MicroReview

Lipid domains in bacterial membranes

Kouji Matsumoto,* Jin Kusaka, Ayako Nishibori and
Hiroshi Hara
Department of Biochemistry and Molecular Biology,
Graduate School of Science and Technology, Saitama
University, 255 Shimo-ohkubo, Sakura, Saitama,
Saitama 338-8570, Japan.

Summary

The recent development of specific probes for lipid
molecules has led to the discovery of lipid domains in
bacterial membranes, that is, of membrane areas dif-
fering in lipid composition. A view of the membrane
as a patchwork is replacing the assumption of lipid
homogeneity inherent in the fluid mosaic model of
Singer and Nicolson (Science 1972, 175: 720–731). If
thus membranes have complex lipid structure, ques-
tions arise about how it is generated and maintained,
and what its function might be. How do lipid domains
relate to the functionally distinct regions in bacterial
cells as they are identified by protein localization
techniques? This review assesses the current knowl-
edge on the existence of cardiolipin (CL) and phos-
phatidylethanolamine (PE) domains in bacterial cell
membranes and on the specific cellular localization of
certain membrane proteins, which include phospho-
lipid synthases, and discusses possible mecha-
nisms, both chemical and physiological, for the
formation of the lipid domains. We propose that bac-
terial membranes contain a mosaic of microdomains
of CL and PE, which are to a significant extent self-
assembled according to their respective intrinsic
chemical characteristics. We extend the discussion to
the possible relevance of the domains to specific
cellular

processes,

including

cell

division

and

sporulation.

Introduction

Until fairly recently, the lipids in the membranes of bacterial
cells were assumed to be homogeneously distributed, and
the fluidity of biological membranes had become generally

accepted, according to the fluid mosaic model by Singer
and Nicolson (1972). However, contradicting the assumed
homogeneity, it is apparent that cell membranes must be
laterally polarized to produce specific environments for
certain membrane proteins, in particular the polar
chemoreceptor proteins and host actin-polymerizing pro-
teins, and proteins involved in cell division at the midcell
and at asymmetrically positioned septa (for a review, see
Shapiro et al., 2002). Furthermore, results indicating lateral
heterogeneity of lipid molecules or lipid domains in the
membranes have emerged from a variety of studies both in
eukaryotic and in prokaryotic cells (for reviews, see Vereb
et al., 2003; Dowhan et al., 2004). In addition to the studies
employing biophysical techniques, microscopic visualiza-
tion of membrane lipids in bacterial cells has reinforced the
view that bacterial membranes do possess structural het-
erogeneity: uneven distribution of fluorescent lipophilic
dyes and selective staining of septal regions has been
observed in mycobacteria (Christensen et al., 1999) and
the distribution of fluorescence in Escherichia coli cells
stained with a lipophilic dye is distinctly uneven (Fishov and
Woldringh, 1999). Finally, unequivocal visualization of car-
diolipin (CL) domains in E. coli and Bacillus subtilis cells
has been accomplished by means of a CL-specific fluores-
cent dye (Mileykovskaya and Dowhan, 2000; Kawai et al.,
2004). Thus, studies with dyes have confirmed the exist-
ence of heterogeneity of phospholipids in bacterial
membranes and have prompted further work on the phos-
pholipid domains and on the relevance of this heterogene-
ity to physiological function.

Laterally heterogeneous distribution of
phospholipids in bacterial membranes

Cardiolipin-rich

domains

were

visualized

with

the

CL-specific fluorescent dye 10-N-nonyl acridine orange
(NAO) in the septal and on the polar membrane regions of
E. coli cells by Mileykovskaya and Dowhan (2000). The
same group proposed a model for the mechanism of
CL-specific staining in which the nonyl group of NAO
inserts between the phosphate groups at the hydrophobic
surface generated by the two outer acyl chains of CL
(Mileykovskaya et al., 2001). The dye forms an array of
parallel skewed stacks on the surface of the hexagonal

Accepted 3 July, 2006. *For correspondence. E-mail koumatsu@
molbiol.saitama-u.ac.jp; Tel. (

[phone]; Fax [phone]

3698.

