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.

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 Gründling 2011) and a thin murein cell wall between them. Usually, the outer leaflet 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,N-trimethylhomoserine (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).

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; MMPE-monomethyl PE; DMPE-dimethyl PE; PC-phosphatidylcholine; LPC-lysophosphatidylcholine; GPC-glycerophosphocholine; PGP-phosphatidylglycerol phosphate; PG-phosphatidylglycerol; CL-cardiolipin; LCL-lysyl-cardiolipin; ACL-alanyl-cardiolipin; LPG-lysyl-phosphatidylglycerol; APG-alanyl-phosphatidylglycerol; ArPG-arginyl-phosphatidylglycerol; PIP-phosphatidylinositol phosphate; PI-phosphatidylinositol; PIM-phosphatidylinositol mannoside; PIM2-phosphatidylinositol dimannoside; Ac1PIM2-acyl PIM2; Ac2PIM2-diacyl PIM2; DGHS-diacylglyceryl homoserine; DGTS-diacylglyceryl-N,N,N-trimethylhomoserine; SQD-sulfoquinovosyl diacylglycerol; GTF-glycosyltransferase.

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; MMPE-monomethyl PE; DMPE-dimethyl PE; PC-phosphatidylcholine; LPC-lysophosphatidylcholine; GPC-glycerophosphocholine; PGP-phosphatidylglycerol phosphate; PG-phosphatidylglycerol; CL-cardiolipin; LCL-lysyl-cardiolipin; ACL-alanyl-cardiolipin; LPG-lysyl-phosphatidylglycerol; APG-alanyl-phosphatidylglycerol; ArPG-arginyl-phosphatidylglycerol; PIP-phosphatidylinositol phosphate; PI-phosphatidylinositol; PIM-phosphatidylinositol mannoside; PIM2-phosphatidylinositol dimannoside; Ac1PIM2-acyl PIM2; Ac2PIM2-diacyl PIM2; DGHS-diacylglyceryl homoserine; DGTS-diacylglyceryl-N,N,N-trimethylhomoserine; SQD-sulfoquinovosyl diacylglycerol; GTF-glycosyltransferase.

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.

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.

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); ClsC-cardiolipin 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.

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); ClsC-cardiolipin 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.

