Chapter 5

Peptidoglycan

Manuel Pazos and Katharina Peters

Abstract The peptidoglycan sacculus is a net-like polymer that surrounds the cyto-
plasmic membrane in most bacteria. It is essential to maintain the bacterial cell shape
and protect from turgor. The peptidoglycan has a basic composition, common to all
bacteria, with species-specific variations that can modify its biophysical properties
or the pathogenicity of the bacteria. The synthesis of peptidoglycan starts in the cyto-
plasm and the precursor lipid II is flipped across the cytoplasmic membrane. The
new peptidoglycan strands are synthesised and incorporated into the pre-existing
sacculus by the coordinated activities of peptidoglycan synthases and hydrolases.
In the model organism Escherichia coli there are two complexes required for the
elongation and division. Each of them is regulated by different proteins from both
the cytoplasmic and periplasmic sides that ensure the well-coordinated synthesis of
new peptidoglycan.

Keywords Peptidoglycan · Sacculus · Lipid II · PBPs · LDTs · TPase · GTase ·

Glycan strands · Cross-linking · Elongation · Septation

Introduction

The peptidoglycan (PG) sacculus is an elastic and net-like polymer that surrounds the
cytoplasmic membrane in most bacteria. It is rigid enough to maintain the species-
specific bacterial cell shape, serving as a scaffold to attach proteins and other poly-
mers, but also porous enough to allow the diffusion of chemical signals, nutrients
and virulence factors. The PG protects the cell from bursting due to its turgor, which
pushes the cytoplasmic membrane towards the cell wall. In Gram-negative bacteria,
a thin and single PG layer is located in the periplasm surrounding the cytoplasmic

M. Pazos (B) · K. Peters

Centre for Bacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle
University, Baddiley-Clark Building, Richardson Road, Newcastle upon Tyne NE2 4AX, UK
e-mail: manuel.pazos@ncl.ac.uk
K. Peters
e-mail: katharina.peters@ncl.ac.uk

© Springer Nature Switzerland AG 2019 127

A. Kuhn (ed.), Bacterial Cell Walls and Membranes, Subcellular
Biochemistry 92, https://doi.org/10.1007/978-3-030-18768-2_5
128 M. Pazos and K. Peters

membrane. In contrast, the PG in Gram-positive bacteria is thicker and multi-layered

with covalently attached cell wall compounds like capsular polysaccharides, cell sur-
face proteins and wall teichoic acids (Fig. 5.1). Bacteria belonging to Rickettsiaceae,
Anaplasmataceae and Mycoplasmataceae families do not have PG. The presence of
PG in Chlamydiaceae has been proven (Pilhofer et al. 2013; Packiam et al. 2015;
Liechti et al. 2016). In order to preserve the structural integrity of the cell envelope
during cell growth and division, the synthetic and hydrolytic PG enzymes must be
coordinated to enlarge and divide the sacculi. According to the current model this
coordination is achieved by multi-enzyme complexes that extend from the cytoplasm
to the outer membrane.
This chapter is mainly focused on the model organism Escherichia coli, but the
reader will find references to other organisms along the different sections. After
describing the PG composition, modifications and its biophysical properties, the
different enzymatic activities and their regulation are presented in the cellular con-
text. The importance of the fluorescent d-amino acids for the field is highlighted,
and a detailed description of the current understanding of the PG substrate flippase
candidates and the unusual ld-transpeptidases (LDTs) is included.

LTA
WTA

LPS
OM

Lpp

PG

Periplasm

CM

Gram-positive Gram-negative

Fig. 5.1 Schematic structure of the cell wall in Gram-positive and Gram-negative bacteria. C, cyto-
plasm; CM, cytoplasmic membrane; PG, peptidoglycan; OM, outer membrane; LPS, lipopolysac-
charide; LTA, lipoteichoic acid; WTA, wall teichoic acid; Lpp, Braun’s lipoprotein
5 Peptidoglycan 129

The Peptidoglycan Composition, Its Modifications and Their

Function

The PG sacculus is an essential macromolecule exclusively found in bacteria. Its

basic structure comprises linear glycan strands that are cross-linked by short pep-
tides (Schleifer and Kandler 1972). The glycan strands, whose length varies between
species, are made out of alternating β-1,4-connected N-acetylglucosamine (GlcNAc)
and N-acetylmuramic acid (MurNAc) residues. MurNAc is a lactic acid ether deriva-
tive of GlcNAc, and the stem pentapeptides are attached to its d-lactyl moiety at
position C3. The stem pentapeptides contain l- and d-amino acids, whereby the lat-
ter is characteristic of bacterial PG. The PG structure shows high diversity between
different species, but it is well conserved in bacteria belonging to the same species.
Modifications in the PG can enhance the bacterial fitness and virulence, providing
resistance against environmental stresses, hydrolytic host enzymes and antimicrobial
agents.

Variations in the Stem Peptide

The amino acid composition of the pentapeptide shows species-specific variations.

Some examples are shown in Fig. 5.2 and described in the following lines. In Gram-
negative bacteria like E. coli the pentapeptide in nascent PG consists of l-Ala-d-iGlu-
mDAP-d-Ala-d-Ala (mDAP: meso-diaminopimelic acid). Two adjacent peptides are
more frequently cross-linked via an amide bond between the carboxyl group of the
d-Ala at position four of one peptide and the ε-amino group of mDAP at position
three of another one. The d-Ala-d-Ala motif of the pentapeptide is the universal
substrate of the PG cross-linking enzymes. The terminal d-Ala is released during the
cross-linking reaction.
In Mycobacterium leprae the first amino acid of the stem peptide is Gly instead of
l-Ala. This modification is suggested to be due to the growth environment (Mahapatra
et al. 2000). In the Gram-positive pathogens Streptococcus pneumoniae and Staphy-
lococcus aureus, and in Mycobacterium tuberculosis the α-carboxyl group of the
d-iso-glutamate on position two of the stem peptide is amidated into d-iso-glutamine
by the essential amidotransferase complex MurT/GatD (Zapun et al. 2013; Morlot
et al. 2018; Münch et al. 2012; Figueiredo et al. 2012). In S. pneumoniae unamidated
glutamate is predominantly found in uncross-linked PG monomers (Bui et al. 2012).
In vitro assays using recombinant pneumococcal PBPs revealed that the amidation
of the stem peptides is needed for an efficient cross-linking reaction (Zapun et al.
2013). In S. aureus the lack of amidation correlates with a decrease in cross-linking
and with a higher susceptibility to antibiotics (Stranden et al. 1997; Boyle-Vavra
et al. 2001; Figueiredo et al. 2012). Thus, understanding the molecular mechanism
of the amidation reaction may help in the development of new therapeutics to tar-
get these important pathogens. Recent studies provide biochemical and structural
130 M. Pazos and K. Peters

GlcNAc – MurNAc GlcNAc – MurNAc GlcNAc – MurNAc GlcNAc – MurNAc

1 L-Ala L-Ala L-Ala L-Ala

2 D-iGlu D-iGlu D-iGlu(NH2) D-iGlu(NH2)

3 mDAP D-Ala 4 mDAP(NH2) D-Ala L-Lys L-Ser L-Ala D-Ala L-Lys (Gly)5 D-Ala

4 D-Ala mDAP 3 D-Ala mDAP(NH2) L-Lys D-Ala L-Lys (Gly)5

D-iGlu 2 D-iGlu D-iGlu(NH2) D-iGlu(NH2)

L-Ala L-Ala L-Ala L-Ala

1
GlcNAc – MurNAc GlcNAc – MurNAc GlcNAc – MurNAc GlcNAc – MurNAc

4-3 cross-link in E. coli 4-3 cross-link in B. subtilis 4-3 cross-link in S. pneumoniae 4-3 cross-link in S. aureus
(major cross-links) with L-Ser-L-Ala bridge with pentaglycine bridge

GlcNAc – MurNAc GlcNAc – MurNAc GlcNAc – MurNAc

L-Ala L-Ala Gly GlcNAc – MurNAc
(NH2)
D-iGlu D-iGlu(NH2) D-iGlu D-Orn D-Ala L-Ala mDAP

L-Hse D-iGlu
mDAP mDAP mDAP(NH2) mDAP(NH2) L-Hse
L-Ala
D-Ala D-iGlu D-Ala D-iGlu(NH2) D-Ala D-iGlu
GlcNAc – MurNAc
L-Ala L-Ala Gly
GlcNAc – MurNAc GlcNAc – MurNAc GlcNAc – MurNAc

3-3 cross-link in E. coli 3-3 cross-link in M. tuberculosis 4-2 cross-link in C. pointsettiae 1-3 cross-link in Acetobacteria
(minor cross-links) with D-ornithine bridge with amidated mDAP

