Institut de Pharmacologie et Biologie Structurale, Centre National de la Recherche Scientifique/Université Paul Sabatier (UMR 5089), 205 route de Narbonne, 31077, Toulouse Cedex 04, France1
Laboratoire des Biomembranes, UMR 8619 CNRS-Université Paris-Sud, 91405 Orsay Cedex, France2
Lehrstuhl für Biotechnologie, Biozentrum der Universität Würzburg, Am Hubland, D-97074 Würzburg, Germany3
Institut Pasteur, Service de Microscopie électronique, 25 rue du Docteur Roux, 75724 Paris Cedex 15, France4
Centre de Génétique Moléculaire, CNRS, 91190 Gif-sur-Yvette, France5
Author for correspondence: M. Daffé. Tel: +33 561 175 569. Fax: +33 561 175 994. e-mail: daffe{at}ipbs.fr
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ABSTRACT |
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Keywords: cell wall, corynebacteria, mycolic acid, polysaccharide, porin
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INTRODUCTION |
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The chemical structure of the cell wall skeleton of corynebacteria, mycobacteria and related genera has been extensively studied (for reviews see Daffé & Draper, 1998 ; Minnikin & Goodfellow, 1980
; Minnikin & ODonnell, 1984
; Minnikin et al., 1978
; Sutcliffe, 1997
); this is formed by a thick meso-diaminopimelic acid-containing peptidoglycan covalently linked to arabinogalactan, which in turn is esterified by long-chain
-alkyl, ß-hydroxy fatty acids. While these fatty acids in mycobacteria, called eumycolic acids, possess a very long chain (C6090) and may contain various oxygen functions in addition to the ß-hydroxyl group (Daffé & Draper, 1998
), mycolic acids found in other actinomycetes consist of homologous mixtures of saturated and unsaturated acids and contain shorter chains, e.g. C4050 in nocardomycolic acids and C2236 in corynomycolic acids (Collins et al., 1982
; Minnikin & Goodfellow, 1980
; Minnikin & ODonnell, 1984
; Minnikin et al., 1978
). Although they are Gram-positive bacteria, corynebacteria, mycobacteria and closely related micro-organisms share with Gram-negative bacteria the property of forming in their envelope an outer barrier that is distinct from the plasma membrane. While this additional barrier in Gram-negative micro-organisms is a typical bilayer of phospholipid and lipopolysaccharide, in mycobacte 9ria and corynebacteria the cell-wall-linked mycolates and corynomycolates certainly participate in this barrier since the disruption of genes that code for mycoloyltransferases, the antigen-85 complex and PS1, respectively, causes a decrease in the amount of cell wall-bound mycolates and affects the permeability of the envelope of the mutants (Jackson et al., 1999
; Puech et al., 2000
). Evidence has also been presented that the chemical structure of mycolic acids plays a role in determining the fluidity and permeability of the mycobacterial cell wall (George et al., 1995
; Liu et al., 1996
; Dubnau et al., 2000
). The existence in mycobacteria, corynebacteria and related genera of an outer membrane diffusion barrier is reinforced by the characterization of cell envelope proteins with pore-forming ability (Kartmann et al., 1999
; Lichtinger et al., 1998
, 1999
; Rieß et al., 1998
; Mukhopadhyay et al., 1997
; Trias et al., 1992
; Trias & Benz, 1994
; Senaratne et al., 1998
).
In all currently proposed models (Minnikin 1982 ; Rastogi, 1991
; Liu et al., 1995
) the outer permeability barrier of mycobacteria consists of a monolayer of mycoloyl residues covalently linked to the cell wall arabinogalactan and includes other lipids which are probably arranged to form a bilayer with the mycoloyl residues. Although no sign of a second lipid bilayer has ever been reported in thin sections of mycobacterial cells (Draper, 1998
), freeze-fractured samples of mycobacteria, corynebacteria and related genera (Barksdale & Kim, 1977
; Benedetti et al., 1984
; Chami et al., 1995
; Takeo et al., 1984
) showed that these organisms had two such planes of weakness in their envelopes, in addition to the expected plasma membrane fracture. A second fracture plane, close to the cell surface of mycobacteria, was observed. The existence of distinct lipid domains in the mycobacterial cell envelope has been indicated by studies using progressive erosion of the envelope (Ortalo-Magné et al., 1996
) or selective lipophilic probes (Christensen et al., 1999
). Freeze-fracture electron microscopy also showed the presence of ordered arrays on the surface of Corynebacterium glutamicum and other Corynebacterium spp. (Chami et al., 1995
; Peyret et al., 1993
; Soual-Hoebeke et al., 1999
); these surface layer proteins (S-layer) have been shown to be composed of the major corynebacteria-secreted protein, PS2 (Chami et al., 1995
; Peyret et al., 1993
; Soual-Hoebeke et al., 1999
). Such crystalline structures are not visible in published freeze-fractured micrographs of mycobacteria despite a singular report of a crystalline protein layer on the outside of Mycobacterium bovis BCG (Lounatmaa & Brander, 1989
).
