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REVIEW ARTICLE
Compartmentalization and spatiotemporal organization of
macromolecules in bacteria
Sutharsan Govindarajan, Keren Nevo-Dinur & Orna Amster-Choder
Department of Microbiology and Molecular Genetics, IMRIC, The Hebrew University – Hadassah Medical School, Jerusalem, Israel
Correspondence: Orna Amster-Choder,
Department of Microbiology and Molecular
Genetics, IMRIC, The Hebrew University –
Hadassah Medical School, PO Box 12272,
Jerusalem 91120, Israel. Tel.: +972 2 675
8460; fax: +972 2 675 7910;
e-mail: [email protected]
Received 28 September 2011; revised 27
June 2012; accepted 28 June 2012.
Final version published online August 2012.
DOI: 10.1111/j.1574-6976.2012.00348.x
Editor: Martin Kupiec
MICROBIOLOGY REVIEWS
Keywords
cell polarity; cell compartmentalization;
subcellular organization; polar cues; RNA
targeting; localized translation.
Abstract
For many years, the bacterial cells were regarded as tiny vessels lacking internal
organization. This view, which stemmed from the scarcity of membranebounded organelles, has changed considerably in recent years, mainly due to
advancements in imaging capabilities. Consequently, despite the rareness of
conventional organelles, bacteria are now known to have an intricate internal
organization, which is vital for many cellular processes. The list of bacterial
macromolecules reported to have distinct localization patterns is rapidly growing. Moreover, time-lapse imaging revealed the spatiotemporal dynamics of
various bacterial macromolecules. Although the regulatory mechanisms that
underlie macromolecules localization in bacterial cells are largely unknown,
certain strategies elucidated thus far include the establishment of cell polarity,
the employment of cytoskeletal proteins, and the use of the membrane properties, that is, curvature, electric potential, and composition, as localization signals. The most surprising mechanism discovered thus far is targeting of certain
mRNAs to the subcellular domains where their protein products are required.
This mechanism relies on localization features in the mRNA itself and does not
depend on translation. Localization of other mRNAs near their genetic loci
suggests that the bacterial chromosome is involved in organizing gene expression. Taken together, the deep-rooted separation between cells with nucleus
and without is currently changing, highlighting bacteria as suitable models for
studying universal mechanisms underlying cell architecture.
Introduction
Clustering of functionally related molecules in a complex
system is a thermodynamically favorable process, because
such an arrangement is expected to yield optimal functioning with reduced disturbances (Zimmerman & Trach,
1991). Membrane-bounded structures, which maintain
the constituents of discrete processes together, represent a
strategy for achieving this type of organization within living systems. The main difference between the two major
domains of life, eukaryotes and prokaryotes, is that,
excluding exceptional cases, prokaryotic cells lack membrane-bounded organelles. Because the differential protein
composition of specific eukaryotic organelles reflects their
function, the scarcity of such organelles in prokaryotes
led to the general assumption that proteins and other
macromolecules are randomly distributed throughout the
FEMS Microbiol Rev 36 (2012) 1005–1022
cell. The bacterial cell wall, the membrane, and the
nucleoid were regarded as exceptions, as these domains
are known to recruit specific proteins. However, the
unexpected observation of cytosolic proteins localizing to
specific subcellular domains, without an apparent physical
structure or a membrane-surrounded organelle (Rudner
& Losick, 2010), surprised cell biologists and questioned
the traditional view of bacterial cells as bags of freely diffusing macromolecules.
Recent advancements in imaging capabilities enable better visualization of bacterial cells and hence the investigation of their internal organization and dynamics. Among
the earlier examples for bacterial proteins that have a distinct subcellular localization were the cell division protein
FtsZ and the constituents of the chemotaxis complex in
Escherichia coli. In both cases, the proteins were initially
visualized by immunoelectron microscopy, FtsZ as a
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1006
discrete ring in the center of dividing cells (Bi & Lutkenhaus, 1991) and the chemoreceptors at the cell poles (Alley
et al., 1992). Visualization of the two proteins was
improved after fusing the proteins under study to fluorescent proteins, such as GFP and yellow fluorescent protein
(YFP), and observing them in live cells (Ma et al., 1996;
Sourjik & Berg, 2000), a technique that became the leading
method for visualization of proteins in the tiny bacterial
cells. Extensive studies that combined fluorescence microscopy with genetic manipulations in three model bacteria,
E. coli, Bacillus subtilis and Caulobacter crescentus, led to
the identification and characterization of several spatiotemporal regulatory mechanisms (Shapiro et al., 2002;
Collier & Shapiro, 2007; Thanbichler & Shapiro, 2008;
Kirkpatrick & Viollier, 2010; Rudner & Losick, 2010; Toro
& Shapiro, 2010). Recently, new experimental approaches
enabled the investigation of mRNA localization in bacterial
cells (reviewed in Broude (2011)), thus revolutionizing our
view of the spatial relationship between the different stages
of gene expression in these allegedly ‘noncompartmentalized’ cells. The observations regarding proteins and RNA
localization, combined with the detection of lipid domains
within the membrane (Matsumoto et al., 2006), the complex arrangement of the chromosome (Toro & Shapiro,
2010), the active movement of the oriC element to the poles
during DNA replication (Toro & Shapiro, 2010), the nonrandom distribution of the ribosomes (Lewis et al., 2000),
and the formation of transcription foci at actively transcribed rRNA genetic locus (Davies & Lewis, 2003)
strengthen the view that clustering and compartmentalization are central themes that underlie spatiotemporal control of molecular processes in bacterial cells.
The list of bacterial macromolecules with distinct subcellular localization is increasing rapidly. Using a highthroughput method for quantitative genome-scale analysis
of protein localization in C. crescentus, Gitai et al. were
able to show that over 10% of the proteins in this
organism, around 10-fold more than previously estimated, display a nonuniform distribution (Werner et al.,
2009). Notably, the number of localization patterns that
have been observed is rather limited, with the majority of
the proteins localizing to the poles, to midcell or demonstrating a patchy distribution along the cell axis (Werner
et al., 2009). Although relatively little is known about the
precise mechanisms that establish the bacterial cell architecture, certain strategies like cell polarity, cytoskeleton
dependence, the contribution of various membrane features, and, recently, mRNA targeting were documented as
being involved in the spatiotemporal control of protein
localization and of protein complex formation. Below, we
will touch upon these mechanisms and their involvement
in macromolecule localization and compartmentalization
in bacteria.
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S. Govindarajan et al.
The involvement of the cytoskeleton in
protein localization
The discovery of bacterial cytoskeletal proteins, for example, the actin-like proteins MreB and ParM, the tubulinlike protein FtsZ, and the intermediate filament protein
crescentin, reinforced the strong similarity between prokaryotes and eukaryotes, thus highlighting the importance
of bacteria as a model system for cellular organization
(Cabeen & Jacobs-Wagner, 2010). Studies in the last decade demonstrated that many functions of eukaryotic cytoskeletal proteins, such as cell division, chromosome
segregation, organelle movement, and cell shape maintenance, are applicable to their prokaryotic homologs (Vats
et al., 2009a, b; Cabeen & Jacobs-Wagner, 2010; de Boer,
2010; Erickson et al., 2010; Jockusch & Graumann, 2011).
In addition to homologs of the eukaryotic cytoskeletal
structures, that is, actin, tubulin, and intermediate filaments, bacteria contain also distinct cytoskeletal proteins,
which are not known to have homologs in eukaryotes
(MinD and ParA), and the list of proteins that form cytoskeletal-like structures in bacteria is still growing (Vats
et al., 2009a, b; Ingerson-Mahar & Gitai, 2012).
Much of what we know about the involvement of
cytoskeletal structures in bacterial protein localization has
come from the studies of FtsZ, which assembles in
midcell to form the Z-ring, eventually leading to the
recruitment of over two dozens of proteins to form the
divisome and the mature septal ring (de Boer, 2010).
However, the role of the tubulin-like FtsZ in protein
localization, extensively reviewed over the years, has been
mostly investigated with respect to cell division. The
actin-like MreB protein, on the other hand, was suggested
to be implicated in localization of proteins that function
in diverse cellular processes, for example, cell shape
determination, cell growth and differentiation, polarity,
motility, chromosome dynamics, and even bacterial
pathogenesis (reviewed in Carballido-Lopez et al., 2006;
Cabeen & Jacobs-Wagner, 2010; Shaevitz & Gitai, 2010).
In addition to regulating protein localization, MreB and
other bacterial cytoskeletal proteins that form filamentlike structures were suggested to play an important role
in localizing large protein complexes and even organelles,
for example, the magnetosome in Magnetospirillum
magneticum (Komeili et al., 2006), the carboxysome in
Synechoccus elongates (Savage et al., 2010), the pilus in
Pseudomonas aeruginosa and Myxococcus xanthus (Cowles
& Gitai, 2010; Mauriello et al., 2010), the stalk in Caulobacter
(Wagner et al., 2005; Divakaruni et al., 2007), and inclusion bodies in E. coli (Rokney et al., 2009). However,
except for cell shape determination, the molecular mechanisms and the possible MreB-interacting partners that are
involved in the different processes are largely unknown.
FEMS Microbiol Rev 36 (2012) 1005–1022
Spatiotemporal organization of bacterial cells
Recently, the use of novel imaging technologies to document MreB localization and its linkage to the cell wall
synthetic machinery opened an interesting and timely discussion on the nature of localization patterns that are
inferred from imaging data. Hence, we chose to focus
below on MreB and its involvement in setting up the cell
morphogenesis.
MreB is widely conserved in rod-shaped bacteria and is
dispensable in several cocci (Carballido-Lopez et al.,
2006). Certain bacterial species, like B. subtilis, encode
more than one MreB paralog (Shaevitz & Gitai, 2010).
The mre locus was first identified in E. coli as encoding
genes that are involved in cell shape maintenance (Wachi
et al., 1987). The idea that MreB functions as a prokaryotic actin-like protein in determining cell morphogenesis
was suggested based on sequence similarity (Bork et al.,
1992) and supported by the in vivo observation of MreB
filamentous structures in B. subtilis (Jones et al., 2001).
The ATP/GTP-dependent polymerization of Thermotoga
maritime MreB1 into filaments in vitro (van den Ent &
Lowe, 2000) reinforced this notion. The MreC, MreD,
Pbp2, RodA, and RodZ proteins, as well as several others,
were shown to associate with the MreB cytoskeletal
structures and to be involved in cell shape determination
(Shiomi et al., 2008; Alyahya et al., 2009; Bendezu et al.,
2009; Shaevitz & Gitai, 2010; van den Ent et al., 2010).
Many studies pointed at the tight linkage between
MreB and cell wall synthesis. The mechanical stiffness
and rigidity of MreB filaments were shown to provide
additional support to the peptidoglycan layer, the major
cell wall component, in maintaining the overall cell shape
(Wang et al., 2010). Moreover, MreB and its homologs
were shown to be directly involved in cell wall synthesis.
By and large, cell wall synthesis in rod-shaped bacteria
can be divided into two stages: cell elongation, which is
linked to the localization of MreB-associated cytoskeleton,
and septal division, which is affected by the FtsZ ring
positioning. Efficient functioning of both processes
depends on the ability of these two proteins to recruit
other proteins involved in the each stage (Mattei et al.,
2010). Using fluorescently labeled vancomycin, an antibiotic that binds to peptidoglycan, cell wall elongation in
B. subtilis was shown to follow the distribution pattern of
Mbl, an MreB homolog, and was suggested to be governed by it (Daniel & Errington, 2003). Both patterns, of
MreB and its homologs and of the cell wall synthetic
machinery, were interpreted as helical structures, although
this interpretation is currently debatable in light of new
data (see below).