Fig. 5.2 Examples of peptides and cross-link types in the peptidoglycan of different species. Ami-
dation of residues is depicted in orange. Interpeptide bridges are framed with a square

insights about the MurT/GatD complex of S. pneumoniae (Morlot et al. 2018) and
S. aureus (Noldeke et al. 2018). The amino acid at position three of the stem peptide
shows the greatest variation. Most Gram-negative bacteria, Mycobacteria and Bacilli
contain an mDAP residue at this position. Bacillus subtilis has an amidated mDAP
(Atrih et al. 1999), due to the action of the amidotransferase AsnB (Dajkovic et al.
2017). In M. tuberculosis AsnB also amidates the mDAP residue, and this modi-
fication is essential for cell growth (Ngadjeua et al. 2018). The amidotransferase
LtsA performs the amidation of the mDAP residue in the PG of Corynebacterium
glutamicum (Levefaudes et al. 2015). Spirochetes, such as Borrelia or Treponema,
have an ornithine residue instead of mDAP (Schleifer and Kandler 1972; Yanagi-
hara et al. 1984). Other species contain at position three different diamino acids like
meso-lanthionine (Fusobacterium nucleatum) and d-Lys (Thermatoga maritima),
or monoamino acids like l-Ala, l-Glu or l-homoserine (reviewed in Vollmer et al.
2008a). However, most Gram-positive bacteria have an l-Lys at position three, which
often carries a linear peptide branch linked to its ε-amino group. These interpeptide
bridges show a great diversity, with sizes varying from two to seven residues and a
wide range of amino acids. In case of S. aureus the FemXAB peptidyltransferases
catalyse the addition of a characteristic Gly5 -interpeptide bridge (Schleifer and Kan-
dler 1972). S. pneumoniae contains branched stem peptides with an l-Ser-l-Ala or
l-Ala-l-Ala dipeptide linked to the ε-amino group of l-Lys. This modification is
added by MurM and MurN (Filipe and Tomasz 2000). The degree of branching and
cross-linking of PG varies between strains. While most pneumococcal strains con-
tain a small percentage of branched peptides, the PG of resistant strains is highly
branched and cross-linked (Garcia-Bustos and Tomasz 1990; Severin et al. 1996).
Most bacteria contain d-Ala-d-Ala at positions four and five of the stem peptide.
5 Peptidoglycan 131

This motif is recognized by vancomycin and other glycopeptide antibiotics that form
a complex with the PG precursors, preventing their incorporation into the sacculus
of Gram-positive bacteria. The replacement of the d-Ala at position five by d-Lac or
d-Ser prevents the binding of the antibiotic to the precursor and mediates resistance
of enterococci (Arthur et al. 1993, 1996).

Variations in the Peptide Cross-Links

The type and extent of the peptide cross-links is different between species. The
most abundant cross-links connect the d-Ala at position four of one stem peptide
with the mDAP (or l-Lys) at position three of another one (4-3 cross-links). They
are synthesized by DD-transpeptidases (dd-TPases). Less frequent are the 4-2 cross-
links, found in Corynebacteria, that connect the d-Ala at position four and the d-iGlu
at position two of adjacent stem peptides. Corynebacterium pointsettiae contains
an l-homoserine at position three of the stem peptide, which is non-reactive to
form cross-links. Therefore, the cross-link starts at d-iGlu and is created via a d-
ornithine bridge (Fig. 5.2) (Schleifer and Kandler 1972). Some bacteria contain a
small amount of 3-3 cross-links, while pathogens like Clostridium difficile and M.
tuberculosis contain predominantly 3-3 cross-links in the PG. These cross-links are
made by ld-transpeptidases (LDTs) that connect two mDAP residues of adjacent
stem peptides (see section “Peptidoglycan Synthesis” for further details on the cross-
linking reactions).
Recently, novel PG structures have been described in Acetobacteria, which prolif-
erate at low pH and produce acetic acid. These modifications include the amidation
of the α-(l)-carboxyl group of mDAP and a novel ld (1-3) cross-link, which con-
nects the l-Ala residue of one stem peptide with the mDAP of another one (Fig. 5.2)
(Espaillat et al. 2016). The enzymes catalyzing both modifications are unknown.

Modifications of the Glycan Strands

The glycan strands can be modified by N-deacetylation and O-acetylation on either

one or both GlcNAc and MurNAc subunits, and by N-glycolylation on MurNAc
(Fig. 5.3) (reviewed in Vollmer 2008; Yadav et al. 2018). The N-glycolylation reac-
tion occurs in the cytoplasm during the synthesis of the PG precursors. The N-
deacetylation and O-acetylation reactions take place outside the cytoplasm, once
the glycan strands have been synthesized. In pathogenic species these modifications
contribute to the survival in the host organism by increasing the resistance to PG
degrading enzymes, preventing cell lysis and the release of bacterial products that
could be detected by the host immune system.
In many Gram-positive and Gram-negative bacteria the PG chains contain an
extra acetyl group linked to the C6–OH group of some MurNAc (O-acetylation). The
132 M. Pazos and K. Peters

CH2 O

CH3 O O CH3
C O C O
O O
CH2 HCCH3 NH CH2
O HO O
O C O C O HO OH
HO Pep CH3 O
NH NH
HCCH3
2,6-N,O- C O C O
1,6-Anhydro ring C O 2,6-N,O-
diacetyl-GlcN CH3 CH3 diacetyl-MurN
at MurNAc Pep

O-acetylation at GlcNAc O-acetylation at MurNAc

CH2OH O CH2OH O
O
HO O
OH
NH O
HCCH3 NH
C O
C O
CH2OH CH3 C O CH2OH
O CH3 O
O Pep HO OH
HO GlcNAc MurNAc O
NH3+ NH3+
HCCH3
Unmodified disaccharide C O
GlcN MurN
Pep
N-deacetylation at GlcNAc
N-deacetylation at MurNAc
CH2OH O
HO OH
O
NH
HCCH3
C O C O
CH2OH
Pep
MurNGlyc
N-glycolylation at MurN

Fig. 5.3 Summary of modifications in the PG glycan strands. The structure of the unmodified
GlcNAc-MurNAc disaccharide (middle) and of selected modifications in the GlcNAc (red) and
MurNAc (blue) are shown. Modifications are highlighted in orange. The O-acetylation of GlcNAc
or MurNAc is reversible. Pep, peptide linked to MurNAc. Modified from Vollmer (2008) and Yadav
et al. (2018)

degree of O-acetylation varies between <20 and 70% depending on the species and
growth conditions. The bulky acetyl group represents a steric hindrance and prevents
the binding of the muramidase lysozyme (Pushkaran et al. 2015), which is secreted
by the host immune system cells to hydrolyse the glycan strands between MurNAc
and GlcNAc. Therefore, PG O-acetylation is a major virulence factor in pathogenic
bacteria. For the O-acetylation of MurNAc, the acetyl moiety is translocated from a
cytoplasmic donor molecule to the periplasm or extracellular space and transferred
to the MurNAc subunit. In Gram-positive bacteria the O-acetyltransferases of the
OatA-type perform both processes (Bera et al. 2005; Bernard et al. 2012), while
Gram-negative bacteria require the coordinated action of multiple enzymes of the
Pat or Pac family (Weadge et al. 2005; Moynihan and Clarke 2010; Dillard and
Hackett 2005). In S. aureus OatA provides resistance to lysozyme (Bera et al. 2005),
protects against killing by macrophages (Shimada et al. 2010), reduces the induction
of pro-inflammatory cytokines and permits reinfection (Sanchez et al. 2017). OatA
homologs have been identified in different Gram-positive species. In S. pneumoniae
5 Peptidoglycan 133

the OatA homolog Adr catalyses the O-acetylation of MurNAc and protects dividing
cells from cleavage by the pneumococcal autolysin LytA. Adr localizes at the septa
and shows mislocalization in some cell division mutants, suggesting an important role
in pneumococcal cell division (Bonnet et al. 2017). In vancomycin-resistant Ente-
rococcus faecalis the vancomycin treatment increases the levels of O-acetylation,
which leads to lysozyme resistance and to an increase in virulence (Chang et al.
2017). O-acetylation of MurNAc blocks the function of the lytic transglycosylases
(LTs), which have the same substrate specificity as lysozyme but catalyse a transg-
lycosylation reaction resulting in the formation of 1,6-anhydro-MurNAc (see section
“Peptidoglycan Hydrolysis and Remodelling” for further information).
In Gram-negative bacteria the level of O-acetylation regulates the activities of
the LTs (Weadge et al. 2005; Weadge and Clarke 2006). Glycan strands can be O-
deacetylated at the MurNAc O-acetyl subunit. The O-acetylesterase Ape reverts the
O-acetylation of MurNAc, and Ape homologs have been found in Gram-positive and
Gram-negative bacteria (Weadge et al. 2005). In Campylobacter jejuni the deletion
of ape1 results in the accumulation of O-acetylated PG, which impairs the cellular
fitness and leads to defects in morphology, motility, biofilm formation and viru-
lence (Ha et al. 2016). This highlights the importance of regulating the degree of
O-acetylation via the activity of O-acetyltransferases and O-acetylesterases for the
bacterial cell. In contrast, the O-acetylation of GlcNAc has been only described in
Lactobacillus plantarum, where it inhibits the major autolysin Acm2 (Bernard et al.
2011).
N-deacetylation of GlcNAc occurs mostly in Gram-positive bacteria (Vollmer and
Tomasz 2000; Boneca et al. 2007; Peltier et al. 2011; Benachour et al. 2012) but
also in some Gram-negative like Shigella flexneri (Kaoukab-Raji et al. 2012). The
PG deacetylase A enzyme (PgdA), identified for the first time in S. pneumonia,
removes the acetyl group at position C2 of the GlcNAc. PgdA mutants are more
susceptible to lysozyme and less virulent (Vollmer and Tomasz 2000). In Listeria
monocytogenes 50% of the GlcNAc residues are deacetylated (Boneca et al. 2007)
and ~23% of the MurNAc residues are O-acetylated at position C6 (Aubry et al.
2011), whereby both modifications enhance synergistically the resistance against
lysozyme. In this organism, PgdA activity is regulated by the cell division protein
GpsB and the PG synthase PBP A1 (Rismondo et al. 2018). The absence of GpsB
increases the lysozyme resistance due to a rise in N-deacetylated muropeptides, and
the absence of PBP A1 reverses this phenotype. This regulation is supported by
in vitro studies suggesting that all three proteins form a complex. In Lactococcus
lactis YvhB catalyses the O-acetylation of MurNAc, and XynD performs the N-
deacetylation of GlcNAc. Both modifications increase the cross-linkage and the cell
wall integrity, leading to acid resistance and to the production of the polycyclic
antibacterial peptide nisin (Cao et al. 2018). In the predatory bacteria Bdellovibrio
bacteriovorus, the PgdA homologs Bd0468 and Bd3279 deacetylate the GlcNAc
residues of the prey PG during predation, making it more susceptible for destruction
at the end of predation (Lambert et al. 2016).
134 M. Pazos and K. Peters

The N-glycolylation of MurNAc subunits exists in Mycobacteria and related Acti-

nomycetales. This modification is added by the cytoplasmic UDP-MurNAc hydrox-
ylase NamH during the synthesis of the UDP-linked precursors, where the N-acetyl
group at the C2 of the muramyl dipeptides is hydroxylated to an N-glycolyl group.
The degree of N-glycolylation varies depending on the bacterial species, and it
changes in response to antibiotics. The deletion of namH increases the suscepti-
bility to lysozyme and β-lactams (Raymond et al. 2005). N-glycolylated MurNAc
activates the innate immunity more than N-acetylated MurNAc, underlining its con-
tribution to the unusual immunogenicity of Mycobacteria (Coulombe et al. 2009)
but not to the pathogenicity of M. tuberculosis infection (Hansen et al. 2014).