Molecular Microbiology (2006) 61(5), 1110–1117

doi:10.1111/j.1365-2958.2006.05317.x

First published online 1 August 2006

© 2006 The Authors
Journal compilation © 2006 Blackwell Publishing Ltd

array that comprises all (four per molecule) acyl chains of
CL (Fig. 1). Septal and polar localization of the fluorescent
domains was observed in B. subtilis cells during exponen-
tial growth, but not in cells carrying a clsA null mutation
blocking CL synthase and lacking measurable levels of CL
(Kawai et al., 2004). In sporulating cells, fluorescent
domains were clearly observed in the polar septal and
engulfment membranes and subsequently in forespore
membranes at different stages during the course of sporu-
lation. Interestingly, spore membranes have a quite high
CL content (Kawai et al., 2006), although its localization in
the membranes is not yet known. The fluorescence images
of NAO in B. subtilis cells seem to be clearer than those
obtained with E. coli, probably due to the simpler envelope
structure of the former. The preferential localization of CL
at the poles of E. coli and B. subtilis cells is consistent with
its enrichment in minicells, which are formed by aberrant
cell division close to the pole (Koppelman et al., 2001 and
our unpublished results).

These findings encouraged us to examine the localiza-

tion of another major phospholipid, phosphatidylethanola-
mine (PE) with the cyclic peptide probe Ro09-0198 (Ro),
which binds specifically to PE (Emoto and Umeda, 2001).

Treatment with biotinylated Ro followed by detection with
tetramethylrhodamine-conjugated streptavidin revealed
that PE is localized in the septal membranes of exponen-
tial growth-phase cells of B. subtilis and in the mem-
branes of the polar septal and the engulfment membranes
and forespore membranes at various stages in sporulat-
ing cells (Nishibori et al., 2005). As mutant cells lacking
PE were not stained, one can be confident that the fluo-
rescence reflects the localization of PE-rich domains in
the septal membranes. A typical example of PE localiza-
tion in B. subtilis cells detected by fluoresceine-labelled
Ro is shown in Fig. 2A. It turns out that the application of
fluoresceine-labelled Ro produces clearer images of the
localization of PE than biotinylated Ro-streptavidin conju-
gated with tetramethylrhodamine (Nishibori et al., 2005).
Figure 2B shows a series of thin sections along the z-axis.
Note the band of intense fluorescence at the septal region
of every thin section, indicating that the intense fluores-
cence band is not an artefact from piling up of weak
fluorescence images but is actually present in the septal
membranes.

In E. coli cells, the fluorescence signal of Ro-bound PE

is distributed uniformly over the cell surface, suggesting
that PE is uniformly distributed over the whole cell mem-
brane (Nishibori et al., 2005). A uniform signal was also
observed in many other Gram-negative bacteria, e.g.
certain strains of Salmonella typhimurium, Pseudomonas

Cardiolipin

NAO

Fig. 1. Proposed arrangement of CL in the presence of NAO. A top
view of the bilayer in which the hexagonal array of large circles
represents the fatty acid chains is shown. The small internal circles
containing P represent the phosphate groups, hydrogen-bonded
tightly by the hydroxyl of the connecting glycerol, above the two
central circles of the four fatty acid chains of CL (red). This tight
array provides room for the NAO molecules (green) to stack in
between the rows of CL head groups. Adapted from Figure 3 of
Mileykovskaya et al. (2001) with the publisher’s permission.

A

B

Fig. 2. Visualization of PE-rich domains in B. subtilis cells with Ro.
A. Wild-type cells (left) and the pssA mutant cells lacking PE (right)
were treated briefly with lysozyme and then stained with
FITC-labelled Ro. Fluorescence images were viewed with a
fluorescence microscope and corresponding phase-contrast images
are also shown below.
B. Wild-type cells were treated with lysozyme and stained with
biotinylated Ro-streptavidin conjugated with tetramethylrhodamine.
A series of images of z-axis sections with a fixed spacing of 0.1

mm

was taken with a confocal laser microscope.

