyme LTA synthase (LtaS) in S. aureus [149]. They showed that S. aureus LtaS synthesizes poly(GroP) LTA and that LTA synthesis is required for bacterial growth and cell division. Interestingly, an ltaS homologue of B. subtilis has been shown to restore LTA synthesis and the growth of LtaS-depleted S. aureus cells [149]. In B. subtilis, four orthologous genes, ltaS (originally yflE), yfnI, yqgS, and yvgJ, of the S. aureus ltaS gene are encoded on the genome [149, 159]. LtaS, YfnI and YqgS are LTA synthases polymerizing the poly(GroP) main chain, while YvgJ is an LTA primase that transfers the initial GroP subunit onto the glycolipid anchor to produce GroP-Glc2-DAG [214] (Fig. 5). Among them, B. subtilis LtaS appears to play a principal role in housekeeping LTA synthesis, since its absence affects cell division during vegetative growth [159]. On the other hand, YfnI is assumed to function especially under conditions of stress and to synthesize a longer LTA polymer in chain length than that synthesized by LtaS [149, 214]. Moreover, YqgS and YvgJ appear to be mainly required for LTA synthesis during the sporulation process [159]. In vitro analyses with the C-terminal catalytic domains of LtaS-type enzymes in S. aureus [215] and B. subtilis [214] have suggested that GroP subunits are derived from the membrane lipid phosphatidylglycerol (PtdGro). In L. monocytogenes, LTA is synthesized by two enzymes, LtaP (Lmo0644) and LtaS (Lmo0927) [211]. LtaP functions as an LTA primase, which produces the GroP-glycolipid intermediate, and LtaS functions as an LTA synthase, which extends the poly(GroP) main chain on this intermediate. (3-8) Phenotypic characterization of LTA-deficient mutants In B. subtilis, it has been reported that an ltaS mutation affects cell division, cell morphogenesis, and divalent cation homeostasis [159]. However, mutations of the other three orthologous genes (yfnI, yqgS and yvgJ) do not affect their cell morphology and cell division [159]. Therefore, products of these three genes appear to play overlapping roles in environmental responses [159, 213]. Transcription of yfnI is under the control of extracytoplasmic sigma factor σM, which is involved in salt stress resistance [216, 217]. In addition, since double mutants of ltaS yqgS and ltaS yvgJ show a lower sporulation frequency compared with that of the wild type, this suggests that YqgS and YvgJ are required for the sporulation process together with LtaS [159]. A quadruple mutant of the ltaS homologue genes abolishes LTA modification and shows slow growth and aberrant filamentous clumps twisted around their long axis [159]. Interestingly, it has also been shown that Mg2+ supplementation and ltaS mutation individually suppress lethality of an mbl mutant [159]. Mbl is one of three actin-like proteins in B.


subtilis and is involved in cell wall synthesis [218]. Therefore, it is thought that LTA is important in scavenging and sequestration of divalent cations including Mg2+, and that loss of the LTA-dependent sequestration of Mg2+ ions reduces the Mg2+-dependency of mbl mutants. S. aureus cells lacking WTA show slight morphological alterations, whereas LTA depletion causes growth arrest and severe morphological defects such as an increase in cell size, partially thickened cell walls, and aberrant cell division [149]. Subsequently, it has been reported that LTA-deficient S. aureus ltaS mutants can grow at 30°C but not at 37°C [160]. In this previous report, the ltaS mutant cells had aberrant cell division and separation, decreased autolysis, and reduced levels of peptidoglycan hydrolases, even at the permissive temperature conditions. In addition, an ltaS tagO double mutant has indicated a synthetic lethal phenotype, suggesting that LTA and WTA compensate for one another in an essential function in S. aureus [160] as well as B. subtilis [159]. Interestingly, growth arrest and aberrant cell morphology of S. aureus LtaS-depleted cells are suppressed by expression of only the B. subtilis ltaS gene, but not of the yfnI, yqgS or yvgJ genes [149]. However, very recently, it has been demonstrated that S. aureus LtaS-depletion is also functionally complemented when the B. subtilis yqgS gene is expressed from a multicopy plasmid [214]. In L. monocytogenes, an ltaS (lmo0927) deletion strain is viable at 30°C but not at 37°C [211]. When the strain is grown even at a permissive temperature, cells exhibit a reduced growth rate and short chain-forming morphology. Growth is ceased and the chain length is increased at 37°C. (3-9) Subcellular localization of LTA and synthetic enzymes Cryo-electron microscopy of frozen-hydrated sections has shown that gram-positive bacteria such as B. subtilis and S. aureus have a periplasmic space between the plasma membrane and a thick cell wall [219, 220], and that LTA is a major component of the B. subtilis periplasm [221]. Moreover, subcellular localization analysis of the glycolipid anchor synthetic enzyme UgtP fused by GFP has shown that the fusion protein is predominantly localized at septa, and that proper septal localization of UgtP is abolished in pgcA or gtaB mutants [222]. Importantly, UgtP, which is a metabolic sensor, localizes to cell division sites in a nutrient-dependent manner and inhibits assembly of the cell division protein FtsZ to control cell size [222]. In addition, two LTA synthases, LtaS and YqgS, fused by GFP, are mainly localized at division sites in both vegetative and sporulating cells [159]. These observations strongly suggest that both glycolipid anchor and LTA


