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membranes displayed variable sensitivity to lipid fluidity, indicating co-existence of .... The fluorescence intensities are normalized to the cells mass (in units of OD420). ...... Loura, L.M.S., de Almeida, R.F.M., and Prieto, M. (2001). Detection and ...
Blackwell Science, LtdOxford, UKMMIMolecular Microbiology 1365-2958Blackwell Publishing Ltd, 200349410671079Original ArticleBacterial membrane domainsS. Vanounou, A. H. Parola and I. Fishov

Molecular Microbiology (2003) 49(4), 1067–1079

doi:10.1046/j.1365-2958.2003.03614.x

Phosphatidylethanolamine and phosphatidylglycerol are segregated into different domains in bacterial membrane. A study with pyrene-labelled phospholipids Sharon Vanounou,1 Abraham H. Parola2 and Itzhak Fishov1* 1 Department of Life Sciences, Ben-Gurion University of the Negev, P. O. B. 653, Beer-Sheva 84105, Israel. 2 Department of Chemistry, Ben-Gurion University of the Negev, P. O. B. 653, Beer-Sheva 84105, Israel. Summary To detect and characterize membrane domains that have been proposed to exist in bacteria, two kinds of pyrene-labelled phospholipids, 2-pyrene-decanoylphosphatidylethanolamine (PY-PE) and 2-pyrenedecanoyl-phosphatidylglycerol (PY-PG) were inserted into Escherichia coli or Bacillus subtilis membrane. The excimerization rate coefficient, calculated from the excimer-to-monomer ratio dependencies on the probe concentration, was two times higher for PY-PE than for PY-PG at 37∞∞C. This was ascribed to different local concentrations rather than to differences in mobility. The extent of mixing between the two fluorescent phospholipids, estimated by formation of their heteroexcimer, was found very low both in E. coli and B. subtilis, in contrast to model membranes. In addition, these two pyrene derivatives exhibited different temperature phase transitions and different detergent extractability, indicating that the surroundings of these phospholipids in bacterial membrane differ in organization and order. Inhibition of protein synthesis, leading to condensation of nucleoid and presumably to dissipation of membrane domains, indeed resulted in increased formation of heteroexcimers, broadening of phase transitions and equal detergent extractability of both probes. It is proposed that in bacterial membranes these phospholipids are segregated into distinct domains that differ in composition, proteo–lipid interaction and degree of order; the proteo-lipid domain being enriched by PE. Introduction Membrane domains broadly speaking are defined as latAccepted 6 May, 2003. *For correspondence. E-mail [email protected]; Tel. (+972) 8 6461368; Fax (+972) 8 6472890/2992.

© 2003 Blackwell Publishing Ltd

eral heterogeneities in the composition or state of a liposome or cellular membrane (Edidin, 1998). A substantial progress was achieved during the last decade both in the characterization of a wide variety of membrane domains (Welti and Glaser, 1994; Bergelson et al., 1995; Edidin, 1997; 2001) and in the development of novel techniques for their investigation (for a recent review see Chattopadhyay, 2001). Membrane domains may be related to certain cellular functions (Edidin, 2001). In Escherichia coli, these membrane domains are proposed to function in major cell cycle events, such as timing of DNA replication and as space marker for cell division (Norris, 1995; Norris and Fishov, 2001), as well as in chromosome segregation (Woldringh, 2002). Indications for membrane heterogeneity in bacteria may be found in the literature (de Leij and Witholt, 1977; Horiuchi et al., 1983; Marty-Mazars et al., 1983; Welby et al., 1996). Different proteins in bacterial membranes displayed variable sensitivity to lipid fluidity, indicating co-existence of physically separated lipid domains of diverse fluidity and composition (Linden et al., 1973; Morrisett et al., 1975). The model suggests that specific membrane domains are formed as a result of the process of coupled transcription, translation and insertion (transertion) of membrane and secreted proteins (Norris, 1995; Binenbaum et al., 1999). Indeed, interfering the transertion process either at the level of transcription, by using rifampicin, or at the level of translation, by using chloramphenicol (Cam) or puromicine, causes both changes in membrane dynamics and nucleoid compaction (Binenbaum et al., 1999). Recently, heterogeneity in order and polarity of bacterial membrane was demonstrated utilizing the solvatochromic properties of the fluorescent membrane probe laurdan (Vanounou et al., 2002). This heterogeneity was ascribed to existence of proteo-lipid and predominantly lipid domains. Membrane patterns with apparent functional implication were observed in E. coli cells and filaments stained with a fluorescent probe FM 4– 64 (Fishov and Woldringh, 1999). These patterns may be related to the large cardiolipin (CL) domains in the septal region and poles of E. coli visualised by use of the fluorescent dye, 10-N-nonylacridine orange, which binds specifically to CL (Mileykovskaya and Dowhan, 2000). These visual data were supported by the finding of CL enrichment in minicells (Koppelman et al., 2001). Beyond visualization, fluorescent PL are widely used to obtain physical

1068 S. Vanounou, A. H. Parola and I. Fishov characteristics of lipid mobility and distribution in model and cellular membrane (e.g. Huijbregts et al., 1996; Loura et al., 2001; Kusba et al., 2002). Lateral heterogeneity can be measured by utilizing the ability of pyrene to form excited dimer (excimer), which is a diffusion-controlled process in fluid membranes (Galla and Sackmann, 1974; Kinnunen et al., 1993; Barenholz et al., 1996; Lehtonen and Kinnunen, 1997). As the two major membrane phospholipids (PL) in E. coli are phosphatidylethanolamine and phosphatidylglycerol (Morein et al., 1996), in the present study we used these pyrene-acyl-chain-labelled phospholipids (Py-PL) in order to characterize their lateral diffusion and distribution when inserted into E. coli membrane. The resistance of phospholipids to gentle extraction by the detergent Triton X-100 is known as a method to reveal the existence of membrane domains (London and Brown, 2000). This was applied in our study in order to assess the heterogeneous distribution of these Py-PL. We compared normal cells with those in which the transertion process was interrupted by Cam and in which domains are presumably dissipated (Binenbaum et al., 1999). A comparison was also performed between results obtained with E. coli and with Bacillus subtilis, Gram-positive bacteria possessing only one membrane. Results Evaluation of labelling conditions Fluorescent derivatives of phospholipids can be easily incorporated into artificial membranes like liposomes by mixing with unlabelled phospholipids (Somerharju et al., 1985). In eukaryotic cells, the cytoplasmic membrane was labelled by spontaneous insertion of the fluorescent lipids from the aqueous phase (Pagano et al., 1991) with the subsequent redistribution into intracellular organelles. This method requires a direct contact between vesicles or micelles containing fluorescent lipid and cellular membrane. In the case of cytoplasmic membrane of Gramnegative bacteria a successful incorporation of fluorescent phospholipids was achieved by permeabilization of the outer membrane or with inverted inner membrane vesicles (Huijbregts et al., 1996). Membranes of a fatty acid auxotroph strain of Micrococcus luteus were successfully labelled metabolically by feeding bacteria with a fatty acid antracene derivative (de Bony et al., 1989). In this case, all kinds of membrane phospholipids are labelled randomly and only the analysis of extracted crosslinked products allowed revealing a compositional heterogeneity in the membrane (de Bony et al., 1989). Our goal was to reach a controlled insertion of a particular fluorescent phospholipid into bacterial membranes that would allow a more detailed characterization of its specific surrounding in situ.

