The two PC biosynthesis pathways and corresponding enzymes (Pcs and PmtA) in A. tumefaciens have been well characterized (Figure 3). Initial work on PC synthesis in Agrobacterium demonstrated incorporation of the ¹⁴C-methyl moiety of SAM into MMPE, DMPE, and PC and ¹⁴C-choline uptake and incorporation into PC (Kaneshiro and Law, 1964; Sherr and Law, 1965; Kaneshiro, 1968). In earlier studies, two distinct Pmts were postulated in A. tumefaciens. A soluble Pmt catalyzing MMPE formation only and a Pmt associated with the particulate cell fraction producing all methylated PE-derivatives (Kaneshiro and Law, 1964). The A. tumefaciens genome, however, contains only a single constitutively expressed pmt gene (pmtA, atu0300) on the circular chromosome (Wessel et al., 2006; Klüsener et al., 2009). The lack of MMPE, DMPE, and PC in a pmtA mutant grown without choline demonstrated that PmtA is the only enzyme responsible for MMPE, DMPE, and PC synthesis via the methylation pathway (Wessel et al., 2006). Purification of recombinant PmtA from the soluble cell fraction suggests that it is a peripheral membrane protein reversibly attaching to its site of action, the membrane (Aktas and Narberhaus, 2009; Aktas et al., 2011a). PmtA is a monomeric small enzyme (~22 kDa) catalyzing the methylation of PE to MMPE, DMPE, and PC. In vitro lipid binding experiments with PmtA revealed strong binding to the anionic lipids PI and PG. Interestingly, overall PmtA activity is stimulated by PG. Association of peripheral proteins with membranes is often mediated via electrostatic interactions with negatively charged PLs such as PG and a similar mechanism is proposed for the A. tumefaciens PmtA enzyme (Figure 4). SAM binding by PmtA occurs only in the presence of its substrates PE, MMPE, DMPE or the end product PC. PG alone does not influence SAM binding suggesting that two distinct binding sites for its substrates or products and for PG exist (Aktas and Narberhaus, 2009).

In vitro PmtA activity is negatively regulated by the end products SAH (via interfering with SAM binding) and by PC. End product-mediated inhibition might also be relevant in vivo to balance proper lipid composition (Aktas and Narberhaus, 2009). Like all Pmt enzymes, PmtA contains a highly conserved N-terminal SAM binding motif [VL(E/D)XGXGXG] (Sohlenkamp et al., 2003). Within this motif, the amino acids E58, G60, G62, and E84 were found to be essential for activity and SAM binding (Aktas et al., 2011a). A. tumefaciens PmtA seems to follow an ordered Bi–Bi reaction mechanism with initial substrate binding followed by a conformational change allowing SAM binding. Subsequently, the methyl group might be transferred to the lipid substrate releasing the first product SAH followed by the release of the methylated lipid product (Aktas et al., 2010).

Bacterial Pmts are classified into Sinorhizobium and Rhodobacter type enzymes. Enzymes belonging to the Sinorhizobium family including A. tumefaciens PmtA, are homologous to rRNA methylases, whereas Rhodobacter-like Pmt enzymes are similar to UbiE, ubiquinone/menaquinone biosynthesis methyltransferases. Similarities between these two Pmt families are restricted to the conserved SAM-binding motif (Sohlenkamp et al., 2003; Aktas et al., 2010; Geiger et al., 2013). The product spectrum of Pmt enzymes varies in different organisms. While A. tumefaciens and S. meliloti pmtA release small amounts of the intermediates MMPE and DMPE, expression of R. sphaeroides pmtA in E. coli exclusively resulted in PC formation (Arondel et al., 1993; de Rudder et al., 2000; Klüsener et al., 2009). The Sinorhizobium type PmtA from X. campestris produces MMPE exclusively and is unable to further methylate it to DMPE and PC (Moser et al., 2014).

Most bacteria contain one Pmt enzyme for all three methylation steps but in some cases several Pmts with different specificities are required (Sohlenkamp et al., 2003). In the soybean symbiont Bradyrhizobium japonicum, PmtA methylates PE to MMPE, which serves as substrate for PmtX1-catalyzed methylation to DMPE and PC. B. japonicum encodes three further Pmt enzymes with distinct specificities (PmtX2-4), which are not expressed under standard laboratory conditions. PmtX1 and PmtX2 are similar to R. sphaeroides PmtA, whereas PmtA, PmtX3 and PmtX4 are homologous to S. meliloti PmtA (Minder et al., 2001; Hacker et al., 2008a,b). Like B. japonicum, Rhizobium leguminosarum, Rhodopseudomonas palustris, and Rhizobium etli seem to encode more than one pmt homolog (López-Lara et al., 2003; Martínez-Morales et al., 2003).

