Microbiology (2004), 150, 2815–2824
DOI 10.1099/mic.0.27223-0
Dynamic localization of membrane proteins in
Bacillus subtilis
A. S. Johnson,3 S. van Horck and P. J. Lewis
School of Environmental and Life Sciences, Biological Sciences, University of Newcastle,
Callaghan, NSW 2308, Australia
Correspondence
P. J. Lewis
[email protected]
Received 8 April 2004
Revised
27 May 2004
Accepted 27 May 2004
The subcellular localization of membrane proteins in Bacillus subtilis was examined by using
fluorescent protein fusions. ATP synthase and succinate dehydrogenase were found to localize
within discrete domains on the membrane rather than being homogeneously distributed around
the cell periphery as expected. Dual labelling of cells indicated partial colocalization of ATP
synthase and succinate dehydrogenase. Further analysis using an ectopically expressed phage
protein gave the same localization patterns as ATP synthase and succinate dehydrogenase,
implying that membrane proteins are restricted to domains within the membrane. 3D reconstruction
of images of the localization of ATP synthase showed that domains were not regular and there
was no bias for localization to cell poles or any other positions. Further analysis revealed that
this localization was highly dynamic, but random, implying that integral membrane proteins
are free to diffuse two-dimensionally around the cytoplasmic membrane.
INTRODUCTION
Biological membranes are not homogeneous structures,
but consist of domains enriched for specific lipids and
proteins often referred to as lipid rafts (Edidin, 2003).
These rafts appear to be highly dynamic structures, in terms
of both lipid/protein content and size (Edidin, 2003).
Although lipid rafts and membrane heterogeneity have
been intensively studied in eukaryotic systems, relatively
little work has been carried out with microbial systems,
probably in part because of their small size. In recent years,
however, convincing evidence has been presented indicating the presence of lipid domains in microbial cytoplasmic
membranes (Fishov & Woldringh, 1999; Mileykovskaya &
Dowhan, 2000; Kawai et al., 2004). In particular, the use of
specific fluorescent probes has indicated that the phospholipid cardiolipin localizes preferentially to the cell poles and
division sites (mid-cell regions) of both Gram-positive and
Gram-negative bacteria (Mileykovskaya & Dowhan, 2000;
Kawai et al., 2004). Further experiments using 2-pyrene
derivatives of the phospholipids phosphatidylethanolamine
(PE) and phosphatidyleglycerol (PG) indicated that the
two lipids segregate into distinct domains in both Grampositive and Gram-negative cell membranes, and that the
PE domains were enriched for protein content (Vanounou
et al., 2003). Despite the similarity of results obtained with
3Present address: Life Therapeutics, PO Box 6126, Frenchs Forest,
NSW 2086, Australia.
Abbreviation: FRAP, fluorescence recovery after photobleaching.
The online version of this paper at http://mic.sgmjournals.org has
supplementary movie files and a supplementary figure.
0002-7223 G 2004 SGM
Gram-positive (Bacillus subtilis) and Gram-negative
(Escherichia coli) model systems, the two organisms have
quite different proportions of common phospholipids in
their cytoplasmic membranes. In B. subtilis PE accounts
for approximately 50 % of phospholipid, PG 15 %, the
lysine ester of PG 2?4 %, cardiolipin (CL) 0?8 %, and monodi- and triglucosyldiacylglycerols 30 % (de Mendoza et al.,
2002), whereas in E. coli PE accounts for about 70–80 % of
phospholipid, PG 15–25 % and CL 5–10 % (Kadner, 1996).
Do integral membrane proteins also segregate into domains
within the cytoplasmic membrane? The presence of chemoreceptors at cell poles is well established (Maddock &
Shapiro, 1993) and this may reflect a preference of these
proteins for acidic lipids such as cardiolipin. Penicillinbinding proteins involved in cell wall synthesis have also
been shown to localize in several patterns, including to
division septa and discrete foci along the long axis of the
cell, reflecting the role of these proteins in specific stages
of cell wall synthesis (Scheffers et al., 2004). However,
analysis of the localization of the E. coli Sec protein secretion machinery, BglF sugar sensor and B. subtilis phage
w29 DNA replication protein p16.7 indicates that these
proteins are homogeneously distributed around the cytoplasmic membrane, with no reported preference for cell
poles or other observable domains (Brandon et al., 2003;
Lopian et al., 2003; Meijer et al., 2001). Certainly, the
images presented in these papers indicate that the localization pattern of these proteins creates a clear outline of the
cytoplasmic membrane.
We have examined the localization of ATP synthase and
succinate dehydrogenase, which carries out steps in both
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
Printed in Great Britain
2815
A. S. Johnson, S. van Horck and P. J. Lewis
the tricarboxylic acid cycle and the electron-transport chain
(as complex II), and re-examined the localization of p16.7
using a series of high-resolution fluorescence microscopy
techniques with fluorescent protein fusions in live B. subtilis
cells. We found that all of the proteins localized around the
cytoplasmic membrane heterogeneously, and appeared to
be free to move randomly throughout the membrane. We
propose that such localization to domains is a general
feature of integral membrane proteins.
