bacterial membrane lipids: diversity in structures and ... ·...

FEMS Microbiology Reviews, fuv008, 40, 2016, 133–159

doi: 10.1093/femsre/fuv008Advance Access Publication Date: 9 April 2015Review Article

REVIEW ARTICLE

Bacterial membrane lipids: diversity in structuresand pathwaysChristian Sohlenkamp∗ and Otto Geiger

Centro de Ciencias Genomicas, Universidad Nacional Autonoma de Mexico, Av. Universidad s/n, Apdo.Postal 565-A, Cuernavaca, Morelos, CP 62210, Mexico∗Corresponding author: Centro de Ciencias Genomicas, Universidad Nacional Autonoma de Mexico, Av. Universidad s/n, Apdo. Postal 565-A,Cuernavaca, Morelos, CP 62210, Mexico. Tel: (+52) 777-3291703; Fax: (+52) 777-3175581; E-mail: [email protected] sentence summary: This review gives an overview about the diversity and distribution of bacterial membrane lipids and the metabolic pathwaysinvolved in their synthesis.Editor: Franz Narberhaus

ABSTRACT

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

Keywords: ornithine lipid; phosphatidylcholine; cardiolipin; phosphatidylinositol; phospholipid; hopanoids;phosphatidylethanolamine

INTRODUCTION

This review tries to open up the world of microbial membranelipids to the reader. We want to give an overview about the in-ventory of membrane lipids present in bacteria belonging tothe different phyla studied. Also, we want to summarize theknown synthesis pathways for the major membrane-forminglipids and discuss how widely these pathways are distributed.However, the present review is not meant to be a complete cata-log of all different unique structures that have been reported inthe literature. Readers looking for this information are referredto 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 lipidspresent in bacterial membranes, leaving membrane lipids fromother microorganisms aside.

Classically, the technique of Gram staining allowed the clas-sification of bacteria as either Gram-positive or Gram-negativebacteria, a differentiation based on their cell wall properties.Gram-positive bacteria such as the firmicutes are character-ized by having a cytoplasmatic membrane and a thick mureincell wall comprising many layers, whereas Gram-negative bac-teria such as proteobacteria are characterized by the presenceof two distinct membranes, called inner and outer membrane(Raetz and Whitfield 2002; Reichmann and Grundling 2011) anda thin murein cell wall between them. Usually, the outer leaflet

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

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134 FEMS Microbiology Reviews, 2016, Vol. 40, No. 1

of the outer membrane is formed by lipid A, the lipophilic an-chor of lipopolysaccharide (LPS) (Raetz and Dowhan 1990; Raetzet al. 2007). Mycobacteria are characterized by the presence of athick lipophilic layer containing mycolic acids on top of the cellwall, that is sometimes called mycobacterial outer membrane(Niederweis et al. 2010). The two latter structures will not be dis-cussed here.

Membranes are formed by amphiphilic lipids which inmost cases and conditions studied are glycerophospho-lipids, composed of two fatty acids, a glycerol moiety, aphosphate group and a variable head group. Examples arephosphatidylethanolamine (PE), phosphatidylglycerol (PG),cardiolipin (CL), lysyl-phosphatidylglycerol (LPG), phosphatidyli-nositol (PI), phosphatidic acid (PA) and phosphatidylserine (PS).Bacteria can also form phosphorus-free membrane lipids suchas ornithine lipids (OLs), sulfolipids, diacylglyceryl-N,N,N-trimethylhomoserine (DGTS), glycolipids (GLs), diacylglycerol(DAG), hopanoids (HOPs) and others. Since the discovery ofthe first pathways for membrane lipid synthesis, a surprisingdiversity and complexity of enzymes and pathways involvedhas emerged. Althoughmicroorganisms are sometimes thoughtof as more limited in the range and types of lipids which theypossess in comparison to plant or animal cells, this view iswholly incorrect as we hope to convince the reader. In thisreview, we will describe the complexity of pathways for thebiosynthesis of phospholipids and phosphorus-free membranelipids and discuss how both types of lipids are metabolicallyconnected.

DISTRIBUTION OF MEMBRANE LIPIDS IN BACTERIA

Membrane lipids in Escherichia coli

As in other areas of microbiology, E. coli has been the stan-dard model also in the study of bacterial membrane lipids formany decades. Escherichia coli accumulates three major mem-brane phospholipids: its predominant lipid is the zwitterionicPE (about 75% of membrane lipids), and additionally it forms theanionic lipids PG (about 20%) and CL (Raetz and Dowhan 1990).Metabolic pathways and lipids known to be present in E. colihavebeen highlighted in red in Fig. 1. In E. coli, two pathways startfrom the central metabolite cytidine diphosphate-diacylglycerol(CDP-DAG): PE is synthesized in two steps by phosphatidylser-ine synthase (Pss) (Figs 1 and 2, reaction 1) and phosphatidylser-ine decarboxylase (Psd) (Figs 1 and 2, reaction 2). Phosphatidyl-glycerolphosphate synthase (PgsA) catalyzes the condensationof glycerol-3-phosphate (G3P) with CDP-DAG leading to the for-mation of PG phosphate (PGP), which is the first step in the syn-thesis of the anionic membrane lipids PG and CL (Figs 1 and3, reaction 4). PGP is dephosphorylated in E. coli by any one ofthree phosphatases (PgpA, PgpB, PgpC) (Figs 1 and 3, reaction5) to form PG (Lu et al. 2011). The CL synthase ClsA can thensynthesize CL from two molecules of PG (Figs 1 and 3, reac-tion 6). Escherichia coli has two more CL synthase (Cls) calledClsB and ClsC. So far ClsB only has been shown to work in invitro assays (Guo and Tropp 2000). ClsC has only been recentlyrecognized as a Cls forming CL from PG and PE (Figs 1 and 3,reaction 7) (Tan et al. 2012). Most of these genes and enzymeswere discovered and characterized already decades ago and thegeneral feeling was that the described set of genes/enzymeswas complete. It is clear now that E. coli presents a relativelysimple membrane composition not representative for the di-versity in membrane lipids present in many other bacteria(Table 1).

Diversity of membrane lipids in bacteria

In addition to (and sometimes instead of) the E. coli lipids PE, PGand CL, other bacteria can form a variety of lipids. Many syn-thesize the methylated PE derivatives monomethyl PE (MMPE),dimethyl PE (DMPE) and phosphatidylcholine (PC). In most bac-teria, MMPE is simply an intermediate in the formation of PC,but in a few species such as Clostridium butyricum, Proteus vul-garis, Yersinia pestis and Xanthomonas campestris, MMPE is the endproduct of the pathway (Goldfine and Ellis 1964; Tornabene 1973;Moser, Aktas and Narberhaus 2014b).

The capacity to form PI, another membrane lipid consid-ered typical for eukaryotes, is widespread in actinomycetes,and several δ-proteobacteria also accumulate it. At least in My-cobacterium tuberculosis, PI (and/or its derivatives) is an essen-tial lipid (Jackson, Crick and Brennan 2000). PI is often furthermodified with mannose residues and fatty acid residues. Thesesimple acyl-phosphatidylinositol mannosides (AcPIMs) can befurther elongated to form higher PIM species and the lipogly-cans lipomannan and lipoarabinomannan which are impor-tant molecules implicated in host–pathogen interactions in thecourse of tuberculosis and leprosy (Berg et al. 2007; Kaur et al.2009; Guerin et al. 2010). Dialkylether-glycero-3-phosphoinositolhas been described recently in the thermophilic Cytophaga-Flavobacterium-Bacteroidetes (CFB) group bacterium Rhodother-mus marinus. This lipid might be also present in Sphingobacteriaand in several α-proteobacteria (Jorge, Borges and Santos 2014).

A few bacteria also accumulate the anionic lipid PS which isan intermediate in the synthesis of PE. Normally Psd is very effi-cient in converting PS into PE, leaving hardly detectable amountsof PS. The δ-proteobacterium Bdellovibrio bacteriovorans is one ex-ample where significant amounts of PS were detected (Nguyen,Sallans and Kaneshiro 2008). PS is also a major lipid in C. bo-tulinum and a few Flavobacterium species (Evans et al. 1998; Lata,Lal and Lal 2012).

The headgroup of PG can be modified by transfer of aminoacids and in most cases the transferred amino acids are ly-sine, alanine or arginine thereby forming LPG, alanyl-PG (APG) orarginyl-PG (ArPG), respectively (Roy, Dare and Ibba 2009; Roy andIbba 2009). Originally, these aminoacyl modifications of PG hadbeen described only in Gram-positive bacteria such as Staphy-lococcus aureus, Bacillus subtilis, Lactococcus plantarum and Listeriamonocytogenes (den Kamp, Redai and van Deenen 1969; Nahaieet al. 1984; Fischer and Leopold 1999), but recently aminoacylatedPG derivativeswere also identified in a fewGram-negative bacte-ria, 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 (Kobayashiet al. 1980). The enzyme responsible for this acyltransferase re-action is not known. Aminoacylated versions of CL with alanineor lysine are formed in different Listeria species (Peter-Katalinicand 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. Severalof them present a DAG backbone, such as glycosylated DAGs,DGTS and the sulfolipid sulfoquinovosyl diacylglycerol (SQD).The presence of glycosylated DAG derivatives (GLs) has beenknown for many years in Gram-positive bacteria, and glycosyl-transferases involved in the synthesis of these lipids were iden-tified (Jorasch et al. 1998). In recent years, the presence of GLshas been described in a few α-proteobacteria such as Mesorhizo-bium loti, Agrobacterium tumefaciens and Rhodobacter sphaeroideswhen these bacteria are grown in media with limiting

Sohlenkamp and Geiger 135

Figure 1. Synthesis of membrane lipids in bacteria. This figure gives an overview about metabolic routes described in the context of membrane lipid formationin bacteria. The reactions catalyzed by the different enzymes are explained in detail in the text. Abbreviations: G3P-glycerol-3-phosphate; LPA-lysophosphatidicacid; 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.

phosphate concentrations (Benning, Huang andGage 1995; Holzlet al. 2005; Holzl and Dormann 2007; Devers et al. 2011; Geskeet al. 2013; Diercks et al. 2015; Semeniuk et al. 2014). There is alarge structural diversity within sugar headgroups with respectto sugar monomers, anomeric forms, linkage between sugarsand covalentmodification of sugars, whichwill not be discussedwithin the present review. Glycophospholipids (GPLs) form aspecial class of DAG-derived GLs. They have been describedmainly in a few Gram-positive bacteria. One important exam-ple is the radiation-resistant Deinococcus radiodurans, where oneof the major membrane lipids is 2′-O-(1,2-diacyl-sn-glycero-3-phospho)-3′-O-(α-galactosyl)-N-D-glyceroyl alkylamine (Ander-son and Hansen 1985).