syntheses predominantly occur at the cell division site. Supporting this idea, both ugtP and ltaS mutations affect cell division [159, 222]. Acknowledgments This work was supported by Grants-in-Aid for Scientific Research (B) (19380047) and (A) (22248008) to J.S., and (C) (19580085 and 23580107) to H.Y., and by a New Energy and Industrial Technology Department Organization (NEDO) grant to J.S. It was also supported by Global COE programs (to J. S.) of the Ministry of Education, Culture, Sports, Science and Technology of Japan. H. Y. was supported by grants from the Kurata Memorial Hitachi Science and Technology Foundation, the Nagase Science and Technology Foundation, and the Research Foundation for the Electrotechnology of Chubu. References 1. Foster, S. J. and Popham, D. L. 2002, Bacillus subtilis and Its Closest Relatives:

from Genes to Cells, Sonenshein A .L., Hoch, J. A., and Losick, R., (Ed.) Washington, DC: American Society for Microbiology, Washington, D. C., 21.

2. Vollmer, W. and Bertsche, U. 2008, Biochim. Biophys. Acta, 1778, 1714. 3. Vollmer, W., Blanot, D., and de Pedro, M. A. 2008, FEMS Microbiol. Rev.,

32, 149. 4. Archibald, A. R., Hancock, L. C., and Harwood, C. R. 1993, Bacillus subtilis and

Other Gram-Positive Bacteria, Sonenshein, A. L., Hoch, J. A., and Losick, R. (Ed.), Washington, DC, American Society for Microbiology Press, 381.

5. Schleifer, K. H. and Kandler, O. 1972, Bacteriological Rev., 36, 407. 6. Höltje, J. V. 1998, Microbiol. Mol. Biol. Rev., 62, 181. 7. Mendelson, N. H. and Thwaites, J. J. 1989, J. Bacteriol., 171, 1055. 8. Harz, H., Burgdorf, K., and Höltje, J.-V. 1990, Anal. Biochem., 190, 120. 9. Vollmer, W. and Seligman, S. J. 2010, Trends Microbiol., 18, 59. 10. Ward, J. 1973, Biochem. J., 133, 395. 11. Braun, V. and Wolff, H. 1970, Eur. J. Biochem., 14, 387. 12. Guariglia-Oropenza, V. and Helmann, J. D. 2011, J. Bacteriol., 193, 6223. 13. Gan, L., Chen, S., and Jensen, G. J. 2008, Proc. Natl. Acad. Sci. U.S.A., 105,

18953. 14. Sekiguchi, J., Akeo, K., Yamamoto, H., Khasanov, F. K., Alonso, J. C. and

Kuroda, A. 1995, J. Bacteriol., 177, 5582. 15. Popham, D. L., Helin, J., Costello, C. E., Setlow, P. 1996, Proc Natl Acad Sci

USA. 93, 15405. 16. Atrih, A., Zollner, P., Allmaier, G., and Foster, S. J. 1996, J. Bacteriol., 178, 6173. 17. Fukushima, T., Yamamoto, H., Atrih, A., Foster, S. J., and Sekiguchi, J. 2002, J.

Bacteriol., 184, 6007.


18. Fukushima, T., Kitajima, T., and Sekiguchi, J. 2005, J. Bacteriol.. 187, 1287. 19. Horsburgh GJ, Atrih A, and Foster SJ. 2003, J Bacteriol., 185, 3813. 20. Vollmer, W., Joris, B., Charlier, P., and Foster, S. 2008, FEMS Microbiol. Rev.,

32, 259. 21. Smith, T. J., Blackman, S. A., and Foster, S. J. 2000, Microbiology, 146, 249. 22. Shida, T. and Sekiguchi, J. 2005, Survival and Death in Bacteria, Yamada, M.

(Ed.), Kerala, Research Signpost, 117. 23. Uehara, T. and Park, J. T. 2008, J. Bacteriol., 190, 3914. 24. Tomioka, S., Nikaido, T., Miyakawa, T., and Matsuhashi, M. 1983, J Bacteriol.,

156, 463. 25. Heidrich, C., Templin, M. F., Ursinus, A., Merdanovic, M., Berger, J., Schwarz,

H., de Pedro, M. A., and Höltje, J. V. 2001, Mol. Microbiol., 41, 167. 26. Priyadarshini, R., de Pedro, M. A., and Young, K. D. 2007, J. Bacteriol., 189,

5334. Role of peptidoglycan amidases in the development and morphology of the division septum in Escherichia coli.

27. Tsui, H. C., Zhao, G., Feng, G., Leung, H. -C. E., and Winkler, M. E. 1994, Mol Microbiol., 11, 189.

28. Uehara, T. and Park, J. T. 2007, J. Bacteriol. 189, 5634. 29. Kerff, F., Petrella, S., Mercier, F., Sauvage, E., Herman, R., Pennartz, A.,

Zervosen, A., Luxen, A., Frère, J.-M., Joris, B., and Charlier, P. 2010, J. Mol. Biol., 397, 249.