From recent advances in molecular biology, it appears that corynebacteria represent a model suitable for functional studies of expression of mycobacterial genes, primarily because (i) they possess the simplest cell wall structure, but one that is functionally and structurally close to those of mycobacteria and related genera; (ii) they are rapid growers and do not form clumps; and (iii) heterologous expression of mycobacterial antigens in C. glutamicum has proved to be effective (Salim et al., 1997 ; Puech et al., 2000
). However, it has to be remembered that, unlike the other related micro-organisms, corynebacteria are widely used in biotechnology for the production of amino acids (Krämer, 1994
); this singular property of corynebacteria suggests that, despite the global similarity of their cell envelope composition with those of related genera, they differ from them in the arrangement of its constituents. To gain further insights into the cell envelope structure of corynebacteria, various representative corynebacterial strains were examined in the present study by combining biochemical analyses and electron microscopy studies. These included the Corynebacterium genus type species C. diphtheriae, the amino-acid-producing strain C. glutamicum, corynomycolate-less strains, i.e. C. amycolatum (Collins et al., 1988
; Barreau et al., 1993
) and defined corynebacterial cell wall mutants (Chami et al., 1995
; Peyret et al., 1993
; Joliff et al., 1992
; Puech et al., 2000
).
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METHODS |
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Isolation of extracellular materials.
Bacteria were grown for 48 h on minimal medium and the culture filtrates were recovered by centrifugation, filtered through a 0·2 µm sterile filter (Nalgene) and concentrated under vacuum to 1/10 of the original volume; the corresponding cells were dried and weighed. Chloroform and methanol were added to a portion of the filtrates to isolate pids as described below and cold ethanol (6 vols) was added to the remaining portion; after centrifugation of the ethanol precipitates, the pellets were extensively dialysed against distilled water and extracellular materials were obtained and analysed for carbohydrate and protein content as described by Lemassu & Daffé (1994) .
Isolation of the crude surface-exposed materials.
Cells grown in either BHI or minimal medium were harvested by centrifugation, extensively washed and gently shaken for 1 min with 10 g glass beads (4 mm diameter) per 2 g of wet cells (Ortalo-Magné et al., 1995 ). The resulting material was then suspended in distilled water and immediately filtered through a 0·2 µm sterile filter (Nalgene) and concentrated under vacuum to 1/10 of the original volume. Cold ethanol (6 vols) was added to a portion of the crude surface-exposed material, and the ethanol-precipitates were extensively dialysed against distilled water, lyophilized, and analysed for their carbohydrate and protein compositions.
CHCl3 and CH3OH were added to the remaining filtrate to give a final one-phase mixture of H2O/CHCl3/CH3OH (0·8:1:2, by vol.; Bligh & Dyer, 1959 ) and lipids were extracted for 1 h at room temperature; a two-phase partition mixture was obtained by adding 1 vol. CHCl3/H2O (1:1, v/v) to the one-phase solution and lipids were recovered in the lower organic phase, dried and analysed by thin-layer chromatography (TLC) as described below.
Isolation, fractionation and analysis of whole-cell lipids.
