Institut de Pharmacologie et de Biologie Structurale, Unité Mixte de Recherche du Centre de National de Recherche Scientifique et de lUniversité Paul Sabatier (UMR 5089), 205 route de Narbonne, 31077 Toulouse cedex 04, France1
Department of Microbiology and Immunology, University of Melbourne, Victoria 3010, Australia2
Institut dExploration Fonctionnelle des Génomes (IFR 109), 118 route de Narbonne, 31062 Toulouse cedex, France3
Author for correspondence: Mamadou Daffé. Tel: +33 561 175 569. Fax: +33 561 175 580. e-mail: mamadou.daffe{at}ipbs.fr
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ABSTRACT |
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Keywords: mycobacteria, cell wall permeability, morphology, ultrastructure, glycolipids
Abbreviations: GPL, glycopeptolipid; MDM, monocyte-derived macrophage; OL, outer layer; SEM, surface-exposed material
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INTRODUCTION |
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The high lipid content of mycobacterial cells, recognized as early as the 1930s, and the extraordinary biological activities associated with some of the purified lipids (for reviews see Daffé & Draper, 1998 ; Goren & Brennan, 1979
), have prompted researchers to devote much effort to the identification of the various types of mycobacterial lipids. Among the characteristic lipids of mycobacteria are the species-specific glycopeptidolipids (GPLs) that typify many non-tuberculous mycobacterial species; these molecules may be subdivided into alkali-stable C-type GPLs and alkali-labile serine-containing GPLs (Daffé & Lemassu, 2000
). The C-type GPLs have been found in saprophytic mycobacteria (Mycobacterium smegmatis), opportunistic pathogens of man (Mycobacterium aviumintracellulare, Mycobacterium scrofulaceum, Mycobacterium peregrinum, Mycobacterium chelonae, Mycobacterium abscessus) and of animals (Mycobacterium lepraemurium, Mycobacterium paratuberculosis, Mycobacterium porcinum, Mycobacterium senegalense) (Brennan, 1988
; Daffé & Lemassu, 2000
) whereas the alkali-labile serine-containing GPLs have been found so far only in Mycobacterium xenopi (Besra et al., 1993
; Rivière & Puzo 1991
). C-type GPLs share a common lipopeptidyl core consisting of a mixture of 3-hydroxy and 3-methoxy C2634 fatty acids (Daffé et al., 1983
) amidated by a tripeptide D-Phe-D-allo-Thr-D-Ala and terminated by L-alaninol (Fig. 1
). They differ from one another by the number and the nature of the saccharide units linked to the hydroxyl group of allo-Thr and/or alaninol (Brennan, 1988
; Daffé & Lemassu, 2000
). These glycosyl units are responsible for both the variability and the specificity of the antigenically distinct serovariants within the M. avium-intracellulare complex and several other C-type GPL-containing mycobacteria (Brennan, 1988
), which implies that these compounds occur on the peripheral bacterial surface, in agreement with their isolation from the outermost compartment of mycobacterial cells (Ortalo-Magné et al., 1996
) and their identification as the receptor of mycobacteriophage D4 (Furuchi & Tokunaga, 1972
; Goren et al., 1972
). Although the role of GPLs in pathogenesis is still unclear (Daffé & Draper, 1998
), they accumulate into the phagosome during bacterial intracellular growth, contributing to the formation of a capsule around the bacteria (Draper, 1974
; Rulong et al., 1991
; Tereletsky & Barrow, 1983
). Less is known, however, about the localization of GPLs in deeper compartments of the mycobacterial cell envelope and their possible contribution to the cell wall permeability barrier. Thanks to the availability of M. smegmatis mutants with transposon insertions in the genes involved in the synthesis of the C-type GPL core (Billman-Jacobe et al., 1999
; Recht et al., 2000
), we addressed the question of the effects of the absence of C-type GPLs on the architecture and surface properties of the cell envelope of M. smegmatis and on the initial interactions of bacteria with host cells.
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METHODS |
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The viability of mycobacterial cells was assessed by i) quantifying the level of isocitrate dehydrogenase, an indicator of autolysis, in the culture filtrates (Raynaud et al., 1998 ) and ii) labelling cells with both propidium iodide and fluorescein diacetate (Cougoule et al., 2002
); the percentages of live (fluorescein-positive, green cells) and dead (propidium iodide, red cells) bacteria were determined by counting at least 100 bacteria.