30. Höltje, J. V., Kopp, U., Ursinus, A., and Wiedemann. B. 1994, FEMS Microbiol. Lett., 122, 159.

31. Jacobs, C., Joris, B., Jamin, M., Klarsov, K., Van Beeumen, J., Mengin-Lecreulx, D., van Heijenoort, J., Park, J. T., Normark, S., and Frère, J. M. 1995, Mol. Microbiol., 15, 553.

32. Korat, B., Mottl, H., Keck, W. 1991, Mol Microbiol., 5, 675. 33. Mottl, H., Keck, W. 1991, Eur J Biochem., 200, 767. 34. Heidrich, C., Ursinus, A., Berger, J., Schwarz, H., and Höltje, J. V. 2002, J.

Bacteriol., 184, 6093. 35. Mottl, H., Terpstra, P., and Keck, W. 1991, FEMS Microbiol. Lett., 62, 213. 36. Popham, D. L. and Young, K. D. 2003, Curr. Opin. Microbiol., 6, 594. 37. Priyadarshini, R., Popham, D. L., and Young, K. D. 2006, J. Bacteriol., 188, 5345. 38. Varma, A. and Young, K. D. 2004, J. Bacteriol., 186, 6768. 39. Young, K. D. 2003, Mol. Microbiol., 49, 571. 40. Clarke, T. B., Kawai, F., Park, S.-Y., Tame, J. R. H., Dowson, C. G., and Roper,

D. I. 2009, Biochemistry, 48, 2675. 41. Meberg, B. M., Paulson, A. L., Priyadarshini, R., and Young, K. D. 2004, J.

Bacteriol., 186, 8326. 42. Tuomanen, E. and Schwartz, J. 1987, J. Bacteriol., 169, 4912. 43. Romeis, T., Höltje, J. -V. 1994, Eur J Biochem., 224, 597. 44. Henderson, T. A., Templin, M., and Young, K. D. 1995, J Bacteriol., 177, 2074. 45. Keck, W., van Leeuwen, A. M., Huber, M., and Goodell, E. W. 1990, Mol.

Microbiol., 4, 209. 46. Firczuk, M. and Bochtler, M. 2007, Biochemistry, 46, 120.


47. Gonzalez-Leiza, S. M., de Pedro, M. A., and Ayala, J. A. 2011, J. Bacteriol., print ahead JB.05764-11.

48. Henderson, T. A., Young, K. D., Denome, S. A., and Elf, P. K. 1997, J. Bacteriol., 179, 6112.

49. Potluri, L., Karczmarek, A., Verheul, J., Piette, A., Wilkin, J.-M., Werth, N., Banzhaf, M., Vollmer, W., Young, K. D., Nguyen-Distèche, M., and den Blaauwen, T. 2010, Mol. Microbiol., 77, 300.

50. Chen, Y., Zhang, W., Shi, Q., Hesek, D., Lee, M., Mobashery, S., and Shoichet, B. K. 2009, J. Am. Chem. Soc., 131, 14345.

51. Baquero, M.-R., Bouzon, M., Quintela, J. C., Ayala, J. A., and Moreno, F. 1996, J. Bacteriol., 178, 7106.

52. Ghosh, A. S., Chowdhury, C., and Nelson, D. E. 2008, Trends Microbiol., 16, 309. 53. Templin, M. F., Ursinus, A., and Höltje, J. V. 1999, EMBO J., 18, 4108. 54. Park, J. T. and Uehara, T. 2008, Microbiol. Mol. Biol. Rev., 72, 211. 55. Cheng, Q., Li, H., Merdek, K., and Park, J. T. 2000, J. Bacteriol., 182, 4836. 56. Vötsch, W. and Templin, M. F. 2000, J. Biol. Chem., 275, 39032. 57. Betzner, A. and Keck, W. 1989, Mol. Gen. Genet., 219, 489. 58. Kraft, A. R., Prabhu, J., Ursinus, A., and Höltje, J. V. 1999, J. Bacteriol., 181, 7192. 59. Ursinus, A., Höltje, J. -V. 1994, J. Bacteriol., 176, 338. 60. Lommatzsch, J., Templin, M., Kraft, A. R., Vollmer, W., Höltje, J.-V. 1997, J.