The second PC synthesis pathway in A. tumefaciens is catalyzed by the Pcs enzyme (Figure 4). Like pmtA, the pcs gene (atu1793) is located on the circular chromosome and is constitutively expressed (Wessel et al., 2006; Klüsener et al., 2009; Wilms et al., 2012). Pcs uses exogenous choline, which is transported via the high-affinity choline transport system ChoXWV. A choXWV-deficient strain is largely impaired in choline transport but can still produce PC when choline is present suggesting alternative choline uptake systems in A. tumefaciens (Aktas et al., 2011b). Similar to A. tumefaciens, the Pcs pathway in S. meliloti and B. abortus rely on exogenous choline delivered by a homologous Cho transport system (de Rudder et al., 1999; Dupont et al., 2004; Herrmann et al., 2013). Choline is a major component of eukaryotic membranes liberated by phospholipases from PC. Large amounts of free choline is found in homogenized plant tissues (Zeisel et al., 2003) and a recent study showed that considerable choline pools are also present on leaf surfaces. Pseudomonas syringae produces PC exclusively via the Pcs pathway and contains three choline transport systems with different specificities (Chen and Beattie, 2008). P. syringae exhibits chemotaxis towards choline and other quaternary amines. Extracellular choline is scavenged by P. syringae and enhances fitness during leaf colonization (Chen et al., 2013).

An A. tumefaciens pcs mutant produces PC via the remaining PmtA pathway and conversely PC production in a pmtA mutant depends on extracellular choline which might be delivered by the host plant. Only a pmtA/pcs double mutant lacks PC excluding alternative PC synthesis pathways in this organism (Wessel et al., 2006). Both A. tumefaciens PC biosynthesis pathways can be functionally reconstituted in E. coli demonstrating that PmtA and Pcs do not require A. tumefaciens specific cofactors or substrates (Klüsener et al., 2009).

The best-characterized Pcs enzyme derives from S. meliloti (de Rudder et al., 1999; Sohlenkamp et al., 2000; Solís-Oviedo et al., 2012). It catalyses the transfer of a phosphatidyl group from CDP-DAG to choline releasing a CMP molecule and PC. Enzyme activity depends on divalent cations like Mn²⁺ or Mg²⁺ and on detergents such as triton X100 (de Rudder et al., 1999). A topological study suggested that sinorhizobial Pcs is an integral membrane protein containing eight TM with N- and C- termini located in the cytosol. Pcs is a member of the CDP-alcohol phosphotransferase (CDP-OH-PT) protein superfamily containing a modified version of a conserved CDP-OH-PT motif (DX₂DGX₂ARX₁₂GX₃GX₃D) characteristic for this enzyme family. Most of the conserved amino acids are located within a cytosolic loop connecting the TM domains II and III and are critical for enzyme activity as shown via mutagenesis (Solís-Oviedo et al., 2012; Geiger et al., 2013). Since the membrane-bound nature of Pcs enzymes has precluded their purification and biochemical characterisation, the precise reaction mechanism of Pcs enzymes is presently unknown but most likely proceeds via a sequential Bi–Bi reaction as in other CDP-OH-PT enzymes (Geiger et al., 2013).

It is not clear why two PC biosynthesis pathways operate simultaneously in Agrobacterium and some other bacteria. Although the Pcs pathway is energetically more favorable than the PE-methylation route, under conditions of choline limitation during competition with other bacteria, the Pmt pathway might be beneficial. In Agrobacterium both pathways seem to be constitutively present. PmtA activity is detected even in the presence of choline, when Pcs is active (Wessel et al., 2006; Klüsener et al., 2009). When two alternative PC synthesis pathways are present in eukaryotes, PC production via PE-methylation is repressed in the presence of choline used by the CDP-choline dependent pathway (Vance and Ridgway, 1988; Vance et al., 1997). It remains to be examined whether PmtA and Pcs pathways produce distinct PC pools with different fatty acyl chains as it is the case in eukaryotes (DeLong et al., 1999). Clearly, PC biosynthesis in Agrobacterium deserves further studies.