The rationale of the method applied here was to facilitate spontaneous insertion of fluorescent phospholipids into the membrane by increasing the concentration of the reaction encounters while slightly perturbing the membrane structure. This perturbation should be gentle and reversible, e.g. fluidization of the membrane caused by an organic solvent (Dombek and Ingram, 1984). Thus, the labelling was performed in a dense bacterial suspension (about 5 ¥ 109 cells ml-1) and at a relatively high probe concentration (about 10-5 M) in a small volume (40 ml). Because the probe was added as a solution in an organic solvent, the latter appeared at a high concentration and served as a chaotropic agent for the membrane. The incubation temperature should be above the phase transition. Excess of the probe and the organic solvent were thoroughly washed out after the labelling. The conditions of labelling were determined by applying the following criteria: (i) effective insertion of the probe; (ii) minimal perturbation of the membrane; and (iii) viability of bacteria after the labelling procedure. At a preliminary step during the design of the method, we have used diphenylhexatriene phosphatidylcholine (DPH-PC) because of its unique advantages. Those are: (i) the strong fluorescence in lipid environment, proportional to the probe concentration (in contrast to pyrene monomers that are also involved in excimerization) and (ii) the sensitive fluorescence anisotropy responses to phospholipid orientation order (Parente and Lentz, 1985). The insertion of DPH-PC into bacterial membranes was indeed facilitated by methanol as seen from the fluorescence intensity dependence (Fig. 1), showing a stepwise increase at 20% methanol. The fluorescence anisotropy of DPH-PC was independent of methanol concentration

Fig. 1. Insertion of the fluorescent probe DPH-PC into E. coli membrane as a function of methanol concentration used in the labelling process (see Experimental procedures for details). The fluorescence intensities are normalized to the cells mass (in units of OD420). © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Bacterial membrane domains 1069 (not shown) and its value, 0.186 ± 0.004, was characteristic for an ordered membrane surrounding (Parente and Lentz, 1985). The anisotropy value for DPH (0.165 ± 0.003) measured in bacteria subjected to the same treatment was also unchanged up to 20% methanol and slightly increased above this concentration. At 20% methanol, DPH fluorescence anisotropy remained unchanged up to 120 min incubation with the solvent. Moreover, with increased probe concentration, excitation of DPH-PC labelled bacterial suspension at 280 nm exhibited a strong quenching of tryptophan fluorescence accompanied by appearance of DPH fluorescence spectrum with a characteristic iso-emissive point (data not shown). It yielded a maximal quenching efficiency of about 40%. This may be ascribed to excitation energy transfer from tryptophans of membrane proteins to closely located fluorescent phospholipids. Another test for the damage caused by the exposure of bacterial cells to methanol was the ability of these cells to grow after such treatment. When bacteria were returned to the growth medium after treatment in 20% methanol, they began growing immediately, although with a lower rate, and recovered to a normal growth rate after two-three generation times (data not shown). The colony counts test indicated that over 90% of methanol-treated cells were viable, i.e. able to form colonies on LB-agar plates. The observed slow growth recovery was caused, perhaps, by a leakage of the low molecular weight constituents during the methanol treatment. Another solvent, ethanol, was found more chaotropic and much more damaging at the same labelling efficiency (data not shown). Thus, 20% methanol was chosen for further labelling procedure as an optimal concentration giving an acceptable probe insertion with a minimal and reversible damage. The efficiency of labeling, determined as the ratio of inserted to added probe concentration (see Experimental procedures), was tested as a function of incubation time in the labelling mixture (not shown) and reached about 10% in an hour for both 1-hexadecanoyl-2-(1-pyrenedecanoyl)sn-glycero-3-phosphoethanolamine (Py-PE) and 1-hexadecanoyl-2-(1-pyrenedecanoyl)-sn-glycero-3-phosphoglycerol (Py-PG). Further, this 10% labelling efficiency was independent of the added Py-PL concentration, i.e. a linear correlation between added Py-PL and that found in the membrane was obtained (data not shown). This allowed us to control the concentration of Py-PL in the membrane for further determination of excimerization rate constants. To find out the distribution of the inserted PL between the two membranes of E. coli, we performed ‘separation’ of inner and outer membranes by the previously used method of hyperosmotic plasmolysis, which was convincing for the localization of DPH and FM 4–64 to the inner membrane (Fishov and Woldringh, 1999). The rhodamine© 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Fig. 2. Fluorescence (left) and phase-contrast (right) images of E. coli, stained with rhodamine-PE as described in Experimental procedures and plasmolyzed by 15% sucrose. Plasmolysis bays on poles can be seen as brighter areas in phase contrast yet they are almost non-fluorescent. Compartments surrounded by inner membrane are dark in phase-contrast and bright in fluorescent images. Images were taken with a cooled ccd camera (Princeton Instruments Inc., model RTE-1317-k-1, Trenton, NJ, USA), mounted on an Olympus BH-2 fluorescence microscope equipped with an oil immersion (¥100) lens and 5x photo-ocular.

labelled PE was used instead of the pyrene derivatives as it is more appropriate for microscopic imaging (pyrene undergoes a fast photobleaching and has a far blue fluorescence). Comparison of fluorescence and phasecontrast images (Fig. 2) reveal that the inner membrane retracted from plasmolysed poles is brightly stained, whereas the outer membrane, retained by the rigid peptidoglycan, is not. It may thus be concluded that the probe is preferentially incorporated into the inner membrane. (The deep plasmolysis bays shown in Fig. 1 in Fishov and Woldringh (1999) were hampered in this case presumably because of the above-mentioned leakage of the inner membrane after methanol treatment.) Concentration dependence of excimerization rate of Py-PL in bacterial membrane Escherichia coli cells labelled with Py-PE or Py-PG by the method described above display a fluorescence spectrum (Fig. 3) characteristic for pyrene derivatives in a hydrophobic environment similar to that reported for liposomes (Somerharju et al., 1985). Since the final probe concentration (calculated per total sample volume) ranged from 0.01 to 0.5 mM, the appearance of excimer fluorescence in the spectrum (lem = 470, Fig. 3) confirms that the probe is highly concentrated in the membrane. The probe mole fraction of total membrane lipids can be estimated as

1070 S. Vanounou, A. H. Parola and I. Fishov between Py-PE and Py-PG (see below Fig. 7 and Table 3), which is in a good agreement with other results (Somerharju et al., 1985). The measured apparent rate constant (eqns 1 and 2) is determined mainly by the excimer lifetime and the second order rate constant for excimer formation. The latter is supposed to be diffusion controlled (Galla et al., 1979; Galla and Hartmann, 1980; Naqvi et al., 2000). On the other hand, the distribution of labelled phospholipids in the membrane may not be homogeneous and thus, the local microscopic concentration may differ essentially from an average value used for calculation. The Py-PL excimer lifetime measured in our conditions (Table 2) was found independent on the polar head-group. The diffusion coefficient can, in principle, be

Fig. 3. Fluorescence spectrum of Py-PE in E. coli membrane. The probe concentration calculated per total sample volume was 2 ¥ 10-7M, determined as described in Experimental procedures.