Bacteriol., 179, 5465. 61. van Straaten, K. E., Barends, T. R. M., Dijkstra, B. W., and Thunnissen, A. W. H.

2007, J. Biol. Chem., 281, 21197. 62. Dijkstra, A. J., Hermann, F., and Keck, W. 1995, FEBS Lett., 366, 115. 63. Engel, H., Smink, A. J., and van Wijngaarden, L., and Keck, W. 1992, J.

Bacteriol., 174, 6394. 64. Ehlert, K., Höltje, J. -V., and Templin, M. F. 1995, Mol Microbiol., 16, 761. 65. Suvorov, M., Lee, M., Hesek, D., Boggess, B., and Mobashery, S. 2008, J. Am.

Chem. Soc., 130, 11878. 66. Dijkstra, A. J. and Keck, W. 1996, Microb Drug Resist., 2, 141–145. 67. Kraft, A. R., Templin, M. F., and Höltje, J. -V. 1998, J. Bacteriol., 180, 3441. 68. Artola-Recolons, C., Carrasco-López, C., Llarrull, L. I., Kumarasiri, M.,

Lastochkin, E., de Ilarduya, I. M., Meindl, K., Usón, I., Mobashery, S., and Hermoso, J. A. 2011, Biochemistry, 50, 2384.

69. Bateman, A. and Bycroft, M. 2000, J. Mol. Biol., 299, 1113. 70. Scheurwater, E. M. and Clarke, A. J. 2008, J. Biol. Chem., 283, 8363. 71. Höltje, J.-V., Mirelman, D., Sharon, N. and Schwarz, U. 1975, J. Bacteriol.,

124, 1067. 72. Uehara, T., Parzych, K. R., Dinh, T., and Bernhardt, T. G. 2010, EMBO J., 29, 1412. 73. Vollmer, W., von Rechenberg, M., and Höltje, J. –V. 1999, J. Biol. Chem.,

274, 6726. 74. Bisicchia, P., Noone, D., Lioliou, E., Howell, A., Quigley, S., Jensen, T., Jarmer,

H., and Devine, K. M. 2007, Mol. Microbiol., 65, 180. 75. Bisicchia, P., Lioliou, E., Noone, D., Salzberg, L. I., Botella, E., Hübner, S.,

Devine, K. M. 2010, Mol. Microbiol., 75, 972.


76. Reith, J. and Mayer, C. 2011, Appl. Microbiol. Biotechnol., 92, 1. 77. Kuroda, A. and Sekiguchi, J. 1990, J. Gen. Microbiol., 136, 2209. 78. Kuroda, A., Imazeki, M., and Sekiguchi, J. 1991, FEMS Microbiol. Lett., 81, 9-14. 79. Nugroho, F. A., Yamamoto, H., Kobayashi, Y., and Sekiguchi, J. 1999, J.

Bacteriol., 181(20):6230-6237. 80. Lewis, K. 2000, Microbiol. Mol. Bio. Rev., 64, 503. 81. Longchamp, P. F., Mauël, C., and Karamata, D. 1994, Microbiology, 140, 1855. 82. Krogh, S., Jorgensen, S. T., and Devine, K. M. 1998, J. Bacteriol., 180, 2110. 83. Regamey, A. and Karamata, D. 1998, Microbiology, 144, 885. 84. Herbold, D. R. and Glaser, L. 1975, J. Biol.Chem., 250, 1676. 85. Herbold, D. R. and Glaser, L. 1975, J. Biol.Chem., 250, 7231. 86. Kuroda, A. and Sekiguchi, J. 1991, J. Bacteriol., 173, 7304. 87. Lazarevic, V., Margot, P., Soldo, B., and Karamata, D. 1992, J. Gen. Microbiol.,

138, 1949. 88. Smith, T. J. and Foster, S. J. 1995, J. Bacteriol., 177, 3855. 89. Blackman, S. A., Smith, T. J., and Foster, S. J. 1998, Microbiology, 144, 73. 90. Kuroda, A., Rashid, M. H., and Sekiguchi, J. 1992, J. Gen. Microbiol.,

138, 1067. 91. Kuroda, A. and Sekiguchi, J. 1992, FEMS Microbiol. Lett., 95, 109. 92. Rashid, M. H., Kuroda, A., and Sekiguchi, J. 1993, FEMS Microbiol. Lett., 112, 135. 93. Kuroda, A. and Sekiguchi, J. 1993, J. Bacteriol., 175, 795. 94. Tokunaga, T., Rashid, M. H., Kuroda A., and Sekiguchi J. 1994, J. Bacteriol.,

176, 5177. 95. Rashid, M. H., Tamakoshi, A., and Sekiguchi, J. 1996, J. Bacteriol., 178, 4861. 96. Kuroda, A., Asami, Y., and Sekiguchi, J. 1993, J. Bacteriol., 175, 6260. 97. Shida, T., Hattori, H., Ise F., and Sekiguchi J. 2000, Biosci. Biotech. Biochem.,

64, 1522. 98. Chastanet, A., Losick, R. 2007, Mol. Microbiol., 64, 139. 99. Morlot, C., Uehara, T, Marquis, K. A., Bernhardt, T. G., and Rudner, D. Z. 2010,

Genes Dev., 24, 411. 100. Gutierrez, J., Smith, R., and Pogliano, K. 2010, J. Bacteriol., 192, 3174. 101. Litzinger S, Duckworth A, Nitzsche K, Risinger C, Wittmann V, and Mayer C.