DGTS is a betaine lipid that occurs in a variety of lowergreen plants, chromophytes, fungi and amoebae. Often there is

reciprocity between PC and DGTS content, meaning that inmostcases when PC is a major lipid no DGTS is detected and viceversa. Only some algae seem to be the exception to the rule andaccumulate PC and DGTS at the same time (Kato et al. 1996). Theoccurrence of DGTS in bacteria is limited. The presence of DGTShas been shown in some α-proteobacteria and DGTS only accu-mulates in these bacteria under conditions of phosphate limi-tation replacing PC (Benning, Huang and Gage 1995; Geiger et al.1999).

Other membrane lipids present in bacteria such as OLs, sul-fonolipids, HOPs or sphingolipids (SLs) do not present a DAGbackbone. OL has not been found in eukaryotes or archaea(Geiger et al. 2010; Vences-Guzman, Geiger and Sohlenkamp2012). It is predicted to be present in many proteobacte-ria, bacteria belonging to the CFB group and actinomycetes.


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-phosphatidylserinedecarboxylase; CL/PE synthase-cardiolipin/phosphatidylethanolamine synthase; CMP-cytidine monophosphate; CDP-cytidine diphosphate.

Interestingly, while in some bacteria OLs are only formed underconditions of phosphate limitation, in other (and often closelyrelated) bacteria OLs are formed constitutively (Geiger et al. 1999;Vences-Guzman et al. 2011, 2013). In a few bacteria, structuralanalogs 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; Ba-trakov et al. 1999; Zhang, Ferguson-Miller and Reid 2009). In re-cent years, several structurally distinct OLs either hydroxylatedor methylated have been described (Gonzalez-Silva et al. 2011;Vences-Guzman et al. 2011, 2013; Vences-Guzman, Geiger andSohlenkamp 2012; Moore et al. 2013; Diercks et al. 2015).

HOPs are pentacyclic triterpenoid lipids that are present ina wide variety of organisms. A large diversity of HOP struc-tures mainly differing in their side chain structures has beendescribed. It is unclear what the functional importance of thisstructural diversity is. Due to their structural similarities, HOPshave been consideredmolecular surrogates of eukaryotic sterols(Saenz et al. 2012a). Consistent with this, it has been observedthat HOPs are involved in enhancing the stability and im-permeability of the bacterial membrane. They are present incyanobacteria, methylotrophic bacteria, actinomycetes, some α-proteobacteria, β-proteobacteria and δ-proteobacteria (Kannen-berg and Poralla 1999; Blumenberg et al. 2009; Seipke and Loria2009; 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 specificHOPs structures arewidely used asmarkers in organic geochem-istry (Welander and Summons 2012; Ricci et al. 2014).

SLs are restricted to a few groups of bacteria: bacteria of theorder Sphingomonadales, several bacteria belonging to the CFBgroup and a few δ-proteobacteria such asMyxococcus xanthus, M.stipitatus, Cystobacter fuscus and Sorangium cellulosum (Eckau, Dilland Budzikiewicz 1984; Stein and Budzikiewicz 1988; Keck et al.2011). Awide variety of SL headgroups has been described, someare sugars whereas others are phosphoethanolamine or inositolderived.

Structural diversity can also be detected in the hydropho-bic moieties of the membrane lipids. Commonly, the hydropho-

bic moieties of amphiphilic membrane lipids are formed by lin-ear fatty acids that can be saturated or unsaturated (contain-ing often one and rarely two or more double bonds). Especiallymany firmicutes, actinomycetes and bacteria of the CFB groupoften contain branched-chain fatty acids, especially of the isoand anteiso types. In many anaerobic bacteria, plasmalogens,1-O-alk-1′-enyl 2-acyl glycerol phospholipids and GLs arepresent (Goldfine 2010). In the deep-branching hyperther-mophilic bacteria Thermotoga and Aquifex, also alkyl residues in-stead of acyl residues can be found. They present acyletherglyc-erol 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 tomake a few generaliza-tions with respect to the membrane lipids present in differentbacterial phyla (Table 1). Almost all α-proteobacteria seem to beable to form PG, CL and PE andmany can also synthesize PC andOLs. Probably as a consequence of their lifestyles, a few can formSQD, DGTS, GLs, LPG, HOPs and even SLs (Benning, Huang andGage 1995; Perzl et al. 1997; Lopez-Lara, Sohlenkamp and Geiger2003; Huang et al. 2009; Palacios-Chaves et al. 2011; Vences-Guzman et al. 2011; Aktas et al. 2014). Most β-proteobacteriacan form PE, PG, CL and OLs, while a few probably can synthe-size the cationic lipid LPG. Lyso-PE, a deacylated version of PEhas also been identified in several β-proteobacteria. Widespreadamong bacteria of the family Burkholderiaceae is the capacity toform PE and/or OLs hydroxylated in the 2-position of a fatty acid(Thiele and Schwinn 1973; Yabuuchi et al. 1995; Taylor, Ander-son andWilkinson 1998; Rahman et al. 2000; Gonzalez-Silva et al.2011). All bacteria belonging to the genus Burkholderia seem to beable to form HOPs (Schmerk, Bernards and Valvano 2011; Mal-ott et al. 2012). Most γ -proteobacteria accumulate PG, CL andPE, while a few can form OLs, PC, HOPs, LPG or APG (Neunlistand Rohmer 1985; Raetz and Dowhan 1990; Conover et al. 2008;Klein et al. 2009; Giles et al. 2011; Lewenza et al. 2011; Moser, Ak-tas and Narberhaus 2014b; Vences-Guzman et al. 2014). Amongthe δ-proteobacteria, the ability to form PG and PE seems to be


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 serveas a precursor for one of the three described Cls pathways or alternatively PG or CL can be (amino-) acylated. CMP-cytidine monophosphate; CDP-cytidine diphos-phate; 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 abun-dance of lipids. PG-phosphatidylglycerol; CL-cardiolipin; PS-phosphatidylserine; PE-phosphatidylethanolamine; PE-2OH-PE hydroxylated inthe 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 ornithineheadgroup and in 2-position of ester-bound fatty acid; NL1-OL hydroxylated in the 2-position of the amide-bound fatty acid; NL2-OL hydrox-ylated 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 onits genome sequence.

Taxonomic group SpeciesMajor membrane lipidsdescribed/predicted References

ProteobacteriaAlpha

Agrobacterium tumefaciens PG, CL, PE, MMPE, DMPE, PC,OL, S2, GL, DGTS, LPG

Geske et al. (2013); Vences-Guzman 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, Lopez-Lara and Geiger (2000); de Rudder,Sohlenkamp and Geiger (1999); Lopez-Lara, Sohlenkamp andGeiger (2003)

Rhizobium tropici PG, CL, PE, MMPE, DMPE, PC,OL, S2, P1, P2, DGTS, LPG

Rojas-Jimenez et al. (2005); Sohlenkamp et al. (2007);Vences-Guzman et al. (2011)

Brucella melitensis PG, CL, PE, PC, OL, LPG Palacios-Chaves et al. (2011); Bukata et al. (2008); Comerciet 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 andGage (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 Zahringer (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)

BetaBurkholderia cenocepacia PG, CL, PE, PE-2OH, OL, P1,

NL1, NL2, LPG, HOPGonzalez-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)

GammaEscherichia 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-Guzman 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, HOPBarridge and Shively (1968); Dees and Shively (1982); Joneset 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,

LPGRing et al. (2009); Caillon, Lubochinsky and Rigomier (1983);Bode et al. (2003)

Bdellovibrio bacteriovorus PG, CL, PS, PE, PT, acyl-PE, SL Muller et al. (2011); Nguyen, Sallans and Kaneshiro (2008);Steiner, Conti and Lester (1973)


Table 1 (Continued.)