1–5% from the known cell mass in a sample and chemical composition of an average E. coli cell (Neidhardt and Umbarger, 1966). This estimation is validated by the similar values of excimer-to-monomer ratio at the same mole fractions of pyrene-phospholipid in liposomes (Jones and Lentz, 1986). The ratio of excimer to monomer increased linearly with concentration of Py-PE and Py-PG as well as concentration of pyrene (Fig. 4), as expected for a diffusioncontrolled reaction (Galla and Sackmann, 1974, Galla and Hartmann, 1980) according to equation: IE k (1) = FE t E k ac I M k FM where IE and IM are fluorescent intensities of the excimer and the monomer, respectively, kFE and kFM – rate constants for excimer and monomer fluorescence, ka – rate constant of excimer formation in the bimolecular reaction, tE – excimer radiative life-time and c – total concentration of pyrene molecules. k (2) k app = FE t E k a k FM The apparent excimerization rate constant, Kapp (eqn 2) was calculated from the slope of the linear fit of excimerto-monomer ratio dependencies on the label concentration (Fig. 4A), results are summarized in Table 1. The high excimerization rate observed for pyrene as compared with Py-PL reflects higher diffusion rate of free pyrene in contrast to the phospholipid-bound. The excimerization rate for Py-PE was twice that for Py-PG. A similar trend was obtained for Gram-positive B. subtilis (Fig. 4B). In phosphatidylcholine liposomes, however, there was no difference in the excimerization rate

Fig. 4. Excimer to monomer ratio as a function of pyrene, Py-PE and Py-PG concentrations in E. coli (A) and B. subtilis (B) membranes. Excimerization rate constants were calculated from the slopes of the linear fits (lines). Concentrations of the probes in the membrane are expressed as total concentration (determined as described in Experimental procedures) normalized to cell biomass. © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Bacterial membrane domains 1071 Table 1. Excimerization rate constants for Pyrene, Py-PE and Py-PG in E. coli grown with glucose or treated with CAM. The rate constant values were determined from linear fit slopes of excimer-to-monomer ratio dependencies on the dye concentration in the membrane (Fig. 4A). Pyrene

Py-PE

Py-PG

Excitation

340 nm

280 nm

340 nm

280 nm

340 nm

280 nm

Glucose CAM

0.48 ± 0.02 0.84 ± 0.04

0.21 ± 0.02 0.49 ± 0.06

0.15 ± 0.01 0.16 ± 0.010

0.12 ± 0.01 0.14 ± 0.04

0.065 ± 0.014 0.041 ± 0.003

0.065 ± 0.015 0.041 ± 0.018

estimated indirectly from the rotational correlation time of pyrene since both are controlled by membrane microviscosity (Parola, 1993; Ben-Shooshan et al., 2002). In order to get information from particular membrane regions of excimerization reaction, the rotational correlation time of excimers formed by each Py-PL was measured (Table 2). These values do not reveal any significant difference in the viscosity of the immediate surroundings of Py-PE and Py-PG excimers. Thus, it may be concluded that (i) the higher excimerization rate obtained for Py-PE is a consequence of its local concentration which is higher than that of Py-PG and (ii) that it cannot be attributed to a lower diffusion rate of Py-PE. This may indicate a specific distribution pattern for each phospholipid in the heterogeneous membrane, arising from the existence of domains.

The last two reactions of heteroexcimer formation should be remarkably restricted if these phospholipids are segregated in the membrane. Varying Py-PE and Py-PG mole fractions while keeping the total concentration of Py-PL constant was used to modulate formation of the heteroexcimer. The measured excimer and monomer fluorescence intensities are actually sums of fluorescence emitted from all kinds of excimers and all kinds of monomers. The apparent rate constant (eqn 2) may therefore be expressed as a function of two homoexcimerization rate constants, kPE and kPG, rate constant for the heteroexcimer formation, kmix (assuming that it is the same for the last two reactions), and the mole fraction of one of the labelled phospholipids, e.g. fPE:

Mixed excimer

The experimental data of Kapp at varying fPE (Fig. 5A) were satisfactorily fitted by eqn 3 and the resulting values of the rate constants are presented in Table 3. The calculated rate constants for the individual phospholipids were close to those obtained from concentration dependencies for each single Py-PL (compare Table 3 and Table 1), but the apparent rate constant for heteroexcimerization appeared to be essentially lower than both. Also shown in Fig. 5A, are predictions for the apparent excimerization rate constant calculated from eqn 3 for two extreme cases: (i) full mixing of the Py-PL, assuming that heteroeximer is formed at a rate equal to the average rate of homoexcimerization (kmix = 1/2(kPE + kPG), upper line); and (b) complete absence of heteroexcimerization (kmix = 0, lower line). It can be seen that the experimental data are very close to the curve predicted for complete segregation of phospholipids. The same type of experiment was performed with Gram-positive B. subtilis (Fig. 6) and in liposomes (Fig. 7). In B. subtilis, like in E. coli, the rate of heteroexcimer formation was four times lower than the excimerization rate for Py-PG, the latter was again slower than that for Py-PE (Fig. 6 and Table 3). In liposomes, in contrast to cells, the heteroexcimerization rate was very close to that of both kinds of homoexcimers (Fig. 7 and Table 3), reflecting a well mixed unrestricted membrane. These results testify that in bacterial cells the two kinds of phospholipids are well segregated so that the probability to form a heteroexcimer is very low.

Kapp = kPE.fPE2 + kPG.(1 - fPE)2 + kmix.fPE.(1 - fPE)

As pointed out in the Introduction, in natural membranes the lateral sorting of phospholipids may be driven by specific interactions with integral and peripheral proteins (Marsh, 1995; Lehtonen and Kinnunen, 1997). The noticed higher local concentration of Py-PE in E. coli membrane suggests an absence of ideal mixing of the two major phospholipids, PE and PG. To examine the degree of mixing, we have tested a possibility of a heteroexcimer formation by pyrene moieties bound to different kinds of PL. When both kinds of Py-PL are present in the membranes, the excimer formation can be a result of four possible reactions between two monomers in excited state, Py*-PE and Py*-PG, and two corresponding unexcited molecules: Py*-PE + Py-PE < = > (Py-PE.Py-PE)* Py*-PG + Py-PG < = > (Py-PG.Py-PG)* Py*-PE + Py-PG < = > (Py-PE.Py-PG)* Py*-PG + Py-PE < = > (Py-PG.Py-PE)*

Table 2. Lifetime t and rotational correlation time r of Py-PE and PyPG in E. coli membrane.

Py-PE Py-PG

t, ns

c2

r, ns

c2

56 ± 1.4 55 ± 0.6

1.8 1.2

1.1 ± 0.15 0.9 ± 0.10

8 9

© 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

(3)

1072 S. Vanounou, A. H. Parola and I. Fishov

Fig. 6. Excimerization rate constants for normal (circles) and for Cam-treated (triangles) B. subtilis cells, labelled with varying mole fractions of Py-PE and Py-PG. Experimental conditions as in Fig. 4. Equation 3 was used to fit the experimental data (see Table 3 for the constant values).

