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Journal of Bacteriology, March 2004, p. 1475-1483, Vol. 186, No. 5
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.5.1475-1483.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Cardiolipin Domains in Bacillus subtilis Marburg Membranes

Fumitaka Kawai, Momoko Shoda, Rie Harashima, Yoshito Sadaie, Hiroshi Hara, and Kouji Matsumoto*

Department of Biochemistry and Molecular Biology, Saitama University, 255 Shimo-ohkubo, Sakura, Saitama, Saitama 338-8570, Japan

Received 31 July 2003/ Accepted 24 November 2003


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ABSTRACT
 
Recently, use of the cardiolipin (CL)-specific fluorescent dye 10-N-nonyl-acridine orange (NAO) revealed CL-rich domains in the Escherichia coli membrane (E. Mileykovskaya and W. Dowhan, J. Bacteriol. 182: 1172-1175, 2000). Staining of Bacillus subtilis cells with NAO showed that there were green fluorescence domains in the septal regions and at the poles. These fluorescence domains were scarcely detectable in exponentially growing cells of the clsA-disrupted mutant lacking detectable CL. In sporulating cells with a wild-type lipid composition, fluorescence domains were observed in the polar septa and on the engulfment and forespore membranes. Both in the clsA-disrupted mutant and in a mutant with disruptions in all three of the paralogous genes (clsA, ywjE, and ywiE) for CL synthase, these domains did not vanish but appeared later, after sporulation initiation. A red shift in the fluorescence due to stacking of two dye molecules and the lipid composition suggested that a small amount of CL was present in sporulating cells of the mutants. Mass spectrometry analyses revealed the presence of CL in these mutant cells. At a later stage during sporulation of the mutants the frequency of heat-resistant cells that could form colonies after heat treatment was lower. The frequency of sporulation of these cells at 24 h after sporulation initiation was 30 to 50% of the frequency of the wild type. These results indicate that CL-rich domains are present in the polar septal membrane and in the engulfment and forespore membranes during the sporulation phase even in a B. subtilis mutant with disruptions in all three paralogous genes, as well as in the membranes of the medial septa and at the poles during the exponential growth phase of wild-type cells. The results further suggest that the CL-rich domains in the polar septal membrane and engulfment and forespore membranes are involved in sporulation.


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INTRODUCTION
 
The bacterial cell membrane is widely recognized as a matrix in which lipid molecules are homogeneously distributed. However, it has been noticed that lipid molecules are heterogeneously distributed in bacterial membranes, and the observations increasingly include those obtained by using fluorescent lipophilic probes. Immunoelectron microscopic observations showing the polar localization of the chemoreceptor complexes in Caulobacter crescentus and Escherichia coli cells provided early indications of membrane heterogeneity (1, 27). By using the lipophilic fluorescent styryl dye FM4-64, laterally uneven distribution of the fluorescence, which could be an indication of heterogeneous distribution of phospholipids in E. coli membranes, was then discovered (14). Recently, the cardiolipin (CL)-specific fluorescent dye 10-N-nonyl-acridine orange (NAO) was used to demonstrate that there are CL-containing domains in E. coli membranes, which were observed mostly in the septal regions and at the poles of the cells (31, 32). The hypothesis that there are CL-containing domains in these regions of E. coli cells was supported by an analysis of the lipid composition of minicells, which consist mainly of polar materials of the envelope (24).

CL in the presence of certain divalent cations, as well as phosphatidylethanolamine, has the potential to form nonbilayer structures, which could introduce discontinuity into the bilayer membrane structures for dynamic membrane functions, such as membrane fusion during cell division, formation of adhesion sites between the outer and inner membranes, integration of proteins into the membrane, and stabilization of membrane proteins (12). Mutants of E. coli lacking either CL or phosphatidylethanolamine are viable, but construction of a mutant lacking both phospholipids is not possible (8, 34, 41), suggesting that a common structural feature of the lipids, the potential to form nonbilayer structures, is required. Additionally, CL and phosphatidylglycerol, both of which have an anionic nature, play a role in recruitment of membrane proteins having positively charged amphitropic {alpha}-helices onto the anionic surface of the membrane (12, 29). In spite of the anticipated roles, the significance of CL in vivo is still clouded by its dispensability (23, 29, 34).

The Bacillus subtilis membrane undergoes dynamic rearrangements, which include formation of polar septa and engulfment and forespore membranes, during the sporulation process in addition to the rearrangements that occur during cell division during vegetative growth. The membranes contain CL (9, 30), which could play important roles in the processes, but little is known about the anticipated roles and the genes responsible for biosynthesis of this compound. B. subtilis has three homologues (ywnE, ywjE, and ywiE) (25) of E. coli cls, the structural gene for CL synthase, but the contribution of these genes to CL synthesis has not been examined previously. Here, we characterized mutants with disruptions in each or all three of these homologues. We found that ywnE plays a dominant role in CL synthesis; thus, ywnE is renamed clsA. We also found a small but significant amount of CL in the sporulation-phase cells of the mutant with disruptions in all three genes. By using NAO staining, we found that B. subtilis cells contain CL-rich domains in the polar septal membrane and in the engulfment and forespore membranes in the sporulation phase, as well as in the membrane of the medial septa and at the poles in vegetative growth phase. We suggest that the CL-containing domains in the former membranes are involved in sporulation.


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MATERIALS AND METHODS
 
Bacterial strains and plasmids. The B. subtilis Marburg and E. coli K-12 strains and the plasmids used in this study are listed in Table 1. The strains with disrupted alleles of clsA, ywjE, and ywiE were constructed as follows. Plasmids pJESPC and pIENEO with the disrupted alleles ywjE1::spc and ywiE2::neo, respectively, were first constructed with pBR322, and wild-type strain 168 was transformed with the plasmid DNA to obtain SDB201 (ywjE1::spc) and SDB202 (ywiE2::neo), respectively. BFS219 (clsA::pMutin4) was then transformed with the chromosomal DNA of SDB202 (ywiE2::neo) to form SDB203 (clsA::pMutin4 ywiE2::neo). Finally, SDB203 was transformed with the DNA of SDB201 (ywjE1::spc) to construct SDB206 (clsA::pMutin4 ywiE2::neo ywjE1::spc). The disrupted alleles in the mutant stains were confirmed by the increase in size by using PCR.