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, alanyl-cardiolipin; APT-phosphoaminopentanetetrol lipid; DEGP-dietherglycerophospholipid; AEGP-acyletherglycerophospholipid; ST-sterols; PL-plasmalogens; 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 PG, CL, PE, MMPE, DMPE, PC, OL, S2, GL, DGTS, LPG Geske et al. (2013); Vences-Guzmán et al. (2013); Aktas et al. (2014); Semeniuk et al. (2014); Wessel et al. (2006
 Sinorhizobium meliloti PG, CL, PE, MMPE, DMPE, PC, OL, DGTS, SQD de Rudder, López-Lara and Geiger (2000); de Rudder, Sohlenkamp and Geiger (1999); López-Lara, Sohlenkamp and Geiger (2003
 Rhizobium tropici PG, CL, PE, MMPE, DMPE, PC, OL, S2, P1, P2, DGTS, LPG Rojas-Jiménez et al. (2005); Sohlenkamp et al. (2007); Vences-Guzmán et al. (2011
 Brucella melitensis PG, CL, PE, PC, OL, LPG Palacios-Chaves et al. (2011); Bukata et al. (2008); Comerci et al. (2006); Conde-Alvarez et al. (2006
 Rhodobacter sphaeroides PG, CL, PE, PC, OL, DGTS, SQD, GL Catucci et al. (2004); Benning et al. (1993); Benning, Huang and Gage (1995); Zhang, Ferguson-Miller and Reid (2009
 Caulobacter crescentus PG, CL, LPG, acyl-PG, GL Contreras, Shapiro and Henry (1978); Jones and Smith (1979); De Siervo and Homola (1980
 Sphingomonas sp. PG, CL, PE, MMPE, DMPE, PC, SL, OL, LPG, DGTS, HOP Kawahara, Kuraishi and Zähringer (1999); Rowe et al. (2000); Huang et al. (2009); Zhang et al. (2005
 Methylobacterium extorquens PG, PE, PC, HOP Bradley et al. (2010); Ostafin et al. (1999); Welander et al. (2010
 Bradyrhizobium diazoefficiens PG, CL, PE, MMPE, DMPE, PC, Bravo et al. (2001); Hacker et al. (2008); Minder et al. (2001); 
 (formerly B. japonicum) HOP Perzl et al. (1997); Silipo et al. (2014
 Rhodopseudomonas palustris PG, CL, PE, PC, HOP Ramana et al. (2012); Welander et al. (2012
Beta   
 Burkholderia cenocepacia PG, CL, PE, PE-2OH, OL, P1, NL1, NL2, LPG, HOP González-Silva et al. (2011); Taylor, Anderson and Wilkinson (1998); Schmerk, Bernards and Valvano (2011); Schmerk et al. (2015
 Ralstonia solanacearum PG, PE, PE-2OH, lyso-PE, OL, LPG Yabuuchi et al. (1995
 Neisseria gonorrhoeae PG, CL, PE, lyso-PE Rahman et al. (2000); Senff et al. (1976); Sud and Feingold (1975
 Bordetella pertussis PG, CL, PE, OL, lyso-PE Kawai and Moribayashi (1982); Kawai, Moribayashi and Yano (1982); Thiele and Schwinn (1973
Gamma   
 Escherichia coli PG, CL, PE Parsons and Rock (2013); Dowhan (2013
 Pseudomonas aeruginosa PG, CL, PE, PC, OL, APG Lewenza et al. (2011); Wilderman et al. (2002); Klein et al. (2009); Pramanik et al. (1990
 Vibrio cholerae PG, CL, PE, OL Giles et al. (2011
 Salmonella typhimurium PG, CL, PE, acyl-PG Ames (1968); Olsen and Ballou (1971
 Serratia proteamaculans PG, CL, PE, OL, NL1 Vences-Guzmán et al. (2014
 Legionella pneumophila PG, CL, PE, MMPE, DMPE, PC Conover et al. (2008
 Xanthomonas campestris PG, CL, PE, MMPE, PC, LPG Moser et al. (2014a); Moser, Aktas and Narberhaus (2014b
 Acidithiobacillus ferroxidans PG, CL, PE, MMPE, DMPE, PC, OL, HOP Barridge and Shively (1968); Dees and Shively (1982); Jones et al. (2012
 Francisella tularensis PG, CL, PE, PC, lyso-PE Li, Wang and Ernst (2011
 Methylococcus capsulatus PG, PE, ST, HOP Lamb et al. (2007); Welander and Summons (2012); Fang, Barcelona and Semrau (2000
Delta   
 Desulfovibrio sp. PG, CL, PS, lyso-PS, PE, OL, DGTS, HOP Makula and Finnerty (1974); Makula and Finnerty (1975); Blumenberg et al. (2009
