Bacterial membrane lipids: diversity in structures and pathways.

FEMS Microbiology Reviews Advance Access published April 9, 2015 FEMS Microbiology Reviews, fuv008 doi: 10.1093/femsre/fuv008 Review Article

REVIEW ARTICLE

Bacterial membrane lipids: diversity in structures and pathways Christian Sohlenkamp∗ and Otto Geiger ´ ´ ´ Centro de Ciencias Genomicas, Universidad Nacional Autonoma de Mexico, Av. Universidad s/n, Apdo. Postal 565-A, Cuernavaca, Morelos, CP 62210, Mexico ∗ Corresponding author: Centro de Ciencias Genomicas, ´ ´ ´ Universidad Nacional Autonoma de Mexico, Av. Universidad s/n, Apdo. Postal 565-A,

Cuernavaca, Morelos, CP 62210, Mexico. Tel: (+52) 777-3291703; Fax: (+52) 777-3175581; E-mail: [email protected] One sentence summary: This review gives an overview about the diversity and distribution of bacterial membrane lipids and the metabolic pathways involved in their synthesis. Editor: Franz Narberhaus

ABSTRACT For many decades, Escherichia coli was the main model organism for the study of bacterial membrane lipids. The results obtained served as a blueprint for membrane lipid biochemistry, but it is clear now that there is no such thing as a typical bacterial membrane lipid composition. Different bacterial species display different membrane compositions and even the membrane composition of cells belonging to a single species is not constant, but depends on the environmental conditions to which the cells are exposed. Bacterial membranes present a large diversity of amphiphilic lipids, including the common phospholipids phosphatidylglycerol, phosphatidylethanolamine and cardiolipin, the less frequent phospholipids phosphatidylcholine, and phosphatidylinositol and a variety of other membrane lipids, such as for example ornithine lipids, glycolipids, sphingolipids or hopanoids among others. In this review, we give an overview about the membrane lipid structures known in bacteria, the different metabolic pathways involved in their formation, and the distribution of membrane lipids and metabolic pathways across taxonomical groups. Keywords: ornithine lipid; phosphatidylcholine; cardiolipin; phosphatidylinositol; phospholipid; hopanoids; phosphatidylethanolamine

INTRODUCTION This review tries to open up the world of microbial membrane lipids to the reader. We want to give an overview about the inventory of membrane lipids present in bacteria belonging to the different phyla studied. Also, we want to summarize the known synthesis pathways for the major membrane-forming lipids and discuss how widely these pathways are distributed. However, the present review is not meant to be a complete catalog of all different unique structures that have been reported in the literature. Readers looking for this information are referred to the book Microbial Lipids by Ratledge and Wilkinson (1988). Lipids come in a diversity of structures and functions. Here, we

focus on polar/amphiphilic lipids, i.e. membrane-forming lipids present in bacterial membranes, leaving membrane lipids from other microorganisms aside. Classically, the technique of Gram staining allowed the classification of bacteria as either Gram-positive or Gram-negative bacteria, a differentiation based on their cell wall properties. Gram-positive bacteria such as the firmicutes are characterized by having a cytoplasmatic membrane and a thick murein cell wall comprising many layers, whereas Gram-negative bacteria such as proteobacteria are characterized by the presence of two distinct membranes, called inner and outer membrane (Raetz and Whitfield 2002; Reichmann and Grundling 2011) and ¨ a thin murein cell wall between them. Usually, the outer leaflet

Received: 5 December 2014; Accepted: 5 March 2015 C FEMS 2015. All rights reserved. For permissions, please e-mail: [email protected]

1

2

FEMS Microbiology Reviews

of the outer membrane is formed by lipid A, the lipophilic anchor of lipopolysaccharide (LPS) (Raetz and Dowhan 1990; Raetz et al. 2007). Mycobacteria are characterized by the presence of a thick lipophilic layer containing mycolic acids on top of the cell wall, that is sometimes called mycobacterial outer membrane (Niederweis et al. 2010). The two latter structures will not be discussed here. Membranes are formed by amphiphilic lipids which in most cases and conditions studied are glycerophospholipids, composed of two fatty acids, a glycerol moiety, a phosphate group and a variable head group. Examples are phosphatidylethanolamine (PE), phosphatidylglycerol (PG), cardiolipin (CL), lysyl-phosphatidylglycerol (LPG), phosphatidylinositol (PI), phosphatidic acid (PA) and phosphatidylserine (PS). Bacteria can also form phosphorus-free membrane lipids such as ornithine lipids (OLs), sulfolipids, diacylglyceryl-N,N,Ntrimethylhomoserine (DGTS), glycolipids (GLs), diacylglycerol (DAG), hopanoids (HOPs) and others. Since the discovery of the first pathways for membrane lipid synthesis, a surprising diversity and complexity of enzymes and pathways involved has emerged. Although microorganisms are sometimes thought of as more limited in the range and types of lipids which they possess in comparison to plant or animal cells, this view is wholly incorrect as we hope to convince the reader. In this review, we will describe the complexity of pathways for the biosynthesis of phospholipids and phosphorus-free membrane lipids and discuss how both types of lipids are metabolically connected.

DISTRIBUTION OF MEMBRANE LIPIDS IN BACTERIA Membrane lipids in Escherichia coli As in other areas of microbiology, E. coli has been the standard model also in the study of bacterial membrane lipids for many decades. Escherichia coli accumulates three major membrane phospholipids: its predominant lipid is the zwitterionic PE (about 75% of membrane lipids), and additionally it forms the anionic lipids PG (about 20%) and CL (Raetz and Dowhan 1990). Metabolic pathways and lipids known to be present in E. coli have been highlighted in red in Fig. 1. In E. coli, two pathways start from the central metabolite cytidine diphosphate-diacylglycerol (CDP-DAG): PE is synthesized in two steps by phosphatidylserine synthase (Pss) (Figs 1 and 2, reaction 1) and phosphatidylserine decarboxylase (Psd) (Figs 1 and 2, reaction 2). Phosphatidylglycerolphosphate synthase (PgsA) catalyzes the condensation of glycerol-3-phosphate (G3P) with CDP-DAG leading to the formation of PG phosphate (PGP), which is the first step in the synthesis of the anionic membrane lipids PG and CL (Figs 1 and 3, reaction 4). PGP is dephosphorylated in E. coli by any one of three phosphatases (PgpA, PgpB, PgpC) (Figs 1 and 3, reaction 5) to form PG (Lu et al. 2011). The CL synthase ClsA can then synthesize CL from two molecules of PG (Figs 1 and 3, reaction 6). Escherichia coli has two more CL synthase (Cls) called ClsB and ClsC. So far ClsB only has been shown to work in in vitro assays (Guo and Tropp 2000). ClsC has only been recently recognized as a Cls forming CL from PG and PE (Figs 1 and 3, reaction 7) (Tan et al. 2012). Most of these genes and enzymes were discovered and characterized already decades ago and the general feeling was that the described set of genes/enzymes was complete. It is clear now that E. coli presents a relatively simple membrane composition not representative for the diversity in membrane lipids present in many other bacteria (Table 1).

Diversity of membrane lipids in bacteria In addition to (and sometimes instead of) the E. coli lipids PE, PG and CL, other bacteria can form a variety of lipids. Many synthesize the methylated PE derivatives monomethyl PE (MMPE), dimethyl PE (DMPE) and phosphatidylcholine (PC). In most bacteria, MMPE is simply an intermediate in the formation of PC, but in a few species such as Clostridium butyricum, Proteus vulgaris, Yersinia pestis and Xanthomonas campestris, MMPE is the end product of the pathway (Goldfine and Ellis 1964; Tornabene 1973; Moser, Aktas and Narberhaus 2014b). The capacity to form PI, another membrane lipid considered typical for eukaryotes, is widespread in actinomycetes, and several δ-proteobacteria also accumulate it. At least in Mycobacterium tuberculosis, PI (and/or its derivatives) is an essential lipid (Jackson, Crick and Brennan 2000). PI is often further modified with mannose residues and fatty acid residues. These simple acyl-phosphatidylinositol mannosides (AcPIMs) can be further elongated to form higher PIM species and the lipoglycans lipomannan and lipoarabinomannan which are important molecules implicated in host–pathogen interactions in the course of tuberculosis and leprosy (Berg et al. 2007; Kaur et al. 2009; Guerin et al. 2010). Dialkylether-glycero-3-phosphoinositol has been described recently in the thermophilic CytophagaFlavobacterium-Bacteroidetes (CFB) group bacterium Rhodothermus marinus. This lipid might be also present in Sphingobacteria and in several α-proteobacteria (Jorge, Borges and Santos 2014). A few bacteria also accumulate the anionic lipid PS which is an intermediate in the synthesis of PE. Normally Psd is very efficient in converting PS into PE, leaving hardly detectable amounts of PS. The δ-proteobacterium Bdellovibrio bacteriovorans is one example where significant amounts of PS were detected (Nguyen, Sallans and Kaneshiro 2008). PS is also a major lipid in C. botulinum and a few Flavobacterium species (Evans et al. 1998; Lata, Lal and Lal 2012). The headgroup of PG can be modified by transfer of amino acids and in most cases the transferred amino acids are lysine, alanine or arginine thereby forming LPG, alanyl-PG (APG) or arginyl-PG (ArPG), respectively (Roy, Dare and Ibba 2009; Roy and Ibba 2009). Originally, these aminoacyl modifications of PG had been described only in Gram-positive bacteria such as Staphylococcus aureus, Bacillus subtilis, Lactococcus plantarum and Listeria monocytogenes (den Kamp, Redai and van Deenen 1969; Nahaie et al. 1984; Fischer and Leopold 1999), but recently aminoacylated PG derivatives were also identified in a few Gram-negative bacteria, examples being Rhizobium tropici and Pseudomonas aeruginosa (Sohlenkamp et al. 2007; Klein et al. 2009; Hebecker et al. 2011; Arendt et al. 2012). The glycerol headgroup of PG can also be acylated (Kobayashi et al. 1980). The enzyme responsible for this acyltransferase reaction is not known. Aminoacylated versions of CL with alanine or lysine are formed in different Listeria species (Peter-Katalinic and Fischer 1998; Fischer and Leopold 1999; Dare et al. 2014). There are also membrane lipids that are not phospholipids, meaning that their structure lacks a phosphate group. Several of them present a DAG backbone, such as glycosylated DAGs, DGTS and the sulfolipid sulfoquinovosyl diacylglycerol (SQD). The presence of glycosylated DAG derivatives (GLs) has been known for many years in Gram-positive bacteria, and glycosyltransferases involved in the synthesis of these lipids were identified (Jorasch et al. 1998). In recent years, the presence of GLs has been described in a few α-proteobacteria such as Mesorhizobium loti, Agrobacterium tumefaciens and Rhodobacter sphaeroides when these bacteria are grown in media with limiting

Sohlenkamp and Geiger

3

Figure 1. Synthesis of membrane lipids in bacteria. This figure gives an overview about metabolic routes described in the context of membrane lipid formation in bacteria. The reactions catalyzed by the different enzymes are explained in detail in the text. Abbreviations: G3P-glycerol-3-phosphate; LPA-lysophosphatidic acid; PA-phosphatidic acid; DAG-diacylglycerol; CDP-DAG-cytidine diphosphate-diacylglycerol; PS-phosphatidylserine; PE-phosphatidylethanolamine; MMPEmonomethyl PE; DMPE-dimethyl PE; PC-phosphatidylcholine; LPC-lysophosphatidylcholine; GPC-glycerophosphocholine; PGP-phosphatidylglycerol phosphate; PGphosphatidylglycerol; CL-cardiolipin; LCL-lysyl-cardiolipin; ACL-alanyl-cardiolipin; LPG-lysyl-phosphatidylglycerol; APG-alanyl-phosphatidylglycerol; ArPG-arginylphosphatidylglycerol; PIP-phosphatidylinositol phosphate; PI-phosphatidylinositol; PIM-phosphatidylinositol mannoside; PIM2 -phosphatidylinositol dimannoside; Ac1 PIM2 -acyl PIM2 ; Ac2 PIM2 -diacyl PIM2 ; DGHS-diacylglyceryl homoserine; DGTS-diacylglyceryl-N,N,N-trimethylhomoserine; SQD-sulfoquinovosyl diacylglycerol; GTFglycosyltransferase.

¨ phosphate concentrations (Benning, Huang and Gage 1995; Holzl ¨ ¨ et al. 2005; Holzl and Dormann 2007; Devers et al. 2011; Geske et al. 2013; Diercks et al. 2015; Semeniuk et al. 2014). There is a large structural diversity within sugar headgroups with respect to sugar monomers, anomeric forms, linkage between sugars and covalent modification of sugars, which will not be discussed within the present review. Glycophospholipids (GPLs) form a special class of DAG-derived GLs. They have been described mainly in a few Gram-positive bacteria. One important example is the radiation-resistant Deinococcus radiodurans, where one of the major membrane lipids is 2 -O-(1,2-diacyl-sn-glycero-3phospho)-3 -O-(α-galactosyl)-N-D-glyceroyl alkylamine (Anderson and Hansen 1985). DGTS is a betaine lipid that occurs in a variety of lower green plants, chromophytes, fungi and amoebae. Often there is

reciprocity between PC and DGTS content, meaning that in most cases when PC is a major lipid no DGTS is detected and vice versa. Only some algae seem to be the exception to the rule and accumulate PC and DGTS at the same time (Kato et al. 1996). The occurrence of DGTS in bacteria is limited. The presence of DGTS has been shown in some α-proteobacteria and DGTS only accumulates in these bacteria under conditions of phosphate limitation replacing PC (Benning, Huang and Gage 1995; Geiger et al. 1999). Other membrane lipids present in bacteria such as OLs, sulfonolipids, HOPs or sphingolipids (SLs) do not present a DAG backbone. OL has not been found in eukaryotes or archaea ´ (Geiger et al. 2010; Vences-Guzman, Geiger and Sohlenkamp 2012). It is predicted to be present in many proteobacteria, bacteria belonging to the CFB group and actinomycetes.

4


Figure 2. PE synthesis in bacteria. PE can be synthesized either via the Pss/Psd pathway or via CL/PE synthase. Pss-phosphatidylserine synthase; Psd-phosphatidylserine decarboxylase; CL/PE synthase-cardiolipin/phosphatidylethanolamine synthase; CMP-cytidine monophosphate; CDP-cytidine diphosphate.

Interestingly, while in some bacteria OLs are only formed under conditions of phosphate limitation, in other (and often closely related) bacteria OLs are formed constitutively (Geiger et al. 1999; ´ et al. 2011, 2013). In a few bacteria, structural Vences-Guzman analogs to OL in which ornithine is replaced by lysine, glycine, serineglycine or glutamine have been described (Tahara et al. 1976; Kawai, Yano and Kaneda 1988; Kawazoe et al. 1991; Batrakov et al. 1999; Zhang, Ferguson-Miller and Reid 2009). In recent years, several structurally distinct OLs either hydroxylated ´ or methylated have been described (Gonzalez-Silva et al. 2011; ´ et al. 2011, 2013; Vences-Guzman, ´ Geiger and Vences-Guzman Sohlenkamp 2012; Moore et al. 2013; Diercks et al. 2015). HOPs are pentacyclic triterpenoid lipids that are present in a wide variety of organisms. A large diversity of HOP structures mainly differing in their side chain structures has been described. It is unclear what the functional importance of this structural diversity is. Due to their structural similarities, HOPs have been considered molecular surrogates of eukaryotic sterols (Saenz et al. 2012a). Consistent with this, it has been observed that HOPs are involved in enhancing the stability and impermeability of the bacterial membrane. They are present in cyanobacteria, methylotrophic bacteria, actinomycetes, some αproteobacteria, β-proteobacteria and δ-proteobacteria (Kannenberg and Poralla 1999; Blumenberg et al. 2009; Seipke and Loria 2009; Bradley et al. 2010; Welander et al. 2010, 2012; Schmerk, Bernards and Valvano 2011; Saenz et al. 2012b; Schmerk et al. 2015 ). HOPs are ubiquitous in sedimentary deposits and specific HOPs structures are widely used as markers in organic geochemistry (Welander and Summons 2012; Ricci et al. 2014). SLs are restricted to a few groups of bacteria: bacteria of the order Sphingomonadales, several bacteria belonging to the CFB group and a few δ-proteobacteria such as Myxococcus xanthus, M. stipitatus, Cystobacter fuscus and Sorangium cellulosum (Eckau, Dill and Budzikiewicz 1984; Stein and Budzikiewicz 1988; Keck et al. 2011). A wide variety of SL headgroups has been described, some are sugars whereas others are phosphoethanolamine or inositol derived. Structural diversity can also be detected in the hydrophobic moieties of the membrane lipids. Commonly, the hydropho-

bic moieties of amphiphilic membrane lipids are formed by linear fatty acids that can be saturated or unsaturated (containing often one and rarely two or more double bonds). Especially many firmicutes, actinomycetes and bacteria of the CFB group often contain branched-chain fatty acids, especially of the iso and anteiso types. In many anaerobic bacteria, plasmalogens, 1-O-alk-1 -enyl 2-acyl glycerol phospholipids and GLs are present (Goldfine 2010). In the deep-branching hyperthermophilic bacteria Thermotoga and Aquifex, also alkyl residues instead of acyl residues can be found. They present acyletherglycerol phospholipids (AEGP) and dietherglycerol phospholipids (DEGP) in addition to DAG phospholipids (Sturt et al. 2004).

A taxonomic view at membrane lipid composition In the following paragraph, we will try to make a few generalizations with respect to the membrane lipids present in different bacterial phyla (Table 1). Almost all α-proteobacteria seem to be able to form PG, CL and PE and many can also synthesize PC and OLs. Probably as a consequence of their lifestyles, a few can form SQD, DGTS, GLs, LPG, HOPs and even SLs (Benning, Huang and ´ Gage 1995; Perzl et al. 1997; Lopez-Lara, Sohlenkamp and Geiger 2003; Huang et al. 2009; Palacios-Chaves et al. 2011; Vences´ et al. 2011; Aktas et al. 2014). Most β-proteobacteria Guzman can form PE, PG, CL and OLs, while a few probably can synthesize the cationic lipid LPG. Lyso-PE, a deacylated version of PE has also been identified in several β-proteobacteria. Widespread among bacteria of the family Burkholderiaceae is the capacity to form PE and/or OLs hydroxylated in the 2-position of a fatty acid (Thiele and Schwinn 1973; Yabuuchi et al. 1995; Taylor, Ander´ son and Wilkinson 1998; Rahman et al. 2000; Gonzalez-Silva et al. 2011). All bacteria belonging to the genus Burkholderia seem to be able to form HOPs (Schmerk, Bernards and Valvano 2011; Malott et al. 2012). Most γ -proteobacteria accumulate PG, CL and PE, while a few can form OLs, PC, HOPs, LPG or APG (Neunlist and Rohmer 1985; Raetz and Dowhan 1990; Conover et al. 2008; Klein et al. 2009; Giles et al. 2011; Lewenza et al. 2011; Moser, Ak´ et al. 2014). Among tas and Narberhaus 2014b; Vences-Guzman the δ-proteobacteria, the ability to form PG and PE seems to be


5

Figure 3. Synthesis of the anionic membrane lipids PG and CL and their acylated derivatives. Only one pathway for PG synthesis has been described. PG then can serve as a precursor for one of the three described Cls pathways or alternatively PG or CL can be (amino-) acylated. CMP-cytidine monophosphate; CDP-cytidine diphosphate; PgsA-phosphatidylglycerolphosphate synthase; PgpA/PgpB/PgpC-phosphatidylglycerolphosphate phosphatase; ClsA-cardiolipin synthase (bacterial type); ClsCcardiolipin synthase; Cls-II-cardiolipin synthase (eukaryotic type); CL/PE synthase-cardiolipin/phosphatidylethanolamine synthase; MprF/LpiA-aminacyl-PG synthase; G3P-glycerol-3-phosphate; Lys-tRNALys -lysyl tRNA charged with the amino acid lysine; tRNALys -uncharged lysyl tRNA.