2010, J Bacteriol., 192, 3132. 102. Fukushima, T., Yao, Y., Kitajima, T., Yamamoto, H., and Sekiguchi, J. 2007,

Mol. Genet. Genomics, 278, 371. 103. Ohnishi, R., Ishikawa, S., and Sekiguchi, J. 1999, J. Bacteriol.. 181, 3178. 104. Margot, P., Pagni, M., and Karamata, D. 1999, Microbiology, 145, 57. 105. Yamamoto, H., Kurosawa, S. and Sekiguchi, J. 2003, J. Bacteriol., 185, 6666. 106. Yamamoto, H., Miyake, Y., Hisaoka, M., Kurosawa, S., and Sekiguchi, J. 2008,

Mol. Microbiol., 70, 297. 107. Margot, P., Wahlen, M., Gholamhoseinian, A., Piggot, P,. and Karamata, D.

1998, J Bacteriol., 180, 749. 108. Ishikawa, S., Y. Hara, R. Ohnishi and J. Sekiguchi. 1998, J. Bacteriol., 180, 2549. 109. Fukushima, T., Afkham, A., Kurosawa, S., Tanabe, T., Yamamoto, H., and

Sekiguchi, J. 2006, J. Bacteriol., 188, 5541.


110. Yamaguchi, H., Furuhata, K., Fukushima, T., Yamamoto, H. and Sekiguchi, J. 2004, J. Biosci. Bioeng., 98, 174.

111. Suzuki, T., and Tahara, Y. 2003, J. Bacteriol., 185, 2379. 112. Kambourova, M., Tangney, M., and Priest, F. G. 2001, Appl. Environ.

Microbiol., 67, 1004. 113. Fukushima, T., Kitajima, T., Yamaguchi, H., Ouyang, Q., Furuhata, K.,

Yamamoto, H., Shida, T., and Sekiguchi, J. 2008, J. Biol. Chem., 283, 11117. 114. Sudiarta, I P., Fukushima, T., and Sekiguchi, J. 2010, J. Biol. Chem., 285, 41232. 115. Margot, P., Mauel, C., and Karamata, D. 1994, Mol. Microbiol., 12, 535. 116. Rashid, M. H., Mori, M., and Sekiguchi, J. 1995, Microbiology, 141, 2391. 117. Horsburgh, G. J., Atrih, A., Williamson, M. P., and Foster, S. J. 2003,

Biochemistry, 42, 257. 118. Chen, Y., Fukuoka, S., and Makino, S. 2000, J. Bacteriol., 182, 1499. 119. Kodama, T., Takamatsu, H., Asai, K., Ogasawara, N., Sadaie, Y., Watabe, K.

2000, J. Biochem., 128, 655. 120. Kodama, T., Takamatsu, H., Asai, K., Kobayashi, K., Ogasawara, N., Watabe, K.

1999, J. Bacteriol., 181, 4584. 121. Chirakkal, H., O'Rourke, M., Atrih, A., Foster, S. J., Moir, A. 2002,

Microbiology, 148, 2383. 122. Moriyama, R., Hattori, A., Miyata, S., Kudoh, S., and Makino S. 1996, J.

Bacteriol., 178, 6059. 123. Moriyama, R., Fukuoka, H., Miyata, S., Kudoh, S., Hattori, A., Kozuka, S.,

Yasuda, Y., Tochikubo, K., Makino, S. 1999, J. Bacteriol., 181, 2373. 124. Boland, F. M., Atrih, A., Chirakkal, H., Foster, S. J., and Moir, A. 2000,

Microbiology, 146, 57. 125. Ishikawa, S., Yamane, K., and Sekiguchi, J. 1998, J. Bacteriol., 180, 1375-1380. 126. Bagyan, I. and Setlow, P. 2002, J. Bacteriol., 184, 1219. 127. Paidhungat, M., Ragkousi, K., Setlow, P. 2001. J. Bacteriol., 183, 4886. 128. Sudiarta, I. P., Fukushima, T., and Sekiguchi, J. 2010, Biochem. Biophy. Res.

Commun., 398, 606. 129. Pagliero, E., Dideberg, O., Vernet, T., and Guilmi, A. M. D. 2005, BMC

Genomics, 6, 19. 130. Shah, I. M. and Dworkin, J. 2010, Mol. Microbiol., 75, 1232. 131. Todd, J. A., Roberts, A. N., Johnstone, K., Piggot, P. J., Winter, G., and Ellar, D.