Taxonomic group SpeciesMajor membrane lipidsdescribed/predicted References

EpsilonHelicobacter 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)

CyanobacteriaSynechococcus sp. PCC 7942 PG, SQD, GL, CL Guler et al. (1996)Synechocystis sp. PCC 6803 PG, SQD, GL, HOP Yuzawa et al. (2014)

DeinococcalesDeinococcus radiodurans GL, GPL Anderson and Hansen (1985); Huang and Anderson (1989,

1995)Chloroflexi (greennon-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); Damste et al. (2007); Huber et al. (1986)Fusobacteria

Fusobacterium mortiferum PG, CL, PE, PL Hagen (1974)Chlamydiae

Chlamydia pneumonia PE, PC, host-derivedmembrane lipids

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

CFB GroupFlavobacterium 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 (greensulfur 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); Berget al. (2007); Laneelle et al. (1990)

Nocardia sp. CL, PE, PI, PIM, OL, SQD, HOP Khuller and Brennan (1972); Trana, Khuller andSubrahmanyam (1980)

Streptomyces coelicolor PG, CL, PE, PI, PIM, OL, HOP Sandoval-Calderon et al. (2009); Jyothikumar et al. (2012); Liuet 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)

PlanctomycetesSingulisphaera 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)

FirmicutesBacillus 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 Trombe, Laneelle and Laneelle (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

lipidsKornspan and Rottem (2012)

conserved and a variety of uncommon lipids can be detected insome cases, such as SLs, PI, HOPs or even sterols. Also, a fewbacteria can form OLs, LPG or DGTS and even phosphatidylthre-onine (PT) has been described in B. bacteriovorus (Makula andFinnerty 1974; Nguyen, Sallans and Kaneshiro 2008; Blumenberget al. 2009; Ring et al. 2009; Keck et al. 2011; Lorenzen et al. 2014).

In general, ε-proteobacteria have a far less complex lipid compo-sition with PE, PG and often CL present (Leach, Harvey and Wait1997; Tannaes, Grav and Bukholm 2000; Tannaes, Bukholm andBukholm 2005; Zhou et al. 2012). The fatty acids present in com-plex 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 andthe GLs monogalactosyl-DAG and digalactosyl-DAG (Guler et al.1996). The presence of CL has not been reported, but severalgenomes encode a putative Cls of the PLD superfamily. HOPs arepresent in several species (Doughty et al. 2009; Saenz et al. 2012b).

The membrane of the radiation-resistant bacterium D. radio-durans has an unusual lipid composition, because a diversity ofGLs and GPLs has been described, whereas the synthesis path-ways of the common phospholipids PG, PE and CL seem to beabsent (Huang and Anderson 1989; Huang and Anderson 1995;Makarova et al. 2001).

In the membranes of the green non-sulfur bacterium Chlo-roflexus aurantiacus, the lipids PG, CL, SQD and several GLs havebeen detected (Kenyon and Gray 1974; Mizoguchi et al. 2013).

The deep-branching hyperthermophiles Aquifex aeolicus andThermotogamaritima present unusual structureswhen comparedto the better studied bacteria. Their membrane lipids are notexclusively DAG based but an important fraction is formed byDEGP and AEGP. Although the presence of ether lipids is con-sidered a typical trait of archaebacteria, Aquifex and Thermo-toga clearly present bacterial-type lipids. The fatty acyl and alkylgroup are in the sn-1- and 2-positions like in bacterial lipids andnot in the sn-2- and 3-positions as in archaea. Little is knownabout their lipid headgroups, but in Aquifex phosphoaminopen-tanetetrol headgroups and in Thermotoga sugar headgroups havebeen described (Manca et al. 1992; Sturt et al. 2004; Damste et al.2007).

Among the bacteria of the CFB group PE, PG, CL and OLsagain seem to be relatively common, but also unusual lipidscan be found: several species can form SLs or sulfonolipids andFlavobacterium/Cytophaga can form glycine-containing and/orserineglycine-containing lipids (Tahara et al. 1976; Kawai, YanoandKaneda 1988; Pitta, Leadbetter and Godchaux 1989; Kawazoeet al. 1991). Dialkylether-glycero-3-phosphoinositol has been de-scribed recently in the thermophilic bacterium R. marinus andmight be also present in Sphingobacteria (Jorge, Borges and San-tos 2014).

The green sulfur bacterium Chlorobium tepidum accumulatesPG, CL, PE and GLs, but apparently does not form SQD, which isin general omnipresent in photosynthetic organisms (Sorensen,Cox and Miller 2008; Mizoguchi et al. 2013).

Within the actinomycetes, many bacteria can form PG, PEand CL and in addition the capacity to synthesize PI and itsmannosylated/acylated derivatives is widespread. Some actino-mycetes are known to form OLs and several more are predictedto be able to formOLs. All species belonging to the genus Strepto-myces seem to be able to formHOPs. Often the fatty acids presentin the complex lipids are branched-chain fatty acids (Khuller andBrennan 1972; Jackson, Crick and Brennan 2000; Nampoothiriet al. 2002; Sandoval-Calderon et al. 2009; Seipke and Loria 2009).

Among the spirochaetes, the distinct genera present differ-ent lipid compositions: Borrelia forms PG, PC and GLs, Treponemaforms PG, PE, CL and PC, and Leptospira forms PE, PG, CL and lyso-PE. Bacteria of the genus Leptospira have also been described toform GLs and are predicted to be able to form OLs (Livermoreand 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 spe-cial type of cell division and intracellular membrane structures(Fuerst and Sagulenko 2011). Within this group are the am-monox bacteria, with their ladderane lipids instead of normalfatty acids (Sinninghe Damste et al. 2005; Boumann et al. 2006).The lipid composition within each genus seems to differ andsome Planctomycetes can form PG, PE, PC, CL and HOPs. Re-

cently, aN,N,N-trimethyl OL has been described in Singulisphaeraacidiphila (Moore et al. 2013).

Among the firmicutes the presence of PE is not universal, butmost seem to be able to form PG and CL. Inmany cases, the pres-ence of aminoacylated PG andCL derivatives has been described.Another feature that is widely distributed is the presence ofGLs and GPLs (White and Frerman 1967; den Kamp, Redai andvan Deenen 1969; Jorasch et al. 1998; Fischer and Leopold 1999;Johnston, Baker and Goldfine 2004; Sauvageau et al. 2012; Meierset al. 2014). Most of the firmicutes studied present branched-chain fatty acids (iso-/anteiso-) in their lipid structure, the ex-ception being the lactobacteria which form linear fatty acids.The latter present straight chain saturated, monounsaturatedand related cyclopropane fatty acids commonly found in Gram-negative bacteria (Ratledge and Wilkinson 1988).

A special group of bacteria is formed by some obligate intra-cellular pathogens that lack the capacity to synthesize all theirmembrane lipids and that obtain a part or all of their lipidsfrom the eukaryotic host. This has been described for Chlamy-dia and forMycoplasma, among others (Hatch andMcClarty 1998;Kornspan and Rottem 2012; Yao et al. 2014).

PATHWAYS FOR PHOSPHOLIPID SYNTHESISIN BACTERIASynthesis 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 beendescribed, the PlsB/PlsC and the PlsX/PlsY/PlsC systems. ThePlsB/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 otherbacteria use the PlsX/PlsY/PlsC system for PA synthesis (Luet al. 2006). CDP-DAG synthase then catalyzes the formationof CDP-DAG from CTP and PA. CDP-DAG is a common pre-cursor used in the synthesis of glycerophospholipids in bacte-ria. The steps from G3P to CDP-DAG are discussed in detail ina recent review by Parsons and Rock (2013). Most of the en-zymes described that use CDP-DAG as substrate belong to ei-ther the CDP-alcohol phosphotransferase family or the PLD su-perfamily. CDP-DAG alcohol phosphotransferases share a com-mon conserved motif (DX2DGX2ARX8GX3DX3D in most cases,DX2DGX2ARX12GX3DX3D in case of Pcs enzymes) (Williams andMcMaster 1998; Sohlenkamp, Lopez-Lara and Geiger 2003; Solıs-Oviedo et al. 2012). Members of this superfamily include phos-phatidylinositolphosphate synthase (Pips), phosphatidylcholinesynthase (Pcs), type II Pss, eukaryote-type cardiolipin synthase(Cls-II) and PgsA. Members of the PLD superfamily present theconserved motif HXKX4DX8G (Liscovitch et al. 2000). This super-family includes for example prokaryote-like Cls, type I Pss andthe CL/PE synthase.

Pathways for PE synthesis

PE is a common membrane lipid in proteobacteria and otherGram-negative bacteria. Its synthesis has been first describedin E. coli (DeChavigny, Heacock and Dowhan 1991). Pss con-denses serine with CDP-DAG leading to the formation of theanionic lipid PS. PS is then decarboxylated by Psd leading tothe formation of the zwitterionic lipid PE (Figs 1 and 2, reac-tions 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 andGeiger 2004). For example, Pss fromE. coli is a type I Pss whereas Pss from B. subtilis and Sinorhizo-bium meliloti are type II Pss (Okada et al. 1994; Matsumoto 1997;Sohlenkamp, de Rudder and Geiger 2004). Despite the fact thatboth types of Pss are unrelated on a sequence level, they bothuse CDP-DAG and L-serine as substrates and form PS and cyti-dine monophosphate (CMP) as products. Type I Pss is restrictedto a few γ -proteobacteria (Enterobacteriaceae, Vibrionaceae, Pas-teurellaceae, Alteromonaceae), whereas type II Pss occurs in allthree kingdoms of life (Sohlenkamp, de Rudder and Geiger 2004).Mutants deficient in pss or psd have been described in severalorganisms, such as E. coli, S. meliloti, A. tumefaciens, Helicobac-ter pylori, etc. (Li and Dowhan 1988; DeChavigny, Heacock andDowhan 1991; Shi et al. 1993; Ge andTaylor 1997;Matsumoto et al.1998; Karnezis et al. 2002; Sohlenkamp, de Rudder and Geiger2004; Vences-Guzman, Geiger and Sohlenkamp 2008). In gen-eral, these mutants are viable under specific growth conditionsand many share a requirement for millimolar bivalent cationsin the growth medium. Psd mutants accumulate PS and in thecase of the S. meliloti Psd mutant, a nodulation phenotype wasobserved (Vences-Guzman, Geiger and Sohlenkamp 2008). Re-cently, Moser et al. (2014a) described a bifunctional CL/PE syn-thase in the plant pathogen X. campestris that catalyzes thesynthesis 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 sameenzyme also functions as a Cls, and as many bacterial Cls, be-longs to the PLD superfamily. In eukaryotes, PE synthesis viathe CDP-ethanolamine pathway has been described, but this re-action has not been shown to be present in prokaryotes so far(Nakashima, Hosaka and Nikawa 1997).