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  		TABLE 1. Bacterial strains and plasmids

The pJESPC and pIENEO plasmids were constructed as follows. For construction of pJESPC, the fragment containing ywjE was first obtained by PCR amplification from B. subtilis 168 chromosomal DNA with primer ywjEEco (5'-TATTgAATTCCTGCTGTTCG-3'), which introduced an EcoRI recognition sequence (underlined) with a mismatch (lowercase letter) starting 56 nucleotides (nt) upstream from the initiation codon, and primer ywjESphAS (5'-TCATCGCATgcGAAACGAACCG-3'), which introduced an SphI recognition sequence (underlined) with two mismatches (lowercase letters) starting 112 nt downstream from the termination codon. The fragment was digested with EcoRI and SphI and then inserted into EcoRI-SphI-digested pBR322. The resulting plasmid was digested with HindIII at a site located 646 nt downstream from the initiation codon, and the HindIII fragment containing spc from pDG1726 (16) was inserted to construct pJESPC.

To construct pIENEO, fragment ywiE-F-B (1,590 bp) containing ywiE that had a newly introduced XbaI site 819 nt downstream from the initiation codon was obtained by a second PCR amplification of the first two PCR products (ywiE-F and ywiE-B) with primers ywiEEco and ywiEHinAS. The ywiE-F product was obtained by PCR amplification from the chromosomal DNA by using primer ywiEEco (5'-ACAGAGGgAATTcAATCAGATTGGAG-3'), which introduced an EcoRI recognition sequence (underlined) with two mismatches (lowercase letters) starting 26 nt upstream from the initiation codon, and primer ywiEXbaAS (5'-TATCTCTagAAAATCCGATGTACG-3'), which introduced an XbaI recognition sequence (underlined) with two mismatches (lowercase letters) starting 829 nt downstream from the initiation codon. The ywiE-B product was obtained from the chromosomal DNA by using primer ywiEXba (5'-GATTTTctAGAGATACACACCTGCGGC-3'), which introduced an XbaI recognition sequence (underlined) with two mismatches (lowercase letters) starting 814 nt downstream from the initiation codon, and primer ywiEHinAS (5'-TTCTCAAgctTACTATACACGGGC-3'), which introduced a HindIII recognition sequence (underlined) with three mismatches (lowercase letters) starting 52 nt downstream from the termination codon. The ywiE-F-B fragment was introduced into EcoRI-HindIII-digested pBR322. The resulting plasmid was digested with XbaI, and then the XbaI fragment containing neo from pBEST502 (21) was inserted to form pIENEO.

Cloning of clsA, ywjE, and ywiE into expression vector pSK6 was performed as follows. The clsA fragment was amplified from B. subtilis wild-type chromosomal DNA by using primer nesNcoI (5'-GGGTTACAccaTGgGTATTTCTTCC-3'), which introduced an NcoI site (underlined) starting 11 nt upstream of the initiation codon GTG with four mismatches (lowercase letters), and primer neasXbaI (5'-CGGGGAAGTcTAgATCACTGACG-3'), which introduced an XbaI site (underlined) with two mismatches (lowercase letters) starting 147 nt downstream from the stop codon. The amplified fragment was digested with NcoI and XbaI and then inserted into NcoI-XbaI-digested pSK6 to construct pMMS1. The ywjE fragment was amplified by using primer jesNcoI (5'-GCCGccATGgAGGTATTTATC-3'), which introduced an NcoI site (underlined) starting 6 nt upstream of the initiation codon with three mismatches (lowercase letters), and primer jeasSphI (5'-GGCCTCAAGCATGCGGTA-3'), which had an SphI site (underlined) starting 435 nt downstream of the termination codon. The amplified fragment was digested with NcoI and SphI and inserted into NcoI-SphI-digested pSK6 to construct pMMS2. The ywiE fragment was amplified by using primer iesNcoI (5'-GAGAGAACCAccATGgTGAAAAGG-3'), which introduced an NcoI site (underlined) starting 12 nt upstream of the initiation codon with three mismatches (lowercase letters), and primer ieasHindIII (5'-CCATTAAAgCTtCCGCATCG-3'), which introduced a HindIII site (underlined) with two mismatches (lowercase letters) starting 176 nt downstream of the stop codon. The amplified fragment was inserted into NcoI-HindIII-digested pSK6 to construct pMMS3. These plasmids were introduced into E. coli SD9 cells to examine their ability to produce CL. PCR amplification was conducted by using the Expand High Fidelity PCR system (Boehringer Mannheim Biochemicals), and other methods used for DNA manipulations have been described previously (42).

Media and bacterial growth. Luria-Bertani (LB) broth contained 1% tryptone (Difco, Detroit, Mich.), 0.5% yeast extract (Difco), and 1% NaCl. Difco sporulation medium (DSM), which contained 0.8% nutrient broth (Difco), 0.1% KCl, 0.025% MgSO4 · 7H2O, 1.0 mM Ca(NO3)2, 10 µM MnCl2, and 1.0 µM FeSO4 (43), was used for cultivation of B. subtilis cells. Synthetic media CI and CII were used for competence development (3). When required, the following supplements were added to the media (per liter); 50 mg of ampicillin (Sigma), 20 mg of neomycin (Wako Pure Chemicals), 50 mg of spectinomycin (Sigma), and 0.3 mg of erythromycin (Sigma). Growth of bacteria was monitored by measuring turbidity with a Klett-Summerson photoelectric colorimeter (no. 54 filter). For membrane depolarization carbonyl cyanamide 3-chlorophenylhydrazone from Sigma was added at a concentration of 10 µM (17). For solid media, 1.5% agar (Difco) was included.