6


Table 1. Diversity of membrane lipids in a selection of eubacteria. The order of appearance of the lipids in the table does not reflect abundance of lipids. PG-phosphatidylglycerol; CL-cardiolipin; PS-phosphatidylserine; PE-phosphatidylethanolamine; PE-2OH-PE hydroxylated in the 2-position of the fatty acid; MMPE-monomethyl PE; DMPE-dimethyl PE; PC-phosphatidylcholine; PA-phosphatidic acid; OL-ornithine lipid; S2-OL hydroxylated in ornithine headgroup; P1-OL hydroxylated in the 2-position of the ester-bound fatty acid; P2-OL hydroxylated in ornithine headgroup and in 2-position of ester-bound fatty acid; NL1-OL hydroxylated in the 2-position of the amide-bound fatty acid; NL2-OL hydroxylated in the 2-positions of the amide- and ester-bound fatty acids; TMOL-N,N,N-trimethyl OL; GCL-glycine-containing lipid; SL-sphingolipid; DGTS-diacylglyceryl-N,N,N-trimethylhomoserine; SQD-sulfolipid sulfoquinovosyl diacylglycerol; SOL-sulfonolipid; PT-phosphatidylthreonine; GL-glycolipid; GPL-glycophospholipid; LPG-lysyl-phosphatidylglycerol; APG-alanyl-phosphatidylglycerol; LCL-lysyl-cardiolipin; ACL, alanylcardiolipin; APT-phosphoaminopentanetetrol lipid; DEGP-dietherglycerophospholipid; AEGP-acyletherglycerophospholipid; ST-sterols; PLplasmalogens; HOP-hopanoids. A lipid name in italics indicates that the respective organism is predicted to be able to form the lipid based on its genome sequence.

Taxonomic group Species

Major membrane lipids described/predicted

References

Proteobacteria Alpha Agrobacterium tumefaciens Sinorhizobium meliloti

Rhizobium tropici Brucella melitensis Rhodobacter sphaeroides Caulobacter crescentus

PG, CL, PE, MMPE, DMPE, PC, OL, S2, GL, DGTS, LPG PG, CL, PE, MMPE, DMPE, PC, OL, DGTS, SQD PG, CL, PE, MMPE, DMPE, PC, OL, S2, P1, P2, DGTS, LPG PG, CL, PE, PC, OL, LPG PG, CL, PE, PC, OL, DGTS, SQD, GL PG, CL, LPG, acyl-PG, GL

Sphingomonas sp.

PG, CL, PE, MMPE, DMPE, PC, SL, OL, LPG, DGTS, HOP Methylobacterium extorquens PG, PE, PC, HOP Bradyrhizobium diazoefficiens PG, CL, PE, MMPE, DMPE, PC, (formerly B. japonicum) HOP Rhodopseudomonas palustris PG, CL, PE, PC, HOP

´ et al. (2013); Aktas et al. Geske et al. (2013); Vences-Guzman (2014); Semeniuk et al. (2014); Wessel et al. (2006) ´ de Rudder, Lopez-Lara and Geiger (2000); de Rudder, ´ Sohlenkamp and Geiger (1999); Lopez-Lara, Sohlenkamp and Geiger (2003) ´ Rojas-Jimenez et al. (2005); Sohlenkamp et al. (2007); ´ et al. (2011) Vences-Guzman Palacios-Chaves et al. (2011); Bukata et al. (2008); Comerci et al. (2006); Conde-Alvarez et al. (2006) Catucci et al. (2004); Benning et al. (1993); Benning, Huang and Gage (1995); Zhang, Ferguson-Miller and Reid (2009) Contreras, Shapiro and Henry (1978); Jones and Smith (1979); De Siervo and Homola (1980) ¨ Kawahara, Kuraishi and Zahringer (1999); Rowe et al. (2000); Huang et al. (2009); Zhang et al. (2005) Bradley et al. (2010); Ostafin et al. (1999); Welander et al. (2010) Bravo et al. (2001); Hacker et al. (2008); Minder et al. (2001); Perzl et al. (1997); Silipo et al. (2014) Ramana et al. (2012); Welander et al. (2012)

Beta Burkholderia cenocepacia

PG, CL, PE, PE-2OH, OL, P1, NL1, NL2, LPG, HOP

Ralstonia solanacearum Neisseria gonorrhoeae

PG, PE, PE-2OH, lyso-PE, OL, LPG PG, CL, PE, lyso-PE

Bordetella pertussis

PG, CL, PE, OL, lyso-PE

Escherichia coli Pseudomonas aeruginosa

PG, CL, PE PG, CL, PE, PC, OL, APG

Vibrio cholerae Salmonella typhimurium Serratia proteamaculans Legionella pneumophila Xanthomonas campestris Acidithiobacillus ferroxidans

PG, CL, PE, OL PG, CL, PE, acyl-PG PG, CL, PE, OL, NL1 PG, CL, PE, MMPE, DMPE, PC PG, CL, PE, MMPE, PC, LPG PG, CL, PE, MMPE, DMPE, PC, OL, HOP PG, CL, PE, PC, lyso-PE PG, PE, ST, HOP

´ Gonzalez-Silva et al. (2011); Taylor, Anderson and Wilkinson (1998); Schmerk, Bernards and Valvano (2011); Schmerk et al. (2015) Yabuuchi et al. (1995) Rahman et al. (2000); Senff et al. (1976); Sud and Feingold (1975) Kawai and Moribayashi (1982); Kawai, Moribayashi and Yano (1982); Thiele and Schwinn (1973)

Gamma

Francisella tularensis Methylococcus capsulatus

Parsons and Rock (2013); Dowhan (2013) Lewenza et al. (2011); Wilderman et al. (2002); Klein et al. (2009); Pramanik et al. (1990) Giles et al. (2011) Ames (1968); Olsen and Ballou (1971) ´ et al. (2014) Vences-Guzman Conover et al. (2008) Moser et al. (2014a); Moser, Aktas and Narberhaus (2014b) Barridge and Shively (1968); Dees and Shively (1982); Jones et al. (2012) Li, Wang and Ernst (2011) Lamb et al. (2007); Welander and Summons (2012); Fang, Barcelona and Semrau (2000)

Delta Desulfovibrio sp. Myxococcus xanthus Stigmatella aurantiaca Bdellovibrio bacteriovorus

PG, CL, PS, lyso-PS, PE, OL, DGTS, HOP PG, CL, PE, SL, PI, PL PG, PE, lyso-PE, PI, SL, ST, OL, LPG PG, CL, PS, PE, PT, acyl-PE, SL

Makula and Finnerty (1974); Makula and Finnerty (1975); Blumenberg et al. (2009) Lorenzen et al. (2014); Ring et al. (2009) Ring et al. (2009); Caillon, Lubochinsky and Rigomier (1983); Bode et al. (2003) Muller et al. (2011); Nguyen, Sallans and Kaneshiro (2008); ¨ Steiner, Conti and Lester (1973)


7

Table 1 (Continued.)

Taxonomic group

Species

Major membrane lipids described/predicted

Helicobacter pylori

PG, CL, PE, lyso-PE

Campylobacter jejuni

PG, PE

References

Epsilon Zhou et al. (2012); Olofsson et al. (2010); Tannaes et al. (2000, 2005) Leach, Harvey and Wait (1997)

Cyanobacteria Synechococcus sp. PCC 7942 PG, SQD, GL, CL Synechocystis sp. PCC 6803 PG, SQD, GL, HOP

Guler et al. (1996) ¨ Yuzawa et al. (2014)

Deinococcus radiodurans

GL, GPL

Anderson and Hansen (1985); Huang and Anderson (1989, 1995)

PG, CL, SQD, GL

Kenyon and Gray (1974); Mizoguchi et al. (2013)

DEGP, AEGP, APT

Jahnke et al. (2001); Sturt et al. (2004)

DEGP, AEGP, GL

Manca et al. (1992); Damste´ et al. (2007); Huber et al. (1986)

PG, CL, PE, PL

Hagen (1974)

PE, PC, host-derived membrane lipids

Yao et al. (2014); Hatch and McClarty (1998)

Deinococcales

Chloroflexi (green non-sulfur bacteria) Chloroflexus aurantiacus Aquificae Aquifex aeolicus Thermotogae Thermotoga maritima Fusobacteria Fusobacterium mortiferum Chlamydiae Chlamydia pneumonia CFB Group

Chlorobium (green sulfur bacteria) Actinobacteria

Flavobacterium johnsoniae PE, PC, OL, P1, SOL, GCL Bacteroidetes thetaiotamicron PG, CL, PS, PE, SL Porphyromonas gingivalis PG, PE, SL

Pitta, Leadbetter and Godchaux (1989); Kawazoe et al. (1991) Smith and Salyers (1981) Nichols and Rojanasomsith (2006); Nichols et al. (2004, 2006); Tavana et al. (2000)

Chlorobium tepidum

PG, CL, PE, GL

Mizoguchi et al. (2013); Sorensen, Cox and Miller (2008)

Mycobacterium tuberculosis

PG, CL, PE, PI, PIM, OL

Nocardia sp.

CL, PE, PI, PIM, OL, SQD, HOP

Streptomyces coelicolor

PG, CL, PE, PI, PIM, OL, HOP

Corynebacterium glutamicum PG, CL, PI, PIM

Khuller et al. (1982); Jackson, Crick and Brennan (2000); Berg ´ et al. (2007); Laneelle et al. (1990) Khuller and Brennan (1972); Trana, Khuller and Subrahmanyam (1980) ´ et al. (2009); Jyothikumar et al. (2012); Liu Sandoval-Calderon et al. (2014) Nampoothiri et al. (2002)

Borrelia burgdorferi Treponema denticola Leptospira interrogans

PG, PC, GL PG, CL, PE, PC PG, PE, CL, Lyso-PE, GL, OL

Wang, Scagliotti and Hu (2004); Ostberg et al. (2007) Kent et al. (2004) Livermore and Johnson (1974); Moribayashi et al. (1991)

Singulisphaera acidiphila Rhodopirellula baltica

PG, PC, TMOL, PA, HOP PG, CL, PE, PC, DGTS

Moore et al. (2013) Bondoso et al. (2014); Roh et al. (2013)

Bacillus subtilis

PG, CL, PE, LPG, GL, GPL

Listeria monocytogenes

PG, CL, LPG, LCL, PI

Staphylococcus aureus Lactobacillus sp. Streptococcus pneumoniae Clostridium perfringens Mycoplasma genitalium

PG, CL, LPG, GPL PG, CL, GL, LPG PG, CL, GL PG, PE, APG, LPG, PL PG, CL, GL, host-derived lipids

Gidden et al. (2009); den Kamp et al. (1969); Salzberg and Helmann (2008); Jorasch et al. (1998) Fischer and Leopold (1999); Dare et al. (2014); Thedieck et al. (2006); Mastronicolis et al. (2008) Oku et al. (2004); White and Frerman (1967) Iwamori et al. (2011); Sauvageau et al. (2012) ´ Laneelle ´ ´ Trombe, and Laneelle (1979); Meiers et al. (2014) Roy and Ibba (2008); Johnston, Baker and Goldfine (2004) Kornspan and Rottem (2012)

Spirochetes

Planctomycetes

Firmicutes

conserved and a variety of uncommon lipids can be detected in some cases, such as SLs, PI, HOPs or even sterols. Also, a few bacteria can form OLs, LPG or DGTS and even phosphatidylthreonine (PT) has been described in B. bacteriovorus (Makula and Finnerty 1974; Nguyen, Sallans and Kaneshiro 2008; Blumenberg et al. 2009; Ring et al. 2009; Keck et al. 2011; Lorenzen et al. 2014).

In general, -proteobacteria have a far less complex lipid composition with PE, PG and often CL present (Leach, Harvey and Wait 1997; Tannaes, Grav and Bukholm 2000; Tannaes, Bukholm and Bukholm 2005; Zhou et al. 2012). The fatty acids present in complex lipids of the proteobacteria are mostly of the linear type, whereas branched-chain fatty acids can only rarely be found.

8


Cyanobacterial membranes mostly contain PG, SQD and the GLs monogalactosyl-DAG and digalactosyl-DAG (Guler et al. ¨ 1996). The presence of CL has not been reported, but several genomes encode a putative Cls of the PLD superfamily. HOPs are present in several species (Doughty et al. 2009; Saenz et al. 2012b). The membrane of the radiation-resistant bacterium D. radiodurans has an unusual lipid composition, because a diversity of GLs and GPLs has been described, whereas the synthesis pathways of the common phospholipids PG, PE and CL seem to be absent (Huang and Anderson 1989; Huang and Anderson 1995; Makarova et al. 2001). In the membranes of the green non-sulfur bacterium Chloroflexus aurantiacus, the lipids PG, CL, SQD and several GLs have been detected (Kenyon and Gray 1974; Mizoguchi et al. 2013). The deep-branching hyperthermophiles Aquifex aeolicus and Thermotoga maritima present unusual structures when compared to the better studied bacteria. Their membrane lipids are not exclusively DAG based but an important fraction is formed by DEGP and AEGP. Although the presence of ether lipids is considered a typical trait of archaebacteria, Aquifex and Thermotoga clearly present bacterial-type lipids. The fatty acyl and alkyl group are in the sn-1- and 2-positions like in bacterial lipids and not in the sn-2- and 3-positions as in archaea. Little is known about their lipid headgroups, but in Aquifex phosphoaminopentanetetrol headgroups and in Thermotoga sugar headgroups have been described (Manca et al. 1992; Sturt et al. 2004; Damste´ et al. 2007). Among the bacteria of the CFB group PE, PG, CL and OLs again seem to be relatively common, but also unusual lipids can be found: several species can form SLs or sulfonolipids and Flavobacterium/Cytophaga can form glycine-containing and/or serineglycine-containing lipids (Tahara et al. 1976; Kawai, Yano and Kaneda 1988; Pitta, Leadbetter and Godchaux 1989; Kawazoe et al. 1991). Dialkylether-glycero-3-phosphoinositol has been described recently in the thermophilic bacterium R. marinus and might be also present in Sphingobacteria (Jorge, Borges and Santos 2014). The green sulfur bacterium Chlorobium tepidum accumulates PG, CL, PE and GLs, but apparently does not form SQD, which is in general omnipresent in photosynthetic organisms (Sorensen, Cox and Miller 2008; Mizoguchi et al. 2013). Within the actinomycetes, many bacteria can form PG, PE and CL and in addition the capacity to synthesize PI and its mannosylated/acylated derivatives is widespread. Some actinomycetes are known to form OLs and several more are predicted to be able to form OLs. All species belonging to the genus Streptomyces seem to be able to form HOPs. Often the fatty acids present in the complex lipids are branched-chain fatty acids (Khuller and Brennan 1972; Jackson, Crick and Brennan 2000; Nampoothiri ´ et al. 2009; Seipke and Loria 2009). et al. 2002; Sandoval-Calderon Among the spirochaetes, the distinct genera present different lipid compositions: Borrelia forms PG, PC and GLs, Treponema forms PG, PE, CL and PC, and Leptospira forms PE, PG, CL and lysoPE. Bacteria of the genus Leptospira have also been described to form GLs and are predicted to be able to form OLs (Livermore and Johnson 1974; Moribayashi et al. 1991; Kent et al. 2004; Wang, Scagliotti and Hu 2004; Ostberg et al. 2007). Planctomycetes form a group of bacteria containing a special type of cell division and intracellular membrane structures (Fuerst and Sagulenko 2011). Within this group are the ammonox bacteria, with their ladderane lipids instead of normal fatty acids (Sinninghe Damste´ et al. 2005; Boumann et al. 2006). The lipid composition within each genus seems to differ and some Planctomycetes can form PG, PE, PC, CL and HOPs. Re-

cently, a N,N,N-trimethyl OL has been described in Singulisphaera acidiphila (Moore et al. 2013). Among the firmicutes the presence of PE is not universal, but most seem to be able to form PG and CL. In many cases, the presence of aminoacylated PG and CL derivatives has been described. Another feature that is widely distributed is the presence of GLs and GPLs (White and Frerman 1967; den Kamp, Redai and van Deenen 1969; Jorasch et al. 1998; Fischer and Leopold 1999; Johnston, Baker and Goldfine 2004; Sauvageau et al. 2012; Meiers et al. 2014). Most of the firmicutes studied present branchedchain fatty acids (iso-/anteiso-) in their lipid structure, the exception being the lactobacteria which form linear fatty acids. The latter present straight chain saturated, monounsaturated and related cyclopropane fatty acids commonly found in Gramnegative bacteria (Ratledge and Wilkinson 1988). A special group of bacteria is formed by some obligate intracellular pathogens that lack the capacity to synthesize all their membrane lipids and that obtain a part or all of their lipids from the eukaryotic host. This has been described for Chlamydia and for Mycoplasma, among others (Hatch and McClarty 1998; Kornspan and Rottem 2012; Yao et al. 2014).

PATHWAYS FOR PHOSPHOLIPID SYNTHESIS IN BACTERIA Synthesis of the common precursor CDP-DAG G3P acyltransferases transfer acyl groups to the sn-1 and sn2 positions of G3P thereby producing 1,2-diacylglycerol-sn-G3P (PA) (Fig. 1). Two different acyltransferase systems have been described, the PlsB/PlsC and the PlsX/PlsY/PlsC systems. The PlsB/PlsC system was first discovered in E. coli (Lightner et al. 1980; Cooper, Jackowski and Rock 1987; Coleman 1990, 1992), and it is mainly present in enterobacteria, whereas most other bacteria use the PlsX/PlsY/PlsC system for PA synthesis (Lu et al. 2006). CDP-DAG synthase then catalyzes the formation of CDP-DAG from CTP and PA. CDP-DAG is a common precursor used in the synthesis of glycerophospholipids in bacteria. The steps from G3P to CDP-DAG are discussed in detail in a recent review by Parsons and Rock (2013). Most of the enzymes described that use CDP-DAG as substrate belong to either the CDP-alcohol phosphotransferase family or the PLD superfamily. CDP-DAG alcohol phosphotransferases share a common conserved motif (DX2 DGX2 ARX8 GX3 DX3 D in most cases, DX2 DGX2 ARX12 GX3 DX3 D in case of Pcs enzymes) (Williams and ´ McMaster 1998; Sohlenkamp, Lopez-Lara and Geiger 2003; Sol´ısOviedo et al. 2012). Members of this superfamily include phosphatidylinositolphosphate synthase (Pips), phosphatidylcholine synthase (Pcs), type II Pss, eukaryote-type cardiolipin synthase (Cls-II) and PgsA. Members of the PLD superfamily present the conserved motif HXKX4 DX8 G (Liscovitch et al. 2000). This superfamily includes for example prokaryote-like Cls, type I Pss and the CL/PE synthase.