Pathways for synthesis of PG and aminoacylated PGderivatives

PgsA is condensing G3P with CDP-DAG leading to the forma-tion of PGP (Figs 1 and 3, reaction 4) (Miyazaki et al. 1985).All PgsA described belong to the CDP-alcohol phosphotrans-ferase family. Mutants deficient in PgsA have been isolated inE. 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 almostall bacterial genomes, one notable exception being D. radiodu-rans (Makarova et al. 2001). PGP is then dephosphorylated by aPGP phosphatase (Figs 1 and 3, reaction 5) (Lu et al. 2011). Theanionic lipid PG can be converted into the cationic lipid LPG ina one-step reaction (Figs 1 and 3, reaction 9). Lysine is trans-ferred from a charged tRNALys to the glycerol headgroup of PG.This biochemical activity was first described in cell-free S. au-reus extracts (Gould and Lennarz 1967; Lennarz, Bonsen and vanDeenen 1967; Nesbitt and Lennarz 1968). The gene encoding theenzyme MprF responsible for LPG formation has first been de-scribed also in S. aureus during a screen for mutants more sus-ceptible than the wildtype to cationic antimicrobial peptides(Peschel et al. 2001). MprF homologs are also present in someGram-negative bacteria and it could be shown that these ho-mologs indeed caused the formation of LPG or APG, respectively(Sohlenkamp et al. 2007; Klein et al. 2009). Homologs of MprFare also responsible for APG formation in C. perfringens and P.aeruginosa and for ArPG formation in Enterococcus faecium (Royand Ibba 2008; Klein et al. 2009; Roy and Ibba 2009). Most MprFhomologs characterized only produce either APG or LPG, butMprF from E. faecium can catalyze the formation of APG, LPG

and ArPG (Roy and Ibba 2009). Further homologs are presentin many α-proteobacteria (A. tumefaciens, several species of thegenera Brucella, Rhizobium), β-proteobacteria (Burkholderia, Ralsto-nia) and γ -proteobacteria (X. campestris, Xylella, several speciesof Pseudomonas and Aeromonas). It is noteworthy that many ifnot most of the organisms predicted to form aminoacyl PGs un-der certain conditions can adopt a lifestyle associated with eu-karyotic 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 cationicantimicrobial peptides, and to various antibiotics such as dapto-mycin, 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 toa set of cationic antimicrobial peptides, to a few antibiotics andchromium ions (Klein et al. 2009; Arendt et al. 2012). Recently,Dare et al. (2014) showed evidence in L. monocytogenes that thefunction of LPG extends beyond contributing to the resistance tocationic peptides. LPG protects against osmotic stress and reg-ulates a motility operon and it is hypothesized that MprF-likeenzymes are involved in the adaptation of bacteria to a broadrange of environmental stresses (Dare et al. 2014). In bacteriaoutside the firmicutes, the genes encoding MprF/LpiA form anoperon with a gene encoding a putative lipase called AcvB/AtvA(Vinuesa et al. 2003; Sohlenkamp et al. 2007). In P. aeruginosa, theAcvB/AtvA homolog PA0919 seems to be responsible for hydrol-ysis of APG (Arendt et al. 2013).

Pathways for synthesis of CL and aminoacylated CLderivatives

For many years, two different pathways for CL synthesis wereknown: (A) the prokaryote-like pathway in which twomoleculesPG react forming CL and glycerol (Figs 1 and 3, reaction 6) and(B) the eukaryote-like pathway in which one molecule PG reactswith CDP-DAG leading to the formation of CL and CMP. As thenames implied, one route was thought to be specific for prokary-otes whereas the other was thought to be specific for eukary-otes. In E. coli, CL is synthesized by ClsA from two molecules ofPG (Nishijima et al. 1988). ClsA from E. coli is a typical prokary-otic Cls, belonging to the PLD superfamily. Sandoval-Calderonet al. (2009) showed that Streptomyces coelicolor and other acti-nomycetes use the eukaryote-like pathway to synthesize CL(Figs 1 and 3, reaction 8). Cls from actinomycetes belongs tothe CDP-alcohol phosphotransferase superfamily. The so-calledprokaryote-like pathway for CL synthesis is not restricted tobacteria, as Trypanosoma cruzei uses a prokaryote-like Cls (be-longing to the PLD superfamily) for CL synthesis (Serricchio andButikofer 2012). Many bacterial genomes present several genesencoding putative Cls; for example, E. coli presents three (clsA,clsB, clsC). Tan et al. (2012) showed that E. coli can synthesizeCL from PG and PE by ClsC (Figs 1 and 3, reaction 7). Interest-ingly, for full Cls activity of ClsC, a second gene called ymdBhas 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 ac-tivity is not clear, but YmdB might involve a stress responseactivation of ClsC. It is unknown how widespread this ClsC-dependent pathway is. Moser et al. (2014a) recently discovered inX. campestris a bifunctional CL/PE synthase belonging to the PLDsuperfamily that synthesizes CL from PG and CDP-DAG, but thatalso catalyzed ethanolamine-dependent PE formation. Interest-ingly, this bifunctional enzyme is larger (70 kDa) than the ear-lier characterized monofunctional Cls enzymes from E. coli and


Arabidopsis thaliana. The gene encoding the bifunctional CL/PEsynthase seems to be restricted to the orders Xanthomonadalesand Pseudomonadales (Moser et al. 2014a). These new activitiesdescribed for two different enzymes of the PLD superfamily(ClsC and CL/PE synthase) imply that maybe other membersof this family present different activities that have not beenfound because people haven’t been looking for them. Bacterialgenomes containmany genes encoding Cls and other PLD super-family members that have not been characterized yet. It is pos-sible that Cls already characterized years ago have additional ac-tivities and this possibility seems worth exploring. The aminoa-cylated derivatives of CL alanyl-CL (ACL) and lysyl-CL (LCL) havebeen described in different Listeria species. An aminoacyl-PGsynthase is required for their formation and if LPG formationis impaired in Listeria, also LCL formation is impaired, mean-ing that there is no dedicated second enzyme required for theaminoacyl transfer to CL (Dare et al. 2014). It is not clear if theamino acid (lysine or alanine) is transferred to CL or if a Cls canuse the aminoacylated PG as substrate and convert it into LCL orACL (Figs 1 and 3, reactions 11a and b).

Pathways for PC synthesis

In eukaryotes, two different pathways for PC synthesis havebeen described, the CDP-choline (or Kennedy) pathway and theN-methylation pathway. In the CDP-choline pathway, choline isphosphorylated by choline kinase and adenosine triphosphate.In a second step, phosphocholine and CTP react to CDP-cholinein a reaction catalyzed by CTP: phosphocholine cytidylyltrans-ferase and finally CDP-choline: 1,2-diacylglycerol cholinephos-photransferase (CPT1) catalyzes a reaction between CDP-cholineand DAG leading to the formation of PC and CMP. In theN-methylation pathway, PE is N-methylated three times lead-ing to the formation of PC. Only few bacteria were knownto be able to synthesize PC and for a long time only the N-methylation pathway for PC synthesis was known in prokary-otes (Kaneshiro and Law 1964; Sohlenkamp, Lopez-Lara andGeiger 2003) (Figs 1 and 4, reactions 12a–c). The genes encodingthe bacterial Pmt enzymes were discovered first in R. sphaeroidesand later in S. meliloti (Arondel, Benning and Somerville 1993;de Rudder, Lopez-Lara and Geiger 2000). Both Pmts share lit-tle sequence similarity and are proposed to belong to two dif-ferent methyltransferase families. The similarity on amino acidlevel is restricted to the putative S-adenosylmethionine (SAM)-binding site and the S. meliloti Pmt shows homology to rRNAmethylases, whereas the R. sphaeroides Pmt shows homologyto ubiquinone/menaquinone methyltransferases. Both methyl-transferases can catalyze all three methylation steps requiredfor PC formation from PE. Later it was observed that the phos-pholipid N-methylation pathway is more complex in Bradyrhizo-bium japonicum and probably other bacteria (Hacker et al. 2008).Bradyrhizobium japonicum presents four methyltransferases thatcan use PE or itsmethylated derivativesMMPE and DMPE as sub-strates (PmtA, PmtX1, PmtX3 and PmtX4) when expressed aloneor together with other Pmts in E. coli. PmtA preferentially con-verts PE to MMPE whereas PmtX1 carries out the subsequentmethylation steps. PmtX3 and PmtX4 canmethylate PE to MMPEand DMPE when expressed in E. coli. The use of multiple Pmt en-zymes has been also described in the yeast Saccharomyces cere-visae which uses two different methyltransferases that togethercause the formation of PC from PE (Kodaki and Yamashita 1987;Kanipes and Henry 1997).

