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Medicine » Endocrinology and Metabolism » "Lipid Metabolism", book edited by Rodrigo Valenzuela Baez, ISBN 978-953-51-0944-0, Published: January 23, 2013 under CC BY 3.0 license. © The Author(s).

Chapter 6

Metabolism of Plasma Membrane Lipids in Mycobacteria and Corynebacteria

By Paul K. Crellin, Chu-Yuan Luo and Yasu S. Morita
DOI: 10.5772/52781

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Mycobacterial plasma membrane and cell wall with flow of key metabolic pathways. Some of the metabolites are exported to the mycomembrane. SLD, small lipid droplet; LD, lipid droplet; FA-CoA, fatty acyl-CoA. See text for other abbreviations used in the figure.
Figure 1. Mycobacterial plasma membrane and cell wall with flow of key metabolic pathways. Some of the metabolites are exported to the mycomembrane. SLD, small lipid droplet; LD, lipid droplet; FA-CoA, fatty acyl-CoA. See text for other abbreviations used in the figure.
Fatty acid biosynthesis pathways in mycobacteria. Point of inhibition by the front-line tuberculosis drug isoniazid is indicated. Product profile of FAS-I is bimodal, and C16-C18-CoA and C24-C26-CoA are produced. Dashed lines indicate that some of the fatty acid products are further utilized for mycolic acid production.
Figure 2. Fatty acid biosynthesis pathways in mycobacteria. Point of inhibition by the front-line tuberculosis drug isoniazid is indicated. Product profile of FAS-I is bimodal, and C16-C18-CoA and C24-C26-CoA are produced. Dashed lines indicate that some of the fatty acid products are further utilized for mycolic acid production.
Glycerolipid/phospholipid biosynthesis pathways. Some pathways such as TAG and PE biosynthesis (shown as green arrows) do not occur in corynebacteria while some others (shown as blue arrows) are known to occur only in corynebacteria. PG is abundant in corynebacteria, but is a minor species in mycobacteria.
Figure 3. Glycerolipid/phospholipid biosynthesis pathways. Some pathways such as TAG and PE biosynthesis (shown as green arrows) do not occur in corynebacteria while some others (shown as blue arrows) are known to occur only in corynebacteria. PG is abundant in corynebacteria, but is a minor species in mycobacteria.
Summary of Lipoglycan Biosynthesis Pathways in Corynebacterineae. A) The mycobacterial pathway and structures for LM-A/LAM are shown, although several steps are inferred from studies in C. glutamicum. A progression from PIM1 to AcPIM2 via AcPIM1 can also occur but is sub-optimal in mycobacteria. AcPIM3 is shown as the substrate for the unidentified flippase but AcPIM2 may be the preferred substrate: AcPIM2 appears to be the flipped intermediate in C. glutamicum. Reactions associated with the PMf and PM-CW subfractions of M. smegmatis are shaded orange and grey, respectively. B) LM-B pathway in C. glutamicum. Its presence in mycobacteria remains to be determined. Intermediates representing Gl-Y and Gl-Z have been detected in very recent studies (Rainczuk et al, submitted). C) PPM serves as the lipid-linked donor of mannose for both pathways. The mechanism of flipping is undetermined. PM, plasma membrane. Other abbreviations are as defined in the text.
Figure 4. Summary of Lipoglycan Biosynthesis Pathways in Corynebacterineae. A) The mycobacterial pathway and structures for LM-A/LAM are shown, although several steps are inferred from studies in C. glutamicum. A progression from PIM1 to AcPIM2 via AcPIM1 can also occur but is sub-optimal in mycobacteria. AcPIM3 is shown as the substrate for the unidentified flippase but AcPIM2 may be the preferred substrate: AcPIM2 appears to be the flipped intermediate in C. glutamicum. Reactions associated with the PMf and PM-CW subfractions of M. smegmatis are shaded orange and grey, respectively. B) LM-B pathway in C. glutamicum. Its presence in mycobacteria remains to be determined. Intermediates representing Gl-Y and Gl-Z have been detected in very recent studies (Rainczuk et al, submitted). C) PPM serves as the lipid-linked donor of mannose for both pathways. The mechanism of flipping is undetermined. PM, plasma membrane. Other abbreviations are as defined in the text.

Metabolism of Plasma Membrane Lipids in Mycobacteria and Corynebacteria

Paul K. Crellin1, Chu-Yuan Luo2 and Yasu S. Morita2

1. Introduction

Bacteria of the Corynebacterineae, a suborder of the Actinobacteria, comprise Mycobacterium, Corynebacterium, Nocardia, Rhodococcus and other genera. This suborder of high GC gram-positive bacteria includes a number of important human pathogens, such as Mycobacterium tuberculosis, Mycobacterium leprae and Corynebacterium diphtheriae, the causative agents of tuberculosis, leprosy and diphtheria, respectively. M. tuberculosis is the most medically significant species, a devastating human pathogen infecting around one-third of the entire human population and responsible for more than 1 million deaths annually. The Corynebacterineae also includes non-pathogenic species such as Mycobacterium smegmatis, a saprophytic species, and Corynebacterium glutamicum, an industrial workhorse for the production of amino acids and other useful compounds. These relatively fast-growing species serve as useful models to study metabolic processes essential to the growth and survival of the slow-growing pathogens.

All these bacteria share a common feature, a distinctive multilaminate cell wall composed of peptidoglycan, complex polysaccharides, and both covalently linked lipids and free lipids/lipoglycans (Fig. 1). Among them, mycolic acids are the hallmark of these species. These long chain α-branched, β-hydroxylated fatty acids are covalently linked to the arabinogalactan polysaccharide layer. This mycolic acid layer is complemented by a glycolipid layer to form an outer “mycomembrane” analogous to the outer membrane of Gram-negative bacteria. [1, 2]. The outer leaflet of the mycomembrane is composed of a variety of lipids including trehalose dimycolates (TDMs), glycopeptidolipids (GPLs), phthiocerol dimycocerosates (PDIMs), sulfolipids, phenolic glycolipids (PGLs), and lipooligosaccharides. Some of these lipids are widely distributed while others are restricted to particular species. For example, TDMs and their structural equivalents are found in both mycobacteria and corynebacteria, while PDIMs and PGLs are restricted to a subset of mycobacteria. The structure and hydrophobic properties of the cell wall make it a potent permeability barrier that is responsible for intrinsic resistance of mycobacteria to an array of host microbiocidal processes, many antibiotics and sterilization conditions [3, 4]. Many of the cell wall components of pathogenic mycobacterial species are essential for pathogenesis and in vitro growth, hampering efforts to characterize the function of individual proteins in their assembly. In contrast, some non-pathogenic species such as C. glutamicum can tolerate the loss of major cell wall components, making them useful model systems for delineating processes involved in the assembly of core cell wall structures.


Figure 1.

Mycobacterial plasma membrane and cell wall with flow of key metabolic pathways. Some of the metabolites are exported to the mycomembrane. SLD, small lipid droplet; LD, lipid droplet; FA-CoA, fatty acyl-CoA. See text for other abbreviations used in the figure.

Studies on mycobacteria and corynebacteria provide a unique opportunity to illustrate the complexity and diversity of lipid metabolic pathways in bacteria. They have a significantly higher lipid content than other bacteria with cell wall lipids comprising ~40% of the dry cell mass. M. tuberculosis produces a diversity of lipids unparalleled in bacteria, from simple fatty acids to highly complex long chain structures such as mycolic acids. It has devoted a significant proportion of its coding capacity to lipid metabolism and produces about 250 enzymes dedicated to fatty acid metabolism, which is around five times the number produced by Escherichia coli [5]. Lipid biosynthesis places a significant metabolic burden on the organism but is ultimately advantageous, allowing M. tuberculosis to survive and replicate in the inhospitable environment of host macrophages. While capable of de novo synthesis, these bacteria also scavenge and degrade host cell membrane lipids to acetyl-CoA, via broad families of β-oxidation and other catabolic enzymes, for incorporation into their own metabolic pathways and to fuel cellular processes.

The plasma membrane provides the platform for lipid metabolism. While some lipid metabolic reactions take place in the cytoplasm or cell wall, the plasma membrane is the pivotal site for the metabolism of lipids. At the same time, this membrane must perform many other functions associated with energy production, nutrient uptake, protein export, and various sensing/signaling reactions. Studies on how these metabolic and cellular processes might be organized within bacterial plasma membranes are in their infancy. Understanding the homeostasis of the plasma membrane is particularly important in Corynebacterineae organisms because this structure must support the high biosynthetic demands of sustaining such a lipid-rich cell wall. In this chapter, we focus our discussion on processes of lipid metabolism that are critical for the biogenesis and maintenance of the plasma membrane, and illustrate the recent progress on our understanding of plasma membrane biogenesis in mycobacteria and corynebacteria.

2. Functions of plasma membrane lipids in mycobacteria and corynebacteria

In this section we will describe the functions of plasma membrane lipids. First, we will describe the functions of major structural phospholipids. We will then describe quantitatively minor lipids, which have important metabolic/physiological functions. Lastly, we will discuss the functions of neutral lipids because their biosynthesis is closely linked to phospholipid metabolism and neutral lipid storage is a critical part of plasma membrane homeostasis.

2.1. Structural lipids

Major structural components of the mycobacterial plasma membrane are phospholipids such as cardiolipin (CL), phosphatidylethanolamine (PE), phosphatidylinositol (PI), and glycosylated PIs (i.e. phosphatidylinositol mannosides (PIM), lipomannans (LM) and lipoarabinomannans (LAM), see below). The ratio of these phospholipids may vary depending on the species and growth conditions [6-8]. For example, one study indicated that CL, PE, and PI/PIMs represent about 37, 32, and 28%, respectively, of the total phospholipids in the plasma membrane in M. smegmatis [9], while another reported the ratio in Mycobacterium phlei to be about 50, 10, and 40% [10]. Phosphatidylglycerol (PG), which is abundant in other bacteria, is a relatively minor species in mycobacteria. Deletion of the PI biosynthetic gene has been shown to be lethal in M. smegmatis [9], indicating that PI or glycosylated PIs are essential for mycobacterial viability. In M. tuberculosis, putative PI synthetase (Rv2612c) and PGP synthetase (Rv2746c, involved in CL synthesis) genes are predicted to be essential [11], while the PS synthetase gene (Rv0436c, involved in PE synthesis) is not [12]. In corynebacteria, major species of phospholipids are PI, PG, CL, and acylphosphatidylglycerol (APG) [13], and PE appears to be absent.

CL is widely found in both prokaryotes and eukaryotes. It forms aggregates within the membrane bilayer. Nonyl acridine orange (NAO) is a fluorescent dye which is proposed to bind the hydrophobic surface created by the CL cluster [14], allowing microscopic visualization of CL domains. Indeed, using NAO, CLs were found to be enriched in septa and poles of actively dividing M. tuberculosis and M. smegmatis cells [15, 16]. CL has a non-bilayer structure [17, 18], and carries a small partially immobilized head group that is more exposed to the aqueous environment than those of other glycerophospholipids [19]. Although the physiological function of CL is unclear, its physical properties may indicate that it provides a platform for membrane-protein interactions. Indeed, some mycobacterial enzymes require CL for activity [20-22], although the molecular basis for these observations has not been clarified. Recent fractionation studies in C. glutamicum revealed that CL (as well as other phospholipids) is enriched in the plasma membrane [23, 24]. However, a large proportion of CL is also found to be associated with the outer membrane [24], suggesting that some of these phospholipids are exported to the outer membrane in corynebacteria. Similarly, CL is released from M. bovis bacillus Calmette-Guerin residing in host phagosomes, and converted to lyso-CL by a host phospholipase A2 [25]. It has been suggested that lyso-CL may influence host immune responses during infection.