METHODS
Strains and growth conditions. Bacterial strains and plasmids
used in this study are listed in Table 1. All cloning was performed
using E. coli DH5a (Gibco-BRL). Unless otherwise stated B. subtilis
strains were cultured in CH medium (Sharpe et al., 1998) supplemented with 0?5 % (w/v) xylose at 37 uC. Transformation of B. subtilis was carried out by the method of Anagnostopoulos & Spizizen
(1961) as modified by Jenkinson (1983) on nutrient agar plates supplemented with the appropriate antibiotics and inducers: chloramphenicol, 5 mg ml21; spectinomycin, 50 mg ml21; xylose 0?5 % (w/
v); IPTG 0?5 mM. The functionality of atpA– and sdhA–gfp fusions
was checked on succinate minimal media plates (SMM: Santana
et al., 1994) supplemented with the appropriate antibiotics and
inducers, and grown at 37 uC.
The vital membrane stain FM4-64 (Molecular Probes) was added to
cultures at a final concentration of 30 ng ml21 30 min prior to
visualization.
DNA manipulations. The complete atpA gene was PCR amplified
using the following primers: forward 59-AGAAGGGGTGAACTCGAGGTGAGCATCAAA-39 and reverse 59-TCATCAGCATTCGAATTCGTTGCTTGGTGC-39. The resulting product was digested with
MunI and EcoRI and the 840 bp 39 fragment inserted into EcoRIdigested pSG1164. One recombinant with the fragment inserted in
the correct orientation was named pNG24 (Table 1).
The 39 1073 bp of sdhA were PCR amplified using the following
primers: forward 59-CGTATTATGGGTACCGAGAATTCATTC-39
and reverse 59-GTTCACTCATGACTCGAGCGCCACCTTCTT-39.
The PCR product was digested with Acc651 and XhoI and inserted
into similarly cut pSG1164, and the recombinant plasmid was named
pNG107 (Table 1).
DNA manipulations for the construction of CFP, YFP and IPTGinducible fusions to atpA and sdhA are indicated in Table 1.
Microscopy and image processing
Image acquisition. Images were obtained using a Zeiss Axioscop 2
epifluorescence microscope fitted with a Quantix 1401ECCD camera
(Photometrics), 100 W Hg lamp source and a 1006 ApoPlan NA
1.4 lens. GFP fluorescence was visualized with set 41018, CFP with
Table 1. Strains and plasmids
Strain or
plasmid
Genotype
Source/construction
Laboratory strain
168 transformed with pNG24
168 transformed with pNG25
168 transformed with pNG107
168 transformed with pNG117
BS24 transformed with pCm : Sp; inserts by double crossover into
cat gene of pNG25 and converts to spcR
BS131 transformed with BS112 chromosomal DNA
110WA
Plasmids
pCm : Sp
pSG1164
pSG1170
pSG1186
pSG1187
pNG24
pNG25
trpC2
trpC2 chr : : pNG24 (atpA–gfp Pxyl–9atpA cat)
trpC2 chr : : pNG25 (atpA–cfp Pxyl–9atpA cat)
trpC2 chr : : pNG107 (sdhA–gfp Pxyl–9sdhA cat)
trpC2 chr : : pNG117 (sdhA–yfp lacI Pspac–9sdhA cat)
trpC2 chr : : pNG25 [atpA–cfp Pxyl–9atpA VpCm : Sp
(catS spcR)]
trpC2 chr : : pNG25 [atpA–cfp Pxyl–9atpA VpCm : Sp
(catS spcR)] chr : : pNG115 (sdhA–yfp Pxyl–9sdhA cat)
trpC2 amyE VpSGW1(Pxyl–p16.7–gfp spc)
bla
bla
bla
bla
bla
bla
bla
pNG107
bla cat Pxyl–9sdhA–gfp
pNG115
bla cat Pxyl–9sdhA–yfp
pNG117
bla cat lacI Pspac–9sdhA–yfp
Bacillus Genetic Stock Center
Lewis & Marston (1999)
Lewis & Marston (1999)
Feucht & Lewis (2001)
Feucht & Lewis (2001)
MunI–EcoRI digest of atpA inserted into EcoRI-cut pSG1164
EcoRI–SpeI digest of pSG1186 inserted into the large fragment of
similarly cut pNG24
Acc651–XhoI digest of 39 terminal portion of sdhA inserted into
similarly cut pSG1164
XhoI–SpeI digest of pSG1187 inserted into the large fragment of
similarly cut pNG107
Acc651–SpeI sdhA–yfp fragment from pNG115 inserted into large
fragment of similarly cut pSG1170
B. subtilis
168
BS23
BS24
BS112
BS121
BS131
BS133
2816
(cat9–spc–9cat)
cat Pxyl–gfp
cat lacI Pspac–gfpUV
cat cfp
cat yfp
cat Pxyl–9atpA–gfp
cat Pxyl–9atpA–cfp
Meijer et al. (2001)
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
Microbiology 150
Localization of membrane proteins in B. subtilis
set 31044V2, YFP with set 41029, and FM4-64 was visualized using
set 61000v2SBX with a 61560 TRITC exciter (all sets from Chroma
Technology). Cells were mounted onto 1?2 % agarose pads as
described by Glaser et al. (1997) or within Gene Frames (Advanced
Biotechnologies) as described by Lewis et al. (2000) for microscopy.