In the late 1990s the PC synthase (Pcs) pathway was discov-ered 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 and4, reaction 13). Pcs belongs to the CDP-alcohol phosphotrans-ferase superfamily. It presents a slightly modified CDP-alcoholphosphotransferase motif that makes the bioinformatic pre-diction of Pcs-encoding genes straightforward (see above). Incontrast, the prediction of N-methyltransferases involved in PCsynthesis is much more difficult. Looking for the presence of ei-ther the Pmt or the Pcs pathway in bacteria using genomic se-quence data, it has been estimated that about 15% of bacteria areprobably able to form PC (Sohlenkamp, Lopez-Lara and Geiger2003; Geiger, Lopez-Lara and Sohlenkamp 2013). The Pcs path-way is widely distributed in α-proteobacteria (mainly Rhodobac-terales and Rhizobiales) and absent in β-proteobacteria. Withinthe γ -proteobacteria, it is present in most Pseudomonas speciesand in the genera Legionella,Halomonas and Francisella andwithinthe δ-proteobacteria only a few strains of Desulfovibrio present aputative pcs gene. Genes encoding Pcs can also be found out-side the proteobacteria. Most noteworthy is probably that moststrains of Borrelia, severalActinomyces sp. strains (Actinomycetales)and several Planctomycetales have Pcs (Geiger, Lopez-Lara andSohlenkamp 2013).

Kent et al. (2004) showed evidence for the presence of a CDP-choline pathway in the spirochaete Treponema denticola. In thecase of Treponema, the first two steps of the CDP-choline path-way are catalyzed by the bifunctional enzyme LicCA responsi-ble for the formation of CDP-choline (Rock et al. 2001; Campbelland Kent 2001; Kwak et al. 2002). Homologs of LicCA are alsopresent in Haemophilus influenzae, different strains of Clostrid-ium 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 stepof the pathway leading to the formation of PC in Treponema hasyet to be identified, but it should catalyze the same reaction asthe Cpt enzyme responsible for the final step of the eukaryoticCDP-choline pathway (Figs 1 and 4, reaction 16). Such a prokary-otic CDP-choline pathway is distinct from the eukaryotic path-way because at least for the first two steps it uses unrelated en-zymes. It is not clear if such a prokaryotic CDP-choline pathwayis present outside the genus Treponema.

Recently, Moser, Aktas and Narberhaus (2014b) found that X.campestris can form PC using glycerolphosphocholine (GPC) as asubstrate for PC formation (Figs 1 and 4, reactions 14 and 15). Asimilar GPC-dependent pathway for PC formation has been de-scribed in the yeast S. cerevisiae (Stalberg et al. 2008). Two acyl-transferase reactions are required to form PC from GPC. In both,yeast and X. campestris only enzymes for the second acyltrans-ferase step from LPC to PC were identified, whereas it is notknown which activity is catalyzing the first step. At present, itis not clear how widespread such a GPC-dependent pathway forPC synthesis is.

Bacterial mutants deficient in PC formation have been de-scribed in several species.Agrobacterium tumefaciensmutants de-ficient in PC formation (pcs−/pmt−) have a defective type IV se-cretion system and do not cause tumor formation on Kalanchoeleaves (Wessel et al. 2006). Legionella pneumophila PC-deficientmutants (pcs−/pmt−) show reduced virulence due to defectsin the type IVB secretion system and macrophage adhesion(Conover et al. 2008). Brucella abortusmutants deficient in PC for-mation (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 andthey are sensitive to hypo- and hyperosmotic growth conditions(Sohlenkamp, Lopez-Lara and Geiger 2003; Geiger, Lopez-Lara


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.


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

Pathway for PI and PIM synthesis

Similar as PC, also PI has been considered a typical eukary-otic lipid. Therefore, it is probably not surprising that there areonly few bacteria that can form PI. PI formation is widespreadwithin the order Actinomycetales, but outside this order only afew 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 andYamashita 1997; Xue et al. 2000). It was thought for many yearsthat the same reaction should be occurring in bacteria, butrecently it was shown for a few bacterial species that a Pipsenzyme catalyzes the reaction between inositol-1-phosphateand CDP-DAG, leading to the formation of phosphatidylinositolphosphate (PIP) (Figs 1 and 5A, reaction 17). In a second step,PIP is dephosphorylated by PIP phosphatase (Pipp) to PI (Moriiet al. 2010, 2014) (Figs 1 and 5A, reaction 18). PI can be furthermodified with mannose residues and acyl groups forming thePIM (Figs 1 and 5A, reactions 19–22). These are the structuralbasis of the lipoglucans lipomannan and lipoarabinomannan(Guerin et al. 2010). PI is essential inMycobacterium (Jackson, Crickand Brennan 2000). The first step of PIM formation in M. tuber-culosis is catalyzed by the mannosyltransferase PimA and con-sists in the transfer of a mannose residue from GDP-mannoseto the 2-position of the myo-inositol ring (Hill and Ballou 1966).Subsequently, the mannosyltransferase PimB′ transfers a man-nose residue from GDP-mannose to the 6-position of the myo-inositol ring of PIM (Guerin et al. 2009). Both PIM1 and PIM2 canbe acylated at the 6-position of the first mannose residue trans-ferred by the acyltransferase Rv2611c, although PIM2 seems tobe the favored substrate (Kordulakova et al. 2003; Guerin et al.2009; Svetlikova et al. 2014). The acyltransferase responsible forthe second acylation step has not been identified. Ac1PIM2 andAc2PIM2 can then be further elongated with additional mannoseresidues thereby forming higher PIM species. Their structuresand synthesis pathways are discussed in detail by Guerin et al.(2009). PI and mannosides derived thereof are also present inmany other genera such as Streptomyces, Nocardia or Corynebac-terium. The PIM structures and their biosyntheses have not beenstudied in detail outside the genus Mycobacterium and it is cer-tainly possible that there are differences.

Recently, a novel pathway for the synthesis of inositolphospholipids was described in the thermophilic bacteriumR. marinum (Jorge, Borges and Santos 2014). L-myo-inositol-1-phosphate cytidylyltransferase catalyzes the formation ofCDP-inositol from inositol-1-phosphate and CTP. Bacterial di-alkylether glycerophosphoinositol synthase (BEPIS) (Fig. 5B, re-action 23) then uses CDP-inositol and dialkylether glycerols toproduce phosphoinositol ether lipids.

PATHWAYS FOR SYNTHESIS OFPHOSPHORUS-FREE MEMBRANE LIPIDS INBACTERIASynthesis of the common precursor DAG

DAG can be produced by a set of metabolic reactions in thecell. Phospholipase C action upon phospholipids will produceDAG and the phosphorylated headgroup, for example PC will

be cleaved into DAG and phosphocholine. The headgroup trans-fer from phospholipids to membrane-derived oligosaccharidesor periplasmic cyclic glucans is also a source of DAG. For ex-ample, phosphoglycerol is transferred from PG to periplasmicglucans, leading to the formation of DAG (Jackson, Bohin andKennedy 1984; Fiedler and Rotering 1985; Zavaleta-Pastor et al.2010; Geiger, Lopez-Lara and Sohlenkamp 2013). Alternatively,PA could be dephosphorylated to DAG by PA phosphatases (Bergand Wieslander 1997; Carman 1997; Zhang et al. 2008; Combaet al. 2013), but it is not clear how relevant this reaction is inbacteria.

Glycolipid (GL) synthesis

GL synthesis usually startswith the transfer of a sugarmonomerfrom 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 themonoglycosyl-DAG leading to more complex GLs. Processiveglycosyltransferases are responsible for the transfer of severalsugars and can for example catalyze three transferase steps,each time using the GL produced by the earlier step as sub-strate (Devers et al. 2011). Some bacterial species produce a widevariety of GLs and in some cases the sugar residues can becovalently modified. Several glycosyltransferases from differentbacteria have been cloned and characterized, whereas notmuchis known about the cellular function of the GLs formed (Joraschet al. 1998; Devers et al. 2011; Geske et al. 2013; Diercks et al. 2015).Readers more interested in GLs and their structures and synthe-ses are referred to a review by Holzl and Dormann (2007).

Sulfoquinovosyl diacylglycerol (SQD) synthesis

A special GL is the anionic lipid SQD which is widely distributedin photosynthetic organisms. Its synthesis has been studied inbacteria and plants. In R. sphaeroides, four genes called sqdA,sqdB, sdqC and sqdD have been identified by genetic analysisto be involved in SQD formation (Benning 1998). SqdA from R.sphaeroides is annotated within the genome sequence as a phos-pholipid/glycerol acyltransferase. SqdB is thought to be involvedin the formation of UDP-sulfoquinovose from UDP-glucose andsulfite. It is the only gene involved in SQD synthesis that has or-thologs in all SQD-producing organisms, including plants whereit is named SQD1. SqdC and SqdD are probably involved in thetransfer of sulfoquinovose to DAG leading to the formation ofSQD (Benning 1998). SqdC is annotated as member of the NAD-dependent epimerase dehydratase family, whereas SqdD is an-notated as a glycosyltransferase. In the cyanobacterium Syne-chococcus sp. PCC 7942, the gene sqdB forms an operon withthe gene encoding the lipid glycosyltransferase SqdX. SqdX waslater shown to be the cyanobacterial SQD synthase (Guler, Es-sigmann and Benning 2000). Plants use an SqdX homolog calledSQD2 for SQD synthesis (Shimojima 2011). Mutants deficient inSQD formation have been described in S. meliloti, Synechococ-cus sp. PCC 7942 and R. sphaeroides (Benning and Somerville1992a,b; Rossak et al. 1995, 1997; Guler et al. 1996; Weissen-mayer, Geiger and Benning 2000). An S. meliloti mutant defi-cient in SQD formation is not affected during nodule develop-ment and function (Weissenmayer, Geiger and Benning 2000).SQD deficiency in R. sphaeroides did not affect photosynthe-sis in any obvious way, but it absence interfered with bacte-rial growth under phosphate-limiting conditions (Benning et al.1993). The Synechococcus mutant deficient in SqdB presentedminor changes in a few photosynthesis parameters. Again,


Figure 5. Synthesis of PI and PI-derivatives. (A) Synthesis of PI and PI-derivatives in Actinomycetes. (B) CDP-inositol-dependent pathway for dialkylether glyc-erophosphoinositol formation described in R. marinus. CMP-cytidine monophosphate; CDP-cytidine diphosphate; GMP-guanosine monophosphate; GDP-guanosinediphosphate; 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.