Pathways for PE synthesis PE is a common membrane lipid in proteobacteria and other Gram-negative bacteria. Its synthesis has been first described in E. coli (DeChavigny, Heacock and Dowhan 1991). Pss condenses serine with CDP-DAG leading to the formation of the anionic lipid PS. PS is then decarboxylated by Psd leading to the formation of the zwitterionic lipid PE (Figs 1 and 2, reactions 1 and 2). Pss enzymes belong to two different families: type I Pss are phospholipase D-like proteins whereas type II


proteins belong to the CDP-alcohol phosphotransferase family (Sohlenkamp, de Rudder and Geiger 2004). For example, Pss from E. coli is a type I Pss whereas Pss from B. subtilis and Sinorhizobium meliloti are type II Pss (Okada et al. 1994; Matsumoto 1997; Sohlenkamp, de Rudder and Geiger 2004). Despite the fact that both types of Pss are unrelated on a sequence level, they both use CDP-DAG and L-serine as substrates and form PS and cytidine monophosphate (CMP) as products. Type I Pss is restricted to a few γ -proteobacteria (Enterobacteriaceae, Vibrionaceae, Pasteurellaceae, Alteromonaceae), whereas type II Pss occurs in all three kingdoms of life (Sohlenkamp, de Rudder and Geiger 2004). Mutants deficient in pss or psd have been described in several organisms, such as E. coli, S. meliloti, A. tumefaciens, Helicobacter pylori, etc. (Li and Dowhan 1988; DeChavigny, Heacock and Dowhan 1991; Shi et al. 1993; Ge and Taylor 1997; Matsumoto et al. 1998; Karnezis et al. 2002; Sohlenkamp, de Rudder and Geiger ´ 2004; Vences-Guzman, Geiger and Sohlenkamp 2008). In general, these mutants are viable under specific growth conditions and many share a requirement for millimolar bivalent cations in the growth medium. Psd mutants accumulate PS and in the case of the S. meliloti Psd mutant, a nodulation phenotype was ´ Geiger and Sohlenkamp 2008). Reobserved (Vences-Guzman, cently, Moser et al. (2014a) described a bifunctional CL/PE synthase in the plant pathogen X. campestris that catalyzes the synthesis of PE from CDP-DAG and ethanolamine (Figs 1 and 2, reaction 3). This is a completely new pathway for PE synthesis, also distinct from pathways described in eukaryotes. The same enzyme also functions as a Cls, and as many bacterial Cls, belongs to the PLD superfamily. In eukaryotes, PE synthesis via the CDP-ethanolamine pathway has been described, but this reaction has not been shown to be present in prokaryotes so far (Nakashima, Hosaka and Nikawa 1997).

Pathways for synthesis of PG and aminoacylated PG derivatives PgsA is condensing G3P with CDP-DAG leading to the formation of PGP (Figs 1 and 3, reaction 4) (Miyazaki et al. 1985). All PgsA described belong to the CDP-alcohol phosphotransferase family. Mutants deficient in PgsA have been isolated in E. coli and in cyanobacteria (Miyazaki et al. 1985; Hagio et al. 2000; Kikuchi, Shibuya and Matsumoto 2000; Matsumoto 2001). A gene encoding a putative PgsA enzyme is present in almost all bacterial genomes, one notable exception being D. radiodurans (Makarova et al. 2001). PGP is then dephosphorylated by a PGP phosphatase (Figs 1 and 3, reaction 5) (Lu et al. 2011). The anionic lipid PG can be converted into the cationic lipid LPG in a one-step reaction (Figs 1 and 3, reaction 9). Lysine is transferred from a charged tRNALys to the glycerol headgroup of PG. This biochemical activity was first described in cell-free S. aureus extracts (Gould and Lennarz 1967; Lennarz, Bonsen and van Deenen 1967; Nesbitt and Lennarz 1968). The gene encoding the enzyme MprF responsible for LPG formation has first been described also in S. aureus during a screen for mutants more susceptible than the wildtype to cationic antimicrobial peptides (Peschel et al. 2001). MprF homologs are also present in some Gram-negative bacteria and it could be shown that these homologs indeed caused the formation of LPG or APG, respectively (Sohlenkamp et al. 2007; Klein et al. 2009). Homologs of MprF are also responsible for APG formation in C. perfringens and P. aeruginosa and for ArPG formation in Enterococcus faecium (Roy and Ibba 2008; Klein et al. 2009; Roy and Ibba 2009). Most MprF homologs characterized only produce either APG or LPG, but MprF from E. faecium can catalyze the formation of APG, LPG

9

and ArPG (Roy and Ibba 2009). Further homologs are present in many α-proteobacteria (A. tumefaciens, several species of the genera Brucella, Rhizobium), β-proteobacteria (Burkholderia, Ralstonia) and γ -proteobacteria (X. campestris, Xylella, several species of Pseudomonas and Aeromonas). It is noteworthy that many if not most of the organisms predicted to form aminoacyl PGs under certain conditions can adopt a lifestyle associated with eukaryotic hosts possibly indicating a role for this lipid in hostsymbiont/pathogen/commensal interactions (Geiger et al. 2010). LPG deficiency leads to an increased susceptibility to cationic antimicrobial peptides, and to various antibiotics such as daptomycin, vancomycin and gentamicin (Ruzin et al. 2003; Nishi et al. 2004; Ernst et al. 2009; Hachmann, Angert and Helmann 2009). APG deficiency in P. aeruginosa caused increased susceptibility to a set of cationic antimicrobial peptides, to a few antibiotics and chromium ions (Klein et al. 2009; Arendt et al. 2012). Recently, Dare et al. (2014) showed evidence in L. monocytogenes that the function of LPG extends beyond contributing to the resistance to cationic peptides. LPG protects against osmotic stress and regulates a motility operon and it is hypothesized that MprF-like enzymes are involved in the adaptation of bacteria to a broad range of environmental stresses (Dare et al. 2014). In bacteria outside the firmicutes, the genes encoding MprF/LpiA form an operon with a gene encoding a putative lipase called AcvB/AtvA (Vinuesa et al. 2003; Sohlenkamp et al. 2007). In P. aeruginosa, the AcvB/AtvA homolog PA0919 seems to be responsible for hydrolysis of APG (Arendt et al. 2013).

Pathways for synthesis of CL and aminoacylated CL derivatives For many years, two different pathways for CL synthesis were known: (A) the prokaryote-like pathway in which two molecules PG react forming CL and glycerol (Figs 1 and 3, reaction 6) and (B) the eukaryote-like pathway in which one molecule PG reacts with CDP-DAG leading to the formation of CL and CMP. As the names implied, one route was thought to be specific for prokaryotes whereas the other was thought to be specific for eukaryotes. In E. coli, CL is synthesized by ClsA from two molecules of PG (Nishijima et al. 1988). ClsA from E. coli is a typical prokary´ otic Cls, belonging to the PLD superfamily. Sandoval-Calderon et al. (2009) showed that Streptomyces coelicolor and other actinomycetes use the eukaryote-like pathway to synthesize CL (Figs 1 and 3, reaction 8). Cls from actinomycetes belongs to the CDP-alcohol phosphotransferase superfamily. The so-called prokaryote-like pathway for CL synthesis is not restricted to bacteria, as Trypanosoma cruzei uses a prokaryote-like Cls (belonging to the PLD superfamily) for CL synthesis (Serricchio and Butikofer 2012). Many bacterial genomes present several genes encoding putative Cls; for example, E. coli presents three (clsA, clsB, clsC). Tan et al. (2012) showed that E. coli can synthesize CL from PG and PE by ClsC (Figs 1 and 3, reaction 7). Interestingly, for full Cls activity of ClsC, a second gene called ymdB has to be co-expressed. YmdB has been shown to be a stressresponsive ribonuclease-binding regulator of RNase III activity (Kim, Manasherob and Cohen 2008). How YmdB affects ClsC activity is not clear, but YmdB might involve a stress response activation of ClsC. It is unknown how widespread this ClsCdependent pathway is. Moser et al. (2014a) recently discovered in X. campestris a bifunctional CL/PE synthase belonging to the PLD superfamily that synthesizes CL from PG and CDP-DAG, but that also catalyzed ethanolamine-dependent PE formation. Interestingly, this bifunctional enzyme is larger (70 kDa) than the earlier characterized monofunctional Cls enzymes from E. coli and

10


Arabidopsis thaliana. The gene encoding the bifunctional CL/PE synthase seems to be restricted to the orders Xanthomonadales and Pseudomonadales (Moser et al. 2014a). These new activities described for two different enzymes of the PLD superfamily (ClsC and CL/PE synthase) imply that maybe other members of this family present different activities that have not been found because people haven’t been looking for them. Bacterial genomes contain many genes encoding Cls and other PLD superfamily members that have not been characterized yet. It is possible that Cls already characterized years ago have additional activities and this possibility seems worth exploring. The aminoacylated derivatives of CL alanyl-CL (ACL) and lysyl-CL (LCL) have been described in different Listeria species. An aminoacyl-PG synthase is required for their formation and if LPG formation is impaired in Listeria, also LCL formation is impaired, meaning that there is no dedicated second enzyme required for the aminoacyl transfer to CL (Dare et al. 2014). It is not clear if the amino acid (lysine or alanine) is transferred to CL or if a Cls can use the aminoacylated PG as substrate and convert it into LCL or ACL (Figs 1 and 3, reactions 11a and b).

Pathways for PC synthesis In eukaryotes, two different pathways for PC synthesis have been described, the CDP-choline (or Kennedy) pathway and the N-methylation pathway. In the CDP-choline pathway, choline is phosphorylated by choline kinase and adenosine triphosphate. In a second step, phosphocholine and CTP react to CDP-choline in a reaction catalyzed by CTP: phosphocholine cytidylyltransferase and finally CDP-choline: 1,2-diacylglycerol cholinephosphotransferase (CPT1) catalyzes a reaction between CDP-choline and DAG leading to the formation of PC and CMP. In the N-methylation pathway, PE is N-methylated three times leading to the formation of PC. Only few bacteria were known to be able to synthesize PC and for a long time only the Nmethylation pathway for PC synthesis was known in prokary´ otes (Kaneshiro and Law 1964; Sohlenkamp, Lopez-Lara and Geiger 2003) (Figs 1 and 4, reactions 12a–c). The genes encoding the bacterial Pmt enzymes were discovered first in R. sphaeroides and later in S. meliloti (Arondel, Benning and Somerville 1993; ´ de Rudder, Lopez-Lara and Geiger 2000). Both Pmts share little sequence similarity and are proposed to belong to two different methyltransferase families. The similarity on amino acid level is restricted to the putative S-adenosylmethionine (SAM)binding site and the S. meliloti Pmt shows homology to rRNA methylases, whereas the R. sphaeroides Pmt shows homology to ubiquinone/menaquinone methyltransferases. Both methyltransferases can catalyze all three methylation steps required for PC formation from PE. Later it was observed that the phospholipid N-methylation pathway is more complex in Bradyrhizobium japonicum and probably other bacteria (Hacker et al. 2008). Bradyrhizobium japonicum presents four methyltransferases that can use PE or its methylated derivatives MMPE and DMPE as substrates (PmtA, PmtX1, PmtX3 and PmtX4) when expressed alone or together with other Pmts in E. coli. PmtA preferentially converts PE to MMPE whereas PmtX1 carries out the subsequent methylation steps. PmtX3 and PmtX4 can methylate PE to MMPE and DMPE when expressed in E. coli. The use of multiple Pmt enzymes has been also described in the yeast Saccharomyces cerevisae which uses two different methyltransferases that together cause the formation of PC from PE (Kodaki and Yamashita 1987; Kanipes and Henry 1997). In the late 1990s the PC synthase (Pcs) pathway was discovered in the root nodule-forming α-proteobacterium S. meliloti

(de Rudder, Sohlenkamp and Geiger 1999; Sohlenkamp et al. 2000) that uses CDP-DAG and choline as substrate (Figs 1 and 4, reaction 13). Pcs belongs to the CDP-alcohol phosphotransferase superfamily. It presents a slightly modified CDP-alcohol phosphotransferase motif that makes the bioinformatic prediction of Pcs-encoding genes straightforward (see above). In contrast, the prediction of N-methyltransferases involved in PC synthesis is much more difficult. Looking for the presence of either the Pmt or the Pcs pathway in bacteria using genomic sequence data, it has been estimated that about 15% of bacteria are ´ probably able to form PC (Sohlenkamp, Lopez-Lara and Geiger ´ 2003; Geiger, Lopez-Lara and Sohlenkamp 2013). The Pcs pathway is widely distributed in α-proteobacteria (mainly Rhodobacterales and Rhizobiales) and absent in β-proteobacteria. Within the γ -proteobacteria, it is present in most Pseudomonas species and in the genera Legionella, Halomonas and Francisella and within the δ-proteobacteria only a few strains of Desulfovibrio present a putative pcs gene. Genes encoding Pcs can also be found outside the proteobacteria. Most noteworthy is probably that most strains of Borrelia, several Actinomyces sp. strains (Actinomycetales) ´ and several Planctomycetales have Pcs (Geiger, Lopez-Lara and Sohlenkamp 2013). Kent et al. (2004) showed evidence for the presence of a CDPcholine pathway in the spirochaete Treponema denticola. In the case of Treponema, the first two steps of the CDP-choline pathway are catalyzed by the bifunctional enzyme LicCA responsible for the formation of CDP-choline (Rock et al. 2001; Campbell and Kent 2001; Kwak et al. 2002). Homologs of LicCA are also present in Haemophilus influenzae, different strains of Clostridium and Streptococcus, but the latter are not known to form PC (Weiser, Love and Moxon 1989; Zhang et al. 1999; Shimizu et al. 2002). The gene/enzyme responsible for catalyzing the final step of the pathway leading to the formation of PC in Treponema has yet to be identified, but it should catalyze the same reaction as the Cpt enzyme responsible for the final step of the eukaryotic CDP-choline pathway (Figs 1 and 4, reaction 16). Such a prokaryotic CDP-choline pathway is distinct from the eukaryotic pathway because at least for the first two steps it uses unrelated enzymes. It is not clear if such a prokaryotic CDP-choline pathway is present outside the genus Treponema. Recently, Moser, Aktas and Narberhaus (2014b) found that X. campestris can form PC using glycerolphosphocholine (GPC) as a substrate for PC formation (Figs 1 and 4, reactions 14 and 15). A similar GPC-dependent pathway for PC formation has been described in the yeast S. cerevisiae (Stalberg et al. 2008). Two acyltransferase reactions are required to form PC from GPC. In both, yeast and X. campestris only enzymes for the second acyltransferase step from LPC to PC were identified, whereas it is not known which activity is catalyzing the first step. At present, it is not clear how widespread such a GPC-dependent pathway for PC synthesis is. Bacterial mutants deficient in PC formation have been described in several species. Agrobacterium tumefaciens mutants deficient in PC formation (pcs− /pmt− ) have a defective type IV secretion system and do not cause tumor formation on Kalanchoe leaves (Wessel et al. 2006). Legionella pneumophila PC-deficient mutants (pcs− /pmt− ) show reduced virulence due to defects in the type IVB secretion system and macrophage adhesion (Conover et al. 2008). Brucella abortus mutants deficient in PC formation (pcs− ) show reduced virulence in mice (Comerci et al. 2006). Sinorhizobium meliloti mutants deficient in PC formation (pcs− /pmt− ) cannot form nodules on their host plant alfalfa and they are sensitive to hypo- and hyperosmotic growth conditions ´ ´ (Sohlenkamp, Lopez-Lara and Geiger 2003; Geiger, Lopez-Lara


11

Figure 4. Synthesis of PC in bacteria. Four different pathways for PC synthesis have been described in bacteria. Pmt-phospholipid N-methyltransferase; Pcsphosphatidylcholine synthase; Cpt1-choline phosphotransferase; CMP-cytidine monophosphate; CDP-cytidine diphosphate; SAM-S-adenosylmethionine; SAHC-Sadenosylhomocysteine; CoA-coenzyme A.

12


and Sohlenkamp 2013). Bradyrhizobium japonicum mutants with a reduced PC content (pmtA− ) can form nodules on their host plant soybean, but they show a reduced nitrogen-fixation activity (Minder et al. 2001).

Pathway for PI and PIM synthesis Similar as PC, also PI has been considered a typical eukaryotic lipid. Therefore, it is probably not surprising that there are only few bacteria that can form PI. PI formation is widespread within the order Actinomycetales, but outside this order only a few bacteria such as Myxococcus have been described to form PI. Eukaryotes form PI using CDP-DAG and inositol as substrates, forming PI in a one-step reaction (Antonsson 1997; Nikawa and Yamashita 1997; Xue et al. 2000). It was thought for many years that the same reaction should be occurring in bacteria, but recently it was shown for a few bacterial species that a Pips enzyme catalyzes the reaction between inositol-1-phosphate and CDP-DAG, leading to the formation of phosphatidylinositol phosphate (PIP) (Figs 1 and 5A, reaction 17). In a second step, PIP is dephosphorylated by PIP phosphatase (Pipp) to PI (Morii et al. 2010, 2014) (Figs 1 and 5A, reaction 18). PI can be further modified with mannose residues and acyl groups forming the PIM (Figs 1 and 5A, reactions 19–22). These are the structural basis of the lipoglucans lipomannan and lipoarabinomannan (Guerin et al. 2010). PI is essential in Mycobacterium (Jackson, Crick and Brennan 2000). The first step of PIM formation in M. tuberculosis is catalyzed by the mannosyltransferase PimA and consists in the transfer of a mannose residue from GDP-mannose to the 2-position of the myo-inositol ring (Hill and Ballou 1966). Subsequently, the mannosyltransferase PimB transfers a mannose residue from GDP-mannose to the 6-position of the myoinositol ring of PIM (Guerin et al. 2009). Both PIM1 and PIM2 can be acylated at the 6-position of the first mannose residue transferred by the acyltransferase Rv2611c, although PIM2 seems to be the favored substrate (Kordulakova et al. 2003; Guerin et al. 2009; Svetlikova et al. 2014). The acyltransferase responsible for the second acylation step has not been identified. Ac1 PIM2 and Ac2 PIM2 can then be further elongated with additional mannose residues thereby forming higher PIM species. Their structures and synthesis pathways are discussed in detail by Guerin et al. (2009). PI and mannosides derived thereof are also present in many other genera such as Streptomyces, Nocardia or Corynebacterium. The PIM structures and their biosyntheses have not been studied in detail outside the genus Mycobacterium and it is certainly possible that there are differences. Recently, a novel pathway for the synthesis of inositol phospholipids was described in the thermophilic bacterium R. marinum (Jorge, Borges and Santos 2014). L-myo-inositol1-phosphate cytidylyltransferase catalyzes the formation of CDP-inositol from inositol-1-phosphate and CTP. Bacterial dialkylether glycerophosphoinositol synthase (BEPIS) (Fig. 5B, reaction 23) then uses CDP-inositol and dialkylether glycerols to produce phosphoinositol ether lipids.

