The biosynthesis of peptidoglycan lipid-linked intermediates

R E V I E W A R T I C L E

Thebiosynthesis of peptidoglycan lipid-linked intermediatesAhmed Bouhss1, Amy E. Trunkfield2, Timothy D.H. Bugg2 & Dominique Mengin-Lecreulx1

1Laboratoire des Enveloppes Bacteriennes et Antibiotiques, Institut de Biochimie et Biophysique Moleculaire et Cellulaire, UMR 8619 CNRS,

Univ Paris-Sud, Orsay, France; and 2Department of Chemistry, University of Warwick, Coventry, UK

Correspondence: Dominique Mengin-

Lecreulx, Laboratoire des Enveloppes

Bacteriennes et Antibiotiques, IBBMC, UMR

8619 CNRS, Universite Paris-Sud, Bat. 430,

91405 Orsay Cedex, France. Tel.: 133 1 69 15

48 41; fax: 133 1 69 85 37 15; e-mail:

[email protected]

Received 10 July 2007; revised 20 September

2007; accepted 24 September 2007.

First published online 11 December 2007.

DOI:10.1111/j.1574-6976.2007.00089.x

Editor: Jacques Coyette

Keywords

peptidoglycan; undecaprenyl phosphate; UppS

synthase; UppP phosphatases; MraY

translocase; MurG transferase.

Abstract

The biosynthesis of bacterial cell wall peptidoglycan is a complex process involving

many different steps taking place in the cytoplasm (synthesis of the nucleotide

precursors) and on the inner and outer sides of the cytoplasmic membrane

(assembly and polymerization of the disaccharide-peptide monomer unit, respec-

tively). This review summarizes the current knowledge on the membrane steps

leading to the formation of the lipid II intermediate, i.e. the substrate of the

polymerization reactions. It makes the point on past and recent data that have

significantly contributed to the understanding of the biosynthesis of undecaprenyl

phosphate, the carrier lipid required for the anchoring of the peptidoglycan

hydrophilic units in the membrane, and to the characterization of the MraY and

MurG enzymes which catalyze the successive transfers of the N-acetylmuramoyl-

peptide and N-acetylglucosamine moieties onto the carrier lipid, respectively.

Enzyme inhibitors and antibacterial compounds interfering with these essential

metabolic steps and interesting targets are presented.

Introduction

Peptidoglycan (murein) is a major heteropolymer of bacter-

ial cell walls that consists in long glycan chains made of

alternating units of N-acetylmuramoyl-peptides (MurNAc-

peptides) and N-acetylglucosamine (GlcNAc) that are cross-

linked together via the short peptide chains (Rogers et al.,

1980; Park, 1996; Vollmer et al., 2008). The main essential

function of this giant cell-sized macromolecule is to protect

cells against the deleterious effects of the internal osmotic

pressure. It also contributes to the maintenance of the

characteristic cell shape and serves as a platform for the

anchoring of other cell envelope components including

proteins (Braun & Sieglin, 1970; Marraffini et al., 2006;

Dramsi et al., 2008) and polysaccharides (Neuhaus &

Baddiley, 2003).

The biosynthesis of peptidoglycan is a complex process

involving many different cytoplasmic and membranes steps

(van Heijenoort, 2001b). The first stage consists in the

formation of the soluble nucleotide precursors, from UDP-

GlcNAc to UDP-MurNAc-pentapeptide. In particular, the

synthesis of the peptide moiety is performed by a series of

enzymes designated as the Mur ligases (MurC, MurD, MurE

and MurF) which are responsible for the respective additions

of L-alanine, D-glutamic acid, meso-diaminopimelic acid

(A2pm) or L-lysine, and D-alanyl-D-alanine to UDP-MurNAc

(Barreteau et al., 2008). As reported by Schleifer & Kandler

(1972) and exemplified in the accompanying reviews

(Barreteau et al., 2008; Vollmer et al., 2008), the structure of

this peptide and the substrate specificity of these enzymes

exhibit some variations in the bacterial world. The membrane

steps then begin with the transfer of the phospho-MurNAc-

pentapeptide moiety from the cytoplasmic precursor to the

membrane acceptor undecaprenyl phosphate (C55-P), a

reaction catalyzed by the transferase MraY (also named

translocase) yielding undecaprenyl-pyrophosphoryl-Mur-

NAc-pentapeptide (lipid I) (Fig. 1). Thereafter, the transferase

MurG catalyzes the transfer of the GlcNAc moiety from UDP-

GlcNAc to lipid I yielding undecaprenyl-pyrophosphoryl-

MurNAc-(pentapeptide)-GlcNAc (lipid II) which, after its

passage through the membrane by a yet unknown mechan-

ism, will be used as the substrate for the polymerization

reactions (van Heijenoort, 2001a, b) (Fig. 1). The C55-P

carrier lipid plays a central role in these steps and is also

FEMS Microbiol Rev 32 (2008) 208–233c� 2007 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

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required for the synthesis of other cell-wall polymers. This

lipid as well as all the enzymes participating in peptidoglycan

synthesis are essential, specific for the bacterial world and

therefore constitute interesting potential targets to be

exploited for the discovery of new antibacterials.

This review summarizes recent advances in the under-

standing of the membrane steps of peptidoglycan synthesis.

In particular, it will focus on recent significant contributions

to the knowledge of the metabolism of the carrier lipid and

on progress made toward the biochemical and structural

characterization of the MraY and MurG activities and the

search of inhibitors of these enzymes.

Undecaprenyl phosphate metabolism

Undecaprenyl phosphate (C55-P), also referred to as bacto-

prenol, is a key lipid involved in the biosynthesis of

peptidoglycan and a variety of other cell-wall polysaccharide

components such as lipopolysaccharides, the enterobacterial

common antigen, capsule polysaccharides, and teichoic

acids (Wright et al., 1967; Scher et al., 1968; Troy et al.,

1971, 1975; Watkinson et al., 1971; Johnson & Wilson, 1977;

Rohr et al., 1977; Reeves et al., 1996; Rick et al., 1998; van

Heijenoort, 2001b; Raetz & Whitfield, 2002; Neuhaus &

Baddiley, 2003). C55-P-linked saccharides are also used for

N-linked protein glycosylation that occurs in certain pro-

karyotes (Glover et al., 2005; Szymanski & Wren, 2005).

That a single lipid participates in the synthesis of various

wall polymers has been earlier considered as a potential

site of control that prevents an imbalance in the formation

of the cell envelope as a whole (Anderson et al., 1972).

Although C55-P remains the classical carrier lipid form

encountered in bacterial world, the lower-size homologs

decaprenyl phosphate (C50-P) and nonaprenyl phosphate

(C45-P) were shown to fulfill this essential function

in mycobacterial species (Kaur et al., 2004; Mahapatra

et al., 2005) and Paracoccus denitrificans (Ishii et al., 1986),

respectively.

In the peptidoglycan pathway, C55-P is needed for the

synthesis and transport of hydrophilic GlcNAc-MurNAc-

peptide monomeric structures across the hydrophobic en-

vironment of the cytoplasmic membrane to the externally

located sites of polymerization. Although this function is

essential, the knowledge of the metabolism of C55-P was still

very limited recently and based on fragmentary data ob-

tained from various bacterial species (Fig. 2). In particular,

only few data on the genes and enzymes involved in steps 2,

3 and 4 were available (Higashi et al., 1970a; Sandermann &

Strominger, 1972; Willoughby et al., 1972; Poxton et al.,

1974; Kalin & Allen, 1979).

Fig. 1. Membrane steps of peptidoglycan

biosynthesis. M, G and the five colored beads

linked to M represent MurNAc, GlcNAc and the

pentapeptide, respectively. C55-PP and C55-P

are for undecaprenyl pyrophosphate and

undecaprenyl phosphate, respectively.

Fig. 2. Metabolism of undecaprenyl phosphate in bacteria. The site

of action of bacitracin, an antibiotic which acts by sequestering of

undecaprenyl pyrophosphate, is indicated. Steps 1–4 are catalyzed

by undecaprenyl pyrophosphate synthase, undecaprenyl pyrophosphate

phosphatase, undecaprenol phosphokinase and undecaprenyl

phosphate phosphatase activities, respectively.

FEMS Microbiol Rev 32 (2008) 208–233 c� 2007 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

209Biosynthesis of peptidoglycan lipid intermediates

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Biosynthesis and recycling of undecaprenylpyrophosphate

The precursor for C55-P, undecaprenyl pyrophosphate

(C55-PP), is synthesized by addition of eight C5 isopentenyl

units (cis[Z]-configuration) onto C15 (all-trans[E])-farnesyl

pyrophosphate (FPP) (Fig. 2). This reaction is catalyzed by

the undecaprenyl pyrophosphate synthase UppS (di-trans,-

poly-cis-decaprenylcistransferase; EC 2.5.1.31), which

belongs to the family of cis-prenyltransferases of group IV

(Ogura & Koyama, 1998). Farnesyl pyrophosphate itself

results from head-to-tail condensation of isopentenyl pyr-

ophosphate with dimethylallyl pyrophosphate, generating

C10 geranyl pyrophosphate, followed by a second condensa-

tion of isopentenyl pyrophosphate, a reaction catalyzed by

the farnesyl pyrophosphate synthase that is a prototype of

the trans-prenyltransferase family (Ogura & Koyama, 1998;

Liang et al., 2002). The structure and mixed E,Z stereo-

chemistry of the C55-prenyl product (Fig. 3), as deduced

from the recent knowledge of the reaction mechanism of the

UppS synthase, confirmed earlier data of mass and nuclear

magnetic resonance spectrometry of the undecaprenol iso-

lated from bacterial membranes (Scher et al., 1968; Gough

et al., 1970).

The UppS synthase had been partially purified and

characterized from several bacteria including Salmonella

newington, Micrococcus luteus, Lactobacillus plantarum,

Bacillus subtilis and Escherichia coli (Christenson et al.,

1969; Baba & Allen, 1978, 1980; Takahashi & Ogura, 1982;

Baba et al., 1985; Fujisaki et al., 1986). Like the enzymes

involved in fatty acid synthesis, UppS is a soluble cyto-

plasmic enzyme that generates a membrane embedded

product (undecaprenyl pyrophosphate). The first uppS gene

was identified much more recently, in 1998 by Ogura’s

group, using a genomic DNA library of M. luteus B-P 26

constructed in E. coli and a screening of recombinant clones

for overexpression of the synthase activity (Shimizu et al.,

1998). Orthologs were subsequently identified in various

Gram-positive and Gram-negative bacterial species (Apfel

et al., 1999) and this gene was demonstrated to be essential

in E. coli (Kato et al., 1999) and Streptococcus pneumoniae

(Apfel et al., 1999). Interestingly, these newly identified

proteins did not exhibit significant sequence similarity with

members of the trans-prenyltransferase family and in parti-

cular they did not carry the characteristic aspartate-rich

DDXXD motif that is involved in substrate binding via a

Mg21 bridge in the latter enzyme family (Chen et al., 1994).