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

DGTS synthesis

Bacteria use two enzymes to synthesize DGTS from DAG.The genes encoding the required activities were first discov-ered in R. sphaeroides (Klug and Benning 2001). The enzymeBtaA responsible for the formation of diacylglyceryl homoser-ine (DGHS) is a SAM/DAG 3-amino-3-carboxypropyltransferase.BtaB is a trifunctional N-methyltransferase related on sequencelevel to bacterial phospholipid N-methyltransferases of the R.sphaeroides family. When BtaA and BtaB from R. sphaeroides orS. meliloti are expressed in E. coli, DGTS is formed (Riekhof, An-dre and Benning 2005a). Based on the distribution of BtaA, it canbe predicted that the capacity to form DGTS is widespread in afew orders of the α-proteobacteria like the Rhodobacterales, Sph-ingomonadales and Rhizobiales. Genes encoding BtaA homologscan also be found in the genomes of a few Planctomycetales, γ -proteobacteria (Thiotrix, Thiomicrospira), cyanobacteria (Mastigo-coleus, Calothrix, Anabaena, Leptolyngba), δ-proteobacteria (Desul-fovibrio), firmicutes (Geobacillus, Brevibacillus), Deinococcus andAcidobacterium. In the alga Chlamydomonas reinhardtii, DGTS syn-thesis is performed by a bifunctional enzyme in which both en-zyme functions are fused (Riekhof, Sears and Benning 2005).The N-terminal domain of the protein is BtaB-like whereas theC-terminal domain is BtaA-like. Similar bifunctional enzymesare also present in fungi and other eukaryotes (Riekhof et al.2014). Sinorhizobiummelilotimutants deficient in DGTS formationwere affected in cell yield under phosphate-limiting conditionswhen the DGTS deficiency was combinedwith a deficiency in OLsynthesis (Lopez-Lara et al. 2005).

OL synthesis

The genes encoding the acyltransferases OlsB and OlsA re-quired for OL synthesis in S. meliloti have been describedfirst by Gao et al. (2004) and Weissenmayer et al. (2002). TheN-acyltransferase OlsB transfers a 3-hydroxy fatty acid from theconstitutive acyl carrier protein AcpP to the α-amino group of or-nithine, thereby forming lyso-OL (LOL) (Fig. 6, reaction 24). Next,theO-acyltransferase OlsA transfers a fatty acyl chain fromAcpPto 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 subsequentS1-modifying reactions (Rojas-Jimenez et al. 2005). Recently, thebifunctional acyltransferase OlsF was described in Serratia pro-teamaculans that is responsible for OL formation in this organ-ism (Fig. 6, reaction 26). OlsF has two acyltransferase domains,and the N-terminal domain was shown to be responsible for theO-acyltransferase reaction whereas the C-terminal domain wasshown to be responsible for the N-acyltransferase reaction re-quired for OL synthesis (Vences-Guzman et al. 2014). OlsBA arepresent in α-, β- and a few γ -proteobacteria and actinomycetes,whereas OlsF is present in γ -, δ- and ε-proteobacteria, and bac-teria of the CFB group (Vences-Guzman et al. 2014). Based on thedistribution of OlsB and OlsF in bacterial genomes, it has beenestimated that about 50% of the sequenced bacteria can formOL (Vences-Guzman et al. 2014). Unmodified OL can be modifiedby the introduction of one or more hydroxyl group or by methy-lation of the δ-amino group in the ornithine headgroup (Rojas-Jimenez et al. 2005; Gonzalez-Silva et al. 2011; Vences-Guzmanet al. 2011; Vences-Guzman, Geiger and Sohlenkamp 2012;Mooreet al. 2013). Three different OL hydroxylases have been described

so far: OlsC was described first in R. tropici and introduces ahydroxyl group in the 2-position of the ester-bound fatty acid(Fig. 6, reaction 27) (Rojas-Jimenez et al. 2005; Vences-Guzmanet al. 2011). OlsD was described first in Burkholderia cenocepaciaand introduces a hydroxyl group at the 2-position in the amide-bound fatty acid and gives rise to the formation of new lipids 1and 2 (NL1, NL2) (Fig. 6, reaction 28) (Gonzalez-Silva et al. 2011;Diercks et al. 2015). OlsE introduces a hydroxyl group in the or-nithine headgroup at a still unknown position (Fig. 6, reaction29) (Vences-Guzman et al. 2011). The methyltransferase respon-sible for the methylation of the δ-amino group of the ornithineheadgroup in the planctomycete S. acidiphila has not been iden-tified (Fig. 6, reactions 30–32) (Moore et al. 2013). OL is proba-bly N-methylated by the unknown methyltransferase leading tothe formation of TMOL (Fig. 6, reaction 30), but it is also possi-ble that free ornithine is N-methylated before TMOL is formed(Fig. 6, reactions 31 and 32). Mutants have been identified inmost genes described to be involved in OL synthesis or modi-fication. Agrobacterium tumefaciens mutants deficient in OlsB orOlsE show accelerated tumor formation in a potato-disc assay(Vences-Guzman et al. 2013). Rhizobium tropici mutants deficientin OlsC are defective in growth at high temperature and at acidpH (Vences-Guzman et al. 2011). Mutants deficient in OlsC and/orOlsE in R. tropici are also defective in nodulation assays usingbean as a host (Vences-Guzman et al. 2011). Mutants deficientin OL formation in S. meliloti showed a slight growth defect un-der phosphate-limiting conditions when at the same time DGTSsynthesis was interrupted (Lopez-Lara et al. 2005). On the otherhand, B. abortus mutants deficient in the formation of OL werenot affected in virulence (Palacios-Chaves et al. 2011). Dierckset al. (2015) could show recently that OL in M. loti contains 80–90% of D-ornithine which has not been observed in other organ-isms. It is not clear whether D-ornithine is also present in OLsof other bacteria.

Sulfonolipid synthesis

Sulfonolipids aremajor lipids in the outermembrane of the glid-ing bacterium Flavobacterium johnsoniae (formerlyCytophaga john-sonae). They contain capninewhich is formed by condensation ofcysteic acid with fatty acyl-CoA under the release of CO2 (White1984; Abbanat et al. 1986). An N-acyltransferase then transfersa fatty acid to capnine leading to the formation of N-acyl cap-nine. Mutants deficient in the synthesis of N-acyl capnine werealso deficient in gliding, so theremight be a connection betweenthe presence of the lipid and the ability to move by gliding (Ab-banat et al. 1986). The genes encoding the enzymes required forsulfonolipid synthesis have not been identified. The proposedstructures of the molecular pathway for sulfonolipid synthesiscan be found in Geiger et al. (2010).

Sphingolipid (SL) synthesis

SLs are ubiquitous components of the eukaryotic cytoplasmicmembranewhere they have important functions in signal trans-duction processes and in the formation of lipid rafts. In con-trast, SLs occur only in a few bacteria, mainly belonging to theCFB group and the Sphingomonadales of the α-proteobacteria, butthey are also present in a few δ-proteobacteria such as Bdellovib-rio, Cystobacter fuscus, My. stipitatus, S. cellulosum and M. xanthus(Keck et al. 2011; Lorenzen et al. 2014). Sequence analysis basedon the presence of genes encoding for putative serine palmi-toyltransferases (SPTs) indicates that SLs might be present ina few α-proteobacteria outside the Sphingomonadales and even


Figure 6. Synthesis of OLs and their hydroxylated or methylated derivatives in bacteria. AcpP-constitutive acyl carrier protein; α-KG-alpha-ketoglutarate. The namesof 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 OLmodifying reactions (Rojas-Jimenez et al. 2005). The formation of new lipids

1 and 2 (NL1, NL2) is due to the enzymatic activity of OlsD.