PE is another major class of glycerophospholipids in mycobacteria. Although PE is generally found in all organisms, it is particularly abundant in bacterial plasma membranes [26]. Mycobacteria are no exception [20], but corynebacteria apparently lack the capacity to synthesize PE [27]. Indeed, PE biosynthetic enzymes, such as PS synthetase and PS decarboxylase, appear to be absent in corynebacterial genomes. Corynebacterium aquaticum has been reported to possess PE [28], but this species was later reclassified as Leifsonia aquatica [29], which belongs to the suborder Micrococcineae of the order Actinomycetales. The functions of PE remain elusive at the molecular level, but it appears to play important roles as a component of the plasma membrane. For example, TBsmr, a small multidrug resistance family protein from M. tuberculosis, shows enhanced catalytic activities when PE is supplemented in a reconstituted liposome [30].

PIs are an important class of phospholipids, and are known to be further modified by extensive glycosylation. The resultant lipoglycans, termed PIMs, LM, and LAM, are essential structural components of mycobacterial and corynebacterial cell walls. Furthermore, in pathogenic species, they have been suggested to perform additional roles in the modulation of host immune responses in favor of the pathogen through myriad effects on macrophages including cytokine production, inhibition of phagosome maturation and apoptosis [31-34]. PIMs are oligo-mannosylated PIs carrying up to 6 mannose residues while LM/LAM carry much longer mannose polymers with arabinan modifications. It remains controversial if these glycolipids are embedded in the plasma membrane or exported to the outer membrane. A recent study suggests that LM/LAM appear to be anchored to both the plasma membrane and outer membrane [35]. In C. glutamicum, the outer membrane and plasma membrane were fractionated on sucrose gradients upon cell lysis, and the analysis of these membrane sub-fractions demonstrated that PIMs, LM and LAM are all enriched in the plasma membrane fraction [23]. Another recent study also suggested that PI/PIMs are major components of the plasma membrane of C. glutamicum [24]. In the latter study, however, substantial amounts of PI/PIMs were detected in the outer membrane as well. The functional significance of these subcellular localizations, as well as the physiological roles of LM/LAM in each of these locations, remain important questions. The structural importance of PIMs remains unclear as well. For example, a pimE-deletion mutant that cannot produce mature PIM6 species (see below) is viable, but shows severe plasma membrane abnormalities [36], suggesting that higher order PIMs may be involved in the maintenance of plasma membrane integrity.

It is notable that some unusual phospholipids have been identified in corynebacteria. APG is an acylated form of PG which is widespread in corynebacteria [37-40], and is a major phospholipid species in Corynebacterium amycolatum. Another interesting phospholipid from C. amycolatum is acyl-phosphatidylinositol (API), which was identified by electrospray ionization mass spectroscopy [41]. C. amycolatum lacks a mycolic acid-based outer membrane, and does not appear to have a fracture plane other than the plasma membrane [42]. Therefore, APG and API are likely to be components of the plasma membrane, and are suggested to play structural roles. Very little is known about their biosynthesis, and acyltransferases responsible for their synthesis remain to be identified for both lipid species.

2.2. Functional lipids

There are some examples of lipids that appear to play no structural roles in the plasma membrane. They often exist in low quantities but play important functional roles. Among these, polyprenol-phospho-sugars function as sugar donors. Two well-studied examples are polyprenol phosphomannose (PPM) and decaprenol phosphoarabinose (DPA). These molecules are the donors of mannose and arabinose, respectively, and their biosynthesis will be discussed in a later section.

PI 3-phosphate, recently identified in both M. smegmatis and C. glutamicum [43], may prove to be another interesting example of a functional lipid. It accumulates only transiently upon stimulation by high concentrations of salt, and behaves as if it is involved in a signaling cascade. However, whether PI 3-phosphate represents a mediator of stress responses remains to be addressed. More recently, lysinylated PG was identified as a minor phospholipid species in M. tuberculosis [44]. The synthesis of lysinylated PG is mediated by LysX and a lysX deletion mutant showed altered phospholipid metabolism and membrane integrity [16, 44], suggesting a regulatory role of lysinylated PG in plasma membrane homeostasis.

Carotenoids are photo-protective pigments and serve to scavenge free radicals or harvest light [45]. Several mycobacterial species are known to produce carotenoids with the notable exception of M. tuberculosis, despite the presence of a carotenoid oxidase in the human pathogen [46]. These hydrophobic pigments are thought to be present in the plasma membrane but whether they play structural roles in addition to a photo-protective role remains to be elucidated.

2.3. Lipid storage for energy and carbon

Neutral lipids are an important reservoir of stored energy and carbon, and their metabolism is closely linked to plasma membrane phospholipid metabolism. Unlike many other bacteria which use polyhydroxyalkanoates as a lipid storage material [47], Actinobacteria use triacylglycerides (TAGs) as a major form of lipid storage, and the presence of TAGs has been reported in Mycobacterium, Streptomyces, Rhodococcus and Nocardia [48-52]. Interestingly, corynebacteria seem to lack the capacity to synthesize TAG, indicating that some lineages of Actinobacteria have eliminated this capacity at some point in their evolution. Recent evidence suggests that M. tuberculosis accumulates TAG-based lipid droplets while residing in macrophages using fatty acids released from host TAGs, and this process is critical for acquiring a dormancy phenotype [53]. Nevertheless, a mutant defective in accumulating TAG remained viable under in vitro dormancy-inducing conditions [54]. These somewhat contradictory observations suggest that our understanding of TAG metabolism in mycobacteria is far from complete. As we illustrate later, there appear to be several redundant genes involved in the final step of TAG synthesis, suggesting that it is an important regulatory step of lipid metabolism in these bacteria.

Cholesterol has recently been suggested to be an alternative form of lipid storage in mycobacteria. Neither mycobacteria nor corynebacteria have the capacity to synthesize cholesterol. However, cholesterol is taken up by M. tuberculosis cells residing in the host, and components of the mce4 operon have been shown to be involved in cholesterol import [55]. Cholesterol catabolism is critical in the chronic phase of animal infection, and a fully functional catabolic pathway is encoded by the M. tuberculosis genome [56]. Furthermore, cholesterol appears to accumulate in the mycobacterial cell envelope, and this might represent a potential form of lipid storage for M. tuberculosis during animal infection [57, 58]. Although the authors of this study suggested that cholesterol accumulates in the outer membrane, it remains possible that the plasma membrane is the true site of accumulation. Therefore, in addition to acting as a lipid storage molecule, cholesterol may play roles in plasma membrane structure and function, and these possibilities await further exploration.

Catabolism of cholesterol, amino acids and odd-chain-length/methyl branched fatty acids produces propionyl-coenzyme A (CoA). Propionate accumulation has been shown to be toxic in various organisms [59-61], and M. tuberculosis has multiple pathways to metabolize propionyl-CoA [62]. Metabolized propionyl-CoA is in part incorporated into TAG [63], and it has been suggested that TAG functions as a sink for reducing equivalents in addition to being a source of carbon and energy.

3. Structure and metabolism of plasma membrane lipids in mycobacteria and corynebacteria

In this section, we will describe the structure and metabolism of various lipids found in the plasma membrane of mycobacteria and corynebacteria in more detail. Lipids are categorized into the following four classes based on their key structural features.

3.1. Fatty acids

M. tuberculosis devotes a large proportion of its coding capacity to genes involved in fatty acid metabolism [5], highlighting the importance of lipids to the organism. Fatty acid metabolism is essential for intracellular survival of the pathogen since it forms the precursors of key membrane components such as plasma membrane phospholipids and outer membrane glycolipids. In particular, mycolic acids, which are very long chain α-alkyl β-hydroxy fatty acids, form the hydrophobic, protective mycomembrane described earlier. M. tuberculosis encodes two distinct enzyme systems for biosynthesis of fatty acids, designated FAS (fatty acid synthase) I and II (Fig. 2). Studies on fatty acid synthesis date back to the 1970s when M. smegmatis was shown to contain both type I fatty acid synthetase (FAS-I), involving a large multifunctional polypeptide, and type II fatty acid synthetase (FAS-II), consisting of a series of distinct enzymes [64]. The key elongation unit is malonyl-CoA, which is produced by acetyl-CoA carboxylase (ACCase) and the M. tuberculosis genome encodes several such enzymes (AccA1-3 and AccD1-6). The resultant malonyl-CoA is incorporated into fatty acids by the two FAS systems.


Figure 2.

Fatty acid biosynthesis pathways in mycobacteria. Point of inhibition by the front-line tuberculosis drug isoniazid is indicated. Product profile of FAS-I is bimodal, and C16-C18-CoA and C24-C26-CoA are produced. Dashed lines indicate that some of the fatty acid products are further utilized for mycolic acid production.

3.1.1. De novo synthesis by FAS-I

Surprisingly, members of the Corynebacterineae use a eukaryote-like FAS-I system for de novo fatty acid synthesis. The single, essential [11], 9.2kb fas gene encodes a 326 kDa protein containing all seven domains necessary to perform the iterative series of reactions: acyl transferase, enoyl reductase, β-hydroxyacyl dehydratase, malonyl transferase, acyl carrier protein, β-ketoacyl reductase, and β-ketoacyl synthase [65, 66]. This very large protein elongates acetyl groups by 2-carbon (acetate) units using acetyl-CoA and malonyl-CoA. Early rounds of elongation yield C16 to C18-CoA products that are used for synthesis of membrane phospholipids or to feed into the FAS-II system. More extensive elongation yields C24-C26 products that ultimately form the α-branch of mycolic acids. Unlike M. tuberculosis, C. glutamicum encodes two fas genes (fasA and fasB) with FasA taking the dominant role [67]. The presence of two Fas proteins may compensate for the lack of a FAS-II system in this organism.

3.1.2. Elongation by FAS-II

The FAS-II system is commonly found in bacteria and plants and, unlike FAS-I, is composed of a series of separate enzymes, each performing one step in the pathway. FAS-II elongates medium chain fatty acids derived from FAS-I using malonyl-CoA, producing C18-C30 fatty acids [68]. FAS-II has been extensively studied in E. coli [69] and orthologs of the fab genes have been identified in mycobacteria. AcpM is a mycobacterial acyl carrier protein (ACP) and plays a key role in transferring acyl groups between the various enzyme components [70]. The seven genes are located in two clusters on the M. tuberculosis chromosome [5], comprising mtfabD-acpM-kasA-kasB-accD6 and mabA-inhA. Initially, the malonate group is transferred from malonyl-CoA to AcpM by the MtFabD protein. Then MtFabH performs a Claisen condensation of malonyl-ACP with acyl-CoA to form β–ketoacyl-ACP. A four-step cycle is then initiated [64] in which:

  1. β–ketoacyl-ACP reductase MabA reduces the β–keto group with concomitant oxidation of NADPH

  2. β-hydroxyacyl-ACP dehydratase dehydrates the β-hydroxyl to enoyl-ACP

  3. enoyl-ACP reductase InhA, a target of the first-line anti-tuberculosis drug isoniazid (INH) [71], reduces enoyl-ACP to acyl-ACP with concomitant oxidation of NADPH

  4. β-ketoacyl-ACP synthase KasA/B elongates acyl-ACP by 2 carbon units, forming β-ketoacyl-ACP, which can feed back into step 1.

