Identification of two lysophosphatidic acid acyltransferase genes with overlapping function in Pseudomonas fluorescens

Méabh Cullinane, Christine Baysse, John P. Morrissey and Fergal O'Gara

BIOMERIT Research Centre, Microbiology Department and Biosciences Institute, National University of Ireland, Cork, Ireland

Correspondence
Fergal O'Gara
f.ogara{at}ucc.ie


   ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Phosphatidic acid (PA) is known to be a crucial phospholipid intermediate in cell membrane biosynthesis. In Escherichia coli, this molecule is produced from lysophosphatidic acid (LPA) by LPA acyltransferase (EC 2.3.1.51), encoded by plsC. E. coli possesses only one such LPA acyltransferase and a plsC mutant is non-permissive for growth at elevated temperatures. This study describes the identification and characterization of two genes from Pseudomonas fluorescens F113 that encode enzymes with LPA acyltransferase activity. One of the genes, hdtS, was previously described, whereas patB is a novel gene. In addition, a putative lyso-ornithine lipid acyltransferase was also identified. All three proteins possess conserved acyltransferase domains and are homologous to PlsC and to LPA acyltransferases identified in Neisseria meningitidis. Functional analysis determined that both HdtS and PatB are functional LPA acyltransferases, as both complemented an E. coli plsC mutant. Mutants lacking each of the putative acyltransferases were constructed and analysed. Growth defects were observed for hdtS and patB single mutants, and a double hdtSpatB mutant could not be constructed. To determine precise roles in phospholipid synthesis, fatty acid methyl ester analysis was carried out. The hdtS mutant displayed a profile consistent with a defect in LPA acyltransferase activity, whereas no such phenotype was observed in the patB mutant, indicating that hdtS encodes the primary LPA acyltransferase in the cell. The presence of at least two genes specifying LPA acyltransferase activity may have implications for the function and survival of P. fluorescens in diverse environments.


Abbreviations: acyl-ACP, acyl-acyl carrier protein; acyl-CoA, acyl-coenzyme A; FAME, fatty acid methyl ester; LPA, lysophosphatidic acid

The GenBank/EMBL/DDBJ accession numbers for the genes reported in this paper are: pasA, DQ088968; olsA, AY876048; patB, AY876049; hdtS, AF286536.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacteria in the genus Pseudomonas are ubiquitous in nature and are commonly found in soil and water habitats. The wide distribution of Pseudomonas has been attributed to its metabolic diversity, as well as its capacity to sense and respond to the diverse environments it may encounter. In general, pseudomonads are considered beneficial microbes, playing important ecological roles in plant health and nutrient cycling. The genus does contain some significant plant pathogens, however, and the opportunistic human pathogen, Pseudomonas aeruginosa, is a major problem in immunocompromised individuals. Some species are also associated with food spoilage, particularly at low temperatures. In contrast, many pseudomonads are benignly associated with plants, and Pseudomonas fluorescens, in particular, can play a major role in promoting plant health. The antagonism of some strains of P. fluorescens to phytopathogenic soil fungi has fuelled interest in the potential of these bacteria to function as natural biocontrol agents, representing an alternative to conventional chemical control methods. Two main factors influence the efficacy of strains in this regard: the ability to colonize the plant rhizosphere to a high level, and the capacity to produce appropriate levels of an anti-fungal metabolite at the correct time (Morrissey et al., 2002; Walsh et al., 2001). Considerable research effort is being devoted to understanding the traits that confer rhizosphere success and environmental adaptability on P. fluorescens (Lugtenberg et al., 2002). In this context, attention has focused on cell membranes and their composition and properties, and it is known that various types of environmental stress, such as temperature and osmotic stress, can cause alterations in the physical properties of membrane lipids (Los & Murata, 2004). Such conditions are frequently encountered by bacteria surviving in changing niches such as the rhizosphere, and so rhizosphere bacteria must constantly be capable of adapting to such stresses in order to survive. The structure and integrity of bacterial membranes must play a pivotal role in the survival and propagation of rhizosphere bacteria. One particular feature of bacterial membranes that has been shown to play a part in rhizosphere function is the phospholipid component. For the plant-associated bacterium Sinorhizobium meliloti, an inability to synthesize phosphatidylcholine (PC) renders the strain unable to form nodules on plant hosts (de Rudder et al., 2000; Sohlenkamp et al., 2003). Similarly, Bradyrhizobium japonicum mutants with reduced PC biosynthesis can only form root nodules with reduced rates of nitrogen fixation (Minder et al., 2001). These examples set a precedent for a biological role for phospholipids in microbial ecology, and raise questions as to the role of phospholipids in other rhizosphere bacteria such as P. fluorescens.