Biophysical Properties of Peptidoglycan

The PG sacculus has essential stress-bearing functions in the cell, which include
maintaining the cellular shape during cell growth and division within changing envi-
ronmental conditions, and constraining the cell volume under turgor. In order to fulfil
these functions, the PG is elastic and stiff at the same time, but also porous to allow
diffusion of small proteins.

Thickness of Peptidoglycan

The osmotic pressure in Gram-negative bacteria like E. coli is up to 3 atm (Cay-

ley et al. 2000) and in Gram-positive bacteria such as B. subtilis and S. aureus is
significantly higher (up to 20 atm) (Whatmore and Reed 1990). The PG provides
mechanical strength to counteract the osmotic pressure in the cell (Höltje 1998;
Koch 1988). The thickness of the PG, which can vary depending on the species,
contributes to cell stiffness. Consistent with that, Gram-positive bacteria contain a
multi-layered and thick PG. The PG of Gram-negative bacteria is a thin and single
layer located between the inner and the outer membranes (Fig. 5.1). A recent study
suggests that mechanical loads are balanced between the outer membrane and the
PG, and postulates a stress-bearing function for the outer membrane during osmotic
changes (Rojas et al. 2018).
Isolated PG sacculi maintain the shape and the size of intact cells. Using atomic
force microscopy (AFM) dried sacculi from E. coli are 3 nm thick, while rehydrated
sacculi are 6 nm thick (Yao et al. 1999). In the same study rehydrated sacculi of
Pseudomonas aeruginosa are only 3 nm thick. These data are in agreement with
measurements performed by cryo-transmission electron microscopy (cryo-TEM) of
frozen-hydrated sections, which preserves native structures (E. coli: 6.35 ± 0.53 nm;
P. aeruginosa: 2.41 ± 0.54 nm) (Matias et al. 2003).
5 Peptidoglycan 135

Cryo-TEM analysis of frozen-hydrated sections of B. subtilis cells revealed two

distinct zones: a 22 nm thick low-density zone surrounding the cytoplasmic mem-
brane and a 33 nm thick high-density zone surrounding the previous one. The high
density zone contains PG, teichoic acids and associated proteins (Matias and Bev-
eridge 2005). A thinner B. subtilis ponA mutant (lacking the bifunctional PG syn-
thase PBP1) shows a 30 nm thick PG (Beeby et al. 2013).

Strength, Stiffness and Elasticity of Peptidoglycan

The mechanical properties of PG have been analysed using different techniques.

The cellular stiffness can be described by the Young’s modulus, also known as the
modulus of elasticity. To determine it, a physical force is applied to a sample (either
cells or isolated sacculi) and the degree by which it can be stretched is measured. If
the sample can withstand the force, it will return to the preloaded size after removing
the force. If the load is too high for the elastic limits of the sample, it will irreversibly
deform and rupture. The elasticity of isolated E. coli sacculi, measured by stretching
them with the tip of the AFM, varies on the hydration state as air dried sacculi are more
rigid than rehydrated ones. Differences in the elastic anisotropy between the long
and the short axis of the sacculus suggests that the more flexible peptide cross-links
are longitudinally orientated and the more rigid glycan strands are circumferentially
orientated (Yao et al. 1999). The proposed architecture is supported by the fact
that changes in the volume of osmotically shocked E. coli cells are mainly due
to changes in cell length but not in cell diameter (van den Bogaart et al. 2007).
Recent analysis of individual E. coli glycan chains by AFM have shown long and
circumferentially orientated glycan chains (200 nm) under normal conditions and
shorter and disordered glycan chains when a spheroid cell shape is induced (Turner
et al. 2018).
Another technique to measure the mechanical properties of bacterial cells is the
Cell Length Analysis of Mechanical Properties (CLAMP), which enables a high
throughput screening. Using this technique the Young’s moduli of Gram-negative
and Gram-positive rod-shaped bacteria are estimated on the basis of initial growth
rates of hydrogel-encapsulated living cells with varying stiffness (Tuson et al. 2012).
A decrease in relative cell elongation over time is observed with increasing stiffness of
the hydrogels. The estimated Young’s modulus for E. coli is fourfold higher than the
result obtained with AFM for hydrated E. coli sacculi (Yao et al. 1999). This differ-
ence can be explained by the fact that the outer membrane also contributes to the cell
wall stiffness. Using the CLAMP technique the Young’s moduli of the cell envelopes
of P. aeruginosa and B. subtilis are similar to E. coli. Several genes affecting cell
stiffness have been identified by measuring the cell growth of hydrogel-encapsulated
mutant cells from the entire KEIO-collection (E. coli single-gene deletion mutants).
These genes have roles in diverse processes, suggesting that cell stiffness is affected
by different cellular functions beside the cell wall-related ones (Auer et al. 2016).
136 M. Pazos and K. Peters

In addition to the thickness, other structural parameters as the amount of cross-

links and the glycan strands length influence the mechanical properties of the PG.
Compared to exponentially growing cells, stationary cells of E. coli contain slightly
more cross-links, higher level of ld cross-links (see section “ld-Transpeptidases”)
and more cross-links between the outer membrane-anchored protein Lpp and the
PG, which might strengthen the cell envelope and presumably enhance the cellular
stiffness (Vollmer and Bertsche 2008). The cell division process of S. aureus changes
the mechanical properties of the cell wall, as the newly formed septum is stiffer
than the surrounding mature cell wall, most likely due to a higher level of cross-
links (Bailey et al. 2014b). A decrease in the cross-linkage of S. aureus PG, by the
absence of PBP4, results in a small reduction of cell wall stiffness (Loskill et al. 2014).
The absence of glucosaminidases, which hydrolyse the bonds between GlcNAc and
MurNAc, increases the cellular stiffness in S. aureus, increasing the glycan chains
length (Wheeler et al. 2015).
Bacteria from diverse phyla produce non-canonical d-amino acids that are incor-
porated into the PG and modify its strength and flexibility. In Vibrio cholerae the
racemase BsrV produces these amino acids (mainly d-Met and d-Leu) during sta-
tionary growth and LDTs catalyse their incorporation into the PG. The bsrV mutant
cells have shorter glycan strands and both, bsrV and ldtA ldtB (encoding for LDTs)
mutants are significantly more sensitive to osmotic shock and accumulate an excess
of PG during stationary phase (Lam et al. 2009; Cava et al. 2011).

Porosity of Peptidoglycan

In early work the pore sizes in B. subtilis and Bacillus licheniformis PG were esti-
mated (radius of 2.5 nm) by measuring the diffusion of different sized proteins out
of the cells, after n-butanol treatment to permeabilize the lipid bilayers (Hughes
et al. 1975). A more recent study using isolated E. coli and B. subtilis sacculi and
fluorescein-labelled dextrans of known molecular weights, determined a similar pore
radius for both sacculi (2.06 nm in E. coli, 2.12 nm in B. subtilis) (Demchick and
Koch 1996). On the basis of these values it was estimated that a globular, hydrophilic
protein with a molecular weight of 25 kDa can freely pass through the relaxed sacculi,
while globular proteins of about 50 kDa should be able to diffuse through stretched
sacculi (Demchick and Koch 1996). A homeostatic mechanism has been postulated
for γ-proteobacteria like E. coli, in which the PG synthesis rate is coupled to the
growth rate of the cell via the PG pores size (Typas et al. 2010).