In this way, the hydrocarbon chain increases by 2 carbons each cycle. Further elongation and processing of the products of FAS-II produces the precursors of the long meromycolate chains that are condensed with the α-branches derived from FAS-I by the large polyketide synthase Pks13 [72]. Reduction of the β–keto group by CmrA forms the mature C60-C90 mycolic acid [73].

3.2. Glycerolipids

Glycerolipids include both nonpolar lipids and polar phospholipids. Their biosynthesis is overlapping and 1,2-diacyl-sn-glycerol 3-phosphate, commonly known as phosphatidic acid (PA), is an important intermediate at the branch point (Fig. 3) [74]. In this section, we focus our discussion on the biosynthesis of PA and its conversion to non-polar lipids. Non-polar lipids are generally divided into three different classes depending on the number of fatty acids attached to glycerol: monoacylglycerol (MAG), diacylglycerol (DAG) and TAG. TAG is a glycerol carrying three fatty acyl chains, and its biosynthesis diverges from phospholipid synthesis after the synthesis of PA. TAG is a major component of lipid droplets, which accumulate in the cytoplasm. How TAG is made in the plasma membrane and incorporated into lipid droplets remains largely unclear. Here, we provide an overview of the TAG metabolic pathway.


Figure 3.

Glycerolipid/phospholipid biosynthesis pathways. Some pathways such as TAG and PE biosynthesis (shown as green arrows) do not occur in corynebacteria while some others (shown as blue arrows) are known to occur only in corynebacteria. PG is abundant in corynebacteria, but is a minor species in mycobacteria.

3.2.1. Biosynthesis of PA

The first step of PA biosynthesis is mediated by glycerol phosphate acyltransferase (GPAT) transferring an acyl chain from acyl-CoA to glycerol-3-phosphate, forming acyl-glycerol 3-phosphate. In general, this reaction produces 1-acyl-sn-glycerol 3-phosphate. However, mycobacteria are unusual in that 2-acyl-sn-glycerol 3-phosphate is used as the main intermediate for the production of PA [75]. Another unusual feature is that oleic acid, an unsaturated fatty acid often found at the sn-2 positions of glycerolipids, is found at the sn-1 position in mycobacteria. Instead, palmitic acid, a saturated fatty acid, is the preferred fatty acid attached to the sn-2 position in mycobacteria [75, 76]. In the second step, acylglycerol phosphate acyltransferase (AGPAT) further transfers a fatty acid from acyl-CoA to 2-acyl-sn-glycerol 3-phosphate, producing PA. PA can be diverted to TAG synthesis, or activated to form cytidine diphosphate-diacylglycerol (CDP-DAG), which is the precursor for the synthesis of phospholipids. Therefore, PA represents an important branch point for the synthesis of TAG and phospholipids [74]. An alternative pathway for PA synthesis is phosphorylation of DAG by DAG kinase, and Rv2252 has been suggested to be involved in this reaction [77]. Disruption of this enzyme results in altered PIM biosynthesis, but precise functions of this metabolic pathway remain unclear.

3.2.2. TAG Biosynthesis

TAG is de novo synthesized by two steps. First, PA is dephosphorylated to become DAG, and this reaction is mediated by phosphatidic acid phosphatase (PAP). PAP was discovered from animal tissues in 1957 by the group of Eugene Kennedy [78], and the gene encoding this activity was recently identified in Saccharomyces cerevisiae [79]. Nothing is known about this enzyme in mycobacteria or corynebacteria. In the second step, diacylglycerol acyltransferase (DGAT) catalyzes the addition of a fatty acyl-CoA to DAG to form TAG. Until recently, little was known about the genes involved in this final step of TAG synthesis in mycobacteria. Analysis of this final step is complicated because there are multiple genes encoding TAG synthetase in mycobacteria and corynebacteria. For example, the M. tuberculosis genome encodes 15 putative TAG synthetase genes [48, 80]. Despite the redundancies, recent studies reported that some of these tgs genes are critical for TAG synthesis in M. tuberculosis [48, 54]. Specifically, TAG synthetases encoded by Rv3130c (tgs1), Rv3734c (tgs2), Rv3234c (tgs3), and Rv3088 (tgs4) have been shown to have TAG synthetase activities [53]. Furthermore, Tgs1 has been demonstrated to be the main contributor to TAG synthesis and lipid droplet formation in M. tuberculosis [53]. More recently, Ag85A, which is known as a mycolyltransferase involved in TDM biosynthesis, was shown to possess DGAT activity [81]. Ag85A is not homologous to other tgs genes, and may represent a novel class of TAG biosynthetic enzymes. TAG not only forms a lipid droplet in the cytoplasm, but also accumulates in the cell wall of mycobacteria [82]. Therefore, Ag85A located in the cell wall might be involved in the production of surface-exposed TAGs.

3.2.3. Utilization of TAG

Under starvation conditions where stored TAG needs to be mobilized for energy production, TAG is catabolized by lipases. In 1977, TAG lipase was purified from stationary phase M. phlei and predicted to have a molecular weight of about 40 kDa [83]. More recently, LipY, encoded by the M. tuberculosis Rv3097c gene, was identified as a TAG lipase [84]. LipY appears to play a critical role in TAG catabolism because a M. tuberculosis lipY deletion mutant cannot utilize accumulated TAG under starvation conditions. Another recent study demonstrated that LipY has a dual localization pattern [85]: while a fraction of LipY was found in the cytoplasm, consistent with its role in the catabolism of intracellular TAG, a significant fraction of LipY was also localized to the outer membrane of the cell wall, indicating that it may be involved in the breakdown of exogenously available TAGs. Indeed, it has been long known that M. tuberculosis depends on fatty acids as a preferred energy source during infection [86], and LipY may well be a critical enzyme for the utilization of host lipids during an M. tuberculosis infection. Another lipase encoded by Rv0183 shows preference for MAG over DAG and TAG, and is localized to the cell wall [87], suggesting its involvement in subsequent reactions of TAG breakdown. However, whether it is involved in degradation of host-derived TAG or intracellular TAG remains to be determined.

3.2.4. Lipid droplet formation

In eukaryotes, lipid droplets form in between the two leaflets of the endoplasmic reticulum membrane [88]. In bacteria, a distinct mechanism of lipid body formation has been proposed. For example, in rhodococci, TAG is formed in the cytoplasmic surface of the plasma membrane. Small lipid droplets are then fused to each other, coated by a monolayer of phospholipids, and released from the surface of the plasma membrane into the cytoplasm as mature lipid droplets [89]. Although no endogenous proteins have been found to associate with lipid droplets in rhodococci or mycobacteria, heterologous expression of known lipid droplet-associated proteins resulted in correct targeting of these proteins to lipid droplets in both R. opacus and M. smegmatis [90, 91], allowing visualization of lipid droplets in these organisms.

3.3. Phospholipids

3.3.1. CDP-DAG

In both eukaryotic and prokaryotic cells, PA is activated by CTP to form CDP-DAG, and this reaction is mediated by CDP-DAG synthase [92]. The synthesis of CDP-DAG commits the pathway to phospholipid biosynthesis, and CDP-DAG is a common precursor for the biosynthesis of all glycerophospholipids in mycobacteria and corynebacteria. The activity of CDP-DAG synthetase is associated with plasma membrane in M. smegmatis, and is possibly encoded by the cdsA (Rv2881c) gene in M. tuberculosis H37Rv [93].

3.3.2. CL

CL is composed of four acyl chains, three glycerols and two phosphates, and is structured in a 1,3-diphosphatidylglycerol configuration [94]. It is a common phospholipid in bacteria, and is one of the abundant phospholipids in mycobacteria and corynebacteria. To initiate CL synthesis, PG phosphate synthase first produces PG phosphate (PGP) using CDP-DAG and glycerol 3-phosphate as substrates. An M. smegmatis strain engineered to overexpress M. tuberculosis PgsA3 (encoded by Rv2746c) was shown to overproduce PG, suggesting that PgsA3 is the PGP synthase [9]. PGP is then converted into PG via PGP phosphatase. Three phosphatases, PgpA, PgpB, and PgpC, have been identified as PGP phosphatases in E. coli [95-97]. Furthermore, Gep4 and PTPMT1 have been identified as PGP phosphatases in yeast and mammals, respectively [98, 99]. Some homologs exist in the genomes of mycobacteria and corynebacteria, but experimental verification of these genes remains to be performed. Typically, the final step of CL synthesis in prokaryotes is mediated by a reaction that utilizes two PG molecules, producing one molecule of CL and one molecule of glycerol. However, in mycobacteria, the eukaryote-like reaction, which utilizes PG and CDP-DAG to produce CL, has been shown to occur [100], and Jackson and colleagues have suggested that PgsA2 might be the enzyme responsible for this reaction [9].

3.3.3. PE

The precise structure of PE was recently reported as 1-O-tuberculostearoyl-2-O-palmitoyl-sn-glycero-3-phosphoethanolamine in M. tuberculosis using tandem mass spectrometry [101]. In the initial step of PE synthesis, PS synthetase transfers serine to CDP-DAG and produces PS. In the second step, PE is produced by decarboxylation of PS mediated by PS decarboxylase. Genes encoding putative PS synthetase (pssA, Rv0436c) and PS decarboxylase (psd, Rv0437c) are found in tandem in the M. tuberculosis genome [9]. However, there is no experimental evidence demonstrating the identities of the genes. In M. smegmatis, PS synthetase and PS decarboxylase activities are enriched in different membrane fractions, which can be distinguished by sucrose gradient sedimentation [102]. However, the significance of differential membrane localization remains to be clarified.

3.3.4. PI

PI is a major phospholipid in both mycobacteria and corynebacteria and forms the anchor for the PIMs, which are substrates for heavy mannosylation to form LMs and additional arabinosylation to produce LAMs. PI is formed by the PI synthase PgsA (Rv2612c) from CDP-DAG and myo-inositol [9, 103]. Inositol is not a common metabolite in bacteria. It is therefore surprising that mycobacteria produce copious amounts of inositol through pathways shared with eukaryotes for incorporation into a range of metabolic pathways (recently reviewed in [104]). The enzyme d-myo-inositol 3-phosphate synthase (Ino1) converts glucose-6-phosphate to d-myo-inositol 3-phosphate, which is dephosphorylated by one of several inositol monophosphatases to produce myo-inositol [105].

3.3.5. PIMs

All Corynebacterineae synthesize PIMs that are important components of the cell envelope. Polar PIM species can also serve as membrane anchors for LM and LAM. Many of the steps of PIM/LM/LAM biosynthesis have now been elucidated [106]. Extensive genetic and biochemical studies have demonstrated that the synthesis of PIMs occurs linearly in mycobacteria with PI as the starting substrate (reviewed in [107]) (Fig. 4A). Early steps of the pathway occur on the cytoplasmic face of the plasma membrane. A PIM biosynthetic membrane, enriched in the early steps, has been purified by sucrose gradient fractionation as a membrane subdomain termed PMf, which is distinct from the bulk plasma membrane [102]. Mannosyltransferases performing the early steps utilize the water-soluble mannose donor, GDP-Man, which can be produced from exogenously acquired mannose or via de novo synthesis from the glycolytic pathway when fructose-6-phosphate is transformed by the enzymes ManA (Rv3255c), ManB (Rv3257c) and ManC (Rv3264c) [108-111], with a degree of redundancy reported at the ManC (NCgl0710) step in C. glutamicum [112]. The first step of PIM synthesis involves mannosylation of the C2-position of the inositol ring of PI by the enzyme PimA (Rv2610c) to form PIM1 [113, 114]. PimA is an essential enzyme in mycobacteria [11, 113] and absent in humans, making it a current target for drug development by several groups. The crystal structure of PimA from M. smegmatis has been solved in complex with GDP and GDP-Man at resolutions of 2.4Å and 2.6Å, respectively [115, 116]. PimA has a typical GT-B fold of glycosyltransferases consisting of two Rossmann-fold domains with a deep fissure at the interface, which contains the active site. Close to the GDP-Man binding site, the N-terminal domain displays a deep pocket containing highly conserved hydrophobic residues. This pocket is proposed to bind the acyl moieties of the acceptor substrate PI [116].