Phospholipid biosynthesis has been most intensively studied in Escherichia coli, which has been a model for studying the genetic and biochemical aspects of lipid metabolism for over 40 years. E. coli has a relatively simple phospholipid composition, consisting mainly of phosphatidylethanolamine (PE), phosphatidylglycerol (PG) and cardiolipin (CL) (DiRusso et al., 1999). In this bacterium, the outer membrane consists of approximately 25 % dry weight phospholipid, while the inner membrane is composed of approximately 40 % (DiRusso et al., 1999). Phospholipid biosynthesis occurs primarily by de novo synthesis on the inner membrane of the cell envelope using glycerol 3-phosphate (G3P) and fatty acids as the main substrates (Cronan & Rock, 1996). Sequentially, two acyltransferase reactions transfer acyl groups from acyl-acyl carrier protein (acyl-ACP) or acyl-coenzyme A (acyl-CoA) to G3P. The first acyltransferase reaction results in the transfer of an acyl group to the sn-1 position of G3P to produce lysophosphatidic acid (LPA), and in E. coli this reaction is catalysed by the enzyme encoded by the plsB gene (Larson et al., 1980; Lightner et al., 1983; Rock & Cronan, 1981). The second step transfers an acyl group to the sn-2 position of LPA to produce phosphatidic acid (PA) and is catalysed by the enzyme LPA acyltransferase, encoded by plsC in E. coli (Coleman, 1990). PA is then converted to PE, PG and CL. Variation in the membrane phospholipid composition comes from both the diversity of phospholipid head groups, and from differences in the chain length and saturation of the acyl groups in the sn-1 and sn-2 positions. When grown at 37 °C, the phospholipids of E. coli contain mainly saturated fatty acids at the sn-1 position (C16 : 0) and unsaturated fatty acid chains (C16 : 1 or C18 : 1) at the sn-2 position (Cronan & Rock, 1996). It has been proposed that the different substrate specificities of G3P acyltransferase (PlsB) and LPA acyltransferase (PlsC) partly account for the acyl chain distribution on the glycerol backbone of membrane phospholipids (Cronan & Rock, 1987). Although in E. coli, a single essential gene, plsC, encodes LPA acyltransferase, in Neisseria meningitidis, two LPA acyltransferases have been identified and characterized (Shih et al., 1999; Swartley et al., 1995). Designated NlaA and NlaB, both proteins share homology to PlsC, possess specific LPA acyltransferase activity in vitro, and can complement a temperature-sensitive E. coli plsC mutant (Shih et al., 1999; Swartley et al., 1995). It has been proposed that N. meningitidis possesses more than one LPA acyltransferase to facilitate a greater diversity of membrane phospholipids (Shih et al., 1999). Based on these findings and the importance of phospholipid structure and composition to environmental adaptation, we have investigated the possible occurrence of multiple LPA acyltransferases in P. fluorescens F113. This strain of P. fluorescens has potential as a biocontrol agent and is capable of inhibiting the growth of Pythium ultimum in vitro and in natural ecosystems (Fenton et al., 1992; Shanahan et al., 1992). In this study, three putative LPA acyltransferases, termed hdtS, olsA and patB, were identified from P. fluorescens F113 and characterized. These genes were localized to different regions in the P. fluorescens F113 chromosome and all shared homology with E. coli PlsC. Analysis of these genes showed they possessed different but related functions. We discuss the implications of these data for the environmental adaptability of P. fluorescens.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacterial strains, plasmids and growth conditions.
Strains and plasmids used in this study are listed in Table 1. All cultures of P. fluorescens F113 and associated mutants were routinely grown on sucrose asparagine (Scher & Baker, 1982), supplemented with 100 µM FeCl3, or LB media (Sambrook et al., 1989) at 28 °C. E. coli strains were routinely grown in LB media at 37 °C with the exception of the E. coli plsC mutant JC201, which was cultivated at 30 °C. Where appropriate, antibiotics were added to growth media at the following concentrations: kanamycin 50 µg ml–1, chloramphenicol 200 µg ml–1 and gentamicin 50 µg ml–1 for P. fluorescens; and kanamycin 50 µg ml–1, chloramphenicol 30 µg ml–1, gentamicin 25 µg ml–1 and tetracycline 12·5 µg ml–1 for E. coli.