Lipids were obtained and analysed as previously described (Puech et al., 2000 ). Briefly, lipids were extracted from wet cells for 16 h with CHCl3/CH3OH (1:1, v/v) at room temperature with continuous stirring; the bacterial residues were re-extracted three times with CHCl3/CH3OH (2:1, v/v) and the organic phases were pooled and concentrated. The crude lipid extracts were partitioned between the aqueous and the organic phases arising from a mixture of CHCl3/CH3OH/H2O (8:4:2, by vol.); the lower organic phases were collected, evaporated to dryness to yield the crude lipid extracts from each strain and comparatively examined by TLC on silica gel-coated plates (G-60, 0·25 mm thickness, Merck) developed with CHCl3/CH3OH (9:1, v/v) or CHCl3/CH3OH/H2O (30:8:1 or 65:25:4, by vol.). Detection of all classes of lipids was performed by spraying the TLC plates with either rhodamine B or 20% H2SO4 in water, the latter followed by heating at 110 °C; glycolipids were revealed by spraying plates with 0·2% anthrone (w/v) in concentrated H2SO4, followed by heating at 110 °C. The DittmerLester reagent (Dittmer & Lester, 1964
) was used for visualizing phosphorus-containing lipids.
In a parallel experiment, surface-exposed lipids were isolated from C. glutamicum cells by extracting bacterial cells with octylglucoside. Briefly, C. glutamicum CGL2005 cells were grown overnight in BHI-rich medium. After extensive washing (twice the culture volume), cells were incubated for 1 h in 25 mM Tris/HCl, pH 6·8 buffer containing 1% octylglucoside. The crude detergent extract was recovered by centrifugation at 8000 r.p.m. for 15 min, extensively dialysed for 24 h against 25 mM Tris/HCl, pH 6·8 buffer, and then centrifuged at 200000 g for 30 min; the pellet was analysed by freeze-fracture electron microscopy.
Quantification of corynomycolic acids was performed as follows. Delipidated cells (1·5 g dry weight) and lipid extracts (100 mg) of the various strains were dried under vacuum prior to weighing and were saponified (Daffé et al., 1983 ). The saponified products were acidified with 20% H2SO4 and the resulting fatty acids were extracted with diethyl ether, converted to methyl esters with diazomethane and dried under vacuum, dissolved in petroleum ether and applied to a Florisil (60100 mesh, Merck) column equilibrated in petroleum ether. The column was irrigated stepwise with increasing concentrations of diethyl ether in petroleum ether. Fractionations were monitored by TLC on silica gel-coated plates using dichloromethane and fractions containing the same lipid compounds (non-hydroxylated fatty acid methyl esters or corynomycolates) were pooled and weighed. Three sequential determinations from separate preparations of delipidated cells were performed (Puech et al., 2000
).
Fatty acid methyl esters from bacteria, delipidated cells and extractable lipids (12 mg) were treated with trimethylsilyl reagents (Sweeley et al., 1963 ) to derivatize hydroxylated components of the mixtures, i.e. corynomycolates, and analysed by GC. The detector response for the various classes of fatty acid methyl esters was determined using authentic samples of C16:0 and C32:0 corynomycolate methyl esters. Identification of non-hydroxylated fatty acid methyl esters and corynomycolate derivatives was achieved by gas chromatographmass spectrometry (GCMS).
Production of cell wall, plasma membrane and cytosol fractions.
The cell fractions of the various corynebacteria were produced as previously described for mycobacteria (Daffé et al., 1990 ; Raynaud et al., 1998
). Wet cells (5 g) were suspended in 20 ml phosphate buffer (50 mM, pH 7·5) and the resulting bacterial suspension was passed through a cell disrupter and then centrifuged at 3000 r.p.m. for 15 min to eliminate unbroken cells; cell walls were recovered from the supernatant by recentrifugation at 10000 g for 1 h. The resulting pellet was extracted with aqueous 2% SDS at 95 °C for 1 h to remove soluble proteins and finally pelleted at 10000 g; the cell walls were washed three times with 80% acetone to remove SDS and lyophilized (Daffé et al., 1990
). The 10000 g supernatant was recentrifuged at 100000 g for 2 h to yield the membrane fraction in the pellet; the supernatant was considered as the cytosol (Raynaud et al., 1998
).
Analysis of the cell wall arabinogalactans.
O-Methylation of cell wall arabinogalactan was performed by suspending lyophilized walls (5 mg) in 3 ml of dimethylsulphoxide; 300 µl of dimethylsulphinyl carbanion was added and the mixture was stirred for 5 h. CH3I was slowly added and the suspension was stirred for 2 h. A mixture of CH3OH/H2O (1:1, v/v) was added and the reaction mixture was dialysed against water. The addition of dimethylsulphoxide, dimethylsulphinyl carbanion and CH3I, followed by dialysis, was repeated four times. The retentate was lyophilized and the per-O-methylated arabinogalactan was hydrolysed with 2 M CF3COOH at 110 °C for 2 h. The hydrolysed products were reduced by NaBH4, acetylated and the resulting partially O-methylated, partially O-acetylated alditols were analysed by GC-MS (Daffé et al., 1990 ).