The construction of expression vectors allowed the purifica-

tion of mg quantities of various UppS in either wild-type or

histidine-tagged form (Shimizu et al., 1998; Apfel et al.,

1999; Pan et al., 2000). The enzyme activity showed both a

detergent (Triton X-100) and MgCl2 dependency (Apfel

et al., 1999), the latter cation being required for the binding

and subsequent condensation of isopentenyl pyrophosphate

(Chen et al., 2002b). UppS has now been characterized in

great detail, both biochemically and structurally, by different

groups. The crystal structure of M. luteus and E. coli

enzymes have been solved, either in apo form or in complex

with Mg21, isopentenyl phosphate, FPP, and product analo-

gues (Fujihashi et al., 2001; Ko et al., 2001; Chang et al.,

2004; Guo et al., 2005). The data showed that not only the

primary but also the three-dimensional (3-D) structure of

cis-prenyltransferases was totally different from that of

trans-prenyltransferases (Takahashi & Koyama, 2006). The

enzyme is a homodimer of 29-kDa subunits and each

monomer is composed of six parallel b-strands forming a

Fig. 3. Structure of the polyprenyl carrier lipid

involved in cell wall biosynthesis.


210 A. Bouhss et al.

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central b-sheet core, which is surrounded by five of the seven

a-helices (Fig. 4). A largely hydrophobic 30 A depth cleft was

observed at the protein surface whose entrance carries

several positively charged residues as well as a ‘structural P

loop’ motif that is characteristic of phosphate recognition

enzymes. A flexible domain located in the vicinity of the H3

a-helix and cleft entrance was identified as an important

region for the catalytic process and the two subunits were

shown to alternate between two different ‘closed’ and ‘open’

conformations (Ko et al., 2001; Chen et al., 2002b), the

‘closed’ form being catalytically active (Chang et al., 2003).

Based on the 3-D structure and site-directed mutagenesis

experiments, a model for substrate binding and a catalytic

mechanism were proposed (Fujikura et al., 2000, 2003; Pan

et al., 2000; Kharel et al., 2001; Takahashi & Koyama, 2006).

How the ultimate product chain length of cis-prenyltrans-

ferases is determined was also investigated in some detail

(Ko et al., 2001; Kharel et al., 2006; Takahashi & Koyama,

2006). Site-directed mutagenesis of E. coli and M. luteus

UppS and a sequence comparison with the C70�120 product

synthesizing eukaryotic cis-prenyltransferases highlighted

some residues that play a critical role in the determination

of the product chain length. It was proposed that charged

residues present at the hinge region of the H3 a-helix might

control the bending direction of the growing hydrophobic

prenyl chain along the hydrophobic interior of the H3 helix

so that the hydrophobic cleft could accommodate the bulk

of the prenyl chain to fit a suitable size during enzymatic

elongation. As the substrate specificity and catalytic proper-

ties of the FPP synthase and cis-prenyltransferases are

varying to some extent in the bacterial world, the size and

stereochemistry of the ultimate polyprenyl product, i.e. the

carrier lipid, are bacterial species specific: undecaprenyl

phosphate in most cases, rarely nonaprenyl phosphate

(C45) (Ishii et al., 1986), both with the classical o,di-

trans,poly-cis conformation, or decaprenyl phosphate (C50)

with an unusual o,trans,octa-cis conformation (Wolucka

et al., 1994; Kaur et al., 2004; Mahapatra et al., 2005) (Fig. 3).

Trace amounts of nona-, deca-, and dodeca-prenyl alcohol

derivatives were always detected together with the predomi-

nant C55-P lipid in membrane extracts, confirming the high

but not absolute specificity of the corresponding UppS

synthase activities observed during in vitro assays (Thorne

& Kodicek, 1966; Higashi et al., 1967, 1970b; Scher et al.,

1968; Gough et al., 1970; Umbreit et al., 1972; Umbreit &

Strominger, 1972b; Apfel et al., 1999).

The bacitracin antibiotic has been shown to inhibit

bacterial cell wall biosynthesis through sequestration of

C55-PP, the product of the UppS synthase, thereby provok-

ing the loss of the cell integrity and lysis (Siewert &

Strominger, 1967; Stone & Strominger, 1971; Storm &

Strominger, 1973). This potent antibiotic produced as a

mixture of related cyclic polypeptides by some strains of

Bacillus licheniformis and B. subtilis is extensively used for

prophylaxis and therapy in food animals. It is clinically

used for treatments of surface tissue infections in combina-

tion with other antimicrobial drugs. Its oral use had

been earlier suggested for the control of vancomycin-

Fig. 4. Crystal structure of undecaprenyl

pyrophosphate synthase dimer (UppS) from

Escherichia coli. The figure was prepared using

PyMol and the atomic coordinates (1JP3)

deposited by Ko et al. (2001). The seven

a-helices and six b-strands are shown in red and

green for subunit A and pink and blue for

subunit B, respectively. The flexible loop

without observable electron densities (residues

72–83) is represented as a dotted line in one

subunit. The hydrophobic cleft at the molecular

surface of each subunit is indicated by an

arrow.



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resistant enterococci (VRE) (O’Donovan et al., 1994)

although without evident success (Mondy et al., 2001;

Hachem & Raad, 2002).

Formation of undecaprenyl phosphate

The dephosphorylation of C55-PP (step 2 in Fig. 2) is

required before the lipid carrier becomes available for use

in the various biosynthetic pathways. This reaction must

also occur after each cycle of polymerization of cell wall

components (e.g. of peptidoglycan) and the release of the

linked saccharides, because the lipid carrier is in most cases

liberated in the pyrophosphate form. However, some excep-

tions exist: for instance, the transfer of 4-amino-4-deoxy-L-

arabinose (L-Ara4N) units to lipid A catalyzed by the ArnT

membrane protein uses undecaprenyl-phosphoryl-L-Ara4N

as the donor substrate and releases the lipid in the C55-P

form (Trent et al., 2001).

The membrane-bound phosphatase catalyzing this reac-

tion had been partially purified from M. luteus by Goldman

& Strominger (1972) about 30 years ago and some of its

properties were investigated. Its optimal pH for activity was

near 7.5, the enzyme did not require any cation, was

stimulated by nonionic Triton detergents, and failed to

hydrolyze isopentenyl pyrophosphate. The gene for this

activity, however, remained to be identified.

In 1993, an E. coli gene whose overexpression resulted in a

decreased susceptibility to bacitracin had been identified

and the authors hypothesized that this gene should encode

an undecaprenol phosphokinase (Cain et al., 1993). This

hypothesis was based on the assumption that a significant

pool of free C55-OH may exist in E. coli membranes, as

earlier demonstrated in some Gram-positive bacteria (Higa-

shi et al., 1970b), that would be directed towards the

formation of C55-P by the overproduced kinase, thereby

reducing cell requirements for C55-PP molecules and the cell

sensitivity to bacitracin. In fact, this question was recently

revisited and the bacA gene product was finally unambigu-

ously proved to be a C55-PP phosphatase (El Ghachi et al.,

2004). The overproduction of the BacA protein, which

allowed E. coli cells to resist to high concentrations of

bacitracin, was correlated with a large (280-fold) increase

of C55-PP phosphatase activity in membranes (El Ghachi

et al., 2004). The increased level of C55-PP phosphatase

activity likely accelerated the conversion of the pool of C55-

PP, the bacitracin target, to C55-P, resulting in an increased

cell resistance to the antibiotic. The 30 kDa protein BacA was

predicted to be an integral membrane protein with eight

transmembrane segments. It was successfully extracted

from cell membranes by the n-dodecyl-b-D-maltoside de-

tergent and purified to near homogeneity in the histidine-

tagged form (El Ghachi et al., 2004). The E. coli enzyme

exhibited a high C55-PP phosphatase activity of ca.

2200 nmol min�1 mg�1 of protein, a value about 7300-fold

higher than the basal activity detected in wild-type cell

membranes. It did not show any detectable C55-OH phos-

phokinase activity. Considering this newly identified func-

tion, it was proposed to rename the bacA gene uppP (El

Ghachi et al., 2004), for undecaprenyl pyrophosphate phos-

phatase (also named undecaprenyl-diphosphatase; EC

3.6.1.27), to follow the nomenclature previously adopted

with uppS that encodes the C55-PP synthase (Apfel et al.,

1999).

The dephosphorylation of C55-PP was predicted to be an

essential metabolic step. The finding that the bacA gene

could be deleted from the chromosome of E. coli (El Ghachi

et al., 2004), Mycobacterium smegmatis (Rose et al., 2004),

Staphylococcus aureus and Streptococcus pneumoniae (Chalk-

er et al., 2000), without loss of viability or any apparent

effect on growth rate or morphology, was therefore quite

surprising. Its deletion however resulted in impaired biofilm

and smegma formation in M. smegmatis, and attenuated

virulence in mouse models of infection in S. aureus and

Streptococcus pneumoniae. All the bacA deletion mutants

showed enhanced susceptibility to bacitracin (Chalker et al.,

2000; El Ghachi et al., 2004; Rose et al., 2004). The

unexpected viability of the bacA deletion mutants suggested

the existence of other cell proteins with C55-PP phosphatase

activity. The detection of a 25% residual phosphatase

activity in the membranes of the E. coli mutant was

consistent with this hypothesis (El Ghachi et al., 2004). No

bacA homologue was found in the genomes of the afore-

mentioned species, indicating that these putative phospha-

tases belonged to a distinct protein family. Although a copy

of bacA gene was found in most bacterial genomes se-

quenced to date (up to three putative bacA orthologs were

found in some species, e.g. Bacillus cereus), it was apparently

absent in some bacteria such as Helicobacter pylori (unpub-

lished data).