Peptidoglycan Precursor Synthesis

The synthesis of the PG precursor starts with the formation of both nucleotide sugar-
linked precursor UDP-GlcNAc and UDP-MurNAc in the cytoplasm (Fig. 5.4) (Bar-
reteau et al. 2008). UDP-GlcNAc is synthesised from fructose 6-phosphate by the
5 Peptidoglycan 137

Outer membrane

LD- LD-
TPase Lpp TPase Lpp exo-
GlcNAc endo- LT
-NH2
MurNAc LT L-Lys-NH2 L-Lys

anhMurNAc
L-Ala
D-Ala
LD- LD- DD-
mDAP TPase EPase
EPase
L-Glu Amidase -NH2
D-iGlu
D-amino acid
Phosphate
LD- DD-
Uridine -NH2 -NH2
CPase CPase
Undecaprenol DD- DD-
TPase TPase

GTase

Periplasm

MraY flippase BacA/PAP2- flippase

Cytoplasmic membrane phosphatases
MurG
Cytoplasm

UTP

GlmU GlmM GlmS Fruc-6-P

PEP
Alr/ MurA Acetyl-CoA
DadX
MurF
DdlA/
ATP MurI
DdlB
NADPH
MurB
ATP

MurE MurD MurC

ATP ATP ATP

Fig. 5.4 Synthetic and hydrolytic reactions occurring during the synthesis and incorporation of
new peptidoglycan in E. coli. Modified from Typas et al. (2011)

enzymes GlmS, GlmM and GlmU (the last reaction requires acetyl-coenzyme A and
uridine triphosphate). UDP-MurNAc is synthesised by the addition of enolpyruvate to
UDP-GlcNAc (by MurA) and the following reduction (by MurB). The pentapeptide
moiety is added to the UDP-MurNAc by MurC, MurD, MurE and MurF enzymes in
a sequence of ATP-dependent reactions that add l-Ala, d-iGlu, mDAP and d-Ala-
d-Ala, respectively. The racemases generate the d-Ala (Alr or DadX) and d-iGlu
(MurI) enantiomers, and the ligases DdlA and DdlB) synthesize the d-Ala-d-Ala
dipeptide required. The first three amino acids (l-Ala-d-iGlu-mDAP) can also be
added as a tripeptide by the ligase Mpl through the “recycling pathway”. Phospho-
MurNAc-pentapeptide is then transferred onto the lipid carrier undecaprenyl phos-
phate by MraY, which crystal structure has been recently solved (Hakulinen et al.
2017), resulting in the formation of undecaprenyl pyrophosphate MurNAc pentapep-
tide (lipid I) and releasing uridine monophosphate (UMP). The subsequent transfer
of GlcNAc to the lipid I by MurG results in the formation of undecaprenyl pyrophos-
phate MurNAc(GlcNAc) pentapeptide (lipid II) and the release of uridine diphosphate
(UDP). In a final step lipid II is translocated (or flipped) from the inner to the outer
leaflet of the cytoplasmic membrane where it is used as substrate for the synthesis of
new PG, releasing the lipid carrier undecaprenol pyrophosphate. Due to the essen-
tiality of the process, these reaction steps are potential targets for antibiotics (Hrast
et al. 2014; Kouidmi et al. 2014; Liu and Breukink 2016).
138 M. Pazos and K. Peters

Lipid II Flippase(s)

The identity of the lipid II flippase(s) is a controversial topic in the field, and two
main candidates to catalyse this reaction has been proposed: FtsW and MurJ.
FtsW is a widely conserved integral membrane protein, member of the SEDS
(shape, elongation, division and sporulation) family, and originally identified from
a filamentous temperature sensitive mutant, showing its essential role for cell divi-
sion (Ishino et al. 1989; Khattar et al. 1994). Different studies have dissected its
10 transmembrane segments, identifying several essential residues required for the
functionality of the protein (Lara and Ayala 2002; Pastoret et al. 2004; Mohammadi
et al. 2014) and its direct interaction with other cell division proteins as FtsN, FtsQ,
PBP3 or PBP1B (Di Lallo et al. 2003; Karimova et al. 2005; Alexeeva et al. 2010;
Fraipont et al. 2011; Leclercq et al. 2017). In vitro experiments have shown that
FtsW binds lipid II (Mohammadi et al. 2011; Leclercq et al. 2017) although with
low affinity (Bolla et al. 2018), and is able to flip fluorescently labelled lipid II in
liposomes and E. coli vesicles (Mohammadi et al. 2011, 2014). However in vivo
experiments do not support its lipid II flippase activity, showing no accumulation of
lipid II at the inner leaflet of the cytoplasmic membrane in FtsW-depleted cells (Lara
et al. 2005; Sham et al. 2014).
MurJ is an integral membrane protein that belongs to the MOP
(multidrug/oligosaccharidyl-lipid/polysaccharide) exporter superfamily of pro-
teins. It was identified by bioinformatic analysis and isolated from a temperature
sensitive mutant (Ruiz 2008; Inoue et al. 2008). It contains 14 transmembrane
segments that adopt a V-shaped structure with a central solvent-exposed cavity
(Kuk et al. 2017; Zheng et al. 2018). Several residues are necessary for the correct
functionality of the protein, including some of the charged residues located in the
central cavity (Butler et al. 2013, 2014; Zheng et al. 2018). The inactivation of MurJ
causes cell shape defects and lysis (Ruiz 2008; Inoue et al. 2008) and, supporting
its flippase activity, an accumulation of lipid II intermediates in the cell (Sham et al.
2014; Qiao et al. 2017; Chamakura et al. 2017; Rubino et al. 2018). Its essential role
can be substituted by other transporters from different species as B. subtilis (MurJ
and Amj) (Meeske et al. 2015) or Streptococcus pyogenes (YtgP) (Ruiz 2009), or
by transporters with a more relaxed substrate specificity (Elhenawy et al. 2016;
Sham et al. 2018). In vitro evidences do not support its lipid II flippase activity in
proteoliposomes and E. coli vesicles (Mohammadi et al. 2011) and show contrary
results on lipid II binding (Leclercq et al. 2017; Bolla et al. 2018).

Lipid Carrier

Undecaprenyl phosphate is the essential and unique lipid carrier used for the syn-
thesis of the bacterial extracellular cell wall polymers as PG, teichoic acids and
O-antigen. It is produced by dephosphorylation of the undecaprenyl pyrophosphate,
5 Peptidoglycan 139

either from de novo synthesis or from the recycling pathway, by membrane embed-
ded phosphatases (BacA or PAP2-type phosphatases) (Manat et al. 2014). In the de
novo synthesis pathway, UppS uses farnesyl pyrophosphate and eight isopentenyl
pyrophosphate molecules to produce undecaprenyl pyrophosphate. In the recycling
pathway, the undecaprenyl pyrophosphate is released after the glycosyltransferase
reaction performed by the PG synthases to incorporate the disaccharide pentapeptide
moiety of the lipid II substrate into the PG glycan chains. In both cases undecaprenyl
phosphate is generated in the outer leaflet of the cytoplasmic membrane, as the active
sites of BacA and the PAP2-type phosphatases PgpB, YbjG and LpxT are oriented
towards the periplasm (Touze et al. 2008; Fan et al. 2014; Tatar et al. 2007; Manat
et al. 2015; El Ghachi et al. 2018; Workman et al. 2018). The lipid carrier has to be
flipped across the membrane, as the synthesis of the PG precursor takes place in the
inner leaflet, but the required protein has not yet been identified.

Peptidoglycan Synthesis

The synthesis of new PG into the sacculus comprises the polymerization of gly-
can chains by glycosyltransferases (GTases) and their incorporation to new or
pre-existing chains through cross-linkage of the stem peptides by transpeptidases
(TPases). There are different TPase reactions depending on the donor muropeptide
(Fig. 5.4). The most abundant reaction occurs when a pentapeptide donor cross-links
the d-Ala at position four with the mDAP at position three of the acceptor muropep-
tide (dd-TPase reaction, 4-3 cross-link). The dd-TPases are named penicillin binding
proteins (PBPs), as β-lactams covalently bind to the catalytic site serine residue, block
the access to the donor muropeptide and therefore inhibit the enzymatic reaction (see
Sauvage et al. 2008 for a detailed description of PBPs). In a less frequent reaction two
muropeptides are cross-linked by their mDAP residues at position three (ld-TPase
reaction, 3-3 cross-link).
The best characterised PG GTases belong to the GT51 family. They contain a
conserved sequence with five motifs, a structure mainly composed of α-helices and
a catalytic glutamate residue. The phosphoglycolipid antibiotic moenomycin binds
to the active site, competing with the newly synthesised glycan chain for the donor
site. E. coli contains four GT51 GTases: a monofunctional GTase (MtgA) and three
bifunctional class A PBPs (encoding also dd-TPase domain) known as PBP1A,
PBP1B and PBP1C (Sauvage et al. 2008). PBP1A and PBP1B are the most important
class A PBPs and both contain a non-catalytic domain (ODD in PBP1A and UB2H
in PBP1B) involved in the regulation of the GTase and TPase activities (Typas et al.
2010). Either PBP1A or PBP1B is required for cell viability (Yousif et al. 1985;
Kato et al. 1985). PBP1A is preferentially involved in the synthesis of PG during
cell elongation and PBP1B during septation. Both proteins are functionally semi-
redundant, meaning that the cell can compensate the absence of each other although
not under all tested conditions (Garcia del Portillo and de Pedro 1990, 1991; Denome
et al. 1999; Pepper et al. 2006; Ranjit and Young 2013). Several in vitro studies have
140 M. Pazos and K. Peters

characterised the activities of both proteins either alone (Barrett et al. 2004; Bertsche
et al. 2005; Born et al. 2006) or in presence of different interacting partners, as in
case of PBP1A and PBP2 (Banzhaf et al. 2012) or LpoA (Typas et al. 2010; Lupoli
et al. 2014), and PBP1B and FtsN (Müller et al. 2007) or LpoB (Typas et al. 2010;
Egan et al. 2014, 2018; Lupoli et al. 2014). The crystal structure of E. coli PBP1B
has been described in complex with different antibiotics (Sung et al. 2009; King et al.
2017). Crystal structures of other bifunctional PBPs from different bacteria are also
available (Jeong et al. 2013; Lovering et al. 2007; Yuan et al. 2007).
E. coli has two essential monofunctional dd-TPases or class B PBPs, PBP2 and
PBP3. Whereas PBP2 is required for cell elongation and its inactivation produces
a spherical phenotype, PBP3 is needed for septation and its inactivation generates
long filamented cells. In both cases cells eventually lyse. Crystal structures of E. coli
PBP3 and H. pylori PBP2 are solved (Sauvage et al. 2014; Contreras-Martel et al.
2017) (see section “Regulation of Peptidoglycan Growth” for further details on the
elongation and septation complexes).
Recently the SEDS family protein RodA has been described as a GTase. It does not
belong to the known GT51 family and is not inhibited by the antibiotic moenomycin.
The crystal structure of RodA from Thermus thermophilus has been solved (Meeske
et al. 2016; Cho et al. 2016; Emami et al. 2017; Sjodt et al. 2018). Future work will
show if the SEDS protein FtsW, essential for cell division, is also a GTase (Taguchi
et al. 2018).