Image acquisition was performed using MetaMorph version 5.0
(UIC); backgrounds were subtracted, and out-of-focus light
removed, using the Nearest Neighbours deconvolution drop-in.
Linescans were performed in MetaMorph and data transferred to
Microsoft Excel for further analysis. Final figures were prepared for
publication using Adobe Photoshop version 7.0.
3D imaging and deconvolution. Image stacks at 20 nm incremental
steps were collected using a Pifoc PI P-721.10 microscope focus
drive (Physik Instrumente) controlled via MetaMorph. Background
subtractions and image stack alignments were carried out in
MetaMorph and image stack deconvolution in AutoDeblur version
6.0 (AutoQuant Imaging). 3D reconstruction was performed in
MetaMorph.
Time lapse. For time-lapse microscopy, exponentially growing cells
were placed onto 1?2 % agarose pads in Gene Frames and a suitable
field of cells was located by phase-contrast microscopy. Cells
were then imaged using the ACQUIRE TIMELAPSE drop-in from
MetaMorph with a sampling interval of 1 min. Stacks were aligned
and images processed as detailed above prior to analysis.
Fluorescence recovery after photobleaching (FRAP). FRAP was
carried out using a Zeiss LSM 510 confocal microscope fitted with
an Orca ER CCD camera (Hammamatsu) using a 1006 PlanApo
NA1.3 objective to view exponentially growing cells mounted on
an inverted agarose pad. GFP fluorescence was viewed using the
488 nm laser line at 10 % full power. A region of interest was
bleached using 40 ms pulses at 40 % power over a 2 s exposure
period. One-second exposures were obtained at the intervals specified in the text during photobleaching recovery. Twelve-bit images
were analysed in MetaMorph.
RESULTS
Fig. 1. Construction of functional GFP fusions to atpA and
sdhA. (A) and (B) show the clockwise arrangement of strains
BS23 (atpA–gfp), BS112 (sdhA–gfp), BS121 (sdhA–yfp) and
BS133 (atpA–cfp and sdhA–yfp) grown on SMM plates supplemented with xylose (A) or IPTG (B), grown at 37 6C. Strains
BS23, BS112 and BS133 show xylose-dependent growth
whereas strain BS121 shows IPTG-dependent growth. (C)
Growth curve of strain BS23 grown in SMM liquid medium in
the presence (&) or absence (m) of 0?5 % (w/v) xylose.
atpA– and sdhA–gfp fusions are functional
It is important to ascertain whether a fluorescent protein
fusion is functional, partially functional or non-functional
when making conclusions about observed localization
patterns. This is particularly important when considering
the localization pattern of fusions involved in the formation of multi-protein complexes such as ATP synthase and
succinate dehydrogenase. Both atpA and sdhA, to which
fusions were made, lie within operons, encoding ATP
synthase and succinate dehydrogenase, respectively, and so
on single crossover of fusions into the chromosome it was
important to ensure the expression of genes downstream
of the insertion through use of either xylose- or IPTGinducible promoters (Lewis & Marston, 1999). The growth
of strains BS23 (atpA–gfp), BS112 (sdhA–gfp), BS121 (sdhA–
yfp) and BS133 (atpA–cfp sdhA–yfp) (Table 1) was tested
on SMM agar plates. With succinate as the only carbon
source, these strains should be dependent on a functional
electron-transport chain and ATP synthesis via ATP synthase. Growth of the strains on SMM supplemented with
either xylose or IPTG is shown in Fig. 1(A, B). Xylose was
required for induction of expression of genes downstream
http://mic.sgmjournals.org
of the fluorescent protein fusions in strains BS23, BS112
and BS133, whereas IPTG was required in strain BS121.
In the presence of xylose, extensive growth of strains
BS23, BS112 and BS133 was observed (Fig. 1A), indicating
expression of the fusion, and the genes downstream in both
the atp and sdh operons led to formation of functional
protein complexes. These results were confirmed by growing strain BS23 in liquid SMM (Fig. 1C). In the presence
of xylose, the fusion strain grew with very similar kinetics
to the control strain (168; td approximately 85 min; not
shown), whereas very little growth was observed in the
absence of xylose. In addition, Western blots of whole-cell
extracts together with cytoplasmic and membrane fractions
showed that there was no detectable degradation of the
fusions, which localized exclusively to the membrane
fraction (data not shown). Virtually no growth of BS121
containing an IPTG-inducible promoter downstream of
an sdhA–yfp insertion was observed (Fig. 1A), indicating
the dependence of these strains on the formation of functional enzyme complexes for growth. Conversely, when
these strains were grown on SMM supplemented with IPTG
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
2817
A. S. Johnson, S. van Horck and P. J. Lewis
instead of xylose, growth was only observed for strain BS121
(Fig. 1B).
Together, these results indicate that functional fusions to
both AtpA and SdhA were constructed. It was possible that
xylose could be used as an alternative carbon source in these
strains and so permit growth by bypassing the requirement for functional electron transport and ATP synthase.