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

Sphingomonadales, SLs aremainly decorated with sugar residues.In Bacteroides species, ceramide phosphorylethanolamine andceramide phosphorylglycerol have been described. The eukary-otic genes involved in SL biosynthesis have identified but littleis known about SL synthesis in bacteria (Dickson 2010; Geiger


et al. 2010; Pata et al. 2010). Only the first step, catalyzed bySPT, is characterized in detail (Lowther et al. 2011, 2012). Eventhe crystal structures of the SPTs from Sphingomonas paucimo-bilis, S. wittichii and Sphingobacterium multivorum have been de-termined (Yard et al. 2007; Ikushiro et al. 2009; Raman et al. 2010).In eukaryotes, acyl-CoA and serine are the substrates for thisenzyme, whereas for the enzyme from S. wittichii acyl-ACP hasbeen suggested as acyl donor, because the gene-encoding SPTforms an operon with a gene encoding a putative ACP (Geigeret al. 2010; Raman et al. 2010). SLs are often 2-hydroxylated andthe α-hydroxylase catalyzing this reaction has been identifiedand characterized from different bacteria (Matsunaga et al. 1997;Ring et al. 2009; Fujishiro et al. 2011). Not much is known aboutthe functions of bacterial SLs. It has been described that someGram-negative bacteria, such as Sp. capsulata, Sp. adhaerens andS. cellulosum, lack LPS in their outer membrane and instead haveglycosphingolipids 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 theinner and outer membrane in proteobacteria. They are of-ten considered bacterial sterol analogs and were first isolatedfrom resins of tropical trees of the genus Hopea (Ourisson,Rohmer and Poralla 1987). HOPs are pentacyclic triterpenic iso-prenoids. Synthesis of the precursors isopentenyl diphosphate(IPP) and dimethylallyl diphosphate (DMPP) occurs dependingon the bacterial species via the mevalonate pathway or via themethylerythrol-4-phosphate pathway (Perez-Gil and Rodrıguez-Concepcion 2013). DMPP is elongated by head-to-tail conden-sation with two molecules IPP to farnesyl diphosphate. Twomolecules farnesyl diphosphate are then condensated head-to-head leading to the formation of squalene. Squalene hopenecyclase (Shc) is responsible for the cyclization of squalene todiplotene or diplopterol (Fig. 7, reaction 33). The synthesis oftetrahymanol, another pentacyclic structure related to HOPs,was also shown to be Shc-dependent in Rhodopseudomonas palus-tris (Welander et al. 2009). Shc is an ortholog of squalene-2,3-epoxide cyclase responsible for the formation of lanosterol fromoxidosqualene. A Shc homolog from firmicutes has been shownto be responsible for the synthesis of sporulenes (Bosak, Losickand Pearson 2008; Kontnik et al. 2008), heptaprenyl lipids thatincrease the resistance of spores to reactive oxygen species.

Starting fromdiploptene elongatedHOPs can be synthesized.Only in recent years, many of the genes/enzymes required forHOP formation have been identified using R. palustris, Methy-lobacterium extorquens and B. cenocepacia asmodel organisms, andthere are still many open questions with respect to the forma-tion of complex HOPs. Often, genes involved in HOP synthesisand modification form operons and cluster within the bacterialgenomes. The radical SAM superfamily enzymeHpnH is respon-sible for the formation of adenosylhopane (Bradley et al. 2010)(Fig. 7, reaction 34). HpnG is responsible for the release of ade-nine from adenosylhopane (Fig. 7, reaction 35). HpnG shows ho-mology to purine nucleoside phosphorylases, but there is still adebate if the reaction product is phosphoribohopane or ribosyl-hopane. It has been suggested that the cyclic ribose opens upwithout the aid of an enzyme (Bradley et al. 2010) (Fig. 7, reac-tion 36). Formylhopane is probably the substrate for the amino-transferase reaction catalyzed by HpnO (Fig. 7, reaction 37) oralternatively it can be reduced to bacteriohopanetetrol (BHT)(Fig. 7, reaction 38). The conversion of ribosylhopane to BHThas been also suggested to happen non-enzymatically, but it

is unclear how the required reduction should be achieved thisway (Schmerk et al. 2015) (Fig. 7, reaction 39). For the forma-tion of BHT glucosamine, the enzymes HpnI and HpnK are re-quired (Bradley et al. 2010; Schmerk et al. 2015) (Fig. 7, reac-tion 40 and 41). HpnI is a glycosyltransferase and HpnK hasbeen suggested to be a deacetylase. HpnJ is a radical SAM su-perfamily member required for the formation of BHT cyclito-lether, but the mechanism of the ring contraction reaction isnot known (Fig. 7, reaction 42). HOPs can also be C6- and C11-unsaturated, but the enzymes responsible for these unsatura-tion reactions have not been identified (Fig. 7, reactions 43 and44). C2 and C3 methylation reactions of HOPs have been de-scribed and the methyltransferases have been identified (Fig. 7,reactions 45 and 46) (Welander et al. 2010; Welander and Sum-mons 2012). The presence of 2-methylhopanoids in sedimen-tary fossils has been used for many years as a biomarker proxyfor 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 biomarkerproxy for aerobic methanotrophs in the past, but it is clear nowthat several proteobacteria and actinomycetes also have a geneencoding homologs of the methyltransferase HpnR required for3-methylhopanoid formation. In addition to the HOP structurespresented here, other HOP structures have been described. InBradyrhizobium, HOPs covalently linked to lipid Ahave been iden-tified (Komaniecka et al. 2014; Silipo et al. 2014). Mutants in differ-ent steps of HOP formation have been characterized. Rhodopseu-domonas palustrismutants deficient in HOP formation presentedaweakened outermembrane andweremore sensitive to pH andtemperature stress (Welander et al. 2009; Doughty et al. 2011).Burkholderia cenocepacia defective in HOP formation displayed in-creased sensitivity to low pH, detergents, various antibiotics anddid not produce flagella (Schmerk, Bernards and Valvano 2011).In contrast, in S. scabies HOPs were not required for tolerance toethanol, oxidative and osmotic stress, high temperature or lowpH (Seipke and Loria 2009).

METABOLIC PATHWAYS CONNECTINGPHOSPHOLIPIDS AND PHOSPHORUS-FREEMEMBRANE LIPIDS

Under phosphorus-limiting conditions, some bacteria canreplace phospholipids such as PE, PG or PC from their mem-branes with phosphorus-free membrane lipids such as DGTS,SQD, GLs or OLs. In S. meliloti, this membrane remodeling isrelatively well studied and some important players have beenidentified. Pulse-chase experiments had suggested that PE andPC are precursors for the synthesis of DGTS. The phospholipaseC SMc00171 from S. meliloti was shown to be responsible forthe conversion of PC to DAG and phosphocholine (Zavaleta-Pastor et al. 2010). DAG can then be used in S. meliloti forthe synthesis of the phosphorus-free lipid DGTS and proba-bly also of the DAG-containing lipid SQD. The expression ofSMc00171 and of several of the enzymes required for SQD andDGTS synthesis is regulated by the transcriptional regulatorPhoB and is induced under phosphate-limiting conditions.Interestingly, the PE methyltransferase from S. meliloti is alsoinduced about 4-fold and the mechanism by which PE is de-graded might be via PC. Genes encoding SMc00171 homologsare present in a few α-proteobacteria (A. tumefaciens, M. loti,B. japonicum, Caulobacter crescentus and R. palustris) and in afew Pseudomonas species. SMc00171-like phospholipases are


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


homologs of LpxH, an enzyme required for lipid A synthesisthat hydrolyses UDP-2,3-diacylglucosamine to UMP andlipid X (2,3-bis-(3R-hydroxy-tetradecanoyl)-αD-glucosamine-1-phosphate) (Babinski, Kanjilal and Raetz 2002a; Babinski, Ribeiroand Raetz 2002b). Genes encoding for SMc00171-like enzymesare absent in intracellular pathogens such as Brucella, Bartonellaand Rickettsia. This absence of a phosphate starvation responseis probably the consequence of an intracellular lifestyle whichis characterized by a stable phosphate supply.

ENVIRONMENTAL CHALLENGES CAUSINGCHANGES IN MEMBRANE LIPID COMPOSITION

Bacteria change their membrane lipid composition in responseto changes in the environment. This adaptation allows bacte-ria to survive unfavorable conditions. Even during the simpleexperiment of growing microorganisms in a flask, the environ-ment the bacteria are sensing is changing constantly, which willbe affecting the membrane lipid composition: nutrient levelsfall, products of metabolism accumulate, the pH of the culturemedium may rise or fall and oxygen levels are changing. Theprobably best-known example of membrane adaptation hap-pens when bacteria are growing at different temperatures. InE. coli and other bacteria, a decrease in temperature causesa decrease in membrane fluidity and bacterial cells respondby increasing the amount of unsaturated fatty acids or fattyacids with analogous properties. An increase in temperaturecauses an increase in fluidity and probably an increase in the oc-currence of discontinuities in the membrane. Bacteria growingat increased temperature usually contain more saturated fattyacids in their membranes (Marr and Ingraham 1962). However,membrane modifications as a response to abiotic stress condi-tions are not restricted to the fatty acid moieties and do alsohappen on headgroup level. The membrane lipid modificationsoccurring can be divided in two types. (1) Existing lipids can bemodified to obtain amembrane with different properties; for ex-ample, the anionic lipid PG and be modified to the cationic lipidLPG simply by transfer of one lysyl group. This modification ofpre-existing membrane lipids has the advantage that it allows aquick response to changes in environmental conditions. (2) Ex-isting lipids are degraded and new lipids with new characteris-tics are synthesized de novo replacing the old lipids. In the fol-lowing, wewill discuss howmembrane lipidmodifications occurunder different stress conditions.