A search in databases for putative membrane phospha-

tases activities identified three proteins of unknown func-

tion that formed the so-called BcrR family: BcrC from

B. licheniformis, YbjG from E. coli, and YwoA from B. subtilis

(El Ghachi et al., 2005). Interestingly, the bcrC gene product

was described as one of the three components of the ABC

transporter system responsible for the protection of

B. licheniformis against the antibiotic it produces, bacitracin

(Podlesek et al., 1995, 2000), and the overexpression of the

E. coli ybjG and B. subtilis ywoA genes were reported to

increase bacitracin resistance in the corresponding bacterial

species (Harel et al., 1999; Cao & Helmann, 2002; Bernard

et al., 2003). Two other E. coli genes displaying similarity

with ybjG that encoded members of the PAP2 phosphatidic

acid-phosphatase family were also identified: yeiU, of un-

known function, and pgpB, encoding one of the two

phosphatidylglycerolphosphate phosphatases (Icho & Raetz,



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1983). All of these candidate proteins were predicted to be

integral membrane proteins and contained a sequence

identical or quite similar to the characteristic phosphatase

signature KX6RP-(X12�54)-PSGH-(X31�54)-SRX5HX3D (El

Ghachi et al., 2005) that had been identified previously by

Stukey & Carman (1997) and Neuwald (1997). Interestingly,

this conserved phosphatase motif was not detected in the

sequence of BacA and the absence of significant sequence

homology between BacA and the above mentioned proteins

clearly indicated that they belonged to two distinct protein

families. The chromosomal ybjG, yeiU and pgpB genes could

be disrupted individually without apparent effect on cell

growth but the coinactivation of the three genes bacA, ybjG

and pgpB was shown to be lethal (El Ghachi et al., 2005). A

thermosensitive conditional triple mutant strain was gener-

ated which lysed at the restrictive temperature due to the

depletion of C55-PP phosphatase activity and arrest of

peptidoglycan synthesis (El Ghachi et al., 2005). It con-

firmed the implication of at least the three proteins BacA,

YbjG and PgpB in the formation of the C55-P carrier lipid in

vivo. As observed with bacA, the overexpression of the

individual ybjG, yeiU and pgpB genes was correlated to an

increased resistance to bacitracin and an increased level of

C55-PP phosphatase activity in membranes. The B. subtilis

ywoA gene product was also subsequently purified and

proved to be a C55-PP phosphatase (Bernard et al., 2005).

Therefore, two different classes of integral membrane

proteins which belong to the BacA and PAP2 phosphatase

families, respectively, could catalyze the dephosphorylation

of C55-PP into C55-P in bacteria. The number of these

proteins could apparently vary from one bacterial species

to another. The situation in E. coli is one BacA protein and at

least two members of the PAP2 family. Most bacterial species

have only one copy of bacA in their genome but some

species seem to express several bacA orthologs (e.g. Bacillus

anthracis, B. cereus) and some others apparently do not

express any (e.g. H. pylori). The situation is similar for

members of the PAP2 family but the precise number of those

proteins that effectively exhibit C55-PP phosphatase activity

remains to be determined for each species.

Interestingly, genes belonging to either of these two

classes were also found in gene clusters conferring antibiotic

resistance in the bacitracin-producing and bacitracin-resis-

tant species. For instance, bcrC and ywoA genes that encode

phosphatases from the PAP2 family were located within

clusters expressing bacitracin efflux systems in B. lichenifor-

mis and B. subtilis strains, respectively (Podlesek et al., 1995;

Cao & Helmann, 2002; Bernard et al., 2003), and one bacA

ortholog was recently found to play a similar essential role

in acquired bacitracin resistance in Enterococcus faecalis

(Manson et al., 2004). C55-PP phosphatase encoding genes

are thus used by bacteria either for generating the essential

carrier lipid C55-P or for depleting cells of the pool of C55-

PP, the bacitracin target, as a mechanism of resistance to this

antibiotic.

The purification and further biochemical characteriza-

tion of these different proteins is now required to discern

phosphatase activities specifically involved in C55-P meta-

bolism from non- or less-specific phosphatases, of distinct

metabolic function, that can also use C55-PP as a substrate.

C55-PP is synthesized at the inner side of the cytoplasmic

membrane but is released at the outer side of the membrane

by the peptidoglycan polymerization machinery. Whether

the dephosphorylation of C55-PP occurs on both membrane

sides or only on one side is at present unknown. A

topological analysis of these different membrane proteins

will help to answer this question. The involvement of either

different phosphatases with catalytic sites orientated to-

wards the cytoplasm and the periplasm or a single phospha-

tase could be envisaged, but in both cases a trans-bilayer

movement of the carrier lipid should occur, suggesting here

also the involvement of a putative flippase (see ‘The possible

existence of a flippase’).

The reaction of dephosphorylation of C55-PP was con-

sidered as an interesting potential target in a search for new

antibiotics. The recent discovery that two classes of enzymes

and multiple orthologs in each class could participate in

this process could render this search more problematic.

However, one efficient way to inhibit this step remains the

sequestration of the C55-PP substrate, as demonstrated with

bacitracin (Siewert & Strominger, 1967; Stone & Strominger,

1971; Storm & Strominger, 1973). An attack of the pyropho-

sphate moiety of C55-PP was also suggested to be part of the

mechanism of action of nisin, a lantibiotic whose primary

target is lipid II (Bonev et al., 2004). It was earlier hypothe-

sized that the reaction of dephosphorylation of C55-PP

could be the site of action of colicin M, a bacteriolytic toxin

produced by some E. coli strains that kills sensitive E. coli

strains and related species (Harkness & Braun, 1989a, b).

This question was recently revisited and colicin M was in

fact identified as an enzyme catalyzing the specific degrada-

tion of lipids I and II peptidoglycan intermediates (El

Ghachi et al., 2006).

Undecaprenol: a storage form of lipid carrier?

The intriguing presence of free undecaprenol (C55-OH) in

bacterial membranes had been earlier reported in Gram-

positive species. More than 90% of the endogenous

C55-isoprenyl lipid of S. aureus was found in this non-

functional alcohol form (Higashi et al., 1970b) and

a similar situation was observed in Enterococcus faecalis

(Umbreit et al., 1972), Listeria plantarum (Thorne & Kodi-

cek, 1966; Gough et al., 1970), and Listeria monocytogenes

(Vilim et al., 1973). It could represent a reserve pool for the

regulation of the C55-P pool, an hypothesis that seemed to



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be corroborated by the detection of two membrane-

associated enzyme activities, undecaprenol phosphokinase

and undecaprenyl phosphate phosphatase, catalyzing

the interconversion of C55-OH and C55-P (steps 3 and 4

in Fig. 2), in some of the latter species (Higashi et al.,

1970a; Sandermann & Strominger, 1972; Willoughby

et al., 1972; Poxton et al., 1974; Kalin & Allen, 1979).

The corresponding genes however remained to be

identified.

Only one report on the characterization of a C55-P

phosphatase had been published to date (Willoughby et al.,

1972) but a potential involvement of nonspecific phospha-

tases that could explain the generation of the large pool of

C55-OH was also suggested (Bohnenberger & Sandermann,

1976). The C55-P phosphatase activity detected in particu-

late fractions from S. aureus did not require any cation, was

optimal at pH 5, and had an apparent Km for C55-P of

1.5mM (Willoughby et al., 1972). All attempts to extract it

from membranes were unsuccessful and the enzyme was

consequently not purified nor further characterized. In the

same report, the authors mentioned the failure to detect a

similar activity in particulate fractions from B. subtilis,

Enterococcus faecalis, M. luteus and E. coli. This activity

observed in S. aureus probably accounted for the stimula-

tion by ATP of the overall rate of peptidoglycan synthesis

observed in vitro with enzyme preparations from this

organism (Anderson et al., 1966).

A C55-OH phosphokinase activity was shown to be

extractable by butanol from S. aureus and Klebsiella aero-

genes membranes (Higashi et al., 1970a; Poxton et al., 1974).

Its activity was optimal at pH around 8.5 and was dependent

on the presence of Mg21 (Higashi et al., 1970a). The enzyme

from S. aureus was purified to near homogeneity and its

molecular weight (MW) was estimated at 14 kDa (Sander-

mann & Strominger, 1972). The estimated Km value was

57mM for both C55-OH and the nucleotide cosubstrate ATP,

and ADP was confirmed as a reaction product. The phos-

phokinase from L. plantarum was partially solubilized by a

variety of methods utilizing Triton X-100 and was charac-

terized in some detail (Kalin & Allen, 1979). Its apparent Km

values for C55-OH and ATP were estimated at 14 mM and

2 mM, respectively. No other nucleoside triphosphate was

shown to substitute for ATP. Interestingly, it was very

recently suggested that the diacylglycerol kinase (DGK)

from Streptococcus mutans could also use C55-OH as an

alternative substrate (Lis & Kuramitsu, 2003). Although this

was not unambiguously established, the physiological sig-

nificance of this putative undecaprenol kinase activity of

DGK was further supported by an increased susceptibility to

bacitracin of the dgk mutant strain as compared with that of

the parental strain. It could thus be hypothesized that the

C55-OH phosphokinase activity that had been purified from

S. aureus membranes about 30 years ago was due, at least in

part, to the DGK enzyme. The S. aureus dgk gene codes for a

114-residues protein with a MW of 12.97 kDa, a value close

to that (14 kDa) earlier estimated for the C55-OH phospho-

kinase from this organism (Sandermann & Strominger,

1972), but no evidence that this protein also exhibits C55-

OH phosphokinase activity has been provided. It should be

noted that the DGK from E. coli was proved to be inactive on

C55-OH (Bohnenberger & Sandermann, 1979; Lis & Kur-

amitsu, 2003). Moreover, C55-OH has never been detected

in E. coli cells membranes, except in nonphysiological

conditions, e.g. following enzymatic degradation of lipid

intermediates by colicin M (El Ghachi et al., 2006). The

existence of a pool of C55-OH and expression of the couple

of kinase and phosphatase that catalyze its interconversion

with C55-P have thus only been demonstrated in a restricted

number of bacterial species. Their effective implication in

the regulation of the pool of C55-P and of its use for the

synthesis of the different cell-wall polymers remain to be

demonstrated.