Single-cell suspensions were prepared with late exponential-phase cultures incubated in 100 ml Sautons medium; the pellicles were harvested by pouring off the medium and were gently shaken for 30 s with 5 g glass beads (4 mm diameter). The declumped cells resulting from this treatment were suspended in 10 ml TS broth plus 0·05% Tween 80 and centrifuged for 10 min 100 g. The OD650 of the supernatant, which contains mainly single cells, was adjusted to 1 OD650 unit and directly used to inoculate the media (100 µl suspension for 50 ml broth, except where indicated otherwise) for the determination of some cell surface properties. Alternatively cells were washed three times with PBS to give a PBS-washed single cell suspension, or stored at -80 °C in the presence of 20% (w/v) glycerol.
MICs were determined by the agar two-fold dilution method in TS agar (bioMérieux). The drug-containing media were inoculated with 100 µl of a dilution (0·01 OD650 unit) of the single cell suspension stored at -80 °C and incubated for 7 days at 37 °C. For all the drugs tested, 99% inhibition of bacterial growth was determined as the MIC of the drug.
Motility.
Surface-spreading assays were adapted from Martinez et al. (1999) ; briefly, Middlebrook 7H9 base medium (Difco) was solidified with 0·4% high grade agarose (Eurogentec). Plates were inoculated into their centre with 10 µl of the 1 OD650 unit PBS-washed single cell suspension (see above). Spreading was evaluated after incubation for 5 days at 37 °C in a humidified incubator by measuring the diameter of the halo of growth formed by the mycobacteria.
Cellular aggregation.
Mycobacterial strains were cultivated with shaking (250 r.p.m.) in TS broth without Tween 80 for 3 days at 37 °C. The bacterial suspensions were centrifuged for 10 min at 100 g to separate the unicellular mycobacteria from the aggregates, which were pelleted (Cougoule et al., 2002 ). The single cells were further recovered by centrifugation for 30 min at 3000 g. Cell pellets were dried and weighed, and the cellular aggregation was calculated, i.e. the percentage of aggregate-containing pellets versus total cell weight.
Congo red accumulation.
The assay (Cangelosi et al., 1999 ) was adapted as follows: mycobacteria were cultivated for 3 days at 37 °C with shaking (250 r.p.m.) in TS broth plus 100 µg Congo red ml-1 and 0·05% Tween 80. Cells were collected by centrifugation (30 min at 3000 g) and washed extensively with distilled water until the supernatant was colourless. Cells were resuspended in 1 ml acetone, vortexed and gently shaken for 2 h at room temperature; cells were then removed by high-speed microcentrifugation and Congo red in the supernatants was spectrophotometrically measured at 488 nm. The Congo red binding was defined as the A488 of the acetone extracts divided by the dry weight (in mg) of the cell pellet.
Hydrophobicity index.
Relative hydrophobicities were assessed by the hexadecane partition procedure (Rosenberg et al., 1980 ). Briefly, 1 OD650 unit PBS-washed single cell suspension (see above) of each strain was mixed with 0·3 ml hexadecane (Avocado) by vortexing for 2 min. The hydrophobicity index was defined as the percentage reduction in the OD650 of the aqueous phase; this reduction was determined for triplicate samples after allowing 15 min for the hydrocarbon phase to rise completely.
Zeta potential.
For determination of the bacterial cell surface charge (Bayer & Sloyer, 1990 ), zeta-potential (
) measurements were performed with 1 OD650 unit PBS-washed single cell suspension (see above) in a zetameter Zetasizer 3000 (Malvern Instruments).
Isolation, fractionation and analysis of the extracellular and surface-exposed components.