Production and analysis of lipoarabinomannans and lipomannan.
Delipidated cells (7 g) were extracted with 100 ml ethanol/water (1/1, v/v) for 2 h at 70 °C; the bacterial residues were recovered by filtration and re-extracted with the same solvent mixture. The two extracts were pooled and dried; then, 100 ml hot phenol/water (1/1, v/v) was added and the mixture was heated for 5 min under continuous stirring followed by partitioning of the two phases. The phenol phase was discarded and the upper phase was extensively washed and dried. The extract was solubilized in water and Triton-X114 (2% w/v) was added to the cooled suspension; the mixture was stirred for 5 min and then heated at 50 °C until two phases had formed. The detergent phase was recovered, diluted by adding 2 ml water and washed three times with CHCl3. The resulting aqueous phase was concentrated to a final volume of 0·5 ml and the macromolecules were precipitated with 6 vols cold ethanol. The precipitate was recovered, resuspended in water (0·5 ml) and the macromolecules were reprecipitated with 6 vols cold ethanol. The resulting pellet was dissolved in 100 mM sodium acetate buffer (pH 4·7), containing 15% (w/w) propan-1-ol and fractionated by hydrophobic interaction chromatography (Leopold & Fisher, 1993 ) by loading the material onto a column of octyl-Sepharose CL-4B (Pharmacia LKB) equilibrated with the buffer; the column was first irrigated with this buffer and then with the buffer containing 20, 30, 40, 50 and 60% (v/v) n-propanol. Fractions were collected in bulk, dialysed and monitored for carbohydrate; a portion of each dialysate was analysed for its carbohydrate and lipid compositions. The remaining portion was analysed by SDS-PAGE; authentic samples of mycobacterial lipoarabinomannan and lipomannan from Mycobacterium bovis BCG, kindly provided by T. Brando (IPBS, Toulouse, France) were used as standard.
Preparation of proteins for SDS-PAGE and Western blot analysis.
Cell wall proteins from the various strains were released by incubating the cell pellet with 2% (w/v) SDS in 50 mM Tris/HCl (pH 6·8) at 100 °C for 2 min (50 µl buffer per equivalent of 0·4 ml bacterial suspension at OD650 1·2); the suspension was centrifuged at 12000 g for 3 min and the supernatant, which contains only cell wall-associated proteins and not cytoplasmic proteins (Peyret et al., 1993 ), was collected. Cell wall proteins from roughly 0·4 ml bacterial suspension at OD650 1, were analysed by SDS-PAGE. The pore-forming proteins were isolated from the culture filtrates and different cell fractions as previously described (Lichtinger et al., 1998
). Proteins were separated by SDS-PAGE (Laemmli, 1970
) with a 4% stacking gel and a 7% or a 10% running gel. Samples were denatured in the presence of 2% SDS in 50 mM Tris/HCl (pH 6·8). After electrophoresis, gels were either stained with Coomassie brillant blue R-250 or blotted onto nitrocellulose membrane (Towbin et al., 1979
). In this latter case proteins were probed with rabbit polyclonal antibodies anti-PS1 (Joliff et al., 1992
), anti-PS2 (Peyret et al., 1993
), or anti-C. glutamicum porin (Lichtinger et al., 1998
). Bands were detected using alkaline-phosphatase-conjugated antibodies and nitroblue tetrazolium (NBT)/5-bromo-4-chloro-3-indoxyl phosphate (BCIP) p-toluidine (Promega) as substates.
Sugar compositional analysis.
The sugar constituents of the various materials were determined after acid hydrolysis either with 2 M CF3COOH at 110 °C for 2 h or with 1 M methanolic HCl at 80 °C for 16 h; the mixture of hydrolysed products was dried, treated with trimethylsilyl reagents (Sweeley et al., 1963 ) to derivatize monosaccharides and analysed by GC for their sugar.
Gas chromatography and mass spectrometry.
