Advertisement
J. Lipid Res.
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1194/jlr.M800384-JLR200 on September 4, 2008

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
M800384-JLR200v1
50/3/477    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rafidinarivo, E.
Right arrow Articles by Daffé, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rafidinarivo, E.
Right arrow Articles by Daffé, M.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Journal of Lipid Research, Vol. 50, 477-490, March 2009
Copyright © 2009 by American Society for Biochemistry and Molecular Biology

Trafficking pathways of mycolic acids: structures, origin, mechanism of formation, and storage form of mycobacteric acids

Elie Rafidinarivo2,*,{dagger}, Marie-Antoinette Lanéelle*,{dagger}, Henri Montrozier*,{dagger}, Pedro Valero-Guillén§, José Astola**, Marina Luquin**, Jean-Claude Promé*,{dagger} and Mamadou Daffé1,*,§

* Université Paul Sabatier (Toulouse III), Institut de Pharmacologie et Biologie Structurale (IPBS), 205 route de Narbonne, 31077 Toulouse Cedex, France
{dagger} Centre National de la Recherche Scientifique (Unité Mixte de Recherche 5089), IPBS, Department ‘Mécanismes Moléculaires des Infections Mycobactériennes,’ 205 route de Narbonne, 31077 Toulouse Cedex, France
§ Departamento de Genética y Microbiologia, Facultad de Medicina y Odontologia, Universidad de Murcia, 30100 Murcia, Spain
** Departament de Genètica y Microbiologia, Facultat de Ciences, Universitat Autonoma de Barcelona, 08193 Bellaterra, Spain
2 Present address of E. Rafidinarivo: Ecole Normale Niveau III, B.P. 881, 101 Antananarivo, Madagascar.

The NMR equipment was funded via European structural funds, CNRS, and the Région Midi-Pyrénées funds as part of the 2000–2006 CPER program.

Published, JLR Papers in Press, September 4, 2008.

1 To whom correspondence should be addressed. e-mail: mamadou.daffe{at}ipbs.fr


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Mycolic acids, the hallmark of mycobacteria and related bacteria, are major and specific components of their cell envelope and essential for the mycobacterial survival. Mycobacteria contain structurally related long-chain lipids, but the metabolic relationships between these various classes of compounds remain obscure. To address this question a series of C35 to C54 nonhydroxylated fatty acids (mycobacteric acids), ketones, and alcohols structurally related to the C70–80 dicyclopropanated or diethylenic mycolic acids were characterized in three mycobacterial strains and their structures compared. The relationships between these long-chain acids and mycolic acids were established by following the in vivo traffic of 14C labeled {alpha}-mycolic acids purified from the same mycobacterial species. The labeling was exclusively found in mycobacteric acids. The mechanism of this degradation was established by incorporation of 18O2 into long-chain lipids and shown to consist in the rupture of mycolic acids between carbon 3 and 4 by a Baeyer-Villiger-like reaction. We also demonstrated that mycobacteric acids occur exclusively in the triacylglycerol (TAG) fraction where one molecule of these acids esterifies one of the three hydroxyl groups of glycerol. Altogether, these data suggest that these compounds represent a pathway of metabolic energy that would be used by mycobacteria in particular circumstances.

Supplementary key words mycobacteria • biosynthesis • catabolism • Baeyer-Villiger reaction • oxidation

Abbreviations: amu, atomic mass unit; BVMO, Baeyer-Villiger monooxygenases; EI, electron impact; MALDI-TOF, MALDI-time-of-flight; MS, mass spectrometry; TAG, triacylglycerol


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Mycolic acids, 2-alkyl-branched, 3-hydroxylated long-chain fatty acids, represent major (up to 40% of cell wall dry mass) and specific constituents of the envelope of members of the genus Mycobacterium (M.) that includes several pathogens such as M. tuberculosis and M. leprae, the causative agents of tuberculosis and leprosy, respectively. These fatty acids play a crucial structural role in the envelope architecture (1–4) and are essential for the mycobacterial growth (5–7). Mycolic acids covalently linked to the cell wall arabinogalactan are organized with other lipids to form an outer barrier with an extremely low fluidity that confers an exceptional low permeability to mycobacteria (8). Trehalose mycolates, found noncovalently attached to the cell wall core, are known to be important for the physiology and virulence of M. tuberculosis (3). In addition, the biosynthesis of mycolic acids is the target of several antituberculous drugs, notably isoniazid (2, 3, 9).

Mycolic acids are found in mycobacterial species as a mixture of structurally related molecules that differ from one another by the nature of the chemical groups at the so-called "proximal" and "distal" positions (relative to the carboxyl group) of their main "meromycolic" chain (Fig. 1 ). The least polar {alpha}-mycolates is ubiquitous in mycobacteria and consist of C74–82 fatty acids that contain two cis cyclopropyl groups or cis/trans double bonds at the proximal and distal positions (Fig. 1); shorter apolar structural analogs (C60–68), called {alpha}'-mycolates, may occur in some rapid-growing species such as M. smegmatis. More polar mycolates are also produced by most mycobacteria. These contain an oxygenated function at the distal position, with a methyl group on the vicinal carbon atom, and a cis or trans (with a methyl group on the vicinal carbon atom) cyclopropyl group or double bond at the proximal position. Depending on the mycobacterial species examined, the oxygenated function may be a methoxy-, a keto-, a hydroxyl-group (in slow-growing species), or an epoxy-ring (in rapid-growing species); in some species, the keto-mycolic acids may be oxidized through a Baeyer-Villiger reaction to yield a wax ester (2, 10). An illustration of the structural diversity of mycolic acids is given by the composition of mycolic acids of the strains under study (i.e., M. brumae, M. aurum, and M. fallax) (Fig. 1).


Figure 1
View larger version (15K):
[in this window]
[in a new window]

  		Fig. 1. Structures of the major mycolic acids from Mycobacterium (M.) brumae and M. fallax (A) and M. aurum (B). M. brumae contains four different types of diethylenic mycolates, namely M0 (x = 15; y = 12; z = 17; l = 19), M1a (x = 15; y = 13; z = 17, 19; l = 19), M1b (x = 15; y = 13; z = 15, 17, 19; l = 19), and M2 (x = 13, 15; y = 10, 12, 14; z = 15, 17, 19; l = 19) (28). M. fallax also contains four main types of diethylenic mycolates; however, this strain contains M1a (x = 15, 17; y = 15, 17; z = 15, 17, 19; l = 19, 21), M1b (x = 15, 17; y = 15, 17; z = 17; l = 19, 21), and M2 (x = 15,17; y = 14, 16, 18; z = 15, 17, 19; l = 19, 21), and is devoid of M0 but contains M3 (x = 13,15; u = 4; v = 12, 14; z = 15, 17, 19; l = 19, 21). In addition, small amounts of tetra-ethylenic homologs occur in this species (31). M. aurum contains {alpha}- (x = 13, 15, 17; y = 12, 14, 16; z = 17, 19; l = 19, 21), keto- and wax-mycolates (x = 13, 15, 17; y + z = 27, 31, 33; l = 19, 21). cis (E) and trans (Z) are relative to the configuration of the double bonds. "Odd" and "even" refer to the chain lengths of the mycolates.

 
Long-chain compounds structurally related to mycolic acids are also found in more or less significant amounts in mycobacteria and related genera (e.g., Nocardia, Corynebacterium); these include long-chain acids, ketones, and alcohols whose general structures are given in Table 1 . Long-chain C40–56 fatty acids that exhibit the structural features of the main chains of mycolic acids have been characterized in various mycobacteria and are known as mycobacteric acids (11–15). Their structures have been shown to be related to those of mycolic acids from the same species. For instance, keto and methoxyl groups were shown to occur in mycobacteric acids and mycolic acids isolated from M. tuberculosis (11); similarly, keto and wax functions were found in both types of fatty acids isolated from M. aurum (16). A series of shorter homologous fatty acids have been also isolated from triacylglycerols (TAGs) of a Nocardia species; these consist in C35–45 fatty acids containing 3 and 4 double bonds as observed in nocardomycolic acids (17). In addition, long-chain ketones and secondary alcohols parented to mycolic acids have been also found in mycobacteria (e.g., the so-called C80 mycolones and mycolols) (18, 19), in nocardia (C50–60 nocardones, nocardols) (20–23), and in corynebacteria (16-hentriacontanone or palmitone) (24, 25). Interestingly, shorter-chain secondary alcohols and ketones (hentriacontan-16-ol and palmitone) have also been characterized in nocardia (26, 27). Because of the above-mentioned similar structural features, it is tempting to postulate biosynthetic or catabolic links between mycolic acids and the other long-chain compounds. To address this question, mycolic acids and their related lipids from M. brumae and M. fallax, two species that contain only one type of mycolates, the so called {alpha}-mycolates (Fig. 1) (28, 29), were isolated, purified, and comparatively analyzed by various complementary analytical methods. In addition, the in vivo trafficking of labeled mycolates in M. aurum, a species known to metabolize lipids, was investigated to validate the conclusions drawn from the structural findings. Importantly, because the elongation unit in the biosynthesis of mycolates is acetate (C2), the major homologs contain an even number carbon atoms. The introduction of a methyl branch or a cyclopropyl group results in the production of mycolates possessing an odd number of carbon atoms whereas that of two cyclopropyl groups or two methyl branches does not change the parity. The number of double bonds in mycolates, usually one in oxygenated compounds and two in {alpha}-mycolates with the notable exception of M. fallax, does not impact on the even carbon number of the chains (Fig. 1). Therefore, the even versus odd ratio of the chain lengths of the structurally related lipids was determinant in the definition of the biosynthetic relationships between the long-chain lipids and the mechanism that governs their catabolism.