Surface-exposed material (SEM) and extracellular compounds were extracted and analysed as previously described (Lemassu et al., 1996 ; Ortalo-Magné et al. 1996
). Briefly, mycobacterial cells were harvested by centrifugation (30 min at 3000 g) and the culture broths were sterilized by filtration through 0·20 µm pore-size sterile filters (Nalgene). Cells were shaken for 1 min with 10 g glass beads (4 mm diameter) per 2 g (wet weight) cells (Ortalo-Magné et al., 1995
), resuspended in distilled water (50 ml per flask) and then the cells were removed by filtration. A portion of the crude filtrate, which contains the SEMs, and of the culture broths, which contains the extracellular materials, were concentrated separately under vacuum, extensively dialysed against distilled water and analysed by colorimetric assays for their carbohydrate content (Dische, 1962
) and protein content by the Lowry method. Chloroform and methanol were added to the remaining portions to obtain partition mixtures composed of chloroform/methanol/water (3:4:3, by vol.); the organic phases were concentrated, washed with water, evaporated to dryness to yield crude lipid extracts, and weighed. The lipid extracts were dissolved in chloroform and analysed by TLC on silica gel Durasil 25-precoated plates (0·25 mm thickness; Macherey-Nagel). The lipids were resolved by TLC run in the following solvent mixtures: petroleum ether/diethyl ether (9:1, v/v) for analysing triacyl glycerols, chloroform/methanol (9:1, v/v) for GPLs and trehalose dimycolates and chloroform/methanol/water (60:30:8 by vol.) for trehalose monomycolates and phospholipids. Sugar-containing compounds (GPLs, trehalose dimycolates, trehalose monomycolates and phosphatidylinositol mannosides) were visualized by spraying the plates with 0·2% anthrone in concentrated sulfuric acid, followed by heating at 110 °C. The DittmerLester reagent (Dittmer & Lester, 1964
) was used to detect phosphorus-containing substances. The ninhydrin reagent was used to reveal the presence of free amino groups and a spray with 10% molybdophosphoric acid in ethanol solution, followed by heating at 110 °C, was used to detect all of the lipid spots, including triacyl glycerols.
Labelling of lipids.
The various classes of extractable and cell surface lipids were quantified by labelling; 1·2 MBq sodium [14C]acetate (Amersham) was added to 100 ml 2-day-old cultures containing mid-exponential-phase bacteria. After 16 h incorporation, the reaction was stopped by centrifugation and the SEMs were isolated by extraction with glass beads (Ortalo-Magné et al., 1995 ). Lipids from these latter materials and those from bead-treated cells were extracted with chloroform/methanol (1:2, v/v). Both types of lipid extracts were analysed by TLC using the solvent mixtures described above; the radioactivity was located and counted on plates using an automatic TLC linear analyser (Berthold LB 2832). Then the lipid spots were visualized by spraying with the appropriate reagents, with heating when necessary.
Permeability assays.
The permeability of the strains of M. smegmatis to chenodeoxycholate was assessed as previously described (Bardou et al., 1998 ; Jackson et al., 1999
). Exponentially grown cells (2-day-old cultures) were first labelled for 16 h with [5,6-3H]uracil (20 µM, 1·85 GBq mmol-1; DuPont NEN) to quantify the biomass present in aliquots used in the accumulation assays. Then, cells were collected by centrifugation and washed with 10 mM HEPES, pH 7·2. Aliquots of labelled cells were counted, dried and weighed to correlate 3H labelling with cell dry weight. Accumulation assays were performed under continuous agitation. [14C]chenodeoxycholate (20 µM, 1·8 GBq mmol-1; DuPont NEN) was added to a 1 ml mixture of HEPES containing about 40 mg 3H-labelled cells. Aliquots (0·1 ml) were removed at different time intervals and added to the top of an Eppendorf centrifuge tube containing 0·25 ml silicon oil/paraffin oil (1:0·2, v/v). Cells were separated from the accumulation medium by centrifugation (13000 g, 1 min). Centrifuged tubes were frozen on dry ice and the pellets were dropped into counting flasks by cutting the cone top. Then the scintillation solution (Aqualuma) was added and the vials were sonicated for 30 min in a water bath to disperse cells.
Transmission electron microscopy.