GC was performed using a Hewlett Packard HP4890A equipped with a fused silica capillary column (25 m length by 0·22 mm internal diameter) containing WCOT OV-1 (0·3 mm film thickness, Spiral). A temperature gradient of 100290 °C at 5 °C min-1, followed by a 10 min isotherm plateau at 290 °C, was used.
GC-MS analysis was conducted on a Hewlett Packard 5890 gas chromatograph connected to a Hewlett Packard 5989A mass spectrometer. Samples were injected in the splitless mode. The column was a 12 m HP-1. A temperature gradient of 100290 °C (8 °C min-1) was used.
Experiments with lipid bilayer membranes.
Black lipid bilayer membranes were formed as described previously (Benz et al., 1978 ). The instrumentation consisted of a Teflon chamber with two aqueous compartments connected by a small circular hole. The hole had a surface area of about 0·5 mm2. Membranes were formed across the hole by painting on a 1% solution of a mixture (molar ratio 4:1) of diphytanoyl phosphatidylcholine (PC) and phosphatidylserine (PS) (Avanti Polar Lipids, Alabaster AL) in n-decane.
Electron microscopy
Transmission electron microscopy.
Bacterial pellets were fixed for 1 h at 4 °C with 2·5% (w/v) glutaraldehyde and 0·05 M lysine in a 0·1 M cacodylate buffer (pH 7·4) containing, or not, 0·075% (w/v) ruthenium red. Cells were washed five times in the same buffer, i.e. with or without 0·075% (w/v) ruthenium red, and postfixed for 1 h at room temperature in 1% (w/v) osmium tetroxide containing, or not, 0·075% (w/v) ruthenium red in cacodylate buffer (pH 7·4), and then rinsed with distilled water. Bacteria were suspended in 1% (w/v) aqueous uranyl acetate for 1 h at room temperature and then washed five times in distilled water. Suspended cells were embedded in 2% molten agar type IX (Sigma) before dehydration through a graded ethanol series. Dehydrated cells were embedded in Spurr medium with intermediate 1,2-epoxypropane infiltration. Blocks were conventionally cut, stained and examined with a Philips CM12 microscope operating under standard conditions.
Freeze-etched preparations for electron microscopy.
Bacterial suspensions were centrifuged at 5000 g. A drop of the pellet was placed between thin copper holders and quenched in liquid propane. The frozen samples were fractured at -125 °C in a vacuum of about 10-7 torr as described by Aggerbeck & Gulik-Krzywicki (1986) . The fractured samples were etched at -100 °C for 3 min at 1·3x10-5 Pa and then replicated with 11·5 nm of deposits of platinum-carbon, and backed with about 20 nm of carbon. The replicas were cleaned overnight with chromic acid, washed with distilled water and observed with a Philips 410 electron microscope.
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RESULTS |
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Chemical analysis of the corynebacterial outermost polysaccharides
Analysis of the acid hydrolysis products originated from the extracellular and surface-exposed macromolecules of the various strains by GC showed that the sugar composition of the outermost constituents of the vast majority of the corynebacterial species analysed was similiar to that in mycobacteria (Lemassu & Daffé, 1994 ; Lemassu et al., 1996
; Ortalo-Magné et al., 1995
); these were 1020% arabinose (Ara), 2035% mannose (Man) and 5070% glucose (Glc). The outermost polysaccharides from C. amycolatum were almost exclusively composed of Glc. To characterize the polysaccharides that are composed of these sugar residues, the crude ethanol precipitates from the extracellular and surface-exposed macromolecules were further investigated; the precipitates were first treated with proteases and then chromatographed on an ion-exchange column (Lemassu & Daffé, 1994
). In both cases, most of the carbohydrate material (>90%) was eluted from the column with the buffer containing no salts (data not shown), indicating that they were composed of neutral substances; gel-filtration chromatography over a Bio-Gel P-10 column led to the isolation of three fractions (Fig. 2a
). The major fraction (peak A) was exclusively composed of Glc and was eluted in the void volume; gel-filtration chromatography over a Sephadex G-200 column of the glucan fraction led to the isolation of a single peak A1 at a position corresponding to an apparent molecular mass of 110 kDa (Fig. 2b
). The second peak of the Bio-Gel P-10 column (peak B, Fig. 2a
) contained Ara and Man (in a molar ratio of 1:1) and exhibited an apparent molecular mass of 13 kDa. The last fraction eluted from the Bio-Gel P-10 column was a large, broad peak (peak C, Fig. 2a
) and contained Ara, Man and Glc; gel-filtration chromatography over a Bio-Gel P-4 column of this material gave two peaks (Fig. 2c
) at positions corresponding to apparent molecular masses of 1·7 kDa (peak C1, glucan, Fig. 2c
) and 1 kDa (peak C2, arabinomannan, Fig. 2c
). A similar pattern was observed when the macromolecules derived from the culture filtrate of C. xerosis were analysed. Thus, like mycobacteria (Lemassu & Daffé, 1994
; Lemassu et al., 1996
; Ortalo-Magné et al., 1995
), C. xerosis elaborates extracellular and surface-exposed polysaccharides composed of a 110 kDa high-molecular-mass glucan and a 13 kDa arabinomannan. In contrast to what was found in mycobacteria, low-molecular-mass arabinomannan and glucan were present in the mixture of polysaccharides from the strains examined.
