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The thick waxy coat of mycobacteria, a protective layer against antibiotics and the host's immune system

Sarah M. Batt, David E. Minnikin, Gurdyal S. Besra

Biochemical Journal · 2020 · Review · CC BY

doi:10.1042/bcj20200194

Abstract

Tuberculosis, caused by the pathogenic bacterium Mycobacterium tuberculosis (Mtb), is the leading cause of death from an infectious disease, with a mortality rate of over a million people per year. This pathogen's remarkable resilience and infectivity is largely due to its unique waxy cell envelope, 40% of which comprises complex lipids. Therefore, an understanding of the structure and function of the cell wall lipids is of huge indirect clinical significance. This review provides a synopsis of the cell envelope and the major lipids contained within, including structure, biosynthesis and roles in pathogenesis.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 26, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

Mycobacterium tuberculosis (Mtb), the causative agent of tuberculosis, was responsible for 1.5 million fatalities worldwide in 2018. Added to this are the growing concerns of a surge in strains resistant to front-line drugs (multidrug resistant; MDR) and those additionally resistant to second-line drugs (extremely drug resistant; XDR) and all known drugs (totally drug resistant; TDR). The distinctive cell envelope of Mtb is robust and waxy, thanks to the rich variety of lipids present, which bestow on this pathogen resilience, a hydrophobic barrier to antibiotic entry and key virulence factors. In view of these facts and since many of the existing antibiotics available for tuberculosis target the synthesis and transport of these complex lipids, an understanding of lipid structures and synthesis in Mtb is a major goal of current research, with promise of novel effective drug targets.

Neither fully Gram-negative nor Gram-positive, the envelope of mycobacteria (Figure 1) is proposed to adopt a remarkable dual membrane structure, with a specialised mycobacterial outer membrane or ‘MOM’, formed as an inner layer of mycolic acids (MAs) and an outer leaflet of free lipids. While the free lipids are not attached and can be stripped from the envelope, the MA layer is covalently linked to the polysaccharide cell wall, a feature not seen in Gram-negative bacteria. This connection forms part of the mycoloylarabinogalactan–peptidoglycan complex (mAGP), consisting of branched arabinose domains esterified on the outer surface with MAs and linked to the peptidoglycan via the galactan helices of the arabinogalactan (AG). The periplasmic space of Mtb is sizeable, allowing for enzymatic reactions to take place and for the material to be processed and translocated to the MOM. The inner membrane of Mtb was initially assumed to be conventional, though lipid studies have since revealed that the phospholipid composition is predominantly phosphatidyl-myo-inositol mannosides (PIMs). More than half is present as PIM2 (with two mannose sugars on the inositol moiety), which appears to be exclusively within the inner leaflet of the inner membrane; while PIM6 (six mannose residues) is thought to occupy the outer leaflet. This unusual arrangement increases the content of fatty acid chains in the inner membrane as Ac1/Ac2PIM2 and Ac1/Ac2PIM6, each has three or four (Ac1 or Ac2) acyl chains. The subsequent tight packing of these chains could lead to a less fluid, more stable membrane with reduced permeability to drugs, coining a new term mycobacterial inner membrane or ‘MIM’. The lipoglycans, lipomannan (LM) and lipoarabinomannan (LAM) have lipid anchors based on PIMs and probably project into the periplasm from the MIM. Some models of the Mtb cell envelope depict a capsule as the outermost layer beyond the MOM, which consists mainly of polysaccharides and proteins, with only small quantities of lipids. This has been observed using cryo-electron microscopy to have a depth of ∼30 nm, but only in cultures grown under optimal conditions.

Mycolic acids

MAs are an abundant feature of the cell envelope of mycobacteria, representing 30% of the dry weight; they provide a hydrophobic permeability barrier to antibiotics and are essential for viability and pathogenesis. Characterised as long-chain α-alkyl-β-hydroxy fatty acids (70–90 carbons), most are covalently bound to the AG, forming the inner leaflet of the MOM. While two-thirds of the AG termini are esterified with MAs, some MAs are present as extractable lipids in the outer leaflet of the MOM, mainly linked to trehalose as trehalose dimycolates (TDM) and trehalose monomycolates (TMM), or as free mycolates (Figure 2b). While these are thought to be mostly intermediates in the attachment of the MA to the AG, there is evidence to suggest that TDM, the so-called ‘cord-factor’ and most highly expressed glycolipid in Mtb, also has dedicated roles in pathogenicity through interactions with the C-type lectin receptor Mincle on the macrophage, which prevents phagosome acidification and promotes granuloma formation. Free mycolates are more abundant in latent phase cells and could also play a role in biofilm formation.

MA structure can be separated into two parts: the meromycolate moiety consists of a long-chain meroaldehyde (up to 56 carbons) and usually has two functional groups, whereas the α-branch is a shorter, saturated alkyl chain (24–26 carbons). There are three classes of MAs in Mtb: α-mycolates are the most abundant and are present with only cis-cyclopropane rings, while oxygenated methoxymycolates and ketomycolates both have subclasses with predominantly either cis- or trans-cyclopropane rings and an adjacent methyl branch (Figure 2a).

The MA chains are packed tightly into parallel arrays perpendicular to the surface of the cell envelope, which may be stabilised by hydrogen bonds between the β-OH groups. Within these arrays, the MAs fold into energetically favourable conformations, modulated by the cis-double bonds and the cyclopropane rings, which introduce kinks into the long chains; stabilising hydrophilic interactions are also formed through the oxygen functions. This enables the keto-MAs to adopt a ‘W-form’ conformation, in which four chains are packed together in parallel. Though the α-MAs and methoxy-MAs can also form a ‘W’ shape, they are more flexible, particularly the α-MAs, adopting more open forms at higher temperatures (Figure 2a). This tight packing provides the MA monolayer with appropriate rigidity, leading to a hydrophobic permeability barrier, which could be regulated by the MAs assuming altered conformations depending on their structure and the ambient temperature. Each mycobacterial species has a specific MA profile, suggesting that the different structural elements have an important biological role, both physiologically and in pathogenicity.

Free lipids

Phthiocerol dimycocerosate (PDIMs) and phenolic glycolipids (PGLs) (Figure 3a,b) are large ( ∼90 carbons), complex, hydrophobic molecules found in the outer leaflet of the MOM. The unusual lipid core consists of a long-chain β-diol (phthiocerol) esterified with two multimethyl branched long-chain fatty acids (mycocerosic acids). Whilst possessing a similar lipid core, PGLs differ through a phenolic residue that forms a link between this core and a chain of up to four saccharide units; the composition of the oligosaccharide is species-specific and confers antigenic properties. The stereochemistry of the diol units of PDIMs and PGLs also vary by species; M. marinum and M. ulecerans both have erythro stereochemistry, while all other strains of mycobacteria have the threo configuration. PDIMs are produced by all virulent strains of Mtb and have important roles in pathogenicity: forming an impermeable barrier on the outer surface of the cell, masking the cell from the host's immune system and lysing the phagosome in order to escape the macrophage and disseminate. Interestingly, PDIMs give a negative advantage to growth in vitro, leading to their spontaneous loss, though they are never absent from clinical strains. This could be partly due to the hydrophobic barrier that PDIMs form, which though useful in the hostile environment of the host, also prevents the diffusion of useful solutes. PE/PPE proteins, a family named for their N-terminal repeated regions of proline-(proline)-glutamate, have recently been shown to compensate for this, forming pores that selectively uptake the necessary nutrients.

PGLs are less prevalent in Mtb isolates, though are present in the Mtb Canetti strain, M. leprae, M. marinum and M. bovis, and could be responsible for the hyper-virulence and drug resistance of the W-Beijing family of Mtb. PGLs affect virulence by suppressing the release of cytokines involved in the inflammatory response of the immune system. In M. leprae, PGLs are also implicated in the Schwann cell invasion and nerve demyelination. The presence of PGLs is directed by a single fused polyketide synthase (PKS), Pks15/1, which is knocked out in strains devoid of PGLs by an insertion that splits the pks15/1 gene in two. PKS proteins are required to condense, extend and modify nascent fatty acids; this particular PKS, Pks15/1, is proposed to extend from the phenol group, producing an intermediate in the synthesis of the phenolphthiocerol moiety, p-hydroxyphenylalkanoate (PHPA). This is extended by the PpsABCDE PKS proteins to phenolphthiocerol. Mycocerosic acid synthase (MAS), a PKS located in the same cluster as the pps operon, produces the methyl-branched mycocerosic acids. The latter two PKS systems are responsible for the synthesis of the lipid core common to PDIMs and PGLs.

Acyl trehaloses, trehalose glycolipids (SGLs) acylated with multimethyl branched fatty acids, are present in the MOM of all virulent strains of mycobacteria. As such, they are important in virulence with a role in blocking phagosome maturation and suppressing the host's immune system by inhibiting T cell proliferation and down-regulating cytokine secretion in activated monocytes. The core structure consists of a central trehalose moiety that is esterified on the 2-position with a saturated fatty acid (C16 or C18); the trehalose is additionally esterified with as many as four multimethyl branched long-chain fatty acids. The simplest forms are the diacyl trehaloses (DATs) (Figure 3c), which are based on mycosanoic (C24) and mycolipanolic (C27) acids. Pentaacyl trehaloses (PATs) (Figure 3d) contain mycolipenates (C27), whereas sulphated SGLs are made up of the longer fatty acids, phthioceranates (C37) and hydroxyphthioceranates (C40), and the trehalose moiety is sulphated at the 2′-position (Figure 3e).

Lipooligosaccharides (LOS) are extremely polar lipids, which are associated with biofilm formation and motility. They consist of a common trehalose-containing saccharide core, acylated on the trehalose moiety with multimethyl branched fatty acids (Figure 3f). The trehalose is further glycosylated with a mono-, or more commonly, an oligosaccharide of 2–6 sugars, which is species-specific. LOS are lacking in many Mtb strains, though are present in Mtb Canetti strain, M. kansasii, M. gordonae, M. smegmatis, M. marinum and a member of the M. fortuitum complex, later classified as M. houstonense. Those strains that do produce LOS are characterised by a smooth appearance, whereas those lacking these glycolipids are rough and have increased hydrophobicity over the outer membrane. LOS synthesis requires the presence of two proximal pks5 gene homologues flanking an associated pap (PKS associated protein) gene. Strains lacking LOS expression appear to have undergone a recombination-deletion event between the two pks5 homologues, losing a pks5-pap segment. Studies have demonstrated that LOS reduces virulence; smooth M. kanasii was unable to establish the chronic infection in the lungs of mice that their rough counterparts could. This could be due to the observed interference by LOS of host-pathogen interactions, perhaps by covering up critical glycolipids, and it has been speculated that this has driven their loss in virulent strains. The associated hydrophobicity detected in mycobacteria lacking LOS may also have the added benefit to the pathogen of enhancing aerosol transmission.

Glycopeptidolipids (GPLs), present in the outer leaflet of non-tuberculosis causing mycobacteria, consist of a lipopeptidyl core, glycosylated on the peptide unit. They are important in biofilm formation, aggregation, motility, cell wall integrity and interactions with the host's immune system. Interestingly, strains producing smaller amounts of GPL are more invasive, denoting a role in pathogenicity. C-type GPLs (Figure 3g), which form a sub-group that are alkaline-stabile, are found in M. smegmatis, M. avium, M. chelonae and M. abscessus; their general structure consists of a core tripeptide-amino alcohol (d-Phe-d-allo-Thr-D-Ala-l-alaninol), esterified with a long fatty acyl chain (C28). The peptide is glycosylated with a 6-deoxytalose on the allo-threonine and a rhamnose unit linked to the alaninol. This basic core can be further acylated, methylated and glycosylated, depending on the strain, producing different serotypes. The other group of GPLs, found in M. xenopi, are serine-containing and alkaline-labile, consisting of a tetrapeptide core with two serines (l-Ser-l-Ser-l-Phe-d-allo-Thr), esterified with a simpler fatty acyl chain (C12) (Figure 3h). The N-terminal serine residue is methylated, acting as the primary glycosylation site for a 6-deoxytalose unit; a di/tetrasaccharide of rhamnose (terminating in a glucose for the tetrasaccharide) is attached to the allo-threonine.