Fluorescence microscopy. NAO (catalog no. A-1372; Molecular Probes) was added to a final concentration of 100 nM to a cell culture in DSM at 37°C that was harvested in the exponential growth phase and in the sporulation phase at 2 and 4 h after the end of log-phase growth (T2 and T4, respectively). After incubation at room temperature for 20 min, the cells were fixed on object slides coated with a layer of 1% (wt/vol) agarose gel in water. FM4-64 (catalog no. T-3166; Molecular Probes) staining was performed similarly by using a concentration of 0.1 to 0.2 µM in DSM. Fluorescence images were viewed by using an ECLIPS E600 fluorescence microscope (Nikon) and a cooled charge-coupled device camera (ORCA-ER; Hamamatsu Photonics Co., Hamamatsu, Japan). Green fluorescence (with excitation at 495 nm and emission at 525 nm) from NAO was detected by using a standard GFP(R)-BP filter unit (excitation at 460 to 500 nm and emission at 510 to 560 nm). Red fluorescence (emission at 640 nm) from NAO was detected by using a set of filters (excitation at 450 to 490 nm and emission at 610 nm). Fluorescence from FM4-64 (excitation at 510 nm and emission at 626 nm) was detected by using a G-2A filter unit (excitation at 510 to 560 nm and emission at 590 nm). To minimize the toxicity of high-energy emission light, the focus was set under phase-contrast conditions, and then fluorescence images were captured shortly after the shift to emission light conditions. The exposure times for green and red fluorescence of NAO were 0.2 to 0.8 and 7.7 s, respectively. The exposure time for FM4-64 fluorescence was 3.3 to 4.5 s. Captured images were processed by using Adobe Photoshop 6.0. The relative intensities of fluorescence were quantified by using NIH Image (Scion Image 4.0.2).

Lipid analysis. Membrane lipids were labeled with 0.5 µCi of [1-14C]acetic acid (57.2 mCi/mmol; Amersham) per ml for more than six generations during cultivation of the mutant and wild-type cells in DSM broth (5 ml) and were harvested at the late exponential phase and T2 and T4 of the sporulation phase. In some cases, including E. coli, cells were cultivated in LB broth with no radioisotope. Lipids were extracted by the method of Bligh and Dyer (2) or by the method developed for sporulating B. subtilis cells by Lacombe and Lubochinsky (26), with minor modifications. For the latter method we incorporated the following acidic treatment into the former method. The treatment included incubation of the cells in suspension with 0.9 M (final concentration) perchloric acid in 1% NaCl at 0°C for 30 min, followed by addition of 1.88 ml of chloroform-methanol (1:2 [vol/vol]) to 0.5 ml of the cell suspension. One-half of each lipid fraction was separated by two-dimensional thin-layer chromatography (TLC) on Silica Gel 60 (Merck, Darmstadt, Germany), first (x dimension) with chloroform-methanol-water (65:25:4 [vol/vol/vol]) and then (y dimension) with chloroform-methanol-acetic acid (65:25:10 [vol/vol/vol]). Spots of 14C-labeled lipid were visualized and quantified with a BAS 1000 bioimaging analyzer (Fuji Photo Film, Tokyo, Japan). In some cases phospholipids were visualized by uniformly spraying the gel with Dittmer-Lester reagent (11) and were quantified with a high-speed TLC scanner (model CS-920; Shimadzu, Kyoto, Japan). The molar percentage of each component was calculated.

Mass spectrometry analysis. The CL fractions that eluted from the CL spots on TLC plates were mixed with 2,5-dihydrobenzoic acid (Aldrich Chemical Co.) to saturation and subjected to mass spectrometry. Matrix-assisted laser desorption ionization—time of flight mass spectra were acquired with a KRATOS KOMPACT MALDI 4 (Shimadzu/Kratos, Tokyo, Japan) equipped with a pulsed nitrogen laser (337 nm) in the linear mode by using an acceleration voltage of 20 kV.


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RESULTS
 
NAO staining of wild-type B. subtilis cells. A previous study with E. coli cells indicated that NAO at a concentration of 100 to 200 nM in the growth medium resulted in staining of the cells having the wild-type phospholipid composition and had no noticeable effect on the growth rate (31). B. subtilis cells also exhibited no noticeable reduction in the growth rate with the same and much higher concentrations (100, 200, and 500 nM) of NAO in LB and DSM media. After staining of wild-type 168 cells in the late exponential growth phase with 100 nM NAO for 20 min at room temperature, the cells were mounted on agarose-coated object slides and viewed with an ECLIPS E600 fluorescence microscope (Nikon) equipped with a standard GFP(R)-BP filter unit (excitation at 460 to 500 nm and emission at 510 to 560 nm). Green fluorescent domains were clearly observed in the septal regions and at the poles (Fig. 1A). A sharp band in the center of the cell was observed in most cells. In some cases two fluorescent dots were located in the center of the cell. The two-dot structures were interpreted to be two-dimensional projections of three-dimensional rings with a small amount of fluorescent material forming an open ring in immunofluorescence studies of the FtsZ ring (10, 40). Sharp bands were interpreted as a ring that contained more fluorescent material in a closed-ring structure. Thus, we concluded that the NAO fluorescent domains were localized in the septal membrane. Membranes at the nascent poles in separating cells were also stained with NAO (Fig. 1A and B). The fluorescence at these poles was therefore considered to be fluorescence from the remnant of the material in the septal membrane. In sporulating cells NAO fluorescent domains were observed in the polar septa and on the engulfment and forespore membranes (Fig. 1B and C). The fluorescence was restricted to these membranes, and the mother cell membrane was not stained with NAO. In order to identify the fluorescence as specific to CL, control experiments with mutants lacking CL were required. However, the gene(s) responsible for CL synthesis in B. subtilis has not been determined.



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  		FIG. 1. Staining of wild-type B. subtilis cells with NAO. Wild-type B. subtilis 168 cells were cultivated in DSM. The cells were harvested during exponential and in the sporulation phase (at T2and T4) and stained with 100 nM NAO (Molecular Probes) for 20 min at room temperature to visualize CL. Fluorescence images of exponential-phase cells (A) and of sporulation-phase cells at T2 (B) and T4 (C) were taken by using the GFP(R)-BP filter unit (excitation at 460 to 500 nm and emission at 510 to 560 nm) as described in Materials and Methods. Corresponding phase-contrast images (D, E, and F) are also shown. The exposure times used for the fluorescence and phase-contrast images were 0.3 and 0.02 s, respectively. The single arrow indicates a sharp fluorescent band in the center of a cell. Two-fluorescent-dot structures in a cell center are indicated by a pair of arrowheads. Regions of NAO-stained nascent poles in cells that just separated are indicated by pairs of arrows.