Recent studies in C. crescentus shed light on how the
cytoplasmic MreB can impact the assembly of the cell
wall, which occurs outside of the inner membrane. The
explanation seems to be provided by the intricate
FEMS Microbiol Rev 36 (2012) 1005–1022
1007
machinery of MreB-interacting partners, which likely
function in efficient channeling of metabolic intermediates and site-directed synthesis of peptidoglycan. In an
elaborate study, the cytoplasmic proteins MurB, MurC,
MurE, MurF, and the inner membrane proteins MurG
and MraF, all functioning in initiating peptidoglycan synthesis, were shown to exhibit a similar localization pattern
that directly depends on a network of MreB–MreD–RodA
interactions (White et al., 2010). This dependence was
validated by perturbation of the observed localization pattern, using the MreB-inhibitor A22; the interactions
between the proteins were confirmed by two-hybrid
assays. Then again, periplasmic proteins involved in the
final stages of peptidoglycan synthesis, including PBP2,
PBP3, MipA, and MltA, were shown to interact with the
transmembrane protein MreC and to exhibit a pattern
which is similar to MreB, that is, a helical path (White
et al., 2010). This study, together with other studies, suggested that the Caulobacter MreB and MreC affect peptidoglycan precursor synthesis and assembly in the
cytoplasm and in the periplasm, respectively (Figge et al.,
2004; Divakaruni et al., 2005; Dye et al., 2005). However,
despite the similar localization patterns, an apparent
interaction between MreC and MreD or MreB was not
demonstrated in two-hybrid experiments (White et al.,
2010). Nevertheless, because such interactions were
reported to occur in B. subtilis (Kruse et al., 2005), it is
still possible that MreB, MreC, and MreD, together with
other interacting proteins, create a common interface for
the cytoplasmic and periplasmic peptidoglycan synthesizing proteins. Intriguingly, in some species, MreB localization depends on MreC, whereas other species have either
MreB or MreC (Kruse et al., 2005), suggesting the
existence of species-specific machineries for cell wall
synthesis.
Recently, a more detailed picture of the subcellular
organization of MreB and the cell wall synthetic machinery has been obtained by three independent studies that
imaged the movement of MreB structures and found it to
be inconsistent with the previous description of MreB as
a continuous helix that extends from pole to pole. Using
high-precision particle tracking (Garner et al., 2011) and
total internal reflection fluorescence microscopy (Dominguez-Escobar et al., 2011), the MreB in B. subtilis was
shown to form discrete patches that move processively
and circumferentially along helical tracks, perpendicularly
to the cell axis. The directional motions of MreB in the
B. subtilis cells were often discontinuous and independent, inconsistent with the existence of a coherent, longrange MreB cytoskeleton. Similarly, the E. coli MreB was
also shown to display heterogeneous spots that rotate
around the long axis of the cell in a persistent manner
(van Teeffelen et al., 2011). The observed localization
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S. Govindarajan et al.
1008
pattern was consistent with the lack of long helical filaments also in this case. Significantly, the dynamic rotational movement of the MreB patches was shown in these
studies not to be caused by MreB polymerization, but
rather by the force exerted by cell wall synthesis. A similar
rotational motion was observed also for components of
the cell wall synthesis complexes. Together, these studies
revealed a mobile, fragmented picture of MreB, rather
than a contiguous helical structure. On the other hand, a
very recent study demonstrated that the chirality of MreB
in rod-shaped bacteria gives rise to a chiral insertion of
peptidoglycan into the cell wall and causes cells to twist
in the same handedness as their MreB during elongational
growth (Wang et al., 2012). Together, these new studies
opened a discussion about other bacterial proteins, whose
organization has previously been interpreted as continuous helices. Additional analyses are required to unequivocally determine to what extent the interpretation of a
continuous vs. a patchy and discontinuous filament is
methodology dependent or, in fact, represents different
patterns that exist in the bacterial cell. The clusters comprising of cytoskeletal elements and the peptidogylcan
synthesizing proteins that move along helical tracks are
schematically presented in Fig. 1.
Quite unexpectedly, partition of MreB in the two
daughter cells during cytokinesis of E. coli, C. crescentus,
and Rhodobacter sphaeroides was shown to involve the
formation of an MreB medial ring in predivisional cells,
which then relocates to the original structures just before
completion of the division process (Figge et al., 2004;
Gitai et al., 2004; Slovak et al., 2005; Vats & Rothfield,
2007). The E. coli MreB cytoskeletal ring was shown to
contain also the MreB-interacting proteins MreC, MreD,
Pbp2, and RodA proteins. Whereas the MreB, MreC,
MreD, and RodA were each able to independently assemble into the cytoskeletal ring, incorporation of Pbp2 into
the ring required MreC (Vats et al., 2009b). This dynamic
process, which is triggered by the membrane association
of the FtsZ cell division protein, apparently leads to equal
partition of MreB and the other cytoskeletal proteins
between the daughter cells.
Summarily, studies thus far established the central role
of bacterial cytoskeleton proteins in determining cell
morphology and in maintaining cell shape. Additional
studies are required to elucidate their involvement in the
spatiotemporal regulation of other processes.
Membrane properties affecting protein
localization
Accumulating evidence for compartmentalization and
nonrandom localization of proteins in bacterial cells suggests the existence of prelocalized cues, which affect the
localization of proteins to certain subcellular domains.
Proteins that are not known to require other proteins for
their localization may be useful for the identification of
these cues. Below, we discuss studies of such proteins,
which have identified certain properties of the cytoplasmic membrane, that is, membrane curvature, membrane
potential, and lipid composition, as localization signals.
Membrane curvature
Fig. 1. Schematic presentation of the cytoskeleton-associated cell
wall synthetic machinery. Shown at the bottom is a helix-like patched
arrangement of clusters (pink balls), which consist of cytoskeletal and
cell wall synthesizing proteins. The overblown cone at the top depicts
a single cluster, composed of cytoplasmic, inner membrane, and
periplasmic proteins, which assemble via intricate successive
interactions with synthesize peptidoglycan.
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Recent studies from Losick’s laboratory identified membrane curvature, which is reflected in cell shape and architecture, as a physical cue for protein localization.
Significantly, both concave (negative curvature) and convex (positive curvature) shapes can act as beacons for
localization of different proteins. The nonuniformity in
membrane curvature is schematically shown in Fig. 2. The
first protein, whose localization was shown to be determined by membrane curvature, was the peripheral membrane protein SpoVM, a small amphipathic alpha helix
peptide, expressed exclusively in the mother cell during
B. subtilis sporulation (Ramamurthi et al., 2006). SpoVM
is tethered to the spore surface and, along with SpoIVA,
forms the spore coat complex. Because genetic and biochemical techniques did not identify any other protein as
necessary for SpoVM localization, it was assumed that
FEMS Microbiol Rev 36 (2012) 1005–1022
1009
Spatiotemporal organization of bacterial cells
(a)
(b)
(c)
Fig. 2. Differences in membrane curvature. (a) In a dividing rodshaped Escherichia coli cell, the highest negative curvature is at the
poles (concave), whereas the constricting midcell marks sites with
moderate negative curvature. (b) In a rod-shaped dividing Bacillus
subtilis cell, the division septum is formed is a relatively sharp angle
relative to the cell axis. Hence, the highest negative curvature is near
the forming septum, whereas the hemisphere poles are characterized
by moderate negative curvature. (C) In a sporulating B. subtilis cell,
the outer surface of the forespore is the only positively curved surface
(convex) in the forespore-engulfing mother cell.
SpoVM recognizes a nonchemical factor that is found
exclusively in the forespore wall. A strong candidate for
such a cue was the positive curvature of the forespore
membrane, because this type of topology is not present
anywhere else in the sporulating cell (Fig. 2c). Imaging of
GFP-tagged SpoVM in cell division–defective mutants
reinforced the notion that positive curvature acts to recruit
SpoVM to the spore coats. In a mutant producing incomplete polar septum and hence devoid of convex-shaped
positive membrane, SpoVM-GFP localizes indiscriminately
to the inner membrane of the mother cell and the outer
membrane of the forespore. SpoVM sensing of positive
curvature was also validated in vitro by its specific binding
to the outer surface of liposomes and in vivo by observing
SpoVM localizing to positively curved membrane structures of the heterologous cells of E. coli and Saccharomyces
cerevisiae. All these results implied that SpoVM is capable
of sensing positive membrane curvature for its localization
(Ramamurthi et al., 2009).
Can negative membrane curvature also act as a cue for
protein localization? This question was addressed by two
research groups that investigated the subcellular localization of DivIVA. The DivIVA protein of B. subtilis is
involved in two important processes, namely, prevention
FEMS Microbiol Rev 36 (2012) 1005–1022
of polar septum formation (Bramkamp et al., 2008) and
segregation of the chromosomal DNA during sporulation
(Ben-Yehuda et al., 2003). Several types of evidence suggested that DivIVA might recognize the negative curvature of the membrane. First, DivIVA-GFP was observed
mainly at the division septa, but also at the poles of
B. subtilis cells (Fig. 2b), and secondly, DivIVA localized
mainly at the poles of heterologous E. coli (Fig. 2a) and
Schizosaccharomyces pombe cells (Lenarcic et al., 2009;
Ramamurthi & Losick, 2009; Eswaramoorthy et al., 2011).
Notably, as opposed to E. coli cells, in which the strongest
negative curvature is at the poles, in B. subtilis cells, the
forming septa, which later becomes the new pole, displays
even sharper membrane curvature than the existing poles
(Ramamurthi, 2010). To further investigate this issue, the
subcellular localization of GFP-tagged DivIVA was monitored under conditions that affect the negative curvature
of B. subtilis membrane. When B. subtilis was grown as
single cells rather than chains, DivIVA was observed only
at the poles. A similar localization pattern was observed
upon prevention of septum formation. Moreover, in cells
that were made spherical, having uniform negative curvature, by lysozyme treatment, DivIVA localized indiscriminately (Lenarcic et al., 2009; Ramamurthi & Losick,
2009). Taken together, certain bacterial proteins are capable of sensing a physical cue, such as membrane curvature, and localize accordingly.
Membrane potential
The inner membrane serves as a preferred subcellular
compartment for protein localization in bacteria. Whereas
the membrane composition and structural features have
been given a lot of attention, one cannot rule out that the
proton motive force (pmf) generated in the membrane
also affects subcellular organization. Generation of pmf in
the membrane is a robust process, which involves
exchange of ions, creation of electric potential, and subsequently, synthesis of the main energy molecule of the cell,
ATP (Ingledew & Poole, 1984). Could membrane potential
affect the localization of proteins to membrane surfaces?
A recent investigation by Strahl & Hamoen (2010)
revealed that pmf might play a significant role in protein
localization. The authors discovered it by serendipity,
when they observed that the use of polylysine-coated
slides resulted in loss of MinD polar localization. Because
it has previously been reported that polylysine may affect
the pmf of bacterial membranes (Katsu et al., 1984), the
possible role of pmf in protein localization was investigated by monitoring localization of several proteins in the
presence of ionophores that dissipate pmf. Indeed, dissipation of pmf led to promiscuous localization of B. subtilis
proteins that are involved in cell division (FtsA, MinCD),
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S. Govindarajan et al.
1010
cell shape regulation (MreB, Mbl, MreBH, and MreCD)
and chromosome segregation (Soj). The observed
mislocalization of certain proteins in E. coli and C. crescentus
because of pmf dissipation indicated that the involvement of pmf in protein localization is widespread among
bacteria.
