School of Environmental and Life Sciences, Biological Sciences, University of Newcastle, Callaghan, NSW 2308, Australia
Correspondence
P. J. Lewis
Peter.Lewis{at}newcastle.edu.au
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ABSTRACT |
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The online version of this paper at http://mic.sgmjournals.org has supplementary movie files and a supplementary figure.
Present address: Life Therapeutics, PO Box 6126, Frenchs Forest, NSW 2086, Australia.
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INTRODUCTION |
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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. Penicillin-binding 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
29 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 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.
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METHODS |
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DNA manipulations.
The complete atpA gene was PCR amplified using the following primers: forward 5'-AGAAGGGGTGAACTCGAGGTGAGCATCAAA-3' and reverse 5'-TCATCAGCATTCGAATTCGTTGCTTGGTGC-3'. The resulting product was digested with MunI and EcoRI and the 840 bp 3' fragment inserted into EcoRI-digested pSG1164. One recombinant with the fragment inserted in the correct orientation was named pNG24 (Table 1).
The 3' 1073 bp of sdhA were PCR amplified using the following primers: forward 5'-CGTATTATGGGTACCGAGAATTCATTC-3' and reverse 5'-GTTCACTCATGACTCGAGCGCCACCTTCTT-3'. 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 IPTG-inducible 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 100x ApoPlan NA 1.4 lens. GFP fluorescence was visualized with set 41018, CFP with 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 100x 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.
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RESULTS |
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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. 2
A). The peaks in the trace correspond to the division septa that contain two membranes and so appear more heavily stained. When exponentially growing cells of strains BS24 (atpAgfp) and BS112 (sdhAgfp) 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
29, 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 p16.7GFP 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 p16.7GFP localization pattern was indistinguishable from that of either SdhAGFP or AtpAGFP (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.
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Fluorescence micrographs of strain BS133 grown in CH medium supplemented with 0·5 % (w/v) xylose at 37 °C are shown in Fig. 3. Fluorescence crossover experiments 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. 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.
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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 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 of recently described actin-like cytoskeletal filaments (see Discussion; Jones et al., 2001
; Kruse et al., 2003
).
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DISCUSSION |
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Dual-labelling experiments with AtpACFP and SdhAYFP 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 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 of minutes, not seconds. Time-lapse movies taken with short (5 s) gaps showed little observable movement of protein domains over a 1015 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 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.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 8 April 2004;
revised 27 May 2004;
accepted 27 May 2004.
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