NAO staining of mutant cells with a disrupted allele of clsA coding for CL synthase. B. subtilis has three candidates for the gene coding for CL synthase, ywiE, ywjE, and ywnE (25). To determine which of these paralogous genes is involved in CL synthesis, lipid was extracted by the Bligh-Dyer method (2) from mutant cells having a pMutin-disrupted allele of each candidate gene cultivated to the mid-exponential growth phase, and the phospholipid composition was examined. No noticeable reduction in CL content was observed in the extracts from the late-log-phase cells of the ywiE and ywjE disruptants, strains BFS1244 and BFS1245, respectively. However, there was no CL in the ywnE disruptant, strain BFS219, suggesting that ywnE is involved in CL synthesis but ywiE and ywjE are not. On the basis of this finding and the results described below ywnE was renamed clsA.

In clsA disruptant cells fluorescence was scarcely detectable (Fig. 2A), as expected because of the lack of CL. In sporulation-phase cells, however, fluorescent domains were observed in the medial and polar septa and engulfment membranes. This might suggest that the clsA-disrupted cells contained CL in the sporulation phase. Since all three gene products have highly conserved duplicated HxK(x)4D(x)6G(x)2N motifs (18, 28), we expected that all three might be active in CL synthesis. The remaining two, ywiE and ywjE, were anticipated to be responsible for minor CL synthesis in the clsA-disrupted cells in the sporulation phase.



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  		FIG. 2. Staining of clsA-disrupted mutant BFS219 cells with NAO. BFS219 cells were cultivated in DSM, harvested during exponential growth and in the sporulation phase at T4, and stained with 100 nM NAO for 20 min. Fluorescence images of exponential-phase cells (A) and sporulation-phase cells at T4 (B) were obtained as described in Materials and Methods. Corresponding phase-contrast images (C and D) are also shown. The exposure times used for the fluorescence and phase-contrast images were 0.3 and 0.02 s, respectively.

CL contents of mutant cells with disrupted clsA and three (clsA, ywiE, and ywjE) disrupted alleles. To prove that the product of ywiE and ywjE has CL synthase activity, these paralogous genes were examined for CL production in E. coli mutant SD9 (pssA1 cls-1) lacking detectable CL (35). Each open reading frame, from its initiation codon obtained by using the new NcoI site created by PCR on the initiation codon, was placed under control of the arabinose promoter of pSK6, a derivative of pSC101 (46), and was expressed in SD9 cells. The cells harboring clsA on plasmid pMMS1 contained CL (4.6% of the total phospholipid) after induction by addition of L-arabinose. The cells harboring ywjE on pMMS2 contained CL that accounted for 2% of the total lipid. The ywjE gene was, therefore, active in CL synthesis. In the cells harboring ywiE on pMMS3, however, we did not detect CL (less than 0.1% of the total phospholipid). When the cells harboring each of these paralogues were incubated at 42°C, expression of clsA and ywjE complemented the temperature-sensitive growth of SD9 cells, although ywjE did this very weakly; however, expression of ywiE did not. The ywiE gene may therefore be inactive in CL synthesis.

To clarify the possible contribution of ywjE and ywiE to CL synthesis, we constructed multiply disrupted mutant strains and examined their lipid compositions by 14C labeling. The drug resistance genes spc and neo were inserted into the unique HindIII site in ywjE and the XbaI site in ywiE, respectively, to construct ywjE1::spc and ywiE2::neo alleles. The disrupted alleles were successively introduced into BFS219 (clsA::pMutin), and multiply disrupted mutant strains were constructed. The triply disrupted mutant, designated SDB206 (clsA::pMutin ywjE1::spc ywiE2::neo), was examined in the studies described below.

Lipids were extracted by the perchloric acid method that was described previously for efficient extraction of CL from sporulating B. subtilis cells (26) from SDB206 and BFS219 cells cultivated in DSM containing [1-14C]acetic acid, and the CL contents of the mutants were examined after two-dimensional TLC on silica gel plates. The results of this lipid extraction analysis indicated that both of the mutant strains contained CL in the sporulation phase (Table 2). The clsA-disrupted mutant cells contained CL (0.3% of the total lipid at T4), although in the exponential growth phase the level was below the detection limit. Note that even in the triply disrupted mutant cells CL accounted for 0.2 and 0.3% of total lipid at T2 and T4, respectively.


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  		TABLE 2. Lipid compositions of strains 168, BFS219, and SDB206

To confirm the presence of CL in sporulating cells of these mutants, we obtained mass spectra of the fraction that eluted from the CL spot on a TLC plate. The region at mass/charge ratios ranging from 1,200 to 1,500 contained peaks of charged CL ions with gross acyl chain compositions ranging from 56 to 72 carbon atoms; this range of fatty acid distribution is the result of CL from B. subtilis (5). In the extracts from SDB206 cells we detected typical peaks of CL species with gross acyl chain compositions that varied from 61 to 69 carbon atoms with zero to three unsaturated bonds. The extract from BSF219 cells produced peaks at similar mass values. These results indicated that both the clsA-disrupted and triply disrupted mutant cells contained CL in the sporulation phase.

Fluorescence of NAO-stained cells of SDB206 with three disrupted alleles (clsA, ywiE, and ywjE). In the cells of the triply disrupted mutant, fluorescence was scarcely detectable during the vegetative growth phase (Fig. 3C-1); this was similar to the results obtained with the cells of the clsA-disrupted mutant (Fig. 2A and Fig. 3B-1). These results contrasted with the results obtained for red fluorescence after staining with FM4-64, a stain used to label plasma membranes of B. subtilis cells approximately uniformly (14, 38); the cell membranes of the mutants were uniformly bright lines and could not be distinguished from those of the wild type (Fig. 3D-1, E-1, and F-1). In the wild-type cells stained with the two dyes the membranes formed uniformly bright red fluorescent lines with FM4-64 (Fig. 3G-1-F) and showed an uneven distribution of green NAO fluorescence (Fig. 3G-1-N). Under such conditions the relative intensity of the FM4-64 fluorescence of the septal membranes was nearly double that of other regions of the membrane, as shown previously (38). The intensities of the NAO fluorescence of the region of the lateral membranes relative to those of the septal membranes were, however, much less (1/8 to 1/10). Similar relative intensity values were obtained for the cells stained with either one of the two dyes. These results indicated that the septal and polar membrane localization of NAO fluorescence was dependent on CL and that it was not caused by cell lysis.