Lipid I biosynthesis

The lipid I is an essential intermediate molecule in the

peptidoglycan biosynthesis pathway (Fig. 1). Its existence

was reported for the first time by Chatterjee & Park (1964)

and Struve & Neuhaus (1965). Afterwards, it was identified

as a lipid intermediate produced by transfer of the phospho-

MurNAc-pentapeptide moiety from UDP-MurNAc-penta-

peptide onto a lipid fraction (Anderson et al., 1965) with

concomitant release of UMP. The elucidation of the lipid

structure had been achieved by Higashi et al. (1967), who

found that the acceptor was a C55 isoprenoid alcohol

phosphate, undecaprenyl phosphate (C55-P). However, as

detailed in ‘Undecaprenyl phosphate metabolism’, the length

and stereochemistry of this carrier lipid appeared to be

slightly different in certain bacterial species, such as

M. smegmatis (Mahapatra et al., 2005) and P. denitrificans

(Ishii et al., 1986). In E. coli, the pool of the lipid I was

estimated at about 700 molecules cell�1 (van Heijenoort

et al., 1992). Such an extremely low pool level was explained

by the fact that this compound is an intermediate in the

pathway whose synthesis and utilization reactions are effi-

ciently coupled.

Translocase I reaction and identification of themraY gene

In 1965, Strominger and Neuhaus laboratories demon-

strated for the first time the transfer of the phospho-

MurNAc-pentapeptide moiety from the soluble UDP

nucleotide precursor onto the C55-P carrier lipid, using

membrane preparations from S. aureus and M. luteus



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(Anderson et al., 1965; Struve & Neuhaus, 1965).

C55-Pþ UDP-MurNAc-pentapeptide2UMPþ C55-PP-MurNAc-pentapeptide ðlipid IÞ

This reaction, which is generally referred to as the

‘transfer reaction’, does not lead to any modification of the

basal MurNAc-pentapeptide structure synthesized by the

Mur synthetases in the cytoplasm. It essentially consists in its

translocation onto the C55 carrier lipid present in the

cytoplasmic membrane. This anchoring is required before

subsequent steps could occur, i.e. the addition of the GlcNAc

residues by the MurG transferase, the passage of the

peptidoglycan monomeric structures through the mem-

brane and finally their polymerization on the outer side of

the cytoplasmic membrane. The enzyme catalyzing this first

transfer/translocation reaction, the phospho-MurNAc-pen-

tapeptide translocase or MraY (E.C. 2.7.8.13), therefore

insures the link between the cytoplasmic and periplasmic

steps of peptidoglycan biosynthesis (van Heijenoort, 2001b;

Bugg et al., 2006).

MraY was also found to be able to exchange radio-

labelled UMP for the unlabelled UMP moiety of UDP-

MurNAc-pentapeptide, consistent with the overall reaction

referred to as the ‘exchange reaction’:

½14C�UMPþUDP-MurNAc-pentapeptide2UMP

þ ½14C�UDP-MurNAc-pentapeptide

The mraY gene had been identified by Ikeda et al. (1991)

within a large cluster of genes, named mra for ‘murein region

A’, located at 2 min on the chromosome map of E. coli. In this

region the genes are tightly packed and appear in the order:

pbpB-murE-murF-mraY-murD-ftsW-murG-murC-ddlB-ftsQ-

ftsA-ftsZ-envA. They all code for proteins involved in pepti-

doglycan biosynthesis and cell division (Mur synthetases

MurC/D/E/F, penicillin-binding protein PBP3; division pro-

teins FtsW/Q/A/Z, MurG transferase, and D-Ala-D-Ala ligase

DdlB). A quite similar organization of this cluster was found

in other bacterial species. Ikeda et al. (1991) showed that the

overexpression of the E. coli mraY gene resulted in an increase

of the UDP-N-acetylmuramoyl-pentapeptide: undecaprenyl-

phosphate phospho-N-acetylmuramoyl-pentapeptide trans-

ferase activity in membranes, demonstrating that this gene

encoded the latter activity. Expression of the mraY gene was

shown to be dependent on the Pmra promoter (Hara et al.,

1997; Mengin-Lecreulx et al., 1998). This gene is essential for

the bacterial viability (Boyle & Donachie, 1998; Thanassi

et al., 2002) and a conditional mraY mutant strain was shown

to accumulate peptidoglycan nucleotide precursors under

restrictive growth conditions (Lara et al., 2005). One copy of

the mraY gene was found in all bacterial genomes sequenced

to date but was not detectable in eukaryotic organisms and

archaebacteria which both lack peptidoglycan. Interestingly,

however, the presence of an mraY gene orthologue was

recently demonstrated in the plant Arabidopsis thaliana

(Mondego et al., 2003). These authors postulated that this

MraY-like product could participate in the biosynthesis of

specific proteoglycan arabinogalactan proteins that reach

peak expression during late flower bud development. It was

also reported that the mraY gene product from A. thaliana

had putative plastid-targeting signals (Machida et al., 2006).

MraY protein, a member of the polyprenyl-phosphate N -acetyl hexosamine 1-phosphatetransferase superfamily

The alignment of MraY orthologue sequences from both

Gram-negative and Gram-positive species available in data-

bases allowed Bouhss et al. (1999) to identify a set of five

well-conserved hydrophilic sequences (I–V) containing 34

invariant amino acid residues (Fig. 5). In the E. coli MraY

sequence these domains were defined as H65-L80 (I), I111-

K133 (II), N189-L200 (III), L251-G275 (IV) and V296-F342

(V). The size of the MraY protein appears to be fairly well

conserved in the bacterial world, the sequences from Gram-

negative species being generally longer than those found in

Gram-positive species (�360 vs. 320 amino acid residues),

owing mainly to an extension at the N-terminal extremity.

Unusually large MraY proteins (�420 residues) are found,

however, in some bacteria such as Bacteroides (Price &

Momany, 2005). The presence of alternating hydrophobic

and hydrophilic segments in its primary structure clearly

suggested that MraY was an integral membrane protein

spanning the cytoplasmic membrane several times (Ikeda

et al., 1991). Moreover, a lipid microenvironment was

shown to be required for the MraY activity (Heydanek &

Neuhaus, 1969; Umbreit & Strominger, 1972a; Geis & Plapp,

1978; Weppner & Neuhaus, 1979). More recently, Bouhss

et al. (1999) determined the two-dimensional membrane

topology of both the E. coli and S. aureus MraY translocases,

using b-lactamase fusion experiments. A common topolo-

gical model was proposed which contained ten transmem-

brane segments joining four periplasmic loops and five

cytoplasmic sequences corresponding to the conserved

hydrophilic patterns I–V (Fig. 5). Both the N- and C-

terminal extremities were located in the periplasmic space.

This model was in agreement with many structural features

predicted from a sequence comparison of MraY ortholo-

gues, strongly suggesting its validity for all eubacterial MraY

proteins (Bouhss et al., 1999).

The cytoplasmic sequences II, III and IV of MraY were

also found in other prokaryotic and eukaryotic proteins

catalyzing the same kind of reaction, namely the transfer of a

lipid phosphate to the b-phosphate of an UDP-linked

hexosamine. These three conserved patterns thus defined

a superfamily of enzymes termed polyprenyl-phosphate



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N-acetylhexosamine-1-phosphate transferases or UDP-Hex-

NAc : polyprenyl-P HexNAc-1-P transferases. In addition to

MraY, this superfamily contained the prokaryotic enzymes

WecA, TagO, WbcO, WbpL and RgpG involved in the

biosynthesis of different cell envelope polymers (enterobac-

terial common antigen, lipopolysaccharide O-antigen, tei-

choic acids or rhamnose-glucose polysaccharide) (Soldo

et al., 2002; Lehrer et al., 2007) and a eukaryotic paralogue,

GPT, involved in protein N-glycosylation (Lehrman, 1994;

Dal Nogare et al., 1998; Burda & Aebi, 1999). These enzymes

share a common membrane-bound acceptor substrate, un-

decaprenyl phosphate in bacteria or dolichyl phosphate in

eukaryotes, but they differ in their selectivity for the soluble

UDP-N-acetyl-hexosamine substrate. The sugar nucleotide

donors are UDP-MurNAc-pentapeptide and UDP-GlcNAc

for MraY and WecA, TagO and GPT, respectively, the MraY

substrate carrying an additional 3-O-lactoyl-pentapeptide

group.

The cytoplasmic sequences I and V are highly specific for

all MraY orthologues and are also present in some bacterial

paralogues with some differences. However, they are not

found in the eukaryotic paralogue sequences (GPTs). Se-

quence and membrane topology analyses revealed that all of

the invariant or highly-conserved residues identified within

MraY, and/or WecA and the eukaryotic GPTs, were located

on the cytoplasmic side of the membrane, consistent with

the active site being orientated towards the cytoplasm

(Bouhss et al., 1999; Lloyd et al., 2004; Lehrer et al., 2007).

The patterns II, III and IV would be involved in the substrate

binding and/or the catalytic process that are common

features in the enzyme superfamily, while the patterns I and

V would be involved in the substrate specificity and in

particular the recognition of the sugar nucleotide substrate.

Purification and biochemical characterization ofMraY

Solubilization, expression, and purification ofMraY

The availability of a pure, stable, soluble and active prepara-

tion of the integral membrane MraY protein was a prere-

quisite to the development of detailed biochemical

investigations. Since 1969, many soluble and active prepara-

tions of MraY were described, extracted from membranes of

various bacteria such as micrococci, S. aureus and E. coli

using detergents and in particular Triton X-100 or CHAPS

(Heydanek & Neuhaus, 1969; Umbreit & Strominger, 1972a;

Pless & Neuhaus, 1973; Brandish et al., 1996a; Breukink

et al., 2003; Lloyd et al., 2004; Stachyra et al., 2004). Upon

detergent extraction, the S. aureus MraY was shown to

require the presence of a phospholipid, either phosphatidyl-

choline, dioleoyl phosphatidyl choline or phosphatidylgly-

cerol (Pless & Neuhaus, 1973). Brandish et al. (1996a)

reported that overexpressed E. coli MraY protein was

preferentially activated by phosphatidylglycerol. All pre-

vious attempts to overexpress significantly and purify any

MraY protein had been unsuccessful. Thus, only partially

purified enzyme preparations were generally used for enzy-

matic assays (Brandish et al., 1996a; Zawadzke et al., 2003;

Lloyd et al., 2004; Stachyra et al., 2004). Recently, a

comparative study performed with recombinant MraY

proteins from E. coli, S. aureus, B. subtilis and Thermotoga

maritima, expressed in the E. coli C43(DE3) host strain,

identified n-dodecyl-b-D-maltoside and N-lauroyl-sarcosine

as the most efficient detergents for the extraction of this

protein from cell membranes (Bouhss et al., 2004). In the

Fig. 5. Membrane topology of the MraY

translocase (Escherichia coli). The topological

model consists in ten transmembrane

segments, four periplasmic loops and five

cytoplasmic sequences corresponding to the

conserved hydrophilic patterns I–V (Bouhss

et al., 1999). Conserved residues by identity

(red) and by similarity (blue) are indicated.