Fatty acid and mycolic acid biosynthesis

The biosynthesis of fatty acids and MAs is carried out by two fatty acid synthases, FAS-I and FAS-II (Figure 4). FAS-I is a multi-domain ‘eukaryotic-like’ fatty acid synthase, a single large protein comprising all the catalytic domains required for fatty acid synthesis. Whereas FAS-II is a ‘prokaryotic-like’ fatty acid synthase, a large complex of discrete enzymes. FAS-I is the only system capable of synthesising fatty acids de novo and uniquely to mycobacteria, the synthesis is bimodal, generating fatty acids with two different chain lengths: C24–C26 (corresponds to the α-branch in MA synthesis) and C16–C18 (extended by FAS-II to produce the meromycolate chain in MA synthesis). These fatty acids are either shuttled to FAS-II for MA production or to other PKS systems for the further extensions and modifications required to manufacture the various complex lipids.

Both FASs extend the nascent acyl chain through sequential additions of acetate (2 carbons) from the extender unit malonyl-Coenzyme A (CoA), though FAS-II requires the malonate to be presented on an acyl carrier protein (AcpM; Rv2244). The first step in fatty acid synthesis is the production of this unit through the carboxylation of acetyl-CoA, catalysed by the acyl-CoA carboxylase complex (ACCase). FabD (Rv2243), a transacylase, next generates the malonyl-AcpM extender unit required by FAS-II, transferring the malonyl group from the phosphopantetheine group of CoA to the same prosthetic group present on the active site serine of the AcpM.

The FAS-I gene present in Mtb (fas; Rv2524c) contains seven domains for fatty acid synthesis: acyl transferase (AT), enoyl reductase (ER), dehydratase (DH), malonyl/palmitoyl transferase (MPT), acyl carrier protein (ACP), β-keto reductase (KR) and β-ketoacyl synthase (KS). Synthesis is initiated with an acetyl group, which is transferred from the CoA to the ACP domain. During synthesis, the growing acyl chain remains bound to this domain.

In the FAS-II system, the discrete enzymes have corresponding functions to the domains of FAS-I. The first step of synthesis in this system is an important link between FAS-I and FAS-II. FabH (Rv0533c), a β-ketoacyl ACP synthase, plays a pivotal role in initiating FAS-II synthesis through the Claisen condensation of the shorter product from FAS-I (acyl-CoA, C16–18) with a malonyl-AcpM. This forms a β-ketoacyl-AcpM, which can then be presented to the FAS-II enzymes, commencing a round of modifications: reduction in the keto group by the keto reductase (MabA; Rv1483), dehydration of the β-hydroxylacyl-AcpM intermediate by the dehydratase heterodimers (HadAB and HadBC; Rv0635–Rv0636 and Rv0636–Rv0637), and finally reduction in the enoyl-AcpM to acyl-AcpM by the enoyl reductase (InhA; Rv1484). After the first cycle, further rounds are initiated by KasA/B (Rv2245 and Rv2246), ketosynthases which then elongate the acyl chain by the addition of two carbons each cycle, forming a β-ketoacyl-AcpM intermediate. The completed Acyl-AcpM undergoes further rounds of elongations, desaturations and also modifications, including cis-/trans-cyclopropanation and methoxy/keto group additions.

How the FAS systems regulate the length of the growing acyl chains is not clear. Though not essential, mutations in kasB result in MAs shorter by 2–4 carbons, suggesting that KasB is required for full-length extensions. The HadBC heterodimer is also associated with later elongation cycles and is similarly expected to play a role in regulating the acyl chain length in FAS-II. The mechanism by which FAS-I is able to synthesise two products with different chain lengths (C16–C18 and C24–C26) is likely to be a complex interplay between products and enzymes. Replacing the M. smegmatis FAS-I system with the Mtb homologue, did not alter the length of the longer fatty acid chains produced in M. smegmatis from the usual C24 to the C26 produced by Mtb; this indicates that regulation is not restricted to FAS-I and may involve interactions between FabH and FAS-II with FAS-I. Endogenous polysaccharides also affect FAS-I product formation, favouring the synthesis of shorter chain fatty acids (C16–C18) as well as increasing the overall rate of synthesis; these polysaccharides, containing 3-O-methyl-d-mannose or 6-O-methyl-d-glucose, are proposed to enhance product release, a rate-limiting step in the synthesis, by forming a complex specifically with the shorter length product and thereby facilitating its release. In the absence of these polysaccharides, the synthesis of the longer fatty acid chain is favoured (C24–C26), which as longer, shows an increased ability to self-interact, forming stable aggregates that also enable product release from FAS-I.

Polyketide synthases (PKS)

The mycobacterial genome boasts a large number of PKS, fundamental in the synthesis of the cell envelope's complex lipids, performing condensations, extensions and modifications of fatty acid chains (Figure 5a). There are three types of PKS systems, with distinct organisations and mechanisms. Multifunctional type I PKSs are large polypeptides with several catalytic domains. Pks12, for example, is the largest encoded polypeptide in Mtb, at 4151 amino acids. There are two varieties of type I PKSs: iterative systems cycle the product several times through the same domains in a mechanism resembling FAS-I, whereas modular PKSs possess a single domain for each catalytic step of the entire synthesis. Type II PKSs are similar to FAS-II, comprising a large complex of enzymes, each responsible for a catalytic step. There are also three type III plant-like PKSs present in the genome of Mtb, condensing enzymes from the chalcone synthase (CHS) superfamily. These enzymes are distinct from the other PKS classes, both in their structure, which is typically a homodimer of KS domains, and mechanism, acting on free acyl-CoA thioesters with no ACP involvement.

PKSs do not carry out de novo fatty acid synthesis, instead using fatty acid precursors from FAS-I. These are usually activated and loaded onto the ACP domain of the PKS by an associated FadD protein. The attached precursor is then modified by the catalytic domains of the PKS, cycling through successive extensions by the KS (β-ketoacyl synthase) domain, usually from a (methyl)malonyl-CoA extender unit via a Claisen-type condensation reaction; reduction in the keto group by the KR (β-keto reductase) domain; dehydration by the DH (dehydratase) domain and reduction in the enoyl group by the ER (enoyl reductase) domain. Some more complex lipids in mycobacteria require several PKSs to synthesise the different lipid components. For instance, in PGL synthesis, Pks15/1 synthesises the PHPA intermediate, PpsABCDE extends this to phenolphthiocerol and mycocerosic acid synthase (MAS), produces the methyl-branched mycocerosic acids that are attached to the diols.

Pks13, an iterative type I PKS, is involved in the synthesis of MAs, joining together the acyl-CoA (C24–26) product of FAS-I with the meromycolyl-AcpM product from FAS-II, via a Claisen-type condensation reaction (Figure 5b). In this process, both substrates are first attached covalently as thioesters to the ACP domains of Pks13. The meromycolyl-AcpM is activated by FadD32 (Rv3801c) to an AMP derivative and loaded on to the N-terminal ACP of Pks13; the acyl-CoA fatty acid is carboxylated, by an ACCase, and loaded onto the C-terminal ACP of Pks13, via the AT domain. The meromycolyl group is then transferred, in the presence of FadD32, to the KS domain of Pks13. The carboxylation step is essential for the Claisen-type reaction that follows, activating the α-carbon and enabling a decarboxylative condensation reaction. This proceeds via a nucleophilic attack by the acidic α-carbon of the carboxyl-acyl group towards the carbonyl group of the meromycolyl chain, releasing CO2 and the meromycolyl chain from the KS domain. The thioesterase (TE) domain of Pks13 next cleaves the α-alkyl β-ketoacyl product from the ACP domain, possibly forming a transient covalent bond with the product via a serine in the active site; the TE domain then catalyses the transfer of the acyl chain onto a trehalose, forming α-alkyl β-ketoacyl trehalose monomycolate (TMMk). The final mature TMM is formed when the keto group is reduced by CmrA (Rv2509).

Trehalose is a significant sugar in mycobacterial lipid production (for review see); in addition to its role in MA synthesis, trehalose constitutes the core of the polyacyl-trehaloses, sulfolipids and LOS. A disaccharide of α(1 → 1)-linked glucose, trehalose is well-known to protect against the stresses of heat, freezing, desiccation and γ-radiation. During MA synthesis, the trehalose could be acting to shield the extreme hydrophobicity of the very long fatty acid chains, enabling export and attachment to the AG. For this reason, it would seem unlikely that the free mycolates found in the MOM, particularly during biofilm formation, are released from Pks13 as free mycolates, but perhaps produced and transported as TMM, and the trehalose removed by enzymatic hydrolysis in the outer membrane. After the MA is attached to the AG by the mycolyltransferases of the antigen 85 complex (Ag85), the trehalose is recycled back into the cytoplasm by the ABC sugar transporter, LpqY-SugA-SugB-SugC (Rv1235–Rv1236–Rv1237–Rv1238); this transporter is essential for virulence and again demonstrates the importance of trehalose to the bacterium.

MmpL/S

MmpL (mycobacterial membrane proteins large) proteins belong to the RND (resistance, nodulation and cell division) superfamily of membrane proteins, which are typically characterised by 12-transmembrane (TM) helices and two large substrate-specific periplasmic loops between TM1–2 and TM7–8. The energy for their activity is derived from a proton gradient across the inner membrane and they usually function as antiporters, coupling the energy from proton movement into the cytoplasm, with substrate efflux. The Mtb genome encodes 13 MmpL proteins, many of which are found in the same loci as PKS proteins and other proteins involved in lipid synthesis. Though MmpL5 and MmpL7 have been shown to have roles in drug resistance and efflux, the primary function of the MmpL proteins is to export the complex lipids synthesised in the cytoplasm across the inner membrane (Figure 5c). Some MmpL proteins (1, 2, 4 and 5) have smaller, transcriptionally coupled accessory proteins, MmpS (mycobacterial membrane proteins small). Other MmpL proteins also require additional membrane proteins; MmpL8 interacts with the membrane protein, Sap and transport by MmpL7 involves the ABC transporter, DrrABC, and lipoprotein, LppX.

Knock-out studies have shown that MmpL3 (Rv0206c) is the only essential MmpL protein, though others are required for virulence. MmpL3 is responsible for transporting the MA precursor, TMM, across the inner membrane. Inhibitors that lead to TMM accumulation in the cytoplasm and simultaneous loss of TDM in the periplasm also selected for spontaneous resistance mutations in mmpL3, findings that have since been confirmed by a mmpL3 conditional mutant in M. smegmatis. Several inhibitors of MmpL3 have recently been found though spontaneous resistance based screening, though some of these are broad-spectrum, targeting bacterial and fungal species lacking MAs, leading to speculation that the real target could be elsewhere. The crystal structure of the homologue from M. smegmatis, however, has demonstrated that many of these inhibitors, including SQ109, do bind directly to MmpL3. The resolved structure consists of a central proton translocating channel, formed between 2 TM helices (TM4 and TM10) that intertwine and are linked by hydrogen-bonding between two sets of Asp-Tyr residues. These interactions, which are essential for proton relay, were disrupted by binding of the inhibitors, all of which co-crystallised in the central pore. While many RND transporters form homo-dimers or -trimers, and a homology model refined with electron microscopy (EM) suggested MmpL3 is a trimer, in the crystal structure MmpL3 was monomeric.