 Myxococcus xanthus PG, CL, PE, SL, PI, PL Lorenzen et al. (2014); Ring et al. (2009
 Stigmatella aurantiaca PG, PE, lyso-PE, PI, SL, ST, OL, LPG Ring et al. (2009); Caillon, Lubochinsky and Rigomier (1983); Bode et al. (2003
 Bdellovibrio bacteriovorus PG, CL, PS, PE, PT, acyl-PE, SL Müller et al. (2011); Nguyen, Sallans and Kaneshiro (2008); Steiner, Conti and Lester (1973
Epsilon   
 Helicobacter pylori PG, CL, PE, lyso-PE Zhou et al. (2012); Olofsson et al. (2010); Tannaes et al. (2000, 2005
 Campylobacter jejuni PG, PE Leach, Harvey and Wait (1997
Cyanobacteria   
 Synechococcus sp. PCC 7942 PG, SQD, GL, CL Güler et al. (1996
 Synechocystis sp. PCC 6803 PG, SQD, GL, HOP Yuzawa et al. (2014
Deinococcales   
 Deinococcus radiodurans GL, GPL Anderson and Hansen (1985); Huang and Anderson (1989, 1995
Chloroflexi (green    
non-sulfur bacteria) Chloroflexus aurantiacus PG, CL, SQD, GL Kenyon and Gray (1974); Mizoguchi et al. (2013
Aquificae   
 Aquifex aeolicus DEGP, AEGP, APT Jahnke et al. (2001); Sturt et al. (2004
Thermotogae   
 Thermotoga maritima DEGP, AEGP, GL Manca et al. (1992); Damsté et al. (2007); Huber et al. (1986
Fusobacteria   
 Fusobacterium mortiferum PG, CL, PE, PL Hagen (1974
Chlamydiae   
 Chlamydia pneumonia PE, PC, host-derived membrane lipids Yao et al. (2014); Hatch and McClarty (1998
CFB Group   
 Flavobacterium johnsoniae PE, PC, OL, P1, SOL, GCL Pitta, Leadbetter and Godchaux (1989); Kawazoe et al. (1991
 Bacteroidetes thetaiotamicron PG, CL, PS, PE, SL Smith and Salyers (1981
 Porphyromonas gingivalis PG, PE, SL Nichols and Rojanasomsith (2006); Nichols et al. (2004, 2006); Tavana et al. (2000
Chlorobium (green    
sulfur bacteria) Chlorobium tepidum PG, CL, PE, GL Mizoguchi et al. (2013); Sorensen, Cox and Miller (2008
Actinobacteria   
 Mycobacterium tuberculosis PG, CL, PE, PI, PIM, OL Khuller et al. (1982); Jackson, Crick and Brennan (2000); Berg et al. (2007); Lanéelle et al. (1990
 Nocardia sp. CL, PE, PI, PIM, OL, SQD, HOP Khuller and Brennan (1972); Trana, Khuller and Subrahmanyam (1980
 Streptomyces coelicolor PG, CL, PE, PI, PIM, OL, HOP Sandoval-Calderón et al. (2009); Jyothikumar et al. (2012); Liu et al. (2014
 Corynebacterium glutamicum PG, CL, PI, PIM Nampoothiri et al. (2002
Spirochetes   
 Borrelia burgdorferi PG, PC, GL Wang, Scagliotti and Hu (2004); Ostberg et al. (2007
 Treponema denticola PG, CL, PE, PC Kent et al. (2004
 Leptospira interrogans PG, PE, CL, Lyso-PE, GL, OL Livermore and Johnson (1974); Moribayashi et al. (1991
Planctomycetes   
 Singulisphaera acidiphila PG, PC, TMOL, PA, HOP Moore et al. (2013
 Rhodopirellula baltica PG, CL, PE, PC, DGTS Bondoso et al. (2014); Roh et al. (2013
Firmicutes   
 Bacillus subtilis PG, CL, PE, LPG, GL, GPL Gidden et al. (2009); den Kamp et al. (1969); Salzberg and Helmann (2008); Jorasch et al. (1998
 Listeria monocytogenes PG, CL, LPG, LCL, PI Fischer and Leopold (1999); Dare et al. (2014); Thedieck et al. (2006); Mastronicolis et al. (2008
 Staphylococcus aureus PG, CL, LPG, GPL Oku et al. (2004); White and Frerman (1967
 Lactobacillus sp. PG, CL, GL, LPG Iwamori et al. (2011); Sauvageau et al. (2012
 Streptococcus pneumoniae PG, CL, GL Trombé, Lanéelle and Lanéelle (1979); Meiers et al. (2014
 Clostridium perfringens PG, PE, APG, LPG, PL Roy and Ibba (2008); Johnston, Baker and Goldfine (2004