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same report, conditions allowing the high-level overexpres-

sion of a MraY protein and, for the first time, its purification

to homogeneity in milligram quantities were described. The

specific activity of the pure B. subtilis MraY protein was

estimated at 1900 nmol min�1 mg�1 (Bouhss et al., 2004).

Substrate specificity and kinetic properties

MraY has two substrates: C55-P and UDP-MurNAc-penta-

peptide. The structure of the lipid substrate is expected to be

conserved in most bacterial species but a few exceptions

exist, as mentioned in ‘Undecaprenyl phosphate metabo-

lism’. The structure of the sugar nucleotide substrate shows

important variations in the bacterial world, particularly in

the peptide moiety (Schleifer & Kandler, 1972; Barreteau

et al., 2008; Vollmer et al., 2008). This peptide is a pentapep-

tide that generally contains at the third position either a

meso-A2pm residue in Gram-negative bacteria (as E. coli)

and Bacillus species or a lysine residue (more rarely

an ornithine) in most Gram-positive bacteria (as S. aureus).

In vivo complementation experiments have shown that the

S. aureus MraY was functional in E. coli and restored growth

of a mraY thermosensitive mutant, indicating that it accepts

the A2pm-containing sugar nucleotide (Bouhss et al., 1999).

Similarly, the overexpression of the S. aureus MurE synthe-

tase in E. coli (which introduces lysine instead of A2pm at

the third position of the nucleotide substrate) resulted in a

massive incorporation of lysine into the peptidoglycan,

demonstrating the efficient utilization of the lysine-contain-

ing UDP-MurNAc-pentapeptide by the E. coli MraY enzyme

(Mengin-Lecreulx et al., 1999). The relatively low specificity

of the MraY translocase and its tolerance toward the

variability of the peptide chain was a well-known character-

istic (Hammes & Neuhaus, 1974) that has been confirmed in

various other circumstances both in vivo and in vitro.

Shorter or longer peptides as well as modified peptides were

shown to be accepted: dipeptides (Ornelas-Soares et al.,

1994), tripeptides (Hammes & Neuhaus, 1974; Pisabarro

et al., 1986; van Heijenoort et al., 1992), tetrapeptides

(Hammes & Neuhaus, 1974), acetylated and dansylated

pentapeptides (Ward & Perkins, 1974; Weppner & Neuhaus,

1977; Brandish et al., 1996a; Stachyra et al., 2004), as well as

hexa- and heptapeptides (Billot-Klein et al., 1997). More

recently, the MraY enzyme from T. maritima was shown to

act on two different substrates in vivo: a D-lysine-containing

UDP-MurNAc-tripeptide and an L-lysine-containing UDP-

MurNAc-pentapeptide, with similar efficiencies (Boniface

et al., 2006).

The replacement of the L-Ala and D-Ala residues at

positions 1 and 4 of the pentapeptide chain, respectively, by

glycine reduced the S. aureus MraY catalytic efficiency by

135-fold (Hammes & Neuhaus, 1974). The use of UDP-

MurNAc-tetrapeptide (lacking one D-Ala) and UDP-Mur-

NAc-tripeptide (lacking two D-Ala) as substrates reduced

the MraY catalytic efficiency by fourfold and

80-fold, respectively, as compared with the nucleotide

pentapeptide. Stickgold & Neuhaus (1967) determined that

5-fluorouracil-substituted UDP-MurNAc-pentapeptide was

utilized at o 2% of the rate of the native substrate and was a

competitive inhibitor (Ki = 0.12 mM) of the transfer reac-

tion, just as 5-fluoro-UMP is a competitive inhibitor

(Ki = 50 mM) of the exchange reaction. However, 20-deoxy

UMP was accepted as a good substrate in the exchange

reaction (Neuhaus, 1971).

Various MraY assays using radiolabeled or fluorescent

substrates, resolution of reaction mixtures by paper or

thin-layer chromatography, or direct measurement using

microplates, have been developed for analyzing the kinetic

properties of this enzyme and for the screening of inhibitors

(Weppner & Neuhaus, 1977; Brandish et al., 1996a, b;

Bouhss et al., 2004; Stachyra et al., 2004). Typical Michae-

lis–Menten kinetics were observed with both purified and

partially purified enzyme preparations (Anderson et al.,

1966; Struve et al., 1966; Stickgold & Neuhaus, 1967;

Hammes & Neuhaus, 1974; Brandish et al., 1996a; Bouhss

et al., 2004; Stachyra et al., 2004). Km values observed for

UDP-MurNAc-pentapeptide were generally in the 5–30 mM

range with nonpurified enzyme preparations. However, a

much higher value of 1 mM was determined with the

purified MraY enzyme from B. subtilis (Bouhss et al., 2004).

The structure and size of the C55-P lipid acceptor

substrate present in the membranes is essentially determined

by the substrate specificity and catalytic properties of the

undecaprenyl pyrophosphate synthase UppS. However,

the E. coli MraY translocase was shown to accept in vitro

heptaprenyl (C35) as well as dodecaprenyl (C60) phosphate

as alternative substrates, with Km values (10–20 mM) similar

to that of the natural substrate (Brandish et al., 1996a).

Breukink et al. (2003) have reported that the Micrococcus

flavus MraY was active in vitro on polyprenyl substrates

containing from two to 25 isoprenyl units, with a maximal

efficiency observed for substrates bigger than C35. The

purified B. subtilis MraY exhibited a Km for C55-P about 10-

fold higher than that of the nonpurified MraYs from other

bacterial species (Bouhss et al., 2004). It could thus be

assumed that the membrane environment affects the recog-

nition of the lipid substrate by the enzyme.

The MraY activity is dependent on the presence of both

mono- and divalent metal ions. The activity of the E. coli

and B. subtilis enzymes was stimulated (two- to fourfold) in

presence of 10–100 mM of potassium or sodium (Brandish

et al., 1996a; Bouhss et al., 2004). Heydanek et al. (1970) also

observed that the S. aureus MraY was stimulated by many

monovalent metal ions such as K1, Rb1, Cs1 and NH41. The

MraY enzyme has an absolute requirement for a divalent

metal ion, particularly Mg21, at 5–40 mM concentrations



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(Heydanek et al., 1970; Bouhss et al., 2004). Mn21 can

replace Mg21 but the activity is decreased by two orders

of magnitude (Bouhss et al., 2004). Lloyd et al. (2004)

proposed that the D115 or D116 aspartate residues of

E. coli MraY, which are located in the conserved hydro-

philic sequence II (Fig. 5) and whose replacement by

Asn leads to loss of catalytic activity, are involved in

Mg21 chelation.

Catalytic mechanism of MraY

In 1969, on the basis of kinetic evidences, Heydanek et al.

(1969) proposed a two-step catalytic mechanism for the

MraY reaction (Fig. 6). It consisted in an attack by a

nonidentified nucleophile residue of MraY on the b phos-

phate of UDP-MurNAc-pentapeptide, generating a covalent

enzyme-phospho-MurNAc-pentapeptide intermediate with

concomitant release of UMP. The second step corresponded

to the attack by an oxyanion from C55-P on the phosphate of

the covalent intermediate, resulting in the formation of lipid I

and regeneration of the native enzyme form. Recently, Lloyd

et al. (2004) proposed that the D267 aspartate residue of

E. coli MraY, which is located in the conserved hydrophilic

sequence III, plays the role of the catalytic nucleophile. Three

observations supported the two-step mechanism: (1) phos-

pho-MurNAc-pentapeptide was formed during the transfer

reaction that likely resulted from the hydrolysis of the

enzyme-linked phospho-MurNAc-pentapeptide intermedi-

ate; (2) the enzyme could catalyze the exchange of UMP with

the UMP moiety of UDP-MurNAc-pentapeptide in the

presence or absence of the cosubstrate C55-P (Pless &

Neuhaus, 1973); (3) dodecylamine inhibited the synthesis of

lipid I and caused the release of the phospho-MurNAc-

pentapeptide.

A major criticism of the experiments performed by

Heydanek et al. that could have resulted in a misinterpreta-

tion of the results was the use of a nonpurified MraY

enzyme. The different observations they made could effec-

tively be attributed to the presence of contaminating en-

zymes (phosphatases) and C55-P lipid acceptor in the

preparation used. An alternative mechanism consists in a

direct attack of the phosphate oxyanion of C55-P onto the bphosphate of UDP-MurNAc-pentapeptide. This would lead

to the formation of lipid I and UMP in only one step (Fig.

6). The recent availability of pure MraY protein (Bouhss

et al., 2004) will allow to revisit this question and elucidate

the catalytic mechanism.

Recent progress on inhibitors of theMraY-catalysed reaction

MraY is the target for several classes of natural product

antibiotics, which in some cases have been studied in detail

through structure/function studies and/or detailed kinetic

or mechanistic studies. The structures of the MraY inhibi-

tors have been reviewed in detail (Kimura & Bugg, 2003;

Dini, 2005; Bugg et al., 2006). This section will briefly survey

the different inhibitor classes, and small molecule inhibitors

arising from structure/function studies.

There are five different classes of uridine-containing

nucleoside natural antibiotic products that target MraY,

illustrated in Fig. 7. The tunicamycins (related to the

streptovirudins and corynetoxins) contain a uridine disac-

charide, attached to a fatty-acyl chain. Tunicamycin is a

reversible, competitive inhibitor of MraY, with Ki 0.6 mM

(Brandish et al., 1996b), but is not useful as an antibacterial

agent, since it is toxic to mammals, through potent inhibi-

tion of GlcNAc-1-P transferase in the dolichol cycle of N-

linked glycoprotein biosynthesis (Heifetz et al., 1979). The

Fig. 6. Proposed MraY translocase reaction mechanisms. The two-step mechanism involves a covalent enzyme-phospho-MurNAc-pentapeptide

intermediate and the single-step mechanism consists in a direct attack of the phosphate oxyanion of C55-P onto the b-phosphate of the nucleotide

substrate.