LD -Transpeptidases

In E. coli only 5–15% of the PG cross-links are 3-3 (Glauner et al. 1988).
This transpeptidase reaction (ld-TPase) is catalysed by the non-essential ld-
transpeptidases (LDTs) (Magnet et al. 2008) that use a disaccharide-tetrapeptide
as acyl donor and form cross-links between two mDAP residues of adjacent stem
peptides. LDTs contain a catalytic cysteine residue (Mainardi et al. 2005) and are
insensitive to β-lactams, with the exception of carbapenems that also target PBPs
(Mainardi et al. 2007). Even though the 3-3 are the minor cross-links in the PG of
most bacteria, the LDTs are promising targets for new antimicrobial drugs against
important pathogens, and therefore they will be discussed in more detail.
Ldtfm , from an ampicillin-resistant Enterococcus faecium strain, was the first dis-
covered LDT with documented ld-TPase activity. In the presence of the antibiotic
E. faecium cells produce exclusively 3-3 cross-links (Mainardi et al. 2000, 2002,
2005). This alternative cross-linkage enables the bypass of PBPs and confers resis-
tance to β-lactam antibiotics, although a dd-CPase reaction is needed to produce
the tetrapeptides that can be used by the LDTs. Since then, Ldt homologues with a
catalytic YkuD like domain (PFAM 03744) have been found and described among
pathogen and non-pathogen Gram-positive and Gram-negative bacteria.
E. coli has six periplasmic LDTs (LdtA-F) with different functions. LdtC and
LdtE contain an N-terminal PG-binding lysin motif (LysM) domain, which is found
5 Peptidoglycan 141

in many cell wall hydrolases (Buist et al. 2008), suggesting that these enzymes may
be active when bound to PG. LdtA, LdtB and LdtC (ErfK, YbiS and YcfS) catalyse
the covalent attachment of the outer membrane lipoprotein Lpp (Braun’s lipoprotein)
to PG, stabilizing the cell envelope (Magnet et al. 2007). These enzymes link the
ε-amino group of the C-terminal lysine of Lpp to the α-carboxyl group of the mDAP
residue in the stem peptide of the PG (Fig. 5.4). Lpp is a small α-helical protein
that with 1 million molecules per cell is the most abundant protein in E. coli (Li
et al. 2014). Lpp is anchored to the outer membrane by a lipid moiety composed of
acyl chains attached to the cysteine residue at its N-terminus. Under normal growth
conditions, about one third of the Lpp molecules are linked to PG (Inouye et al.
1972). LdtD and LdtE (YcbB and YnhG) form 3-3 cross-links (Fig. 5.4) (Magnet
et al. 2008). The expression of ldtD is regulated by the Cpx pathway, which mediates
adaption to cell envelope stress (Bernal-Cabas et al. 2015). In response to Cpx-
activated conditions the production of LdtD is upregulated resulting in increased 3-3
cross-linkage, suggesting a role for LdtD under cellular stress conditions (Bernal-
Cabas et al. 2015; Delhaye et al. 2016). Furthermore, LdtD is twice the size of the
other E. coli LDTs and shares lower sequence similarity with them (Sanders and
Pavelka 2013). LdtF (YafK) is an Ldt homolog protein with yet unknown enzymic
function, although a role in biofilm formation has been suggested in pathogenic E.
coli (Sheikh et al. 2001). The deletion of all six ldt genes leads to a lack of 3-3 cross-
links and a reduction in Lpp-PG attachment, indicating the non-essentiality of these
enzymes (Peters et al. 2018). Remarkably, the ampicillin resistant E. coli strain M1
shows elevated levels of the alarmone (p)ppGpp and is able to replace the dd-TPase
activity of the PBPs by production of the β-lactam-insensitive LdtD (Hugonnet et al.
2016). This finding shows a new mode of PG polymerization in E. coli requiring the
GTase activity of PBP1B, the dd-CPase activity of PBP5 and the ld-TPase activity
of LdtD (Hugonnet et al. 2016).
LDTs play important roles in different pathogenic bacteria. The formation of 3-3
cross-links is essential for the virulence of the pathogen Salmonella enterica serovar
Typhi. In this organism the secretion of the typhoid toxin depends on the activity of the
N-acetyl-β-d-muramidase TtsA, which hydrolyses the 3-3 cross-links synthesised by
the LDT YcbB (Geiger et al. 2018). LDTs from E. coli and E. faecium are inhibited by
copper ions at sub-millimolar concentrations, likely through the binding to the thiol
group of the catalytic cysteine residue and its activity inhibition. The resulting lack
of 3-3 cross-links and the decrease in the Lpp-PG attachment impair the robustness
of the cell envelope (Peters et al. 2018). Additionally this inhibition counteracts the
LDT-mediated β-lactam resistance of E. coli and E. faecium strains (Peters et al.
2018). The PG of C. difficile and M. tuberculosis is predominantly 3-3 cross-linked,
with several LDTs encoded in each organism, suggesting an important role for these
enzymes (Peltier et al. 2011; Lavollay et al. 2008, 2011; Sutterlin et al. 2018). The
deletion of two of the three LDTs in C. difficile leads to a significant decrease in
3-3 cross-links, partially compensated by an increase in 4-3 cross-links by PBPs,
and a less cross-linked PG (Peltier et al. 2011). Attempts to inactivate the third
LDT have been unsuccessful. Out of the five LDT homologues in M. tuberculosis
(LdtMt1 –LdtMt5 ), LdtMt2 is the dominant one, as its inactivation causes altered colony
142 M. Pazos and K. Peters

morphology, loss of virulence and increased susceptibility to amoxicillin (Gupta

et al. 2010). The ld-TPase activity of LdtMt2 requires an amidated mDAP in the stem
peptide, catalysed by the amidotransferase AsnB (Ngadjeua et al. 2018). LdtMt1 ,
LdtMt3, LdtMt4 and LdtMt5 are all of similar size and share an amino acid sequence
identity of 28–36% with LdtMt2 . The crystal structure of LdtMt5 shows structural
and functional differences to the other LDTs, since it contains a different catalytic
site and a proline rich C-terminal domain. Cells lacking a functional LdtMt5 displays
aberrant growth and increased sensitivity to osmotic shock and certain carbapenems,
indicating its critical role in maintaining the cell wall integrity (Brammer et al. 2015).

Peptidoglycan Hydrolysis and Remodelling

E. coli cells coordinate the synthesis of PG with the cleavage of the pre-existing
PG material by dedicated hydrolases to incorporate new PG into the sacculi. PG
hydrolases are also involved in autolysis, maturation, turnover and recycling of PG,
showing substrate and PG-linkage specificity. According to the cleavage site they
can be classified as N-acetylmuramoyl-l-Alanine amidases (amidases), carboxypep-
tidases, endopeptidases and lytic transglycosylases (van Heijenoort 2011; Vollmer
et al. 2008b) (Fig. 5.4). Further information about the PG-recycling and β-lactamase
induction pathways can be found in different reviews (Park and Uehara 2008; Juan
et al. 2017; Dik et al. 2018).

N-Acetylmuramoyl-L-Alanine Amidases

Amidases specifically cleave the amide bond linking the l-Ala residue of the stem
peptide to the MurNAc subunit of the muropeptide. E. coli contains five different ami-
dases, which are grouped in two superfamilies based on the amino acid sequence sim-
ilarity: AmiA, AmiB and AmiC (pfam: amidase 3) and AmpD and AmiD (pfam: ami-
dase 2). AmiA, AmiB and AmiC exhibit specificity for MurNAc substrates, AmpD
for 1,6-anhydro-MurNAc, and AmiD cleave both MurNAc and 1,6-anhydro-MurNAc
substrates.
AmiA, AmiB and AmiC are exported to the periplasm via the Tat system (AmiA
and AmiC) and through the Sec translocon (AmiB) (Ize et al. 2003; Bernhardt and
de Boer 2003). Although AmiB and AmiC are recruited to the septal position during
cell division, and AmiA remains dispersed throughout the cell periplasm (Bernhardt
and de Boer 2003; Peters et al. 2011), all of them perform redundant reactions that
compensate the absence of the others. The inactivation of all three amidases results
in cell chains, in which daughter cells are not separated from each other. Single
and double mutants have milder phenotypes, showing shorter cell chains (Heidrich
et al. 2001). Genetic evidences indicate a partially redundant role of the Rcs and
Cpx stress responses in supporting the growth and viability of the triple mutant
5 Peptidoglycan 143

(Yakhnina et al. 2015). The activity of the amidases is dependent on the presence
of two LytM-domain containing proteins, EnvC and NlpD, as the inactivation of
both genes shows the same cell chain phenotype than the triple amiABC mutant.
Their LytM-domains do not contain the residues required for the coordination of
the catalytic Zn2+ ion and lack the ion itself (Uehara et al. 2009, 2010; Peters et al.
2013), typical from the metallo-peptidase family M23 (Pfam: Peptidase_M23). EnvC
activates AmiA and AmiB in the outer-leaflet of the cytoplasmic membrane, and the
outer membrane-anchored lipoprotein NlpD activates AmiC (Uehara et al. 2010). The
structure of AmiC (Rocaboy et al. 2013) and studies on AmiB show the presence
of a conserved α-helix blocking the access to the catalytic site (Yang et al. 2012).
Conformational changes induced by the interaction with the activator enable the open
access to the active site (Yang et al. 2012) (see section “Regulation of Peptidoglycan
Growth” for further details on EnvC and NlpD functioning).
AmpD is a cytoplasmic Zn2+ -dependent amidase required for the PG-recycling
process during bacterial cell growth. The specificity for 1,6-anhydro-MurNAc con-
taining substrates (Jacobs et al. 1995) prevents AmpD of interfering with the synthesis
of cell wall precursors. The inactivation of AmpD leads to the accumulation of its
substrate 1,6-anhydro-MurNAc-l-Ala-d-iGlu-mDAP (Jacobs et al. 1994).
AmiD is an outer membrane-anchored lipoprotein with Zn2+ -dependent amidase
activity. It cleaves the amide bond of intact PG or soluble fragments containing
tri-, tetra- or pentapeptides, regardless of the presence or absence of 1,6-anhydro-
MurNAc (Uehara and Park 2007; Pennartz et al. 2009). AmpD and AmiD are the
only proteins with specificity for 1,6-anhydro-MurNAc substrates (Uehara and Park
2007). A reaction mechanism has been proposed based on the crystal structures of
AmiD, alone and in complex with either the 1,6-anhydro-MurNAc-tripeptide or the
tripeptide (Kerff et al. 2010). AmiD is not required for cell separation (Uehara and
Park 2007).