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Table 1. Bacterial strains and plasmids

 
DNA and RNA manipulations.
Primers used are listed in Table 2. Restriction digests, ligations, transformations and agarose gel electrophoresis were performed as described by Sambrook et al. (1989). All restriction enzymes were obtained from Roche Pharmaceuticals. Small-scale plasmid DNA isolation was performed using the Qiagen Plasmid Mini-kit (Qiagen) according to the manufacturer's instructions. Prior to cloning, PCR products were purified either from agarose gels using the Qiagen Gel Extraction kit or directly from solution using the Qiagen PCR purification kit and cloned into the pCR2.1TOPO plasmid using the TA cloning kit (Invitrogen) according to the manufacturer's specifications. Plasmids were mobilized into Pseudomonas strains either by biparental mating using E. coli GJ23 (Van Haute et al., 1983) or triparental mating using the helper plasmid pRK2013 (Figurski & Helinski, 1979). Total RNA was isolated from 7x109 bacterial cells grown to an OD600 of 0·8 using the RNeasy Total RNA isolation kit (Qiagen) according to the manufacturer's instructions. The RNA was treated with RQ1 RNase-free DNase I (Promega) and quantified via spectrophotometry, and its integrity was checked by TBE agarose gel electrophoresis. PCR analysis using primers specific to multiple genes ensured that no DNA was left in the sample post-DNase I treatment. For semi-quantitative analysis of the fliC gene, cDNA was synthesized by the RT SuperScript II (Invitrogen) with 5 µg RNA and 400 U reverse transcriptase for 1 h at 37 °C. After RNase treatment for 1 h at 37 °C (RNase I, 0·2 U µl–1), cDNA was purified using the DNA purification kit from Qiagen. PCR reactions were carried out using 100 ng cDNA template for 22–26 cycles for the FliF and FliR primers and 30–35 cycles for the MC10F and MC10R primer pair. The pasA gene (GenBank accession no. DQ088968), which is constitutively transcribed in P. fluorescens F113, was used as an internal control for the RT-PCR experiments. Subsequent analysis was carried out using the Phoretix 1D Advanced software, version 5.0 (Nonlinear Dynamics).


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Table 2. Primers used in this study

Underlined sequences denote artificial restriction enzyme sites; italicized sequences denote an artificial ribosome-binding site.

 
Determination of nucleotide sequence and sequence analysis.
Nucleotide sequencing was performed by Lark Technologies. The sequence data were assembled using the DNASTAR software package and analysed using the university of wisconsin genetic computer group (gcg) program FASTA (Pearson & Lipman, 1988) and BLAST (Altschul et al., 1990) at the National Center for Biotechnology Information (NCBI). Multiple sequence alignments were performed using the CLUSTALW program (Chenna et al., 2003).

Cloning of P. fluorescens F113 olsA and patB.
Using online databases, the region surrounding the olsA gene in several Pseudomonas species was analysed and, based on sequence similarity, primers MC12F and MC12R were designed. Following PCR analysis on P. fluorescens F113 genomic DNA, the resulting 2·7 kb PCR product was gel-purified and introduced into pCR2.1TOPO using the TA cloning kit. The PCR product was subsequently sequenced using primers M13F, M13R, MC12F, MC12R, NlaF, NlaR, Nla2F and Nla2R to obtain the full sequence of olsA and the upstream gene. The GenBank accession number for olsA is AY876048. Cloning of the patB gene was carried out as follows: based on sequence similarity, primers MC26F and MC26R were designed to amplify the region containing the putative patB gene. The resulting 740 bp PCR product was gel purified and introduced into pCR2.1TOPO using the TA cloning kit to form plasmid pCR2.1patB. The patB ORF was subsequently sequenced using the M13F and M13R primer pair. The GenBank accession number for patB is AY876049.