PATHWAYS FOR SYNTHESIS OF PHOSPHORUS-FREE MEMBRANE LIPIDS IN BACTERIA Synthesis of the common precursor DAG DAG can be produced by a set of metabolic reactions in the cell. Phospholipase C action upon phospholipids will produce DAG and the phosphorylated headgroup, for example PC will

be cleaved into DAG and phosphocholine. The headgroup transfer from phospholipids to membrane-derived oligosaccharides or periplasmic cyclic glucans is also a source of DAG. For example, phosphoglycerol is transferred from PG to periplasmic glucans, leading to the formation of DAG (Jackson, Bohin and Kennedy 1984; Fiedler and Rotering 1985; Zavaleta-Pastor et al. ´ 2010; Geiger, Lopez-Lara and Sohlenkamp 2013). Alternatively, PA could be dephosphorylated to DAG by PA phosphatases (Berg and Wieslander 1997; Carman 1997; Zhang et al. 2008; Comba et al. 2013), but it is not clear how relevant this reaction is in bacteria.

Glycolipid (GL) synthesis GL synthesis usually starts with the transfer of a sugar monomer from activated sugars like UDP-glucose, UDP-galactose or GDPmannose to DAG, leading to the formation of a monoglycosylDAG. Subsequently, additional sugars can be transferred to the monoglycosyl-DAG leading to more complex GLs. Processive glycosyltransferases are responsible for the transfer of several sugars and can for example catalyze three transferase steps, each time using the GL produced by the earlier step as substrate (Devers et al. 2011). Some bacterial species produce a wide variety of GLs and in some cases the sugar residues can be covalently modified. Several glycosyltransferases from different bacteria have been cloned and characterized, whereas not much is known about the cellular function of the GLs formed (Jorasch et al. 1998; Devers et al. 2011; Geske et al. 2013; Diercks et al. 2015). Readers more interested in GLs and their structures and synthe¨ ¨ ses are referred to a review by Holzl and Dormann (2007).

Sulfoquinovosyl diacylglycerol (SQD) synthesis A special GL is the anionic lipid SQD which is widely distributed in photosynthetic organisms. Its synthesis has been studied in bacteria and plants. In R. sphaeroides, four genes called sqdA, sqdB, sdqC and sqdD have been identified by genetic analysis to be involved in SQD formation (Benning 1998). SqdA from R. sphaeroides is annotated within the genome sequence as a phospholipid/glycerol acyltransferase. SqdB is thought to be involved in the formation of UDP-sulfoquinovose from UDP-glucose and sulfite. It is the only gene involved in SQD synthesis that has orthologs in all SQD-producing organisms, including plants where it is named SQD1. SqdC and SqdD are probably involved in the transfer of sulfoquinovose to DAG leading to the formation of SQD (Benning 1998). SqdC is annotated as member of the NADdependent epimerase dehydratase family, whereas SqdD is annotated as a glycosyltransferase. In the cyanobacterium Synechococcus sp. PCC 7942, the gene sqdB forms an operon with the gene encoding the lipid glycosyltransferase SqdX. SqdX was later shown to be the cyanobacterial SQD synthase (Guler, Es¨ sigmann and Benning 2000). Plants use an SqdX homolog called SQD2 for SQD synthesis (Shimojima 2011). Mutants deficient in SQD formation have been described in S. meliloti, Synechococcus sp. PCC 7942 and R. sphaeroides (Benning and Somerville 1992a,b; Rossak et al. 1995, 1997; Guler et al. 1996; Weissen¨ mayer, Geiger and Benning 2000). An S. meliloti mutant deficient in SQD formation is not affected during nodule development and function (Weissenmayer, Geiger and Benning 2000). SQD deficiency in R. sphaeroides did not affect photosynthesis in any obvious way, but it absence interfered with bacterial growth under phosphate-limiting conditions (Benning et al. 1993). The Synechococcus mutant deficient in SqdB presented minor changes in a few photosynthesis parameters. Again,


13

Figure 5. Synthesis of PI and PI-derivatives. (A) Synthesis of PI and PI-derivatives in Actinomycetes. (B) CDP-inositol-dependent pathway for dialkylether glycerophosphoinositol formation described in R. marinus. CMP-cytidine monophosphate; CDP-cytidine diphosphate; GMP-guanosine monophosphate; GDP-guanosine diphosphate; Pips-phosphatidylinositolphosphate synthase; Pipp-phosphatidylinositolphosphate phosphatase; PimA-phosphatidylinositol mannosyltransferase; PimB’-phosphatidylinositolmannoside mannosyltransferase; AcylT-phosphatidylinositoldimannoside acyltransferase; PIM2 -PI dimannoside; Ac1 PIM2 -acyl PIM2 ; Ac2 PIM2 -diacyl PIM2 ; BEPIS-bacterial dialkylether glycerophosphoinositol synthase.

14


similarly to the Rhodobacter mutant it showed reduced growth under phosphate-limiting conditions (Guler et al. 1996). ¨

DGTS synthesis Bacteria use two enzymes to synthesize DGTS from DAG. The genes encoding the required activities were first discovered in R. sphaeroides (Klug and Benning 2001). The enzyme BtaA responsible for the formation of diacylglyceryl homoserine (DGHS) is a SAM/DAG 3-amino-3-carboxypropyltransferase. BtaB is a trifunctional N-methyltransferase related on sequence level to bacterial phospholipid N-methyltransferases of the R. sphaeroides family. When BtaA and BtaB from R. sphaeroides or S. meliloti are expressed in E. coli, DGTS is formed (Riekhof, Andre and Benning 2005a). Based on the distribution of BtaA, it can be predicted that the capacity to form DGTS is widespread in a few orders of the α-proteobacteria like the Rhodobacterales, Sphingomonadales and Rhizobiales. Genes encoding BtaA homologs can also be found in the genomes of a few Planctomycetales, γ proteobacteria (Thiotrix, Thiomicrospira), cyanobacteria (Mastigocoleus, Calothrix, Anabaena, Leptolyngba), δ-proteobacteria (Desulfovibrio), firmicutes (Geobacillus, Brevibacillus), Deinococcus and Acidobacterium. In the alga Chlamydomonas reinhardtii, DGTS synthesis is performed by a bifunctional enzyme in which both enzyme functions are fused (Riekhof, Sears and Benning 2005). The N-terminal domain of the protein is BtaB-like whereas the C-terminal domain is BtaA-like. Similar bifunctional enzymes are also present in fungi and other eukaryotes (Riekhof et al. 2014). Sinorhizobium meliloti mutants deficient in DGTS formation were affected in cell yield under phosphate-limiting conditions when the DGTS deficiency was combined with a deficiency in OL ´ synthesis (Lopez-Lara et al. 2005).

OL synthesis The genes encoding the acyltransferases OlsB and OlsA required for OL synthesis in S. meliloti have been described first by Gao et al. (2004) and Weissenmayer et al. (2002). The N-acyltransferase OlsB transfers a 3-hydroxy fatty acid from the constitutive acyl carrier protein AcpP to the α-amino group of ornithine, thereby forming lyso-OL (LOL) (Fig. 6, reaction 24). Next, the O-acyltransferase OlsA transfers a fatty acyl chain from AcpP to the LOL leading to the synthesis of OL (Fig. 6, reaction 25). This unmodified OL is sometimes also called S1 (substrate 1), a name initially describing its role as substrate in subsequent ´ S1-modifying reactions (Rojas-Jimenez et al. 2005). Recently, the bifunctional acyltransferase OlsF was described in Serratia proteamaculans that is responsible for OL formation in this organism (Fig. 6, reaction 26). OlsF has two acyltransferase domains, and the N-terminal domain was shown to be responsible for the O-acyltransferase reaction whereas the C-terminal domain was shown to be responsible for the N-acyltransferase reaction re´ et al. 2014). OlsBA are quired for OL synthesis (Vences-Guzman present in α-, β- and a few γ -proteobacteria and actinomycetes, whereas OlsF is present in γ -, δ- and -proteobacteria, and bac´ et al. 2014). Based on the teria of the CFB group (Vences-Guzman distribution of OlsB and OlsF in bacterial genomes, it has been estimated that about 50% of the sequenced bacteria can form ´ et al. 2014). Unmodified OL can be modified OL (Vences-Guzman by the introduction of one or more hydroxyl group or by methylation of the δ-amino group in the ornithine headgroup (Rojas´ ´ ´ Jimenez et al. 2005; Gonzalez-Silva et al. 2011; Vences-Guzman ´ Geiger and Sohlenkamp 2012; Moore et al. 2011; Vences-Guzman, et al. 2013). Three different OL hydroxylases have been described

so far: OlsC was described first in R. tropici and introduces a hydroxyl group in the 2-position of the ester-bound fatty acid ´ ´ (Fig. 6, reaction 27) (Rojas-Jimenez et al. 2005; Vences-Guzman et al. 2011). OlsD was described first in Burkholderia cenocepacia and introduces a hydroxyl group at the 2-position in the amidebound fatty acid and gives rise to the formation of new lipids 1 ´ and 2 (NL1, NL2) (Fig. 6, reaction 28) (Gonzalez-Silva et al. 2011; Diercks et al. 2015). OlsE introduces a hydroxyl group in the ornithine headgroup at a still unknown position (Fig. 6, reaction ´ et al. 2011). The methyltransferase respon29) (Vences-Guzman sible for the methylation of the δ-amino group of the ornithine headgroup in the planctomycete S. acidiphila has not been identified (Fig. 6, reactions 30–32) (Moore et al. 2013). OL is probably N-methylated by the unknown methyltransferase leading to the formation of TMOL (Fig. 6, reaction 30), but it is also possible that free ornithine is N-methylated before TMOL is formed (Fig. 6, reactions 31 and 32). Mutants have been identified in most genes described to be involved in OL synthesis or modification. Agrobacterium tumefaciens mutants deficient in OlsB or OlsE show accelerated tumor formation in a potato-disc assay ´ et al. 2013). Rhizobium tropici mutants deficient (Vences-Guzman in OlsC are defective in growth at high temperature and at acid ´ et al. 2011). Mutants deficient in OlsC and/or pH (Vences-Guzman OlsE in R. tropici are also defective in nodulation assays using ´ et al. 2011). Mutants deficient bean as a host (Vences-Guzman in OL formation in S. meliloti showed a slight growth defect under phosphate-limiting conditions when at the same time DGTS ´ synthesis was interrupted (Lopez-Lara et al. 2005). On the other hand, B. abortus mutants deficient in the formation of OL were not affected in virulence (Palacios-Chaves et al. 2011). Diercks et al. (2015) could show recently that OL in M. loti contains 80– 90% of D-ornithine which has not been observed in other organisms. It is not clear whether D-ornithine is also present in OLs of other bacteria.

Sulfonolipid synthesis Sulfonolipids are major lipids in the outer membrane of the gliding bacterium Flavobacterium johnsoniae (formerly Cytophaga johnsonae). They contain capnine which is formed by condensation of cysteic acid with fatty acyl-CoA under the release of CO2 (White 1984; Abbanat et al. 1986). An N-acyltransferase then transfers a fatty acid to capnine leading to the formation of N-acyl capnine. Mutants deficient in the synthesis of N-acyl capnine were also deficient in gliding, so there might be a connection between the presence of the lipid and the ability to move by gliding (Abbanat et al. 1986). The genes encoding the enzymes required for sulfonolipid synthesis have not been identified. The proposed structures of the molecular pathway for sulfonolipid synthesis can be found in Geiger et al. (2010).

Sphingolipid (SL) synthesis SLs are ubiquitous components of the eukaryotic cytoplasmic membrane where they have important functions in signal transduction processes and in the formation of lipid rafts. In contrast, SLs occur only in a few bacteria, mainly belonging to the CFB group and the Sphingomonadales of the α-proteobacteria, but they are also present in a few δ-proteobacteria such as Bdellovibrio, Cystobacter fuscus, My. stipitatus, S. cellulosum and M. xanthus (Keck et al. 2011; Lorenzen et al. 2014). Sequence analysis based on the presence of genes encoding for putative serine palmitoyltransferases (SPTs) indicates that SLs might be present in a few α-proteobacteria outside the Sphingomonadales and even


15

Figure 6. Synthesis of OLs and their hydroxylated or methylated derivatives in bacteria. AcpP-constitutive acyl carrier protein; α-KG-alpha-ketoglutarate. The names of the lipids S1 (substrate 1) and S2 (substrate 2) were initially describing their roles as substrates in OlsC-dependent OL modifying reactions, whereas the names P1 ´ (product 1) and P2 (product 2) were describing their roles as products of OlsC-dependent OL modifying reactions (Rojas-Jimenez et al. 2005). The formation of new lipids 1 and 2 (NL1, NL2) is due to the enzymatic activity of OlsD.

in a few β- and γ -proteobacteria. A variety of SL headgroups has been described in bacteria, such as ceramides, ceramide phosphoethanolamines, glucosyl sphingenine and ceramide phosphoinositols (Eckau, Dill and Budzikiewicz 1984; Stein and Budzikiewicz 1988; Keck et al. 2011; Lorenzen et al. 2014). In the

Sphingomonadales, SLs are mainly decorated with sugar residues. In Bacteroides species, ceramide phosphorylethanolamine and ceramide phosphorylglycerol have been described. The eukaryotic genes involved in SL biosynthesis have identified but little is known about SL synthesis in bacteria (Dickson 2010; Geiger

16


et al. 2010; Pata et al. 2010). Only the first step, catalyzed by SPT, is characterized in detail (Lowther et al. 2011, 2012). Even the crystal structures of the SPTs from Sphingomonas paucimobilis, S. wittichii and Sphingobacterium multivorum have been determined (Yard et al. 2007; Ikushiro et al. 2009; Raman et al. 2010). In eukaryotes, acyl-CoA and serine are the substrates for this enzyme, whereas for the enzyme from S. wittichii acyl-ACP has been suggested as acyl donor, because the gene-encoding SPT forms an operon with a gene encoding a putative ACP (Geiger et al. 2010; Raman et al. 2010). SLs are often 2-hydroxylated and the α-hydroxylase catalyzing this reaction has been identified and characterized from different bacteria (Matsunaga et al. 1997; Ring et al. 2009; Fujishiro et al. 2011). Not much is known about the functions of bacterial SLs. It has been described that some Gram-negative bacteria, such as Sp. capsulata, Sp. adhaerens and S. cellulosum, lack LPS in their outer membrane and instead have glycosphingolipids as functional replacements (Kawahara et al. 2000, 2001; Keck et al. 2011).

Hopanoid (HOP) synthesis HOPs are widespread in bacteria and can be present in the inner and outer membrane in proteobacteria. They are often considered bacterial sterol analogs and were first isolated from resins of tropical trees of the genus Hopea (Ourisson, Rohmer and Poralla 1987). HOPs are pentacyclic triterpenic isoprenoids. Synthesis of the precursors isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMPP) occurs depending on the bacterial species via the mevalonate pathway or via the ´ methylerythrol-4-phosphate pathway (Perez-Gil and Rodr´ıguez´ 2013). DMPP is elongated by head-to-tail condenConcepcion sation with two molecules IPP to farnesyl diphosphate. Two molecules farnesyl diphosphate are then condensated head-tohead leading to the formation of squalene. Squalene hopene cyclase (Shc) is responsible for the cyclization of squalene to diplotene or diplopterol (Fig. 7, reaction 33). The synthesis of tetrahymanol, another pentacyclic structure related to HOPs, was also shown to be Shc-dependent in Rhodopseudomonas palustris (Welander et al. 2009). Shc is an ortholog of squalene-2,3epoxide cyclase responsible for the formation of lanosterol from oxidosqualene. A Shc homolog from firmicutes has been shown to be responsible for the synthesis of sporulenes (Bosak, Losick and Pearson 2008; Kontnik et al. 2008), heptaprenyl lipids that increase the resistance of spores to reactive oxygen species. Starting from diploptene elongated HOPs can be synthesized. Only in recent years, many of the genes/enzymes required for HOP formation have been identified using R. palustris, Methylobacterium extorquens and B. cenocepacia as model organisms, and there are still many open questions with respect to the formation of complex HOPs. Often, genes involved in HOP synthesis and modification form operons and cluster within the bacterial genomes. The radical SAM superfamily enzyme HpnH is responsible for the formation of adenosylhopane (Bradley et al. 2010) (Fig. 7, reaction 34). HpnG is responsible for the release of adenine from adenosylhopane (Fig. 7, reaction 35). HpnG shows homology to purine nucleoside phosphorylases, but there is still a debate if the reaction product is phosphoribohopane or ribosylhopane. It has been suggested that the cyclic ribose opens up without the aid of an enzyme (Bradley et al. 2010) (Fig. 7, reaction 36). Formylhopane is probably the substrate for the aminotransferase reaction catalyzed by HpnO (Fig. 7, reaction 37) or alternatively it can be reduced to bacteriohopanetetrol (BHT) (Fig. 7, reaction 38). The conversion of ribosylhopane to BHT has been also suggested to happen non-enzymatically, but it

is unclear how the required reduction should be achieved this way (Schmerk et al. 2015) (Fig. 7, reaction 39). For the formation of BHT glucosamine, the enzymes HpnI and HpnK are required (Bradley et al. 2010; Schmerk et al. 2015) (Fig. 7, reaction 40 and 41). HpnI is a glycosyltransferase and HpnK has been suggested to be a deacetylase. HpnJ is a radical SAM superfamily member required for the formation of BHT cyclitolether, but the mechanism of the ring contraction reaction is not known (Fig. 7, reaction 42). HOPs can also be C6- and C11unsaturated, but the enzymes responsible for these unsaturation reactions have not been identified (Fig. 7, reactions 43 and 44). C2 and C3 methylation reactions of HOPs have been described and the methyltransferases have been identified (Fig. 7, reactions 45 and 46) (Welander et al. 2010; Welander and Summons 2012). The presence of 2-methylhopanoids in sedimentary fossils has been used for many years as a biomarker proxy for the presence of cyanobacteria and oxygenic photosynthesis, but it is clear now that in addition to cyanobacteria several αproteobacteria and acidobacteria can form 2-methylated HOPs. In contrast, 3-methylhopanoids have been used as a biomarker proxy for aerobic methanotrophs in the past, but it is clear now that several proteobacteria and actinomycetes also have a gene encoding homologs of the methyltransferase HpnR required for 3-methylhopanoid formation. In addition to the HOP structures presented here, other HOP structures have been described. In Bradyrhizobium, HOPs covalently linked to lipid A have been identified (Komaniecka et al. 2014; Silipo et al. 2014). Mutants in different steps of HOP formation have been characterized. Rhodopseudomonas palustris mutants deficient in HOP formation presented a weakened outer membrane and were more sensitive to pH and temperature stress (Welander et al. 2009; Doughty et al. 2011). Burkholderia cenocepacia defective in HOP formation displayed increased sensitivity to low pH, detergents, various antibiotics and did not produce flagella (Schmerk, Bernards and Valvano 2011). In contrast, in S. scabies HOPs were not required for tolerance to ethanol, oxidative and osmotic stress, high temperature or low pH (Seipke and Loria 2009).