Lipid domains in bacterial membranes

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© 2006 The Authors
Journal compilation © 2006 Blackwell Publishing Ltd, Molecular Microbiology, 61, 1110–1117

putida, Azotobacter vinelandii and Proteus vulgaris. In
many Bacillus species, including Bacillus polymixa, Bacil-
lus amyloliquefaciens and Brevibacillus brevis, Ro-bound
PE showed septal PE localization, as in B. subtilis. The
different distribution of the signal between the Gram-
negative and Gram-positive bacteria might imply that PE
plays different physiological roles in the two bacterial
types.

The Ro probe binds at the cleavage furrow of dividing

Chinese hamster ovary (CHO) cells and it has been sug-
gested that PE, which usually resides in the inner leaflet of
the plasma membrane, is exposed on the outer leaflet of
the membrane of the cleavage furrow at the final stage of
cytokinesis (Emoto and Umeda, 2001). The outer leaflet
of the plasma membrane of yeasts has been probed for
PE using similar techniques (Iwamoto et al., 2004). The
PE signals are located at the bud neck of late mitotic
stage, large-budded Saccharomyces cerevisiae cells. In
the fission yeast Schizosaccharomyces pombe, PE is
located at the division plane of late mitotic cells and at one
or both poles of mononucleated cells, suggesting that PE
exposure at the region of cell division is a common
feature. In addition, the use of a fluorescent probe, filipin,
indicated sterol localization to the site of cell division in the
fission yeast (Wachtler et al., 2003).

Septal localization of phospholipid synthases and
generation of lipid domains

The septal localization of both PE- and CL-rich domains
directed our interest to the subcellular localization of the
enzymes involved in PE and CL synthesis. The committed
step in PE synthesis in B. subtilis is catalysed by
phosphatidylserine synthase (PssA). Its reaction product,
phosphatidylserine, is then converted to PE. Phosphati-
dylglycerophosphate synthase (PgsA) catalyses the com-

mitted step for the synthesis of phosphatidylglycerol (PG),
which is then used by CL synthase (ClsA) to produce CL.
All GFP fusions to these enzymes were septally localized,
even when expression levels (and hence fluorescence
intensity) were low (Nishibori et al., 2005). Thus, these
enzymes probably concentrate at the septum under
natural conditions.

Attempts to localize other B. subtilis enzymes involved

in lipid synthesis (Table 1) have yielded interesting results
(Nishibori et al., 2005). GFP fusions to several phospho-
lipid synthases were localized to the septum in a thick,
bright fluorescence band. These synthases include CdsA,
which produces CDP-diacylglycerol, Psd, which converts
phosphatidylserine into PE, MprF, which transfers lysine
to PG to produce lysyl-PG, and UgtP, which is responsible
for glucolipid synthesis. Their distribution thus differed
from the uniform distribution of the membrane proteins
AtpC, a subunit of ATP synthase, and SecY, the uniform
membrane distribution of GFP fusions of which has been
shown by H. Takamatsu and T. Kobayashi (pers. comm.)
and has also been confirmed in our laboratory. It also
differed from the uniform cytoplasmic localization of
GpsA, which catalyses the production of glycerol
3-phosphate. It has recently been suggested that a
complex of lipid synthesis factors, including the acylcarrier
protein, YbgC (which exhibits thioesterase activity on
acyl-CoA derivatives), PssA and sn-glycerol 3-phosphate
acyltransferase (PlsB), resides in the E. coli inner mem-
brane (Gully and Bouveret, 2006). It seems likely that the
septally localized lipid synthases are also integrated in
such a lipid synthesis complex for co-ordinated lipid
metabolism.