Fig. 5. Apparent excimerization rate constants for normal (A) and Cam-treated (B) E. coli cells, labelled with both Py-PE and Py-PG. The mole fraction of each Py-PL was varied as shown, while keeping the total probe concentration constant (3 ¥ 10-6M during labelling). The points represent an average value of four independent experiments. To get the apparent excimerization rate constants for homoand heteroexcimers (shown in Table 3), the experimental data were fitted using eqn 3 (continuous line). The dashed lines are generated from the same equation, assuming that the excimerization rate constant for the heteroexcimer is zero (lower line) or equal to the average of the rate constant values for Py-PE and Py-PG (upper line).

The effect of inhibition of protein synthesis on excimerization rates Treatment of bacteria with chloramphenicol was suggested to dissipate putative membrane domains formed by coupled transcription, translation and insertion of membrane proteins (Binenbaum et al., 1999). This domain dissipation should reduce the degree of order in the membrane, previously observed as a decrease in the average membrane viscosity (Binenbaum et al., 1999). Pyrene excimerization rate was two times higher in Camtreated cells (Table 1), supporting this result. However, the

Fig. 7. Excimerization rate constants for multilamellar liposomes, labelled with varying mole fractions of Py-PE and Py-PG. Experimental conditions and fits as in Fig. 4.

apparent rate of Py-PE excimerization did not change and that of Py-PG was even lower in E. coli treated with Cam in comparison with normal cells (Table 1). Moreover, the heteroexcimer formation was also close to zero (Fig. 5B, Table 3). A different result was obtained with B. subtilis where excimerization rate of Py-PG increased remarkably with almost no change in Py-PE excimerization; the rate constant for heteroexcimerization was elevated to the average value for the two Py-PL (Table 3). Energy transfer from intrinsic tryptophan to the pyrene probes Membrane-embedded pyrene probes can serve as good © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Bacterial membrane domains 1073 Table 3. Excimerization rate constants for homo- and heteroexcimers of Py-PE and Py-PG in bacterial membranes and in liposomes. E. coli

B. subtilis

Membrane Treatment



+Cam



+Cam

Liposomes –

kPE kPG kmix

0.172 ± 0.012 0.098 ± 0.013 0.033 ± 0.018

0.175 ± 0.007 0.062 ± 0.007 -0.016 ± 0.01

0.496 ± 0.023 0.164 ± 0.023 0.039 ± 0.034

0.57 ± 0.086 0.40 ± 0.086 0.48 ± 0.130

4.71 ± 0.12 4.13 ± 0.12 3.44 ± 0.18

acceptors for energy transfer from membrane proteins due to the well-overlapping spectra of tryptophan fluorescence and pyrene excitation, similar to DPH as described above. Quenching of tryptophan fluorescence was less effective by free pyrene (about 30%) than that by DPHPC presumably due to lower extinction coefficient of pyrene and a bulkier molecule. Quenching by Py-PL was much weaker (about 10%). That may be ascribed to a stronger sterical hindrance. With this level of quenching, the accuracy of measurement does not allow the detection of any differences in the accessibility of proteins to various phospholipids. An alternative approach can be an estimation of phospholipid mobility in the lipid boundary of membrane proteins (Engelke et al., 1994). This was done by measuring pyrene excimerization rate when excited through tryptophans, i.e. only the probe molecules located in the range of energy transfer from protein tryptophans (Engelke et al., 1994). The excimerization rate constants for pyrene and Py-PL upon excitation at 285 nm, presented in Table 1, show over 50% decreased mobility of pyrene and slightly decreased excimerization of Py-PE but not of Py-PG in glucose grown cells in comparison with those of directly excited probes. In Cam-treated cells, the difference in excimerization of Py-PE excited either at 340 or at 285 nm was less pronounced. It can be concluded that, in general, the membrane is essentially more condensed in the vicinity of proteins as exhibited by pyrene, which is supposed to be randomly distributed. Moreover, the mobility of boundary Py-PE is slightly restricted in contrast to that of Py-PG.

the final transition to the gel state of the entire membrane, consistent with the transition temperature detected by other probes (Vanounou et al., 2002). A higher temperature for the first phase transition detected with Py-PE as compared with that detected with Py-PG is another indication for a different localization at distinct membrane domains of each pyrene-phospholipid in contrast to the randomly distributed pyrene. The apparent activation energy for excimerization of pyrene in the liquid membrane (above 33∞C) was similar to that observed in liquidcrystalline liposomes (Daems et al., 1985). It decreased at lower temperatures presumably due to exclusion of pyrene into the liquid phase co-existing with the expanding gel phase (Galla and Sackmann, 1974) and again increased in the pure gel phase below 18∞C. The strikingly low and almost identical activation energies for PyPE excimerization in both liquid-crystalline and gel phases suggests that the reaction is not diffusion-controlled, meaning that a fraction of Py-PE excimers are static. The apparent activation energy for Py-PG excimerization was also low compared with pure lipid vesicles (Somerharju et al., 1985), but it increased below the

Multiple phase transitions in bacterial membrane detected by Py-PL excimerization The ratio of excimer to monomer was measured as a function of temperature in E. coli cells labelled with pyrene (Fig. 8), Py-PE and Py-PG (Fig. 9A). The break points in the Arrhenius plots, indicating phase transition temperatures, and the activation energies for excimer formation are given in Table 4. The three phase transition temperatures observed for free pyrene are differently sensed by pyrene-phospholipids: each of the phospholipids displays only one of the two high-temperature transitions (at about 25∞C for Py-PG and 34∞C for Py-PE), while the lowest break point at 18.5∞C was common for all three probes. The lowest temperature most probably reflects © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Fig. 8. Temperature dependence of the apparent excimerization rate constant of pyrene in E. coli membrane. Normal (closed circles) or Cam-treated (open circles) E. coli cells were labelled with 5 ¥ 10-7M pyrene and the excimer/monomer ratio was measured at different temperatures as described in Experimental procedures. The lines drawn through the experimental points are the best linear fit. The vertical bars mark the breaks in the line slope and the corresponding temperatures are shown in Table 4.

1074 S. Vanounou, A. H. Parola and I. Fishov

Fig. 10. Per cent of unextractable Py-PE (circles) and Py-PG (triangles) in normal (closed symbols) or Cam-treated (open symbols) E. coli cells. The cells were labelled with fluorescent phospholipids and subjected to extraction by different concentrations of Triton as described in Experimental procedures.

tion energy above 33.6∞C (7.4 kJ mole-1, Fig. 8).

was

essentially

lower

Extractability of Py-PL from labelled cells

Fig. 9. Temperature dependencies of the apparent excimerization rate constants of Py-PE (circles) and Py-PG (triangles) in the membrane of normal (A) and Cam-treated E. coli cells (B). Cells were labelled with Py-PL to a final probe concentration in the sample of 10-7M and the excimer/monomer ratio was measured at different temperatures as described in Experimental procedures. The lines drawn through the experimental points are the best linear fit. The vertical bars mark the breaks in the line slope and the corresponding temperatures are shown in Table 4.

phase transition (Table 4). In Cam-treated cells, phase transitions were not detectable by Py-PL excimerization (Fig. 9B). The excimerazation was characterized by low activation energies, 9.0 and 13.0 kJ mole-1 for Py-PE and Py-PG respectively. Phase transitions detected by free pyrene were the same as in normal cells, but the activa-