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mureidomycins (related to the pacidamycins and napsamy-

cins) are peptidyl nucleoside natural products, containing a

30-deoxyuridine sugar attached via an enamide linkage to an

unusual peptide chain. The peptide chain contains an

N-methyl 2,3-diaminobutyric acid residue, and a urea

linkage to a C-terminal aromatic amino acid, which can be

meta-tyrosine, Trp, or Phe. Mureidomycin A is a slow-

binding inhibitor of E. coli MraY, with K�i = 2.2 nM (Brand-

ish et al., 1996a). The enamide functional group is not

essential for inhibition, and a range of dihydro-pacidamycin

analogues have been synthesized and tested for biological

activity (Boojamra et al., 2001). The mechanism of action of

mureidomycin A has been studied, and it was proposed that

the amino-terminus binds to the Mg21 cofactor-binding

site, and is positioned by an N-methyl amide cis-amide bond

(Howard & Bugg, 2003). A uridinyl dipeptide analogue of

mureidomycin A retained biological activity against Pseudo-

monas putida (Howard & Bugg, 2003).

The liposidomycins are fatty acyl nucleosides, whose

structures contain a sulfated aminoglycoside residue. Lipo-

sidomycin B is a slow-binding inhibitor (K�i = 80 nM)

of solubilized E. coli MraY (Brandish et al., 1996b).

A synthetic analogue containing the aminoribofuranose

monosaccharide attached to the 50 position of uridine

showed moderate levels of inhibition (IC50 = 50mM)

against translocase I when assayed in toluene-permeabilized

E. coli cells (Dini et al., 2000; Stachyra et al., 2004). Further

structure–function studies in this series of compounds

(riburamycins) gave more active inhibitors, and a synthetic

analogue containing a C12-fatty-acyl chain showed antibac-

terial activity against S. aureus (Dini et al., 2002; Stachyra

et al., 2004).

The muraymycins, reported by Wyeth Research in 2002,

contain an aminoribofuranoside monosaccharide, attached

to a short peptide chain, containing a urea linkage to a

C-terminal amino acid (McDonald et al., 2002). Members of

this family were potent inhibitors of MraY in vitro

(IC50 = 0.027mg mL�1), showed antibacterial activity

against S. aureus (MIC = 2–16mg mL�1) and enterococci

(MIC = 16–64mg mL�1) and were able to protect mice

against S. aureus infection (ED50 = 1.1 mg kg�1) (McDonald

et al., 2002). Synthetic analogues lacking the aminoribofur-

anoside showed reduced activity in vitro, but retained some

antibacterial activity (Yamashita et al., 2003).

Capuramycin, a further uridine nucleoside antibiotic, is a

potent inhibitor of MraY in vitro (IC50 = 0.017mg mL�1),

and a methylated derivative showed antibacterial activity

against M. smegmatis (MIC = 2–16mg mL�1) (Muramatsu

et al., 2003). Acylation derivatives of capuramycin showed

very potent activity (MIC = 0.06 mg mL�1) against several

Mycobacterium strains (Hotoda et al., 2003).

Genetic studies by Young and coworkers showed that

MraY was also the target for the bacteriolytic E protein from

bacteriophage fX174 (Bernhardt et al., 2000). The E protein

is a 91-amino acid protein, containing a transmembrane

domain. The killing action of E also required a host protein

Fig. 7. Structure of MraY translocase inhibitors.



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SlyD, a peptidyl-prolyl isomerase. Recent studies on

the mechanism of action of E showed that a 37-amino

acid peptide containing the transmembrane domain of

E was a potent inhibitor of membrane-bound MraY

(IC50 = 0.8 mM), but did not inhibit solubilized MraY, unlike

the small molecule inhibitors (Mendel et al., 2006). It has

been proposed that E inhibits MraY via a protein–protein

interaction, blocking the formation of protein–protein

interactions between MraY and other cell wall assembly

proteins in the cytoplasmic membrane at cell division

(Mendel et al., 2006).

Interestingly, it was recently demonstrated that colicin M

was an enzyme acting by specifically targeting and destroy-

ing the peptidoglycan lipid intermediates, thereby provok-

ing the arrest of peptidoglycan synthesis and cell lysis (El

Ghachi et al., 2006). The cleavage site was located between

the C55 and pyrophosphoryl groups, as demonstrated

in vitro by appropriate assays and in vivo by the observation

that colicin M-treated cells accumulated C55-OH, a lipid

form that is normally not detectable in E. coli cell mem-

branes (El Ghachi et al., 2006). To the authors’ knowledge

this is the first example of a mechanism of inhibition of

peptidoglycan biosynthesis occurring via enzymatic degra-

dation of its precursors, in this case the MraY and MurG

lipid reaction products.

Lipid II biosynthesis

The translocase II reaction and identification ofthe murG gene

The translocase II (MurG) catalyses the second membrane

associated step of peptidoglycan synthesis (Fig. 1). This

glycosyl transferase of the GT-B superfamily transfers the

GlcNAc moiety from UDP-GlcNAc to the C4 hydroxyl

group of lipid I to form a b-linked disaccharide (lipid II). It

was originally identified in E. coli but has since been

recognized in all other bacteria that make peptidoglycan.

The murG gene was first discovered by Salmond et al.

(1980) who identified, cloned and mapped the gene within

the mra region of E. coli. The nucleotide sequence of the

E. coli murG coding region (Mengin-Lecreulx et al., 1990)

revealed an ORF of 1065 nucleotides theoretically coding for

a moderately hydrophobic 37.8 kDa protein. When cultures

of a thermosensitive murG mutant strain (GS58) growing

exponentially at 30 1C were shifted to the nonpermissive

temperature of 42 1C, cells progressively lost their rod shape,

became ovoid with a greatly increased volume and finally

lysed (Mengin-Lecreulx et al., 1991). This mutant accumu-

lated significant amounts of UDP-MurNAc-pentapeptide,

UDP-GlcNAc and to a lesser extent a lipid compound as

labelled A2pm-containing precursors, suggesting that the

mutational block was in a membrane step. The fact that the

ratio of the two lipid intermediates I and II, which had

previously been reported to be between 0.3 : 1 and 0.6 : 1 in

E. coli strains (Ramey & Ishiguro, 1978), was reversed and

reached the considerably high value of 8.2 : 1 in this mutant

clearly indicated that it was the second membrane step

that was altered. It was consequently deduced that murG

encoded the UDP-GlcNAc : MurNAc-(pentapeptide)-

pyrophosphoryl-undecaprenol GlcNAc transferase (EC

2.4.1.227) which catalyses the formation of lipid II from

lipid I. This protein was recently shown to interact with the

MraYand MreB proteins and was suggested to participate in

two multi-protein complexes involved in cell elongation and

cell division, respectively (Aaron et al., 2007; Mohammadi

et al., 2007; den Blaauwen et al., 2008).

Purification and biochemical characterizationof MurG

Crouvoisier et al. (1999) first reported the overproduction,

solubilization and purification of the protein. MurG was

solubilized using salt or detergent from the cell membranes

and purified using anion exchange for the wild-type protein

or Ni21 affinity for the His-tagged protein. Ha et al. (2000)

also reported a similar procedure where the protein was

solubilized with detergent directly from cell pellets of E. coli

and purified using cation exchange followed by gel filtration.

van den Brink-van der Laan et al. (2003) showed that the

cardiolipin content of E. coli cell membranes was increased

following overexpression of MurG and that lipid vesicles

copurified with MurG. The activity of MurG was increased

in the presence of cardiolipin, suggesting specific interac-

tions of the protein with phospholipids and in particular

with cardiolipin.

Using optimal conditions in HEPES buffer, pH 7.9,

supplemented with 5 mM MgCl2 or MnCl2, Ha et al.

(1999) determined the kinetic parameters for E. coli MurG.

Because lipid I is almost impossible to isolate from bacterial

cells and difficult to handle, a synthetic lipid I analogue was

used. The assay used radiolabelled UDP-[14C]GlcNAc and a

biotin-labelled lipid I analogue. A biotin capture technique

was used to sequester products from the reaction which

were then counted for bound radioactivity. The parameters

were KUDP-GlcNAc = 58� 30 mM, Klipid I analogue = 37� 4 mM,

kcat = 16� 2 min�1 in the presence of MgCl2. Using a

different (C35) lipid I analogue and a new assay based

on HPLC, Auger et al. (2003) showed that a high con-

centration (35%) of dimethylsulfoxide was necessary

for maximal enzyme activity. The kinetic constants

they determined in these conditions were: Km UDP-GlcNAc

= 150� 20mM, Km lipid I analogue = 2.8� 1 mM, kcat = 56

� 5 min�1. Ha et al. (1999) also investigated the acceptor

and donor substrate specificity for MurG. The results

showed that a biotin-labelled UDP-MurNAc-pentapeptide



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was also a substrate for the enzyme with a relative rate of c.

20% when compared with the biotin-labelled lipid I analo-

gue. The results also revealed that the enzyme was sensitive

to the identity of the nucleotide, and required the presence

of the diphosphate linkage. The enzyme was inhibited by

UDP, but not by UMP, nor any other nucleotide dipho-

sphate. The enzyme also showed high specificity for the

equatorial stereochemistry at the C4 position of the donor.

Therefore, UDP-N-acetylgalactosamine failed to show in-

hibitory activity even at millimolar concentrations and

showed very little donor activity. Liu et al. (2003) reported

the development of a continuous fluorescence coupled

enzymatic assay in which the formation of UDP was coupled

to a pyruvate kinase-lactate dehydrogenase assay and the

fluorescence signal of NADH monitored. This group tested a

variety of lipid I analogues, with various substituents repla-

cing the undecaprenyl moiety, as substrates. MurG accepted

all of them with Km values of around 20–50 mM but there

were, however, large differences in the specific activity (kcat).

Acceptor substrates with long saturated linear alkyl chains

were better substrates than the natural lipid I demonstrating

an increase in kcat from 11� 2 min�1 for the natural

substrate to up to 180� 13 min�1 for an analogue with a

C14H29 chain.