 Mycoplasma genitalium PG, CL, GL, host-derived lipids Kornspan and Rottem (2012
Taxonomic group Species Major membrane lipids described/predicted References 
Proteobacteria   
Alpha   
 Agrobacterium tumefaciens PG, CL, PE, MMPE, DMPE, PC, OL, S2, GL, DGTS, LPG Geske et al. (2013); Vences-Guzmán et al. (2013); Aktas et al. (2014); Semeniuk et al. (2014); Wessel et al. (2006
 Sinorhizobium meliloti PG, CL, PE, MMPE, DMPE, PC, OL, DGTS, SQD de Rudder, López-Lara and Geiger (2000); de Rudder, Sohlenkamp and Geiger (1999); López-Lara, Sohlenkamp and Geiger (2003
 Rhizobium tropici PG, CL, PE, MMPE, DMPE, PC, OL, S2, P1, P2, DGTS, LPG Rojas-Jiménez et al. (2005); Sohlenkamp et al. (2007); Vences-Guzmán et al. (2011
 Brucella melitensis PG, CL, PE, PC, OL, LPG Palacios-Chaves et al. (2011); Bukata et al. (2008); Comerci et al. (2006); Conde-Alvarez et al. (2006
 Rhodobacter sphaeroides PG, CL, PE, PC, OL, DGTS, SQD, GL Catucci et al. (2004); Benning et al. (1993); Benning, Huang and Gage (1995); Zhang, Ferguson-Miller and Reid (2009
 Caulobacter crescentus PG, CL, LPG, acyl-PG, GL Contreras, Shapiro and Henry (1978); Jones and Smith (1979); De Siervo and Homola (1980
 Sphingomonas sp. PG, CL, PE, MMPE, DMPE, PC, SL, OL, LPG, DGTS, HOP Kawahara, Kuraishi and Zähringer (1999); Rowe et al. (2000); Huang et al. (2009); Zhang et al. (2005
 Methylobacterium extorquens PG, PE, PC, HOP Bradley et al. (2010); Ostafin et al. (1999); Welander et al. (2010
 Bradyrhizobium diazoefficiens PG, CL, PE, MMPE, DMPE, PC, Bravo et al. (2001); Hacker et al. (2008); Minder et al. (2001); 
 (formerly B. japonicum) HOP Perzl et al. (1997); Silipo et al. (2014
 Rhodopseudomonas palustris PG, CL, PE, PC, HOP Ramana et al. (2012); Welander et al. (2012
Beta   
 Burkholderia cenocepacia PG, CL, PE, PE-2OH, OL, P1, NL1, NL2, LPG, HOP González-Silva et al. (2011); Taylor, Anderson and Wilkinson (1998); Schmerk, Bernards and Valvano (2011); Schmerk et al. (2015
 Ralstonia solanacearum PG, PE, PE-2OH, lyso-PE, OL, LPG Yabuuchi et al. (1995
 Neisseria gonorrhoeae PG, CL, PE, lyso-PE Rahman et al. (2000); Senff et al. (1976); Sud and Feingold (1975
 Bordetella pertussis PG, CL, PE, OL, lyso-PE Kawai and Moribayashi (1982); Kawai, Moribayashi and Yano (1982); Thiele and Schwinn (1973
Gamma   
 Escherichia coli PG, CL, PE Parsons and Rock (2013); Dowhan (2013
 Pseudomonas aeruginosa PG, CL, PE, PC, OL, APG Lewenza et al. (2011); Wilderman et al. (2002); Klein et al. (2009); Pramanik et al. (1990
 Vibrio cholerae PG, CL, PE, OL Giles et al. (2011
 Salmonella typhimurium PG, CL, PE, acyl-PG Ames (1968); Olsen and Ballou (1971
 Serratia proteamaculans PG, CL, PE, OL, NL1 Vences-Guzmán et al. (2014
 Legionella pneumophila PG, CL, PE, MMPE, DMPE, PC Conover et al. (2008
 Xanthomonas campestris PG, CL, PE, MMPE, PC, LPG Moser et al. (2014a); Moser, Aktas and Narberhaus (2014b
 Acidithiobacillus ferroxidans PG, CL, PE, MMPE, DMPE, PC, OL, HOP Barridge and Shively (1968); Dees and Shively (1982); Jones et al. (2012
 Francisella tularensis PG, CL, PE, PC, lyso-PE Li, Wang and Ernst (2011
 Methylococcus capsulatus PG, PE, ST, HOP Lamb et al. (2007); Welander and Summons (2012); Fang, Barcelona and Semrau (2000
Delta   
 Desulfovibrio sp. PG, CL, PS, lyso-PS, PE, OL, DGTS, HOP Makula and Finnerty (1974); Makula and Finnerty (1975); Blumenberg et al. (2009