Next, O-6 mannosylation of the myo-inositol ring is performed by the cytoplasmic α-mannosyltransferase PimB’ (Rv2188c) [117, 118] resulting in the formation of PIM2. While the mycobacterial enzyme Rv0557 was originally assigned this function [119], later studies showed that Rv2188c was the true PimB (designated PimB’) [118], with Rv0557 being renamed MgtA and included in the LM-B pathway [120, 121] (see below). PimB’ is an essential enzyme in mycobacteria [11, 117] and the crystal structure of the equivalent corynebacterial enzyme has been solved at high resolution complexed with nucleotide [122]. In corynebacteria, PIM2 is detected mainly in its mono-acylated form, but in mycobacteria PIM2 accumulates in the cell envelope as monoacyl (AcPIM2) and diacyl (Ac2PIM2) forms, the former produced by the acyltransferase Rv2611c [123] which acts optimally on the product of the PimB’ reaction [117].

The next enzyme in the pathway, PimC, has been identified in M. tuberculosis strain CDC1551 and could produce trimannosylated PIMs [124]. However, the absence of this enzyme in other strains indicates redundancy at this step and the enzymes involved remain to be identified, as does the putative “PimD” protein. Flipping of PIM intermediates from the cytoplasmic face of the membrane to the periplasm is thought to occur at this point of the pathway, but the precise intermediate and transporter involved are also undefined.

AcPIM4 species can be further mannosylated to form more polar PIMs in reactions thought to take place on the periplasmic side of the cytoplasmic membrane. These reactions are performed by glycosyltransferases that require a lipid sugar donor in the form of PPM, since these reactions are amphomycin-sensitive [36, 125-128]. In mycobacteria, AcPIM4 is proposed to be a branch point for synthesis of polar PIM end products and LM/LAM. PimE (Rv1159) has been shown to elongate AcPIM4 with one or more α1-2 linked mannoses to form AcPIM6 [36]. This polytopic membrane protein has sequence similarities with eukaryotic PIG-M mannosyltransferases and localizes to a cell wall-associated plasma membrane subdomain in M. smegmatis, termed PM-CW, to which enzymatic activities of AcPIM4-6 synthesis are enriched [102]. Its catalytic activity was successfully mapped to a conserved aspartate residue in the first outer loop of the protein. Whether PimE also forms AcPIM6 is unknown. Interestingly, no PimE orthologue is present in C. glutamicum and Ac/Ac2PIM6 do not accumulate in this species, so the formation of Ac/Ac2PIM5/6 appears to be a mycobacterium-specific side-branch of the pathway.

Surprisingly, studies in M. smegmatis have revealed a role for a lipoprotein in PIM/LAM synthesis, since lpqW mutants produce reduced levels of LM/LAM [129]. As the only non- enzymatic component of the pathway identified to date, LpqW has been proposed to have a regulatory role at the bifurcation point of the pathways, as well as a functional connection with PimE, since mutations in the pimE gene can bypass the requirement for LpqW [130]. Very recent studies have implicated the corynebacterial ortholog of LpqW in the LM-B pathway as well (Rainczuk et al, submitted). Structural studies on the M. smegmatis LpqW revealed a scaffold similar to substrate binding proteins associated with ABC transporters, which has evolved to fulfill a new role in the regulation of PIM/LAM biosynthesis [131].

3.3.6. LM/LAM

A subpopulation of PIMs (AcPIM4 in mycobacteria [128, 129, 132] and AcPIM2 in corynebacteria [133]) can be extended with chains of α1-6 linked mannose to form LM that is further modified with a number of single α1-2 mannose side chains [134-136]. MptB is a PPM-dependent mannosyltransferase involved in extending AcPIM2 to form the proximal α1-6 mannan backbone of LM in C. glutamicum but is redundant in M. smegmatis [133], indicating additional complexity at this step in mycobacteria. Further elongation is performed by MptA (Rv2174) [135, 136], with MptC (Rv2181) required for addition of α1-2 linked Man side chains [106, 134, 137, 138]. Further additions of arabinose units by EmbC (Rv3793), AftC (Rv2673), AftD (Rv0236c) and at present unidentified α1-5 arabinofuranosyltransferases, result in the formation of mature LAM [139-142]. In M. tuberculosis and other pathogenic mycobacteria, additional mannose capping is present [143], synthesized by the enzymes MptC [137] and Rv1635c [144]. Alternatively, M. smegmatis LAM is capped with inositol phosphate [145].

While the general PIPIMLMLAM pathway is conserved in corynebacteria, a second pathway of lipoglycan biosynthesis exists in which a sub-population of LM lipoglycans is assembled on a glucopyranosyluronic acid diacylglycerol (Gl-A, GlcADAG) glycolipid anchor [118, 121, 133]. In this pathway (Fig. 4B), Gl-A is first mannosylated by MgtA (NCgl0452 in C. glutamicum and previously termed PimB, see above) forming mannosyl-glucuronic acid diacylglycerol (Gl-X, ManGlcADAG), which is subjected to further α1-6 (backbone) and α1-2 (side unit) mannosylation resulting in LM [121]. This pathway shares some PPM-dependent enzymes with the PI-based LM pathway including MptB, since an mptB (NCgl1505) mutant of C. glutamicum fails to produce intermediates beyond Gl-X or AcPIM2 [133]. For clarity, the PI-based LM pool has been designated LM-A while this second pathway produces LM-B [121, 146], the major LM pool in C. glutamicum [118]. While C. glutamicum produces LAMs via LM-A using a similar pathway to mycobacteria, its LAMs are smaller and structurally distinct, with more extensive mannosylation and singular Araf capping [147].


Figure 4.

Summary of Lipoglycan Biosynthesis Pathways in Corynebacterineae. A) The mycobacterial pathway and structures for LM-A/LAM are shown, although several steps are inferred from studies in C. glutamicum. A progression from PIM1 to AcPIM2 via AcPIM1 can also occur but is sub-optimal in mycobacteria. AcPIM3 is shown as the substrate for the unidentified flippase but AcPIM2 may be the preferred substrate: AcPIM2 appears to be the flipped intermediate in C. glutamicum. Reactions associated with the PMf and PM-CW subfractions of M. smegmatis are shaded orange and grey, respectively. B) LM-B pathway in C. glutamicum. Its presence in mycobacteria remains to be determined. Intermediates representing Gl-Y and Gl-Z have been detected in very recent studies (Rainczuk et al, submitted). C) PPM serves as the lipid-linked donor of mannose for both pathways. The mechanism of flipping is undetermined. PM, plasma membrane. Other abbreviations are as defined in the text.

3.4. Prenol lipids

Polyprenol phosphate (Pol-P) is a key carrier lipid in synthesis of the core structures of the mycobacterial cell wall, including peptidoglycan and arabinogalactan. Unlike most bacteria, mycobacteria contain multiple types of Pol-P. For example, M. smegmatis produces decaprenol phosphate (C50, Dec-P, [148]) and heptaprenol phosphate (C35, Hep-P, [149]). Polyprenols are thought to be synthesized via the condensation of two C5 lipids, isopentenyl diphosphate [150-152] and dimethylallyl diphosphate derived from the mevalonate-independent methylerythritol 4-phosphate (MEP) pathway. These reactions are catalyzed by prenol diphosphate synthases and two M. tuberculosis proteins, Rv2361c and Rv1086, acting sequentially, are thought to fulfill this role [148, 149, 153-155]. Finally, the C50 decaprenol diphosphate is dephosphorylated to produce Dec-P by an unknown phosphatase.

3.4.1. Polyprenol-phospho-sugars PPM

PPM, a β-d-mannosyl-1-monophosphoryldecaprenol, is utilized by periplasmic mannosyltransferases for synthesis of polar PIM species and LMs [106]. C35/C50-P-Manp is formed by the PPM synthase Ppm1 (Rv2051c) from GDP-Manp and Pol-P [126, 127] (Fig. 4C). In M. tuberculosis Ppm1 consists of two domains: a C-terminal catalytic domain and a N-terminal membrane anchor with 6 transmembrane helices. However, the equivalent domains of the M. smegmatis PPM synthase are two distinct, but interacting, proteins [156]. The importance of PPM in cell wall synthesis has been highlighted by analysis of a ppm1 mutant in C. glutamicum that failed to produce PPM, resulting in severe defects in lipoglycan biosynthesis. While the mutant could synthesize PIM2 species, all downstream products (LM, LAM) were absent, indicating their reliance on the PPM donor [127]. These findings were consistent with others showing that amphomycin, an antibiotic specific for PPM-dependent polymerases, blocked the PIM pathway at the PIM2 or PIM3 stage [125, 128]. The product of the Rv3779 gene has also been implicated in PPM synthesis but its role remains unclear [157, 158]. DPA

DPA is the only known donor of arabinose (Ara) for mycobacterial cell wall synthesis, contributing Araf units to arabinogalactan and LAM [106] with concomitant release of Pol-P. The Araf portion is derived from the pentose-phosphate pathway [159-161]. 5-phosphoribose 1-diphosphate is transferred to Dec-P by the Rv3806c gene product [162] and the resultant Dec-P-β-d-5-phosphoribose is dephosphorylated to form Dec-P-β-d-ribose. Oxidation of the 2’ hydroxyl is followed by a reduction reaction to form DPA. This two-step epimerization reaction is catalyzed by the combined activities of DprE1 (Rv3790) and DprE2 (Rv3791) [163]. Since DPA is the sole donor of Araf residues for mycobacterial cell wall synthesis, this pathway is of interest for drug development. Indeed, DprE1 is the target of dinitrobenzamide derivatives (DNBs) [164] and a set of nitro-compounds related to DNBs, the nitro-benzothiazinones (BTZ), a class of compounds with nanomolar anti-M. tuberculosis activities but minimal host-cell toxicity [165-167]. The essential nature of DprE1 in species beyond M. tuberculosis [168] reinforces it as a “magic” drug target [169]. The crystal structure of M. tuberculosis DprE1 complexed with BTZ inhibitors has been reported very recently, revealing the mode of inhibitor binding [170].