Using synthetic substrate analogues and products con-

taining different length lipid chains, as well as a synthetic

dead-end acceptor analogue, Chen et al. (2002a) showed

that MurG follows an ordered Bi-Bi mechanism in which

UDP-GlcNAc binds first (Fig. 8). The enzyme likely utilizes a

mechanism that involves partial participation of the lone

pair on the sugar ring oxygen and therefore an oxocarbe-

nium-ion-like transition state. Evidence exists that supports

the procedure of glycosyltransferase-mediated reactions

through an oxocarbenium-ion-like transition state, similar

to that proposed for similar enzymes (Singh et al., 1987; Kim

et al., 1988; Takayama et al., 1999). Using a synthetic,

radiolabelled-analogue of lipid II, Auger et al. (2003)

reported for the first time that the reaction catalyzed by

MurG was reversible.

3-D structure of the MurG protein

The 1.9 A crystal structure of E. coli MurG (Ha et al., 2000)

revealed that the free enzyme consists of two protein

molecules in an asymmetric unit. Each protein chain has

two domains separated by a cleft which is c. 20 A deep and

18 A across at its widest point. Each domain adopts a

a/b open sheet motif which is characteristic of domains that

bind nucleotides. The N-terminal domain contains seven

parallel b-strands and six a-helices. The C-terminal domain

contains six parallel b-strands and eight a-helices including

one irregular bipartite helix that connects the N-domain to

the first b-strand of the C-domain. The structural homology

between the two domains is high, despite low sequence

homology. A subsequent, 2.5 A crystal structure of

E. coli MurG complexed with the donor substrate, UDP-

GlcNAc, was reported in 2003 (Hu et al., 2003a) (Fig. 9).

Before this, no X-ray crystal structures containing intact

substrates had been obtained for any of the NDP-glycosyl-

transferase superfamily. The substrate bound enzyme was

reported to have only a 16 A wide cleft, c. 2 A narrower than

the free enzyme. UDP-GlcNAc binds in both protein sub-

units along with four glycerol molecules and 121 water

molecules. Only one of the UDP-GlcNAc molecules is in

Fig. 8. Proposed MurG reaction mechanism. The model predicts a

deprotonation of the C4 hydroxyl group of the MurNAc moiety of lipid I

by a residue of MurG (probably His18). The oxyanion thus generated

then attacks the C1 of GlcNAc of the nucleotide substrate to form the

oxocarbenium-ion-like transition state. Finally, UDP is released generat-

ing lipid II. R = �C(CH3)CO-L-Ala-g-D-Glu-meso-A2pm-D-Ala-D-Ala,

Un = undecaprenyl.



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the correct orientation for catalytic activity and is presumed

to be the Michaelis complex where the UDP is displaced by

the incoming nucleophile. MurG contains a sequence motif

that is found in most members of the GT-B superfamily.

This a/b/a subunit has two a-helices located near to the cleft

between the domains. The UDP-GlcNAc substrate makes

several contacts to these helices and also to the loops that

connect them to the adjacent b-strands. The GlcNAc moiety

of the donor substrate makes contacts with the invariant

residues of MurG through hydrogen bonding interactions

between the backbone amide of the A263 residue, the side

chain amides of N291 and N127 and the C4 hydroxyl group

of GlcNAc. Interactions also exist between the side chain

amide of Q287 to both the C3 and C4 hydroxyl groups of

GlcNAc. The catalytic base appears to be histidine H18

situated 9.52 A across the domain cleft in line with the

anomeric bond. The contacts between the enzyme and the

diphosphate are purely hydrogen bonding. S191 has been

shown to be an important residue whose mutation to

alanine affects all kinetic parameters including an increase

in the Km for lipid I binding from 0.053 to 0.179 mM. This

serine residue is located on a GGS loop that is conserved in

all MurG homologs which moves up towards the donor

substrate as it binds. Because MurG is believed to utilise a

sequential mechanism in which lipid I binds following UDP-

GlcNAc (Chen et al., 2002a), it is thought that conforma-

tional changes in the GGS loop may play a role in the

adjustments required for lipid I binding, as well as directly

contributing to UDP-GlcNAc binding. The uracil nucleotide

is anchored in a pocket by hydrogen bonds from the back-

bone amide of I244 to the N3H and O4 atoms. There is also a

possible interaction between the uracil O2 atom and R163.

The F243 residue also rotates, as the substrate binds, to cap

the binding pocket. Contacts between the enzyme and the

ribose 20 and 30 hydroxyl groups exist through hydrogen

bonds from E268. To evaluate their role in the activity of the

E. coli MurG enzyme, 13 residues that are invariant or highly

conserved in the MurG enzyme family (T15, H18, Y105,

H124, E125, N127, N134, S191, N198, R260, E268, Q288,

N291) and located within or very close to the active site

were recently submitted to site-directed mutagenesis

(Hu et al., 2003a; Crouvoisier et al., 2007). Most of these

mutations resulted in a great loss of enzyme activity, con-

sistent with the interactions observed in the crystal structures

of MurG alone or in complex with UDP-GlcNAc (Ha

et al., 2000; Hu et al., 2003a). The important role of these

residues either in substrate binding/catalysis or maintenance

of the protein’s conformation was confirmed (Hu et al.,

2003a; Crouvoisier et al., 2007). Interestingly, a hydro-

phobic patch has been identified in the MurG structure,

consisting of I74, L78, F81, W84 and W115 residues, which

appears to be the site of interaction between this extrinsic

membrane protein and the phospholipid bilayer (Ha et al.,

2000).

Recent progress on inhibitors of the MurGreaction

The cyclic lipoglycodepsipeptide antibiotic ramoplanin has

excellent activity against a wide range of Gram-positive

bacteria. It was demonstrated that ramoplanin forms com-

plexes with both lipids I and II (Reynolds & Somner, 1990;

Fig. 9. Structure of Escherichia coli MurG

complexed with UDP-GlcNAc. This structure,

drawn with Swiss PDB Viewer, shows the

interactions between UDP-GlcNAc and the

enzyme active site. Binding of the lipid I

substrate is presumed to take place in the

cleft between the two domains of the MurG

structure. From Hu et al. (2003a).



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Lo et al., 2000; Helm et al., 2002; Hu et al., 2003b), however,

ramoplanin was also shown to directly interact with MurG

although the exact mechanism of inhibition is not known

(Fang et al., 2006).

Using the most active lipid I analogue substrate they had

identified, Liu et al. (2003) developed a screening for MurG

inhibitors. The potential inhibitors screened were vancomy-

cin, moenomycin and two chloro-biphenyl derivatives of

vancomycin. All of these compounds were found to inhibit

the enzyme with one of the vancomycin derivatives being

particularly potent. These known inhibitors of the bacterial

cell wall biosynthesis pathway, however, cannot penetrate

the bacterial cell membrane and so they do not encounter

MurG in vivo because the enzyme localizes on the cytoplas-

mic side of the cell membrane (Bupp & van Heijenoort,

1993).

A recent publication by Helm et al. (2003) detailed the

high throughput screening of almost 50 000 compounds

from a commercial library as inhibitors of MurG. The assay

employed in the screening used an N-acyl fluoresceinated

UDP-GlcNAc analogue instead of the natural substrate. As

MurG was added to the labelled substrate, an increase in

polarization occurred indicating that the substrate had

bound. The compounds were then screened at 25 mg mL�1

for in vitro activity as competitive inhibitors by looking for a

decrease in polarization. Eleven compounds were found to

reproducibly inhibit the enzyme by 50% or more. Seven of

these compounds contained a five-membered, nitrogen

containing heterocyclic core with an alkyl or aryl substituent

at N-1 and an arylidine substituent at the 3 position. A later

publication by Hu et al. (2004) utilized the same assay

to screen a further 64 000 compounds from a variety of

different compound libraries, including commercial and

diversity-orientated synthesis (DOS) libraries. Fifty-five

compounds were found to exhibit 4 40% inhibition of

MurG at a concentration of 2.5mg mL�1. Interestingly, 31 of

the 55 compounds were found to contain one of four

common cores (Fig. 10). It is not known exactly how these

compounds bind to MurG and antibacterial activity has not

yet been reported.

A direct continuous fluorescence assay for MurG based

on fluorescence resonance energy transfer (FRET) was

recently described (Li & Bugg, 2004). It used a dansyl-

labelled lipid I (lex = 500 nm, lem = 550 nm) and a fluores-

cent UDP-GlcNAc analogue (lex = 290 nm, lem = 500 nm)

as substrates for MurG. A linear relationship between

enzyme concentration and the rate of increase in fluores-

cence was observed, indicating that an efficient energy

transfer occurred as the substrates were converted to pro-

ducts by the enzyme. This assay is believed to be the first

direct continuous fluorescence assay for MurG and may

be very useful for determining the activity of potential

inhibitors.

Enzymatic modification of lipid II in thebacterial world

The peptidoglycan structure of many bacteria, especially

Gram-positive species, contains an additional peptide cross-

link between the residue (L-Lys or meso-A2pm in most cases)

at position 3 of the pentapeptide chain and the D-alanine

residue at position 4 of the cross-linked strand. There is

significant variation in the type and structure of these cross-

links across the bacterial kingdom (Schleifer & Kandler,

1972; Vollmer et al., 2008). Some examples are shown in

Fig. 11.

The addition of the cross-link amino acids occurs most

commonly at the level of the lipid II intermediate (Fig. 11),

although in Weisselia viridescens it has been shown that it

occurs at the level of the nucleotide precursor (Maillard

et al., 2005). Kamiryo & Matsuhashi (1972) showed that the

formation of the penta-glycine cross-link in S. aureus

required glycyl tRNA and lipid II . More recently, the

femABX genes have been implicated in the addition of the

penta-glycine bridge (Rohrer & Berger-Bachi, 2003): FemX

is responsible for addition of the first glycine; FemA the next

two glycines; and FemB the final two glycines (Rohrer &

Berger-Bachi, 2003). The FemABX reactions have recently

been reconstituted using purified recombinant proteins,

with lipid II and glycyl tRNA as substrates (Schneider et al.,

2004). There is a crystal structure of S. aureus FemA, which

contains a pair of extended a-helices, similar in structure to

a motif found in seryl tRNA synthetase that interacts with

the tRNA substrate (Benson et al., 2002). A crystal structure

of W. viridescens FemX complexed with its substrate UDP-

MurNAc-pentapeptide has been determined, which reveals

enzyme–substrate binding interactions (Biarrotte-Sorin

et al., 2004). Because aminoacyl tRNAs are found only in

the cytoplasm, the lipid II modification reactions must

occur on the cytoplasmic side of the membrane, after the

Fig. 10. General structures of MurG inhibitors. From Hu et al. (2004).