In contrast to S. meliloti and R. sphaeroides, which produce only minor amounts of OLs under phosphate-replete conditions, A. tumefaciens accumulates significant amounts of two different OLs namely OLS1 and OLS2 even under full nutrient supply (Geske et al., 2013; Vences-Guzmán et al., 2013). In A. tumefaciens the OLs are composed of the fatty acids C16 3OH and C19:0 cyclo as shown by mass spectrometry analyses. OLS2 is the hydroxylated form of OLS1 containing the hydroxyl group within the ornithine moiety (Geske et al., 2013; Vences-Guzmán et al., 2013; Figure 5). Agrobacterium encodes the three ols homologs olsA (atu0355), olsB (atu0344), and olsE (atu0318) on the circular chromosome (Vences-Guzmán et al., 2012). olsE and olsB mutants in the A. tumefaciens A208 strain revealed that olsB is essential for formation of both OLS1 and OLS2 whereas olsE is only required for OLS2 synthesis (Figure 6). Heterologous expression of olsE resulted in OLS2 formation providing further evidence that OlsE is the hydroxylase responsible for OLS2 formation. Thus, the first step in OL synthesis in Agrobacterium is mediated by the acyltransferase OlsB forming ornithine to the lyso-ornithine lipid (LOL). Subsequently, LOL might be acylated via the putative OlsA to form OLS1. OLS2 formation is completed by hydroxylation of the ornithine moiety by OlsE (Vences-Guzmán et al., 2013).

Under low phosphate conditions, both OLs accumulate to a total amount of 45–50% whereas the total PL content decreases in A. tumefaciens A208. A putative Pho box is located in the promoter region of olsB suggesting PhoB-induced expression under phosphate starvation (Geske et al., 2013; Vences-Guzmán et al., 2013). In the olsB mutant, lack of OLs seems to be compensated by an increase in DGTS and GL accumulation under phosphate reduced conditions (Vences-Guzmán et al., 2013).

Ornithine lipids production under phosphate starvation conditions seems to vary in different Agrobacterium strains. In contrast to A. tumefaciens A208, the total amount of OLs does not change under phosphate-limiting conditions in A. tumefaciens C58C1 but the degree of hydroxylation is ninefold increased (Geske et al., 2013; Vences-Guzmán et al., 2013). Deviations in the experimental setups such as the growth media and the precise phosphate concentrations might account for these differences.

Glycolipids contain carbohydrate residues, which are glycosidically bound to the 3-position of a sn-1,2-DAG (Shaw, 1970). Different GLs are produced in bacteria under phosphate starvation. The photosynthetic bacterium R. sphaeroides produces the unique GL glucosylgalactosyl-DAG (GGD) with α-glucose (1→4)-linked to β-galactose (Benning et al., 1995). A series of GLs found in the nitrogen-fixing symbiont Mesorhizobium loti differs from the rhodobacterial GL. M. loti produces the GLs GGD, digalactosyl-DAG (DGD), and different molecular species of triglycosyl-DAG with various combinations of galactose and glucose in the head. All of the sugars are in β-configuration and (1→6)-linked to each other. Additionally, M. loti contains two further GLs with yet unknown head groups (Devers et al., 2011).

A. tumefaciens produces under phosphate deprivation four different GLs and DGTS accounting to 35% of the total lipids (Geske et al., 2013). The GLs have been identified recently as GGD and DGD with a β-configuration and monoglucosyl-DAG (MGlcD) and glucuronosyl-DAG (GlcAD) with a α-configuration (Geske et al., 2013; Semeniuk et al., 2014). The relative amount of these lipids in A. tumefaciens C58C1 is given in Figure 6B. Similar to M. loti, GGD and DGD are synthesized in A. tumefaciens via a processive glycosyltransferase namely Pgt (Figure 6) by a successive transfer of glucosyl and/or galactosyl residues to DAG. Functional characterisation of Pgt in E. coli and Pichia pastoris and overexpression in Agrobacterium revealed a broad substrate specificity concerning the glycosyl acceptor (DAG or ceramides) and sugar residues (uridine diphospho, UDP-galactose or UDP-glucose). However, Pgt favors DAG over ceramide and UDP-galactose over UDP-glucose (Hölzl et al., 2005). The promoter region of pgt contains a predicted Pho box suggesting an induced Pgt synthesis upon phosphate limitation mediated via the PhoR-PhoB system. A pgt mutant lacks GGD and DGD but the remaining lipids accumulate wild type-like (Geske et al., 2013).