However, the lack of growth of strain BS121 on SMM
supplemented with xylose, and growth of the same strain on
SMM supplemented with the non-metabolizable galactose
analogue IPTG, indicate that this does not occur.
Heterogeneous localization of integral
membrane proteins
The vital membrane stain FM4-64 has been used for
many years as an indicator of the cell boundaries and cell
membrane in cell biology and shows the membrane to be
a smooth homogeneous structure enveloping the cytoplasm (e.g. Pogliano et al., 1999; Lewis et al., 2000). When
exponentially growing cells of strain 168 (Table 1) were
stained with FM4-64 a homogeneous band outlining the
cells was observed, and this was confirmed when a linescan
through the top border of the cells was performed (Fig. 2A).
The peaks in the trace correspond to the division septa
that contain two membranes and so appear more heavily
Fig. 2. Subcellular localization of membrane proteins. The lefthand panels show fluorescence micrographs and the right-hand
panels show linescans through the uppermost membrane
margin of chains of four cells from the corresponding micrograph. Arrows indicate the start points for the linescans. (A)
Cells stained with the membrane dye FM4-64; (B) cells expressing the atpA–gfp fusion; (C) cells expressing the sdhA–gfp
fusion; (D) cells expressing the p16.7–gfp fusion.
2818
stained. When exponentially growing cells of strains BS24
(atpA–gfp) and BS112 (sdhA–gfp) were visualized using the
same image processing techniques a much more heterogeneous staining pattern was observed (Fig. 2B, C). To
test the prevalence of this phenomenon, the localization of
a completely unrelated integral membrane protein, p16.7
from the B. subtilis phage w29, was tested. In the absence
of other phage proteins, p16.7 is simply a small integral
membrane protein with no known role in B. subtilis
physiology. The localization of a p16.7–GFP fusion has
been previously reported and shown to be confined to the
cytoplasmic membrane in a pattern very similar to FM4-64
(Meijer et al., 2001). However, we found that the p16.7–
GFP localization pattern was indistinguishable from that
of either SdhA–GFP or AtpA–GFP (Fig. 2D), suggesting
that this punctate distribution pattern was most likely a
general feature for the localization of integral membrane
proteins. An explanation for the apparently different localization pattern for p16.7 found by us and by Meijer et al.
(2001) is given in the Discussion.
Although the above results indicate that membrane proteins preferentially localize to submembranous domains,
they do not tell us whether this localization is to the same
or different domains. The rather punctate pattern of distribution was intriguing, and it was possible that electron
transport (succinate dehydrogenase) and ATP synthesis
were closely juxtaposed within the cell due to their connected roles in energy generation. There is some circumstantial evidence for close juxtapositioning of these enzymes,
as it is possible to isolate respiratory chain supercomplexes
containing various combinations of components of the
electron-transport chain and ATP synthase (Schägger,
2002). In order to address this issue, dual-labelled cells
were constructed containing atpA–cfp and sdhA–yfp fusions
(strain BS133; Table 1). In this strain the fusions were
inserted by single crossover into the chromosomal locus,
similar to their respective single-labelled strains, with
expression of genes downstream of both atpA and sdhA
dependent on the presence of xylose (see Fig. 1A, B). For
reasons we cannot explain, this appeared to be the only
combination of fusions that gave reproducibly bright
signals, despite numerous attempts to construct strains
in which one fusion was expressed ectopically from the
amyE locus or when one of the fusions was placed under
the control of the IPTG-inducible Pspac promoter, which
we know produces functional fusions (Fig. 1A, B) and
gives bright signals in single-labelled strains (not shown).
Nevertheless, chromosomal PCR and fluorescence microscopy checks indicated that strain BS133 contained the
correct constructs in the right chromosomal context, and
growth on SMM supplemented with xylose indicated that
the strain produced functional fluorescent protein fusions
(Fig. 1A).
Fluorescence micrographs of strain BS133 grown in CH
medium supplemented with 0?5 % (w/v) xylose at 37 uC
are shown in Fig. 3. Fluorescence crossover experiments
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
Microbiology 150
Localization of membrane proteins in B. subtilis
Fig. 3. Colocalization of AtpA–CFP and SdhA–YFP. (A) AtpA–CFP fluorescence; (B) SdhA–YFP fluorescence; (C) an
overlay of the two signals. AtpA–CFP has been pseudocoloured red and SdhA–YFP pseudocoloured green. A linescan
around the pair of rightmost cells indicated with the arrow using the same colour assignment is shown in (D); the arrows
above the linescan indicate regions of signal colocalization.