X = S, O.



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MurG-catalysed reaction, but before flipping to the cell

surface.

Dipeptide Ala-Ala cross-links are found in Enterococcus

faecalis (Bouhss et al., 2001), and in strains of highly

penicillin-resistant Streptococcus pneumoniae (Garcia-Bus-

tos & Tomasz, 1990), in the latter case linked to the murMN

genes (Filipe & Tomasz, 2000). The murMN genes are found

in all pneumococci, but specific sequences of murM are

found in highly resistant strains (Filipe et al., 2000). Two

transferases involved in the formation of the Ala–Ala cross-

link in Enterococcus faecalis have also been identified, which

are able to add Ala-tRNA to UDP-MurNAc-pentapeptide

in vitro (Bouhss et al., 2002). The Streptococcus pneumoniae

MurM reaction has recently been reconstituted in vitro using

recombinant MurM, lipid II, and alanyl-tRNA (Lloyd et al.,

2007). MurM from a penicillin-resistant strain was found to

show 80-fold higher kcat/Km than MurM from a sensitive

strain, providing an explanation for the higher level of cross-

links in the resistant strains (Lloyd et al., 2007). The ATP-

dependent enzyme responsible for addition of D-aspartic

acid to L-lysine residue in Enterococcus faecium has also been

identified (Bellais et al., 2006).

In a number of bacterial strains, the D-glutamic acid

residue at position 2 of the pentapeptide chain is amidated,

to form D-isoglutamine. In S. aureus, it is known that this

amidation is dependent upon ATP and ammonia, and that

either lipid I or II can be amidated in vitro (Siewert &

Strominger, 1968). A membrane-bound amidating activity

has also been observed in Bacillus stearothermophilus that is

able to amidate the nucleotide precursor (Linnett & Stromin-

ger, 1974). Lipid II isolated from M. smegmatis has been

found to contain several modifications: the MurNAc residue

is modified as N-glycolylmuramic acid, or as de-acylated

muramic acid; the diaminopimelic acid residue is amidated;

and the terminal D-alanine residue is methylated (Mahapatra

et al., 2005). It appears that a number of modification

reactions occur in particular bacterial strains, most com-

monly at the level of the lipid-linked intermediates (Fig. 11).

In S. aureus, the acceptor substrate of sortase A that catalyzes

anchoring of surface proteins to peptidoglycan was identified

as lipid II (Perry et al., 2002). These different modifications

should be taken into account when considering the lipid II as

a potential target for new antibacterial agents (Breukink & de

Kruijff, 2006).

Chemical and enzymatic synthesis oflipids I and II and analogues

In order to reconstitute in vitro the lipid-linked steps of

peptidoglycan biosynthesis, it was necessary to prepare the

Fig. 11. Structure of lipid II and its main modifications in bacterial world. R = H or COOH for Lys and A2pm, respectively; D-Asx = D-Asp or D-Asn.



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lipid-linked intermediates. Because they are present natu-

rally in very low abundance, it was not feasible to isolate

them from bacterial cells in useful quantities (van Heije-

noort et al., 1992; Guan et al., 2005). Chemical syntheses

were published for lipid I (VanNieuwenhze et al., 2001) and

for lipid II (Ye et al., 2001; VanNieuwenhze et al., 2002),

however the synthetic routes were long and proceeded in

fairly low overall yield. Auger et al., (1997, 2003) first

succeeded for the hemi-synthesis of functional analogues of

both lipids I and II. Chemical synthesis was used to prepare

soluble analogues of lipid I containing 10-carbon (Ha et al.,

1999) and 20-carbon chains (Cudic et al., 2001), which were

used as soluble substrates for MurG in vitro (Lazar & Walker,

2002).

Alternatively, lipids I and II can be prepared enzymati-

cally. The cytoplasmic precursor UDP-MurNAc-pentapep-

tide (containing meso-A2pm at position 3) can be

accumulated in B. subtilis using a cell wall synthesis medium

(Lugtenberg et al., 1972), and isolated in 50 mg quantities.

High level expression and purification of enzymes MurA-F

from E. coli or Pseudomonas aeruginosa and their use for the

preparation of the cytoplasmic intermediates in vitro have

been reported (Reddy et al., 1999; El Zoeiby et al., 2001;

Bouhss et al., 2004). An efficient procedure for the

conversion of UDP-MurNAc-pentapeptide into lipid I or

lipid II was published by Breukink et al. (2003), that used

membranes prepared from M. flavus, a species which

has elevated levels of MraY and MurG enzyme activities.

Using this procedure, they reported the preparation of

up to 50 mg quantities of lipids I or II. The availability of

purified C55-PP synthase UppS, C55-PP phosphatase

BacA, MraY and MurG proteins also facilitated the synthesis

of different forms of lipids I and II, with a labelling in

either the C55 or the pentapeptide moiety (El Ghachi et al.,

2006).

The e-amino group of meso-A2pm or L-Lys at position

3 of the pentapeptide chain can be chemically modified

by fluorophores such as dansyl. The N-dansyl analogue was

used to develop a fluorescence assay for MraY, for enzyme

kinetic studies (Brandish et al., 1996a), and for high

throughput inhibition assays (Stachyra et al., 2004). The

N-dansylated lipid II analogue was prepared and was shown

to give rise to fluorescence changes upon treatment with

E. coli PBP1b (Schwartz et al., 2002). An alternative

labelling strategy was recently published, in which the

D-alanine residue at position 4 or 5 can be replaced by

D-cysteine, by enzymatic synthesis from D-Ala-D-Cys or

D-Cys-D-Ala, and the thiol side chain of D-Cys can be

labelled by thiol-selective reagents such as pyrene maleimide

(Schouten et al., 2006). The S-labelled UDP-MurNAc-pen-

tapeptide analogues prepared in this way were converted

into fluorescent lipid I and II analogues, using M. flavus

membranes.

Possible existence of a flippase

Lipid II, biosynthesized on the cytoplasmic face of the

membrane, must somehow be flipped onto the extracellular

face of the membrane (Fig. 1). The possible existence of a

‘flippase’ protein capable of catalysing this process has been

discussed for many years (Weppner & Neuhaus, 1978; Ehlert

& Holtje, 1996) but no gene has been identified to date.

Analysis of fluorescently labelled peptidoglycan precursors

in S. aureus membranes by fluorescence energy transfer have

earlier concluded that the rate of unassisted flipping was

much too low to account for the rate of peptidoglycan

synthesis required at cell division (Weppner & Neuhaus,

1978), therefore it was generally assumed that there must be

some protein assistance for the flipping of lipid II across the

membrane. In fact, the biochemical process was only

recently demonstrated experimentally, in particular by ana-

lysis of the transmembrane transport of fluorescent lipid II

through model and bacterial membranes (Breukink et al.,

1999; van Dam et al., 2007). It was shown that lipid II flop

did not occur spontaneously and was not obligatory coupled

to lipid II synthesis. This suggested the involvement of one

or more integral membrane proteins or membrane-asso-

ciated proteins in this process. Based on their results, the

authors further hypothesized a coupling of the translocation

of lipid II with the subsequent reactions of polymerization

(transglycosylation) catalyzed by the penicillin-binding pro-

teins (van Dam et al., 2007).

The existence of ABC transporters that are able to catalyse

similar types of membrane transport events suggests that the

putative flippase might be similar to this protein family, for

which there is now structural information (Chang & Roth,

2001). The bacterial wzx gene was suggested as a candidate

flippase gene in lipopolysaccharide biosynthesis, on the basis

of gene knockout studies (Liu et al., 1996). A putative

flippase gene involved in N-linked protein glycosylation in

eukaryotic cells was identified, based upon gene knockout

studies (Helenius et al., 2002). The pglK gene encoding an

ABC transporter responsible for N-linked protein glycosyla-

tion in Campylobacter jejuni was shown to complement a

wzx deficiency in E. coli O-antigen biosynthesis, although

the corresponding gene products showed no sequence

similarity (Alaimo et al., 2006). In contrast, van Dam et al.

(2007) found that the flipping of lipid II was not dependent

upon ATP, which would argue against the involvement of an

ABC transporter. These different studies suggest that there is

a flippase protein for peptidoglycan biosynthesis, but its

identity and precise mechanism of action are yet to be

determined.

Concluding remarks

In summary, the understanding of the biochemistry of the

lipid-linked intermediates of bacterial peptidoglycan



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biosynthesis has increased considerably in the last few years.

Several genes encoding the phosphatase and kinase enzymes

involved in undecaprenyl phosphate metabolism have now

been identified. The corresponding enzymes are potential

new targets for antibacterial action. Translocase MraY has

been successfully purified, and is the target for a number of

natural product and synthetic inhibitors. There is an X-ray

crystal structure for glycosyltransferase MurG, which will

facilitate the development of novel inhibitors for this

enzyme. Both MraY and MurG are attractive targets for

antibacterial development, and methods for high-through-

put screening of both enzymes have been developed. The

putative ‘flippase’ activity responsible for transport of lipid

intermediate II across the cytoplasmic membrane remains

an interesting target which has not yet been identified. A

number of enzymatic modifications of lipid II by the

FemABX/MurMN proteins have now been reconstituted;

their role in bacterial cell growth and antibiotic resistance

mechanisms can now be studied. Further modifications of

lipid-linked intermediates take place in certain microorgan-

isms, and the ability to prepare lipids I and II in useful

quantities will allow more detailed studies on new modifica-

tion reactions. Availability of lipid II will also allow more

detailed biochemical studies on the transglycosylase and

transpeptidase reactions catalysed by the penicillin-binding

proteins in future years. How all these different membrane

proteins are organized in the membrane and possibly

interact together in relation to the cell elongation and cell

division processes also remains to be elucidated.

Acknowledgements

The authors thank Didier Blanot and Thierry Touze for

critical reading of the manuscript. This work was supported

by the Centre National de la Recherche Scientifique and by

the European Community (FP6 projects COBRA, LSHM-

CT-2003-503335, and EUR-INTAFAR, LSHM-CT-2004-

512138). AET was supported by a PhD studentship from

the Engineering and Physical Sciences Research Council.

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The biosynthesis of peptidoglycan lipid-linked intermediates

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