MmpL protein expression is controlled by transcriptional repressors, which are in turn regulated by the availability of fatty acids; MmpL3 is regulated by three TetR regulators: Rv3249c, Rv0302 and Rv1816, all of which dissociate from DNA upon binding palmitic acid, relieving transcriptional repression. Phosphorylation of the large C-terminal domain possessed by MmpL3 has also been implicated as a post-translational regulation system.

Phosphatidyl-myo-inositol mannosides (PIMs), lipomannan (LM) and lipoarabinomannan (LAM)

These lipoglycans (Figure 6) comprise a phosphatidyl-myo-inositol (PI) core decorated with mannose residues (Manp, pyranose ring form), with additional branches of arabinose (Araf, furanose ring) in LAM. Acyl chains on the inositol, glycerol and mannose moieties provide lipid anchors into the inner and possibly also the outer membranes, though the inner membrane is perhaps their principal location as LM and LAM possess the same lipid anchors as the PIMs, which share this location. They are ubiquitous in mycobacteria, with important roles in stability, permeability and cell division. LM and LAM are also important in virulence, modulating interactions with the immune system; LAM, in particular, plays a key role in the infection life cycle of Mtb, blocking phagosome maturation and therefore avoiding lysis and antigen presentation, enabling Mtb to avoid the host's defences whilst providing a habitable environment in which to replicate. Synthesis proceeds via PIM → LM → LAM, yet PIM2 and PIM6 are not only intermediates in the synthesis of LM and LAM, but end products in their own right, forming much of the cytoplasmic membrane, enhancing the stability and reducing drug permeability. In fact, PIM6 is unlikely to be intermediary, possessing two α(1 → 2) linked mannose residues not present in LM and LAM; instead PIM4 is the likely branch point between the two pathways. Structurally, in PIM2 (Figure 6a), the 2 and 6 positions of the myo-inositol ring each have a single linked mannose residue, whereas in PIM6 (Figure 6b), a chain of five mannose units resides at the 6-position in addition to the mannose at position 2. Both LM and LAM possess a core chain of 21–34 α(1 →6) linked mannose residues from position 6 of the myo-inositol ring, dotted with single α(1 → 2) linked mannose units (Figure 6c). LAM has an additional arabinan domain of highly branched arabinose residues, similar to AG (Figure 6d). The non-reducing termini of these arabinose branches can be capped mannose residues to form ManLAM.

The biosynthesis of PIMs, LM and LAM is compartmentalised, which is achieved by the presence of both soluble, membrane-associated glycosyltransferases (GTs) in the cytoplasm (from the GT-A/B superfamily) and membrane-bound GTs (from the GT-C superfamily) in the periplasm. This enables the initial glycosylation of the lower PIMs to occur within the cell; the lipid-anchored PI sits on the cytoplasmic face of the inner membrane, with the inositol moiety pointing inwards. Here, mannose is transferred from a GDP-Mannose sugar donor, initially with α(1 → 2) linkage to position 2 of the myo-inositol by PimA (Rv2610c), forming PIM1; and then with α(1 → 6) linkage to position 6 by PimB’ (Rv2188c), forming PIM2 (Figure 6a). An acyl chain (palmitate) is transferred by PatA (Rv2611c) to position 6 of the mannose added by PimA, forming Ac1PIM2, which has three acyl chains; position 3 of the myo-inositol ring can also be acylated by an unknown acyl transferase to Ac2PIM2, a tetra-acylated form, though LM and LAM are predominantly triacylated. PIM2 either remains in the inner leaflet of the cytoplasm where it forms the majority of the lipid content, or is further mannosylated on position 6 with α(1 → 6) linkages to PIM4. The GT(s) responsible for this has yet to be identified; PimC was found in the genome of Mtb CDC1551 as a possible candidate for the mannosylation to PIM3, though there is no homologue in some strains of mycobacteria (Mtb H37Rv and M. smegmatis) and a knock-out in M. bovis did not reduce the levels of PIMs, LM or LAM, indicating that another pathway must exist.

The biosynthesis of the higher PIMs, LM and LAM occurs outside of the cytoplasm, though it is not known whether the juncture for this is PIM2, PIM3 or PIM4. The PIM is flipped across the inner membrane so that the inositol and mannose moieties now point into the periplasm. A candidate for this transport is the putative ABC transporter, Rv1747, a knock-out of which had low levels of PIMs and showed reduced virulence. However, this mutant still had WT levels of LM and LAM, suggesting the presence of an additional transport mechanism. In the periplasm, membrane-bound GTs transfer further mannose units from the lipid-linked donor, polyprenyl-mannose. The formation of PIM6 from PIM4 (Figure 6b) involves the addition of two α(1 →2) linked mannose residues, the first or both of which are transferred by PimE (Rv1159). In LM and LAM biosynthesis, the α(1 → 6) linked mannose core is extended from PIM4 by MptB (Rv1459c), followed by MptA (Rv2174), an order determined by knock-outs. The deletion of MptA in C. glutamicum resulted in a truncated LM and halted LAM production, which was complemented by Rv2174 or the C. glutamicum homologue. The deletion of MptB in C. glutamicum completely abolished all LM and LAM synthesis, which was also successfully complemented. A similar knock-out of MptB in M. smegmatis, however, did not affect LM/LAM production, allowing for some redundancy in this pathway. The α(1 → 2) linked mannose residues that decorate the mannose core, producing mature LM, are added by MptC (Rv2181), a knock-out of which abolished all LM synthesis in M. smegmatis, though still produced a truncated version of LAM; the immature LM is speculated to be either degraded or processed into LAM. The regulation of the branch point of PIM4 to either PIM6 or LM is controlled by LpqW (Rv1166). Disruption of the corresponding gene in M. smegmatis reduced LM/LAM synthesis, producing an unstable mutant that restored LM/LAM levels through a spontaneous mutation disrupting pimE, which simultaneously abolished the dependent PIM6 pathway. A similar deletion in Corynebacterium glutamicum, which lacks a PimE homologue and higher PIMs, was stabilised by a point mutation in mptB. MptB, which is required for the initial mannosylation of the LM core, was subsequently shown to have reduced activity in the absence of LpqW and authors speculate that this mechanism could regulate MptB and subsequently LM synthesis. GDP could also be a regulator, it has been shown to inhibit early PIM synthesis and reduce levels of the polyprenyl-mannose substrate, perhaps by reversing the synthesis of this substrate to GDP-Mannose.

The arabinan domain of LAM is similar to that of arabinoagalactan (AG), sharing the same linkages and branched core of 18–22 arabinose units, though LAM is more variable in structure and is characterised by the presence of more linear chains at the reducing end. AG is terminated by a distinctive branched hexa-arabinoside unit (β-d-Araf(1 → 2)α-d-Araf]2-3,5-α-d-Araf-(1 → 5)-α-d-Araf), whereas LAM also has the linear tetra-arabinoside motif (Araf4; a chain of four α(1 → 5) linked Araf). This is unsurprising since both domains are constructed by a similar set of membrane-bound arabinosyltransferases, using the lipid-linked donor, decaprenylphosphoryl-d-arabinose (DPA). During synthesis, the mannan core of the LM is first primed, though the arabinosyltransferase responsible and the location of this attachment are not known; while the O-2 of the mannose was originally implicated, more recent studies suggest that the linkage is to the O-6 position, which would restrict the attachment of the arabinan domain to the very end of the mannan core. Clarification requires further analysis with longer chain saccharides. After the initial priming, EmbC (Rv3793) adds an arabinan core of 12–15 arabinose residues with α(1 → 5) linkage. AftC (Rv2673), which has α(1 → 3) activity, introduces 3,5-arabinose branches in both LAM and AG synthesis. AftD (Rv0236c) has been shown to have similar activity to AftC and may also introduce branch points, though α(1 → 5) activity in AG synthesis has also been demonstrated. EmbC further extends these branch points by up to 8–9 arabinose units with α(1 → 5) linkage. Finally, AftB (Rv3805c) adds the terminal arabinose residue, with β(1 → 2) linkage, to the non-reducing ends of the arabinan.

Another difference in the arabinan domains of LAM and AG is the nature of the reducing end. In AG, most are esterified with MAs, while LAM is capped with mannose, producing ManLAM. This mannose capping is species-specific: in Mtb and other slow-growing virulent strains, the LAM is capped with up to three linear α(1 → 2) linked mannose residues (Manp) on the 5-position of the terminal arabinose; in faster-growing mycobacteria, such as M. smegmatis, much of the LAM is uncapped, while a portion is capped with phosphoinositide (PILAM); M. leprae has much reduced mannose capping and M. chelonae has none at all. Capping is initiated by CapA (Rv1635c), which adds the first mannose residue with α(1 → 5) linkage. Further residues are next attached by MptC, the mannosyltransferase responsible for adding the α(1 → 2) mannose to the mannan core.

Cell surface labelling, using either biotin followed by lipid extraction or antibodies and cryo-electron microscopy, has shown the presence of arabinan mannan (AM; non-acylated LAM), and also PIMs, LM and LAM, on the surface of the mycobacterial envelope; the PIMs and LM/LAM have lipid anchors and are thought to sit within the lipids of the MOM, projecting into the capsule, while the AM lacks this anchor and is free within the capsular layer. However, this issue is debateable and not always reproducible; a study using atomic force microscopy with immunogold labelling, for example, was only able to demonstrate the presence of surface-exposed LAM after treatment with drugs that target the cell wall. Also, if PIMs, LM and LAM do indeed locate to the MOM, a yet undiscovered transport system would be required to extract these lipoglycans from their inner membrane anchoring and then relocate them across the periplasmic space and through the lipids of the MOM, a system hypothesised to be similar to the one used by Escherichia coli in the transport of LPS. The presence of PIMs, LM and LAM in the MOM would certainly be reassuring, since surface exposure is essential for their observed roles in infection and interactions with the host's immune system, especially since the PIMs, LM and LAM located in the inner membrane were proven not to be surface-exposed. Studies with purified ManLAM and PILAM demonstrated that the mannose cap blocks the fusion of the phagosome and expression of pro-inflammatory cytokines, though these results were not replicated with live bacteria deleted of capA and lacking mannose caps. Similar mutants did, however, demonstrate that ManLAM is recognised by Dectin-2, initiating the expression of pro-inflammatory cytokines. LM has also been shown to activate the Toll-like receptor 2 (TLR2), triggering the innate immune system. This response is proportional to the chain length of the mannan core but is reduced with LAM, possibly because the mannan core is obscured by the arabinan domain; truncations of the arabinan domain of LAM restored this activation. While initial stimulation of the TLR2 activates the immune response, prolonged stimulation in macrophages has been shown to have the opposite effect, preventing antigen presentation by major histocompatibility complex class II molecules (MHC). This could be a crucial part of the mechanism employed by Mtb to evade the host's immune system and establish a latent phase infection in macrophages.

Mycobacterial antigens and CD1 presentation

For many years, it had been assumed that proteins are the sole antigens, activating T cells via the T cell receptor (TCR), and lipids were limited to stimulating the innate immune system. More recently, it has been recognised that many mycobacterial lipids, including mycolates, PIM, LAM and phosphatidylglycerol (PG), activate T cells through the CD1 (cluster of differentiation molecules) presentation pathway (Table 1). CD1 proteins are similar to MHC molecules, which present peptide antigens on antigen-presenting cells (APCs) to conventional T cells via the TCR, though CD1 presentation is limited to CD1-restricted T cells. There are five human CD1 proteins: CD1a, -b, -c and -d are membrane proteins involved in presenting lipids and are expressed on APCs, such as B cells, Langerhans cells and dendritic cells as well as cortical thymocytes. CD1e is a soluble protein that processes the glycolipids in order to increase the versatility of the CD1 proteins. CD1e functions by binding directly to the glycolipids and aiding their enzymatic digestion, for example by α-mannosidase in PIM6 presentation by CD1b.