confirmed that there was no detectable crossover of CFP
signal into the YFP channel and vice versa (not shown). The
fluorescence patterns of both CFP-labelled ATP synthase
and YFP-labelled succinate dehydrogenase appear very
similar to that observed for the single-labelled strains
(compare Fig. 2B and C with Fig. 3A and B). The ATP
synthase signal was pseudocoloured red, while the succinate
dehydrogenase was pseudocoloured green in the overlay
shown in Fig. 3(C). Usually, when red and green signals
of approximately equal intensity coincide in overlays, a
yellow colour is observed. While yellow can be seen in the
overlay, there are clearly regions of either green or red signal,
implying that there was relatively little colocalization of
the signals. However, careful examination of the linescan
shown in Fig. 3(D) indicates that there were substantial
regions of signal overlap, but that there was considerable
variation in the intensity of red or green signal at some
regions of overlap that may have given rise to the appearance
of a red or green region in the overlay in Fig. 3(C) (see
regions arrowed in Fig. 3D). However, the linescan also
shows that the signals are not perfectly coincident, with
considerable regions of signal heterogeneity. Analysis of
multiple fields of cells indicated a mean level of 62 % signal
overlap with respect to the number of AtpA-CFP peaks.
http://mic.sgmjournals.org
Overall, we feel these results are consistent with the
localization of both proteins to submembranous regions,
with both enzymes present in similar, at least partially
overlapping, domains.
Integral membrane proteins localize in large
amorphous patches
A more detailed analysis of the localization of integral
membrane proteins was carried out using image stack
deconvolution and 3D reconstruction. Results are presented
for the AtpA–GFP fusion, but SdhA and p16.7 fusions have
similar localization patterns.
A 3D reconstruction of ATP synthase localization in a pair
of cells in a series of orientations is shown in Fig. 4(A) and
as an animation in supplementary movie 1 with the online
version of this paper at http://mic.sgmjournals.org. The
3D rendition clearly shows heterogeneously distributed
ATP synthase over the membranes. There is no clearly
ordered organization of the enzyme; rather it appears to be
distributed in a series of discrete foci and larger heterogeneous patches. A more detailed analysis of the image
stacks was used to confirm this observation. Three image
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
2819
A. S. Johnson, S. van Horck and P. J. Lewis
of recently described actin-like cytoskeletal filaments (see
Discussion; Jones et al., 2001; Kruse et al., 2003).
Protein mobility within the membrane
Fig. 4. 3D localization of ATP synthase. (A) A contrast-inverted
3D rendering of ATP synthase localization. The numbers indicate the degrees of rotation of the image. (B) Three processed
image slices taken from near the top (red), middle (green) and
bottom (blue) of the cell. An overlay of the three slices is
shown in the fourth panel. (C) A linescan around the margin of
the overlaid image; the arrow indicates the start point of the
scan. Colour coding is the same as in (B). In (B) and (C), ‘a’
and ‘b’ refer to a diffuse domain and discrete focus, discussed
in more detail in the text.
slices taken from positions indicated in the top right panel
corners are shown in Fig. 4(B). A slice from towards the
top of the cell is shown in red, from the middle in green,
and towards the bottom in blue. These three slices were
overlaid (Fig. 4B bottom right panel) and a linescan made
around the periphery of the pair of cells (Fig. 4C). The
arrows in Fig. 4(B) and (C) indicate the position where
the linescan starts. A region we interpret as representing
a large patch of ATP synthase that extends rightwards
from the top of the cell to the bottom is indicated by the
‘a’ in Fig. 4(B, C). A more defined region that we interpret
as representing a focus is indicated by the ‘b’ in Fig. 4(B, C).
This analysis confirmed the heterogeneity of localization
of ATP synthase and there was no evidence for a specific
pattern of protein localization, such as aggregation at cell
poles/mid-cells or in spirals that could reflect the distribution
2820
Given the heterogeneous localization of proteins in the
membrane, we wanted to know whether the observed
localization represented protein restricted to specific
domains within the cell membrane, or whether proteins
were free to diffuse around the membrane. Time-lapse
movies of the AtpA–GFP fusion strain were obtained at
1 min intervals over 30 min. A full movie sequence is
provided in supplementary movie 2 with the online version
of this paper (http://mic.sgm.journals.org), and a more
detailed analysis of a 4 min period of the time-course is
shown in Fig. 5. It is clear from both the movie and Fig. 5
that the localization of AtpA–GFP is highly dynamic
within the membrane. Images taken at 0, 2 and 4 min
into the time-lapse are shown in red, green and blue,
respectively in Fig. 5(A) along with an overlay of the three
images. The lack of coincidence of the signals in the overlay
panel presents a graphic illustration of the movement of
AtpA–GFP over the imaging period. A linescan around the
cell, starting at the point indicated by the arrow, is shown
in Fig. 5(B). The movement of a small focus of AtpA–GFP,
marked ‘a’, ‘b’ and ‘c’ in the red, green and blue panels,
respectively, was mapped for further analysis (Fig. 5A, B).
This particular focus appears to start near the cell pole (a),
diffuse slightly and move away from the pole (b) then move
back towards the pole (c), suggesting that movement was
due to random diffusion through the membrane. However,
interpretation of these images is complicated, with unequivocal assignment of foci in each panel difficult. It may
be that the focus mapped in ‘b’ and ‘c’ starts at ‘a9’ rather
than ‘a’ (Fig. 5A, B). If this is the case, then the time-lapse
images suggest that movement is unidirectional within the
membrane. However, it is not possible to draw such
conclusions from this experiment as it is unlikely that all
the images were obtained in exactly the same focal plane.