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  		FIG. 3. Staining of triply disrupted mutant SDB206 cells with NAO and FM4-64. Wild-type strain 168 (A-1 to A-3 and D-1 to D-3), strain BFS219 (B-1 to B-3 and E-1 to E-3), and strain SDB206 (C-1 to C-3 and F-1) cells were cultivated in DSM, harvested during exponential growth(A-1, B-1, C-1, D-1, E-1, and F-1) and in the sporulation phase at T2 (A-2, B-2, C-2, D-2, and E-2) and at T4 (A-3, B-3, C-3, D-3, and E-3), and stained with NAO (A-1 to A-3, B-1 to B-3, and C-1 to C-3) or FM4-64 (D-1 to D-3, E-1 to E-3, and F-1) as described in the legend to Fig. 1. Corresponding phase-contrast images are below the stained images. The exposure times used for the NAO fluorescence and phase-contrast images were 0.2 to 0.8 and 0.02 s, respectively. The exposure times used for the FM4-64 fluorescence images were 3.3 to 4.5 s. The images of the wild-type cells (G-1-N and G-1-F) stained with both NAO and FM4-64 were obtained by using a GFP(R)-BP filter unit (G-1-N) and a G-2A filter unit (G-1-F) to detect the fluorescence of NAO and FM4-64, respectively. The exposure times used for the NAO and FM4-64 fluorescence images in panels G-1-N and G-1-F were 0.8 and 1.9 s, respectively.

In the sporulation-phase cells of the mutants, however, the fluorescent domains of NAO appeared in the medial septa and at the poles 2 h after the end of exponential growth (T2) (Fig. 3A-2, B-2, and C-2) and then on the engulfment and forespore membranes (Fig. 3A-3, B-3, and C-3). The intensity of the fluorescence in these mutants seemed to be weaker than that in the wild type (Fig. 3A-3), which probably reflected the low CL content. Note that the formation of the polar septa and engulfment membranes was delayed for ca. 1 h in these mutant cells.

NAO binding to CL results in dimerization of the two dye molecules in a stacking form in which the molecules are in close proximity, and thus the emitted peak of the fluorescence for the monomer (at 525 nm) shifts to red (at 640 nm) due to the metachromatic effect (36). Thus, red fluorescence emission indicates labeling of CL domains in the membrane (15, 36). When the GFP(R)-BP filter unit was replaced with a set of filters for red fluorescence (excitation at 450 to 490 nm and emission at 610 nm), red fluorescence was observed in the same regions (Fig. 4). The red fluorescence was weaker than the green fluorescence, which is consistent with the results of a spectral analysis of the NAO-CL interaction which showed that the red fluorescence is approximately 14-fold weaker than the green fluorescence (36). The brightness of the red fluorescence was thus intensified with Photoshop 6.0 for comparison with the localization of the green fluorescence. The red fluorescent domains in the septa and at the poles of the wild-type cells colocalized with the green fluorescence (Fig. 4A-2 and A-3). The images in polar septa and on engulfment membranes in sporulation phase (T4) were colocalized as well (Fig. 4B-2 and B-3). In the mutant cells the red fluorescence domains were observed in the sporulation phase, and they were colocalized with the green fluorescence domains (Fig. 4C-2 and C-3), indicating that the green fluorescence of the mutant cells was from NAO bound to CL in the membranes. These results confirmed the notion, based on TLC and mass spectrometry analyses, that both the clsA-disrupted and triply disrupted mutant cells contained CL in the sporulation phase.



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  		FIG. 4. Red fluorescence from NAO bound to CL of stationary-phase cells of mutant SDB206. Wild-type strain 168 cells during vegetative growth (A-1 to A-3) and in the sporulation phase at T4 (B-1 to B-3) and SDB206 cells in the sporulation phase at T4 (C-1 to C-3) were harvested and stained with NAO as described in the legend to Fig. 1. Green fluorescence (emission at 525 nm) (A-1, B-1, and C-1) and red fluorescence (emission at 640 nm) (A-2, B-2, and C-2) were detected by using a GFP(R)-BP filter unit (excitation at 460 to 500 nm and emission at 510 to 560 nm) and a filter unit with excitation at 450 to 490 nm and emission at 610 nm, respectively, as described in Materials and Methods. The exposure times used for green and red fluorescence were 0.3 and 7.7 s, respectively. (A-3, B-3, and C-3) Colocalization of green and red fluorescence images.

In the mutant cells the appearance of the polar septa and engulfment membranes was delayed during the sporulation process (Fig. 3). We therefore examined the frequency of heat-resistant cells during sporulation. In the case of the mutant strains the frequency of heat-resistant cells that could form colonies after 80°C treatment was lower in the later stage of sporulation. The frequencies of heat-resistant spores of SDB206 and BFS219 at 24 h after sporulation initiation were 30 and 50%, respectively, of the frequency of heat-resistant spores of the wild type. These results suggest that the CL-rich domains in the membranes are involved in a process required for sporulation.


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DISCUSSION
 
Examination with the fluorescent dye NAO showed that B. subtilis cells contain CL-specific fluorescence domains in the septa and at the poles in the exponential growth phase and in the polar septa and on the engulfment and forespore membranes during the sporulation phase. The specificity of NAO for CL in these experiments was supported by examination of the mutant strains with defects in putative CL synthase genes (clsA, ywiE, and ywjE). Fluorescence was not observed in the clsA-disrupted mutant cells that lacked detectable CL in the exponential growth phase. During sporulation the CL content increased in both wild-type and clsA-disrupted mutant cells (Table 2), which is analogous to the findings obtained for E. coli, in which the CL content increased during the stationary phase (7, 34, 44). The increase in the CL content during the sporulation phase is consistent with the results of ß-galactosidase transcriptional fusion experiments which indicated that both clsA and ywjE exhibited the maximal activity, although the activity of ywjE was quite low, in the early stationary phase and that ywiE had almost no activity (unpublished results). Indeed, the clsA and ywjE mRNA levels exhibited 2.1- and 11.7-fold {sigma}E-dependent increases, respectively (13). The increase in transcription is analogous to the increase in E. coli CL synthase activity, which increased in the stationary phase (20). The product of ywjE is thus anticipated to produce CL in sporulating cells of the clsA-disrupted mutant, although it was not effective under the conditions which we examined (Table 2). In spite of its highly conserved duplicated HxK(x)4D(x)6G(x)2N motifs for bacterial CL synthase (18, 28), the product of ywiE may not be involved in CL synthesis. This molecule may have a role in the heat shock response since the level of the transcript of ywiE increases after a heat shock (19).