The expression of the CD1 proteins is regulated by the TLR, via interleukin-1β, a pathway that is triggered upon Mtb infection. The lipid antigen binds to CD1 either directly at the cell surface or within the endosome, after the antigen or whole bacteria has been phagocytosed into the cell, a mechanism that enables additional processing by CD1e. If binding occurs in the endosome, the CD1-lipid complex is then trafficked to the cell surface in order to present the antigen to the TCR of CD1-restricted T cells. CD1b and CD1c are better adapted to binding to lipids within endosomal compartments, whereas CD1a is more suited to binding to antigens at the cell surface. Studies have shown that there are higher levels of CD1-restricted T cells present in individuals exposed to Mtb and their activation by CD1 presenting molecules is thought to function in the host defences. The T cell response to this activation is not known in humans, though in vitro effector mechanisms have been demonstrated to include the secretion of alpha and gamma interferon and granulysin production, which results in lysis of infected cells and macrophage activation. CD1d is recognised by CD1d-restricted natural killer (NK) T cells, which activate both the innate and the adaptive immune system, with responses that include proliferation and cytokine secretion, including gamma interferon and interleukin-4. Invariant NKT cells are also important in granuloma formation and are recruited to the site by lipid antigens, though the evidence suggests that CD1d is not involved in their presentation.

Conclusions

Mtb commands a complex array of lipids, many of which are crucial to its pathogenicity and survival, with roles in stability, drug permeability, cell division, biofilm formation, infection and host interactions. Many of the lipids present, such as the MAs and the lipid anchors of the PIMs, are packed together tightly in parallel arrangements, a feature that greatly enhances structural stability, while providing a generous hydrophobic barrier that reduces permeability to drugs. Free lipids provide yet more hydrophobic coating, constituting the outer layer of the MOM, an unconventional second membrane. The distinctiveness of the Mtb cell envelope does not end here as the bulk of the complex lipids present form part of much larger complexes, mAGP and LM/LAM, structures essential for viability that are unique to the Corynebacterium genus.

While it might be easy to consider the cell envelope to be a rigid, unchanging entity, it is in fact dynamic. For instance, the conformation of the MAs is temperature dependent. Also the lipid content of the envelope varies according to the infectivity cycle (for review see). PIMs are up-regulated during early infection, while LM and LAM are more likely to predominate during latent phase; the high levels of LAM expressed in the granulomas may guard against the host's defences, masking the mannan core. Prolonged stimulation of the TLR2 by LM could also assist in establishing and maintaining the granulomas, by inhibiting antigen presentation in the macrophages and thus evading the host's immune response. MAs, as an essential component of the cell wall, are synthesised in line with growth, though their structural composition is thought to vary throughout. The TDM and TMM levels, however, decrease in stationary phase, accompanied by an increase in the levels of free mycolates, which could be an initial step in biofilm formation. In addition, PDIM is highly synthesised in actively growing cells, but not in stasis, correlating with the crucial role that it plays during active infection.

Many of the proteins involved in lipid synthesis and transport are essential and as such represent ideal drug targets. Indeed, many the current antibiotics already inhibit some of the enzymes involved, including isoniazid (INH) and ethionamide (ETH), both of which target InhA, abolishing MA biosynthesis; ethambutol inhibits the arabinosyltransferases involved in assembling the branched arabinan domains of both LAM and AG, blocking MA attachment to the latter; the recently approved second-line drugs delamanid and pretomanid also affect MA synthesis, though the exact targets are not known at present; and not to mention the plethora of compounds in the pipe-line that look to be inhibiting MA transport by MmpL3. Future research into the area would benefit by studying the MmpL proteins, key lipid transporters present in the inner membrane, one of which is essential for survival (MmpL3) and many more are required for pathogenicity. Searching for the enzymes missing from some of the biosynthesis pathways could also prove fruitful, as would structural data and in vitro assay development for all appropriate targets. Today much of the current drug discovery works backwards from the drug to the target; though in the future, as our knowledge of this organism grows and enzyme assays are developed, screening could become increasingly tailored to the target. The cell envelope of Mtb is truly unique, contributing to its triumph as the most successful pathogen of our time. An understanding of its structure and biosynthesis, therefore, is crucial in the search for new drugs to tackle the ever-growing tide of resistance.

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    Besra G.S., Minnikin D.E., Sharif A. and Stanford J.L. (1990 ) Characteristic new members of the phthiocerol and phenolphtiocerol families from Mycorbacterium ulcerans. FEMS Microbiol. Lett. 66, 11–13

    Not in our library10.1111/j.1574-6968.1990.tb03964.x

  56. 56.

    Maskens K., Minnikin D.E. and Polgar N. (1966 ) Studies relating to phthiocerol. Part VI. Stereochemical studies. J. Chem. Soc. Chem. Commun., 2113–2115

    Not in our library10.1039/j39660002107

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    Not in our library10.1074/jbc.m100662200

  58. 58.

    Wang Q., Boshoff H.I.M., Harrison J.R., Ray P.C., Green S.R., Wyatt P.G. et al. (2020 ) PE/PPE proteins mediate nutrient transport across the outer membrane of Mycobacterium tuberculosis. Science 367, 1147–1151

    Not in our library10.1126/science.aav5912

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    Murry J.P., Pandey A.K., Sassetti C.M. and Rubin E.J. (2009 ) Phthiocerol dimycocerosate transport is required for resisting interferon-γ-independent immunity. J. Infect. Dis. 200, 774–782

    Not in our library10.1086/605128

  60. 60.
    The Cell Wall Lipid PDIM Contributes to Phagosomal Escape and Host Cell Exit of Mycobacterium tuberculosis · 2017
    No notice
  61. 61.

    Domenech P. and Reed M.B. (2009 ) Rapid and spontaneous loss of phthiocerol dimycocerosate (PDIM) from Mycobacterium tuberculosis grown in vitro: implications for virulence studies. Microbiology 155, 3532–3543

    Not in our library10.1099/mic.0.029199-0

  62. 62.

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    Daffé M., Lacave C., Lanéelle M.-A. and Lanéelle G. (1987 ) Structure of the major triglycosyl phenol-phthiocerol of Mycobacterium tuberculosis (strain Canetti). Eur. J. Biochem. 167, 155–160

    Not in our library10.1111/j.1432-1033.1987.tb13317.x

  64. 64.

    Hunter S.W. and Brennan P.J. (1981 ) A novel phenolic glycolipid from Mycobacterium leprae possibly involved in immunogenicity and pathogenicity. J. Bacteriol. 147, 728–735

    Not in our library10.1128/jb.147.3.728-735.1981

  65. 65.

    Villé C. and Gastambide-Odier M. (1970 ) Le 3-O-méthyl-l-rhamnose, Sucre isolé du mycoside G de Mycobacterium marinum. Carbohydr. Res. 12, 97–107

    Not in our library10.1016/s0008-6215(00)80229-3

  66. 66.

    Jarnagin J.L., Brennan P.J. and Harris S.K. (1983 ) Rapid identification of Mycobacterium bovis by a thin-layer chromatographic technique. Am. J. Vet. Res. 44, 1920–1922 PMID:

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  67. 67.

    Ng V., Zanazzi G., Timpl R., Talts J.F., Salzer J.L., Brennan P.J. et al. (2000 ) Role of the cell wall phenolic glycolipid-1 in the peripheral nerve predilection of Mycobacterium leprae. Cell 103, 511–524

    Not in our library10.1016/s0092-8674(00)00142-2

  68. 68.

    Rambukkana A., Zanazzi G., Tapinos N. and Salzer J.L. (2002 ) Contact-dependent demyelination by Mycobacterium leprae in the absence of immune cells. Science 296, 927–931

    Not in our library10.1126/science.1067631

  69. 69.

    He W., Soll C.E., Chavadi S.S., Zhang G., Warren J.D. and Quadri L.E.N. (2009 ) Cooperation between a coenzyme A-independent stand-alone initiation module and an iterative type I polyketide synthase during synthesis of mycobacterial phenolic glycolipids. J. Am. Chem. Soc. 131, 16744–16750

    Not in our library10.1021/ja904792q

  70. 70.

    Azad A.K., Sirakova T.D., Fernandes N.D. and Kolattukudy P.E. (1997 ) Gene knockout reveals a novel gene cluster for the synthesis of a class of cell wall lipids unique to pathogenic mycobacteria. J. Biol. Chem. 272, 16741–16745

    Not in our library10.1074/jbc.272.27.16741

  71. 71.

    Trivedi O.A., Arora P., Vats A., Ansari M.Z., Tickoo R., Sridharan V. et al. (2005 ) Dissecting the mechanism and assembly of a complex virulence mycobacterial lipid. Mol. Cell 17, 631–643

    Not in our library10.1016/j.molcel.2005.02.009

  72. 72.

    Fujiwara N. (1997 ) [Distribution of antigenic glycolipids among Mycobacterium tuberculosis strains and their contribution to virulence]. Kekkaku 72, 193–205 PMID:

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  73. 73.

    Munoz M., Laneelle M.A., Luquin M., Torrelles J., Julian E., Ausina V. et al. (1997 ) Occurrence of an antigenic triacyl trehalose in clinical isolates and reference strains of Mycobacterium tuberculosis. FEMS Microbiol. Lett. 157, 251–259

    Not in our library10.1016/s0378-1097(97)00483-7

  74. 74.

    Ortalo-Magné A., Lemassu A., Lanéelle M.A., Bardou F., Silve G., Gounon P., Marchal G. and Daffé M. (1996 ) Identification of the surface-exposed lipids on the cell envelopes of Mycobacterium tuberculosis and other mycobacterial species. J. Bacteriol. 178, 456–461

    Not in our library10.1128/jb.178.2.456-461.1996

  75. 75.
    High Content Phenotypic Cell-Based Visual Screen Identifies Mycobacterium tuberculosis Acyltrehalose-Containing Glycolipids Involved in Phagosome Remodeling · 2010
    No notice
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    Lee K.-S., Dubey V.S., Kolattukudy P.E., Song C.-H., Shin A.-R., Jung S.-B., et al. (2007 ) Diacyltrehalose of Mycobacterium tuberculosis inhibits lipopolysaccharide- and mycobacteria-induced proinflammatory cytokine production in human monocytic cells. FEMS. Microbiol. Lett. 267, 121–128

    Not in our library10.1111/j.1574-6968.2006.00553.x

  77. 77.

    Saavedra R., Segura E., Leyva R., Esparza L.A. and López-Marı´n L.M. (2001 ) Mycobacterial Di-O-Acyl-Trehalose inhibits mitogen- and antigen-induced proliferation of murine T cells in vitro. Clin. Diagn. Lab. Immunol. 8, 1081

    Not in our library10.1128/cdli.8.6.1-91-1088.2001

  78. 78.

    Saavedra R., Segura E., Tenorio E.P. and Lopez-Marin L.M. (2006 ) Mycobacterial trehalose-containing glycolipid with immunomodulatory activity on human CD4+ and CD8+ T-cells. Microbes Infect. 8, 533–540

    Not in our library10.1016/j.micinf.2005.08.005

  79. 79.

    Besra G.S., Bolton R.C., McNeil M.R., Ridell M., Simpson K.E., Glushka J. et al. (1992 ) Structural elucidation of a novel family of acyltrehaloses from Mycobacterium tuberculosis. Biochemistry 31, 9832–9837

    Not in our library10.1021/bi00155a040

  80. 80.

    Bertozzi C. and Schelle M (2008 ) Sulfated Metabolites from M. tuberculosis: sulfolipid-1 and beyond In The Mycobacterial Cell Envelope (Daffe M. and Reyrat J.-M., eds), pp. 291–304, American Society for Microbiology, Washington, DC

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  81. 81.