The compositional heterogeneity of the membrane may also be manifested by a different local sensitivity to solubilization, e.g. by existence of so-called detergent-resistant domains (London and Brown, 2000). Assuming that the partition of the fluorescent phospholipids is similar to their natural analogues, the extractability of Py-PL inserted into bacterial membranes could be different if they are located in different domains. The resistance of Py-PE and Py-PG inserted into normal and Cam-treated cell membranes to cold extraction by Triton X-100 is presented in Fig. 10. In the range of detergent concentrations from 0.05 to 0.1%, the fraction of unextracted Py-PE was twice higher than that of Py-PG in normal cells. This difference was, however, eliminated by Cam treatment. At higher Triton concentrations or at room temperature (not shown), the extraction of both phospholipids was almost complete. Notably, examination of membrane viscosity of bacteria subjected to cold 0.1% Triton extraction showed

Table 4. Temperatures of phase transitions and activation energies for excimer formation revealed by pyrene, Py-PE and Py-PG in E. coli membrane.

Pyrene Py-PE Py-PG

T1, ∞C

T2, ∞C

T3, ∞C

E1, kJ/mol

E2, kJ/mol

E3, kJ/mol

E4, kJ/mol

33.6 33.7

25.0

18.5 18.5 17.4

29.8 10.3 17.0

11.0 16.5

18

23.7 9.3 22.3

25–21.5

6.1

© 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Bacterial membrane domains 1075 about 30% increase in DPH fluorescence anisotropy (data not shown), indicating that the more fluid membrane fraction was extracted. Thus, it may be concluded that Py-PE is preferentially partitioned into a more viscose (ordered) membrane region, presumably proteo-lipid domain.

Discussion Method of labelling Our aim was to insert fluorescent phospholipid into bacterial membranes under controlled conditions, thus, metabolic labeling used by de Bony et al. (1989) couldn’t accomplish our demand. On the other hand, spontaneous insertion of phospholipids in the inner membrane couldn’t occur because of shielding by the outer membrane. That is why either cells with the outer membrane permeabilized by EDTA treatment or inner membrane vesicles were employed for insertion of fluorescent phospholipids (Huijbregts et al., 1996). The rationale of our labelling method was to facilitate the rate of phospholipid insertion into the membrane by increasing concentrations of the ‘reactants’ along with a gentle and reversible perturbation of the membrane. The chosen procedure included incubation of a concentrated cell suspension (2 ¥ 108 cells/ 40 ml) with varied concentrations of Py-PL (as 10-6 ∏ 10-5 M solution in 10 ml of methanol) during one hour at 37∞C followed by two to three times washing in PBS. These conditions of labelling affected neither the membrane order (as measured by DPH fluorescence anisotropy) nor the ability of bacteria to grow. Several indications confirm that the label is inserted into the membrane: (i) extensive washing of excess free label didn’t change the fluorescence intensity of the cell suspension. (ii) The relatively high value of fluorescence anisotropy for DPH-PC in these cells (0.186 ± 0.004) is close to that reported for liposomes (Parente and Lentz, 1985) and is characteristic for an ordered membrane. (iii) The total concentration of Py-PL in the suspension of the labelled cells is about 0.1 mM, one that is too low for a noticeable formation of excimers in a homogeneous solution. Appearance of the excimer peak in the fluorescence spectra (Fig. 3) is thus indicative of a much higher local concentration of the label partitioned into the membrane. (iv) Quenching of tryptophan fluorescence by DPH-PC was about 40%, implying the close vicinity between the membrane proteins and the labelled phospholipids (Engelke et al., 1994). (v) Excimerization rates of Py-PL excited through tryptophans were similar to those measured by direct excitation, again indicating their close proximity which is a prerequisite for the observed efficient energy transfer. (vi) Temperature phase transitions displayed by Py-PL (Fig. 9A) are similar to those revealed by other lipophillic labels like DPH and laurdan (Vanounou © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

et al., 2002). These results strongly support our claim that the fluorescent phospholipids were successfully inserted into bacterial membranes. The mole fraction of Py-PL in the membrane was evaluated as 1–5% and should not affect membrane organization. This value seems to be reliable, because the ratio IEx/IM in bacterial membranes (from 0.1 to 0.4, Fig. 4) is close to that reported for liposomes labelled at the same probe mole fraction (Jones and Lentz, 1986). In addition, the localization of fluorescent phospholipids in the plasma membrane was microscopically visualized using rhodamine labelled phopholipids (Fig. 2). Segregation of Py-PE and Py-PG in bacterial membrane The major result that points to segregation between the two labelled phospholipids, Py-PE and Py-PG, is the very low heteroexcimer formation both in E. coli and B. subtilis in contrast to liposomes (Figs 5A, 6 and 7, Table 3). A possible reason for the absence of complete mixing could be asymmetrical distribution of PE and PG between two leaflets of the membrane. This possibility seems unlikely as it was shown that in bacterial membranes phospholipids undergo a fast transmembrane flip-flop (Huijbregts et al., 1996). The similarity in the results obtained with Gram-positive B. subtilis that has only one membrane, supports the claim that in E. coli the two probes are segregated within the same membrane rather than partitioned into different membranes, outer and inner. It can be assumed that there are at least two types of domains in the bacterial plasma membrane: one enriched with PyPE and another enriched with Py-PG. The domain that is enriched with Py-PE is characterized by a higher local probe concentration, a conclusion that is derived from a higher excimerization rate of Py-PE than that of Py-PG (Fig. 4), whereas rotational correlation time and fluorescence life time were the same for the two phospholipids (Table 2). Moreover, the degree of order of these domains is different as expressed by the different temperatures of phase transitions: 34∞C for Py-PE and 25∞C for Py-PG (Fig. 9A and Table 4). Activation energy calculated for PyPE excimerization was unusually low both above and below phase transition (Table 4). This may be ascribed to the formation of static excimers of Py-PE due to their higher local concentration. Our results reveal that the interaction of membrane proteins with Py-PE may be stronger than those with PyPG: (i) excimerization rate of Py-PE excited through protein tryptophans was 20% lower than with direct excitation yet no difference was observed for the mobility of Py-PG (Table 1). (ii) Resistance to extraction by detergent was different for Py-PE (maximal extraction 75%) and for Py-PG (extraction about 90%). Taking the phospholipid/protein stoichiometry in the E. coli membrane as