As indicated by the 3D reconstruction images in Fig. 4, a
small change in focal plane could result in focusing on a
very different distribution pattern for AtpA–GFP. In order
to resolve the issue of whether movement within the membrane was random or directed, we performed fluorescence
recovery after photobleaching (FRAP) experiments using
a confocal microscope system. Following bleaching of a
portion of a cell, the recovery of fluorescence could be
monitored to determine if it is uni- or bi-directional.
Unidirectional recovery would imply that Atp–GFP moved
around the observed plane of the cell in one direction,
whereas bi-directional recovery implies random movement
of Atp–GFP.
The results of a FRAP experiment are shown in Fig. 6.
Images of cells before, immediately following, 3 and
6 min after photobleaching are shown in Fig. 6(A).
Linescans from the top right cell in Fig. 6(A) are shown
in Fig. 6(B), with the arrow indicating the start of the
linescan. Linescans were made down the left-hand (L) and
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
Microbiology 150
Localization of membrane proteins in B. subtilis
Fig. 5. ATP synthase is highly mobile within the membrane. (A) Images of ATP synthase localization taken at the start (red),
and 2 min (green) and 4 min (blue) into a time-lapse series. An overlay of the three images is shown in the fourth panel. (B) A
linescan around the margin of the overlaid image, starting at the point indicated by the arrow. Colour coding is the same as in
(A). Letters ‘a’, ‘a9’, ‘b’ and ‘c’ are referred to in more detail in the text.
right-hand (R) sides of the cell and are shown in Fig. 6(B).
The colour scheme used in Fig. 6(A) is retained in Fig. 6(B)
for comparative purposes with the region that was bleached
lying to the right of the dashed line in the linescans. The
cells indicated by ‘a’ in Fig. 6(A) were totally bleached,
and the lack of fluorescence recovery in these cells indicates
that there was no detectable de novo synthesis of AtpA–
GFP over the course of this experiment.
Following photobleaching, the intensity of fluorescence in
the bottom half of the analysed cell dropped considerably
(compare the red and green panels in Fig. 6A and the red
and green linescans in Fig. 6B). At 6 min following photobleaching, fluorescence recovery throughout the cell appeared
to be largely complete (cyan panel and linescan in Fig. 6A
and B). The image taken 3 min following photobleaching
represents an intermediate stage between bleaching and
recovery (blue panel and linescan Fig. 6A and B). Comparison of the linescans through the left- and right-hand side
of the cell indicated that the rate of fluorescence recovery
was approximately the same on both sides (Fig. 6B), which
was observed in all FRAP experiments. This result suggests
that AtpA–GFP movement within the membrane is most
likely due to random diffusion.
DISCUSSION
We have shown in this study that microbial membrane
proteins appear to be restricted to domains rather than
http://mic.sgmjournals.org
being homogeneously distributed within the cytoplasmic
membrane. This appears to be a general property, which
was analysed with GFP-tagged subunits of ATP synthase
(AtpA) and succinate dehydrogenase (SdhA) as well as the
B. subtilis phage w29 protein p16.7. All the proteins appeared
to have a similar heterogeneous pattern of localization
that was strikingly different from the distribution of total
membrane when observed with the membrane stain FM464. The localization pattern for p16.7 has previously been
reported and described as being similar to that of FM4-64
(Meijer et al., 2001). Indeed, in this work we used the same
strain, 110WA, to observe p16.7 localization. However, we
do not feel that our results contradict those previously
reported; rather, our analysis and interpretation has been
more focused on the signal heterogeneity. In addition, our
images were processed to remove out-of-focus light (see
Methods) whereas Meijer et al. (2001) used unprocessed
images. Both sets of results indicate that p16.7 is distributed
within the cytoplasmic membrane, and it is possible to
observe signal heterogeneity in the insert box of Fig. 5 in
Meijer et al. (2001), indicating that strain 110WA gives
similar results in our hands to those of Meijer et al. (2001).
Dual-labelling experiments with AtpA–CFP and SdhA–
YFP fusions suggested that membrane proteins localize
to approximately similar submembranous domains, as
observed by the partial colocalization of AtpA-CFP and
ShdA-YFP signals (Fig. 3). However, the degree of colocalization of these signals also indicated that it is very
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
2821
A. S. Johnson, S. van Horck and P. J. Lewis
Fig. 6. FRAP of the AtpA–GFP fusion. (A) A group of cells analysed in a FRAP experiment. Cells prior to bleaching are
shown in red, immediately after bleaching in green, 3 min post-bleach in blue, and 6 min post-bleach in cyan. (B) Linescans of
the cell indicated by an arrow in (A) down the left and right hand margins, indicated by L and R, respectively in (A). The colour
coding is the same as in (A). The letter ‘a’ indicates a pair of cells that were totally photobleached and failed to redevelop
fluorescence during the experiment (see text for more details). The dotted line in the linescans in (B) indicates the approximate
rightmost limit of photobleaching.
unlikely that ATP synthase and succinate dehydrogenase
aggregate together in specific ‘energy-generating’ domains,
but rather that membrane proteins segregate to domains
that are approximately similar. Heterogeneities observed
between the two signals could be due to differences in
phospholipid preferences for the membrane complexes
and/or the levels of expression of the genes. ATP synthase
fusions were extremely bright in comparison to the other
fusions, possibly indicating a higher level of expression
of this operon, and it has been reported that the level of
expression of the atp operon is extremely high (Santana
et al., 1994).