How does pmf affect protein localization? Because the
proteins that exhibited delocalization upon pmf dissipation
by ionophores require ATP for their activity, and because
such a treatment can result in a significant decrease in ATP
levels in the cell, a drop in ATP could theoretically account
for the observed effect on protein localization. By repeating
the experiments in strains impaired in ATP synthesis or
with non-ATP-binding mutant proteins, the effect of pmf
was shown to be independent of the ATP levels in the cell
(Strahl & Hamoen, 2010). The next question to be
addressed was which of the two components of pmf, transmembrane electric potential (DΨ) or transmembrane
chemical potential (DpH), is involved in protein localization. Using ionophores that specifically modify each pmf
component, it is has been shown that DΨ is the key factor
that affects localization of proteins. How does DΨ influence
protein localization? Dissection analysis of MinD revealed
that its C-terminal amphipathic helix is sensitive to disturbance of membrane potential. Moreover, binding of this
amphipathic helix to lipids was shown to depend on DΨ in
vitro, raising the possibility that a change in membrane
potential results in a subtle conformational change of the
amphipathic helix that enhances its binding (Strahl &
Hamoen, 2010). On the other hand, membrane fluidity,
previously shown to depend on DΨ (Schaffer & Thiele,
2004), has also been demonstrated to affect MinD binding
(Mazor et al., 2008a), suggesting that membrane potential
might change the membrane lipid properties, which facilitates the insertion of amphipathic helices. Further studies
are required to elucidate the exact molecular mechanism
that underlies the effect of DΨ on protein localization.
Finally, protein localization was shown to be affected
by depletion of oxygen, which is the major electron
acceptor that influences membrane potential (Strahl &
Hamoen, 2010). The authors demonstrated that the morphological changes, which depletion of oxygen induces in
bacterial cells, correlate with delocalization of cell division
and cytoskeletal proteins. These results suggest that the
changes in bacterial cell morphology observed in natural
habitats, in which oxygen levels fluctuate constantly, can
be explained by the effect of membrane potential on
protein localization.
Membrane lipid composition
Studies on bacterial membrane properties revealed that
the phospholipids are not homogenously distributed.
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Rather, some of them were observed as microdomains
within the membrane, which can be compared to the
lipid rafts in eukaryotes. This discovery was made possible by the use of efficient lipid-binding fluorescent dyes
(Matsumoto et al., 2006). Two well-characterized lipids
that form distinct microdomain in bacterial membranes
are cardiolipin and phosphatidylglycerol (PG). Whereas
cardiolipin was shown to concentrate in the poles and
septum of E. coli cells (Fig. 3a) (Mileykovskaya & Dowhan, 2009), PG was observed as spirals in B. subtilis cells
(Fig. 3b) (Barak et al., 2008).
The lateral heterogeneity of lipids in the bacterial
membranes, prompted scientists to examine the possibility that this arrangement acts as a mechanism for protein
localization, that is, certain proteins localize by binding to
specific lipids. ProP, an osmosensory transporter protein
from E. coli, was the first protein shown to depend on
the polarly concentrated cardiolipin for its localization
(Romantsov et al., 2007). It has been shown that increasing osmolarity increases the cardiolipin content in the
poles; this cardiolipin microdomain recruits ProP to
the poles; ProP then functions to import osmolytes into
the cell. Finally, polar localization of ProP was shown to
be independent of its expression level but correlated with
the proportion of polarly localized cardiolipin (Romantsov
et al., 2007). The same research group that studied ProP
(a)
(b)
Fig. 3. Nonrandom distribution of lipids in the bacterial membrane.
(a) Schematic presentation of cardiolipin (pink phospholipids)
enrichment at the bacterial cell poles. Cardiolipin was suggested to
promote polar localization of the ProP and MscS cytoplasmic
membrane proteins in Escherichia coli (blue balls). (b) The scheme
depicts the helical distribution of PG within the bacterial inner
membrane.
FEMS Microbiol Rev 36 (2012) 1005–1022
1011
Spatiotemporal organization of bacterial cells
polar localization later identified MscS, an E. coli membrane protein that forms a part of the mechanosensitive
channels, also as a cardiolipin-dependent polar-localizing
protein (Romantsov et al., 2010). Figure 3a illustrates the
mechanism of cardiolipin-dependent polar localization of
ProP and MscS.
Lipid composition was suggested to be involved also in
the spatial organization of the Min proteins, which are
involved in the inhibition of polar septum formation
(Mileykovskaya et al., 2003). It has long been known that
MinCDE proteins of E. coli oscillate from pole to pole in
a dynamic manner (Vats et al., 2009a, b; Lenz & Sogaard-Andersen, 2011). Several studies of the Min system
indicated that the lipid composition of the membrane has
a direct influence on dynamic localization of the Min
proteins and hence on the regulation of cell division. The
dimeric ATP-bound form of the MinD protein, which
tethers to the membrane through its C-terminal amphipathic alpha helix, designated membrane-targeting
sequence (MTS), is stabilized by anionic lipids (Mazor
et al., 2008a, b). The pole-to-pole oscillation of MinD
follows a helical path, and it has been shown by fluorescence resonance energy transfer analysis that the MinD–
GFP helices co-localize with the spirals of the anionic
phospholipid PG in B. subtilis cells. Furthermore, MinD
spirals were not observed in mutants lacking PG (Barak
et al., 2008). MinE, another Min protein, which activates
MinD ATPase activity and thereby releases MinD–ADP
from the membrane, was also recently shown to bind
directly to the membrane by electrostatic interactions
(Hsieh et al., 2010).
Other proteins that whose localization was shown to
depend on lipid composition include DnaA (Sekimizu &
Kornberg, 1988), FtsA (Barak et al., 2008), and the Sec
translocon system (Gold et al., 2010). Because DnaA and
FtsA also possess an MTS domain, it was assumed that
binding of these proteins to the anionic-rich membrane
follows a similar pattern as that of MinD (Barak et al.,
2008). In the case of the Sec system, cardiolipin was
shown to not only promote dimerization of SecYEG, but
to also increase the ATPase activity of SecA (Gold et al.,
2010). Finally, diverse bacteria were shown to harbor
homologs of flotillins, which are ‘raft markers’ found
exclusively in lipid rafts within the membrane of eukaryotic cells (Zhang et al., 2005; Donovan & Bramkamp,
2009; Lopez & Kolter, 2010). YuaG, a flotilin homolog
from B. subtilis, renamed FloT because of its similarity to
the eukaryotic flotillin-1, was shown to localize in discrete
foci along the bacterial membrane in a dynamic fashion.
Although YuaG co-isolates with anionic phospholipids,
its in vivo localization was shown not to depend exclusively on these phospholipids (Donovan & Bramkamp,
2009). The lipids associated with the bacterial rafts were
FEMS Microbiol Rev 36 (2012) 1005–1022
suggested to be polyisoprenoids (Lopez & Kolter, 2010).
Components of various physiological processes, such as
biofilm formation, protein secretion, and the onset of
sporulation, were proposed to be organized in lipid rafts
(Donovan & Bramkamp, 2009; Lopez & Kolter, 2010).
Taken together, the data that accumulated thus far suggest that lipid microdomains are important higher-order
structures that directly affect localization of certain proteins, thus impinging on the regulation of diverse cellular
processes in bacteria.
Unidentified polar cues for protein
recruitment
Cellular asymmetry is critical for the life cycle of many
types of bacteria. Cell polarity, which apparently underlies
the establishment of asymmetry, is often visually manifested by the existence of polar organelles, such as pili or
flagella (Ebersbach & Jacobs-Wagner, 2007; Dworkin,
2009; Shapiro et al., 2009). Alternatively, spatial and
temporal pole-to-pole oscillations of proteins can establish polarity, for example, the Ras-like G proteins that set
up the polarity of motility proteins in M. xanthus cells
(Kirkpatrick & Viollier, 2010; Lenz & Sogaard-Andersen,
2011). Polarity is also established in cells lacking discernible polar organelles, such as E. coli cells, in which
specialized protein complexes localize at or near the
poles, despite their apparent symmetrical morphology.
Polarization in these cells, which stems from the different
division history of two poles (old vs. new), is seemingly
exploited for the positioning of protein complexes
(Janakiraman & Goldberg, 2004). The list of proteins and
complexes that localize to the poles of rod-shaped bacteria is growing rapidly, emphasizing the importance of this
domain for cell architecture and organization. Polar
positioning of many of these proteins was shown to be
crucial to fundamental cellular processes and for orchestrating sophisticated regulatory processes (Kirkpatrick &
Viollier, 2010). For example, it has recently been suggested that the polarly localized histidine kinase CckA in
C. crescentus is regulated in a protected zone near the
pole, without the use of membrane-enclosed compartments (Tsokos et al., 2011). Spatial regulation is also of
importance for bacterial pathogenesis, as many virulence
factors localize to the poles (Goldberg et al., 1993; Scott
et al., 2001; Judd et al., 2005; Jain et al., 2006; Jaumouille
et al., 2008; Carlsson et al., 2009; Bowman et al., 2010).
Although several of the polarly localized proteins were
suggested to be positioned by one of the localization
mechanisms discussed above, a large number of proteins
are targeted to the poles by a yet unidentified mechanism
(s). Careful examination of the list of polarly localized
proteins reveals that some of them belong to closely
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S. Govindarajan et al.
1012
related signaling pathways, which need to communicate
to generate an optimal cellular response. Two such linked
pathways, which exhibit a similar spatial organization, are
the phosphotransferase (PTS) and the chemotaxis systems
(see schematic presentation of the spatial organization of
both systems in Fig. 4). The chemotaxis system senses
nutrients or repellents availability in the environment and
controls bacterial taxis along chemical gradients accordingly (Porter et al., 2011). The PTS system determines the
hierarchical uptake of carbon sources from the environment and consequently adjusts cell metabolism via the
control of global regulatory systems, such as carbon
catabolite repression and inducer exclusion (Deutscher,
2008). Together, the PTS and chemotaxis systems sense
various nutrients and determine the metabolic status of
the cell. Both systems were extensively investigated, but
the mechanism that underlies their cross talk remained
unknown. A hint about their cooperativity was provided
by the suggested interaction between key components of
(a)
the two systems, that is, the general PTS protein EI and
the chemotaxis histidine kinase CheA (Lux et al., 1995).
Knowledge that accumulated in recent years regarding the
subcellular localization of the components of both systems suggests a spatial rationale for their cross talk
(Lopian et al., 2010; Amster-Choder, 2011).
The chemotaxis system of E. coli is one of the best
studied signal transduction systems, and the proteins that
compose this pathway were among the first to be
observed at the bacterial cell poles. Detailed information
regarding the interactions and the mechanism underlying
chemotaxis has been reviewed elsewhere (Roberts et al.,
2010; Sourjik & Armitage, 2010). Here, we will highlight
only the information that accumulated regarding the
localization strategies of the chemotaxis proteins. The
E. coli chemotaxis system is comprised of five different
transmembrane chemoreceptors (Tar, Tsr, Tap, Trg and Aer),
a histidine kinase (CheA), an adaptor protein (CheW), a
response regulator (CheY), a phosphatase (CheZ), a methyl
(b)
Fig. 4. Spatial organization of the PTS and the chemotaxis systems in Escherichia coli. (a) Schematic presentation of the PTS sugar transport
system. The general PTS proteins, EI and HPr, localize to the E. coli cell poles (top). Upon the addition of PTS sugars to the growth medium, HPr
is phosphorylated by EI; HPr-P is released from the poles and distributes in the cell (bottom); a fraction of HPr-P gets near the membrane to
phosphorylate the PTS sugar permeases, allowing them to translocate the PTS sugars into the cell and to phosphorylate them. (b) Schematic
presentation of the chemotaxis system. The methyl-accepting chemotaxis proteins (MCPs) form large clusters at the E. coli cell poles. The
chemotaxis proteins (for simplicity, only CheW, CheA, and CheY are shown) associate with the polar MCP clusters to form the chemotaxis
complexes (top); the small response regulator CheY is phosphorylated by the histidine kinase CheA; CheY-P migrates from the poles to the
membrane, where it interacts with proteins in the flagellar motor (bottom). Binding of CheY-P to the flagellar motor changes the rotation from
counterclockwise to clockwise and hence the swimming mode from ‘run’ (top) to ‘tumble’ (bottom).