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Distribution of the corynomycoloyltransferase PS1 and the S-layer constituent PS2 in corynebacteria
It was recently demonstrated that one of the two major secreted protein from C. glutamicum, PS1, possesses a mycoloyltransferase activity in this species (Puech et al., 2000 ). However, like the other major secreted S-layer constituent, PS2, the occurrence of PS1 in different corynebacteria has not been investigated. To determine the distribution of PS1, proteins extracted from the outermost cell envelope compartment were analysed by SDS-PAGE. Compared to those of mycobacteria (Ortalo-Magné et al., 1995
), the protein profiles of corynebacteria were less complex (Fig. 4a
, consisting only of a few major bands, notably three bands with electrophoretic migrations similar to those of PS1 and PS2 (in the 6070 kDa region). To characterize these proteins further, Western blot analyses were performed on them using polyclonal antibodies directed against PS1 and PS2. The anti-PS1 antibodies reacted strongly with polypeptide bands exhibiting the corresponding molecular mass in C. glutamicum (positive control, Fig. 4b
, lane 1), one strain of C. xerosis (ATCC 9016; Fig. 4b
, lane 3) and C. amycolatum (Fig. 4b
, lane 5), and gave a faint reaction with proteins from C. diphtheriae (Fig. 4b
, lane 4) and the type strain of C. xerosis (Fig. 4b
, lane 2). In these latter strains, stronger reactions were observed with other polypeptides which may share some epitopes with PS1. Although the corynomycoloyltransferase activity of PS1 (Puech et al., 2000
) was not investigated, these data suggest the presence of PS1 in all the strains examined, including in those devoid of corynomycolates, i.e. C. amycolatum (Collins et al., 1988
; Barreau et al., 1993
). In contrast, the anti-PS2 antibodies reacted only with proteins originated from the PS2+ parent strain of C. glutamicum (positive control); proteins from the PS2- strain CGL2025 of C. glutamicum (cspB-disrupted mutant, negative control), as well as those from the other corynebacterial strains examined did not react with the anti-PS2 antibodies (data not shown). Although the production of PS2 was shown to greatly depend on the carbon source (Soual-Hoebeke et al., 1999
), the failure to detect this protein was not attributed to the growth medium used to analyse the other corynebacterial strains since all the strains examined were grown on the same BHI medium, which had been shown to allow C. glutamicum strain CGL2005 to produce a highly ordered surface layer composed of PS2 (Chami et al., 1995
; Peyret et al., 1993
). It was thus concluded that the S-layer component PS2 was absent from most of the strains examined herein.
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Non-covalently associated cell envelope lipids
To explain the occurrence of the cell wall fracture plane in strains exhibiting a very low cell-wall-associated corynomycolate content, we postulated that other lipids may replace corynomycoloyl residues in the inner monolayer of the outer membrane, based on the structural similarities between C3236 corynomycolate-containing molecules, e.g. TDCM and TMCM (with two parallel C1618 chains), and C1618-containing lipids such as phospholipids. Therefore, non-covalently associated lipids were extracted from bacterial cells grown with organic solvents and weighed (Table 3). Interestingly, strains of C. amycolatum and the type strain of C. xerosis contained 36% of extractable lipids whereas the other corynebacterial strains produced significantly more lipids. TLC analysis of these lipids showed that their composition was similar to that found in the extracellular and surface-exposed materials; TDCM, TMCM and phospholipids were the main substances detectable in all the strains, except C. amycolatum, where only phospholipids were observed. Compared to the outermost lipid materials, however, many more phospholipids were detected in bacterial lipids, as expected from their partiality for plasma membrane. It was thus concluded that the presence of significant amounts of extractable lipids, e.g. TDCM and TMCM, probably contributes to the occurrence of a cell wall fracture plane in corynebacteria.