Two-dimensional TLC analysis of the lipid extracted from 14C-labeled cells indicated that in the triply disrupted mutant CL accounted for 0.2 and 0.3% of total lipid at T2 and T4, respectively, in the sporulation phase (Table 2). Mass spectrometry analysis of the material eluted from the CL spot supported the identification. The red fluorescence in the stationary-phase cells of the triply disrupted mutant (Fig. 4) confirmed that CL was present in the polar septa and on the engulfment and forespore membranes, since NAO binding to CL results in dimerization of the two dye molecules in a stacking form in which the molecules are in close proximity, which shifts the emitted peak of green fluorescence for the monomer (at 525 nm) to red (at 640 nm) due to the metachromatic effect (36). What enzyme is responsible for production of CL in the triply disrupted mutant cells? This enzyme could be the product of pss, phosphatidylserine synthase, since a significant amount of CL was synthesized (0.15% of the total phospholipid as determined by 3 min of pulse-labeling with 32P) in E. coli null pssA cls double-mutant cells harboring a complementing B. subtilis pss gene on the pBR322 plasmid (41), similar to CL production in E. coli cls mutants by the product of E. coli pssA (34). The activity of the enzyme or the side reaction may be activated to produce CL during the early sporulation phase. The increase in the CL content during the sporulation phase may also be explained in part by the inactivity of a putative CL-specific phosphodiesterase (phospholipase D) that depends on ATP, as suggested by examination of stationary-phase cells of E. coli (6, 44).

CL-containing domains are visualized with NAO in the septal regions and at the poles of E. coli cells (31), indicating that the phospholipid distribution in the membrane is heterogeneous. This conclusion was supported by an examination showing that E. coli minicells, which consist mainly of polar materials of the envelope, are rich in CL (24). In our work, we obtained evidence of heterogeneous distribution of CL in the gram-positive bacterium B. subtilis. In eukaryotic cells loading and retention of NAO on membranes are thought to be dependent on the membrane potential of mitochondria, since the intensity and staining pattern of the cells are affected by depolarization treatment (22). However, isolated mitochondria that are devoid of membrane potential after fixation in glutaraldehyde are stained bright by NAO (22). Fixation of B. subtilis cells in 4% paraformaldehyde did not alter the pattern of NAO fluorescence (data not shown). Depolarization treatment of B. subtilis cells by addition of 10 µM carbonyl cyanamide 3-chlorophenylhydrazone did not affect the staining pattern, although the intensity of the fluorescence was slightly reduced (data not shown), as observed for eukaryotic cells (22), indicating that the pattern of NAO fluorescence reflects a spatially heterogeneous distribution of CL-rich domains in B. subtilis membranes. The CL-rich domains were also found in the sporulating cells in the polar septa and on the engulfment and forespore membranes and not only in the cells in the exponential growth phase. The biological significance of the CL-rich domains in sporulation was illustrated by using mutants with retarded emergence of the polar septal and engulfment membranes that was accompanied by a low frequency of heat-resistant cells.

What is the role of the CL-rich domains in these membranes? We can imagine two distinct roles that CL-rich domains play in the membranes. First, CL could contribute to form a nonbilayer structure, which is thought to be required for the formation of septa and for engulfment, since it facilitates a dynamic phase shift that causes formation of nonbilayer structures in the presence of certain divalent cations (mainly Ca2+, Mg2+, or Sr2+) under physiological conditions (12). The polar septal membranes that develop into engulfment membranes may require CL for dynamic progression, since their appearance was retarded in the sporulating cells of the mutants (Fig. 3). Development of the polar septa may depend on a level of CL that increases in the early sporulation phase. Second, CL could contribute to recruitment of peripheral membrane proteins to localize to specific regions of the cell membranes, and it could also contribute to maintenance of the optimal activity of these proteins. The former type of contribution has recently been described for MinD, which localizes in a horseshoe structure on the membrane at the poles of E. coli cells (39, 45). MinD, with its C-terminal amphiphilic {alpha}-helix bound to liposomes containing anionic phospholipids, has a higher affinity for CL-enriched liposomes (33, 47). The second type of contribution is also feasible, as essential interactions of CL with integral membrane proteins (e.g., cytochrome c oxidase) are well known (12) and an interaction with CL of a Rhodobacter sphaeroides type II reaction center has been demonstrated by X-ray crystallography (48). Hence, some of the sporulation-phase-specific proteins involved in the formation of the polar septal and the engulfment membranes (37) in B. subtilis could require CL-dependent localization and/or maintenance for optimal activity.


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ACKNOWLEDGMENTS
 
We thank Marie-Françoise Hullo and Patrick Stragier for mutant strains, Hiroaki Takahashi, Ayako Nishibori, and Satoko Fuchizawa for assistance, and Fujio Kawamura, Hideaki Nanamiya (St. Paul's University), and Kei Asai for guidance concerning fluorescence microscopy. We also thank Roy H. Doi for reviewing the manuscript. Thanks are also due to Isao Shibuya, Hiroshi Matsuzaki, Junichi Ohnishi, Shigeki Moriya (Nara Institute), and Junichi Sekiguchi (Shinshu University) for encouragement and helpful discussions.