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FEMS Microbiol Rev 36 (2012) 1005–1022
1013
Spatiotemporal organization of bacterial cells
transferase (CheR), and a methyl esterase (CheB). Orchestration of all these proteins’ activity determines the
locomotion directionality of the bacterium (Fig. 4b).
Extensive studies revealed that this coordination is
achieved by clustering the chemotaxis proteins at the
poles. Clusters of the chemoreceptors, coupled with
CheW and CheA, form the core of the chemotaxis polar
complex (Sourjik, 2004). Localization of the other Che
proteins was known to be mediated by their association
with the core complex, but the mechanism that directs
the core complex to the poles remained an enigma (Sourjik & Armitage, 2010). Recently, two models were suggested to explain the formation of the polar complex.
One model suggests that stochastic self-assembly of receptors is the mechanistic reason for cluster formation.
According to this model, receptor clusters are formed
either by the assembly of newly synthesized receptors with
already existing clusters or by the formation of new
nucleation centers at places of high concentration of the
newly synthesized proteins. The interplay between these
two assembling patterns was proposed to determine the
spatial distribution of receptor clusters in the membrane
(Thiem & Sourjik, 2008). It was also suggested that the
localization of lateral clusters marks the future division
sites and hence the poles to be of the bacterial cell
(Thiem et al., 2007). The stochastic self-assembly model
was also supported by visualization of cluster assembly
using super-resolution light microscopy (Greenfield et al.,
2009). The second model suggests that an active process
of partitioning, rather than self-assembly is involved in
the localization of the chemoreceptor. Time-lapse analysis
of newly synthesized Tsr, a major chemoreceptor, fused
with GFP revealed that the receptors are first synthesized
in a helical fashion in the membrane; these newly synthesized receptors then migrate within the membrane to
form large clusters at the poles. The helically distributed
lateral receptor clusters co-localizes with the subcellular
spirals of the Sec proteins of E. coli, suggesting that the
Sec secretion system is involved in spatial partitioning of
the chemoreceptors (Shiomi et al., 2006). Further studies
using FRAP analysis revealed that the chemotaxis polar
clusters are relatively stable over the chemotactic signaling
period, with few components, mainly CheY, exhibiting a
dynamic behavior and shuttling between the polar complex and the flagellar motors, which are randomly distributed throughout the E. coli membrane (Schulmeister
et al., 2008).
In the case of the PTS system, the two general PTS
proteins, EI and HPr, were recently shown to localize to
the cell poles. Studies of ectopically expressed EI and HPr
in a pts deletion mutant revealed that their polar localization is independent of each other. The EI polar foci are
relatively stable, whereas HPr was observed to relocate
FEMS Microbiol Rev 36 (2012) 1005–1022
from pole to the cytoplasm upon sugar stimulation. HPr
relocation explains its ability to transfer a phosphoryl
group from EI to the soluble domain of the different
permeases of PTS sugars, which either localize near the
cell membrane or are distributed throughout the cytoplasm (Fig. 4a) (Lopian et al., 2010). The PTS system was
also shown to cross talk with other regulatory systems by
spatial modulation of their components (Lopian et al.,
2010). The cues that recruit the PTS components to the
poles are not known. Moreover, the factors that are
responsible for limiting the diffusion of the soluble EI
protein and for maintaining it at the pole, subsequent to
its recruitment to this domain, are yet to be identified.
The picture that emerges from the studies described
above is that the PTS and the chemotaxis systems utilize
a similar strategy for information processing and signal
transduction, that is, they localize their respective ‘command centers’ to the poles and rely on small proteins,
HPr and CheY, respectively, that shuttle between the
poles and other cellular domains, where they execute their
function (Fig. 4). It is reasonable to assume that the similar spatial organization of the two systems provides a
basis for their cross talk, that is, their spatial proximity
may facilitate communication between their components
to coordinate their actions and generate an optimal metabolic response (Amster-Choder, 2011). It remains to be
studied whether recruitment of the chemotaxis and PTS
systems to the poles is interdependent.
Localized translation
The question of where translation of mRNA transcripts
takes place in the cell has been investigated over the
years, with novel techniques changing our understanding
of this issue time and again. The first observation which
suggested that translation might occur at specific cellular
compartments was the detection of ribosomes bound to
the membrane of the endoplasmic reticulum (ER) half a
century ago (Palade, 1958). In the 1970s, Sabatini et al.
detected mRNA molecules that remained attached to the
ER membranes even after the removal of the ribosomes
(Lande et al., 1975). On the basis of these results, it has
been suggested that mRNA is involved in assembling
membrane-bound polysomes, which actively synthesize
proteins that are secreted via the ER membrane (Lande
et al., 1975). In the early 1980s, evidence for the signal
recognition particle (SRP) model accumulated. These
studies suggested that many proteins localize via a signal
peptide to the ER (Walter & Blobel, 1980; Gilmore et al.,
1982; Walter & Blobel, 1982) or to the inner membrane
(Poritz et al., 1988; Bernstein et al., 1989; Romisch et al.,
1989), usually while the nascent chain is still associated
with the ribosome. The SRP model established a link
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1014
between mRNA localization and protein targeting and
encouraged scientists to pursue imaging of intracellular
distribution of mRNAs. An asymmetric distribution of
specific mRNAs in the cytoplasm was first visualized in
the early 1980s by in situ hybridization to detect b-actin
mRNA in ascidians embryos (Jeffery et al., 1983). In the
last 10 years, monitoring mRNA trafficking in live cells
became possible because of the development of RNA
labeling methods by fluorescent RNA-binding proteins
(Bertrand et al., 1998; Beach et al., 1999). Hence, the current view is that intracellular localization of mRNA is a
common mechanism to regulate spatial and temporal
protein expression in most eukaryotic model organisms,
thus controlling the development and physiology of these
cells (Donnelly et al., 2010).
Is the concept of localized translation relevant for
prokaryotes, or did the mechanisms that enable these
pathways coevolve with the nucleus? In fact, protein
localization via mRNA targeting has not been considered
to occur in bacteria because of the lack of nucleus and to
the coupling between transcription and translation
(Ramamurthi, 2010). Therefore, although components of
the bacterial mRNA degradosome machinery, as well as
the ribosomes, were shown to have specific intracellular
distributions (Herskovits & Bibi, 2000; Taghbalout &
Rothfield, 2007; Khemici et al., 2008), the localization of
mRNA in bacterial cells remained poorly characterized.
Very recent studies of mRNA localization in bacterial cells
shed new light on this long-neglected field. These studies
made use of the main tools that were used for visualizing
mRNA in eukaryotic cells, which were adapted to bacterial cells. Below is a short description of these tools, their
adaptation to bacteria, and the novel findings regarding
mRNA localization in bacterial cells.
Methods for studying mRNA localization
The two main tools for mRNA visualization are fluorescent nucleic acid probes, which enable the direct detection of specific nucleic acid sequences (fluorescent in situ
hybridization, FISH), and fluorescent RNA-binding proteins, which mark the location of transcripts that are
bound by these proteins. Owing to the improvement in
imaging technologies and the development of better fluorescent reagents, both methods have been recently applied
to detect RNA in bacterial cells, despite the inherent limitations, that is, the small cell size and the low copy number of mRNAs per cell (five copies or fewer) (Taniguchi
et al., 2010).
The in situ hybridization technique has been extensively
used over the years to visualize DNA and later RNA with
ever-increasing sensitivity. Methods for improved labeling
allowed direct detection of RNA with fluorescent singleª 2012 Federation of European Microbiological Societies
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S. Govindarajan et al.
strand probes (Kislauskis et al., 1993). The synthesis of
locked nucleic acid significantly increased the hybridization capabilities of the probes and allowed for a higher
sensitivity and specificity (Obika et al., 1997; Koshkin
et al., 1998). In recent years, a large variety of indirect
and direct labeling tools have been developed, allowing
one to choose from a broad spectrum of detection methods.
Compared with FISH, the methodology that enables
monitoring of mRNA trafficking in live cells is fairly
new. It is based on the use of RNA-binding proteins,
which are fused fluorescent proteins, and requires tagging of the target RNA with an array of aptamers or
binding motifs. Binding of the fluorescent RNA-binding
protein to its binding sites on the target RNA enables
the visualization of the RNA-protein complex. The most
widely used RNA-binding protein is the MS2 phage coat
protein, which in many cases is simply referred to as
MS2 protein. This method was first applied to visualize
the Ash1 mRNA in yeast (Bertrand et al., 1998; Beach
et al., 1999). Visualization of mRNA by the MS2 protein
was adjusted to E. coli by Golding and Cox, who fused
a tandem dimer of the MS2 protein to the N-terminus
of an enhanced version of GFP and placed it under the
control of an inducible promoter (Golding & Cox,
2004).
The background signal, which is generated by the use
of intact fluorescence proteins to label RNA, triggered the
development of several bimolecular fluorescence complementation (BiFC) systems. The BiFC methodology is
based on formation of a fluorescent complex through the
association of two fragments of a fluorescent protein,
which are brought together because of interaction
between proteins that are fused into these fragments. This
methodology, developed to monitor protein–protein
interaction (Hu et al., 2002; Michnick, 2003), has been
adjusted for observing RNA–protein interaction by Rackham and Brown (Rackham & Brown, 2004). In the latter
case, the two fragments of a spilt fluorescence protein are
fused into two different RNA-binding proteins. Two adjacent sets of aptamers, which are recognized by the RNAbinding proteins, are then introduced into the target
RNA. Interaction of the RNA-binding proteins with their
target RNA sites brings the two fragments of the spilt
fluorescence protein together, close enough to reassemble
and to generate a fluorescent signal. This variation in the
BiFC methodology, which was described by Rackham and
Brown as a trimolecular fluorescence complementation
(TriFC) system, has been further developed over the last
few years and was adapted to several organisms including
bacteria (Valencia-Burton et al., 2007; Yiu et al., 2011). A
more detailed description of this methodology has been
reviewed elsewhere (Broude, 2011).
FEMS Microbiol Rev 36 (2012) 1005–1022
Spatiotemporal organization of bacterial cells
The major advantage of in situ hybridization is that the
targets are nonmodified RNA or DNA; its drawback is
that it can only be used to image dead and fixed cells and
therefore does not allow monitoring of real-time dynamics of the nucleic acids. The main advantage of labeling
RNA by fluorescent proteins is that it allows viewing of
mRNA in live cells and therefore enables real-time measurements of the target RNA dynamics; the drawbacks are
that the target RNA is modified, and an array of protein
molecules is being attached to it. Hence, mRNA localization should ideally be studied by comparing data
obtained with the two approaches.