    Goren M.B. and Brennan P.J. (1979 ) Mycobacterial lipids: chemistry and biologic activities In Tuberculosis (Youmans G.P., ed.), pp. 63–193, W.B. Saunders Company, Philadelphia, PA

    Not in our library

  82. 82.

    Ren H., Dover L.G., Islam S.T., Alexander D.C., Chen J.M., Besra G.S. et al. (2007 ) Identification of the lipooligosaccharide biosynthetic gene cluster from Mycobacterium marinum. Mol. Microbiol. 63, 1345–1359

    Not in our library10.1111/j.1365-2958.2007.05603.x

  83. 83.

    Hunter S.W., Jardine I., Yanagihara D.L. and Brennan P.J. (1985 ) Trehalose-containing lipooligosaccharides from mycobacteria: structures of the oligosaccharide segments and recognition of a unique N-acylkanosamine-containing epitope. Biochemistry 24, 2798–2805 PMID:

    Not in our library10.1021/bi00332a030

  84. 84.

    Gilleron M., Vercauteren J. and Puzo G. (1994 ) Lipo-oligosaccharidic antigen from Mycobacterium gastri. Complete structure of a novel C4-branched 3,6-dideoxy-.alpha.-xylo-hexopyranose. Biochemistry 33, 1930–1937

    Not in our library10.1021/bi00173a041

  85. 85.

    Daffe M., McNeil M. and Brennan P.J. (1991 ) Novel type-specific lipooligosaccharides from Mycobacterium tuberculosis. Biochemistry 30, 378–388

    Not in our library10.1021/bi00216a011

  86. 86.

    Hunter S.W., Murphy R.C., Clay K., Goren M.B. and Brennan P.J. (1983 ) Trehalose-containing lipooligosaccharides. A new class of species-specific antigens from Mycobacterium. J. Biol. Chem. 258, 10481–10487

    Not in our library

  87. 87.

    Saadat S. and Ballou C.E. (1983 ) Pyruvylated glycolipids from Mycobacterium smegmatis. Structures of two oligosaccharide components. J. Biol. Chem. 258, 1813–1818 PMID:

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  88. 88.

    Minnikin D.E., Ridell M., Wallerstrom G., Besra G.S., Parlett J.H., Bolton R.C. et al. (1989 ) Comparative studies of antigenic glycolipids of mycobacteria related to the leprosy bacillus. Acta Leprol. 7, 51–54 PMID:

    Not in our library

  89. 89.

    Burguière A., Hitchen P.G., Dover L.G., Kremer L., Ridell M., Alexander D.C. et al. (2005 ) Losa, a key glycosyltransferase involved in the biosynthesis of a novel family of glycosylated acyltrehalose lipooligosaccharides from Mycobacterium marinum. J. Biol. Chem. 280, 42124–42133

    Not in our library10.1074/jbc.m507500200

  90. 90.

    Besra G.S., McNeil M.R. and Brennan P.J. (1992 ) Characterization of the specific antigenicity of Mycobacterium fortuitum. Biochemistry 31, 6504–6509

    Not in our library10.1021/bi00143a021

  91. 91.

    Tsang A.Y., Barr V.L., McClatchy J.K., Goldberg M., Drupa I. and Brennan P.J. (1984 ) Antigenic relationships of the Mycobacterium fortuitum-Mycobacterium chelonae complex. Int. J. Syst. Bacteriol. 34, 35–44

    Not in our library10.1099/00207713-34-1-35

  92. 92.

    Belisle J.T. and Brennan P.J. (1989 ) Chemical basis of rough and smooth variation in mycobacteria. J. Bacteriol. 171, 3465–3470

    Not in our library10.1128/jb.171.6.3465-3470.1989

  93. 93.

    Jankute M., Nataraj V., Lee O.Y.-C., Wu H.H.T., Ridell M., Garton N.J. et al. (2017 ) The role of hydrophobicity in tuberculosis evolution and pathogenicity. Sci. Rep. 7, 1315

    Not in our library10.1038/s41598-017-01501-0

  94. 94.

    Boritsch E.C., Frigui W., Cascioferro A., Malaga W., Etienne G., Laval F., et al. (2016 ) pks5-recombination-mediated surface remodelling in Mycobacterium tuberculosis emergence. Nat. Microbiol. 1, 15019

    Not in our library10.1038/nmicrobiol.2015.19

  95. 95.

    Villeneuve C., Etienne G., Abadie V., Montrozier H., Bordier C., Laval F. et al. (2003 ) Surface-exposed glycopeptidolipids of Mycobacterium smegmatis specifically inhibit the phagocytosis of mycobacteria by human macrophages. identification of a novel family of glycopeptidolipids. J. Biol. Chem. 278, 51291–51300

    Not in our library10.1074/jbc.m306554200

  96. 96.

    Etienne G., Villeneuve C., Billman-Jacobe H., Astarie-Dequeker C., Dupont M.-A. and Daffe M. (2002 ) The impact of the absence of glycopeptidolipids on the ultrastructure, cell surface and cell wall properties, and phagocytosis of Mycobacterium smegmatis. Microbiology 148, 3089–3100

    Not in our library10.1099/00221287-148-10-3089

  97. 97.

    Recht J., Martinez A., Torello S. and Kolter R. (2000 ) Genetic analysis of sliding motility in Mycobacterium smegmatis. J. Bacteriol. 182, 4348–4351

    Not in our library10.1128/jb.182.15.4348-4351.2000

  98. 98.

    Barrow W.W., Davis T.L., Wright E.L., Labrousse V., Bachelet M. and Rastogi N. (1995 ) Immunomodulatory spectrum of lipids associated with Mycobacterium avium serovar 8. Infect. Immun. 63, 126–133

    Not in our library10.1128/iai.63.1.126-133.1995

  99. 99.

    Byrd T.F. and Lyons C.R. (1999 ) Preliminary characterization of a Mycobacterium abscessus mutant in human and murine models of infection. Infect. Immun. 67, 4700–4707

    Not in our library10.1128/iai.67.9.4700-4707.1999

  100. 100.

    Howard S.T., Rhoades E., Recht J., Pang X., Alsup A., Kolter R. et al. (2006 ) Spontaneous reversion of Mycobacterium abscessus from a smooth to a rough morphotype is associated with reduced expression of glycopeptidolipid and reacquisition of an invasive phenotype. Microbiology 152, 1581–1590

    Not in our library10.1099/mic.0.28625-0

  101. 101.

    Catherinot E., Clarissou J., Etienne G., Ripoll F., Emile J.-F., Daffe M. et al. (2007 ) Hypervirulence of a rough variant of the Mycobacterium abscessus type strain. Infect. Immun. 75, 1055–1058

    Not in our library10.1128/iai.00835-06

  102. 102.

    Bozic C.M., McNeil M., Chatterjee D., Jardine I. and Brennan P.J. (1988 ) Further novel amido sugars within the glycopeptidolipid antigens of Mycobacterium avium. J. Biol. Chem. 263, 14984–14991 PMID:

    Not in our library

  103. 103.

    Laneelle G. and Asselineau J. (1968 ) Structure d'un glycoside de peptidolipide isolé d'une mycobactérie. Eur. J. Biochem. 5, 487–491

    Not in our library10.1111/j.1432-1033.1968.tb00396.x

  104. 104.

    Patterson J.H., McConville M.J., Haites R.E., Coppel R.L. and Billman-Jacobe H. (2000 ) Identification of a methyltransferase from Mycobacterium smegmatis involved in glycopeptidolipid synthesis. J. Biol. Chem. 275, 24900–24906

    Not in our library10.1074/jbc.m000147200

  105. 105.

    Ripoll F., Deshayes C., Pasek S., Laval F., Beretti J.-L., Biet F., et al. (2007 ) Genomics of glycopeptidolipid biosynthesis in Mycobacterium abscessus and M. chelonae. BMC Genom. 8, 114

    Not in our library10.1186/1471-2164-8-114

  106. 106.

    Besra G.S., McNeil M.R., Rivoire B., Khoo K.H., Morris H.R., Dell A. et al. (1993 ) Further structural definition of a new family of glycopeptidolipids from Mycobacterium xenopi. Biochemistry 32, 347–355

    Not in our library10.1021/bi00052a043

  107. 107.

    Riviere M. and Puzo G. (1991 ) A new type of serine-containing glycopeptidolipid from Mycobacterium xenopi. J. Biol. Chem. 266, 9057–9063 PMID:

    Not in our library

  108. 108.

    Peterson D.O. and Bloch K. (1977 ) Mycobacterium smegmatis fatty acid synthetase. Long chain transacylase chain length specificity. J. Biol. Chem. 252, 5735–5739 PMID:

    Not in our library

  109. 109.

    Kikuchi S., Rainwater D.L. and Kolattukudy P.E. (1992 ) Purification and characterization of an unusually large fatty acid synthase from Mycobacterium tuberculosis var. bovis BCG. Arch. Biochem. Biophys. 295, 318–326

    Not in our library10.1016/0003-9861(92)90524-z

  110. 110.

    Bloch K. (1977 ) Control mechanisms for fatty acid synthesis in Mycobacterium smegmatis. Adv. Enzymol. Relat. Areas Mol. Biol. 45, 1–84

    Not in our library10.1002/9780470122907.ch1

  111. 111.

    Mdluli K., Slayden R.A., Zhu Y., Ramaswamy S., Pan X., Mead D. et al. (1998 ) Inhibition of a Mycobacterium tuberculosis β-Ketoacyl ACP synthase by isoniazid. Science 280, 1607

    Not in our library10.1126/science.280.5369.1607

  112. 112.

    Cronan J.E.J. and Waldrop G.L. (2002 ) Multi-subunit acetyl-CoA carboxylases. Prog. Lipid Res. 41, 407–435

    Not in our library10.1016/s0163-7827(02)00007-3

  113. 113.

    Kremer L., Nampoothiri K.M., Lesjean S., Dover L.G., Graham S., Betts J. et al. (2001 ) Biochemical 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. 276, 27967–27974

    Not in our library10.1074/jbc.m103687200

  114. 114.

    Fernandes N.D. and Kolattukudy P.E. (1996 ) Cloning, sequencing and characterization of a fatty acid synthase-encoding gene from Mycobacterium tuberculosis var. bovis BCG. Gene 170, 95–99

    Not in our library10.1016/0378-1119(95)00842-x

  115. 115.

    Choi K.H., Kremer L., Besra G.S. and Rock C.O. (2000 ) Identification and substrate specificity of beta -ketoacyl (acyl carrier protein) synthase III (mtFabH) from Mycobacterium tuberculosis. J. Biol. Chem. 275, 28201–28207

    Not in our library10.1074/jbc.m003241200

  116. 116.

    Marrakchi H., Ducasse S., Labesse G., Montrozier H., Margeat E., Emorine L. et al. (2002 ) Maba (FabG1), a Mycobacterium tuberculosis protein involved in the long-chain fatty acid elongation system FAS-II. Microbiology 148, 951–960

    Not in our library10.1099/00221287-148-4-951

  117. 117.

    Sacco E., Covarrubias A.S., O'Hare H.M., Carroll P., Eynard N., Jones T.A. et al. (2007 ) The missing piece of the type II fatty acid synthase system from Mycobacterium tuberculosis. Proc. Natl. Acad. Sci. U.S.A. 104, 14628–14633

    Not in our library10.1073/pnas.0704132104

  118. 118.

    Banerjee A., Dubnau E., Quemard A., Balasubramanian V., Um K.S., Wilson T. et al. (1994 ) Inha, a gene encoding a target for isoniazid and ethionamide in Mycobacterium tuberculosis. Science 263, 227–230

    Not in our library10.1126/science.8284673

  119. 119.