View this table:
[in this window]
[in a new window]

  		TABLE 1. General formulae of long-chain lipids structurally related to mycolic acids and comparison of odd/even ratios of their chain lengths in Mycobacterium (M.) fallax

 

Figure 2
View larger version (18K):
[in this window]
[in a new window]

  		Fig. 2. Trafficking in mycolic acids pathways. A: Biosynthesis of mycolic acids. The long-chain acid at the origin of the meromycolic chain (R1-CH2-COOH with an even number of carbon atoms) is first activated (X), in the present case as an acyl AMP derivative. The R2-CO-X (X = CoA) with 20, 22, or 24 carbon atoms at the origin of the {alpha}-branch chain in mycolates, is carboxylated as alkylmalonate. The condensation between the two activated derivatives gives the β-keto ester, the reduction of which leads to mature mycolic acids (with an even number of carbon atoms) that exist either as esters (X = trehalose or cell arabinogalactan). B: Degradation of mycolic acids. The β-keto ester intermediate in the synthesis of mycolates (with an even number of carbon atoms) and/or derived from the oxidation of mycolates would be decarboxylated (either as a consequence of the chemical hydrolysis of lipids or by a decarboxylase) to yield mycolones (with an odd number of carbon atoms). The β-keto esters, and possibly mycolones would be then oxidized, through the addition of an oxygen atom near the keto group by a putative Baeyer-Villiger monoxygenase, to yield a wax. The cleavage of the ester linkage would give primary alcohols (with an odd number of carbon atoms) whose oxidation would be at the origin of the C44–56 and their probably β-oxidation C35–44 series of mycobacteric acids. These latter substances were isolated both as free fatty acids and triacyl glycerol containing one mycobacteroyl residue (R1-CO) and two moles of conventional (C16–18) fatty acyl substituents (R3-CO). The positions of the fatty acyl residues in the triacyl glycerol are arbitrary. Bold character O labels correspond to 18O incorporated in lipids. "Even" and "odd" refer to the number of carbon atoms of lipid chains.

 

   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Strains and media
The type strain of M. brumae Collection Institut Pasteur (CIP) 103465T (28) was grown in Sauton's medium as surface pellicles at 37°C for 3 weeks. That of M. fallax CIP 8139T (29) was grown in Middlebrook 7H9 medium supplemented with 0.5% casitone and 1% glucose (Difco, Detroit, MI) at 30°C under agitation. M. aurum strain A+ was obtained from Pr H. David (Institut Pasteur, Paris) and was grown on Middlebrook 7H9 medium (Difco) supplemented with 0.5% casitone and glucose 1% at 37°C under agitation.

Lipid extraction and purification from M. brumae
Complex lipids were extracted from wet cells with CHCl3/CH3OH 1:2, 1:1, and 2:1 (v/v), successively. The crude lipid extracts were pooled and then fractionated by column chromatography on Florisil 100–200 mesh equilibrated in petroleum ether, using a gradient of diethyl ether in petroleum ether as eluent. TAGs were eluted with petroleum ether/ether 90:10 (v/v). Free fatty acids were eluted from the column with diethyl ether containing 0.1% acetic acid. The separation was monitored by TLC and infrared spectroscopy for the detection of trans double bonds at 965 cm–1.

Saponification of complex lipids
Lipids were saponified using a 5% KOH solution in methanol/benzene 8:2 (v/v) for 7 h under reflux. After acidification of the mixture and extraction of lipids with diethyl ether, the organic phase was dried and treated with diazomethane. The resulting long-chain lipids (i.e., ketones, esters, and alcohols) were fractionated by column chromatography on Florisil as previously described.

Isolation of lipids from M. fallax
Cells (17.5 g dry weight obtained from 10l of cultures) were extracted with CHCl3/CH3OH as previously indicated to yield 3.1 g crude lipid extract. After saponification, acidification, and methylation, lipids were fractionated on a Florisil column as previously described, and the separation was monitored by TLC. Mycolones (20 mg), mycobacteric esters (20 mg), mycolols (20 mg), and long-chain primary alcohols (10 mg) were obtained, in addition to {alpha}-mycolates (1.7 g) and fatty esters that were the main lipids of the mixture.

TLC
TLC was performed on silica gel coated plates (G-60 0.25 mm thickness, Merck, Darmstadt, Germany) developed with either CH2Cl2 or with an appropriate combination of petroleum ether/diethyl ether. Lipids (fatty esters, ketones alcohols, and TAGs) were located by spraying either Rhodamine B (0.01% in sodium dihydrogen phosphate 0.25 M) or molybdophosphoric acid (10% solution in ethanol), followed by charring. Glycolipids were detected with anthrone after elution with CHCl3/CH3OH/H2O 65:25/4 (v/v/v). For preparative TLC, the same solvents were used but the detection was obtained by spraying Rhodamine B and the visualization was performed under the UV light. The compounds were then scraped off the plates and extracted with diethyl ether and the ethereal solution was evaporated to dryness. The stain was eliminated by passing through a Florisil Pasteur pipette (1 cm length) that was irrigated with diethyl ether. Fatty acid methyl esters were further fractionated according to their double bond content on AgNO3-impregnated silica gel TLC plates developed with CH2Cl2; visualization was performed by spraying a 0.2% ethanolic dichlorofluorescein solution and the compounds were scraped off the plates and eluted with diethyl ether.

Oxidative cleavage of double bonds
Ethylenic lipids were cleaved by permanganate-periodate oxidation (30, 31). Briefly, lipids (around 20 mg) dissolved in benzene (0.6 ml) were added to a mixture of t-butanol (8 ml), aqueous 2.10–2 M sodium carbonate (2.4 ml), distilled water (1.2 ml) and periodate permanganate (5.4 ml). This last solution contained 0.1 M sodium periodate and potassium permanganate 2.5 10–2 M in water. The oxidation was carried out at 30°C for 2 days with constant shaking. The reaction was stopped by additing sodium metabisulphite until discoloration. The solution was acidified with 20% sulfuric acid, and the resulting oxidation products were extracted with diethyl ether and the solvent evaporated. The crude extract was methylated with diazomethane and purified on preparative TLC with CH2Cl2 as developing solvent. Compounds were identified by GC-MS analysis according to their retention times and their fragmentation patterns (32).

Preparation of anilide derivatives of mycobacteric acids
Anilide derivatives were prepared as follows: the carbonyl group was first activated by adding 0.5 ml of a 20% solution of oxalyl chloride (Fluka) in anhydrous benzene to fatty acids (5 mg). The reaction was left at room temperature for 2 h, followed by heating at 50°C for 1 h. After evaporation of the solvent under nitrogen, 0.35 ml of a 20% solution of aniline (Aldrich) in anhydrous benzene (20-fold excess) was added to the acyl chloride. After 1 h at room temperature, the reaction mixture was heated at 70°C for 30 min. The solvent was evaporated, and the residue was dissolved in diethyl ether and the excess of aniline was removed by washing the diethyl ether phase with 1 M HCl. Anilides were analyzed by TLC with hexane/dioxane 9:1 (v/v) as the running solvent and detected with Rhodamine B before high-pressure liquid chromatography.

Preparation of pyrrolidide derivatives of mycobacteric acids
Fatty acid methyl esters (1 mg) were dissolved in 1 ml of freshly distilled pyrrolidine (Aldrich chemicals Co., WI) and 0.1ml acetic acid. The mixture was heated at 100°C for 30 min and cooled to room temperature. The amide so-formed was taken up in CHCl3, and the organic phase was washed with 1 M HCl and water (33). After evaporation, the purity of the derivatives was checked by TLC.

Pyrolysis of mycolic acid methyl esters
The pyrolytic reaction was realized at 300°C under vacuum and the oxidation of the yielded meroaldehydes was performed as previously described (34). The resulting meromycolic acids were methylated and purified by preparative TLC as described for long-chain lipids.