3.4.2. Carotenoids

Carotenoids are isoprenoid pigments widely distributed in biology and mostly based on C40-polyene. Synthesis of these pigments has been poorly studied in mycobacteria but they have proven useful for taxonomic and identification purposes. Mycobacterial pigments are generally yellow or orange and most have been confirmed as carotenoids [171]. While carotenoid genetics has been best studied in plants, the key enzymes of the pathway have been identified in bacteria, including mycobacteria [172-174]. There are two classes of carotenoids in bacteria, carotenes and xanthophylls, the latter of which contain oxygen. Both classes are composed of eight isoprenoid units with a long central chain of double bonded carbons. A consensus pathway for carotenoid biosynthesis in bacteria has been elucidated with orthologues of key enzymes identified in several species of mycobacteria [175]. As described above for prenol lipids, the carotenoid pathway begins with isopentenyl diphosphate and dimethylallyl diphosphate derived from the MEP pathway [176]. Head-to-tail condensation of these terpenes produces geranylgeranyl pyrophosphate (GGPP) due to the activity of GGPP synthase (CrtE). Condensation of two GGPP molecules [177] is followed by desaturation to phytoene by phytoene synthase (CrtB). Phytoene desaturase (CrtI) converts phytoene to lycoprene followed by cyclization to β-carotene by lycoprene cyclase (CrtY) [178].

4. Concluding remarks

Lipid metabolism in mycobacteria and corynebacteria is a highly complex network of catabolic and anabolic reactions. While the metabolic pathways and many of the enzymes involved have been actively elucidated over the past decade, substantial efforts are still needed to draw a comprehensive map of lipid metabolism in these organisms. In particular, our understanding of regulatory mechanisms of lipid metabolism is currently at an early stage. In addition, there are very few studies describing the interactions between multiple metabolic pathways of lipid biosynthesis. One promising approach for the comprehensive understanding of lipid metabolism is lipidomics, which is the study of lipid biosynthetic and catabolic pathways at a global level [179]. In the past, metabolic pathways have generally been examined in isolation without consideration of how different pathways might interact with, and influence, one another. Since the plasma membrane is a shared platform for most lipid biosynthetic pathways, and some donors are shared between different pathways (see above), it seems unlikely that the various pathways are truly independent. Recent advances in mass spectrometry (e.g. MALDI-MS, ESI-MS), nuclear magnetic resonance spectroscopy and associated computational methods have fuelled the development of this field [180]. Members of the Corynebacterineae, with their extensive lipid repertoires and complex metabolic pathways, would seem to be ideal targets to assess the true potential of lipidomics technologies. Recently, appropriate databases and methods for detection and identification of all major lipid classes of M. tuberculosis from a single crude extract have been developed [181, 182]. Very recently, a lipidomics profiling platform has been reported that uses high-performance liquid chromatography/mass spectrometry to resolve more than 12,000 molecules from M. tuberculosis [183]. These exciting advances provide a basis for future studies on the regulation of lipid metabolism and may allow, for the first time, a true appreciation of the interactive lipid networks of mycobacteria and corynebacteria.