The method for preparation of samples for transmission electron microscopy was based on the procedures of Paul & Beveridge (1992) . M. smegmatis mc2155 and the GPL-deficient mutant TM99 were grown for 72 h in both the LB broth and Sautons medium, then subcultured into fresh broths by making a 1/100 dilution. The fresh broth was incubated overnight and cells were harvested by centrifugation (8000 g, 10 min), and washed twice in PBS. The resulting pellets were fixed in 2·5% (w/v) glutaraldehyde, 0·05% (w/v) ruthenium red in cacodylate buffer for 2 h in the dark at room temperature. Cells were washed three times in cacodylate buffer (0·1 M, pH 6·8), postfixed for 2 h in the dark in 1% (w/v) osmium tetroxide, 0·05% (w/v) ruthenium red and then washed twice each in cacodylate buffer and in water. Cells were dehydrated through a graded ethanol series of 10, 20, 30, 40, 50, 60, 70, 80 and 95% for 5 min each then washed twice for 15 min each in 100% ethanol, then twice for 15 min each in propylene oxide. Cells were suspended in 1:1 propylene oxide/Spurr resin for 2 h in an open tube. After infiltration overnight, the tubes were opened for 2 h; then samples were transferred to 100% Spurr resin and left overnight. Resin was replenished the next morning and samples were left to cure at 60 °C overnight. Blocks were thin-sectioned on a Reichert-Jung microtome and mounted on copper grids. Sections were poststained with uranyl acetate and Reynolds lead citrate. Microscopy was performed on a JEOL 120 EX electron microscope.
Phagocytosis.
Macrophages were obtained by isolating monocytes from human peripheral blood as previously described by Astarie-Dequeker et al. (1999) ; these monocyte-derived macrophages (MDMs) were cultured at 37 °C in 5% CO2 for 6 to 7 days on sterile glass coverslips in 24-well tissue culture plates (5x105 cells per well) containing RPMI medium supplemented with 10% inactivated foetal calf serum and antibiotic; the culture medium was renewed on the third day. Before use, MDMs were washed twice with fresh serum-free RPMI and equilibrated for 20 min at 37 °C in 5% CO2. Single cells of M. smegmatis were prepared and fluorescently labelled with FITC as previously described (NDiaye et al., 1998
). MDMs were incubated with M. smegmatis in serum-free RPMI, for the time indicated, and then washed twice with fresh medium to remove unbound particles. Phagocytosis of FITC-stained bacteria was determined as previously described (Peyron et al., 2000
). Briefly, MDMs were fixed with 3·7% paraformaldehyde in PBS containing 15 mM sucrose, pH 7·4, for 20 min at room temperature. After neutralization with 50 mM NH4Cl, extracellular mycobacteria were labelled with rabbit polyclonal antibodies directed against mycobacteria (1/50), revealed by TRITC-conjugated secondary antibodies. MDMs containing at least one FITC-stained bacterium (
100 cells) were counted in duplicate samples.
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RESULTS |
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Most prokaryotic cells present a negative surface charge, a parameter that may contribute to the adhesive properties of cells (Van Loosdrecht et al., 1987 ; Bendinger et al., 1993
). Although C-type GPLs of M. smegmatis are neutral lipids (Daffé et al., 1983
), the absence of these molecules may induce a change in the distribution of surface-exposed lipids of the mutant, resulting in a relative abundance of phospholipids already present in the outermost compartment of the envelope (Ortalo-Magné et al., 1996
) whose phosphate groups may influence the surface net charge of cells. Because electrophoresis allows the examination of large populations of microorganisms and has been demonstrated to be the most appropriate method among those experimentally examined for measuring the net charge of bacteria cells (Bayer & Sloyer, 1990
), strain TM99 and its parent strain were compared by this technique. No significant difference was found between the negative zeta-potential displayed by the two strains (Table 1
).
Internalization of the mps-disrupted mutant
M. smegmatis is internalized by MDMs through receptors that include the mannose receptor (Astarie-Dequeker et al., 1999 ). To investigate the consequence of the phenotypic changes induced by the absence of C-type GPLs from the surface of strain TM99 on the nonopsonic phagocytosis of the bacterial cells, the kinetics of internalization of individualized cells from the wild-type and mutant strains were compared (Fig. 3
). A high rate of phagocytosis of both strains of M. smegmatis (70%) was observed when bacterial cells were incubated with MDMs for 90 min. Interestingly, while only 5% of MDMs internalized the wild-type strain in the early 15 min, mutant cells were phagocytosed by 40% of the MDMs during the same period; a significant difference in the uptake rate of individual bacterial cells from the two strains was still observed up to 1 h after the addition of bacteria. After this time period, however, the percentages of MDMs having internalized both types of mycobacterial cells were very similar. This observation indicated that the absence of C-type GPLs from the bacterial cell surface induced a very rapid internalization of M. smegmatis cells by MDMs.