RNA localization in bacteria
The first to look at mRNA in bacteria were Golding &
Cox (2004), who used the MS2 system to study the
motion of model RNAs. These authors found that RNA
motion is limited, in most cases, in space to near the center or the quarter points of the cell. This localized motion
is consistent with Brownian motion of RNA polymers
tethered to their template DNA, which in this case was
an F-plasmid, previously shown to be distributed at the
center and quarter points of the E. coli cell (Gordon
et al., 1997; Pogliano, 2002). Occasionally, some RNAs
exhibited a polymer chain dynamics that can be ascribed
to transcription elongation, whereas in other cases, the
transcripts were observed as freely diffusing in the cytoplasm (Golding & Cox, 2004). However, the histogram of
the RNA location implied that in the latter case the
mRNAs lingered near the poles (Golding & Cox, 2004).
This behavior can be explained by hydrodynamic coupling between the RNA and the cell wall, which would be
expected to give an effective pair potential and to
decrease the diffusion rate, causing the transcripts to
spend more time near the cell poles (Lin et al., 2002;
Golding & Cox, 2004). In a later study, Broude et al. used
the elF4A split protein, a variation in the BiFC methodology, to explore the localization of three different RNAs in
E. coli, encoding lacZ, 5S rRNA, or a short artificial
untranslated RNA (Valencia-Burton et al., 2007). The
three mRNAs were encoded from the same plasmid, but
showed different localization patterns: the lacZ mRNA
was distributed evenly in the cytoplasm, the 5S RNA
transcripts localized to foci, similarly to ribosomes localization to sites outside the nucleoid, and the short
untranslated RNA was observed primarily at the cell
poles. Moreover, using time-lapse imaging, the localization pattern of the untranslated RNA transcripts was
shown to change dramatically over time. The above studies were pioneering in the field of mRNA localization in
bacterial cells and demonstrated that mRNAs can diffuse
and specifically localize in these cells.
FEMS Microbiol Rev 36 (2012) 1005–1022
1015
Further demonstration of mRNA localization in bacteria was provided by three thought-provoking studies,
published in 2009. In the first, Broude et al. extended
their study of RNA localization in live E. coli cells by the
use of the elF4A split protein. In addition to localization
of RNA at the mid and quarter points of the cell, they
observed RNA that localized laterally along the cell axis,
in a pattern suggesting the existence of ordered helical
RNA structures (Valencia-Burton et al., 2009). In the second study, Russel and Keiler used FISH to investigate for
the first time the localization pattern of an endogenous
bacterial RNA, the tmRNA in C. crescentus. The tmRNA,
a ubiquitous small RNA, which participates in transtranslation by entering the ribosome and helping with the
addition of a peptide tag on a nascent polypeptide, was
shown to localize in a helix-like pattern within the cell.
The same localization pattern was revealed for the
tmRNA-binding protein, SmpB, fused to GFP. SmpB
demonstrated a helical pattern also in the absence of
tmRNA, but not vice versa, suggesting that SmpB is
responsible for the localization of tmRNA (Russell &
Keiler, 2009). In the third study, Schleifer et al. documented
the localization of nifH, which codes for the enzyme dinitrogenase reductase that acts in the process of dinitrogen
fixation. The nifH RNA, detected by FISH in Klebsiella
oxytoca and Azotobacter vinelandii cells, exhibited an
uneven distribution, especially in K. oxytoca cells, in
which the mRNA localized to one or both poles (Pilhofer
et al., 2009). This localization pattern might correlate
with the subcellular regions in which dinitrogen fixation
takes place, suggesting localized translation of NifH via
targeting of the nifH RNA transcripts. Together, these
three studies reinforced the notion that RNA is not
randomly localized in bacterial cells.
Recently, two different models for mRNA localization
in bacteria were suggested, based on the use of FISH and
live cell imaging (Fig. 5a and b). Jacobs-Wagner et al.
imaged several mRNAs expressed from the chromosome
of C. crescentus, as well as lacZ transcripts in E. coli
(Montero Llopis et al., 2010). They observed very limited
dispersion of the transcripts from their site of transcription during their lifetime (Fig. 5b). It is worth mentioning that the E. coli lacZ distribution observed in this
study differed from the pattern observed by Broude et al.
(Valencia-Burton et al., 2007), maybe due to differences
in the mRNA expression systems, from the chromosome
or from a plasmid, respectively, or due to the different
visualization methodologies that have been applied in the
two studies. On the basis of their results, Jacobs-Wagner
et al. suggested that bacteria use the chromosome layout
to limit post-transcriptional processes, that is, translation
and potentially mRNA decay, to a defined spatial space,
thus increasing the subcellular concentration of the
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S. Govindarajan et al.
1016
(a)
(b)
Fig. 5. Patterns of mRNAs localization observed in bacteria.
(a) Several mRNAs in Escherichia coli were shown to localize in a
translation-independent manner to the subcellular domains where
their protein products are required, that is, to the poles, the
cytoplasm or the membrane. (b) Several mRNAs in Caulobacter
crescentus and E. coli were shown to localize near the chromosomal
loci from which they are transcribed (one transcribed gene is depicted
as a white band in the double helix). This distribution pattern can be
explained by translation of the mRNA that is still tethered to the DNA
during the process of transcription.
encoded proteins and their probability to interact and to
form complexes. The restriction in mRNA diffusion and
in ribosome motility that the authors documented in
C. crescentus can be explained by the coupling between
transcription and translation in bacteria, that is, translation of an mRNA that is still tethered to the DNA can
limit its diffusion. Additionally, the presence of multiple
ribosomes that translate one mRNA, and the nascent
polypeptides emerging from each ribosome may limit
both mRNA and ribosome mobility. Significantly, JacobsWagner et al. show that the C. crescentus RNase E appears
associated with chromosome, unlike the E. coli RNase E
that has been observed in a helix-like pattern and in association with the membrane (Taghbalout & Rothfield,
2007, 2008). One interesting implication of this model is
that the chromosome organization can provide a mean to
compartmentalize complex formation, thereby explaining
clustering of genes that encode interacting proteins in
bacterial chromosomes. The idea that chromosome organization is important and essential for complex formation
was suggested by several studies, which demonstrated that
full complementation of gene disruption occurred only
when the complementing genes were introduced into the
original chromosomal site, or when the entire operon was
expressed from an ectopic site (Jones et al., 2001; Defeu
Soufo & Graumann, 2004). Then again, other bacterial
proteins, expressed from plasmids or from ectopic chromosomal sites, were shown to localize, function, and
assemble into macromolecular complexes properly (Kain
et al., 1991; Guerout-Fleury et al., 1996; Lopian et al.,
2010) making gene order seem less critical. Still, it is
important to keep in mind that none of these studies
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compared the fitness cost of expressing a gene from different locations.
A different picture emerges from the study of AmsterChoder et al., who detected mRNAs at particular positions within E. coli cells. In addition to FISH and live cell
imaging, the authors also monitored the presence of
specific mRNAs in the cytosolic or in the membrane fractions of cells, after their disruption, by reverse transcription and PCR amplification. Three types of localization
patterns were observed: (1) a helix-like pattern in the
cytoplasm; (2) around the cell circumference, that is, at
or near the cytoplasmic membrane, and (3) at the cell
poles (Fig. 5a) (Nevo-Dinur et al., 2011). Significantly,
localization of the mRNAs that have been looked at correlated with the position of their future protein products.
Moreover, targeting of transcripts was shown to be in
tight correlation with the requirements for complex formation, that is, an mRNA of a transcription factor was
detected either at the membrane, when co-expressed with
its cognate membrane sensor with which it forms a complex, or at the cell poles, where it associates with another
protein, when expressed by itself. On the basis of these
results, the authors suggested that, like in eukaryotic cells,
localization of bacterial proteins can be achieved via targeting of their mRNA transcripts. Up until now, the prevailing view has been that protein localization in bacteria
depends solely on targeting signals within the protein,
because of the lack of nucleus and the coupling between
transcription and translation. However, Amster-Choder
et al. have shown that bacterial mRNAs can localize in a
translation-independent manner because of cis-acting
sequences within the mRNAs. A hierarchy in zip codes
localization capacity was observed: a membrane-encoding
sequence was found to be dominant to a hydrophilicencoding sequence in determining transcript localization.
Intriguingly, the targeting signals seem to be conserved
across the eukaryotic–prokaryotic divide, because transcripts encoding a transmembrane protein from Drosophila melanogaster localized to the inner membrane when
expressed in E. coli (Nevo-Dinur et al., 2011). This observation might be explained by the results of a recent bioinformatic study, which showed that the sequences of
mRNAs that encode membrane-spanning domains of
integral membrane proteins in prokaryotes as well as in
eukaryotes are enriched in uracils (Prilusky & Bibi, 2009),
a feature that may serve as a membrane-targeting signal.
In a very recent study, Gueiros-Filho et al. detected
another pattern of mRNA localization, that is, to the
forming septum in midcell. The authors have found that
ComN, which plays a role in the post-transcriptional control of the late competence comE operon in B. subtilis,
promotes accumulation of its target comE mRNA to
septal and polar sites (Dos Santos et al., 2012). On the
FEMS Microbiol Rev 36 (2012) 1005–1022
Spatiotemporal organization of bacterial cells
basis of their results, the authors speculated that localized
translation of ComE proteins, facilitated by mRNA-mediated
targeting of their encoding mRNA, may be required for
efficient competence development.
The above results suggest that mRNA targeting might be
important for localized translation in bacteria. This is in
line with the reported localization of ribosomes in B. subtilis and in E. coli to regions outside of the nucleoid, as
opposed to RNA polymerase, which resides principally
within the nucleoid (Lewis et al., 2000; Bakshi et al., 2012).
In fact, based on the evident separation of the nucleoid and
the transcription machinery from the translation machinery, Weisshaar et al. suggested that, at least in E. coli and
probably also in B. subtilis, most translation is not coupled
to transcription; rather, complete mRNA molecules find
their way to the ribosome-rich regions (Bakshi et al.,
2012). As mentioned before, the limited dispersion of
mRNA in C. crescentus is in correlation with the dispersion
of both the ribosomes and the DNA throughout the cytoplasm in this organism (Montero Llopis et al., 2010).
RNA targeting may also correlate with other posttranscriptional processes, such as mRNA decay. This suggestion is in agreement with the detection of components
of the E. coli degradosome, RNase E and RNA helicase B,
near the membrane and in a helix-like pattern (Taghbalout & Rothfield, 2007; Khemici et al., 2008). The imaging of Hfq, an RNA-binding protein that plays a role in
post-transcriptional regulation, and of RodZ, an RNAbinding cytoskeleton anchoring protein, close to the bacterial membrane (Diestra et al., 2009; Mitobe et al., 2011)
suggests that RNA localization might be linked to different regulatory mechanisms in the cell. Again, the limited
distribution of the transcripts in C. crescentus correlates
with the distribution of RNaseE in this organism (Montero Llopis et al., 2010). The parallels between the subcellular distribution of mRNA transcripts and of the various
post-translational machineries strongly suggest a link
between cell architecture and function.
Regardless of the different models that emerge for
mRNA distribution, both models agree that, despite the
scarcity of membrane-bounded organelles, bacterial cells
have the capacity to spatially organize mRNA molecules.
It is possible that the two models operate in different
organisms or even coexist in the same organism. The field
of mRNA localization in bacteria is still in its infancy,
and additional studies are required to unravel the relevant
mechanisms.