Lytic Transglycosylases

Lytic transglycosylases (LTs) are periplasmic N-acetylmuramidases that catalyse the

cleavage of the β-1,4 glycosidic bond between MurNAc and GlcNAc, generating a 1,6-
anhydro bond in the MurNAc. The resultant muropeptides containing 1,6-anhydro-
MurNAc are transported from periplasm to cytoplasm, via the transmembrane protein
AmpG. The cleavage reaction performed by LTs can take place at the end of the glycan
chains (exolytic activity) or within them (endolytic activity). E. coli contains eight
LTs (MltA-MltG and Slt). LTs are functionally redundant and the absence of six of
them (MltA-E and Slt) does not affect cell viability although generates cell chain
phenotype (Heidrich et al. 2002). RlpA, recently described as an interacting partner
of the cell division protein FtsK in E. coli (Berezuk et al. 2018), is considered a LT
based on its activity in P. aeruginosa (Jorgenson et al. 2014). Slt is a periplasmic
LT, MltA-F are outer membrane-anchored lipoproteins, and MltG is anchored to
the outer-leaflet of the cytoplasmic membrane. LTs are classified based on their
144 M. Pazos and K. Peters

domain architecture, belonging to family 1 (Slt, family 1A; MltC, family 1B; MltE,
family 1C; MltD, family 1D; MltF, family 1E), family 2 (MltA), family 3 (MltB)
and YceG-family (MltG). Slt is described in more detail in the following lines, and
further information about LTs can be found in different reviews (van Heijenoort
2011; Dik et al. 2017; Alcorlo et al. 2017).
Slt has a main exolytic activity performed on non cross-linked muropeptides
containing stem peptides, although a low endolytic activity has been described (Lee
et al. 2013, 2018). It has a “doughnut-shaped” structure, as shown in complex with the
specific inhibitor bulgecin or the 1,6-anhydro-muropeptide (Thunnissen et al. 1994,
1995; van Asselt et al. 1999). Recent works with P. aeruginosa Slt and Neisseria
meningitidis LtgA suggest that conformational rearrangements on the active site take
place during the hydrolytic reactions (Lee et al. 2018; Williams et al. 2018). In E. coli
Slt interacts with the PG synthases PBP1B, PBP1C, PBP2 and PBP3, and with the
PG endopeptidase PBP7 (Romeis and Höltje 1994b; von Rechenberg et al. 1996).
In the absence of slt, or protein inactivation by addition of bulgecin, cells do not
show different phenotype or alter the PG composition but modified sensitivity to
certain β-lactams. Inactivation of PBP3 in slt mutant cells produces bulges and rapid
lysis (Templin et al. 1992), instead of cell filamentation and lysis as in WT cells
(Spratt 1975). The sensitivity to the PBP2-specific inhibitor mecillinam in slt cells
depends on the FtsZ protein levels, being more sensitive at WT levels (Templin et al.
1992) and more resistant at high levels (Vinella et al. 1993). The hypersensitivity
to mecillinam has been described as a consequence of the accumulation of newly
synthesised PG glycan chains and its misincorporation into the sacculus by LDTs. In
WT cells the non cross-linked new PG glycan chains are removed by Slt, contributing
to the increased PG turnover by the β-lactam mecillinam stress response (Cho et al.
2014).

Endopeptidases

Endopeptidases (EPases) catalyse the cleavage of the amide bonds between amino
acids from different stem peptides. According to the substrate specificity, they are
classified as dd-endopeptidases (dd-EPases) and ld-endopeptidases (ld-EPases).
dd-EPases cleave the bond formed by the 4-3 cross-links, in E. coli between d-Ala
at position four from one stem peptide and mDAP at position three from a different
stem peptide. ld-EPases show specificity for the bond formed by the 3-3 cross-links,
in E. coli between the mDAP residues at position three of different stem peptides.
E. coli contain seven proteins encoding for dd-EPase activity, three of them are
sensitive to β-lactams (the class C PBPs PBP4, PBP7 and AmpH) and four are
insensitive to β-lactams (MepA, MepH, MepM and MepS). The absence of each
single protein does not change the cell phenotype, indicating functional redundancy
between them. Recently it has been shown that the overproduction of MepA, MepM,
PBP7 or MepS confers resistance to mecillinam through the stimulation of the PG
synthesis by PBP1B (Lai et al. 2017).
5 Peptidoglycan 145

PBP4 is a periplasmic protein that shows partial association with the cytoplas-
mic membrane (Korat et al. 1991; Jacoby and Young 1988; Leidenix et al. 1989). In
addition to the dd-EPase activity, PBP4 also shows dd-carboxypeptidase (dd-CPase)
activity (see section “Carboxypeptidases”), using either soluble muropeptides or iso-
lated PG as substrate (Korat et al. 1991; Li et al. 2004; Clarke et al. 2009). The crystal
structure of the protein alone and bound to different antibiotics shows three different
domains and the formation of a soluble dimer (Kishida et al. 2006). Residues close
to the catalytic serine encoded in the domain I are required for the accommodation
of the mDAP residue of the stem peptide substrate (Clarke et al. 2009). In vivo stud-
ies based on the combination of gene deletions suggest a role for PBP4 in different
cell processes including cell morphology, separation of daughter cells and biofilm
formation (Meberg et al. 2004; Priyadarshini et al. 2006; Gallant et al. 2005).
The periplasmic protein PBP7 shows a loose association with the cytoplasmic
membrane, which can be abolished by high salt treatment (Romeis and Höltje 1994a).
A C-terminal truncated variant, identified as PBP8, led to confusion in the past
(Henderson et al. 1994). PBP7 shows dd-EPase activity when isolated PG is used
as substrate, but not in case of purified dimeric muropeptides (Romeis and Höltje
1994a). It interacts with Slt, which enhances its LTase activity (Romeis and Höltje
1994b). The inactivation or overproduction of PBP7 does not generate any cellular
defect (Henderson et al. 1995). Combined inactivation of PBP7, PBP4 and PBP5
enhances the morphological defects caused by the absence of PBP5 (main dd-CPase
in E. coli) (Meberg et al. 2004), and the additional inactivation of AmpH activates
the Rcs phosphorelay and Cpx stress responses (Evans et al. 2013).
AmpH is a periplasmic protein associated to the cytoplasmic membrane, despite
the lack of any membrane anchoring domain. In contrast to PBP4 and PBP7, high
salt treatment does not dissociate the protein from the membrane fraction, requiring
detergent for that purpose. AmpH displays dd-EPase and dd-CPase activities using
both intact sacculi and isolated dimeric muropeptides as substrates, and weak β-
lactamase activity (Gonzalez-Leiza et al. 2011). Inactivation of AmpH does not
cause any defect in the cell, unless it is combined with other mutations producing
morphological changes (Henderson et al. 1997) or stress response activation (Evans
et al. 2013). A role in PG recycling and remodelling has been suggested based on
the wide range of substrates used by AmpH.
MepA is a periplasmic protein belonging to the LAS metallopeptidase family
(lysostaphin-type enzymes, d-alanyl-d-alanine CPase, and sonic hedgehog protein),
which is characterised for the presence of a Zn2+ -binding site in the catalytic domain.
It shows dd-EPase activity on intact sacculi and isolated muropeptides (Keck and
Schwarz 1979; Firczuk and Bochtler 2007) and ld-EPase activity on intact sac-
culi (Engel et al. 1992). The conserved metal ligands are required for the correct
folding and catalytic activity of the protein (Firczuk and Bochtler 2007), which
can be inhibited by DNA, lipoteichoic acid and metal-chelating agents (Tomioka
and Matsuhashi 1978; Keck and Schwarz 1979). Inactivation or overproduction
of MepA does not cause any change in the PG composition or cell phenotype
146 M. Pazos and K. Peters

(Iida et al. 1983; Keck et al. 1990). The combined inactivation of PBP4, PBP7 and
MepA does not cause any defect in the cell, but enhances the cell chain phenotype
due to the absence of amidases and LTs (Heidrich et al. 2002).
MepH, MepM and MepS, in contrast to the previously described EPases, are
redundantly essential as their absence cause cell lysis (Singh et al. 2012). MepH
is a periplasmic protein of the NlpC/P60 peptidase superfamily with the conserved
Cys-His-His catalytic triad (Aramini et al. 2008). MepM is a bitopic cytoplasmic
membrane protein of the metallopeptidase family M23, containing the characteris-
tic LytM domain and the catalytic Zn2+ -binding site. Both proteins show dd-EPase
activity against isolated muropeptides and, partially, intact sacculi (Singh et al. 2012).
MepS is an outer membrane-anchored lipoprotein belonging to the NlpC/P60 pep-
tidase superfamily, which shows dd-EPase and weak ld-CPase activity against iso-
lated muropeptides but not against intact sacculi (Singh et al. 2012). MepS protein
levels are higher during exponential phase, being modulated by the proteolytic sys-
tem NlpI-Prc (Singh et al. 2015). NlpI is an outer membrane-anchored lipoprotein
containing tetratricopeptide-repeats that facilitates the interaction between MepS
and the periplasmic protease Prc (Singh et al. 2015; Su et al. 2017). A reaction
mechanism for the degradation of MepS has been proposed based on the crystal
structure and biophysical and mutational analyses of the NlpI-Prc complex (Su et al.
2017). The impairment of this proteolytic system alters the morphology of the cells
at high-salinity growth conditions (Kerr et al. 2014), changes the PG dynamics in
stationary phase, decreases the Lpp-PG attachment and increases the formation of
outer membrane vesicles (Schwechheimer et al. 2015).