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Composition of the SEM of the mps-disrupted mutant
Even if GPLs are not important for the architecture of the outermost layer, they are assumed to be abundant and known to be surface-located. If so, the absence of GPLs from the surface of mutant cells should induce the exposure of the substances (such as trehalose dimycolates) that are known to be located in deeper compartments of the cell envelope of the parent strain (Ortalo-Magné et al., 1996 ). To address this question, the nature and relative amounts of the SEM of the mutant were determined and compared to those of the parent strain (Table 2
). Gentle shaking of the bacterial cells with glass beads, a technique that declumps aggregates by extracting the amorphous material covering cells as observed by scanning electron microscopy (Ortalo-Magné et al., 1995
, 1996
), was used. The amount of SEM extracted with glass beads from the surface of the mutant was similar to that obtained from the parent strain (15 to 16% of the cell dry weight). The carbohydrate and protein contents of the SEMs from the two strains were comparable. The percentages of carbohydrate in the SEMs of the parent and the mutant strains were estimated to 13·2% and 13·8% of the dry weight, respectively; the sugar compositions of the SEMs from both origins were similar and consisted of arabinose, mannose, xylose and glucose (Lemassu et al., 1996
). Protein represented 59·3% and 75·4% of the dry weight of the SEMs of the parent and mutant strains, respectively. Surface lipids represent a very minor fraction (0·020·1% bacterial dry weight) of mycobacteria grown on synthetic Sautons medium (Lemassu et al., 1996
; Ortalo-Magné et al., 1996
). Consequently, bacterial cells of the two strains grown to exponential phase cells were labelled with sodium [14C]acetate before extracting SEMs to easily quantify surface-exposed lipids. In contrast, larger amounts of lipids were extracted from the SEMs of both strains grown on LB medium. They represented 3·4% and 2·3% bacterial dry weight of the parent and mutant strains, respectively. C-type GPLs represented 85% of the radioactivity found in the surface-exposed lipids from the wild-type strain grown on both media and were not detected in lipids from strain TM99 (Table 2
), as expected. Lipids extracted from the parent and mutant strains after bead treatment represented 10·5 and 11·2% of the cell dry weight, respectively. The lipid composition of the materials originated from both strains was similar, except for the absence of C-type GPLs, and consisted of triacyl glycerols, trehalose monomycolates, trehalose dimycolates and phospholipids. Importantly, it has to be noted that trehalose dimycolates, which were not detected in the surface-exposed lipids from the parent strain, were also absent from the SEM of strain TM99. This data indicated that the absence of GPLs from the mutant strain does not induce the exposure of lipids that are buried in deeper compartments of the cell envelope.
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Ultrastructural features of the mps-inactivated strain TM99
Cells from M. smegmatis strain mc2155 and its isogenic TM99 mutant were grown in both LB and Sautons media and examined by electron microscopy (Fig. 4). Ultrathin sections of cells were stained with ruthenium red, a stain that strongly reacts with the surface of mycobacteria (Rastogi et al., 1984
). Examination of thin sections revealed a cell envelope structure of M. smegmatis strain mc2155 (Fig. 4A
) composed of (i) a plasma membrane, (ii) a thick internal electron-dense layer, (iii) an electron-transparent layer and (iv) a thick electron-dense outer layer (OL); the space observed between the plasma membrane and the thick electron-dense layer corresponds to the hypothetical periplasmic space (Daffé & Draper, 1998
). This ultrastructural appearance is similar to that previously found in other mycobacterial species (Daffé & Draper, 1998
; Draper, 1982
; Paul & Beveridge, 1992
; Rastogi et al., 1986
). In sharp contrast, the thick electron-dense OL stained with ruthenium red and observed on the thin sections of the parent M. smegmatis strain mc2155 (Fig. 4A
) was unlabelled in those of the cells from the isogenic mps-inactivated strain TM99 (Fig. 4B
). However, the thickness of the electron-transparent layer of the mutant roughly corresponded to the sum of the thickness of the electron-transparent layer and that of the OL of the parent strain. These data suggested that the absence of staining of the OL of the mutant strain was not due to the loss of this layer but, rather, may be attributed to absence of GPLs from the surface of the mutant cells which would bind ruthenium red. This conclusion was further supported by the observation of the thickness of this layer in adjacent cells (Fig. 4C
). That the staining of OL in the parent strain by ruthenium red is attributable to GPLs is supported by the correlation between the ultrastructural appearance of stained cells and the amounts of surface-exposed GPLs. When large amounts of GPLs are present on the cell surface of the parent strain (growth in the LB medium, see above), the whole surface of the bacilli is stained by the dye that also interacts with GPLs budding from the cell surface and with an appearance of filamentous and rope-like structures (Fig. 4D
). When cells exposed less amounts of GPLs on their surface (growth in Sautons medium, see above) they are stained less by ruthenium red (Fig. 4E
). These data establish that the dye binds to the OL through interactions with surface-exposed GPLs.