1 - C Hoffmann, A Leis, M Niederweis, J. M Plitzko, H Engelhardt, Disclosure of the mycobacterial outer membrane: cryo-electron tomography and vitreous sections reveal the lipid bilayer structure. Proc Natl Acad Sci U S A. 200810539637
2 - B Zuber, M Chami, C Houssin, J Dubochet, G Griffiths, M Daffe, Direct visualization of the outer membrane of mycobacteria and corynebacteria in their native state. J Bacteriol. 2008190567280
3 - C. E Barry, rd, Mdluli K. Drug sensitivity and environmental adaptation of mycobacterial cell wall components. Trends Microbiol. 1996427581
4 - M Daffe, P Draper, The envelope layers of mycobacteria with reference to their pathogenicity. Adv Microb Physiol. 199839131203
5 - S. T Cole, R Brosch, J Parkhill, T Garnier, C Churcher, D Harris, et alDeciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature. 199839353744
6 - A. K Nandedkar, Comparative study of the lipid composition of particular pathogenic and nonpathogenic species of Mycobacterium. J Natl Med Assoc. 1983756974
7 - A Subramoniam, D Subrahmanyam, Light-induced changes in the phospholipid composition of Mycobacterium smegmatis ATCC 607. J Gen Microbiol. 198212841921
8 - G. K Khuller, R Taneja, N Nath, Effect of fatty acid supplementation on the lipid composition of Mycobacterium smegmatis ATCC 607, grown at 27 degrees and 37 degrees C. J Appl Bacteriol. 198354638
9 - M Jackson, D. C Crick, P. J Brennan, Phosphatidylinositol is an essential phospholipid of mycobacteria. J Biol Chem. 2000275300929
10 - Y Akamatsu, S Nojima, Separation and analyses of the individual phospholipids of mycobacteria. J Biochem. 1965574309
11 - C. M Sassetti, D. H Boyd, E. J Rubin, Genes required for mycobacterial growth defined by high density mutagenesis. Mol Microbiol. 2003487784
12 - G Lamichhane, M Zignol, N. J Blades, D. E Geiman, A Dougherty, J Grosset, et alA postgenomic method for predicting essential genes at subsaturation levels of mutagenesis: application to Mycobacterium tuberculosis. Proc Natl Acad Sci U S A. 200310072138
13 - G Yague, M Segovia, P. L Valero-guillen, Phospholipid composition of several clinically relevant Corynebacterium species as determined by mass spectrometry: an unusual fatty acyl moiety is present in inositol-containing phospholipids of Corynebacterium urealyticum. Microbiology. 2003149167585
14 - E Mileykovskaya, W Dowhan, R. L Birke, D Zheng, L Lutterodt, T. H Haines, Cardiolipin binds nonyl acridine orange by aggregating the dye at exposed hydrophobic domains on bilayer surfaces. FEBS Lett. 200150718790
15 - E Maloney, S. C Madiraju, M Rajagopalan, M Madiraju, Localization of acidic phospholipid cardiolipin and DnaA in mycobacteria. Tuberculosis (Edinb). 2011Suppl 1:S1505
16 - E Maloney, S Lun, D Stankowska, H Guo, M Rajagoapalan, W. R Bishai, et alAlterations in phospholipid catabolism in Mycobacterium tuberculosis lysX mutant. Front Microbiol. 2011
17 - M Dahlberg, Polymorphic phase behavior of cardiolipin derivatives studied by coarse-grained molecular dynamics. J Phys Chem B. 20071117194200
18 - G. L Powell, D Marsh, Polymorphic phase behavior of cardiolipin derivatives studied by 31P NMR and X-ray diffraction. Biochemistry. 19852429028
19 - R. N Lewis, D Zweytick, G Pabst, K Lohner, R. N Mcelhaney, Calorimetric, x-ray diffraction, and spectroscopic studies of the thermotropic phase behavior and organization of tetramyristoyl cardiolipin membranes. Biophys J. 200792316677
20 - T Imai, Y Kageyama, J Tobari, Mycobacterium smegmatis malate dehydrogenase: activation of the lipid-depleted enzyme by anionic phospholipids and phosphatidylethanolamine. Biochim Biophys Acta. 1995124618996
21 - T Kimura, J Tobari, Participation of flavin-adenine dinucleotide in the activity of malate dehydrogenase from Mycobacterium avium. Biochim Biophys Acta. 196373399405
22 - J Tobari, Requirement of flavin adenine dinucleotide and phospholipid for the activity of malate dehydrogenase from Mycobacterium avium. Biochem Biophys Res Commun. 196415504
23 - C. H Marchand, C Salmeron, Bou Raad R, Meniche X, Chami M, Masi M, et al. Biochemical disclosure of the mycolate outer membrane of Corynebacterium glutamicum. J Bacteriol. 201219458797
24 - R Bansal-mutalik, H Nikaido, Quantitative lipid composition of cell envelopes of Corynebacterium glutamicum elucidated through reverse micelle extraction. Proc Natl Acad Sci U S A. 2011108153605
25 - K Fischer, D Chatterjee, J Torrelles, P. J Brennan, S. H Kaufmann, U. E Schaible, Mycobacterial lysocardiolipin is exported from phagosomes upon cleavage of cardiolipin by a macrophage-derived lysosomal phospholipase A2. J Immunol. 2001167218792
26 - K Murzyn, T Rog, M Pasenkiewicz-gierula, Phosphatidylethanolamine-phosphatidylglycerol bilayer as a model of the inner bacterial membrane. Biophys J. 2005881091103
27 - P. J Brennan, D. P Lehane, The phospholipids of corynebacteria. Lipids. 197164019
28 - G. K Khuller, P. J Brennan, Further studies on the lipids of corynebacteria. The mannolipids of Corynebacterium aquaticum. Biochem J. 197212736973
29 - K. I Suzuki, M Suzuki, J Sasaki, Y. H Park, K. K Komagata, Leifsonia gen. nov., a genus for 2,4-diaminobutyric acid-containing actinomycetes to accommodate "Corynebacterium aquaticum" Leifson 1962 and Clavibacter xyli subsp. cynodontis Davis et al. 1984. J Gen Appl Microbiol. 19994525362
30 - K Charalambous, D Miller, P Curnow, P. J Booth, Lipid bilayer composition influences small multidrug transporters. BMC Biochem. 2008
31 - D Chatterjee, K. H Khoo, Mycobacterial lipoarabinomannan: an extraordinary lipoheteroglycan with profound physiological effects. Glycobiology. 1998811320
32 - J Nigou, M Gilleron, M Rojas, L. F Garcia, M Thurnher, G Puzo, Mycobacterial lipoarabinomannans: modulators of dendritic cell function and the apoptotic response. Microbes Infect. 2002494553
33 - G. R Strohmeier, M. J Fenton, Roles of lipoarabinomannan in the pathogenesis of tuberculosis. Microbes Infect. 1999170917
34 - A Vercellone, J Nigou, G Puzo, Relationships between the structure and the roles of lipoarabinomannans and related glycoconjugates in tuberculosis pathogenesis. Front Biosci. 1998e14963
35 - S Pitarque, G Larrouy-maumus, B Payre, M Jackson, G Puzo, J Nigou, The immunomodulatory lipoglycans, lipoarabinomannan and lipomannan, are exposed at the mycobacterial cell surface. Tuberculosis (Edinb). 2008885605
36 - Y. S Morita, C. B Sena, R. F Waller, K Kurokawa, M. F Sernee, F Nakatani, et alPimE is a polyprenol-phosphate-mannose-dependent mannosyltransferase that transfers the fifth mannose of phosphatidylinositol mannoside in mycobacteria. J Biol Chem. 20062812514355
37 - M. P Lechevalier, C Debievre, H Lechevalier, Chemotaxonomy of aerobic actinomycetes- phospholipid composition. Biochem Syst Ecol. 1977524960
38 - G Yague, M Segovia, P. L Valero-guillen, Acyl phosphatidylglycerol: a major phospholipid of Corynebacterium amycolatum. FEMS Microbiol Lett. 199715112530
39 - T Niepel, H Meyer, V Wray, W. R Abraham, Intraspecific variation of unusual phospholipids from Corynebacterium spp. containing a novel fatty acid. J Bacteriol. 199818046507
40 - N Mazzella, J Molinet, A. D Syakti, A Dodi, P Doumenq, J Artaud, et alBacterial phospholipid molecular species analysis by ion-pair reversed-phase HPLC/ESI/MS. J Lipid Res. 200445135563
41 - P. L Valero-guillen, G Yague, M Segovia, Characterization of acyl-phosphatidylinositol from the opportunistic pathogen Corynebacterium amycolatum. Chem Phys Lipids. 20051331726
42 - V Puech, M Chami, A Lemassu, M. A Laneelle, B Schiffler, P Gounon, et alStructure of the cell envelope of corynebacteria: importance of the non-covalently bound lipids in the formation of the cell wall permeability barrier and fracture plane. Microbiology. 2001147136582
43 - Y. S Morita, Y Yamaryo-botte, K Miyanagi, J. M Callaghan, J. H Patterson, P. K Crellin, et alStress-induced synthesis of phosphatidylinositol 3-phosphate in mycobacteria. J Biol Chem. 20102851664350
44 - E Maloney, D Stankowska, J Zhang, M Fol, Q. J Cheng, S Lun, et alThe two-domain LysX protein of Mycobacterium tuberculosis is required for production of lysinylated phosphatidylglycerol and resistance to cationic antimicrobial peptides. PLoS Pathog. 2009e1000534.
45 - P. D Fraser, P. M Bramley, The biosynthesis and nutritional uses of carotenoids. Prog Lipid Res. 20044322865
46 - D Scherzinger, E Scheffer, C Bar, H Ernst, S Al-babili, The Mycobacterium tuberculosis ORF Rv0654 encodes a carotenoid oxygenase mediating central and excentric cleavage of conventional and aromatic carotenoids. FEBS J. 2010277466273
47 - A Steinbuchel, H. E Valentin, Diversity of bacterial polyhydroxyalkanoic acids. FEMS Microbiol Lett. 199512821928
48 - J Daniel, C Deb, V. S Dubey, T. D Sirakova, B Abomoelak, H. R Morbidoni, et alInduction of a novel class of diacylglycerol acyltransferases and triacylglycerol accumulation in Mycobacterium tuberculosis as it goes into a dormancy-like state in culture. J Bacteriol. 2004186501730
49 - H. M Alvarez, A Steinbuchel, Triacylglycerols in prokaryotic microorganisms. Appl Microbiol Biotechnol. 20026036776
50 - H. M Alvarez, R Kalscheuer, A Steinbuchel, Accumulation and mobilization of storage lipids by Rhodococcus opacus PD630 and Rhodococcus ruber NCIMB 40126. Appl Microbiol Biotechnol. 20005421823
51 - H. M Alvarez, H Luftmann, R. A Silva, A. C Cesari, A Viale, M Waltermann, et alIdentification of phenyldecanoic acid as a constituent of triacylglycerols and wax ester produced by Rhodococcus opacus PD630. Microbiology. 2002148140712
52 - E. R Olukoshi, N. M Packter, Importance of stored triacylglycerols in Streptomyces: possible carbon source for antibiotics. Microbiology. 1994Pt 4):931-43.
53 - J Daniel, H Maamar, C Deb, T. D Sirakova, P. E Kolattukudy, Mycobacterium tuberculosis uses host triacylglycerol to accumulate lipid droplets and acquires a dormancy-like phenotype in lipid-loaded macrophages. PLoS Pathog. 2011e1002093.
54 - T. D Sirakova, V. S Dubey, C Deb, J Daniel, T. A Korotkova, B Abomoelak, et alIdentification of a diacylglycerol acyltransferase gene involved in accumulation of triacylglycerol in Mycobacterium tuberculosis under stress. Microbiology. 2006152271725
55 - A. K Pandey, C. M Sassetti, Mycobacterial persistence requires the utilization of host cholesterol. Proc Natl Acad Sci U S A. 2008105437680
56 - J. E Griffin, J. D Gawronski, M. A Dejesus, T. R Ioerger, B. J Akerley, C. M Sassetti, High-resolution phenotypic profiling defines genes essential for mycobacterial growth and cholesterol catabolism. PLoS Pathog. 2011e1002251.
57 - A Brzostek, J Pawelczyk, A Rumijowska-galewicz, B Dziadek, J Dziadek, Mycobacterium tuberculosis is able to accumulate and utilize cholesterol. J Bacteriol. 2009191658491
58 - Y Av-gay, R Sobouti, Cholesterol is accumulated by mycobacteria but its degradation is limited to non-pathogenic fast-growing mycobacteria. Can J Microbiol. 20004682631
59 - L. D Wright, H. R Skeggs, Reversal of sodium propionate inhibition of Escherichia coli with beta-alanine. Arch Biochem. 1946103836
60 - M Brock, W Buckel, On the mechanism of action of the antifungal agent propionate. Eur J Biochem. 2004271322741
61 - K Maruyama, H Kitamura, Mechanisms of growth inhibition by propionate and restoration of the growth by sodium bicarbonate or acetate in Rhodopseudomonas sphaeroides S. J Biochem. 19859881924
62 - S Savvi, D. F Warner, B. D Kana, J. D Mckinney, V Mizrahi, S. S Dawes, Functional characterization of a vitamin B12-dependent methylmalonyl pathway in Mycobacterium tuberculosis: implications for propionate metabolism during growth on fatty acids. J Bacteriol. 2008190388695
63 - A Singh, D. K Crossman, D Mai, L Guidry, M. I Voskuil, M. B Renfrow, et alMycobacterium tuberculosis WhiB3 maintains redox homeostasis by regulating virulence lipid anabolism to modulate macrophage response. PLoS Pathog. 2009e1000545.
64 - K Bloch, Control mechanisms for fatty acid synthesis in Mycobacterium smegmatis. Adv Enzymol Relat Areas Mol Biol. 197745184
65 - N. D Fernandes, P. E Kolattukudy, Cloning, sequencing and characterization of a fatty acid synthase-encoding gene from Mycobacterium tuberculosis var. bovis BCG. Gene. 1996170959