Carboxypeptidases

Carboxypeptidases (CPases) remove the terminal residues from stem peptides. E. coli
encodes for six non-essential dd-CPases (PBP4, PBP4b, PBP5, PBP6, PBP6b and
AmpH) that remove the d-Ala residue from pentapeptides, and one cytoplasmic ld-
CPase (LdcA) that removes the d-Ala residue from tetrapeptides. PBP4 and AmpH
also show dd-EPase activity and were described in the section “Endopeptidases”.
PBP4b is a non-essential protein, showing sequence homology to AmpH and
weak dd-CPase activity on artificial substrate (Vega and Ayala 2006) but not on
natural isolated muropeptides (J. Ayala, unpublished results mentioned in Sauvage
et al. 2008).
PBP5 is one of the best characterised and most abundant PBP in the cell. It is a
membrane protein anchored to the outer leaflet of the cytoplasmic membrane by its
C-terminal amphipathic helix. It shows sequence similarities with PBP4, PBP6 and
certain β-lactamases, and has been recently described to dimerise (Meiresonne et al.
2017). The crystal structure of a soluble form of PBP5 shows the presence of an
N-terminal domain (domain I) including the catalytic site and a C-terminal domain
(domain II) containing a hydrophobic surface and the mentioned membrane-binding
helix. The domain I contains a -loop-like region, similar to the one found in certain
5 Peptidoglycan 147

β-lactamases, which is essential for keeping the β-lactam resistance and the cell
shape (Dutta et al. 2015; Kar et al. 2018). The function of the domain II is not well
known, although it seems to be required for the stabilization and full function of the
protein. It shows dd-CPase activity against soluble muropeptides and cross-linked
and uncross-linked PG. Although non-essential, the loss of PBP5 activity causes cell
shape defects and an increase in the pentapeptide content in the PG. These shape
defects are enhanced and lead to the formation of branches when combined with
the absence of other PBPs. Variations in the level of the cell division protein FtsZ
and in the formation of the FtsZ-ring structure also enhance the cell shape defects
(Varma and Young 2004; Varma et al. 2007; Potluri et al. 2012a). PBP5 localizes to
areas of ongoing PG synthesis, meaning along the lateral cell wall and at division
sites before septation starts (preseptal sites) (Potluri et al. 2010). The absence of the
membrane binding helix does not affect the catalytic activity of the protein, although
the resultant soluble PBP5 does not localize at preseptal sites or restore the phenotype
in mutant cells, as also observed for inactive PBP5 variants (Nelson and Young 2000;
Nelson et al. 2002; Potluri et al. 2010). Overproduction of PBP5 causes non-viable
spherical cells (Markiewicz et al. 1982), suggesting a role for PBP5 in regulation
of the pentapeptide subunits required for the formation of cross-links by dd-TPase.
The combination of the dd-CPase activity of PBP5, the ld-TPase activity of LdtD
and the GTase activity of PBP1B has been described to bypass the dd-TPase activity
of the PBPs (Hugonnet et al. 2016).
PBP6 and PBP6b have strong sequence and structural similarities with PBP5,
including the C-terminal membrane anchor that can substitute the one from PBP5
(Nelson et al. 2002). PBP6 shows weaker activity than PBP5 against similar substrates
(Amanuma and Strominger 1980). In the absence of PBP5, the main dd-CPase
activity is performed by PBP4 at neutral pH, and by PBP6b at low pH. The role of
PBP6b at low pH is also supported by its ability to maintain the WT cell shape in
the absence of PBP5 (Peters et al. 2016).

Regulation of Peptidoglycan Growth

During the E. coli cell cycle the PG sacculus is enlarged and split through the enzy-
matic activities encoded in the protein complexes named elongasome and divisome,
which facilitate cell elongation and cell division, respectively. Both, synthetic and
hydrolytic activities are required for the incorporation of the new PG material, as pro-
posed in the 3-for-1 growth model (Höltje 1998). The control of these activities, from
the cytoplasm by the cytoskeletal proteins and from the periplasmic side by outer
membrane and periplasmic proteins, ensures the precise and coordinated synthesis
of new PG. The composition and functioning of both elongasome and divisome are
largely unknown. However, the protein interactions described in the literature sup-
port the formation of these multienzymatic complexes, although it is unlikely that all
the interactions take place at the same time (Fig. 5.5) (Typas et al. 2011; Egan and
Vollmer 2013). The presence of large sets of seemingly redundant enzymes, under
148 M. Pazos and K. Peters

Amidases
LTs
EPases
LpoA LpoB

RodA PBP1A PBP1B PBP3 FtsW Tol-Pal

PBP2
PBP1A / PBP1B FtsN FtsQLB FtsK FtsEX
RodZ MreCD
ZipA / FtsA*-FtsN FtsA ZipA

MreB FtsZ FtsZ

Elongasome Preseptal PG Divisome

Fig. 5.5 Components of the multienzymatic complexes suggested for new PG incorporation during
cell growth and division in E. coli. The hydrolytic enzymes (amidases, LTs and EPases) are not
specified, as their coordination with the synthetic activities remains largely unknown in each case.
LTs, lytic transglycosylases; EPases, endopeptidases

laboratory standard conditions, may be required to ensure a robust peptidoglycan

growth under the different environmental conditions (Pazos et al. 2017).

Regulation of Peptidoglycan Synthesis by the Cytoskeletal

Proteins

The elongasome is required for the elongation and shape maintenance of rod-shape
cells. It contains the synthetic activities of RodA (GTase), PBP2 (TPase) and PBP1A
(GTase and TPase). The activity of the elongasome is regulated by the cytoskeletal
protein MreB and the membrane-associated MreC, MreD and RodZ proteins. MreB
is an actin homolog that polymerizes in an ATP-dependent manner into dynamic
double antiparallel filaments (van den Ent et al. 2014), which localize on the inner-
leaflet of the cytoplasmic membrane in a curvature-dependent way and rotate along
the short cell axis as patches (Garner et al. 2011). MreB binds directly to the mem-
brane through its N-terminal amphiphatic helix (Salje et al. 2011). The disruption
of these filaments has an impact on the cell shape, including the eventual spheri-
cal phenotype obtained after inactivation of MreB (Kruse et al. 2005). The MreB
inhibitor A22 competes for the ATP binding pocket of MreB, which is not able to
polymerize and remains homogeneously localized in the cytoplasm. The presence of
A22-blocked MreB monomers reduces the characteristic growth heterogeneity and
generates cell wall growth at the poles (Ursell et al. 2014), which were previously
suggested to be inert (de Pedro et al. 1997). Cell poles are enriched in cardiolipin
and phosphatidylglycerol, which prevent the localization of MreB filaments but not
of MreB monomers (Kawazura et al. 2017). The fact that MreB motion requires an
5 Peptidoglycan 149

active elongasome, and the elongasome can function in the absence of an active MreB,
suggests a passive role of MreB in guiding the elongasome (van Teeffelen et al. 2011;
Ursell et al. 2014). Single particle tracking data showed that these protein complexes
are very dynamic, as observed by the different motion of PBP2 and MreB (Lee et al.
2014), the presence of MreB-like slower subpopulations of RodA and PBP2 (Cho
et al. 2016) or the fast and diffusive behaviour of the class A PBPs (Cho et al. 2016;
Lee et al. 2016), which can be modified either by specific inhibitors or activators
(Lee et al. 2016). MreB localization requires a functional Sec-translocon system for
the correct insertion of RodZ into the membrane (Govindarajan and Amster-Choder
2017; Rawat et al. 2015). RodZ is required for the attachment of MreB filaments to
the cytoplasmic membrane, modulating their localization in the cell (Colavin et al.
2018; Bratton et al. 2018). Further information about MreB and associated proteins
can be found in reviews (Errington 2015; Shi et al. 2018).
Despite the essentiality of MreB and the elongasome, the overproduction of the
tubulin homolog FtsZ is able to restore cell viability, but not rod-shape, upon deletion
of any of them (Bendezu and de Boer 2008). Although the mechanism of this FtsZ-
mediated rescue is not clear, it is known that both MreB and FtsZ interact directly at
the cell division site and that this interaction is essential for cell division but not for
rod-shape maintenance. An impaired interaction is lethal for the cell, which shows
neither preseptal nor septal PG synthesis (Fenton and Gerdes 2013). Preseptal PG is
synthesised at the future division site before any constriction is visible, in a PBP3-
independent manner. It is considered as a transition stage between cell elongation and
cell division, or an early cell division stage (Nanninga 1991; de Pedro et al. 1997).
So far, in E. coli only the cell division proteins FtsZ and ZipA, and either PBP1A or
PBP1B are described as essential proteins for preseptal PG synthesis. Several proteins
from the elongasome or divisome are not required, e.g. RodA, FtsA, FtsEX, FtsK
or FtsQ (Potluri et al. 2012b). ZipA links the cytosolic Z-ring and the PG synthases
PBP1A or PBP1B, an essential role that can be compensated by the mutant FtsA*
presumably bound to FtsN (Fig. 5.5) (Pazos et al. 2018). In the absence of both ZipA
and FtsN, the cells are not viable and do not synthesise preseptal PG. Further work
is required to study the essentiality of the preseptal PG synthesis for the correct cell
septation.
The divisome is involved in the septation and separation of the two daughter
cells. Several proteins are required for the correct assembly and functioning of the
divisome and many described interactions support the hypothesis of a multiprotein
complex (Fig. 5.5) (reviewed in Egan and Vollmer 2013). The synthesis of the septal
PG is carried out by PBP1B (GTase and TPase) and PBP3 (TPase). Controversial
results about the SEDS family protein FtsW propose that either it is a lipid II flippase
or a GTase. Future work will clarify its role. The cell division protein FtsZ is the
scaffold in which the divisome is build up. FtsZ polymerizes into a ring-like structure,
termed the Z-ring. FtsZ polymerization and depolymerization requires the binding
and hydrolysis of GTP, respectively. These filaments are bound to the inner-leaflet of
the cytoplasmic membrane by the membrane-attached proteins FtsA and ZipA. The
GTP hydrolysis generates conformational changes in the FtsZ filaments that adopt
a curved shape (Lu et al. 2000), which has been proposed to supply the constriction
150 M. Pazos and K. Peters