    Schaeffer M.L., Agnihotri G., Volker C., Kallender H., Brennan P.J. and Lonsdale J.T. (2001 ) Purification and biochemical characterization of the Mycobacterium tuberculosis beta-ketoacyl-acyl carrier protein synthases KasA and KasB. J. Biol. Chem. 276, 47029–47037

    Not in our library10.1074/jbc.m108903200

  120. 120.

    Kremer L., Dover L.G., Carrere S., Nampoothiri K.M., Lesjean S., Brown A.K. et al. (2002 ) Mycolic acid biosynthesis and enzymic characterization of the beta-ketoacyl-ACP synthase A-condensing enzyme from Mycobacterium tuberculosis. Biochem. J. 364, 423–430

    Not in our library10.1042/bj20011628

  121. 121.

    Glickman M.S. (2003 ) The mmaA2 gene of Mycobacterium tuberculosis encodes the distal cyclopropane synthase of the alpha-mycolic acid. J. Biol. Chem. 278, 7844–7849

    Not in our library10.1074/jbc.m212458200

  122. 122.

    Barkan D., Rao V., Sukenick G.D. and Glickman M.S. (2010 ) Redundant function of cmaA2 and mmaA2 in mycobacterium tuberculosis cis cyclopropanation of oxygenated mycolates. J. Bacteriol. 192, 3661–3668

    Not in our library10.1128/jb.00312-10

  123. 123.

    Slayden R.A. and Barry C.E. III (2002 ) The role of KasA and KasB in the biosynthesis of meromycolic acids and isoniazid resistance in Mycobacterium tuberculosis. Tuberc. Edinb. Scotl. 82, 149–160

    Not in our library10.1054/tube.2002.0333

  124. 124.

    Marrakchi H., Bardou F. and Daffé M (2008 ) A comprehensive overview of mycolic acid structure and biosynthesis In The Mycobacterial Cell Envelop (Daffe M. and Reyrat J.-M., eds), pp. 41–62, American Society for Microbiology, Washington, DC

    Not in our library

  125. 125.

    Zimhony O., Vilcheze C. and Jacobs W.R.J. (2004 ) Characterization of Mycobacterium smegmatis expressing the Mycobacterium tuberculosis fatty acid synthase I (fas1) gene. J. Bacteriol. 186, 4051–4055

    Not in our library10.1128/jb.186.13.4051-4055.2004

  126. 126.

    Papaioannou N., Cheon H.-S., Lian Y. and Kishi Y. (2007 ) Product-regulation mechanisms for fatty acid biosynthesis catalyzed by Mycobacterium smegmatis FAS I. Chembiochem 8, 1775–1780

    Not in our library10.1002/cbic.200700380

  127. 127.

    Banis R.J., Peterson D.O. and Bloch K. (1977 ) Mycobacterium smegmatis fatty acid synthetase. Polysaccharide stimulation of the rate-limiting step. J. Biol. Chem. 252, 5740–5744 PMID:

    Not in our library

  128. 128.

    Wood W.I., Peterson D.O. and Bloch K. (1977 ) Mycobacterium smegmatis fatty acid synthetase. A mechanism based on steady state rates and product distributions. J. Biol. Chem. 252, 5745–5749

    Not in our library

  129. 129.

    Cole S.T., Brosch R., Parkhill J., Garnier T., Churcher C., Harris D., et al. (1998 ) Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393, 537–544

    Not in our library10.1038/31159

  130. 130.

    Moore B. and Hopke J. (2001 ) Discovery of a new bacterial polyketide biosynthetic pathway. Chembiochem 2, 35–38

    Not in our library10.1002/1439-7633(20010105)2:1<35::aid-cbic35>3.0.co;2-1

  131. 131.

    Schröder J. (1997 ) A family of plant-specific polyketide synthases: facts and predictions. Trends Plant Sci. 2, 373–378

    Not in our library10.1016/s1360-1385(97)87121-x

  132. 132.

    Trivedi O.A., Arora P., Sridharan V., Tickoo R., Mohanty D. and Gokhale R.S. (2004 ) Enzymic activation and transfer of fatty acids as acyl-adenylates in mycobacteria. Nature 428, 441–445

    Not in our library10.1038/nature02384

  133. 133.

    Léger M., Gavalda S., Guillet V., van der Rest B., Slama N., Montrozier H. et al. (2009 ) The dual function of the Mycobacterium tuberculosis FadD32 required for mycolic acid biosynthesis. Chem. Biol. 16, 510–519

    Not in our library10.1016/j.chembiol.2009.03.012

  134. 134.

    Gande R., Gibson K.J.C., Brown A.K., Krumbach K., Dover L.G., Sahm H. et al. (2004 ) Acyl-CoA carboxylases (accD2 and accD3), together with a unique polyketide synthase (Cg-pks), are key to mycolic acid biosynthesis in corynebacterianeae such as Corynebacterium glutamicum and Mycobacterium tuberculosis. J. Biol. Chem. 279, 44847–44857

    Not in our library10.1074/jbc.m408648200

  135. 135.

    Portevin D., de Sousa-D'Auria C., Montrozier H., Houssin C., Stella A., Lanéelle M.-A. et al. (2005 ) The Acyl-AMP ligase FadD32 and AccD4-containing Acyl-CoA carboxylase are required for the synthesis of mycolic acids and essential for mycobacterial growth: identification of the carboxylation product and determination of the Acyl-CoA carboxylase components. J. Biol. Chem. 280, 8862–8874

    Not in our library10.1074/jbc.m408578200

  136. 136.

    Gavalda S., Leger M., van der Rest B., Stella A., Bardou F., Montrozier H., et al. (2009 ) The Pks13/FadD32 crosstalk for the biosynthesis of mycolic acids in Mycobacterium tuberculosis. J. Biol. Chem. 284, 19255–19264

    Not in our library10.1074/jbc.m109.006940

  137. 137.

    Gande R., Dover L.G., Krumbach K., Besra G.S., Sahm H., Oikawa T. et al. (2007 ) The two carboxylases of Corynebacterium glutamicum essential for fatty acid and mycolic acid synthesis. J. Bacteriol. 189, 5257

    Not in our library10.1128/jb.00254-07

  138. 138.

    Takayama K., Wang C. and Besra G.S. (2005 ) Pathway to synthesis and processing of mycolic acids in Mycobacterium tuberculosis. Clin. Microbiol. Rev. 18, 81

    Not in our library10.1128/cmr.18.1.81-101.2005

  139. 139.

    Gavalda S., Bardou F., Laval F., Bon C., Malaga W., Chalut C. et al. (2014 ) The polyketide synthase Pks13 catalyzes a novel mechanism of lipid transfer in Mycobacteria. Chem. Biol. 21, 1660–1669

    Not in our library10.1016/j.chembiol.2014.10.011

  140. 140.

    Bhatt A., Brown A.K., Singh A., Minnikin D.E. and Besra G.S. (2008 ) Loss of a mycobacterial gene encoding a reductase leads to an altered cell wall containing beta-oxo-mycolic acid analogs and accumulation of ketones. Chem. Biol. 15, 930–939

    Not in our library10.1016/j.chembiol.2008.07.007

  141. 141.

    Lea-Smith D.J., Pyke J.S., Tull D., McConville M.J., Coppel R.L. and Crellin P.K. (2007 ) The reductase that catalyzes mycolic motif synthesis is required for efficient attachment of mycolic acids to arabinogalactan. J. Biol. Chem. 282, 11000–11008

    Not in our library10.1074/jbc.m608686200

  142. 142.

    Nobre A., Alarico S., Maranha A., Mendes V. and Empadinhas N. (2014 ) The molecular biology of Mycobacterial trehalose in the quest for advanced tuberculosis therapies. Microbiology 160, 1547–1570

    Not in our library10.1099/mic.0.075895-0

  143. 143.

    Hatzios S.K., Schelle M.W., Holsclaw C.M., Behrens C.R., Botyanszki Z., Lin F. et al. (2009 ) Papa3 is an acyltransferase required for polyacyltrehalose biosynthesis in Mycobacterium tuberculosis. J. Biol. Chem. 284, 12745–12751

    Not in our library10.1074/jbc.m809088200

  144. 144.

    Kumar P., Schelle M.W., Jain M., Lin F.L., Petzold C.J., Leavell M.D. et al. (2007 ) Papa1 and PapA2 are acyltransferases essential for the biosynthesis of the Mycobacterium tuberculosis virulence factor sulfolipid-1. Proc. Natl. Acad. Sci. U.S.A. 104, 11221

    Not in our library10.1073/pnas.0611649104

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    Jackson M., Stadthagen G. and Gicquel B. (2007 ) Long-chain multiple methyl-branched fatty acid-containing lipids of Mycobacterium tuberculosis: biosynthesis, transport, regulation and biological activities. Tuberc. Edinb. Scotl. 87, 78–86

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    Bhatt K., Gurcha S.S., Bhatt A., Besra G.S. and Jacobs W.R.J. (2007 ) Two polyketide-synthase-associated acyltransferases are required for sulfolipid biosynthesis in Mycobacterium tuberculosis. Microbiology 153, 513–520

    Not in our library10.1099/mic.0.2006/003103-0

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    Besra G.S., Khoo K.H., Belisle J.T., McNeil M.R., Morris H.R., Dell A. et al. (1994 ) New pyruvylated, glycosylated acyltrehaloses from Mycobacterium smegmatis strains, and their implications for phage resistance in mycobacteria. Carbohydr. Res. 251, 99–114

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  148. 148.

    Lee J., Lin E.-W., Lau U.Y., Hedrick J.L., Bat E. and Maynard H.D. (2013 ) Trehalose glycopolymers as excipients for protein stabilization. Biomacromolecules 14, 2561–2569

    Not in our library10.1021/bm4003046

  149. 149.
    Trehalose Metabolism: From Osmoprotection to Signaling · 2009
    No notice
  150. 150.

    Webb K.M. and DiRuggiero J. (2012 ) Role of Mn2+ and compatible solutes in the radiation resistance of thermophilic bacteria and archaea. Archaea 2012, 845756

    Not in our library10.1155/2012/845756

  151. 151.

    Belisle J.T., Vissa V.D., Sievert T., Takayama K., Brennan P.J. and Besra G.S. (1997 ) Role of the major antigen of Mycobacterium tuberculosis in cell wall biogenesis. Science 276, 1420

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  152. 152.

    Kalscheuer R., Weinrick B., Veeraraghavan U., Besra G.S. and Jacobs W.R.J. (2010 ) Trehalose-recycling ABC transporter LpqY-SugA-SugB-SugC is essential for virulence of Mycobacterium tuberculosis. Proc. Natl. Acad. Sci. U.S.A. 107, 21761–21766

    Not in our library10.1073/pnas.1014642108

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    Saier M.H.J., Tam R., Reizer A. and Reizer J. (1994 ) Two novel families of bacterial membrane proteins concerned with nodulation, cell division and transport. Mol. Microbiol. 11, 841–847

    Not in our library10.1111/j.1365-2958.1994.tb00362.x

  154. 154.

    Chim N., Torres R., Liu Y., Capri J., Batot G., Whitelegge J.P. et al. (2015 ) The structure and interactions of periplasmic domains of crucial MmpL membrane proteins from Mycobacterium tuberculosis. Chem. Biol. 22, 1098–1107

    Not in our library10.1016/j.chembiol.2015.07.013

  155. 155.

    Ma D., Cook D.N., Alberti M., Pon N.G., Nikaido H. and Hearst J.E. (1993 ) Molecular cloning and characterization of acrA and acrE genes of Escherichia coli. J. Bacteriol 175, 6299–6313

    Not in our library10.1128/jb.175.19.6299-6313.1993

  156. 156.

    Seeger M.A., Schiefner A., Eicher T., Verrey F., Diederichs K. and Pos K.M. (2006 ) Structural asymmetry of AcrB trimer suggests a peristaltic pump mechanism. Science 313, 1295–1298

    Not in our library10.1126/science.1131542

  157. 157.