J. 1986, J. Bacteriol., 167, 257. 132. Atrih, A., Bacher, G., Allmaier, G., Williamson, M. P., and Foster, S. J. 1999, J.

Bacteriol., 181, 3956. 133. Buchanan, C. E. and Ling, M. –L. 1992, J. Bacteriol., 174, 1717. 134. Popham, D. L., Gilmore, M. E., and Setlow, P. 1999, J. Bacteriol., 181, 126. 135. Wu, J. –J., Schuch, R., and Piggot, P. J., 1992, J. Bacteriol., 174, 4885. 136. Pedersen, L. B., Murray, T., Popham, D. L., and Setlow, P. 1998, J. Bacteriol.,

180, 4967. 137. Sauvage, E., Duez,, C., Herman, R., Kerff, F., Petrella, S., Anderson, J. W.,

Adediran, S. A., Pratt, R. F., Frère, L. -M., and Charlier, P. 2007, J. Mol. Biol., 371, 528.


138. Popham, D. L. and Setlow, P. 1993, J. Bacteriol., 175, 2917. 139. Palomino, M. M., Sanchez-Rivas, C., and Ruzal, S. M. 2009, Res. Microbiol.,

160, 117. 140. Chen, R., Guttenplan, S. B., Blair, K. M., and Kearns, D. B. 2009, J. Bacteriol.,

191, 5775. 141. Bisicchia, P., Noone, D., Lioliou, E., Howell, A., Quigley, S., Jensen, T., Jarmer,

H., and Devine, K. M. 2007, Mol. Microbiol., 65, 180. 142. Carballido-Lopez, R., Formstone, A., Li, Y., Ehrich, S. D., Noirot, P., and

Errington, J., 2006, Developmental Cell, 11, 399. 143. Yamamoto, H., Hashimoto, M., Higashitsuji, Y., Harada, H., Hariyama, N.,

Takahashi, L., Iwashita, T., Ooiwa, S., and Sekiguchi, J. 2008, Mol. Microbiol., 70, 168.

144. Londoño-Vallejo, J. A., Fréhel, C., Stragier, P. 1997, Mol Microbiol. 24, 29. 145. Sun, Y. L., Sharp, M. D., Pogliano, K. 2000, J. Bacteriol., 182, 2919. 146. Bhavsar, A.P., and Brown, E.D. 2006, Mol. Microbiol., 60, 1077. 147. Archibald, A.R., Armstrong, J.J., Baddiley, J., and Hay, J.B. 1961, Nature, 191, 570. 148. Lazarevic, V., Pooley, H.M., Mauël, C., and Karamata, D. 2002, Biopolymers,

Vol. 5, Polysaccharides I: Polysaccharides from Prokaryotes, Vandamme, E.J., DeBaets, S., and Steinbüchel, A. (Ed.), Weinheim: Wiley- VCH, 465.

149. Gründling, A., and Schneewind, O. 2007, Proc. Natl. Acad. Sci. U S A, 104, 8478. 150. Rahman, O., Dover, L.G., and Sutcliffe, I.C. 2009, Trends. Microbiol., 17, 219. 151. Sutcliffe, I.C. 2011, Mol. Microbiol., 79, 553. 152. Reichmann, N.T., and Gründling, A. 2011, FEBS Microbiol. Lett., 319, 97. 153. Soldo, B., Lazarevic, V., Pagni, M., and Karamata, D. 1999, Mol. Microbiol., 31, 795. 154. Brown, S., Meredith, T., Swoboda, J., and Walker, S. 2010, Chem. Biol., 17, 1101. 155. Fischer, W., Mannsfeld, T., and Hagen, G. 1990, Biochem. Cell Biol., 68, 33. 156. Pollack, J.H., and Neuhaus, F.C. 1994, J. Bacteriol., 176, 7252. 157. Lazarevic, V., and Karamata, D. 1995, Mol. Microbiol., 16, 345. 158. Soldo, B., Lazarevic, V., and Karamata, D. 2002, Microbiology, 148, 2079. 159. Schirner, K., Marles-Wright, J., Lewis, R.J., and Errington, J. 2009, EMBO J., 8, 830. 160. Oku, Y., Kurokawa, K., Matsuo, M., Yamada, S., Lee, B.L., and Sekimizu, K.