Concluding remarks
Studies published in the last decade completely changed
our view of bacterial cells. We have learnt that the
division to cells with and without membrane-enclosed
FEMS Microbiol Rev 36 (2012) 1005–1022
1017
organelle, first and foremost a nucleus, does not imply
that the two main kingdoms of cells differ in having
intricate internal organization. The mechanisms that are
responsible for cell architecture in bacteria are currently
unraveling. In this review, we described several strategies
that help establish the bacterial cell architecture and regulate vital processes spatially and temporally. Nevertheless,
there are obviously other strategies that we did not cover
in this review or that have not been discovered yet. This
hypothesis is supported by the finding that in addition to
the obvious subcellular domains, that is, the membrane,
the chromosome milieu, the poles, and the septum, there
are other separate physical compartments in bacterial
cells. One example is the metabolosomes, which are metabolically active structures that are bound by a proteinaceous shell and are located within the cytoplasm (Yeates
et al., 2008). New studies shed light on the composition, production, and structure of these protein-based
microcompartments, which act as simple organelles by
compartmentalizing functionally related enzymes that
compose specific metabolic pathways (Parsons et al.,
2010; Tanaka et al., 2010). The existence of such organelles, evidently developed to cope with metabolic stress,
draws parallels with eukaryotic organelles and expands
our view of cell compartmentalization. Undoubtedly, the
development of super-resolution microscopes and novel
fluorescent reagents will lead to the discovery of additional nonconventional subcellular compartments.
The thermodynamic rationale behind cell compartmentalization is valid for bacteria as it is for eukaryotes. In
fact, clustering of functionally related molecules and colocalization of signaling pathways seems a requirement
considering the high macromolecules concentration in
exponentially growing bacterial cells that, apparently,
limits diffusion rates in this environment (Zimmerman &
Trach, 1991). The large number of factors and complexes,
including pathogenesis factors, observed at the poles of
bacterial cells suggests that the poles have evolved,
because of their special characteristics, for example, composition, geometry, or structure, to serve as hubs that
allows for the association of components of complex systems, for the coordination of bacterial signaling systems
and, in general, for enabling a tighter regulation (AmsterChoder, 2011).
The demonstration that bacteria can localize RNAs to
subcellular regions in the cell revolutionized our view of
gene expression in prokaryotes. Similarly, unexpected was
the finding that localization of bacterial transcripts may
occur in a translation-independent manner. Once again,
the orthodox distinction between prokaryotes and eukaryotes is not valid anymore, as it is clear that the mechanistic
basis for separating transcription and translation did not
coevolve with the nucleus, but, rather, has developed in the
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
1018
primitive bacterial cell. All in all, these results raise numerous questions regarding the mechanisms underlying RNA
localization: How do the mRNA transcripts localize to the
different subcellular regions? In eukaryotes, mRNA localization was suggested to occur by facilitated diffusion or by
active transport along cytoskeletal filaments (Czaplinski &
Singer, 2006; Palacios, 2007). The average short half-life of
bacterial mRNAs (Bernstein et al., 2002), together with the
short generation time of bacterial cells, suggests that active
mechanisms are involved in mRNA targeting, at least in
some cases. Careful monitoring of single molecules in live
cells should be applied to distinguish between the different
possibilities. New tools are constantly being developed for
observing RNA within cells, for example, the RNA aptamers that bind fluorophores mimic GFP (Paige et al.,
2011). In the case that active mechanisms are involved, the
factors that recognize the mRNA zip codes and help transport the transcripts to their destination will need to be
identified. The role of mRNA targeting in post-transcriptional processes is another relevant question. In this
respect, it is worth mentioning that the recent documentation of binding of the membrane-anchored cytoskeletal
protein RodZ to certain mRNAs in Shigella (Mitobe et al.,
2011) provoked the authors to suggest that RodZ provides
a scaffold for post-transcriptional regulation. Finally, how
do sRNA and other regulatory noncoding RNA localize?
All these questions await further investigation using
advanced methodologies.
Summarily, the evidence that accumulated thus far
about the intricate organization of bacterial cells makes
them suitable models for studying spatial regulation of cellular processes and for elucidating the mechanisms that
underlie cellular architecture in higher organisms. Efforts
to unravel these mechanisms will pay off, as they will reveal
universal strategies that are relevant to all types of cells, as
well as bacterial-specific mechanisms, that can provide targets for the development of novel antibacterial drugs.
Acknowledgements
We thank all past and present members of the laboratory
for fruitful discussions over the years. This work was supported by the Israel Science Foundation founded by the
Israel Academy of Sciences and Humanities.
References
Alley MR, Maddock JR & Shapiro L (1992) Polar localization
of a bacterial chemoreceptor. Genes Dev 6: 825–836.
Alyahya SA, Alexander R, Costa T, Henriques AO, Emonet T
& Jacobs-Wagner C (2009) RodZ, a component of the
bacterial core morphogenic apparatus. P Natl Acad Sci USA
106: 1239–1244.
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
S. Govindarajan et al.
Amster-Choder O (2011) The compartmentalized vessel: the
bacterial cell as a model for subcellular organization (a tale
of two studies). Cell Logist 1: 77–81.
Bakshi S, Siryaporn A, Goulian M & Weisshaar JC (2012)
Superresolution imaging of ribosomes and RNA polymerase
in live Escherichia coli cells. Mol Microbiol 85: 21–38.
Barak I, Muchova K, Wilkinson AJ, O’Toole PJ & Pavlendova
N (2008) Lipid spirals in Bacillus subtilis and their role in
cell division. Mol Microbiol 68: 1315–1327.
Beach DL, Salmon ED & Bloom K (1999) Localization and
anchoring of mRNA in budding yeast. Curr Biol 9: 569–578.
Bendezu FO, Hale CA, Bernhardt TG & de Boer PA (2009) RodZ
(YfgA) is required for proper assembly of the MreB actin
cytoskeleton and cell shape in E. coli. EMBO J 28: 193–204.
Ben-Yehuda S, Rudner DZ & Losick R (2003) RacA, a
bacterial protein that anchors chromosomes to the cell
poles. Science 299: 532–536.
Bernstein HD, Poritz MA, Strub K, Hoben PJ, Brenner S &
Walter P (1989) Model for signal sequence recognition from
amino-acid sequence of 54K subunit of signal recognition
particle. Nature 340: 482–486.
Bernstein JA, Khodursky AB, Lin PH, Lin-Chao S & Cohen SN
(2002) Global analysis of mRNA decay and abundance in
Escherichia coli at single-gene resolution using two-color
fluorescent DNA microarrays. P Natl Acad Sci USA 99:
9697–9702.
Bertrand E, Chartrand P, Schaefer M, Shenoy SM, Singer RH
& Long RM (1998) Localization of ASH1 mRNA particles
in living yeast. Mol Cell 2: 437–445.
Bi EF & Lutkenhaus J (1991) FtsZ ring structure associated
with division in Escherichia coli. Nature 354: 161–164.
Bork P, Sander C & Valencia A (1992) An ATPase domain
common to prokaryotic cell cycle proteins, sugar kinases,
actin, and hsp70 heat shock proteins. P Natl Acad Sci USA
89: 7290–7294.
Bowman GR, Lyuksyutova AI & Shapiro L (2010) Bacterial
polarity. Curr Opin Cell Biol 23: 71–77.
Bramkamp M, Emmins R, Weston L, Donovan C, Daniel RA
& Errington J (2008) A novel component of the divisionsite selection system of Bacillus subtilis and a new mode of
action for the division inhibitor MinCD. Mol Microbiol 70:
1556–1569.
Broude NE (2011) Analysis of RNA localization and
metabolism in single live bacterial cells: achievements and
challenges. Mol Microbiol 80: 1137–1147.
Cabeen MT & Jacobs-Wagner C (2010) The bacterial
cytoskeleton. Annu Rev Genet 44: 365–392.
Carballido-Lopez R, Formstone A, Li Y, Ehrlich SD, Noirot P
& Errington J (2006) Actin homolog MreBH governs cell
morphogenesis by localization of the cell wall hydrolase
LytE. Dev Cell 11: 399–409.
Carlsson F, Joshi SA, Rangell L & Brown EJ (2009) Polar
localization of virulence-related Esx-1 secretion in
mycobacteria. PLoS Pathog 5: e1000285.
Collier J & Shapiro L (2007) Spatial complexity and control of
a bacterial cell cycle. Curr Opin Biotechnol 18: 333–340.
FEMS Microbiol Rev 36 (2012) 1005–1022
Spatiotemporal organization of bacterial cells
Cowles KN & Gitai Z (2010) Surface association and the MreB
cytoskeleton regulate pilus production, localization and
function in Pseudomonas aeruginosa. Mol Microbiol 76:
1411–1426.
Czaplinski K & Singer RH (2006) Pathways for mRNA
localization in the cytoplasm. Trends Biochem Sci 31: 687–693.
Daniel RA & Errington J (2003) Control of cell morphogenesis
in bacteria: two distinct ways to make a rod-shaped cell.
Cell 113: 767–776.
Davies KM & Lewis PJ (2003) Localization of rRNA synthesis
in Bacillus subtilis: characterization of loci involved in
transcription focus formation. J Bacteriol 185: 2346–2353.
de Boer PA (2010) Advances in understanding E. coli cell
fission. Curr Opin Microbiol 13: 730–737.
Defeu Soufo HJ & Graumann PL (2004) Dynamic movement of
actin-like proteins within bacterial cells. EMBO Rep 5: 789–794.
Deutscher J (2008) The mechanisms of carbon catabolite
repression in bacteria. Curr Opin Microbiol 11: 87–93.
Diestra E, Cayrol B, Arluison V & Risco C (2009) Cellular
electron microscopy imaging reveals the localization of the
Hfq protein close to the bacterial membrane. PLoS ONE 4:
e8301.
Divakaruni AV, Loo RR, Xie Y, Loo JA & Gober JW (2005)
The cell-shape protein MreC interacts with extracytoplasmic
proteins including cell wall assembly complexes in
Caulobacter crescentus. P Natl Acad Sci USA 102:
18602–18607.
Divakaruni AV, Baida C, White CL & Gober JW (2007) The
cell shape proteins MreB and MreC control cell
morphogenesis by positioning cell wall synthetic complexes.
Mol Microbiol 66: 174–188.
Dominguez-Escobar J, Chastanet A, Crevenna AH, Fromion V,
Wedlich-Soldner R & Carballido-Lopez R (2011) Processive
movement of MreB-associated cell wall biosynthetic
complexes in bacteria. Science 333: 225–228.
Donnelly CJ, Fainzilber M & Twiss JL (2010) Subcellular
communication through RNA transport and localized
protein synthesis. Traffic 11: 1498–1505.
Donovan C & Bramkamp M (2009) Characterization and
subcellular localization of a bacterial flotillin homologue.
Microbiology 155: 1786–1799.
Dos Santos VT, Bisson-Filho AW & Gueiros-Filho FJ (2012)
DivIVA-mediated polar localization of ComN, a posttranscriptional regulator of B. subtilis. J Bacteriol, 194:
3661–3669.
Dworkin J (2009) Cellular polarity in prokaryotic organisms.
Cold Spring Harb Perspect Biol 1: a003368.
Dye NA, Pincus Z, Theriot JA, Shapiro L & Gitai Z (2005)
Two independent spiral structures control cell shape in
Caulobacter. P Natl Acad Sci USA 102: 18608–18613.
Ebersbach G & Jacobs-Wagner C (2007) Exploration into the
spatial and temporal mechanisms of bacterial polarity.
Trends Microbiol 15: 101–108.
Erickson HP, Anderson DE & Osawa M (2010) FtsZ in
bacterial cytokinesis: cytoskeleton and force generator all in
one. Microbiol Mol Biol Rev 74: 504–528.
FEMS Microbiol Rev 36 (2012) 1005–1022
1019
Eswaramoorthy P, Erb ML, Gregory JA, Silverman J, Pogliano
K, Pogliano J & Ramamurthi KS (2011) Cellular
architecture mediates DivIVA ultrastructure and regulates
min activity in Bacillus subtilis. mBio 2: e00257–11.