Catalytic hydrogenation of double bonds in long-chain alcohols
Around 5 mg of lipids were dissolved in 1ml of hexane in the presence of 2 mg of platinum oxide. Hydrogenation was carried out under shaking for 2 h at room temperature. The mixture was then filtered off to remove the catalyst. The solvent was evaporated under vacuum. Electron impact (EI)-MS analysis showed that the hydrogenation was complete in these conditions.

Incorporation of 14C acetate in long-chain lipids of M. brumae
To a growing culture of M. brumae, 1 µCi ml–1 of 1-14C acetate (2GBq mmole–1) were added. After 90 min, the medium was discarded and 30 ml of fresh Sauton's medium was added. At various time intervals (0, 24, 48, 72, 96, and 120 h) a loopful of cells was collected and saponified. Fatty acids were methylated and the profile of long-chain compounds (alcohols, ketones, and fatty esters) was analyzed by TLC with CH2Cl2 as developing solvent. The TLC plates were read using a PhosphoImager after a 48-h exposure.

Preparation of labeled 1-14C mycolic acids of M. aurum
One µCi ml–1 of [1,2-14C] sodium acetate (50–60 mCi mmol–1) was added to a 72-h grown 100-ml culture. After a 90-min growth under shaking, bacteria were harvested by centrifugation and saponified.

Biotransformation of 14C-labeled mycolic acids by M. aurum
An ethereal solution of 14C-labeled mycolic acids (0.5 ml, specific radioactivity of 4.106 cpm mg–1) was added to a 100 ml culture medium. Bacteria were then inoculated as a 1% suspension after the removal of diethyl ether and collected by centrifugation after a 72-h growth.

Growth under 18O atmosphere
A 4-day culture of M. fallax was added to 250 ml medium in a vacuum flask to obtain a 1% cell suspension. Two vacuum 500 ml flasks were used in parallel: one for the assay and the second for the control. Each flask was filled with 250 ml culture medium and inoculated to obtain a 1% cell suspension. After a 5-day growth, the air was partly removed from each flask under a slight vacuum. Approximately 150 ml of 18O2 (99% purity) was introduced in the assay flask and the same amount of 16O2 was filled in the control flask. Both flasks were hermetically closed and the cultures were allowed to grow at 30°C for 48 h. Bacteria were then collected by centrifugation and saponified. In order to estimate the amount of 18O uptake, the relative abundances between m/z 803 and 805 were more precisely measured by multichannel detection.

Instrumentation
Infrared spectra were recorded using a Perkin-Elmer model FTIR 1600 apparatus. Samples were analyzed as a film between two NaCl discs.

The optical rotations of purified molecules at 589 nm ([{alpha}]D) were determined with a Perkin-Elmer model 141 polarimeter. Samples were dissolved in CHCl3 (5 mg ml–1).

Nuclear magnetic resonance (NMR) spectra of the purified samples dissolved in CDCl3 (100% D) were recorded using a Brucker Avance 600 MHz spectrometer equipped with a 5 mm TCi cryoprobe at 298 K. Chemical shifts values were relative to the internal CHCl3 resonance (at 7.27 ppm).

High-pressure liquid chromatography was carried out on a Waters 6000A apparatus equipped with a µ Porasil (0.39 x 30 cm) column for adsorption chromatography; the elution was performed with hexane/dioxane 9/1 (v/v). The flow rate was 0.5 ml min–1, and the detection was followed using an UV detector Waters 450 at 242 nm for chromophoric derivatives. All solvents were degassed by filtration under vacuum.

GC-MS was performed on a Hewlett-Packard 5989A mass selective detector connected to a Hewlett-Packard 5890 gas chromatograph. The temperature of the direct inlet probe was set at 270°C, the source temperature at 300°C, and the electron energy was 70 eV.

EI-MS was performed using a VG AutoSpec (Fison) instrument fitted with an EI ion source. The accelerating voltage was 8 kV, the electron energy was 70 eV, the source temperature was set at 250°C. The temperature of the solid probe was ramped from 100°C to 300°C in 10 min (35). The same instrument (provided with a caesium gun, 25 kV) was employed for fast-atom bombardment-MS in positive and negative modes; the accelerating voltage was 8 kV and the matrix used was m-nitrobenzyl alcohol.

MALDI-time of flight (MALDI-TOF) mass spectrometry (MS) was performed on a 4700 Analyzer mass spectrometer (Applied Biosystems, Framingham, MA) equipped with a Nd:YAG laser YAG (355 nm wave length) operating by pulses of 500 ps with a frequency of 200 Hz. Twenty-five hundred shots were accumulated in positive ion mode, and MS data were acquired using the instrument default calibration. Lipid samples were dissolved in CHCl3 at a concentration of 1 mM, and were directly spotted onto the target plate as 0.5 µl droplets, followed by the addition of 0.5 µl of 2-5-dihydroxybenzoic acid (10 mg ml–1) in CHCl3/CH3OH (1:1, v/v) (36). Samples were allowed to crystallize at room temperature.

The radioactivity was located using an automatic TLC linear analyzer Berthold LB 2832 or a PhosphoImager.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Mycobacteria contain long-chain lipids whose structures are related to those of mycolic acids. They consist on long-chain fatty acids, primary and secondary alcohols, and ketones (Fig. 2 ) (Table 1). As they are present in small amounts, usually less than 1% of saponified lipids, their detection on TLC is not easy. Accordingly and in order to visualize them and to follow their relationships with mycolic acids, exponentially grown cells of mycobacterial species were labeled with 14C-acetate for 90 min. The medium was then discarded and replaced by a fresh medium and the labeling was followed for various times (0 h, 24 h, 48 h, 72 h, 96 h, and 10 days) by taking off a loopfull of bacteria. After saponification of cells, the resulting lipid extracts were methylated and analyzed by TLC using CH2Cl2 as developing solvent. Besides mycolates (compounds e, Fig. 3 ) and C16–24 fatty esters (compounds c), which are the expected major constituents of the mixtures, mycobacteria accumulate substantiate amounts of a variety of long-chain compounds (Fig. 3). Compound b, which corresponds to long-chain fatty esters, was also detected after methylation of M. brumae native lipids extracted from cells by organic solvents, indicating that these substances exist also as free acids.


Figure 3
View larger version (48K):
[in this window]
[in a new window]

  		Fig. 3. Radio-TLC of the saponification products from M. brumae. Cells were labeled with 14C acetate for 90 min, then the medium was discarded and a fresh Sauton's medium was added. At various time points (i.e., 0 h, 24 h, 48 h, 72 h, 96 h, and 10 days (lanes 1, 2, 3, 4, 5, and 6, respectively) aliquots were taken up and saponified. Lipids were extracted, methylated, and analyzed by TLC using dichloromethane as eluent. (a) mycolones, (b) mycobacteric acid methyl esters, (c) C16–24 fatty acid methyl esters, (d) mycolols, (e) methyl mycolates. (f) indicates the Rf of primary long-chain alcohols that are not detected in significant amounts in M. brumae but present in M. fallax. See Table 1 for general formula of the compounds. The arrow indicates the solvent front.

 
Structural features of the long-chain fatty acids from M. brumae
To determine the structure of compounds b, free fatty acids were isolated by column chromatography of the extractable lipids from M. brumae. Following methylation with diazomethane, the main component of this fraction exhibited a mobility higher than that of nonhydroxylated fatty acid methyl esters (compound c) as observed for compound b on TLC using CH2Cl2 as developing solvent. Infrared spectroscopy of the methyl ester derivatives showed the characteristic bands of nonhydroxylated fatty esters: absorption bands assigned to C-H stretching at 2,921 and 2,845 cm–1, to C = O stretching of carboxylic esters at 1,745 cm–1 and to long-chain deformations at 722 cm–1. The presence of an intense band at 965 cm–1 was indicative of the occurrence of trans double bond(s) and suggestive of a methyl branch at their allylic position and, as a consequence, of negative optical rotation values, as found in mycolic acids (37). Consistently, a negative molecular rotation ([M]D = –42°) was measured for the purified ester.

1H-NMR analysis of the methyl esters of the purified free fatty acids from M. brumae (compound b) showed the occurrence of characteristic signal resonances of long-chain fatty acids (Table 2 ); these include signals attributable to terminal CH3- (at 0.89 ppm, triplet, 3H), to -CH2- (at 1.28 ppm, broad signal), to -CH2-CH2-COO- (at 1.65 ppm, multiplet), and to -CH2-COO- (at 2.38 ppm, multiplet). More specific resonances were observed at 5.35 ppm (assigned to ethylenic protons, two multiplets, 2H), at 2.04 ppm (-CH - CH = CH-) and 0.96 and 0.99 ppm (attributed to CH3- adjacent to double bonds). Successive uncouplings of vicinal CH protons allowed the identification of direct neighboring groups, whereas the analysis of the 1H homonuclear 2D correlated spectroscopy spectrum further extended the attributions (data not shown). For instance, a cross-peak was observed between the proton resonance at 5.35 ppm (CH = CH-) and that at 2.04 ppm (-CH-CH = CH-); the latter resonance correlated with and that at 0.99 ppm (CH3- adjacent to double bonds). This allowed the location of a methyl branch at the {alpha}-position of a trans double bond. The main data deduced from the analysis of the 1H-NMR and 13C-NMR spectra are consigned in Table 2 and consistent with the compounds being a mixture of long-chain ethylenic fatty esters with methyl branch(es) adjacent to trans double bond(s), as observed in the merochains of mycolic acids (28).