Although polar and septal localization of CL has been

demonstrated in E. coli, efforts to localize CL synthase
have been less successful. GFP-CL synthase chimeras
constructed so far were not functional and their fluores-

Table 1. Cellular localization of the product of the genes involved in lipid synthesis in B. subtilis and E. coli.

Gene

Function

Localization in B. subtilis cells

gpsA

sn-Glycerol 3-phosphate dehydrogenase (glycerol phosphate synthase)

Cytoplasmic

plsB*

sn-Glycerol 3-phosphate acyltransferase

–*

plsC (yhdO)

1-Acylglycerol 3-phosphate acyltransferase

Septal ?

cdsA

CDP-diacylglycerol synthetase

Septal

pgsA

Phosphatidylglycerophosphate synthase

Septal

clsA (ywnE)

Cardiolipin synthase

Septal

ywjE

Cardiolipin synthase

Septal

ywiE

Similar to cardiolipin synthase

Septal

mprF (yfiX)**

Lysylphosphatidylglycerol synthase

Septal

pssA

Phosphatidylserine synthase

Septal

psd

Phosphatidylserine decarboxylase

Septal

dgkA

Diacylglycerol kinase

Septal

ugtP (ypfP)**

UDP-glucose:diacylglycerol glucosyltransferase

Septal***

Genes and their functions and cellular localization of the products are from Nishibori et al. (2005). An asterisk indicates that the gene is absent in
B. subtilis and double asterisk indicates absence in E. coli. The triple asterisk indicates that the septal localization appears as a two-dot structure
that is thickest near the edge of the septal face. The question mark indicates that the localization may not be confined to the septal membranes
(unpublished results). Note that no specific localization of the enzymes in E. coli cells has been shown.

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K. Matsumoto, J. Kusaka, A. Nishibori and H. Hara

© 2006 The Authors

Journal compilation © 2006 Blackwell Publishing Ltd, Molecular Microbiology, 61, 1110–1117

cence was observed in the cytoplasm. No specific local-
ization of PgsA, the enzyme responsible for the preceding
reaction in E. coli cells, has been found either; the fluo-
rescent GFP-PgsA chimera was observed as dots distrib-
uted around the periphery of the cells (our unpublished
data).

How can phospholipid synthases be targeted to the

septal membranes? FtsZ-depletion experiments indicate
that the phospholipid synthase localization depends on
FtsZ and exclude the possibility that they become local-
ized before FtsZ ring assembly (Nishibori et al., 2005).
Thus, localization probably follows or is concurrent with
the assembly of cell division proteins in order to
synthesize phospholipid membranes in concert with the
synthesis of peptidoglycan at the leading edge of the
invaginating envelope. Two-hybrid analyses of B. subtilis
lipid synthases with cell division proteins and envelope
proteins are in progress. Preliminary results suggest pos-
sible interactions of the lipid synthases with some cell
division proteins and envelope proteins (our unpublished
results in collaboration with the group of Dr H. Yoshikawa,
Tokyo University of Agriculture). The lipid synthases pre-
sumably have a specific region(s) responsible for the
septal localization (or interaction with certain cell division
proteins), leading one to wonder about the consequences
of their inactivation. Furthermore, as the septal localiza-
tion of the phospholipid synthases in B. subtilis cells
implies that most phospholipids are synthesized mainly at
the septal membranes, there must be a mechanism that
prevents PE and CL diffusion into the lateral membranes.
This mechanism need not to be a physical barrier. It could
be that the diffusion of lipid molecules is essentially free
but that some lipids associate preferentially with much
larger structures that themselves are localized while other
lipids go elsewhere (V. Norris, pers. comm.).

It should be noted that the GFP-labelled version of

UgtP, responsible for glucolipid synthesis, appears as a
‘two dot’ structure that is thickest near the edge of the
septal face. It seems likely that it actually forms a ring
structure, like FtsZ, and thus differs from the phospholipid
synthases, which form a band on the septal membranes.
UgtP is probably not an integral membrane protein, as it
does not have a membrane-spanning region (Nishibori
et al., 2005), in contrast to phospholipid synthases, which
have several membrane-spanning regions. This property
of UgtP might have some relation to the difference in its
localization pattern on the septal membranes.