 Myxococcus xanthus PG, CL, PE, SL, PI, PL Lorenzen et al. (2014); Ring et al. (2009
 Stigmatella aurantiaca PG, PE, lyso-PE, PI, SL, ST, OL, LPG Ring et al. (2009); Caillon, Lubochinsky and Rigomier (1983); Bode et al. (2003
 Bdellovibrio bacteriovorus PG, CL, PS, PE, PT, acyl-PE, SL Müller et al. (2011); Nguyen, Sallans and Kaneshiro (2008); Steiner, Conti and Lester (1973
Epsilon   
 Helicobacter pylori PG, CL, PE, lyso-PE Zhou et al. (2012); Olofsson et al. (2010); Tannaes et al. (2000, 2005
 Campylobacter jejuni PG, PE Leach, Harvey and Wait (1997
Cyanobacteria   
 Synechococcus sp. PCC 7942 PG, SQD, GL, CL Güler et al. (1996
 Synechocystis sp. PCC 6803 PG, SQD, GL, HOP Yuzawa et al. (2014
Deinococcales   
 Deinococcus radiodurans GL, GPL Anderson and Hansen (1985); Huang and Anderson (1989, 1995
Chloroflexi (green    
non-sulfur bacteria) Chloroflexus aurantiacus PG, CL, SQD, GL Kenyon and Gray (1974); Mizoguchi et al. (2013
Aquificae   
 Aquifex aeolicus DEGP, AEGP, APT Jahnke et al. (2001); Sturt et al. (2004
Thermotogae   
 Thermotoga maritima DEGP, AEGP, GL Manca et al. (1992); Damsté et al. (2007); Huber et al. (1986
Fusobacteria   
 Fusobacterium mortiferum PG, CL, PE, PL Hagen (1974
Chlamydiae   
 Chlamydia pneumonia PE, PC, host-derived membrane lipids Yao et al. (2014); Hatch and McClarty (1998
CFB Group   
 Flavobacterium johnsoniae PE, PC, OL, P1, SOL, GCL Pitta, Leadbetter and Godchaux (1989); Kawazoe et al. (1991
 Bacteroidetes thetaiotamicron PG, CL, PS, PE, SL Smith and Salyers (1981
 Porphyromonas gingivalis PG, PE, SL Nichols and Rojanasomsith (2006); Nichols et al. (2004, 2006); Tavana et al. (2000
Chlorobium (green    
sulfur bacteria) Chlorobium tepidum PG, CL, PE, GL Mizoguchi et al. (2013); Sorensen, Cox and Miller (2008
Actinobacteria   
 Mycobacterium tuberculosis PG, CL, PE, PI, PIM, OL Khuller et al. (1982); Jackson, Crick and Brennan (2000); Berg et al. (2007); Lanéelle et al. (1990
 Nocardia sp. CL, PE, PI, PIM, OL, SQD, HOP Khuller and Brennan (1972); Trana, Khuller and Subrahmanyam (1980
 Streptomyces coelicolor PG, CL, PE, PI, PIM, OL, HOP Sandoval-Calderón et al. (2009); Jyothikumar et al. (2012); Liu et al. (2014
 Corynebacterium glutamicum PG, CL, PI, PIM Nampoothiri et al. (2002
Spirochetes   
 Borrelia burgdorferi PG, PC, GL Wang, Scagliotti and Hu (2004); Ostberg et al. (2007
 Treponema denticola PG, CL, PE, PC Kent et al. (2004
 Leptospira interrogans PG, PE, CL, Lyso-PE, GL, OL Livermore and Johnson (1974); Moribayashi et al. (1991
Planctomycetes   
 Singulisphaera acidiphila PG, PC, TMOL, PA, HOP Moore et al. (2013
 Rhodopirellula baltica PG, CL, PE, PC, DGTS Bondoso et al. (2014); Roh et al. (2013
Firmicutes   
 Bacillus subtilis PG, CL, PE, LPG, GL, GPL Gidden et al. (2009); den Kamp et al. (1969); Salzberg and Helmann (2008); Jorasch et al. (1998
 Listeria monocytogenes PG, CL, LPG, LCL, PI Fischer and Leopold (1999); Dare et al. (2014); Thedieck et al. (2006); Mastronicolis et al. (2008
 Staphylococcus aureus PG, CL, LPG, GPL Oku et al. (2004); White and Frerman (1967
 Lactobacillus sp. PG, CL, GL, LPG Iwamori et al. (2011); Sauvageau et al. (2012
 Streptococcus pneumoniae PG, CL, GL Trombé, Lanéelle and Lanéelle (1979); Meiers et al. (2014
 Clostridium perfringens PG, PE, APG, LPG, PL Roy and Ibba (2008); Johnston, Baker and Goldfine (2004
 Mycoplasma genitalium PG, CL, GL, host-derived lipids Kornspan and Rottem (2012

