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

Research Signpost
37/661 (2), Fort P.O.
Trivandrum-695 023
Kerala, India
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]
116
Junichi Sekiguchi & Hiroki Yamamoto
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, grampositive 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 glycerolphosphate-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 penicillinbinding 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 Lconfiguration), 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 γglutamyl bond. In E. coli, m-A2pm-NH2 and m-A2pm cross-linkage as well as D-Alam-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; DGlu, 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-Lalanine amidase; 4, LD-endopeptidase; 5, DL-endopeptidase; 6, DD-endopeptidase; 7,
DD-carboxypeptidase; and 8, LD-carboxypeptidase.
118
Junichi Sekiguchi & Hiroki Yamamoto
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 (Oacetylation 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.
Cell wall structure
119
Figure 2. Spore peptidoglycan (cortex) structure of B. subtilis. δ indicates the deltalactam 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
Table 1. Cell wall (murein) hydrolases in E. coli.
120
Junichi Sekiguchi & Hiroki Yamamoto
Table 1. Continued
Cell wall structure
121
Table 2. Cell wall (murein) hydrolases and paralogs in B. subtilis
122
Junichi Sekiguchi & Hiroki Yamamoto
Table 2. Continued
Cell wall structure
123
Table 2. Continued
124
Junichi Sekiguchi & Hiroki Yamamoto
Table 2. Continued
Cell wall structure
125
126
Junichi Sekiguchi & Hiroki Yamamoto
[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, DDendopeptidase which cleaves the D-Ala-meso-A2pm cross-linkage (mesoA2pm 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
Cell wall structure
127
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 DDendopeptidase 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 (DDcarboxypeptidase 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-NAcetylglucosaminidase 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-anhMurNAcpeptides 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
128
Junichi Sekiguchi & Hiroki Yamamoto
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 Cterminal 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 twocomponent 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 domaincontaining 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
Cell wall structure
129
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 –[GlcNAcMurNAc]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
130
Junichi Sekiguchi & Hiroki Yamamoto
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-Glumeso-A2pm (DL-), and cross-linking D-Ala-meso-A2pm (DD-) linkages (Fig.
1), and these cleaving endopeptidases have been found in B. subtilis. LDendopeptidase 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]. DLendopeptidase 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 Nterminal (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 Nacetylmuramidase 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 Lglutamic 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
Cell wall structure
131
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-Nacetylglucosaminidase), LytG (exo-N-acetylglucosaminidase), glycohydrolase
family
18
(endo-N-acetylglucosaminidase)
and
NagZ
(exo-Nacetylglucosaminidase) (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 cortexhydrolyzing amidase [122, 123], but muropeptide analysis suggests that SleB
132
Junichi Sekiguchi & Hiroki Yamamoto
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 (Nterminal 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 twodomain 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 Nterminal. 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 twocomponent system, YycFG (WalRK) [74, 75]. In contrast, the DLendopeptidase 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
Cell wall structure
133
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, PBPC3, 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].
134
Junichi Sekiguchi & Hiroki Yamamoto
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, glycerolphosphate, 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 Nacetylmuramyl residue in peptidoglycan via a linkage unit, which consists of
Cell wall structure
135
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 1phosphate, 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
136
Junichi Sekiguchi & Hiroki Yamamoto
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, undecaprenylpyrophosphate, 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 phosphatelimiting 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
Cell wall structure
137
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
138
Junichi Sekiguchi & Hiroki Yamamoto
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 galactosylglucosyl-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 GalGlcDAG, 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
Cell wall structure
139
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 LtaSdepleted 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 GroPGlc2-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.
140
Junichi Sekiguchi & Hiroki Yamamoto
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 LTAdeficient 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
Cell wall structure
141
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.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
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.
Vollmer, W. and Bertsche, U. 2008, Biochim. Biophys. Acta, 1778, 1714.
Vollmer, W., Blanot, D., and de Pedro, M. A. 2008, FEMS Microbiol. Rev.,
32, 149.
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.
Schleifer, K. H. and Kandler, O. 1972, Bacteriological Rev., 36, 407.
Höltje, J. V. 1998, Microbiol. Mol. Biol. Rev., 62, 181.
Mendelson, N. H. and Thwaites, J. J. 1989, J. Bacteriol., 171, 1055.
Harz, H., Burgdorf, K., and Höltje, J.-V. 1990, Anal. Biochem., 190, 120.
Vollmer, W. and Seligman, S. J. 2010, Trends Microbiol., 18, 59.
Ward, J. 1973, Biochem. J., 133, 395.
Braun, V. and Wolff, H. 1970, Eur. J. Biochem., 14, 387.
Guariglia-Oropenza, V. and Helmann, J. D. 2011, J. Bacteriol., 193, 6223.
Gan, L., Chen, S., and Jensen, G. J. 2008, Proc. Natl. Acad. Sci. U.S.A., 105,
18953.
Sekiguchi, J., Akeo, K., Yamamoto, H., Khasanov, F. K., Alonso, J. C. and
Kuroda, A. 1995, J. Bacteriol., 177, 5582.
Popham, D. L., Helin, J., Costello, C. E., Setlow, P. 1996, Proc Natl Acad Sci
USA. 93, 15405.
Atrih, A., Zollner, P., Allmaier, G., and Foster, S. J. 1996, J. Bacteriol., 178, 6173.
Fukushima, T., Yamamoto, H., Atrih, A., Foster, S. J., and Sekiguchi, J. 2002, J.
Bacteriol., 184, 6007.
142
Junichi Sekiguchi & Hiroki Yamamoto
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.
Cell wall structure
143
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.
144
76.
77.
78.
79.
Junichi Sekiguchi & Hiroki Yamamoto
Reith, J. and Mayer, C. 2011, Appl. Microbiol. Biotechnol., 92, 1.
Kuroda, A. and Sekiguchi, J. 1990, J. Gen. Microbiol., 136, 2209.
Kuroda, A., Imazeki, M., and Sekiguchi, J. 1991, FEMS Microbiol. Lett., 81, 9-14.
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.
Cell wall structure
145
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.
146
Junichi Sekiguchi & Hiroki Yamamoto
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.
Cell wall structure
147
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.
148
Junichi Sekiguchi & Hiroki Yamamoto
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.