The chemical basis for the generation of lipid
domains

How do the lipid molecules form domains in membranes
that are fluid? Both lipid–lipid interactions and lipid–
membrane protein interactions are suspected to induce

the formation of microdomains, which comprise at most
some tens of molecules of a specific lipid (for a review,
see Edidin, 1997). The following properties of CL and PE
might account for the formation of labile microdomains
through lipid–lipid interactions.

The polar head of the PE molecule has both a cationic

amine residue and an anionic phosphate residue. Each
amine and unesterified phosphate oxygen can participate
in two short distance intermolecular hydrogen bonds. The
ethanolamine groups thus form a linkage between phos-
phorous groups of adjacent PE molecules producing a
very compact, rigid head-group network at the bilayer
surface (Elder et al., 1977; Hauser et al., 1981; Boggs,
1987), giving PE substantially higher T

m

values than phos-

phatidylcholine, which has an identical acyl chain struc-
ture (reviewed in London and Brown, 2000). This compact
head-group network of PE might well suffice to explain PE
microdomain formation. Interactions with membrane pro-
teins (both transmembrane and peripheral membrane
proteins) might then modify the lipid organization and
further stabilize the head-group network of PE, as sug-
gested by Edidin (1997).

Cardiolipin, which has a double-glycerophospholipid

structure connected with a glycerol residue, has a rather
unique head group, with a tightly locked, surprisingly
small configuration and only one negative charge (Haines
and Dencher, 2002; this result contradicts the depiction
found in textbooks). The two phosphates in the head
group trap a proton and are locked in a bicyclic array held
together by the hydroxyl residue of the glycerol, which
connects the two halves of the CL molecule. This small
polar head group makes for a tighter packing of hydro-
phobic acyl chains between CL molecules by van der
Waals force interactions than is found in lipids with larger
polar head groups. It has also been suggested (Haines
and Dencher, 2002) that interaction between adjacent
head groups creates a compact array of CL molecules
(Fig. 1), which becomes manifest in the presence of asso-
ciated NAO arrays (Mileykovskaya et al., 2001). Head
groups of PG, the third major phospholipid in E. coli, inter-
act by an extensive network of hydrogen bonds, ionic
bonds

and

co-ordination

bonds

between

glycerol

hydroxyls and the unesterified phosphate oxygen both in
the anhydrous crystal and in the hydrated gel state
(Boggs, 1987; Pascher et al., 1987). In fact, PG is
perhaps segregated into distinct domains, which are dif-
ferent in their composition and proteo-lipid interaction,
according to studies using pyren-labelled phospholipids in
both B. subtilis and E. coli membranes (Vanounou et al.,
2003). This extensive and tight network of PG molecules
might cause exclusion of CL molecules to produce
patches of CL in the membranes, as the small and tightly
locked head group of CL cannot interact with PG. Certain
membrane proteins with an affinity to CL might also have

Lipid domains in bacterial membranes

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© 2006 The Authors
Journal compilation © 2006 Blackwell Publishing Ltd, Molecular Microbiology, 61, 1110–1117

a role in stabilizing the patches of CL. The head-group
array formed within patches of CL might allow these mem-
brane proteins to bind specifically to the patches. The
observed CL-rich domains of NAO fluorescence in the
polar and septal regions should be rich in such micro-
domains stabilized by membrane proteins.