Plasmid constructions.
To construct plasmid pBBR1MCShdtS, an 815 bp region surrounding the hdtS ORF was amplified using primers MC15F and MC7R. Primer MC15F contains an artificial ribosome-binding site at the 5' end while MC7R contains an Asp718 site. The resultant PCR product was cloned into the pCR2.1TOPO vector to form pCR2.1hdtS, retrieved by digestion with EcoRV and Asp718, and cloned into EcoRV/Asp718-cut pBBR1MCS. Plasmid pBBR1MCSolsA was obtained by subcloning the olsA gene into the multiple cloning site of pBBR1MCS as follows: the olsA ORF was PCR-amplified from P. fluorescens F113 DNA using primers MCER and MCKF, which contain EcoRV and Asp718 restriction enzyme sites, respectively. The 1·05 kb PCR product obtained was cloned into the pCR2.1TOPO vector to form pCR2.1olsA, retrieved by digestion with EcoRV and Asp718, and cloned into EcoRV/Asp718-cut pBBR1MCS. To construct plasmid pBBR1MCSpatB, a 733 bp region surrounding the patB ORF was amplified using primers MC32F and MC32R, which contain EcoRV and Asp718 restriction enzyme sites, respectively. Primer MC32F also contains an artificial ribosome-binding site at the 5' end. The resultant PCR product was digested with the appropriate enzyme and cloned into EcoRV/Asp718-cut pBBR1MCS. pBBR1MCSplsC was obtained by subcloning the plsC gene from E. coli using primers MC22F and MC22R. The resultant 844 bp PCR product was cloned into the pCR2.1TOPO vector, retrieved by digestion with EcoRV and Asp718, and cloned into EcoRV/Asp718-cut pBBR1MCS.

P. fluorescens F113 mutant constructions.
The P. fluorescens F113 hdtS mutant was obtained as follows: a 397 bp fragment of the hdtS ORF was PCR-amplified using primers MC9F and MC9R, which contain XbaI and PstI restriction enzyme sites, respectively. The PCR product was cloned into XbaI/PstI-cut pK18mob to form pK18mobhdtS and the vector introduced into P. fluorescens F113 by triparental mating, selecting for a single crossover event. The P. fluorescens F113 patB mutant was obtained as follows: a 402 bp fragment of the patB ORF was PCR-amplified using primers MC30F and MC30R. The PCR product obtained was cloned into the pCR2.1TOPO vector and excised with EcoRI. The resultant fragment was cloned into alkaline phosphatase (Promega)-treated EcoRI-cut pK18mob to form pK18mobpatB and the vector introduced into P. fluorescens F113 by triparental mating, selecting for a single crossover event. All single mutants were complemented via introduction of the appropriate gene in trans.

Fatty acid methyl ester (FAME) analysis.
Alterations in the fatty acid compositions of the membrane phospholipids from P. fluorescens F113 and associated mutants were measured via FAME analysis, carried out by Microcheck Inc.

Motility analysis.
Swimming ability was analysed on tryptone swim plates [1 % (w/v) tryptone, 0·5 % (w/v) NaCl, 0·3 % (w/v) agar]. These plates were inoculated using a sterile toothpick with bacteria grown overnight on LB agar. Motility was assessed qualitatively by examining the circular turbid zone formed by bacterial cells migrating from the inoculation point at 24 h post-inoculation.

E. coli plsC complementation experiments.
Plasmids pBBR1MCShdtS, pBBR1MCSolsA and pBBR1MCSpatB were transformed into the E. coli plsC mutant strain JC201. To test complementation on plates, the E. coli parent strain (JC200), JC201 and JC201 harbouring each of the three P. fluorescens genes were grown on LB plates overnight at 42 °C. For liquid complementation assays, the strains were grown in LB media with shaking for 1 h at 30 °C before shifting to the non-permissive temperature of 42 °C, at which the growth rate was measured spectrophotometrically for a further 8 h.


   RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Identification of two putative LPA acyltransferases in P. fluorescens
In silico analysis of draft genomes from P. fluorescens Pf0-1 (www.jgi.doe.gov), P. fluorescens SBW25 (www.sanger.ac.uk) and P. fluorescens Pf-5 (www.tigr.org) showed the presence of three putative LPA acyltransferases with homology to the LPA acyltransferase (EC 2.3.1.51) from E. coli encoded by plsC. One of these genes, hdtS, was previously identified as a putative N-acyl homoserine lactone synthase in P. fluorescens F113 (Laue et al., 2000), but the other two genes are novel and previously uncharacterized in P. fluorescens. For reasons that are outlined below, we designated one of these genes olsA and the second patB (Pseudomonas acyltransferase). Based on conserved genomic sequences, degenerate PCR primers were designed that allowed the amplification of olsA and patB from P. fluorescens F113 and the genes were cloned and sequenced. Protein sequence alignment shows conservation of motifs between the P. fluorescens proteins and characterized LPA acyltransferases from E. coli (PlsC) and N. meningitidis (NlaB and NlaA). Interestingly, however, these motifs are also present in the Sinorhizobium meliloti lyso-ornithine lipid acyltransferase, olsA (Weissenmayer et al., 2002) (Fig. 1). The key motifs, designated IPB002123A and IPB002123B (DBGET, GenomeNet), are highly conserved in prokaryotic and eukaryotic LPA acyltransferases and have been suggested to constitute an acyl-CoA/acyl-ACP binding site (Shih et al., 1999; West et al., 1997). All three Pseudomonas proteins have homology to E. coli PlsC, with HdtS showing even higher levels of homology to the PlsC orthologue in N. meningitidis, NlaB. At the amino acid level, HdtS shows 20 % amino acid identity to PlsC and 34 % identity to NlaB. OlsA shows 11 % identity to PlsC, 20 % identity to the putative LPA acyltransferase NlaA and 25 % identity to the S. meliloti lyso-ornithine lipid acyltransferase OlsA. Analysis of the genomic region flanking P. fluorescens olsA revealed the presence of a gene immediately upstream with strong homology to olsB from S. meliloti, a gene that encodes another enzyme in the ornithine lipid biosynthetic pathway (Gao et al., 2004). Furthermore, RT-PCR analysis indicated that this upstream gene and olsA are co-transcribed in P. fluorescens F113 (data not shown). In combination, these data provide strong evidence that this gene in P. fluorescens is a lyso-ornithine lipid acyltransferases and justify its designation olsA. The third protein, PatB, shows 40 % identity at the amino acid level to PlsC and low levels of homology to other putative LPA acyltransferase outside the conserved motifs. This in silico analysis suggested that HdtS and PatB Pseudomonas enzymes are putative LPA acyltransferases, but that OlsA is not: a hypothesis that we tested experimentally.



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Fig. 1. Alignment of membrane lipid acyltransferases. Proteins included are LPA acyltransferases from E. coli (EcoPlsC) and N. meningitidis (NmNlaA and NmNlaB), a lyso-ornithine lipid acyltransferase from S. meliloti (SmOlsA) and HdtS (PfluHdtS), OlsA (PfluOlsA) and PatB (PfluPatB) from P. fluorescens F113. All proteins possess two consensus domains (boxed) identified in phospholipid and glycerol acyltransferases (blocks IPB002123A and IPB002123B). Accession numbers for P. fluorescens F113 genes are as follows: hdtS, AF286536; olsA, AY876048; and patB, AY876049.

 
Functional complementation of an E. coli plsC mutant by P. fluorescens LPA acyltransferases
To test the activity of these putative LPA acyltransferases from P. fluorescens, each gene was introduced in trans into E. coli plsC mutant JC201. This strain carries a Ts allele of plsC and is unable to grow at 42 °C (Coleman, 1990). Previously, it was demonstrated that both nlaA and nlaB from N. meningitidis could complement this mutation (Shih et al., 1999; Swartley et al., 1995). To determine whether this is also true of hdtS, patB or olsA, E. coli strain JC201 was transformed with a plasmid carrying the appropriate gene expressed from a constitutive promoter, and transconjugants were grown at the restrictive temperature (42 °C). On plate assays, it was found that hdtS fully restored the wild-type phenotype, patB appeared to give partial complementation, and olsA did not complement the mutation (Fig. 2a). This experiment was repeated in broth cultures (Fig. 2b). E. coli strains carrying the Pseudomonas genes were pre-grown at 30 °C and then transferred to 42 °C. The same result was observed: JC201[olsA] showed no complementation, JC201[patB] partially complemented; but the complemented strain failed to grow at the same rate or to the same final OD600 as wild-type strains, and the growth of JC201[hdtS] was indistinguishable from JC200. To ensure that the different degrees of complementation were not a consequence of weak expression, RT-PCR analysis was performed and it was shown that all Pseudomonas genes were expressed well in the plasmid-carrying E. coli strains (data not shown).