    Murakami S., Nakashima R., Yamashita E., Matsumoto T. and Yamaguchi A. (2006 ) Crystal structures of a multidrug transporter reveal a functionally rotating mechanism. Nature 443, 173–179

    Not in our library10.1038/nature05076

  158. 158.

    Domenech P., Reed M.B. and Barry C.E. III (2005 ) Contribution of the Mycobacterium tuberculosis MmpL protein family to virulence and drug resistance. Infect. Immun. 73, 3492–3501

    Not in our library10.1128/iai.73.6.3492-3501.2005

  159. 159.

    Milano A., Pasca M.R., Provvedi R., Lucarelli A.P., Manina G., Luisa de Jesus Lopes Ribeiro A. et al. (2009 ) Azole resistance in Mycobacterium tuberculosis is mediated by the MmpS5–MmpL5 efflux system. Tuberculosis 89, 84–90

    Not in our library10.1016/j.tube.2008.08.003

  160. 160.

    Halloum I., Viljoen A., Khanna V., Craig D., Bouchier C., Brosch R. et al. (2017 ) Resistance to thiacetazone derivatives active against Mycobacterium abscessus involves mutations in the MmpL5 transcriptional repressor MAB_4384. Antimicrob. Agents Chemother. 61, e02509-02516

    Not in our library10.1128/aac.02509-16

  161. 161.

    Hartkoorn R.C., Uplekar S. and Cole S.T. (2014 ) Cross-resistance between clofazimine and bedaquiline through upregulation of MmpL5 in Mycobacterium tuberculosis. Antimicrob. Agents Chemother. 58, 2979–2981

    Not in our library10.1128/aac.00037-14

  162. 162.

    Seeliger J.C., Holsclaw C.M., Schelle M.W., Botyanszki Z., Gilmore S.A., Tully S.E. et al. (2012 ) Elucidation and chemical modulation of sulfolipid-1 biosynthesis in Mycobacterium tuberculosis. J. Biol. Chem. 287, 7990–8000

    Not in our library10.1074/jbc.m111.315473

  163. 163.

    Sulzenbacher G., Canaan S., Bordat Y., Neyrolles O., Stadthagen G., Roig-Zamboni V., et al. (2006 ) Lppx is a lipoprotein required for the translocation of phthiocerol dimycocerosates to the surface of Mycobacterium tuberculosis. EMBO J. 25, 1436–1444

    Not in our library10.1038/sj.emboj.7601048

  164. 164.

    Grzegorzewicz A.E., Pham H., Gundi V.A.K.B., Scherman M.S., North E.J., Hess T., et al. (2012 ) Inhibition of mycolic acid transport across the Mycobacterium tuberculosis plasma membrane. Nat. Chem. Biol. 8, 334–341

    Not in our library10.1038/nchembio.794

  165. 165.

    Tahlan K., Wilson R., Kastrinsky D.B., Arora K., Nair V., Fischer E. et al. (2012 ) SQ109 targets MmpL3, a membrane transporter of trehalose monomycolate involved in mycolic acid donation to the cell wall core of Mycobacterium tuberculosis. Antimicrob. Agents Chemother. 56, 1797

    Not in our library10.1128/aac.05708-11

  166. 166.
    MmpL Genes Are Associated with Mycolic Acid Metabolism in Mycobacteria and Corynebacteria · 2012
    No notice
  167. 167.

    La Rosa V., Poce G., Canseco J.O., Buroni S., Pasca M.R., Biava M. et al. (2012 ) Mmpl3 is the cellular target of the antitubercular pyrrole derivative BM212. Antimicrob. Agents Chemother. 56, 324

    Not in our library10.1128/aac.05270-11

  168. 168.

    Zhang B., Li J., Yang X., Wu L., Zhang J., Yang Y., et al. (2019 ) Crystal structures of membrane transporter MmpL3, an anti-TB drug target. Cell 176, 636–648.e13

    Not in our library10.1016/j.cell.2019.01.003

  169. 169.

    Sennhauser G., Bukowska M.A., Briand C. and Grutter M.G. (2009 ) Crystal structure of the multidrug exporter MexB from Pseudomonas aeruginosa. J. Mol. Biol. 389, 134–145

    Not in our library10.1016/j.jmb.2009.04.001

  170. 170.

    Murakami S., Nakashima R., Yamashita E. and Yamaguchi A. (2002 ) Crystal structure of bacterial multidrug efflux transporter AcrB. Nature 419, 587–593

    Not in our library10.1038/nature01050

  171. 171.

    Kumar N., Su C.-C., Chou T.-H., Radhakrishnan A., Delmar J.A., Rajashankar K.R. et al. (2017 ) Crystal structures of the Burkholderia multivorans hopanoid transporter HpnN. Proc. Natl. Acad. Sci. U.S.A. 114, 6557–6562

    Not in our library10.1073/pnas.1619660114

  172. 172.

    Belardinelli J.M., Yazidi A., Yang L., Fabre L., Li W., Jacques B., et al. (2016 ) Structure-function profile of MmpL3, the essential mycolic acid transporter from Mycobacterium tuberculosis. ACS Infect. Dis. 2, 702–713

    Not in our library10.1021/acsinfecdis.6b00095

  173. 173.

    Delmar J.A., Chou T.-H., Wright C.C., Licon M.H., Doh J.K., Radhakrishnan A., et al. (2015 ) Structural basis for the regulation of the MmpL transporters of Mycobacterium tuberculosis. J. Biol. Chem. 290, 28559–28574

    Not in our library10.1074/jbc.m115.683797

  174. 174.

    Chou T.-H., Delmar J.A., Wright C.C., Kumar N., Radhakrishnan A., Doh J.K., et al. (2015 ) Crystal structure of the Mycobacterium tuberculosis transcriptional regulator Rv0302. Protein Sci. 24, 1942–1955

    Not in our library10.1002/pro.2802

  175. 175.

    Galagan J.E., Minch K., Peterson M., Lyubetskaya A., Azizi E., Sweet L., et al. (2013 ) The Mycobacterium tuberculosis regulatory network and hypoxia. Nature 499, 178–183

    Not in our library10.1038/nature12337

  176. 176.

    Prisic S., Dankwa S., Schwartz D., Chou M.F., Locasale J.W., Kang C.-M. et al. (2010 ) Extensive phosphorylation with overlapping specificity by Mycobacterium tuberculosis serine/threonine protein kinases. Proc. Natl. Acad. Sci. U.S.A. 107, 7521–7526

    Not in our library10.1073/pnas.0913482107

  177. 177.

    Pitarque S., Larrouy-Maumus G., Payre B., Jackson M., Puzo G. and Nigou J. (2008 ) The immunomodulatory lipoglycans, lipoarabinomannan and lipomannan, are exposed at the mycobacterial cell surface. Tuberculosis (Edinb) 88, 560–565

    Not in our library10.1016/j.tube.2008.04.002

  178. 178.

    Parish T., Liu J., Nikaido H. and Stoker N.G. (1997 ) A Mycobacterium smegmatis mutant with a defective inositol monophosphate phosphatase gene homolog has altered cell envelope permeability. J. Bacteriol. 179, 7827

    Not in our library10.1128/jb.179.24.7827-7833.1997

  179. 179.

    Patterson J.H., Waller R.F., Jeevarajah D., Billman-Jacobe H. and McConville M.J. (2003 ) Mannose metabolism is required for mycobacterial growth. Biochem. J. 372, 77–86

    Not in our library10.1042/bj20021700

  180. 180.

    Fukuda T., Matsumura T., Ato M., Hamasaki M., Nishiuchi Y., Murakami Y., et al. (2013 ) Critical roles for lipomannan and lipoarabinomannan in cell wall integrity of mycobacteria and pathogenesis of tuberculosis. MBio 4, e00472–e00412

    Not in our library10.1128/mbio.00472-12

  181. 181.

    Fratti R.A., Chua J., Vergne I. and Deretic V. (2003 ) Mycobacterium tuberculosis glycosylated phosphatidylinositol causes phagosome maturation arrest. Proc. Natl. Acad. Sci. U.S.A. 100, 5437–5442

    Not in our library10.1073/pnas.0737613100

  182. 182.

    Besra G.S., Morehouse C.B., Rittner C.M., Waechter C.J. and Brennan P.J. (1997 ) Biosynthesis of mycobacterial lipoarabinomannan. J. Biol. Chem. 272, 18460–18466

    Not in our library10.1074/jbc.272.29.18460

  183. 183.

    Khoo K.H., Dell A., Morris H.R., Brennan P.J. and Chatterjee D. (1995 ) Structural definition of acylated phosphatidylinositol mannosides from Mycobacterium tuberculosis: definition of a common anchor for lipomannan and lipoarabinomannan. Glycobiology 5, 117–127

    Not in our library10.1093/glycob/5.1.117

  184. 184.

    Severn W.B., Furneaux R.H., Falshaw R. and Atkinson P.H. (1998 ) Chemical and spectroscopic characterisation of the phosphatidylinositol manno-oligosaccharides from Mycobacterium bovis AN5 and WAg201 and Mycobacterium smegmatis mc2 155. Carbohydr. Res. 308, 397–408

    Not in our library10.1016/s0008-6215(98)00108-6

  185. 185.

    Hunter S.W. and Brennan P.J. (1990 ) Evidence for the presence of a phosphatidylinositol anchor on the lipoarabinomannan and lipomannan of Mycobacterium tuberculosis. J. Biol. Chem. 265, 9272–9279 PMID:

    Not in our library

  186. 186.

    Chatterjee D., Bozic C.M., McNeil M. and Brennan P.J. (1991 ) Structural features of the arabinan component of the lipoarabinomannan of Mycobacterium tuberculosis. J. Biol. Chem. 266, 9652–9660 PMID:

    Not in our library

  187. 187.

    Chatterjee D., Lowell K., Rivoire B., McNeil M.R. and Brennan P.J. (1992 ) Lipoarabinomannan of Mycobacterium tuberculosis. Capping with mannosyl residues in some strains. J. Biol. Chem. 267, 6234–6239 PMID:

    Not in our library

  188. 188.

    Liu J. and Mushegian A. (2003 ) Three monophyletic superfamilies account for the majority of the known glycosyltransferases. Protein Sci. 12, 1418–1431

    Not in our library10.1110/ps.0302103

  189. 189.

    Morita Y.S., Velasquez R., Taig E., Waller R.F., Patterson J.H., Tull D. et al. (2005 ) Compartmentalization of lipid biosynthesis in Mycobacteria. J. Biol. Chem. 280, 21645–21652

    Not in our library10.1074/jbc.m414181200

  190. 190.

    Hill D.L. and Ballou C.E. (1966 ) Biosynthesis of mannophospholipids by Mycobacterium phlei. J. Biol. Chem. 241, 895–902 PMID:

    Not in our library

  191. 191.

    Guerin M.E., Kaur D., Somashekar B.S., Gibbs S., Gest P., Chatterjee D. et al. (2009 ) New insights into the early steps of phosphatidylinositol mannoside biosynthesis in mycobacteria: PimB’ is an essential enzyme of Mycobacterium smegmatis. J. Biol. Chem. 284, 25687–25696

    Not in our library10.1074/jbc.m109.030593

  192. 192.

    Kordulakova J., Gilleron M., Mikusova K., Puzo G., Brennan P.J., Gicquel B. et al. (2002 ) Definition of the first mannosylation step in phosphatidylinositol mannoside synthesis. PimA is essential for growth of mycobacteria. J. Biol. Chem. 277, 31335–31344

    Not in our library10.1074/jbc.m204060200

  193. 193.