View this table:
[in this window]
[in a new window]

  		TABLE 2. Assignments of the main resonance signals of the NMR spectra of the mycobacteric acids from M. brumae

 
Analyses of the fatty acids, as free acids and as methyl ester derivatives by MALDI-TOF MS were unsuccessful. Molecular mass data were obtained by fast-atom bombardment and EI-MS of the free acids and were consistent with the occurrence of a family of C37–45 fatty acids of odd and even chain lengths (i.e., mycobacteric acids); the mass peaks of the major homologs were seen at 546, 560, 574 m/z and corresponded to fatty acids containing C37, C38, C39 with two unsaturations.

Localization of the double bonds in mycobacteric acids of M. brumae
Fatty acids methyl esters were submitted to oxidative cleavage and the resulting fragments, monocarboxylic and dicarboxylic acids, were methylated, purified by preparative TLC, and analyzed by GC-MS. The major monocarboxylic methyl ester was identified as a 2-methyl-octadecanoate (molecular mass peak at 312 m/z and base peak at 88 m/z); minor linear and branched homologs containing 15, 17, 19, and 21 carbon atoms were also found. This result allowed us to localize the double bond at the {omega}-18 position of the molecules, as found in M. brumae mycolic acids (Fig. 1A) and in agreement with the occurrence of a methyl branch at {omega}-17 position and adjacent to a (trans) double bond, as revealed by NMR and infrared spectroscopies.

The dicarboxylic methyl esters arising from the oxidation of the double bonds consisted of a complex mixture of linear molecules with 12, 13, and 14 carbons and compounds with shorter chain lengths that contained branched and linear molecules. As these substances corresponded to the chemical entities between the two double bonds and those located between the proximal double bond and the carboxyl group, it was not possible to precisely assign the identified dicarboxylic acids to one of the two classes. To circumvent this problem and to confirm the molecular masses of the dicarboxylic acids, the native purified fatty acid mixture was derivatized as pyrrolidides and analyzed by EI-MS (33). A regular series of peaks with 14 atomic mass units (amu) interval corresponded to the primary fragmentation of the polymethylenic chain. All ions retained the pyrrolidide group. Interestingly, when a double bond is present in the molecule, a gap of 12 amu was observed between the two consecutive mass peaks. The position of the double bond can be deduced from a rule defined for mono- and polyenoic acid pyrrolidides (33).

The mass spectrum of M. brumae pyrrolidide derivatives of mycobacteric acids exhibited molecular ion peaks at 585, 599, 613, 627, and 641 m/z, which corresponded to the molecular masses of diethylenic C36 to C41 compounds (Table 3 ); the major mass peak was seen at 627 m/z and corresponded to a C39 diunsaturated pyrrolidide. However, due to the presence of carbon chain homologs and the possible presence of double bond isomers possessing or not methyl branch, no clear conclusion could be drawn from the analysis of the mixture. Consequently, the pyrrolidide derivatives were fractionated by argentation TLC, a technique that separates substances according to the number and configuration of the double bonds (28, 31), isolated and analyzed as above. The EI mass spectrum of the major purified compound showed an intense molecular ion at 627 m/z, the expected value for the C39:2 pyrrolidide, with a base peak at 113 m/z that is originated from a McLafferty rearrangement (Fig. 4A ). The presence of an intense peak at 402 m/z, attributable to the fragment bearing the methyl branch, indicated the occurrence of a methyl branch at the adjacent position of a (trans) double bond. Oxidative cleavage of this fraction, followed by methylation, yielded three compounds (Fig. 4A). GC-MS analysis of these molecules led to the identification of methyl esters of: i) 2-methyl octadecanoate (molecular ion at 312 m/z and base peak at 88 m/z); ii) tetradecane-1,14- dioate (base peak at 98 m/z, due to a double cleavage (38), and molecular ion at 286 m/z); and iii) a C6 pyrrolidide (molecular mass at 213 m/z and base peak at 113 m/z) with additional peaks at 182 m/z (M-31) and 154 (Fig. 4A). Altogether these results established the structure of the major homolog of free mycobacteric acids of M. brumae as a 4,21-dimethyl-5,19-di-(trans)-enoyl-heptatriacontanoic acid (compound E0, Fig. 4B). Among the other homologs separated by argentation TLC, compounds E1a (Fig. 4B) related to M1a mycolic acids, were identified. Compounds containing one cis double bound in the distal part of the molecule and one trans double bond in the proximal portion of the molecule were also characterized (compound E1b, Fig. 4B). Detailed NMR analyses led us to propose for some of them the occurrence of a methyl group at the proximal part of the molecule, namely at C3, in agreement with both 1H-NMR prediction (Table 2, data not shown). The most polar compound on argentation TLC was identified as a mixture of 5,21- and 7,23-di-enoyl pyrrolidides. Indeed, the configuration of the two double bonds was shown to be cis, and no methyl branch was detected by EI-MS and NMR analyses of the pyrrolidide derivatives (compounds E2, Fig. 4B). It is important to note, however, that although all the structures established for the mycobacteric acids E0, E1a, E1b, and E2 of M. brumae (Fig. 4B) are related, respectively, to mycolic acids M0, M1a, M1b, and M2 (Fig. 1), the fact that the major homolog E0 has an odd chain length (C39) was unexpected because mycolic acids (and their main merochains) that contain two methyl branches or two cyclopropyl rings have been observed to possess an even chain length in accordance with the proposed mechanism of synthesis of trans double bonds (39).


View this table:
[in this window]
[in a new window]

  		TABLE 3. Chain lengths of M. brumae mycobacteric acid pyrrolidide derivatives

 

Figure 4
View larger version (13K):
[in this window]
[in a new window]

  		Fig. 4. Structures of the major homologs of mycobacteric acids from M. brumae. A: Illustration of the strategy used to establish the structures of mycobacteric acid derivatives deduced from the data from the electron impact (EI)-mass spectra of the pyrrolidide derivatives and from the characterization by GC-MS analyses of the fragments obtained after oxidative cleavage. The C39 H74 O2 acid (E0) is given as an example. B: Structures of the major homologs of the mycobacteric acids: E0 refers to acids with 2 trans double bonds, E1a to molecules with trans and cis double bonds, E1b to substances with cis and trans double bonds and E2 to compounds with two cis double bonds.

 
Identification of the natural forms of mycobacteric acids of M. brumae
The linked forms of the long-chain acids were investigated by fractionating the crude lipid extract by chromatography on a Florisil column and seeking for compounds with trans double bonds by infrared spectroscopy. The 965 cm–1 absorption band that indicates the occurrence of trans double bonds was exclusively found associated with fractions eluted with petroleum ether/ether. Analysis of these fractions by TLC showed that they were composed of two compounds migrating in the region of TAGs: one of these, TAG2, exhibited an Rf similar to that of the standard tripalmitin, whereas the other, TAG1, was less polar. The two TAGs were purified by preparative TLC, and their infrared spectra were similar to that of tripalmitin except for the presence of an intense band at 965 cm-1 in the spectrum of TAG1. Expectedly, the molecular rotation ([M]D) of this triglyceride was negative (–40°), consistent with the contribution of the chiral methyl group adjacent to trans double bonds (28).

The 1H-NMR spectra of TAG1 and TAG2 were compared with that of the commercial tripalmitin. They shared characteristic signals of triglycerides, notably those assignable to the proton resonances of the fatty acyl moiety at {delta} 0.85 (terminal CH3, 9H, triplet), 1.23 (-CH2-, broad), 1.58 (-CH2-CH2-COO, 6H, distorted triplet), 2.28 (-CH2-COO, 6H, two overlapping triplets). In addition, proton resonances attributable to those of methylene(s) and methine(s) bearing hydroxyl groups esterified by fatty acyl residues were observed at 4.09, 4.13, 4.25, and 4.28 ppm and assigned to those of positions 1 and 3 of glycerol (-CH2-OOC); the resonance of the proton at position 2 of glycerol (-CH-OOC) was observed at {delta} 5.25. The spectrum of TAG1 also contained specific proton resonances of ethylenic long-chain esters observed at {delta} 5.25–5.35 (-CH = CH-), 2.04 [-CH = CH-CH(CH3)-CH2-], 0.96 [-CH = CH-CH(CH3)-CH2-] and 0.99 [-CH = CH-CH(CH3)-CH2-COO]. All the attributions were established by 2D homo- and heteronuclear correlated spectroscopy, demonstrating that TAG1 corresponded to a TAG that contained one or more fatty acyl residue(s) containing methyl branch(es) adjacent to the double bond(s).