1076 S. Vanounou, A. H. Parola and I. Fishov 100/1 (2.35 ¥ 106 protein molecules per cell, 7.5% of them in the inner membrane, and 22.106 lipids per cell, 75% in the inner membrane (Neidhardt and Umbarger, 1966), we calculated that roughly 12–15 molecules of either PE or PG are tightly protein-bound (that is, unextractable by detergent in Cam-treated cells) and at least 10 additional PE molecules confined to its domain (unextractable in normal cells); the other 75 molecules are easily extractable. Summing up, the differences between Py-PE and PyPG mentioned above may be attributed to the localization of Py-PE within a close vicinity of the membrane proteins, i.e. boundary lipid domain (Parola, 1993). This may explain the formation of static excimers of locally concentrated and partially immobilized probe molecules, exhibiting higher phase transition temperature and higher resistance to detergent extraction. Distinctly however, PyPG molecules exhibit characteristics of protein free phospholipid surroundings, similar to that of homogeneous liposomes. Effect of inhibition of protein synthesis According to the transertion model, inhibition of protein synthesis is suggested to dissipate putative membrane domains formed by coupled transcription, translation and insertion of membrane proteins (Binenbaum et al., 1999). In Cam-treated E. coli, this dissipation is reflected in decreased average membrane fluidity as demonstrated previously by DPH fluorescence anisotropy (Binenbaum et al., 1999). Furthermore, analysis of spectral properties of laurdan revealed that the membrane in such cells is characterized by a decreased polarity and higher homogeneity and, hence, co-operativity (Vanounou et al., 2002). The strongly increased pyrene excimerization rate (Table 1) in Cam-treated cells is consistent with these findings. The lowered excimerization rate of Py-PG in these cells (Table 1) may be explained either by its decreased local concentration or by a decreased connectivity in the dissipated PG domain(s). Decreased connectivity is supported by reduced activation energy (13 kJ mol-1) of Py-PG excimerization in Cam-treated E. coli cells. The low activation energy could be attributed to static excimers formed between Py-PG moieties that lack sensitivity to phase transitions (Fig. 9B), similar to that of protein-bound Py-PE. The degree of percolation properties (connection/disconnection) of the membrane may influence in-plane homo- and hetero-dimerization reactions (Thompson et al., 1995; Vaz, 1995) and explain also the unexpected decrease in the formation of mixed excimers in Cam-treated E. coli (Fig. 4B). In B. subtilis, however, the PL composition may be rather different, with a dominant fraction of acidic PL (about 80%, Bishop et al., 1967; Op den Kamp et al., 1969; Minnikin et al., 1972;

Shohayeb and Chopra, 1985; Guffanti et al., 1987). We thus speculate that in B. subtilis PE domain dissipation could increase connectivity, leading to the observed increase in PG excimerization rate in the acidic bulk phase (Table 3). Furthermore, the mixed excimer will be more readily formed (Fig. 6 and Table 3) because of increased border length of the larger number of smaller PE domains (as the result of dissipation) in the ‘sea’ of acidic PL phase. A more rigorous explanation requires an additional detailed knowledge on the composition of putative proteolipid domains, mechanism of their formation and dissipation. These studies are underway in our laboratory. Conclusions We assume that added Py-PL are distributed in bacterial membranes according to hydrophobic matching and polar interactions. Because the fatty acid composition of the probes used is identical, we suppose that the distribution obtained is determined mainly by the polar interactions, influenced, in turn, by protein-lipid binding specificity. It can thus be summed up that in bacterial membranes different native phospholipids are also segregated in distinct domains that differ in composition, proteo–lipid interaction and degree of order. The proteo-lipid domain is enriched with PE. The results obtained here with the pyrene labelled phospholipids are consistent and supportive to those derived from the laurdan studies (Vanounou et al., 2002). They lend further support to the existence of membrane domains with a presumed role in bacterial cell cycle. Whereas imaging studies reveal spatial correlation with bacterial cell cycle (Fishov and Woldringh, 1999; Mileykovskaya and Dowhan, 2000), the mechanism of membrane domain formation, composition and their functional role in bacterial cell cycle still awaits further investigations.

Experimental procedures Bacterial strains and growth conditions Strains of E. coli B/r H266 and B. subtilis OI1 (ilvC1, leu-1) were grown in M-9 minimal salt medium supplemented with 0.4% glucose, and with 0.4% glucose and 1% casein hydrolysate respectively. Bacteria were cultivated at 37∞C with vigorous shaking (Gyrotory Water Bath Shaker, Model G76, New Brunswick Scientific, Edison, NJ). Absorbance as a measure of biomass concentration (OD420; Novaspec II, Pharmacia LKB), cell counts and size distributions (Coulter Counter, Model ZM, Coulter Electronics Ltd, England, 30 m orifice) were used to follow growth and to characterize the physiological state. The fluorescent phospholipids 1-hexadecanoyl-2-(1-pyrenedecanoyl)-sn-glycero-3-phosphoglycerol, ammonium salt (Py-PG) and 1-hexadecanoyl-2-(1-pyrenedecanoyl)-sn-glycero-3-phosphoethanolamine (Py-PE) were purchased from Molecular Probes (Eugene, OR). © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Bacterial membrane domains 1077 Labelling procedure and fluorescence measurements The membrane structure and dynamics depend on its composition, specific interactions between components (proteoprotein, proteo-lipid and lipid-lipid), functional state of proteins and electrochemical gradients across the membrane. Formaldehyde fixation was used in our studies for sample preparation to avoid any changes caused by enzymatic activities during labeling and measurement. The fluorescent probe(s) is therefore expected to detect changes in phospholipid or protein composition and organization, implying that the measured parameters are attributed to a non-active membrane without gradients. Formaldehyde, as a monoaldehyde, inhibits primarily protein function by modifying essential amino groups (Griffiths, 1993 and references cited therein). Although it is able to form methylene cross-links, this reaction can involve mostly various parts of the same protein or different proteins in existing complexes due to the very short link (about 1.5 Å). As was demonstrated in our previous publications (Binenbaum et al., 1999; Fishov and Woldringh, 1999), formaldehyde fixation (at the low, 0.25%, concentration used) does not affect membrane order measured by DPH. It does however, allow revealing dynamic changes in the membrane induced by modulation of the functional state before fixation. It also preserves the presumptive domain structure visualized by the staining pattern of FM 4–64. Samples of bacterial cultures (2–5 ml) either at steadystate growth or treated with 100 mg ml-1 Cam for 1 h were fixed by formaldehyde (0.25% final concentration), collected on polycarbonate membrane filters (0.2 mm pore size, Poretics, USA) and washed with PBS (phosphate buffered saline, pH 7.4). Filters were frozen in liquid nitrogen. For labeling, filters were thawed, bacteria were resuspended in PBS (OD420 = 0.22) and centrifuged. The fluorescent phospholipids (10 ml of methanol solution at a desired concentration) were added to the cell pellet (40 ml) and the mixture was gently homogenized by short exposure (about 10 s) in a water-bath sonicator. After 80 min incubation at 37∞C, PBS was added up to 1 ml and the cells were washed twice with PBS by centrifugation for 10 min to remove excess, nonincorporated label. Finally, the labelled cells were suspended in PBS to OD420 about 0.2 and used for the spectral measurements. The evaluation of labeling conditions is described in Results. For labelling with pyrene or DPH we followed the previously described method (Binenbaum et al., 1999): 1 ml of cells suspension (OD420 = 0.25) was incubated for 5– 45 min at 37∞C in the presence of 10-8-10-7 M dye added from a stock solution in ethanol or tetrahydrofuran; the final solvent concentration never exceeded 0.5%. Steady-state fluorescence spectra were measured with a Perkin-Elmer LS50B spectrofluorometer (Perkin-Elmer, Beaconsfield, England). Fluorescence intensity and anisotropy of DPH and DPH-PC were measured at 430 nm with excitation at 360 nm exactly as described (Binenbaum et al., 1999). Pyrene and Py-PL were excited at 340 nm and fluorescence intensity of monomers and excimers were detected at 377 and 470 nm, respectively. Temperature was controlled to ± 0.2∞C with a water circulating bath at 37∞C or was changed stepwise by about 2∞C, descending from high to low temperature; the sample was allowed to equilibrate for at least 10 min at each temperature with stirring. Time-resolved measurements of fluorescence and rotational correlation time © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

were performed using multifrequency phase modulation spectrofluorometry, already described before (Parola, 1993; Zamai et al., 1998). The detailed experimental conditions are indicated in the text and legends.