METABOLIC PATHWAYS CONNECTING PHOSPHOLIPIDS AND PHOSPHORUS-FREE MEMBRANE LIPIDS Under phosphorus-limiting conditions, some bacteria can replace phospholipids such as PE, PG or PC from their membranes with phosphorus-free membrane lipids such as DGTS, SQD, GLs or OLs. In S. meliloti, this membrane remodeling is relatively well studied and some important players have been identified. Pulse-chase experiments had suggested that PE and PC are precursors for the synthesis of DGTS. The phospholipase C SMc00171 from S. meliloti was shown to be responsible for the conversion of PC to DAG and phosphocholine (ZavaletaPastor et al. 2010). DAG can then be used in S. meliloti for the synthesis of the phosphorus-free lipid DGTS and probably also of the DAG-containing lipid SQD. The expression of SMc00171 and of several of the enzymes required for SQD and DGTS synthesis is regulated by the transcriptional regulator PhoB and is induced under phosphate-limiting conditions. Interestingly, the PE methyltransferase from S. meliloti is also induced about 4-fold and the mechanism by which PE is degraded might be via PC. Genes encoding SMc00171 homologs are present in a few α-proteobacteria (A. tumefaciens, M. loti, B. japonicum, Caulobacter crescentus and R. palustris) and in a few Pseudomonas species. SMc00171-like phospholipases are


Figure 7. Synthesis of HOPs in bacteria. Shc-squalene hopene cyclase; α-KG-alpha-ketoglutarate; Glu-glutamate; UDP-GlcNAc-UDP-N-acetylglucosamine.

17

18


homologs of LpxH, an enzyme required for lipid A synthesis that hydrolyses UDP-2,3-diacylglucosamine to UMP and lipid X (2,3-bis-(3R-hydroxy-tetradecanoyl)-αD-glucosamine-1phosphate) (Babinski, Kanjilal and Raetz 2002a; Babinski, Ribeiro and Raetz 2002b). Genes encoding for SMc00171-like enzymes are absent in intracellular pathogens such as Brucella, Bartonella and Rickettsia. This absence of a phosphate starvation response is probably the consequence of an intracellular lifestyle which is characterized by a stable phosphate supply.

ENVIRONMENTAL CHALLENGES CAUSING CHANGES IN MEMBRANE LIPID COMPOSITION Bacteria change their membrane lipid composition in response to changes in the environment. This adaptation allows bacteria to survive unfavorable conditions. Even during the simple experiment of growing microorganisms in a flask, the environment the bacteria are sensing is changing constantly, which will be affecting the membrane lipid composition: nutrient levels fall, products of metabolism accumulate, the pH of the culture medium may rise or fall and oxygen levels are changing. The probably best-known example of membrane adaptation happens when bacteria are growing at different temperatures. In E. coli and other bacteria, a decrease in temperature causes a decrease in membrane fluidity and bacterial cells respond by increasing the amount of unsaturated fatty acids or fatty acids with analogous properties. An increase in temperature causes an increase in fluidity and probably an increase in the occurrence of discontinuities in the membrane. Bacteria growing at increased temperature usually contain more saturated fatty acids in their membranes (Marr and Ingraham 1962). However, membrane modifications as a response to abiotic stress conditions are not restricted to the fatty acid moieties and do also happen on headgroup level. The membrane lipid modifications occurring can be divided in two types. (1) Existing lipids can be modified to obtain a membrane with different properties; for example, the anionic lipid PG and be modified to the cationic lipid LPG simply by transfer of one lysyl group. This modification of pre-existing membrane lipids has the advantage that it allows a quick response to changes in environmental conditions. (2) Existing lipids are degraded and new lipids with new characteristics are synthesized de novo replacing the old lipids. In the following, we will discuss how membrane lipid modifications occur under different stress conditions. Growth under acidic conditions provokes a modification in membrane lipids. In R. tropici, two membrane-modifying activities have been described that occur under condition of acid stress and that allow the bacteria to prosper under these conditions. The operon lpiA/atvA is transcriptionally induced after acid shock (Vinuesa et al. 2003). LpiA is an LPG synthase homologous to MprF responsible for formation of the cationic lipid LPG (Sohlenkamp et al. 2007). The second gene of the operon encodes the putative lipase AtvA. This operon structure is also present in P. aeruginosa. Pseudomonas aeruginosa MprF is responsible for APG formation and the AtvA homolog PA0919 has been shown to cause the hydrolysis of APG and formation of PG and alanine. Thus, it can be hypothesized that AtvA from R. tropici will hydrolyze LPG to lysine and PG, but at the moment it is not clear how this should lead to an increased acid resistance of the bacteria. This modification of the anionic membrane lipid PG to the cationic (LPG) or zwitterionic (APG) membrane lipids has been shown to be important in resistence to cationic peptides and antibiotics. Another lipid modification described in R. tropici that is

known to have importance under acidic growth conditions is the ´ 2-hydroxylation of OL by OlsC described above (Rojas-Jimenez ´ et al. 2011) (Fig. 6, reaction 27). An et al. 2005; Vences-Guzman α-hydroxyl group is introduced into the ester-bound fatty acid of OL, analogous to the 2-hydroxylation of lipid A-bound fatty acids in Salmonella typhimurium and the 2-hydroxylation of SLs (Matsunaga et al. 1997; Gibbons et al. 2000, 2008; Ring et al. 2009; Fujishiro et al. 2011). It is thought that the presence of the additional hydroxyl group allows the formation of hydrogen bonds that increase the lateral interactions between lipid molecules and stabilize the leaflet structure (Nikaido 2003). Consistent with this hypothesis, R. tropici mutants deficient in OlsC and lacking the 2-hydroxylated OL termed P1 show a severe growth defect at ´ et al. 2011). a pH lower than 4.5 (Vences-Guzman The 2-hydroxylation of OL also seems to play a role in the response to increased growth temperature in R. tropici and B. cepacia. In R. tropici, a mutant deficient in the 2-hydroxylase OlsC grows like the wildtype at 30◦ C, but grows significantly slower than the wildtype at 37 and 42◦ C, whereas B. cepacia accumulates increased amounts of hydroxylated OL and hydroxylated PE at increased growth temperatures. Again, increased lateral interactions between lipid molecules due to the extra hydroxyl groups might be responsible for this phenotype (Taylor, Ander´ et al. 2011). son and Wilkinson 1998; Vences-Guzman During the response to high osmotic pressures, the anionic membrane lipid CL has been shown to play an important role in different bacteria. Increased osmotic pressure causes an increase in CL formation in E. coli and R. sphaeroides (Catucci et al. 2004; Tsatskis et al. 2005). In E. coli, CL is located to the cellular poles where the proline transporter ProP is embedded in CLenriched regions of the cytoplasmic membrane. ProP is denoted an osmosensory transporter because it is activated by increasing osmolarity and causes the accumulation of the compatible solute proline allowing bacterial growth at increased osmotic pressure. The activation threshold of ProP correlates with the amount of CL present in the membrane and its presence controls the amount and activity of the proline transporter ProP (Romantsov et al. 2007, 2008; Romantsov, Guan and Wood 2009). Many soil bacteria remodel their membranes in response to conditions of phosphate limitation. Plants and microbes mostly live in environments where available phosphate is a growthlimiting factor. Apparently, some bacteria can replace their phospholipids with membrane lipids devoid of phosphorus. This remodeling has been studied in some detail in R. sphaeroides, S. meliloti, A. tumefaciens and M. loti (Benning et al. 1993; Benning, Huang and Gage 1995; Geiger et al. 1999; Zavaleta-Pastor et al. 2010; Devers et al. 2011; Geske et al. 2013; Diercks et al. 2015). The phospholipids PE and PC are actively degraded and are replaced by the DAG-derived phosphorus-free membrane lipids DGTS, SQD and GLs and by the aminolipid OL. This remodeling is not a minor process because PC makes up for about 60% of membrane lipids when bacteria are cultivated on complex medium, whereas in minimal medium under limiting phosphate conditions DGTS increases to about 60% (Geiger et al. 1999). It was suggested that phospholipids are functionally replaced by phosphorus-free membrane lipids under conditions where phosphate becomes limiting thereby making the phosphate pool present in the membrane accessible for other cellular processes such as nucleic acid synthesis. It is thought that PC is replaced by DGTS, both being zwitterionic lipids with bulky headgroups, that PG is replaced by SQD, both being anionic lipids and that PE is replaced by OLs, both being zwitterionic lipids with smaller headgroups. There is evidence supporting these hypotheses: Recently, Riekhof and collaborators


constructed a S. cerevisiae strain that had replaced its PC with DGTS. They showed that essential processes of membrane biogenesis and organelle assembly were functional in this strain (Riekhof et al. 2014). On an experimental level, this is probably the best hint that DGTS can functionally replace PC in the membrane. In cyanobacteria, which form PG, SQD and neutral GLs under phosphate-replete conditions, PG is decreased under phosphate limitation whereas SQD is increased. In A. tumefaciens, different OLs are formed constitutively meaning that their synthesis does not depend on phosphate starvation but also occurs in phosphate-replete complex medium. When an A. tumefaciens mutant deficient in OlsB is constructed, PE levels increase ´ et al. 2013). drastically (Vences-Guzman

OPEN QUESTIONS Bacteria account for most of the diversity of life on our planet and they have been around for billions of years. During this time they have adapted to all types of environments on Earth. From this point of view, it is probably not surprising that bacteria present such a rich diversity of lipid structures and pathways involved in the formation of membrane lipids when compared to eukaryotes. When studying the diversity of bacterial membrane lipids, the ultimate goal probably should be to have a complete catalog of the lipids that are formed when bacteria are growing in their natural habitats and understand their functions within the physiology of the organisms. Although we have come a long way, there is still much work to do. What are the problems and what has to be done to fill these information gaps? (I) Only a few bacteria can be easily grown in the laboratory in pure cultures. Most bacteria of the environment still escape our attempts to grow them in the laboratory. This means that almost everything we know about bacterial membrane lipids comes from a relatively small set of easily culturable bacteria. We can probably expect to find several unknown lipid structures and unknown synthesis pathways in these unculturable bacteria. One possibility to untap part of this variety of bacterial membrane lipids is to combine culturomics approaches with lipidomics (Lagier et al. 2012). Culturomics is a systematic approach to find growth conditions of so-far unculturable bacteria. Once this or these conditions are defined, the organism’s membrane lipid composition can be studied. Examples of recently discovered lipid structures include a trimethylated OL in Planctomycetes, diacylserinophospholipids from Verrucomicrobia and diol-based lipids in thermophilic bacteria (Moore et al. 2013; Lagutin et al. 2014; Anders et al. 2015). (II) Currently, we have a basic knowledge about the lipid composition of major model bacteria and many of the genes and enzymes involved in their synthesis. However, even the well-studied E. coli still has hidden secrets and we are probably more ignorant than we want to realize. For decades lipids biochemists had the idea that not much new was to be discovered in E. coli, but recently, Tan et al. (2012) identified the Cls ClsC that synthesizes CL from the precursors PG and PE, something that was completely unheard of. It belongs to the PLD superfamily, which together with the CDP-alcohol phosphotransferase superfamily (Pips, Pcs, PgsA, type-II PssA or the eukaryotic type Cls) represents a large fraction of the known enzymes involved in bacterial phospholipid synthesis and turnover. Even in well-studied organisms such as E. coli, S. meliloti or St. coelicolor, there are proteins belonging to one of these two superfamilies

19

with unassigned functions. Almost all CDP-alcohol phosphotransferases characterized play a role in phospholipid synthesis, the only known exceptions being a few enzymes involved in the synthesis of inositol-based compatible solute in hyperthermophilic bacteria and archaea (Brito et al. 2011; Nogly et al. 2014). Although there might be redundancy with respect to some activities, bacteria might form other lipids that have not yet been described, and/or new enzymatic activities are waiting to be discovered. Examples are the above-mentioned ClsC or the CL/PE synthase from X. campestris (Tan et al. 2012; Moser et al. 2014a). A systematic mining the wealth of genome sequence data should not be restricted to both protein superfamilies, but include also the search for other genes/enzymes such as acyltransferases involved in membrane lipid synthesis, turnover and degradation. It can be expected that many new structures and activities will be identified. (III) Bacteria are cultured in the laboratory under conditions that in most cases do not reflect the natural habitat. The membrane lipid composition of bacteria is not a constant but is responding to the environmental conditions. If we want to understand what role the membrane (lipids) play for the functioning of a bacterium but also during the interaction with other bacteria or eukaryotes, it is essential to study the bacteria within their habitat. Experimental approaches should be developed and improved that allow the study of bacterial membrane lipids in the natural environment or try to take the habitat to the laboratory. That means go to the sites where the bacteria are growing, sample and if possible study in situ or bring part of the habitat to the laboratory and study it there. One possibility is to combine the study of environmental lipid samples as for example geobiochemists do when analyzing samples from marine sediments, with metagenomic studies. The metagenomic study will inform us what bacteria are present in the habitat and the lipid analysis will show what membrane lipids are present. In this model metadata such as the environmental conditions can be included, so that a modeling should be possible. Finally, the perfect experimental approach would be to reconstitute the natural environment in the lab and use site-directed mutants in this experimental setup to follow how changes or conditions affect membrane composition and interactions between organisms.

FUNDING Research in our lab was supported by grants from Consejo ´ ´ Nacional de Ciencia y Tecnolog´ıa-Mexico (CONACyT-Mexico; ´ General de Asuntos del Per153200, 178359) and Direccion ´ ´ ´ sonal Academico-Universidad Nacional Autonoma de Mexico (DGAPA-UNAM; PAPIIT IN202413, IN203612). Conflict of interest. None declared.

REFERENCES Abbanat DR, Leadbetter ER, Godchaux W, et al. Sulfonolipids are molecular determinants of gliding motility. Nature 1986;324:367–9. ¨ Aktas M, Danne L, Moller P, et al. Membrane lipids in Agrobacterium tumefaciens: biosynthetic pathways and importance for pathogenesis. Front Plant Sci 2014;5:109.

20


Ames GF. Lipids of Salmonella typhimurium and Escherichia coli: structure and metabolism. J Bacteriol 1968;95:833–43. Anders H, Power JF, MacKenzie AD, et al. Limisphaera ngatamarikiensis gen. nov. sp. nov., a thermophilic, pink-pigmented coccus isolated from subaqueous mud of a geothermal hotspring in New Zealand. Int J Syst Evol Micr 2015, doi:10.1099/ijs.0.00006310.1099/ijs.0.000063. Anderson R, Hansen K. Structure of a novel phosphoglycolipid from Deinococcus radiodurans. J Biol Chem 1985;260:12219–23. Antonsson B. Phosphatidylinositol synthase from mammalian tissues. Biochim Biophys Acta 1997;1348:179–86. Arendt W, Groenewold MK, Hebecker S, et al. Identification and characterization of a periplasmic aminoacylphosphatidylglycerol hydrolase responsible for Pseudomonas aeruginosa lipid homeostasis. J Biol Chem 2013;288:24717–30. ¨ Arendt W, Hebecker S, Jager S, et al. Resistance phenotypes mediated by aminoacyl-phosphatidylglycerol synthases. J Bacteriol 2012;194:1401–16. Arondel V, Benning C, Somerville CR. Isolation and functional expression in Escherichia coli of a gene encoding phosphatidylethanolamine methyltransferase (EC 2.1.1.17) from Rhodobacter sphaeroides. J Biol Chem 1993;268:16002–8. Babinski KJ, Kanjilal SJ, Raetz CR. Accumulation of the lipid A precursor UDP-2,3-diacylglucosamine in an Escherichia coli mutant lacking the lpxH gene. J Biol Chem 2002a;277:25947– 56. Babinski KJ, Ribeiro AA, Raetz CR. The Escherichia coli gene encoding the UDP-2,3-diacylglucosamine pyrophosphatase of lipid A biosynthesis. J Biol Chem 2002b;277:25937–46. Barridge JK, Shively JM. Phospholipids of the thiobacilli. J Bacteriol 1968;95:2182–5. Batrakov SG, Nikitin DI, Mosezhnyi AE, et al. A glycine-containing phosphorus-free lipoaminoacid from the gram-negative marine bacterium Cyclobacterium marinus WH. Chem Phys Lipids 1999;99:139–43. Benning C. Biosynthesis and function of the sulfolipid sulfoquinovosyl diacylglycerol. Annu Rev Plant Phys 1998;49: 53–75. Benning C, Beatty JT, Prince RC, et al. The sulfolipid sulfoquinovosyldiacylglycerol is not required for photosynthetic electron transport in Rhodobacter sphaeroides but enhances growth under phosphate limitation. P Natl Acad Sci USA 1993;90:1561–5. Benning C, Huang ZH, Gage DA. Accumulation of a novel glycolipid and a betaine lipid in cells of Rhodobacter sphaeroides grown under phosphate limitation. Arch Biochem Biophys 1995;317:103–11. Benning C, Somerville CR. Identification of an operon involved in sulfolipid biosynthesis in Rhodobacter sphaeroides. J Bacteriol 1992a;174:6479–87. Benning C, Somerville CR. Isolation and genetic complementation of a sulfolipid-deficient mutant of Rhodobacter sphaeroides. J Bacteriol 1992b;174:2352–60. Berg S, Kaur D, Jackson M, et al. The glycosyltransferases of Mycobacterium tuberculosis—roles in the synthesis of arabinogalactan, lipoarabinomannan, and other glycoconjugates. Glycobiology 2007;17:35–56R. Berg S, Wieslander A. Purification of a phosphatase which hydrolyzes phosphatidic acid, a key intermediate in glucolipid synthesis in Acholeplasma laidlawii A membranes. Biochim Biophys Acta 1997;1330:225–32. Blumenberg M, Oppermann BI, Guyoneaud R, et al. Hopanoid production by Desulfovibrio bastinii isolated from oilfield formation water. FEMS Microbiol Lett 2009;293:73–8.