To firmly establish the structure of TAG1, we determined the molecular masses of the triglyceride constituents of M. brumae by MALDI-TOF mass spectroscopy (Fig. 5 ). Compared with the spectrum of the commercial tripalmitin (Fig. 5A), which contained a major sodiated molecular ion peak observed at 829 m/z expected for a [C51 H98 O6 Na], that of the TAG fraction of M. brumae was more complex. Two series of peaks were seen in its mass spectrum: one with of molecular ion peaks in the range of 773 to 1,051 m/z and the other massif at 1,105 to 1,344 m/z. The two massifs corresponded to the masses of TAG2 and TAG1, respectively, as demonstrated by the analysis of the mass spectra of the purified compounds (Fig. 5B, C). The spectrum of TAG2 exhibited a series of intense peudomolecular ion peaks from 801 to 967 m/z, differing one another by 28 amu, and corresponding to the mass values of triglycerides composed of fatty acid substituents with C14–22. (Fig. 5B). The mass spectrum of the purified TAG1 contained pseudomolecular ion peaks between 1,105 and 1,344 m/z (Fig. 5C). The major peak, seen at 1,147 m/z [C74 H140 06 Na], corresponded to the mass of a TAG molecule bearing one mole of C39:2 and two moles of C16:0 fatty acyl residues. The specific location of mycobacteric acids in TAG, a storage compound, suggested that they correspond to the degradation products of mycolic acids; consistent with this hypothesis is the observation that they have the same structural features as mycolic acids but with shorter and different ratios of odd versus even chain lengths.


Figure 5
View larger version (20K):
[in this window]
[in a new window]

  		Fig. 5. MALDI-time-of-flight (TOF) spectra of triacylglycerols (TAGs). Mass spectra of the commercial tripalmitin (A), the product isolated from M. brumae (TAG2) with a Rf similar to that of triplamitin (B) and the compound from M. brumae with a higher Rf (TAG1) (C). The Rfs of TAG1 and TAG2 from M. aurum are shown in Fig. 5A.

 
Comparative structural analysis of the meromycolic acids and compounds derived from the metabolism of mycolic acids of M. fallax
To determine the possible biosynthetic relationships between mycolic acids and the structurally related long-chain ketones, secondary and primary alcohols (compounds a, d, and f, respectively, Fig. 3) we directly compared their ratios of odd versus even chain lengths, a clear indication of catabolism, to those of the meroaldehyde chains obtained by pyrolitical cleavage of mycolic acids. For this purpose, M. fallax was chosen because this species accumulated more ketones and alcohols, notably compound f, than did M. brumae. The various types of mycolates from M. fallax were separated by argentation TLC as M1a and M1b, M2, and M3 (Fig. 1A). Each class of mycolic acids was then pyrolysed to yield the corresponding meroaldehydes. The same argentation TLC strategy was applied to the long-chain compounds. After purification, mycolic acids and their structurally related minor lipids were analyzed and their structures compared. The results from MS analyses allowed calculating the ratios of odd/even chain lengths for each class of compounds (Fig. 1) (Table 1). Interestingly, the chain lengths of the meroaldehydes from mycolic acid pyrolysis were one carbon longer than those of the long-chain mycobacteric esters. Consistently, the ratio of odd versus even chains was reversed in the mycobacteric acids compared with mycolic acids and the corresponding meroaldehydes, mycolones, and mycolols (Fig. 1) (Table 1). This observation was also valid for long-chain primary alcohols. Furthermore, the meromycolic acids resulting from the reduction of the meroaldehydes exhibited carbon numbers between 50 and 55, whereas mycobacteric acids and long-chain primary alcohols contained a larger pool of homologs with shorter chain lengths, from C35–44 and C44–56, respectively. Altogether, these data supported the hypothesis that mycobacteric acids (b) and the primary alcohols (f) derive from the mero aldehyde chain, whereas mycolones (a) and their reduced form (d) probably result from the decarboxylation of a β-keto ester.

Degradation of exogenic mycolic acids by M. aurum
The structural analogies between mycobacteric acids and the meromycolic chains suggest that the former compounds derive from the degradation of mycolic acids after the cleavage of the mycoloyl chain at the level of the {alpha}-branched/β-hydroxylated structure, the so-called mycolic unit. To test this hypothesis, the strain A+ of M. aurum that is known to degrade exogenic lipids (40) was used. Accordingly, uniformly 14C labeled {alpha}-mycolic acids were prepared by growing strain A+ of M. aurum on a medium supplemented with 14C acetate. The labeled {alpha}-mycolates were purified and added to the growth medium of the same strain and their degradation was followed. Around 30% to 60% of the labeling was found in the extractable lipids. The labeling was observed in the TAG fraction and most exclusively in TAG1 (Fig. 6A ), whereas no labeling was detected in phospholipids, trehalose mycolates, and bacterial residues. This labeled fraction was isolated by preparative TLC, saponified, and the resulting fatty acids were converted into their anilides derivatives and separated by normal phase liquid chromatography in two fractions, LC1 and LC2 (Fig. 6B). The anilide derivatives of the two fractions were analyzed by field desorption mass spectrometry, a technique that produces molecular ions without fragmentation of the carbon chain. Analysis of the mass spectra showed that the compounds consisted in a mixture of long-chain acids composed of both odd and even chain lengths (Table 4 ). LC2 was composed of acyl anilides having C35 to C44 acyl moieties, the most abundant corresponding to a C39:2 acyl group (molecular mass at 649 m/z), whereas LC1 consisted of homologs with 44 to 54 carbon atoms, with the main constituents being C49:2 and C50:2 acyl groups (molecular masses at 789 and 803 m/z). The metabolic products under study contained the two cyclopropyl rings found at the proximal and distal positions of the {alpha}-mycolates of M. aurum (Fig. 1B), consistent with a structural relationship between the latter compounds and the long-chain fatty acids LC1 and LC2 found in TAG1.


Figure 6
View larger version (8K):
[in this window]
[in a new window]

  		Fig. 6. Radio-TLC and -HPLC of the TAG fraction of M. aurum. Bacteria were grown in a medium containing 14C {alpha}-mycolate from M.aurum for 72 h under agitation. Cells were then extracted with chloroform/methanol and the resulting lipids were separated on a Florisil column. A: Radio-TLC of the TAG fraction eluted from the column with petroleum ether/ether 85:15 (v/v) and analyzed by TLC with the same solvent system. TAG1 and TAG2 correspond, respectively, to TAGs that contained long (C34–55) and moderate (C16–24) fatty acids chains. The mass spectra of the corresponding substances from M. brumae are shown in Fig. 4A, B, respectively. B: Labeling (dotted lines) and UV ({lambda} = 242 nm) profiles of anilide derivatives of the fatty acids obtained by saponification of the TAG1 fraction. The acid derivatives were fractionated using a µPorasil HPLC column (0.39 x 30 cm) eluted with hexane/dioxane (9:1, v/v); a flow rate of 1 ml/min was applied. LC1 and LC2 correspond to the two classes of long-chain fatty acid anilide derivatives.

 

View this table:
[in this window]
[in a new window]

  		TABLE 4. Molecular masses and chain lengths of the mycobacteric acids obtained from the degradation of exogenous 14C-labeled {alpha}-mycolic acids from M. aurum

 
Incorporation of 18O in long-chain alcohols of M. fallax
Two processes of cleavage of mycolic acids may lead to these labeled mycobacteric acid dead products: i) a retro-Claisen reaction, the reverse process of mycolic acid condensation, that would lead to compounds with an even number of carbon atoms, as occurring during the biosynthesis of meromycolates; and ii) a Baeyer-Villiger oxidation (i.e., oxidation of a keto group by the addition of molecular oxygen to give a wax that would yield upon hydrolysis long-chain fatty alcohols possessing one carbon less than the meromycolates and an alkymalonate) (Fig. 2). The ratio odd versus even chain lengths of mycobacteric acids characterized from M. brumae and M. fallax (see previous discussion) did not support the retro-Claisen hypothesis. To test the Baeyer-Villiger oxidation hypothesis, M. fallax was preferred to M. brumae and M. aurum for the occurrence and abundance of all the lipids. Accordingly, cells of M. fallax were incubated under either 18O2 or 16O2 atmosphere, and the reaction was stopped by saponification. Lipids were isolated, and mycolic acids and alcohols were purified on AgNO3-impregnated TLC and analyzed by EI-MS. The spectra of alcohols from both conditions were compared. For normal growth conditions, the major molecular ion was seen at 742 m/z, the expected mass of a C52:2 alcohol. When cells were grown in presence of 18O2, a significant increase of the M+2 ion peak was observed at m/z 744; the same mass shift was also found for the other homologs (Fig. 7A ). This increase in the abundance of the M+2 isotope strongly suggested the incorporation of one 18O atom into the long-chain alcohols. To ascertain that this increase was due to the incorporation of 18O isotope, and not to the gain of hydrogen atom during the assay, catalytic hydrogenation was performed and the resulting saturated alcohols were trimethlysilylated to improve the abundance of high mass ions by EI-MS (Fig. 7B). In the high mass range, intense M-15 ions (loss of a methyl group from the trimethylsilyl substituent) of the major homologs of alcohols were obtained. Upon growth in the presence of 16O, the M-15 ion from the C52 homolog was seen at 803 m/z. Under incubation in an 18O atmosphere, the relative intensity of the ion at 805 m/z increased dramatically, indicating that the increase of two amu in the M-15 ion was readily due to the incorporation of an 18O atom into the molecules.