force in liposomes (Osawa et al. 2008; Osawa and Erickson 2013). The cellular
localization of the Z-ring is regulated by the Min system, the nucleoid occlusion,
and the Ter region of the chromosome (reviewed in Schumacher 2017; Wettmann
and Kruse 2018; Bailey et al. 2014a). The placement of the Z-ring at mid-cell is
essential to generate two identical daughter cells after septum synthesis and cleavage.
Septal PG synthesis is guided and driven by the dynamic FtsZ polymers, which move
around the ring structure by treadmilling (Yang et al. 2017; Bisson-Filho et al. 2017).
Remarkably, in the round-shaped bacteria S. aureus the recruitment of the proposed
lipid II flippase MurJ to the divisome speeds up the synthesis of septal PG and makes it
independent of FtsZ-treadmilling (Monteiro et al. 2018). In E. coli, FtsZ is not present
at the division site during the whole process of septation, as it moves to the future cell
division sites before constriction is finished (Soderstrom et al. 2014, 2016). Instead of
that, the cell division proteins FtsN and PBP3, both spatially separated from the FtsZ
ring, remain at division site until septum synthesis is completed (Soderstrom et al.
2016, 2018). PBP3 activity, but not FtsZ, has been shown to be the main regulator of
septum closure (Coltharp et al. 2016), leading to different models about the source of
constrictive force (Xiao and Goley 2016; Schoenemann and Margolin 2017; Holden
2018).

Regulation of Peptidoglycan Synthesis from the Periplasmic

Side

E. coli contains periplasmic proteins that regulate the activities of the PG multienzy-
matic complexes. LpoA and LpoB are two outer membrane-anchored lipoproteins
that activates the synthetic activities of PBP1A and PBP1B, respectively. The pres-
ence of one of these proteins is essential for cell viability, in a similar way to PBP1A
and PBP1B, suggesting that the Lpo’s are required for the proper functioning of the
PBPs (Typas et al. 2010; Paradis-Bleau et al. 2010). The regions involved in the
interaction with their cognate PBPs are identified (Jean et al. 2014; Egan et al. 2014;
King et al. 2014; Sathiyamoorthy et al. 2017). In vitro studies have shown that LpoA
stimulates the TPase activity of PBP1A, and LpoB stimulates both GTase and TPase
activities of PBP1B (Lupoli et al. 2014; Egan et al. 2014, 2018). The identification
of PBP1B mutants able to bypass the in vivo requirement of LpoB support the role
of LpoB in PBP1B activation (Markovski et al. 2016). It has been hypothesised that
the activation by Lpo’s could regulate the PG enlargement depending on the sacculus
pores size, which might be different depending on the cellular growth rate (Typas
et al. 2010).
During cell division several proteins have been shown to regulate and coordinate
the activity of the PG synthases and hydrolases. In the next lines some recent results
are briefly mentioned, including the Tol-Pal complex, FtsN and PG amidases. The
Tol-Pal complex is a well conserved system in Gram negative bacteria and includes
cytoplasmic membrane, periplasmic and outer membrane proteins. These proteins
5 Peptidoglycan 151

interact with each other and, in the presence of proton motive force, maintain the
stability of the outer membrane (Cascales et al. 2000, 2001, 2002). TolA and CpoB
interact directly with PBP1B and regulate its synthetic activities. TolA enhances the
GTase activity of PBP1B, either in presence or absence of LpoB, and CpoB decreases
the TPase activity of PBP1B in presence of LpoB without interfering with the GTase
activation (Gray et al. 2015; Egan et al. 2018). The CpoB effect can be reverted by
TolA (Gray et al. 2015).
FtsN is considered the septation trigger, as it is mainly recruited at late stages of
cell division just before constriction starts. This recruitment is due to the binding
of its periplasmic SPOR (sporulation-related repeat) domain to PG glycan chains
(Ursinus et al. 2004) lacking the stem peptides, suggesting that it follows the action
of amidases (Yahashiri et al. 2015). The cytoplasmic region of FtsN is required
for an earlier recruitment of FtsN to midcell by the interaction with FtsA (Busiek
et al. 2012; Busiek and Margolin 2014; Pichoff et al. 2015). Whereas favouring the
FtsA-FtsN interaction the cell bypasses different impairments in other cell division
proteins (Pichoff et al. 2018), point mutations in FtsA abolish the essentiality of FtsN
(Bernard et al. 2007). The essentiality of FtsN seems to be encoded in a small peptide
from the periplasmic domain (Gerding et al. 2009; Liu et al. 2015), which would
be involved in the eventual activation of septation. FtsN interacts with different cell
division proteins including the PG synthases PBP3 and PBP1B, and the potential PG
synthase FtsW (Di Lallo et al. 2003; Müller et al. 2007; Alexeeva et al. 2010), and
in vitro experiments have shown that FtsN enhances both GTase and TPase activities
of PBP1B (Müller et al. 2007). Due to the non-essentiality of PBP1B, the activation
of PBP3 and/or FtsW is likely to be the final target of FtsN. Genetic evidences have
shown that the conserved cell division proteins FtsQLB are involved in this activation
pathway (Liu et al. 2015; Tsang and Bernhardt 2015).
E. coli contains three PG amidases (AmiA, AmiB and AmiC) required for splitting
the PG sacculi of two daughter cells during septation, although only AmiB and AmiC
are recruited to the division site. Their catalytic activities are regulated by protein
activators: AmiA and AmiB by EnvC, and AmiC by NlpD. The correct localization
of both activators is essential for the temporal and spatial regulation of the amidase
activities. EnvC is localized at preseptal positions (Peters et al. 2011) by the cell
division ATP-binding cassette complex FtsEX, in which FtsE is the cytoplasmic
nucleotide-binding protein and FtsX is the integral membrane component that binds
to EnvC (de Leeuw et al. 1999; Yang et al. 2011). Conformational changes in FtsX,
driven by the ATPase activity of FtsE, are essential for the amidase activation by EnvC
(Yang et al. 2011). NlpD, as EnvC, it is located at the division site before septal PG
is synthesised (Peters et al. 2011). Additional proteins, as the Tol-Pal complex and
YraP, are required for the activation of AmiC by NlpD (Tsang et al. 2017).
152 M. Pazos and K. Peters

Fluorescent D-Amino Acids as Tools

The recent development of new fluorescent d-amino acids (FDAAs), combined with
the continuous improvement of microscopic techniques, enables the visualization of
PG synthesis, remodelling and dynamics in high resolution. These powerful tools are
used to study PG synthesis processes during cell elongation and division in diverse
Gram-positive and Gram-negative bacteria. The FDAAs designed by VanNieuwen-
hze’s and Brun’s labs mimic the acyl acceptor during the PG synthesis reaction and are
thought to be incorporated into the stem peptides through a d-amino-exchange reac-
tion performed by either dd-TPase (Lupoli et al. 2011) or dd-CPase and ld-TPase
activities (Cava et al. 2011; Hsu et al. 2017) (Fig. 5.4). Based on their specificity,
FDAAs efficiently label the active PG synthesis sites with minimal cell toxicities
(Kuru et al. 2012). FDAAs have been successfully used in many PG studies of
diverse species, providing new insights about PG synthesis, recycling and turnover
(for a detailed review about FDAAs, including an updated list of studies using them
(see Radkov et al. 2018)).

Concluding Remarks

In the last years we have gained insight into many different aspects of the bacterial
cell wall, but we are still far from understand this complex cellular structure. The
introduction of new tools and technologies (as FDAAs, high resolution microscopy
or high-throughput genetic screenings), combined with the multidisciplinary con-
tribution to the field, is currently boosting the knowledge of the field. Besides the
new biophysical and structural approaches, the better understanding of the essential
proteins roles might lead to the identification of new antimicrobial drug targets, for
example the LDTs. New antimicrobial drugs are required to reduce the dramatic
spread of multidrug resistant bacteria. In terms of basic knowledge, the formation
and coordination of the elongasome and divisome complexes remain elusive. The
further characterisation of the transition between both elongation and division, and
the following septum formation and cell constriction will aid the comprehension of
this complex but exciting macromolecule, the peptidoglycan.

Acknowledgements The authors thank Professor Waldemar Vollmer at Newcastle University for
critical reading this manuscript.
5 Peptidoglycan 153

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