66 - K Bloch, D Vance, Control mechanisms in the synthesis of saturated fatty acids. Annu Rev Biochem. 19774626398
67 - E Radmacher, L. J Alderwick, G. S Besra, A. K Brown, K. J Gibson, H Sahm, et alTwo functional FAS-I type fatty acid synthases in Corynebacterium glutamicum. Microbiology. 200515124217
68 - J. M Odriozola, J. A Ramos, K Bloch, Fatty acid synthetase activity in Mycobacterium smegmatis. Characterization of the acyl carrier protein-dependent elongating system. Biochim Biophys Acta. 197748820717
69 - C. O Rock, J. E Cronan, Escherichia coli as a model for the regulation of dissociable (type II) fatty acid biosynthesis. Biochim Biophys Acta. 19961302116
70 - L Kremer, K. M Nampoothiri, S Lesjean, L. G Dover, S Graham, J Betts, et alBiochemical characterization of acyl carrier protein (AcpM) and malonyl-CoA:AcpM transacylase (mtFabD), two major components of Mycobacterium tuberculosis fatty acid synthase II. J Biol Chem. 20012762796774
71 - H Marrakchi, , G Laneelle, , A Quemard, . Inh, , a target of the antituberculous drug isoniazid, is involved in a mycobacterial fatty acid elongation system, FAS-II. Microbiology. 2000;146:289-96.
72 - D Portevin, De Sousa-D’Auria C, Houssin C, Grimaldi C, Chami M, Daffe M, et al. A polyketide synthase catalyzes the last condensation step of mycolic acid biosynthesis in mycobacteria and related organisms. Proc Natl Acad Sci U S A. 20041013149
73 - D. J Lea-smith, J. S Pyke, D Tull, M. J Mcconville, R. L Coppel, P. K Crellin, The reductase that catalyzes mycolic motif synthesis is required for efficient attachment of mycolic acids to arabinogalactan. J Biol Chem. 2007282110008
74 - K Athenstaedt, G Daum, Phosphatidic acid, a key intermediate in lipid metabolism. Eur J Biochem. 1999266116
75 - H Okuyama, Y Kameyama, M Fujikawa, K Yamada, H Ikezawa, Mechanism of diacylglycerophosphate synthesis in mycobacteria. J Biol Chem. 197725266826
76 - H Okuyama, T Kankura, S Nojima, Positional distribution of fatty acids in phospholipids from mycobacteria. J Biochem. 1967617327
77 - R. M Owens, F. F Hsu, VanderVen BC, Purdy GE, Hesteande E, Giannakas P, et al. M. tuberculosis Rv2252 encodes a diacylglycerol kinase involved in the biosynthesis of phosphatidylinositol mannosides (PIMs). Mol Microbiol. 200660115263
78 - S. W Smith, S. B Weiss, E. P Kennedy, The enzymatic dephosphorylation of phosphatidic acids. J Biol Chem. 195722891522
79 - G. S Han, W. I Wu, G. M Carman, The Saccharomyces cerevisiae Lipin homolog is a Mg2+-dependent phosphatidate phosphatase enzyme. J Biol Chem. 200628192108
80 - R Kalscheuer, A Steinbuchel, A novel bifunctional wax ester synthase/acyl-CoA:diacylglycerol acyltransferase mediates wax ester and triacylglycerol biosynthesis in Acinetobacter calcoaceticus ADP1. J Biol Chem. 2003278807582
81 - A. A Elamin, M Stehr, R Spallek, M Rohde, M Singh, The Mycobacterium tuberculosis Ag85A is a novel diacylglycerol acyltransferase involved in lipid body formation. Mol Microbiol. 201181157792
82 - A Ortalo-magne, A Lemassu, M. A Laneelle, F Bardou, G Silve, P Gounon, et alIdentification of the surface-exposed lipids on the cell envelopes of Mycobacterium tuberculosis and other mycobacterial species. J Bacteriol. 199617845661
83 - J. L Paznokas, A Kaplan, Purification and properties of a triacylglycerol lipase from Mycobacterium phlei. Biochim Biophys Acta. 197748740521
84 - C Deb, J Daniel, T. D Sirakova, B Abomoelak, V. S Dubey, P. E Kolattukudy, A novel lipase belonging to the hormone-sensitive lipase family induced under starvation to utilize stored triacylglycerol in Mycobacterium tuberculosis. J Biol Chem. 2006281386675
85 - K. C Mishra, C De Chastellier, Y Narayana, P Bifani, A. K Brown, G. S Besra, et alFunctional role of the PE domain and immunogenicity of the Mycobacterium tuberculosis triacylglycerol hydrolase LipY. Infect Immun. 20087612740
86 - E. J Munoz-elias, J. D Mckinney, Carbon metabolism of intracellular bacteria. Cell Microbiol. 200681022
87 - K Cotes, R Dhouib, I Douchet, H Chahinian, A De Caro, F Carriere, et alCharacterization of an exported monoglyceride lipase from Mycobacterium tuberculosis possibly involved in the metabolism of host cell membrane lipids. Biochem J. 200740841727
88 - M Waltermann, A Steinbuchel, Neutral lipid bodies in prokaryotes: recent insights into structure, formation, and relationship to eukaryotic lipid depots. J Bacteriol. 2005187360719
89 - M Waltermann, A Hinz, H Robenek, D Troyer, R Reichelt, U Malkus, et alMechanism of lipid-body formation in prokaryotes: how bacteria fatten up. Mol Microbiol. 20055575063
90 - J Hanisch, M Waltermann, H Robenek, A Steinbuchel, Eukaryotic lipid body proteins in oleogenous actinomycetes and their targeting to intracellular triacylglycerol inclusions: Impact on models of lipid body biogenesis. Appl Environ Microbiol. 200672674350
91 - J Hanisch, M Waltermann, H Robenek, A Steinbuchel, The Ralstonia eutropha H16 phasin PhaP1 is targeted to intracellular triacylglycerol inclusions in Rhodococcus opacus PD630 and Mycobacterium smegmatis mc2155, and provides an anchor to target other proteins. Microbiology. 2006152327180
92 - W Dowhan, CDP-diacylglycerol synthase of microorganisms. Biochim Biophys Acta. 1997134815765
93 - J Nigou, G. S Besra, Cytidine diphosphate-diacylglycerol synthesis in Mycobacterium smegmatis. Biochem J. 200236715762
94 - J Lecocq, C. E Ballou, On the structure of cardiolipin. Biochemistry. 1964397680
95 - T Icho, C. R Raetz, Multiple genes for membrane-bound phosphatases in Escherichia coli and their action on phospholipid precursors. J Bacteriol. 198315372230
96 - C. R Funk, L Zimniak, W Dowhan, The pgpA and pgpB genes of Escherichia coli are not essential: evidence for a third phosphatidylglycerophosphate phosphatase. J Bacteriol. 199217420513
97 - Y. H Lu, Z Guan, J Zhao, C. R Raetz, Three phosphatidylglycerol-phosphate phosphatases in the inner membrane of Escherichia coli. J Biol Chem. 2011286550618
98 - C Osman, M Haag, F. T Wieland, B Brugger, T Langer, A mitochondrial phosphatase required for cardiolipin biosynthesis: the PGP phosphatase Gep4. EMBO J. 201029197687
99 - J Zhang, Z Guan, A. N Murphy, S. E Wiley, G. A Perkins, C. A Worby, et alMitochondrial phosphatase PTPMT1 is essential for cardiolipin biosynthesis. Cell Metab. 201113690700
100 - A. K Mathur, P. S Murthy, G. S Saharia, T. A Venkitasubramanian, Studies on cardiolipin biosynthesis in Mycobacterium smegmatis. Can J Microbiol. 1976223548
101 - Ter Horst BSeshadri C, Sweet L, Young DC, Feringa BL, Moody DB, et al. Asymmetric synthesis and structure elucidation of a glycerophospholipid from Mycobacterium tuberculosis. J Lipid Res. 201051101722
102 - Y. S Morita, R Velasquez, E Taig, R. F Waller, J. H Patterson, D Tull, et alCompartmentalization of lipid biosynthesis in mycobacteria. J Biol Chem. 20052802164552
103 - M Salman, J. T Lonsdale, G. S Besra, P. J Brennan, Phosphatidylinositol synthesis in mycobacteria. Biochim Biophys Acta. 1999143643750
104 - Y. S Morita, T Fukuda, C. B Sena, Y Yamaryo-botte, M. J Mcconville, T Kinoshita, Inositol lipid metabolism in mycobacteria: biosynthesis and regulatory mechanisms. Biochim Biophys Acta. 2011181063041
105 - J Nigou, G. S Besra, Characterization and regulation of inositol monophosphatase activity in Mycobacterium smegmatis. Biochem J. 200236138590
106 - A. K Mishra, N. N Driessen, B. J Appelmelk, G. S Besra, Lipoarabinomannan and related glycoconjugates: structure, biogenesis and role in Mycobacterium tuberculosis physiology and host-pathogen interaction. FEMS Microbiol Rev. 201135112657
107 - M. E Guerin, J Kordulakova, P. M Alzari, P. J Brennan, M Jackson, Molecular basis of phosphatidyl-myo-inositol mannoside biosynthesis and regulation in mycobacteria. J Biol Chem. 20102853357783
108 - Y Ma, R. J Stern, M. S Scherman, V. D Vissa, W Yan, V. C Jones, et alDrug targeting Mycobacterium tuberculosis cell wall synthesis: genetics of dTDP-rhamnose synthetic enzymes and development of a microtiter plate-based screen for inhibitors of conversion of dTDP-glucose to dTDP-rhamnose. Antimicrob Agents Chemother. 200145140716
109 - T. R Mccarthy, J. B Torrelles, MacFarlane AS, Katawczik M, Kutzbach B, Desjardin LE, et al. Overexpression of Mycobacterium tuberculosis manB, a phosphomannomutase that increases phosphatidylinositol mannoside biosynthesis in Mycobacterium smegmatis and mycobacterial association with human macrophages. Mol Microbiol. 20055877490
110 - B Ning, A. D Elbein, Purification and properties of mycobacterial GDP-mannose pyrophosphorylase. Arch Biochem Biophys. 199936233945
111 - J. H Patterson, R. F Waller, D Jeevarajah, H Billman-jacobe, M. J Mcconville, Mannose metabolism is required for mycobacterial growth. Biochem J. 20033727786
112 - A. K Mishra, K Krumbach, D Rittmann, S. M Batt, O. Y Lee, De S, et alDeletion of manC in Corynebacterium glutamicum results in a phospho-myo-inositol mannoside- and lipoglycan-deficient mutant. Microbiology. 2012158190817
113 - J Kordulakova, M Gilleron, K Mikusova, G Puzo, P. J Brennan, B Gicquel, et alDefinition of the first mannosylation step in phosphatidylinositol mannoside synthesis. PimA is essential for growth of mycobacteria. J Biol Chem. 20022773133544
114 - X Gu, M Chen, Q Wang, M Zhang, B Wang, H Wang, Expression and purification of a functionally active recombinant GDP-mannosyltransferase (PimA) from Mycobacterium tuberculosis H37Rv. Protein Expr Purif. 2005424753
115 - M. E Guerin, A Buschiazzo, J Kordulakova, M Jackson, P. M Alzari, Crystallization and preliminary crystallographic analysis of PimA, an essential mannosyltransferase from Mycobacterium smegmatis. Acta Crystallograph Sect F Struct Biol Cryst Commun. 20056151820
116 - M. E Guerin, J Kordulakova, F Schaeffer, Z Svetlikova, A Buschiazzo, D Giganti, et alMolecular recognition and interfacial catalysis by the essential phosphatidylinositol mannosyltransferase PimA from mycobacteria. J Biol Chem. 20072822070514
117 - M. E Guerin, D Kaur, B. S Somashekar, S Gibbs, P Gest, D Chatterjee, et alNew insights into the early steps of phosphatidylinositol mannoside biosynthesis in mycobacteria: PimB’ is an essential enzyme of Mycobacterium smegmatis. J Biol Chem. 20092842568796
118 - D. J Lea-smith, K. L Martin, J. S Pyke, D Tull, M. J Mcconville, R. L Coppel, et alAnalysis of a new mannosyltransferase required for the synthesis of phosphatidylinositol mannosides and lipoarabinomannan reveals two lipomannan pools in Corynebacterineae. J Biol Chem. 2008283677382
119 - M. L Schaeffer, K. H Khoo, G. S Besra, D Chatterjee, P. J Brennan, J. T Belisle, et alThe pimB gene of Mycobacterium tuberculosis encodes a mannosyltransferase involved in lipoarabinomannan biosynthesis. J Biol Chem. 19992743162531
120 - A. K Mishra, S Batt, K Krumbach, L Eggeling, G. S Besra, Characterization of the Corynebacterium glutamicum deltapimB’ deltamgtA double deletion mutant and the role of Mycobacterium tuberculosis orthologues Rv2188c and Rv0557 in glycolipid biosynthesis. J Bacteriol. 2009191446572
121 - R. V Tatituri, P. A Illarionov, L. G Dover, J Nigou, M Gilleron, P Hitchen, et alInactivation of Corynebacterium glutamicum NCgl0452 and the role of MgtA in the biosynthesis of a novel mannosylated glycolipid involved in lipomannan biosynthesis. J Biol Chem. 2007282456172
122 - S. M Batt, T Jabeen, A. K Mishra, N Veerapen, K Krumbach, L Eggeling, et alAcceptor substrate discrimination in phosphatidyl-myo-inositol mannoside synthesis: structural and mutational analysis of mannosyltransferase Corynebacterium glutamicum PimB’. J Biol Chem. 20102853774152
123 - J Kordulakova, M Gilleron, G Puzo, P. J Brennan, B Gicquel, K Mikusova, et alIdentification of the required acyltransferase step in the biosynthesis of the phosphatidylinositol mannosides of Mycobacterium species. J Biol Chem. 20032783628595
124 - L Kremer, S. S Gurcha, P Bifani, P. G Hitchen, A Baulard, H. R Morris, et alCharacterization of a putative alpha-mannosyltransferase involved in phosphatidylinositol trimannoside biosynthesis in Mycobacterium tuberculosis. Biochem J. 200236343747
125 - G. S Besra, C. B Morehouse, C. M Rittner, C. J Waechter, P. J Brennan, Biosynthesis of mycobacterial lipoarabinomannan. J Biol Chem. 1997272184606