A growing number of proteins have been shown to

localize to the septal and polar membranes in bacterial
cells (Lybarger and Maddock, 2001; Lai et al., 2004). It is
possible therefore that some of them, having an affinity for
the head-group array of CL, might help stabilize patches
of the lipid. E. coli protein MurG, a peripheral membrane
N-acetylglucosamine transferase involved in murein
synthesis, interacts preferentially with CL, and its over-
expression results in formation of vesicles enriched with
CL at the poles of cells (van den Brink-van der Laan
et al., 2003). MurG is thus a candidate CL microdomain-
stabilizing protein. Another one is monoglucosyl diacyl-
glycerol synthase, a glucosyltransferase from Achole-
plasma laidlawii with an affinity for acidic phospholipids
that preferentially localizes to the cell poles when it is
expressed in E. coli cells (Wikström et al., 2004).

In summary, the observations and considerations

above give us a new view of bacterial membrane
surfaces. They consist of a mosaic of small patches or
microdomains of phospholipids with a specific polar head
group, the maintenance of which might be supported by
certain membrane proteins. The conditions required for
their formation are probably similar to those for the forma-
tion in eukaryotic cell membranes of sphingolipid rafts that
are promoted by both hydrogen bonding between –OH
and –NH in the polar head groups (Boggs, 1987) and the
van der Waals forces of long, saturated acyl chains
(Edidin, 1997; London and Brown, 2000; Kobayashi et al.,
2001).

Physiological roles of lipid domains

The propensity of CL and PE to form non-bilayers in cell
division and sporulation

Phosphatidylethanolamine and CL domains in B. subtilis
cells tend to occur in the same regions (Nishibori et al.,
2005). What, then, might be the roles of the CL and PE
domains in the septal membranes and in the membranes
of sporulating cells? It appears that the propensity to form
non-bilayer structures conferred by small polar head
groups (Dowhan, 1997) is essential here. At the initial
stage of cell division, the small radius of curvature of the
division site, on the leading edge, requires a lipid with a
small head group in the concave region of the outer
monolayer (Fig. 3). However, as invagination proceeds to
decrease the diameter of the ring, the constraints become
dominated by the convexity of the monolayer. However,

the packing constraints (and, hence, the nature of the
lipids) in the inner monolayer of the bilayer membranes
are totally opposite (Norris et al., 2002). Fusion and
fission of bilayer membranes might require a lipid to take
on a non-bilayer structure (Cullis and de Kruijff, 1979). If
this is correct, then cells are faced with the problem of
ensuring a supply of appropriate lipids at the division site.
It seems likely that the septal localization of phospholipid
synthases can meet this need by serving lipids of appro-
priate shape at proper times and sites during the cell
division process; indeed, this might, in fact, be the major
reason for the septal localization of the majority of the
phospholipid synthases.

In sporulating cells, PE and CL domains are also found

in the polar septa and on the engulfment and forespore
membranes. During the sporulation process, membranes
undergo dynamic transformations, including the formation
of the asymmetric septal membrane, engulfment and
finally fusion (Fig. 4). Development of these sporulation-
specific membranes probably requires that the lipids
involved be able to form non-bilayers as described above.
The significance in sporulation of CL has been demon-
strated by examining mutants lacking CL, which show
retarded emergence of polar septal and engulfment mem-
branes and a reduced frequency of heat-resistant spores
(Kawai et al., 2004). The process of activation of sigma E
and sigma F during the sporulation phase involves the
asymmetric septal membranes (Feucht et al., 1996;
Schujman et al., 1998), which should be rich in CL, and
the activation of the former has been shown to require de
novo fatty acid synthesis (Schujman et al., 1998). Thus,
some of the factors responsible for the activation, which
are on the septal membranes, might require CL to work
properly (this requirement is not necessarily the propen-
sity to form non-bilayer structures).

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e

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v

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:

T

Fig. 3. Changes in the radius of curvature of the cytoplasmic
membrane during cell division at the division site. As the contractile
ring narrows, the region of the division site changes from a
concave to a convex conformation. Shapes of phospholipids
needed for the curvatures of monolayers of the membranes in the
concave and convex structures are shown above. The right panel
is adapted from Figure 1 of Norris et al. (2002) with the publisher’s
permission.