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 Cytophaga-Flavobacterium-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 phosphate concentrations (Benning, Huang and Gage 1995; Hölzl et al.2005; Hölzl and Dörmann 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-3-phospho)-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-Guzmán, Geiger and Sohlenkamp 2012). It is predicted to be present in many proteobacteria, bacteria belonging to the CFB group and actinomycetes. 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; Vences-Guzmán et al.2011, 2013). In a few bacteria, structural 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 (González-Silva et al.2011; Vences-Guzmán et al.2011, 2013; Vences-Guzmán, Geiger and 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 α-proteobacteria, 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 hydrophobic 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; López-Lara, Sohlenkamp and Geiger 2003; Huang et al.2009; Palacios-Chaves et al.2011; Vences-Guzmán et al.2011; Aktas et al.2014). Most β-proteobacteria 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, Anderson and Wilkinson 1998; Rahman et al.2000; González-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, Aktas and Narberhaus 2014b; Vences-Guzmán et al.2014). Among the δ-proteobacteria, the ability to form PG and PE seems to be 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, ε-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.

Cyanobacterial membranes mostly contain PG, SQD and the GLs monogalactosyl-DAG and digalactosyl-DAG (Güler 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; Damsté 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.2002; Sandoval-Calderón et al.2009; Seipke and Loria 2009).