Figure 7
View larger version (10K):
[in this window]
[in a new window]

  		Fig. 7. EI-MS analysis of long-chain primary alcohols from M. fallax cultures incubated in presence of 18O. A: Partial EI-mass spectra of M. fallax long-chain alcohols. Cells were grown under normal conditions (16 O atmosphere) (a) and under 18O atmosphere (b). Ions at m/z 742 and 744 correspond to a C52 alcohol. B: Partial-EI-mass spectra of M. fallax long-chain alcohols after catalytic reduction of double bonds, followed by silylation. (c) normal conditions; (d) 18O atmosphere. Ions at m/z 803 and m/z 817 correspond to [M-15]+. Ions at m/z 805 and m/z 819 correspond to [M-15] + 2 (from 18O).

 
To determine the specificity of the incorporation of the 18O into long-chain compounds, a comparative analysis of structurally related lipids, such as mycolones and mycolic acids, was performed. Accordingly, the oxygenated lipids were purified by argentation TLC and the EI spectra of the secondary alcohols derived from mycolones (i.e., mycolols) and those of mycolic acids were compared. Interestingly, comparable EI-MS profiles were obtained for these compounds grown under either 18O or 16O (data not shown). Consequently, the absence of incorporation of 18O in the hydroxyl groups of mycolols and mycolic acids proved the specificity of incorporation of 18O into the long-chain primary alcohols. It was thus concluded that the long-chain primary alcohols derived from the Baeyer-Villiger oxidation of mycolic acids through a keto intermediate (Fig. 2). Moreover, an alkylmalonate was identified in the GC-MS profile of volatile fatty esters (fragmentation ions m/z 132 and 145), in agreement with this mechanism.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The presence of long-chain fatty acids structurally related to mycolic acids is known for long to occur in mycobacteria. Two size ranges of chain lengths have been generally found, namely C36–47 fatty acids for which the name of mycobacteric acids has been proposed, and the other with chains between C50–56 (11–15). Both classes of long-chain fatty acids contain the structural features observed in the mycolic acids of the same species (i.e., cyclopropyl rings, double bonds, and/or oxygenated functions) (11, 13, 16). They have been isolated from a large variety of mycobacteria [e.g., M. tuberculosis (11, 12), M. smegmatis (15), M. kansasii (41), and from a species of Nocardia (17)]. In the present work, a series of C35–39 acids with odd and even numbers of carbon atoms and containing trans and cis double bonds was characterized and shown to be structurally related to the mycolic acids of M. brumae (28). Two pools of fatty acids, with C39 and C51 as the main terms, were also characterized in M. fallax and were shown to contain double bonds located at the same positions as in the meromycolic chain of the mycolic acids of this species. The similarities between the structural features of these mycobacteric acids and those of mycolic acids were a priori compatible with the former products being either the precursor or degradation compounds of the latter molecules. In this connection, a role for this type of long-chain fatty acids as mycolic acid precursors in growing cells has been suggested (41, 42). What seems to support this hypothesis is the fact that a cell-free system from M. tuberculosis was shown in earlier studies to synthesize saturated and unsaturated C30–40 and C48–56 fatty acids, the latter exhibiting chain lengths similar to those of the meromycolic chain (14). Similarly, of the two condensing enzymes KasA and KasB, which are components of the fatty acid synthase II system implicated in the biosynthesis of the main (mero) chains of mycolic acids, characterized in M. tuberculosis, KasA specifically elongates palmitate into long-chain fatty acids averaging 40 carbons in length; KasB, in the presence of KasA, produces longer fatty acids averaging C54 and structurally related to the meromycolic acids (43). However, if mycobacteric acids are implicated in a biosynthetic process, it is puzzling to observe that compounds characterized in the present work and in past studies contain a ratio even versus odd chains different from that of the aldehydes obtained by pyrolytical cleavage of {alpha}-mycolic acids.

To arrive to a documented working model about the origin of long-chain fatty acids, we followed the degradation of uniformly 14C labeled dicyclopropanated {alpha}-mycolic acids of M. aurum by a growing culture of M. aurum, a strain known to degrade long-chain compounds (40). Upon saponification of the cells, the labeling was exclusively found in fatty acids with chain averaging C39 and C51, as demonstrated by the presence of 14C radiolabeled mycobacteric acids. Interestingly, the two series of labeled fatty acids were shown to derive from the meromycolic chain of the {alpha}-mycolic acids, as they contain the two cyclopropyl groups. These results indicated that mycobacteric acids result from the degradation of the mero mycolic chain, implying first the cleavage of the mycolic acid unit to liberate the mero chain, followed by shortening the so-formed long chain by β-oxidation. At least two possibilities of cleavage of the mycolic acid unit would yield the meromycolate chain. One of them implies a mechanism similar to a retro-Claisen reaction, the reverse of the mycolic condensation. Such a reaction has been chemically performed with β-keto acyl trehalose, the product of the mycolic condensation (44, 45). Upon the growth of M. aurum cells in the presence of labeled mycolates, however, we were unable to detect radiolabeled acyl trehalose among the complex lipids; instead, the only radiolabeled complex lipid observed was apolar TAGs. Importantly, the retro-Claisen reaction would lead to mycobacteric acids analogous of the biosynthetic precursors, the meromycolic acids, in terms of parity of chain lengths. Because the meromycolic chains of the {alpha}-mycolic acids used in this experiment contained an even carbon atom number, it could be postulated that all long-chain acids so-formed and subsequently degraded by β-oxidation would retain an even carbon number. The ratio odd versus even chain lengths of the long-chain labeled molecules we characterized does not support this hypothesis. Alternatively, the rupture of the mycolic acid unit may be realized through a biological process similar to a Baeyer-Villiger reaction. Such a reaction consists in the insertion of one oxygen atom in the C-C bond adjacent to a keto group, yielding an ester function (wax). To perform such a process the reaction substrate would be a β-keto ester or a ketone (Fig. 2). The chain cleavage of the so-formed wax by esterases would occur between carbons 3 and 4 in the case of a keto ester and should produce, on the one hand, a long-chain alcohol that would contain one carbon less than the mero-chain and, on the other hand, an alkylmalonate (Fig. 2). The primary alcohol could be further dehydrogenated into a long-chain fatty acid that would be the substrate of β-oxidation. The chain lengths of mycobacteric acids with the main odd numbered 39 and 51 carbon terms and the characterization of the C22-alkymalonate in the lipids of saponification support this hypothesis (Fig. 2).

An efficient and specific insertion of one oxygen atom into an organic substrate is easily realized by specific enzymes, the flavoprotein-dependent enzymes [e.g., flavin monooxygenases and Baeyer-Villiger monooxygenases (BVMO)]. These enzymes play a role in the removal of toxic ketones and participate to the degradation of hydrocarbons by microorganisms. Flavin monooxygenases and BVMO have almost identical sequences (46), and several monooxygenases have been characterized in Corynebacterinae. For example, a putative BVMO has been detected in M. avium (47), a species containing keto- and its Baeyer-Villiger oxidation product, wax-mycolic acids (48–50). Interestingly, M. tuberculosis contains seven genes with BMVO motif (51). Their roles remain to be established but it was recently shown that a BMVO (Rv3854c) is responsible for the activation and therefore the efficacy of the widely used antituberculous prodrug ethionamide (52, 53). In the present work, the demonstration that the 18O labeling was exclusively found in long-chain primary alcohols parented to the mero mycolic chain, with the expected chain lengths, and not in other structurally related oxygenated long-chain lipids examined, clearly establishes that a key step in the degradation of mycolic acids is catalyzed by a Baeyer-Villiger-like process. The ability of these enzymes to convert relatively hydrophobic substrates and their membrane association fits well with a function in mycolic acid degradation. Future biochemical studies should clarify the role of oxygenases in mycobacterial physiology. Interestingly, because no BVMO have been identified in plants, animals, and humans, the identification of new classes of substrates and the corresponding reactivity as Baeyer-Villiger oxidation open new opportunities for developing chemotherapy for treating mycobacterial infections.