1114

K. Matsumoto, J. Kusaka, A. Nishibori and H. Hara

© 2006 The Authors

Journal compilation © 2006 Blackwell Publishing Ltd, Molecular Microbiology, 61, 1110–1117

The significance of PE, however, has not been clarified

in B. subtilis cells as PE-defective mutant cells show no
obvious change in phenotype (Matsumoto et al., 1998).
Whatever the role of PE might be, it appears that CL or
other lipids with similar properties can replace it.

The acidic nature of CL and recruitment of membrane
proteins

The negative charge of CL recruits peripheral membrane
proteins to the membranes. This has been amply illus-
trated by the examples of DnaA, FtsY, GlpD, PssA and
SecA (Dowhan, 1997; Matsumoto, 2001; Walz et al.,
2002). Basic residues of these proteins are assumed to
interact

with

the

negative

charge

of

the

acidic

phospholipid. The MinD regulator of cell division site
selection is also included in this category of proteins
(Mileykovskaya et al., 2003; Dowhan et al., 2004) and will
be discussed briefly in the next section.

Many proteins have been localized at the polar mem-

branes (Lybarger and Maddock, 2001; Lai et al., 2004),
and some of them might also be recruited by the negative
charge of CL. It has been suggested that in the invasive
bacterium Shigella, where IcsA localized at the pole
nucleates host actin filaments to form a comet-like tail that
propels the bacterium forward to penetrate a neighbour-
ing cell, certain specific receptors exist on the membrane
at the old pole, which induce the nascent IcsA polypeptide
to be secreted there (Brandon et al., 2003; for a review
see Pugsley and Buddelmeijer, 2004). Such receptor pro-
teins and other proteins involved in similar polar targeting

might be recruited to the poles by the negative charge of
CL.

Possible relevance to regulation of cell division
machinery

The septal localization of domains enriched in particular
phospholipids suggests their relevance to the cell division
process. Besides meeting membrane curvature require-
ments and forming non-bilayer structures during the
process (as discussed above), they might be involved in
the functional regulation of the cell division machinery.
The cell division process begins with the formation of a
ring composed of polymerized FtsZ protein (Weiss, 2004).
The position of the ring formation is confined to the midcell
by a combination of nucleoid occlusion and Min systems,
which inhibit the Z-ring assembly in the vicinity of the
nucleoids and of the poles respectively.

The MinC division inhibitor is recruited to the cytoplas-

mic membrane by the ATP-bound form of the MinD
protein. In E. coli cells, the MinE topological specificity
factor drives MinD to oscillate rapidly from pole to pole
along helical arrays by stimulating the ATPase activity of
MinD. This oscillation maximizes the time-averaged con-
centration of MinC at the poles and minimizes it at the
midcell. MinD interacts preferentially with acidic phospho-
lipids (Mileykovskaya et al., 2003). The acidic phospho-
lipid content of the membrane affects the affinity of MinD
for the membrane and modulates its dynamic behaviour.
In a pssA-null mutant that completely lacks zwitterionic
PE and contains only acidic phospholipids, MinD does not
oscillate from pole to pole but forms compact spots which
randomly migrate around the thus partially division-
defective filamentous cell (Mileykovskaya et al., 2003).
This leads to the conclusion that polar localization of
CL-rich domains in the wild-type cell plays an important
role in MinD oscillation by promoting assembly of MinD
oligomers at the poles (Dowhan et al., 2004; Mileyk-
ovskaya and Dowhan, 2005), although it does not simply
define the MinD distribution in the absence of MinE, as
MinD is distributed almost evenly around the cell periph-
ery in a minE mutant. We note that the interaction of MinD
with the membrane depends on its conserved C-terminal
membrane-targeting sequence (MTS). This sequence
forms an amphipathic

a-helix with a preference for acidic

phospholipids because of basic residues on the hydro-
philic side (Szeto et al., 2003).