    Lea-Smith D.J., Martin K.L., Pyke J.S., Tull D., McConville M.J., Coppel R.L. et al. (2008 ) Analysis of a new mannosyltransferase required for the synthesis of phosphatidylinositol mannosides and lipoarbinomannan reveals two lipomannan pools in corynebacterineae. J. Biol. Chem. 283, 6773–6782

    Not in our library10.1074/jbc.m707139200

  194. 194.

    Korduláková J., Gilleron M., Puzo G., Brennan P.J., Gicquel B., Mikušová K. et al. (2003 ) Identification of the required acyltransferase step in the biosynthesis of the phosphatidylinositol mannosides of Mycobacterium species. J. Biol. Chem. 278, 36285–36295

    Not in our library10.1074/jbc.m303639200

  195. 195.

    Albesa-Jove D., Svetlikova Z., Tersa M., Sancho-Vaello E., Carreras-Gonzalez A., Bonnet P., et al. (2016 ) Structural basis for selective recognition of acyl chains by the membrane-associated acyltransferase PatA. Nat. Commun. 7, 10906

    Not in our library10.1038/ncomms10906

  196. 196.

    Kremer L., Gurcha S.S., Bifani P., Hitchen P.G., Baulard A., Morris H.R. et al. (2002 ) Characterization of a putative alpha-mannosyltransferase involved in phosphatidylinositol trimannoside biosynthesis in Mycobacterium tuberculosis. Biochem. J. 363, 437–447

    Not in our library10.1042/bj3630437

  197. 197.

    Morita Y.S., Patterson J.H., Billman-Jacobe H. and McConville M.J. (2004 ) Biosynthesis of mycobacterial phosphatidylinositol mannosides. Biochem. J. 378, 589–597

    Not in our library10.1042/bj20031372

  198. 198.
    Mycobacterium tuberculosis universal stress protein Rv2623 interacts with the putative ATP binding cassette (ABC) transporter Rv1747 to regulate mycobacterial growth. · 2017
    Not checked
  199. 199.

    Spivey V.L., Whalan R.H., Hirst E.M.A., Smerdon S.J. and Buxton R.S. (2013 ) An attenuated mutant of the Rv1747 ATP-binding cassette transporter of Mycobacterium tuberculosis and a mutant of its cognate kinase, PknF, show increased expression of the efflux pump-related iniBAC operon. FEMS Microbiol. Lett. 347, 107–115

    Not in our library10.1111/1574-6968.12230

  200. 200.

    Morita Y.S., Sena C.B.C., Waller R.F., Kurokawa K., Sernee M.F., Nakatani F. et al. (2006 ) Pime is a polyprenol-phosphate-mannose-dependent mannosyltransferase that transfers the fifth mannose of phosphatidylinositol mannoside in mycobacteria. J. Biol. Chem. 281, 25143–25155

    Not in our library10.1074/jbc.m604214200

  201. 201.

    Mishra A.K., Alderwick L.J., Rittmann D., Wang C., Bhatt A., Jacobs Jr W.R. et al. (2008 ) Identification of a novel α(1→6) mannopyranosyltransferase MptB from Corynebacterium glutamicum by deletion of a conserved gene, NCgl1505, affords a lipomannan- and lipoarabinomannan-deficient mutant. Mol. Microbiol. 68, 1595–1613

    Not in our library10.1111/j.1365-2958.2008.06265.x

  202. 202.

    Mishra A.K., Alderwick L.J., Rittmann D., Tatituri R.V.V., Nigou J., Gilleron M. et al. (2007 ) Identification of an α(1→6) mannopyranosyltransferase (MptA), involved in Corynebacterium glutamicum lipomanann biosynthesis, and identification of its orthologue in Mycobacterium tuberculosis. Mol. Microbiol. 65, 1503–1517

    Not in our library10.1111/j.1365-2958.2007.05884.x

  203. 203.

    Mishra A.K., Krumbach K., Rittmann D., Appelmelk B., Pathak V., Pathak A.K. et al. (2011 ) Lipoarabinomannan biosynthesis in Corynebacterineae: the interplay of two α(1→2)-mannopyranosyltransferases MptC and MptD in mannan branching. Mol. Microbiol. 80, 1241–1259

    Not in our library10.1111/j.1365-2958.2011.07640.x

  204. 204.

    Kaur D., Berg S., Dinadayala P., Gicquel B., Chatterjee D., McNeil M.R. et al. (2006 ) Biosynthesis of mycobacterial lipoarabinomannan: Role of a branching mannosyltransferase. Proc. Natl. Acad. Sci. U.S.A. 103, 13664

    Not in our library10.1073/pnas.0603049103

  205. 205.

    Kovacevic S., Anderson D., Morita Y.S., Patterson J., Haites R., McMillan B.N.I. et al. (2006 ) Identification of a novel protein with a role in lipoarabinomannan biosynthesis in mycobacteria. J. Biol. Chem. 281, 9011–9017

    Not in our library10.1074/jbc.m511709200

  206. 206.

    Crellin P.K., Kovacevic S., Martin K.L., Brammananth R., Morita Y.S., Billman-Jacobe H. et al. (2008 ) Mutations in pimE restore lipoarabinomannan synthesis and growth in a Mycobacterium smegmatis lpqW mutant. J. Bacteriol. 190, 3690–3699

    Not in our library10.1128/jb.00200-08

  207. 207.

    Rainczuk A.K., Yamaryo-Botte Y., Brammananth R., Stinear T.P., Seemann T., Coppel R.L. et al. (2012 ) The lipoprotein LpqW is essential for the mannosylation of periplasmic glycolipids in Corynebacteria. J. Biol. Chem. 287, 42726–42738

    Not in our library10.1074/jbc.m112.373415

  208. 208.

    Shi L., Berg S., Lee A., Spencer J.S., Zhang J., Vissa V. et al. (2006 ) The carboxy terminus of embC from Mycobacterium smegmatis mediates chain length extension of the arabinan in lipoarabinomannan. J. Biol. Chem. 281, 19512–19526

    Not in our library10.1074/jbc.m513846200

  209. 209.

    Treumann A., Xidong F., McDonnell L., Derrick P.J., Ashcroft A.E., Chatterjee D. et al. (2002 ) 5-methylthiopentose: a new substituent on lipoarabinomannan in Mycobacterium tuberculosis. J. Mol. Biol. 316, 89–100

    Not in our library10.1006/jmbi.2001.5317

  210. 210.

    Torrelles J.B., Khoo K.-H., Sieling P.A., Modlin R.L., Zhang N., Marques A.M. et al. (2004 ) Truncated structural variants of lipoarabinomannan in Mycobacterium leprae and an ethambutol-resistant strain of Mycobacterium tuberculosis. J. Biol. Chem. 279, 41227–41239

    Not in our library10.1074/jbc.m405180200

  211. 211.

    Turnbull W.B., Shimizu K.H., Chatterjee D., Homans S.W. and Treumann A. (2004 ) Identification of the 5-methylthiopentosyl substituent in Mycobacterium tuberculosis lipoarabinomannan. Angew. Chem. Int. Ed. 43, 3918–3922

    Not in our library10.1002/anie.200454119

  212. 212.

    McNeil M.R., Robuck K.G., Harter M. and Brennan P.J. (1994 ) Enzymatic evidence for the presence of a critical terminal hexa-arabinoside in the cell walls of Mycobacterium tuberculosis. Glycobiology 4, 165–174

    Not in our library10.1093/glycob/4.2.165

  213. 213.

    Wolucka B.A., McNeil M.R., de Hoffmann E., Chojnacki T. and Brennan P.J. (1994 ) Recognition of the lipid intermediate for arabinogalactan/arabinomannan biosynthesis and its relation to the mode of action of ethambutol on mycobacteria. J. Biol. Chem. 269, 23328–23335 PMID:

    Not in our library

  214. 214.

    Chatterjee D., Khoo K.H., McNeil M.R., Dell A., Morris H.R. and Brennan P.J. (1993 ) Structural definition of the non-reducing termini of mannose-capped LAM from Mycobacterium tuberculosis through selective enzymatic degradation and fast atom bombardment-mass spectrometry. Glycobiology 3, 497–506

    Not in our library10.1093/glycob/3.5.497

  215. 215.

    Angala S.K., McNeil M.R., Zou L., Liav A., Zhang J., Lowary T.L. et al. (2016 ) Identification of a novel mycobacterial arabinosyltransferase activity which adds an arabinosyl residue to α-d-mannosyl residues. ACS Chem. Biol. 11, 1518–1524

    Not in our library10.1021/acschembio.6b00093

  216. 216.

    Zhang N., Torrelles J.B., McNeil M.R., Escuyer V.E., Khoo K.-H., Brennan P.J. et al. (2003 ) The Emb proteins of mycobacteria direct arabinosylation of lipoarabinomannan and arabinogalactan via an N-terminal recognition region and a C-terminal synthetic region. Mol. Microbiol. 50, 69–76

    Not in our library10.1046/j.1365-2958.2003.03681.x

  217. 217.

    Birch H.L., Alderwick L.J., Appelmelk B.J., Maaskant J., Bhatt A., Singh A. et al. (2010 ) A truncated lipoglycan from mycobacteria with altered immunological properties. Proc. Natl. Acad. Sci. U.S.A. 107, 2634–2639

    Not in our library10.1073/pnas.0915082107

  218. 218.

    Birch H.L., Alderwick L.J., Bhatt A., Rittmann D., Krumbach K., Singh A. et al. (2008 ) Biosynthesis of mycobacterial arabinogalactan: identification of a novel alpha(1 → 3) arabinofuranosyltransferase. Mol. Microbiol. 69, 1191–1206

    Not in our library10.1111/j.1365-2958.2008.06354.x

  219. 219.

    Skovierova H., Larrouy-Maumus G., Zhang J., Kaur D., Barilone N., Kordulakova J., et al. (2009 ) Aftd, a novel essential arabinofuranosyltransferase from mycobacteria. Glycobiology 19, 1235–1247

    Not in our library10.1093/glycob/cwp116

  220. 220.
    AftD functions as an α1 → 5 arabinofuranosyltransferase involved in the biosynthesis of the mycobacterial cell wall core · 2018
    Not checked
  221. 221.

    Jankute M., Alderwick L.J., Noack S., Veerapen N., Nigou J. and Besra G.S. (2017 ) Disruption of Mycobacterial AftB results in complete loss of terminal β(1 → 2) arabinofuranose residues of lipoarabinomannan. ACS Chem. Biol. 12, 183–190

    Not in our library10.1021/acschembio.6b00898

  222. 222.

    Khoo K.-H., Dell A., Morris H.R., Brennan P.J. and Chatterjee D. (1995 ) Inositol phosphate capping of the nonreducing termini of lipoarabinomannan from rapidly growing strains of Mycobacterium. J. Biol. Chem. 270, 12380–12389

    Not in our library10.1074/jbc.270.21.12380

  223. 223.

    Guérardel Y., Maes E., Elass E., Leroy Y., Timmerman P., Besra G.S. et al. (2002 ) Structural study of lipomannan and lipoarabinomannan from Mycobacterium chelonae: presence of unusual components with α1,3-mannopyranose side chainS. J. Biol. Chem. 277, 30635–30648

    Not in our library10.1074/jbc.m204398200

  224. 224.

    Dinadayala P., Kaur D., Berg S., Amin A.G., Vissa V.D., Chatterjee D. et al. (2006 ) Genetic basis for the synthesis of the immunomodulatory mannose caps of lipoarabinomannan in Mycobacterium tuberculosis. J. Biol. Chem. 281, 20027–20035

    Not in our library10.1074/jbc.m603395200

  225. 225.

    Appelmelk B.J., den Dunnen J., Driessen N.N., Ummels R., Pak M., Nigou J. et al. (2008 ) The mannose cap of mycobacterial lipoarabinomannan does not dominate the Mycobacterium-host interaction. Cell. Microbiol. 10, 930–944

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