This work was supported in part by grants-in-aid for scientific research from the Ministry of Education, Science and Culture of Japan.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biochemistry and Molecular Biology, Saitama University, 255 Shimo-ohkubo, Sakura, Saitama, Saitama 338-8570, Japan. Phone: 81-48-858-3406. Fax: 81-48-858-3698. E-mail: koumatsu{at}molbiol.saitama-u.ac.jp. Back


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REFERENCES
 
    1
  1. Alley, M. R. K., J. R. Maddock, and L. Shapiro. 1993. Requirement of the carboxyl terminus of a bacterial chemoreceptor for its targeted proteolysis. Science 259:1754-1757.[Abstract/Free Full Text]
    2. 2
  2. Ames, G. F. 1968. Lipids of Salmonella typhimurium and Escherichia coli: structure and metabolism. J. Bacteriol. 95:833-843.[Abstract/Free Full Text]
    3. 3
  3. Anagnostopoulos, C., and I. Crawford. 1961. Transformation studies on the linkage of markers in the tryptophan pathway in Bacillus subtilis. Proc. Natl. Acad. Sci. USA 47:378-390.[Free Full Text]
    4. 4
  4. Bolivar, F., R. L. Rodriguez, P. J. Greene, M. C. Betrach, H. L. Heyneker, H. W. Boyer, H. W. Crosa, and S. Falkow. 1977. Construction and characterization of new cloning vehicles. II. A multi-purpose cloning system. Gene 2:95-113.[Medline]
    5. 5
  5. Clejan, S., T. A. Krulwich, K. R. Mondrus, and D. Seto-Young. 1986. Membrane lipid composition of obligately and facultatively alkalophilic strains of Bacillus spp. J. Bacteriol. 168:334-340.[Abstract/Free Full Text]
    6. 6
  6. Cole, R., and P. Proulx. 1975. Phospholipase D activity of gram-negative bacteria. J. Bacteriol. 124:1148-1152.[Abstract/Free Full Text]
    7. 7
  7. Cronan, J. E., Jr., and C. O. Rock. 1987. Biosynthesis of membrane lipids, p. 474-497. In F. C. Neidhardt, J. L. Ingraham, K. B. Low, B. Magasanik, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology. American Society for Microbiology, Washington, D.C.
    8. 8
  8. DeChavigny, A., P. N. Heacock, and W. Dowhan. 1991. Sequence and inactivation of the pss gene of Escherichia coli. Phosphatidylethanolamine may not be essential for cell viability. J. Biol. Chem. 266:5323-5332.[Abstract/Free Full Text]
    9. 9
  9. de Mendoza, D., G. E. Schujman, and P. S. Aguilar. 2002. Biosynthesis and function of membrane lipids, p. 43-55. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and its closest relatives. ASM Press, Washington, D.C.
    10. 10
  10. den Blaauwen, T., N. Buddelmeijer, M. E. G. Aarsman, C. M. Hameete, and N. Nanninga. 1999. Timing of FtsZ assembly in Escherichia coli. J. Bacteriol. 181:5167-5175.[Abstract/Free Full Text]
    11. 11
  11. Dittmer, J. C., and R. L. Lester. 1964. A simple, specific spray for the detection of phospholipids on thin layer chromatograms. J. Lipid Res. 5:126-127.
    12. 12
  12. Dowhan, W. 1997. Molecular basis for membrane phospholipid diversity: why are there so many lipids? Annu. Rev. Biochem. 66:199-232.[CrossRef][Medline]
    13. 13
  13. Eichenberger, P., S. T. Jensen, E. M. Conlon, C. van Ooij, J. Silvaggi, J.-E. Gonzalez-Pastor, M. Fujita, S. Ben-Yehuda, P. Stragier, J. S. Liu, and R. Losick. 2003. The {sigma}E regulon and the identification of additional sporulation genes in Bacillus subtilis. J. Mol. Biol. 327:945-972.[CrossRef][Medline]
    14. 14
  14. Fishov, I., and C. L. Woldringh. 1999. Visualization of membrane domains in Escherichia coli. Mol. Microbiol. 32:1166-1172.[CrossRef][Medline]
    15. 15
  15. Gallet, P. F., A. Maftah, J. M. Petit, M. Denis-Gay, and R. Julien. 1995. Direct cardiolipin assay in yeast using the red fluorescence emission of 10-N-nonyl acridine orange. Eur. J. Biochem. 228:113-119.[Medline]
    16. 16
  16. Guerout-Fleury, A. M., K. Shazand, N. Frandsen, and P. Stragier. 1995. Antibiotic-resistance cassettes for Bacillus subtilis. Gene 167:335-336.[CrossRef][Medline]
    17. 17
  17. Guffanti, A. A., S. Clejan, L. H. Falk, D. B. Hicks, and T. A. Krulwich. 1987. Isolation and characterization of uncoupler-resistant mutants of Bacillus subtilis. J. Bacteriol. 169:4469-4478.[Abstract/Free Full Text]
    18. 18
  18. Guo, D., and B. E. Tropp. 2000. A second Escherichia coli protein with CL synthase activity. Biochim. Biophys. Acta 1483:263-274.[Medline]
    19. 19
  19. Helmann, J. D., M. F. W. Wu, P. A. Kobel, F.-J. Gamo, M. Wilson, M. M. Morshedi, M. Navre, and C. Paddon. 2001. Global transcriptional response of Bacillus subtilis to heat shock. J. Bacteriol. 183:7318-7328.[Abstract/Free Full Text]
    20. 20
  20. Hiraoka, S., H. Matsuzaki, and I. Shibuya. 1993. Active increase in cardiolipin synthesis in the stationary growth phase and its physiological significance in Escherichia coli. FEBS Lett. 336:221-224.[CrossRef][Medline]
    21. 21
  21. Itaya, M., K. Kondo, and T. Tanaka. 1989. A neomycin resistance gene cassette selectable in a single copy state in the Bacillus subtilis chromosome. Nucleic Acids Res. 17:4410.[Free Full Text]
    22. 22
  22. Jacobson, J., M. R. Duchen, and S. J. R. Heales. 2002. Intracellular distribution of the fluorescent dye nonyl acridine orange responds to the mitochondrial membrane potential: implications for assays of cardiolipin and mitochondrial mass. J. Neurochem. 82:224-233.[CrossRef][Medline]
    23. 23
  23. Kikuchi, S., I. Shibuya, and K. Matsumoto. 2000. Viability of an Escherichia coli pgsA null mutant lacking detectable phosphatidylglycerol and cardiolipin. J. Bacteriol. 182:371-376.[Abstract/Free Full Text]
    24. 24