Determination of actual concentration of the label in membranes The amount of the probe inserted into the membrane was measured by dissolving the washed cells in 1% SDS (final concentration) at 80∞C for 10 min. In the resulting clear solution, the probe concentration was derived from fluorescence intensity at 377 nm (no excimers were detectable) and calibration curves obtained for SDS solutions of each labelled phospholipid. The found label concentration was normalized by the cell mass in the sample (in units of OD 420) to get an estimation of actual concentration in the membrane.

Preparation of liposomes Multilamellar liposomes were prepared according to a conventional method (Somerharju et al., 1985). Lipids (soyabean phosphatidylcholine, Sigma) were dissolved in chlorophorm/ methanol 9:1 v/v and pyrene phospholipids were added to this solution to reach a desired molar fraction; the lipids were taken to dryness under vacuum for several hours. The dry lipids were dispersed in PBS by vortexing or by brief waterbath sonication at 45∞C.

Extraction with Triton X-100 Escherichia coli, labelled with Py-PL as described above, were preincubated on ice for 20 min and then Triton X-100 (Sigma) was added to final concentrations from 0.02 to 0.15% v/v. After additional 5 min incubation, the cells were washed twice with cold PBS by centrifugation for 10 min at 4∞C. The percent of pyrene-phospholipids resistant to Triton extraction was calculated from the probe concentrations in cells before and after detergent extraction, determined by the SDS dissolution method described above.

Acknowledgements We are indebted to Mrs Emilia Klyman for her dedicated technical assistance. We are grateful to Dr Chithra Hariharan for help in Global analysis. This work was supported by the Seed Grant of the Wise President for R and D at BGU (to I.F. and A.H.P.), by the grant #9027/98 from Israeli Academy of Sciences for purchasing the K-2 Multifrequency phase modulation spectrofluorometer (to A.H.P. and I.F) and partially by the James-Frank Foundation for Laser–Matter Interaction (to A.H.P).

References Barenholz, Y., Cohen, T., Haas, E., and Ottolenghi, M. (1996) Lateral organization of pyrene-labelled lipids in bilayers as determined from the deviation from equilibrium between

1078 S. Vanounou, A. H. Parola and I. Fishov pyrene monomers and excimers. J Biol Chem 271: 3085– 3090. Ben-Shooshan, I., Kessel, A., Ben-Tal, N., Cohen-Luria, R., and Parola, A.H. (2002) On the regulatory role of dipeptidyl peptidase IV (CD = adenosine deaminase complexing protein) on adenosine deaminase activity. Biochim Biophys Acta 1587: 21–30. Bergelson, L.O., Gawrisch, K., Feretti, J.A., and Blumenthal, R., eds. (1995) Special issue on domain organization in biological membranes. Mol Membr Biol 12: 1–162. Binenbaum, Z., Parola, A.H., Zaritsky, A., and Fishov, I. (1999) Transcription- and translation-dependent changes of membrane dynamics in bacteria: testing the transertion model for domain formation. Mol Microbiol 32: 1173– 1182. Bishop, D.G., Rutberg, L., and Samuelson, B. (1967) The chemical composition of the cytoplasmic membrane of Bacillus subtilis. Eur J Biochem 2: 448–453. de Bony, J., Lopez, A., Gilleron, M., Welby, M., Lane’elle, G., Roussea, B., et al. (1989) Transverse and lateral distribution of phospholipids and glycolipids in the membrane of the bacterium Micrococcus luteus. Biochemistry 28: 3728– 3737. Chattopadhyay, A. (ed.) (2001) Dynamics of organized molecular assemblies: from micelles to cells. J Fluoresc (Special Issue) 11: 139–246. Daems, D., Van den Zegel, M., Bones, N., and De Schryver, F.C. (1985) Fluorescence decay of pyrene in small and large unilamellar L, alpha-dipalmitoylphosphatidylcholine vesicles above and below the phase transtion temperature. Eur Biophys J 12: 97–105. Dombek, K.M., and Ingram, L.O. (1984) Effects of ethanol on the Escherichia coli plasma membrane. J Bacteriol 157: 233–239. Edidin, M. (1997) Lipid microdomains in cell surface membranes. Curr Opin Struct Biol 7: 528–532. Edidin, M. (1998) Defining and imaging membrane domains. In Search of a New Biomembrane Model. Vol. 49. Mouritsen, O.G., and Andersen, O.S. (eds). Copenhagen: Munksgaard, Biologiske Skrifter, The Royal Danish Academy of Sciences and Letters, pp. 19–21. Edidin, M. (2001) Shrinking patches and slippery rafts: scales of domains in the plasma membrane. Trends Cell Biol 11: 492–496. Engelke, M., Behmann, T., Ojeda, F., and Diehl, H.A. (1994) Heterogeneity of microsomal membrane fluidity: evaluation using intrinsic tryptophan energy transfer to pyrene probes. Chem Phys Lipids 72: 35–40. Fishov, I., and Woldringh, C.L. (1999) Visualization of membrane domains in Escherichia coli. Mol Microbiol 32: 1166– 1172. Galla, H.-J., and Hartmann, W. (1980) Excimer-forming lipids in membrane research. Chem Phys Lipids 27: 199– 219. Galla, H.-J., and Sackmann, E. (1974) Lateral diffusion in the hydrophobic region of membranes: use of pyrene excimers as optical probes. Biochim Biophys Acta 339: 103–115. Galla, H.-J., Hartmann, W., Theilen, U., and Sackmann, E. (1979) On two-dimensional passive random walk in lipid bilayers and fluid path-ways in biomembranes. J Memb Biol 48: 215–236.