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 lyso-PE. 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 Damsté 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. Recently, 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 branched-chain 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 Gram-negative 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 sn-2 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 (DX2DGX2ARX8GX3DX3D in most cases, DX2DGX2ARX12GX3DX3D in case of Pcs enzymes) (Williams and McMaster 1998; Sohlenkamp, López-Lara and Geiger 2003; Solís-Oviedo 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 HXKX4DX8G (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-Guzmán, 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 observed (Vences-Guzmán, Geiger and Sohlenkamp 2008). Recently, 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 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 host-symbiont/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 prokaryotic Cls, belonging to the PLD superfamily. Sandoval-Calderón 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 stress-responsive 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 ClsC-dependent 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 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 N-methylation pathway for PC synthesis was known in prokaryotes (Kaneshiro and Law 1964; Sohlenkamp, López-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, López-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).

Figure 4.

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

Figure 4.

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

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, López-Lara and Geiger 2003; Geiger, López-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, López-Lara and Sohlenkamp 2013).

Kent et al. (2004) showed evidence for the presence of a CDP-choline 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, López-Lara and Geiger 2003; Geiger, López-Lara 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 myo-inositol 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. Ac1PIM2 and Ac2PIM2 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.

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; Ac1PIM2-acyl PIM2; Ac2PIM2-diacyl PIM2; BEPIS-bacterial dialkylether glycerophosphoinositol synthase.

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; Ac1PIM2-acyl PIM2; Ac2PIM2-diacyl PIM2; BEPIS-bacterial dialkylether glycerophosphoinositol synthase.

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-inositol-1-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, López-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 GDP-mannose to DAG, leading to the formation of a monoglycosyl-DAG. 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 syntheses are referred to a review by Hölzl and Dörmann (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 NAD-dependent 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 (Güler, Essigmann 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; Güler et al.1996; Weissenmayer, 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, similarly to the Rhodobacter mutant it showed reduced growth under phosphate-limiting conditions (Güler 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 (López-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-Jiménez 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 required for OL synthesis (Vences-Guzmán et al.2014). OlsBA are present in α-, β- and a few γ-proteobacteria and actinomycetes, whereas OlsF is present in γ-, δ- and ε-proteobacteria, and bacteria of the CFB group (Vences-Guzmán et al.2014). Based on the distribution of OlsB and OlsF in bacterial genomes, it has been estimated that about 50% of the sequenced bacteria can form OL (Vences-Guzmán et al.2014). Unmodified OL can be modified by the introduction of one or more hydroxyl group or by methylation of the δ-amino group in the ornithine headgroup (Rojas-Jiménez et al.2005; González-Silva et al.2011; Vences-Guzmán et al.2011; Vences-Guzmán, Geiger and Sohlenkamp 2012; Moore 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-Jiménez et al.2005; Vences-Guzmán et al.2011). OlsD was described first in Burkholderia cenocepacia and introduces a hydroxyl group at the 2-position in the amide-bound fatty acid and gives rise to the formation of new lipids 1 and 2 (NL1, NL2) (Fig. 6, reaction 28) (González-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 29) (Vences-Guzmán et al.2011). The methyltransferase responsible 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 (Vences-Guzmán et al.2013). Rhizobium tropici mutants deficient in OlsC are defective in growth at high temperature and at acid pH (Vences-Guzmán et al.2011). Mutants deficient in OlsC and/or OlsE in R. tropici are also defective in nodulation assays using bean as a host (Vences-Guzmán et al.2011). Mutants deficient in OL formation in S. meliloti showed a slight growth defect under phosphate-limiting conditions when at the same time DGTS synthesis was interrupted (López-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.

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-Jiménez et al.2005). The formation of new lipids 1 and 2 (NL1, NL2) is due to the enzymatic activity of OlsD.

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-Jiménez et al.2005). The formation of new lipids 1 and 2 (NL1, NL2) is due to the enzymatic activity of OlsD.

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 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 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 (Pérez-Gil and Rodríguez-Concepción 2013). DMPP is elongated by head-to-tail condensation with two molecules IPP to farnesyl diphosphate. Two molecules farnesyl diphosphate are then condensated head-to-head 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-2,3-epoxide 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.

Figure 7.

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

Figure 7.

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

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 C11-unsaturated, 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 α-proteobacteria 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 (Zavaleta-Pastor 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 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-1-phosphate) (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-Jiménez et al.2005; Vences-Guzmán et al.2011) (Fig. 6, reaction 27). An 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 a pH lower than 4.5 (Vences-Guzmán et al.2011).

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, Anderson and Wilkinson 1998; Vences-Guzmán et al.2011).

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 CL-enriched 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 growth-limiting 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 drastically (Vences-Guzmán et al.2013).

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?

  • 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).

  • 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 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.

  • 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 Tecnología-México (CONACyT-México; 153200, 178359) and Dirección General de Asuntos del Personal Académico-Universidad Nacional Autónoma de México (DGAPA-UNAM; PAPIIT IN202413, IN203612).

Conflict of interest. None declared.

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