Synthesis of MGlcD and the acidic GlcAD in A. tumefaciens is catalyzed by a single promiscuous glycosyltransferase namely Agt encoded by atu2297 (Figure 6). Enzyme assays with recombinant Agt in E. coli protein extracts provided evidence that Agt uses UDP-glucose and UDP-glucuronic acid as sugar donors for MGlcD and GlcAD synthesis, repectively (Figure 6; Semeniuk et al., 2014). A. tumefaciens Agt is the first described glycosyltransferase using sugars with different chemistry. An A. tumefaciens agt mutant is deficient in MGlcD and GlcAD formation and loss of these GLs is compensated by a twofold increase in GGD and DGD. Remarkably, while DGTS and all other PLs are not influenced in the agt mutant, PC amount is strongly reduced. Deletion of both pgt and agt genes results in the loss of all GLs, which is compensated by a strong DGTS accumulation. Similar to the single agt mutant, the PC content of the double mutant is strongly reduced. One reason might be that PC is degraded to provide DAG for GGD/DGD synthesis in case of the agt mutant or for DGTS synthesis in the double mutant. It is unclear, however, why specifically PC and no other PL is turned over to supply DAG for the synthesis of phosphate-free lipids. Another reason might be that reduction of the bilayer-stabilizing PC in membranes missing the acidic GlcAD is necessary to sustain membrane structure and fluidity (Semeniuk et al., 2014). In S. meliloti, loss of the acidic glycolipid SQD is compensated by an increase of the anionic and bilayer-forming lipid PG (Weissenmayer et al., 2000).

Since loss of all GLs has no impact on growth and virulence even under phosphate-limited conditions, A. tumefaciens seems to compensate the lack of all GLs by DGTS (Geske et al., 2013; Semeniuk et al., 2014). A S. meliloti mutant deficient in all phosphate-free lipids shows decreased growth under phosphate starvation but is not influenced in nodule formation on its host alfalfa (López-Lara et al., 2005) suggesting that these lipids function as bulk membrane lipids. Whether lack of all GLs and DGTS impacts A. tumefaciens physiology and virulence remains to be seen.

The acidic GlcAD in A. tumefaciens might be the counterpart of the glycolipid SQD which is absent in Agrobacterium but widespread in photosynthetic organisms and present in a few non-photosynthetic bacteria such as some rhizobia (López-Lara et al., 2003). The role of SQD in these organisms is still unclear. It has been speculated that SQD might have a special role in photosynthesis or is required for nodule formation and nitrogen fixation. However, SQD-free mutants of the photosynthetic purple bacterium R. sphaeroides and the nitrogen fixing S. meliloti are not compromised in photosynthesis and symbiosis, respectively suggesting no general function of bacterial SQD in these processes (Benning et al., 1993; Weissenmayer et al., 2002; López-Lara et al., 2003).

DGTS-(N,N,N,-trimethyl)homoserine is a betaine-ether linked glycerolipid abundant in membranes of plants, algae, and fungi and is found in a few bacteria (Dembitsky, 1996; López-Lara et al., 2003). In Agrobacterium membranes DGTS is a major non-phosphorus lipid (~20 mol%) during phosphate starvation (Figure 6B). Similar to PC, DGTS is a zwitterionic lipid containing a quaternary amino head group (Figure 5). It has been observed that the content of PC and DGTS within a cell is reciprocal. Organisms containing major amounts of PC produce only traces of DGTS and vice versa (Geiger et al., 2010). The structural similarity and the inverse relationship between DGTS and PC concentrations led to the speculation that these two lipids are functionally interchangeable (López-Lara et al., 2003; Geiger et al., 2010; Devers et al., 2011).