Bode HB, Zeggel B, Silakowski B, et al. Steroid biosynthesis in prokaryotes: identification of myxobacterial steroids and cloning of the first bacterial 2,3(S)-oxidosqualene cyclase from the myxobacterium Stigmatella aurantiaca. Mol Microbiol 2003;47:471–81. Bondoso J, Albuquerque L, Lobo-da-Cunha A, et al. Rhodopirellula lusitana sp. nov. and Rhodopirellula rubra sp. nov., isolated from the surface of macroalgae. Syst Appl Microbiol 2014;37:157–64. Bosak T, Losick RM, Pearson A. A polycyclic terpenoid that alleviates oxidative stress. P Natl Acad Sci USA 2008;105:6725–9. Boumann HA, Hopmans EC, vande Leemput I, et al. Ladderane phospholipids in anammox bacteria comprise phosphocholine and phosphoethanolamine headgroups. FEMS Microbiol Lett 2006;258:297–304. Bradley AS, Pearson A, Saenz JP, et al. Adenosylhopane: the first intermediate in hopanoid side chain biosynthesis. Org Geochem 2010;41:1075–81. Bravo JM, Perzl M, Hartner T, et al. Novel methylated triterpenoids of the gammacerane series from the nitrogen-fixing bacterium Bradyrhizobium japonicum USDA 110. Eur J Biochem 2001;268:1323–31. Brito JA, Borges N, Vonrhein C, et al. Crystal structure of Archaeoglobus fulgidus CTP:inositol-1-phosphate cytidylyltransferase, a key enzyme for di-myo-inositol-phosphate synthesis in (hyper)thermophiles. J Bacteriol 2011;193:2177– 85. Bukata L, Altabe S, de Mendoza D, et al. Phosphatidylethanolamine synthesis is required for optimal virulence of Brucella abortus. J Bacteriol 2008;190:8197–203. Caillon E, Lubochinsky B, Rigomier D. Occurrence of dialkyl ether phospholipids in Stigmatella aurantiaca DW4. J Bacteriol 1983;153:1348–51. Campbell HA, Kent C. The CTP:phosphocholine cytidylyltransferase encoded by the licC gene of Streptococcus pneumoniae: cloning, expression, purification, and characterization. Biochim Biophys Acta 2001;1534:85–95. Carman GM. Phosphatidate phosphatases and diacylglycerol pyrophosphate phosphatases in Saccharomyces cerevisiae and Escherichia coli. Biochim Biophys Acta 1997;1348:45–55. Catucci L, Depalo N, Lattanzio VM, et al. Neosynthesis of cardiolipin in Rhodobacter sphaeroides under osmotic stress. Biochemistry 2004;43:15066–72. Coleman J. Characterization of Escherichia coli cells deficient in 1acyl-sn-glycerol-3- phosphate acyltransferase activity. J Biol Chem 1990;265:17215–21. Coleman J. Characterization of the Escherichia coli gene for 1-acylsn-glycerol-3-phosphate acyltransferase (plsC). Mol Gen Genet 1992;232:295–303. Comba S, Menendez-Bravo S, Arabolaza A, et al. Identification and physiological characterization of phosphatidic acid phosphatase enzymes involved in triacylglycerol biosynthesis in Streptomyces coelicolor. Microb Cell Fact 2013;12:9. Comerci DJ, Altabe S, de Mendoza D, et al. Brucella abortus synthesizes phosphatidylcholine from choline provided by the host. J Bacteriol 2006;188:1929–34. Conde-Alvarez R, Grillo´ MJ, Salcedo SP, et al. Synthesis of phosphatidylcholine, a typical eukaryotic phospholipid, is necessary for full virulence of the intracellular bacterial parasite Brucella abortus. Cell Microbiol 2006;8:1322–35. Conover GM, Mart´ınez-Morales F, Heidtman MI, et al. Phosphatidylcholine synthesis is required for optimal function of Legionella pneumophila virulence determinants. Cell Microbiol 2008;10:514–28.


Contreras I, Shapiro L, Henry S. Membrane phospholipid composition of Caulobacter crescentus. J Bacteriol 1978;135:1130–6. Cooper CL, Jackowski S, Rock CO. Fatty acid metabolism in snglycerol-3-phosphate acyltransferase (plsB) mutants. J Bacteriol 1987;169:605–11. Damste´ JS, Rijpstra WI, Hopmans EC, et al. Structural characterization of diabolic acid-based tetraester, tetraether and mixed ether/ester, membrane-spanning lipids of bacteria from the order Thermotogales. Arch Microbiol 2007;188:629–41. Dare K, Shepherd J, Roy H, et al. LysPGS formation in Listeria monocytogenes has broad roles in maintaining membrane integrity beyond antimicrobial peptide resistance. Virulence 2014;5:534–46. ´ de Rudder KE, Lopez-Lara IM, Geiger O. Inactivation of the gene for phospholipid N-methyltransferase in Sinorhizobium meliloti: phosphatidylcholine is required for normal growth. Mol Microbiol 2000;37:763–72. de Rudder KE, Sohlenkamp C, Geiger O. Plant-exuded choline is used for rhizobial membrane lipid biosynthesis by phosphatidylcholine synthase. J Biol Chem 1999;274:20011–6. De Siervo AJ, Homola AD. Analysis of Caulobacter crescentus lipids. J Bacteriol 1980;143:1215–22. DeChavigny A, Heacock PN, Dowhan W. Sequence and inactivation of the pss gene of Escherichia coli. Phosphatidylethanolamine may not be essential for cell viability. J Biol Chem 1991;266:5323–32. Dees C, Shively JM. Localization of quantitation of the ornithine lipid of Thiobacillus thiooxidans. J Bacteriol 1982;149:798–9. den Kamp JA, Redai I, van Deenen LL. Phospholipid composition of Bacillus subtilis. J Bacteriol 1969;99:298–303. Devers EA, Wewer V, Dombrink I, et al. A processive glycosyltransferase involved in glycolipid synthesis during phosphate deprivation in Mesorhizobium loti. J Bacteriol 2011;193:1377–84. Dickson RC. Roles for sphingolipids in Saccharomyces cerevisiae. Adv Exp Med Biol 2010;688:217–31. Diercks H, Semeniuk A, Gisch N, et al. Accumulation of novel glycolipids and ornithine Lipids in Mesorhizobium loti under phosphate deprivation. J Bacteriol 2015;197:497–509. Doughty DM, Coleman ML, Hunter RC, et al. The RND-family transporter, HpnN, is required for hopanoid localization to the outer membrane of Rhodopseudomonas palustris TIE-1. P Natl Acad Sci USA 2011;108:E1045–51. Doughty DM, Hunter RC, Summons RE, et al. 2-Methylhopanoids are maximally produced in akinetes of Nostoc punctiforme: geobiological implications. Geobiology 2009;7:524–32. Dowhan W. A retrospective: use of Escherichia coli as a vehicle to study phospholipid synthesis and function. Biochim Biophys Acta 2013;1831:471–94. Eckau H, Dill D, Budzikiewicz H. Novel ceramides from Cystobacter fuscus (Myxobacterales). Z Naturforsch C 1984;39:1–9. Ernst CM, Staubitz P, Mishra NN, et al. The bacterial defensin resistance protein MprF consists of separable domains for lipid lysinylation and antimicrobial peptide repulsion. PLoS Pathog 2009;5:e1000660. Evans RI, McClure PJ, Gould GW, et al. The effect of growth temperature on the phospholipid and fatty acyl compositions of non-proteolytic Clostridium botulinum. Int J Food Microbiol 1998;40:159–67. Fang J, Barcelona MJ, Semrau JD. Characterization of methanotrophic bacteria on the basis of intact phospholipid profiles. FEMS Microbiol Lett 2000;189:67–72. Fiedler W, Rotering H. Characterization of an Escherichia coli mdoB mutant strain unable to transfer sn-1-phosphoglycerol

21

to membrane-derived oligosaccharides. J Biol Chem 1985;260:4799–806. Fischer W, Leopold K. Polar lipids of four Listeria species containing L-lysylcardiolipin, a novel lipid structure, and other unique phospholipids. Int J Syst Bacteriol 1999;49(Pt 2): 653–62. Fuerst JA, Sagulenko E. Beyond the bacterium: planctomycetes challenge our concepts of microbial structure and function. Nat Rev Microbiol 2011;9:403–13. Fujishiro T, Shoji O, Nagano S, et al. Crystal structure of H2 O2 dependent cytochrome P450SPα with its bound fatty acid substrate. J Biol Chem 2011;286:29941–50. Gao JL, Weissenmayer B, Taylor AM, et al. Identification of a gene required for the formation of lyso-ornithine lipid, an intermediate in the biosynthesis of ornithine-containing lipids. Mol Microbiol 2004;53:1757–70. Ge Z, Taylor DE. The Helicobacter pylori gene encoding phosphatidylserine synthase: sequence, expression, and insertional mutagenesis. J Bacteriol 1997;179:4970–6. ´ ´ Geiger O, Gonzalez-Silva N, Lopez-Lara IM, et al. Amino acidcontaining membrane lipids in bacteria. Prog Lipid Res 2010;49:46–60. ´ Geiger O, Lopez-Lara IM, Sohlenkamp C. Phosphatidylcholine biosynthesis and function in bacteria. Biochim Biophys Acta 2013;1831:503–13. ¨ Geiger O, Rohrs V, Weissenmayer B, et al. The regulator gene phoB mediates phosphate stress-controlled synthesis of the membrane lipid diacylglyceryl-N,N,N-trimethylhomoserine in Rhizobium (Sinorhizobium) meliloti. Mol Microbiol 1999;32: 63–73. ¨ Geske T, Vom Dorp K, Dormann P, et al. Accumulation of glycolipids and other non-phosphorous lipids in Agrobacterium tumefaciens grown under phosphate deprivation. Glycobiology 2013;23:69–80. Gibbons HS, Lin S, Cotter RJ, et al. Oxygen requirement for the biosynthesis of the S-2-hydroxymyristate moiety in Salmonella typhimurium lipid A. J Biol Chem 2000;275: 32940–9. Gibbons HS, Reynolds CM, Guan Z, et al. An inner membrane dioxygenase that generates the 2-hydroxymyristate moiety of Salmonella lipid A. Biochemistry 2008;47:2814–25. Gidden J, Denson J, Liyanage R, et al. Lipid compositions in Escherichia coli and Bacillus subtilis during growth as determined by MALDI-TOF and TOF/TOF mass spectrometry. Int J Mass Spectrom 2009;283:178–84. Giles DK, Hankins JV, Guan Z, et al. Remodelling of the Vibrio cholerae membrane by incorporation of exogenous fatty acids from host and aquatic environments. Mol Microbiol 2011;79:716–28. Goldfine H. The appearance, disappearance and reappearance of plasmalogens in evolution. Prog Lipid Res 2010;49:493–8. Goldfine H, Ellis ME. N-methyl groups in bacterial lipids. J Bacteriol 1964;87:8–15. ´ ´ Gonzalez-Silva N, Lopez-Lara IM, Reyes-Lamothe R, et al. The dioxygenase-encoding olsD gene from Burkholderia cenocepacia causes the hydroxylation of the amide-linked fatty acyl moiety of ornithine-containing membrane lipids. Biochemistry 2011;50:6396–408. Gould RM, Lennarz WJ. Biosynthesis of aminoacyl derivatives of phosphatidylglycerol. Biochem Bioph Res Co 1967;26:512–5. Guerin ME, Kaur D, Somashekar BS, et al. New insights into the early steps of phosphatidylinositol mannoside biosynthesis in mycobacteria: PimB’ is an essential enzyme of Mycobacterium smegmatis. J Biol Chem 2009;284:25687–96.

22


Guerin ME, Kordulakova J, Alzari PM, et al. Molecular basis of phosphatidyl-myo-inositol mannoside biosynthesis and regulation in mycobacteria. J Biol Chem 2010;285:33577–83. Guler S, Essigmann B, Benning C. A cyanobacterial gene, sqdX, ¨ required for biosynthesis of the sulfolipid sulfoquinovosyldiacylglycerol. J Bacteriol 2000;182:543–5. ¨ Guler S, Seeliger A, Hartel H, et al. A null mutant of Synechococcus ¨ sp. PCC7942 deficient in the sulfolipid sulfoquinovosyl diacylglycerol. J Biol Chem 1996;271:7501–7. Guo D, Tropp BE. A second Escherichia coli protein with CL synthase activity. Biochim Biophys Acta 2000;1483:263–74. Hachmann AB, Angert ER, Helmann JD. Genetic analysis of factors affecting susceptibility of Bacillus subtilis to daptomycin. Antimicrob Agents Ch 2009;53:1598–609. Hacker S, Sohlenkamp C, Aktas M, et al. Multiple phospholipid Nmethyltransferases with distinct substrate specificities are encoded in Bradyrhizobium japonicum. J Bacteriol 2008;190:571– 80. Hagen PO. Lipids of Sphaerophorus ridiculosis: plasmalogen composition. J Bacteriol 1974;119:643–5. Hagio M, Gombos Z, Varkonyi Z, et al. Direct evidence for requirement of phosphatidylglycerol in photosystem II of photosynthesis. Plant Physiol 2000;124:795–804. Hatch GM, McClarty G. Phospholipid composition of purified Chlamydia trachomatis mimics that of the eucaryotic host cell. Infect Immun 1998;66:3727–35. Hebecker S, Arendt W, Heinemann IU, et al. Alanylphosphatidylglycerol synthase: mechanism of substrate recognition during tRNA-dependent lipid modification in Pseudomonas aeruginosa. Mol Microbiol 2011;80:935–50. Hill DL, Ballou CE. Biosynthesis of mannophospholipids by Mycobacterium phlei. J Biol Chem 1966;241:895–902. ¨ ¨ Holzl G, Dormann P. Structure and function of glycoglycerolipids in plants and bacteria. Prog Lipid Res 2007;46:225–43. ¨ Holzl G, Leipelt M, Ott C, et al. Processive lipid galactosyl/glucosyltransferases from Agrobacterium tumefaciens and Mesorhizobium loti display multiple specificities. Glycobiology 2005;15:874–86. Huang HD, Wang W, Ma T, et al. Sphingomonas sanxanigenens sp. nov., isolated from soil. Int J Syst Evol Micr 2009;59:719–23. Huang Y, Anderson R. Structure of a novel glucosaminecontaining phosphoglycolipid from Deinococcus radiodurans. J Biol Chem 1989;264:18667–72. Huang Y, Anderson R. Glucosyl diglyceride lipid structures in Deinococcus radiodurans. J Bacteriol 1995;177:2567–71. Huber R, Langworthy TA, Konig H, et al. Thermotoga-maritima sp-nov represents a new genus of unique extremely thermophilic eubacteria growing up to 90-degrees-C. Arch Microbiol 1986;144:324–33. Ikushiro H, Islam MM, Okamoto A, et al. Structural insights into the enzymatic mechanism of serine palmitoyltransferase from Sphingobacterium multivorum. J Biochem 2009;146:549–62. Iwamori M, Sakai A, Minamimoto N, et al. Characterization of novel glycolipid antigens with an α-galactose epitope in lactobacilli detected with rabbit anti-Lactobacillus antisera and occurrence of antibodies against them in human sera. J Biochem 2011;150:515–23. Jackson BJ, Bohin JP, Kennedy EP. Biosynthesis of membranederived oligosaccharides: characterization of mdoB mutants defective in phosphoglycerol transferase I activity. J Bacteriol 1984;160:976–81. Jackson M, Crick DC, Brennan PJ. Phosphatidylinositol is an essential phospholipid of mycobacteria. J Biol Chem 2000;275:30092–9.

Jahnke LL, Eder W, Huber R, et al. Signature lipids and stable carbon isotope analyses of Octopus Spring hyperthermophilic communities compared with those of Aquificales representatives. Appl Environ Microb 2001;67:5179–89. Johnston NC, Baker JK, Goldfine H. Phospholipids of Clostridium perfringens: a reexamination. FEMS Microbiol Lett 2004;233: 65–8. Jones DE, Smith JD. Phospholipids of the differentiating bacterium Caulobacter crescentus. Can J Biochem 1979;57:424–8. Jones DS, Albrecht HL, Dawson KS, et al. Community genomic analysis of an extremely acidophilic sulfur-oxidizing biofilm. ISME J 2012;6:158–70. Jorasch P, Wolter FP, Zahringer U, et al. A UDP glucosyltransferase from Bacillus subtilis successively transfers up to four glucose residues to 1,2-diacylglycerol: expression of ypfP in Escherichia coli and structural analysis of its reaction products. Mol Microbiol 1998;29:419–30. Jorge CD, Borges N, Santos H. A novel pathway for the synthesis of inositol phospholipids uses CDP-inositol as donor of the polar head group. Environ Microbiol 2014, DOI:10.1111/14622920.12734. Jyothikumar V, Klanbut K, Tiong J, et al. Cardiolipin synthase is required for Streptomyces coelicolor morphogenesis. Mol Microbiol 2012;84:181–97. Kaneshiro T, Law JH. Phosphatidylcholine synthesis in Agrobacterium tumefaciens. I. Purification and properties of a phosphatidylethanolamine N-methyltransferase. J Biol Chem 1964;239:1705–13. Kanipes MI, Henry SA. The phospholipid methyltransferases in yeast. Biochim Biophys Acta 1997;1348:134–41. Kannenberg EL, Poralla K. Hopanoid biosynthesis and function in bacteria. Naturwissenschaften 1999;86:168–76. Karnezis T, Fisher HC, Neumann GM, et al. Cloning and characterization of the phosphatidylserine synthase gene of Agrobacterium sp. strain ATCC 31749 and effect of its inactivation on production of high-molecular-mass (1–>3)-betaD-glucan (curdlan). J Bacteriol 2002;184:4114–23. Kato M, Sakai M, Adachi K, et al. Distribution of betaine lipids in marine algae. Phytochemistry 1996;42:1341–45. Kaur D, Guerin ME, Skovierova H, et al. Chapter 2: biogenesis of the cell wall and other glycoconjugates of Mycobacterium tuberculosis. Adv Appl Microbiol 2009;69:23–78. ¨ Kawahara K, Kuraishi H, Zahringer U. Chemical structure and function of glycosphingolipids of Sphingomonas spp and their distribution among members of the α-4 subclass of Proteobacteria. J Ind Microbiol Biot 1999;23:408–13. Kawahara K, Lindner B, Isshiki Y, et al. Structural analysis of a new glycosphingolipid from the lipopolysaccharide-lacking bacterium Sphingomonas adhaesiva. Carbohyd Res 2001;333: 87–93. Kawahara K, Moll H, Knirel YA, et al. Structural analysis of two glycosphingolipids from the lipopolysaccharide-lacking bacterium Sphingomonas capsulata. Eur J Biochem 2000;267: 1837–46. Kawai Y, Moribayashi A. Characteristic lipids of Bordetella pertussis: simple fatty acid composition, hydroxy fatty acids, and an ornithine-containing lipid. J Bacteriol 1982;151:996–1005. Kawai Y, Moribayashi A, Yano I. Ornithine-containing lipid of Bordetella pertussis that carries hemagglutinating activity. J Bacteriol 1982;152:907–10. Kawai Y, Yano I, Kaneda K. Various kinds of lipoamino acids including a novel serine-containing lipid in an opportunistic pathogen Flavobacterium—their structures and biological activities on erythrocytes. Eur J Biochem 1988;171:73–80.