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Fig. 2. Complementation analysis of E. coli plsC mutant with hdtS, olsA or patB in trans. (a) Complementation analysis of E. coli plsC mutant with hdtS, olsA or patB in trans via plate assays. Expression of the hdtS gene in this mutant restores growth (plsC[hdtS]). Expression of the patB gene partially restores growth (plsC[patB]), while the olsA gene does not complement for this mutation (plsC[olsA]). E. coli JC200, the parent strain of JC201, was used as a control (WT). (b) Expression of hdtS, olsA or patB in trans in an E. coli plsC mutant. Strains were grown at 30 °C for 1 h and then shifted to 42 °C. Growth was measured spectrophotometrically at 1 h intervals for a further 8 h. The E. coli plsC mutant (JC201) cannot grow at 42 °C ({bullet}). Expression of the hdtS gene in this mutant restores growth by providing acyltransferase activity in trans ({square}). Expression of the patB gene can partially complement for growth ({blacktriangleup}), while expression of olsA in trans does not restore growth to a plsC mutant ({circ}). E. coli JC200, the parent strain of JC201, was used as a control ({blacksquare}).

 
Comparison of membrane fatty acid profiles in wild-type and mutant strains of P. fluorescens
To assess the potential roles of HdtS and PatB in membrane phospholipid biosynthesis, strains with disruptions in each of these genes were constructed and total fatty acid profiles were examined (Table 3). The most striking result was that mutation of hdtS led to a shift in the ratio of C16 : C18 fatty acids. The wild-type strain contained approximately 66 % C16 fatty acids and 14 % C18 fatty acids, whereas the hdtS mutant possessed approximately 37 % C16 fatty acids and 48 % C18 fatty acids. In particular, there was a shift to the unsaturated fatty acid C18 : 1. This C16 : C18 shift was reversed by introduction of either the Pseudomonas hdtS gene or the E. coli plsC gene in trans. Expression of patB in the hdtS mutant did not restore the ratio to that of the wild-type. Unlike deletion of hdtS, deletion of patB did not cause any significant alteration in the relative proportion of fatty acids. Multiple efforts and strategies to construct a hdtSpatB mutant strain were unsuccessful, leading to the conclusion that this is likely to constitute a lethal combination. In summary, these data support the idea that HdtS is a functional LPA acyltransferase, with a substrate specificity for shorter-chain acyl groups. In the absence of HdtS, another enzyme, probably PatB, with a substrate specificity for longer-chain fatty acids, can carry out the LPA acyltransferase reaction.


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Table 3. Fatty acid profile of bacterial cells as determined by FAME analysis

The numbers represent the percentage of each fatty acid in a total cell extract of P. fluorescens strains. Values are representative of multiple experiments. Major changes in fatty acid profiles are represented in bold type. Deletion of hdtS significantly alters the membrane fatty acid composition, whereas deletion of patB has a minimal impact on the fatty acid profile of the bacterium. Introduction of either hdtS or plsC, but not patB, in trans into the hdtS mutant restores the fatty acid profile to wild-type.

 
LPA acyltransferase mutants display varied growth and motility phenotypes
LPA acyltransferase is an essential activity; therefore, the mutant strains were assessed to determine whether any phenotypes were evident. Growth of the mutant strains was examined over a range of growth media and at different temperatures (Fig. 3). The hdtS mutant showed a significantly impaired growth rate under all conditions tested (Fig. 3a). The wild-type phenotype was fully restored by expression of either hdtS (data not shown) or E. coli plsC (Fig. 3a) in trans, confirming the functional equivalence of the proteins. At 30 °C, the patB mutant displayed a very minor reduction in growth rate, but this was greatly exacerbated when the strain was grown at 38 °C (Fig. 3b). Growth analysis at 12 °C did not show any exacerbated growth defect for the hdtS mutant, and patB mutant grew as wild-type (data not shown).



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Fig. 3. Growth kinetics of hdtS and patB mutants of P. fluorescens F113. (a) Growth kinetics at 30 °C. Strains were grown in LB media and the OD600 measured spectrophotometrically at 2 h intervals. Whereas F113patB ({blacktriangleup}) shows only a marginal reduction in growth compared to F113WT ({blacksquare}), F113hdtS ({circ}) shows a reduced growth rate. Expressing the plsC gene in trans ({bullet}) in F113hdtS restores the growth rate to wild-type levels. (b) Growth of P. fluorescens F113 and F113patB in LB media at 38 °C. Growth rate was measured spectrophotometrically at 2 h intervals. The patB mutant ({blacktriangleup}) shows a reduction in growth compared to F113WT ({blacksquare}) at this temperature. All growth curves were carried out in triplicate with similar results.