DGTS-(N,N,N,-trimethyl)homoserine synthesis in R. sphae-roides and S. meliloti occurs via the BtaA/B system (Klug and Benning, 2001; López-Lara et al., 2005). BtaA is a SAM/DAG 3-amino-3-carboxypropyl transferase that converts DAG to DAG-homoserine (DGHS) using SAM as homoseryl donor. Subsequently, DGHS is threefold methylated via BtaB, a SAM:DAG-homoserine-N-methyltransferase, to DGTS. Expression of the sinorhizobial btaA and btaB genes is PhoB regulated. BtaA (atu2119) and BtaB (atu2120) homologs which have not been characterized yet are encoded in the A. tumefaciens genome suggesting a similar DGTS biosynthesis and regulation (Yuan et al., 2006; Figure 6). In A. tumefaciens, DGTS and GL accumulation under phosphate limitation also seems to be controlled not only on transcriptional level of the responsible biosynthesis genes but also via DAG substrate availability. A PlcP homolog, encoded by atu1649 in the A. tumefaciens genome (Geske et al., 2013) suggests a similar membrane remodeling mechanism as described in S. meliloti (Zavaleta-Pastor et al., 2010; Geiger et al., 2013). Interestingly, phosphate starvation results not only in the replacement of PLs by non-phosphorus lipids in A. tumefaciens but also in changes in fatty acid composition of DAG and PLs with a shift from 18:1 to 19:0 cyclo fatty acids (Geske et al., 2013). Whereas under full nutrition PLs are mainly composed of 18:1 (50–60%) fatty acids and contain low proportions of 19:0 cyclo (20 and 40% in PC) fatty acid, phosphate limitation results in a decrease in 18:1 (10%) and a strong increase in 19:0 cyclo (60%) fatty acids (Geske et al., 2013). A. tumefaciens codes for a putative cyclopropane fatty acid (CFA) synthase presumably responsible for this modification (Geske et al., 2013). Cyclopropanation of pre-existing unsaturated fatty acids is widespread in bacteria and maximal activity is observed during stationary phase. The biological role of CFA containing lipids in bacteria is not fully understood. Accumulation of CFAs in E. coli is correlated with acid tolerance and seems to be important for pathogenic bacteria–host interactions as shown for Mycobacterium tuberculosis (Chang and Cronan, 1999; Glickman et al., 2000; Zhang and Rock, 2009). A twofold increase in CFA content under phosphate starvation and acid conditions is also observed in S. meliloti. Here, two CFA synthases have been described, with Cfa1 essential for cyclopropanation of fatty acids under tested conditions. Both cfa genes are not required for symbiotic nitrogen fixation in S. meliloti (Saborido Basconcillo et al., 2009). Whether cyclopropanated lipids are required for A. tumefaciens virulence remains to be determined.

Although the typical eukaryotic membrane lipid PC is rarely found in bacteria it is a main constituent of A. tumefaciens inner and outer membranes suggesting an important role for this organism (Klüsener et al., 2009). Indeed, loss of PC causes different physiological defects. A PC-deficient mutant is impaired in growth on solid medium at elevated temperatures and is unable to grow in the presence of the anionic detergent SDS. Furthermore, it is less motile and produces larger amounts of surface-attached biomass (Klüsener et al., 2009). The motility defect is explained by reduced flagellar proteins (FlaA and FlaB) in minimal medium (Klüsener et al., 2009, 2010). The most striking phenotype of a PC-deficient mutant is its defect in tumor formation due to loss of the VirB/D4 Type 4 SS (T4SS) essential for T-DNA transfer (Wessel et al., 2006). In response to plant stimuli the two component system VirA/G controls the expression of 11 transcriptional units, among them the virB and virD operons encoding the T4SS. The homodimeric histidine kinase VirA is anchored in the inner membrane. Plant-released signals, e.g., phenolic compounds are recognized by a cytoplasmic linker domain whereas acidic pH and monosaccharides are perceived by the periplasmic domain (Nair et al., 2011). The global response to PC-deficiency in A. tumefaciens as determined by proteomics and transcriptomics shows that the VirA/G-controlled vir gene expression under virulence-induced conditions is drastically reduced thus explaining the absence of the T4SS (Klüsener et al., 2010). Only a limited set of other genes coding for membrane-related proteins were changed in the absence of PC. Expression of virG in the PC-deficient mutant was also dramatically reduced suggesting that lack of virulence gene induction is due to low virG expression. Since the loss of vir gene expression in a PC-deficient mutant cannot be complemented by expression of a plasmid-encoded wild type virG but by a constitutively active VirG, it seems that a non-functional VirA sensor kinase is responsible for the loss of virulence gene expression in the PC-lacking Agrobacterium mutant. These observations suggest that signal transduction between VirA and VirG is impaired in the absence of PC, possibly due to limitations in membrane insertion or folding of VirA (Figure 7). It remains to be seen whether the observed phenotypic defects in the PC-deficient mutant are PC-specific or a consequence of altered bulk physico-chemical properties of the membrane in the absence of PC. The structural organization of membranes is defined by the physical properties and shape of membrane lipids. Cylindrical-shaped lipids such as PG or PC are bilayer-forming lipids whereas cone-shaped lipids such as PE are considered non-bilayer forming lipids (van Meer et al., 2008). However, non-bilayer lipids can form bilayer-structures depending on solvent conditions, alkyl chain composition, and temperature.