Kawazoe R, Okuyama H, Reichardt W, et al. Phospholipids and a novel glycine-containing lipoamino acid in Cytophaga johnsonae Stanier strain C21. J Bacteriol 1991;173:5470–5. Keck M, Gisch N, Moll H, et al. Unusual outer membrane lipid composition of the gram-negative, lipopolysaccharidelacking myxobacterium Sorangium cellulosum So ce56. J Biol Chem 2011;286:12850–9. Kent C, Gee P, Lee SY, et al. A CDP-choline pathway for phosphatidylcholine biosynthesis in Treponema denticola. Mol Microbiol 2004;51:471–81. Kenyon CN, Gray AM. Preliminary analysis of lipids and fatty acids of green bacteria and Chloroflexus aurantiacus. J Bacteriol 1974;120:131–8. Khuller GK, Brennan PJ. The polar lipids of some species of Nocardia. J Gen Microbiol 1972;73:409–12. Khuller GK, Taneja R, Kaur S, et al. Lipid composition and virulence of Mycobacterium tuberculosis H37Rv. Aust J Exp Biol Med 1982;60(Pt 5):541–7. Kikuchi S, Shibuya I, Matsumoto K. Viability of an Escherichia coli pgsA null mutant lacking detectable phosphatidylglycerol and cardiolipin. J Bacteriol 2000;182:371–6. Kim KS, Manasherob R, Cohen SN. YmdB: a stress-responsive ribonuclease-binding regulator of E. coli RNase III activity. Gene Dev 2008;22:3497–508. Klein S, Lorenzo C, Hoffmann S, et al. Adaptation of Pseudomonas aeruginosa to various conditions includes tRNAdependent formation of alanyl-phosphatidylglycerol. Mol Microbiol 2009;71:551–65. Klug RM, Benning C. Two enzymes of diacylglyceryl-O-4’-(N,N,N,trimethyl)homoserine biosynthesis are encoded by btaA and btaB in the purple bacterium Rhodobacter sphaeroides. P Natl Acad Sci USA 2001;98:5910–5. Kobayashi T, Nishijima M, Tamori Y, et al. Acyl phosphatidylglycerol of Escherichia coli. Biochim Biophys Acta 1980;620:356–63. Kodaki T, Yamashita S. Yeast phosphatidylethanolamine methylation pathway. Cloning and characterization of two distinct methyltransferase genes. J Biol Chem 1987;262: 15428–35. Komaniecka I, Choma A, Mazur A, et al. Occurrence of an unusual hopanoid-containing lipid A among lipopolysaccharides from Bradyrhizobium species. J Biol Chem 2014;289:35644–55. Kontnik R, Bosak T, Butcher RA, et al. Sporulenes, heptaprenyl metabolites from Bacillus subtilis spores. Org Lett 2008;10:3551–4. Kordulakova J, Gilleron M, Puzo G, et al. Identification of the required acyltransferase step in the biosynthesis of the phosphatidylinositol mannosides of mycobacterium species. J Biol Chem 2003;278:36285–95. Kornspan JD, Rottem S. The phospholipid profile of mycoplasmas. J Lipids 2012;2012:640762. Kwak BY, Zhang YM, Yun M, et al. Structure and mechanism of CTP:phosphocholine cytidylyltransferase (LicC) from Streptococcus pneumoniae. J Biol Chem 2002;277: 4343–50. Lagier JC, Armougom F, Million M, et al. Microbial culturomics: paradigm shift in the human gut microbiome study. Clin Microbiol Infec 2012;18:1185–93. Lagutin K, MacKenzie A, Houghton KM, et al. The identification and quantification of phospholipids from thermus and meiothermus bacteria. Lipids 2014;49:1133–41. Lamb DC, Jackson CJ, Warrilow AG, et al. Lanosterol biosynthesis in the prokaryote Methylococcus capsulatus: insight into the evolution of sterol biosynthesis. Mol Biol Evol 2007;24: 1714–21.

23

´ ´ Laneelle MA, Prome´ D, Laneelle G, et al. Ornithine lipid of Mycobacterium tuberculosis: its distribution in some slow- and fast-growing mycobacteria. J Gen Microbiol 1990;136:773–8. Lata P, Lal D, Lal R. Flavobacterium ummariense sp. nov., isolated from hexachlorocyclohexane-contaminated soil, and emended description of Flavobacterium ceti Vela et al. 2007. Int J Syst Evol Micr 2012;62:2674–9. Leach S, Harvey P, Wait R. Changes with growth rate in the membrane lipid composition of and amino acid utilization by continuous cultures of Campylobacter jejuni. J Appl Microbiol 1997;82:631–40. Lennarz WJ, Bonsen PP, van Deenen LL. Substrate specificity of O-L-lysylphosphatidylglycerol synthetase. Enzymatic studies on the structure of O-L-lysylphosphatidylglycerol. Biochemistry 1967;6:2307–12. Lewenza S, Falsafi R, Bains M, et al. The olsA gene mediates the synthesis of an ornithine lipid in Pseudomonas aeruginosa during growth under phosphate-limiting conditions, but is not involved in antimicrobial peptide susceptibility. FEMS Microbiol Lett 2011;320:95–102. Li QX, Dowhan W. Structural characterization of Escherichia coli phosphatidylserine decarboxylase. J Biol Chem 1988;263:11516–22. Li Y, Wang X, Ernst RK. A rapid one-step method for the characterization of membrane lipid remodeling in Francisella using matrix-assisted laser desorption ionization time-offlight tandem mass spectrometry. Rapid Commun Mass Sp 2011;25:2641–8. Lightner VA, Larson TJ, Tailleur P, et al. Membrane phospholipid synthesis in Escherichia coli. Cloning of a structural gene (plsB) of the sn-glycerol-3-phosphate acyltransferase. J Biol Chem 1980;255:9413–20. Liscovitch M, Czarny M, Fiucci G, et al. Phospholipase D: molecular and cell biology of a novel gene family. Biochem J 2000;345(Pt 3):401–15. Liu W, Sakr E, Schaeffer P, et al. Ribosylhopane, a novel bacterial hopanoid, as precursor of C35 bacteriohopanepolyols in Streptomyces coelicolor A3(2). Chembiochem 2014;15:2156–61. Livermore BP, Johnson RC. Lipids of the Spirochaetales: comparison of the lipids of several members of the genera Spirochaeta, Treponema, and Leptospira. J Bacteriol 1974;120:1268–73. ´ Lopez-Lara IM, Gao JL, Soto MJ, et al. Phosphorus-free membrane lipids of Sinorhizobium meliloti are not required for the symbiosis with alfalfa but contribute to increased cell yields under phosphorus-limiting conditions of growth. Mol Plant Microbe In 2005;18:973–82. ´ Lopez-Lara IM, Sohlenkamp C, Geiger O. Membrane lipids in plant-associated bacteria: their biosyntheses and possible functions. Mol Plant Microbe In 2003;16:567–79. Lorenzen W, Bozhuyuk KA, Cortina NS, et al. A comprehensive insight into the lipid composition of Myxococcus xanthus by UPLC ESI-MS. J Lipid Res 2014;55:2620–33. Lowther J, Charmier G, Raman MC, et al. Role of a conserved arginine residue during catalysis in serine palmitoyltransferase. FEBS Lett 2011;585:1729–34. Lowther J, Naismith JH, Dunn TM, et al. Structural, mechanistic and regulatory studies of serine palmitoyltransferase. Biochem Soc T 2012;40:547–54. Lu YH, Guan Z, Zhao J, et al. Three phosphatidylglycerolphosphate phosphatases in the inner membrane of Escherichia coli. J Biol Chem 2011;286:5506–18. Lu YJ, Zhang YM, Grimes KD, et al. Acyl-phosphates initiate membrane phospholipid synthesis in Gram-positive pathogens. Mol Cell 2006;23:765–72.

24


Makarova KS, Aravind L, Wolf YI, et al. Genome of the extremely radiation-resistant bacterium Deinococcus radiodurans viewed from the perspective of comparative genomics. Microbiol Mol Biol R 2001;65:44–79. Makula RA, Finnerty WR. Phospholipid composition of Desulfovibrio species. J Bacteriol 1974;120:1279–83. Makula RA, Finnerty WR. Isolation and characterization of an ornithine-containing lipid from Desulfovibrio gigas. J Bacteriol 1975;123:523–9. Malott RJ, Steen-Kinnaird BR, Lee TD, et al. Identification of hopanoid biosynthesis genes involved in polymyxin resistance in Burkholderia multivorans. Antimicrob Agents Ch 2012;56:464–71. Manca MC, Nicolaus B, Lanzotti V, et al. Glycolipids from Thermotoga maritima, a hyperthermophilic microorganism belonging to Bacteria domain. Biochim Biophys Acta 1992;1124: 249–52. Marr AG, Ingraham JL. Effect of temperature on the composition of fatty acids in Escherichia coli. J Bacteriol 1962;84: 1260–7. Mastronicolis SK, Arvanitis N, Karaliota A, et al. Coordinated regulation of cold-induced changes in fatty acids with cardiolipin and phosphatidylglycerol composition among phospholipid species for the food pathogen Listeria monocytogenes. Appl Environ Microb 2008;74:4543–9. Matsumoto K. Phosphatidylserine synthase from bacteria. Biochim Biophys Acta 1997;1348:214–27. Matsumoto K. Dispensable nature of phosphatidylglycerol in Escherichia coli: dual roles of anionic phospholipids. Mol Microbiol 2001;39:1427–33. Matsumoto K, Okada M, Horikoshi Y, et al. Cloning, sequencing, and disruption of the Bacillus subtilis psd gene coding for phosphatidylserine decarboxylase. J Bacteriol 1998;180: 100–6. Matsunaga I, Yokotani N, Gotoh O, et al. Molecular cloning and expression of fatty acid α-hydroxylase from Sphingomonas paucimobilis. J Biol Chem 1997;272:23592–6. Meiers M, Volz C, Eisel J, et al. Altered lipid composition in Streptococcus pneumoniae cpoA mutants. BMC Microbiol 2014;14:12. Minder AC, de Rudder KE, Narberhaus F, et al. Phosphatidylcholine levels in Bradyrhizobium japonicum membranes are critical for an efficient symbiosis with the soybean host plant. Mol Microbiol 2001;39:1186–98. Miyazaki C, Kuroda M, Ohta A, et al. Genetic manipulation of membrane phospholipid composition in Escherichia coli: pgsA mutants defective in phosphatidylglycerol synthesis. P Natl Acad Sci USA 1985;82:7530–4. Mizoguchi T, Harada J, Yoshitomi T, et al. A variety of glycolipids in green photosynthetic bacteria. Photosynth Res 2013;114:179–88. Moore EK, Hopmans EC, Rijpstra WI, et al. Novel mono-, di-, and trimethylornithine membrane lipids in northern wetland planctomycetes. Appl Environ Microb 2013;79:6874–84. Moribayashi A, Goto N, Arimitsu Y, et al. Lipids and fatty-acids of Leptospira interrogans serovar copenhageni virulent-strain Shibaura. Jpn J Med Sci Biol 1991;44:87–97. Morii H, Ogawa M, Fukuda K, et al. A revised biosynthetic pathway for phosphatidylinositol in Mycobacteria. J Biochem 2010;148:593–602. Morii H, Ogawa M, Fukuda K, et al. Ubiquitous distribution of phosphatidylinositol phosphate synthase and archaetidylinositol phosphate synthase in Bacteria and Archaea, which contain inositol phospholipid. Biochem Bioph Res Co 2014;443:86–90.

Moser R, Aktas M, Fritz C, et al. Discovery of a bifunctional cardiolipin/phosphatidylethanolamine synthase in bacteria. Mol Microbiol 2014a;92:959–72. Moser R, Aktas M, Narberhaus F. Phosphatidylcholine biosynthesis in Xanthomonas campestris via a yeast-like acylation pathway. Mol Microbiol 2014b;91:736–50. Muller FD, Beck S, Strauch E, et al. Bacterial predators possess ¨ unique membrane lipid structures. Lipids 2011;46:1129–40. Nahaie MR, Goodfellow M, Minnikin DE, et al. Polar lipid and isoprenoid quinone composition in the classification of Staphylococcus. J Gen Microbiol 1984;130:2427–37. Nakashima A, Hosaka K, Nikawa J. Cloning of a human cDNA for CTP-phosphoethanolamine cytidylyltransferase by complementation in vivo of a yeast mutant. J Biol Chem 1997;272:9567–72. Nampoothiri KM, Hoischen C, Bathe B, et al. Expression of genes of lipid synthesis and altered lipid composition modulates L-glutamate efflux of Corynebacterium glutamicum. Appl Microbiol Biot 2002;58:89–96. Nesbitt JA, 3rd, Lennarz WJ. Participation of aminoacyl transfer ribonucleic acid in aminoacyl phosphatidylglycerol synthesis. I. Specificity of lysyl phosphatidylglycerol synthetase. J Biol Chem 1968;243:3088–95. Neunlist S, Rohmer M. Novel hopanoids from the methylotrophic bacteria Methylococcus capsulatus and Methylomonas methanica. (22S)-35-aminobacteriohopane-30,31,32,33,34pentol and (22S)-35-amino-3 beta-methylbacteriohopane30,31,32,33,34-pentol. Biochem J 1985;231:635–9. Nguyen NA, Sallans L, Kaneshiro ES. The major glycerophospholipids of the predatory and parasitic bacterium Bdellovibrio bacteriovorus HID5. Lipids 2008;43:1053–63. Nichols FC, Riep B, Mun J, et al. Structures and biological activity of phosphorylated dihydroceramides of Porphyromonas gingivalis. J Lipid Res 2004;45:2317–30. Nichols FC, Riep B, Mun J, et al. Structures and biological activities of novel phosphatidylethanolamine lipids of Porphyromonas gingivalis. J Lipid Res 2006;47:844–53. Nichols FC, Rojanasomsith K. Porphyromonas gingivalis lipids and diseased dental tissues. Oral Microbiol Immun 2006;21:84–92. Niederweis M, Danilchanka O, Huff J, et al. Mycobacterial outer membranes: in search of proteins. Trends Microbiol 2010;18:109–16. Nikaido H. Molecular basis of bacterial outer membrane permeability revisited. Microbiol Mol Biol R 2003;67:593–656. Nikawa J, Yamashita S. Phosphatidylinositol synthase from yeast. Biochim Biophys Acta 1997;1348:173–8. Nishi H, Komatsuzawa H, Fujiwara T, et al. Reduced content of lysyl-phosphatidylglycerol in the cytoplasmic membrane affects susceptibility to moenomycin, as well as vancomycin, gentamicin, and antimicrobial peptides, in Staphylococcus aureus. Antimicrob Agents Ch 2004;48:4800–7. Nishijima S, Asami Y, Uetake N, et al. Disruption of the Escherichia coli cls gene responsible for cardiolipin synthesis. J Bacteriol 1988;170:775–80. Nogly P, Gushchin I, Remeeva A, et al. X-ray structure of a CDPalcohol phosphatidyltransferase membrane enzyme and insights into its catalytic mechanism. Nat Commun 2014;5: 4169. Okada M, Matsuzaki H, Shibuya I, et al. Cloning, sequencing, and expression in Escherichia coli of the Bacillus subtilis gene for phosphatidylserine synthase. J Bacteriol 1994;176:7456–61. Oku Y, Kurokawa K, Ichihashi N, et al. Characterization of the Staphylococcus aureus mprF gene, involved in lysinylation of phosphatidylglycerol. Microbiology 2004;150:45–51.


Olofsson A, Vallstrom A, Petzold K, et al. Biochemical and functional characterization of Helicobacter pylori vesicles. Mol Microbiol 2010;77:1539–55. Olsen RW, Ballou CE. Acyl phosphatidylglycerol. A new phospholipid from Salmonella typhimurium. J Biol Chem 1971;246: 3305–13. Ostafin M, Haber J, Doronina NV, et al. Methylobacterium extorquens strain P14, a new methylotrophic bacteria producing poly-beta-hydroxybutyrate (PHB). Acta Microbiol Pol 1999;48:39–51. Ostberg Y, Berg S, Comstedt P, et al. Functional analysis of a lipid galactosyltransferase synthesizing the major envelope lipid in the Lyme disease spirochete Borrelia burgdorferi. FEMS Microbiol Lett 2007;272:22–9. Ourisson G, Rohmer M, Poralla K. Prokaryotic hopanoids and other polyterpenoid sterol surrogates. Annu Rev Microbiol 1987;41:301–33. ´ Palacios-Chaves L, Conde-Alvarez R, Gil-Ram´ırez Y, et al. Brucella abortus ornithine lipids are dispensable outer membrane components devoid of a marked pathogen-associated molecular pattern. PLoS One 2011;6:e16030. Parsons JB, Rock CO. Bacterial lipids: metabolism and membrane homeostasis. Prog Lipid Res 2013;52:249–76. Pata MO, Hannun YA, Ng CK. Plant sphingolipids: decoding the enigma of the Sphinx. New Phytol 2010;185:611–30. ´ ´ M. Metabolic plasticity for isoPerez-Gil J, Rodr´ıguez-Concepcion prenoid biosynthesis in bacteria. Biochem J 2013;452:19–25. Perzl M, Muller P, Poralla K, et al. Squalene-hopene cyclase from Bradyrhizobium japonicum: cloning, expression, sequence analysis and comparison to other triterpenoid cyclases. Microbiology 1997;143(Pt 4):1235–42. Peschel A, Jack RW, Otto M, et al. Staphylococcus aureus resistance to human defensins and evasion of neutrophil killing via the novel virulence factor MprF is based on modification of membrane lipids with L-lysine. J Exp Med 2001;193: 1067–76. Peter-Katalinic J, Fischer W. α-D-glucopyranosyl-, D-alanyl- and L-lysylcardiolipin from gram-positive bacteria: analysis by fast atom bombardment mass spectrometry. J Lipid Res 1998;39:2286–92. Pitta TP, Leadbetter ER, Godchaux W, 3rd. Increase of ornithine amino lipid content in a sulfonolipid-deficient mutant of Cytophaga johnsonae. J Bacteriol 1989;171:952–7. Pramanik BN, Zechman JM, Das PR, et al. Bacterial phospholipid analysis by fast atom bombardment mass spectrometry. Biomed Environ Mass 1990;19:164–70. Raetz CR, Dowhan W. Biosynthesis and function of phospholipids in Escherichia coli. J Biol Chem 1990;265:1235–8. Raetz CR, Reynolds CM, Trent MS, et al. Lipid A modification systems in gram-negative bacteria. Annu Rev Biochem 2007;76:295–329. Raetz CR, Whitfield C. Lipopolysaccharide endotoxins. Annu Rev Biochem 2002;71:635–700. Rahman MM, Kolli VS, Kahler CM, et al. The membrane phospholipids of Neisseria meningitidis and Neisseria gonorrhoeae as characterized by fast atom bombardment mass spectrometry. Microbiology 2000;146(Pt 8):1901–11. Raman MC, Johnson KA, Clarke DJ, et al. The serine palmitoyltransferase from Sphingomonas wittichii RW1: an interesting link to an unusual acyl carrier protein. Biopolymers 2010;93:811–22. Ramana VV, Chakravarthy SK, Raj PS, et al. Descriptions of Rhodopseudomonas parapalustris sp. nov., Rhodopseudomonas harwoodiae sp. nov. and Rhodopseudomonas

25

pseudopalustris sp. nov., and emended description of Rhodopseudomonas palustris. Int J Syst Evol Micr 2012;62: 1790–8. Ratledge C, Wilkinson SG. Microbial Lipids. London, San Diego: Academic Press, 1988. Reichmann NT, Grundling A. Location, synthesis and function ¨ of glycolipids and polyglycerolphosphate lipoteichoic acid in Gram-positive bacteria of the phylum Firmicutes. FEMS Microbiol Lett 2011;319:97–105. Ricci JN, Coleman ML, Welander PV, et al. Diverse capacity for 2-methylhopanoid production correlates with a specific ecological niche. ISME J 2014;8:675–84. Riekhof WR, Andre C, Benning C. Two enzymes, BtaA and BtaB, are sufficient for betaine lipid biosynthesis in bacteria. Arch Biochem Biophys 2005a;441:96–105. Riekhof WR, Naik S, Bertrand H, et al. Phosphate starvation in fungi induces the replacement of phosphatidylcholine with the phosphorus-free betaine lipid diacylglycerylN,N,N-trimethylhomoserine. Eukaryot Cell 2014;13: 749–57. Riekhof WR, Sears BB, Benning C. Annotation of genes involved in glycerolipid biosynthesis in Chlamydomonas reinhardtii: discovery of the betaine lipid synthase BTA1Cr . Eukaryot Cell 2005;4:242–52. Ring MW, Schwar G, Bode HB. Biosynthesis of 2-hydroxy and isoeven fatty acids is connected to sphingolipid formation in myxobacteria. Chembiochem 2009;10:2003–10. Rock CO, Heath RJ, Park HW, et al. The licC gene of Streptococcus pneumoniae encodes a CTP:phosphocholine cytidylyltransferase. J Bacteriol 2001;183:4927–31. Roh SW, Lee HW, Yim KJ, et al. Rhodopirellula rosea sp. nov., a novel bacterium isolated from an ark clam Scapharca broughtonii. J Microbiol 2013;51:301–4. ´ Rojas-Jimenez K, Sohlenkamp C, Geiger O, et al. A ClC chloride channel homolog and ornithine-containing membrane lipids of Rhizobium tropici CIAT899 are involved in symbiotic efficiency and acid tolerance. Mol Plant Microbe In 2005;18:1175–85. Romantsov T, Guan Z, Wood JM. Cardiolipin and the osmotic stress responses of bacteria. Biochim Biophys Acta 2009;1788:2092–100. Romantsov T, Helbig S, Culham DE, et al. Cardiolipin promotes polar localization of osmosensory transporter ProP in Escherichia coli. Mol Microbiol 2007;64:1455–65. Romantsov T, Stalker L, Culham DE, et al. Cardiolipin controls the osmotic stress response and the subcellular location of transporter ProP in Escherichia coli. J Biol Chem 2008;283: 12314–23. Rossak M, Schafer A, Xu N, et al. Accumulation of sulfoquinovosyl-1-O-dihydroxyacetone in a sulfolipiddeficient mutant of Rhodobacter sphaeroides inactivated in sqdC. Arch Biochem Biophys 1997;340:219–30. Rossak M, Tietje C, Heinz E, et al. Accumulation of UDPsulfoquinovose in a sulfolipid-deficient mutant of Rhodobacter sphaeroides. J Biol Chem 1995;270:25792–7. Rowe NJ, Tunstall J, Galbraith L, et al. Lipid composition and taxonomy of [Pseudomonas] echinoides: transfer to the genus Sphingomonas. Microbiology 2000;146(Pt 11):3007–12. Roy H, Dare K, Ibba M. Adaptation of the bacterial membrane to changing environments using aminoacylated phospholipids. Mol Microbiol 2009;71:547–50. Roy H, Ibba M. RNA-dependent lipid remodeling by bacterial multiple peptide resistance factors. P Natl Acad Sci USA 2008;105:4667–72.