 
A role for HdtS in motility was also determined (Fig. 4a). Analysis of the hdtS mutant on 0·3 % agar swim plates was carried out: the size of the halo produced by a bacterial sample on these plates is a measure of motility. The absence of a halo in the hdtS mutant established that the mutant is defective in flagellar-mediated motility. This could be a consequence of lack of expression of the flagella biosynthetic genes or of the inability of the flagella to assemble correctly. RT-PCR analysis showed that the major flagellin subunit, fliC, was still transcribed in the hdtS mutant, suggesting that the phenotype may be due to improper insertion of the flagella into the bacterial membrane (Fig. 4b). Examination of the strains under the light microscope indicated that motility is not completely abolished in the hdtS mutant, although a qualitative reduction was apparent. Supplying either hdtS (Fig. 4a) or plsC (data not shown) in trans complemented this phenotype. Mutating the patB gene had no effect on bacterial motility (data not shown).



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Fig. 4. Motility analysis of P. fluorescens F113 and hdtS mutant. (a) Motility was assessed on 0·3 % (w/v) agar plates and is directly proportional to the halo size produced by each strain. The hdtS mutant (F113hdtS) shows a reduced swimming phenotype compared to wild-type (F113WT). This phenotype is restored by expressing the hdtS gene in trans (F113hdtS[hdtS]). (b) RT-PCR analysis of fliC transcription in F113WT and F113hdtS showed no significant difference in expression of this gene between these two strains. RT-PCR analysis was carried out as described in Methods and the pasA gene, known to be constitutively expressed in P. fluorescens F113, was used as an internal standard. Band intensity was measured using the Phoretix 1D Advanced software version 5.0 (Nonlinear Dynamics).

 

   DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
This study describes the characterization of two putative LPA acyltransferases in P. fluorescens as well as identification of a putative lyso-ornithine lipid acyltransferase. Although there is only one dedicated LPA acyltransferase in E. coli, multiple LPA acyltransferases have been identified in other bacteria. For example, N. meningitidis possesses two functional LPA acyltransferase genes, termed nlaA and nlaB (Philipp et al., 1996; Shih et al., 1999). Multiple LPA acyltransferases have also been identified among some eukaryotes. For instance, two such proteins have been found in the crop plant Limnanthes douglasii, and in that case there is tissue-specific expression, with one protein restricted to developing seeds (Brown et al., 1995).

The evidence establishing HdtS as an LPA acyltransferase, functionally analogous to PlsC in E. coli, is substantial. The two most compelling pieces of data are the following: first, hdtS and plsC genes can cross-complement and functionally replace each other; and second, a hdtS mutant shows alterations in cellular fatty acid composition consistent with an alteration in LPA acyltransferase activity. The hdtS gene was previously postulated to encode an N-acyl homoserine lactone (HSL) synthase (Laue et al., 2000), but our data suggest that this is not its physiological function in P. fluorescens. In addition to the experiments mentioned above, we analysed HSL production in the hdtS mutant and found that this mutant makes normal levels of C6-HSL (data not shown). We also failed to detect HSLs in E. coli expressing the construct used in this study. Based on the combined evidence, it seems likely that the previous findings reflected non-specific acyltransferase activity in E. coli under the particular growth conditions used in that study. A similar conclusion has been reached with studies of the homologous gene, PA0005, in P. aeruginosa (C. Baysse, unpublished data). As LPA acyltransferase activity is an essential activity, and as the hdtS gene is not essential, it follows that another protein(s) must be capable of carrying out the function, analogous to NlaA and NlaB in N. meningitidis. In silico analysis of complete genome sequences identified OlsA and PatB as the only likely candidates for this role. Bioinformatic analysis suggests that olsA encodes a lyso-ornithine lipid acyltransferase. Although absolute confirmation that this is the true function of this enzyme requires further experimental evidence, the findings that an olsA mutant showed no changes in membrane fatty acid profiles (data not shown) and that the olsA gene cannot complement an E. coli plsC mutant are consistent with this designation. In contrast, the patB gene partially complements an E. coli plsC mutation, and our inability to make the hdtSpatB mutant suggests that in the absence of HdtS, PatB carries out the LPA acyltransferase reaction. Thus, PatB is an LPA acyltransferase, although this may not be its primary function or activity.