Figge RM, Divakaruni AV & Gober JW (2004) MreB, the cell
shape-determining bacterial actin homologue, co-ordinates
cell wall morphogenesis in Caulobacter crescentus. Mol
Microbiol 51: 1321–1332.
Garner EC, Bernard R, Wang W, Zhuang X, Rudner DZ &
Mitchison T (2011) Coupled, circumferential motions of the
cell wall synthesis machinery and MreB filaments in B.
subtilis. Science 333: 222–225.
Gilmore R, Walter P & Blobel G (1982) Protein translocation
across the endoplasmic reticulum. II. Isolation and
characterization of the signal recognition particle receptor.
J Cell Biol 95: 470–477.
Gitai Z, Dye N & Shapiro L (2004) An actin-like gene can
determine cell polarity in bacteria. P Natl Acad Sci USA
101: 8643–8648.
Gold VA, Robson A, Bao H, Romantsov T, Duong F &
Collinson I (2010) The action of cardiolipin on the bacterial
translocon. P Natl Acad Sci USA 107: 10044–10049.
Goldberg MB, Barzu O, Parsot C & Sansonetti PJ (1993)
Unipolar localization and ATPase activity of IcsA, a Shigella
flexneri protein involved in intracellular movement.
J Bacteriol 175: 2189–2196.
Golding I & Cox EC (2004) RNA dynamics in live Escherichia
coli cells. P Natl Acad Sci USA 101: 11310–11315.
Gordon GS, Sitnikov D, Webb CD, Teleman A, Straight A,
Losick R, Murray AW & Wright A (1997) Chromosome and
low copy plasmid segregation in E. coli: visual evidence for
distinct mechanisms. Cell 90: 1113–1121.
Greenfield D, McEvoy AL, Shroff H, Crooks GE, Wingreen
NS, Betzig E & Liphardt J (2009) Self-organization of the
Escherichia coli chemotaxis network imaged with superresolution light microscopy. PLoS Biol 7: e1000137.
Guerout-Fleury AM, Frandsen N & Stragier P (1996) Plasmids
for ectopic integration in Bacillus subtilis. Gene 180: 57–61.
Herskovits AA & Bibi E (2000) Association of Escherichia coli
ribosomes with the inner membrane requires the signal
recognition particle receptor but is independent of the signal
recognition particle. P Natl Acad Sci USA 97: 4621–4626.
Hsieh CW, Lin TY, Lai HM, Lin CC, Hsieh TS & Shih YL (2010)
Direct MinE-membrane interaction contributes to the proper
localization of MinDE in E. coli. Mol Microbiol 75: 499–512.
Hu CD, Chinenov Y & Kerppola TK (2002) Visualization of
interactions among bZIP and Rel family proteins in living
cells using bimolecular fluorescence complementation. Mol
Cell 9: 789–798.
Ingerson-Mahar M & Gitai Z (2012) A growing family: the
expanding universe of the bacterial cytoskeleton. FEMS
Microbiol Rev 36: 256–267.
Ingledew WJ & Poole RK (1984) The respiratory chains of
Escherichia coli. Microbiol Rev 48: 222–271.
Jain S, van Ulsen P, Benz I, Schmidt MA, Fernandez R,
Tommassen J & Goldberg MB (2006) Polar localization of
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
1020
the autotransporter family of large bacterial virulence
proteins. J Bacteriol 188: 4841–4850.
Janakiraman A & Goldberg MB (2004) Recent advances on the
development of bacterial poles. Trends Microbiol 12: 518–525.
Jaumouille V, Francetic O, Sansonetti PJ & Tran Van Nhieu G
(2008) Cytoplasmic targeting of IpaC to the bacterial pole directs
polar type III secretion in Shigella. EMBO J 27: 447–457.
Jeffery WR, Tomlinson CR & Brodeur RD (1983) Localization
of actin messenger RNA during early ascidian development.
Dev Biol 99: 408–417.
Jockusch BM & Graumann PL (2011) The long journey: actin
on the road to pro- and eukaryotic cells. Rev Physiol
Biochem Pharmacol 161: 67–85.
Jones LJ, Carballido-Lopez R & Errington J (2001) Control of
cell shape in bacteria: helical, actin-like filaments in Bacillus
subtilis. Cell 104: 913–922.
Judd PK, Kumar RB & Das A (2005) Spatial location and
requirements for the assembly of the Agrobacterium
tumefaciens type IV secretion apparatus. P Natl Acad Sci
USA 102: 11498–11503.
Kain KC, Orlandi PA & Lanar DE (1991) Universal promoter
for gene expression without cloning: expression-PCR.
Biotechniques 10: 366–374.
Katsu T, Tsuchiya T & Fujita Y (1984) Dissipation of
membrane potential of Escherichia coli cells induced by
macromolecular polylysine. Biochem Biophys Res Commun
122: 401–406.
Khemici V, Poljak L, Luisi BF & Carpousis AJ (2008) The
RNase E of Escherichia coli is a membrane-binding protein.
Mol Microbiol 70: 799–813.
Kirkpatrick CL & Viollier PH (2010) Poles apart: prokaryotic
polar organelles and their spatial regulation. Cold Spring
Harb Perspect Biol 3: a006809.
Kislauskis EH, Li Z, Singer RH & Taneja KL (1993) Isoformspecific 3′-untranslated sequences sort alpha-cardiac and
beta-cytoplasmic actin messenger RNAs to different
cytoplasmic compartments. J Cell Biol 123: 165–172.
Komeili A, Li Z, Newman DK & Jensen GJ (2006)
Magnetosomes are cell membrane invaginations organized
by the actin-like protein MamK. Science 311: 242–245.
Koshkin AA, Singh SK, Nielsen P, Rajwanshi VK, Kumar R,
Meldgaard M, Olsen CE & Wengel J (1998) LNA (locked
nucleic acids): synthesis of the adenine, cytosine, guanine,
5-methylcytosine, thymine and uracil bicyclonucleoside
monomers, oligomerisation, and unprecedented nucleic acid
recognition. Tetrahedron 54: 3607–3630.
Kruse T, Bork-Jensen J & Gerdes K (2005) The morphogenetic
MreBCD proteins of Escherichia coli form an essential
membrane-bound complex. Mol Microbiol 55: 78–89.
Lande MA, Adesnik M, Sumida M, Tashiro Y & Sabatini DD
(1975) Direct association of messenger RNA with
microsomal membranes in human diploid fibroblasts. J Cell
Biol 65: 513–528.
Lenarcic R, Halbedel S, Visser L et al. (2009) Localisation of
DivIVA by targeting to negatively curved membranes.
EMBO J 28: 2272–2282.
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
S. Govindarajan et al.
Lenz P & Sogaard-Andersen L (2011) Temporal and spatial
oscillations in bacteria. Nat Rev Microbiol 9: 565–577.
Lewis PJ, Thaker SD & Errington J (2000)
Compartmentalization of transcription and translation in
Bacillus subtilis. EMBO J 19: 710–718.
Lin BH, Cui BX, Lee JH & Yu J (2002) Hydrodynamic
coupling in diffusion of quasi-one-dimensional Brownian
particles. Europhys Lett 57: 724–730.
Lopez D & Kolter R (2010) Functional microdomains in
bacterial membranes. Genes Dev 24: 1893–1902.
Lopian L, Elisha Y, Nussbaum-Shochat A & Amster-Choder O
(2010) Spatial and temporal organization of the E. coli PTS
components. EMBO J 29: 3630–3645.
Lux R, Jahreis K, Bettenbrock K, Parkinson JS & Lengeler JW
(1995) Coupling the phosphotransferase system and the
methyl-accepting chemotaxis protein-dependent chemotaxis
signaling pathways of Escherichia coli. P Natl Acad Sci USA
92: 11583–11587.
MaX, Ehrhardt DW & Margolin W (1996) Colocalization of
cell division proteins FtsZ and FtsA to cytoskeletal structures
in living Escherichia coli cells by using green fluorescent
protein. P Natl Acad Sci USA 93: 12998–13003.
Matsumoto K, Kusaka J, Nishibori A & Hara H (2006)
Lipid domains in bacterial membranes. Mol Microbiol 61:
1110–1117.
Mattei PJ, Neves D & Dessen A (2010) Bridging cell wall
biosynthesis and bacterial morphogenesis. Curr Opin Struct
Biol 20: 749–755.
Mauriello EM, Mouhamar F, Nan B, Ducret A, Dai D,
Zusman DR & Mignot T (2010) Bacterial motility
complexes require the actin-like protein, MreB and the Ras
homologue, MglA. EMBO J 29: 315–326.
Mazor S, Regev T, Mileykovskaya E, Margolin W, Dowhan W
& Fishov I (2008a) Mutual effects of MinD-membrane
interaction: I. Changes in the membrane properties induced
by MinD binding. Biochim Biophys Acta 1778: 2496–2504.
Mazor S, Regev T, Mileykovskaya E, Margolin W, Dowhan W
& Fishov I (2008b) Mutual effects of MinD-membrane
interaction: II. Domain structure of the membrane enhances
MinD binding. Biochim Biophys Acta 1778: 2505–2511.
Michnick SW (2003) Protein fragment complementation
strategies for biochemical network mapping. Curr Opin
Biotechnol 14: 610–617.
Mileykovskaya E & Dowhan W (2009) Cardiolipin membrane
domains in prokaryotes and eukaryotes. Biochim Biophys
Acta 1788: 2084–2091.
Mileykovskaya E, Fishov I, Fu X, Corbin BD, Margolin W &
Dowhan W (2003) Effects of phospholipid composition on
MinD-membrane interactions in vitro and in vivo. J Biol
Chem 278: 22193–22198.
Mitobe J, Yanagihara I, Ohnishi K, Yamamoto S, Ohnishi M,
Ishihama A & Watanabe H (2011) RodZ regulates the posttranscriptional processing of the Shigella sonnei type III
secretion system. EMBO Rep 12: 911–916.
Montero Llopis P, Jackson AF, Sliusarenko O, Surovtsev I,
Heinritz J, Emonet T & Jacobs-Wagner C (2010) Spatial
FEMS Microbiol Rev 36 (2012) 1005–1022
Spatiotemporal organization of bacterial cells
organization of the flow of genetic information in bacteria.
Nature 466: 77–81.
Nevo-Dinur K, Nussbaum-Shochat A, Ben-Yehuda S &
Amster-Choder O (2011) Translation-independent
localization of mRNA in E. coli. Science 331: 1081–1084.
Obika S, Nanbu N, Hari Y, Morio K-I, In Y, Ishida T &
Imanishi T (1997) Synthesis of 2′-O,4′-C-methyleneuridine
and -cytidine. Novel bicyclic nucleosides having a fixed
C3′-endo sugar puckering. Tetrahedron Lett 38: 8735–8738.
Paige JS, Wu KY & Jaffrey SR (2011) RNA mimics of green
fluorescent protein. Science 333: 642–646.
Palacios IM (2007) How does an mRNA find its way?
Intracellular localisation of transcripts. Semin Cell Dev Biol
18: 163–170.
Palade GE (1958) In Microsomal Particles and Protein Synthesis.
Pergamon Press, New York.
Parsons JB, Frank S, Bhella D, Liang M, Prentice MB,
Mulvihill DP & Warren MJ (2010) Synthesis of empty
bacterial microcompartments, directed organelle protein
incorporation, and evidence of filament-associated organelle
movement. Mol Cell 38: 305–315.
Pilhofer M, Pavlekovic M, Lee NM, Ludwig W & Schleifer KH
(2009) Fluorescence in situ hybridization for intracellular
localization of nifH mRNA. Syst Appl Microbiol 32: 186–192.