126 - S. S Gurcha, A. R Baulard, L Kremer, C Locht, D. B Moody, W Muhlecker, et alPpm1, a novel polyprenol monophosphomannose synthase from Mycobacterium tuberculosis. Biochem J. 200236544150
127 - K. J Gibson, L Eggeling, W. N Maughan, K Krumbach, S. S Gurcha, J Nigou, et alDisruption of Cg-Ppm1, a polyprenyl monophosphomannose synthase, and the generation of lipoglycan-less mutants in Corynebacterium glutamicum. J Biol Chem. 20032784084250
128 - Y. S Morita, J. H Patterson, H Billman-jacobe, M. J Mcconville, Biosynthesis of mycobacterial phosphatidylinositol mannosides. Biochem J. 200437858997
129 - S Kovacevic, D Anderson, Y. S Morita, J Patterson, R Haites, B. N Mcmillan, et alIdentification of a novel protein with a role in lipoarabinomannan biosynthesis in mycobacteria. J Biol Chem. 200628190117
130 - P. K Crellin, S Kovacevic, K. L Martin, R Brammananth, Y. S Morita, H Billman-jacobe, et alMutations in pimE restore lipoarabinomannan synthesis and growth in a Mycobacterium smegmatis lpqW mutant. J Bacteriol. 200819036909
131 - Z Marland, T Beddoe, L Zaker-tabrizi, I. S Lucet, R Brammananth, J. C Whisstock, et alHijacking of a substrate-binding protein scaffold for use in mycobacterial cell wall biosynthesis. J Mol Biol. 200635998397
132 - G. S Besra, P. J Brennan, The mycobacterial cell wall: biosynthesis of arabinogalactan and lipoarabinomannan. Biochem Soc Trans. 19972584550
133 - A. K Mishra, L. J Alderwick, D Rittmann, C Wang, A Bhatt, W. R Jacobs, Jr., et alIdentification of a novel alpha(1-->6) mannopyranosyltransferase MptB from Corynebacterium glutamicum by deletion of a conserved gene, NCgl1505, affords a lipomannan- and lipoarabinomannan-deficient mutant. Mol Microbiol. 2008681595613
134 - D Kaur, S Berg, P Dinadayala, B Gicquel, D Chatterjee, M. R Mcneil, et alBiosynthesis of mycobacterial lipoarabinomannan: role of a branching mannosyltransferase. Proc Natl Acad Sci U S A. 2006103136649
135 - D Kaur, M. R Mcneil, K. H Khoo, D Chatterjee, D. C Crick, M Jackson, et alNew insights into the biosynthesis of mycobacterial lipomannan arising from deletion of a conserved gene. J Biol Chem. 20072822713340
136 - A. K Mishra, L. J Alderwick, D Rittmann, R. V Tatituri, J Nigou, M Gilleron, et alIdentification of an alpha(1-->6) mannopyranosyltransferase (MptA), involved in Corynebacterium glutamicum lipomannan biosynthesis, and identification of its orthologue in Mycobacterium tuberculosis. Mol Microbiol. 200765150317
137 - D Kaur, A Obregon-henao, H Pham, D Chatterjee, P. J Brennan, M Jackson, Lipoarabinomannan of Mycobacterium: mannose capping by a multifunctional terminal mannosyltransferase. Proc Natl Acad Sci U S A. 2008105179737
138 - C. B Sena, T Fukuda, K Miyanagi, S Matsumoto, K Kobayashi, Y Murakami, et alControlled expression of branch-forming mannosyltransferase is critical for mycobacterial lipoarabinomannan biosynthesis. J Biol Chem. 20102851332636
139 - H. L Birch, L. J Alderwick, A Bhatt, D Rittmann, K Krumbach, A Singh, et alBiosynthesis of mycobacterial arabinogalactan: identification of a novel alpha(1-->3) arabinofuranosyltransferase. Mol Microbiol. 2008691191206
140 - L. J Alderwick, G. S Lloyd, H Ghadbane, J. W May, A Bhatt, L Eggeling, et alThe C-terminal domain of the arabinosyltransferase Mycobacterium tuberculosis EmbC is a lectin-like carbohydrate binding module. PLoS Pathog. 2011e1001299.
141 - H Skovierova, G Larrouy-maumus, J Zhang, D Kaur, N Barilone, J Kordulakova, et alAftD, a novel essential arabinofuranosyltransferase from mycobacteria. Glycobiology. 200919123547
142 - N Zhang, J. B Torrelles, M. R Mcneil, V. E Escuyer, K. H Khoo, P. J Brennan, et alThe Emb proteins of mycobacteria direct arabinosylation of lipoarabinomannan and arabinogalactan via an N-terminal recognition region and a C-terminal synthetic region. Mol Microbiol. 2003506976
143 - D Chatterjee, S. W Hunter, M Mcneil, P. J Brennan, Lipoarabinomannan. Multiglycosylated form of the mycobacterial mannosylphosphatidylinositols. J Biol Chem. 1992267622833
144 - B. J Appelmelk, den Dunnen J, Driessen NN, Ummels R, Pak M, Nigou J, et al. The mannose cap of mycobacterial lipoarabinomannan does not dominate the Mycobacterium-host interaction. Cell Microbiol. 20081093044
145 - K. H Khoo, A Dell, H. R Morris, P. J Brennan, D Chatterjee, Inositol phosphate capping of the nonreducing termini of lipoarabinomannan from rapidly growing strains of Mycobacterium. J Biol Chem. 1995270123809
146 - A. K Mishra, C Klein, S. S Gurcha, L. J Alderwick, P Babu, P. G Hitchen, et alStructural characterization and functional properties of a novel lipomannan variant isolated from a Corynebacterium glutamicum pimB’ mutant. Antonie Van Leeuwenhoek. 20089427787
147 - R. V Tatituri, L. J Alderwick, A. K Mishra, J Nigou, M Gilleron, K Krumbach, et alStructural characterization of a partially arabinosylated lipoarabinomannan variant isolated from a Corynebacterium glutamicum ubiA mutant. Microbiology. 200715326219
148 - B. A Wolucka, M. R Mcneil, E De Hoffmann, T Chojnacki, P. J Brennan, Recognition of the lipid intermediate for arabinogalactan/arabinomannan biosynthesis and its relation to the mode of action of ethambutol on mycobacteria. J Biol Chem. 19942692332835
149 - K Takayama, H. K Schnoes, E. J Semmler, Characterization of the alkali-stable mannophospholipids of Mycobacterium smegmatis. Biochim Biophys Acta. 197331621221
150 - A. M Bailey, S Mahapatra, P. J Brennan, D. C Crick, Identification, cloning, purification, and enzymatic characterization of Mycobacterium tuberculosis 1-deoxy-d-xylulose 5-phosphate synthase. Glycobiology. 20021281320
151 - R. K Dhiman, M. L Schaeffer, A. M Bailey, C. A Testa, H Scherman, D. C Crick, Deoxy-d-xylulose 5-phosphate reductoisomerase (IspC) from Mycobacterium tuberculosis: towards understanding mycobacterial resistance to fosmidomycin. J Bacteriol. 20051878395402
152 - A Argyrou, J. S Blanchard, Kinetic and chemical mechanism of Mycobacterium tuberculosis 1-deoxy-deoxy-d-xylulose-5-phosphate isomeroreductase. Biochemistry. 200443437584
153 - K Takayama, D. S Goldman, Enzymatic synthesis of mannosyl-1-phosphoryl-decaprenol by a cell-free system of Mycobacterium tuberculosis. J Biol Chem. 197024562517
154 - B. A Wolucka, E De Hoffmann, The presence of beta-d-ribosyl-1-monophosphodecaprenol in mycobacteria. J Biol Chem. 1995270201515
155 - B. A Wolucka, E De Hoffmann, Isolation and characterization of the major form of polyprenyl-phospho-mannose from Mycobacterium smegmatis. Glycobiology. 1998895562
156 - A. R Baulard, S. S Gurcha, J Engohang-ndong, K Gouffi, C Locht, G. S Besra, In vivo interaction between the polyprenol phosphate mannose synthase Ppm1 and the integral membrane protein Ppm2 from Mycobacterium smegmatis revealed by a bacterial two-hybrid system. J Biol Chem. 200327822428
157 - H Scherman, D Kaur, H Pham, H Skovierova, M Jackson, P. J Brennan, Identification of a polyprenylphosphomannosyl synthase involved in the synthesis of mycobacterial mannosides. J Bacteriol. 2009191676972
158 - H Skovierova, G Larrouy-maumus, H Pham, M Belanova, N Barilone, A Dasgupta, et alBiosynthetic origin of the galactosamine substituent of arabinogalactan in Mycobacterium tuberculosis. J Biol Chem. 20102854134855
159 - J. S Klutts, K Hatanaka, Y. T Pan, A. D Elbein, Biosynthesis of d-arabinose in Mycobacterium smegmatis: specific labeling from d-glucose. Arch Biochem Biophys. 200239822939
160 - M Scherman, A Weston, K Duncan, A Whittington, R Upton, L Deng, et alBiosynthetic origin of mycobacterial cell wall arabinosyl residues. J Bacteriol. 1995177712530
161 - M. S Scherman, L Kalbe-bournonville, D Bush, Y Xin, L Deng, M Mcneil, Polyprenylphosphate-pentoses in mycobacteria are synthesized from 5-phosphoribose pyrophosphate. J Biol Chem. 1996271296528
162 - H Huang, M. S Scherman, D Haeze, W Vereecke, D Holsters, M Crick, DC, et al. Identification and active expression of the Mycobacterium tuberculosis gene encoding 5-phospho-d-ribose-1-diphosphate: decaprenyl-phosphate 5-phosphoribosyltransferase, the first enzyme committed to decaprenylphosphoryl-d-arabinose synthesis. J Biol Chem. 20052802453943
163 - K Mikusova, H Huang, T Yagi, M Holsters, D Vereecke, D Haeze, W, et al. Decaprenylphosphoryl arabinofuranose, the donor of the d-arabinofuranosyl residues of mycobacterial arabinan, is formed via a two-step epimerization of decaprenylphosphoryl ribose. J Bacteriol. 200518780205
164 - T Christophe, M Jackson, H. K Jeon, D Fenistein, M Contreras-dominguez, J Kim, et alHigh content screening identifies decaprenyl-phosphoribose 2’ epimerase as a target for intracellular antimycobacterial inhibitors. PLoS Pathog. 2009e1000645.
165 - V Makarov, G Manina, K Mikusova, U Mollmann, O Ryabova, B Saint-joanis, et alBenzothiazinones kill Mycobacterium tuberculosis by blocking arabinan synthesis. Science. 20093248014
166 - V Makarov, O. B Riabova, A Yuschenko, N Urlyapova, A Daudova, P. F Zipfel, et alSynthesis and antileprosy activity of some dialkyldithiocarbamates. J Antimicrob Chemother. 20065711348
167 - M. R Pasca, G Degiacomi, A. L Ribeiro, F Zara, P De Mori, B Heym, et alClinical isolates of Mycobacterium tuberculosis in four European hospitals are uniformly susceptible to benzothiazinones. Antimicrob Agents Chemother. 20105416168
168 - P. K Crellin, R Brammananth, R. L Coppel, Decaprenylphosphoryl-beta- d-ribose 2’-epimerase, the target of benzothiazinones and dinitrobenzamides, is an essential enzyme in Mycobacterium smegmatis. PLoS One. 2011e16869.
169 - G Manina, M. R Pasca, S Buroni, E De Rossi, G Riccardi, Decaprenylphosphoryl-beta- d-ribose 2’-epimerase from Mycobacterium tuberculosis is a magic drug target. Curr Med Chem. 2010173099108
170 - S. M Batt, T Jabeen, V Bhowruth, L Quill, P. A Lund, L Eggeling, et alStructural basis of inhibition of Mycobacterium tuberculosis DprE1 by benzothiazinone inhibitors. Proc Natl Acad Sci U S A. 2012109113549
171 - I Tarnok, Z Tarnok, Carotenes and xanthophylls in mycobacteria. II. Lycopene, alpha- and beta-carotene and xanthophyll in mycobacterial pigments. Tubercle. 19715212735
172 - M Houssaini-iraqui, M. H Lazraq, S Clavel-seres, N Rastogi, H. L David, Cloning and expression of Mycobacterium aurum carotenogenesis genes in Mycobacterium smegmatis. FEMS Microbiol Lett. 19926923944
173 - L Ramakrishnan, H. T Tran, N. A Federspiel, S Falkow, A crtB homolog essential for photochromogenicity in Mycobacterium marinum: isolation, characterization, and gene disruption via homologous recombination. J Bacteriol. 199717958628
174 - M Viveiros, P Krubasik, G Sandmann, M Houssaini-iraqui, Structural and functional analysis of the gene cluster encoding carotenoid biosynthesis in Mycobacterium aurum A+. FEMS Microbiol Lett. 200018795101
175 - J. A Robledo, A. M Murillo, F Rouzaud, Physiological role and potential clinical interest of mycobacterial pigments. IUBMB Life. 201163718
176 - S. W Kim, J. D Keasling, Metabolic engineering of the nonmevalonate isopentenyl diphosphate synthesis pathway in Escherichia coli enhances lycopene production. Biotechnol Bioeng. 20017240815
177 - M Ito, Y Yamano, C Tode, A Wada, Carotenoid synthesis: retrospect and recent progress. Arch Biochem Biophys. 20094832248
178 - N Misawa, Y Satomi, K Kondo, A Yokoyama, S Kajiwara, T Saito, et alStructure and functional analysis of a marine bacterial carotenoid biosynthesis gene cluster and astaxanthin biosynthetic pathway proposed at the gene level. J Bacteriol. 1995177657584
179 - M. R Wenk, The emerging field of lipidomics. Nat Rev Drug Discov. 20054594610
180 - M. R Wenk, Lipidomics: new tools and applications. Cell. 201014388895
181 - E Layre, L Sweet, S Hong, C. A Madigan, D Desjardins, D. C Young, et alA comparative lipidomics platform for chemotaxonomic analysis of Mycobacterium tuberculosis. Chem Biol. 201118153749
182 - M. J Sartain, D. L Dick, C. D Rithner, D. C Crick, J. T Belisle, Lipidomic analyses of Mycobacterium tuberculosis based on accurate mass measurements and the novel "Mtb LipidDB". J Lipid Res. 20115286172
183 - C. A Madigan, T. Y Cheng, E Layre, D. C Young, M. J Mcconnell, C. A Debono, et alLipidomic discovery of deoxysiderophores reveals a revised mycobactin biosynthesis pathway in Mycobacterium tuberculosis. Proc Natl Acad Sci U S A. 2012109125762