An obvious question is why does P. fluorescens have two enzymes to carry out a reaction that is catalysed by one in E. coli? Following several previous studies, it has been suggested that the rationale for multiple LPA acyltransferases is to permit the synthesis of a greater, or perhaps regulated, range of membrane phospholipids (Brown et al., 2002; Shih et al., 1999). In the case of N. meningitidis, the data suggest that NlaA may preferentially specify shorter acyl side chains at the sn-2 position, whereas NlaB would specify longer chain acyl groups. In fact, data from our study indicate that the opposite situation prevails in Pseudomonas: HdtS appears to favour shorter (C16) side chains over longer (C18) side chains. The precise role of patB remains to be determined, as analysis of fatty acid profiles in mutant strains did not establish its function. One explanation might be that these genes are differentially expressed under certain environmental conditions and our single-condition assay failed to represent this. Although we did establish by RT-PCR that all three genes are expressed in P. fluorescens (data not shown), comprehensive expression studies will be required to address this issue.

Although all three putative acyltransferases possess conserved domains, OlsA displays some divergence from HdtS and PatB. Both HdtS and PatB contain the peptide sequences NHQS and PEGTR, whereas OlsA contains the peptide sequences NHVS and PEGTT at these two conserved sites (Fig. 1). Similar variation was previously noted for NlaA and NlaB in N. meningitidis: NlaA containing the peptide sequences NHVS and PEART at the two conserved sites, and NlaB containing the sequences KHQS and PEGTR. Although it remains to be determined whether these particular sequence changes are responsible for altered substrate specificity, it has been found that slight changes in the amino acid sequence of LPA acyltransferase proteins are sufficient to alter activity or substrate specificity (Nagiec et al., 1993; West et al., 1997). This may explain some apparent anomalies, such as why N. meningitidis NlaA can complement the E. coli plsC mutant but P. fluorescens OlsA cannot.

The potential significance of phospholipid biosynthesis in plant-associated bacteria has previously been highlighted. In particular, phosphatidylcholine, which is a significant component of Pseudomonas membranes, has been proposed to play an important role in plant–microbe associations (Lopez-Lara et al., 2003). Of a range of in vitro phenotypes assessed, impaired growth in the hdtS and patB mutants, and reduced motility in the hdtS mutant were ecologically relevant phenotypes observed. Furthermore, it is also important to recognize that bacteria in natural ecosystems will be exposed a wider range of stresses and environmental changes than was assessed in this study. The demonstration that P. fluorescens possesses at least two LPA acyltransferases is the first indication that this bacterium may have regulatory mechanisms to vary one aspect of membrane phospholipid structure. It will be very interesting to determine whether other mechanisms also exist, and what role they play in environmental adaptation. Indeed, the finding that P. fluorescens also possesses an enzyme specifying the synthesis of ornithine-containing lipids suggests an additional mode of varying membrane structure. With specific regard to this study, some intriguing questions remain to be answered. Perhaps foremost is the relationship between the LPA acyltransferases, their genetic regulation, and their substrate specificity. Exploring gene regulation is complicated by the observation that in P. fluorescens, as in N. meningitidis, these genes are located in operons with genes that appear to be functionally unlinked (Shih et al., 1999; Swartley & Stephens, 1995). In addition, it is curious that, although PatB bears highest homology to E. coli PlsC, HdtS appears to be the functional equivalent of this protein in P. fluorescens F113. With the growing realization of the potential significance of membrane structure for bacterial cell function, answering these questions will provide an opportunity to explore this issue in the context of an ecologically relevant and diverse bacterial species, P. fluorescens. The potential for regulated alteration of phospholipids and other membrane components is suggestive of a novel bacterial strategy for environmental adaptation and interaction with plants. Future studies will address the biology of this as well as potential applications for environmental biotechnology.


   ACKNOWLEDGEMENTS
 
The authors would like to thank Pat Higgins for technical assistance and Dr Max Dow and Dr Louise Mark for helpful discussions. Our thanks to Professor David Stephens for supplying us with E. coli JC200 and E. coli JC201. We also acknowledge the very helpful comments of referees in the review stage of this manuscript. This work was supported in part by grants awarded by the European Commission (QLK3-CT-2000-31759, QLTK3-CT-2001-00101 to F. O. G. and J. P. M.), Enterprise Ireland (SC/02/517 to J. P. M.; SC/02/520 to F. O. G.; postgraduate scholarship to M. C.), the Higher Education Authority of Ireland (PRTLI programmes to F. O. G. and J. P. M.), Science Foundation Ireland (02/IN.1/B1261 and 04/BR/B0597 to F. O. G.), Health Research Board (RP/2004/145 to F. O. G.), and the Irish Research Council for Science, Engineering and Technology (Embark post-doctoral fellowship to C. B.).


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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Received 9 February 2005; revised 1 June 2005; accepted 6 June 2005.



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