We do not know yet whether loss of other PLs such as PE, PG, or CL in A. tumefaciens causes similar effects on physiology and VirA/G-mediated signal transduction. It has been shown that PE can act as molecular chaperone for proper folding and function of membrane proteins such as the lactose permease LacY in E. coli (Bogdanov et al., 1999; Bogdanov and Dowhan, 2012). Replacement of PE by PC during reconstitution of the ABC multidrug exporter HorA from Lactobacillus brevis into membrane vesicles altered the orientation of TM helices and abolished transport function (Gustot et al., 2010). The effect of PC depletion on membrane proteins (Klüsener et al., 2010) suggests that PC and probably other PLs play a role in membrane protein homeostasis in A. tumefaciens.

It is important to note that the requirement of PC for productive host–microbe interactions is not restricted to A. tumefaciens. PC-deficient S. meliloti mutants are unable to establish nitrogen-fixing symbiosis with their host plant alfalfa (Sohlenkamp et al., 2003). Reduced PC levels in B. japonicum, the symbiont of the soybean Glycine max cause formation of nodules with impaired nitrogen fixation activity (Minder et al., 2001). A PC-deficient mutant of the intracellular human pathogen Legionella pneumophila shows lowered cytotoxicity and adhesion to the host cell. Loss of PC affects the Dot/Icm T4SS, system which delivers virulence factors into the cytosol of infected cells and is required for intracellular growth (Conover et al., 2008). In P. syringae PC is essential for secretion of the HrpZ harpin effector protein possibly due to a non-functional T3SS (Xiong et al., 2014). B. abortus, the causative agent of brucellosis produces PC via the Pcs pathway. A pcs mutant is defective in PC formation and attenuated in virulence when assayed in the mouse model (Comerci et al., 2006). PC is not generally critical for physiology or microbe–host interactions. Loss of PC in the opportunistic pathogen P. aeruginosa did not affect physiology and virulence (Malek et al., 2012). It is important to note here that PC is only a minor (~4%) component of P. aeruginosa membranes (Geiger et al., 2013).

Although various bacteria deficient in OL biosynthesis have been characterized, the function of OLs still is largely unclear. OLs have been implicated in high-temperature tolerance in Burkholderia cepacia (Taylor et al., 1998). In Bordetella pertussis and Flavobacterium meningosepticum OLs are involved in hemagglutination and stimulation of macrophages (Kawai and Yano, 1983; Kawai and Akagawa, 1989; Kawai et al., 1999). In Rhodobacter capsulatus OL is critical for optimal yields of cytochrome c (Aygun-Sunar et al., 2006). In Gram-negative bacteria OLs are enriched in the outer membrane. It has been postulated that the zwitterionic OLs increase outer membrane stability via stabilization of the negative charges of LPS. Hydroxylation of OLs often correlates with bacterial stress response (Vences-Guzmán et al., 2012). It is speculated that the additional OH group increases hydrogen bonding between the lipid molecules as shown for the 2-hydroxylated lipid A in Salmonella typhimurium. This would decrease the membrane fluidity and make it less permeable (Gibbons et al., 2000; Vences-Guzmán et al., 2012).

In A. tumefaciens A208 grown at low temperatures (15°C) unmodified OLS1 is completely hydroxylated to OLS2 suggesting a role of this modified OL in temperature stress. However, lack of both OLs has no impact on growth even under high osmolarity or at low temperature. Interestingly, A. tumefaciens A208 mutants devoid of OLS2 induce about 1 week earlier tumors and consequently, the tumor size is increased compared to wild type induced tumors (Vences-Guzmán et al., 2013). OLs share a 3-acyl-oxyacylamide structure with lipid A of Gram-negative bacteria, which is an elicitor in plant–microbe interactions (Scheidle et al., 2005; Silipo et al., 2008; Madala et al., 2011). It has been speculated that hydroxylated OLs cause a plant defense response, which might be lowered in the absence of OLs thus explaining accelerated tumor formation (Figure 7).

The role of OLs in plant interaction cannot be generalized and it seems that OLs have different functions in different bacteria. In contrast to A. tumefaciens, the two modified OLs P1 and P2 are necessary for a successful symbiotic interaction in the nitrogen-fixing symbiont R. tropici CIAT899, which is highly tolerant to different environmental stresses (Rojas-Jiménez et al., 2005). The OL in S. meliloti is required for normal growth under phosphate-limiting conditions but not necessary for symbiotic performance (López-Lara et al., 2005).