3D reconstruction of ATP synthase distribution indicated
that protein domains varied between small concentrated
foci to rather large diffuse domains that spread over a
significant proportion of the cell (Fig. 4 and supplementary
movie 1). These imaging experiments were performed in
live, non-fixed cells and so it is possible that the domains
were moving during the image acquisition process. This
did probably occur, although we feel that movement was
not large during the acquisition process. At 200 ms exposures, 60 images could be acquired in little over 12 s. As
seen in Figs 5 and 6, movement was monitored over a period
2822
of minutes, not seconds. Time-lapse movies taken with
short (5 s) gaps showed little observable movement of
protein domains over a 10–15 s period (see supplementary
Fig. S1 at http://mic.sgm.journals.org). Therefore, we feel
the results presented in Fig. 4 and supplementary movie
1 do provide a reasonable snapshot of ATP synthase and
other integral membrane protein distribution around the
membrane.
We also analysed the dynamic distribution of ATP synthase in more detail, and time-lapse microscopy experiments showed that the irregular protein domains were
highly mobile within the membrane (Fig. 5). It was not
possible to conclude from these experiments whether movement within the membrane was unidirectional (specific) or
bidirectional (random). Further experiments (Fig. 6) monitoring the recovery of fluorescence around the membrane
in cells where half the membrane was bleached indicated
that movement of the protein domains was most likely by
random diffusion.
In eukaryotic cells, lipid rafts are known to have contacts
with the actin cytoskeleton via actin-binding proteins such
as annexins (Edidin, 2003), and many proteins are known
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
Microbiology 150
Localization of membrane proteins in B. subtilis
to interact with actin filaments, including enzymes involved
in glycolysis (Minaschek et al., 1992). It has recently become
clear that bacteria also contain filamentous structures composed of proteins that belong to the actin superfamily, and
that these cytoskeletal filaments form regular helical structures that are probably closely juxtaposed with the cytoplasmic membrane (Jones et al., 2001; Kruse et al., 2003).
If there were discrete ‘energy-generating domains’ within
the cell, it is possible that glycolysis, the tricarboxylic acid
cycle, electron transport and ATP synthase could be closely
juxtaposed. However, we found no evidence for the proteins examined in this study localizing in a pattern consistent with a connection with cytoskeletal filaments. In
addition, we found our protein patches to be highly mobile
within the membrane, with substantial movements observed
over a period of a few minutes. FRAP of the Mbl cytoskeleton in B. subtilis occurred over a period of tens of
minutes (Carballido-López & Errington, 2003), indicating
that the dynamics of protein movement in membranes and
cytoskeletons occurs on different timescales. Most likely,
integral membrane proteins are simply inserted into the
membrane directly via translation from ribosomes with
little or no further connection with the cytoplasm. We do
know that ribosomes preferentially localize towards the
cell poles (Lewis et al., 2000), and membrane protein
localization does not reflect a similar polar bias. However,
once inserted into the membrane, it is clear that most
proteins (or those that do not have a specific function
dependent on their subcellular localization) are free to
diffuse within the membrane and so would have a
localization pattern unrelated to their site of synthesis.
In conclusion, integral membrane protein localization is
probably dependent on the lipid distribution within the
membrane, and is highly dynamic and random. Given the
apparent preference of these proteins for submembranous
domains, it will be interesting to determine whether formation of these protein-rich domains is correlated with lipid
raft distribution.
ACKNOWLEDGEMENTS
de Mendoza, D., Schujman, G. E. & Aguilar, P. S. (2002).
Biosynthesis and function of membrane lipids. In Bacillus subtilis
and its Closest Relatives: from Genes to Cells, pp. 43–56. Edited by
A. L. Sonenshine, J. A. Hoch & R. Losick. Washington, DC:
American Society for Microbiology Press.
Edidin, M. (2003). The state of lipid rafts: from model membranes
to cells. Annu Rev Biophys Biomol Struct 32, 257–283.
Feucht, A. & Lewis, P. J. (2001). Improved plasmid vectors for the
production of multiple fluorescent protein fusions in Bacillus subtilis.
Gene 264, 289–297.
Fishov, I. & Woldringh, C. (1999). Visualization of membrane
domains in Escherichia coli. Mol Microbiol 32, 1166–1172.
Glaser, P., Sharpe, M. E., Raether, B., Perego, M., Ohlsen, K. &
Errington, J. (1997). Dynamic, mitotic-like behaviour of a bacterial
protein required for accurate chromosome partitioning. Genes Dev
11, 1160–1168.
Jenkinson, H. (1983). Altered arrangement of proteins in the spore
coat of a germination mutant of Bacillus subtilis. J Gen Microbiol 129,
1945–1958.
Jones, L. J. F., Carballido-López, R. & Errington, J. (2001). Control
of cell shape in bacteria: helical, actin-like filaments in Bacillus
subtilis. Cell 104, 913–922.