26


Roy H, Ibba M. Broad range amino acid specificity of RNAdependent lipid remodeling by multiple peptide resistance factors. J Biol Chem 2009;284:29677–83. Ruzin A, Severin A, Moghazeh SL, et al. Inactivation of mprF affects vancomycin susceptibility in Staphylococcus aureus. Biochim Biophys Acta 2003;1621:117–21. Saenz JP, Sezgin E, Schwille P, et al. Functional convergence of hopanoids and sterols in membrane ordering. P Natl Acad Sci USA 2012a;109:14236–40. Saenz JP, Waterbury JB, Eglinton TI, et al. Hopanoids in marine cyanobacteria: probing their phylogenetic distribution and biological role. Geobiology 2012b;10:311–9. Salzberg LI, Helmann JD. Phenotypic and transcriptomic characterization of Bacillus subtilis mutants with grossly altered membrane composition. J Bacteriol 2008;190:7797– 807. ´ M, Geiger O, Guan Z, et al. A eukaryote-like Sandoval-Calderon cardiolipin synthase is present in Streptomyces coelicolor and in most actinobacteria. J Biol Chem 2009;284:17383–90. Sauvageau J, Ryan J, Lagutin K, et al. Isolation and structural characterisation of the major glycolipids from Lactobacillus plantarum. Carbohyd Res 2012;357:151–6. Schmerk CL, Bernards MA, Valvano MA. Hopanoid production is required for low-pH tolerance, antimicrobial resistance, and motility in Burkholderia cenocepacia. J Bacteriol 2011;193: 6712–23. Schmerk CL, Welander PV, Hamad MA, et al. Elucidation of the Burkholderia cenocepacia hopanoid biosynthesis pathway uncovers functions for conserved proteins in hopanoid-producing bacteria. Environ Microbiol 2015;17: 735–50. Seipke RF, Loria R. Hopanoids are not essential for growth of Streptomyces scabies 87–22. J Bacteriol 2009;191:5216–23. Semeniuk A, Sohlenkamp C, Duda K, et al. A bifunctional glycosyltransferase from Agrobacterium tumefaciens synthesizes monoglucosyl and glucuronosyl diacylglycerol under phosphate deprivation. J Biol Chem 2014;289:10104–14. Senff LM, Wegener WS, Brooks GF, et al. Phospholipid composition and phospholipase A activity of Neisseria gonorrhoeae. J Bacteriol 1976;127:874–80. Serricchio M, Butikofer P. An essential bacterial-type cardiolipin synthase mediates cardiolipin formation in a eukaryote. P Natl Acad Sci USA 2012;109:E954–61. Shi W, Bogdanov M, Dowhan W, et al. The pss and psd genes are required for motility and chemotaxis in Escherichia coli. J Bacteriol 1993;175:7711–4. Shimizu T, Ohtani K, Hirakawa H, et al. Complete genome sequence of Clostridium perfringens, an anaerobic flesh-eater. P Natl Acad Sci USA 2002;99:996–1001. Shimojima M. Biosynthesis and functions of the plant sulfolipid. Prog Lipid Res 2011;50:234–9. Silipo A, Vitiello G, Gully D, et al. Covalently linked hopanoidlipid A improves outer-membrane resistance of a Bradyrhizobium symbiont of legumes. Nat Commun 2014;5:5106. Sinninghe Damste´ JS, Rijpstra WI, Geenevasen JA, et al. Structural identification of ladderane and other membrane lipids of planctomycetes capable of anaerobic ammonium oxidation (anammox). FEBS J 2005;272:4270–83. Smith RD, Salyers AA. Incorporation of leucine into phospholipids of Bacteroides thetaiotaomicron. J Bacteriol 1981;145:8–13. Sohlenkamp C, de Rudder KE, Geiger O. Phosphatidylethanolamine is not essential for growth of Sinorhizobium meliloti on complex culture media. J Bacteriol 2004;186:1667–77.

¨ Sohlenkamp C, de Rudder KE, Rohrs VV, et al. Cloning and characterization of the gene for phosphatidylcholine synthase. J Biol Chem 2000;275:27500. Sohlenkamp C, Galindo-Lagunas KA, Guan Z, et al. The lipid lysyl-phosphatidylglycerol is present in membranes of Rhizobium tropici CIAT899 and confers increased resistance to polymyxin B under acidic growth conditions. Mol Plant Microbe In 2007;20:1421–30. ´ Sohlenkamp C, Lopez-Lara IM, Geiger O. Biosynthesis of phosphatidylcholine in bacteria. Prog Lipid Res 2003;42:115–62. Sol´ıs-Oviedo RL, Mart´ınez-Morales F, Geiger O, et al. Functional and topological analysis of phosphatidylcholine synthase from Sinorhizobium meliloti. Biochim Biophys Acta 2012;1821:573–81. Sorensen PG, Cox RP, Miller M. Chlorosome lipids from Chlorobium tepidum: characterization and quantification of polar lipids and wax esters. Photosynth Res 2008;95:191–6. Stalberg K, Neal AC, Ronne H, et al. Identification of a novel GPCAT activity and a new pathway for phosphatidylcholine biosynthesis in S. cerevisiae. J Lipid Res 2008;49:1794–806. Stein J, Budzikiewicz H. Bacterial Components .36. Ceramide-1Phosphoethanolamines from Myxococcus stipitatus. Z Naturforsch B 1988;43:1063–7. Steiner S, Conti SF, Lester RL. Occurrence of phosphonosphingolipids in Bdellovibrio bacteriovorus strain UKi2. J Bacteriol 1973;116:1199–211. Sturt HF, Summons RE, Smith K, et al. Intact polar membrane lipids in prokaryotes and sediments deciphered by highperformance liquid chromatography/electrospray ionization multistage mass spectrometry—new biomarkers for biogeochemistry and microbial ecology. Rapid Commun Mass Sp 2004;18:617–28. Sud IJ, Feingold DS. Phospholipids and fatty acids of Neisseria gonorrhoeae. J Bacteriol 1975;124:713–7. Svetlikova Z, Barath P, Jackson M, et al. Purification and characterization of the acyltransferase involved in biosynthesis of the major mycobacterial cell envelope glycolipid– monoacylated phosphatidylinositol dimannoside. Protein Expres Purif 2014;100:33–9. Tahara Y, Kameda M, Yamada Y, et al. New lipid—ornithine and taurine-containing cerilipin. Agr Biol Chem 1976;40:243–4. Tan BK, Bogdanov M, Zhao J, et al. Discovery of a cardiolipin synthase utilizing phosphatidylethanolamine and phosphatidylglycerol as substrates. P Natl Acad Sci USA 2012;109:16504–9. Tannaes T, Bukholm IK, Bukholm G. High relative content of lysophospholipids of Helicobacter pylori mediates increased risk for ulcer disease. FEMS Immunol Med Mic 2005;44:17–23. Tannaes T, Grav HJ, Bukholm G. Lipid profiles of Helicobacter pylori colony variants. APMIS 2000;108:349–56. Tavana AM, Korachi M, Boote V, et al. Phospholipid analogues of Porphyromonas gingivalis. J Appl Microbiol 2000;88:791–9. Taylor CJ, Anderson AJ, Wilkinson SG. Phenotypic variation of lipid composition in Burkholderia cepacia: a response to increased growth temperature is a greater content of 2hydroxy acids in phosphatidylethanolamine and ornithine amide lipid. Microbiology 1998;144:1737–45. Thedieck K, Hain T, Mohamed W, et al. The MprF protein is required for lysinylation of phospholipids in listerial membranes and confers resistance to cationic antimicrobial peptides (CAMPs) on Listeria monocytogenes. Mol Microbiol 2006;62:1325–39. Thiele OW, Schwinn G. The free lipids of Brucella melitensis and Bordetella pertussis. Eur J Biochem 1973;34:333–44.


Tornabene TG. Lipid composition of selected strains of Yersinia pestis and Yersinia pseudotuberculosis. Biochim Biophys Acta 1973;306:173–85. Trana AK, Khuller GK, Subrahmanyam D. Metabolism of phospholipids in Nocardia polychromogenes. J Gen Microbiol 1980;116:89–92. ´ ´ Trombe´ MC, Laneelle MA, Laneelle G. Lipid composition of aminopterin-resistant and sensitive strains of Streptococcus pneumoniae. Effect of aminopterin inhibition. Biochim Biophys Acta 1979;574:290–300. Tsatskis Y, Khambati J, Dobson M, et al. The osmotic activation of transporter ProP is tuned by both its C-terminal coiled-coil and osmotically induced changes in phospholipid composition. J Biol Chem 2005;280:41387–94. ´ Vences-Guzman MA, Geiger O, Sohlenkamp C. Sinorhizobium meliloti mutants deficient in phosphatidylserine decarboxylase accumulate phosphatidylserine and are strongly affected during symbiosis with alfalfa. J Bacteriol 2008;190:6846–56. ´ MA, Geiger O, Sohlenkamp C. Ornithine lipids Vences-Guzman and their structural modifications: from A to E and beyond. FEMS Microbiol Lett 2012;335:1–10. ´ MA, Guan Z, Bermudez-Barrientos Vences-Guzman JR, et al. ´ Agrobacteria lacking ornithine lipids induce more rapid tumour formation. Environ Microbiol 2013;15:895–906. ´ MA, Guan Z, Escobedo-Hinojosa WI, et al. DisVences-Guzman covery of a bifunctional acyltransferase responsible for ornithine lipid synthesis in Serratia proteamaculans. Environ Microbiol 2014, DOI:10.1111/1462-2920.12562. ´ MA, Guan Z, Ormeno-Orrillo E, et al. HydroxyVences-Guzman lated ornithine lipids increase stress tolerance in Rhizobium tropici CIAT899. Mol Microbiol 2011;79:1496–514. Vinuesa P, Neumann-Silkow F, Pacios-Bras C, et al. Genetic analysis of a pH-regulated operon from Rhizobium tropici CIAT899 involved in acid tolerance and nodulation competitiveness. Mol Plant Microbe In 2003;16:159–68. Wang XG, Scagliotti JP, Hu LT. Phospholipid synthesis in Borrelia burgdorferi: BB0249 and BB0721 encode functional phosphatidylcholine synthase and phosphatidylglycerolphosphate synthase proteins. Microbiology 2004;150:391–7. Weiser JN, Love JM, Moxon ER. The molecular mechanism of phase variation of H. influenzae lipopolysaccharide. Cell 1989;59:657–65. ´ Weissenmayer B, Gao JL, Lopez-Lara IM, et al. Identification of a gene required for the biosynthesis of ornithine-derived lipids. Mol Microbiol 2002;45:721–33. Weissenmayer B, Geiger O, Benning C. Disruption of a gene essential for sulfoquinovosyldiacylglycerol biosynthesis in Sinorhizobium meliloti has no detectable effect on root nodule symbiosis. Mol Plant Microbe In 2000;13:666–72. Welander PV, Coleman ML, Sessions AL, et al. Identification of a methylase required for 2-methylhopanoid production and implications for the interpretation of sedimentary hopanes. P Natl Acad Sci USA 2010;107:8537–42. Welander PV, Doughty DM, Wu CH, et al. Identification and characterization of Rhodopseudomonas palustris TIE1 hopanoid biosynthesis mutants. Geobiology 2012;10: 163–77. Welander PV, Hunter RC, Zhang L, et al. Hopanoids play a role in membrane integrity and pH homeostasis in Rhodopseudomonas palustris TIE-1. J Bacteriol 2009;191: 6145–56.

27

Welander PV, Summons RE. Discovery, taxonomic distribution, and phenotypic characterization of a gene required for 3-methylhopanoid production. P Natl Acad Sci USA 2012;109:12905–10. ¨ Wessel M, Klusener S, Godeke J, et al. Virulence of Agrobac¨ terium tumefaciens requires phosphatidylcholine in the bacterial membrane. Mol Microbiol 2006;62:906–15. White DC, Frerman FE. Extraction, characterization, and cellular localization of the lipids of Staphylococcus aureus. J Bacteriol 1967;94:1854–67. White RH. Biosynthesis of the sulfonolipid 2-amino-3-hydroxy15-methylhexadecane-1-sulfonic acid in the gliding bacterium Cytophaga johnsonae. J Bacteriol 1984;159:42–6. Wilderman PJ, Vasil AI, Martin WE, et al. Pseudomonas aeruginosa synthesizes phosphatidylcholine by use of the phosphatidylcholine synthase pathway. J Bacteriol 2002;184:4792–9. Williams JG, McMaster CR. Scanning alanine mutagenesis of the CDP-alcohol phosphotransferase motif of Saccharomyces cerevisiae cholinephosphotransferase. J Biol Chem 1998;273:13482–7. Xue HW, Hosaka K, Plesch G, et al. Cloning of Arabidopsis thaliana phosphatidylinositol synthase and functional expression in the yeast pis mutant. Plant Mol Biol 2000;42:757–64. Yabuuchi E, Kosako Y, Yano I, et al. Transfer of two Burkholderia and an Alcaligenes species to Ralstonia gen. Nov.: Proposal of Ralstonia pickettii (Ralston, Palleroni and Doudoroff 1973) comb. Nov., Ralstonia solanacearum (Smith 1896) comb. Nov. and Ralstonia eutropha (Davis 1969) comb. Nov. Microbiol Immunol 1995;39:897–904. Yao J, Abdelrahman YM, Robertson RM, et al. Type II fatty acid synthesis is essential for the replication of Chlamydia trachomatis. J Biol Chem 2014;289:22365–76. Yard BA, Carter LG, Johnson KA, et al. The structure of serine palmitoyltransferase; gateway to sphingolipid biosynthesis. J Mol Biol 2007;370:870–86. Yuzawa Y, Shimojima M, Sato R, et al. Cyanobacterial monogalactosyldiacylglycerol-synthesis pathway is involved in normal unsaturation of galactolipids and lowtemperature adaptation of Synechocystis sp. PCC 6803. Biochim Biophys Acta 2014;1841:475–83. Zavaleta-Pastor M, Sohlenkamp C, Gao JL, et al. Sinorhizobium meliloti phospholipase C required for lipid remodeling during phosphorus limitation. P Natl Acad Sci USA 2010;107:302–7. Zhang JR, Idanpaan-Heikkila I, Fischer W, et al. Pneumococcal licD2 gene is involved in phosphorylcholine metabolism. Mol Microbiol 1999;31:1477–88. Zhang X, Ferguson-Miller SM, Reid GE. Characterization of ornithine and glutamine lipids extracted from cell membranes of Rhodobacter sphaeroides. J Am Soc Mass Spectr 2009;20: 198–212. Zhang Y, Yang Z, Huang X, et al. Cloning, expression, and characterization of a thermostable PAP2L2, a new member of the type-2 phosphatidic acid phosphatase family from Geobacillus toebii T-85. Biosci Biotechnol Biochem 2008;72: 3134–41. Zhang YQ, Chen YG, Li WJ, et al. Sphingomonas yunnanensis sp. nov., a novel gram-negative bacterium from a contaminated plate. Int J Syst Evol Mic 2005;55:2361–4. Zhou P, Hu R, Chandan V, et al. Simultaneous analysis of cardiolipin and lipid A from Helicobacter pylori by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Mol Biosyst 2012;8:720–5.

Crystal structures of membrane lipids.

Structures of archaebacterial membrane lipids.

Diversity of bacterial manipulation of the host ubiquitin pathways.

Membrane lipids in Agrobacterium tumefaciens: biosynthetic pathways and importance for pathogenesis.

Interaction of lead and bacterial lipids.

Lipids in bacterial taxonomy - a taxonomist's view.

Fluidity of bacterial membrane lipids monitored by intramolecular excimerization of 1.3-di(2-pyrenyl)propane.

Bacterial flagellar diversity and significance in pathogenesis.

The diversity of membrane transporters encoded in bacterial arsenic-resistance operons.

Anesthetic action of membrane lipids.

Polyunsaturated fats, membrane lipids and animal longevity.

Biosynthesis of membrane lipids in rat axons.

Erythrocyte membrane lipids in Friedreich's ataxia.

Conformation of beta hairpins in protein structures: classification and diversity in homologous structures.

Sclerotic rings in mosasaurs (Squamata: Mosasauridae): structures and taxonomic diversity.

Polyunsaturated Lipids Regulate Membrane Domain Stability by Tuning Membrane Order.

Anionic lipids and the maintenance of membrane electrostatics in eukaryotes.

Erythrocyte membrane lipids and cationic transport systems in men.

Lipids and immunoinflammatory pathways of beta cell destruction.

Bacterial Strain Diversity Within Wounds.

BacDive--the Bacterial Diversity Metadatabase.

Lipid peroxidation in membrane lipids and action of glutathione peroxidase.

Plasma membrane lipids and their role in fungal virulence.

Mucosa-associated bacterial diversity in necrotizing enterocolitis.