To evaluate the possible contribution of non-covalently bound lipids in the cell wall permeability barrier, the outermost material of C. glutamicum was extracted with octylglucoside; this method did not affect the integrity of bacterial cells, as judged by freeze-fracture microscopy (data not shown). The crude detergent extract, which contained mostly polysaccharides and lipids, was extensively dialysed to eliminate the detergent and to reconstitute organized structures; it was then centrifuged and the pellet was analysed by freeze-fracture electron microscopy (Fig. 5g). Homogeneous smooth vesicles whose diameter varied from 50 to 500 nm were observed throughout the samples analysed, indicating that the material examined spontaneously forms liposomes upon dialysis of the detergent; analysis of the lipid material by TLC showed that it primarily consisted of trehalose dicorynomycolates, with small amounts of phospholipid and trehalose monocorynomycolates. It follows then that the non-covalently bound lipids from the cell envelope of corynebacteria are strong candidates for participating in a membrane-like structure.
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DISCUSSION |
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As far as the ultrastructural appearance is concerned, the cell envelope of corynebacteria is similar to that of mycobacteria, consisting of a typical plasma membrane bilayer, a thick electron-dense layer (EDL), an electron-transparent layer (ETL) and an outer layer (OL). The electron density of the EDL makes it likely that it contains the cell wall peptidoglycan, which possesses charged groups able to bind metallic stains used in electron microscopy. The EDL is surrounded by a thin ETL which is traditionally considered to consist of mycolic residues, based on the transparency of this layer to electrons; indeed, the occurrence of eumycoloyl, nocardomycoloyl, or corynomycololyl residues covalently linked to the cell wall arabinogalactan of mycobacteria and related genera, on the one hand, and the effect of the removal of lipids from isolated mycobacterial cell walls, which causes the disappearance of the layer, on the other hand, support this interpretation. Surprisingly, however, this layer was seen in all corynebacterial species examined, including C. amycolatum, which is devoid of corynomycolates (Fig. 1); indeed, this observation could be explained by the replacement on the arabinogalactan of C. amycolatum of corynomycoloyl residues by normal-chain fatty acyl residues, i.e. C1618. This hypothesis was ruled out by the analysis of the purified cell walls of the bacterium, where no fatty acyl substituent could be detected, indicating that the ETL was not synonymous with a lipid layer. Furthermore, the thickness of this layer in corynebacteria is comparable to that measured in mycobacteria (67 nm). Assuming a linearity between the thickness of the hydrocarbon portion and the number of carbon atoms, the theoretical length of the C5060 main alkyl chain, the so-called meromycolate chain, of eumycoloyl residues is estimated to be 6·5 nm (see Daffé & Draper, 1998
) and that of corynomycoloyl residues (C16) should be one-third of that observed in mycobacteria. Clearly, further studies are warranted to decipher the chemical nature of the ETL in corynebacteria and mycobacteria. The OL of the corynebacterial strains examined were heavily stained with ruthenium red, as previouly seen for mycobacteria (Rastogi et al., 1986
) but in apparent conflict with a previous report (Rastogi et al., 1984
). Although it is not clear what bacterial components react with the dye, it is evident that these are not acidic polysaccharides as previously claimed (Rastogi et al., 1984
), primarily because the outermost carbohydrate constituents of corynebacteria (this study) and mycobacteria (Ortalo-Magné et al., 1995
; Lemassu et al., 1996
) were found almost exclusively to be neutral substances.