We showed that mycobacteric acids occurred either free or esterifying glycerol, solely as TAG but neither as trehalose lipids nor in phospholipids. MS analysis further demonstrated that the main homologs consisted of TAGs bearing one mycobacteric acyl residue per molecule. Interestingly, structurally related apolar nonhydroxylated fatty acids have been recently characterized in TAGs from various other mycobacteria, which include both slow growers (e.g., M. kansasii, M. gastri, and M. bovis BCG) and rapid growers (e.g., M. smegmatis mc2155 and M. thermoresistibile) (41). TAG fractions of M. smegmatis and M. bovis have been shown to be extremely asymmetric molecules, having a series of very long fatty acids located on position 3 of glycerol, in contrast to phospholipids where C20–26 fatty acyl substituents were absent (54). TAGs containing significant amounts of C37, C39, and C41 acids with odd chain lengths have been previously found in a Nocardia species that contained C56–58 linear ethylenic mycolic acids. More polar mycobacteric acids, with additional oxygen functions such as keto and wax, were also characterized in the TAG fractions of mycobacteria producing keto- and wax- mycolates: 2-octadecanol, a secondary alcohol found in wax mycolic acids, has been identified after saponification of TAG from M. aurum (16). The proposed storage role of TAGs may be advantageous for mycobacteria in vivo and relevant to the survival in a nonreplicating state. This concept is supported by a report on the upregulation of diacylglycerol transferases involved in TAG biosynthesis in stationary phase of M. tuberculosis cultures and by the observation that TAGs accumulate in dormant-like cultures of the bacillus (55). Likewise, M. kansasii was reported to accumulate TAGs and a so-called monomeromycoloyl diacylglycerol (similar to TAG1 characterized in the present study) during the late growth phase (41).


   ACKNOWLEDGMENTS
 
The authors are grateful to Drs Anne Lemassu and Françoise Laval (IPBS, Toulouse) and Jose Rodríguez (Servicio de Apoyo a la Investigacion, Universidad de Murcia, Spain) for NMR and fast-atom bombardment-MS analyses, respectively. We are indebted to Dr Sabine Gavalda (IPBS, Toulouse) for her great help in the preparation of figures.

Submitted on July 22, 2008
Revised on September 3, 2008


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
  1. Asselineau, C., J. Asselineau, G. Lanéelle, and M. A. Lanéelle. 2002. The biosynthesis of mycolic acids by Mycobacteria: current and alternative hypotheses. Prog. Lipid Res. 41: 501–523.[CrossRef][Medline]

  2. Barry III, C. E., R. E. Lee, K. Mdluli, A. E. Sampson, B. G. Schroeder, R. A. Slayden, and Y. Yuan. 1998. Mycolic acids: structure, biosynthesis and physiological functions. Prog. Lipid Res. 37: 143–179.[CrossRef][Medline]

  3. Daffé, M., and P. Draper. 1998. The envelope layer of mycobacteria with reference to their pathogenicity. Adv. Microb. Physiol. 39: 131–203.[Medline]

  4. Takayama, K., C. Wang, and G. S. Besra. 2005. Pathway to synthesis and processing of mycolic acids in Mycobacterium tuberculosis. Clin. Microbiol. Rev. 18: 81–101.[Abstract/Free Full Text]

  5. Vilchèze, C., H. R. Morbidoni, T. R. Weisbrod, H. Iwamoto, M. Kuo, J. C. Sacchettini, and W. R. Jacobs. 2000. Inactivation of the inhA-encoded fatty acid synthaseII (FASII) enoyl-carrier protein reductase induces accumulation of the FASI end products and cell lysis of Mycobacterium smegmatis. J. Bacteriol. 182: 4059–4067.[Abstract/Free Full Text]

  6. Portevin, D., C. de Souza-D'Auria, C. Houssin, C. Grimaldi, M. Chami, M. Daffé, and C. Guilhot. 2004. A polyketide synthase catalyses the last condensation step of mycolic acid biosynthesis in mycobacteria and related organisms. Proc. Natl. Acad. Sci. USA. 101: 314–319.[Abstract/Free Full Text]

  7. Portevin, D., C. de Souza-D'Auria, H. Montrozier, C. Houssin, A. Stella, M. A. Lanéelle, F. Bardou, C. Guilhot, and M. Daffé. 2005. The acyl-AMP ligase FAD32 and AccD4-containing acylCoA carboxylase are required for the synthesis of mycolic acids and essential for mycobaterial growth: identification of the carboxylation product and determination of the acyl-coA carboxylase components. J. Biol. Chem. 280: 8862–8874.[Abstract/Free Full Text]

  8. Brennan, P. J., and H. Nikaido. 1995. The cell envelope of mycobacteria. Annu. Rev. Biochem. 64: 29–63.[CrossRef][Medline]

  9. Banerjee, A., E. Dubnau, A. Quémard, V. Balasubramanian, K. S. Um, T. Wilson, D. Collins, G. de Lisle, and W. R. Jacobs. 1994. inhA, a gene encoding a target for isoniazid and ethionamide in Mycobacterium tuberculosis. Science. 263: 227–230.[Abstract/Free Full Text]

  10. Marrakchi, H., F. Bardou, M. A. Lanéelle, and M. Daffé. 2008. A comprehensive overview of mycolic acid structure and biosynthesis. In The mycobacterial cell envelope. M. Daffé and J. M. Reyrat, editors. ASM Press Washington DC. 41–62.

  11. Promé, J. C., C. Asselineau, and J. Asselineau. 1966. Acides mycobactériques, nouveaux acides isolés des lipids des mycobactéries. C. R. Acad. Sci. 263: 448–451.

  12. Takayama, K., and N. Qureshi. 1978. Isolation and characterization of the monounsaturated long chain fatty acids of Mycobacterium tuberculosis. Lipids. 13: 575–579.[CrossRef][Medline]

  13. Qureshi, N., K. Takayama, and H. K. Schnoes. 1980. Purification of C30–C56 fatty acids from Mycobacterium tuberculosis. J. Biol. Chem. 255: 182–189.[Free Full Text]

  14. Qureshi, N., N. Sathyamoorthy, and K. Takayama. 1984. Biosynthesis of C30 to C56 fatty acids by an extract of Mycobacterium tuberculosis H37 Ra. J. Bacteriol. 157: 46–52.[Abstract/Free Full Text]

  15. Sathyamoorthy, N., N. Qureshi, and K. Takayama. 1985. Purification and characterization of C28–55 fatty acids from Mycobacterium smegmatis. Can. J. Microbiol. 31: 214–219.[Medline]

  16. Lanéelle, M. A., C. Lacave, M. Daffé, and G. Lanéelle. 1988. Mycolic acids of Mycobacterium aurum. Structure and biogenetic implications. Eur. J. Biochem. 177: 631–635.[Medline]

  17. Vacheron, M. J., and G. Michel. 1971. Présence d'acides de poids moléculaire élevé dans les triglycérides isolés des parois cellulaires de Nocardia kirovani. C. R. Hebd. Seances Acad. Sci. 273: 1778–1781.

  18. Asselineau, J. 1954. Sur la tuberculénone, cétone insaturée isolée des lipides de souches humaines de Mycobacterium tuberculosis. C. R. Acad. Sci. 239: 1561–1563.

  19. Miquel, A. M., B. C. Das, and A. H. Etémadi. 1966. Sur la structure des {alpha}-kansamycolones, cétones à haut poids moléculaire isolées de Mycobacterium kansasii. Bull. Soc. Chim. Biol. (Paris). 47: 2095–2104.

  20. Bordet, C., and G. Michel. 1966. Isolement et structure des nocardones, cétones à haut poids moléculaire de Nocardia asteroides. C. R. Acad. Sci. 262: 1810–1812.

  21. Lanéelle, M. A. 1966. Sur la relation entre des cétones à haut poids moléculaires isolées de Nocardia brasiliensis et les acides nocardomycoliques. C. R. Acad. Sci. 263: 560–563.

  22. Bordet, C., and G. Michel. 1966. Les nocardols, alcools à haut poids moléculaire de Nocardia asteroides. C. R. Acad. Sci. 262: 1294–1296.