Kadner, R. J. (1996). Cytoplasmic membrane. In Escherichia coli
and Salmonella: Cellular and Molecular Biology 2nd edn, pp 58–87.
Edited by F. C. Neidhardt and others. Washington DC: American
Society for Microbiology.
Kawai, F., Shoda, M., Harashima, R., Sadaie, Y., Hara, H. &
Matsumoto, K. (2004). Cardiolipin domains in Bacillus subtilis
Marburg membranes. J Bacteriol 186, 1475–1483.
Kruse, T., Møller-Jensen, J., Løbner-Oelsen, A. & Gerdes, K.
(2003). Dysfunctional MreB inhibits chromosome segregation in
Escherichia coli. EMBO J 22, 5283–5292.
Lewis, P. J. & Marston, A. L. (1999). GFP vectors for controlled
expression and dual labelling of protein fusions in Bacillus subtilis.
Gene 227, 101–109.
Lewis, P. J., Thaker, S. D. & Errington, J. (2000). Compart-
mentalization of transcription and translation in Bacillus subtilis.
EMBO J 19, 710–718.
Lopian, L., Nussbaum-Schochat, A., O’Day-Kerstein, K.,
Wright, A. & Amster-Choder, O. (2003). The BglF sensor
recruits the BglG transcription regulator to the membrane
and releases it on stimulation. Proc Natl Acad Sci U S A 100,
7099–7104.
Maddock, J. R. & Shapiro, L. (1993). Polar location of the
This work was supported by the Australian Research Council through
grants A00105184 and DP0449482 and University of Newcastle RMC
grants to P. L. P. L. acknowledges the help of Scott Merrington and
Eileen McLaughlin with confocal imaging and FRAP experiments.
chemoreceptor complex in the Escherichia coli cell. Science 19,
1717–1723.
Meijer, W. J. J., Serna-Rico, A. & Salas, M. (2001). Characterisation
of the bacteriophage w29-encoded protein p16.7: a membrane
protein involved in phage DNA replication. Mol Microbiol 39,
731–746.
Mileykovskaya, E. & Dowhan, W. (2000). Visualization of phos-
REFERENCES
transformation in Bacillus subtilis. J Bacteriol 81, 741–746.
pholipid domains in Escherichia coli by using the cardiolipinspecific fluorescent dye 10-N-nonyl acridine orange. J Bacteriol 182,
1172–1175.
Brandon, L. D., Goehring, N., Janakirman, A., Yan, A. W., Wu, T.,
Beckwith, J. & Goldberg, M. B. (2003). IcsA, a polarly localized
Minaschek, G., Groschel-Stewart, U., Blum, S. & Bereiter-Hahn, J.
(1992). Microcompartmentation of glycolytic enzymes in cultured
autotransporter with an atypical signal peptide, uses the Sec
apparatus for secretion, although the Sec apparatus is circumferentially distributed. Mol Microbiol 50, 45–60.
Pogliano, J., Osborne, N., Sharp, M. D., Abanes-De Mello, A.,
Perez, A., Sun, Y. L. & Pogliano, K. (1999). A vital stain for studying
Anagnostopoulos, C. & Spizizen, J. (1961). Requirements for
Carballido-López, R. & Errington, J. (2003). The bacterial cyto-
skeleton: in vivo dynamics of the actin-like protein Mbl of Bacillus
subtilis. Dev Cell 4, 19–28.
http://mic.sgmjournals.org
cells. Eur J Cell Biol 58, 418–428.
membrane dynamics in bacteria: a novel mechanism controlling
septation during Bacillus subtilis sporulation. Mol Microbiol 31,
1149–1159.
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
2823
A. S. Johnson, S. van Horck and P. J. Lewis
Santana, M., Ionescu, M. S., Vertes, A., Longin, R., Kunst, F.,
Danchin, A. & Glaser, P. (1994). Bacillus subtilis F0F1 ATPase: DNA
Sharpe, M. E., Hauser, P. M., Sharpe, R. G. & Errington, J.
(1998). Bacillus subtilis cell cycle as studied by microscopy:
sequence of the atp operon and characterization of atp mutants.
J Bacteriol 176, 6802–6811.
Schägger, H. (2002). Respiratory chain supercomplexes of mitochondria and bacteria. Biochim Biophys Acta 155, 154–159.
Scheffers, D.-J., Jones, L. J. F. & Errington, J. (2004). Several distinct
localization patterns for penicillin-binding proteins in Bacillus
subtilis. Mol Microbiol 51, 749–764.
constancy of cell length at initiation of DNA replication
and evidence for active nucleoid partitioning. J Bacteriol 180,
547–555.
2824
Vanounou, S., Parola, A. H. & Fishov, I. (2003). Phosphatidyl-
ethanolamine and phosphatidylglycerol are segregated into different
domains in bacterial membrane. A study with pyrene-labelled
phospholipids. Mol Microbiol 49, 1067–1079.
Downloaded from www.microbiologyresearch.org by
IP: 88.99.165.207
On: Mon, 31 Jul 2017 18:32:39
Microbiology 150