Consistent with the phylogenetic relatedness between mycobacteria and corynebacteria is the observation that the main structural features of the major polysaccharides of the two genera were very similar. No gross structural difference was found between the non-covalently associated cell envelope polysaccharides, i.e. the 110 kDa glucan, 13 kDa arabinomanan and lipoglycans, characterized in this study and those published for mycobacteria (Lemassu & Daffé, 1994 ; Lemassu et al., 1996
; Ortalo-Magné et al., 1995
; Hunter et al., 1986
); in addition, low-molecular-mass glucan and arabinomannan, not previously described in mycobacteria, were identified in corynebacteria. The backbone of the major cell wall polysaccharide, arabinogalactan, was also found to be held in common between mycobacteria (Daffé et al., 1990
, 1993
) and corynebacteria (Abou-Zeid et al., 1982
; this study). In contrast to mycobacteria, however, while the mycobacterial arabinogalactans are almost exclusively composed of Ara and Gal residues (Daffé et al., 1993
), significant amounts of Man and Glc residues may be found covalently linked to arabinan and/or galactan segments, a situation encountered in other actinomycetes such as rhodococci and nocardiae (Daffé et al., 1993
). As in these latter genera, the glycosyl-linkage composition of arabinogalactan in corynebacteria may vary according to the species examined.
Although they are Gram-positive bacteria, corynebacteria and closely related micro-organisms share with Gram-negative bacteria the property of possessing in their envelopes an outer barrier that is distinct from the plasma membrane. While this additional barrier in Gram-negative micro-organisms is a typical bilayer of phospholipid and lipopolysaccharide, in mycobacteria and corynebacteria the cell-wall-linked mycolates and corynomycolates certainly contribute to this barrier, since the disruption of genes that code for mycoloyltransferases causes a decrease in the amount of cell wall-bound mycolates and corynomycolates, and affects the permeability of the envelope of the mutants (Jackson et al., 1999 ; Puech et al., 2000
). This outer permeability barrier also includes non-covalently linked lipids which are probably arranged to form a bilayer with the corynomycoloyl residues, as proposed for mycobacteria (Minnikin 1982
; Rastogi, 1991
; Liu et al., 1995
). As expected from these data, freeze-fractured samples of mycobacteria, corynebacteria and related genera showed that these organisms possess, in addition to the expected plasma membrane fracture, a major plane of weakness in their cell walls, close to the cell surface of mycobacteria (Chami et al., 1995
; Benedetti et al., 1984
). The present study extends this concept by showing that the various corynebacterial strains that are devoid of corynomycolates, i.e. C. amycolatum, exhibit only one fracture plane, which occurs in their plasma membrane, whereas the corynomycolate-containing strains exhibit a cell wall fracture plane, with the notable exception of the type strain of C. xerosis. Paradoxically, this strain possesses a relatively high content of covalently linked cell wall corynomycolates (Table 3
) but most cells exhibit only a plasma membrane fracture plane, indicating that the corynomycolate content is not sufficient to explain the occurrence of the cell wall fracture plane. Furthermore, in most corynebacterial species examined, notably the csp1-inactivated mutant of C. glutamicum (CGL2022), the amount of covalently linked cell wall corynomycolates (Table 3
) is less than the 0·085 µmol of mycolic acids necessary for forming a monolayer which would cover the 230 cm2 bacterial cell surface area (calculated for 1 mg of bacterial dry weight; see Nikaido et al., 1995
). To reconcile this fact with the occurrence of the second fracture plane in the cell envelope of most of the corynomycolate-containing species, we reasoned that other non-covalently linked lipids, e.g. trehalose dicorynomycolate, may interact with the covalently bound cell wall corynomycolates to form a monolayer which would cover the bacterial cell surface area. This assumption was based first on the structural analogy between C3236 corynomycolate-containing molecules, whose two C1618 chains would be involved in hydrophobic interactions in a monolayer model, and C1618-containing complex lipid compounds. In this respect, it has been shown that, like phospholipids, suspensions of synthetic trehalose dicorynomycolates exhibited well-defined transition phases and that these suspensions in a fluid state were clearly more active on mitochondria than in a gel state (Durand et al., 1979
). Secondly, and more importantly, we showed that trehalose dicorynomycolates are capable of forming bilayers. Consequently, a quantification of lipid extracts from the various corynebacterial strains was undertaken (Table 3
); this led to the demonstration that strains exhibiting a second fracture plane contained more extractable lipids than those lacking the cell wall fracture plane. Based on this observation, we speculate (Fig. 7
) that in most corynebacteria, non-covalently linked lipid molecules are involved in the monolayer that is traditionally composed of mycolates. Thus, although the cell envelopes of corynebacteria and mycobacteria have several chemical and physical properties in common, the two organisms may be different in terms of the intimate arrangement of their constituents.
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Received 30 October 2000;
revised 30 January 2001;
accepted 1 February 2001.