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DISCUSSION |
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In M. smegmatis, the phenotypic changes were associated with the absence of binding of ruthenium red to the surface of the mutant cells, as observed by transmission electron microscopy. This data implies that the surface-exposed C-type GPLs of M. smegmatis are the substances that react with ruthenium red to give the electron-dense appearance of the outermost cell envelope layer. The absence of these compounds, however, does not cause the disorganization of the outer layer since no specific release of surface constituents into the culture broth of the mutant was observed. Consistent with this observation, compounds such as trehalose dimycolates that are located in deeper compartments of the cell envelope of the parent strain (Ortalo-Magné et al., 1996 ) were not surface-exposed in the mutant. These data reinforce the credibility of the recent model (Daffé & Draper, 1998
) in which the surface of mycobacteria is composed of polysaccharides, proteins and lipids, as opposed to models in which the surface consists of a lipid layer (Brennan & Nikaido, 1995
; Liu et al., 1995
; Minnikin, 1982
). Nevertheless, at the interface between the bacterial and host cells, surface-exposed GPLs may play a role in the internalization of C-type GPL-containing non-tuberculous mycobacteria by macrophages which are believed to use glycoconjugates as ligands of many of their receptors (Ehlers & Daffé, 1998
). Accordingly, phagocytosis of the wild-type strain and its isogenic mps-inactivated strain of M. smegmatis by human MDMs was investigated. As expected from the existence of numerous receptors capable of internalizing mycobacteria, both strains were found in phagosomes but the kinetics of phagocytosis of the mutant was much more rapid than that of the parent strain. This result suggests that either GPLs may mask some as yet undefined ligands that interact with highly efficient macrophage receptor(s) or, alternatively, the bacterial cell surface hydrophobicity may play an important role in the rate of internalization of bacteria. Further studies are warranted to determine the precise role of GPLs in the internalization of mycobacteria.
GPLs are the major extractable lipids of M. smegmatis and other non-tuberculous mycobacterial species containing these molecules and a large portion of these substances are localized in the deeper compartments of the wild-type M. smegmatis envelope, presumably in the mycolic acid-containing asymmetric bilayer. It was therefore important to investigate the consequence of the absence of GPLs on cell wall architecture, especially its implication in the outer permeability barrier. In all currently proposed models (Brennan & Nikaido, 1995 ; Daffé & Draper, 1998
; Liu et al., 1995
; Minnikin, 1982
; Rastogi, 1991
) 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 (Barksdale & Kim, 1977
; Benedetti et al., 1984
; 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 cell wall-linked mycolates certainly participate in this barrier since the disruption of a gene that encodes a mycoloyltransferase, namely antigen-85C, causes a decrease in the amount of cell wall-bound mycolates and affects the permeability of the envelope of the mutant (Jackson et al., 1999
). 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
). To date, however, the implication of non-covalently bound lipids in the wall bilayer has been demonstrated for only phthiocerol dimycocerosates of M. tuberculosis (Camacho et al., 2001
). The present work demonstrated that the absence of C-type GPLs from the cell envelope of M. smegmatis has a profound effect on the uptake of chenodeoxycholate, a hydrophobic molecule that diffuses through lipid domains of the mycobacterial cell wall (Liu et al., 1996
; Yuan et al., 1997
). Based on the localization of cell wall fracture plane in mycobacteria (Barksdale & Kim, 1977
; Benedetti et al., 1984
; Takeo et al., 1984
), far from the cell surface, it is likely that the GPLs located in inner compartments of the cell envelope, but not surface-exposed GPLs, contribute to the permeability barrier.
In conclusion, in addition to be distinctive markers of numerous mycobacterial species that may help in the diagnosis of non-tuberculous diseases and for taxonomic purposes, GPLs are also involved in the wall permeability barrier that certainly plays a role in bacterial physiology.
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ACKNOWLEDGEMENTS |
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Received 19 March 2002;
revised 3 June 2002;
accepted 6 June 2002.