  24. Koppelman, C.-M., T. Den Blaauwen, M. C. Duursma, R. M. A. Heeren, and N. Nanninga. 2001. Escherichia coli minicell membranes are enriched in cardiolipin. J. Bacteriol. 183:6144-6147.[Abstract/Free Full Text]
    25. 25
  25. Kunst, F., N. Ogasawara, I. Moszer, A. M. Albertini, et al. 1997. The complete genome sequence of the Gram-positive bacterium Bacillus subtilis. Nature (London) 390:249-256.[CrossRef][Medline]
    26. 26
  26. Lacombe, C., and B. Lubochinsky. 1988. Specific extraction of bacterial cardiolipin from sporulationg Bacillus subtilis. Biochim. Biophys. Acta 961:183-187.[Medline]
    27. 27
  27. Maddock, J. R., and L. Shapiro. 1993. Polar location of the chemoreceptor complex in the Escherichia coli. Science 259:1717-1723.[Abstract/Free Full Text]
    28. 28
  28. Matsumoto, K. 1997. Phosphatidylserine synthase from bacteria. Biochim. Biophys. Acta 1348:214-227.[Medline]
    29. 29
  29. Matsumoto, K. 2001. Dispensable nature of phosphatidylglycerol in Escherichia coli: dual roles of anionic phospholipids. Mol. Microbiol. 39:1427-1433.[CrossRef][Medline]
    30. 30
  30. Matsumoto, K., M. Okada, Y. Horikoshi, H. Matsuzaki, T. Kishi, M. Itaya, and I. Shibuya. 1998. Cloning, sequencing, and disruption of Bacillus subtilis psd gene coding for phosphatidylserine decarboxylase. J. Bacteriol. 180:100-106.[Abstract/Free Full Text]
    31. 31
  31. Mileykovskaya, E., and W. Dowhan. 2000. Visualization of phospholipid domains in Escherichia coli by using the cardiolipin-specific fluorescent dye 10-N-nonyl acridine orange. J. Bacteriol. 182:1172-1175.[Abstract/Free Full Text]
    32. 32
  32. Mileykovskaya, E., W. Dowhan, R. L. Birke, D. Zheng, L. Lutterodt, and T. H. Haines. 2001. Cardiolipin binds nonyl acridine orange by aggregating the dye at exposed hydrophobic domains on bilayer surfaces. FEBS Lett. 507:187-190.[CrossRef][Medline]
    33. 33
  33. Mileykovskaya, E., I. Fishov, X. Fu, B. D. Corbin, W. Margolin, and W. Dowhan. 2003. Effects of phospholipid composition on MinD-membrane interactions in vitro and in vivo. J. Biol. Chem. 278:22193-22198.[Abstract/Free Full Text]
    34. 34
  34. Nishijima, S., Y. Asami, N. Uetake, S. Yamagoe, A. Ohta, and I. Shibuya. 1988. Disruption of the Escherichia coli cls gene responsible for cardiolipin synthesis. J. Bacteriol. 170:775-780.[Abstract/Free Full Text]
    35. 35
  35. Ohta, A., T. Obara, Y. Asami, and I. Shibuya. 1985. Molecular cloning of the cls gene responsible for cardiolipin synthesis in Escherichia coli and phenotypic consequences of its amplification. J. Bacteriol. 163:506-514.[Abstract/Free Full Text]
    36. 36
  36. Petit, J. M., O. Huet, P. F. Gallet, A. Maftah, M. H. Ratinaud, and R. Julien. 1994. Direct analysis and significance of cardiolipin transverse distribution in mitochondrial inner membranes. Eur. J. Biochem. 220:871-879.[Medline]
    37. 37
  37. Piggot, P. J., and R. Losick. 2002. Sporulation genes and intercompartmental regulation, p. 483-517. In A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and its closest relatives. ASM Press, Washington, D.C.
    38. 38
  38. Pogliano, J., N. Osborne, M. D. Sharp, A. Abanes-De Mello, A. Perez, Y. L. Sun, and K. Pogliano. 1999. A vital stain for studying membrane dynamics in bacteria: a novel mechanism controlling septation during Bacillus subtilis sporulation. Mol. Microbiol. 31:1149-1159.[CrossRef][Medline]
    39. 39
  39. Raskin, D. M., and P. A. de Boer. 1999. Rapid pole-to-pole oscillation of a protein required for directing division to the middle of Escherichia coli. Proc. Natl. Acad. Sci. USA 96:4971-4976.[Abstract/Free Full Text]
    40. 40
  40. Rueda, S., M. Vicente, and J. Mingorance. 2003. Concentration and assembly of the division ring proteins FtsZ, FtsA, and ZipA during the Escherichia coli cell cycle. J. Bacteriol. 185:3344-3351.[Abstract/Free Full Text]
    41. 41
  41. Saha, S. K. 1996. Biosynthetic regulations and biological functions of phospholipids in Escherichia coli. Ph.D. thesis. Saitama University, Saitama, Japan.
    42. 42
  42. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
    43. 43
  43. Schaeffer, P., H. Ionesco, A. Ryter, and Balassa. 1965. La sporulation de Bacillus subtilis: etude genetique et physiologique, p. 553-563. In Mechanismes de regulation des activites cellulaires microorganismes. Centre National de la Recherche Scientifique, Paris, France.
    44. 44
  44. Shibuya, I. 1992. Metabolic regulations and biological functions of phospholipids in Escherichia coli. Prog. Lipid Res. 31:245-299.[CrossRef][Medline]
    45. 45
  45. Suefuji, K., R. Valluzzi, and D. RayChaudhuri. 2002. Dynamic assembly of MinD into filament bundles modulated by ATP, phospholipids, and MinE. Proc. Natl. Acad. Sci. USA 99:16776-16781.[Abstract/Free Full Text]
    46. 46
  46. Suzuki, M., H. Hara, and K. Matsumoto. 2002. Envelope disorder of Escherichia coli cells lacking phosphatidylglycerol. J. Bacteriol. 184:5418-5425.[Abstract/Free Full Text]
    47. 47
  47. Szeto, T. H., S. L. Rowland, L. I. Rothfield, and G. F. King. 2002. Membrane localization of MinD is mediated by a C-terminal motif that is conserved across eubacteria, archaea, and chloroplasts. Proc. Natl. Acad. Sci. USA 99:15693-15698.[Abstract/Free Full Text]
    48. 48
  48. Wakeham, M., R. Sessions, M. R. Jones, and P. K. Fyfe. 2001. Is there a conserved interaction between cardiolipin and the type II bacterial reaction center? Biophys. J. 80:1395-1405.[Medline]

Journal of Bacteriology, March 2004, p. 1475-1483, Vol. 186, No. 5
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.5.1475-1483.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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