  23. Bordet, C., and G. Michel. 1969. Structure and biogenesis of high molecular weight lipids from Nocardia asteroides. Bull. Soc. Chim. Biol. (Paris). 51: 527–548.[Medline]

  24. Pudles, J., and E. Lederer. 1954. Sur l'isolement et la constitution de l'acide corynomycolénique et de deux cétones des lipides du bacille diphtérique. Bull. Soc. Chim. Biol. (Paris). 36: 759–777.[Medline]

  25. Asselineau, J. 1961. Sur la composition des lipides de Corynebacterium diphtheriae. Biochim. Biophys. Acta. 54: 359–360.[Medline]

  26. Bordet, C., and G. Michel. 1964. Isolation of a new alcohol, 16-hentriacontanol from the lipids of Nocardia brasiliensis. Bull. Soc. Chim. Biol. (Paris). 46: 1101–1112.[Medline]

  27. Lanéelle, M. A., J. Asselineau, and G. Castelnuovo. 1965. Etudes sur les mycobactéries et les nocardiae IV Composition des lipides de Mycobacterium rhodocrous, Mycobacterium pellegrino sp et de quelques souches de nocardiae. Ann. Inst. Pasteur (Paris). 108: 69–82.[Medline]

  28. Luquin, M., V. Ausina, V. Vincent-Lévy-Frébault, M. A. Lanéelle, F. Belda, M. Garcia-Barcelo, G. Prats, and M. Daffé. 1993. Mycobacterium brumae sp. nov., a rapidly growing, non photochromogenic Mycobacterium. Int. J. Syst. Bacteriol. 43: 405–413.[Abstract/Free Full Text]

  29. Lévy-Frébault, V., E. Rafidinarivo, J. C. Promé, J. Gandry, H. Boisvert, and H. L. David. 1983. Mycobacterium fallax sp. nov. Int. J. Syst. Bacteriol. 33: 336–343.[Abstract/Free Full Text]

  30. Von Rudloff, E. 1956. Periodate-permanganate oxidations V. Oxidation of lipids in media containing organic solvents. Can. J. Chem. 34: 1413–1418.[CrossRef]

  31. Rafidinarivo, E., J. C. Promé, and V. Lévy-Frébault. 1985. New kinds of unsaturated mycolic acids from Mycobacterium fallax. Chem. Phys. Lipids. 36: 215–228.[CrossRef]

  32. Odham, G., and E. Senhagen. 1972. Fatty acids. In Biochemical applications of Mass Spectrometry. G. R. Waller, editor. Willey-Interscience. New York. 211–228.

  33. Andersson, B. A., and R. T. Holman. 1974. Pyrrolidides for mass pectrometric determination of the position of the double bond in monounsaturated fatty acids. Lipids. 9: 185–190.[Medline]

  34. Yuan, Y., D. Mead, B. G. Schroeder, Y. Zhu, and C. E. Barry III. 1998. The biosynthesis of mycolic acids in Mycobacterium tuberculosis. Enzymatic methyl(ene) transfer to acyl carrier protein bound meromycolic acid. J. Biol. Chem. 273: 21282–21290.[Abstract/Free Full Text]

  35. Quémard, A., M. A. Lanéelle, H. Marrakchi, D. Promé, E. Dubnau, and M. Daffé. 1997. Structure of a hydroxymycolic acid potentially involved in the synthesis of oxygenated mycolic acids of the Mycobacterium tuberculosis complex. Eur. J. Biochem. 250: 758–763.[Medline]

  36. Laval, F., R. Haites, F. Movahedzadeh, A. Lemassu, C. Y. Wong, N. Stoker, H. Billman-Jacobe, and M. Daffé. 2008. Investigating the function of the putative mycolic-acid methyltransferase UmaA: divergence between the Mycobacterium smegmatis and Mycobacterium tuberculosis proteins. J. Biol. Chem. 283: 1419–1427.[Abstract/Free Full Text]

  37. Daffé, M., M. A. Lanéelle, and C. Lacave. 1991. Structure and stereochemistry of mycolic acids from Mycobacterium marinum and Mycobacterium ulcerans. Res. Microbiol. 142: 397–403.[Medline]

  38. Rhyage, R., and E. Stenhagen. 1959. Mass spectrometry studies III. Esters of saturated dibasic acids. Ark. Kemi. 14: 497–509.

  39. Lederer, E. 1969. Some problems concerning biological C-alkylation reactions and phytosterol biosynthesis. Q. Rev. Chem. Soc. 23: 453–481.[CrossRef]

  40. Promé, D., C. Lacave, B. Montsarrat, H. David, and J. C. Promé. 1983. Conversion of sterols and triterpenes by mycobacteria. I Formation of progesterone and 1-dehydroprogesterone from Mycobacterium aurum, strain A+. Biochim. Biophys. Acta. 753: 60–64.[Medline]

  41. Kremer, L., C. de Chastellier, G. Dobson, K. J. C. Gibson, P. Bifani, S. Balor, J. P. Gorvel, C. Locht, D. E. Minnikin, and G. S. Besra. 2005. Identification and structural characterization of an unusual mycobacterial monomeromycoloyl-diacylglycerol. Mol. Microbiol. 57: 1113–1126.[CrossRef][Medline]

  42. Lacave, C., M. A. Lanéelle, M. Daffé, H. Montrozier, M. P. Rols, and C. Asselineau. 1987. Etude structurale et métabolique des acides mycoliques de Mycobacterium fortuitum. Eur. J. Biochem. 163: 369–378.[Medline]

  43. Slayden, R. A., and C. E. Barry. 2002. The role of KasA and KasB in the biosynthesis of meromycolic acids and isoniazid resistance in Mycobacterium tuberculosis. Tuberculosis (Edinb.). 82: 149–160.[CrossRef][Medline]

  44. Aurelle, H., and J. C. Promé. 1980. The reaction of 3-oxo fatty acid esters with trehalose derivatives in an alkaline medium. Tetrahedron Lett. 21: 3277–3280.[CrossRef]

  45. Promé, J. C., R. W. Walker, and C. Lacave. 1974. Condensation of two molecules of palmitic acid in Corynebacterium diphtheriae: formation of a beta-keto ester of trehalose. C.R. Acad Sci (Paris). 278: 1065–1068.

  46. Van Berkel, W. J. H., N. M. Kamerbeek, and M. W. Fraaije. 2006. Flavoprotein monooxygenase, a diverse class of oxidative biocatalysts. J. Biotechnol. 124: 670–689.[CrossRef][Medline]

  47. Thierry, D., V. Vincent, F. Clément, and J. L. Guesdon. 1993. Isolation of specific DNA fragments of Mycobacterium avium and their possible use in diagnosis. J. Clin. Microbiol. 31: 1048–1054.[Abstract/Free Full Text]

  48. Etemadi, H. A., and J. Gashe. 1965. On the biogenetic origin of 2-eicosanol and 2-octadecanol of Mycobacterium avium. Bull. Soc. Chim. Biol. (Paris). 47: 2095–2104.[Medline]

  49. Lanéelle, M. A., and G. Lanéelle. 1970. Structure d'acides mycoliques et d'un intermédiaire dans la biosynthèse d'acides mycoliques dicarboxyliques. Eur. J. Biochem. 12: 296–300.[Medline]

  50. Toriyama, S., I. Imaizumi, I. Tomiyasu, M. Masui, and I. Yano. 1982. Incorporation of 18O into long-chain secondary alcohols derived from ester mycolic acids in Mycobacterium phlei. Biochim. Biophys. Acta. 712: 427–429.

  51. Fraaije, M. W., N. M. Kamerbeek, W. J. H. van Berkel, and D. B. Janssen. 2002. Identification of a Baeyer-Villiger monooxygenase sequence motif. FEBS Lett. 518: 43–47.[CrossRef][Medline]

  52. Fraaije, M. W., N. M. Kamerbeek, A. J. Heidekamp, R. Fortin, and D. B. Janssen. 2004. The prodrug activator EtaA from Mycobacterium tuberculosis is a Baeyer-Villiger monooxygenase. J. Biol. Chem. 279: 3354–3360.[Abstract/Free Full Text]

  53. Snajdora, R., G. Grogan, and M. D. Mihovilovic. 2006. Resolution of fused bicyclic ketones by a recombinant biocatalyst expressing the Baeyer-Villiger monooxygenase gene Rv3049c from Mycobacterium tuberculosis H37Rv. Bioorg. Med. Chem. Lett. 16: 4813–4817.[CrossRef][Medline]

  54. Walker, R. W., H. Barakat, and J. G. C. Hung. 1970. The positional distribution of fatty acids in the phospholipids and triglycerides of Mycobacterium smegmatis and M. bovis BCG. Lipids. 5: 684–691.[Medline]

  55. Daniel, J., C. Deb, V. S. Dubey, T. D. Sirakova, B. Abomoelak, H. R. Morbidoni, and P. E. Kolattukudy. 2004. Induction of a novel class of diacylglycerol acyltransferases and triacyl glycerol accumulation in Mycobacterium tuberculosis as it goes into a dormancy-like state in culture. J. Bacteriol. 186: 5017–5030.[Abstract/Free Full Text]


Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?



This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
M800384-JLR200v1
50/3/477    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rafidinarivo, E.
Right arrow Articles by Daffé, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rafidinarivo, E.
Right arrow Articles by Daffé, M.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Journal of Biological Chemistry 
 Molecular and Cellular Proteomics   ASBMB Today 
Advertisement
spacer
Advertisement
Advertisement