Growth under acidic conditions provokes a modification inmembrane lipids. In R. tropici, two membrane-modifying activ-ities have been described that occur under condition of acidstress and that allow the bacteria to prosper under these con-ditions. The operon lpiA/atvA is transcriptionally induced afteracid shock (Vinuesa et al. 2003). LpiA is an LPG synthase homolo-gous to MprF responsible for formation of the cationic lipid LPG(Sohlenkamp et al. 2007). The second gene of the operon encodesthe putative lipase AtvA. This operon structure is also presentin P. aeruginosa. Pseudomonas aeruginosa MprF is responsible forAPG formation and the AtvA homolog PA0919 has been shownto cause the hydrolysis of APG and formation of PG and alanine.Thus, it can be hypothesized that AtvA from R. tropici will hy-drolyze LPG to lysine and PG, but at the moment it is not clearhow this should lead to an increased acid resistance of the bac-teria. This modification of the anionic membrane lipid PG to thecationic (LPG) or zwitterionic (APG) membrane lipids has beenshown to be important in resistence to cationic peptides and an-tibiotics. Another lipid modification described in R. tropici that is

known to have importance under acidic growth conditions is the2-hydroxylation of OL by OlsC described above (Rojas-Jimenezet al. 2005; Vences-Guzman et al. 2011) (Fig. 6, reaction 27). Anα-hydroxyl group is introduced into the ester-bound fatty acidof OL, analogous to the 2-hydroxylation of lipid A-bound fattyacids 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 addi-tional hydroxyl group allows the formation of hydrogen bondsthat increase the lateral interactions between lipid moleculesand stabilize the leaflet structure (Nikaido 2003). Consistentwiththis hypothesis, R. tropici mutants deficient in OlsC and lackingthe 2-hydroxylated OL termed P1 show a severe growth defect ata pH lower than 4.5 (Vences-Guzman et al. 2011).

The 2-hydroxylation of OL also seems to play a role in theresponse to increased growth temperature in R. tropici and B.cepacia. In R. tropici, a mutant deficient in the 2-hydroxylase OlsCgrows like the wildtype at 30◦C, but grows significantly slowerthan the wildtype at 37 and 42◦C, whereas B. cepacia accumu-lates increased amounts of hydroxylated OL and hydroxylatedPE at increased growth temperatures. Again, increased lateralinteractions between lipid molecules due to the extra hydroxylgroups might be responsible for this phenotype (Taylor, Ander-son and Wilkinson 1998; Vences-Guzman et al. 2011).

During the response to high osmotic pressures, the anionicmembrane lipid CL has been shown to play an important rolein different bacteria. Increased osmotic pressure causes an in-crease 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 cellu-lar poles where the proline transporter ProP is embedded in CL-enriched regions of the cytoplasmic membrane. ProP is denotedan osmosensory transporter because it is activated by increas-ing osmolarity and causes the accumulation of the compatiblesolute proline allowing bacterial growth at increased osmoticpressure. The activation threshold of ProP correlates with theamount of CL present in the membrane and its presence con-trols 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 toconditions of phosphate limitation. Plants and microbes mostlylive in environments where available phosphate is a growth-limiting factor. Apparently, some bacteria can replace theirphospholipidswithmembrane lipids devoid of phosphorus. Thisremodeling has been studied in some detail in R. sphaeroides,S. meliloti, A. tumefaciens and M. loti (Benning et al. 1993; Ben-ning, Huang and Gage 1995; Geiger et al. 1999; Zavaleta-Pastoret al. 2010; Devers et al. 2011; Geske et al. 2013; Diercks et al.2015). The phospholipids PE and PC are actively degraded andare replaced by the DAG-derived phosphorus-free membranelipids DGTS, SQD and GLs and by the aminolipid OL. This re-modeling is not a minor process because PC makes up for about60% of membrane lipids when bacteria are cultivated on com-plexmedium,whereas inminimalmediumunder limiting phos-phate conditions DGTS increases to about 60% (Geiger et al.1999). It was suggested that phospholipids are functionally re-placed by phosphorus-free membrane lipids under conditionswhere phosphate becomes limiting thereby making the phos-phate pool present in the membrane accessible for other cel-lular processes such as nucleic acid synthesis. It is thoughtthat PC is replaced by DGTS, both being zwitterionic lipids withbulky headgroups, that PG is replaced by SQD, both being an-ionic lipids and that PE is replaced by OLs, both being zwitte-rionic lipids with smaller headgroups. There is evidence sup-porting these hypotheses: Recently, Riekhof and collaborators


constructed a S. cerevisiae strain that had replaced its PC withDGTS. They showed that essential processes of membrane bio-genesis and organelle assembly were functional in this strain(Riekhof et al. 2014). On an experimental level, this is probablythe best hint that DGTS can functionally replace PC in themem-brane. In cyanobacteria, which form PG, SQD and neutral GLsunder phosphate-replete conditions, PG is decreased underphosphate limitation whereas SQD is increased. In A. tumefa-ciens, different OLs are formed constitutively meaning that theirsynthesis does not depend on phosphate starvation but also oc-curs in phosphate-replete complex medium. When an A. tume-faciensmutant deficient in OlsB is constructed, PE levels increasedrastically (Vences-Guzman et al. 2013).

OPEN QUESTIONS

Bacteria account for most of the diversity of life on our planetand they have been around for billions of years. During thistime they have adapted to all types of environments on Earth.From this point of view, it is probably not surprising that bacte-ria present such a rich diversity of lipid structures and pathwaysinvolved in the formation ofmembrane lipidswhen compared toeukaryotes. When studying the diversity of bacterial membranelipids, the ultimate goal probably should be to have a completecatalog of the lipids that are formed when bacteria are growingin their natural habitats and understand their functions withinthe physiology of the organisms. Although we have come a longway, there is still much work to do. What are the problems andwhat has to be done to fill these information gaps?

(I) Only a few bacteria can be easily grown in the labora-tory in pure cultures. Most bacteria of the environmentstill escape our attempts to grow them in the laboratory.This means that almost everything we know about bacte-rial membrane lipids comes from a relatively small set ofeasily culturable bacteria. We can probably expect to findseveral unknown lipid structures and unknown synthesispathways in these unculturable bacteria. One possibility tountap part of this variety of bacterial membrane lipids isto combine culturomics approaches with lipidomics (Lagieret al. 2012). Culturomics is a systematic approach to findgrowth conditions of so-far unculturable bacteria. Once thisor these conditions are defined, the organism’s membranelipid composition can be studied. Examples of recently dis-covered lipid structures include a trimethylated OL in Planc-tomycetes, diacylserinophospholipids from Verrucomicrobiaand diol-based lipids in thermophilic bacteria (Moore et al.2013; Lagutin et al. 2014; Anders et al. 2015).

(II) Currently, we have a basic knowledge about the lipid com-position of major model bacteria and many of the genesand enzymes involved in their synthesis. However, even thewell-studied E. coli still has hidden secrets and we are prob-ably more ignorant than we want to realize. For decadeslipids biochemists had the idea that not much new was tobe discovered in E. coli, but recently, Tan et al. (2012) iden-tified the Cls ClsC that synthesizes CL from the precur-sors PG and PE, something that was completely unheardof. It belongs to the PLD superfamily, which together withthe CDP-alcohol phosphotransferase superfamily (Pips, Pcs,PgsA, type-II PssA or the eukaryotic type Cls) represents alarge fraction of the known enzymes involved in bacterialphospholipid synthesis and turnover. Even in well-studiedorganisms such as E. coli, S. meliloti or St. coelicolor, thereare proteins belonging to one of these two superfamilies

with unassigned functions. Almost all CDP-alcohol phos-photransferases characterized play a role in phospholipidsynthesis, the only known exceptions being a few enzymesinvolved in the synthesis of inositol-based compatible so-lute in hyperthermophilic bacteria and archaea (Brito et al.2011; Nogly et al. 2014). Although there might be redun-dancy with respect to some activities, bacteria might formother lipids that have not yet been described, and/or newenzymatic activities are waiting to be discovered. Examplesare the above-mentioned ClsC or the CL/PE synthase fromX. campestris (Tan et al. 2012; Moser et al. 2014a). A system-atic mining the wealth of genome sequence data shouldnot be restricted to both protein superfamilies, but includealso the search for other genes/enzymes such as acyltrans-ferases involved inmembrane lipid synthesis, turnover anddegradation. It can be expected that many new structuresand activities will be identified.

(III) Bacteria are cultured in the laboratory under conditionsthat in most cases do not reflect the natural habitat. Themembrane lipid composition of bacteria is not a constantbut is responding to the environmental conditions. If wewant to understand what role the membrane (lipids) playfor the functioning of a bacterium but also during the in-teraction with other bacteria or eukaryotes, it is essentialto study the bacteria within their habitat. Experimental ap-proaches should be developed and improved that allow thestudy of bacterial membrane lipids in the natural environ-ment or try to take the habitat to the laboratory. Thatmeansgo to the sites where the bacteria are growing, sample andif possible study in situ or bring part of the habitat to thelaboratory and study it there. One possibility is to combinethe study of environmental lipid samples as for examplegeobiochemists do when analyzing samples from marinesediments, with metagenomic studies. The metagenomicstudywill informuswhat bacteria are present in the habitatand the lipid analysis will show what membrane lipids arepresent. In this model metadata such as the environmen-tal conditions can be included, so that amodeling should bepossible. Finally, the perfect experimental approach wouldbe to reconstitute the natural environment in the lab anduse site-directed mutants in this experimental setup to fol-low how changes or conditions affect membrane composi-tion and interactions between organisms.

FUNDING

Research in our lab was supported by grants from ConsejoNacional de Ciencia y Tecnologıa-Mexico (CONACyT-Mexico;153200, 178359) and Direccion General de Asuntos del Per-sonal Academico-Universidad Nacional Autonoma de Mexico(DGAPA-UNAM; PAPIIT IN202413, IN203612).

Conflict of interest. None declared.

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