2009, J. Bacteriol., 191, 141. 161. Wecke, J., Perego, M., and Fischer, W. 1996, Microb. Drug Resist., 2, 123. 162. Steen, A., Buist, G., Leenhouts, K.J., El Khattabi, M., Grijpstra, F., Zomer, A.L.,

Venema, G., Kuipers, O.P., and Kok, J. 2003, J. Biol. Chem., 278, 23874. 163. Fedtke, I., Mader, D., Kohler, T., Moll, H., Nicholson, G., Biswas, R., Henseler,

K., Götz, F., Zähringer, U., and Peschel, A. 2007, Mol. Microbiol., 65, 1078. 164. Schlag, M., Biswas, R., Krismer, B., Kohler, T., Zoll, S., Yu, W., Schwarz, H.,

Peschel, A., and Götz, F. 2010, Mol. Microbiol., 75, 864. 165. Fedtke, I., Götz, F., and Peschel, A. 2004, Int. J. Med. Microbiol., 294,189. 166. Seo, H.S., Michalek, S.M., and Nahm, M.H. 2008, Infect. Immun., 76, 206. 167. Morath, S., Geyer, A., and Hartung, T. 2001, J. Exp. Med., 193, 393. 168. Weidenmaier, C., Kokai-Kun, J.F., Kristian, S.A., Chanturiya, T., Kalbacher, H.,

Gross, M., Nicholson, G., Neumeister, B., Mond, J.J, and Peschel, A. 2004, Nat. Med., 10, 243.


169. Fittipaldi, N., Sekizaki, T., Takamatsu, D., Harel, J., Domínguez-Punaro Mde, L., Von Aulock, S., Draing, C., Marois, C., Kobisch, M., and Gottschalk, M. 2008, Infect. Immun., 76, 3587.

170. Weidenmaier, C., McLoughlin, R.M., and Lee, J.C. 2010, PLoS One, 5, e13227. 171. Gross, M., Cramton, S.E., Götz, F., and Peschel, A. 2001, Infect. Immun.,

69, 3423. 172. Lazarevic, V., Soldo, B., Médico, N., Pooley, H., Bron, S., and Karamata, D.

2005, Appl. Environ. Microbiol., 71, 39. 173. Heptinstall, S., Archibald, A.R., and Baddiley, J. 1970, Nature, 225, 519. 174. Hyyrylainen, H.L., Vitikainen, M., Thwaite, J., Wu, H., Sarvas, M., Harwood,

C.R., Kontinen, V.P., and Stephenson, K. 2000, J. Biol. Chem., 275, 26696. 175. Kristian, S.A., Lauth, X., Nizet, V., Goetz, F., Neumeister, B., Peschel, A., and

Landmann R. 2003, J. Infect. Dis., 188, 414. 176. Kovács, M., Halfmann, A., Fedtke, I., Heintz, M., Peschel, A., Vollmer, W.,

Hakenbeck, R., and Brückner, R. 2006, J. Bacteriol., 188, 5797. 177. Koprivnjak, T., Weidenmaier, C., Peschel, A., and Weiss, J.P. 2008, Infect.

Immun., 76, 2169. 178. Kohler, T., Weidenmaier, C., and Peschel, A. 2009, J. Bacteriol., 191, 4482. 179. Neuhaus, F.C., and Baddiley, J. 2003, Microbiol. Mol. Biol. Rev., 67, 686. 180. Freymond, P.P., Lazarevic, V., Soldo, B., and Karamata, D. 2006, Microbiology,

152, 1709. 181. Brown, S., Zhang, Y.H., and Walker, S. 2007, Chem. Biol., 15, 12. 182. Ginsberg, C., Zhang, Y.H., Yuan, Y., and Walker, S. 2006, ACS Chem. Biol., 17, 25. 183. Bhavsar, A.P., Truant, R., and Brown, E.D. 2005, J. Biol. Chem., 280, 36691. 184. Pooley, H.M., Abellan, F.X., and Karamata, D. 1992, J. Bacteriol., 174, 646. 185. Schertzer, J.W., and Brown, E.D. 2003, J. Biol. Chem., 278, 18002. 186. Pooley, H.M., Abellan, F.X., and Karamata, D. 1991, J. Gen. Microbiol.,

137, 921. 187. Allison, S.E., D'Elia, M.A., Arar, S., Monteiro, M.A., and Brown, E.D. 2011, J.

Biol. Chem., 286, 23708. 188. Soldo, B., Lazarevic, V., Margot, P., and Karamata, D. 1993, J. Gen. Microbiol.,

139, 3185. 189. Perego, M., Glaser, P., Minutello, A., Strauch, M.A., Leopold, K., and Fischer,

W. 1995, J. Biol. Chem., 270, 15598. 190. Kawai, Y., Marles-Wright, J., Cleverley, R.M., Emmins, R., Ishikawa, S.,

Kuwano, M., Heinz, N., Bui, N.K., Hoyland, C.N., Ogasawara, N., Lewis, R.J., Vollmer, W., Daniel, R.A., and Errington, J. 2011, EMBO J. doi: 10.1038/emboj.2011.358. [Epub ahead of print].