The FtsA protein, which functions to tether FtsZ poly-

mers to the membrane, has the conserved amphipathic
MTS in its C-terminus. This MTS also has basic residues
on the hydrophilic side of the helix and is functionally
interchangeable with that of MinD (Pichoff and Lutken-
haus, 2005). Although the question has not yet been
examined, it can be reasonably expected that FtsA also

O

A

N

-

L

C

P

F

G

-

A

s

l

C

Fig. 4. Need for non-bilayer-forming lipids during sporulation.
Dynamic rearrangement of the membranes in engulfment and
fusion is illustrated (left). Sporulation septal, engulfment and
forespore membranes are rich in CL, which has a propensity to
form non-bilayer membranes, as evidenced by NAO staining
(middle). The figure is adapted from Figure 1C of Kawai et al.
(2004) with the publisher’s permission. Localization of CL synthase
(ClsA)–GFP fusion protein on the sporulation septal and forespore
membranes (right: fluorescence images and corresponding figures
of the fluorescence are illustrated below).

Lipid domains in bacterial membranes

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Journal compilation © 2006 Blackwell Publishing Ltd, Molecular Microbiology, 61, 1110–1117

preferentially interacts with acidic phospholipids. This
might have significance for the function of FtsA, consid-
ering the localization of the CL-rich domains in the septal
region.

In B. subtilis, which does not have MinE, the topological

specificity factor of the Min system is the coiled-coil
protein DivIVA. It is targeted to the cell poles by an as yet
unknown mechanism, independent of FtsZ and other divi-
sion proteins (Harry and Lewis, 2003) and recruits MinD,
which then recruits MinC, to the poles. These proteins
persist there without oscillation and block polar division.
Surprisingly, DivIVA is targeted to the cell poles in E. coli
and even in S. pombe (Edwards et al., 2000). It has been
proposed that this protein might be attracted by a physical
property of the poles and that the targeting signal for
division sites is conserved across eukaryotes and
prokaryotes. It is intriguing that the nascent division sites
and poles of these organisms are enriched in CL and PE
(B. subtilis), CL (E. coli), and PE and sterols (S. pombe;
cf. Wachtler et al., 2003; Iwamoto et al., 2004), all confer-
ring a non-bilayer-forming propensity on the membrane.

Concluding remarks

The standard concept of the ‘fluid’ and ‘mosaic’ architec-
ture of membranes (Singer and Nicolson, 1972) that has
been depicted in the textbooks of biochemistry assumes
that lipid molecules are homogeneously distributed in
the membranes and that integral membrane proteins
resemble icebergs floating unencumbered in a two-
dimensional lipid sea. However, the evidence that lipid
molecules in eukaryotic membranes are segregated into
regions (microdomains) of specific lipid molecules or spe-
cific composition is accumulating (see the latest review,
Engelman, 2005). The view of prokaryotic membranes is
also changing to one where a mosaic of small patches or
microdomains of phospholipids with a specific polar head
group, CL and PE, is to a significant extent self-
assembled according to their respective intrinsic chemical
characteristics, and possibly maintained with the support
of certain membrane proteins. Thus, contradicting the pre-
viously assumed homogeneous distribution, bacterial
membranes are patchy with regions of specific lipid mol-
ecules and one must admit that, for lipid membranes, the
fluid is a mosaic.

Acknowledgements

We thank Professor Emeritus Isao Shibuya and Professor
Yoshito Sadaie for discussion and encouragement. We also
thank Tony Pugsley for a critical reading of this manuscript.
Thanks are also due to Yoshinori Hara, Tomohiro Hayakawa,
Toshihide Kobayashi, Eugenia Mileykovskaya, Vic Norris,
Akinori Ohta, Satoshi Shuto, Hiromu Takamatsu, Masato

Umeda, Akihiro Yoshida and Hirofumi Yoshikawa for their
helpful discussions, suggestions and encouragement. Due to
the concise nature of this Microreview, we were unable to cite
all relevant studies regarding this area of research.

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