Pogliano J (2002) Dynamic cellular location of bacterial
plasmids. Curr Opin Microbiol 5: 586–590.
Poritz MA, Siegel V, Hansen W & Walter P (1988) Small
ribonucleoproteins in Schizosaccharomyces pombe and
Yarrowia lipolytica homologous to signal recognition
particle. P Natl Acad Sci USA 85: 4315–4319.
Porter SL, Wadhams GH & Armitage JP (2011) Signal
processing in complex chemotaxis pathways. Nat Rev
Microbiol 9: 153–165.
Prilusky J & Bibi E (2009) Studying membrane proteins through
the eyes of the genetic code revealed a strong uracil bias in
their coding mRNAs. P Natl Acad Sci USA 106: 6662–6666.
Rackham O & Brown CM (2004) Visualization of RNAprotein interactions in living cells: FMRP and IMP1 interact
on mRNAs. EMBO J 23: 3346–3355.
Ramamurthi KS (2010) Protein localization by recognition of
membrane curvature. Curr Opin Microbiol 13: 753–757.
Ramamurthi KS & Losick R (2009) Negative membrane
curvature as a cue for subcellular localization of a bacterial
protein. P Natl Acad Sci USA 106: 13541–13545.
Ramamurthi KS, Clapham KR & Losick R (2006) Peptide
anchoring spore coat assembly to the outer forespore
membrane in Bacillus subtilis. Mol Microbiol 62:
1547–1557.
Ramamurthi KS, Lecuyer S, Stone HA & Losick R (2009)
Geometric cue for protein localization in a bacterium.
Science 323: 1354–1357.
Roberts MA, Papachristodoulou A & Armitage JP (2010)
Adaptation and control circuits in bacterial chemotaxis.
Biochem Soc Trans 38: 1265–1269.
Rokney A, Shagan M, Kessel M, Smith Y, Rosenshine I &
Oppenheim AB (2009) E. coli transports aggregated proteins
FEMS Microbiol Rev 36 (2012) 1005–1022
1021
to the poles by a specific and energy-dependent process.
J Mol Biol 392: 589–601.
Romantsov T, Helbig S, Culham DE, Gill C, Stalker L &
Wood JM (2007) Cardiolipin promotes polar localization of
osmosensory transporter ProP in Escherichia coli. Mol
Microbiol 64: 1455–1465.
Romantsov T, Battle AR, Hendel JL, Martinac B & Wood JM
(2010) Protein localization in Escherichia coli cells:
comparison of the cytoplasmic membrane proteins ProP,
LacY, ProW, AqpZ, MscS, and MscL. J Bacteriol 192:
912–924.
Romisch K, Webb J, Herz J, Prehn S, Frank R, Vingron M &
Dobberstein B (1989) Homology of 54K protein of signalrecognition particle, docking protein and two E. coli
proteins with putative GTP-binding domains. Nature 340:
478–482.
Rudner DZ & Losick R (2010) Protein subcellular localization
in bacteria. Cold Spring Harb Perspect Biol 2: a000307.
Russell JH & Keiler KC (2009) Subcellular localization of a
bacterial regulatory RNA. P Natl Acad Sci USA 106:
16405–16409.
Savage DF, Afonso B, Chen AH & Silver PA (2010) Spatially
ordered dynamics of the bacterial carbon fixation
machinery. Science 327: 1258–1261.
Schaffer E & Thiele U (2004) Dynamic domain formation in
membranes: thickness-modulation-induced phase
separation. Eur Phys J E Soft Matter 14: 169–175.
Schulmeister S, Ruttorf M, Thiem S, Kentner D, Lebiedz D &
Sourjik V (2008) Protein exchange dynamics at
chemoreceptor clusters in Escherichia coli. P Natl Acad Sci
USA 105: 6403–6408.
Scott ME, Dossani ZY & Sandkvist M (2001) Directed polar
secretion of protease from single cells of Vibrio cholerae via
the type II secretion pathway. P Natl Acad Sci USA 98:
13978–13983.
Sekimizu K & Kornberg A (1988) Cardiolipin activation of
dnaA protein, the initiation protein of replication in
Escherichia coli. J Biol Chem 263: 7131–7135.
Shaevitz JW & Gitai Z (2010) The structure and function of
bacterial actin homologs. Cold Spring Harb Perspect Biol 2:
a000364.
Shapiro L, McAdams HH & Losick R (2002) Generating and
exploiting polarity in bacteria. Science 298: 1942–1946.
Shapiro L, McAdams HH & Losick R (2009) Why and how
bacteria localize proteins. Science 326: 1225–1228.
Shiomi D, Yoshimoto M, Homma M & Kawagishi I (2006)
Helical distribution of the bacterial chemoreceptor via
colocalization with the Sec protein translocation machinery.
Mol Microbiol 60: 894–906.
Shiomi D, Sakai M & Niki H (2008) Determination of
bacterial rod shape by a novel cytoskeletal membrane
protein. EMBO J 27: 3081–3091.
Slovak PM, Wadhams GH & Armitage JP (2005) Localization
of MreB in Rhodobacter sphaeroides under conditions
causing changes in cell shape and membrane structure.
J Bacteriol 187: 54–64.
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
1022
Sourjik V (2004) Receptor clustering and signal processing in
E. coli chemotaxis. Trends Microbiol 12: 569–576.
Sourjik V & Armitage JP (2010) Spatial organization in
bacterial chemotaxis. EMBO J 29: 2724–2733.
Sourjik V & Berg HC (2000) Localization of components of
the chemotaxis machinery of Escherichia coli using
fluorescent protein fusions. Mol Microbiol 37: 740–751.
Strahl H & Hamoen LW (2010) Membrane potential is
important for bacterial cell division. P Natl Acad Sci USA
107: 12281–12286.
Taghbalout A & Rothfield L (2007) RNaseE and the other
constituents of the RNA degradosome are components of
the bacterial cytoskeleton. P Natl Acad Sci USA 104: 1667–
1672.
Taghbalout A & Rothfield L (2008) RNaseE and RNA
helicase B play central roles in the cytoskeletal
organization of the RNA degradosome. J Biol Chem 283:
13850–13855.
Tanaka S, Sawaya MR & Yeates TO (2010) Structure and
mechanisms of a protein-based organelle in Escherichia coli.
Science 327: 81–84.
Taniguchi Y, Choi PJ, Li GW et al. (2010) Quantifying E. coli
proteome and transcriptome with single-molecule sensitivity
in single cells. Science 329: 533–538.
Thanbichler M & Shapiro L (2008) Getting organized–how
bacterial cells move proteins and DNA. Nat Rev Microbiol 6:
28–40.
Thiem S & Sourjik V (2008) Stochastic assembly of
chemoreceptor clusters in Escherichia coli. Mol Microbiol 68:
1228–1236.
Thiem S, Kentner D & Sourjik V (2007) Positioning of
chemosensory clusters in E. coli and its relation to cell
division. EMBO J 26: 1615–1623.
Toro E & Shapiro L (2010) Bacterial chromosome
organization and segregation. Cold Spring Harb Perspect Biol
2: a000349.
Tsokos CG, Perchuk BS & Laub MT (2011) A dynamic
complex of signaling proteins uses polar localization to
regulate cell-fate asymmetry in Caulobacter crescentus. Dev
Cell 20: 329–341.
Valencia-Burton M, McCullough RM, Cantor CR & Broude
NE (2007) RNA visualization in live bacterial cells using
fluorescent protein complementation. Nat Methods 4:
421–427.
Valencia-Burton M, Shah A, Sutin J et al. (2009)
Spatiotemporal patterns and transcription kinetics of
induced RNA in single bacterial cells. P Natl Acad Sci USA
106: 16399–16404.
van den Ent F & Lowe J (2000) Crystal structure of the cell
division protein FtsA from Thermotoga maritima. EMBO J
19: 5300–5307.
van den Ent F, Johnson CM, Persons L, de Boer P & Lowe J
(2010) Bacterial actin MreB assembles in complex with cell
shape protein RodZ. EMBO J 29: 1081–1090.
van Teeffelen S, Wang S, Furchtgott L, Huang KC, Wingreen
NS, Shaevitz JW & Gitai Z (2011) The bacterial actin MreB
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
S. Govindarajan et al.
rotates, and rotation depends on cell-wall assembly. P Natl
Acad Sci USA 108: 15822–15827.
Vats P & Rothfield L (2007) Duplication and segregation of
the actin (MreB) cytoskeleton during the prokaryotic cell
cycle. P Natl Acad Sci USA 104: 17795–17800.
Vats P, Yu J & Rothfield L (2009a) The dynamic nature of the
bacterial cytoskeleton. Cell Mol Life Sci 66: 3353–3362.
Vats P, Shih YL & Rothfield L (2009b) Assembly of the MreBassociated cytoskeletal ring of Escherichia coli. Mol Microbiol
72: 170–182.
Wachi M, Doi M, Tamaki S, Park W, Nakajima-Iijima S &
Matsuhashi M (1987) Mutant isolation and molecular
cloning of mre genes, which determine cell shape, sensitivity
to mecillinam, and amount of penicillin-binding proteins in
Escherichia coli. J Bacteriol 169: 4935–4940.
Wagner JK, Galvani CD & Brun YV (2005) Caulobacter
crescentus requires RodA and MreB for stalk synthesis and
prevention of ectopic pole formation. J Bacteriol 187:
544–553.
Walter P & Blobel G (1980) Purification of a membraneassociated protein complex required for protein
translocation across the endoplasmic reticulum. P Natl Acad
Sci USA 77: 7112–7116.
Walter P & Blobel G (1982) Signal recognition particle
contains a 7S RNA essential for protein translocation across
the endoplasmic reticulum. Nature 299: 691–698.
Wang S, Arellano-Santoyo H, Combs PA & Shaevitz JW
(2010) Actin-like cytoskeleton filaments contribute to cell
mechanics in bacteria. P Natl Acad Sci USA 107: 9182–9185.
Wang S, Furchtgott L, Huang KC & Shaevitz JW (2012)
Helical insertion of peptidoglycan produces chiral ordering
of the bacterial cell wall. P Natl Acad Sci USA 109: E595–
E604.
Werner JN, Chen EY, Guberman JM, Zippilli AR, Irgon JJ &
Gitai Z (2009) Quantitative genome-scale analysis of protein
localization in an asymmetric bacterium. P Natl Acad Sci
USA 106: 7858–7863.
White CL, Kitich A & Gober JW (2010) Positioning cell wall
synthetic complexes by the bacterial morphogenetic proteins
MreB and MreD. Mol Microbiol 76: 616–633.
Yeates TO, Kerfeld CA, Heinhorst S, Cannon GC & Shively
JM (2008) Protein-based organelles in bacteria:
carboxysomes and related microcompartments. Nat Rev
Microbiol 6: 681–691.
Yiu H-W, Demidov VV, Toran P, Cantor CR & Broude NE
(2011) RNA detection in live bacterial cells using fluorescent
protein complementation triggered by interaction of two
RNA aptamers with two RNA-binding peptides.
Pharmaceuticals 4: 494–508.
Zhang HM, Li Z, Tsudome M, Ito S, Takami H & Horikoshi
K (2005) An alkali-inducible flotillin-like protein from
Bacillus halodurans C-125. Protein J 24: 125–131.
Zimmerman SB & Trach SO (1991) Estimation of
macromolecule concentrations and excluded volume
effects for the cytoplasm of Escherichia coli. J Mol Biol 222:
599–620.
FEMS Microbiol Rev 36 (2012) 1005–1022