191. Pereira, M.P., D'Elia, M.A., Troczynska, J., and Brown, E.D. 2008, J. Bacteriol., 190, 5642.

192. Schirner, K., Stone, L.K., and Walker, S. 2011, ACS Chem. Biol., 20, 407. 193. Bhavsar, A.P., Erdman, L.K., Schertzer, J.W., Brown, E.D. 2004, J. Bacteriol.,

186, 7865. 194. D'Elia, M.A, Millar, K.E., Beveridge, T.J., and Brown, E.D. 2006, J. Bacteriol.,

188, 8313.


195. D'Elia, M.A., Pereira, M.P., Chung, Y.S., Zhao, W., Chau, A., Kenney, T.J., Sulavik, M.C., Black, T.A., and Brown, E.D. 2006, J. Bacteriol., 188, 4183.

196. D'Elia, M.A., Millar, K.E., Bhavsar, A.P., Tomljenovic, A.M., Hutter, B., Schaab, C., Moreno-Hagelsieb, G., and Brown, E.D. 2009, Chem. Biol., 16, 548.

197. Atilano, M.L., Pereira, P.M., Yates, J., Reed, P., Veiga, H., Pinho, M.G., and Filipe, S.R. 2010, Proc. Natl. Acad. Sci. U S A, 107, 18991.

198. Weidenmaier, C., Peschel, A., Xiong, Y.Q., Kristian, S.A., Dietz, K., Yeaman, M.R., and Bayer, A.S. 2005, J. Infect. Dis., 191, 1771.

199. Mobley, H.L., Koch, A.L., Doyle, R.J., and Streips, U.N. 1984, J. Bacteriol., 158, 169.

200. Clarke-Sturman, A.J., Archibald, A.R., Hancock, I.C., Harwood, C.R., Merad, T., and Hobot, J.A. 1989, J. Gen. Microbiol., 135, 657.

201. Merad, T., Archibald, A.R., Hancock, I.C., Harwood, C.R., and Hobot, J.A. 1989, J. Gen. Microbiol., 135, 645.

202. Daniel, R.A., and Errington, J. 2003, Cell, 113, 767. 203. Tiyanont, K., Doan, T., Lazarus, M.B., Fang, X., Rudner, D.Z., and Walker, S.

2006, Proc. Natl. Acad. Sci. U S A, 103, 11033. 204. Kawai, Y., Daniel, R.A., and Errington, J. 2009, Mol. Microbiol., 71, 1131. 205. Kawai, Y., Asai, K., and Errington, J. 2009, Mol. Microbiol., 73, 719. 206. Formstone, A., Carballido-López, R., Noirot, P., Errington, J., and Scheffers, D.J.

2008, J. Bacteriol., 190, 1812. 207. Jorasch, P., Wolter, F.P., Zähringer, U., and Heinz, E. 1998, Mol. Microbiol., 29, 419. 208. Kiriukhin, M.Y., Debabov, D.V., Shinabarger, D.L., and Neuhaus, F.C. 2001, J.

Bacteriol., 183, 3506. 209. Gründling, A., and Schneewind, O. 2007, J. Bacteriol., 189, 2521. 210. Uchikawa, K., Sekikawa, I., and Azuma, I. 1986, J. Bacteriol., 168, 115. 211. Webb, A.J., Karatsa-Dodgson, M., and Gründling, A. 2009, Mol. Microbiol., 74, 299. 212. Peschel, A., Otto, M., Jack, R.W., Kalbacher, H., Jung, G., and Götz, F. 1999, J.

Biol. Chem., 274, 8405. 213. Hether, N.W., and Jackson, L.L. 1983, J. Bacteriol., 156, 809. 214. Wörmann, M.E., Corrigan, R.M., Simpson, P.J., Matthews, S.J., and Gründling,

A. 2011, Mol. Microbiol., 79, 566. 215. Karatsa-Dodgson, M., Wörmann, M.E., and Gründling, A. 2010, J. Bacteriol.,

192, 5341-5349. 216. Jervis, A.J., Thackray, P.D., Houston, C.W., Horsburgh, M.J., and Moir, A. 2007,

J. Bacteriol., 189, 4534. 217. Eiamphungporn, W., and Helmann, J.D. 2008, Mol. Microbiol., 67, 830. 218. Jones, L.J., Carballido-López, R., and Errington, J. 2001, Cell, 104, 913. 219. Matias, V.R., Beveridge, T.J. 2005, Mol. Microbiol., 56, 240. 220. Matias, V.R., Beveridge, T.J. 2006, J. Bacteriol., 188, 1011. 221. Matias, V.R., and Beveridge, T.J. 2008, J. Bacteriol., 190, 7414. 222. Weart, R.B., Lee, A.H., Chien, A.C., Haeusser, D.P., Hill, N.S., and Levin, P.A.

2007, Cell, 130, 335.

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4-3. Cell wall structure of E. coli and B. subtilis...Cell wall structure 117 approximately 10 times thicker than in the E. coli cell wall, and therefore the B. subtilis cell wall