Griffiths, G. (1993) Fine Structure Immunocytochemistry. Heidelberg: Springer Verlag, p. 39. Guffanti, A.A., Clejan, S., Falk, L.H., Hicks, D.B., and Krulwich, T.A. (1987) Isolation and characterization of uncoupler resistant mutants of Bacillus subtilis. J Bacteriol 169: 4479–4485. Horiuchi, S., Marty-Mazars, D., Tai, P.C., and Davis, B.D. (1983) Localization and quantitation of proteins characteristic of the complexed membrane of Bacillus subtilis. J Bacteriol 154: 1215–1221. Huijbregts, R.P., de Kroon, A.I., and de Kruijff, B. (1996) Rapid transmembrane movement of C6-NBD-labeled phospholipids across the inner membrane of Escherichia coli. Biochim Biophys Acta 1280: 41–50. Jones, M.E., and Lentz, B.R. (1986) Phospholipid lateral organization in synthetic membranes as monitored by pyrene-labeled phospholipids: effects of temperature and prothrombin fragment binding. Biochem 25: 567–574. Kinnunen, P.K.J., Koiv, A., and Mustonen, P. (1993) Pyrenelabeled lipids as fluorescent probes in studies in biomembranes and membrane models. In Fluorescence Spectroscopy. Wolfbeis, O.S. (ed.) Berlin: Springer-Verlag, pp. 159– 169. Koppelman, C.-M., Den Blaauwen, T., Duursma, M.C., Heeren, R.M.A., and Nanninga, N. (2001) Escherichia coli minicell membranes are enriched in cardiolipin. J Bacteriol 183: 6144–6147. Kusba, J., Li, L., Gryczynski, I., Piszczek, G., Johnson, M., and Lakowicz, J.R. (2002) Lateral diffusion coefficients in membranes measured by resonance energy transfer and a new algorithm for diffusion in two dimensions. Biophys J 82: 1358–1372. Lehtonen, J.Y., and Kinnunen, P.K. (1997) Evidence for phospholipid microdomain formation in liquid crystalline liposomes reconstituted with Escherichia coli lactose permease. Biophys J 72: 1247–1257. de Leij, L.D., and Witholt, B. (1977) Structural heterogeneity of the cytoplasmic and outer membranes of Escherichia coli. Biochem Biophys Acta 471: 92–104. Linden, C.D., Wright, K.L., McConnell, H.M., and Fox, C.F. (1973) Latteral phase separations in membrane lipids and the mechanism of sugar transport in Escherichia coli. Proc Natl Acad Sci USA 70: 2271–2275. London, E., and Brown, D.A. (2000) Insolubility of lipids in triton X-100: physical origin and relationship to sphingolipid/cholesterol membrane domains (rafts). Biochim Biophys Acta 1508: 182–195. Loura, L.M.S., de Almeida, R.F.M., and Prieto, M. (2001) Detection and characterization of membrane microheterogeneity by resonance energy transfer. J Fluorescence 11: 197–209. Marsh, D. (1995) Specifity of lipid–protein interactions. In Biomembranes. Vol. 1. Lee, A.G. (ed). Greenwich: JAI Press, pp. 137–186. Marty-Mazars, D., Horiuchi, S., Tai, P.C., and Davis, B.D. (1983) Proteins of ribosome-bearing and free-membrane domains in Bacillus subtilis. J Bacteriol 154: 1381–1388. Mileykovskaya, E., and Dowhan, W. (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. © 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

Bacterial membrane domains 1079 Minnikin, D.E., Abdolrahimzadeh, H., and Baddiley, J. (1972) Variation of polar lipid composition of Bacillus subtilis (Marburg) with different growth conditions. FEBS Lett 27: 16– 18. Morein, S., Andersson, A.-S., Rilfors, L., and Lindblom, G. (1996) Wild-type Escherichia coli cells regulate the membrane lipid composition in a ‘window’ between gel and nonlamellar structures. J Biol Chem 271: 6801–6809. Morrisett, J.D., Pownall, H.J., Plumlee, R.T., Smith, L.C., Zehner, Z.E., Esfahani, M., and Wakil, S.J. (1975) Multiple thermotropic phase transition in E.coli membrane lipid. J Biol Chem 250: 6969–6976. Naqvi, K.R., Martins, J., and Melo, E. (2000) Recipes for analyzing diffusion-controlled reactions in two dimensions: time-resolved and steady-state measurements. J Phys Chem B 104: 12035–12038. Neidhardt, F.C., and Umbarger, H.E. (1966) Chemical composition of Escherichia coli. In Escherichia Coli and Salmonella. Cellular and Molecular Biology, 2nd edn. Neidhardt, F.C., Curtiss, R., III, Ingraham, J.L., Lin, E.C.C., Low, K.B., Jr, Magasanik, B., et al. (eds). Washington, DC: American Society for Microbiology Press, pp. 13–28. Norris, V. (1995) Hypothesis: chromosome seperation in E. coli involves autocatalytic gene expression, transertion and membrane-domain formation. Mol Microbiol 16: 1051– 1057. Norris, V., and Fishov, I. (2001) Hypothesis: membrane domains and hyperstructures control bacterial division. Biochimie 83: 91–97. Op den Kamp, J.A.F., Redai, I., and van Deenen, L.L.M. (1969) Phospholipid composition of Bacillus subtilis. J Bacteriol 99: 298–303. Pagano, R.E., Martin, O.C., Kang, H.C., and Haugland, R.P. (1991) A novel fluorescent ceramide analogue for studying membrane traffic in animal cells: accumulation at the golgi apparatus results in alterd spectral properties of the sphingolipids precursor. J Cell Biol 113: 1267–1279. Parente, R.A., and Lentz, B.R. (1985) Advantages and limitaions of 1-palmitoyl-2-[[2[4(6-phenyl-trans-1,3,5-hexa-

© 2003 Blackwell Publishing Ltd, Molecular Microbiology, 49, 1067–1079

trienyl) phenyl]ethyl]carbonyl]-3-sn-phosphatidylcholine as a fluorescent membrane probe. Biochem 24: 6178–6185. Parola, A.H. (1993) Mammalian membranes: structure and function. In Biomembranes, Physical Aspects. Shinitzky, M., (ed.). New York: Balaban Publishers VCH, pp. 159–227. Shohayeb, M., and Chopra, I.J. (1985) Composition of membranes from whole cells and minicells of Bacillus subtillis. J Gen Microbiol 131: 345–354. Somerharju, P.J., Virtanen, J.A., Eklund, K.K., Vainio, P., and Kinnunen, P.K.J. (1985) 1-Palmitoyl-2-pyrenedecanoyl glycerophospholipids as membrane probes: evidence for regular distribution in liquid-crystalline phosphatidylcholine bilayers. Biochem 24: 2773–2781. Thompson, T.E., Sankaram, M.B., Biltonen, R.L., Marsh, D., and Vaz, W.L. (1995) Effects of domain structure on in– plane reactions and interactions. Mol Membr Biol 12: 157– 162. Vanounou, S., Pines, D., Pines, E., Parola, A.H., and Fishov, I. (2002) Coexistence of domains with distinct order and polarity in fluid bacterial membranes. Photochem Photobiol 76: 1–11. Vaz, W.L. (1995) Percolation properties of two-component, two-phase phospholipid bilayers. Mol Membr Biol 12: 39– 43. Welby, M., Poquet, Y., and Tocanne, J.F. (1996) The spatial distribution of phospholipids and glycolipids in the membrane of the bacterium Micrococcus luteus varies during the cell cycle. FEBS Lett 384: 107–111. Welti, R., and Glaser, M. (1994) Lipid domains in model and biological membranes. Chem Phys Lipids 73: 121–137. Woldringh, C.L. (2002) The role of co-transcriptional translation and protein translocation (transertion) in bacterial chromosome segregation. Mol Microbiol 45: 17–29. Zamai, M., Caiolfa, V.R., Pines, D., Pines, E., and Parola, A.H. (1998) Nature of interaction between basic fibroblast growth factor and the antiangiogenic drug 7,7-(carbonylbis[imino-N-methyl-4,2-pyrrolecarbonylimino[N-methyl-4,2pyrrole]-carbonylimino])-bis-(1,3-naphthalene disulfonate). Biophys J 75: 672–682.